Methods
in
Molecular Biology™
Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK
For further volumes: http://www.springer.com/series/7651
Chloroplast Research in Arabidopsis Methods and Protocols, Volume II Edited by
R. Paul Jarvis Department of Biology, University of Leicester, Leicester, UK
Editor R. Paul Jarvis, PhD Department of Biology University of Leicester University Road Leicester, LE1 7RH United Kingdom
[email protected]
ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-236-6 e-ISBN 978-1-61779-237-3 DOI 10.1007/978-1-61779-237-3 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011932678 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)
Preface Chloroplasts are green plastids found in land plants, algae, and some protists. They are the unique site for the reactions of photosynthesis in such cells, and thus chloroplasts are responsible for much of the world’s primary productivity. As photosynthesis is the only significant mechanism of energy-input into living cells, these organelles are essential for the survival of plants and animals alike. Consequently, agriculture is wholly dependent upon the photosynthesis that takes place in chloroplasts. Moreover, many other important cellular activities occur uniquely inside chloroplasts or in other non-photosynthetic types of plastid. These activities include the production of starch, amino acids, fatty acids, lipids, terpenoids, purine and pyrimidine bases, and colourful pigments in fruits, flowers, and leaves, as well as key aspects of nitrogen and sulphur metabolism. Many products of these biosynthetic processes are vital components of mammalian diets or offer opportunities for industrial exploitation. Advances in our understanding of plastid biogenesis will facilitate the manipulation and exploitation of these processes and aid improvements in the quantity or quality of the various products. Over the years, chloroplast biology has been studied in a variety of different organisms, based on technical considerations. Such work has undoubtedly led to major advances in the field, but has had the significant disadvantage that findings made using different experimental systems or species are not always directly cross-comparable. The relatively recent adoption of Arabidopsis thaliana as the model organism of choice for plant science research, across the globe, has led to its emergence as a pre-eminent system for research on chloroplasts and other types of plastid. The availability of genomic sequence resources and extensive germplasm collections for Arabidopsis, as well as its amenability to molecular genetic analysis, have all contributed to this change. This book (together with its partner, Volume I) aims to bring together in a single location some of the most important, modern techniques and approaches for chloroplast research, with the unifying theme of Arabidopsis as the model system. Within the confines of this remit, we have produced a book that is relatively broad in its scope, and which many Arabidopsis researchers and biotechnologists with a general interest in chloroplasts, plastids, or related processes might use as an aid to their work. In essence, it is a book for Arabidopsis integrative biologists with a general focus on chloroplast and plastid research. In spite of the central position afforded to Arabidopsis, many of the presented methods can be applied to other experimental organisms with minimal modification. Leicester, UK
R. Paul Jarvis
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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Part I Multiprotein Complexes and Protein-Protein Interactions 1 One- and Two-Dimensional Blue Native-PAGE and Immunodetection of Low-Abundance Chloroplast Membrane Protein Complexes . . . . . . . . . . . . . . . Shingo Kikuchi, Jocelyn Bédard, and Masato Nakai 2 Analysis of Thylakoid Protein Complexes by Two-Dimensional Electrophoretic Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sari Sirpiö, Marjaana Suorsa, and Eva-Mari Aro 3 Preparation of Multiprotein Complexes from Arabidopsis Chloroplasts Using Tandem Affinity Purification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Charles Andrès, Birgit Agne, and Felix Kessler 4 Studying Interactions Between Chloroplast Proteins in Intact Plant Cells Using Bimolecular Fluorescence Complementation and Förster Resonance Energy Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jodi Maple and Simon G. Møller 5 Studying Chloroplast Protein Interactions In Vitro: An Overview of the Available Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Joanna Tripp and Enrico Schleiff
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Part II Omics and Large-Scale Analyses 6 Proteome Databases and Other Online Resources for Chloroplast Research in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Diogo Ribeiro Demartini, Célia Regina Carlini, and Jay J. Thelen 7 Use of Transcriptomics to Analyze Chloroplast Processes in Arabidopsis . . . . . . . . . 117 Tatjana Kleine and Dario Leister 8 Use of Non-aqueous Fractionation and Metabolomics to Study Chloroplast Function in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Peter Geigenberger, Axel Tiessen, and Jörg Meurer 9 Chloroplast Phenomics: Systematic Phenotypic Screening of Chloroplast Protein Mutants in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Yan Lu, Linda J. Savage, and Robert L. Last
Part III Proteomics and Suborganellar Fractionation 10 Preparation of Envelope Membrane Fractions from Arabidopsis Chloroplasts for Proteomic Analysis and Other Studies . . . . . . . . 189 Daniel Salvi, Lucas Moyet, Daphné Seigneurin-Berny, Myriam Ferro, Jacques Joyard, and Norbert Rolland
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11 Preparation of Stroma, Thylakoid Membrane, and Lumen Fractions from Arabidopsis thaliana Chloroplasts for Proteomic Analysis . . . . . . . . . . . . . . . . Michael Hall, Yogesh Mishra, and Wolfgang P. Schröder 12 Preparation of Plastoglobules from Arabidopsis Plastids for Proteomic Analysis and Other Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Celine Besagni, Lucia Eugeni Piller, and Claire Bréhélin 13 Preparation and Proteomic Analysis of Chloroplast Ribosomes . . . . . . . . . . . . . . . . Kenichi Yamaguchi 14 The Workflow for Quantitative Proteome Analysis of Chloroplast Development and Differentiation, Chloroplast Mutants, and Protein Interactions by Spectral Counting . . . . . . . . . . . Giulia Friso, Paul Dominic B. Olinares, and Klaas J. van Wijk 15 Use of Phosphoproteomics to Study Posttranslational Protein Modifications in Arabidopsis Chloroplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anne Endler and Sacha Baginsky
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Part IV Photosynthesis and Biochemical Analysis 16 Use of a Pulse-Amplitude Modulated Chlorophyll Fluorometer to Study the Efficiency of Photosynthesis in Arabidopsis Plants . . . . . . . . . . . . . . . Matthew D. Brooks and Krishna K. Niyogi 17 Gas Exchange Measurements for the Determination of Photosynthetic Efficiency in Arabidopsis Leaves . . . . . . . . . . . . . . . . . . . . . . . . . Giles Johnson and Erik Murchie 18 Measurement of the DpH and Electric Field Developed Across Arabidopsis Thylakoids in the Light . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steven M. Theg and Curtis Tom 19 Measurement of Chloroplast ATP Synthesis Activity in Arabidopsis . . . . . . . . . . . . Aleel K. Grennan and Donald R. Ort 20 Methods for Analysis of Photosynthetic Pigments and Steady-State Levels of Intermediates of Tetrapyrrole Biosynthesis . . . . . . . . . . Olaf Czarnecki, Enrico Peter, and Bernhard Grimm 21 Analysis of Starch Metabolism in Chloroplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carmen Hostettler, Katharina Kölling, Diana Santelia, Sebastian Streb, Oliver Kötting, and Samuel C. Zeeman 22 Analysis of Lipid Content and Quality in Arabidopsis Plastids . . . . . . . . . . . . . . . . . Anna Maria Zbierzak, Peter Dörmann, and Georg Hölzl Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Contributors Birgit Agne • Abteilung Pflanzenbiochemie, Martin-Luther-Universität Halle-Wittenberg, Halle, Germany Charles Andrès • Plant Physiology Laboratory, University of Neuchatel, Neuchatel, Switzerland Eva-Mari Aro • Molecular Plant Biology, Department of Biochemistry and Food Chemistry, University of Turku, Turku, Finland Sacha Baginsky • Institut für Biochemie und Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Halle, Saale, Germany Jocelyn Bédard • Institute for Protein Research, Osaka University, Suita, Osaka, Japan Celine Besagni • Laboratoire de Physiologie Végétale, Université de Neuchâtel, Neuchâtel, Switzerland Claire Bréhélin • Laboratoire de Biogenèse Membranaire, UMR 5200, CNRS-Université Victor Segalen Bordeaux 2, Bordeaux, France Matthew D. Brooks • Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA; Division of Physical Biosciences, Lawrence Berkeley National Laboratory, Berkeley, CA, USA Célia Regina Carlini • Department of Biophysics, Center of Biotechnology, Universidade Federal do Rio Grande do Sul, Porto Alegre, RS, Brazil Olaf Czarnecki • Institute of Biology/Plant Physiology, Humboldt University Berlin, Berlin, Germany Diogo Ribeiro Demartini • Department of Biophysics, Center of Biotechnology, Universidade Federal do Rio Grande do Sul, Porto Alegre, RS, Brazil Peter Dörmann • Institute of Molecular Physiology and Molecular Biotechnology of Plants (IMBIO), University of Bonn, Bonn, Germany Anne Endler • Institut für Biochemie und Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Halle, Saale, Germany Myriam Ferro • Laboratoire Etude de la Dynamique du Protéome, CEA, INSERM, Université de Grenoble, Grenoble, France Giulia Friso • Department of Plant Biology, Cornell University, Ithaca, NY, USA Peter Geigenberger • Plant Metabolism, Department Biologie I, Ludwig-Maximilians-Universität München, Planegg-Martinsried, Germany Aleel K. Grennan • Department of Plant Biology, University of Illinois, Urbana, IL, USA Bernhard Grimm • Institute of Biology/Plant Physiology, Humboldt University Berlin, Berlin, Germany Michael Hall • Department of Biological Chemistry, Institute of Chemistry and Umeå Plant Science Centre (UPSC), Umeå University, Umeå, Sweden Georg Hölzl • Institute of Molecular Physiology and Molecular Biotechnology of Plants (IMBIO), University of Bonn, Bonn, Germany ix
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Carmen Hostettler • Department of Biology, ETH Zurich, Zurich, Switzerland Giles Johnson • Life Sciences, University of Manchester, Manchester, UK Jacques Joyard • Laboratoire de Physiologie Cellulaire Végétale, CNRS, CEA, INRA, Université de Grenoble, Grenoble, France Felix Kessler • Plant Physiology Laboratory, University of Neuchatel, Neuchatel, Switzerland Shingo Kikuchi • Institute for Protein Research, Osaka University, Suita, Osaka, Japan Tatjana Kleine • Lehrstuhl für Molekularbiologie der Pflanzen (Botanik), Department Biologie I, Ludwig-Maximilians-Universität, Planegg-Martinsried, Germany Katharina Kölling • Department of Biology, ETH Zurich, Zurich, Switzerland Oliver Kötting • Department of Biology, ETH Zurich, Zurich, Switzerland Robert L. Last • Department of Biochemistry and Molecular Biology, and Department of Plant Biology, Michigan State University, East Lansing, MI, USA Dario Leister • Lehrstuhl für Molekularbiologie der Pflanzen (Botanik), Department Biologie I, Ludwig-Maximilians-Universität, Planegg-Martinsried, Germany Yan Lu • Department of Biological Sciences, Western Michigan University, Kalamazoo, MI, USA Jodi Maple • Centre For Organelle Research, Universitetet i Stavanger, Stavanger, Norway Jörg Meurer • Department Biologie I, Biozentrum der LMU München, Planegg-Martinsried, Germany Yogesh Mishra • Department of Biological Chemistry, Institute of Chemistry and Umeå Plant Science Centre (UPSC), Umeå University, Umeå, Sweden Simon G. Møller • Centre For Organelle Research, Universitetet i Stavanger, Stavanger, Norway Lucas Moyet • Laboratoire de Physiologie Cellulaire Végétale, CNRS, CEA, INRA, Université de Grenoble, Grenoble, France Erik Murchie • Division of Plant and Crop Sciences, School of Biosciences, University of Nottingham, Sutton Bonington, UK Masato Nakai • Institute for Protein Research, Osaka University, Suita, Osaka, Japan Krishna K. Niyogi • Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA; Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA Paul Dominic B. Olinares • Department of Plant Biology, and Department of Chemistry and Chemical Biology, Cornell University, Ithaca, NY, USA Donald R. Ort • Photosynthesis Research Unit, U.S. Department of Agriculture-Agricultural Research Service, Urbana, IL, USA; Department of Plant Biology, University of Illinois, Institute for Genomic Biology, Urbana, IL, USA Enrico Peter • Institute of Biology/Plant Physiology, Humboldt University Berlin, Berlin, Germany Lucia Eugeni Piller • Laboratoire de Physiologie Végétale, Université de Neuchâtel, Neuchâtel, Switzerland
Contributors
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Norbert Rolland • Laboratoire de Physiologie Cellulaire Végétale, CNRS, CEA, INRA, Université de Grenoble, Grenoble, France Daniel Salvi • Laboratoire de Physiologie Cellulaire Végétale, CNRS, CEA, INRA, Université de Grenoble, Grenoble, France Diana Santelia • Department of Biology, ETH Zurich, Zurich, Switzerland Linda J. Savage • Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, USA Enrico Schleiff • Molecular Cell Biology of Plants, Centre of Membrane Proteomics, Cluster of Excellence Macromolecular Complexes, Goethe-University, Frankfurt, Germany Wolfgang P. Schröder • Department of Biological Chemistry, Institute of Chemistry and Umeå Plant Science Centre (UPSC), Umeå University, Umeå, Sweden Daphné Seigneurin-Berny • Laboratoire de Physiologie Cellulaire Végétale, CNRS, CEA, INRA, Université de Grenoble, Grenoble, France Sari Sirpiö • Molecular Plant Biology, Department of Biochemistry and Food Chemistry, University of Turku, Turku, Finland Sebastian Streb • Department of Biology, ETH Zurich, Zurich, Switzerland Marjaana Suorsa • Molecular Plant Biology, Department of Biochemistry and Food Chemistry, University of Turku, Turku, Finland Jay J. Thelen • Department of Biochemistry and Interdisciplinary Plant Group, Christopher S. Bond Life Sciences Center, University of Missouri, Columbia, MO, USA Steven M. Theg • Department of Plant Biology, University of California Davis, Davis, CA, USA Axel Tiessen • Departamento de Ingenieria Genetica, CINVESTAV, Irapuato, Mexico Curtis Tom • Department of Plant Biology, University of California Davis, Davis, CA, USA Joanna Tripp • Molecular Cell Biology of Plants, Goethe-University, Frankfurt, Germany Klaas J. van Wijk • Department of Plant Biology, Cornell University, Ithaca, NY, USA Kenichi Yamaguchi • Division of Biochemistry, Nagasaki University, Nagasaki, Japan Anna Maria Zbierzak • Institute of Molecular Physiology and Molecular Biotechnology of Plants (IMBIO), University of Bonn, Bonn, Germany Samuel C. Zeeman • Department of Biology, ETH Zurich, Zurich, Switzerland
Part I Multiprotein Complexes and Protein–Protein Interactions
Chapter 1 One- and Two-Dimensional Blue Native-PAGE and Immunodetection of Low-Abundance Chloroplast Membrane Protein Complexes Shingo Kikuchi, Jocelyn Bédard, and Masato Nakai Abstract Blue native polyacrylamide gel electrophoresis (BN-PAGE) is a powerful method for separating protein complexes from biological membranes under native conditions. BN-PAGE provides much higher resolution than gel filtration or sucrose density gradient centrifugation, and it can be used to estimate the molecular mass of protein complexes. First, membrane protein complexes need to be solubilized with a mild nonionic detergent such as digitonin or dodecyl maltoside. Coomassie brilliant blue G-250, a negatively charged dye that binds to the surface of the solubilized complexes, is then added so these can be resolved according to their size by non-denaturing (native) electrophoresis. BN-PAGE can be combined with a second dimension SDS-PAGE step (two-dimensional (2D)-BN/SDS-PAGE), so that the subunits making up these complexes are also separated according to their size. Here, we present our 2D-BN/SDSPAGE method, and subsequent immunoblotting method, for the detection of relatively low-abundance proteins from plant chloroplasts. Key words: Blue native polyacrylamide gel electrophoresis, Chloroplast, Arabidopsis, Digitonin, Membrane protein complexes, Immunoblotting
1. Introduction Blue native polyacrylamide gel electrophoresis (BN-PAGE) was developed by Schägger and von Jagow (1, 2) and has been widely used for the analysis of membrane protein complexes from mitochondria, chloroplasts, and bacteria (3). BN-PAGE provides much higher resolution than size exclusion chromatography or sucrose density gradient centrifugation, and further provides a more reliable assessment of the molecular mass of protein complexes (2). In addition to membrane protein complexes, water-soluble protein complexes can also be separated by BN-PAGE.
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_1, © Springer Science+Business Media, LLC 2011
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The BN-PAGE method is summarized as follows: (1) Isolated membranes or organelles are solubilized using a mild, nonionic detergent such as digitonin or dodecyl maltoside; (2) Coomassie brilliant blue G-250 (CBB-G) is added to the solubilized sample; and (3) protein complexes are separated in a non-denaturing polyacrylamide gradient gel. The key step that makes this method unique is the addition of CBB-G to solubilized membrane protein complexes before electrophoresis. CBB-G is a negatively charged molecule that binds to the surface of protein complexes and displaces the detergent molecules, in most cases, without affecting the stability of the complexes. During electrophoresis, because of the negative charge conveyed by the bound CBB-G, protein complexes migrate toward the anode and are resolved according to their size. BN-PAGE allows the separation of protein complexes in the molecular mass range of ~20 kDa to several thousand kilodaltons. In addition, the subunit composition of the protein complexes separated by a one-dimensional (1D) BN-PAGE can be analyzed by performing a second dimension electrophoresis step. This second electrophoresis step is carried out in the presence of the strong, denaturing ionic detergent sodium dodecyl sulfate (SDS) and, as a consequence, is similar to a standard SDS-PAGE analysis (4). In this two-dimensional BN/SDS-PAGE (2D-BN/SDS-PAGE) analysis, the denatured subunits of protein complexes are aligned in vertical columns, allowing their visualization by staining or immunodetection. This, therefore, makes 2D-BN/SDS-PAGE a very powerful technique for the characterization of membrane protein complexes. To date, the two photosystems, the light-harvesting complexes, the ATP synthase, the cytochrome b6f complex, and other membrane complexes from plant chloroplasts have been well characterised by both 1D-BN-PAGE and 2D-BN/SDS-PAGE (5–10). In these cases, subunits of these abundant protein complexes can be visualized by Coomassie staining or silver staining. However, if the amount of proteins of interest is low, immunoblotting is required. Recently, we have used 1D-BN-PAGE and 2D-BN/ SDS-PAGE followed by immunoblotting for the analysis of some of the translocon complexes at the outer and inner envelope membranes of chloroplasts (TOC and TIC) (11). We have also adapted 1D-BN-PAGE and 2D-BN/SDS-PAGE for a detailed characterization of radiolabeled translocation intermediates (12). Applying the 1D-BN-PAGE and 2D-BN/SDS-PAGE and subsequent immunoblotting methods for analyses of proteins from the model plant Arabidopsis is especially advantageous and, therefore, highly recommended for the following reasons: (1) relatively small amounts of proteins (in most cases less than 100 mg) are generally sufficient for a single electrophoretic analysis; (2) a large number of publically available mutant lines and various transgenic lines can be used for comparative analyses with the methods to
1 One- and Two-Dimensional Blue Native-PAGE…
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examine their effects on the biogenesis and/or assembly of the complex of interest; (3) the completely sequenced genome information and growing numbers of various proteomic data, both of which are also available in public databases, may help to identify proteins after the electrophoretic separation. In this chapter, we would like to introduce our 1D-BN-PAGE and 2D-BN/SDS-PAGE techniques and subsequent immunoblotting protocols to visualize relatively low-abundance proteins from plant chloroplasts.
2. Materials 2.1. First-Dimension BN-PAGE
1. AB mix: 48% (w/v) acrylamide/1.5% (w/v) bisacrylamide (49.5% T, 3% C); store at 4°C.
2.1.1. S tock Solutions
2. 3× Gel buffer: 150 mM Bis-Tris–HCl, pH 7.0, and 1.5 M aminocaproic acid; store at 4°C. 3. 10× Cathode buffer: 0.5 M Tricine and 150 mM Bis-Tris–HCl, pH 7.0; store at room temperature. 4. 10× Anode buffer: 0.5 M Bis-Tris–HCl, pH 7.0; store at room temperature. 5. 5% CBB-G solution: 5% (w/v) Serva blue G (Coomassie brilliant blue G-250, Serva, Germany) in 50 mM Bis-Tris–HCl, pH 7.0, and 0.5 M aminocaproic acid. Filter through a 0.2-mm membrane filter using a microcentrifuge; store at 4°C. 6. Water-saturated butanol; store at room temperature.
2.1.2. Freshly Prepared Solutions
We prepare these solutions on the day of the experiment. 1. 1× Cathode buffer containing CBB-G and 1× anode buffer: dissolve 0.1 g (0.02%) of Serva blue G (Coomassie brilliant blue G-250) completely in 500 mL of 1× cathode buffer. These buffers should be precooled (4°C) before use. 2. 10% (w/v) ammonium persulfate (APS) solution. 3. 5% (w/v) water-soluble digitonin solution (see Note 1). 4. Solubilization buffer: 50 mM Bis-Tris–HCl, pH 7.0, 0.5 M aminocaproic acid, 10% (w/v) glycerol, 1% (w/v) water-soluble digitonin, and 1% (v/v) protease inhibitor cocktail for plant extracts (Sigma, P-9599) (see Note 2). Detergents other than digitonin can be used for BN-PAGE (see Notes 3 and 4). 5. Molecular mass standards for BN-PAGE: ferritin solution (>50 mg/mL, ferritin from equine spleen; store at 4°C) and bovine serum albumin (BSA) solution (2% (w/v); store at 4°C) are diluted to give final concentrations of 2 and 0.2 mg/mL, respectively, in 50 mM Bis-Tris–HCl, pH 7.0, 0.5 M
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a minocaproic acid, 10% (w/v) glycerol, 1% (w/v) water-soluble digitonin, and 1% (v/v) protease inhibitor cocktail. The detergent and protease inhibitor cocktail are added so that the conditions in all samples are the same for electrophoresis. 2.1.3. O ther Materials
1. N,N,N,N ¢-tetramethylethylenediamine (TEMED). 2. Gradient maker GR-40 (Advantec, Japan; present code CHG042AA) (see Note 5); equipped with 75-cm tube (diameter = 1–2 mm). 3. A pair of glass gel plates (16 × 16 cm) with 1-mm-thick spacer. 4. Electrophoresis tank suitable for 16 × 16-cm gel plates such as the Bio Craft BE-122 system (Tokyo, Japan). 5. Hamilton syringe. 6. Ultracentrifuge (e.g., Hitachi CS120GX with the S100AT3 rotor). 7. Optional: Vacuum desiccator.
2.2. SecondDimension SDS-PAGE
1. 30% Acrylamide/bis solution: 29.2% (w/v) acrylamide and 0.8% (w/v) bisacrylamide (30% T, 2.6% C); store at 4°C.
2.2.1. S tock Solutions
2. 4× LGB: 1.5 M Tris–HCl, pH 8.8, and 0.4% (w/v) sodium dodecyl sulfate (SDS); store at room temperature. 3. 4× UGB: 0.5 M Tris–HCl, pH 6.8, and 0.4% (w/v) SDS; store at room temperature. 4. 10× Running buffer: 0.25 M Tris, 1.92 M glycine, and 1% (w/v) SDS; store at room temperature. 5. Water-saturated butanol; store at room temperature.
2.2.2. Freshly Prepared Solutions
1. 10% (w/v) APS solution. 2. SDS denaturing buffer: 3.3% (w/v) SDS, and 4% (v/v) 2-mercaptoethanol in 65 mM Tris–HCl, pH 6.8 (see Notes 6 and 7). 3. Agarose sealing buffer: 0.5% (w/v) high-quality agarose, 25 mM Tris, 192 mM glycine, and 0.1% (w/v) SDS.
2.2.3. O ther Materials
1. TEMED. 2. Pre-stained protein marker. 3. Pairs of glass gel plates (16 × 16 cm) with 1.5-mm-thick spacer. 4. 13-cm sized cooking knife which has a sharp, smooth, and curved cutting edge. 5. Flat-end forceps (13 cm). 6. Hybridization oven and bottles. 7. Electrophoresis tank suitable for 16 × 16 cm gel plates such as the Bio Craft BE-122 system (Tokyo, Japan).
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2.3. Western Blotting
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1. Trans-Blot cell system (Bio-Rad, tank blotting system). 2. Polyvinylidene difluoride (PVDF) membrane (Millipore) (see Note 8). 3. Transfer buffer: 100 mM Tris, 192 mM glycine, and 20% (v/v) methanol. 4. TBS-T: 25 mM Tris–HCl, pH 7.5, 137 mM NaCl, 2.7 mM KCl, and 0.05% (w/v) Tween 20. 5. 5% Milk in TBS-T: 10 g skimmed milk is dissolved in 200 mL TBS-T. Stir for more than 20 min at room temperature. 6. Enhanced chemiluminescence (ECL) system (GE Healthcare) and ECL plus system (GE Healthcare) are used for the visualization of immunodecorated proteins.
3. Methods Blue native-PAGE separates membrane protein complexes in a native state. Therefore, all steps of sample preparation should be performed under cold conditions (4°C). The selection of a suitable detergent to solubilize the membrane protein complexes is important. Nonionic detergents such as digitonin, dodecyl maltoside, and Triton X-100 have been successfully used for BN-PAGE (see Note 3). We recommend the use of digitonin as a first choice since it may prevent the loss of protein subunits which are only weakly associated to the complex of interest. Digitonin is considered to be one of the mildest detergents for solubilization. The subunits of protein complexes resolved by BN-PAGE can be size-separated by second-dimension SDS-PAGE. However, before this, denaturation of protein complexes with SDS and 2-mercaptoethanol is essential. After denaturation, a BN-PAGE gel lane is overlaid on top of the stacking gel of the second-dimension SDS-PAGE gel. To facilitate this procedure, the first-dimension BN-PAGE is carried out using a 1-mm-thick gel, and the seconddimension SDS-PAGE is carried out using thicker, 1.5-mm gels. When doing immunoblotting, although blots can be produced directly from 1D-BN-PAGE gels, we strongly recommend to also use 2D-BN/SDS-PAGE blots to confirm whether a detected signal on 1D-BN-PAGE blots corresponds to the true specific target protein of the antibody and not an unspecific cross-reacting protein. The 2D-BN/SDS-PAGE blots enable this by showing the actual size of the detected protein, whereas this information cannot be obtained from 1D-BN-PAGE blots. 3.1. Pouring the Gradient Gel
The given recipe is suitable for a pair of gel plates with 16 × 16 cm dimensions and with 1-mm thick spacers. We normally make a 4–14% linear gradient gel and a 3% stacking gel. This setup allows
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Table 1 Composition of BN-PAGE gels Stacking gel
Gradient separation gel
3% T
4% T
14% T
AB (49.5% T, 3% C)
364 mL
1.2 mL
4.2 mL
3× Gel buffer
2 mL
5 mL
5 mL
Glycerol
–
–
3 g
10% APS
50 mL
50 mL
50 mL
TEMED
5 mL
5 mL
5 mL
Total volume
6 mL
15 mL
15 mL
Note: Gel volumes sufficient to prepare two gels
the separation of protein complexes in the molecular mass range of ~50 kDa to several thousand kilodaltons. 1. Assemble a pair of glass plates with 1-mm-thick spacers. 2. Prepare 4 and 14% separation gel solutions without TEMED and APS as described in Table 1. 3. Optional: Degas both solutions under a vacuum desiccator for 5 min with gentle stirring. 4. Precool the gradient maker and the pair of glass plates in a cold chamber (4°C). Put the gradient maker on an upper shelf of the cold chamber, and put the glass plates on another shelf approximately 50 cm lower. Connect the gradient maker and the pair of glass plates via a 75-cm tube (see Fig. 1). 5. Add TEMED and APS to the two gel solutions as described in Table 1 with gentle stirring on ice. Cooling is required to avoid premature polymerization. 6. Pour 7.5 mL of the 4% gel solution into the chamber furthest from the exit of the gradient maker. Open the valve that separates the two chambers and fill the bridge part between the two chambers with the 4% gel solution. Then, close the valve. 7. Pour 7.5 mL of the 14% gel solution into the chamber closest to the exit of the gradient maker. 8. If the gradient maker used has an exit valve, open the exit valve. Air in the tube is pulled from the bottom of the tube using a syringe to make the gel solution flow. Open the valve separating the two chambers. 9. Pour the gradient gel solution from the top of the glass plates by gravity flow in the cold chamber. This takes about 10 min.
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Fig. 1. A setup for making a gradient gel. A gradient maker GR-40 is connected to a pair of assembled glass gel plates (16 × 16 cm) via a 75-cm tube in a cold chamber (4°C). The gel solution is poured by gravity flow.
10. Overlay the gel with water-saturated butanol. Let the gel polymerize at room temperature for approximately 1 h (see Note 9). 11. After polymerization of the separating gel, discard the overlying butanol and wash the space above the gel several times with deionized water. 12. Prepare a 3% stacking gel solution as described in Table 1. 13. Add TEMED and APS to the stacking gel solution as described in Table 1 with gentle stirring. Pour the stacking gel solution
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and insert an appropriate comb (e.g., a comb with 0.5-cm wide and 2-cm long teeth is suitable to load 40 mL of sample per well). 3.2. Solubilization of the Chloroplasts
In this section, we introduce our method to prepare protein extracts for BN-PAGE from total chloroplasts. Chloroplast sub-fractions such as a thylakoid membrane fraction can be used as starting material (see Chapter 2, Vol. 2). For analysis of envelope membrane proteins, although an isolated envelope membrane fraction can be used as starting material, solubilization from total chloroplasts usually provides satisfactory resolution for the purpose of immuno blotting. 1. Isolation of intact chloroplasts is essentially done by standard methods as described in Chapter 17, Vol. 1 and by others (11, 13). After chloroplast aliquots (0.1 mg chlorophyll) are pelleted at 9,500 × g for 1 min using a standard cooling microcentrifuge, the supernatant is removed using an aspirator. For storage, freeze the pellets without the supernatant in liquid nitrogen and store at −80°C. 2. Resuspend pellets in 200 mL of the solubilization buffer to give a final concentration of 0.5 mg chlorophyll/mL (both fresh and frozen pellets can be used). This chlorophyll concentration normally corresponds to 1–5 mg protein/mL. 3. Incubate the samples on ice for at least 10 min with occasional mixing to allow solubilization. 4. Insoluble material is removed by ultracentrifugation at 100,000 × g for 10 min. Residual insoluble material in the samples may cause smearing during the electrophoretic separation. However, prolonged ultracentrifugation should be avoided since sedimentation of large protein complexes may occur. After ultracentrifugation, transfer the clarified supernatant to a new tube while taking care not to carry over any insoluble material. 5. Add 1 mL of the 5% CBB-G solution to 40 mL of the supernatant (see Note 10). 6. Spin down at 9,500 × g for 1 min.
3.3. First-Dimension BN-PAGE
1. Prepare 500 mL of 1× cathode buffer containing 0.02% CBBG, and 500 mL of 1× anode buffer by diluting the stock solutions. Precool the solutions in the cold chamber (4°C). 2. After polymerization of the stacking gel, carefully remove the comb. Place a piece of white adhesive tape on the surface of the glass plate that is exposed to the cathode buffer such that it allows the visualization of the gel wells in the dark-blue cathode buffer during loading of the samples.
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3. Pour the anode buffer into the lower chamber of an electrophoresis apparatus and assemble the gel onto the apparatus. Pour the cathode buffer into the upper chamber of the electrophoresis apparatus. Precool the setup in the cold chamber. 4. Load 40 mL of the protein samples supplemented with CBB-G into the gel wells with a Hamilton syringe. For estimating the size of the protein complexes, load 40 mL of the molecular mass standards (described in Subheading 2.1.2) in one or more wells (see Note 11). 5. Run electrophoresis overnight (12–14 h) at 100 V (constant voltage) in the cold chamber (4°C) (see Note 12). 6. Stop electrophoresis when the thick blue dye front has reached the bottom of the separation gel. 3.4. Transfer of BN-PAGE Gel Lanes onto the SecondDimension SDS-PAGE Gels
This section describes the transfer of BN-PAGE gel lanes to the second-dimension SDS-PAGE gel. When proteins are to be immunoblotted directly from the BN-PAGE gel (i.e., without performing second-dimension SDS-PAGE), the BN-PAGE gel is incubated in SDS denaturing buffer for 30 min at 37°C (see Notes 6 and 7) and then immunoblotted as described in Subheading 3.6. 1. Assemble two or more pairs of glass plates (16 × 16 cm dimensions) with 1.5-mm-thick spacers (see Note 13). 2. For two 1.5-mm-thick gels, prepare a 12.5% separation gel solution by mixing 20.8 mL of 30% acrylamide/bis solution, 12.5 mL of 4× LGB, 16.5 mL of deionized water, 250 mL of 10% APS, and 25 mL of TEMED (see Note 14). 3. Pour 22.5 mL of the solution into each pair of glass plates to make gels of 9–10 cm in height. Overlay with water-saturated butanol. The gel should polymerize in about 60 min. 4. Discard the overlying butanol and wash the space above the gel several times with deionized water. 5. For two gels, prepare a 6% stacking gel solution by mixing 2 mL of 30% acrylamide/bis solution, 2.5 mL of 4× UGB, 5.5 mL of deionized water, 50 mL of 10% APS, and 5 mL of TEMED. 6. Pour the stacking gel solution, leaving sufficient space to insert a first-dimension BN-PAGE gel lane above the stacking gel, between two glass plates. Overlay with water-saturated butanol. The gel should polymerize within 30 min. 7. Discard the overlying butanol and wash the space for the BN-PAGE gel lane several times with deionized water. 8. Pre-warm SDS denaturing buffer to 37°C. 9. Disassemble the first-dimension BN-PAGE gel plate and remove the stacking gel.
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10. Cut out individual lanes of the BN-PAGE gel with a 13-cm-sized cooking knife. Since sliding of the knife may cause unexpected breakage of the gel, we move the blade in a rocking motion, putting sufficient pressure to produce a clean cut (see Notes 15 and 16). 11. Fill a large tray (e.g., 30 × 23 cm metal tray) with (~1.5 L) deionized water. Put the glass plate with excised lanes under water. Float individual lanes one by one using flat-end forceps in the water to prevent unwanted breakage of the lanes. 12. Pick up the excised lanes by catching the bottom side (14%) using flat-end forceps and incubate in 50 mL of SDS denaturing buffer with constant rotation for 30 min at 37°C (see Note 6). We use a hybridization oven and bottles (normally used for Southern/Northern blotting) for this procedure. Prepare one bottle for each lane to avoid cross-contamination. 13. During the incubation, prepare agarose sealing buffer: dissolve 0.5 g of high-quality agarose in 90 mL of deionized water by boiling (see Note 17). After the temperature drops below 50°C, add 10 mL of 10× running buffer. Keep the agarose sealing buffer as liquid by gentle stirring. 14. Apply 20 mL of pre-stained protein marker to small pieces of filter paper (5 × 5 mm) and allow them to dry. 15. Empty the entire contents of a bottle into the large tray filled with deionized water. Pick up the SDS-treated gel lane by catching the bottom side (14%) using flat-end forceps, and place it on the stacking gel by sliding it between the two glass plates. Avoid leaving any space or air bubbles between the BN-PAGE gel lane and the stacking gel. Repeat this procedure for each BN-PAGE gel lane/bottle. By emptying the contents of each bottle into a tray of fresh water, unwanted breakage of the gel lane may be avoided. Also, proteins that diffused out of the gel are diluted using excess water, and therefore, background of subsequent immunodetection can be reduced. Furthermore, dissipation of 2-mercaptoethanol to the ambient air can be minimized. 16. Remove excess water using filter paper and place filter papers containing protein markers precisely at both ends of the BN-PAGE gel lane (see Note 18). 17. Once the temperature of agarose sealing buffer has decreased to about 40°C, embed the BN-PAGE gel lane and filter papers in agarose sealing buffer by pouring sufficient buffer to fill the empty space above the stacking gel. Because some membrane proteins may aggregate when they become denatured by heating, avoid embedding the gel lanes with excessively warm agarose sealing buffer.
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1. Prepare the 1× running buffer by diluting 10× stock. Assemble the electrophoresis apparatus. 2. Run the electrophoresis by standard methods at room temperature. If an unwanted increase in temperature occurs, cooling is required to prevent the aggregation of membrane proteins. 3. A representative example of a silver-stained 2D-BN/SDSPAGE gel prepared with total chloroplasts is shown in Fig. 2.
3.6. Transfer of Proteins to PVDF Membrane and Immunodetection
1. Cut PVDF membranes and filter papers to fit the dimensions of the gel. 2. Wet PVDF membranes in methanol and preincubate the membranes for a few minutes in transfer buffer. 3. Assemble Trans-Blot cell system (tank blotting system; see Note 19) as described in the manufacturer’s instruction manual.
Fig. 2. Resolution of total chloroplast protein complexes from Arabidopsis by 2D-BN/SDSPAGE. An Arabidopsis chloroplast pellet was solubilized in solubilization buffer containing 1% dodecyl maltoside to give a final concentration of 0.5 mg chlorophyll/mL. After ultracentrifugation, the supernatant was subjected to 4–14% BN-PAGE in the first dimension and 15% SDS-PAGE in the second dimension. Proteins were visualized by silver staining. Molecular mass markers for first-dimension BN-PAGE are ferritin (880 and 440 kDa) and bovine serum albumin (132 and 66 kDa). Major protein complexes (photosystem I (PSI) and rubisco) are indicated by arrows. The authors thank Midori Imai for providing these data.
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4. Transfer the proteins to the PVDF membrane at constant voltage (40 V) for 3–4 h on ice. The 1.5-mm-thick gels require longer transfer times than standard 1.0-mm-thick gels. 5. Following transfer, wash the membranes with methanol to remove CBB-G. 6. Equilibrate the membranes with TBS-T for more than 2 min. 7. Block the membranes by incubation in 5% skimmed milk in TBS-T for 1 h. 8. Carry out primary and secondary antibody incubations in 5% skimmed milk in TBS-T with specific antibodies following standard immunodetection procedures (see Note 20). 9. Visualization of specific proteins is carried out using the enhan ced chemiluminescence (ECL) system for normal-abundance proteins, or the ECL plus system for low-abundance proteins. 10. Representative examples of a number of Western blots from 2D-BN/SDS-PAGE are shown in Fig. 3.
Fig. 3. Detection of specific proteins by immunoblotting after 2D-BN/SDS-PAGE. An Arabidopsis chloroplast pellet was solubilized in solubilization buffer containing 1% digitonin to give a final concentration of 0.5 mg chlorophyll/mL. After ultracentrifugation, the supernatant was subjected to 2D-BN/SDS-PAGE. Proteins were transferred to PVDF membranes and then immunodecorated with anti-pea Toc75, -Arabidopsis Toc33, -pea Tic110, -pea Tic55, or -pea Hsp93 (ClpC) antibodies. Immunodecorated proteins were visualized by the enhanced chemiluminescence (ECL) system exposed to X-ray films.
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4. Notes 1. Digitonin is a natural product and crude digitonin is practically insoluble in water. To obtain reproducible results, we use the water-soluble fraction of digitonin. Water-soluble digitonin can be purified as described (14). Also, high-purity digitonin is available commercially from a small number of manufacturers. 2. When the chloroplast surface has been treated with trypsin, trypsin inhibitor should be included in the solubilization buffer. Residual trypsin often degrades the protein of interest under non-denaturing BN-PAGE conditions. 3. An appropriate detergent should be selected and the concentration used for solubilization should be optimized. A broad survey of the combinations of detergents and organelles/ prokaryotes previously used for BN-PAGE was done by Krause (3). 4. Addition of salt (from 150 mM to 1 M NaCl) to the solubilization buffer often enhances the solubilization of membrane protein complexes (3, 12). 5. In general, it is difficult to make reproducible linear gradient in a small scale. However, in our experience, the gradient maker GR-40, which has two inverted conical chambers and a builtin mixing device, allows us to pour gradient gels with greater reproducibility (see Fig. 1). 6. Sufficient denaturation of proteins separated in the BN-PAGE gel is required both for the transfer to PVDF membrane and for the SDS-PAGE as the second dimension. However, since some membrane proteins tend to aggregate when heated even in the presence of SDS, we normally incubate the BN-PAGE gel at 37°C in the SDS denaturing buffer. If the proteins of interest do not aggregate when heated, the BN-PAGE gel can be incubated at higher temperature (e.g., at 80°C) to facilitate efficient denaturation. 7. When performing immunoblotting directly from a first-dimension BN-PAGE, we and others have experienced that sometimes the expected protein bands are not detected. This is probably due to insufficient denaturation of protein complexes and often happens especially if the denaturing buffer used contains relatively low concentrations of SDS (e.g., ~0.1%, the standard SDS concentration used for 1× SDS-PAGE running buffer). If protein complexes or proteins are not denatured sufficiently, they may be only poorly transferred from the BN-PAGE gel to the PVDF membrane, or even if they are transferred efficiently, the epitopes normally recognized by the antibodies may not be surface exposed and therefore remain inaccessible. The use of
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high concentrations of SDS (3.3%) and 2-mercaptoethanol (4%) in the SDS denaturing buffer we have been using probably alleviates this problem. 8. Use of Millipore PVDF membrane is recommended since it has very smooth surface on both sides. PVDF membranes from other companies may not be of suitable quality. 9. Polymerization of the gel is completed when a layer of water appears between the gel surface and the butanol layer. 10. Changing the ratio between detergent and Coomassie may affect the stability of the protein complex of interest. In some cases, optimization may be required. 11. Plan to cut out at least one marker lane from the BN-PAGE gel after the electrophoresis run is complete for CBB staining. 12. In the original articles describing the BN-PAGE technique (1, 2), it is recommended that when running a BN-PAGE gel for electroblotting, the blue cathode buffer is replaced by fresh buffer lacking CBB-G after one-third of the whole run has been completed, because CBB-G is believed to inhibit the transfer process. We normally omit this step since we have not seen any problems when using the tank or “wet” blotting technique. 13. One SDS-PAGE gel is required for each BN-PAGE gel lane to be analyzed. For comparative analysis, second-dimension gels have to be prepared with BN-PAGE lanes excised from the same gel. 14. Concentration of the separation gel can be changed in order to obtain good resolution of your protein(s) of interest. 15. The use of a sharp knife ensures that smooth cuts can be made when excising gel lanes. If the edge of the excised BN-PAGE lane is notched due to inappropriate cutting, protein bands after 2D-BN/SDS-PAGE might be less focused. 16. BN-PAGE gel lanes can be stored at 4°C for a few days before performing second-dimension electrophoresis. 17. To prevent unwanted air bubbles, agarose should be melted in deionized water by boiling using a microwave oven prior to adding the SDS. 18. Placing the pre-stained markers at both ends of the BN-PAGE gel lanes on the second-dimension SDS-PAGE gels helps to determine the exact position of the gel lanes on the 2D-BN/ SDS-PAGE blots (produced in Subheading 3.6). The prestained marker lanes can also be used to align a CBB-stained BN-PAGE marker lane horizontally with the blots to determine the native position of the complexes as precisely as possible on the immunoblots.
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19. We prefer to use the tank blotting system for generally, in our experience, better transfer efficiency when compared with that of the semi-dry blotting system. 20. If nonspecific immunoblot signals are obtained, with your antibodies, the antibodies should be purified, especially prior to use with 1D-BN-PAGE blots. For a short and simple antibody purification method, we recommend the “blot-affinity purification” method (15). References 1. Schägger, H., and von Jagow, G. (1991) Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 199, 223–231. 2. Schägger, H., Cramer, W. A., and von Jagow, G. (1994) Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis. Anal. Biochem. 217, 220–230. 3. Krause, F. (2006) Detection and analysis of protein-protein interactions in organellar and prokaryotic proteomes by native gel electrophoresis: (membrane) protein complexes and supercomplexes. Electrophoresis 27, 2759–2781. 4. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 5. Kügler, M., Jänsch, L., Kruft, V., Schmitz, U. K., and Braun, H.-P. (1997) Analysis of the chloroplast protein complexes by blue-native polyacrylamide gel electrophoresis (BN-PAGE). Photosynth. Res. 53, 35–44. 6. Poetsch, A., Neff, D., Seelert, H., Schägger, H., and Dencher, N. A. (2000) Dye removal, catalytic activity and 2D crystallization of chloroplast H+−ATP synthase purified by blue native electrophoresis. Biochim. Biophys. Acta 1466, 339–349. 7. Heinemeyer, J., Eubel, H., Wehmhöner, D., Jänsch, L., and Braun, H.-P. (2004) Proteomic approach to characterize the supramolecular organization of photosystems in higher plants. Phytochemistry 65, 1683–1692. 8. Suorsa, M., Regel, R. E., Paakkarinen, V., Battchikova, N., Herrmann, R. G., and Aro,
E.-M. (2004) Protein assembly of photosystem II and accumulation of supercomplexes in the absence of low molecular mass subunits PsbL and PsbJ. Eur. J. Biochem. 271, 96–107. 9. Aro, E.-M., Suorsa, M., Rokka, A., Allahverdiyeva, Y., Paakkarinen, V., Saleem, A., Battchikova, N., and Rintamäki, E. (2005) Dynamics of photosystem II: a proteomic approach to thylakoid protein complexes. J. Exp. Bot. 56, 347–356. 10. Rokka, A., Suorsa, M., Saleem, A., Battchikova, N., and Aro, E.-M. (2005) Synthesis and assembly of thylakoid protein complexes: multiple assembly steps of photosystem II. Biochem. J. 388, 159–168. 11. Kikuchi, S., Hirohashi, T., and Nakai, M. (2006) Characterization of the preprotein translocon at the outer envelope membrane of chloroplasts by blue native PAGE. Plant Cell Physiol. 47, 363–371. 12. Kikuchi, S., Oishi, M., Hirabayashi, Y., Lee, D. W., Hwang, I., and Nakai, M. (2009) A 1-megadalton translocation complex containing Tic20 and Tic21 mediates chloroplast protein import at the inner envelope membrane. Plant Cell 21, 1781–1797. 13. Bruce, B. D., Perry, S., Froehlich, J., and Keegstra, K. (1994) In vitro import of proteins into chloroplasts. In, Plant Molecular Biology Manual, Vol. J1 (Gelvin, S.B., and Schilperoot, R. A., eds.) Kluwer Academic Publishers, Belgium, pp. 1–15. 14. Mori, H., Summer, E. J., Ma, X., and Cline, K. (1999) Component specificity for the thylakoidal Sec and Delta pH-dependent protein transport pathways. J. Cell Biol. 146, 45–55. 15. Tang, W.-J. Y. (1993) Blot-affinity purification of antibodies. Methods Cell Biol. 37, 95–104.
Chapter 2 Analysis of Thylakoid Protein Complexes by Two-Dimensional Electrophoretic Systems Sari Sirpiö, Marjaana Suorsa, and Eva-Mari Aro Abstract Photosynthetic machinery in the thylakoid membrane is prone to modifications depending on environmental, developmental, and morphological parameters. Such plasticity in the composition of the thylakoid membrane protein complexes guarantees efficient function of the photosynthetic machinery. In this chapter, we describe methods for separation of thylakoid membrane protein complexes at high resolution by two-dimensional gel electrophoretic systems. Solubilization of the thylakoid membrane protein complexes either by dodecylmaltoside or digitonin is described first. Then, two partially overlapping methods, blue native gel electrophoresis and high-resolution clear native gel electrophoresis, are demonstrated to separate the individual protein complexes. Finally, denaturing SDS-polyacrylamide gel electrophoresis is used to reveal the protein composition of each complex. Critical points in all protocols are addressed and representative examples of the composition of Arabidopsis thaliana thylakoid membrane protein complexes are shown. Key words: Chloroplast, Electrophoresis, Native gel, Photosynthesis, Protein complex, Thylakoid, Two-dimensional gel electrophoresis
1. Introduction Two-dimensional (2D) gel electrophoresis is a useful tool to resolve the composition of the thylakoid membrane protein complexes. For 2D gel electrophoresis, the protein complexes are first solubilized from the membrane, then separated in a native gel, followed by denaturing sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), which further separates the distinct subunits of the complexes. Blue native gel electrophoresis (1) has been successfully applied for the first dimension separation of the thylakoid membrane protein complexes of various photosynthetic organisms. Prior to blue native gel electrophoresis, thylakoid membranes are solubilized with mild
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_2, © Springer Science+Business Media, LLC 2011
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detergent (e.g., dodecyl maltoside [DM] or digitonin). Subsequently, Coomassie G-250 dye is added to the sample. Coomassie dye introduces a negative charge to the protein complexes and, thus, enhances migration and reduces aggregation of the protein complexes during electrophoresis. The Coomassie dye is omitted from clear native gel electrophoresis, which is thus more suitable for in-gel catalytic activity assays and for detection and quantification of proteins tagged with fluorescent dyes. However, the lack of the negative charge often results to less efficient separation of the protein complexes than in blue native gels. Recently, a variation of the clear native gel electrophoresis, in which mild anionic and nonionic detergents are used to introduce a negative charge to the protein complexes, was introduced (2). The advantage of the modified protocol is that it enables better solubilization and enhanced resolution of the protein complexes (2). Here, we describe optimized methods for 2D separation of Arabidopsis thaliana thylakoid membrane protein complexes by using both blue native and the improved clear native gel electrophoresis method. High diversity of the various photosystem (PS) II assemblies is a distinct feature of the higher plant thylakoid membrane in natural environments. Two extreme examples of such are the PSII-light harvesting complex (LHC) II supercomplex and the monomeric PSII complex lacking CP43, which represent the most active PSII complex and a transient assembly intermediate of PSII undergoing repair cycle, respectively (3). In addition, several other supercomplexes, either permanent or temporary, can be found in the thylakoid membrane of higher plants. For example, the PSI–LHCI complex is capable of interacting with LHCII in low light conditions, thus forming a state transition-specific supercomplex (4). Dynamic alterations in the composition of the thylakoid membrane protein complexes guarantee maximal photosynthetic efficiency under fluctuating environmental conditions. Analysis of thylakoid protein complexes by various 2D electrophoretic systems provides an excellent tool for monitoring the structure–function relationship of the thylakoid membrane, and thus reliable separation and resolution techniques are necessary to guarantee the best possible understanding of the photosynthetic apparatus.
2. Materials 2.1. Thylakoid Membrane Isolation
1. Grinding buffer: 50 mM 4-(2-hydroxyethyl)piperazine-1- ethanesulfonic acid (HEPES)–KOH, pH 7.5, 330 mM sorbitol, 2 mM ethylenediaminetetraacetic acid (EDTA), 1 mM MgCl2 (store at −20°C). 5 mM ascorbate, 0.05% (w/v) bovine serum albumin, and 10 mM natrium fluoride are added to the solution prior to use (see Note 1).
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2. Shock buffer: 50 mM HEPES–KOH, pH 7.5, 5 mM sorbitol, 5 mM MgCl2 (store at −20°C). 10 mM natrium fluoride is added to the solution prior to use if protein phosphorylation is to be addressed. 3. Storage buffer: 50 mM HEPES–KOH, pH 7.5, 100 mM sorbitol, 10 mM MgCl2 (store at −20°C). 10 mM natrium fluoride is added to the solution prior to use if protein phosphorylation is to be addressed. 4. Homogenizer (e.g., Ultra-Turrax). 2.2. Casting the Gradient Gel 2.2.1. Stock Solutions and Reagents
1. Acrylamide stock: 48% (w/v) acrylamide, 1.5% (w/v) bisacrylamide solution (store at room temperature [RT]). Caution: Toxic. 2. Gel buffer (3×): 150 mM Bis Tris, pH 7.0, 1.5 M e-amino-ncaproic acid (store at 4°C). 3. 75% (w/v) glycerol solution in water (store at 4°C). 4. Ammonium persulfate (APS): 5% (w/v) solution in water. 5. N,N,N,N ¢-tetramethylethylenediamine (TEMED). Caution: Toxic.
2.2.2. Working Solutions to Be Freshly Prepared Prior to Use
1. Heavy solution (12.5% (w/v) acrylamide): 0.530 mL of acrylamide stock, 0.7 mL of 3× gel buffer, 0.560 mL of 75% (w/v) glycerol, 0.290 mL of water, 2 mL of TEMED, and 11 mL of 5% (w/v) APS. 2. Light solution (5% (w/v) acrylamide): 0.212 mL of acrylamide stock, 0.7 mL of 3× gel buffer, 0.140 mL of 75% (w/v) glycerol, 1.028 mL of water, 2 mL of TEMED, and 11 mL of 5% (w/v) APS. 3. Stacking gel (4% (w/v) acrylamide): 0.121 mL of acrylamide stock, 0.5 mL of 3× gel buffer, 0.860 mL of water, 3 mL of TEMED, and 16 mL of 5% (w/v) APS.
2.2.3. Instruments
1. Gel caster (e.g., Hoefer dual gel caster, Amersham Biosciences). 2. 10 × 8 cm plates (one aluminium plate to disperse heat evenly, one glass plate), spacers (0.75 mm), and comb. 3. Gradient maker (e.g., Hoefer SG5, Amersham Biosciences). 4. Pump for gradient gel casting (e.g., Ismatec), flow rate 0.5 mL/ min. 5. Silicone tube (inner diameter 2 mm), needle (inner diameter 0.6 mm). 6. Dust-free paper (e.g., Whatman chromatography paper 3MM CHR).
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2.3. Sample Preparation for Blue Native Gel Electrophoresis 2.3.1. Stock Solutions and Reagents
1. 50BTH40G solution: 50 mM Bis Tris–HCl, pH 7.0, 40% (w/v) glycerol (store at 4°C). 2. 10 mg/mL Pefabloc SC (4-[2-aminoethyl]-benzenesulfonyl fluoride hydrochloride [AEBSF]; Roche) solution in water (store at −20°C). 3. Serva Blue G buffer: 100 mM Bis Tris–HCl, pH 7.0, 0.5 M e-amino-n-caproic acid, 30% (w/v) sucrose, 50 mg/mL Serva Blue G (store at −20°C) (see Note 2). 4. 10% (w/v) n-dodecyl b-d-maltoside (DM) solution in water (store at −20°C). 5. 10% (w/v) digitonin, high purity (Calbiochem), solution in water (store at −20°C) (see Note 3). Caution: Toxic. 6. 200 mM natrium fluoride solution in water (prepare fresh). Caution: Toxic.
2.3.2. Working Solutions to Be Freshly Prepared Prior to Use
1. 25BTH20G solution: 50 mL of 50BTH40G, 42.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, and 5 mL of 200 mM natrium fluoride. 2. Buffer for empty wells (if unused wells exist): 50 mL of 50BTH40G, 10 mL of 10% (w/v) DM or 10% (w/v) digitonin, 40 mL of water, and 10 mL of Serva Blue G buffer. 3. 2% DM: 50 mL of 50BTH40G, 27.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, and 20 mL of 10% (w/v) DM. 4. 4% digitonin: 50 mL of 50BTH40G, 2.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, 5 mL of 200 mM natrium fluoride, and 40 mL of 10% (w/v) digitonin.
2.4. Sample Preparation for High-Resolution Clear Native Gel Electrophoresis
1. 50BTH40G solution: 50 mM Bis Tris–HCl, pH 7.0, 40% (w/v) glycerol (store at 4°C).
2.4.1. Stock Solutions and Reagents
4. 10% (w/v) digitonin, high purity (Calbiochem), solution in water (store at −20°C) (see Note 3). Caution: Toxic.
2. 10 mg/mL Pefabloc SC (AEBSF, Roche) solution in water (store at −20°C). 3. 10% (w/v) DM solution in water (store at −20°C).
5. 200 mM natrium fluoride solution in water (prepare fresh). Caution: Toxic. 2.4.2. Working Solutions to Be Freshly Prepared Prior to Use
1. 25BTH20G solution: 50 mL of 50BTH40G, 42.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, and 5 mL of 200 mM natrium fluoride. 2. Buffer for empty wells (if unused wells exist): 50 mL of 50BTH40G, 10 mL of 10% (w/v) DM or 10% (w/v) digitonin, and 40 mL of water. 3. 2% DM: 50 mL of 50BTH40G, 27.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, and 20 mL of 10% (w/v) DM.
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4. 4% digitonin: 50 mL of 50BTH40G, 2.5 mL of water, 2.5 mL of 10 mg/mL Pefabloc, 5 mL of 200 mM natrium fluoride, and 40 mL of 10% (w/v) digitonin. 2.5. Separation of Thylakoid Membrane Protein Complexes by Gradient Gel Electrophoresis 2.5.1. Stock Solutions and Reagents 2.5.2. Instruments
1. Anode buffer (10×): 0.5 M Bis Tris–HCl, pH 7.0 (store at 4°C). 2. Cathode buffer for blue native (1×): 50 mM Tricine, 15 mM Bis Tris–HCl, pH 7.0, 0.01% (w/v) Serva Blue G (store at 4°C) (see Note 2). 3. Cathode buffer for high-resolution clear native (1×): 15 mM Bis Tris–HCl, pH 7.0, 50 mM Tricine, 0.05% (w/v) sodium deoxycholate, 0.01% (w/v) DM (store at 4°C).
1. Running system Biosciences).
(e.g.,
Hoefer
SE
250,
Amersham
2. Cooling thermostats (e.g., Lauda), cooling output at least −10°C. 3. Power supply. 2.6. Solubilization of the Gradient Gel Strips
1. Laemmli buffer: 138 mM Tris–HCl, pH 6.8, 6 M urea, 22.2% (v/v) glycerol and 4.3% (w/v) SDS (VWR BDH Prolabo) (store at 20°C) (see Note 4). 2. 2-mercaptoethanol (see Note 5). 3. Rocker (rocking mixer).
2.7. SDSPolyacrylamide Gel Electrophoresis 2.7.1. Stock Solutions and Reagents
1. Separation buffer: 1.5 M Tris–HCl, pH 8.8 (store at 20°C). 2. Stacking buffer: 0.5 M Tris–HCl, pH 6.8 (store at 20°C). 3. 50% (w/v) acrylamide/1.33% (w/v) bisacrylamide solution (store at 20°C). Caution: Toxic. 4. Urea. 5. 20% (w/v) SDS (VWR BDH Prolabo) in water (store at 20°C) (see Note 4). Caution: Toxic. 6. TEMED. Caution: Toxic. 7. APS: 10% (w/v) solution in water. 8. Water–isopropanol solution (1:1) (store at 20°C). 9. Running buffer (10×): 250 mM Tris base, 1.9 M glycine, 1% (w/v) SDS (store at 20°C) (see Note 4). 10. 0.5% (w/v) agarose in SDS-PAGE running buffer (1×) (see Note 4).
2.7.2. Working Solutions to Be Freshly Prepared Prior to Use
1. Separation gel (15% (w/v) acrylamide, 6 M urea): 8.1 mL of separation buffer, 10.6 mL of acrylamide/bisacrylamide solution, 12.7 g of urea, 6.2 mL of water, 0.7 mL of 20% (w/v) SDS, 280 mL of 10% (w/v) APS, and 28 mL of TEMED.
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2. Stacking gel (6% (w/v) acrylamide, 6 M urea): 2.5 mL of stacking buffer, 1.2 mL of acrylamide/bisacrylamide solution, 3.6 g of urea, 3.45 mL of water, 2 mL of 20% (w/v) SDS, 100 mL of 10% (w/v) APS, and 10 mL of TEMED. 2.7.3. Instruments
1. Gel casting stand (e.g., Bio-Rad Protean II xi). 2. Glass plates (22.5 × 20 cm and 20 × 20 cm), 1-mm spacers, and 1 mm 2D comb. 3. Running system (e.g., Bio-Rad protean II xi). 4. Cooling thermostats (e.g., Lauda). 5. Power supply. 6. Dust-free paper (e.g., Whatman Chromatography paper 3MM CHR).
3. Methods 3.1. Thylakoid Membrane Isolation
All steps should be performed under very dim light at 4°C, and the samples and buffers should be kept on ice in order to guarantee the static state of the sample. Plants are capable of detecting all visible wavelengths of light, so do not use, for example, a green light source during isolation. 1. Grind fresh leaves gently (e.g., 5 × 2 s pulses with homogenizer) in ice-cold grinding buffer. 2. Filter the suspension through two layers of Miracloth and centrifuge at 5,000 × g at 4°C for 4 min. 3. Suspend the pellet in shock buffer to break down the cells and centrifuge at 5,000 × g at 4°C for 4 min. 4. In order to remove remnants of the shock buffer, resuspend the pellet into storage buffer and centrifuge at 5,000 × g at 4°C for 4 min. 5. Resuspend the pellet into small aliquot of storage buffer. 6. Thylakoid samples can be stored in high concentration at −80°C for later use. Freezing step should be as quick as possible, hence liquid nitrogen is recommended (see Note 6).
3.2. Casting the Gradient Gel
Instructions assume the use of a Hoefer gel caster with 10 × 8 cm plates and Hoefer SG5 gradient maker, but the protocol is easily adaptable to other systems. 1. Prepare the heavy and light solutions on ice to prevent premature polymerization of acrylamide. Immediately after adding TEMED and APS to the heavy and light solutions, transfer
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1.80 mL of heavy solution to the right chamber (which is closer to the pump) of the gradient maker, and 1.72 mL of light solution to the left chamber (see Note 7). 2. Open the tube between the chambers of the gradient maker and turn the pump on (flow rate: 0.5 mL/min). Use a magnetic stirrer in the heavy chamber to mix the heavy and light solutions during casting. The pump flow transfers the solutions via a silicone tube to the needle, which is placed between the plates. With this system the acrylamide gradient will be ranging from 12.5% (bottom of the gel) to 5% (top of the gel), but if required, the acrylamide concentration in the gradient gel can easily be adjusted. Use the pump (with lowered flow rate) to overlay the gel with a layer of water in order to guarantee an even surface of the gel. Allow the gradient gel to polymerize for 2 h at 20°C without disturbing the gradient. Before casting the stacking gel, dry the surface of the gradient gel with dust-free paper. 3. Immediately after adding TEMED and APS to the stacking gel solution, pipette the stacking solution on top of the gradient gel and insert a comb to generate wells for the samples. Allow the stacking gel to polymerize for 30 min at 20°C. After polymerization, the gel can be used immediately or it can be stored at +4°C for several days, covered with wet hand towels and wrapped with a plastic film. 3.3. Sample Preparation for Blue Native Gel Electrophoresis
1. Suspend thylakoid membranes (see Note 8) in ice cold 25BTH20G buffer to a final chlorophyll (Chl) concentration of 1.0 mg/mL (keep samples at 4°C in dim light) (see Note 6). 2. Add an equal volume of detergent solution, either 2% (w/v) DM or 4% (w/v) digitonin, and solubilize the thylakoids in darkness for 5 min. When using DM, let the sample solubilize on ice, and for digitonin, mix the sample continuously (e.g., in a shaker) in room temperature (see Note 6). 3. Remove traces of insolubilized material by centrifugation at 18,000 × g at 4°C for 20 min (keep samples at 4°C in dim light). If not removed, insolubilized material is a potential source of streaking. Avoid formation of the air bubbles because the detergent bubbles hamper the injection of the sample to the gel. 4. Prior to loading, supplement the supernatant with 1/10 volume of Serva Blue G buffer to introduce a negative charge. 5. If there are empty wells, fill them with an equal amount of empty well buffer. This is required for even migration of the samples.
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3.4. Sample Preparation for High-Resolution Clear Native Gel Electrophoresis
1. Resuspend thylakoid membranes (see Note 8) in ice cold 25BTH20G buffer to a final Chl concentration of 1.0 mg/mL (keep samples at 4°C in dim light) (see Note 6). 2. Add an equal volume of detergent, either 2% (w/v) DM or 4% (w/v) digitonin, and solubilize the thylakoids for 5 min (keep samples at 4°C in darkness) (see Note 6). When using DM, let the sample solubilize on ice, and for digitonin, mix the sample vigorously (e.g., by vortexing) in room temperature. 3. Remove traces of insolubilized material by centrifugation at 18,000 × g at 4°C for 20 min (keep samples at 4°C in dim light) prior to loading of the samples. If not removed, insolubilized material is a potential source of streaking. Avoid formation of the air bubbles because the detergent bubbles hamper the injection of sample to the gel. 4. If there are empty wells, fill them with an equal amount of empty well buffer. This is required for even migration of the samples.
3.5. Separation of Thylakoid Membrane Protein Complexes by Gradient Gel Electrophoresis
When ready to run the gel, remove the comb and assemble the gel into the running system. Wash the sample wells with a syringe filled with cathode buffer. Fill the upper (cathode) and lower (anode) buffer chambers with the appropriate 1× buffer. Inject the samples into the wells. Perform the electrophoresis at 0°C with gradually increasing the voltage as follows: 75 V for 30 min, 100 V for 30 min, 125 V for 30 min, 150 V for 1 h, 175 V for 30 min, followed by 200 V until the sample reaches the end of the gel (Fig. 1) (see Note 9).
3.6. Solubilization of the Gradient Gel Strips
Excise the strips after electrophoresis and incubate them in Laemmli buffer containing 5% (v/v) 2-mercaptoethanol for 1 h at 20°C to solubilize the protein complexes. For solubilization, place each 1D gel strip in separate 5-mL plastic tube containing 2 mL of solubilization buffer. In order to guarantee sufficient solubilization, place the tubes horizontally to a rocker. Alternatively, strips can be stored in a 5-mL plastic tube in deep freezer for later use (see Note 10).
3.7. SDSPolyacrylamide Gel Electrophoresis
This protocol assumes the use of Bio-Rad Protean II xi Cell gel electrophoresis system but can easily be adjusted for other electrophoresis systems as well. In one 1 × 160 × 160 mm SDS-PAGE gel, two 0.75-mm native gel strips can be run in the second dimension. 1. Prepare the separation gel (15% (w/v) acrylamide, 6 M urea) by mixing separation buffer, acrylamide/bisacrylamide solution, urea, and water until urea has dissolved. Add SDS, APS, and TEMED. Pour the gel, leaving space for a stacking gel, and overlay with water–isopropanol (1:1, v/v) solution to constitute even surface of the gel. Allow the gel to polymerize for 2 h at 20°C. After polymerization, wash the top of the gel five times
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Fig. 1. Separation of thylakoid membrane protein complexes by blue native and high-resolution clear native gel electrophoretic systems. Solubilization of the thylakoid membrane is demonstrated both with n-dodecyl b-d-maltoside (DM) and digitonin (see text for further details). Electrophoresis was performed at 0°C with gradually increasing voltage. Photosystem (PS) II and I complexes together with various combinations of light harvesting complexes (LHC), as well as the chloroplast NAD(P) H dehydrogenase (NDH) and the cytochrome (Cyt) b6f dimer are resolved. Note that when digitonin is used as a solubilization agent, a lot of material does not penetrate into separation gel but instead remains in the stacking gel (dashed lines). sc supercomplex.
with water to remove remnants of isopropanol and then dry with dust-free paper. 2. Prepare the stacking gel (6% (w/v) acrylamide, 6 M urea) by mixing stacking buffer, acrylamide/bisacrylamide solution, urea, and water until urea has dissolved. Add SDS, APS, and TEMED. Pour the stacking gel on top of the separation gel and insert the 2D comb (suitable for gel strips). The stacking gel should polymerize within 30 min at 20°C. 3. Once the stacking gel has polymerized, remove the comb and wash the well first with water and then with SDS-PAGE running buffer to remove impurities. 4. Use a spatula to transfer the strip from the tube to the top of the SDS-PAGE gel. Seal the strip with 0.5% (w/v) agarose in SDS-PAGE running buffer (see Note 11). 5. Add running buffer (1×) to the upper and lower chambers of the gel unit and run the gel over night at 15°C. 6. After electrophoresis, the protein spots of the two-dimensional gel may be directly stained, for example with Coomassie Brilliant Blue R-250, SYPRO®Ruby, or silver nitrate stain (Fig. 2), allowing visualization of all the proteins, or with ProQ® Diamond phosphoprotein gel stain to allow visualization of phosphorylated proteins, or be processed further for different purposes (e.g., for Western blotting).
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Fig. 2. Two-dimensional gel-based separation of thylakoid membrane protein complexes solubilized either with n-dodecyl b-d-maltoside (DM) or digitonin (see Fig. 1). The blue native gel electrophoresis strips (see Fig. 1) were cut out, solubilized with the Laemmli solubilization buffer supplemented with 5% 2-mercaptoethanol and placed horizontally on top of the SDS-PAGE gel. Silver nitrate staining is used for visualization of the proteins. Major photosynthetic protein complexes, photosystems (PS) II and I together with various combinations of light harvesting complexes (LHC), as well as the cytochrome (Cyt) b6f dimer and the ATP synthase are indicated. sc supercomplex.
4. Notes 1. Ascorbate protects proteins from oxidation, and bovine serum albumin is used to diminish the effect of proteases. Add these protectants always to thylakoid grinding buffers. In contrast, natrium fluoride is a protein phosphatase inhibitor. It should be added to all buffers if protein phosphorylation is addressed. Weigh these chemicals to the thylakoid isolation buffers prior to use.
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2. In order to dissolve Serva Blue G completely, mix the solution vigorously for several hours. 3. Digitonin is a detergent, which solubilizes lipids. Digitonin is less-effective solubilization agent than DM. The primary target of digitonin within thylakoid membrane is the stroma lamellae. Before use, purify digitonin according to manufacturer’s recommendation. 4. SDS is used for denaturation of proteins by disrupting noncovalent bonds, and it also gives a negative charge to polypeptides. For the best result, we strongly recommend the using of VWR BDH Prolabo SDS in all 2D electrophoresis solutions. The use of 2D incompatible SDS tends to cause vertical streaking of protein spots in 2D SDS-PAGE gels. 5. 2-mercaptoethanol is used for denaturation of proteins by reducing the disulfide bonds. Mercaptoethanol loses its activity by time, and if vertical streaking of protein spots in 2D SDSPAGE gels is evident, we recommend to purchase a new reagent. 6. Thylakoid membrane protein complexes in higher plants, especially the super and megacomplexes, are easily disassembled during freezing and thawing cycles of thylakoid samples and upon sample preparation for native gel electrophoresis. Thus, avoid repeated freezing and thawing of the thylakoid membrane samples and perform the sample preparation for native gel electrophoresis rapidly under very dim light at 4°C. 7. Prior to the casting of the gradient gel, check that a gel cassette, composed of one aluminium plate, one glass plate and two 0.75-mm spacers, is waterproof. Also, test the fluent operation of the pump with water. If the gel cassette is leaking or the flow of the pump is uneven, the continuity of the gradient might be disturbed. 8. For the best result, proteins are loaded on native gels in amounts corresponding to 30–80 mg of protein per well (mini gels). Overloading might cause aggregation of the sample. 9. The background of the blue native gel can be destained during the run by replacing the cathode buffer with a colorless cathode buffer (without Serva Blue G) when 125 V has been reached. This allows better visualization of the protein complexes. 10. If frozen strips are used, melt them at room temperature before solubilization with Laemmli buffer. 11. Avoid using too hot agarose in order to prevent the melting of the gel strip and subsequent streaking of the protein spots.
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References 1. Kügler, M., Jänsch, L., Kruft, V., Schmitz, U. K., and Braun, H.-P. (1997) Analysis of the chloroplast protein complexes by blue-native polyacrylamide gel electrophoresis (BN-PAGE). Photosynth. Res. 53, 35–44. 2. Wittig, I., Karas, M., and Schägger, H. (2007) High resolution clear native electrophoresis for in-gel functional assays and fluorescence studies of membrane protein complexes. Mol. Cell. Proteomics 7, 1215–1225. 3. Aro, E.M., Suorsa, M., Rokka, A., Allahverdiyeva, Y., Paakkarinen, V., Saleem, A., Battchikova,
N., and Rintamäki, E. (2005) Dynamics of photosystem II: A proteomic approach to thylakoid protein complexes. J. Exp. Bot. 56, 347–356. 4. Pesaresi, P., Hertle, A., Pribil, M., Kleine, T., Wagner, R., Strissel, H., Ihnatowicz, A., Bonardi, V., Scharfenberg, M., Schneider, A., Pfannschmidt, T., and Leister, D. (2009) Arabidopsis STN7 kinase provides a link between short- and long-term photosynthetic acclimation. Plant Cell 8, 2402–2423.
Chapter 3 Preparation of Multiprotein Complexes from Arabidopsis Chloroplasts Using Tandem Affinity Purification Charles Andrès, Birgit Agne, and Felix Kessler Abstract Since its first description in 1998 (Rigaut et al., Nat Biotech 17:1030–1032, 1999), the TAP method, for Tandem Affinity Purification, has become one of the most popular methods for the purification of in vivo protein complexes and the identification of their composition by subsequent mass spectrometry analysis. The TAP method is based on the use of a tripartite tag fused to a target protein expressed in the organism of interest. A TAP tag has two independent binding regions separated by a protease cleavage site, and therefore allows two successive affinity purification steps. The most common TAP tag consists of two IgG binding repeats of Protein A from Staphylococcus aureus (ProtA) separated from a calmodulin-binding peptide by a Tobacco Etch Virus (TEV) protease cleavage site. Using the TAP method, native protein complexes can be purified efficiently with a reduced contaminant background when compared to single step purification methods. Initially developed in the yeast model system, the TAP method has been adapted to most common model organisms. The first report of the purification of protein complexes from plant tissue by the TAP method was published in 2004 by Rohila et al. (Plant J 38:172–181, 2004). The synthetic TAP tag gene described in this study has been optimized for use in plants, and since then, has been successfully used from single gene analyses to high-throughput studies of whole protein families (Rohila et al., PLoS ONE 4:e6685, 2009). Here, we describe a TAP tag purification method for the purification of protein complexes from total Arabidopsis extracts, that we employed successfully using a TAP-tagged chloroplast outer envelope protein. Key words: Protein complex purification, TAP tag, Chloroplast proteomics
1. Introduction This TAP method is based on the use of transgenic Arabidopsis thaliana plants, stably transformed with a gene coding the protein of interest fused to a TAP tag at its N or C terminus. Several variations of the TAP tag have been proposed since its inception, and these are summarized in Fig. 1. The tags may differ in their affinity domains as well as in their cleavage sites; for a review, see
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_3, © Springer Science+Business Media, LLC 2011
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bait
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S-tag TEV
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TEV RGH-6His
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bait
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HTB tag BIO
3C 9His
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Fig. 1. Current TAP tags described in the literature. The lengths of the schematic drawings are proportional to the molecular mass of the tags. Consider that increasing bulk of a tag may lead to steric effects. The original TAP tag has a size of about 20 kDa (1) while the smallest TAP tag is FLAG HA at 3 kDa (Sigma–Aldrich). The other sizes are PTP 18 kDa (2), TAPa 32 kDa (3), SF 4 kDa (4), S3S 4 kDa (5), GS 19 kDa (6), HBH 10 kDa, HTB 13 kDa (7, 8), and HPM 14 kDa (9).
ref. 10. The size of the affinity tags varies from 3 kDa for an FLAG-HA tag, to 32 kDa for the ProtA-His-Myc tag (TAPa tag). In our experiments, we make use of Arabidopsis thaliana plants stably transformed with a gene fused to the TAP tag version of Rohila et al. (11, 12); this is the original TAP tag but optimized for use in plants, consisting of two IgG-binding ProtA repeats, a TEV cleavage site, and a calmodulin-binding peptide (CBP). The choice of the tag, as well as that of the protein terminus to attach it to, are critical points in the experimental design; for a review, see ref. 13. Some proteins might require a free N or C terminus to assure correct targeting, processing, and/or functional activity. In general, the TAP protocol implies two affinity purification steps separated by a protease cleavage step to achieve a high degree
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of purity (Fig. 2). However, in some cases good success may be achieved by carrying out only the first affinity purification step. For the identification of protein complex components by Western blotting and mass spectrometry, this short version of the protocol may be sufficient. The short protocol may also be used if the yield is more important than purity. If, however, highly purified complexes are the objective of the purification, as required for cryoelectron microscopy, functional assays, or identification of associated molecules, such as RNA or lipids, the complete protocol should be followed. With the classical TAP tag, the purification of complexes is achieved through two affinity purifications: firstly, the binding of the Protein A (a cell surface protein from Staphylococcus aureus) to IgG fixed on beads; secondly, the binding of the CBP (the CBP derived from muscle myosin light chain kinase 2 [MLCK2]) to calmodulincoated beads. Each affinity purification step may be carried out by either batch or column procedures. Elution of a TAP-tagged protein from the IgG beads can be done by breaking the strong IgG-ProtA interaction with a standard low pH elution buffer or, by TEV protease cleavage if the calmodulin-affinity chromatography step is to be carried out (Fig. 2). CBP binding to calmodulin beads can be disrupted specifically for sample elution, by incubating with a divalent cation chelator. This strong, Ca2+-dependent affinity allows for a high level of stringency during the washing steps. Chloroplasts, the focus organelle of this book, have seven subcompartments: the two chloroplast envelope membranes and the intermembrane space, the thylakoid membranes and lumen, the stroma, and finally plastoglobule lipid droplets; so, the proteins of interest may either be soluble or associated with one of the chloroplast membrane structures. In this chapter, we present different ways of sample preparation differing in the solubilization process of the membranes. For the purification of soluble protein complexes, the plant tissue is homogenized in a detergent-free buffer. Detergent can be added if the localization of a protein is unclear and may either be in a membrane or soluble compartment. For the purification of membrane-bound protein complexes, the plant tissue is first homogenized in a detergent-free buffer, in order to recover intact membranes by ultracentrifugation; in a second step, the membranes are solubilized using a detergent-containing buffer. These two procedures are summarized in Fig. 3, for both soluble and membrane-bound proteins, and Fig. 4, for membrane-bound proteins only. Quality control during the purification experiment can be carried out by Western blotting. Commercial antibodies are available against all of the commonly used affinity tags, and are extremely helpful to monitor solubilization, affinity, and cleavage steps. In addition, antibodies against known, or presumed, members of a
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first affinity purification
TEV elution
second affinity purification
SDS-PAGE gel separation followed by in-gel digestion
Mass spectrometry analysis
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Homogenisation (grinding buffer with or without detergent) Ultracentrifugation
Recovery of supernatant Incubation with IgG beads IgG beads washing and recovery TEV cleavage elution Incubation with calmodulin beads washing and recovery EGTA elution
SDS-PAGE, in-gel tryptic digest, mass spectrometry
SDS-PAGE, in-gel tryptic digest, mass spectrometry
Fig. 3. Protocol for the purification of protein complexes composed of soluble and membrane proteins. The short protocol stops at the dashed line.
protein complex are excellent tools to evaluate the biological relevance of the samples obtained. The complexes obtained by the TAP tag purification are often analyzed by mass spectrometry to identify the protein components. This type of analysis is highly dependent on the facilities available at the host institution; here, we only describe one efficient way to prepare the samples (in-gel tryptic digest). The proposed TAP tag purification protocol could also be efficiently used in combination with other protocols in this book, like those described in Chapters 10–13, Vol. 2 for the prefractionation of the plant samples, or those in Chapter 1, Vol. 2 for the analysis of purified complexes by native gel electrophoresis. Fig. 2. Principle of the TAP method for the identification of protein complex components. General scheme of the TAP method applied to the identification of the components of a protein complex. Protein complexes (represented by grouped heptagonal shapes in the diagram) are purified from chloroplast extracts by two consecutive affinity purifications and an intervening TEV protease cleavage step. The large spheres shown at each purification step in the diagram represent the two different affinity matrices that bind to the tagged protein. The components of the complex are separated by SDS–PAGE prior to in-gel tryptic digestion followed by identification by mass spectrometry.
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Homogenisation (detergent-free grinding buffer) Ultracentrifugation Recovery of membrane pellet Membrane solubilisation (buffer with detergent)
Ultracentrifugation Recovery of supernatant
Incubation with IgG beads IgG beads washing and recovery TEV cleavage elution Incubation with calmodulin beads washing and recovery EGTA elution
SDS-PAGE, in-gel tryptic digest, mass spectrometry
SDS-PAGE, in-gel tryptic digest, mass spectrometry
Fig. 4. Protocol for the purification of protein complexes composed of membrane-bound proteins. The short protocol stops at the dashed line.
2. Materials 2.1. Plant Culture
1. In vitro medium (½ MS): 0.8% (w/v) Phyto Agar (Duchefa, Haarlem, The Netherlands) containing 0.5× Murashige and Skoog (MS) medium, including vitamins (Duchefa), and 0.8% (w/v) sucrose; pH 5.8 (adjust pH with KOH). 2. Soil: Rasenerde Top Dressing (Ricoter AG, Aarberg, Switzerland).
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2.2. Preparation of Home-Made HsIgG CNBr-Activated Sepharose 4B
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1. Purified human immunoglobulin G (HsIgG) (MP Biomedicals, Irvine, CA, USA). 2. Coupling buffer: 0.1 M NaHCO3, pH 8.5 (adjust pH with 0.1 M Na2CO3). 3. Cyanogen bromide (CNBr)-activated sepharose 4B (GE Healthcare, Chalfont St. Giles, UK; 17-0430-01). 4. 1 mM and 0.1 M HCl (4°C). 5. Sintered glass filter (porosity G3; Schott AG, Mainz, Germany) and side-arm flask for vacuum suction. 6. Blocking buffer: 0.1 M Tris–HCl, pH 8.0. 7. NaCl coupling buffer: Coupling buffer containing 1 M NaCl. 8. 0.1 M glycine–HCl, pH 2.8. 9. 0.2 M glycine–HCl, pH 2.8. 10. Phosphate-buffered saline (PBS): 4.3 mM Na2HPO4, 1.4 mM KH2PO4, 137 mM NaCl, 2.7 mM KCl (the final solution should be pH 7.3). 11. 1 M (~6.5% [w/v]) NaN3 (sodium azide) (see Note 1).
2.3. Preparation of Commercial HsIgG-Sepharose Fast Flow Beads
1. HsIgG-Sepharose Fast Flow (GE Healthcare; 17-0969-01).
2.3.1. Cross-Linking of HsIgG-Sepharose Fast Flow Beads (14) (see Note 2)
4. 0.2 M ethanolamine, pH 8; adjust pH with NaOH.
2. 0.2 M sodium borate, pH 9; adjust pH with NaOH. 3. Dimethyl pimelimidate (DMP; solid) (Sigma–Aldrich, St. Louis, MO, USA). 5. Washing buffer: 1 M NaCl, 50 mM Tris–HCl, pH 8. 6. PBS (see Subheading 2.2). 7. Storage buffer: 0.01% (w/v) NaN3 in PBS (see Note 1).
2.3.2. Equilibration of HsIgG-Sepharose Fast Flow Beads
1. 0.5 M acetic acid (HAc), pH 3.4 (see Note 3). 2. Tris-buffered saline (TBS): 10 mM Tris–HCl, pH 7.5, 150 mM NaCl. 3. 0.1 M glycine–HCl, pH 3.0.
2.4. Affinity Purification of TAP-Tagged Proteins and Associated Complexes (see Note 4)
1. Grinding buffer (GB): 100 mM NaCl, 50 mM Tris–HCl, pH 7.5, 0.5% (v/v) Triton X-100, 1 mM phenylmethanesulfonyl fluoride (PMSF), 5 mM NaF, 0.2% (v/v) Plant Protease Inhibitor Cocktail (Sigma–Aldrich) (see Notes 5 and 6).
2.4.1. Protein Extraction
4. Ultracentrifuge with rotor and suitable ultracentrifuge tubes.
2. Cold mortar and pestle. 3. Miracloth (Merck, Darmstadt, Germany). 5. Bradford assay reagent (e.g., Bio-Rad Protein Assay; Bio-Rad, Hercules, CA, USA).
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2.4.2. Affinity Purifications
1. Mobicol spin columns with 35-mm pore-size filter (Mobitec, Goettingen, Germany) or equivalent for small samples less than 1 g of fresh weight; alternatively, Bio-Spin columns (BioRad) or equivalent for larger samples. 2. 20× TEV buffer : 1 M Tris–HCl, pH 8.0, 10 mM ethylenediaminetetraacetic acid (EDTA), 20 mM dithiothreitol (DTT); add detergeant according to the concentration used in the previous step to avoid precipitation of insoluble proteins during elution steps. 3. AcTEV™ protease (Invitrogen, Carlsbad, CA, USA). 4. Nickel-nitrilotriacetic acid (Ni-NTA) Agarose (Qiagen, Venlo, The Netherlands). 5. Acidic elution buffer (AEB): 0.1 M glycine–HCl, pH 3.0, 0.1% (v/v) Triton X-100. 6. 1 M CaCl2. 7. Calmodulin-binding buffer (CBB): 10 mM Tris–HCl, pH 7.9, 100 mM NaCl, 2 mM CaCl2, 10 mM b-mercaptoethanol, 0.1% (v/v) Triton X-100. 8. Calmodulin–Agarose beads (Sigma–Aldrich). 9. Calmodulin wash buffer (CWB): 10 mM Tris–HCl, pH 7.9, 100 mM NaCl, 0.1 mM CaCl2, 10 mM b-mercaptoethanol, 0.1% (v/v) Triton X-100. 10. Calmodulin elution buffer (CEB): 10 mM Tris–HCl, pH 7.9, 10 mM b-mercaptoethanol, 0.1% (v/v) Triton X-100, 100–200 mM potassium acetate (KAc), 5–20 mM ethylene glycol-bis(2-aminoethylether)-N,N,N ¢,N ¢-tetraacetic acid (EGTA) (see Note 7).
2.5. Analysis of the Eluate (see Note 8)
1. High-purity methanol. 2. High-purity chloroform. 3. Vacuum centrifuge (e.g., SpeedVac).
2.5.1. Methanol/ Chloroform Protein Precipitation 2.5.2. Preparative Gel and SYPRO Ruby Staining
1. NuPAGE (Novex, Invitrogen) 4–12% Bis-Tris–HCl, pH 6.4, polyacrylamide gels, including sample buffer (NuPAGE LDS sample buffer) and NuPAGE reducing agent (see Note 9). 2. Fix solution: 50% (v/v) ethanol, 7% (v/v) acetic acid. 3. SYPRO Ruby Protein Gel Stain (Invitrogen). 4. Wash solution: 10% (v/v) methanol, 7% (v/v) acetic acid. 5. Basic imaging system, such as the Bio-Rad Gel Doc with an SYPRO Ruby/Texas Red filter, 630BP30, 62 mm (170–8076). Alternatively, for better results use a laser-scanning instrument equipped with lasers that emit at 450, 473, 488, or 532 nm.
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1. Acetonitrile (ACN). 2. Reducing buffer: 10 mM DTT, 100 mM AmBic, pH 8 (see Note 10). 3. Alkylation buffer: 55 mM iodoacetamide (IAM) in 100 mM AmBic, pH 8. 4. 100, 50, and 20 mM NH4HCO3, pH 8 (AmBic). 5. Trypsin digestion buffer (TDB): 12.5 ng/mL trypsin (sequencegrade modified porcine trypsin; Promega, Madison, WI, USA) in 50 mM AmBic, pH 8. 6. 5% (v/v) formic acid, 50% (v/v) ACN.
3. Methods 3.1. Plant Culture
1. Sow Arabidopsis seeds on standard autoclaved soil and incubate for 48 h at 4°C in the dark (stratification); after that period, move the trays to a growth chamber with “short day” lighting conditions (8/16 h photoperiod at 120 mmol/m2/s, 21°C) for 3–4 weeks. Sow the seeds at high density, around 2 mm apart. 2. Alternatively, sow the seeds on ½ MS plates and grow the plants as described above.
3.2. Preparation of Home-Made HsIgG Sepharose Beads
1. Reconstitute 50 mg of HsIgG in 10 mL of coupling buffer; store at 4°C until initiating the coupling reaction. 2. Preparing the medium. Weigh out 3.75 g of CNBr-activated sepharose 4B powder and suspend in 100 mL of 1 mM HCl. 3. Incubate for 30 min at room temperature (RT). 4. Transfer the swollen sepharose to a sintered glass filter and wash under mild vacuum with 8 × 100 mL of 0.1 M HCl (the wash solution should be precooled to 4°C). 5. Resuspend the sepharose in a small volume of 0.1 M HCl (4°C) and transfer to a 50-mL conical tube (to make sure not to loose any sepharose beads, wash the sintered glass filter twice with 0.1 M HCl and transfer to the same tube). 6. Spin at 400 × g for 5 min at 4°C. 7. Carefully remove the supernatant with a pipette (do not decant). 8. Fill up to 50 mL with coupling buffer; mix briefly, and then spin at 400 × g for 5 min at 4°C. 9. Coupling the ligand. Add the 10 mL HsIgG in coupling buffer (from step 1) to the sepharose, mix briefly, and then spin at 400 × g for 5 min at 4°C. 10. To monitor the coupling efficiency, take a small sample of supernatant. Determination of the starting protein concentration can be done by measuring absorbance at 280 nm (OD280).
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11. Rotate the mixture end-over-end for 2 h at RT or overnight at 4°C (do not use magnetic stirrers as these can damage the sepharose beads). 12. Spin at 400 × g for 5 min at 4°C. 13. Carefully take off the supernatant and determine the OD280 value. Comparison of the protein content before and after the coupling reaction (OD280) allows one to estimate the coupling efficiency; typically, a tenfold decrease of the OD280 value indicates a good coupling efficiency. 14. Fill up the tube containing the sepharose to 50 mL with coupling buffer, mix briefly, and then spin at 400 × g for 5 min at 4°C. 15. Blocking of remaining active CNBr groups. Wash the sepharose twice with blocking buffer (add buffer, mix briefly, spin at 400 × g for 5 min, and then remove the supernatant). 16. Fill up the tube containing the sepharose to 50 mL with fresh blocking buffer. 17. Rotate the mixture end-over-end for 2 h at RT. 18. Spin at 400 × g for 5 min at 4°C. 19. Washing. Remove the supernatant and wash the sepharose with a total volume of 200 mL of NaCl coupling buffer (keep at 4°C). Afterward, carry out four additional washes as outlined in steps 20–23 below. 20. Wash with 100 mL of 0.1 M glycine–HCl, pH 2.8 (4°C). 21. Wash with 100 mL of 0.2 M glycine–HCl, pH 2.8 (4°C). 22. Wash with 200 mL of ultrapure water (4°C). 23. Wash with 200 mL of PBS buffer (4°C) (see Note 11). 24. Resuspend the sepharose in 20 mL of PBS and as a preservative add NaN3 to a final concentration of 0.01% (see Note 1). 25. Store at 4°C. 26. By rule of thumb, use 10 mL of HsIgG sepharose beads suspension per gram of initial fresh weight of plant tissue. 1. Wash the HsIgG-Sepharose Fast Flow beads twice with 10 volumes of 0.2 M sodium borate (see Note 12).
3.3. Alternative: Preparation of Commercial HsIgGSepharose Fast Flow Beads
2. Centrifuge for 5 min at 3,000 × g.
3.3.1. Cross-Linking of HsIgG-Sepharose Fast Flow Beads (14)
4. Mix for 30 min at RT on a shaker.
3. Resuspend the beads in 1 volume of 0.2 M sodium borate and add enough solid DMP to bring the final concentration to 20 mM. 5. Stop the reaction by briefly washing the beads twice in 10 volumes of 0.2 M ethanolamine; between each wash step,
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pellet the beads as described in step 2 above. Then, incubate the beads for 2 h at RT in 10 volumes of fresh 0.2 M ethanolamine on a shaker. 6. Wash the beads once briefly with 10 volumes of washing buffer, and then incubate them for 30 min in 10 volumes of fresh washing buffer at RT. 7. Wash the beads twice briefly in 10 volumes of PBS. 8. Wash the beads once briefly in 10 volumes of storage buffer (see Notes 1 and 13). 9. Pellet the beads as described in step 2 above, and then resuspend them for storage in 5 volumes of storage buffer (see Notes 14 and 15). Store the beads at 4°C for up to 12 months or until required. 3.3.2. Equilibration of HsIgG-Sepharose Fast Flow Beads (see Note 16)
1. Transfer 2 × 200 mL aliquots of the cross-linked HsIgG Sepharose Fast Flow beads (see Subheading 3.3.1, step 9) to two 1.5-mL Eppendorf tubes (see Note 17). 2. Centrifuge at 100 × g and 4°C for 1 min. 3. Remove the supernatants and discard them. 4. Wash the beads briefly and gently in 1 mL of ice-cold 0.5 M HAc, pH 3.4. After the wash, pellet the beads as described in step 2 above, discarding the supernatant. Then, similarly conduct eight additional washes as described in steps 5–9 below. 5. Wash with 1 mL of ice-cold TBS. 6. Wash with 1 mL of ice-cold 0.5 M HAc, pH 3.4. 7. Wash with 1 mL of ice-cold TBS. 8. Wash three times with 1 mL of ice-cold 0.1 M glycine–HCl, pH 3.0. 9. Wash twice with 1 mL of ice-cold TBS. 10. Resuspend the beads in 1 mL of ice-cold TBS, mix gently, then close the tubes and seal with Parafilm. Store at 4°C until use (see Note 18).
3.4. Affinity Purification of TAP-Tagged Proteins and Associated Complexes
1. Grind plant material with 3 mL of GB per gram of fresh tissue using an ice-cold mortar with pestle (see Note 19).
3.4.1. Protein Extraction (see Fig. 3)
4. Centrifuge the filtrate for 10 min at 1,500 × g at 4°C.
2. Incubate the homogenate on a turning-wheel mixer for 20 min at 4°C (for efficient solubilization). 3. Filter the homogenate through two layers of Miracloth. 5. Transfer the supernatant to cold ultracentrifuge tubes. 6. Centrifuge for 1 h at 100,000 × g at 4°C in an ultracentrifuge (see Note 19).
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7. Transfer the supernatant into new cold tubes (one tube per plant sample and phase; see Note 19). 8. Determine the protein concentration of the samples using the Bradford assay and calculate the total protein content of each sample (see Notes 20 and 21). The typical concentration range is between 1 and 5 mg/mL (i.e., a yield of 4–20 mg protein per gram of tissue). 9. Prepare new tubes with equal protein amounts of each sample for affinity purification in Subheading 3.4.2. Retain a small “Load” sample for later analysis (see Note 22). 3.4.2. First Affinity Purification
1. To the tube(s) containing aliquoted protein extract, add 10 mL of IgG sepharose beads per gram of initial fresh tissue (one may use either home-made beads from Subheading 3.2 or cross-linked commercial beads from Subheading 3.3). In our case, we found that home-made beads leaked less IgG and were therefore preferable. 2. Incubate for 2 h at 4°C on a turning-wheel mixer (see Note 23). 3. Centrifuge at 100 × g and 4°C for 1 min. 4. Transfer the supernatant to a new tube. Retain a small “Flowthrough” sample for later analysis (see Note 24). 5. Wash the beads twice with a volume of ice-cold GB equal to the initial volume of plant extract used in step 1 above, as follows. Add GB, mix gently, spin at 100 × g for 1 min, and remove the supernatant; then, add GB again, mix gently, spin at 100 × g for 1 min, and take off the supernatant. Finally, add GB again (see Note 25). 6. Transfer the beads with the GB into the spin columns. 7. Wash the beads five times with ice-cold GB. Each wash should employ 50 volumes of ice-cold GB (1 volume being the amount of beads used in step 1). For each wash, close both ends of the spin columns and invert them a few times; then, open both ends of the columns, place them in 2-mL Eppendorf collecting tubes, and centrifuge at 100 × g and 4°C for 1 min, to remove the GB buffer through the filter.
3.4.3. TEV Elution
1. For each 10 mL of beads (in an appropriate spin column, which should be closed at the base with the supplied plug and sealed with Parafilm on top), add 1.5 mL of 20× TEV buffer, 5 U of TEV protease, and complete to 30 mL with ultrapure H2O. 2. Close the spin column lid and then incubate overnight at 4°C or for 2 h at 16°C on a turning-wheel mixer. We do not normally remove the His-tagged TEV protease. However, it is possible to remove the TEV-protease from the sample using Ni–NTA Agarose; this can be done immediately after the overnight
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incubation with TEV protease, for an additional incubation period of 30 min under the same conditions. 3. Carefully open both ends of the spin column and place it into a new Eppendorf tube. 4. Spin at 100 × g and 4°C for 1 min. 5. Without removing the column, add 100 mL of cold 1× TEV buffer. 6. Immediately spin at 100 × g and 4°C for 1 min. 7. Collect the “Eluate” (see Note 26). 8. After the TEV elution, one may carry out an additional acidic elution to check efficiency of the TEV cleavage. This is done by repeating the elution step with 100 mL of AEB instead of TEV buffer. 3.4.4. Second Affinity Purification (see Note 27)
1. To the TEV eluate, add: 1 volume of CBB and 3 mL of 1 M CaCl2 per mL of TEV eluate. 2. Transfer this to a spin column containing 1 mL of Calmodulin– Agarose beads per 2 mL of IgG beads used in Subheading 3.4.2. 3. Seal the column and then incubate for 2 h at 4°C on a turningwheel mixer. 4. Spin the column in a collection tube with both ends open at 100 × g and 4°C for 1 min, to collect the “Calmodulin flowthrough” containing unbound proteins. 5. Close the bottom of the column and then add 500 mL of CBB to wash the beads. Close the top of the column and invert three times. 6. Spin the column in the same collecting tube with both ends open at 100 × g and 4°C for 1 min. 7. Close the bottom of the column and then add 300 mL of CWB to wash again the beads. Close the top of the column, and invert three times 8. Spin the column in a new collection tube with both ends open at 100 × g and 4°C for 1 min, to collect the “Calmodulin wash”. 9. Elute at 4°C with twice 100 mL of each concentration of CEB; for each elution solution, proceed essentially as in steps 3–5 above, but with an incubation time of only 3 min (see Note 28). These are the “Calmodulin eluate” samples.
3.5. Analysis of the Eluates 3.5.1. Methanol/ Chloroform Protein Precipitation (16) (see Note 29)
1. At room temperature, add 2.4 volumes of methanol and 0.8 volumes of chloroform to the protein samples to be precipitated for analysis. For example, one may wish to precipitate the “Load” (total protein extract), “Flow-through” (unbound protein), “Wash” (contaminants), and “Eluate” (isolated proteins of interest) samples (see Note 30).
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2. Vortex the sample briefly. 3. Add 3.2 volumes of ultrapure water. 4. Vortex again briefly. 5. Centrifuge at RT for 1 min at 13,000 × g. 6. Remove aqueous (upper) phase. Take care not to disturb the interphase containing the proteins. 7. Add the same volume of methanol as in the first step. 8. Vortex the sample briefly. 9. Centrifuge at RT for 5 min at 13,000 × g. 10. Remove the supernatant carefully using a pipette. 11. Dry the protein pellet (either air-dry or use a SpeedVac) and then resuspend in a volume of sample buffer corresponding to the capacity of the wells of the gel to be used (~20 mL) and taking into account loading equivalents of ~25 mg of protein. 3.5.2. Preparative Gel and SYPRO Ruby Staining
1. Load the samples on a Bis-Tris–HCl buffered (pH 6.4) 4–12% polyacrylamide gel. Load an equivalent of 25 mg of protein for “Load,” “Flow through,” and IgG beads “Wash” steps; load the entire volume of the other samples (including the TEV “Eluate,” and the “Calmodulin flow-through,” “Calmodulin wash,” and “Calmodulin eluate” samples). 2. Run the gel for 40 min at 20 V/cm. 3. Incubate the gel at RT for 30 min to overnight in 20 volumes of fix solution, preferably in darkness and without shaking. 4. Repeat step 3 once for 10 min using fresh fix solution. 5. Wash the gel three times at RT for 10 min with ultrapure water under gentle shaking. 6. Incubate the gel in SYPRO Ruby protein gel stain (the amount of stain used should be 10 times the gel volume) in a closed clean plastic container, in a shaking water bath at 80°C for 30 min (see Note 31). 7. Wash the gel with wash solution (10 times the gel volume) at RT for 30 min in a clean container (see Note 32). 8. Wash the gel for 5 min twice in ultrapure water (see Note 33). 9. Visualize and photograph the gel (see Note 34).
3.5.3. In-Gel Digestion
1. Excise the band of interest from the stained gel, and cut it into 1-mm cubes using a clean razor blade on a clean glass surface. Transfer the pieces to an Eppendorf tube. Remove all remaining liquid with a pipette. 2. Add 25–35 mL of ACN to cover the gel pieces, and incubate for 10 min at RT to dehydrate and shrink the gel pieces.
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3. Remove the ACN with a pipette and then SpeedVac to dryness for 10 min. 4. Swell the gel particles by adding 150 mL of reducing buffer and incubating for 1 h at 56°C. 5. Cool the sample to RT. Replace the reducing buffer with 150 mL of alkylation buffer and then incubate for 45 min at RT in the dark with occasional vortexing. 6. Remove the solution and then wash the gel pieces with 150 mL of 100 mM AmBic. Incubate for 10 min at RT without shaking. 7. Remove the AmBic solution with a pipette and add 150 mL of ACN to dehydrate the gel pieces. Incubate for 10 min at RT. 8. Repeat the washing step (steps 6 and 7) once, then remove the ACN and SpeedVac to dryness for 10 min. 9. Place the tubes in an ice water bath and swell the gel particles by adding 25–35 mL of TDB and incubating for 45 min. 10. Remove the excess of TDB and cover the gel pieces with 50 mM AmBic to keep them wet during cleavage. Incubate overnight at 37°C. 11. Centrifuge briefly at 13,000 × g in a microfuge to spin down the gel pieces. Transfer the supernatant into a collection tube (a new 200-mL Eppendorf tube). 12. Add 20 mL of 20 mM AmBic to cover the gel pieces. Incubate for 10 min at RT. Transfer the supernatant to the collection tube used in step 11. 13. Add 25 mL of 5% formic acid, 50% ACN to the gel pieces and incubate for 20 min at RT. 14. Centrifuge briefly at 13,000 × g and then transfer the formic acid/ ACN solution to the same PCR collecting tube used in step 11. 15. Repeat the peptide extraction (steps 13 and 14) twice more (i.e., these steps should be done a total of three times), combining all of the solution into the same tube. 16. Dry the combined sample in a SpeedVac to complete dryness. Store at −20°C until analysis can commence or proceed to mass spectrometry analysis.
4. Notes 1. Sodium azide is highly toxic; handle with precaution. 2. DTT present in the standard TEV protease buffer could break the disulfide bonds linking IgGs onto the beads; therefore, cross-linking of the IgG is proposed to avoid IgG leakage during the cleavage step. Alternatively, an elution buffer either
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without or with very low concentrations of DTT could be used for TEV cleavage. 3. Add 286 mL of acetic acid to 10 mL of ultrapure H2O (this gives a pH of 2.6); then, adjust to pH 3.4 by adding 0.5 M ammonium acetate (NH4Ac). 4. All buffers have to be refrigerated and used at 4°C. 5. Detergent and detergent concentration for solubilization of membrane proteins should be chosen with regard to the TAPtagged protein to be purified. 6. Prepare GB freshly as PMSF is not stable in aqueous solutions. 7. Composition of the CEB has to be adjusted for each complex analysis. The concentrations of KAc and EGTA may range, respectively, between 100 and 200 mM, and between 5 and 20 mM (e.g., see Note 28). 8. Always use ultrapure water (Milli-Q, 18 MW cm) to avoid sample contamination. 9. Self-poured SDS–PAGE gels are suitable, but gradient gels should be preferred as they provide better separation of proteins over a wider range of molecular masses. Additionally, precast gels offer excellent reproducibility, and therefore allow better comparison between experiments. 10. Reducing buffer is used to avoid oxidation of disulfides bridges. 11. Washing could be done by centrifugation or under mild vacuum in a sintered glass funnel; the pH of the last washing supernatant should be controlled and should be the same as the pH of the PBS buffer used for washing. 12. Usually the volume of beads to be washed is 200 mL, allowing one to work in 2-mL Eppendorf tubes. 13. Steps 8 and 9 (i.e., addition of sodium azide) can be avoided if the beads are to be used immediately. 14. A sample of the beads (non-cross-linked and cross-linked) should be denatured in SDS–PAGE sample buffer (containing DTT) and analyzed on an SDS–PAGE gel for comparison. Ideally, if cross-linking was successful, no IgG subunits should be observed by SDS–PAGE, showing that the DTT was without effect on the IgG, and that the TEV digestion requiring DTT can be conducted safely. 15. Sodium azide is a powerful biocide preventing bacterial contamination in the prepared beads. 16. Each washing step consists of the removal of the liquid phase, the addition of the washing solution, followed by centrifugation at 100 × g at 4°C for 1 min.
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17. Use a cut pipette tip to avoid the loss of beads at the edge of the tip. 18. Check the pH of the last supernatant with pH paper. It should be neutral; if it is not, repeat the TBS washing until the pH is neutral. 19. If the TAP-tagged protein of interest is known to be associated with membranes, it is possible to perform the first steps before the first ultracentrifugation using grinding buffer that lacks any added detergent (e.g., Triton X-100) (see Fig. 4). The membrane pellet obtained by ultracentrifugation can then be homogenized in grinding buffer with an appropriate nonionic detergent for membrane protein solubilization (e.g., Triton X-100 or another, such as digitonin or dodecylmaltoside). A second ultracentrifugation step is required to remove the insolubilized material (see Fig. 4). 20. Some detergents are not compatible with the Bradford assay. In this case, another method to determine the protein concentration (e.g., bicinchoninic acid [BCA] protein assay) has to be used. 21. Alternatively, sample volumes can be adjusted to chlorophyll concentration. Extraction is done by adding 1 mL of 80% (v/v) acetone to 5 mL of the sample. Total chlorophyll is determined by measuring absorbance at 652 nm, and its concentration is calculated according to Arnon (17). 22. Take a sample equivalent to 5% of the total volume for each tube and precipitate the protein by the methanol/chloroform method (see Subheading 3.5.1). This is the “Load” sample that will be used for Western blot analysis to provide an indication of the success of the protein extraction. 23. Overnight incubation should be possible but needs to be tested in each case. 24. Take a sample equivalent to 5% of the total volume for each tube and precipitate the proteins by methanol/chloroform method (see Subheading 3.5.1). This is the “Flow-through” sample that will be used for Western blot analysis to provide an indication of the yield of the binding phase. 25. It may be helpful to retain the supernatants as IgG beads “Wash” samples for later analysis by Western blotting. 26. Cross-linking of IgGs is normally sufficient to avoid significant IgG leaching. Nonetheless, one may consider using protein A sepharose (e.g., CL-4B, GE Healthcare) to remove any leached IgGs from the TEV eluate after this step. 27. This second affinity purification is intended to increase the purity of the isolated complexes; in many cases, MS analysis may be carried out with satisfactory results after TEV elution.
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28. Elution can be done with CEB containing increasing concentrations of KAc and EGTA. For example, first 100 mM KAc, 5 mM EGTA, then 150 mM KAc, 10 mM EGTA, and finally 200 mM KAc, 20 mM EGTA. 29. Protein precipitation is required when the sample volume is too large to be directly loaded on a gel. Additionally, an aliquot of 5% for each sample can be precipitated independently for Western blot analysis. For precipitation, first complete the aliquot to 200 mL with ultrapure water. 30. Take care of the size of the tubes used for precipitation; volumes up to a maximum of 250 mL can be precipitated in 2-mL Eppendorf tubes. 31. The incubation can be done overnight at RT, resulting in a higher fluorescence signal, but also a higher background. 32. Transferring the gel into a new container avoids heating the methanol-containing wash solution. 33. This washing step is included to protect the imaging device. 34. The SYPRO Ruby stained gel can be visualized under UV or blue light. SYPRO Ruby dye has two excitation maxima at 280 and 450 nm, for a fluorescence emission at 610 nm. SYPRO Ruby stained bands are conveniently visualized and excised using a gel documentation system, such as a Bio-Rad Gel Doc with a SYPRO Ruby/Texas Red filter. This instrument allows one to take a high-quality picture of the gel, and with the UV transilluminator in “preparation” mode, outside the instrument, the bands can be excised precisely. For optimum detection and resolution, use a laser-scanning instrument for the documentation of the gel. References 1. Rigaut, G., Shevchenko, A., Rutz, B., Wilm, M., Mann, M., and Seraphin, B. (1999) A generic protein purification method for protein complex characterization and proteome exploration. Nat. Biotech. 17, 1030–1032. 2. Schimanski, B., Nguyen, T. N., and Gunzl, A. (2005) Highly efficient tandem affinity purification of trypanosome protein complexes based on a novel epitope combination. Eukaryotic Cell 4, 1942–1950. 3. Rubio, V., Shen, Y., Saijo, Y., Liu, Y., Gusmaroli, G., Dinesh-Kumar, S. P., and Deng, X. W. (2005) An alternative tandem affinity purification strategy applied to Arabidopsis protein complex isolation. Plant J. 41, 767–778. 4. Gloeckner, C., Boldt, K., and Ueffing, M. (2009) Strep/FLAG tandem affinity purification (SF-TAP) to study protein interactions. Current Protoc. Protein Sci. 57, 19.20.1-19.20.19.
5. Lehmann, R., Meyer, J., Schuemann, M., Krause, E., and Freund, C. (2009) A novel S3S-TAP-tag for the isolation of T-cell interaction partners of adhesion and degranulation promoting adaptor protein. Proteomics 9, 5288–5295. 6. Burckstummer, T., Bennett, K. L., Preradovic, A., Schutze, G., Hantschel, O., Superti-Furga, G., and Bauch, A. (2006) An efficient tandem affinity purification procedure for interaction proteomics in mammalian cells. Nat. Methods 3, 1013–1019. 7. Tagwerker, C., Flick, K., Cui, M., Guerrero, C., Dou, Y., Auer, B., Baldi, P., Huang, L., and Kaiser, P. (2006) A tandem affinity tag for twostep purification under fully denaturing conditions. Mol. Cell. Proteomics 5, 737–748. 8. Tagwerker, C., Zhang, H., Wang, X., Larsen, L. S. Z., Lathrop, R. H., Hatfield, G. W., Auer,
3 Preparation of Multiprotein Complexes from Arabidopsis… B., Huang, L., and Kaiser, P. (2006) HB tag modules for PCR-based gene tagging and tandem affinity purification in Saccharomyces cerevisiae. Yeast 23, 623–632. 9. Graumann, J., Dunipace, L. A., Seol, J. H., McDonald, W. H., Yates, J. R., Wold, B. J., and Deshaies, R. J. (2004) Applicability of tandem affinity purification MudPIT to pathway proteomics in yeast. Mol. Cell. Proteomics 3, 226–237. 10 Xu, X., Song, Y., Li, Y., Chang, J., Zhang, H., and An, L. (2010) The tandem affinity purification method: An efficient system for protein complex purification and protein interaction identification. Protein Expr. Purif. 72, 149–156. 11. Rohila, J. S., Chen, M., Cerny, R., and Fromm, M. E. (2004) Improved tandem affinity purification tag and methods for isolation of protein heterocomplexes from plants. Plant J. 38, 172–181. 12. Rohila, J. S., Chen, M., Chen, S., Chen, J., Cerny, R. L., Dardick, C., Canlas, P., Fujii, H., Gribskov, M., Kanrar, S., Knoflicek, L.,
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Stevenson, B., Xie, M., Xu, X., Zheng, X., Zhu, J.-K., Ronald, P., and Fromm, M. E. (2009) Protein-protein interactions of tandem affinity purified protein kinases from rice. PLoS ONE 4, e6685. 13. Li, Y. (2010) Commonly used tag combinations for tandem affinity purification. Biotechnol. Appl. Biochem. 55, 73–83. 14. Harlow, E., and Lane, D. (1998) Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA, pp. 323–325. 15. Shevchenko, A., Tomas, H., Havlis, J., Olsen, J. V., and Mann, M. (2006) In-gel digestion for mass spectrometric characterization of proteins and proteomes. Nat. Protoc. 1, 2856–2860. 16. Wessel, D., and Flügge, U. I. (1984) A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal. Biochem. 138, 141–143. 17. Arnon D.I. (1949) Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta vulgaris. Plant Physiol. 24, 1–15.
Chapter 4 Studying Interactions Between Chloroplast Proteins in Intact Plant Cells Using Bimolecular Fluorescence Complementation and Förster Resonance Energy Transfer Jodi Maple and Simon G. Møller Abstract Protein–protein interactions play crucial roles in the execution of many cellular functions, including those in plastids. Identifying and characterising protein–protein interactions can yield valuable information regarding the function of a protein and can also contribute towards understanding protein–protein interaction networks in plastids, thereby contributing to a better understanding of cellular processes. Here, we describe the planning and experimental procedures required to perform both bimolecular fluorescence complementation and Förster resonance energy transfer assays to detect protein–protein interactions. Arabidopsis is well-suited for microscopy and its small size facilitates live cell imaging, enabling observation of protein–protein interactions in living chloroplasts. The methods described in this chapter can be used to analyse protein–protein interactions of two known proteins and to dissect interacting protein domains. Key words: FRET, BiFC, Protein–protein interaction, Yellow fluorescent protein, Cyan fluorescent protein
1. Introduction Biological processes are executed by proteins that, to a large extent, depend on interactions with other proteins for their activity. In chloroplasts, protein–protein interactions are essential for almost all processes, from the formation of the protein import machinery, which permits the import of nuclear-encoded proteins into plastids, to the division machinery, which enables the organelle to divide, to the formation of the photosynthetic complexes, which enables the conversion of solar energy into sugar, to mention but a few (1).
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_4, © Springer Science+Business Media, LLC 2011
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Protein–protein interactions can be classified into different types depending on their specificity (proteins may interact with a specific partner or with an entire class of proteins), strength (proteins may interact transiently, allowing one protein to be modified, or may form part of a more stable protein complex), and composition (proteins may interact to form homo- and hetero-oligomers). The consequences of such interactions can vary greatly and identifying and characterising protein–protein interactions in plants can lead to a greater understanding of the mechanisms of biological processes and define networks of interacting proteins within the organelle. Many powerful technologies exist to identify and characterise individual protein–protein interactions (see Chapter 5, Vol. 2). Well-established systems include co-immunoprecipitation (co-IP), the yeast-two hybrid system and newer approaches, such as protein chip arrays (2). Additionally, computational methods have been developed to predict protein–protein interactions (3). However, each of these approaches has disadvantages. For example, the preparation of cell extracts for co-IP experiments may disrupt the physical conditions under which a given pair of proteins interacts. Similarly, heterologously expressed plastid proteins in yeast, as part of yeast two-hybrid assays, may not reflect the in planta situation. Indeed, some protein–protein interactions are dependent on their native subcellular compartment, on correct folding, and/or on modifications within the plastid. Furthermore, the identification of protein– protein interactions in heterologous systems or in silico requires additional verification within the environment of the plastid. Finally, the suborganellar localisation of an observed protein– protein interaction cannot be deduced using the above-mentioned systems. Bimolecular fluorescence complementation (BiFC) and Förster resonance energy transfer (FRET) assays are both fluorophorebased assays that can be used to identify, validate, and characterise protein–protein interactions in living cells (4–9). Both techniques have two main advantages (1) Protein–protein interactions are observed and analysed within their correct cellular environment under physiological conditions and (2) the correct subcellular localisation of the protein–protein interaction can be visualised. Both FRET and BiFC require that the proteins are in close proximity and provide strong evidence for the close association of (although not necessarily the direct interaction between) two proteins of interest, making them valuable techniques for the study of protein– protein interactions in chloroplasts in vivo (10–12). BiFC assays are based on the reconstitution of a fluorescent complex when two non-fluorescent fragments of a fluorescent protein are brought together by an interaction between two proteins fused to these fragments (Fig. 1a) (4). BiFC assays have been reported for protein partners up to 7 nm apart (13). BiFC assays are suitable to detect weakly associated and short-lived interactions
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Fig. 1. Schematic of BiFC and FRET assays. (a) BiFC assay: Protein A and protein B are fused to the N-terminal residues (N; amino acid 1–154) and C-terminal residues (C; amino acid 155–238) of YFP. Interaction of A and B will bring N and C into close proximity, allowing YFP to reconstitute into its native structure and emit a fluorescent signal. (b) FRET assay: Protein A and protein B are fused to full-length CFP (donor fluorophore) and YFP (acceptor fluorophore). Interaction of A and B will bring CFP and YFP into close proximity and FRET will occur. FRET manifests itself both by quenching of donor fluorescence and by an increase in acceptor fluorescence emission.
since the complementation of two fluorescent fragments in BiFC is stable (14). Additionally, only standard fluorescence microscope equipment is required, making BiFC a popular technique. However, BiFC assays are not suitable for monitoring interactions in real time, and the assay is not reversible. FRET is based on the non-radiative transfer of energy from an excited fluorophore (donor molecule) to a second fluorophore (acceptor molecule) (Fig. 1b). For this energy transfer to take place, the molecules must be in close proximity (less than the Förster distance of 5–6 nm) (15). When FRET occurs, there is a decrease in the fluorescence intensity of the donor (donor quenching) and an increase in the fluorescence intensity of the acceptor (sensitised emission) (Fig. 1b). With digital imaging techniques, it is possible to quantify FRET by measuring the changes in the donor/ acceptor fluorescence (5). The most common FRET fluorophore
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pair used in chloroplast biology is cyan fluorescent protein (CFP; donor molecule) and yellow fluorescent proteins (YFP; acceptor molecule), which offer benefits over other fluorescent protein combinations in that there is no significant crosstalk (bleed through) with autofluorescence (16). FRET can be used to quantify the distance between two molecules and also enables the detection of protein–protein interactions in real time. However, FRET requires specialised filter sets and data processing, making it technically more challenging than BiFC. Here, we present an overview of both techniques and provide experimental protocols for expressing fusion proteins in plants systems, as well as for BiFC and FRET data acquisition. Despite the focus on Arabidopsis and tobacco chloroplasts, these techniques are easily adaptable to other plant systems, making them very versatile.
2. Materials 2.1. Vector Construction for BiFC and FRET Experiments
1. Plant compatible BiFC and FRET vectors: pWEN18, pWEN15, pWEN-NY, pWEN-CY, pBA002 (10, 17). 2. Antibiotics suitable for the bacterial strains and plasmids used at the following concentrations: ampicillin (50 mg/mL), spectinomycin (50 mg/mL), kanamycin (50 mg/mL). 3. LB media: 10 g/L bacto-tryptone, 10 g/L NaCl, 5 g/L yeast extract; pH adjusted to 7.5 with NaOH. 4. Plasmid Midiprep kit to isolate plasmid DNA from recombinant Escherichia coli cultures.
2.2. Co-transformation and Co-expression of Fusion Proteins in Leaf Cells
1. Arabidopsis thaliana or Nicotiana tabacum: Arabidopsis rosette leaves of 2–4 week-old non-bolting plants grown at 22°C in a 16 h light/8 h dark cycle. Tobacco leaves of 4–8 week old plants grown at 24°C in a 16 h light/8 h dark cycle. 2. Murashige–Skoog (MS) media: 4.4 g/L MS salts, 1% (w/v) sucrose, 0.8% (w/v) plant agar; pH adjusted to 5.8 with NaOH. 3. Agrobacterium tumefaciens strain suitable for transformation, such as GV3101. 4. Infiltration medium: Dissolve 4.88 g 2-(N-morpholino)ethanesulfonic acid (MES) in 400 mL of H2O, then adjust pH to 5.6 using NaOH. Subsequently, add 2.5 g glucose, 0.123 g Na2HPO4, 0.156 g NaH2PO4, and adjust the final volume to 475 mL. Autoclave and then add 25 mL of AB salts: 373 mM NH4Cl, 24 mM MgSO4, 40 mM KCl, 1.36 mM CaCl2, 0.18 mM FeSO4.
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5. 50 mM acetosyringone. Dissolve powder in 2–3 mL of 70% (v/v) ethanol and make up to final concentration in H2O. 6. Sterile single-use syringes (1 mL). 7. PDS-1000/He™ helium biolistic particle delivery system and bombardment consumables: macrocarrier holders, rupture discs 1,100 psi, stopping screens, macrocarriers, 0.6 mm gold (Bio-Rad Laboratories, USA). 8. Microcentrifuge. 9. 2.5 M CaCl2. Filter sterilise (filter, 0.22 mm) and then store at 4°C for no more than 2 weeks. 10. 0.1 M free-base spermidine. To dissolve the spermidine, heat the vial at 37°C for 5 min. Aliquot 15.8 mL of spermidine into 1.5-mL Eppendorf tubes and store at −70°C. Before use, thaw one vial and add 984.2 mL of sterile H2O. Mix and aliquot the 0.1 M solution into 0.5-mL Eppendorf tubes. Store 0.1 M stocks at −20°C. 11. Cold 100% ethanol. 12. 70% (v/v) ethanol. 13. Nescofilm. 2.3. Microscopy
1. Epifluorescence microscope equipped with an appropriate camera, for example the Nikon DS-Qi1 with DS-L2 Controller (Nikon, Japan). 2. Objective lenses: A water objective is optimal since the high water content in plant cells means that this lens is matched to the refractive index of the sample, for example a 60× magnification CFI Plan Apo VC60WI (Nikon). 3. Appropriate filter sets, for example: CFP (exciter S436/10, emitter S470/30) and YFP (exciter HQ500/20, emitter S535/30) (Chroma Technologies, USA). 4. Image acquisition software, for example OpenLab (Improvision, UK) (see Note 1). 5. Microscope coverslips (22 × 40 × 0.1 mm) and microscope slides (75 × 25 mm).
3. Methods The methods outlined below describe (1) the design of BiFC and FRET experiments and construction of suitable vectors, (2) the transient expression of fusion proteins in Arabidopsis or tobacco leaf cells, and (3) data acquisition in BiFC and FRET assays.
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Fig. 2. Plant compatible vectors required for FRET and BiFC assays. Genes of interest can be cloned into the multiple cloning site (XhoI and KpnI) to generate in-frame fusions to the N termini of the fluorophores. The fusions are driven by the cauliflower mosaic virus 35S promoter (35S). The presence of restriction enzyme recognition sites downstream of the fluorophores allows easy subcloning of the fusion cassettes to binary vectors. G10; ten glycine linker sequence. Numbers in subscript indicate amino acid residues.
3.1. Planning and Vectors Construction for BiFC and FRET Experiments
Outlined below is the rhetoric behind the vectors required for BiFC and FRET assays and the necessary controls for each assay. The first step in both BiFC and FRET assays is to generate fusions of the proteins predicted to interact (for example, protein A and protein B) to the N-terminal of the full length or truncated fluorophores, and many different vectors are available (see Note 2). In this laboratory, vectors are adapted from Kost et al. (17) and are expressed under the control of cauliflower mosaic virus 35S promoter (Fig. 2; see Note 3). The principle for cloning genes of interest as fusions to YFP and CFP genes follows standard cloning protocols for the insertion of genes of interest, whereby full-length or fragments of cDNAs are cloned through restriction and ligation. The correct orientation, reading frame, and sequence must be confirmed by DNA sequencing.
3.1.1. BiFC Vectors
In BiFC, the genes of interest are cloned into the pWEN-NY and pWEN-CY (Fig. 2) vectors to generate translational fusions between the proteins that are predicted to interact and the N-terminal residues (amino acid 1–154) and C-terminal residues (amino acid 155–238) of the YFP protein, respectively (10). When the two fusions are expressed in living cells, interaction of the candidate proteins will bring the two halves of YFP into close proximity, allowing YFP to reconstitute into its native structure and emit
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its fluorescent signal (Fig. 1a) (4). It is recommended to perform the BiFC assays with the transgenes as fusions to both the N- and C-terminal fragments of YFP (for example, to express the pWENNY/geneA + pWEN-CY/geneB and pWEN-NY/geneB + pWENCY/geneA) (see Note 4). If the assay is designed to test for homo-dimerisation, then the gene of interest must be cloned into both vectors. 3.1.2. FRET Vectors
FRET requires that proteins that are postulated to interact are fused independently to CFP and YFP and co-expressed in living cells. This is achieved by cloning the genes of interest into pWEN15 and pWEN18, to generate fusions to the N-terminal of CFP and YFP, respectively (17). Interaction of the candidate proteins brings the fluorophores into close spatial proximity and excitation energy can be non-radiatively transferred from the donor to the acceptor fluorophore (Fig. 1b). It is recommended that the FRET assay is carried out using both proteins as a donor, fused to CFP (for example, to co-express the pWEN18/geneA + pWEN15/geneB and pWEN18/ geneB + pWEN15/geneA) (see Note 4). If the assay is designed to analyse homo-dimerisation, then the gene of interest must be cloned into both the CFP- and the YFP-containing vectors.
3.1.3. Design of Controls for BiFC and FRET Assays
It is necessary to include both positive and negative controls in any BiFC and FRET experiment. For both assays, the positive controls can take the form of known interacting proteins, and this is useful to validate the technique in the laboratory and ensure that the overall microscope parameters are correct (see Note 5). There are several possibilities for negative controls and they require careful consideration: in both BiFC and FRET assays, the result is not absolute and must be compared to suitable negative control(s) to give meaningful results. The recommended negative control is designed by mutating/deleting the site of interaction between the two candidate proteins (Fig. 3b). Providing that the mutation does not affect the localisation or expression level of the fusion protein, this serves as the most appropriate negative control. It is also possible to select a negative control protein that localises to the same subcellular localisation, but is known not to interact with the candidate protein (Fig. 3c), or to use the fluorophores/fluorophore fragments in isolation (Fig. 3d).
3.2. Co-transformation and Co-expression of Fusion Proteins in Leaf Cells
Described below are two protocols for transient co-expression of the fusion proteins in Arabidopsis or tobacco leaf chloroplasts (see Note 6). In addition, protoplast transfection and the generation of stable transgenic tobacco or Arabidopsis lines are suitable approaches (see Note 7). Before embarking on a BiFC or a FRET experiment, it is advisable to list all combinations of vectors to be co-transformed into the Arabidopsis or tobacco so that the appropriate amount of leaves can be prepared (see Note 8).
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Fig. 3. Typical experimental design to analyse the interaction between protein A and protein B. (a) Samples co-expressing protein A and protein B fused to the N- and C-terminal halves of YFP are compared to (b) samples co-expressing either a mutated form of protein B, in which the site of interaction has been mutated/deleted, or (c) an endogenous protein known not to interact with protein A (protein X), or (d) the fluorophore fragment alone (in this case, the fluorophore fragment must be targeted to the same compartment of the cell as protein A). In principle, the same control proteins can be used for both BiFC and FRET assays. 3.2.1. Bombardment of Leaf Cells
Biolistic transformation (particle bombardment) allows direct gene transfer and transient expression in a broad range of tissues. This technique involves accelerating DNA-coated gold particles (the microcarriers) directly into intact tissues. 1. Before preparing the gold stocks, the consumables for shooting should be sterilised. Wash the rupture discs, stopping screens, and macrocarriers (five of each per shooting) briefly in 100% ethanol to remove dust and leave them to dry thoroughly. Then, place the macrocarriers in the macrocarrier holders. Place the leaves on MS medium in Petri dishes and label each plate with the constructs to be bombarded. 2. Vortex the microcarrier stock (60 mg/mL in 50% [v/v] glycerol) (see Note 9) thoroughly for 5 min to resuspend and disrupt agglomerated particles and then immediately transfer 20 mL (for each vector(s) to be bombarded) to a labelled, pre-chilled 1.5-mL centrifuge tube. 3. Sequentially add 12 mL of plasmid DNA (1 mg/mL), 10 mL of 2.5 M CaCl2, and 8 mL of 0.1 M spermidine to the microcarriers and mix by pipetting up and down 20 times between each addition. The two plasmids to be co-bombarded should be added in a 1:1 ratio with a total volume of 12 mL.
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4. Vortex the mixture for 3 min. The mixture can then be incubated on ice for up to 1 h. 5. To wash, pellet the microcarrier/DNA complexes by pulse centrifugation for 2 s at maximum speed in a microcentrifuge. Carefully remove the supernatant without disturbing the pellet, add 100 mL of 70% ethanol and invert the tube gently ten times. Repeat this wash step with 100 mL of 100% ethanol. Finally, resuspend the particles in 20 mL of 100% ethanol. Use immediately. 6. Spread 4 mL of particles (microcarrier/DNA complexes) over the central 1 cm of the macrocarrier, air-dry and use immediately for biolistic transformation. 7. The biolistic transformation is carried out using a Bio-Rad PDS-1000/He™ Helium biolistic particle delivery system, in which a burst of helium gas accelerates the microcarriers into the sample (see Note 10). The Petri dish containing the Arabidopsis or tobacco leaf is placed on target shelf 3, 6 cm from the macrocarrier holder, and the transformations are performed using 1,100 psi rupture discs under 25 in Hg vacuum. 8. After bombardment, seal the Petri dishes with Nescofilm, wrap in foil, and incubate at 24°C for 16–48 h before image acquisition (see Note 11). 3.2.2. Infiltration of Leaf Cells
The Agrobacterium infiltration method is a modified version of the protocol described by Yang et al. (18) and requires that the expression cassettes are transferred to a binary vector, for example pBA002 (17). For infiltration, the leaves can be attached to the plant or can be prepared in Petri dishes as for the bombardment approach (see Subheading 3.2.1). 1. Agrobacterium is individually transformed with each of the binary vectors (see Note 12). 2. The day prior to infiltration initiate a 5 mL starter cultures of Agrobacterium from bacteria growing on LB-agar plates. Grow the cultures in LB containing spectinomycin (50 mg/mL) at 28–30°C in an orbital shaking incubator at 250 rpm. 3. The following morning pellet 1 mL of the overnight culture in a microcentrifuge and resuspended in 10 mL of infiltration medium. Grow at 28–30°C in an orbital shaking incubator at 250 rpm for 5–8 h. 4. Adjust the bacterial densities to an OD600 of 0.2 prior to infiltration (see Note 13). 1 mL of bacterial culture is sufficient to infiltrate four segments of leaf. For co-infiltration of two constructs, Agrobacterium containing each plasmid should be grown separately and the two strains mixed in a 1:1 ratio after
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the OD600 of each culture has been adjusted to 0.2. Immediately prior to infiltration add 50 mL of 50 mM acetosyringone per 1 mL of culture. 5. Inject the leaves using a plastic 1-mL syringe without a needle on the abaxial side of the leaf (see Note 14). Keep the infected plants under standard greenhouse conditions for 16–48 h before image acquisition (see Note 11). 3.3. Analysis of BiFC and FRET Assays
Below, we outline the procedures for the preparation of the leaf tissue for microscopy and the visualisation and analysis of both FRET and BiFC experiments. A standard fluorescence or confocal microscope can be used. The intention of this chapter is not to give a detailed introduction to fluorescent microscopy. It is, however, important for the user to understand the principles of image acquisition and the pitfalls involved. It is critical to keep imaging times to a minimum since photo-oxidative damage may occur. This is especially important when imaging CFP since the appropriate excitation light causes an increase in the levels of autofluorescence.
3.3.1. Preparation of Leaf Tissue for Microscopy
1. Cut a small section (approximately 1–2 cm2) of the leaf tissue from either the infiltrated or bombarded plant. 2. Place the leaf on the microscope slide abaxial side up with a drop of sterile water. Cover with a coverslip and gently apply pressure over the surface of the leaf to remove air bubbles from the surface. It may be necessary to add more water to the slide. To ensure that the coverslip stays in place, tape the short edges of the coverslip to the slide.
3.3.2. BiFC Assays
The negative controls should be analysed to determine the levels of background signal using a suitable YFP filter set. Subsequently, BiFC samples are analysed for the presence of a YFP signal. If a fluorescent signal is detected in samples co-expressing protein A and protein B fused to the two halves of YFP, which is significantly higher than the control proteins expressed at the same level, then this is a strong indication that the two proteins interact. If there is no difference between the intensity of the signals seen between the samples and the negative control, then any signal observed is likely due to the non-specific interactions between the fusion proteins. It is conceivable that the non-fluorescent protein fragments are able to form fluorescent complexes with low efficiency, even in the absence of a specific interaction. It is important to note that if no signal is seen, this is not proof that the proteins of interest do not interact (see Note 15). Examine all samples and capture images from both the negative and positive controls. The exposure settings must be the same when acquiring images for the negative and positive controls.
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Various methods exist to measure FRET from the changes in donor and acceptor emission. A convenient and standard way to measure FRET is to quantify the emission of the acceptor upon donor excitation (acceptor-sensitised emission). This method is also referred to as the 3-image FRET technique since in addition to the measurement of acceptor emission, one must also account for bleedthrough caused by both CFP and YFP into the FRET filter set (that is the emission from the donor into the acceptor channel, and also for the direct excitation of the acceptor) (5, 19). The bleed-through values are normalised fractions between 0.0 and 1.0 and are essentially the fractional “bleed” of donor and acceptor into the FRET channel. These values are calculated once and used to correct the FRET channel pixel intensities in all subsequent FRET experiments carried out under the same experimental conditions. It is critical that the same settings and exposure times are maintained for each filter set during image acquisition of all samples. It is convenient to use a macro that allows the automated acquisition of images (see Note 16). Described below are the steps required to measure acceptor emission using the “Openlab FRET module” (Improvision). This FRET module captures three images for each data point: a donor image (excite with CFP filter, CFP dichroic, emit with CFP filter), an acceptor image (excite with YFP filter, YFP dichroic, emit with YFP filter), and a FRET image (excite with CFP filter, CFP dichroic, emit with YFP filter). 1. First, acquire images with all three filter sets of non-transformed cells to measure background levels. 2. Calculate the donor bleed-through constant by producing two images (a FRET image and a donor image) from a control sample transformed with the donor only. In this scenario, there is a real CFP signal but no YFP. The following calculation is performed: Donor bleed-through = (intensity of cell in FRET image)/ (intensity of cell in donor image). 3. Calculate the acceptor bleed-through constant by producing two images (a FRET image and an acceptor image) from a control sample transformed with the acceptor only. In this scenario, there is a real YFP signal but no CFP. The following calculation is performed: Acceptor bleed-through = (intensity of cell in FRET image)/ (intensity of cell in acceptor image). 4. Next, perform the FRET experiments on samples co-expressing the CFP and YFP vectors (see Note 17). The FRET module captures three sets of images: the donor, FRET, and acceptor images. These images are background subtracted and then, based on the intensity of the images in the donor and acceptor
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channels, and on the two bleed-through constants, the level of fluorescence expected in the FRET channel as a result of bleedthrough is calculated: Total bleed-through = (donor bleed-through constant × intensity of donor in donor channel) + (acceptor bleed-through constant × intensity of acceptor in acceptor channel). FRET is then measured as the relative increase in fluorescence in the FRET channel over the expected background from the bleed-through: FRET = (signal in FRET channel)/(total bleed-through). This correction method is performed by the software on a pixelby-pixel basis for each set of three images. The FRET signal generated can be false coloured using the pallet in Openlab (Improvision). If no FRET signal is observed, this is not empirical proof that the proteins do not interact (see Note 15).
4. Notes 1. The minimal requirements of image acquisition software are to operate the shutters to allow fast and accurate acquisition times. 2. Fusion proteins must be generated using the N terminus of the fluorescent protein(s). This is important so as not to block the transit peptide to allow import of the proteins into the chloroplast. The stop codon must be removed from the cDNA and the cDNA must be in frame with the YFP or CFP coding sequence to generate a successful translational fusion. 3. Sometimes, the use of the endogenous promoter to drive expression of the fusion proteins may reduce over-expressionrelated artefacts. Alternatively, a relatively low and controllable level of expression can be achieved using inducible promoters. 4. Because FRET and BiFC are highly dependent on the proximity and orientation of the fluorophore/fluorophore fragments, performing the experiment with different fusion proteins can increase the chances of successfully detecting the protein– protein interaction. 5. The proteins that are selected for positive controls should localise to the same compartment of the plant cell, and be expressed at similar levels, to the proteins of interest. 6. The protocols described are suitable for both Arabidopsis and tobacco leaves. The use of Arabidopsis allows one to analyse protein–protein interactions in both wild-type and mutant
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ackgrounds. However, transformation efficiencies are higher b in tobacco, and many Arabidopsis proteins have been demonstrated to localise and interact in tobacco cell chloroplasts (10–12), thus making this a very convenient system for BiFC and FRET assays. 7. High co-transformation efficiency greatly facilitates analysis of BiFC and FRET assays. Bombardment (see Subheading 3.2.1) provides a very quick and easy method of transiently coexpressing two constructs in chloroplasts and is favoured in this laboratory. However, if the efficiency of transformation is low, then the fusion cassettes should be transferred to binary vectors and the constructs transformed by infiltration (see Subheading 3.2.2), which will result higher numbers of cotransformed cells. Protoplast transformation is also an extremely effective method for the co-expression of constructs in Arabidopsis cells (see Chapter 4, Vol. 1). 8. Healthy Arabidopsis or tobacco is a vital starting point for any infiltration or bombardment experiment. The use of old or stressed plants can dramatically reduce the transformation efficiency and may also increase the levels of background during microscopy. 9. To prepare the microcarriers, weigh 60 mg of microcarriers in 1 mL of 70% (v/v) ethanol (freshly prepared). Vortex well for 5 min and incubate at room temperature for 15 min. Spin for 2 s in a microcentrifuge and remove the supernatant. Add 1 mL of sterile water and vortex well for 1 min; allow to settle for 1 min and then spin again for 2 s. Remove the supernatant and add 1 mL of sterile 50% (v/v) glycerol. The microcarriers can be stored at −20°C. Poor preparation of the microcarriers can lead to the formation of gold clumps that will damage/kill the cells on bombardment. 10. The manufacturer’s protocols should be referred to for safe operation of the PDS-1000/He™ Helium biolistic particle delivery system. 11. Samples can be observed any time after approximately 16 h. Timing is vital because if samples are incubated for too long, the levels of fusion protein can accumulate and increase the risk of non-specific interactions and/or false localisations. 12. The binary vectors can be transferred to the Agrobacterium using the freeze–thaw technique or by electroporation (20). 13. The density of the Agrobacterium cultures used for infiltration can affect the efficiency of transformation. We have found that OD600 = 0.2 provides consistent results; however, successful transformations have been carried out with bacteria diluted to OD600 = 0.02.
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14. Before infiltration, label each leaf or plant with the vector(s) to be infiltrated for identification. Place the tip of the syringe on the abaxial side of the leaf and place a gloved finger directly on the opposite side of the leaf. Press down gently on the plunger until the infiltration solution can be seen to diffuse through the air spaces in the leaf. Protective clothing, facemask, and eyewear should be used in case the infiltration solution sprays out of the syringe. 15. A negative BiFC or FRET assay is not empirical proof that two proteins do not interact. It is possible that the fusion to the YFP or CFP fragments has hindered the interaction, that the fusion has affected the structure of the protein or, in the case of BiFC, that the YFP fragments are unable to associate. 16. Software modules are available from many major microscope and software companies to allow automated acquisition of FRET images with each filter sets. Some macros allow acquisition of each image with the optimal settings for each filter set. It is also convenient to select a region of interest within a field of view, to limit the analysis to one cell. FRET modules are available from Improvision, Olympus, Nikon Instruments, Carl Zeiss and Leica Microsystems, to mention but a few. 17. FRET requires that cells with similar intensity levels of CFP and YFP are selected. In this laboratory, acquisition times are normally in the range of 200–1,000 ms.
Acknowledgements The authors would like to thank Daniela Gargano for constructive comments on this manuscript. This work was supported by Stavanger Health Research and The Norwegian Research Council. References 1. Lopez-Juez, E., and Pyke, K. A. (2005) Plastids unleashed: their development and their integration in plant development. Int. J. Dev. Biol. 49, 557–577. 2. Shoemaker, B. A., and Panchenko, A. R. (2007) Deciphering protein-protein interactions. Part I. Experimental techniques and databases. PLoS Comput. Biol. 3, e42. 3. Shoemaker, B. A., and Panchenko, A. R. (2007) Deciphering protein-protein interactions. Part II. Computational methods to predict protein and domain interaction partners. PLoS Comput. Biol. 3, e43.
4. Hu, C. D., Chinenov, Y., and Kerppola, T. K. (2002) Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Mol. Cell 9, 789–798. 5. Gordon, G. W., Berry, G., Liang, X. H., Levine, B., and Herman, B. (1998) Quantitative fluorescence resonance energy transfer measurements using fluorescence microscopy. Biophys. J. 74, 2702–2713. 6. Lakowicz, J. R. (1999) Principles of Fluorescence Spectroscopy, 2nd edn., Plenum Publishing Corp., New York, USA.
4 Studying Interactions Between Chloroplast Proteins… 7. Clegg, R. M. (1996) Fluorescence resonance energy transfer. In, Fluorescence Imaging Spectroscopy and Microscopy, Vol. 137 (Wang, X. F., and Herman, B., eds.) John Wiley and Sons Inc., New York, USA, pp. 179–252. 8. Förster, T. (1965) Delocalized excitation and excitation transfer. In, Modern Quantum Chemistry, Vol. 3 (Sinanoglu, O., ed.) Academic Press Inc., New York, USA, pp. 93–137. 9. Kerppola, T. K. (2006) Design and implementation of bimolecular fluorescence complementation (BiFC) assays for the visualization of protein interactions in living cells. Nat. Protoc. 1, 1278–1286. 10. Maple, J., Aldridge, C., and Moller, S. G. (2005) Plastid division is mediated by combinatorial assembly of plastid division proteins. Plant J. 43, 811–823. 11. Maple, J., Vojta, L., Soll, J., and Moller, S. G. (2007) ARC3 is a stromal Z-ring accessory protein essential for plastid division. EMBO Rep. 8, 293–299. 12. Fujiwara, M. T., Nakamura, A., Itoh, R., Shimada, Y., Yoshida, S., and Moller, S. G. (2004) Chloroplast division site placement requires dimerization of the ARC11/AtMinD1 protein in Arabidopsis. J. Cell Sci. 117, 2399–2410. 13. Fan, J. Y., Cui, Z. Q., Wei, H. P., Zhang, Z. P., Zhou, Y. F., Wang, Y. P., and Zhang, X. E. (2008) Split mCherry as a new red bimolecular fluorescence complementation system for visual-
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izing protein-protein interactions in living cells. Biochem. Biophys. Res. Commun. 367, 47–53. 14. Kerppola, T. K. (2006) Visualization of molecular interactions by fluorescence complementation. Nat. Rev. Mol. Cell Biol. 7, 449–456. 15. Patterson, G. H., Piston, D. W., and Barisas, B. G. (2000) Forster distances between green fluorescent protein pairs. Anal. Biochem. 284, 438–440. 16. Pollok, B. A., and Heim, R. (1999) Using GFP in FRET-based applications. Trends Cell Biol. 9, 57–60. 17. Kost, B., Spielhofer, P., and Chua, N. H. (1998) A GFP-mouse talin fusion protein labels plant actin filaments in vivo and visualizes the actin cytoskeleton in growing pollen tubes. Plant J. 16, 393–401. 18. Yang, Y., Li, R., and Qi, M. (2000) In vivo analysis of plant promoters and transcription factors by agroinfiltration of tobacco leaves. Plant J. 22, 543–551. 19. Xia, Z., and Liu, Y. (2001) Reliable and global measurement of fluorescence resonance energy transfer using fluorescence microscopes. Biophys. J. 81, 2395–2402. 20. An, G., Ebert, P., Mitra, A., Ha, S. B. (1988) Binary vectors. In, Plant Molecular Biology Manual (Gelvin, S. B., Schilperoort, R.A., and Verma, D. P. S., eds.) Kluwer Academic Publishers, Dordrecht, Netherlands, pp. A3/1-A3/19.
Chapter 5 Studying Chloroplast Protein Interactions In Vitro: An Overview of the Available Methods Joanna Tripp and Enrico Schleiff Abstract The analysis of protein–protein interactions is essential for the understanding of the molecular events in enzymatic pathways, signaling cascades, or transport processes in the chloroplast. A large variety of methods are available, which range from qualitative assays allowing for screening for new interaction partners, and semiquantitative assays allowing for a rough description of the interaction between two partners, to quantitative assays that permit detailed determination of kinetic and thermodynamic parameters. We summarize the available technologies, describe their range of applications and pitfalls, and give some examples from chloroplast research. The described techniques are generic and thereby important and useful to study the interaction network of proteins in Arabidopsis thaliana. In addition, we refer the reader to detailed protocols published elsewhere for each method. Key words: Chloroplast, Protein–protein interactions, Protein transport
1. Introduction Protein–protein interactions are analyzed to reconstruct the molecular events in enzymatic pathways, signal cascades, or transport processes. Many methods can be applied, and their selection is defined by the questions to be answered. Such questions can range from the identification of novel binding partners to the analysis of affinities and structural changes during interactions. Thus, methods to analyze protein-protein interactions range from qualitative assays such as immunoprecipitation and pull-down (see Subheading 2 below), which were refined in the tandem affinity purification (TAP) technique (see Chapter 3, Vol. 2), blue native polyacrylamide gel electrophoresis (BN-PAGE) (see Chapters 1 and 2, Vol. 2), chemical crosslinking approaches (see Chapter 18, Vol. 1), or yeast two-hybrid
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analysis (see Subheading 2), through semiquantitative assays such as dot-blot binding assays and size-exclusion analysis (see Subheading 3), to quantitative assays such as isothermal titration calorimetry, surface plasmon resonance and surface mirror techniques, or analytical ultracentrifugation (see Subheading 4). In the following sections, a variety of methods are described, their application range and pitfalls are discussed, and some examples from chloroplast research fields are presented. A summary of the described methods can be found in Table 1.
2. Qualitative Analysis of Protein–Protein Interactions
2.1. Hybrid ProteinBased Analysis of Interactions
Qualitative approaches are usually applied to identify new interaction partners, and herein they are powerful and applicable. However, qualitative analysis of protein interactions does not allow conclusions about affinities of the complexes formed. At best, different mutants of an interaction partner can be compared in their binding efficiency. Hence, all qualitative experiments provide some basic information about the interaction, but their results have to be confirmed by other experiments. Among many other qualitative techniques described in other chapters (see Chapters 18, Vol. 1, Chapters 1 and 3, Vol. 2), hybrid marker systems are used to explore interactions. Two methods have become popular for studying the interactions between components and substrates of the chloroplastic import machinery, the yeast two-hybrid (1, 2) and the split-ubiquitin system (3). Both the yeast two-hybrid and the split-ubiquitin systems have the advantage that they (1) allow studying protein–protein interactions in a cellular environment and (2) can be used for screenings of cDNA libraries for unknown interaction partners. In many cases, authors argue about a native environment to rationalize this approach. This argument, however, has to be taken with care. The classical yeast two-hybrid system (see below) analyzes the interaction of proteins within the nucleoplasm, creating an environment that is not comparable to the environment of typical chloroplast proteins in terms of pH and salt concentrations. Similarly, the split-ubiquitin system allows studying the interaction of proteins in the lipid environment of yeast membranes, which naturally do not resemble chloroplast membranes with respect to their lipid composition. In turn, the interactions can also be influenced by yeast proteins, for example by chaperones. Both strategies involve the generation of fusion constructs (see below). These fusions can lead to masking of binding sites or misfolding of the protein. Thus, care has to be taken when designing the fusion constructs. For the split-ubiquitin system, it has to be
Qualitative dot blot
Pull-down
Immobilization required; inaccurate Inaccurate
KD, koff, kon
Fluorescence based approaches b
KD, koff, kon
Immobilization required Labeling required
Sedimentation coeffi- Slow cient, hydrodynamic properties, molar mass, KD, 10 nM– KD, DG, DS, DH, DCP Large sample 100 mM requirement
nM–mM
KD, stoichiometry
SPR/resonant mirror pM–mM
ITC
Analytical ultracentrifugation
mM–mMa
KD, kon
–
No membrane proteins; fusion required Fusion required Immobilization required Immobilization required Immobilization required
Limitations
b
a
(60–67)
(44, 53–59)
(30, 42, 46–52)
(19, 31, 38–45)
(28–32, 36, 37)
(28–32, 35)
(1, 10, 13, 17, 21–27)
(3, 7, 8) (11, 12, 14–20)
(1, 2, 4–6)
References
Allows studying interactions (32, 68–72, 74–81) in bulk and at the singlemolecule level
No labeling; thermodynamic parameters can be determined Small sample requirement
No labeling; very suitable for homooligomeric interactions
No labeling; most suitable for homooligomeric interactions
Fast, easy comparison of mutants
Fast
Fast and easy
Suitable for screening Fast and easy
Suitable for screening
Advantages
Dependent on the detector As outlined in Subheading 4.5, fluorescence-based approaches can measure interactions at the single-molecule level
Quantitative
SEC
–
–
Split ubiquitin – Immunoprecipitation
Parameter set –
KD range –
Yeast two-hybrid
Semiquantitative Semiquantitative dot blot
Qualitative
Method
Table 1 Features and application range of methods for studying protein interactions
5 Studying Chloroplast Protein Interactions In Vitro… 69
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taken into account that the fused fragments have to be exposed to the cytosolic side of the membrane: again, the appropriate design of the constructs is crucial for the success of the experiment. These limitations and pitfalls have to be kept in mind while interpreting the results. Not surprisingly, the false-positive rate of yeast two-hybrid assays was estimated to be about 50% in largescale screenings (4). However, both systems have proven to be powerful when carefully controlled. The classical yeast two-hybrid system is based on the fusion of the putative binding partners to the two domains of the Gal4 transcription factor, namely, the DNA-binding domain (BD) and the activation domain (AD). When these two domains come together in the course of interaction of the binding partners, transcription of a reporter gene is initiated, which leads to survival on selective media. For detailed protocols, the reader is referred to (5, 6). The yeast two-hybrid system has been developed mainly for studying the interactions of soluble proteins. For membrane proteins, the transmembrane domains have to be removed. With this approach, interactions of the stromal domains of the inner envelope translocon components, Tic40 and Tic110, as well as of Tic40 and Hsp93, could be demonstrated (1, 2). The split-ubiquitin system was developed to study membrane protein interactions. The ubiquitin protein is split into an N-terminal part (Nub) and a C-terminal part (Cub). The Nub region is mutated in such a way that it cannot assemble spontaneously with Cub (NubG). Further, a transcriptional activator is fused to the C-terminus of Cub. These modified halves are fused to the two putative binding partners. If NubG and Cub assemble upon interaction of the putative binding partners, ubiquitin-specific proteases cleave Cub, leading to the release of the transcriptional activator, which results in expression of a reporter gene. Detailed protocols for the split-ubiquitin system and a summary of yeast-based methods for the analysis of membrane protein interactions can be found in refs. 7 and 8, respectively. The suitability of the split-ubiquitin system for studying interactions of thylakoid membrane proteins has been demonstrated by the example of the thylakoid Sectranslocase subunits SecY and SecE (3). Interactions of the SecY/E proteins with the thylakoid membrane insertase Alb3, and of Alb3 with reaction center proteins of PSI and PSII were shown, indicating the important function of Alb3 in the assembly of these membrane protein complexes. For the analysis of the heterodimerization of the Arabidopsis precursor protein import receptors atToc33 and atToc159 in the outer envelope membrane of chloroplasts, the split-ubiquitin system was adapted for application in plant protoplasts (9) (note that the “at” prefix in each case denotes species of origin: Arabidopsis thaliana). Here, the interaction was detected by cleavage of GFP from the Cub moiety. The authors aimed to explore the interaction
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of membrane inserted proteins, but could not detect membrane insertion of the atToc33 receptor. In addition, a high background cleavage was detected, possibly due to unspecific proteolytic cleavage of the Cub fusion protein. Another disadvantage of the system is that the success of the experiment is dependent on protoplast transformation, which can vary in its quality, and on the detection of the cleavage product by Western blot analysis. Nevertheless, the expansion of the hybrid technology to the more native plant system opens new perspectives for the analysis of chloroplastic proteins. Here, the system comes close to approaches with fluorescence tags, such as bimolecular fluorescence complementation (BiFC) and Förster resonance energy transfer (FRET) (see Chapter 4, Vol. 2). 2.2. Immuno precipitation and Pull-Down Assays
In cell biology, immunoprecipitation (IP) and pull-down assays are widely used for the identification of new binding partners or for the direct assessment of protein–protein interactions. One binding partner (“bait”) is immobilized on a solid support and binding of the other (“prey”) is determined. Identification of new binding partners can be achieved by gel electrophoresis followed by mass spectrometry (10). For analysis of interactions between known partners, the prey can be detected by immunodecoration or by radioactive labeling (11–13). In IP assays, antibodies are used to isolate proteins and characterise protein complexes. Immobilization of the antibody can be achieved by using Protein A- or Protein G-Sepharose. Detailed protocols can be found in refs. 11, 12. Protein G, although more expensive, has the advantage that it binds to a higher variety of antibody classes (11). When analyzing the bound proteins by immunodetection, the presence of antibody heavy and light chains can be very disruptive. This problem can be circumvented by the cross-linking of the antibody to Protein A- or Protein G-Sepharose, or by direct coupling of the antibody to amino-reactive resins (14). The latter method has the advantage that it eliminates the need for Protein A or Protein G, but requires purification of the antibody. As control experiments, IPs with preimmune serum (15–17) or with an antibody not recognizing the bait (15), and IPs with cell extracts not containing the bait, are recommended. It has to be kept in mind that interactions might be hampered if the epitope recognized by the antibodies is located in the binding surface of the bait. Numerous examples can be found in the literature describing the successful application of IP in the field of chloroplast research. These include experiments describing (1) the association of radioactively labeled chloroplastic precursor proteins with components of the cytoplasmic chaperone system in wheat germ extract or reticulocyte lysate (18, 19), (2) the association of plastidic components of the import machinery with radiolabeled precursor proteins (15), and (3) the assembly of components of the plastidic import machinery in the presence (15, 16) or absence of precursor
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proteins (20). IP of membrane-associated translocon components requires membrane solubilization prior to the experiment, and for this Triton X-100 (16, 20) and decylmaltoside (15, 17, 19) are the most frequently used detergents for plastidic membranes. To investigate the formation of Tic/Toc translocon supercomplexes in the chloroplast envelope, in the presence and absence of precursor proteins, a sequential immunoprecipitation strategy was used in ref. 16. For this aim, chloroplasts were cross-linked to saturating concentrations of a modified version of the precursor of the small subunit of Rubisco (preSSU) containing a single cysteine in its transit peptide (pS-1), or left untreated. Following isolation of envelope membranes and solubilization with Triton X-100, the extract was applied sequentially to IgG-Sepharose coated with antibodies against Toc34, Toc159, and Tic110. Eluates and flowthrough were analyzed by SDS-PAGE and immunoblotting. The results revealed stable association of the outer membrane translocon components Toc75, Toc34, and Toc159, whereas only a minor fraction of the inner membrane components Tic110, Tic20, and Tic22 associated with the Toc complex, irrespective of the presence or absence of precursor. Interestingly, Tic110, Tic20, and Tic22 did not seem to stably associate in the absence of Toc components. The guanosine triphosphate (GTP)-dependence of the association of the Toc core components, Toc75, Toc34, and Toc159, was addressed in ref. 17. Here, immunoprecipitation experiments were performed with outer envelope vesicles solubilized with decylmaltoside in the presence of guanosine diphosphate (GDP) or the nonhydrolyzable GTP-analog guanylyl-imidodiphosphate (GMPPNP). Immunoprecipitation was carried out with antisera against Toc34 or Toc159 (both of which are GTPases), followed by incubation with Protein A-Sepharose. In the absence of nucleotides or in the presence of GTP, stable association of Toc159 and Toc34, as well as of Toc34 and Toc75, could be detected, and this association was reduced in the presence of GDP. By contrast, the association of Toc159 and Toc75 was nucleotide-independent. The results revealed the regulation of the assembly of the Toc complex by GTP-binding and -hydrolysis. In pull-down experiments, the bait protein is directly bound to a solid support. For this aim, an affinity tag, such as glutathione S-transferase (GST) or hexahistidine (His) can be fused to the bait protein, allowing purification via glutathione or Ni-nitrilotriacetic acid (NTA) matrices. For protocols, the reader is referred to (10, 13). The tagged protein is usually recombinantly produced (e.g., in bacteria, insect cells, or yeast), purified, and can subsequently be incubated with in vitro translated, radioactively labeled proteins (21, 22), cell lysates (23), or purified protein (1); alternatively, it may be directly expressed in the target organism (24). An in vivo strategy for the purification of protein complexes by tandem affinity purification is described in Chapter 3, Vol. 2. The bait protein can
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also be immobilized by covalent coupling to the matrix, for example via cross-linking of cysteine residues, amine or thiol groups (25). Irrespective of the immobilization strategy, some limitations common to all solid-phase binding assays should be considered when designing the experiment. First, false-positive results are commonly due to high protein concentrations in the assay mixture and/or protein denaturation on the support, which enhances the probability of unspecific hydrophobic interactions between proteins to be investigated. To verify the specificity of the assay, the following control experiments can be performed: (1) Competition of binding to immobilized bait protein by soluble bait protein or another protein competing for the same binding site (20, 23). (2) Site-directed mutagenesis or deletion of functionally important amino-acids or domains (26, 27). (3) Addition of effectors, e.g., nucleotides, regulatory proteins, or precursors (1, 17, 21, 27), which alter the interaction of binding partners. False negative results or decreased binding efficiency may also be caused by the high density of the immobilized protein at the surface of the support (steric hindrance). In the case of IP experiments, the result might be influenced by the epitope recognized by the antibodies used, as they might target the interaction surface for the prey and thereby either compete for the interaction or only precipitate unbound bait proteins. Furthermore, the requirement of many wash steps poses a disadvantage for the determination of binding constants. Hence, solid-phase binding assays are usually not suitable for detecting transient, low-affinity (i.e., high off-rate) interactions. Using pull-down assays, interactions between Tic110, Tic40, and Hsp93 required for precursor translocation across the inner envelope were elucidated (1). For this aim, GST-tagged versions of the whole Tic40 stromal hydrophilic domain containing both the tetratricopeptide repeat (TPR) and Hip/Hop subdomains, or the TPR and Hip/Hop subdomains alone (GST-Tic40S, -Tic40TPR, and -Tic40Hip/Hop, respectively), were recombinantly expressed and purified. By incubation with the Tic110 stromal domain fused to a C-terminal His-tag (Tic110S-His6), and subsequent recovery with glutathione resin, the TPR-domain of Tic40 could be identified as the binding site for Tic110. The affinity between Tic110SHis6 and GST-Tic40S was found to be increased in presence of transit peptides. Nucleotide dependent association of GST-Tic40S with the stromal chaperone Hsp93 was demonstrated using Histagged Hsp93 (Hsp93-His6) preincubated with adenosine triphosphate (ATP), adenosine phosphate (ADP), or the nonhydrolyzable ATP analog adenylyl-imidodiphosphate (AMP-PNP). The binding reaction was carried out in the presence of precursor of ferredoxin (prFD) transit peptides and Tic110S-His6. Binding of Hsp93 to Tic40 was higher in the ATP- or AMP-PNP-bound states, suggesting that Tic40 might function in stimulating Hsp93 ATPase activity.
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The assembly of structurally distinct Toc complexes was demonstrated by Ivanova et al. (20) using C-terminally His-tagged, recombinant versions of the two Arabidopsis thaliana isoforms of the Toc34 receptor, atToc33 (atToc33G) and atToc34 (atToc34G); these recombinant receptors lacked their transmembrane domains and were immobilized on an Ni-NTA matrix. Following incubation with in vitro translated, 35S-labeled atToc120, atToc132, or atToc159 (all of which are Arabidopsis isoforms of the other main precursor receptor, Toc159), the amount of bound, radiolabeled protein was determined by SDS-PAGE and phosphorimager analysis. While atToc159 showed a higher binding preference for atToc33G than either atToc132 or atToc120, atToc120 and atToc132 bound to atToc34 more strongly than atToc159 did. It can be speculated that the differential assembly of the aforementioned isoforms of the Toc34 and Toc159 receptor GTPases into distinct Toc complexes reflects distinct affinities for different precursor proteins and, therefore, distinct targeting pathways. 2.3. Qualitative Dot-Blot Analysis
For the identification of protein binding motifs within proteins, a peptide blot is a very efficient and convenient method (28). In general, the sequence of the target protein is dissected into 13 amino-acid-long peptides, with an overlap of 10 amino acids between adjacent peptides. The peptides are spotted onto a cellulose membrane (29) and incubated with the binding partner supposed to interact with the target protein. The interaction can be probed by immunodecoration using antibodies raised against the binding partner, or a tag fused to it (29–31). Alternatively, the binding partner may be labeled fluorescently or radioactively (32). The binding partner has to be recombinantly produced (e.g., in bacteria, insect cells, or yeast) (29–31, 33, 34) and purified before application. By defining the experimental conditions, it has to be ensured that binding of the interaction partner to the cellulose is insignificant. Furthermore, one has to consider that only secondary structure elements are formed within the short peptides, and that peptides of high hydrophobicity might tend to aggregate. To exclude the possibility of artificial binding caused by the immobilization of the peptide, it is important to confirm the determined binding region by further experiments. Nevertheless, using the proper controls, the peptide blot assay can be very useful in the identification of putative binding sites. The peptide blot analysis was used to determine sites within the sequence of the phosphate carrier, which are recognized by Toc64 lacking the TPR domain (Toc64DTPR) (35). Here, the interaction was performed in the presence of 0.05% Tween 20, 100 mM KCl, 0.05% bovine serum albumin (BSA) to reduce the background binding. Two binding sites were identified, and the corresponding peptides were able to compete for the interaction of Toc64DTPR with another substrate protein.
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The dot-blot analysis can also be performed with entire proteins or other substrates coupled to the surface of cellulose membranes. However, these approaches are often used for semiquantitative analysis of interactions (see Subheading 3.1).
3. Semiquantitative Analysis of Protein–Protein Interactions
3.1. Semiquantitative Dot-Blot Analysis
Semiquantitative experiments give an estimate of the parameters describing the interaction between two compounds. However, due to technical limitations, these techniques usually do not allow the determination of all parameters. For the dot-blot experiments described below (see Subheading 3.1), the dissociation of the interacting protein during the wash steps cannot be controlled to its final extent. Similarly, during size-exclusion chromatography (see Subheading 3.2), complexes can dissociate because of the dilution during protein separation. Nevertheless, these methods allow the determination of apparent dissociation constants, which can be compared between different substrates or receptors. As described in Subheading 2.2, in dot-blot analysis, a substrate immobilized on a membrane is incubated with a soluble binding partner (28, 29). The detection of the binding partner can be carried out by immunodecoration or by radioactive or fluorescence labeling. The assay can be calibrated by spotting defined amounts of purified binding partner onto the membrane. Two different variants of the assay can be performed. In the first variant, a dilution series of one protein is immobilized on the membrane support and incubated with a constant amount of the soluble binding partner. Alternatively, the amount of the spotted protein may be kept constant while varying the concentration of the soluble binding partner. For practical reasons, the first approach is used more often. In general, it has to be noted that protein denaturation and/or steric hindrance caused by immobilization may have a major influence on the assay quality. Furthermore, dot-blot assays have the disadvantage that the quantification can be performed only in relation to the amount of spotted protein, and not in relation to its concentration. Dot-blot analysis was performed to study the interaction of the outer envelope protein CHUP1 (chloroplast unusual positioning1) and profilin (32). Different amounts of profilin (as interaction partner) and BSA (as control) ranging from 0.1 to 4 mg were spotted onto a nitrocellulose membrane. The blot was blocked with 0.3% skimmed milk and 0.03% egg albumin in phosphate-buffered saline. The bait CHUP1 was produced by in vitro transcription and translation in wheat germ extract in the presence of 35S-methionine. The remaining radioactive methionine was removed by a G25 Sephadex column. Radioactively labeled CHUP1 was incubated with the
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membrane, and the binding was visualized by autoradiography. The experimental result was verified by immobilizing the recombinantly produced C-terminal portion of CHUP1 (CHUP1-CT), followed by incubation with profilin. Profilin was subsequently visualized by immunodecoration with commercial antibodies. Such double experiments have proven to be very helpful in differentiating between specific binding and nonspecific interaction with the blot surface. A similar approach was used for determination of nucleotidedependent homo- and heterodimerization of the GTPase domains of the Toc34 and Toc159 preprotein receptors (30, 31). The test protein was preloaded with nucleotides and spotted onto nitrocellulose membranes using a 96-well vacuum manifold (Bethesda Research Laboratories). The nitrocellulose membranes were saturated with 0.3% low-fat milk powder with 0.03% BSA and subsequently incubated with purified GST-atToc33/34 or GST-psToc159 (GST is used here as a purification tag; the “at” and “ps” prefixes indicate species of origin: Arabidopsis thaliana and Pisum sativum). Background binding was controlled by incubation of GSTatToc33/34 or GST-psToc159 with saturated nitrocellulose membranes spotted with BSA. After two washes (10 min), the amount of bound protein was determined by immunodetection with GST antibodies. Background staining caused by unspecific binding of GST antibodies was determined by immunodetection of membranes with spotted proteins without adding the soluble interaction partner. Intensities were quantified by densitometry. A similar approach can be performed using immobilized metabolites or lipids; the latter is known as a Fat Western or Protein Lipid Overlay (PLO) assay (36). Here, the lipids are dissolved in a mixture of chloroform–methanol–water and spotted onto Hybond-C or polyvinylidene fluoride (PVDF) membranes. The membrane is subsequently blocked. One has to take care that the blocking solution is fatty-acid-free. However, in most protocols a detergent such as Tween 20 is present in the blocking solution to avoid hydrophobic associations of the protein of interest with the membrane. The membranes are subsequently incubated with the protein in solution, typically with different concentrations. After washing the blots, the amount of bound protein can be quantified. Such a system was used to investigate the association of chloroplastic precursor proteins with different chloroplast lipids (37). Different concentrations of lipids were spotted onto PVDF membranes, which were subsequently saturated with 0.25% fattyacid-free BSA for 1 h. Different mutants of the precursor of ferredoxin:NADP+-oxidoreductase (FNR) were in vitro transcribed and translated in the presence of 35S-methionine, diluted into 0.25% fatty-acid-free BSA and 1 mM cold methionine, and incubated for 1 h at 20°C while rotating the blot. The blot was washed three
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times and the binding was analyzed and quantified by phosphorimaging. The assay was calibrated by spotting defined amounts of the translation product onto the membrane. For a bimolecular interaction as denoted in Eq. 1 with A being the spotted protein and B being the protein in the soluble phase, the dissociation constant (KD) is defined by Eq. 2.
A + B AB
(1)
K D = [A] × [B] / [AB] = koff / kon
(2)
Thereby, kon and koff denote the association (on) and dissociation (off) rate constants. The concentration of [AB] is determined in the assay and corresponds to the amount of bound partner ([B]). The concentration of unbound A and B can be expressed as the difference between the respective starting concentration ([A]0 or [B]0) and the concentration of the complex ([AB]), as denoted in Eqs. 3 and 4.
[B] = [B]0 − [AB]
(3)
[A] = [A]0 − [AB]
(4)
By substitution, we can rewrite Eqs. 2–5.
K Dapp = ([A]0 − [AB])× ([B]0 − [AB]) / [AB]
[AB] = −{(K Dapp + [A]0 + [B]0 )2 / 4 − [A]0 × [B]0 }1/ 2 + (K Dapp + [A]0 + [B]0 ) / 2
(5a)
(5b)
However, with Eq. 5, one can only determine the apparent dissociation constant (KD app), since the washing steps cause a reequilibration of the bound protein between the solid and immobile phase and, therefore, its partial removal. Furthermore, the number of available binding sites on the component A is hard to estimate owing to inhomogeneous orientation of the spotted protein and steric hindrance. 3.2. Size-Exclusion Chromatography
Size-exclusion chromatography (SEC) is commonly used both for preparative (i.e., protein purification) and analytical purposes (38, 39). The method is based on gradual separation of molecules according to their hydrodynamic radius, which is, for globular proteins, roughly proportional to their molecular weight. Often, this approach is combined with static light scattering for a more precise determination of the molecular weight (40). Thus, this simple method is generally used for the determination of the oligomeric state of self-associating proteins. At the same time, it is possible to estimate the affinity for the formation of binary complexes (homo- or heterodimers). To this end, SEC is performed at different concentrations
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of one interaction partner (A) while the concentration of the other (B) is kept constant. The ratio between the free (unbound) and complexed species A is calculated after integration of peaks corresponding to the particular species in the resulting elution profile (usually recorded as absorbance at 280 nm). One problem occurring while analyzing heterodimers is the distinct absorption of the two interaction partners. Hence, the total absorption for each compound has to be determined in the absence of the other partner. Some practical limitations of this technique should be considered. First, one should be aware that interactions (electrostatic, van der Waals, hydrophobic) between the sample and the column material might occur, leading to anomalous separation profiles. This can, in most cases, be avoided by the proper choice of the column material. As a consequence, the migration behavior of each of the interaction partners has to be determined beforehand for an optimal choice. Second, for optimal resolution, the sample volume is restricted to £5% of the column volume and the protein(s) of interest must, therefore, be available at a high concentration (usually ³1 mg/ml), which might, in some cases, lead to protein aggregation and precipitation. Addition of stabilizing osmolytes such as glycerol is usually applied to decrease the aggregation, even though this compromises the resolution of SEC due to their high viscosity. Third, the titration experiments are applicable only for interactions with moderate dissociation constants, since the lower detection limit is defined by the sensitivity of the optical cell recording the absorbance. And fourth, SEC requires long separation times resulting in a progressive separation of different oligomeric and monomeric species. In addition, the sample becomes diluted. As a consequence, a constant dissociation through readjustment of the chemical equilibration can occur. This problem becomes more apparent during the analysis of low-affinity interactions, often causing anomalous, “smearing” elution profiles. To circumvent this problem, one of the interaction partners can be added into the separation buffer. Dissociation caused by dilution is the most severe problem for the estimation of precise dissociation constants by this technique. In this case, Eq. 1 becomes unidirectional for the dissociation. For an analysis of homodimerization, one can formulate a kinetic Eq. 6 for the decay of the dimeric species Ad, with [Am]0 or [Ad]0 being the concentration of the monomer or homodimer prior to application to SEC. Hence, the increase of the monomeric species can be formulated as presented in Eq. 7.
[A d ] = [A d ]0 × exp {−koff t }
(6)
d[A m ]/dt = koff × 12 × [A d ]
(7)
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Solving this equation, one obtains a running-time-dependent relation for the final monomeric concentration [Am]f or the final dimeric concentration [Ad]f (Eqs. 8 and 9, respectively).
(
)
[A m ]f = [A m ]0 + [A d ]0 × 1 − exp {−koff t } / 2
(8)
[A d ]f = ([A]0 − [A m ]f ) / 2
(9)
Thus, even without the knowledge of the initial values of [Am]0 or [Ad]0, one can estimate the off rate koff by determination of [Am]f or [Ad]f at two or more different flow rates. In Eq. 8, [Am]0 can be replaced by [A]0 − 2 × [Ad]0, leaving only [Ad]0 and koff as the free parameter. If the separation window of the SEC material is well chosen, this technique allows the determination of all three parameters, namely, [A]f, [B]f, and [AB]f. Assuming a slow off rate, one can approximate [X]f = [X] and thereby the determination of the dissociation constant can be performed by Eq. 2. In cases where one of the interaction partners is added to the separation buffer to obtain a constant concentration, one has to use Eq. 5. The dissociation constant for homodimerization can be estimated using Eq. 10.
{
}
[A d ] = (K D + 4 × [A]0 − K D (K D + 8 × [A]0 )
1/ 2
)/ 4
(10)
SEC was applied successfully to the analysis of dimerization of the soluble GTPase domains of Toc159 (41) and Toc34 family members (31, 41, 42) due to the high yield of recombinant proteins and their solubility. The behavior of dimerization in different environments, e.g., under different pH conditions, was compared (43). For atToc33, the apparent KD for homodimerization was estimated as outlined above (31). SEC is also commonly used to verify the association of individual proteins in heterooligomeric complexes (19, 44). If the molecular weight of all components occurring in a complex is known, SEC may be used to estimate the relative stoichiometry of individual proteins in the complex (45).
4. Quantitative Analysis of Protein–Protein Interactions
The description of the molecular mechanisms of biological processes involving protein–protein interactions requires the knowledge of the physical parameters of these interactions. This is relevant as many parameters influence the kinetics and the affinity of protein– protein interactions. For instance, protein–protein interactions are often sensitive to environmental changes such as pH or salt
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conditions, to the availability of cofactors such as metabolites, nucleotides, or other proteins, or to posttranslational modifications of interaction partners. However, the influence of these parameters only becomes obvious when significant changes of the dissociation constant KD, the association rate kass, or the dissociation rate kdiss are observed. Here, the authors would like to define two terms, which are often misused in the literature. In contrast to the dissociation rate, the association rate kon determined within an experiment is concentration dependent. For calculation of the concentration-independent association rate kass, Eq. 11 has to be used. As a consequence, the association rate kon has to be determined at different ligand concentrations [L] to obtain kass.
kon = kass [L] + kdiss
(11)
Another term that is often misused is affinity. Many authors wrongly equate changes of the association Ka or dissociation constant KD with changes of affinity. By definition, the chemical affinity (A) is the change of the Gibbs or Helmholtz energy. The affinity in a nonequilibrated system is described by Eq. 12 where R is the gas constant, T is the temperature (in Kelvin), and Qr is the reaction quotient.
A = −∆G = −∆G 0 − RT ln {Q r }
(12)
However, for biological systems, the term standard affinity (Ast) is used, describing the affinity in equilibrium (with Ka in M-1 and KD in M):
1 Ast = −∆G 0 = RT ln {K a } = RT ln KD
(13)
The change of the affinity between protein 1 and protein 2 is thereby given by Eq. 14,
∆Ast = −∆∆G 0 = RT ln{K a1 / K a2} = RT ln{K D2 / K D1},
(14)
where the fold of affinity change is defined by ln{Ka1}/ ln{Ka2} = ln{1/KD1}/ln{1/KD2}. These relations have to be taken into account when discussing affinities of interactions. The quantitative techniques described in this section allow the analysis of biophysical parameters of interactions, such as stoichiometry of complexes, binding affinities, equilibrium association and dissociation constants, and kinetic rate constants. All of these techniques measure interactions in an isolated system. 4.1. Analytical Ultracentrifugation
Analytical ultracentrifugation (AU) has become a versatile technique for the study of protein–protein interactions. Monitoring the sedimentation of protein-complexes through UV absorbance
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and interference optics in the analytical ultracentrifuge allows characterization of the molecular mass, hydrodynamic properties, and binding constants of protein complexes in solution, without the need for labeling or immobilization. Two basic methods are distinguished in AU: the sedimentation velocity (SV) method and the sedimentation equilibrium (SE) method. Detailed protocols for both methods are given in refs. 46, 47. In the SV method, protein complexes are sedimented at high velocity. The motion of a concentration boundary formed by depletion of the protein at the meniscus of the sample cell is recorded as a function of time. Analyses of the obtained sedimentation velocity data can be performed using the program SEDFIT (48), allowing the calculation of sedimentation coefficient distributions, expressed in Svedberg units. Parameters such as the solvent viscosity h and density r required for determination of the sedimentation coefficient can be determined using the program SEDNTERP (49) for a wide variety of solutions. Likewise, the program can be used for calculation of the partial specific volume v of the protein from its amino-acid sequence. The SV method can be used for the determination of stoichiometry and dissociation constants of protein complexes in the nanomolar to millimolar range, as well as their shape. Applications in the field of chloroplast protein import are so far restricted to SV analyses of Toc receptor homodimerization (1) of mutants (42, 50) and (2) in different nucleotide loading states (30). These data provided important insights into the regulation of Toc34 homodimerization and the function of a structure resembling a GTPase activating protein (GAP) arginine finger involved in dimer formation. For the investigation of the dimerization of Toc34 by analytical ultracentrifugation, several concentrations were tested, revealing concentration-dependent dimerization. The R133A mutation in the arginine finger of pea Toc34 extending into the GTP-binding pocket of the dimerization partner was shown to prevent dimerization (50). For investigation of the nucleotide dependence of dimerization, Toc34 preloaded with GDP or the nonhydrolyzable GTP analog GMP-PNP was used in SV studies (30). The results suggested that the nucleotide load has no effect on dimerization. In these experiments, dissociation constants could be derived from the sedimentation velocity data using the program SEDPHAT (51). The SE method allows determination of dissociation constants and molecular mass. In SE, the sample is centrifuged at lower speed, until sedimentation and diffusion are in equilibrium. SE experiments were shown to be also suitable for studying the mass of solubilized membrane protein complexes in detergent micelles (52). 4.2. Isothermal Titration Calorimetry
Isothermal titration calorimetry (ITC) allows measurement of the thermodynamic parameters of protein interactions by measuring the heat changes resulting from the interaction of two binding
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partners. In contrast to other techniques, ITC does not require labeling or immobilization of the binding partners. The ITC instrument contains two identical cells: a reference cell and a sample cell containing one of the binding partners. The other binding partner is injected through a syringe, initiating the binding reaction, which leads to the release or absorption of heat. This causes a temperature difference between the sample and the reference cell. The measurement is based on recording the power (m cal/s) it takes to keep the temperature between the sample cell and a reference cell constant. Equation 15 describes the relation between cumulative heat of binding and the association constant Ka = 1/KD, where Q is the heat evolved on addition of ligand, V0 is the volume of the cell (known), DH is the enthalpy of binding per mole of ligand, [M] is the total concentration of the receptor, and [L]f is the free ligand concentration.
Q = V0 × ∆H × [M] × K a × [L]f / (1 + K a × [L]f )
(15)
Equation 15 can be transformed into Eq. 16a, considering more than one binding site (n) and the total ligand concentration [L]t.
Q = n × V0 × ∆H × [M] / 2 × {(1 + [L]t / n / [M] + 1 / n / [M] / K a ) − X 1/ 2}
X = (1 + [L]t / n / [M] + 1 / n / [M] / K a )2 − 4 × [L]t / n / [M]
(16a)
(16b)
By Eq. 16a, the association constant Ka, the binding enthalpy DH, and the binding stoichiometry n can be directly determined. From these parameters, the Gibbs energy DG and the entropy DSt can be calculated using Eq. 17.
∆G = −RT ln K a
(17a)
T ∆S = RT ln K a + ∆H
(17b)
DH provides important information about the strength of the interaction between the binding partners compared to those with the solvent, while DS is influenced by two parameters, desolvation and changes in conformation. Even if two binding reactions exhibit the same changes in their Gibbs energy, different contributions of DH and DS indicate different binding mechanisms. By performing experiments at different temperatures, information about heat capacity changes (DCP) can be obtained (Eq. 18), and this can be related to the change in surface area buried upon formation of a bimolecular interface (53).
∆C p =
d∆H dT
(18)
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Since the ITC instrument measures also the heat of dilution generated during injection of the reactant in the sample cell, it is indispensable to perform a control experiment, where the binding partner is injected into buffer solution. The obtained values can be subsequently subtracted. In general, the buffer composition in the syringe and the cells needs to be exactly the same to avoid unspecific heat effects. Detailed protocols and detailed explanations of the calculations can be found in refs. 54, 55. Ligands with dissociation constants in the range KD 10 nM–100 mM are best suited for ITC experiments. However, adaptations of the experimental design allow measuring high affinity binding reactions. One binding partner can be prebound to a competitive ligand with lower affinity so that the other binding partner has to displace the competitor from the binding site (displacement titration) (56). ITC has been successfully applied for the characterization of interactions in the chloroplastic signal recognition particle (cpSRP)mediated pathway, which is involved in targeting photosynthetic proteins to the thylakoid membrane. Here, details such as binding affinities and stoichiometry of the interactions of the chromodomains of the cpSRP subunit SRP43 with SRP53 (57, 58), and of SRP43 and SRP53 with the soluble C-terminal domain of Alb3 (25), could be elucidated. Furthermore, binding affinities of the ankyrin repeats in SRP43 to a signal sequence in its substrate lightharvesting chlorophyll protein (LHCP), the L18 peptide, and critical residues for interactions were determined (59). 4.3. Surface Plasmon Resonance or Resonant Mirror Techniques
Surface plasmon resonance (SPR) or resonant mirror techniques are suitable for highly accurate quantitative analyses allowing for determination of binding affinity and kinetics (60, 61). By performing experiments under different temperature conditions, determination of thermodynamic parameters of an interaction is possible. In both techniques, one interaction partner is immobilized on a coated optical sensor chip, and interactions are monitored by using different optical detection systems. In a typical experiment, the ligand in solution is added to the immobilized binding partner, and binding is determined by measuring refractive index changes caused by accumulation of macromolecules close to the sensor surface. After removal of the ligand in solution, dissociation is recorded over time. The two systems differ not only in the optical detection system but more importantly in the design of the sample chamber. In the most common SPR systems such as Biacore (Uppsala, Sweden), a flow system is used, which allows a constant delivery of the ligand and buffer to the sensor surface. In resonant mirror devices (e.g., IAsys, Cambridge, UK), a stirred cuvette system is used. Since the concentration of the ligand in solution should not alter during binding, and the ligand should be rapidly removed after dissociation,
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the volume of the cuvette should be kept large in comparison to the amount of immobilized binding partner, which should be kept minimal. So-called mass transport effects can affect the accuracy of the measurement (62). Mass transport of the ligand from the bulk of the solution to the sensor can be slowed down, e.g., by properties of the immobilization matrix on the sensor surface, thus influencing the binding kinetics. Mass transport effects should be taken into account in the data analysis and can be estimated by computer prediction. For SPR sensor chips and resonant mirror cuvettes, different immobilization strategies are possible which include covalent linkage (e.g., amine or thiol coupling) as well as noncovalent binding methods (e.g., Ni-NTA, antibodies) (60). Covalent immobilization via side-chain amino groups that usually occur at several positions in each protein molecule may denature the bait protein or alter its molecular dynamics. Furthermore, the orientation of the immobilized protein cannot be controlled. If possible, it is, therefore, advantageous to introduce one single cysteine residue at either terminus of the protein and immobilize it, for example, via a maleimide function. The problem can also be circumvented by using His- or GST-tagged bait proteins, which are bound to covalently coupled Ni-NTA groups or GST-antibodies. For studying membrane proteins in a lipid environment, biosensor surfaces were engineered that allow reconstitution of proteins into immobilized lipid bilayers (63). To obtain reliable results for kinetic analyses, four to six ligand concentrations spanning a range from 0.1- to 10-times the KD should be used in the experiment. In general, the useful KD range for SPR is picomolar to high micromolar, and association and dissociation rate constants can be determined in the following ranges: kon = 103 − 108 M−1 s−1, koff = 10−5 – 1 s−1 (64). For investigation of interactions between the cpSRP subunits SRP43 and SRP54 using SPR (65), GST–cpSRP43 constructs with deletions in the chromodomains and adjacent regions, or GST as a control, were bound to anti-GST-antibody covalently linked to the sensor chip. Recombinant cpSRP54M-His was passed over the decorated chip in at least six different concentrations. Association and dissociation rates and KD were determined, leading to the conclusion that chromodomain 2 (CD2) provides the major binding site for SRP54. The resonant mirror technique was used for studying the GTPdependent preprotein and transit peptide binding properties of pea Toc34 (psToc34) and two Toc34 homologs from Arabidopsis (atToc33 and atToc34) (66, 67). For this aim, a cuvette with coupled Ni-NTA was used, to which recombinantly produced, His-tagged psToc34, atToc33, and atToc34 lacking their transmembrane domains (psToc34DTM, atToc33DTM, and atToc34DTM) were bound. As ligands, recombinantly produced preproteins (preSSU [Rubisco small subunit] and preOE33 [oxygen evolving complex 33 kDa subunit]) and synthetic peptides corresponding to the
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preSSU transit peptide were used. In this way, important information about transit peptide recognition by psToc34, as well as about the GTP-dependence of precursor binding by atToc33 and atToc34, could be gained. 4.4. FluorescenceBased Approaches
The techniques introduced in this section are based on measuring changes in parameters of fluorescence emission, such as polarization, intensity, lifetime, and emission energy, that occur upon binding of a fluorescently labeled protein to its interaction partner. For labeling, a fluorophore has to be selected that is sensitive to changes in the environment. The fluorophore is usually covalently attached to thiol- or amino-groups, ideally to a single one (68). Contacts of the fluorophore with the solvent cause quenching effects due to, e.g., motions of the fluorophore, a higher dielectric environment, or collisions with heavy atom quenchers added to the solution (69, 70). When a protein complex is formed, the quenching is reduced. Fluorescence anisotropy makes use of the effect that a fluorophore excited with polarized light emits polarized light. When the fluorophore is in contact with the solution, its tumbling will reduce the polarization of the emitted light. Upon binding of the interaction partner, the fluorophore will tumble more slowly and the polarization of the emitted light will increase. All of the above-mentioned parameters can be used to determine association and dissociation constants of protein–protein interactions. For studying the interaction of the actin modifying protein profilin with the C-terminus of the chloroplast outer envelope protein CHUP1 (CHUP1-CT), primary amine residues in profilin were labeled with the fluorophore dansyl chloride (32). After titration of CHUP1-CT or actin to dansyl-profilin, the increase in fluorescence intensity was measured in a fluorometer with an excitation wavelength of 337 nm. Emission was recorded at 450 nm. From the data, a binding curve could be generated that allowed the determination of dissociation constants as described in ref. 71. Fluorescence anisotropy was applied for the investigation of the interaction of cpSRP43 and LHCP, an interaction that is thought to be essential not only for preventing aggregation, but also mediating disaggregation of the substrate LHCP (72). For the interaction studies, a cysteine residue in LHCP was labeled with fluorescein-5¢-maleimide and the binding of fluorescein-labeled LHCP to different concentrations of cpSRP43, cpSRP54, cpSRP, and FtsY (the cpSRP receptor) was measured. Fluorescence was excited at 450 nm, and fluorescence anisotropy was recorded at 524 nm. An increase in fluorescence anisotropy could be detected upon addition of cpSRP43 or cpSRP. Fitting of the data as described in ref. 72 allowed the determination of apparent dissociation constants (KD) of 97 and 138 nM, respectively. The specificity of binding was verified by competition experiments with the L18 peptide, the binding site for SRP43 in LHCP.
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Inhomogeneities of the sample, if not recognized, can lead to misinterpretations of the data. In recent years, many new fluorescence-based techniques have evolved that allow the observation of interactions on the single-molecule level and have thus overcome this problem. These techniques are discussed in the next section. 4.5. Single-Molecule Approaches as a Future Perspective
The methods described above make it possible to qualitatively assess protein–protein interactions and (some of them) determine their kinetic and thermodynamic parameters. However, most cellular processes, such as protein translocation, involve multiple sequential or simultaneous steps, which are triggered by a variety of effector molecules. Due to the transient nature of these interactions, multi-step processes are often hidden in a “black box.” In a classical reductionistic approach, the interactions between individual proteins are investigated separately and the findings are integrated in a complex interaction network scheme (73). However, the limitations imposed by such reductionism are obvious – the interplay with other components involved in the process is gated out and it is almost impossible to obtain a time-resolved picture. The recently emerging single-molecule techniques open new perspectives for future analyses of such stepwise processes. In single-molecule fluorescence approaches, the interacting proteins and their effectors can be labeled with FRET-pair fluorophores, and the interaction can be monitored in real time by recording the fluorescence signals of individual molecules and calculating the resulting FRET efficiencies (74). It has to be noted, however, that the time resolution of the method is limited to 1 ms. Above that limit, the kinetic parameters of dynamic interactions can be determined (75). The work of Zhao and coworkers (76) demonstrated the potential of the method by analyzing the conformational states occurring during leucine transport by the LeuT protein, a bacterial membrane protein. To date, analysis of intermolecular interactions is technically limited to three colors (77, 78) but an extension to four (or more) steps appears to be, in principle, possible (79). Another fluorescence technique that has been applied on the singlemolecule level is fluorescence correlation spectroscopy (FCS) (80). FCS allows real-time measurement of the diffusion of fluorescent molecules by detecting fluorescence fluctuations and allows determination of molecular parameters such as diffusion coefficients, concentration, and molecular interactions. An improvement in sensitivity was achieved by the development of fluorescence crosscorrelation spectroscopy (FCCS). In FCCS, both interaction partners are labeled with different fluorophores. If the two proteins interact, a coincidence in movement of the two particles will be detected. The use of FCCS has been recently extended to the use of more than two colors. More advanced setups combine detection of single-molecule fluorescence with force-sensitive techniques such as magnetic or
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optical tweezers (81). In that way, the coincidence between force generation and the interaction of two (or more) fluorescently labeled molecules can be visualized. It is obvious that such systems might be applied to study the coupling of protein–protein interactions and/or nucleotide binding and hydrolysis cycles that are required for unidirectional passage of a precursor protein through the protein-conducting channel of the chloroplast envelope, or in other chloroplast processes. References 1. Chou, M.-L., Chu, C.-C., Chen, L. J., Akita, M., and Li, H.-M. (2006) Stimulation of transit-peptide release and ATP hydrolysis by a cochaperone during protein import into chloroplasts. J. Cell Biol. 175, 893–900. 2. Bédard J, Kubis S, Bimanadham S, and Jarvis P. (2007) Functional similarity between the chloroplast translocon component, Tic40, and the human co-chaperone, Hsp70-interacting protein (Hip). J. Biol. Chem. 282, 21404–21414. 3. Pasch, J. P., Nickelsen, J., and Schünemann, D. (2005) The yeast split-ubiquitin system to study chloroplast membrane protein interactions. Appl. Microbiol. Biotechnol. 69, 440–447. 4. Sprinzak, E., Sattath, S., and Margalit, H. (2003) How reliable are experimental protein-protein interaction data? J. Mol. Biol. 327, 919–923. 5. Maple, J., and Møller, S. G. (2007) Yeast twohybrid screening. Methods Mol. Biol. 362, 207–223. 6. Garcia-Cuellar, M. P., Mederer, D., and Slany, R. K. (2009) Identification of protein interaction partners by the yeast two-hybrid system. Methods Mol. Biol. 538, 347–367. 7. Kittanakom, S., Chuk, M., Wong, V., Snyder, J., Edmonds, D., Lydakis, A., Zhang, Z., Auerbach, D., and Stagljar, I. (2009) Analysis of membrane protein complexes using the splitubiquitin membrane yeast two-hybrid (MYTH) system. Methods Mol. Biol. 548, 247–271. 8. Stagljar, I., and Fields, S. (2002) Analysis of membrane protein interactions using yeastbased technologies. Trends Biochem. Sci. 27, 559–563. 9. Rahim, G., Bischof, S., Kessler, F., and Agne, B. (2009) In vivo interaction between atToc33 and atToc159 GTP-binding domains demonstrated in a plant split-ubiquitin system. J. Exp. Bot. 60, 257–267. 10. Brymora, A., Valova. V. A., and Robinson, P. J. (2004) Protein-protein interactions identified by pull-down experiments and mass spectrometry. Curr. Protoc. Cell Biol. 22, 17.5.1–17.5.51.
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of MARCKS-related protein (MRP) in solution. Journal Biol. Chem. 271, 26794–26802. 72. Jaru-Ampornpan, P., Shen, K., Lam, V. Q., Ali, M., Doniach, S., Jia, T. Z., and Shan, S.-O. (2010) ATP-independent reversal of a membrane protein aggregate by a chloroplast SRP subunit. Nature Struct. Mol. Biol. 17, 696–703. 73. Sommer, M. S., and Schleiff, E. (2009) Molecular interactions within the plant TOC complex. Biol. Chem. 390, 739–744. 74. Roy, R., Hohng, S., and Ha, T. (2008) A practical guide to single-molecule FRET. Nat. Methods 5, 507–516. 75. Mickler, M., Hessling, M., Ratzke, C., Buchner, J., and Hugel, T. (2009) The large conformational changes of Hsp90 are only weakly coupled to ATP hydrolysis. Nat. Struct. Mol. Biol. 16, 281–286. 76. Zhao, Y., Terry, D., Shi, L., Weinstein, H., Blanchard, S. C., and Javitch, J.A. (2010) Single-molecule dynamics of gating in a
neurotransmitter transporter homologue. Nature 465, 188–193. 77. Hohng, S., Joo, C., and Ha, T. (2004) Singlemolecule three-color FRET. Biophys J. 87, 1328–1337. 78. Lee, N. K., Kapanidis, A. N., Koh, H. R., Korlann, Y., Ho, S. O., Kim, Y., Gassman, N., Kim, S. K., and Weiss, S. (2007) Three-color alternating-laser excitation of single molecules: monitoring multiple interactions and distances. Biophys. J. 92, 303–312. 79. Heilemann, M., Tinnefeld, P., Sanchez Mosteiro, G., Garcia Parajo, M., Van Hulst, N. F., and Sauer, M. (2004) Multistep energy transfer in single molecular photonic wires. J. Am. Chem. Soc. 126, 6514–6515. 80. Haustein, E., and Schwille, P. (2004) Singlemolecule spectroscopic methods. Curr. Opin. Struct. Biol. 14, 531–540. 81. Mickler, M., Schleiff, E., and Hugel, T. (2008) From biological towards artificial molecular motors. Chemphyschem 9, 1503–1509.
Part II Omics and Large-Scale Analyses
Chapter 6 Proteome Databases and Other Online Resources for Chloroplast Research in Arabidopsis Diogo Ribeiro Demartini, Célia Regina Carlini, and Jay J. Thelen Abstract Proteomics aimed at addressing subcellular fractions, such as chloroplasts, are a complex challenge. In the past few years, several studies in different laboratories have identified and, more recently, quantified, thousands of proteins within whole chloroplasts or chloroplast fractions. A considerable number of these studies are available for querying, using online resources, such as databases containing the proteins identified, encoding genes, acquired spectra, and phosphopeptides. The main purpose of this review is to identity and highlight useful features of these online resourses, mainly focused in proteomics databases related to chloroplast research in Arabidopsis thaliana. Several web sites were consulted. Among them, 11 were selected and discussed herein. The databases were classified into Plastid Databases, General Organelle Proteome Databases, and General Arabidopsis Proteome Databases. Special care was taken to present information regarding protein identification, protein quantification, and data integration. A selected list of online resources is presented in two tables. The databases analyzed are a useful source of information for researchers in the plastid organelle and plant proteomics fields. Key words: Arabidopsis, Databases, Proteomics, Chloroplasts, Mass spectrometry, Organelle
1. Introduction Proteomics is a powerful approach to identify thousands of proteins en masse and is, therefore, particularly useful for comprehensive characterization of in vitro-purified organelles such as plastids (1–5) which contain approximately 80–100 plastid-encoded proteins and about 2,500–3,500 predicted nucleus-encoded proteins (4). Plastids are “cell-specific organelles” which acquired specialized functions during the evolutionary development of this organelle
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(6). In addition to studies on whole plastids, several publications report the protein composition of sub-plastidial fractions (4, 5, 7– 9). In nearly all of these studies, leaf chloroplasts are the system du jour, undoubtedly because there is little difficulty purifying them to homogeneity in milligram quantities necessary for near-complete cataloging of proteins. Presently, large-scale studies using proteomics are moving past the simple identification of proteins within a cell or an organelle. Sophisticated, quantitative comparisons are now commonly being performed to elucidate the dynamics of complex regulatory networks and pathways within “omics” datasets (10). Proteomics methodologies are able to access a large set of events within cellular and subcellular compartments, clarifying the associated phenomena under study at that time (10). In a typical bottom-up proteomic study, the workflow consists of extraction of intact proteins, optional fractionation (e.g., SDS–PAGE, twodimensional gels [2D gels]), protein digestion, and liquid chromatography typically online to tandem mass spectrometry (LC-MS/MS). MS/MS spectra must then be mined using one of several possible search algorithms against a translated genome or protein database (11). As with all “omics” approaches, data mining and bioinformatics are essential to proteomics; this is perhaps the most rapidly evolving aspect of proteomics workflows. Despite ongoing advancements at each of the steps, analysis of all proteins in an organelle or organism is still a challenging task (12). Complex proteomics workflows, and the integration and interpretation of comparative proteomics data were deeply explored by Lisacek et al. (11). Bioinformatics tools for proteomics can be used for qualitative (protein cataloging) and quantitative (protein quantitation) data analysis, and such analyses are becoming more reliable with the advance of higher resolution mass spectrometers (13). Dissemination of these data is now routinely achieved through online database resources, and several published studies on chloroplasts are available in public web databases. Depending on the volume and type of data contained in the databases, it is possible to compare and describe complex metabolic pathways and biological processes. Using this approach, for example, details of the plastidial isoprenoid (4) and lipid metabolic pathways (5) are being elucidated with the recent mapping of several low-abundant proteins within these databases. The main purpose of this review is to explore databases related to plastid research in Arabidopsis thaliana. Since the number of databases that exclusively fit this criterion was not expansive, we decided to extend this review to include Arabidopsis databases pertaining to proteomics and other plant organelles. The key points of each database are highlighted and a list of links is provided.
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2. Methods 2.1. Database Searches
The search for Arabidopsis scientific databases was performed using Scopus™ (http://www.scopus.com), PubMed (http://www.ncbi. nlm.nih.gov/pubmed/), ISI Web of Knowledge™ (http://www. wokinfo.com), and Google™ (http://www.google.com). Keywords used for searching were “chloroplast,” “Arabidopsis,” “database,” “on-line,” “proteomic,” “mass spectrometry,” “knockout,” “metabolomic,” “microarray,” and “organelle.” Boolean operators AND and OR were used for all except Google™ searches. Publication titles, abstracts, and keywords were analyzed to further explore relevant results. The papers or web sites (in case of Google) retrieved containing the desired words were analyzed. Several databases were retrieved but not analyzed since they were not related to the main purpose of this chapter. Publications related to the release, or important updates, of the databases presented were selected. Finally, the databases were selected in this order of priority: –– Plastid Proteome Databases: databases related to chloroplast proteome studies, primarily pertaining to Arabidopsis thaliana. –– Plant Organelle Databases: databases devoted to plant organelle proteomics. In this case, Arabidopsis thaliana was one of many plants studied. –– General Arabidopsis Proteome Databases: databases devoted to proteome studies using Arabidopsis thaliana. Databases were not specifically devoted to chloroplasts or organelle proteomics. Phosphoproteomics-related databases are included in this section.
2.2. Database Analyses
Databases that were relevant to this review were analyzed and summarized, emphasizing unique or interesting features. A brief introduction for each one is provided. The experimental approaches used either for plastid purification or for building the databases are briefly explained, when it was critical for the results obtained. All databases presented were carefully queried and analyzed for functionality, exploring the features presented in each main menu. Each database’s menus, and links contained therein, are italicized in this chapter for improved clarity. The first databases presented are the Plastids Databases, followed by General Organelle Databases, and finally General Arabidopsis Proteome Databases. Particular focus is given to proteomics databases.
3. Plastid Databases 3.1. AT_CHLORO Database (14)
(http://www.grenoble.prabi.fr/at_chloro/) The AT_CHLORO database condenses information from 1,323 nonredundant proteins identified from purified subfractions
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of Arabidopsis thaliana chloroplasts. Purification of the chloroplast subcompartments and the number of proteins detected is briefly described. For further experimental details, see refs. 15, 16. Samples from three plastid subcompartments (envelope, thylakoids, and stroma) were either digested in solution or pre-fractionated by standard SDS-PAGE, prior to trypsin digestion and MS analyses (14). Proteins detected in the envelope, stroma, and thylakoid fractions were quantified by label-free spectral counting. Analysis of chloroplast envelopes resulted in the identification of 487 proteins, combining the results from a non-treated extraction procedure and a NaOH-treated membrane fraction. This prefractionation approach proved beneficial as it is well known that high-abundance proteins can mask minor proteins. To ensure that minor proteins could be detected in the stroma fraction, ammonium sulfate precipitation was performed to remove RuBisCO (40–50% of stroma proteins from the stroma fraction) which was recovered in the 60% salt cut and analyzed separately. A total of 260 LC-MS/MS analyses were performed only for the stromal portion of this study. In total, 483 proteins were identified in the stroma fraction. In the case of thylakoids, either alkaline or organic solvent extraction protocols (15, 16) were employed. A total of 63 samples were obtained and 129 LC-MS/MS analyses performed. In total, 220 thylakoid proteins were identified. On the main page of the AT_CHLORO database, there is a brief description about the experiments performed to build the database, important references, and contact information. The user has the options to search for one particular protein or a set of proteins by loading the protein accession number or name. The AT_CHLORO database top main menu contains six options: All, Envelope, Stroma, Thylakoids, Search Result, and Search. When a search is performed clicking one of these options, the web page will reload and present a table listing information about the proteins detected in that plastid subfraction. Among 19 different characteristics for each protein include the accession number, protein length (number of amino acids), and abundance based on spectral counts. Partial information for the same protein can be retrieved from different sources, such as The Arabidopsis Information Resource (17), TAIR (localization and description; http://www.tair.org); the Plant Proteome Database (PPDB) (1) (Pi, molecular weight, curated localization; http://ppdb.tc.cornell.edu/); and Plant Membrane Protein Database (18), Aramemnon (number of transmembrane helices; http://aramemnon.botanik. uni-koeln.de/). Figure 1 presents an example when the Envelope menu is accessed. One interesting feature of this database is that it compares the percentage of localization for each protein based on spectral counting, and presents the total spectral count for a given protein (Fig. 1: ENV SC; STR SC; THY SC). From this page, it is possible to export data in five different formats: *.csv (comma-separated
Fig. 1. Screenshot of a result page listing some of the proteins extracted from chloroplast envelopes deposited in the AT_CHLORO database (http://www.grenoble.prabi.fr/at_chloro/). Spectral counting (SC) quantification is presented in such a way that the protein quantification can be compared among the envelope (ENV), stroma (STR), and thylakoid (THY) subfractions.
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values), *.xls (workable in Microsoft Excel™, for example), *.ods (OpenDocument Spreadsheet), *.pdf (portable document format), and *.xml (eXtensible Markup Language). Once the user selects one particular protein, the next window displays the curated annotation for the protein. In this page, the database gives full information such as description, putative function, localization, quantification (spectral count), and references (without links). The database presents the nonredundant peptides for that protein, including Mascot™ score, retention time, monoisotopic mass, amino acid sequence, and number of MS/MS counts for each peptide. On the top of this page, information about this protein from different databases can be obtained (a new window opens) in their respective links. The AT_CHLORO database provides an easy way to compare the abundance of proteins in different chloroplast subfractions. 3.2. Chloroplast Function Database (19)
(http://rarge.psc.riken.jp/a/chloroplast/) The Chloroplast Function Database provides a large-scale collection and phenotype analysis of single gene knockout lines for nucleus-encoded plastid proteins (19). Specifically, the database contains molecular and phenotype analysis of 3,246 lines containing the Ds/Spm (Dissociator/Suppressor–mutator) transposon or T-DNA-tagged insertions which disrupt the protein-coding regions of 1,369 proteins. Analyses were performed with 3-week-old Arabidopsis seedlings grown in agar plates. Detailed information regarding the database are available in the paper published by Myouga et al. (19). It is possible to search by keyword, locus ID, line ID (typing one option), or phenotype (selecting one or more of the available options). Regardless, the result page is a table containing the line number, identification of the disrupted locus, and protein/phenotype descriptions. Accessing the Phenotype Number hyperlink presents the identification information, containing the locus number, descriptions, and links to TAIR and Arabidopsis thaliana transfactor and cis-element prediction database (ATTED-II; http:// atted.jp/) (20). The tests presented in the result page are divided according to the generation analyzed (F3, F4, etc.) or in case of T-DNA-tagged lines, by the result of PCR tests. The number of plants with visible phenotypes, plants resistant to antibiotics, and plants sensitive to antibiotics is presented in the Ratio row, respectively. Phenotype classification of the mutants is shown in the Trait row and in Remarks row, containing additional comments about the mutants. In some cases, images are available and can be viewed in high resolution by clicking on them (Fig. 2). The homozygous insertion lines are available through the Arabidopsis Biological Resource Center (ABRC, http://abrc.osu.edu/) and RIKEN BioResource Center (http://www.brc.riken.jp/inf/en/index. shtml). Finally, the user has the option to download all the data contained in the database, from the List menu.
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Fig. 2. Partial screenshot of a result page from the Chloroplast Function Database (http://rarge.psc.riken.jp/a/chloroplast/). Protein description, links, and phenotype images are presented. In this example, the search was done by keyword, typing in one line number, which is indicated on the top.
3.3. The Plastid Protein Database (21)
(http://www.plprot.ethz.ch/) The Plastid Protein Database (plprot) comprises information from proteomes of different plastid types analyzed in several largescale proteomics experiments. The first version of the database had 2,043 proteins (21). The source of chloroplasts was Arabidopsis thaliana leaves, 7 weeks old (22, 23). The proteins from etioplasts deposited in the database were extracted from red bell pepper (Capsicum annuum L.) fruits (24), and the proteins from proplastids were obtained from tobacco BY-2 cell culture (25). The user can obtain experimental information about the plastids analyzed, by clicking on the respective plastid subtype in the main page. There are two modules: “Tools for proteome data integration” and “Select database to search in.” In “Tools for proteome data integration,” the user has the option to Basic Local Alignment Search Tool (BLAST) (26) one query sequence or upload a file with multiple sequences. The result page of a batch BLAST search is not exportable. In the next module, “Select database to search in,” it is possible to search for proteins among all plastid proteins deposited in the database. The user can choose the organism that will be used for the search query. The database being searched is always shown at the top of each page in this module. Protein keyword can be input and upon completion of the search, a new window opens with the information available for that particular protein. A maximum of
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eight fields are presented in the output page, including Gene ID, TargetP predictions (http://www.cbs.dtu.dk/services/TargetP/) (27), functional categories, and amino acid sequences. However, there is no option to export the data. In this module, there is a 2D-gel electrophoresis etioplast/chloroplast proteome map. The search for a target protein will result in a highlighted spot in a 2D gel. Using the option to increase the display resolution makes the spots more visible. When the result page for a particular protein is presented, the best homologs in the different plastid subtypes are also presented. 3.4. The Plant Proteome Database (1, 28)
(http://ppdb.tc.cornell.edu/) Originally, the PPDB was devoted to proteins from peripheral and integral thylakoid membranes from Arabidopsis thaliana leaf chloroplasts. Data on 159 proteins were compiled in the first release of the database (2004). Reverse-phase HPLC and 1D and 2D electrophoresis gels followed protein extraction and peptide fractionation (salts, detergents, and organic solvents) (1). Nine different options, shown in the left menu, are available to search in the database: Accession, Gene Name & Annotations, Proteome Experiments, Comparative Proteomics, Subcellular Proteomes,
Fig. 3. Screenshot of The Plant Proteome Database (PPDB, http://ppdb.tc.cornell.edu/). The image shows the search window obtained when a user clicks on Subcellular Proteomes, on the left menu. In this case, the number in parentheses beside the plastid subfraction name indicates the number of proteins in that fraction. Users can access the specific location by clicking on the relevant small circle.
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Protein Function, Proteomics Publications, Post-Translation Modifications, and Biochemical Pathways (Fig. 3). The Accession option allows the user to search using accession IDs for Arabidopsis thaliana and Zea mays. The gene/protein name formats accepted are AtGI (from TAIR, Arabidopsis thaliana Gene Index) and ZmGI (from TIGR, The Maize Gene Index). In this window, it is possible to choose several options for the output table. Detailed explanations regarding the Display Options are available when the user clicks on the question mark (?). Under Proteome Experiments, it is possible to select one of five species, two cell types, and two subcellular fractions (thylakoids or stroma). The Display Options are the same as those in Accession. The result window either for Proteome Experiments or for Accession is the same, and displays the standard annotation and accession number of the proteins that matched the search criteria. Once the user selects one protein, another window opens and shows additional information from ten different prediction tools, such as TargetP, Predotar (http://urgi.versailles.inra.fr/predotar/predotar. html) (29), PFAM (http://pfam.sanger.ac.uk) (30), and others. There are links to different databases for that particular query ID, which are TAIR, POGs/PlantRBP (Putative Orthologous Groups), http://pogs.uoregon.edu/ (31), AtProteome (http://fgcz-atproteome.unizh.ch/) (32), SUBA II (SUB-cellular location database for Arabidopsis proteins, http://suba.plantenergy.uwa.edu.au/) (33), and PhosPhAt (The Arabidopsis Protein Phosphorylation Site Database, http://phosphat.mpimp-golm.mpg.de/) (34, 35). In addition, related genes from Oryza sativa and Z. mays are presented with their respective percent identities, similarities, E-values, and TargetP prediction, if applicable. The Details of the experimental sources is a useful feature of this database, showing a number which refers to the experimental approach used when the protein was detected, presented in a table. There is a link corresponding to each number, which explores in more detail that particular experiment. It is possible to click on Spot and ask for Details, retrieving the peptides identified for the selected protein. A pop-up window gives the protein sequence, with identified peptides highlighted and showing the predicted chloroplast transit peptide underlined. In cases where the protein was identified in several different experiments, the data with the five highest Mascot scores are presented. There is information on the experimental identification (Details of experimental sources), number of unique peptides, and the biological sample where the protein was detected.
4. General Organelle Databases 4.1. The Plant Organelle Database 2 (36, 37)
(http://podb.nibb.ac.jp/Organellome/)
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The Plant Organelle Database 2 (PODB2) is an image and movie database. It contains microscopic pictures and movies from cellular structures (organelles). There is information from 25 organelles, including plastids from Arabidopsis thaliana and other species. The objective is to understand organelle dynamics through high-quality videos and images. The available information comes from different studies in the organelle research field. There are 82 available videos (*.mov format), including 17 on plastids, among which nine are related to Arabidopsis thaliana plastids. The database displays five options in the main menu on the top of the page: Organelles Movie Database, Organellome Database, Functional Analysis Database, External Links, and Organelle Word (Japanese). It also provides a list of databases (external links) assembled into sub-menus: Genomics, Transcriptomics, Proteomics, Organelles, Analytical Tools, and others. In the Organelles Movie Database menu, it is possible to search by the organism (common name), organelle (by selecting within available options in a menu), gene accession number, and keyword. Once the results are presented in a refreshed window, the Video Titles are presented. When the user clicks on the Title, a new page opens, presenting a list of information about that particular experiment, including a brief description. Important data such as developmental stage, type of microscope used, frames per second, micrometer per pixel, colors of fluorescence probes, and publication source (shown on the bottom of the page) are displayed in this page. The Functional Analysis Database is a compilation of protocols applied to different experimental approaches. Biochemical assays, Detection and analysis of proteins, and Tissue and organelle isolation are part of a list with ten categories containing 91 different protocols. 4.2. The Arabidopsis Subcellular Database (33)
(http://suba.plantenergy.uwa.edu.au/) The Arabidopsis Subcellular Database (SUBA II) uses information provided by several different sources to predict subcellular location of proteins. Presently, there are 2,576 entries based on chimeric fusion studies, representing 1,674 distinct proteins from 826 publications. Additional data from 72 different publications add 10,143 entries based on subcellular proteomic studies, and comprises 4,685 distinct proteins. All cited publications can be downloaded in *.txt format (in fact, references are in Research Information Systems (RIS) format; in this case, the file can be imported into most reference manager software packages) or in *.xls format. The references are linked to each paper’s full text, through either ISI Web of Knowledge™ or PubMed. Currently, more than 6,000 nonredundant proteins can be accessed in SUBA II database. TAIR, Swiss-Prot (http://ca.expasy.org/sprot) (38), and AmiGO (http://www.geneontology.org/) (39) are used to extract protein annotation. To search the database, the user can choose among ten different prediction tools and 14 different subcellular
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fractions, including plastids. A list of references (from 1990 to 2010) is available in another searchable field. Combining keyword, prediction tool, reference, and all other possibilities that SUBA II offers, the user can cross-check all these available information using Boolean operators to retrieve one specific protein. This is a distinct advantage of this database. A detailed tutorial can be downloaded in *.pdf format from the Help menu. Once the query protein is submitted, the result page presents a table containing the AGI accession number, protein annotation (retrieved from TAIR), and 24 more options that can be selected by the user for each protein by clicking on its AGI number. The results can be saved in *.xls format. In a new window, the protein length, amino acid sequence, molecular mass, description, and subcellular localization are presented. Nearly all retrieved information are linked to a reference, a very helpful feature of this database. There are references describing subcellular localization for proteins based on MS/MS data, annotation results, and prediction tools. The result page also presents hydropathy plots, which allows visualization of hydrophobicity patterns over the length of specific peptide sequences (40). Finally, there is a link to the National Center for Biotechnology Information (NCBI) BLAST web page to monitor current annotation.
5. General Arabidopsis Proteome Databases 5.1. Arabidopsis Seed Proteome (41–45)
(http://www.seed-proteome.com/) The database is devoted to seed development studies using proteomics approaches. In the menu Protein Maps, a sub-menu, Seed Germination Proteins, presents different 2D gels, which map proteins in five different developmental stages of the seed, from 0 h up to 24 h of seed germination. There are five developmental stages analyzed: Constant during germination (0 h), Decrease during germination sensu stricto (0 h), Decrease during radicle protusion (0 h), Increase during germination sensu stricto (24 h), and Increase during radicle protusion (48 h). The first three have dry seed as reference, and the remaining stages are 1 and 2 days imbibed seeds, respectively. For each of these developmental stages, the user can click on a protein spot and a new window opens describing that particular protein, divided into four sections: Essential, Data, Sequences, and Coordinates. Information such as molecular masses and theoretical and experimental isoeletric point are displayed. There is a link to the search result for that particular protein (from Mascot), and the peptides detected with their respective masses are presented in the Sequences option.
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In the Protein Catalog menu, proteins are listed in alphabetical order, and the same information as retrieved from the 2D gels can be found in this link as well. Once a particular protein is selected, the same page with the protein’s information is loaded. Protocols in the main menu presents seven sub-menus with general information on 2D electrophoresis, protein staining techniques, de novo “synthesized” proteome, and preparation of protein extracts. Within the Protocols link, there is a long list of useful websites on proteomics resources, seed-related websites, and even news on international meetings. 5.2. AtProteome Database (32)
(http://fgcz-atproteome.unizh.ch/) A proteome map for Arabidopsis thaliana is presented in the AtProteome database. Six different organs/samples were analyzed (cell culture, flowers, leaves, roots, seeds, and siliques). Data were collected in a linear trap quadruple (LTQ)-ion trap mass spectrometer. As a result, more than 85,000 unique peptides deduced from about 790,000 collected mass spectra resulted in the identification of 13,029 proteins (32). There are four criteria to search for a particular protein, and there are two main windows displaying results. The user types the name of the protein of interest or one keyword into the Protein Search, Proteotypic Peptide Search, or Protein Quantification options. When the search is completed through Protein Search or Proteotypic Peptide Search, the first result window presents a table containing the gene model, description, molecular weight (MW), isoeletric point (pI), number of amino acids (RES), number or theoretical tryptic peptides (TTP), and the number of distinct peptides detected for the query protein (Qty Peptides). The user can input either the AtGI numbers (gene model) or keywords in the Protein Quantification module. In this case, after clicking Search, the protein quantification performed through the individual modified APEXfactor is presented in a color code (46), hierarchically clustered for each organ analyzed. Independent of the criteria adopted in the search, full information about the query protein will be displayed when the user clicks either Gene Model (AtGI number) or Description. The next window shows information on the Protein Sequence, Peptide Browser, Peptides, and Spectrum Summary. The Peptide Browser section containing full information results on a protein is particularly noteworthy (Fig. 4). The number of spectra identified for one particular peptide is indicated in a white-to-blue color gradient. The peptides are aligned to the cDNA, and coding sequence is presented in the same window, which is easily identified. A full list of identified peptides is presented as a table in the Peptides section. It is possible to select the organ analyzed to retrieve information about the protein. The table presents the peptide sequence, including its start position in the protein sequence and tissues in which it was identified and quantified. A bar graph summarizes all the information
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Fig. 4. Screenshot from the results window for RuBisCO Large Subunit (ATCG00490) in the AtProteome database (http:// fgcz-atproteome.unizh.ch/). The information displayed consists of the protein amino acid sequence of the first splice variant of the protein, a link section, the Peptide Browser, and the detected peptides sorted by their position in the protein (N-terminal end first, http://fgcz-atproteome.unizh.ch/help.pdf.).
presented in this table, including a spectrum count report for the query protein in all samples checked. If the user clicks in one particular peptide in the Peptide Browser section, it will be highlighted in the table (orange), and more information becomes available including the Spectrum Report for that peptide. In this case, a new list presents the samples where that peptide was detected and the peptide sequence, including modifications, in a code mode. The code can be found in the Help menu. Detailed information about the cross-correlation score (Xcorr), search result discriminated score (fval, PeptideProphet discriminant score), delta CN score (dCN), and false discovery rate (fdr) are given just below the peptide sequence. By clicking on the sequence, the MS/MS spectrum for that peptide is presented in a new window. Several options are available in the MS/MS spectrum window, such as exporting in different formats and editing the spectrum itself (or scaling). Beyond this, b and y ions are marked in the m/z axis, in a color code mode easily identified. By dragging the mouse over each
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individual peak in the MS/MS spectrum, m/z, intensity, and mass delta values are presented. The Proteotypic Peptide Search is another advantage of this database which displays unambiguously detected peptides for one particular protein of interest (47). The search criterion is the same as any other one previously detailed, but the result window will retrieve only a list of peptides (proteotypic) and the samples where they were detected. If the user clicks on one peptide for a query protein, the same detailed result window shows up containing the Peptide Browser, Peptides, Spectrum Summary, and Spectrum Information. The Help menu has more options for researchers interested in going deeper into this database. 5.3. The Arabidopsis Protein Phosphorylation Site Database (PhosPhAt) (34, 35)
(http://phosphat.mpimp-golm.mpg.de/index.html) The interactive PhosPhAt is devoted to Arabidopsis thaliana protein phosphorylation. It offers three ways to search for a particular protein: Basic Search, Advanced Search, and Prediction. In Basic Search, the user can input the accession number (AGI), peptide sequence, or protein description. In the case of Advanced Search, there are 12 “designated items” which can be cross-checked using Boolean operators. Among these items, the instrument, the tissue, and compartment can be chosen (plastids are not included as compartments). Once the user drags the designated item to the Query Panel, some options (Compartment, Enrichment, Instrument, Tissue, PubMed, and MapMan Bin) will present as a drop-down list. The database provides a powerful way to search the data, similar to the one provided by the SUBA II database. After the search is finished, the results identify the proteins by the accession number and list 12 characteristics including the phosphorylated peptide, precursor ion, and charge state. It is also possible to check which engine was used for search (Mascot™, SEQUEST™, etc.) with statistics related. If a spectrum and protein quantification are available, it will be indicated in the last columns on the right side (Fig. 5). Full information about the phosphorylated peptide and protein will be retrieved when the user clicks on the respective row with the results while still displaying the previous window. By clicking on the accession number, the next window brings the protein sequence and the phosphorylated peptides will be highlighted. Dragging the mouse over the highlighted amino acids retrieves the phosphorylation site score and PFAM domain information. In this page, a full description of the protein and links to SUBA II, TAIR, ATTED-II, Aramemnon, and GABI Primary Database (GabiPD, http://www.gabipd.org/) (48) are available. The results of all performed searches are kept as “worksheets” at the top of the page, which makes it simple to go back, if necessary.
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Fig. 5. Screenshot from a search result page obtained at The Arabidopsis Protein Phosphorylation Site Database (PhosPhAt, http://phosphat.mpimp-golm.mpg.de/index.html). Advanced search options are presented in the left menu. Phosphorylated peptides are initially listed, and further information is obtained by clicking on Details (as shown). Peptide parameters displays the instrument used for MS analysis, the tissue analyzed, and the enrichment performed (TiO2, GaO2, etc.), which can also be used for searching. In case one spectrum is available, it is presented. By clicking on Protein prediction, the amino acid sequence with the phosphorylated peptide(s) highlighted is shown.
5.4. Proteomics of Oilseeds Database (49, 50)
(http://www.oilseedproteomics.missouri.edu/) The database released in 2004 contains expression data of proteins during the seed filling stage of seed development for Arabidopsis thaliana, Brassica napus (oilseed rape), Glycine max (soybean), and Ricinis communis (castor). The top menu in the website presents the plants used in the study. A brief introduction about each plant is given once the user clicks on the respective plant. The methodology used to identify the proteins during the seed filling stage for all plants analyzed is also described. In the case of Arabidopsis thaliana seed filling, five stages were analyzed: 5, 7, 9, 11, and 13 days after flowering. Two different pI ranges, pI 4–7 and pI 3–10, were used for 2D difference gel electrophoresis (DIGE). There is a link to the DIGE gel and once the user clicks on this link, a high-resolution image of each gel is presented. In the 2D gel electrophoresis window, it is possible to check the protein annotation and accession number of each spot; it is presented in the top of the page. Full information about any protein in the gel can be obtained by clicking on it. A new window opens and displays the Expression Profile window, containing the Expression Data and Mass Spectrometry Data (LC-MS/MS). In this page, Expression Data presents the relative volume of the spot (average of three biological replicates), and the standard deviation for each particular protein along the developmental stages is presented. The information can be seen either in a time-course graph or in a time-
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course table. In the castor analyses, two 2D gels (pI 4–7 and pI 3–10) are presented, and the data displayed once the user clicks on a spot, as in the Arabidopsis thaliana case. The time course for castor samples was 2, 3, 4, 5, and 6 weeks after flowering. Oilseed rape and soybean were analyzed in a similar way, using high-resolution 2D-gel electrophoresis. The intervals of the analyses were from 2 to 6 weeks after flowering, in 1-week intervals. Two 2D gels are presented (pI 4–7 and pI 3–10), and similar to the Arabidopsis data, they contain information about the spots, both by hovering over the 2D spot and by clicking it, which reveals Expression and MS data. In this case, the results present peptide mass fingerprint assignments by both matrix-assisted laser desorption/ionization (MALDI-TOF) analyses and LC-MS/MS assignments. Additional information about the proteins includes molecular mass, pI, and the database used for the search, and in cases where the search retrieves an ortholog protein, the Species column presents it. In all plants analyzed, it is also possible to verify all proteins based on functional classification. At the bottom of each page, there is a clickable functional classification table. If the user selects one particular class, the detected protein is displayed within its functional class, and the spot number of the 2D gel or DIGE is presented. Finally, the database also offers to the user a set of protocols such as the popular phenol extraction protocol for plant proteins, protein quantification, 2D- and SDS-PAGE protocols, and trypsin digestion protocols. 5.5. Plant Protein Phosphorylation Database (51)
(http://digbio.missouri.edu/p3db/) The Plant Protein Phosphorylation Database (P3DB) condenses phosphoproteomics information from three different plants at this time: Arabidopsis thaliana, Medicago truncatula, and O. sativa. On the left side of the main page, there are eight different options: Search, Blast, Browse, Tools, Download, Submit, Help, and Contact. In the Search menu, it is possible to search for phosphoprotein, phosphopeptides, and protein description. In addition, the user can restrict the source organism and research group of reference. Since the database is still growing, it is being constantly updated as additional data become available. It is possible to employ the BLAST tool for either phosphoproteins or phosphopeptides within the database. The sequences must be inputted in FASTA format (a text-based type of file in which either base pairs or amino acids are represented in singleletter codes; each sequence contained in the file is divided into header and sequence itself). It is possible to select the organism as well and the desired E-value threshold for a phosphorylation hit. There are few examples that help the user employ this tool and interpret the output. After the BLAST search is completed, the result page presents one table with the results, sorted by E-threshold
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Fig. 6. Screenshot of a search result page of the Plant Protein Phosphorylation Database (P3DB, http://digbio.missouri.edu/ p3db/). Full information about each particular protein can be obtained when the user clicks on the protein Description. The phosphopeptide position within the amino acid sequence will be presented.
value (smallest to largest). As in a regular BLAST, a Query protein is aligned with the Subject protein; in addition, the phosphorylated amino acid is highlighted in red. It is possible to click on the phosphopeptide presented in the results page, after the BLAST search is completed, and a new window containing the Peptide Source, Reference (linked to NCBI), Protein Description, Charge State, and Mass Error opens. Access to the mass spectrum that led to the phosphorylation site assignment is also available. The Browse option brings two options to the user: list all phosphoproteins and list phosphoproteins according to the organism (Fig. 6). In either case, the result window is the same as the one provided in the Search result page, in which the protein is listed with its Annotation, Peptide Source, Publication Source, and Accession. The entire database can be downloaded for in-house applications, and several tools are available (http://digbio.missouri. edu/p3db/tools/), such as DecoyDBCreator, a decoy database creator; PhosSite, which helps processing large-scale phosphoproteomics data; P3-Motif, a phosphorylation motif prediction tool; and SpotLink, a 2D-gel web-based creator. In summary, the P3DB archives all published plant protein phosphorylation data acquired from high-resolution mass spectrometry instrumentation, and unpublished phosphorylation sites from devel-
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oping Arabidopsis seed. The noteworthy features of this site include the successful archiving of high-quality protein phosphorylation data from multiple plant species and the ability to BLAST search a protein for “similar” or conserved phosphorylation sites.
6. Concluding Remarks The main purpose of this chapter was to provide a collection of web database resources to support biochemical and cell biological research for the model plant Arabidopsis thaliana. Also, we provide a simple but directed analysis of each database presented, with emphasis on unique features of each site. Table 1 summarizes the
Table 1 Databases analyzed in this chapter* Database URL addresses and contacts
References
Organisms
Key features
Arabidopsis Seed Proteome http://www.seedproteome.com/
(41–45)
A. thaliana; B. vulgaris
2D gels from different developmental stages of seed germination Link to Mascot results List of protocols presented
AT_CHLORO Database http://www.grenoble.prabi. fr/at_chloro/, norbert.
[email protected], myriam.
[email protected]
(14)
A. thaliana
Data can be presented according to plastid subfraction Presents the peptides detected and compares one protein in different fractions based on spectral counting Retention time for each peptide is shown
AtProteome http://fgcz-atproteome. unizh.ch/,
[email protected]
(32)
A. thaliana
Protein quantification and spectrum report for all peptides, including modification when presented Peptide browser Complete information exportable Search for proteotypic peptide
Chloroplast Function Database http://rarge.psc.riken.jp/ a/chloroplast/,
[email protected]
(19)
A. thaliana
Search according to the phenotype High-resolution pictures of different A. thaliana phenotypes Although brief, Help menu is complete All data can be downloaded
Plant Protein Phosphorylation Database (P3DB) http://digbio.missouri.edu/ p3db/, thelenj@missouri. edu,
[email protected]
(51)
A. thaliana; M. truncatula; O. sativa
Database can be downloaded Integrates multiple phosphorylation site studies Phosphopeptide BLAST browser (continued)
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Table 1 (continued) Database URL addresses and contacts
References
(49, 50) Proteomics of Oilseeds Database http://www.oilseedproteomics. missouri.edu/
Organisms
Key features
A. thaliana; B. napus; R. communis
Three oilseed plants analyzed in a large-scale 2D-gel approach Proteins functionally classified Clickable spots in 2D gels of all stages of development analyzed
The Arabidopsis Protein Phosphorylation Site Database (PhosPhAt) http://phosphat.mpimpgolm.mpg.de/index.html/
(34, 35)
A. thaliana
Devoted to phosphoproteomics in A. thaliana Spectrum and quantification presented Phosphopeptide(s) indicated in the protein sequence Powerful search engine
The Arabidopsis Subcellular Database (SUBA II) http://suba.plantenergy.uwa. edu.au/, hmillar@cyllene. uwa.edu.au, tontis@iinet. net.au,
[email protected]. edu.au
(33)
A. thaliana
Powerful search criteria which allow the user easily find what is being searched for 14 organelles, including plastids References are fully linked and can be downloaded in *.xls or *.txt (RIS) formats
The Plant Organelle Database (36, 37) 2 (PODB2) http://podb.nibb.ac.jp/ Organellome/, podb@ nibb.ac.jp
A. cepa; A. thaliana; N. tabacum; P. patens; S. oleracea
25 organelles, including plastids High-quality videos and pictures with linked references Presents a long list of useful websites
The Plant Proteome Database (1, 28) (PPDB) http://ppdb.tc.cornell.edu/,
[email protected], qisun@ tc.cornell.edu
A. thaliana; Z. mays
Results from prediction tools are linked to their websites There are 11 organelles including plastids that are fractionated into several subfractions Links to references Ability to BLAST protein sequences Full explanation about the database in the index page
The Plastid Protein Database (plprot) http://www.plprot.ethz.ch/
A. thaliana; C. annuum; N. tabacum; O. sativa
Brief experimental approaches for all plastids isolated BLAST query protein or FASTA file against the database Easy plastid-type comparison
(21)
* The web site addresses and contact information (extracted from the main pages or contact pages) are presented in the first column. Respective references are indicated in the second column. Organisms contained in the databases are shown in the third column. The Key features column presents database highlights
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Table 2 Websites referenced in this article* Database
References
Description
Arabidopsis Coexpression Data Mining Tools http://www.arabidopsis.leeds. ac.uk/act/index.php
(48)
Data mining tools to analyze gene coexpression in A. thaliana
ARTRA – Arabidopsis and Transcriptome Microarray http://artra.kazusa.or.jp/
(52)
Contains transcriptomics sequences and gene-specific sequences for DNA microarray and RNAi knockdown
AtPIN – Arabidopsis thaliana Protein Interaction Network http://bioinfo.esalq.usp.br/atpin/
(53)
Database devoted to protein–protein interactions. It integrates five interaction datasets with TAIR and SUBA II
CATMA – The Complete Arabidopsis Transcript Microarray http://www.catma.org/
(54)
Comprehensive DNA tag repertoire for protein-encoding genes
ChloroplastDB: The Chloroplast Genome Database http://chloroplast.cbio.psu.edu/
(55)
Contains genomic protein, DNA and RNA sequences, gene locations, RNA-editing sites, putative protein families, and alignments for fully sequenced plastid genomes
ExPASY Proteomics Server http://www.expasy.ch/
(38)
Several tools and databases for proteomics research
GabiPD – GABI Primary dabatase http://www.gabipd.org/
(48)
The main aim of the database is to integrate and make available through the web the relevant primary data from the GABI projects
TAIR – The Arabidopsis Information Resource http://www.arabidopsis.org/
(17)
Database of genetics and molecular biology for Arabidopsis thaliana. Several tools available and exportable
The Arabidopsis Unannotated Secreted Peptide Database http://peptidome.missouri.edu/
(56)
Identification of candidate peptide-encoding genes currently not annotated
*References and a brief description of each site are listed
information about the databases analyzed in detail. Table 2 provides a list of useful websites related to proteomics approaches, not only for chloroplasts but also for other organelles or for Arabidopsis thaliana as the organism of study. We hope the information compiled will be useful for both beginners and experienced researchers in plant proteomics.
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Chapter 7 Use of Transcriptomics to Analyze Chloroplast Processes in Arabidopsis Tatjana Kleine and Dario Leister Abstract The vast majority of the several thousands of chloroplast proteins are encoded by nuclear genes. Regulation of their expression involves control of their transcription, and thus requires the transmission of information from chloroplast to nucleus (retrograde signalling). The most powerful approach to the analysis of the transcriptional regulation of chloroplast functions involves RNA hybridization to microarrays representing almost all nuclear genes of Arabidopsis thaliana, followed by statistical data analysis. This chapter provides detailed protocols for the preparation of RNA for microarray experiments, in particular the widely used Affymetrix ATH1 array. Finally, the use of the publicly available program Robin for statistical data analysis, as well as approaches to confirm microarray data, is introduced. Key words: Arabidopsis, Chloroplast, Retrograde signalling, Affymetrix, Microarray analysis
1. Introduction Chloroplasts are endosymbiotic descendants of cyanobacteria-like prokaryotes (1). Owing to the massive relocation of organelle genes to the nucleus during evolution, the majority of their several thousand different proteins are now encoded by nuclear genes; for a review, see ref. 2. The relatively few genes remaining in the chloroplast code for proteins involved in organelle gene expression (OGE) and photosynthesis. Hence, chloroplast multiprotein complexes, such as ribosomes and photosystems, are actually mosaics of subunits encoded by nuclear and chloroplast genes. Therefore, coordination of gene expression between the two compartments is required to ensure appropriate assembly of the multiprotein complexes. Mechanisms have evolved that permit direct control of OGE by the products of nuclear genes (“anterograde signalling”). These, in turn, rely on “retrograde signalling” from the chloroplast, which conveys information on the organelle’s developmental R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_7, © Springer Science+Business Media, LLC 2011
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and metabolic state to the nucleus. In consequence, nuclear gene expression (NGE) is modified in accordance with the physiological status of the chloroplast; for reviews, see refs. 3–5. In addition to its photosynthetic function, the chloroplast is involved in the metabolism of numerous essential compounds, including amino acids, fatty acids, lipids, nucleotides, and vitamins. It is the site of synthesis of tetrapyrroles (such as chlorophyll, heme, and the cofactor of phytochromes), and of isoprenoids and carotenoids, such as xanthoxin, the precursor of abscisic acid. Exposure to stresses, e.g., high light levels, can lead to increased accumulation of reactive oxygen species, predominantly in chloroplasts, mitochondria, and peroxisomes (6). Disturbances in any of these processes may trigger retrograde signalling. Moreover, it is well documented that mutations affecting structural components of the photosynthetic machinery, thylakoid electron flow, or chloroplast functions in general give rise to changes in the transcription of nuclear and chloroplast genes (7–13). The crucial questions are as follows: (1) how is the physiological state of the chloroplast monitored, and (2) how is this information transduced into alterations in NGE and OGE? In spite of more than 25 years of research (14, 15), the fundamental mechanisms underlying chloroplast signalling remain elusive. Very few primary target genes of chloroplast signalling have yet been identified. Furthermore, signalling mutants and marker genes are lacking that would enable one to discriminate between the different possible pathways and work out how they interact. In addition, it is not known whether NGE and OGE are regulated sequentially or in parallel. To dissect how NGE and OGE are influenced by interorganellar signalling networks, extensive microarray analyses of appropriate mutants and states are required, together with approaches employing inducible RNA interference (RNAi) to transiently induce lesions in specific chloroplast functions. Initially, custom-made gene-sequence-tag macroarrays, such as one covering 2,661 nuclear genes for chloroplast proteins and 631 genes encoding non-chloroplast proteins (16), were employed to exhaustively characterise gene expression changes in wild-type Arabidopsis plants exposed to various conditions and stresses, and in mutants defective in diverse chloroplast functions (16, 17). Nowadays, commercially available high-density oligonucleotide microarray systems, such as the GeneChip® Arabidopsis ATH1 Genome Array, designed in collaboration with The Institute for Genomic Research (TIGR) and containing more than 22,500 probe sets that permit the simultaneous detection of approximately 24,000 transcripts (http://www.affymetrix.com), constitute a powerful technology for global gene expression profiling and are a standard method in nearly every laboratory. When it comes to data analysis, the researcher is confronted with an embarras de richesses. Various statistical methods and programs for processing the raw data and analyzing the results exist.
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The statistics environment R (http://www.r-project.org) provides a wide variety of tools (including classical statistical tests, timeseries analysis, classification, and clustering) and graphical techniques. R, together with the BioConductor project (http://www. bioconductor.org) (18), provides a powerful platform for microarray data analysis and quality assessment. However, BioConductor operates via a text console and sophisticated analysis requires programming skills. Other microarray analysis programs such as GeneSpring or GeneMaths XT are only available commercially and might be too expensive for users who need to process relatively few data sets. Most publicly available programs are either not intuitive, offer only parts of the analysis workflow or are Web-based. Among the intuitive, non-commercial software packages available for statistical analysis of microarray expression data are FlexArray (http:// genomequebec.mcgill.ca/FlexArray) and the recently published program Robin (19). While FlexArray is a Microsoft Windows software package, the developers of Robin also offer a MacOSX installer package. Robin provides a Java-based graphical user interface to the R/BioConductor functions for the analysis of both two-colour and single-channel microarrays, and the user is guided through all steps of the analysis including quality assessment, evaluation, and experiment design. Here, we describe how to prepare RNA samples for array analysis and apply Robin to data analysis, and how to confirm directly the validity of microarray results.
2. Materials 2.1. Plant Growth and Harvesting
1. Arabidopsis thaliana wild-type (and mutant) seeds. 2. Greenhouse or growth chamber. 3. Soil and fertilizer. 4. Multitrays.
2.2. RNA Preparation
1. Mortar and pestle. 2. Liquid nitrogen. 3. RNeasy Plant Mini Kit (Qiagen, Hilden, Germany). 4. RNase-Free DNase Set (Qiagen). 5. Spectrophotometer.
2.3. Quantification and Integrity Assessment of the RNA
1. 50× Tris–acetate–EDTA (TAE) buffer: 2 M Tris base, 2 M acetic acid, and 50 mM ethylenediaminetetraacetic acid (EDTA). 2. 5× Gel loading buffer: 15% (w/v) Ficoll, 50 mM EDTA, 0.5% (w/v) sodium dodecyl sulphate (SDS), 1/50 vol. 50× TAE, 0.025% (w/v) bromophenol blue, and 0.025% (w/v) xylene cyanol.
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3. 5 mg/mL Ethidium bromide. 4. Agarose gel electrophoresis apparatus. 5. Gel documentation system. 2.4. Analysis of Microarrays: Identification of Differentially Expressed Genes Involved in Chloroplast Processes
1. The open-source software MapMan (http://mapman.gabipd. org/web/guest/mapman-version-3.5.0) (20).
2.5. Confirmation of Microarrays: Real-Time PCR Analysis
1. iScript cDNA Synthesis Kit (Bio-Rad Laboratories, Hercules, CA, USA).
2. The open-source microarray processing software Robin (http://mapman.gabipd.org/web/guest/robin) (19). 3. Spreadsheet program (e.g., Microsoft Excel).
2. RNase-free H2O (e.g., that of the RNeasy Plant Mini Kit). 3. 2× SYBR Green Supermix (e.g., Bio-Rad Laboratories). 4. Primer pairs (each 5 pmol/mL) for the genes of interest and the reference gene. 5. Optical 96-well reaction plates. 6. Optical adhesive film. 7. Real-time PCR cycler system with a standard 96-well block module.
3. Methods 3.1. Plant Growth and Harvesting
To mimic natural conditions it is best to grow the plants on soil. However, investigating a mutant that cannot survive on soil may require the use of media plates to ensure sugar supply. It is important to focus on biological replicates (do three!), rather than technical replicates (see Note 1). Strict control of growth conditions (light intensity and spectrum, moisture and temperature) is important for reproducible results. When working with Arabidopsis under non-stressing conditions, levels of white light should be maintained at around 100 mmol photons/m2/s, temperature should be kept between 18 and 22°C, and humidity held at 50–70%. Even in a climate chamber, growth conditions are variable. Thus, when different genotypes are being investigated, it is best to grow the plants randomly distributed in individual pots on multitrays. For each replicate and genotype or condition, material should be harvested from at least ten different plants. Take care to harvest leaves of the same age (see Note 2). The plant material should be snap-frozen immediately upon harvesting.
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When handling RNA samples special precautions should be taken to avoid degradation by adventitious RNases. Use gloves, as well as nuclease-free reagents and plastic ware. In the following protocol, RNA is extracted using the Qiagen RNeasy Plant Mini Kit. Other RNA extraction protocols, such as the Trizol reagent method, may also be used as long as RNA quality and quantity is ensured. 1. Using a pestle, grind plant material thoroughly in a mortar in the presence of liquid nitrogen. Decant tissue powder and liquid nitrogen into an RNase-free, liquid-nitrogen-cooled, 1.5-mL microcentrifuge tube. No more than 100 mg of plant material should be used (see Note 3). 2. Extract the RNA using the Qiagen RNeasy Plant Mini Kit. Add 450 mL of buffer RLT (ensure that 10 mL of b-mercaptoethanol is added per 1 mL of buffer RLT) to the tissue powder and vortex vigorously. 3. Transfer the lysate to a QIAshredder spin-column placed in a 2-mL collection tube and centrifuge for 2 min at full speed. Transfer the liquid phase of the flow-through to a new microcentrifuge tube, taking care not to disturb the pellet of cell debris. 4. Add 0.5 volume of ethanol (96–100%) to the cleared lysate and mix immediately by pipetting. 5. Transfer the sample, including any precipitate that may have formed, to an RNeasy spin-column placed in a 2-mL collection tube. Close the lid and centrifuge for at least 15 s at 8,000 × g. Discard the flow-through. If the sample volume exceeds 700 mL, centrifuge successive aliquots in the same RNeasy spin column. Discard the flow-through after each centrifugation. 6. Add 350 mL of buffer RW1 to the RNeasy spin-column. Close the lid gently and centrifuge for at least 15 s at 8,000 × g. Discard the flow-through. 7. To degrade DNA trapped on the column membrane, first add 10 mL of DNase I stock solution to 70 mL of buffer RDD, mix by gently inverting the tube (do not vortex), and centrifuge briefly (see Note 4). 8. Add 80 mL of the DNase I incubation mix directly to the RNeasy spin column membrane and leave on the benchtop (20–30°C) for 15 min. 9. Add 350 mL of buffer RW1. Close the lid gently and centrifuge for at least 15 s at 8,000 × g. Discard the flow-through. 10. Add 500 mL of buffer RPE (ensure that ethanol is added to buffer RPE before the first use) to the RNeasy spin column. Close the lid gently and centrifuge for at least 15 s at 8,000 × g to wash the spin column membrane. Discard the flowthrough.
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11. Add 500 mL of buffer RPE to the RNeasy spin column. Close the lid gently and centrifuge for 2 min at 8,000 × g to wash the column membrane. 12. Carefully remove the RNeasy spin column from the collection tube, ensuring that the column does not come into contact with the flow-through. Place the RNeasy spin column into a new 2-mL collection tube and discard the old collection tube with the flow-through. Close the lid gently and centrifuge at full speed for 1 min. 13. Place the RNeasy spin column in a new 1.5-mL collection tube. Add 30–50 mL of RNase-free water directly to the spincolumn membrane. Close the lid gently, let stand for 1 min, and centrifuge for 1 min at 8,000 × g to elute the RNA. RNA should be stored at −20°C or −80°C for short-term or longterm storage, respectively. 3.3. Quantification and Integrity Assessment of the RNA
The extracted total RNA is quantified spectrophotometrically at 260 nm (A260). Dilute the sample in a buffer at neutral pH and be sure to calibrate the spectrophotometer with the same solution used for dilution. To ensure significance, A260 readings should exceed 0.15. An absorbance of 1 unit at 260 nm of the RNA solution corresponds to a concentration of 40 mg/mL (see Note 5). The purity of the RNA is estimated from the ratio of the readings at 260 and 280 nm (A260/A280). Ideally, this should be ~2, but values between 1.7 and 2.1 are acceptable. The integrity and size distribution of the purified total RNA are often checked by denaturing formaldehyde-agarose gel electrophoresis or by using an Agilent 2100 Bioanalyzer. These methods provide enhanced sensitivity. However, RNA integrity can be adequately checked by electrophoresis on TAE or TBE gels under non-denaturing conditions. 1. Prepare a 1.5% agarose gel by adding the appropriate amount of agarose to 1× TAE buffer. For minigels, often no more than 50 mL is needed. 2. Heat in a microwave (make sure that the solution does not boil over!). 3. Allow to cool to 50–60°C and add 1 mL of ethidium bromide per 50 mL of gel. 4. Pour the liquid gel into the gel apparatus and place the comb at the cathode end. 5. For gel electrophoresis, overlay the completely cooled and settled gel with 1× TAE. If the gel must be stored, it can be moistened with some 1× TAE and wrapped in plastic wrap. 6. Remove the comb after covering the gel with electrophoresis buffer.
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Fig. 1. Representative agarose gel of total RNA extracted from Arabidopsis thaliana. Approximately 2 mg of RNA was electrophoresed on a 1.5% non-denaturing agarose gel and stained with ethidium bromide. Bands were assigned to individual rRNAs according to The Arabidopsis Information Resource (TAIR).
7. Add appropriate amounts of RNase-free H2O and gel loading buffer to 1–3 mg aliquots of the RNA samples, heat at 70°C for 1 min, and place on ice. 8. Load the gel with the prepared RNA samples. 9. Run the gel at 5 V/cm constant current until the bromophenol blue runs approximately two thirds of the way down the gel. 10. Take a picture of the gel with a gel documentation system. 11. The integrity of the RNA can now be assessed by inspecting the gel picture (for a sample picture, see Fig. 1). The ribosomal RNAs should appear as sharp bands (see Note 6). If the ribosomal RNA bands in a specific sample are not sharp, but appear as a smear towards smaller sized RNAs, it is likely that the sample has suffered major degradation and should not be used for microarray analysis. 3.4. Analysis of Microarrays: Identification of Differentially Expressed Genes Involved in Chloroplast Processes
Many companies and institutions will carry out chip hybridization at a reasonable cost. European scientists might choose to send their RNA samples to the NASC Affymetrix Service (http://affy.arabidopsis.info/prep.html). For the new Affymetrix RNA amplification protocol, it is sufficient to send 100–500 ng of RNA (see Note 7). This allows one to retain sufficient amounts of the prepared RNA for confirmatory experiments such as Northern blot analyses or real-time PCR. To check the quality of the hybridized chips and to analyze the data with Robin, the probe results files (CEL files) will be needed.
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1. Download the open-source software MapMan (http://mapman. gabipd.org/web/guest/mapman-version-3.5.0) (20). 2. Download the open-source microarray processing software Robin (http://mapman.gabipd.org/web/guest/robin) (19). Windows users should be sure to download the installer package that matches the Java Virtual Machine (JVM) installed on their system, irrespective of the underlying operating system. If a 32-bit JVM is installed on a 64-bit Windows 7 system, the 32-bit installer package is needed. 3. Create a project folder with all the CEL files you want to analyze. 4. Open the Robin program and choose “Affymetrix GeneChip® microarray experiment”. Clicking on “Start new project” will open a browser menu in which you select the created project folder. The subfolders “detailed results”, “plots”, “qualitychecks”, and “source” are automatically created, but the CEL files are not imported, yet. Click “Continue”. 5. Then, click “Add”, and the browser menu opens again. Select the CEL files from the project folder and click “Next” to import them into the Robin program. 6. Select all quality-checking tools for the analysis of the microarrays. If you are not experienced in analyzing microarray data, do not select “edit expert options”. The program uses by default RMA (robust multi-array average) (21) to normalize the chips (see Note 8), BH (Benjamini–Hochberg false discovery rate) (22) for the p-value correction method (see Note 9), and the linear model package limma (http://www. statsci.org/smyth/pubs/limma-biocbook-reprint.pdf) as the analysis strategy (see Note 10). 7. Clicking “Next” starts the calculations and after some calculation time (a matter of minutes) a window with the results of the quality check appears. Some of the quality assessment functions may have issued warnings. Clicking on the warning icon opens an info panel. Individual chips can be excluded from further analyses by checking the “Exclude” box (see Note 11). 8. Go to “Next”. The chips are assigned to groups of biological replicates by selecting the input files in the left panel and clicking “Add selected”. Allocate a name to each group of replicates (such as “mutant”, “wild type”, “treated”, “untreated”). Groups can be added or deleted by selecting “Add group” or “Delete group”. 9. The graphical experiment designer appears upon clicking “Next”. A dialogue box in the left panel explains how to set up the comparisons by CTRL-click-dragging arrows from one group to another one. If only a single experimental factor is varied in the experiment, define direct comparisons between the groups by dragging an arrow from, for example, the “mutant” to the “wild type” box (see Note 12). Here again,
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do not use the “expert settings” for data analysis. The program uses by default RMA, BH, and nestedF as a multiple testing strategy. The normalized data are saved, the log-fold change minimum is set to 1, and the p-value cut-off is set to 0.05. 10. Clicking “Next” starts the calculations and an annotation file menu appears after some calculation time (see Note 13). Choose “Ath_AFFY_ATH1_TAIR9_Jan2010_m02” as the annotation file (see Note 14). If the right mapping file was chosen, clicking “Annotate” opens a window with “Mapping successful”. Clicking “OK” opens a window with information about the location to which the results were saved and offers options to exit, restart, or modify the current experiment or to view data in MapMan. We recommend that one directly takes advantage of the implementation of MapMan in Robin. To view data in MapMan, make sure that the MapMan communication server is running. 11. After choosing the option to directly display the data in MapMan, the data format must be specified. Check that the default options are okay. Clicking “OK” starts the transfer of the data to MapMan, after which you exit the Robin program. 12. The MapMan program displays genomics data sets as diagrams of functional classifications (“bins”), which are available in the form of mapping files that contain entries for each probe set. To map the data, choose the desired classification in the drop-down menu under “Experiments” and double-click on the pathway. To get a comprehensive overview of the changes in the main chloroplast processes, select “Metabolism Overview”. This mapping file includes the light reactions, photorespiration, and tetrapyrrole biosynthesis. Furthermore, “Regulation Overview” includes information about hormones such as abscisic acid, and redoxlinked data sets (e.g., heme, thioredoxin, and glutaredoxin). The pathway “Protein targeting” displays components of the chloroplast import apparatus. Finally, in the “Primary Metabolism menu”, select “Photosynthesis”, “Amino Acids: N-metabolism”, “Amino Acids: Sulphate-assimilation”, and “Sugar and Sugar Derivatives: Sucrose-Starch”. Differentially regulated genes are displayed in boxes on the basis of a blue-to-red-scale. Mousing over the box displays detailed information about the gene. 13. The Robin program automatically saves all files relevant to the experiment into the initially created project folder as text (txt) files. To access and work with that information, one must import these files into a spreadsheet program. 14. Information about the log2-fold change of all genes in all mutants/treatments and their annotation is provided in the results.txt.annotated file. The subfolder “detailed_results” contains files with information about present/absent calls of the probe sets, and the mean and raw RMA-normalized expression
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values. The full tables contain the complete statistical results for the different mutants or treatments. Columns with the unique identifier for the probe sets, the log2-fold change values, the average normalized expression value, and the raw and BH-corrected p-values are provided (see Note 15). These files do not contain information about the annotation. Thus, merge the files containing information about the annotation, the present and absent calls, and the full tables: (1) Open the full table, save as a new file, and sort the “ID column” in ascending order. (2) Open the results.txt.annotated file, copy the “Description” column and paste it into the new file. (3) Open the PAcalls. table file and paste the columns containing information about present (P) and absent (A) calls into the new file. 15. Sort the log2-fold change column in descending order and identify the transcripts whose levels have changed (i.e., with log2-fold change of at least 1 or −1) and which, in addition, fulfil the BH p-value cut-off and are called “present” (see Note 16). 3.5. Confirmatory Experiments: RealTime Quantitative PCR Analyses
Whenever possible, design and use primers that flank intron sites of your gene of interest to discriminate amplification of genomic DNA. For instance, PerlPrimer (http://perlprimer.sourceforge. net) (23) can be used for this purpose. The size of the amplified products should be in the range of 100–300 bp. Table 1 offers a list of tested primers that amplify transcripts of photosynthesis, reactive oxygen species (ROS) marker and reference genes. Before running the real-time PCR experiment, it is advisable to check that the primer pair amplifies a product with the expected size. The primer pair should also be tested with a dilution series of the cDNA in real-time PCR and a standard curve plotted. A dilution factor of 10 ideally results in a crossing point value of 3.3 less than that of the original solution; this means the slope of the standard curve is ideally close to −3.3. Under experimental conditions, a range of −3.3 ± 0.3 is acceptable. The standard curve should also be linear,
Table 1 List of primer pairs tested and suitable for real-time PCR ATG number
Description
Primer pair (5¢–3¢)
AT4G36800
RCE1
AT2G37620
ACTIN 1
AT3g41768
18S rRNA
CTGTTCACGGAACCCAATTCa GGAAAAAGGTCTGACCGACA TTCACCACCACAGCAGAGCc ACCTCAGGACAACGGAATCG TCAACTTTCGATGGTAGGATAGTGc CCGTGTCAGGATTGGGTAATTT
Reference genes
(continued)
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Table 1 (continued) ATG number
Description
Primer pair (5¢–3¢)
ATCG00350
PSAA
AT5g64040
PSAN
ATCG00020
PSBA
AT1g06680
PSBP1
ATCG00480
ATPB
AT3G54890
LHCA1
AT!G61520
LHCA3
AT1G29920
LHCB1.1
AT1G29910
LHCB1.2
AT3G27690
LHCB2.4
AT5G54270
LHCB3
AT1G15820
LHCB6
AACCAATTTCTAAACGCTGGb TGATGATGTGCTATATCGGT CTACACTCTTCTCCACTGCTd CTCCACTTGTTGCCAATCTC GTGCCATTATTCCTACTTCTGb AGAGATTCCTAGAGGCATACC ATGGAGATGGGTTCAAAGTGd TAGGAGGTAATTAACCTGAGAG TATCGCCCAAATCATTGGTCd ACTCATAGCTACAGCTCTAACTC GAACTCGCTTATGAGCTGTGd GTCAAACCCAAAGTCACCAG GGGTTAGAGAAGGGTTTGGCb GAGGATGGCGAGCATAGC AGAGTCGCAGGAAATGGGb AAGCCTCTGGGTCGGTAG CCGTGAGCTAGAAGTTATCCc GTTTCCCAAGTAATCGAGTCC GCCATCCAACGATCTCCTCa TGGTCCGTACCAGATGCTCa CTCGGATCTCCCAAGTACACd CATGGATCACCTCAAGAGCT GGTGTGGCATGGTTTGAAGCd CGATTGAGAATCCGGGTTGAAG
Photosynthesis genes
ROS marker genes AT2G43510 AT1G26380 AT4G34410 AT3G01140 AT4G23290
CTTAGTCATTTCCGATGTGCCc GCATCTTCCACCTTTAGCTC FAD-LINKED TGTCGCTAACAAATTCCCTGc OXIDOREDUCTASE ATTATCTCCATCAGCTCATCGG AP2 DOMAIN CGTCTTCAGTTTCATCTCCTc TRANSCRIPTION FACTOR TCATCATATTCATCCACTCCTC MYB106 GCGCTTACAGAACCTAAACAGc CATGGCGATATGATCATGATCAG SER/THR KINASE-LIKE GGATACGCTGTTTCTAGGAGc PROTEIN GATCATCCGTAGCATCATCTG DEFENSIN-LIKE PROTEIN
(continued)
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Table 1 (continued) ATG number
Description
Primer pair (5¢–3¢)
AT3G22840
ELIP1
AT4G14690
ELIP2
AT1G77490
tAPX
AT3G02730
THIOREDOXIN F1
AT3G15360
THIOREDOXIN M4
AT1G50320
THIOREDOXIN X
AT3G26060
PEROXIREDOXIN Q
CGTTGCCGAAGTCACCATa AATCCAACCATCGCTAAACG CACCACAAATGCCACAGTCTa TGCTAGTCTCCCGTTGATCC AAACCTGAGACAAAGTACACGAc CTCTGCATAGTTCTTGAATGAAGG GTACACTCAATGGTGTGGTCc CTTCCTTGACAACCTTGTTATCC CGTCGAAGTACCAAATCTGTCc GAATTTGAACTTCCCTGCGA AAGCCTTATCTCAGGAATATGGc TGAAGAGAATGAAATGCGGT ATCTTTGCCAAGGTTAACAAGGc GAGTCTCTGAAAGCACAAGC
Others
Primer pair published in ref. 28 Primer pair published in ref. 10 c Primer pair published in ref. 29 d Not published a
b
with an R2 value close to 0.99. The real-time PCR efficiency (E) of one cycle in the exponential phase is calculated as: E = 10−1/slope. Thus, the standard curve gives information about the PCR efficiency of the tested primer pair. Ideally, it is 2 (equal to 100%). If cDNA synthesis is performed with a mixture of oligo(dT) primers and random hexamer primers, nuclear as well as organellar RNA will be reverse-transcribed in an efficient way. The iScript cDNA Synthesis Kit provides an easy-to-use solution for two-step RT-PCR. The iScript reverse transcriptase is RNAse H+, resulting in greater sensitivity than RNAse H-enzymes and already contains RNase inhibitor. The 5× iScript reaction mix contains reaction buffer, dithiothreitol, oligo(dT)-primers, random hexamer primers, and dNTP mix. 1. Set up the following reaction for cDNA synthesis (see Note 17): RNase-free H2O
15 − x mL
5× iScript reaction mix
4 mL
1 mg RNA
x mL
iScript reverse transcriptase
1 mL
2. Mix carefully and centrifuge briefly. 3. Incubate in a PCR cycler for 5 min at 25°C, 30 min at 42°C, and 5 min at 85°C, and then hold at 10°C.
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4. Proceed with the real-time PCR on the cDNA samples using the 2× SYBR Green Supermix (see Note 18): 2× SYBR Green Supermix
10 mL
Forward primer (5 pmol/mL)
2 mL
Reverse primer (5 pmol/mL)
2 mL
H2O
6 mL
cDNA or H2O
2 mL
5. Prepare enough master mix (containing all reagents but no cDNA) for the number of samples + 1. Pipette triplicates of each cDNA sample and use H2O as a blank control in the fourth reaction per primer pair. This means that if a master mix for wild type and one mutant should be pipetted, for example, a master mix for eight samples is appropriate: 2× SYBR Green Supermix
80 mL
Forward primer (5 pmol/mL)
16 mL
Reverse primer (5 pmol/mL)
16 mL
H2O
48 mL
6. For each reaction, pipette 18 mL of the master mix into a well in the optical 96-well reaction plate. 7. Add the cDNA or (in one well per primer pair) 2 mL of H2O (see Note 19). 8. Centrifuge briefly in a microtiter plate centrifuge to collect the samples on the base of each well. 9. Seal the plate tightly with the optical adhesive film. Avoid trapping air bubbles! 10. Program the real-time PCR machine. For the primer combinations listed in Table 1 the following program yields the best results: 95°C
5 min (“Hot start” of the DNA polymerase)
40 cycles consisting of: 95°C
10 s
55°C
30 s
72°C
20 s
Then, 1 cycle consisting of: 95°C
1 min
55°C
1 min
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Melting curve: Heat from 55°C to 95°C in 1°C steps, holding at each step for 10 s 11. To analyze the results of the finished run we recommend using a mathematical model, in which the relative expression ratio is calculated from the real-time PCR efficiencies and threshold cycle (CT) deviations (24) (see Note 20). The ∆CT between the mean CT of the gene of interest for the wild type (or control condition) and that for the mutant or challenging condition (sample) is calculated: ∆C T (gene of interest) = mean C T(gene of interest, control) − mean C T(gene of interest, sample) .
The same is done for the reference gene: ∆C T (reference gene) = mean C T(reference gene, control) − mean C T(reference gene, sample) .
Finally, the relative expression is calculated based on the efficiencies (E) and the ∆CT values: Ratio = (E gene of interest ∆C T (gene of interest) ) / (E reference gene ∆C T (reference gene) ) (see Note 21).
4. Notes 1. Analysis of technical replicates will lead to an artificially inflated number of differentially expressed genes. 2. It is best to grow quadruplicates instead of triplicates in case something should go wrong during the later procedures. 3. Weighing the tissue is the most accurate way to determine the amount. Weigh the empty 1.5-mL microcentrifuge tube, then fill it with the plant material. Allow the liquid nitrogen to evaporate, but do not allow the tissue to thaw. Weigh the filled microcentrifuge tube and calculate the difference. 4. Prepare DNase I stock solution before using the RNase-Free DNase Set for the first time. To avoid loss of DNase I, do not open the vial, but inject 550 mL of RNase-free water into it using an RNase-free needle and syringe. Mix gently by inverting the vial, and do not vortex. For long-term storage
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of DNase I, divide it into aliquots; these can be stored at −20°C for up to 9 months. Thawed aliquots can be stored at 2–8°C for up to 6 weeks. Do not refreeze the aliquots after thawing. 5. It may be difficult to determine small amounts of RNA of a low concentration photometrically. The use of a NanoDrop spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA) solves this problem. 6. The rule that the 28S rRNA band should be twice as intense as the 18S band in intact total RNA holds for vertebrate rRNA, but does not necessarily apply to invertebrates and plants. The overwhelming majority of these have a so-called “hidden break” in the 28S rRNA (25). 7. A workflow describing the steps starting from reverse transcription of the RNA and finally resulting in the hybridized chip is presented in the user manual of the GeneChip® 3¢ IVT Express Kit (http://microarray.csc.mrc.ac.uk/downloads/3%27%20 IVT%20Express%20Manual.pdf). 8. We recommend the article by Bolstad et al. (26) for further reading and background information about normalization of high-density oligonucleotide arrays. 9. A review that describes the progress made in assessing the false discovery rate, as well as the major conceptual developments that followed, is presented in ref. 27. 10. Robin also offers the rank product-based analysis, but this option is limited to the comparison of only two experimental conditions. 11. Chips of very poor quality can have a disproportionate effect on the final results, in terms of the lists of differentially expressed genes. Robin offers a range of quality check plots that cover many different levels of the chip data quality. Ideally, all chips in a given experiment should have comparable signal intensity distributions even before normalization. This is visualized in the analysis of signal intensity distribution plots. The false-colour images of probe level model weights point to technical problems caused, for example, by washing, dust on the chip, or scanner malfunction. Chips showing a consistently increased RLE (relative logarithmic expression) and/or NUSE (normalized unscaled standard error) are likely to be of low quality. Information about RNA degradation is extracted from the probe sets representing the genes. The probes are ordered from the 5¢ to the 3¢ end. Generally, RNA degradation is more active at the 5¢ terminus; thus, signal intensities of the probes closer to this terminus are weaker. If they are too low, Robin issues a
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warning. In the “scatter plots” Robin plots pair-wise comparisons of the normalized expression values of all combinations of two chips. These are useful for assessing whether two replicate chips show similar behaviour. If they do, the points should be arranged on a diagonal line. Replicate chips that do not show this behaviour suggest a problem, e.g., accidentally swapped or mislabelled samples, technical problems, or strong RNA degradation. Finally, chips that show similar expression profiles should cluster together in the Cluster Dendrogram and the Principal Components Analysis (PCA) Plot. 12. If more than one experimental factor is varied (e.g., mutants and different treatments), groups can be combined into metagroups, and comparisons between metagroups are made by dragging connections between them. 13. If chips that provoked a warning in the quality check were not excluded, a second warning summary will appear. To get an initial impression of potentially differentially regulated genes, one may click “Ignore and continue”. 14. If the mapping file is not already offered in the scroll-down menu, download the m02 mapping file via http://mapman. gabipd.org/web/guest/mapmanstore under “Mappings”, “Arabidopsis thaliana”, “Affymetrix, Ath_AFFY_ATH1_ TAIR9_Jan2010 download”. 15. The top 100 tables contain the same data columns as the full tables, but do not fulfil the p-value and/or log2-fold change cut-offs. 16. If a mutant is investigated in which the transcript of the mutated gene is known to be down-regulated, checking whether the transcript is down-regulated in the results table provides a quick indication of the quality of the data. 17. It is possible to adjust the reaction volume to as little as 5 mL, adding only 250 ng of RNA. 18. The SYBR Green Supermix already contains each dNTP at 0.4 mM, iTaq DNA polymerase, MgCl2, SYBR Green I, and fluorescein. The user manual suggests using a final volume of 50 mL, but it is sufficient to set up a 20-mL reaction. 19. For the primers listed in Table 1, often a 1:10 dilution of the cDNA reaction gives results in the linear range. 20. The threshold cycle is the first cycle in the real-time PCR in which the fluorescence signal of a sample exceeds its background fluorescence. Using relative quantification, the CT values of two samples (e.g., wild type and mutant) are compared and the ratio is calculated. It is important, indeed crucial, to choose as a reference a gene whose expression is known not to vary between the samples investigated.
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21. The efficiencies calculated from the standard curves can be used here. However, it is more accurate to calculate the efficiencies from each run. The LinRegPCR program, which is available on request from the author (email:
[email protected]. nl; subject: LinRegPCR), performs linear regression on the Log(fluorescence) per cycle number data to calculate PCR efficiencies for each sample.
Acknowledgements We thank Paul Hardy for critical reading of the manuscript, Michael Scharfenberg for preparing Figure 1 and the DFG (FOR 804) and EC (ITN “COSI”) for funding. References 1. Timmis, J. N., Ayliffe, M. A., Huang, C. Y., and Martin, W. (2004) Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nat. Rev. Genet. 5, 123–135. 2. Kleine, T., Maier, U. G., and Leister, D. (2009) DNA transfer from organelles to the nucleus: the idiosyncratic genetics of endosymbiosis. Annu. Rev. Plant Biol. 60, 115–138. 3. Kleine, T., Voigt, C., and Leister, D. (2009) Plastid signalling to the nucleus: messengers still lost in the mists? Trends Genet. 25, 185–192. 4. Woodson, J. D., and Chory, J. (2008) Coordination of gene expression between organellar and nuclear genomes. Nat. Rev. Genet. 9, 383–395. 5. Pogson, B. J., Woo, N. S., Forster, B., and Small, I. D. (2008) Plastid signalling to the nucleus and beyond. Trends Plant Sci. 13, 602–609. 6. Apel, K., and Hirt, H. (2004) Reactive oxygen species: metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 55, 373–399. 7. Bonardi, V., Pesaresi, P., Becker, T., Schleiff, E., Wagner, R., Pfannschmidt, T., Jahns, P., and Leister, D. (2005) Photosystem II core phosphorylation and photosynthetic acclimation require two different protein kinases. Nature 437, 1179–1182. 8. Fey, V., Wagner, R., Bräutigam, K., Wirtz, M., Hell, R., Dietzmann, A., Leister, D., Oelmüller, R., and Pfannschmidt, T. (2005) Retrograde plastid redox signals in the expression of nuclear genes for chloroplast proteins of Arabidopsis thaliana. J. Biol. Chem. 280, 5318–5328.
9. Ihnatowicz, A., Pesaresi, P., Varotto, C., Richly, E., Schneider, A., Jahns, P., Salamini, F., and Leister, D. (2004) Mutants for photosystem I subunit D of Arabidopsis thaliana: effects on photosynthesis, photosystem I stability and expression of nuclear genes for chloroplast functions. Plant J. 37, 839–852. 10. Pesaresi, P., Hertle, A., Pribil, M., Kleine, T., Wagner, R., Strissel, H., Ihnatowicz, A., Bonardi, V., Scharfenberg, M., Schneider, A., Pfannschmidt, T., and Leister, D. (2009) Arabidopsis STN7 kinase provides a link between short- and long-term photosynthetic acclimation. Plant Cell 21, 2402–2423. 11. Pfannschmidt, T., Nilsson, A., and Allen, J. F. (1999) Photosynthetic control of chloroplast gene expression. Nature 397, 625–628. 12. Laloi, C., Stachowiak, M., Pers-Kamczyc, E., Warzych, E., Murgia, I., and Apel, K. (2007) Cross-talk between singlet oxygen- and hydrogen peroxide-dependent signaling of stress responses in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 104, 672–677. 13. Lee, K. P., Kim, C., Landgraf, F., and Apel, K. (2007) EXECUTER1- and EXECUTER2dependent transfer of stress-related signals from the plastid to the nucleus of Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 104, 10270–10275. 14. Oelmüller, R., Levitan, I., Bergfeld, R., Rajasekhar, V. K., and Mohr, H. (1986) Expression of nuclear genes as affected by treatments acting on the plastids. Planta 168, 482–492. 15. Börner, T., Metzlaff, M., Koch-Siemenroth, A., Steiner, K., and R., Hagemann (1985)
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T. Kleine and D. Leister Chloroplast control of the accumulation of nuclear-DNA encoded proteins. In, 1st International Congress on Plant Molecular Biology, Savannah, GA, USA. Richly, E., Dietzmann, A., Biehl, A., Kurth, J., Laloi, C., Apel, K., Salamini, F., and Leister, D. (2003) Covariations in the nuclear chloroplast transcriptome reveal a regulatory masterswitch. EMBO Rep. 4, 491–498. Biehl, A., Richly, E., Noutsos, C., Salamini, F., and Leister, D. (2005) Analysis of 101 nuclear transcriptomes reveals 23 distinct regulons and their relationship to metabolism, chromosomal gene distribution and co-ordination of nuclear and plastid gene expression. Gene 344, 33–41. Gentleman, R. C., Carey, V. J., Bates, D. M., Bolstad, B., Dettling, M., Dudoit, S., Ellis, B., Gautier, L., Ge, Y., Gentry, J., Hornik, K., Hothorn, T., Huber, W., Iacus, S., Irizarry, R., Leisch, F., Li, C., Maechler, M., Rossini, A. J., Sawitzki, G., Smith, C., Smyth, G., Tierney, L., Yang, J. Y., and Zhang, J. (2004) Bioconductor: open software development for computational biology and bioinformatics. Genome Biol. 5, R80. Lohse, M., Nunes-Nesi, A., Krüger, P., Nagel, A., Hannemann, J., Giorgi, F. M., Childs, L., Osorio, S., Walther, D., Selbig, J., Sreenivasulu, N., Stitt, M., Fernie, A. R., and Usadel, B. (2010) Robin: an intuitive wizard application for R-based expression microarray quality assessment and analysis. Plant Physiol. 153, 642–651. Thimm, O., Blasing, O., Gibon, Y., Nagel, A., Meyer, S., Krüger, P., Selbig, J., Müller, L. A., Rhee, S. Y., and Stitt, M. (2004) MAPMAN: a user-driven tool to display genomics data sets onto diagrams of metabolic pathways and other biological processes. Plant J. 37, 914–939.
21. Irizarry, R. A., Hobbs, B., Collin, F., BeazerBarclay, Y. D., Antonellis, K. J., Scherf, U., and Speed, T. P. (2003) Exploration, normalization, and summaries of high density oligonucleotide array probe level data. Biostatistics 4, 249–264. 22. Benjamini, Y., and Hochberg, Y. (1995) Controlling the false discovery rate - a practical and powerful approach to multiple testing. J. R. Stat. Soc. Series B Methodol. 57, 289–300. 23. Marshall, O. J. (2004) PerlPrimer: cross-platform, graphical primer design for standard, bisulphite and real-time PCR. Bioinformatics 20, 2471–2472. 24. Pfaffl, M. W. (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29, e45. 25. Ishikawa, H. (1977) Evolution of ribosomal RNA. Comp Biochem Physiol B 58, 1–7. 26. Bolstad, B. M., Irizarry, R. A., Astrand, M., and Speed, T. P. (2003) A comparison of normalization methods for high density oligonucleotide array data based on variance and bias. Bioinformatics 19, 185–193. 27. Benjamini, Y. (2010) Discovering the false discovery rate. J. R. Stat. Soc. Series B Stat. Methodol. 72, 405–416. 28. Kleine, T., Kindgren, P., Benedict, C., Hendrickson, L., and Strand, A. (2007) Genome-wide gene expression analysis reveals a critical role for CRYPTOCHROME1 in the response of Arabidopsis to high irradiance. Plant Physiol. 144, 1391–1406. 29. Voigt, C., Oster, U., Börnke, F., Jahns, P., Dietz, K. J., Leister, D., and Kleine, T. In-depth analysis of the distinctive effects of norflurazon implies that tetrapyrrole biosynthesis, organellar gene expression and ABA cooperate in the GUN-type of plastid signalling. Physiol. Plant. 138, 503–519.
Chapter 8 Use of Non-aqueous Fractionation and Metabolomics to Study Chloroplast Function in Arabidopsis Peter Geigenberger, Axel Tiessen, and Jörg Meurer Abstract Chloroplasts are the chemical factories of plant cells because they are able to fix inorganic carbon and convert it to a wide-range of photoassimilates that are exported to the cytosol and other sub-cellular compartments. If the regulation of these processes is to be understood, the in vivo concentrations of a large number of metabolites have to be measured in all of these compartments separately. Sophisticated analytical approaches and continued advances in the technology of mass spectrometry coupled to a variety of fractionation and separation techniques allow the reliable analysis of a comprehensive complement of metabolites in photosynthetic tissues. Metabolomic approaches allow the multi-parallel analysis of a widerange of metabolic intermediates and have been used for rapid phenotyping of different genotypes and environmental effects in plants. In addition to this, methods have been developed to analyse metabolite levels in different sub-cellular compartments of plant cells. Here, we describe methods for sub-cellular fractionation of Arabidopsis leaves using a non-aqueous density gradient technique, sample preparation suitable for metabolite profiling using gas-chromatography-mass spectrometry, and calculation of subcellular metabolite concentrations. Key words: Arabidopsis, Chloroplast, Non-aqueous density gradient fractionation, Sub-cellular biochemical analysis, GC-MS, Metabolomics, Metabolic regulation of photosynthesis
1. Introduction In contrast to other genomics technologies, which operate on a single class of chemicals such as DNA or RNA, metabolite profiling operates on a much larger range of molecular species that have widely divergent physical and chemical properties. Therefore, no unique “master” method exists that allows quantification of all metabolites in plant samples (1, 2). Metabolic profiling can be regarded as the sum of several analytical methods requiring different equipments and separation technologies for analysing the different families of chemical compounds found in plants. Gas-chromatography mass spectrometry (GC-MS) allows the qualitative identification and R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_8, © Springer Science+Business Media, LLC 2011
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robust quantification of a few hundred metabolites in a single chromatographic run. It provides a snapshot-type view of plant metabolism and a particularly comprehensive coverage of sugaralcohols, amino acids, and organic acids (3). Nevertheless, GC-MS data typically do not include some important metabolites of central metabolism such as nucleotide-phosphates or sugar-dinucleotides. It is therefore important to use different analytical techniques complementarily to GC-MS such as high-performance liquid chromatography (HPLC), spectrophotometry in the ultraviolet and visible range (UV–VIS), and capillary electrophoresis (CE). Metabolic profiling has been used as a powerful tool for the functional analysis of Arabidopsis mutants and the characterisation of complex changes in different genetic conditions and environmental situations (4–10), as well as for discovering metabolic biomarkers for phenotype prediction in medical and agricultural science (11–13). More recently, GC-MS-based metabolite profiling was used to analyse distinct metabolic states during photosynthetic acclimation to different light conditions (14) and limiting CO2 concentrations (15). Detailed characterisation of the interaction of chloroplast metabolism with other metabolic compartments requires analysis of metabolites at a sub-cellular level. Most metabolic profiling approaches use samples derived from bulks of cells and tissues. Thus, such results provide only an average across all sub-cellular compartments and cell types. To overcome this limitation, a nonaqueous fractionation (NAF) method has been developed, which allows leaf material to be separated into a series of experimental fractions enriched in material deriving from different compartments (16–19). The in vivo distribution of the metabolites between chloroplast, cytosol, and vacuole can then be quantitatively estimated by comparison with the activities of marker enzymes, which are confined to a single compartment in vivo. The method has also been used successfully in heterotrophic potato tubers (6, 20) and developing seeds (data not shown). The NAF technique consists of the following stepwise procedures: leaf material is immediately frozen in liquid N2, homogenised to a fine powder (<5 mm average particle size), lyophilised, re-suspended in an organic solvent, ultrasonicated, filtered through a nylon net (30 mm), and subjected to density gradient centrifugation in a mixture of non-aqueous liquids (heptane and tetrachlorethylene). Four to six fractions are collected, and marker enzymes and metabolites are measured. The distribution of metabolites in different sub-cellular compartments can then be calculated by solving an array of mathematical equations applying the method of least squares using the solver function of Microsoft Excel. In this chapter, we provide a detailed protocol for NAF of Arabidopsis leaves. We describe harvesting, fractionation, and sample preparation procedures. We also describe the measurement of some marker enzymes and provide a protocol for chemical
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derivatisation suitable for downstream GC-MS-based metabolite profiling. Finally, we explain the required calculations that need to be done in order to obtain estimates about sub-cellular metabolite concentrations.
2. Materials 2.1. Sampling, Freeze-Stop, Homogenisation, and Lyophilisation of Leaf Material
1. Cotton gloves and goggles. 2. Plastic forceps, metal scissors, and plastic sieve for collecting tissue. 3. Aluminium bags to store leaf tissue (15 cm × 10 cm). 4. Dewars, for use with liquid N2 (KGW-Isotherm, Karlsruhe, Germany). 5. Liquid nitrogen. 6. −80°C Ultra-deep freezer. 7. Oscillating ball-mill, MM 400 (Retsch, Haan, Germany). 8. Stainless steel containers (20 mL volume; Retsch) and metal mill balls (20 mm). 9. Metal spoon to transfer frozen powder to Falcon tubes. 10. Polypropylene tubes, 50 mL (Falcon tubes, BD Biosciences, Franklin Lake, NJ, USA). 11. Aluminium holder (20 cm × 20 cm × 5 cm) to put 50-mL Falcon tubes inside the lyophiliser. The holder should be precooled in liquid N2. 12. Rubber bands and Miracloth tissue to close Falcon tubes. 13. Lyophiliser system (Christ, Osterode, Germany), with 2 horsepower (2HP) power oil vacuum pump, refrigerated compartment (−50°C), and manual or magnetic ventilation. The lyophiliser needs to be connected to a dry N2 gas supply to ventilate it. 14. Polycarbonate desiccators (diameter 40 cm), containing silica gel and phosphorus pentoxide drying agent with a moisture indicator (Sicapent, Merck, Darmstadt, Germany). 15. Kitchen sealable plastic bags for storage of Falcon tubes containing dry tissue powder. 16. −20°C Deep freezer.
2.2. Ultrasonication and Non-aqueous Density Gradient Fractionation
1. Molecular sieve (pore size, 0.4 nm), with moisture indicator (Merck; catalogue number 1.06108.0250). 2. Tetrachlorethylene (Merck Uvasol grade) (see Note 1). 3. n-Heptane (Merck Uvasol grade) (see Note 1). 4. Microbalance for weighing up to 3 kg with accuracy down 100 mg (e.g., Sartorius or Metler).
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5. Tetrachlorethylene/heptane mixture (v/v 66:34; d = 1.3 g/ cm3). Mix 1,069 g of tetrachlorethylene (~660 mL) with 231 g of n-heptane (~340 mL). Use a microbalance to weight solvents in a brown glass bottle. Prepare at least 1 day in advance and add molecular sieve. 6. Polypropylene tubes, 50 and 15 mL (Falcon tubes). 7. At least two medium-sized polycarbonate desiccators (diameter 40 cm), containing silica gel. 8. Small foam stopper (e.g., those that come in aluminium enzyme containers), to be used with the sonicator needle in order to seal the Falcon tube and keep it water vapour free. 9. Ultrasonicator (Sonopuls HD 200 with microtip MS 73/D; Bandelin, Berlin, Germany). 10. Ear protection (to be used during sonication). 11. Metal stator and four universal holder grips for holding the Falcon tubes in the right position during sonication, gradient formation, and collection of fractions. 12. Aluminium block (20 cm × 20 cm × 5 cm) with appropriate holes for holding 50 mL Falcons. This is for cooling the tubes during sonication. The block should be pre-cooled to −20°C. 13. Plastic sieve of polyester monolen (pore size, 30 mm; Neolab, Heidelberg, Germany; catalogue number 2-4072). 14. Plastic funnel (15-cm diameter). 15. Refrigerated centrifuge with swing-out rotor for Falcon tubes (e.g., Megafuge 5810 R; Eppendorf, Hamburg, Germany). 16. Gradient pump system (Biologic system, Bio-Rad, Hercules, CA, USA), with programmable peristaltic pump (Econo Pump, Bio-Rad) and gradient mixer for two solvents. 17. Metal needle (1.5-mm thick and 20-cm long), for pumping the solvent into the bottom of the tube during gradient formation (Neolab). 18. Approximately, 5 m of silicon tube (0.8-mm inner diameter and 0.8-mm thick wall; Bio-Rad; catalogue number 731 8210). 19. Approximately, 5 m of silicon tube (1.6-mm inner diameter and 0.8-mm thick wall; Bio-Rad; catalogue number 731 8211). 20. Approximately, 4 m of teflon tube (1.6-mm inner diameter and 1.6-mm thick wall; Bio-Rad; catalogue number 750 0603). 21. Set of male and female luer-locks for coupling all silicon and teflon tubes (Bio-Rad; catalogue number 731 8220). 22. Set of micro-pipettes (200 mL, 1,000 mL, and 5 mL). 23. Polypropylene micro-tubes, 2 mL (Eppendorf Safe-Lock). 24. Large polycarbonate desiccator (diameter 60 cm), containing silica gel, with connection to vacuum pump.
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25. Teflon membrane vacuum pump (Laboport, KNF Neuberger, Freiburg, Germany). 2.3. Sample Preparation and Measurement of Marker Enzyme Activities
1. Enzyme extraction buffer: 50 mM 4-(2-hydroxyethyl) piperazine-1-ethanesulfonic acid (HEPES)–KOH, pH 7.5, 10% (w/v) glycerol, 5 mM MgCl2, 1 mM ethylenediaminetetra acetic acid (EDTA), 2 g/L polyvinylpolypyrrolidone (PVPP), 0.1% (v/v) Triton X-100, 1 mM dithiothreitol (DTT), 1 mM e-aminocaproic acid, 1 mM benzamidine, 1.5 mM phenylmethylsulfonyl fluoride (PMSF). 2. NAP-5 columns (Pharmacia, GE Healthcare, Chalfont St. Giles, UK) (loading volume, 0.5 mL; elution volume, 1 mL). 3. Small plastic or glass beads (~1-mm diameter). 4. Analytical balance, with accuracy down to 0.1 mg (e.g., Sartorius or Metler). 5. Thermomixer Comfort (Eppendorf). 6. Polypropylene micro-tubes, 2 mL (Eppendorf Safe-Lock). 7. 80% (v/v) ethanol. 8. Solutions and reagents for measurement of marker–enzyme activities. The pipetting/measuring schemes for all stock solutions/reagents are shown in Tables 1–3. These stocks should be stored at 4°C or −20°C, according to the manufacturer’s instructions. (a) Measurement of the activities of the cytosolic UDPglucose pyrophosphorylase (UGPase; Table 1): 500 mM
Table 1 Stock solutions and reagents required for measurement of the cytosolic UDP-glucose pyrophosphorylase (UGPase) Final concentration
Stock concentration/ metabolites
Master-mix recipe for 50 samples (total volume = 10 mL)
H2O
–
7.73 mL
100 mM Tris–HCl, pH 8
500 mM
2 mL
2 mM MgCl2
100 mM
0.2 mL
20 mM G1,6 bP
10 mM
0.5 mM NADP
787.4 g/mol
3.94 mg
3 mM UDP-Gluc
610.3 g/mola
18.31 mg
1.6 U/mL PGM (from rabbit)
0.4 U/mL
40 mL
1 U/mL G6P-DH (from yeast)
0.7 U/mL
14.29 mL
20 mL a
Molar mass of molecule. These items are added to the Master-mix by weight
a
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Table 2 Stock solutions and reagents required for measurement of the plastidial ADP-glucose pyrophosphorylase (AGPase)
Final concentration
Stock concentration/ metabolites
Master-mix recipe for 50 samples (total volume = 10 mL)
H2O
–
4.94 mL
50 mM HEPES–KOH, pH 7.8, 5 mM MgCl2
100/10 mM
5 mL
10 mM G1,6 bP
10 mM
10 mL
3 mM PGA
230 g/mola
6.9 mg
3 mM DTT
154.3 g/mola
4.63 mg
0.5 mM NADP
a
787.4 g/mol
3.94 mg
1 mM ADP-Gluc
633.3 g/mola
6.33 mg
1.6 U/mL PGM (from rabbit)
0.4 U/mL
40 mL
0.7 U/mL G6P-DH (from yeast)
0.7 U/mL
10 mL
Molar mass of molecule. These items are added to the Master-mix by weight
a
Table 3 Stock solutions required for measurement of the vacuolar a-mannosidase Final concentration
Stock concentration Per assay (mL) Per blank (mL)
50 mM citrate-KOH, pH 4.5
75 mM
100
100
Sample extract (fraction)
–
20
20 (of H2O)
30
30
100
100
5 mM pNP-a-Man
(starts reaction) 25 mM
0.4 M Tris–borate, pH 9.8 (stops reaction) 0.8 M
Tris–HCl, pH 8; 100 mM MgCl2; 10 mM glucose-1,6bisphosphate (G1,6 bP); nicotinamide adenine dinucleotide phosphate (NADP; molecular weight [MW], 787.4 g/mol); UDP-glucose (UDP-Gluc; MW, 610.3 g/ mol); 0.4 U/mL (200 U/mg) phosphoglucomutase (PGM), from rabbit (Roche, Basel, Switzerland; catalogue number 108375); and, 0.7 U/mL (140 U/mg) glucose6-phosphate-dehydrogenase (G6P-DH), from yeast (Roche; catalogue number 127671).
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(b) Measurement of the activities of the plastid ADP-glucose pyrophosphorylase (AGPase; Table 2): 100 mM HEPES– KOH, pH 7.8, 10 mM MgCl2; 10 mM G1,6 bP; 3-phosphoglycerate (PGA; MW, 230 g/mol); DTT (MW, 154.3 g/mol); NADP (MW, 787.4 g/mol); ADP-glucose (ADP-Gluc; MW, 633.3 g/mol); 0.4 U/mL PGM (Roche); and, 0.7 U/mL G6P-DH (Roche). (c) Measurement of the activities of the vacuolar a-mannosidase (Table 3): 75 mM citrate-KOH, pH 4.5; 25 mM p-nitrophenyl-d-a-mannopyranoside (pNP-a-Man); and 0.8 M Tris–borate, pH 9.8. 9. Microplate spectrophotometer (e.g., Anthos HT3 or Biotek). 10. Flat-bottom microplates made from UV–VIS-transparent polystyrene (96 well) (e.g., Sarstedt or Greiner). 2.4. Sample Extraction for Gas Chromatographic Analysis
1. 100% methanol (gradient grade for liquid chromatography; Merck), pre-cooled at −20°C. 2. 0.2 g/L Ribitol in distilled water (dH2O) (Sigma, St. Louis, MO, USA). 3. Thermomixer Comfort (Eppendorf). 4. Glass vial (GL14, Schott, Mainz, Germany). 5. 100% Chloroform (for liquid chromatography; Merck), pre-cooled at −20°C. 6. Centrifuge (e.g., Eppendorf Megafuge 5810 R with swing-out rotor). 7. Glass tube suitable for GC-MS analysis. 8. Vacuum concentrator (e.g., Maxi-Dry Lyo, Heto-Holten, Allerød, Denmark). 9. Argon gas, purity level 5.0 (Air Liquide, Düsseldorf Germany). 10. −80°C Ultra-deep freezer. 11. Desiccant (orange silica gel; Carl Roth, Karlsruhe, Germany).
2.5. Chemical Derivatisation of the Samples for GC-MS
1. Methoxyamination solution: 20 mg/mL methoxyamine hydrochloride in 100% pyridine at room temperature (RT). Use the solution only when freshly prepared (see Note 2). 2. Polypropylene micro-tubes, 1.5 mL (Eppendorf Safe-Lock). 3. Thermomixer Comfort (Eppendorf). 4. N-Methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA reagent; Macherey-Nagel, Düren, Germany; catalogue number 2458978-4). Store the reagent in 1-mL tubes at 4°C (see Note 3). 5. Retention-time standards; e.g., a mixture of alkanes with chain lengths from C8 to C20 (Sigma; catalogue number 04070).
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1. Pegasus time-of-flight (TOF) mass spectrometer (Leco Instruments, St. Joseph, MI, USA).
2.6. GC-TOF-MS Metabolite Profiling
2. Gas chromatograph, 7890A, split/splitless injector (Agilent, Santa Clara, CA, USA). 3. Autosampler, MPS 2XL dual-rail (Gerstel, Mülheim an der Ruhr, Germany); the configuration comprises an agitator, two 54-sample trays with cooling for 2-mL vials, a fast wash station for two solvents, and 2-mL syringes mounted on the two robotic autosampler arms. 4. MDN-35 capillary column, 30 m × 0.32 mm, 0.25-mm film thickness (Macherey-Nagel). 5. Helium carrier gas, purity level 5.0 (Air Liquide). 6. ChromaTOF chromatography processing and mass spectral deconvolution software, version 4.00 or higher (Leco Instruments). 1. Computer with spreadsheet software (Microsoft Excel or Open Office Calc).
2.7. Calculation of Sub-cellular Metabolite Concentrations
2. Regression analysis software (Solver). The add-in function Solver needs to be installed in Microsoft Excel. The Solver function can find the minimum of the quadratic differences (least squares) between theoretical and observed distributions. Examples of data tables are given in Tables 4–6. Prepare an
Table 4 Representation of an Excel spreadsheet showing marker enzymes and compartments required for regression analysis A
B
1
C
D
Vacuole (mannosidase)
Plastid (AGPase)
Cytosol/Mitochondria (UGPase)
100.00%
100.00%
100.00%
Compartments
2
Gradient 11
3
Total aliquot t
4
Fraction p
26.16%
3.63%
9.34%
5
Fraction 0
21.90%
4.62%
17.59%
6
Fraction 1
17.84%
5.54%
26.45%
7
Fraction 2
6.98%
46.70%
28.03%
8
Fraction 3
0.24%
20.24%
12.04%
a
A random example of gradient no. 11 is shown
a
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Excel spreadsheet containing Tables 4–6. Use precisely the coordinates given in each table. Table 4 starts with column A, line 1; Table 5 starts with column A, line 20; and, Table 6 starts with column H, line 20. Enter the formulae in the indicated cells, and replace the data with your own data. Prepare one Excel file for each gradient (in this example, it was gradient number 11) and make separate Excel sheets for each metabolite measured in the gradient (in this example, it was aspartate).
Table 5 Representation of an Excel spreadsheet showing metabolite values required for regression analysis A
B
C
20
Gradient 11a
Amount in Metabolite fraction (% of total in (nmol/g DW) aliquot)
21
Total 5,205.14 aliquot t
100% = B21/B$21
123% = J23 × B3 + J24 × C3 + J25 × D3
22
Fraction p
619.80
12% = B22/B$21
12% = J23 × B4 + J24 × C4 + J25 × D4
0.000000158 = (C22 − D22) × (C22 − D22)
23
Fraction 0
773.33
15% = B23/B$21
15% = J23 × B5 + J24 × C5 + J25 × D5
0.000000235 = (C22 − D22) × (C22 − D22)
24
Fraction 1
1,129.27
22% = B24/B$21
22% = J23 × B6 + J24 × C6 + J25 × D6
0.000000037 = (C22 − D22) × (C22 − D22)
25
Fraction 2
2,115.17
41% = B25/B$21
40% = J23 × B7 + J24 × C7 + J25 × D7
0.000014786 = (C22 − D22) × (C22 − D22)
26
Fraction 3
795.15
15% = B26/B$21
16% = J23 × B8 + J24 × C8 + J25 × D8
0.000074231 = (C22 − D22) × (C22 − D22)
Sum of quadratic differences
8.94467E-05 = sum(E22:E26)
28 A random example of gradient no. 11 is shown
a
D
E
Theoretical value predicted by formula
Quadratic differences
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Table 6 Representation of an Excel spreadsheet showing fitted values obtained by regression analysis for the metabolite aspartate H
I
20
Results:
Gradient 11
21
Sub-cellular compartmentation of:
22
J
K
Absolute values (nmol/g DW)
Best fit (relative% of homogenate)
Rel. amount found in fractions
a
23
Vacuole
384.60 = K23 × B$21
9.09%
7.39% = J23 × J$31
24
Plastid
1,065.16 = K24 × B$21
25.17%
20.46% = J24 × J$31
25
Cytosol/mitochondria
3,755.38 = K25 × B$21
88.74%
72.15% = J25 × J$31
26 27 28
31
Solver finds minimum value for cell E28 by modifying cells J23, J24, and J25 Sum
5,205.14 = sum (I23:I25)
122.99% = sum (J23:J25)
100.00% = sum (K23:K25)
A random example of gradient no. 11 is shown
a
3. Methods In the following section, a detailed protocol is provided for sampling, NAF, and sub-cellular metabolite analysis of Arabidopsis leaves. The flow diagram in Fig. 1 provides an overview of the main steps of the experimental procedure. While this protocol is also applicable for leaves of other species, sub-cellular analysis of heterotrophic tissues requires some modifications, which are detailed elsewhere for potato tubers (6, 20). We do not provide details on the GC-TOF MS-based metabolite profiling itself, such as instrument settings and evaluating of the resultant chromatograms. The reader is referred a recent publication (3), where this has been documented in great detail.
8 Use of Non-aqueous Fractionation and Metabolomics… Harvesting Resuspension Sonication
Homogenisation
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Lyophilisation
Filtering 30 mm nylon net
Concentration Resuspension
Fractionation density gradient
A: Heptane
gradient pump
B: C2Cl4
needle
80% C2Cl4
~35 mL Total aliquots t 4 200 µL
100% C2Cl4
~4 mL sample
centrifugation 1 h >4000 g
Non Aqueous Fractionation Fractions
Data analysis Best fit distribution Excel Solver function
Marker enzyme measurements
1/3
Metabolite measurements
2/3
3 2 1 0 p
amyloplast vacuole pellet
Fig. 1. Flow-chart providing a schematic overview of the experimental procedures. Each fraction has to be divided into two parts for measuring marker enzymes and metabolites separately. The post-centrifugation pattern shown is for heterotrophic tissue, but the procedures are similar for leaf tissue.
NAF is neither attempting to isolate whole organelles nor intact membranes. It only attempts to obtain a partial enrichment of microscopic particles (random pieces of sub-cellular compartments) in any of the density fractions, so that the actual concentration of a given metabolite/activity in a given sub-cellular compartment can be calculated by linear regression. For this, leaves are frozen in liquid nitrogen, lyophilised, and broken into fragments of approximately <5 mm size. These fragments are separated according to their specific density. After freeze drying, the vacuolar fragments which initially contained high amounts of water and minerals and low amounts of proteins will become relatively more dense than the cytosolic particles which have a different composition of proteins, minerals, sugars, and lipids. The density gradient that is formed by the linear mixture of non-aqueous solvents allows partial enrichment of material from a given sub-cellular compartment. The sub-cellular particles are physically separated in water-free solvents (non-aqueous solvents) so that metabolic activity is quenched and the redistribution of metabolites and enzymes is prevented. This means that metabolites and enzymes stay together during the NAF, and that they can be quantitatively recovered (biomolecules are chemically stable in a water-free environment). Metabolites such as starch, cellulose, and chlorophyll, or enzymes such as ADPglucose pyrophosphorylase (AGPase) and UDP-glucose pyrophosphorylase (UGPase), can be used as markers for a given sub-cellular
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compartment. The use of extremely low temperatures first and the subsequent exclusion of water (lyophilisation and then non-aqueous solvents maintained water-free with molecular sieves) prevents metabolic activity during the whole fractionation procedure (16). The procedure is suitable to study metabolites and proteins that are present in the polar phase of the cells. A proportion of hydrophobic proteins, lipids, and membrane components might get dissolved in the organic solvents employed during the whole NAF procedure. Nevertheless, compounds as hydrophobic as chlorophyll are still relatively stable after the fractionation and can be obtained in the different fractions with an acceptable recovery. 3.1. Sampling, Freeze-Stop, Homogenisation, and Lyophilisation of Leaf Material
1. Plants should be grown under controlled conditions with respect to humidity, temperature, light quality and quantity, photoperiod, nutrients, and pathogen control. Since metabolite levels can show strong diurnal variations (e.g., starch), leaf samples should be taken at specific time-points during the day. Metabolite levels also vary considerably between different parts of a single leaf (basal and apical parts) and between different leaves of a single plant. Care should be taken to sample leaves of similar developmental stages and, if only parts of a leaf are sampled, to sample always the same part of the leaf (see Note 4). The most frequent procedure is to make a bulk of all source leaves of Arabidopsis using the “forceps harvesting procedure” (see Note 5). 2. To stop leaf metabolism as quickly as possible, leaves should be plunged directly into liquid nitrogen using the “forceps harvesting procedure” (see Note 5). The turnover rate of metabolites during leaf photosynthesis can be exceptionally fast (~1 s), making it essential that a rapid quench is achieved. During the freeze-stop, light conditions should be kept unaltered. The frozen material should be kept under liquid N2 or can be stored in a freezer at −80°C for up to 3 months until use. 3. To homogenise the frozen leaves into a fine powder, frozen leaf material (5–10 g fresh weight) is placed into a stainless steel container, containing metal mill balls and pre-cooled with liquid N2, which is subsequently fixed into an oscillating ballmill (Retsch mill). The material is then homogenised at 20 Hz for 2 min (see Note 6). The homogenised material is subsequently transferred to 50-mL Falcon tubes placed in liquid N2 (see Note 7). 4. Place the 50-mL Falcon tubes containing the homogenous frozen powder (bits ca. 5 mm) into a pre-cooled aluminium holder inside the lyophiliser which has been pre-cooled under −50°C. During lyophilisation, cover the Falcon tubes with a paper tissue or Miracloth attached with an elastic rubber band. Do not close them with hermetic caps when the tubes are inside the lyophiliser; keep the Falcon caps for closing the tubes
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hermetically when taking samples out of the lyophiliser. The material should be dried at −50°C and 0.03 Torr for ~70 h, before ventilating the lyophiliser with dry N2 gas (see Note 8). Ensure that the temperature of the sample rises to approximately room temperature (RT) before releasing the vacuum or opening the chamber. When there is no water left in the sample, sublimation stops and the sample approximates chamber temperature. Never interrupt your runs, release the vacuum, or add new samples to the lyophiliser while you are using it (see Note 9). Patiently wait 3 days until lyophilisation is complete. 5. After lyophilisation, close each Falcon tube with the respective cap as fast as possible. Wear plastic gloves for this step; the hand of a nervous scientist with sweating hands can be a sufficient source of water vapour to affect the dry powder and spoil your NAF gradient. The Falcon tubes containing the completely dried material with closed caps are then transferred to a desiccator containing silica gel and a moisture indicator. The dried leaf material can be used immediately for fractionation or be stored inside the desiccator at RT for 1–3 days or in sealed plastic bags containing silica gel at −20°C until use (storage <1 month). When using samples after cold storage, put the samples within the sealed bag inside the desiccator until the sample reaches RT (see Notes 10–12). 3.2. Ultrasonication and Non-aqueous Density Gradient Fractionation
1. Resuspension of particles. Add 20 mL of tetrachlorethylene/ heptane mixture (v/v 66:34; d = 1.3 g/cm3) to the homogenised material in the Falcon tube and mix gently until a homogenous suspension is formed. Try to keep the tubes always closed with a hermetic cap, and work as much as possible inside the desiccator. 2. Sonication. Place the Falcon tube in the sonicator (the needle of the sonicator should be ~2 cm deep into the solution). In order to maintain the tube closed, use a foam stopper with a hole for the ultrasonic needle. Place the Falcon tube in a precooled metal block to avoid overheating during sonication. Sonicate for 2 min (in 10 s steps with 2 s breaks) with a power setting of 60%. After sonication, close the tube rapidly with a clean and dry cap and place the tube back into the desiccator. 3. Filtration. In order to remove particles that are too big, the powder suspension is filtered through a <30 mm monolen sieve attached to a plastic funnel into a new Falcon tube. Do this step inside the desiccator. Shake the funnel in order to speed up the procedure. Wash the filter with 10–20 mL of heptane. 4. Precipitation/concentration. Heptane is a “light” solvent (d = 0.68 g/cm3), and tetrachlorethylene is a “heavy” solvent (d = 1.62 g/cm3). Precipitate your particles by adding heptane up to ~40 mL in the Falcon tube. Shake the tube well and centrifuge at 5,000 × g for 10 min at ~10°C.
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5. During centrifugation, turn on the peristaltic pump (Econo Pump) for the gradient formation and “equilibrate” it with the water-free solvents. To do this, place tetrachlorethylene in B (in a Falcon tube with a small hole in the stopper and molecular sieves) and heptane in A (also in a Falcon with a holedstopper and molecular sieve); run 66% B (0.5 mL/min [do not pump faster, as this is the maximum for the diameter of the tubes]) if you are working with leaves (80% B may be more suitable for other sample types, such as potato tubers), until the whole system is filled and there are NO air bubbles. Check that all the tubes are working fine and that there are no leaks. Organic solvents harm the silicone tubes after a few uses (flexible silicon tubes are used for the peristaltic pump, while hard teflon tubes are used for the rest of the system). If the pump has problems, check all tubes and fittings. The silicon tube at the peristaltic pump should be changed approximately every four gradients. 6. After centrifugation (step 4 above), wait until the tubes reach RT and then discard the supernatant (“halogen waste”). Resuspend the particles in 5 mL of tetrachlorethylene/heptane mixture (66:34). 7. Gradient formation: (a) Transfer 4× aliquots of 200 mL from your homogenous particle suspension into 4× 2-mL Eppendorf tubes using cut yellow tips, and label them appropriately (these are called total aliquots); the reason to take four total aliquots is to have at least two replicas for measuring enzymes and metabolites separately. Use a micropipette to add 800 mL of heptane to each one, mix well, and then centrifuge them at 14,000 × g for 5 min at room temperature. Discard the supernatants and keep the pellets. Later they will be dried together with the fractions (see steps h–j below). (b) Determine the exact volume of the rest of your sample solution (~5 mL) and transfer it into a new Falcon tube. The volume is very important for subsequent calculations of recovery values. The total aliquots can give you an estimate of the total amount applied to the gradient if you know all the volumes accurately. It is also important that you determine the dry weight of the total aliquots. By this, you can calculate results of metabolite concentrations on a dry weight basis. The ratio of volumes multiplied by the dry weight (DW) of the aliquot pellet gives a correction factor that is specific for each individual gradient (e.g., 0.2 g DW × 200 mL/5,000 mL). The recovery refers to the ratio of the value obtained in the total aliquot and the value obtained in the gradient (the sum of all fractions).
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(c) Close the tube with the stopper containing a small hole and pass the metal needle through it until it reaches the bottom of the tube. Programme the gradient as follows: 60 min (~30 mL) 66–100% B (1.3–1.62 g/cm3) + 10 min (~5 mL) 100% B for Arabidopsis leaves. Alternatively, for other sample types such as potato tubers, the following may be more appropriate: 60 min (~30 mL) 80–100% B (1.43–1.62 g/cm3) + 20 min (~10 mL) 100% B. Run the gradient so that it forms below the sample in the Falcon tube. Always equilibrate the pump system immediately before usage and ensure that there is enough solvent for the whole pumping time! (d) When the gradient is finished, close the Falcon tube with a new hermetic cap and handle the Falcon tube very carefully. Avoid strong movements since the gradient is very unstable and can mix very easily. During the gradient formation, prepare a balancing Falcon tube (“weight tara”) for the centrifugation; use a Falcon tube containing the same volume (~40 mL) of 66:34 solvent mixture for the balance, as water is not “heavy” enough. (e) Centrifugation. Place the tubes in the pre-cooled centrifuge with swing-out rotor and centrifuge at 4°C. Turn off the rotor brake and select minimal acceleration and deceleration. Centrifuge at 4,000 rpm (~5,000 × g) for 60 min at least. (f) Fraction collection. After centrifugation, place the gradients (CAREFULLY) in the desiccator and wait (as always) until they reach RT before opening the caps. Securely place the Falcon tubes at eye height in a universal holder grip. Observe the particles as they have accumulated in the gradient (see Fig. 2). Divide each gradient into 3–6 fractions. Mark the boundaries of the fractions to be taken with a pencil on the wall of the Falcon tube. Use a 1-mL pipette with a cut blue tip to collect the fractions by hand. Collect fractions from top to bottom (collect first fraction 3 then fraction 2 and so on). The pellet fraction p is collected at the end. Transfer each of the fractions (which should on average be ~2–6 mL) into separate 15-mL Falcon tubes (fractions p, 0, 1, 2, 3). (g) Add heptane to the Falcon tubes to fill them up to ~12 mL. Close the caps and mix well. Each fraction must then be divided into two parts for measuring the marker enzymes (~1/3 of the volume: 4 mL) and the metabolites (~2/3 of the volume: 8 mL). For the marker enzyme samples, transfer equal aliquots (2 × 2 mL) into two 2-mL Eppendorf Safe-Lock tubes. Keep the remaining 8 mL in the Falcon tubes for measuring metabolites. Depending on the family
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Before Sample
After centrifugation
Fractions
4 3 2
Plastidial Thylakoid Plastidial Stromal Cytosol
1
Vacuolar
0 p LEAF
Cytosol
3 2 1
Amyloplast Cytosol Vacuolar
0 p TUBER
Fig. 2. Fraction positions in different tissues and schematic representation of tubes containing the density gradient. Particles move to a respective density layer after centrifugation yielding a partial enrichment of enzymes/metabolites of a given sub-cellular compartment. The enrichment of markers is shown for Arabidopsis leaf samples and also for potato tubers as comparison. Vacuolar marker enzymes are enriched at the bottom (heavy fractions), whereas plastidial and cytosolic markers are enriched in the upper layers (light fractions). A band in the middle area can be formed when a great amount of starch is present (e.g., in potato tuber samples). A critical point of the NAF procedure is how to take manually the fractions to get a good enrichment of sub-cellular compartments. The separation of the vacuole is always very good. Most critical is the separation of AGPase and UGPase. Pay special attention when taking fractions 1–3.
of metabolites to be analysed, use separate aliquots for GC-MS or HPLC measurements (which require different extraction protocols: methanol or trichloroacetic acid extraction, respectively). (h) The exact volumes used for subsampling are later needed for the calculations. Label all the tubes appropriately. Remember to add heptane (filling up to ~1.2 mL) also to the four aliquots taken before the fractionation (i.e., the total aliquots taken in steps a and b)! (i) Concentration of fractions. Precipitate the particles by centrifuging at maximum speed (4,000 rpm in the Eppendorf swing-out rotor; equivalent to 5,000 × g) for 10 min at 10°C. Samples in 2-mL Eppendorf tubes are centrifuged separately in a microcentrifuge (14,000 rpm equivalent to ~14,000 × g). Remove the supernatants (halogen waste) and keep the pellets. (j) Place the pellet samples (Falcon tubes for metabolites and Eppendorf tubes for enzymes and total aliquots) in the desiccator containing silica gel and connected to a vacuum pump. Keep the samples under vacuum until they are completely dry (~4 h). Dried fractions can be stored in boxes with silica gel in a plastic bag at −20°C.
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We recommend that investigators take the following general precautions during the course of this work. Keep all the tubes closed and within the desiccator when they are not in use, and never open cold tubes. Avoid having any water solutions nearby and always keep your gloves dry. Work under the fume hood and wear gloves. Solvents are hazardous and produce headaches because of the bad smell. 3.3. Sample Preparation and Measurement of Marker Enzyme Activities
1. Prepare ~10 mL enzyme extraction buffer per sample. 2. Equilibrate the NAP-5 columns with 3× bed volumes (~3 mL) of enzyme extraction buffer and keep them cool (on ice or inside the cold room). 3. Select all your marker enzyme Eppendorf tubes with dry pellets. Weigh the tubes with an analytical balance (after extraction, weigh the empty tube again, so that you can later calculate the weight of the dry pellet). Add 500 mL of enzyme extraction buffer to each sample (total aliquot and fractions p, 0, 1, 2, and 3). The dry pellets of leaf samples are very difficult to dissolve; therefore, add some small plastic or glass beads (~1 mm diameter) to facilitate resuspension of the particles. Vortex very well until all the material is in solution (10 min in a Thermomixer at 4°C). Keep your samples always at ice temperature. 4. Centrifuge at maximum speed (14,000 × g) in a microfuge for 5 min at 4°C. Use the supernatants for enzyme measurements. Enzyme extraction pellets can be used for starch determination; store pellets frozen at −20°C, or add 80% (v/v) ethanol right away. 5. Desalting. Transfer 500 mL of the supernatant extract (from step 4 above) to the pre-equilibrated NAP-5 columns (in the cold room, at 4°C), and elute immediately with 1 mL of enzyme extraction buffer. Keep the eluates in new labelled 2-mL Eppendorf tubes on ice. Do not let extracts stand for longer than 60 min. Measure marker enzymes right away (without freezing the samples). After the first round of marker enzyme measurements, the extracts can be frozen in liquid N2 and stored for several weeks at −80°C for a second round of marker/enzyme measurements. 6. Enzyme measurement. To check if the NAF gradients were good, measure the following activities in all fractions right after enzyme extraction: AGPase (plastid), UGPase (cytosol), and a-mannosidase (vacuole). Prepare activity assays and buffers before you extract the enzymes (see Tables 1–3). Measure enzyme activities in 96-well microtitre format, as described in steps 7–9 below. 7. To measure UGPase, add 2–5 mL of each fraction to 200 mL of Master-mix (see Table 1). Start the reaction by adding 2 mL of
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100 mM pyrophosphate (PPi) to each well (final PPi concentration, 1 mM). Perform measurements at 340 nm, with kinetic readings every 30 s. 8. To measure AGPase, add 20–50 mL of each fraction to 200 mL of Master-mix (see Table 2). Start the reaction by adding 2 mL of 100 mM PPi to each well (final PPi concentration, 1 mM). Perform measurements at 340 nm, with kinetic readings every 30 s. 9. To measure a-mannosidase, use the following previously described method (16) (see Table 3). To 100 mL of citrateKOH buffer, add 20 mL of sample (fraction) and 30 mL of p-nitrophenyl-d-a-mannoside. Stop the blank wells immediately with 100 mL of 0.8 M borate buffer, without leaving time for activity. Incubate the samples at 30°C for 40 min when using Arabidopsis leaf material (or for 20 min when using potato tuber samples). After adding the borate buffer to stop the reactions, follow the absorbance in the microplate for 10 min at 405 nm. Use the mean absorbance value after 10 min for the calculations. The absorption coefficient of the reaction product, p-nitrophenyl, is 18.5 mL/cm × mmol. 10. Calculate the relative distribution of the enzymes in each fraction and the recovered activity from the aliquot taken before separation (see Tables 4–6). For a more detailed description of this, see Subheading 3.6 below. 11. Optional: If the gradients are judged to be good, one may consider measuring additional markers, like other enzymes, chlorophyll content, or protein amount quantified by immunology (using stored samples at −80°C). Examples of marker enzyme enrichments in different samples are given in Fig. 3. 3.4. Sample Extraction for Gas Chromato graphic Analysis
In the following section, a procedure is described for GC-MS-based metabolic profiling. Use aliquots of the metabolite samples for this purpose (from Subheading 3.2, step 7j). Use additional aliquots for other profiling approaches (e.g., HPLC or UV–VIS spectrometry) that require other extraction protocols. 1. Add 1.5 mL of pre-cooled methanol to each sample (total aliquot t and fractions p, 0, 1, 2, and 3), and vortex for 15 s. 2. Add 60 mL of internal standard Ribitol to each tube and vortex for 15 s. 3. Shake the vials at 70°C for 10 min at 1,300 rpm using a Thermomixer. 4. Centrifuge the tubes for 10 min at 14,000 × g in a microcentrifuge at 4°C. 5. Carefully transfer each supernatant to a glass vial (Schott GL14).
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a 50% mannosidase AGPase
activity (% of total found)
40%
UGPase 30%
PFP citrate synthase
20%
10%
0%
b
p
0
1 fraction
2
3
60%
Vacuole
50%
Cytosol
Plastid Mito
recovery
40% 30% 20% 10% 0%
37a p
37a 0
37a 1
37a 2
37a 3
37a 4
fraction
Fig. 3. Examples of marker enzyme enrichments in different samples. (a) Marker–enzyme distribution in potato tubers. Averaged data from a representative tuber gradient is presented, divided into five different fractions (p, 0, 1, 2, 3). (b) Marker–enzyme distribution in Arabidopsis leaves. Data from a representative Arabidopsis leaf gradient (no. 37a) are presented, divided into six different fractions (p, 0, 1, 2, 3, 4). Leaf marker enzymes were measured twice and SD are presented. Note the significant differences between vacuolar, cytosolic, and plastidic compartments. Note also the similarity between the levels of cytosolic and mitochondrial markers. When no significant enrichment can be achieved between fractions for marker enzymes, then the relevant sub-cellular compartments have to be assumed as one pooled compartment.
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6. To each vial, add 0.75 mL of pre-cooled chloroform and then 1.5 mL of cold dH2O (4°C), and vortex again for 15 s. 7. Centrifuge the tubes for 15 min at 2,200 × g at 4°C. 8. Transfer 150 mL from the upper polar phase into a glass tube suitable for GC-MS analysis. 9. As a backup, keep a second aliquot in a separate tube. You can also take a larger aliquot (e.g., 500 mL) in the case that you are interested in low-abundance metabolites. 10. Vacuum-dry the samples at RT (e.g., in a Maxi-Dry Lyo for approximately 10 h). 11. Fill the tubes with argon gas (see Note 13). 12. Keep the tubes in a closed box containing a desiccant and store them at −80°C (for no longer than 1 week). 3.5. Chemical Derivatisation of the Samples for GC-MS
For gas chromatography (GC), the volatility, thermal stability, and chromatographic mobility of mostly polar organic metabolites like organic acids, amino acids, sugars, and fatty acids has to be increased. This is achieved by chemical derivatisation. For GC-MS equipment, derivatisation is done by oxime formation and subsequent trimethylsilylation, which reduces the dipole–dipole interaction since it replaces hydrogen on polar groups with trimethylsilyl groups. For other detection methods, such as flame ionisation detector (GC-FID), other derivatisation procedures might be more appropriate. It has to be considered that volatile derivatives of many secondary metabolites are unstable and, therefore, cannot be detected by GC-MS. Here, we describe the procedures of sample preparation and derivatisation for metabolic profiling. For detailed downstream GC-TOF-MS metabolite profiling, such as instrument settings and evaluating the resultant chromatograms, the reader should refer to Lisec et al. (3). The following protocol for derivatisation is adapted from Lisec et al. (3) and can be performed by hand using accurate micropipettes. However, for better reproducibility, it is recommended to use an HPLC autosampler module or a pipetting robot. 1. After removing the samples from the freezer (see Subheading 3.4, step 12) dry them again in a vacuum concentrator for 30 min. Samples should be totally free of water since it interferes with the chemical derivatisation procedure. 2. For each of 10 mg of dry weight in the samples, add 40 mL of methoxyamination solution to the sample and keep the tube closed (see Note 2). 3. As a control, add 40 mL of methoxyamination solution to an empty 1.5-mL Eppendorf tube. 4. Place the tubes in a shaker (Thermomixer) for 2 h at 37°C.
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5. Mix MSTFA reagent with the retention-time standards (suitable retention standards might be a mixture of alkanes with chain lengths from C8 to C20). 6. Add 70 mL of the MSTFA mixture prepared in step 5 to the samples after the incubation in step 4. 7. Mix the samples well and place the tubes again in the shaker for 2 h at 37°C. 8. After the incubation, keep the samples at 4°C and protected from light. Do not store the samples, but use them directly for GC-MS analysis. Place the samples in the GC autosampler for the metabolite measurements (inject 1–10 mL of sample). 3.6. Calculation of Sub-cellular Metabolite Concentrations
1. Inject your sample, usually 1 mL, with high or low concentration of metabolite in splitless or split mode, respectively, into the gas chromatograph connected with mass spectrometer and the autosampler. Perform chromatography with an MDN-35 capillary column using helium carrier gas. For deconvolution of the chromatogram, use the ChromaTOF software. Analyse GC-MS peak areas and obtain quantitative GC-MS data per metabolite and per sample with respect to the Ribitol internal standard. 2. Metabolite distribution. Measure the percentage of markers and metabolites in each fraction and calculate the recovery from the aliquot taken before separation. See Tables 4–6 for examples. 3. Calculations of sub-cellular distributions. The distribution of your marker enzymes in the fractions has to be determined experimentally (see Subheading 3.3 and the example results in Table 4). Similarly, the distribution of metabolites in the fractions has also to be determined experimentally (see Subheading 3.5 and the example results in Table 5). The relative distribution of metabolites in the sub-cellular compartments (not in the enriched fractions, but extrapolated to the pure cytosol, vacuole, and plastid) needs to be calculated using the Solver add-in function of Microsoft Excel. By finding the best fit (minimum value for the sum of all squared differences, the so-called “least squared method” in statistics), the theoretical distribution of the metabolite in each sub-cellular compartment can be obtained (see the example best fit in Table 6). The Excel formulae that need to be defined in the corresponding cells are indicated in Tables 5 and 6. The solver is used to find the minimum value for cell E28 (Table 5) by modifying cells J23, J24, and J25 (Table 6). The solver procedure needs also to be defined so as to not accept negative values for any of the fitted cells J23:J25. This means that the mathematical negative solutions are excluded. This is practically achieved by defining the following restrictions for the cells in the solver dialogue box in Excel: J23 ³ 0, J24 ³ 0, and J25 ³ 0 (Table 6).
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Define all formulae and restrictions and run the solver and accept the best fit result. The values you obtain in cells I23:I25 (Table 6) are your final result that you can then publish as your sub-cellular distribution (NAF data). The value you obtain in cell D21 (Table 5) or cell I31 (Table 6) is the recovery values that you can also report as a reliability measure. Ideally, metabolite recovery should be between 80 and 120%. 4. Calculations of sub-cellular volumes. The sub-cellular structure of the tissue can be analysed via transmission electron microscopy (TEM) (6, 20). Representative parts of the tissue need to be photographed and the pictures have to be printed onto DIN A4 paper. The paper has to be hand cut with scissors and the relative surface of sub-cellular compartments (vacuole, cytosol, plastid, mitochondria, nucleus, cell wall, and apoplast) can be determined by weighing the paper pieces with an analytical balance. The percentage weight equals the percentage surface. Given a high number of samples and pictures analysed (>30), the value for the relative volume of the sub-cellular compartments is very close to the measured value of the relative surface of the sub-cellular compartments in the pictures (6). This applies regardless of the magnification of the TEM pictures. Low-magnification pictures (700×) in which several cells could be visualised should be preferentially used for the determination of most compartments. High-magnification pictures (>1,000×) need to be used to determine the volume of mitochondria which are not always visible in the lowmagnification pictures. The relative volumes (% of total) can be converted to absolute volumes per unit of mass by taking into account the specific gravity (mass per unit of volume). For example, for growing potato tuber tissue, it is 1.017 g FW/mL or 0.3 g DW/mL (20) (FW, fresh weight; DW, dry weight). The concentration of metabolites in the sub-cellular compartments can be calculated by dividing the amount of metabolite in a given compartment (nmol/g DW) by the volume of the sub-cellular compartment (mL/g DW). 3.7. Concluding Remarks
The NAF technique is a work-intensive procedure and requires a lot of investment (time for learning and training, effort for method establishment and data analysis, and finance for obtaining required equipment and materials, etc.). However, it is the only currently available method that can be used to reliably calculate the subcellular distribution of metabolites in plant tissues. For measuring the sub-cellular distribution of proteins and enzymes only (which do not rapidly redistribute during fractionation), standard aqueous fractionation methods are much easier to implement. The whole NAF procedure can be considered as a crown method of plant biochemistry. Students should master many analytical methods before they can expect to be successful with NAF.
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Beginners should first learn to make extractions, measure metabolites, assay a variety of different enzymes, master Beer–Lambert law, and be professional in the usage of microplates, spectrophotometers, HPLC, and GC-MS before attempting NAF as described in this chapter. Many scientists have attempted to do NAF believing that it is a standard cookbook method, but have failed at one or the other step. Some scientists underestimate the time required for mastering the method (>6 months) and others have encountered difficulties at the very end, when analysing the data, because the entire relevant information set was not collected during fractionation, or because it is hard to understand the mathematics behind the linear regression formula for calculating sub-cellular metabolite levels.
4. Notes 1. All solvents must be kept water-free with molecular sieves (prepare solvents and solvent mixtures at least 2 days before use). 2. Wear suitable gloves and eye/face protection and work under the fume hood because methoxyamine hydrochloride causes skin burns. 3. The reagent is very toxic and should be handled accordingly under the fume hood. Wear suitable hand gloves and eye/face protection. 4. Fractionation of chloroplasts relies on the high content of thylakoid lipids and stromal proteins (Rubisco), as well as on the high content of starch inside the chloroplast. Dark-adapted leaves with reduced amounts of starch yield a different fractionation pattern than light-adapted leaf samples with starch-containing chloroplasts. The same is true for Arabidopsis mutant plants with altered plastidial and thylakoid structure. 5. The forceps harvesting procedure is as follows: cut rosette plants with scissors at the base of the root while still maintained in the light. Use the plastic forceps to hold the rosette keeping the leaves blades at one side of the forceps and the petioles and rosette stem on the other side of the forceps. Holding the forceps closed, rapidly submerge individual plants inside a small Dewar containing clean liquid N2. By using the forceps procedure, one can freeze the whole plant within 1 s while simultaneously getting rid of the stem and leaf petioles. Use the plastic sieve to collect leaf tissue into a properly labelled aluminium bag. Other harvesting procedures typical for molecular biological purposes are inapplicable for studying chloroplast metabolism. One should avoid moving plants from the growth chamber to the laboratory, cutting the leaves individually, washing them with
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water to get rid of the soil, and harvesting the material slowly (>10 s is too slow). Sterility is much less important for biochemistry than rapidity and maintenance of a cool temperature. 6. Efficient homogenisation using a liquid N2-cooled ball-mill is necessary to break the leaves into small fragments of around 1–5 mm of size, which are enriched in material from a given compartment. The use of mortar and pestle does not allow getting particles that are small enough for NAF. Care must be taken not to thaw the material during homogenisation. During homogenisation, wear cotton gloves and goggles to protect yourself and the sample! 7. The Falcon tubes should not be full. Fill up to 20 mL of frozen powder into the tubes. Prefill the tubes with some liquid N2, for further cooling the powder. The easiest way is to place the open 50-mL Falcon tubes in a metal holder inside a polystyrene box filled with liquid N2. Use the same metal holder during lyophilisation (see Note 8). 8. Lyophilisation. Read carefully the instructions for the lyophiliser equipment and become familiar with the phenomenon of sublimation and temperature–pressure curves. Pre-cool the aluminium holder with liquid N2. Place the Falcon tubes (with powder plus nitrogen) in the metal holder in a horizontal position into the lyophiliser, so that the powder covers most of the wall of the Falcon tube. Lyophilisation takes at least 2 days (~60 h). Do not prepare too much of the same material at once, as sometimes problems might arise during the lyophilisation procedure and the sample gets lost. Never interrupt the vacuum before the drying process is completely finished. 9. Ensure that the vacuum in the lyophiliser drops rapidly enough before the temperature in the chamber reaches approximately −5°C. Check all the junctions of the chamber (they must be clean and dry) and ensure that all valves are closed. If the vacuum does not drop fast enough, be prepared to place your samples as quickly as possible back into liquid N2. 10. Moisture condensation of cold samples can be a severe obstacle for successful fractionation. Try to avoid leaving the tubes with lyophilised powder outside the desiccator, especially when they are still cold. The powder is VERY hygroscopic and might attract water even in the closed Falcon tubes! 11. Approximately 3–5 g dry weight start-up material is needed per gradient. Perform at least three repetitions with independently harvested material (biological replicates). The reason for so much starting material per gradient is because most of the material is retained in the 30-mm sieve. There is no advantage of using particles larger than whole cells (>30 mm), since no subcellular enrichment can be achieved. In the ideal case, particles
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are so small that they only contain material from a single compartment. In addition, the fractionation in 3–6 samples and the dual use of each fraction for metabolite and enzyme assays increases the amount of tissue needed. At least 2 g dry weight post-sieve material should be loaded onto each gradient. 12. Run one or two gradients in 1 day and measure marker enzymes the next day. Do not run many gradients on consecutive days because the distribution of markers needs to be controlled each time. The optimal collection of fractions can only be performed in an iterative process of observing the distribution of particles in the gradient, collecting fractions of different size, and measuring the distribution of markers. Try to remember how you did collect the fractions and how the markers were distributed/ enriched. The NAF procedure requires a lot of learning and practice (a new student needs at least 6 months of practice and hard work to obtain useful enriched fractions and reliable NAF data). All solvents and samples must be kept water-free at all possible times. Keep all the tubes closed and within the desiccator when not using them, and never open cold tubes. Avoid having any water solutions nearby, and always keep your gloves dry. Work under the fume hood and wear gloves, as the solvents used are hazardous. 13. Overlaying aliquots with argon gas avoids oxidative degradation of the metabolites by air components.
Acknowledgments We are grateful to the Deutsche Forschungsgemeinschaft for support (grants Ge 878/1-1 and 5-1 and SFB TR1). References 1. Sumner, L. W., Mendes, P., and Dixon, R. A. (2003) Plant metabolomics: large-scale phytochemistry in the functional genomics era. Phytochemistry 62, 817–836. 2. Trethewey, R. N. (2004) Metabolite profiling as an aid to metabolic engineering in plants. Curr. Opin. Plant Biol. 7, 96–201. 3. Lisec, J., Schauer, N., Kopka, J., Willmitzer, L., and Fernie, A.R. (2006) Gas chromatography mass spectrometry-based metabolite profiling in plants. Nat. Protoc. 1, 387–396. 4. Fiehn, O., Kopka, J., Dörmann, P., Altmann, T., Trethewey, R.N., and Willmitzer, L. (2000) Metabolite profiling for plant functional genomics. Nat. Biotechnol. 18, 1157–1161.
5. Roessner, U., Luedemann, A., Brust, D., Fiehn, O., Linke, T., Willmitzer, L., and Fernie, A. (2001) Metabolic profiling allows comprehensive phenotyping of genetically or environmentally modified plant systems. Plant Cell 13, 11–29. 6. Farre, E. M., Tiessen, A., Roessner, U., Geigenberger, P., Trethewey, R. N., and Willmitzer, L. (2001) Analysis of the compartmentation of glycolytic intermediates, nucleotides, sugars, amino acids and sugar alcohols in potato tubers using a non-aqueous fractionation method. Plant Physiol. 127, 685–700. 7. Fernie, A. R., Trethewey, R. N., Krotzky, A. J., and Willmitzer, L. (2004) Metabolite profiling:
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P. Geigenberger et al. from diagnostics to systems biology (Innovation). Nat. Rev. Mol. Cell Biol. 5, 763–769. Kopka, J., Fernie, A., Weckwerth, W., Gibon, Y., and Stitt, M. (2004) Metabolite profiling in plant biology: platforms and destinations. Genome Biol. 5, 109. Messerli, G., Nia, V. P., Trevisan, M., Kolbe, A., Schauer, N., Geigenberger, P., Chen, J., Davison, A. C., Fernie, A. R., and Zeeman, S. C. (2007) Rapid classification of phenotypic mutants of Arabidopsis via metabolite fingerprinting. Plant Physiol. 143, 1484–1492. van Dongen, J. T., Fröhlich, A., RamirezAguilar, S., Schauer, N., Fernie, A. R., Erban, A., Kopka, J., Clark, J., Langer, A., and Geigenberger, P. (2009) Transcript and metabolite profiling of the adaptive response to mild decreases in oxygen concentration in the roots of Arabidopsis plants. Ann. Bot. 103, 269–280. Meyer, R.C., Steinfath, M., Lisec, J., Becher, M., Witucka-Wall, H., Törjek, O., Fiehn, O., Eckardt, A., Willmitzer, L., Selbig, J., and Altmann, T. (2007) The metabolic signature related to high plant growth rate in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 104, 4759–4764. Jacobsen, M., Mattow, J., Repsilber, D., and Kaufmann, S. H. (2008) Novel strategies to identify biomarkers in tuberculosis. Biol. Chem. 389, 487–495. Steinfath, M., Strehmel, N., Peters, R., Groth, D., Hummel, J., Steup, M., Selbig, J., Kopka, J., Geigenberger, P., and Van Dongen, J. T. (2010) Discovering plant metabolic biomarkers for phenotype prediction using an untargeted approach. Plant Biotechnol. J. 8, 1–12. Bräutigam, K., Dietzel, L., Kleine, T., Ströher, E., Wormuth, D., Dietz, K.-J., Radke, D., Wirtz, M., Hell, R., Dörmann, P., Nunes-Nesi, A., Schauer, N., Fernie, A. R., Oliver, S. N., Geigenberger, P., Leister, D., Pfannschmidt, T. (2009) Dynamic plastid redox signals integrate gene expression and metabolism to
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induce distinct metabolic states in photosynthetic acclimation. Plant Cell 21, 2715–2732. Renberg, L., Johansson, A. I., Shutova, T., Stenlund, H., Aksmann, A., Raven, J. A., Gardeström, P., Moritz, T., and Samuelsson, G. (2010) A Metabolomic Approach to Study Major Metabolite Changes during Acclimation to Limiting CO2 in Chlamydomonas reinhardtii. Plant Physiol. 154, 187–196. Stitt, M., Lilley, R. M., Gerhardt, R., and Heldt, H. W. (1989) Metabolite levels in specific cells and subcellular compartments of plant leaves. Methods Enzymol. 174, 518–550. Heineke, D., Lohaus, G., and Winter, H. (1997) Compartmentation of C/N metabolism. In, A Molecular Approach to Primary Metabolism in Higher Plants (Foyer, C. H., and Quick, W. P., eds.) Tailor and Francis Ltd., London, UK, pp. 205–217. Fettke, J., Eckermann, N., Tiessen, A., Geigenberger, P., and Steup, M. (2005) Identification, subcellular localisation and biochemical characterisation of water-soluble heteroglycans (SHG) in leaves of Arabidopsis thaliana L.: distinct SHG reside in the cytosol and in the apoplast. Plant J. 43, 568–585. Fettke, J., Eckermann, N., Poeste, S., Pauly, M., Tiessen, A., Geigenberger, P., and Steup, M. (2006) Analysis of cytosolic heteroglycans from leaves of transgenic potato (Solanum tuberosum L.) plants that under- or overexpress the pho-2 phosphorylase isozyme. Plant Cell Physiol. 46, 1987–2004. Tiessen, A., Hendriks, J. H. M., Stitt, M., Branscheid, A., Gibon, Y., Farré, E. M., and Geigenberger, P. (2002) Starch synthesis in potato tubers is regulated by post-translational redox-modification of ADP-glucose pyrophosphorylase: a novel regulatory mechanism linking starch synthesis to the sucrose supply. Plant Cell 14, 2191–2213.
Chapter 9 Chloroplast Phenomics: Systematic Phenotypic Screening of Chloroplast Protein Mutants in Arabidopsis Yan Lu, Linda J. Savage, and Robert L. Last Abstract As part of a project to analyze chloroplast functional networks systematically, we have subjected mutants in >3,200 nuclear genes predicted to encode chloroplast-targeted proteins in Arabidopsis thaliana (http:// www.plastid.msu.edu) to parallel phenotypic assays. Detailed methods are presented for the various assays being used in this project to study chloroplast biology. These include morphological analysis of plants, chloroplasts, and seeds using controlled vocabulary. Metabolites synthesized in the chloroplast such as starch, amino acids, and fatty acids are analyzed in groups according to their chemical properties. As an indicator for the relative composition of seed storage oil and proteins, the carbon and nitrogen contents are determined by an elemental analyzer. The methods in this chapter describe how the assays are configured to run in relatively high throughput, maximizing data quality. Key words: Amino acid, Seed composition, Chloroplast, Chlorophyll fluorescence, Fatty acid, Genotyping, Morphology, Starch, Systems biology, Reverse genetics, Phenomics
1. Introduction While reverse genetics approaches are commonly used to study the function of individual or small groups of genes, the approach often yields disappointing results due to the small number of mutants and assays employed. This approach also has limited ability to detect broader phenotypic syndromes and cross-pathway interactions. To try to overcome these factors and gain information for a large number of nuclear genes and the networks in which they function, it is desirable to test large numbers of mutants for diverse phenotypes. The Chloroplast 2010 project (http://www.plastid.msu.edu) makes use of recent advances in phenotyping technologies to screen
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a large number of mutants in a systematic manner (1). The reference dicot plant Arabidopsis thaliana has large collections of sequenceindexed insertional mutants (2–6) and these are well suited to use in large-scale phenotypic screens (1, 7–9). For projects that aim to analyze large numbers of samples for many phenotypes, sample tracking becomes challenging. The use of laboratory information management systems (LIMS) minimizes tracking error when processing thousands of samples (1). Screening a large collection of insertion mutants with a dozen chloroplast-oriented phenotypic assays would permit the detection of phenotypic syndromes in the mutants and genetic networks among chloroplast-target genes (1, 8). These phenotypic assays are modified to suit the requirements of high-throughput mutant screening, as described in the following sections. 1.1. Plant Care and Harvesting Schedule
Growth conditions, water and nutrient supplies, and harvesting time impact metabolite concentrations in harvested tissues. To minimize variations caused by these factors, protocols should be used consistently during seed planting, plant maintenance, and tissue harvesting.
1.2. Whole Plant Morphology
Assessment of whole plant morphology provides an overall measure of the structure, growth, and development of the plants. Evaluating plant morphology (and the morphology of chloroplasts and seeds) with controlled vocabulary ensures consistency of phenotypic descriptions and the ability to conduct consistent searches of the database (http://bioinfo.bch.msu.edu/2010_LIMS).
1.3. Leaf Starch Assay
Transitory starch is stored inside the chloroplast during the day when carbon assimilation takes place and is metabolized at night. While much is known about the starch synthesis and its regulation, starch degradation is still relatively poorly understood (10). Iodine staining of leaf disks to detect starch content provides a highthroughput method to identify novel genes that regulate starch synthesis and degradation (1, 11).
1.4. Leaf Amino Acid Assay
Amino acids contribute to many aspects of plant physiology, including nitrogen transport and storage, yet large gaps still exist in our understanding of the pathways and regulation of amino acids biosynthesis and catabolism (12, 13). Direct amino acid profiling is a powerful tool to identify genes controlling the regulation of amino acid biosynthesis, catabolism, and transport. This is done by extracting free amino acids and subjecting samples to highthroughput high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) (1, 12, 14).
1.5. Leaf Fatty Acid Assay
In plants, chloroplasts play a central role in fatty acid and acyl lipid biosynthesis, and much of the resultant lipid products are assembled into thylakoid membrane lipids. Fatty acid methyl ester (FAME)
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profiling is a robust diagnostic method to discover genes regulating fatty acid biosynthesis, turnover, and lipid biosynthesis (15, 16). The procedure uses hot methanolic HCl to release fatty acids from leaf lipids and simultaneously convert the fatty acids to corresponding FAME (1, 17). FAME content is subsequently analyzed using gas chromatography-flame ionization detection (GC-FID). 1.6. Chloroplast Morphology
Chloroplast morphology analysis can reveal genes of various functions, especially those involved in chloroplast division (18). The major processes include (a) fixing and macerating the base and apex of expanded leaves, and (b) analyzing samples with an inverted microscope and polarization contrast optics (1, 18).
1.7. DNA Archiving and Genotyping
The value of data generated by reverse genetics projects is critically dependent on the identification of available homozygous mutants (8). To ensure proper interpretation of phenotypic data of T-DNA insertion mutants, it is important to archive DNA and genotype all individual mutants. Because of the large numbers of samples in the Chloroplast 2010 project, a relatively high-throughput genotyping assay was developed (8).
1.8. Seed Morphology
Arabidopsis seed plastids are photoheterotrophic and thus carry out a wide range of metabolism (19). Morphology and coloration are easy-to-score phenotypes of mature seeds, which can be associated with compositional changes in seeds (20, 21). Seed phenotypes are visually inspected with a stereomicroscope and recorded using controlled vocabulary (1).
1.9. Seed Amino Acid Assay
Due to differential expression of genes in amino acid biosynthesis and degradation, mutants with altered amino acid composition in leaf do not always have similar amino acid pattern in seeds and vice versa (12–14, 22). Parallel analysis of leaf and seed amino acids in mutants is performed to identify genes that regulate long-distance transport of amino acids, and the storage of amino acids and their precursors into developing seeds (1, 12–14). The procedure for seed free amino acid analysis is similar to that for leaf amino acids.
1.10. Seed C and N Assay
Triacylglycerols and storage proteins are the major seed storage components in Arabidopsis. The production of triacylglycerols and storage proteins depends upon the biosynthesis of fatty acid and amino acid precursors in the plastid. Because ~90% of seed nitrogen is in storage proteins and >50% seed carbon is in oil (calculated from data in ref. 19), measurement of seed C/N ratio provides a convenient method to estimate the relative composition of storage oil and proteins in seeds (21). The procedure involves (a) accurate weighing of desiccated seeds with a microbalance, and (b) quantification of C and N contents with an elemental analyzer (1).
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1.11. Seed Starch Assay
In Arabidopsis seeds, starch accumulates in developing seeds and is then degraded later in development when lipids and proteins are synthesized (20, 23). Iodine staining of seed coat starch is a highthroughput method to identify genes regulating starch accumulation and mobilization in seeds (1).
1.12. Chlorophyll Fluorescence Assay
Analysis of in vivo chlorophyll fluorescence is a powerful, noninvasive technique to identify mutations affecting photosynthesis (24–26). This assay is done by simultaneous measurement of 12 plants with a pulse-amplitude modulated (PAM) fluorescence imaging system (1, 27).
1.13. Systematic Data Analysis
Screening several thousand mutants for more than 80 morphological, metabolic, and physiological traits requires systematic data normalization and conversion. This is done by converting the concentration of metabolites to median-adjusted z-scores and converting morphological and qualitative data to numeric codes (1, 28).
2. Materials 2.1. Plant Care and Harvesting Schedule
1. 32-Plug press-fill trays and 2.5″ (length) × 2.5″ (width) × 3.5″(height) press-fill pots (Hummert International, Earth City, MO, USA), barcoded. 2. 22″ (length) × 11″ (width) × 2.5″ (height) Perma-Nest plant trays (Growers Supply, Dexter, MI, USA), barcoded. 3. 22″ (length) × 11″ (width) × 3″ (height) clear plastic humidity domes (Growers Supply). 4. Redi-earth plug and seedling mix (SUN GRO Horticulture, Bellevue, WA, USA). 5. Fine-grade vermiculite (Thermo-O-Rock East, New Eagle, PA, USA). 6. Arabidopsis nutrient solution: 5 mM KNO3, 2.5 mM KH2PO4, 2 mM MgSO4, 2 mM Ca(NO3)2, 50 nM FeNaethylenediaminetetraacetic acid (EDTA), and 0.5 mL/L micronutrients (see below and Note 1). 7. Arabidopsis micronutrients: 70 mM H3BO3, 10 nM CoCl2, 5 mM CuSO4, 14 mM MnCl2, 200 nM NaMoO4, 10 mM NaCl, and 1 mM ZnSO4 (see Note 1). 8. Aracon bases and tubes (Lehle Seeds, Round Rock, TX, USA). 9. Matrix 1.6-mL 2D barcoded storage tubes and racks (Thermo Fisher Scientific, Waltham, MA, USA). 10. Secador™ 4.0 auto-desiccator cabinets (Bel-Art Products, Pequannock, NJ, USA).
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2.2. Whole Plant Morphology
1. Rosette size measuring guide (circles of different diameters printed on a clear transparency).
2.3. Leaf Starch Assay
1. Cork borer #2, 5.5-mm internal diameter (Cole-Parmer Instrument, Vernon Hills, IL, USA). 2. CoolSafe system for microplates (Diversified Biotech, Boston, MA, USA), −20°C. 3. 96-Well 360-mL flat-bottom polystyrene microplates and lids (Corning Inc., Corning, NY, USA), barcoded. 4. Aluminium sealing films (Excel Scientific, Victorville, CA, USA). 5. Killing solution: 80% (v/v) ethanol and 5% (v/v) formic acid. Store at room temperature (RT). 6. 80% (v/v) ethanol. Store at RT. 7. Lugol’s IKI solution: 5.7 mM iodine and 43.4 mM KI. The reagent is light sensitive; wrap the bottle with foil and store at 4°C.
2.4. Leaf Amino Acid Assay
1. 3-mm stainless steel beads (CCR Products, West Hartford, CT, USA) in 2-mL barcoded microfuge tubes (1 bead/tube). 2. Prefrozen (−80°C) 96-tube rack. 3. S2200 Dual Head paint shaker (Hero Products Group, Delta, BC, Canada). 4. Single use (50 mL) aliquots of 1 mM l-Phe-a,b,b,2,3,4,5,6-d8 (Cambridge Isotope Laboratories, Andover, MA, USA). Store at −80°C. 5. Single use (50 mL) aliquots of 10 mM dithiothreitol (DTT). Store at −80°C. 6. Matrix 1,250 mL 8-channel electronic pipette with expandable tip spacing (Thermo Fisher Scientific). 7. 1.1-mL MicroTubes™ strips, MicroCaps™ strips, MicroRacks™ (Dot Scientific, Burton, MI, USA).
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8. 96-Well MultiScreen® Solvinert filter plate with 0.45 mM lowbinding hydrophilic polytetrafluoroethylene (PTFE) membrane and 96-well v-bottom collection plate (Millipore, Billerica, MA, USA). 9. Single use (2 mL) aliquots of 9 mM l-Val-2,3,4,4,4,5,5,5-d8 (Cambridge Isotope Laboratories). Store at −80°C. 10. 96-Well full skirt PCR plates (Denville Scientific), barcoded. 11. Aluminium sealing films (Excel Scientific). 12. Silicone sealing mats for 96-well microplates (Axygen Scientific, Union City, CA, USA).
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13. Single use (~120 mL) aliquots of 0.01, 0.05, 0.1, 0.5, 1, 5, 10, 50, and 100 mM amino acids standards containing 9 mM DTT, 0.9 mM Phe-d8, 0.9 mM Val-d8, and varying concentrations of 20 protein amino acids, GABA, anthranilate, homo-Ser, Hyp, and S-methyl Met (see Note 2). 14. 12 mm (outer diameter) × 32 mm (length) 2-mL wide-opening screw-top vials (Agilent Technologies, Santa Clara, CA, USA). 15. Screw caps with PTFE and rubber septa for above vials (Agilent Technologies). 16. 200-mL conical bottom inserts for above vials (SUN SRI, Rockwood, TN, USA). 17. Symmetry® C18 3.5 mM (particle size), 2.1 mm (inner diameter) × 100 mm (length) analytical column (Waters, Milford, MA, USA). 18. 0.5-mM stainless steel high-pressure inline solvent filter and 0.5 mM (porosity) 0.062″ (inner diameter) × 0.062″ (thickness) × 0.25″ (outer diameter) stainless steel frits (IDEX Health and Science, Oak Harbor, WA, USA). 19. One-piece PEEK Direct-Connect™ column coupler for Waters fittings (Grace Davison Discovery Science, Deerfield, IL, USA). 20. LC-20 AD HPLC and SIL-5000 auto-injector (Shimadzu Scientific Instruments, Columbia, MD, USA). 21. Quattro micro™ API mass spectrometer (Waters). 22. 1 mM perfluoroheptanoic acid. 23. Acetonitrile. 2.5. Leaf Fatty Acid Assay
1. 13 mm (outer diameter) × 100 mm (length) 8-mL borosilicate glass culture tubes with screw-cap finish (VWR International, West Chester, PA, USA), barcoded. 2. 13 mm (outer diameter) × 14 mm (length) phenolic screw caps with PTFE/rubber liner (Kimble Chase, Vineland, NJ, USA). 3. 0.01″-thick 12-mm PTFE septa (SUN SRI). 4. Methanol and a 10-mL 33-mm-neck size bottle-top dispenser (VWR International). 5. 1 mg/mL pentadecanoic acid (Sigma–Aldrich, St. Louis, MO, USA) in toluene. 6. 10 mg/mL butylated hydroxytoluene (Sigma–Aldrich) in methanol. 7. Methylation reagent: 1 N methanolic HCl, 5 mg/mL pentadecanoic acid (from 1 mg/mL stock), and 10 mg/mL butylated hydroxytoluene (from 10 mg/mL stock) in methanol. Mix
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thoroughly and use the same day, dispensing with a 5-mL glass dispenser (Barnstead Internationals, Dubuque, IA, USA). 8. 0.9% (w/v) NaCl and a 20-mL dispenser (Thermo Fisher Scientific). 9. Heptane and a 1-mL glass dispenser (Barnstead Internationals). 10. 300-mL extra long (90 mm) pipette tips (Denville Scientific). 11. 12 mm (outer diameter) × 32 mm (length) 2-mL wide-opening screw-top vials (Agilent Technologies), barcoded. 12. Screw caps with PTFE/rubber septa for above vials (Agilent Technologies). 13. 150-mL inserts for above vials (Agilent Technologies). 14. Agilent 7683B series injector and 6890 series GC system with a flame ionization detector (Agilent Technologies). 15. J & W DB-23 0.25-mm (internal diameter), 30-m (length), 0.25-mM (film size) capillary column (Agilent Technologies). 2.6. Chloroplast Morphology
1. 32-tube CRYO-SAFE™ maxi coolers and lids (Bel-Art Products), 4°C. 2. 3.5% (w/v) glutaraldehyde: diluted from 50% grade I (Sigma– Aldrich). The diluted solution is stable for up to 1 week at 4°C. 3. Matrix 1,250-mL 8-channel electronic pipette with expandable spacing (Thermo Fisher Scientific). 4. GyroMini™ nutating mixer (Labnet International, Edison, NJ, USA). 5. 0.1 M EDTA, pH 9.0. 6. Microscope with polarization contrast optics, e.g., Leica DMI 3000B inverted microscope with a 40 × 0.75 HCX PL Fluotar objective lens and a DFC320 digital camera (Leica Microsystems, Bannockburn, IL, USA). 7. 1.2-mm-thick 3″ (length) × 1″ (width) plain micro slides (VWR International). 8. 18 mm × 18 mm No. 1.5 micro coverslips (VWR International).
2.7. DNA Archiving and Genotyping
1. FTA® plantsaver cards (Whatman, Florham Park, NJ, USA), barcoded. These cards are intended for archiving DNA from leaf tissue for PCR analysis. 2. Porcelain pestle, any size (CoorsTek, Golden, CO, USA). 3. 1.2-mm Harris UNI-CORE™ punch (Whatman). 4. 2.5″ × 3″ Harris cutting mat (Whatman). 5. 96-Well full skirt PCR plates (USA Scientific, Ocala, FL, USA), barcoded. 6. FTA® purification reagent (Whatman).
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7. TE0.1 buffer: 10 mM Tris and 0.1 mM EDTA, pH 8.0. 8. REDTaq® ReadyMix™ PCR mix (Sigma–Aldrich). 9. Gene-specific primers (LP and RP) are designed using SALK T-DNA Primer Design Tool, http://signal.salk.edu/ tdnaprimers.2.html with the Ext5 value changed to 200. They are manufactured in 96-well plates at Integrated DNA Technologies (Coralville, IA, USA), 10 nmol per well, and lyophilized. Primers are in alphanumeric order with all LP primers in one plate and all RP primers in another plate. 2.8. Seed Morphology
1. 96-Well 360-mL flat-bottom polystyrene microplates and lids (Corning Inc.), barcoded. 2. Microscope paper stage guide/image background (see Note 3). 3. Stereomicroscope with polarizing lens, e.g., Leica MZ12.5 high-performance stereomicroscope (Leica Microsystems) with 1× objective lens and SPOT Insight Color 3.2.0 digital camera (Diagnostic Instruments, Sterling Heights, MI, USA).
2.9. Seed Amino Acid Assay
1. 1-mL 96-well deep-well microplate (VWR) and lid containing one 3-mm stainless steel bead (CCR Products). 2. All other materials are the same as in Subheading 2.4, items 3–23.
2.10. Seed C and N Assay
1. Nalgene 280-mm (outer diameter) polycarbonate desiccator and 230-mm (outer diameter) ceramic-metal composite plate (Thermo Fisher Scientific). 2. Indicating drierite (W. A. Hammond Drierite, Xenia, OH, USA). 3. XP26 DeltaRange® microbalance (Mettler Toledo, Columbus, OH, USA). 4. 5 mm (diameter) × 9 mm (length) tin capsules (CE Elantech, Lakewood, NJ, USA). 5. 96-Well 360-mL flat-bottom polystyrene microplates and lids (Corning Inc.), barcoded. 6. Carlo Erba® NC2100 elemental analyzer (Thermo Fisher Scientific) with zero blank autosampler (Costech Analytical Technologies, Valencia, CA, USA).
2.11. Seed Starch Assay
1. 96-Well 360-mL flat-bottom polystyrene microplates and lids (Corning Inc.), barcoded. 2. Aluminium sealing films (Excel Scientific). 3. 80% (v/v) ethanol. Store at RT.
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4. IKI solution: 0.67% (w/v) iodine and 3.33% (w/v) KI. The reagent is light sensitive; wrap the bottle with foil and store at 4°C. 2.12. Chlorophyll Fluorescence Assay
1. 5.25″ (length) × 5.25″ (width) 12-plug com-pack beddingplant containers (Hummert International) for 22″ (length) × 11″(width) × 2.5″ (height) flats. The containers or subflats are barcoded. 2. All other growth materials are the same as in Subheading 2.1, items 2–7. 3. MAXI version of IMAGING-PAM M-series chlorophyll fluorescence system (Heinz Walz GmbH, Effeltrich, Germany) and AVT Dolphin camera (Allied Vision Technologies GmbH, Stadtroda, Germany) in a dark room.
2.13. Systematic Data Analysis
Some data analysis approaches described in this chapter are performed with JMP 8.0 statistical software (SAS Institute).
3. Methods 3.1. Plant Care and Harvesting Schedule
To accommodate high-throughput sample processing procedures that involve the use of 96-well plates, plants are grown in two sets of Homozygous seeds from ABRC Grow 2 plants under 16/8 Plant morph Genotyping Bulk seeds, sibling 1
Seed assays Seed morph Seed AA Seed C/N Seed starch
Bulk seeds, sibling 2
Grow 1 plant under 12/12 for leaf assays Plant morph Morning starch Leaf AA Fatty acids Afternoon starch Chloroplast morph DNA archiving
Grow 1 plant under 12/12 for leaf assays Plant morph Morning starch Leaf AA Fatty acids Afternoon starch Chloroplast morph DNA archiving
Putative mutants Database Queries
Fig. 1. Mutant analysis pipeline.
Seed assays Seed morph Seed AA Seed C/N Seed starch
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three 32-plug flats (1, 28). Each mutant line is therefore planted in duplicate. Seeds harvested from plants grown under the 16/8-h photoperiod are used for seed assays and are sown for growth and leaf assays in a 12/12-h photoperiod (Fig. 1). The irradiance is 100 mmol photons/m2/s with a mix of cool white fluorescent and incandescent bulbs; the temperature is 21°C. Seeds are sown on Redi-earth plug and seedling mix topped with a thin layer of vermiculite. The sown seeds are stratified at 4°C in the dark for 3–4 days. To stagger the tissue harvest dates and thus facilitate rapid collection of each set of leaf tissue, sets of 96 pots (3 flats) are moved to the same growth chamber on successive days (days 3 and 4 after sowing). After 7 days in the growth chamber, the humidity domes are taken off and seedlings are thinned to one plant per pot. Plants are watered by filling the tray to excess with deionized water on Mondays and Wednesdays and Arabidopsis nutrient solution on Fridays. Two hours after watering, excess water or nutrient solution is poured off. The six flats are rotated and the orientation of each flat switched each time they are watered or fertilized. This watering procedure reduces plant-to-plant phenotypic variability by ensuring that the soil mix for each plant reaches a similar level of saturation. Plants growing under a 16/8-h photoperiod are photographed and scored for whole plant morphology after 24 days in the growth chamber. On the following day, one leaf per plant is harvested for DNA archiving and genotyping. Aracon bases and tubes are placed on the plants after the plants start flower stalk elongation. Watering is stopped after ~10 weeks of growth, and intact plants are allowed to dry for 3 weeks more in the growth chamber, maintaining the flat rotation protocol, before seeds are harvested and stored. Seed boxes are stored in auto-desiccator cabinets at ~30% relative humidity in the cold room (8°C). Plants growing under a 12/12-h photoperiod are photographed and scored for whole plant morphology after 29 days in the growth chamber. On the following day, leaf tissues are harvested for morning starch (leaf #8, 8:00–8:40 A.M.), leaf amino acid (leaf #7, 9:00– 10:15 A.M.), leaf fatty acid (leaves #5 and #6, 12:30–2:30 P.M.) and afternoon starch (leaf #9, 4:00–4:40 P.M.). It should be noted that leaf numbers are counted from the newest expanding leaf (leaf #1, Fig. 2). To minimize light effects on metabolite concentration, plants are placed under a tabletop fluorescent light fixture (~100 mmol photons/m2/s at leaf surface) during harvest for leaf starch, amino acid, and fatty acid assays. Leaf tissues for chloroplast morphology are harvested 4 days later, and leaves for DNA archiving are harvested ³5 days later (see Note 4). To maximize accuracy in data tracking, every flat, pot, plate, tube, vial, DNA archiving card, seed stock, and storage box is barcoded and data associations are tracked in a relational database (1, 28). 3.2. Whole Plant Morphology
While each plant in a flat experiences approximately the same environment, it is expected that there will be flat-to-flat or within-flat
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Fig. 2. Leaf numbers 1–14 of a typical 29-day-old Arabidopsis plant grown at a 12/12-h photoperiod. Leaves #5–6 are for fatty acid analysis; leaf #7 is for amino acid profiling; and leaves #8–9 are for morning and afternoon starch staining, respectively. Four days later, leaf #4 grows to a mature leaf and is harvested for chloroplast morphology. Any healthy, non-senescent leaf can be used for DNA archiving and genotyping.
variation in the microenvironment. For whole plant morphology, phenotypes are scored compared to the “average” of flat-mates. The whole flat is assessed first to determine the flat norm. Morphology traits are scored using the following controlled vocabulary (1, 28): 1. Rosette size (diameter): Normal (4.0–6.0 cm), 1 (2.5–3.9 cm), 2 (1.25–2.4 cm), or 3 (≤1.24 cm). Hold the measuring guide (Fig. 3) over the plant, with the center directly over the center of the rosette. Observe all leaves; any leaf extending beyond the line determines the score entered. 2. Inflorescence: visible or not visible. 3. Leaf color: lighter, normal, or darker. 4. Leaf color variation: evenly colored or color variation. If the leaves appear to have color variation, determine the color (light green, dark green, necrotic, purple, yellow, white, or normal) and location (apex, margin, vein, or mottled) of the variation. 5. Leaf number: less, normal, or more. Count the leaves, determine a normal range for that flat, and then compare the number of leaves for each individual plant to the flat norm (see Note 5). 6. Leaf shape: normal or abnormal. Examples for leaves of abnormal shape include curled, flat, narrow, rolled, round, serrated,
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Fig. 3. Overlay of rosette size measuring guide and a typical 29-day-old Arabidopsis plant grown at a 12/12-h photoperiod.
succulent, wilted, or wrinkled leaves, or leaves with a pointed apex or short petiole (see Note 6). 7. Mature leaf size: smaller, normal, or larger. 8. Trichomes: present or absent. 3.3. Leaf Starch Assay
1. Harvest one leaf disk from leaf #8 (morning starch) with a cork borer immediately after growth chamber lights come on. Harvest leaf #9 (afternoon starch) after 8 h of light. In each case, place disks into a microplate chilled with CoolSafe system. 2. After harvesting, add 200 mL of killing solution to each well with an 8-channel pipette, push leaf disks that are not submerged into the liquid with toothpicks, and seal the wells with a piece of aluminium sealing film. 3. Incubate the plate in an oven at 80°C for 10 min. After incubation, cool at RT for 2 min. 4. Replace the killing solution with 200 mL of 80% ethanol, incubate at 80°C for 5 min, and remove 80% ethanol.
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5. Stain the leaf disks in 200 mL of Lugol’s IKI solution at RT for 3 min (1, 11), and remove IKI solution. 6. Rinse the leaf disks with 200 mL of distilled water. 7. Add another 200 mL of distilled water and incubate at 80°C for 15 min (see Note 7). 8. After incubation, cool at RT for 2 min. Make sure that the leaf disks lay flat and then photograph the plate on a light box. Score individual leaf disk as starch excess or normal for morning starch (see Note 8). For afternoon starch, score individual leaf disk as starchless or normal. 3.4. Leaf Amino Acid Assay
1. Harvest the blade only (no petiole) of leaf #7 from each plant and record the sample fresh weight. Cut the leaf into 2–3 pieces, put into a 2-mL microfuge tube containing a steel bead, and place the tube immediately in a 96-tube rack chilled by surrounding below and around the sides with dry ice. Leaf samples can be stored at −80°C until processing. 2. Quickly and keeping samples frozen, place a thin layer of plastic foam inside the cover for 96-tube rack, cover the samples, and secure the assembly with heavy-duty tape. Shake the samples for 2 min on the paint shaker while still frozen (see Note 9). Keep the samples on dry ice afterward. 3. Spin down the frozen tissue powders at ~5,000 × g for 10 s at −9°C and keep on ice afterward. 4. Make 45 mL extraction reagent containing 1 mM Phe-d8 and 10 mM DTT with distilled water and concentrated stocks. 5. Add 400 mL of extraction reagent to each sample tube and shake manually for 10 s. 6. Briefly spin down samples and incubate in a water bath at 90°C for 5 min. 7. Briefly spin down samples again and transfer them to MicroTubes™ strips to separate sample solutions from the beads. 8. Centrifuge at 3,220 × g for 5 min at 4°C with a swinging bucket centrifuge. 9. Pre-wet filter plate: add 100 mL of distilled water to each well on a filter plate and centrifuge the filter plate with the collection plate at 2,000 × g for 5 min at 4°C. 10. Place the pre-wetted filter plate on a new collection plate, transfer 300 mL of supernatant from step 9 to the pre-wetted filter plate, and centrifuge at 2,000 × g for 5 min at 4°C. 11. Add 10 mL of 9 mM Val-d8 to each well of two 96-well plates (see Note 10). 12. Transfer 90 mL of filtrate from step 11 to each of the 96-well plates and seal the plates with aluminium sealing film.
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13. Vortex and spin down sample solutions. Sample plates can be stored at −80°C until assaying. 14. On the day of HPLC-MS/MS analysis, thaw one of the sample plates, vortex, centrifuge, and replace the aluminium film with a silicone sealing mat. 15. Thaw one set of amino acid standards and transfer them into screw-cap vials with inserts. 16. Place standards and the sample plate in the sample storage unit of Shimadzu SIL-5000 auto-injector and condition the column for 30 min with 98% 1 mM perfluoroheptanoic acid (A) and 2% acetonitrile (B). 17. Analyze standards and samples on the Shimadzu/Waters HPLC-MS/MS system (1, 14), at the following conditions: HPLC: Injection volume: 10 mL. Mobile phases: (A) 1 mM perfluoroheptanoic acid in water and (B) acetonitrile. Total HPLC run time: 6 min. Flow rate: 0.3 mL/min. Gradient: 0–0.09 min, 98% A, 2% B; 0.1–2.29 min, 80% A, 20% B; 2.3–4.09 min, 60% A, 40% B; 4.1–6 min, 98% A, 2% B. Mass Spectrometer: Electrospray ionization: positive ion mode. Capillary voltage: 3.0 kV. Cone voltage: 26 V. Source temperature: 110°C. Desolvation temperature: 350°C. Note that for improved accuracy for amino acids other than Phe and Val, isotopically-labeled amino acid standard may be added (14). 3.5. Leaf Fatty Acid Assay
1. Place a PTFE liner in each phenolic screw cap. To wash sample tubes, dispense 1 mL of methanol in each tube, cap, vortex, discard used methanol, and then air-dry tubes and caps in a fume hood. 2. On the day of harvesting, dispense 1 mL of methylation reagent to washed tubes and cap tightly. 3. Preheat three 2-block heaters at 80°C. 4. Harvest the blade of leaf #5 or #6 of each plant, record the sample fresh weight, put into a glass tube containing methylation reagent, cap tightly, and place the tube into a preheated heat blocker. Be sure that the leaf is completely immersed.
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5. Incubate all samples in the heat blockers at 80°C for 30 min, remove from the heater, and allow to cool to RT. 6. In a fume hood, add 1.0 mL 0.9% NaCl and 0.15 mL heptanes and vortex each tube well. 7. Centrifuge the sample tubes at 200 × g for 10 min at RT with a swinging bucket centrifuge. 8. Transfer the upper heptane layer (~75 mL) to GC vials with inserts, using a pipette and extra long tips. Samples can be stored at 4°C or −20°C if you do not plan to assay them on the same day. 9. Load the GC vials on the autosampler of the Agilent GC system and analyze the samples with GC-FID (1, 17), at the following conditions: Injector: Volume: 1 mL. Mode: split ratio 30:1. Temperature: 270°C. Oven: Initial temperature: 140°C. Temperature ramp: 10°C/min. Final temperature: 260°C (hold for 3 min). Detector: Temperature: 270°C. 3.6. Chloroplast Morphology
1. Harvest a newly expanded leaf, including the petiole. Excise the base (petiole) and the apex of the blade into 2 mL green and blue microfuge tubes, respectively. Cap and store the tubes in prechilled coolers until they can be processed, but not longer than 1 h. 2. Transfer the sample tubes to 96-tube racks. Working in a fume hood, add 900 mL of 3.5% glutaraldehyde to each tube. 3. Incubate samples at RT for 2 h on nutating mixers. 4. Working in a fume hood, remove the glutaraldehyde from each tube. Be careful not to aspirate the leaf sample. 5. Add 1 mL of 0.1 M EDTA to each tube. Be sure that the leaf sample is completely immersed. 6. Incubate the samples in an oven at 55°C for 2 h. 7. Cool at RT for 10 min. Store fixed samples at 4°C. Fixed samples must be used within 3 days. 8. On the day of slide preparation and sample scoring, add a small drop of distilled water to a microslide. Remove a fixed tissue sample from the tube and place it in the water drop. Cover the sample with a coverslip.
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Fig. 4. Examples of images corresponding to controlled terms used in chloroplast morphology. (a) Leaf base; normal chloroplast number, shape, and size. (b) Leaf base; less, amorphous, bigger chloroplasts. (c) Leaf base; less, dumbbell and amorphous, bigger chloroplasts. (d) Leaf base; less, elongated and amorphous, bigger chloroplasts. (e) Leaf base; more, wrinkled chloroplasts. (f) Leaf apex; normal chloroplast number, shape, and size. (g) Leaf apex; less, amorphous, bigger chloroplasts. (h) Leaf apex; less, dumbbell and amorphous, bigger chloroplasts. (i) Leaf apex; less, amorphous, bigger chloroplasts. (j) Leaf apex; more, wrinkled chloroplasts.
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9. Tap the top of the coverslip above the specimen gently with the eraser-end of a clean, unused pencil to crush the specimen to a fine green haze. Blot excess water. 10. Invert the sample and view with a 40× objective on a Leica DMI 3000B microscope (1, 18). Scan the entire slide specimen and examine the chloroplasts within the cells. Locate a representative area of several isolated cells to photograph. If abnormal chloroplasts are seen, capture additional images (see Fig. 4 for examples of images). 11. Chloroplast morphology traits scored using controlled vocabulary include the following: (a) Chloroplast number: less, normal, or more. Estimate the number of chloroplasts per cell. Cells at the base or apex of a mature wild-type leaf blade contain 80–100 chloroplasts (see Note 11). (b) Chloroplast shape: normal or abnormal. Normal chloroplasts are small, round, and grass green. Abnormal chloroplasts may be amorphous, dumbbell shaped, elongated, heterogeneously shaped (more than one type), or wrinkled. (c) Chloroplast size: smaller, normal, or larger. 3.7. DNA Archiving and Genotyping
To save freezer space and simplify sample preparation procedure, DNA samples from each plant are archived on FTA cards. To process the large number of archived DNA samples, genotyping can be done with high-throughput “first-pass” reactions followed by “second-pass” reactions (8). In “first-pass” screening, both mutant line sibling samples are pooled together to reduce the number of PCRs required. This approach only distinguishes homozygotes from any non-homozygotes. “Second-pass” assays distinguish between heterozygotes and wild-type alleles for the samples that do not appear to be homozygous. 1. Excise a healthy leaf from each plant and place the leaf on the matrix of a FTA card. Replace the cover sheet and crush each sample completely using a porcelain pestle by rubbing and tapping. 2. Allow the FTA card to air-dry at RT for at least 1 h before punching DNA samples. The archived samples can be stored at RT indefinitely in desiccated boxes. “First-pass” genotyping: 3. Use a 1.2-mm punch and a cutting mat to take disks from FTA cards (see Note 12). Two sibling DNA samples are placed into the same well of a 96-well PCR plate; they will be genotyped with two gene-specific primers LP and RP. One wild-type DNA sample is placed in the adjacent well; it will be genotyped with the same two gene-specific primers as a positive control.
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4. Wash all disks with 100 mL of FTA purification reagent two times, with 5-min incubation at RT for each wash. 5. Wash all disks with 100 mL of TE0.1 twice, with 5-min incubation at RT for each wash. Be sure to remove the final TE0.1 wash completely. 6. Allow the disks to air-dry completely for 1 h at RT or for 20 min at 56°C. 7. The dried samples are genotyped using REDTaq® ReadyMix™ PCR mix and gene-specific primers at a final concentration of 0.1 nM. The following thermal profile is used: 95°C for 10 min; 35 cycles of 95°C for 2 min, 56°C for 1 min, 72°C for 2 min; and 72°C for 10 min; hold at 4°C. 8. PCR result is analyzed by agarose gels. If the two sibling punches do not yield a PCR product but the wild-type sample does, then both siblings are considered to be homozygous. “Second-pass” genotyping: 9. Take disks from FTA cards as in step 1. Siblings are analyzed in separate wells. Each sibling is amplified using two separate reactions: one with the gene-specific primers LP and RP, and one using RP and a T-DNA insert-specific primer for SALK T-DNA mutant lines (LBa1, 5¢-TGGTTCACGTAGTGGGCC ATCG-3¢). One wild-type DNA sample is also assayed with the same LP and RP as a positive control. 10. DNA samples are washed and PCRs done as in steps 4–8. 3.8. Seed Morphology
1. Tap seeds from an individual seed stock into one well of a microplate to form a single layer. View all the seed stocks in a box at one time by quickly scanning through all the wells under a stereomicroscope. This is to establish the norm for the seed box, which contains seeds harvested from 96 plants grown at the same time and in the same growth chamber. The phenotype of a seed stock is always compared to other seed stocks in the same seed box. 2. To assess and photograph an individual sample, tap 40–50 seeds of a seed stock onto the sample placement guide, remove chaff, and distribute seeds evenly with blunt-tip forceps. 3. Examine the seeds using both the microscope and computer monitor, and photograph them (1). Score the seeds based on the characteristics of the majority of the sample’s seeds, i.e., ignore phenotypes represented by less than 10 seeds per ~50 seed sample. Morphology traits scored using controlled vocabulary include the following:
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(a) Seed color of the population: homogeneous or heterogeneous. If ~25% of the seeds have a different color, the entire population is scored as heterogeneous. (b) Seed coat color: lighter, normal, or darker. (c) Seed coat color variation: evenly colored or color variation. (d) Seed coat surface: normal or abnormal. Abnormal seed coat surfaces may be dull or shiny. (e) Seed morphology of the population: homogeneous or heterogeneous. If ~25% of the seeds have a different shape or size, the entire population is scored as heterogeneous. (f) Seed shape: normal or abnormal. Abnormal shaped seeds may be aborted, elongated, round, or wrinkled. (g) Seed size: smaller, normal, or larger. 3.9. Seed Amino Acid Assay
1. Record the dry weight of ~7.5 mg of seeds after removing non-seed material and transfer via a funnel into a deep-well microplate with a steel bead in each well. Aliquoted seeds can be stored in a humidity-controlled container in the cold room until processing. 2. Make 45 mL extraction reagent containing 1 mM Phe-d8 and 10 mM DTT. 3. Add 400 mL extraction reagent to each well with an 8-channel pipette. Cover the deep-well plate with the lid and then press each dome in the lid down firmly to make sure none of the wells leak. 4. Shake the plate on the paint shaker for 5 min and then keep on ice. 5. Briefly centrifuge the samples and incubate the plate in a water bath at 90°C for 5 min (see Note 13). 6. Centrifuge at 3,220 × g for 10 min at 4°C with a swinging bucket centrifuge. 7. Carefully transfer ~350 mL supernatant to MicroTubes™ strips (see Note 14). 8. Centrifuge at 3,220 × g for 10 min at 4°C. 9. Pre-wet the filter plate as described in step 9 in Subheading 3.4. 10. Carefully transfer 300 mL of supernatant from step 9 to prewetted filter plate and centrifuge at 2,000 × g for 50 min at 4°C (see Note 15). 11. Aliquot 10 mL of 9 mM Val-d8 and 90 mL of filtrate from step 10 to each well of two 96-well plates, seal, vortex, centrifuge, and store as described in steps 11–13 in Subheading 3.4.
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12. Analyze seed amino acid samples with HPLC-MS/MS as described in steps 14–17 for leaf amino acid analysis (see Subheading 3.4). 3.10. Seed C and N Assay
1. Aliquot ~12 mg of seeds (remove chaff) from each seed stock in the same seed box to decapped 2-mL microfuge tubes. 2. Place the rack of seed samples in a dessicator with drierite and dry the samples under vacuum for 3 days. 3. Weigh ~10 mg of dried seeds on a microbalance (accurate to 0.01 mg), transfer all seeds to a tin capsule, and fold the capsule into a ball (see Note 16 and http://www.biology.duke.edu/ jackson/devil/sampleprep.html). Make sure that weighed seeds do not spill and the tin ball does not leak. It is very important that seeds in the same sample set are weighed at equal dryness. To prevent dried seed samples from imbibing moisture from the air once they are removed from the dessicator, samples are prepared in small batches, and waiting samples are kept dry under vacuum. 4. The seed samples are analyzed by the Duke Environmental Stable Isotope Laboratory (http://www.biology.duke.edu/ jackson/devil/). The C and N contents are quantified by combusting the seeds at 1,200°C (1) in an elemental analyzer.
3.11. Seed Starch Assay
1. Tap a monolayer of seeds from each seed stock into one well of a microplate. It should be noted that the set of seed samples that is used for establishing the norm for assessing seed morphology (see Subheading 3.8) can be reused for this assay. 2. Add 200 mL of 80% ethanol with an 8-channel pipette and seal the wells with a piece of aluminium sealing film. 3. Incubate the plate in an oven at 80°C for 20 min. 4. Cool the plate at RT for 2 min. 5. Attach a Pasteur pipette to a water aspirator (with a liquid trap in between), turn on the aspirator, and aspirate the 80% ethanol from each well (see Note 17). 6. Stain the seeds in 200 mL of IKI solution at RT for 3 min (1, 23) and then remove IKI solution by aspiration. 7. To rinse the seeds, add 200 mL of distilled water with an 8-channel pipette and remove by aspiration. Repeat this step once. 8. Add another 200 mL of distilled water, photograph the plate with a white background, and immediately score individual seed samples as starch excess or normal.
3.12. Chlorophyll Fluorescence Assay
Plants used for this assay are grown for 3 weeks in 12-plug beddingplant containers (subflats) so that each subflat can be analyzed with
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the MAXI version of IMAGING-PAM M-series chlorophyll fluorescence system (1, 27). The 12-h light growth conditions are the same as those for leaf assays (see Subheading 3.1). 1. On the morning of the assay, add a thin layer of vermiculite to the soil around the plant to mask any algal growth, which would otherwise cause high background fluorescence. Blow away excess vermiculite on plants. 2. Dark-adapt the plants for 20 min. 3. Determine maximum photochemical efficiency of PSII (Fv/Fm) before high light and non-photochemical quenching (NPQ) using the chlorophyll fluorescence system. The following light program is used: 1 min of 3 mmol photons/m2/s (measuring light), a flash of 2,800 mmol photons/m2/s (saturating light), 3 min of 533 mmol photons/m2/s (actinic light), and a second flash of saturating light. 4. Treat plants with high light (1,500–1,700 mmol photons/m2/s) for 3 h. 5. Dark-adapt the plants for 20 min and determine Fv/Fm after high light. 6. Transfer recovery plants to the original growth chamber for 2 days. 7. Add another thin layer of vermiculite to the soil, dark-adapt the plants, and determine Fv/Fm after recovery. 8. View and compare images (Fv/Fm or NPQ) for all subflats within a flat, and enter a cutoff value for Fv/Fm (before high light, after high light, or after recovery) or NPQ, which must then be used for all subflats within that flat (1). Plants with one or more values below the corresponding cutoff value appear red and are considered putative hits. 3.13. Systematic Data Analysis
Putative mutants from morphological and qualitative assays can be identified by querying the database using controlled vocabulary. For quantitative assays, raw data are normalized and converted before mutant identification (1, 28). Procedures for quantitative data normalization are described below: 1. Compute mol% and nmol/g fresh weight of leaf fatty acids, mol% and nmol/g fresh weight of leaf and seed amino acids, and seed C/N ratio for each sample. 2. Calculate median for each flat and median absolute deviation (MAD) for each plant in the flat (see Note 18). MAD is given by the equation MAD = 1.482 × medi (|xi – median|), where xi is the value of each individual measurement and medi is the median of the n absolute values of the deviations about the median (1).
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3. Compute z-score by the equation zi = (xi − median)/MAD. Putative mutants from quantitative assays can be identified by specifying z-score cutoff values for any metabolites analyzed. Generally, a z-score cutoff value of at least 3 or -2 for two siblings gives a reasonable number of putative mutants, although the variance for different metabolites differs. Mutants with more subtle phenotypes may be sought by using reduced stringency criteria with the potential result of more false-positive results to evaluate by rescreening. To look for phenotypic syndromes of the mutants and connections across different physiological processes, data from morphological and qualitative assays can be systematically converted to numeric codes and merged with quantitative z-score data (1, 28). The broad phenotypic patterns can be visualized by clustering methods, such as hierarchical clustering and k-means clustering, with JMP 8.0 software (1). The degree of correlation between pairs of traits or genes can be evaluated by correlation analysis, such as parametric Pearson correlation method and nonparametric Spearman’s correlation method in JMP 8.0 (1).
4. Notes 1. Arabidopsis nutrient solution is made from stock solutions. Make up all stock solutions in distilled water. The final watering solution may be made up in deionized water. To prevent the formation of insoluble precipitates, do not pipette concentrated solutions all together before adding water. Instead, fill a large carboy 1/3–1/2 full, pipette the nutrient stocks directly into the carboy, slosh around to mix, fill to the line, and slosh around to mix again. Some components in Arabidopsis micronutrient solution are toxic as powders or concentrated solutions. 2. Phe-d8 is used to normalize extraction and Val-d8 is used to monitor loading accuracy. Their concentrations in the standards are the same as those in leaf or seed samples – 0.9 mM. To simplify preparation of standards and to calculate amino acid concentrations in the samples, the final concentrations for the 20 protein amino acids, GABA, anthranilate, Homo-Ser, Hyp, and S-methyl Met in the standards are 0.9× of the stated concentration. 3. This is a 10-cm diameter circle of white paper. An arrow and a black closed circle with a 1.2-cm diameter are printed near the center of the paper. Seeds are placed on the black circle and the arrow helps you center the seeds in the field of view in the microscope. 4. Leaf tissues for chloroplast morphology are harvested in the morning (8:00–10:00 A.M.) when starch levels are low. The
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presence of large starch granules makes the chloroplast appear wrinkled, especially for starch-excess mutants. 5. Sometimes a plant with smaller leaves may misleadingly appear as though it has fewer leaves. 6. Remember that smaller plants have leaves that are naturally not as elongated as larger plants, but this does not necessarily mean that the shape should be scored as “round.” Become familiar with the range of normal phenotypes for a range of plant sizes. 7. This incubation step reduces background staining. 8. Iodine staining of starch is light and time sensitive. It is important to photograph and score starch phenotype immediately. 9. The paint shaker is custom modified to grind samples: the inner sides of the clamps are filled with 1 inch thick foam sheet. To get better grinding results, secure sample plates near the edge of the clamps so the agitation arc is larger. For frozen leaf samples, after grinding for 1 min, flip the sample plate horizontally by 180° and grind for another minute. Grinding can also be done with a GenoGrinder or a similar system. 10. Although samples are split into two identical plates, only one plate is typically used. Occasionally problems happen during HPLC-MS/MS and samples from the extra plate can be used for a repeat instrument run. 11. Cells at the leaf base are large and rectangular and the chloroplasts spread through several distinct planes. Therefore, focus through several view planes to estimate the number of chloroplasts. 12. The static charge on plastic labware and gloves can repel the disks. To reduce the static electricity, PCR plates can be placed on damp paper towels. 13. During the incubation of seed amino acid samples at 90°C, the edges of the mat may curl slightly. Make sure that the sample plate does not float in the water bath and that the sealing mat is above water so that bath water does not get into the sample wells. 14. Seed residues may clog pipette tips. Avoid letting pipette tips touch seed pellets at the bottom of the wells during pipetting. 15. It takes much longer to filter seed plates than leaf plates. 16. The work area, microbalance, and utensils need to be clean. Wipe utensils thoroughly with methanol and allow them to dry briefly. 17. Aspiration of liquid off Arabidopsis seeds is tricky. It is easier to use a Pasteur pipette connected with a water aspirator than an 8-channel pipette. 18. Compared to mean and standard deviation, median and MAD are not as sensitive to extreme values. Therefore, the z-scores
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for identifying putative mutants are calculated from median and MAD of each flat.
Acknowledgments The authors thank Kathleen M. Imre for configuring the chlorophyll fluorescence assay and Imad Ajjawi for configuring the fatty acid assay. We are grateful to the many project members who contributed to the establishment and refinement of these protocols, including the many undergraduate students involved in the project. This work was supported by the US National Science Foundation 2010 Project Grants MCB-0519740 and DBI-0619489 for LC-MS equipment. References 1. Lu, Y., Savage, L. J., Ajjawi, I., Imre, K. M., Yoder, D. W., Benning, C., DellaPenna, D., Ohlrogge, J., Osteryoung, K. W., Weber, A. P. M., Wilkerson, C. G., and Last, R. L. (2008) New connections across pathways and cellular processes: industrialized mutant screening reveals novel associations between diverse phenotypes in Arabidopsis. Plant Physiol. 146, 1482–1500. 2. Sussman, M. R., Amasino, R. M., Young, J. C., Krysan, P. J., and Austin-Phillips, S. (2000) The Arabidopsis knockout facility at the University of Wisconsin-Madison. Plant Physiol. 124, 1465–1467. 3. Sessions, A., Burke, E., Presting, G., Aux, G., McElver, J., Patton, D., Dietrich, B., Ho, P., Bacwaden, J., Ko, C., Clarke, J. D., Cotton, D., Bullis, D., Snell, J., Miguel, T., Hutchison, D., Kimmerly, B., Mitzel, T., Katagiri, F., Glazebrook, J., Law, M., and Goff, S. A. (2002) A high-throughput Arabidopsis reverse genetics system. Plant Cell 14, 2985–2994. 4. Alonso, J. M., Stepanova, A. N., Leisse, T. J., Kim, C. J., Chen, H. M., Shinn, P., Stevenson, D. K., Zimmerman, J., Barajas, P., Cheuk, R., Gadrinab, C., Heller, C., Jeske, A., Koesema, E., Meyers, C. C., Parker, H., Prednis, L., Ansari, Y., Choy, N., Deen, H., Geralt, M., Hazari, N., Hom, E., Karnes, M., Mulholland, C., Ndubaku, R., Schmidt, I., Guzman, P., Aguilar-Henonin, L., Schmid, M., Weigel, D., Carter, D. E., Marchand, T., Risseeuw, E., Brogden, D., Zeko, A., Crosby, W. L., Berry, C. C., and Ecker, J. R. (2003) Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301, 653–657. 5. Kuromori, T., Hirayama, T., Kiyosue, Y., Takabe, H., Mizukado, S., Sakurai, T., Akiyama, K., Kamiya, A., Ito, T., and Shinozaki,
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K. (2004) A collection of 11,800 single-copy Ds transposon insertion lines in Arabidopsis. Plant J. 37, 897–905. O’Malley, R. C., and Ecker, J. R. (2010) Linking genotype to phenotype using the Arabidopsis unimutant collection. Plant J. 61, 928–940. Kuromori, T., Wada, T., Kamiya, A., Yuguchi, M., Yokouchi, T., Imura, Y., Takabe, H., Sakurai, T., Akiyama, K., Hirayama, T., Okada, K., and Shinozaki, K. (2006) A trial of phenome analysis using 4000 Ds-insertional mutants in gene-coding regions of Arabidopsis. Plant J. 47, 640–651. Ajjawi, I., Lu, Y., Savage, L. J., Bell, S. M., and Last, R. L. (2010) Large-scale reverse genetics in Arabidopsis: case studies from the Chloroplast 2010 Project. Plant Physiol. 152, 529–540. Myouga, F., Akiyama, K., Motohashi, R., Kuromori, T., Ito, T., Iizumi, H., Ryusui, R., Sakurai, T., and Shinozaki, K. (2010) The Chloroplast Function Database: a large-scale collection of Arabidopsis Ds/Spm- or T-DNAtagged homozygous lines for nuclear-encoded chloroplast proteins, and their systematic phenotype analysis. Plant J. 61, 529–542. Lu, Y., and Sharkey, T. D. (2006) The importance of maltose in transitory starch breakdown. Plant Cell Environ. 29, 353–366. Yu, T. S., Kofler, H., Hausler, R. E., Hille, D., Flügge, U. I., Zeeman, S. C., Smith, A. M., Kossmann, J., Lloyd, J., Ritte, G., Steup, M., Lue, W. L., Chen, J., and Weber, A. (2001) The Arabidopsis sex1 mutant is defective in the R1 protein, a general regulator of starch degradation in plants, and not in the chloroplast hexose transporter. Plant Cell 13, 1907–1918.
9 Chloroplast Phenomics: Systematic Phenotypic Screening of Chloroplast Protein… 12. Jander, G., Norris, S. R., Joshi, V., Fraga, M., Rugg, A., Yu, S. X., Li, L. L., and Last, R. L. (2004) Application of a high-throughput HPLC-MS/MS assay to Arabidopsis mutant screening; evidence that threonine aldolase plays a role in seed nutritional quality. Plant J. 39, 465–475. 13. Gu, L. P., Jones, A. D., and Last, R. L. (2010) Broad connections in the Arabidopsis seed metabolic network revealed by metabolite profiling of an amino acid catabolism mutant. Plant J. 61, 579–590. 14. Gu, L., Jones, A. D., and Last, R. L. (2007) LC-MS/MS assay for protein amino acids and metabolically related compounds for largescale screening of metabolic phenotypes. Anal. Chem. 79, 8067–8075. 15. Ohlrogge, J., and Browse, J. (1995) Lipid biosynthesis. Plant Cell 7, 957–970. 16. Somerville, C., Browse, J., Jaworski, J. G., and Ohlrogge, J. (2000) Lipids. In, Biochemistry and Molecular Biology of Plants (Buchanan, R. B., Gruissem, W., and Jones, R., eds.) American Society of Plant Physiology Press, Rockville, MD, USA. 17. Browse, J., McCourt, P. J., and Somerville, C. R. (1986) Fatty acid composition of leaf lipids determined after combined digestion and fatty acid methyl ester formation from fresh tissue. Anal. Biochem. 152, 141–145. 18. Osteryoung, K. W., Stokes, K. D., Rutherford, S. M., Percival, A. L., and Lee, W. Y. (1998) Chloroplast division in higher plants requires members of two functionally divergent gene families with homology to bacterial ftsZ. Plant Cell 10, 1991–2004. 19. Ruuska, S. A., Schwender, J., and Ohlrogge, J. B. (2004) The capacity of green oilseeds to utilize photosynthesis to drive biosynthetic processes. Plant Physiol. 136, 2700–2709.
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20. Baud, S., Boutin, J. P., Miquel, M., Lepiniec, L., and Rochat, C. (2002) An integrated overview of seed development in Arabidopsis thaliana ecotype WS. Plant Physiol. Biochem. 40, 151–160. 21. Li, Y. H., Beisson, F., Pollard, M., and Ohlrogge, J. (2006) Oil content of Arabidopsis seeds: The influence of seed anatomy, light and plant-toplant variation. Phytochemistry 67, 904–915. 22. Less, H., and Galili, G. (2008) Principal transcriptional programs regulating plant amino acid metabolism in response to abiotic stresses. Plant Physiol. 147, 316–330. 23. Caspar, T., Lin, T. P., Kakefuda, G., Benbow, L., Preiss, J., and Somerville, C. (1991) Mutants of Arabidopsis with altered regulation of starch degradation. Plant Physiol. 95, 1181–1188. 24. Maxwell, K., and Johnson, G. N. (2000) Chlorophyll fluorescence – a practical guide. J. Exp. Bot. 51, 659–668. 25. Müller, P., Li, X. P., and Niyogi, K. K. (2001) Non-photochemical quenching. A response to excess light energy. Plant Physiol. 125, 1558–1566. 26. Kramer, D. M., Johnson, G., Kiirats, O., and Edwards, G. E. (2004) New fluorescence parameters for the determination of QA redox state and excitation energy fluxes. Photosynth. Res. 79, 209–218. 27. Lu, Y., Hall, D. A., and Last, R. L. (2011). A small zinc finger thylakoid protein plays a role in maintenance of photosystem II in Arabidopsis thaliana. Plant Cell. First Published on May 17, 2011; doi: 10.1105/ tpc.111.085456. 28. Lu, Y., Savage, L. J., Larson, M. D., Wilkerson, C. G., and Last, R. L. (2011). Choloroplast 2010: a database for large-scale phenotypic screening of Arabidopsis mutants. Plant Physiol. 155, 1589–1600.
Part III Proteomics and Suborganellar Fractionation
Chapter 10 Preparation of Envelope Membrane Fractions from Arabidopsis Chloroplasts for Proteomic Analysis and Other Studies Daniel Salvi, Lucas Moyet, Daphné Seigneurin-Berny, Myriam Ferro, Jacques Joyard, and Norbert Rolland Abstract Plastids are semiautonomous organelles restricted to plants and protists. These plastids are surrounded by a double membrane system, or envelope. These envelope membranes contain machineries to import nuclear-encoded proteins, and transporters for ions or metabolites, but are also essential for a range of plastid-specific metabolisms. Targeted semiquantitative proteomic investigations have revealed specific cross-contaminations by other cell or plastid compartments that may occur during chloroplast envelope purification. This article describes procedures developed to recover highly purified envelope fractions starting from Percoll-purified Arabidopsis chloroplasts, gives an overview of possible cross-contaminations, provides some tricks to limit these cross-contaminations, and lists immunological markers and methods that can be used to assess the purity of the envelope fractions. Key words: Chloroplast, Chloroplast envelope, Cross-contamination, Stroma, Thylakoids, Mass spectrometry, Proteome
1. Introduction Plastids are semiautonomous organelles found in plants and protists. They are universally considered to have originated from endosymbiotic cyanobacteria. In plant leaves, plastids differentiate into chloroplasts and become photosynthetically active. In addition to photosynthesis, chloroplasts are also the place of various additional functions that are critical for cell and plant life, such as nitrogen and sulfur assimilation, and syntheses of vitamins, lipids, pigments, amino acids, hormone precursors, etc. Chloroplasts have a specific suborganellar organization (see Fig. 1). They are R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_10, © Springer Science+Business Media, LLC 2011
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Fig. 1. Main compartments of the chloroplast.
surrounded by a two-membrane system or envelope. This envelope system is composed of the inner and outer membranes and of an intermembrane space located between these two layers. The chloroplasts also contain a soluble phase, called the stroma, and an internal membrane system, called the thylakoids. Within these thylakoid vesicles is found another suborganellar fraction called the thylakoid lumen. While containing their own genome, plastids only synthesize less than a 100 proteins and must import 2–3,000 nucleus-encoded proteins synthesized outside the organelle. Translocation of most of these nucleus-encoded proteins across the envelope is achieved by the joint action of TOC and TIC translocons, located, respectively, in the outer and inner membranes of the plastid envelope (1). The envelope is also involved in the controlled exchange of a variety of ions and metabolites between the cytosol and the chloroplast (2), and it is the site of specific biosynthetic functions such as the synthesis of plastid membrane components (glycerolipids, pigments, prenylquinones) or chlorophyll breakdown (3–5). The soluble phase (stroma) contains enzymes required for the photosynthesis reactions (the Calvin cycle), the synthesis of amino acids or vitamins, and the plastid transcription and translation machineries. Within the stroma are also found stacks of thylakoids, the suborganelle membrane system where the light phase of photosynthesis takes place. According to their various subplastidial localizations (inner or outer envelope membranes, intermembrane space, stroma, thylakoid membrane, or lumen), chloroplast proteins have specific functions in the organelle. In order to get access to the protein content of these different plastid subcompartments, protocols exist that allow the collection of highly pure fractions of envelope, stroma, or thylakoids. Targeted semiquantitative proteomic investigations have revealed specific cross-contaminations by other cell compartments (e.g., plasma membrane, tonoplast, mitochondrial membranes) or plastid subfractions (i.e., stroma and thylakoid membranes) that may occur during chloroplast envelope purification.
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The purpose of this article is to provide a detailed protocol to purify envelope membranes starting from Percoll-purified Arabidopsis chloroplasts, and to accurately evaluate the contamination of these purified envelope fractions with markers from other cell or chloroplast compartments that were revealed to contaminate the envelope fractions by mass spectrometry-based analyses.
2. Materials 2.1. Growth of Arabidopsis Plants
1. Arabidopsis rosette leaves are obtained from 3- to 4-week-old Arabidopsis thaliana plantlets (see Note 1). Four to six cases containing such Arabidopsis plantlets are expected to provide 400–500 g of rosette material. 2. Large (30 cm × 45 cm) plastic cases filled with compost and water. 3. Arabidopsis thaliana seeds. These should be sown onto the surface of the compost by scattering them carefully at a high density (around 30 mg of seeds for a whole case). 4. Growth rooms providing a 12-h light cycle, set at 23°C (day)/18°C (night) with a light intensity of 150 mmol/m2/s.
2.2. Purification of Chloroplasts from Arabidopsis Leaves
1. Muslin or cheesecloth, 80-cm-large. 2. Nylon blutex (50 mm aperture) (Tripette et Renaud, Sailly Saillisel, France). 3. Beakers (500 mL, 1 L, and 5 L). 4. Ice and ice buckets. 5. Pipettes (1 and 10-mL). 6. Percoll (GE Healthcare, USA). 7. Motor-driven blender, three speeds, 1 gallon (3.785 L) capacity (Waring blender). 8. Superspeed refrigerated centrifuge (Sorvall RC5), with the following rotors (and corresponding tubes): fixed-angle rotors GS-3 (6 × 500-mL plastic bottles) and SS-34 (8 × 50-mL polypropylene tubes); swinging-bucket rotor HB-6 (6 × 50-mL polycarbonate tubes). Equivalent alternative equipment (e.g., from Beckman) may also be used. 9. Leaf grinding medium: 20 mM Tricine-KOH, pH 8.4, 0.4 M sorbitol, 10 mM ethylenediaminetetraacetic acid (EDTA), 10 mM NaHCO3, and 0.1% (w/v) bovine serum albumin (BSA, defatted). 10. Chloroplast washing medium: 20 mM Tricine-KOH, pH 7.6, 0.4 M sorbitol, 5 mM MgCl2, and 2.5 mM EDTA. This solution
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should also contain the following protease inhibitors when long-term storage of protein samples is required: 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM benzamidine, and 0.5 mM e-amino caproic acid. 11. Percoll gradient solution. Mix 1 volume of Percoll with 1 volume of medium containing 40 mM Tricine-KOH, pH 7.6, 0.8 M sorbitol, 10 mM MgCl2, and 5 mM EDTA, to obtain a 50% (v/v) Percoll/0.4 M sorbitol solution. 2.3. Purification of Envelope Membranes from Arabidopsis Chloroplasts
1. Hypotonic medium for chloroplast lysis: 10 mM 3-(N-morpholino) propane sulfonic acid (MOPS)-NaOH, pH 7.8, 4 mM MgCl2. This solution should also contain the following protease inhibitors when long-term storage of protein samples is required: 1 mM PMSF, 1 mM benzamidine, and 0.5 mM e-amino caproic acid. 2. Sucrose gradient solutions for chloroplast fractionation: 10 mM MOPS-NaOH, pH 7.8, 4 mM MgCl2, and 0.3 M, 0.6 M, or 0.93 M sucrose. 3. Membrane washing medium (to wash chloroplast envelope and thylakoid membranes): 10 mM MOPS-NaOH, pH 7.8, 1 mM PMSF, 1 mM benzamidine, and 0.5 mM e-amino caproic acid. 4. Preparative refrigerated ultracentrifuge (Beckman L7), with a SW 41 Ti rotor (6 × 13.2-mL Ultraclear tubes), or equivalent. 5. Microcentrifuge (Eppendorf 5415D or equivalent) placed in a cold room with 1.5-mL plastic tubes. 6. Branson Sonifier model S-250D (or equivalent), with 3-mm microtip and ice bucket. 7. Nitrogen (or argon) gas supply (from cylinder) with gas pressure regulator connected to a Pasteur pipette via a plastic tube.
2.4. SDS-PAGE and Protein Transfer to Nitrocellulose
1. Gel electrophoresis apparatus (Bio-Rad Protean 3 or equivalent), with the various accessories needed for protein separation by electrophoresis (combs, plates, and casting apparatus). 2. Acrylamide stock: 30% (w/v) acrylamide, 0.8% (w/v) bisacrylamide. Dissolve 300 g of acrylamide and 8 g of bisacrylamide in H2O to 1 L. Alternatively, a ready-to-use acrylamide– bisacrylamide solution may be employed (e.g., Acrylamide-bis 30% (37.5:1), Merck Chemicals, Darmstadt, Germany). 3. SDS stock solution: 20% (w/v) sodium dodecyl sulfate (SDS). Dissolve 2 g of SDS in H2O to 10 mL. Store at room temperature. Alternatively, a ready-to-use 20% (w/v) SDS stock solution may be employed (e.g., Euromedex, Mundolsheim, France).
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4. 4× Laemmli stacking gel buffer: 0.5 M Tris–HCl, pH 6.8. Dissolve 363 g of Tris in H2O to 900 mL, adjust to pH 6.8 at 25°C with concentrated HCl, and make up volume to 1 L. Store at room temperature. 5. 8× Laemmli resolving gel buffer: 3 M Tris–HCl, pH 8.8. Dissolve 60.6 g of Tris in H2O to 900 mL, adjust to pH 8.8 at 25°C with concentrated HCl, and make up volume to 1 L. Store at room temperature. 6. Stacking gel (5% acrylamide). Mix 0.83 mL of 30% acrylamide/ 0.8% bisacrylamide stock solution, 1.25 mL of 4× Laemmli stacking gel buffer, 2.8 mL of H2O, 25 mL of 20% (w/v) SDS, 5 mL of N,N,N¢,N¢-tetramethylethylenediamine (TEMED), and 50 mL of 10% (w/v) ammonium persulfate (dissolve 1 g of ammonium persulfate in H2O to 10 mL; store at 4°C and prepare fresh every month). The total volume will be 4.96 mL (sufficient for two 7-cm-long gels). 7. Single acrylamide resolving gels (10, 12 or 15% acrylamide). (1) For a 10% acrylamide gel, mix 3.3 mL of 30% acrylamide/0.8% bisacrylamide stock solution, 1.25 mL of 8× Laemmli resolving gel buffer, 5.3 mL of H2O, 50 mL of 20% (w/v) SDS, 4 mL of TEMED, and 0.1 mL of 10% (w/v) ammonium persulfate. (2) For a 12% acrylamide gel, mix 4 mL of 30% acrylamide/0.8% bisacrylamide stock solution, 1.25 mL of 8× Laemmli resolving gel buffer, 4.6 mL of H2O, 50 mL of 20% (w/v) SDS, 4 mL of TEMED, and 0.1 mL of 10% (w/v) ammonium persulfate. (3) For a 15% acrylamide gel, mix 5 mL of 30% acrylamide/0.8% bisacrylamide stock solution, 1.25 mL of 8× Laemmli resolving gel buffer, 3.6 mL of H2O, 50 mL of 20% (w/v) SDS, 4 mL of TEMED, and 0.1 mL of 10% (w/v) ammonium persulfate. In each case, the total volume should be ~10 mL (sufficient for two 7-cm-long gels). 8. 4× Loading buffer for protein solubilization: 200 mM Tris– HCl, pH 6.8, 40% (v/v) glycerol, 4% (w/v) SDS, 0.4% (w/v) bromophenol blue, and 100 mM dithiothreitol. 9. Gel reservoir buffer: 38 mM glycine, 50 mM Tris, and 0.1% (w/v) SDS. Prepare about 400 mL for each reservoir. 10. Gel staining medium: acetic acid–isopropanol–water, 10/25/ 65 (v/v/v), supplemented with 2.5 g/L of Coomassie Brilliant Blue R250. Store in clean and closed bottles. 11. Gel destaining medium: acetic acid–ethanol–water, 7/40/53 (v/v/v). 2.5. Western Blots
1. System for protein transfer to nitrocellulose membranes (including central core assembly, holder cassette, filter paper (e.g., 3MM, Whatman, Maidstone, UK), fiber pads, and cooling unit).
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2. Protein transfer medium. To make this solution, dilute gel reservoir buffer (see Subheading 2.4) with ethanol to obtain a 20% (v/v) final ethanol concentration. The final buffer composition is as follows: 30.4 mM glycine, 40 mM Tris, 0.08% (w/v) SDS, and 20% (v/v) ethanol. Prepare about 800 mL for each experiment. 3. Nitrocellulose membranes (BA85, Schleicher and Schuell, Germany, or equivalent). 4. TBST (Tris-buffered saline with Triton): 0.15 M NaCl, 50 mM Tris–HCl, pH 7.5, and 0.05% (w/v) Triton X-100. 5. Milk-containing TBST. To make this solution, supplement TBST with 50 g/L of fat-free milk powder. 6. Anti-H+-ATPase (P-type) antibody (6, 7) raised against the plasma membrane H+-ATPase of Nicotiana plumbaginifolia (used at a 1:250 dilution). 7. Anti-TIP antibody (8) raised against a tobacco (Nicotiana tabacum) tonoplast protein (used at a 1:2,000 dilution). 8. Anti-Nad9 antibody (9) raised against an extrinsic protein of the wheat (Triticum aestivum) mitochondrial inner membrane (used at a 1:2,000 dilution). 9. Anti-Tom40 antibody (7, 10) raised against an outer membrane protein from Vicia faba mitochondria (used at a 1:1,000 dilution). 10. Anti-T subunit of the glycine-decarboxylase complex (11) raised against a matrix protein from pea (Pisum sativum) mitochondria (used at a 1:10,000 dilution). 11. Anti-HMA1 antibody (7) raised against a protein from the inner envelope membrane of Arabidopsis chloroplasts (used at a 1:1,000 dilution). 12. Anti-ceQORH antibody (12) raised against a protein from the inner envelope membrane of Arabidopsis chloroplasts (used at a 1:5,000 dilution). 13. Anti-KARI antibody (13) raised against a soluble protein from the stroma of spinach (Spinacia oleracea) chloroplasts (used at a 1:1,000 dilution). 14. Anti-GAPDH antibody (14) raised against a soluble protein of the chloroplast stroma from Chlamydomonas reinhardtii (used at a 1:5,000 dilution). 15. Anti-LHCP antibody (15, 16) raised against a thylakoid membrane protein from Chlamydomonas reinhardtii chloroplasts (used at a 1:25,000 dilution). 16. Solution A: 90 mM P-coumaric acid (14 mg/mL in dimethyl sulfoxide [DMSO]).
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17. Solution B: 250 mM luminol (3-aminophthalhydrazide) (44 mg/mL in DMSO). 18. 100 mM Tris–HCl, pH 8.5. 19. Chemiluminescence adapted films (Amersham Hyperfilm ECL, GE Healthcare), and a suitable exposure cassette. 20. Developer and fixer solutions, for film development under red safe-light in a dark room.
3. Methods (see Fig. 2) 3.1. Purification of Chloroplasts from Arabidopsis Leaves
All operations should be carried out at 0–5°C. 1. Prior to the experiment, prepare six tubes each containing 30 mL of a 50% Percoll/0.4 M sorbitol solution. Preform Percoll gradients for chloroplast purification by centrifugation at 38,700 × g for 55 min (Sorvall SS-34 rotor) (see Note 2). Store the tubes containing preformed Percoll gradients in the cold room until use. 2. Harvest 400–500 g of rosette leaves (see Note 3). Wash them with deionized water. Blot the washed leaves on paper tissue and transfer them into a cold room for the next step. 3. Homogenize the leaf material (for 400–500 g of leaves, use 2 L of leaf grinding medium) two times, for 2 s each time, in a Waring blender at low speed (see Note 4). Filter rapidly the homogenate through 4–5 layers of muslin and one layer of nylon blutex. 4. Distribute equally the filtered suspension into six bottles for centrifugation (500 mL each) and centrifuge them at 2,070 × g for 2 min (Sorvall GS 3 rotor) (see Note 5). 5. Suck up the supernatant with a water pump and carefully resuspend each pellet, containing a crude chloroplast fraction, by addition of a minimal volume (36 mL final volume) of chloroplast washing medium (use a spatula to gently resuspend the organelles). 6. Load the chloroplast suspension (6 mL per tube) on the top of the preformed Percoll gradients. Centrifuge the gradients at 13,300 × g for 10 min (Sorvall swinging-bucket HB-6 rotor) (see Note 6). At the conclusion of this step, aspirate the upper part of the gradient (see Note 7) and then recover intact chloroplasts (a broad, dark-green band in the lower part of the gradient; see Note 8) with a pipette. 7. Dilute three- to fourfold the intact chloroplast suspension with 200–300 mL of chloroplast washing medium. Centrifuge the suspension at 2,070 × g for 2 min (Sorvall SS-34 rotor).
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8. Recover each pellet, containing washed, purified intact chloroplasts, for chloroplast envelope preparation. At this stage, the yield of intact chloroplasts is 50–60 mg of protein (see Note 9). 3.2. Fractionation of Arabidopsis Chloroplasts to Purify the Envelope Fraction
All operations should be carried out at 0–5°C. 1. Prior to the experiment, prepare six tubes (13.2 mL, Ultraclear, Beckman) with sucrose gradients each consisting of three layers: 3 mL of 0.93 M, 2.5 mL of 0.6 M, and 2 mL of 0.3 M sucrose. Each layer should be carefully overlaid with a pipette (see Note 10) on top of the previous layer, starting with the densest one (0.93 M, at the bottom) and finishing with the lightest one. Store the tubes in the cold room until use. 2. Lyse the purified and washed intact chloroplasts (obtained as described in Subheading 3.1) by adding to the pellets hypotonic medium (adjust for a final total volume of 21 mL for all six pellets) containing protease inhibitors. 3. Load the lysed chloroplasts (3.5 mL per tube) on top of the sucrose gradients. Centrifuge the tubes at 70,000 × g for 1 h (Beckman SW41-Ti rotor) (see Note 11). After centrifugation, the envelope membranes and the thylakoids are present as a yellow band at the 0.93–0.6 M sucrose interface and a darkgreen band at the bottom of the tube, respectively. The soluble fraction containing the stroma remains on the top of this gradient. 4. To recover the soluble stromal proteins, carefully remove the upper part of the gradient with a pipette (3 mL should be recovered from each gradient tube) and store it in liquid nitrogen (see Note 12). 5. To purify envelope membranes, recover the yellow band containing the envelopes with a pipette, dilute the suspension three- to fourfold in hypotonic medium (containing protease inhibitors), and concentrate the membranes as a pellet by centrifugation at 110,000 × g for 1 h (Beckman SW 41 Ti rotor). 6. Add a minimal volume of membrane washing medium (containing protease inhibitors) to the envelope pellet. Take an aliquot for protein amount determination (see Note 13). Store envelope membrane preparations in liquid nitrogen. 7. Thylakoids can be diluted in membrane washing medium (containing protease inhibitors) and stored in liquid nitrogen (see Note 14). 8. From such preparations, an average of ~30 mg of stroma proteins, ~20 mg of thylakoid proteins, and ~300 mg of envelope proteins can be obtained (see Note 15).
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3.3. Separation of Envelope Proteins by 1D SDS-PAGE (see Note 16)
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1. Prior to the experiment, prepare slab gels for protein electrophoresis (see Note 17). Assemble the gel apparatus according to the manufacturer’s specifications (see Note 18). Prepare the different gel solutions (stacking gel, and 10, 12, or 15% acrylamide separation gel). The volumes to be used are determined by gel dimensions and, therefore, by the specifications of the apparatus. 2. Several dilutions (a factor of 3 for every dilution step) of the protein samples might be performed to estimate cross-contaminations during further Western blot analyses (see Note 19 and Fig. 2). If Western blotting is to be conducted (see Subheading 3.4), the Coomassie staining step (step 5 below) should be omitted. 3. Add 4× loading buffer to the samples. Heat the protein samples at 95°C for 5 min to solubilize the proteins. Place protein samples (20 mL) into gel slots by means of a pipette. Load the molecular weight markers in another slot. 4. Set the conditions for the electrophoresis at 150 V (~20 V/cm). Run the gels for 1 h at room temperature (until the bromophenol blue dye reaches the lower part of the gel). 5. After electrophoresis, remove the gels from the apparatus; place them in plastic boxes in the presence of gel staining medium. Shake the box gently for 30 min. Pour off the staining solution and replace it with the gel destaining medium. Shake the box gently for 15 min. Repeat the washing step once or twice. Typical results are shown in Fig. 2 (step 4).
3.4. Immunological Markers: Western Blot Analyses (see Note 20)
Having access to highly purified chloroplast envelope fractions is a prerequisite to answer many essential biological questions. However, while they are well-established, the protocols used to purify plastid subcompartments cannot totally exclude cross-contaminations originating from other plastids and cell compartments (see Fig. 2). An inventory of the origins of these cross-contaminations (see Table 1) has recently been performed using semiquantitative proteomic approaches (13). The aim of the following assays is to assess marker proteins deriving from these cell or plastid compartments in purified envelope proteins. The presence of these markers should be tested in purified envelope extracts, as well as in crude cell extract and, when available, corresponding preparations of other cellular (plasma membrane, mitochondrial membranes, tonoplast, etc.) or chloroplastic (stroma, thylakoids) compartments, and then compared to assess cross-contamination levels. Western blots should be performed after separation of proteins by SDS-PAGE (see Subheading 3.3). After gel migration, transfer the proteins onto a nitrocellulose membrane using a gel transfer apparatus.
Step1: growth of Arabidopsis plants and collection of plant leaves Step 2: purification of chloroplasts on Percoll gradient Soluble components of the cell: cytosol, mitochondrial matrix, chloroplast stroma, vacuolar proteins…
Percoll gradients: preformed from 50% Percoll / 0.4 M sorbitol solution
Broken chloroplasts Intact mitochondria Intact chloroplasts Intact cells, cell debris, cell walls, nuclei, starch, DNA…
Step 3: fractionation of chloroplasts on sucrose gradient Soluble proteins from the mitochondrial matrix Stroma
Sucrose gradient, 3 sucrose layers: 2 mL of 0.3 M 2.5 mL of 0.6 M 3 mL of 0.93 M
Light membrane vesicles: plasma mb., mitochondrial mb., tonoplast, ER, Golgi, light thylakoid vesicles (stroma lamellae), plastoglobules… Envelope Thylakoids Nuclei, starch, DNA, fusion of thylakoid and envelope vesicles…
Step 4: SDS-PAGE to visualize abundant marker proteins kDa 118 85
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20
M: CE: Cp: E: S: T:
Protein markers Crude leaf extract Crude chloroplast Envelope: triose-P/Pi translocator (TPT) Stroma: large subunit of Rubisco (RBCL) Thylakoids: light harvesting complex proteins (LHCPs)
Step 5: SDS-PAGE + western blots to evaluate cross contaminations E M Ratio 1 µg 10
Thylakoids 1/100 1/60 1/30 1/10 1/6 1/3 1 0.1 0.17 0.33 1 1.67 3.33 10
SDS-PAGE TPT
Western blot
LHCP
E: Envelope proteins M: Protein markers Thylakoids: Various dilutions of the purified thylakoid fraction LHCPs: Light harvesting complex proteins
LHCPs antibody (1/25,000)
Fig. 2. Origin of potential cross-contaminations occurring during chloroplast and envelope purification processes. A scheme of the steps used to purify chloroplast envelope membranes from Arabidopsis thaliana, and to evaluate cross-contamination of envelope preparation with other cell or chloroplast compartments, is shown. The location of proteins or cell compartments that might contaminate chloroplast (step 2) and envelope (step 3) preparations is supplied (in italics). Step 4 is required to visualize abundant markers of the envelope (TPT, a 30 kDa phosphate/triose-phosphate translocator), stroma (RBCL, the 45 kDa large subunit or Rubisco), and thylakoids (LHCPs, 26 kDa light harvesting complex proteins). Western blots can be performed to evaluate the cross-contamination of chloroplast (Cp) and envelope (E) preparations with marker proteins from other cell compartments that will also be detected in the crude leaf extract (CE). Step 5 is a combination of SDS-PAGE and Western blots that needs to be performed to estimate cross-contaminations of the purified envelope fractions with other chloroplast compartments (e.g., thylakoid contamination of envelope preparations is approximately 3%, as revealed by the LHCP blot shown). This last step can be reproduced with a marker for any other purified cell or plastid compartment, depending on the origin of the suspected cross-contamination.
14.8
69
23
S + E
14
87
S
Mit
14.7
713
2,067
S + E
8.9
1,683
S
1.7
321
Th
Mit
0.13
24
Per
0.64
4
Per
0.13
25
PM
0.64
4
PM
1.36
258
Ton
2.73
17
Ton
0.26
50
Cyt
3.69
23
Cyt
– 10 3 nd
52.49 25.04 14.45 8.03
– 10 3 nd
79.31 12.67 6.08 1.94 100
%
%
100
WB
%
When considering spectral counting, and in agreement with the levels of cross-contamination estimated using Western blot (WB) experiments, envelope membranes appear to contain up to 10% stroma proteins and 6% thylakoid membranes proteins (13), but do not contain detectable amounts of PM or mitochondrial marker proteins (7) The upper part of the table provides information about numbers (and origins) of proteins detected in purified envelope fractions (13). The lower part of the table gives sums of spectra (spectral counts) detected for every peptide deriving from these proteins. Note that, if one only considers protein numbers, this overestimates cross-contamination levels. For example, cytosol contamination is relatively high (3.69%) if one calculates the ratio of detected cytosolic proteins to the number of all proteins detected in the purified envelope fraction. On the contrary, if one compares the spectral count corresponding to peptides deriving from these few cytosolic proteins, to the spectral count corresponding to peptides deriving from all the proteins present in the purified envelope fraction, this ratio is very low (0.26%). When comparing data obtained from Western blots with the ones obtained from semiquantitative proteomics, it appears that spectral counting better reflects the cross-contamination of envelope fractions than protein numbers. Some proteins were found to be shared between envelope and thylakoids (Th + E) or envelope and stroma (S + E), and their presence within both compartments was above measured cross-contamination levels and thus resulting from dual localization. For calculation, these proteins were classified in the compartment (envelope, stroma or thylakoid) in which they were more abundant. This explains why proteins shared by envelope and thylakoids can be found either in the envelope lane or in the thylakoid lane (Th + E). The same principle was applied to proteins shared by envelope and stroma (S + E) The abbreviations used in the table are as follows: IM inner envelope membrane, OM outer envelope membrane, E? suspected envelope localization, Th + E shared by dual localization in thylakoid and envelope membranes, S + E shared by dual localization in the stroma and the envelope, S stroma, Th thylakoids, Mit mitochondria, Per peroxysomes, PM plasma membrane, Ton tonoplast, Cyt cytosol, Nuc nucleus, WB refers to cross-contaminations estimated from Westerns blots (7, 13), nd not detected
14
828
1,810
Th + E
0.05
2.83
535
E?
%
9.78
1,850
OM
10
46.2
8,734
IM
Nonplastid
Thylakoids
Stroma
Envelope
Spectral counting
8.19
51
Th
0.32
13.3
39
44
Th + E
%
9.31
58
E?
2
4.01
25
OM
Nonplastid
Thylakoids
28.4
177
Envelope
Stroma
IM
Protein numbers
Table 1 Evaluation of the cross-contaminations of envelope membrane fractions extracted from Arabidopsis chloroplasts using semiquantitative proteomic data
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1. Prepare the cassette as follows. Add the following successively: 1 × fiber pad, 3 × filter papers, the gel, a nitrocellulose membrane, 3 × filter papers, and 1 × fiber pad. Then, insert the sandwich into the holder cassette (the membrane should be placed beside the positive electrode). 2. Insert the cassette in the central core assembly unit (together with the cooling unit). 3. Perform the transfer for 2 h at 80 V in protein transfer medium, at room temperature. 4. Recover the nitrocellulose membrane. The following incubation and washing steps (steps 5–10 below) require agitation on a rocking plate. 5. Rinse the nitrocellulose membrane with TBST for 10 min. 6. Saturate the nitrocellulose membrane with milk-containing TBST. Leave it for at least 1 h at room temperature. 7. Add the primary antibody diluted in milk-containing TBST. Leave it for 3 h at room temperature or 12 h at 4°C (see Note 21). 8. Wash the nitrocellulose membrane three times, for 10 min each time, with TBST. 9. Add the secondary antibody diluted at 1/10,000 in TBST (see Note 22). Leave it for 1.5 h at room temperature. 10. Wash the nitrocellulose membrane three times, for 10 min each time, with TBST. Then, proceed to detect the chemiluminescent signal as described in the following steps. 11. Mix 3 mL of 100 mM Tris–HCl, pH 8.5, with 13.3 mL of Solution A. 12. Mix 3 mL of 100 mM Tris–HCl, pH 8.5, with 30 mL of Solution B. 13. Combine and mix together the two above solutions (prepared in steps 11 and 12 above) in a dark room. 14. Incubate the nitrocellulose membrane for 1 min in the previously prepared mixture (the chemiluminescence substrate solution). 15. Expose to film for a few seconds and up to several minutes depending on the detected signal. 16. Incubate the film successively in the developer solution (for 1–3 min, depending on the signal to noise ratio), in water (for 10 s), and in the fixer solution (for 2 min). Rinse the film in water and dry it. Typical results are shown in Fig. 2 (step 5).
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4. Notes 1. The procedures described in this article were applied efficiently to the three most frequently used Arabidopsis ecotypes (Landsberg erecta, Columbia, and Wassilewskija). It is tempting to start with large amounts of mature material (large rosettes with big leaves). It, however, appears that staring from younger Arabidopsis leaves (3- to 4-week-old) improves yield, purity, and integrity of the purified chloroplasts. In our hands, isolation of envelope membranes from older leaves was unsuccessful. 2. Vertical rotors can easily be used to obtain preformed Percoll gradients and to subsequently purify chloroplasts (17). 3. The number of starch granules present in chloroplasts is critical for the preparation of intact chloroplasts: chloroplasts containing large starch grains will generally be broken during centrifugation (17). Therefore, prior to the experiment, the plants can be kept in a dark and cold room (4°C) to reduce the amount of starch. A good way to proceed is to place the plants under such conditions at the beginning of the afternoon prior to the day of the experiment. 4. It is critical to limit the grinding process to 2 s. It can be frustrating to limit the grinding and thus to apparently “lose” a lot of plant material. However, while longer blending strongly improves the yield of recovered chlorophyll, it also increases the ratio of broken chloroplasts. When large amounts of broken chloroplasts are present in the suspension, this definitively affects the efficiency of the Percoll gradient (see Fig. 2) in separating intact and broken organelles. 5. It is essential to equilibrate two-by-two on a balance the different tubes prior to centrifugation. 6. It is recommended to disconnect the brake or to use the automatic rate controller (if available) to prevent mixing of the gradients at the critical stage of deceleration. 7. It is important to remove carefully the top content of the tube by aspiration with a water pump. This allows the removing of soluble proteins derived from various cell compartments, including the cytosol, mitochondrial matrix, chloroplast stroma, and vacuole, that were copurified with the chloroplasts during earlier steps (see Fig. 2). Broken chloroplasts are present in the upper part of the gradient as a broad band and must be removed by aspiration. This part of the gradient also contains remaining intact mitochondria that will be removed with the broken chloroplasts (see Fig. 2). Then, recover the intact chloroplasts with
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a pipette. To limit breakage of chloroplasts, if using the blue tip of an Eppendorf pipette, cut the tip with a razor blade so that the hole will have a 2-mm diameter. 8. A small pellet (containing cell pieces, large debris, starch, DNA, nuclei, etc.; see Fig. 2) is found at the bottom of the Percoll gradient. It is, thus, essential to use recovering conditions that will limit aspiration of this pellet with the intact chloroplasts. Note that nuclei are not broken during the osmotic shock used to fractionate intact chloroplasts. Thus, when some nuclei are taken with the intact chloroplasts at this step, they will later be recovered within thylakoid fractions, at the bottom of the sucrose gradient (see Fig. 2). 9. Percoll-purified chloroplasts are largely devoid of contamination deriving from other cell compartments. However, since some minor contaminants can be present, caution must be taken when assigning a protein to a subcellular location, even though the cell compartment of interest can be highly purified. Considering the high sensitivity of present mass spectrometers, it is not surprising, using proteomic analyses, to detect minute amounts of a few extraplastidial contaminants, which are major proteins in their respective subcellular compartment. In every case, other complementary approaches are necessary to assert the subcellular localization of a protein (immuno-localization of proteins, expression of GFP fusions in planta coupled to confocal microscopy, etc.). 10. The use of a peristaltic pump to prepare the sucrose gradients is recommended (avoiding mixing of the layers), since some expertise is needed to load the different layers by hand. 11. It is recommended to disconnect the brake to prevent mixing of the gradients at the critical stage of deceleration. 12. The proteins from the stroma will be recovered in the hypotonic medium used to break the chloroplasts and this fraction will, thus, contain protease inhibitors (i.e., 1 mM PMSF, 1 mM benzamidine, and 0.5 mM e-amino caproic acid). Further desalting of these soluble proteins may be performed using G-25 columns (e.g., PD-10, GE Healthcare) if required. Note that if intact mitochondria are recovered from the Percoll gradient step (due to incomplete removal of the upper part of the Percoll gradient containing broken chloroplasts; see Note 7 and Fig. 2), these mitochondria will also be ruptured by the osmotic shock used to fractionate the chloroplasts, and soluble proteins from the mitochondrial matrix will copurify with soluble proteins from the stroma (see Fig. 2). It is, however, important to note that while the detection of genuine mitochondrial proteins in the purified chloroplast stroma may be the result of cross-contamination, the “dual targeting” of
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some proteins known to be imported into both plastids and mitochondria might also explain their presence (18). 13. Protein contents of fractions are estimated using the Bio-Rad Protein Assay reagent (19). 14. The major possible contaminations of crude thylakoid membrane preparations derive from envelope membrane proteins (see Fig. 2 and Table 1). It is estimated that 50% of the envelope membrane vesicles are recovered in the crude thylakoid fraction. However, since the ratio of envelope to thylakoid membranes is 1/50 in the chloroplast, these contaminations will be limited to less than 1% of the crude thylakoid membrane fraction. Alternatively, heavy components of the cell might be recovered within the thylakoid pellet if originally aspirated with intact chloroplasts from the bottom of the Percoll gradient (see Note 8 and Fig. 2). 15. At this stage, the major possible contaminants of envelope preparations are soluble stroma proteins and light vesicles of thylakoid membranes. Being the most likely source of membrane contamination of the purified envelope fraction (13, 16), thylakoid cross-contamination needs to be precisely assessed. The yellow color of purified envelope vesicles should first indicate that this membrane system contains almost no chlorophyll and, therefore, very few contaminating thylakoid vesicles. By Western blot analyses using antibodies raised against LHCP, Ferro et al. (16) demonstrated that several independent Arabidopsis envelope preparations contained an average of 3% thylakoid proteins. These data are in agreement with the 6% cross-contamination of envelope membrane preparation with thylakoid membranes (Table 1) as estimated using spectral counting (13). Since the envelope membranes are at the interface of two soluble media (i.e., the cytosol and the stroma), and since soluble proteins from both sides of the membrane might be trapped within membrane vesicles during chloroplast rupture (during osmotic shock), envelope contamination with these soluble fractions might be expected. However, cytosolic contaminations are barely detected within purified envelope fractions (Table 1) and mostly result from the detection of cytosolic ribosomal subunits (13). On the contrary, stroma proteins might represent up to 10% of purified envelope fractions, as estimated from Western blot analyses (13) or spectral counting (Table 1). Various treatments (Na2CO3, NaCl, chloroform–methanol, NaOH) of envelope membranes were shown to extract proteins that are rather weakly associated with the membrane including most soluble contaminants from the stroma (16, 20, 21). It might, therefore, be recommended to try several of these treatments to remove abundant soluble contaminants. However, one has to consider that such treatments also
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remove genuine envelope proteins, i.e., the less hydrophobic envelope membrane proteins. In other words, these approaches should be limited to experiments aiming to analyze intrinsic and hydrophobic envelope membrane proteins that will remain within treated envelope vesicles. 16. Classical proteomic approaches are based on the use of twodimensional (2D) gel electrophoresis that proved to be very efficient for the analysis of soluble proteins or peripheral membrane proteins. On the contrary, this approach is rather inefficient at analyzing intrinsic membrane proteins. Therefore, thylakoid and envelope fractions are preferably analyzed by SDS-PAGE, while further proteomic studies targeted to the stroma (or the thylakoid lumen) can either rely on SDS-PAGE or 2D-gel electrophoresis. 17. We routinely use the procedure described by Chua (22) to separate membrane proteins by SDS-PAGE. This article describes in detail all stock solutions, and medium for stacking and separation gels. 18. We use a Bio-Rad apparatus, with 7-cm-long gels. 19. As a first step, SDS-PAGE analyses should be used to study the purified envelope fraction to detect known abundant markers associated with this fraction and a lack of markers deriving from other plastidic or cellular compartments (Fig. 2, step 4). If cross-contaminations are revealed, additional SDS-PAGE analyses should then include experiments aiming to quantify these cross-contamination levels. For this, it is essential to use dilutions of protein samples deriving from those cell compartments that were shown to cross-contaminate the purified envelope fractions (Fig. 2, step 5). In order to evaluate these cross-contamination levels using Western blots, one method would be to quantify the relative strength of the fluorescence or chemiluminescence signal (using appropriate tools and materials) detected in the envelope fraction, and compare it with that obtained in the fraction suspected to contaminate the envelope fraction. Another option relies on the dilution of the purified protein sample corresponding to the compartment that is suspected to contaminate envelope fraction (see Fig. 2, step 5), and on the search for a dilution that provides similar signals in both samples (10). 20. Follow the instructions for saturation (blocking) and incubation of the membrane with primary and secondary antibodies provided by the manufacturer. 21. Several dilutions of the primary antibodies should be tested to determine the best signal–noise ratio. 22. In our case, since the used primary antibodies were obtained from rabbit antisera, the secondary antibody is an anti-rabbit-IgG
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antibody coupled to the horseradish peroxidase for further detection. However, the secondary antibody has to be adapted to the primary antibody (which may be from rabbit, mouse, goat, guinea pig, etc.) and the detection procedure to be used (alkaline phosphatase or horseradish peroxidase). References 1. Jarvis, P. (2008) Targeting of nucleus-encoded proteins to chloroplasts in plants. New Phytol. 179, 257–285. 2. Linka, N., and Weber, A. P. (2010) Intracellular metabolite transporters in plants. Mol. Plant 3, 21–53. 3. Block, M. A., Douce, R., Joyard, J., and Rolland, N. (2007) Chloroplast envelope membranes: a dynamic interface between plastids and the cytosol. Photosynth. Res. 92, 225–244 4. Joyard, J., Ferro, M., Masselon, C., SeigneurinBerny, D., Salvi, D., Garin, J., and Rolland, N. (2009) Chloroplast proteomics and the compartmentation of plastidial isoprenoid biosynthetic pathways. Mol. Plant 2, 1154–1180. 5. Joyard, J., Ferro, M., Masselon, C., SeigneurinBerny, D., Salvi, D., Garin, J., and Rolland, N. (2010) Chloroplast proteomics highlights the subcellular compartmentation of lipid metabolism. Prog. Lipid Res. 49, 128–158 6. Morsomme, P., Dambly, S., Maudoux, O., and Boutry, M. (1998) Single point mutations distributed in 10 soluble and membrane regions of the Nicotiana plumbaginifolia plasma membrane PMA2 H+−ATPase activate the enzyme and modify the structure of the C-terminal region. J. Biol. Chem. 273, 34837–34842. 7. Seigneurin-Berny, D., Gravot, A., Auroy, P., Mazard, C., Kraut, A., Finazzi, G., Grunwald, D., Rappaport, F., Vavasseur, A., Joyard, J., Richaud, P., and Rolland, N. (2006) HMA1, a new Cu-ATPase of the chloroplast envelope, is essential for growth under adverse light conditions. J. Biol. Chem. 281, 2882–2892. 8. Gerbeau, P., Guclu, J., Ripoche, P., and Maurel, C. (1999) Aquaporin Nt-TIPa can account for the high permeability of tobacco cell vacuolar membrane to small neutral solutes. Plant J. 18, 577–587. 9. Combettes, B., and Grienenberger, J. M. (1999) Analysis of wheat mitochondrial complex I purified by a one-step immunoaffinity chromatography. Biochimie 81, 645–653. 10. Perryman, R. A., Mooney, B., and Harmey, M. A. (1995) Identification of a 42-kDa plant mitochondrial outer membrane protein, MOM42,
11.
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14.
15.
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involved in the import of precursor proteins into plant mitochondria. Arch. Biochem. Biophys. 316, 659–664. Vauclare, P., Macherel, D., Douce, R., and Bourguignon, J. (1998) The gene encoding T protein of the glycine decarboxylase complex involved in the mitochondrial step of the photorespiratory pathway in plants exhibits features of light induced genes. Plant Mol. Biol. 37, 309–318. Miras, S., Salvi, D., Piette, L., SeigneurinBerny, D., Grunwald, D., Garin, J., Reinbothe, C., Joyard, J., Reinbothe, S., and Rolland, N. (2007) Toc159- and Toc75-independent import of a transit sequence-less precursor into the inner envelope of chloroplasts. J. Biol. Chem. 282, 29482–29492. Ferro, M., Brugière, S., Salvi, D., SeigneurinBerny, D., Court, M., Moyet, L., Ramus, C., Miras, S., Mellal, M., Le Gall, S., KiefferJaquinod, S., Bruley, C., Garin, J., Joyard, J., Masselon, C., and Rolland, N. (2010) AT_ CHLORO, a comprehensive chloroplast proteome database with subplastidial localization and curated information on envelope proteins. Mol. Cell. Proteomics 9, 1063–1084. Seigneurin-Berny, D., Salvi, D., Dorne, A.-J., Joyard, J., and Rolland, N. (2008) Percollpurified and photosynthetically active chloroplasts from Arabidopsis thaliana leaves. Plant Physiol. Biochem. 46, 951–955. Vallon, O., Bulte, L., Dainese P., Olive, J., Bassi, R., and Wollman, F. A. (1991) Lateral redistribution of cytochrome b6/f complexes along thylakoid membranes upon state transitions. Proc. Natl. Acad. Sci. USA 88, 8262–8266. Ferro, M., Salvi, D., Brugière, S., Miras, S., Kowalski, S., Louwagie, M., Garin, J., Joyard, J., and Rolland, N. (2003) Proteomics of the chloroplast envelope membranes from Arabidopsis thaliana. Mol. Cell. Proteomics 2, 325–345. Douce, R., and Joyard, J. (1982) Purification of the chloroplast envelope. In, Methods in Chloroplast Molecular Biology (Edelman, M., Hallick, R., and Chua, N.H., eds.) Elsevier, Amsterdam, North-Holland, pp. 139–256.
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18. Carrie, C., Giraud, E., and Whelan, J. (2009) Protein transport in organelles: dual targeting of proteins to mitochondria and chloroplasts. FEBS J. 276, 1187–1195. 19. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of proteindye binding. Anal. Biochem. 72, 248–254. 20. Seigneurin-Berny, D., Rolland, N., Garin, J., and Joyard, J. (1999) Differential extraction of hydrophobic proteins from chloroplast envelope membranes: a subcellular-specific proteomic
approach to identify rare intrinsic membrane proteins. Plant J. 19, 217–228. 21. Ferro, M., Salvi, D., Rivière-Rolland, H., Vermat, T., Seigneurin-Berny, D., Grunwald, D., Garin J., Joyard, J., and Rolland, N. (2002) Integral membrane proteins of the chloroplast envelope: identification and subcellular localization of new transporters. Proc. Natl. Acad. Sci. USA 99, 11487–11492. 22. Chua, N. H. (1980) Electrophoretic analysis of chloroplast proteins. Methods Enzymol. 69, 434–436.
Chapter 11 Preparation of Stroma, Thylakoid Membrane, and Lumen Fractions from Arabidopsis thaliana Chloroplasts for Proteomic Analysis Michael Hall, Yogesh Mishra, and Wolfgang P. Schröder Abstract For many studies regarding important chloroplast processes such as oxygenic photosynthesis, fractionation of the total chloroplast proteome is a necessary first step. Here, we describe a method for isolating the stromal, the thylakoid membrane, and the thylakoid lumen subchloroplast fractions from Arabidopsis thaliana leaf material. All three fractions can be isolated sequentially from the same plant material in a single day preparation. The isolated fractions are suitable for various proteomic analyses such as simple mapping studies or for more complex experiments such as differential expression analysis using two-dimensional difference gel electrophoresis (2D-DIGE) or mass spectrometry (MS)-based techniques. Besides this, the obtained fractions can also be used for many other purposes such as immunological assays, enzymatic activity assays, and studies of protein complexes by native-polyacrylamide gel electrophoresis (native-PAGE). Key words: Arabidopsis thaliana, Lumen isolation, Thylakoid isolation, Stroma isolation, Yeda press
1. Introduction The chloroplast of Arabidopsis has been predicted to contain more than 2,000 proteins (1). This number is approximately within the range of resolution of a large 2D-PAGE, and it might in theory be possible to resolve the entire chloroplast proteome in one 2D-electrophoresis experiment. However, such a global approach normally only exposes a small fraction of the expected proteins; see for instance (2). One of the main reasons for this is that the chloroplast contains six protein containing compartments: the outer envelope, envelope membrane space, inner envelope, stroma, thylakoid membrane, and the thylakoid lumen (Fig. 1). The subcellular
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_11, © Springer Science+Business Media, LLC 2011
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Fig. 1. A schematic drawing of a chloroplast, showing the complex architecture of the membranes found in this organelle. The size of the organelle is roughly 10 × 1 mM, and thus, it is important to use nylon cloth with a mesh size of 20 mM to ensure that the organelle passes through but that unbroken cells and nuclei are filtered out.
proteomes of each of these compartments pose their individual challenges. For instance, for the envelope, a low amount of material is usually available and it mainly consists of hydrophobic membrane proteins, which in part are difficult to class as chloroplast proteins because transit peptides are not present; for the stroma, dynamic range problems because of the high abundance of ribulose-bisphosphate carboxylase (RuBisCo); for thylakoid membranes, hydrophobic membrane proteins and dynamic range problems because of the high abundance of PSII and light harvesting complex (LHC) proteins; for the thylakoid lumen, low abundance of lumenal proteins as compared to other chloroplast proteins. Thus, in four of these compartments, the proteins are integral components or alternatively more or less strongly extrinsically associated with the membrane. The standard 2D-PAGE methodology is known to be inadequate in resolving these types of proteins. Another reason for the low recovery is that the chloroplast contains two dominating proteins, the stromal located RuBisCo and the thylakoid membrane located LHC proteins, which constitute more than 50% of the total protein content in the chloroplast. Thus, the dynamic range is another challenge for a global approach based on 2D-PAGE or other proteomic methods.
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One way to overcome these problems is to fractionate the chloroplast, which so far has been the preferred way for most researchers working on the chloroplast proteome. Fractionation of the chloroplast will not only increase the amount of identified proteins but also enable making a first localization analysis of proteins within the chloroplast. By combining proteomes of fractionated chloroplasts from various studies and MS techniques, the Arabidopsis SUBA database has experimentally verified 1,110 chloroplast proteins (3). The starting material for any kind of fractionation is Arabidopsis chloroplasts. In general, compared to spinach and pea, the Arabidopsis chloroplast tends to be much more fragile, and the protocols developed for spinach cannot be directly transferred to the new model organism Arabidopsis. We normally use soil-grown plants, but hydroponically or plate-grown plants can also be used for isolation. There are now several methods for obtaining highly intact Arabidopsis chloroplasts that are import-competent (4, 5); see also Chapters 17, 18, and 20, Vol. 1. The crucial step in obtaining high purity is a Percoll gradient centrifugation step. The drawback of this procedure is that the yield of highly intact chloroplasts is quite low even if several gradients are applied. Another problem is that the residual amount of Percoll could interfere with the fractionation steps. To increase speed and yields, we normally omit the Percoll gradient step, and instead perform two quick lowspeed (1,000 × g) centrifugations. This will give high yields of reasonably pure and intact chloroplasts. The reason that this is adequate is that in the fractionation steps that follow for thylakoid membranes and the thylakoid lumen several additional washing steps are included to ensure that contaminants are removed. In the case of stroma isolation, we suggest that Percoll purified chloroplasts are used if a highly pure stroma fraction is required, although the method for stroma isolation described here may be sufficient in many cases. The stroma of the chloroplast is often incorrectly considered to be an aqueous solution even though the protein concentration in this compartment has been estimated to be more than 20 mg protein/mL (6). Normally, the stroma fraction is obtained by classical hypotonic lysis of chloroplasts (see for instance refs. 7, 8), and its oligomeric form, i.e., various complexes and/or subcomplexes formed in the stroma, was analyzed by loading isolated stroma onto a native PAGE (7). The majority of enzymes found in this fraction are involved in the Calvin cycle, but proteins involved in protein, starch, lipid, amino acid, and pigment synthesis are also located in this compartment. This means that a quite large variety of proteins can be detected in this fraction, even though the dominating proteins are the large and small subunits of RuBisCo. It should be mentioned that this protein can be removed by immunoprecipitation or an immunoaffinity chromatography step.
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Unfortunately, thylakoid membrane proteins are much more difficult than stroma and thylakoid lumen fractions to separate and to analyze experimentally on 2D-PAGE, because of their hydrophobic nature, resulting in insolubility and adsorption, and incompatibilities of ionic detergents with mass spectrometry. Different experimental strategies for large-scale identification of membrane proteins have been explored (9). Chloroplasts in green algae and higher plants contain photosynthetic thylakoid membranes with four major multisubunit protein complexes (photosystem I [PSI], photosystem II [PSII], ATP synthase, and cytochrome b6f complexes), each with multiple cofactors. These four complexes are composed of at least 70 different proteins which perform the photosynthetic reactions (10–12). For 2D-PAGE analysis of thylakoid membrane fractions we use a combination of DHCP (diheptanoyl phosphatidylcholine, which has been shown to effectively solubilize biological membranes (13, 14)) and Triton X-100. DHPC is a phospholipid with a short fatty acyl chain of seven carbon atoms, which has no net charge and is stable over a wide pH range of 4–10. After obtaining thylakoid membranes, the last step described in this chapter is the isolation of the lumen fraction. This compartment was originally considered to not contain that many proteins and to mainly serve as space for the production of the proton gradient needed for ATP synthase and for the balancing of some counter ions. The first in-depth isolation and analysis of the thylakoid lumen fraction to be published was from spinach (6); other publications since then are focused on Arabidopsis (15) and pea (16). These studies showed that this compartment contains a unique proteome, clearly different from the stroma. These original studies have been followed by other work (17, 18). In these studies, the lumen fraction is obtained by disruption (Yeda press) of isolated thylakoid membranes, without salts present to obtain the most hydrophilic lumen proteins, or alternatively in the presence of salts to release membrane proteins interacting with the inside of the thylakoid membrane. The chloroplast fractionation procedure for stroma, thylakoids, and lumen reported in detail here has been successfully used in proteomic studies by us (6, 15, 18, 19) as well as other groups (8, 20–25).
2. Materials 2.1. Plant Material and Chloroplast Extraction
1. 4–8-week-old Arabidopsis thaliana plants, grown under short day conditions on soil for maximal yield of leaf material. 2. Chloroplast extraction buffer: 20 mM N-(tris[hydroxymethyl]) methylglycine (Tricine)–NaOH, pH 8.4, 300 mM sorbitol,
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10 mM KCl, 10 mM Na-ethylenediaminetetraacetic acid (EDTA), 0.25% (w/v) bovine serum albumin (BSA) (add just before use), 4.5 mM sodium ascorbate (add just before use), 5 mM l-cysteine (add just before use) (see Notes 1 and 2). Store at 4°C. 3. Chloroplast wash buffer: 20 mM N-2-hydroxyethyl piperazine-N ¢-2-ethanesulfonic acid (HEPES)–NaOH, pH 7.8, 300 mM sorbitol, 10 mM KCl, 2.5 mM Na-EDTA, 5 mM MgCl2. Store at 4°C. 4. Homogenizer/blender with sharp blades, a nylon cloth with mesh size 20 mM (see Note 3), a 3-L Erlenmeyer flask, a large funnel, a soft artist’s paintbrush, and ice. 5. Approximately twenty 40 mL uncapped centrifuge tubes and a corresponding cooled centrifuge with rotor capable of 1,000– 20,000 × g, such as a Beckman JA-20 or Sorvall SS-34 rotor. 6. Cold 80% (v/v) acetone for determination of chlorophyll concentration. Store at −20°C. 2.2. Isolation of Chloroplast Stroma Proteins
1. Cooled centrifuge equipment with rotor. 2. 40-mL glass Potter homogenizer with pestle A (Tissue grind tube, Kontes Glass Company) and a soft artist’s paintbrush. 3. Osmotic shock buffer: 10 mM Na-pyrophosphate-NaOH, pH 7.8. Store at 4°C. 4. Amicon Ultra-15 10K/3K ultrafiltration concentrators or similar (Millipore, Billerica, MA, USA). 5. Cooled ultracentrifuge, rotor, and corresponding centrifuge tubes capable of a relative centrifugal force of 100,000 × g. 6. Bradford reagents for protein concentration determination.
2.3. Isolation of Thylakoid Membranes
1. Centrifuge equipment according to Subheading 2.1, step 5, above. 2. 40 mL glass Potter homogenizer with pestle A (Tissue grind tube, Kontes Glass Company) and a soft artist’s paintbrush. 3. Osmotic shock buffer: 10 mM Na-pyrophosphate-NaOH, pH 7.8. Store at 4°C. 4. Thylakoid wash buffer I: 2 mM Tricine–HCl, pH 7.8, 300 mM sucrose. Store at 4°C. 5. Thylakoid wash buffer II: 30 mM NaH2PO4–NaOH, pH 7.8, 5 mM MgCl2, 50 mM NaCl, 100 mM sucrose, 1 mM Na-EDTA. Store at 4°C. 6. Kitchen wire mesh sieve and liquid nitrogen for flash freezing of thylakoid membranes.
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Fig. 2. Use of the Yeda press. One crucial point when breaking the thylakoid membranes is the Yeda press, which uses shearing forces to break the thylakoids and release the lumen fraction for isolation. Ensure that the Yeda press is correctly assembled, and be careful as high pressure (100 bar) is applied. Carefully open and close the outlet repeatedly and collect the ruptured thylakoids in the centrifuge tube.
2.4. Isolation of Thylakoid Lumen Proteins
1. Yeda press high-pressure steel chamber (originally obtained from LINCA-Lamon Instrumentation Co., Israel; see Note 4), nitrogen gas flask at a minimum pressure of 100 bar, and protective eye wear (see Fig. 2). 2. Cooled ultracentrifuge, rotor, and corresponding centrifuge tubes capable of a relative centrifugal force of 200,000 × g.
2.5. Preparation of Protein Samples for 2D Gel Electrophoresis
1. 100% acetone. Store at −20°C. 2. 2D Quant kit (GE Healthcare, Uppsala, Sweden). 3. Rehydration buffer for soluble proteins: 7 M urea, 2 M thiourea (see Note 5), 4% (w/v) 3-([3-cholamidopropyl] dimethylammonio)-1-propanesulfonate (CHAPS), 20 mM 1,4-dithiothreitol (DTT), 0.5% (v/v) IPG buffer (GE Healthcare), 0.002% (w/v) bromophenol blue. Prepare the buffer with all components except for the IPG buffer, divide into 500-mL aliquots and store at −20°C. Just before use, thaw an aliquot and add the desired IPG buffer to a final concentration of 0.5% (v/v). 4. Rehydration buffer for thylakoid membranes: 7 M urea, 2 M thiourea (see Note 5), 15 mM DHPC (1,2-diheptanoyl-snglycero-3-phosphatidyl choline, Avanti Polar-Lipids Inc.;
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catalogue number 850306), 0.5% (v/v) Triton X-100, 20 mM DTT, 0.4% (v/v) IPG buffer (GE Healthcare), 0.002% (w/v) bromophenol blue. Prepare the buffer with all components except for the IPG buffer, divide into 500-mL aliquots and store at −20°C. Just before use, thaw an aliquot and add the desired IPG buffer to a final concentration of 0.5% (v/v).
3. Methods Preparation of different chloroplast fractions for proteomic analysis can usually be performed in a sequential manner from the same plant material. First, intact chloroplasts are isolated from leaf material by homogenization followed by differential centrifugation and the chloroplasts are then broken by osmotic shock. This results in a soluble fraction, the chloroplast stroma, and a second fraction containing the thylakoid membrane. The thylakoid membranes are typically washed six times to remove contaminating chloroplast stroma proteins as well as peripheral membrane proteins. Finally, the soluble thylakoid lumen proteins can be isolated by Yeda press rupture of the isolated thylakoid membranes. It is important to realize that the thylakoid lumen proteins are of quite low abundance in comparison to the stromal and thylakoid proteins so that a large amount of leaf material (typically 60–120 g) is required to acquire usable amounts of thylakoid lumen fraction for proteomic analysis. If the goal is only to purify stroma and thylakoid membrane fractions, a lesser amount of leaf material can be used. It is important that all preparation is performed in a cold room at 4°C and that all buffers, labware, centrifuges, and rotors are properly cooled well in advance of the start of the preparation. A full preparation, starting from leaf harvesting and including isolation of the stroma, thylakoid membrane, and thylakoid lumen fractions typically requires 10–12 h of laboratory work. 3.1. Preparation of Chloroplasts
1. The day before isolation starts all buffers and labware should be prepared and stored at 4°C, including centrifuge tubes and rotors. Note that BSA, sodium ascorbate, and l-cysteine are not added to the preparation medium until just before starting the preparation (see Note 6). 2. Harvest Arabidopsis leaf material from 4 to 8 week old plants (see Note 7). For a full preparation, i.e., if isolation of all fractions (stroma, thylakoid and lumen) is desired, 60–120 g fresh weight of leaves is used. When preparing only stroma or thylakoid proteins, it is possible to use a lesser amount of starting material if this is limiting.
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3. Add to the chloroplast extraction buffer just before starting: BSA to 0.25% (w/v), sodium ascorbate to 4.5 mM, and l-cysteine to 5 mM. Place a large Erlenmeyer flask (2–3 L) on ice and then place a funnel with four layers of nylon cloth (mesh size 20 mM) in the flask. 4. Pour 170 mL of chloroplast extraction buffer and 20 g of leaf material into a homogenizer (see Note 8). Homogenize for 5 × 1 s. 5. Let the homogenate pass through the nylon cloth into the flask that is standing on ice. Shaking the funnel containing the cloth, or prewetting with buffer, may be needed to make the homogenate flow through. Do not squeeze the homogenate through the nylon mesh; instead, patiently wait for it to pass through. Repeat step 4 and this step for each 20 g batch of leaf material and optionally retrieve nonhomogenized material from the nylon cloth and rehomogenize it to increase yield. 6. Divide the homogenate in 40-mL centrifuge tubes on ice (see Note 9). Centrifuge at 1,000 × g, 4°C, for 2 min. With each tube, carefully pour off (and discard) the supernatant and immediately place the pellet on ice. 7. To each tube, add a volume of chloroplast wash buffer equivalent to that used previously of chloroplast extraction buffer (i.e., 170 mL divided by the number of tubes) and carefully resuspend the chloroplast pellet with a soft brush. Centrifuge at 1,000 × g, 4°C, for 2 min. Carefully pour off the supernatant and immediately place the pellet on ice. 8. Resuspend each chloroplast pellet in a small volume of chloroplast wash buffer, approximately 2–3 mL, and thereafter pool all pellets together. 9. Determine the chlorophyll concentration in acetone in three replicates according to (26) (see Note 10). Typically, a 1:500 dilution of the sample in 80% acetone will give suitable absorbance values. 3.2. Isolation of Chloroplast Stroma Proteins
1. Dilute the chloroplast suspension by adding osmotic shock buffer so that the final concentration of chlorophyll (Chl) becomes 0.2 mg Chl/mL. This will rupture the chloroplasts by osmotic shock, releasing the soluble stromal proteins and the thylakoid membranes. 2. Homogenize the suspension once in a 40-mL glass Potter homogenizer using pestle A. Divide the homogenized suspension in 40-mL centrifuge tubes on ice and centrifuge at 7,500 × g, 4°C, for 5 min. 3. Collect the supernatant containing the soluble stroma proteins in Falcon tubes or any other suitable container and store on ice.
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4. If the intention is to isolate thylakoid membrane and/or thylakoid lumen fractions, continue with Subheading 3.3 (Isolation of thylakoid membranes) immediately. Continue with the stroma fraction according to the steps below, either in parallel or after completing the thylakoid membrane/lumen isolations. 5. To remove remaining thylakoid material, centrifuge the stroma protein sample at 100,000 × g, 4°C, for 1 h in an ultracentrifuge. Make sure that you use centrifuge tubes and a rotor with the proper specifications and that the tubes are at least three-quarters full. After centrifugation transfer the supernatant to Falcon tubes or any other suitable container and store on ice. 6. Concentrate the stromal protein sample using Amicon Ultra15 10K or 3K ultrafiltration devices at 4°C to a final volume of 1–5 mL. When working with large volumes of stroma sample, this will require repeated concentrations. 7. Determine the protein concentration using an appropriate assay, such as the Bradford assay with BSA as standard. 8. Aliquot the protein and store at −80°C or use immediately for subsequent experiments. An approximate yield of stroma protein fraction is 1–2 mg protein per 10 g Arabidopsis leaf starting material. The protein composition of a typical stromal fraction prepared by this method is shown in Fig. 3. 3.3. Isolation of Thylakoid Membranes
1. Resuspend each thylakoid membrane pellet obtained in Subheading 3.2, step 4, in 25 mL of osmotic shock buffer using a brush. Homogenize the suspension once in a 40-mL glass Potter homogenizer using pestle A and thereafter centrifuge at a speed of at least 7,500 × g, 4°C, for 5 min. Discard the supernatant. 2. Repeat step 1 once more. 3. Resuspend each thylakoid membrane pellet in 25 mL of thylakoid wash buffer I using a brush. Homogenize the suspension once in a 40-mL glass Potter homogenizer using pestle A and thereafter centrifuge at 7,500 × g, 4°C, for 5 min. Discard the supernatant carefully, as the pellet obtained in this step is quite loose. 4. Repeat step 3 once more. 5. Resuspend each thylakoid membrane pellet in 25 mL of thylakoid wash buffer II using a soft brush. Homogenize the suspension once in a 40-mL glass Potter homogenizer using pestle A and thereafter centrifuge at 7,500 × g, 4°C, for 5 min. Discard the supernatant. 6. Repeat step 5 once more.
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Fig. 3. SDS-PAGE of the obtained chloroplast subfractions compared to the intact chloroplast. As seen, the whole chloroplast (lane A) and stroma fraction (lane B) are dominated by the strong RuBisCo band at ~50 kDa. Meanwhile, the thylakoid fraction (lane C) is dominated by the LHC proteins and the lumen (lane D) by plastocyanin and the extrinsic PSII proteins.
7. Resuspend each thylakoid membrane pellet in a small volume of thylakoid wash buffer II, approximately 0.5 mL per pellet, and then pool all pellets together. 8. Determine the chlorophyll concentration in three replicates according to ref. 26 (see Note 10). Typically, a 1:500 dilution of the sample in 80% acetone will give suitable absorbance values. The chlorophyll concentration should now be approximately 3–4 mg Chl/mL, which is a suitable concentration for preparing the chloroplast lumen fraction according to Subheading 3.4. 9. The thylakoid membrane fraction can now either be used for preparing the thylakoid lumen fraction according to Subheading 3.4 or it can be flash-frozen in liquid nitrogen and stored at −80°C (see Note 11). To freeze the thylakoids, partially submerge a metal sieve in a container filled with liquid nitrogen (for example a kitchen wire mesh sieve in an ice box containing liquid nitrogen) and use a Pasteur pipette to drop the thylakoid suspension one drop at a time into the kitchen sieve partially submerged in liquid nitrogen. Make sure that
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the thylakoid suspension is continuously mixed so that all drops have the same composition and concentration. Collect the flash frozen thylakoid drops from the kitchen sieve and transfer them to 1.5-mL test tubes (4–5 drops per tube) and immediately store at −80°C. The protein composition of a typical thylakoid membrane fraction prepared by this method is shown in Fig. 3. 3.4. Isolation of Thylakoid Lumen Proteins
1. Prepare thylakoid membranes at 3–4 mg Chl/mL according to Subheading 3.3. If the chlorophyll concentration is lower than this, the thylakoid suspension can be recentrifuged in the same way as previously (see Subheading 3.3, step 5) and then resuspended in a smaller volume of thylakoid wash buffer II. 2. Keep the thylakoid suspension on ice for 15–60 min. 3. Place the thylakoid suspension in a precooled Yeda press chamber, ensure the sample collection valve is closed, and then fit the chamber to a flask of nitrogen gas. Wearing eye protection, apply 100 bar pressure to the thylakoids. Turn of the gas flow and then very carefully open the sample collection valve while holding a 50-mL Falcon tube at the outlet. Only open it so much that the thylakoid suspension slowly seeps out from the pressure chamber. Opening it too fast will result in vast overpressure and spraying the room with thylakoid material. 4. Centrifuge the fragmented thylakoid membranes at 200,000 × g, 4°C, for 1 h in an ultracentrifuge. Transfer the supernatant to a new centrifuge tube and centrifuge again at 200,000 × g, 4°C, for 1 h. Carefully transfer the supernatant to a 15-mL Falcon tube and store on ice. The pellet of ruptured thylakoid membranes may if desired be resuspended in thylakoid wash buffer II and flash-frozen in liquid nitrogen as described in Subheading 3.3, step 9. 5. Determine the protein concentration using an appropriate assay, such as the Bradford assay with BSA as standard. Typical concentrations following the procedure above are 0.2–0.4 mg protein/mL and an approximate yield of lumen protein fraction is 2–4 mg protein per 100 g Arabidopsis leaf starting material. 6. Aliquot the protein and store at −80°C or use immediately for subsequent experiments. The protein composition of a typical thylakoid lumen fraction prepared by this method is shown in Fig. 3.
3.5. Preparation of Protein Samples for 2D Gel Electrophoresis
1. Thaw an appropriate amount of chloroplast stroma, thylakoid membrane, or thylakoid lumen sample for 2D gel electrophoresis (see Note 12). Add four volumes of ice-cold 100% acetone to one volume of sample (depending on the sample
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volume, it may be necessary to divide the sample in several 1.5-mL Eppendorf tubes). Mix the tubes and thereafter store the tubes for at least 2 h at −20°C. 2. Centrifuge the precipitated proteins for 10 min at maximum speed in a tabletop microcentrifuge. 3. Discard the supernatant and thereafter wash the protein pellets with ice-cold 100% acetone, using the same volume as the total volume in step 1. Do not resuspend the pellet, but rather rinse it a few times by inverting the tube or by gently pipetting. Centrifuge again for 10 min at maximum speed, remove the supernatant, and then repeat this step twice for a total of three washes. 4. Allow the pellets to air-dry so that any residual acetone is evaporated. Do this by simply letting the tubes stand open on the laboratory bench for 5–10 min. Make sure that all the acetone has evaporated. 5. Resuspend the chloroplast stroma or thylakoid lumen protein pellet in an appropriate volume of rehydration buffer for soluble proteins by pipetting (for example 450 mL of rehydration buffer for one 24 cm IPG strip from GE Healthcare). For thylakoid membranes, resuspend the pellet in an appropriate volume of rehydration buffer for thylakoid membranes. 6. Allow the sample to solubilize on the bench for 30–60 min and thereafter centrifuge the sample at maximum speed in a tabletop microcentrifuge at room temperature for 10 min (see Note 13). Transfer the supernatant to a fresh tube. 7. The sample is now ready for your preferred IEF gradient for the first dimension of a 2D-PAGE separation. The protein composition of typical thylakoid membrane, stroma, and lumen fractions prepared by these methods and subsequently separated by 2D-PAGE are shown in Fig. 4.
4. Notes 1. BSA is added to absorb fatty acids that are liberated during homogenization of the sample, which have been shown to inhibit for instance PSII activity. Ascorbate and L-cysteine are added to prevent oxidation of proteins. 2. All buffers and equipment needed for extraction are preferably prepared 1 day in advance and stored in the cold room where the whole extraction will be performed. 3. The nylon mesh is a critical material for the purity of the extraction. The mesh can be obtained from several suppliers
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Fig. 4. Typical 2D-PAGE (pH range 3–11) results using fractions prepared with the protocols described here. 2D-PAGE of the stroma (a), thylakoid membrane (b), and thylakoid lumen (c) fractions, showing that they all have very distinct and different protein “fingerprint” patterns.
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(e.g., Millipore) and with several pore sizes. A pore size of 20 mM will prevent unbroken cells, nuclei, and other larger fragments from passing through, but will let the 1–10 mM sized chloroplasts to pass. The nylon mesh cannot be replaced by cotton wool or tape. The mesh should also be handled with care to ensure that no holes are punched into it, leading to contamination. 4. LINCA Lamon Instrumentation Co., Ltd., Tel-Aviv, Israel has produced Yeda press equipment for many years and has provided most of the equipment found in laboratories around the world. However, it has recently come to our knowledge that this company has closed down. The authors are now currently investigating whether the production of new instruments could be undertaken in Sweden. Please feel free to contact us regarding this matter. 5. Thiourea is a toxic compound and requires special permission to handle in some countries. 6. The volume and temperature of the added components can be neglected, as the final volume is so large. 7. Because the preparation takes several hours, it is recommended to start early in the morning, just at the end of a dark period. Leaves can either be harvested directly before starting the preparation or the evening before. If harvesting is done the evening before, the leaves can be stored in sealed plastic bags, in darkness at 4°C overnight. 8. When selecting the type of homogenizer, the most important parameter is the sharpness of the blades. An unsharp homogenizer will tear apart chloroplast and thylakoid membranes, thereby decreasing yield. A good industrial type homogenizer is recommended. 9. Optionally, larger tubes can be used at this initial stage to save time and tube handling; for example, 250 mL tubes can be used when performing a large preparation. 10. Into an Eppendorf tube containing 995 mL of 80% ice-cold (−20°C) acetone, pipette 5 mL of sample and incubate the sample on ice for 15 min. To obtain correct measurements, ensure that sample adhering to the outside of the pipette tip is wiped off against the tube or against a tissue prior to dilution of the sample in 80% acetone. Centrifuge the sample for 15 min at maximum speed in a tabletop microcentrifuge to pellet proteins. The absorbance (A) of the supernatant, containing the extracted pigments, is measured at 646.6 and 663.6 nm. According to Porra et al. (26), the total chlorophyll concentration is calculated using the formula Chl a + b = 17.76 A646.6 + 7.34 A663.6 multiplied by the dilution factor.
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11. Before freezing the thylakoid membranes, it can be practical to adjust the chlorophyll concentration to 1.0 mg Chl/mL with thylakoid wash buffer II, simply because this is a handy working concentration for many subsequent experiments, but the thylakoids can also be frozen at higher concentrations such as 3–4 mg Chl/mL. It is also important to note that frozen thylakoids cannot be used for preparation of the thylakoid lumen fraction; this preparation must be performed using fresh thylakoid material. 12. The amount of sample required will depend on the number and type of 2D gels which are to be analyzed. Using 24 × 20 × 0.1 cm gels, suitable amounts of chloroplast stroma fraction are in the range of 400–600 mg protein per gel, while for the thylakoid lumen fraction 50–80 mg protein per gel gives high resolution. For thylakoid membrane samples, it is convenient to approximate the protein concentration by multiplying the chlorophyll concentration by eight at this stage, and later making an exact protein quantification using a 2D Quant kit (GE Healthcare, Uppsala, Sweden). Typically, 400–600 mg protein (50–75 mg chlorophyll) per 2D gel is suitable for thylakoid membrane samples using the protocol provided here. 13. Thylakoid membranes are usually more difficult to solubilize than soluble protein samples, and it is, therefore, a good idea to gently pipette the sample up and down a few times every 15 min or so.
Acknowledgments Thomas Kieselbach for the photo shown in Fig. 2, suggestions and comments on this chapter, the Lawski foundation for postdoctoral support to Y.M., the Lars Hiertas Memorial Fund for financial support to M.H., and the Swedish Research Council for financial support to W.P.S. References 1. Richly, E., and Leister, D. (2004) An improved prediction of chloroplast proteins reveals diversities and commonalities in the chloroplast proteomes of Arabidopsis and rice. Gene 329, 11–16. 2. Wilson, K. A., McManus, M. T., Gordon, M. E., and Jordan, T. W. (2002) The proteomics of senescence in leaves of white clover, Trifolium repens (L). Proteomics 2, 1114–1122. 3. Heazlewood, J. A., Verboom, R. E., TontiFilippini, J., Small, A., and Millar, A. H. (2007)
SUBA: the Arabidopsis Subcellular Database. Nucleic Acids Res. 35, 213–218. 4. Aronsson, H., and Jarvis, P. (2002) A simple method for isolating import-competent Arabidopsis chloroplasts. FEBS Lett. 529, 215–220. 5. Schulz, A., Knoetzel, J., Scheller, H. V., and Mant, A. (2004) Uptake of a fluorescent dye as a swift and simple indicator of organelle intactness: import-competent chloroplasts from soil-grown Arabidopsis. J. Histochem. Cytochem. 52, 701–704.
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6. Kieselbach, T., Hagman, Å., Andersson, B., and Schröder, W. P. (1998) The thylakoid lumen of chloroplasts: isolation and characterization. J. Biol. Chem. 273, 6710–6716. 7. Peltier, J.B., Cai, Y., Sun, Q., Zabrouskov, V., Giacomelli, L., Rudella, A., Ytterberg, A. J., Rutschow, H., and van Wijk, K. J. (2006) The oligomeric stromal proteome of A. thaliana chloroplasts. Mol. Cell. Proteomics 5, 114–133. 8. Goulas, E., Schubert, M., Kieselbach, T., Kleczkowski, L. A., Gardeström, P., Schröder, W. P., and Hurry, V. (2006) The chloroplast lumen and stromal proteomes of Arabidopsis thaliana show differential sensitivity to shortand long-term exposure to low temperature. Plant J. 47, 720–734. 9. Wu, C., and Yates, J. R. (2003) The application of mass spectrometry to membrane proteomics. Nat. Biotech. 21, 262–267. 10. Ort, D. R., and Yocum, C. F. (1996) Oxygenic Photosynthesis: The Light Reactions. Kluwer Academic Publishers, Dordrecht, The Netherlands. 11. Rochaix, J. D., Goldschmidt-Clermont, M., and Merchant, S. (1998) The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas. Advances in Photosynthesis and Respiration, Vol. 7. Kluwer Academic Publishers, Dordrecht, The Netherlands. 12. Wollman, F. A., Minai, L., and Nechushtai, R. (1999) The biogenesis and assembly of photosynthetic proteins in thylakoid membranes. Biochim. Biophys. Acta 1411, 21–85. 13. Hauser, H. (2000) Short-chain phospholipids as detergents. Biochim. Biophys. Acta 1508, 164–181. 14. Mishra, Y., Hall, M., Kieselbach, T., Jansson, S., and Schröder W. P. (2010) Dissection of the real developmental acclimation strategies of Arabidopsis thaliana by comparing indoor and outdoor plants: a comparative sub cellular proteomics study (manuscript in preparation). 15. Kieselbach, T., Bystedt, M., Hynds, P., Robinson C., and Schröder, W. P. (2000) A peroxidase homologue and a novel plastocyanin located by proteomics to the Arabidopsis chloroplast thylakoid lumen. FEBS Lett. 480, 271–276. 16. Peltier, J. B., Friso, G., Kalume, D. E., Roepstorff, P., Nilsson, F., Adamska, I., and van Wijk, K. J. (2000). Proteomics of the chloroplast: systematic identification and targeting analysis of lumenal and peripheral thylakoid proteins. Plant Cell 12, 319–341. 17. Peltier, J. B., Emanuelsson, O., Kalume, D. E., Ytterberg, J., Friso, G., Rudella, A., Liberles, D. A., Soderberg, L., Roepstorff, P., von Heijne, G., and van Wijk, K. J. (2002) Central
functions of the lumenal and peripheral thylakoid proteome of Arabidopsis determined by experimentation and genome-wide prediction. Plant Cell 14, 211–236. 18. Schubert, M., Petersson, U. A., Haas, B. J., Funk, C., Schröder, W. P., and Kieselbach, T. (2002) Proteome map of the chloroplast lumen of Arabidopsis thaliana. J. Biol. Chem. 277, 8354–8365. 19. Hall, M., Mata-Cabana, A., Åkerlund, H-E., Florencio, F. J., Schröder, W. P., Lindahl, M., and Kieselbach, T. (2010) Thioredoxin targets of the plant chloroplast lumen and their implications for plastid function. Proteomics 10, 987–1001. 20. Bally, J., Paget, E., Droux, M., Job, C., Job, D., and Dubald, M. (2008) Both the stroma and thylakoid lumen of tobacco chloroplasts are competent for the formation of disulphide bonds in recombinant proteins. Plant Biotechnol. J. 6, 46–61. 21. Edvardsson, A., Shapiguzov, A., Petersson, U. A., Schröder, W. P., and Vener, A. V. (2007) Immunophilin AtFKBP13 sustains all peptidylprolyl isomerase activity in the thylakoid lumen from Arabidopsis thaliana deficient in AtCYP202. Biochemistry 46, 9432–9442. 22. Edvardsson, A., Eshaghi, S., Vener, A. V., and Andersson, B. (2003) The major peptidyl- prolyl isomerase activity in thylakoid lumen of plant chloroplasts belongs to a novel cyclophilin TLP20. FEBS Lett. 542, 137–141. 23. Ingelsson, B., Shapiguzov, A., Kieselbach, T., and Vener, A. V. (2009) Peptidyl-prolyl isomerise activity in chloroplast thylakoid lumen is a dispensable function of immunophilins in Arabidopsis thaliana. Plant Cell Physiol. 50, 1801–1814. 24. Romano, P. G., Edvardsson, A., Ruban, A. V., Andersson, B., Vener, A. V, Gray J. E., and Horton P. (2004) Arabidopsis AtCYP20-2 is a light-regulated cyclophilin-type peptidyl- prolylcis-trans isomerase associated with the photosynthetic membranes. Plant Physiol. 134, 1244–1247. 25. Shapiguzov, A., Edvardsson, A., and Vener, A. V. (2006) Profound redox sensitivity of peptidyl-prolyl isomerase activity in Arabidopsis thylakoid lumen. FEBS Lett. 580, 3671–3676. 26. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Determination of accurate extinction coefficients and simultaneous equations for assaying chlorophylls a and b extracted with four different solvents: verification of the concentration of chlorophyll standards by atomic absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394.
Chapter 12 Preparation of Plastoglobules from Arabidopsis Plastids for Proteomic Analysis and Other Studies* Celine Besagni, Lucia Eugeni Piller, and Claire Bréhélin Abstract Plastoglobules are particles specifically located inside different types of plastids. They mainly contain lipids and proteins and are physically attached to thylakoids. Proteomic studies have underlined the role of plastoglobules in diverse plastid metabolic pathways, such as those producing vitamin K, vitamin E, and carotenoids, and have implicated them in plant response to stress. This chapter describes the isolation of pure and intact plastoglobules from Arabidopsis leaves. The procedure starts with the isolation of intact chloroplasts by centrifugation on a Percoll gradient. Plastoglobules are then separated from the plastid membranes by flotation on a sucrose gradient. Finally, the purity of the plastoglobule fraction is verified by immunoblotting. Key words: Plastoglobules, Sucrose gradient, Chloroplast preparation, Arabidopsis, Immunoblot, Percoll gradient, Flotation, Protein precipitation, Plastid membranes
1. Introduction Plastoglobules are lipoprotein particles present in different plastid types such as proplastids, chloroplasts, chromoplasts, and gerontoplasts. Plastoglobules are mainly composed of isoprenoids, neutral lipids, and proteins (1). They are physically attached to thylakoids via the outer half of the lipid bilayer of the thylakoid, which surrounds plastoglobules (2). Long viewed as passive lipid droplets, plastoglobules have been receiving increasing attention in the last half decade. Indeed, proteomic studies of plastoglobules (3, 4) have demonstrated their active role in plastid biology. This idea is reinforced by the fact that plastoglobule size and number vary depending on plastid type and environmental conditions. Currently, a growing body of evidence suggests that plastoglobules are *Celine Besagni and Lucia Eugeni Piller have contributed equally to this chapter. R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_12, © Springer Science+Business Media, LLC 2011
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involved in various metabolic pathways such as those leading to vitamin K (5), tocopherol (3, 6), and carotenoids (4), and in plant response to stress by accumulating antioxidants (e.g., tocopherols) and sequestrating toxic molecules (e.g., fatty acid phytyl esters) (7). In addition, studies of the plastid ultrastructure suggested that plastoglobule size and number are regulated in correlation with the fitness of the thylakoid membranes, putatively playing the role of a reservoir for thylakoid membrane lipids (1, 8). Besides their physiological roles, plastoglobules may present a biotechnological interest for molecular farming by providing a hydrophobic environment necessary for the production of certain proteins, associated with a simple purification procedure (3). The identification of components (proteins or lipids) specifically localized in plastoglobules relies on preparation of a plastoglobule fraction exempt of any contamination. Such contamination could originate either from light particles present in other compartments of the cell (e.g., oil bodies) or from plastid membranes. Thus, the preparation of pure plastoglobules depends on two critical steps: (1) isolation of intact plastids on Percoll gradients and (2) accurate separation of plastid membranes by flotation on sucrose gradients. The range of sucrose concentrations (5–45%) that is used for the second step allows the separation of plastoglobules from envelope membranes, which is generally not achieved with standard protocols designed for plastid membrane preparation. In addition, because plastoglobules are physically attached to thylakoids, the procedure must include a step to achieve a good separation of plastoglobules from thylakoids. Finally, the purity of the plastoglobule fraction is verified by immunoblotting, and plastoglobules are then available for proteomic analysis or other studies.
2. Materials 2.1. Arabidopsis Culture
1. Universal soil (Ricoter, Switzerland). 2. Culture trays (30 × 50 cm), with propagator lids. 3. Solbac biological control agent (Andermatt Biocontrol, Switzerland); dilute 1/400 (v/v) in tap water. 4. Phytotron: Percival AR-66L (CLF Plant Climatics GmbH, Germany) or similar, providing the following growth conditions: 8 h of light per day, 120 mmol photons/m2/s, 20°C, 60% humidity.
2.2. Preparation of Intact Chloroplasts and Purification of Plastoglobules
1. HB buffer: 450 mM sorbitol, 20 mM Tricine–KOH, pH 8.4, 10 mM ethylenediaminetetraacetic acid (EDTA) (see Note 1), 10 mM NaHCO3, 1 mM MnCl2, 5 mM Na-ascorbate, 0.05% (w/v) bovine serum albumin (BSA) fraction V, and 1 mM
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phenylmethylsulfonyl fluoride (PMSF) (see Notes 2 and 3). Prepare freshly (see Note 4). 2. RB buffer (8×) stock: 2.4 M sorbitol, 160 mM Tricine–KOH, pH 7.6, 20 mM EDTA (see Note 1), and 40 mM MgCl2. Working buffer is prepared by diluting 100 mL of RB buffer (8×) in 700 mL of water. Store at −20°C (see Note 5). 3. 40% (v/v) and 85% (v/v) Percoll solutions: mix 40 or 85 mL of Percoll (pH 8.5–9.5; Sigma, MO, USA) with 12.5 mL of RB buffer (8×) and adjust volume to 100 mL with deionized water. These solutions can be kept for several months at −20°C. 4. 80% (v/v) acetone diluted in water. This can be stored for several weeks at room temperature (see Note 6). 5. TrE buffer: prepare 10× stock solution with 50 mM Tricine– KOH, pH 7.5, 2 mM EDTA, and 2 mM dithiothreitol (DTT) (see Note 7). For use, 100 mL of TrE (10×) is diluted in 900 mL of water. Buffer can be stored at −20°C for several months. 6. Sucrose (0.6 M) dissolved in TrE buffer (1×). The solution can be stored at −20°C for several months. 7. Protease inhibitor cocktail (e.g., for Plant Cell and Tissue Extracts, Sigma). 8. 45, 38, 20, 15, and 5% (w/v) sucrose solutions: sucrose is dissolved with TrE buffer (10×) and autoclaved deionized water to obtain desired sucrose concentration in TrE (1×). The solutions can be stored at −20°C for several months. 9. Centrifuges: superspeed refrigerated centrifuge (e.g., Sorvall RC-5B, Thermo Scientific, MA, USA), with fixed-angle rotor (e.g., Sorvall SLA1500) and corresponding plastic 250 mL bottles (Nalgene, NY, USA), or with swinging-bucket rotor (e.g., Sorvall HB-6) and corresponding open polycarbonate 50 mL tubes (Nalgene); refrigerated bench-top centrifuge (e.g., Eppendorf 5810R), with swinging-bucket rotor (e.g., Eppendorf A4-62) and capped polypropylene 50 mL tubes (Falcon, BD biosciences, CA, USA); ultracentrifuge (e.g., Beckman L7, Beckman Coulter Inc., CA, USA), with swinging-bucket rotor (e.g., Beckman SW 28) and UltraClearTM SW28 tubes (25 × 89 mm, Beckman). 10. Waring blender homogenizer. 11. Miracloth and cheesecloth. 12. 15- and 50-mL Potter-Elvehjem tissue grinders with Teflon pestles. 13. Spectrophotometer.
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2.3. Protein Precipitation and SDSPolyacrylamide Gel Electrophoresis
1. Sample buffer (SB 4×) stock: 200 mM Tris–HCl, pH 6.8, 400 mM DTT, 8% (w/v) sodium dodecyl sulfate (SDS), 0.4% (w/v) bromophenol-blue, and 50% (w/v) glycerol. 100 mL of SB (4×) should be diluted in 300 mL of water and then used to resuspend dried proteins. 2. Stock solutions for running gel: 40% (w/v) acryl/bisacrylamide (37.5/1); 2 M Tris–HCl, pH 8.8; 20% (w/v) SDS; N,N,N¢,N¢-tetramethylethylenediamine (TEMED); 10% (w/v) ammonium persulfate (APS) prepared in water and stored frozen in single-use aliquots (200 mL) at −20°C (see Note 8). 3. Stock solutions for stacking gel: 40% (w/v) acryl/bisacrylamide (37.5/1); 0.5 M Tris–HCl, pH 6.8; 20% (w/v) SDS; TEMED; and 10% (w/v) APS. 4. Running buffer (5× stock solution): 100 mM Tris, 1 M glycine, pH 8.3, and 0.5% (w/v) SDS. Store at room temperature. For use, dilute 100 mL of 5× stock with 900 mL of water. 5. SDS-PAGE Molecular Weight Standards, Broad Range (Bio-Rad, CA, USA), or similar. 6. Microcentrifuge. 7. Speed Vacuum Concentrator 5301 (Eppendorf, Germany), or similar. 8. Electrophoresis system for SDS-PAGE (e.g., PerfectBlue Dual Gel System Twin ExWS with glass plates (20 × 10 cm), a 20-teeth comb, and two 0.8-mm spacers; PeqLab Biotechnologies GmbH, Germany). 9. Power supply (e.g., Power Pac 300, Bio-Rad).
2.4. Verification of Plastid Membrane Fractions Purity by Immunoblotting
1. Nitrocellulose membrane (e.g., Protran R, 0.45 mm pore size, Whatman, UK). 2. Blotting paper (e.g., 3MM, Whatman). 3. Transfer tank (e.g., Trans-Blot TM Cell, Bio-Rad). 4. Rocking platform (e.g., Stuart SSM3 3D gyratory rocker, Bibby Scientific Limited, UK). 5. Transfer buffer: 15 mM NaH2PO4, 0.05% (v/v) SDS, and 20% (v/v) ethanol. Prepare freshly. 6. AmidoBlack staining solution: 45% (v/v) ethanol, 10% (v/v) glacial acetic acid, and 0.1% (w/v) AmidoBlack 10B (Merck, NJ, USA). Store at room temperature (see Note 9). 7. Destaining solution: 40% (v/v) ethanol, 10% (v/v) acetic acid. Store at room temperature. 8. PBS buffer (10×) stock: 1.4 M NaCl, 27 mM KCl, 0.1 M Na2HPO4, and 20 mM KH2PO4. Dilute 100 mL with 900 mL of water to obtain 1× PBS for use.
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9. TBS buffer (10×) stock: 10 mM Tris–HCl, 0.15 M NaCl, pH 7.5. Dilute 100 mL with 900 mL of water to obtain 1× TBS for use. 10. Blocking buffer: 5% (w/v) nonfat dry milk in 1× PBS or 1× TBS. 11. Primary antibodies: (1) the anti-AtPGL35 serum (Agrisera, Sweden), specific to plastoglobules, has been raised against the Arabidopsis thaliana plastoglobulin AtPGL35. Use it diluted 1:3,000 in PBS with 5% (w/v) nonfat dry milk; (2) the antiLHCB2 serum (Agrisera) is a marker of thylakoid membranes and recognizes chlorophyll a/b-binding proteins of the light harvesting antenna complex II (LHCII). Use it diluted 1:3,000 in PBS, 5% (w/v) nonfat dry milk; (3) the anti-Toc75 serum (Agrisera) is specific to the plastid envelope membrane and has been raised against the 75 kDa component of the translocon at the outer envelope membrane of chloroplasts (TOC). Use it diluted 1:3,000 in TBS, 5% (w/v) nonfat dry milk. Sera should be stored at −20°C; however, an aliquot of around 50 mL can be kept at 4°C for daily use. 12. Secondary antibody: horseradish peroxidase-coupled goat antirabbit IgG (Bio-Rad) diluted 1:3,000 in PBS or TBS with 5% (w/v) nonfat dry milk. 13. Enhanced chemiluminescent (ECL) reagent: 0.1 M Tris–HCl, pH 8.5, 0.2 mM p-coumaric acid (Fluka, Switzerland), 1.25 mM 3-aminophthalhydrazide (Luminol, Fluka) (see Note 10). 14. 3% (w/w) H2O2: the H2O2 stock is diluted in water and the solution is stored at 4°C for several weeks. 15. Chemiluminescent detection in a dark room: Hyperfilm ECL 18 × 24 cm X-ray film (GE Healthcare, NJ, USA) placed in an X-ray film cassette (e.g., Rego Gollwitzer GmbH, Germany) and developed in a tabletop processor (e.g., SRX-101A, Konica Minolta, Japan). Alternatively, the luminescent signal may be monitored using a luminescence imaging device (e.g., ChemiDock system and the QuantityOne software, both from Bio-Rad). 16. Stripping buffer: 62.5 mM Tris–HCl, pH 6.8, 2% (w/v) SDS, 100 mM b-mercaptoethanol (see Note 11).
3. Methods 3.1. Arabidopsis Culture
1. Sow Arabidopsis thaliana seeds on six trays filled with universal soil (see Note 12). The soil is first watered with Solbac solution to prevent the growth of insect larvae. Trays are covered with transparent plastic propagator lids (or cling film) to ensure appropriate humidity for germination.
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2. After 2 days at 4°C, place the trays in a phytotron for 6–8 weeks. Lids are removed after 2 weeks to avoid algae and/or fungi development. 3.2. Preparation of Intact Chloroplasts and Purification of Plastoglobules
We describe here the procedure we routinely follow to obtain pure Arabidopsis plastoglobules. Each step of the protocol below should be performed at 4°C to preserve the integrity of the chloroplasts, and to avoid protein degradation. When the starting material, and consequently, the amount of isolated chloroplasts, are limiting, smaller sucrose gradients can be prepared in appropriate tubes by proportionally reducing the volume of each sucrose solutions, as exemplified in Vidi et al. (9). 1. Prior to harvesting, place plants in the dark for 24–48 h to avoid starch accumulation (see Note 13). 2. Harvest Arabidopsis leaves with scissors or a scalpel, weigh them in a beaker (see Note 14), and then maintain them in chilled water for 30 min (see Note 15). 3. Prepare six Percoll gradients as follows. Start by pouring 15 mL of 40% Percoll solution into a 50-mL open tube. Then, with a glass Pasteur pipette, carefully introduce 5 mL of 85% Percoll solution below the 40% Percoll layer. Keep the gradients at 4°C. 4. Using a Waring blender homogenizer, grind the leaves three times in 500 mL of cold HB buffer (5 s at high strength and then two times 3 s at low strength) (see Note 16). 5. Filter the homogenate immediately through two layers of cheesecloth and one layer of Miracloth placed in a funnel on top of an Erlenmeyer flask. Gently squeeze the homogenate inside the cheesecloth to extract most of the liquid. 6. Divide the filtrate between three or four 250-mL bottles and then centrifuge for 10 min at 1,075 × g in an SLA1500 fixedangle rotor. 7. Gently resuspend each pellet with 1–2 mL of 1× RB buffer (see Note 17) and pool the crude chloroplast extracts in a 50-mL Falcon tube. If necessary, add additional RB buffer to resuspend any residual pellet material, and at the end, rinse the tubes with small amounts of additional RB. Gently mix the suspension by inverting the tube and then load 2–3 mL of crude chloroplast extract onto each Percoll gradient. 8. Centrifuge the gradients for 10 min at 13,600 × g in an HB-6 swinging-bucket rotor (see Note 18). At the end of the centrifugation, intact chloroplasts are located at the interface between the 85 and 40% Percoll phases. The green ring situated in the upper part of the gradient corresponds to broken chloroplasts and should be eliminated.
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9. Aspirate most of the 40% Percoll layer with a vacuum aspirator, delicately collect the band of intact chloroplasts with a plastic pipette (with large opening), and transfer to a 50-mL Falcon tube. 10. Distribute chloroplasts from all gradients among two 50-mL Falcon tubes and dilute with 10 volumes of RB buffer (1×). Gently invert the tube to mix. 11. Centrifuge the suspension 2 min at 2,600 × g in a swingingbucket rotor (A4-62). 12. Carefully decant the supernatant as the pellet is very loose. 13. Resuspend the pellet in 5 mL of TrE buffer (1×) and quantify the chlorophyll (see Note 19). 14. Adjust the sample volume to 50 mL with TrE buffer (1×) to wash the chloroplasts, and then centrifuge for 10 min at 2,600 × g in a swinging-bucket rotor (A4-62). 15. Carefully decant the supernatant and then resuspend (see Note 20) the chloroplast pellet with 0.6 M sucrose/TrE to a concentration of 2–3 mg/mL of chlorophyll. Add a cocktail of antiproteases to preserve protein integrity. 16. Incubate on ice for 10 min and then freeze at −80°C for at least 1 h (see Note 21). 17. Thaw the chloroplast suspension and dilute with 2 volumes of TrE buffer (1×). 18. Homogenize chloroplast suspension for at least 20 strokes in a 50-mL Potter homogenizer and then transfer the homogenate into UltraClear SW28 tubes (see Note 22). 19. Carefully balance the tubes and then perform ultracentrifugation at 100,000 × g in a swinging-bucket rotor for 1 h (see Note 23). 20. Remove the supernatant, which corresponds to the stroma, and resuspend the pellet of total membrane (which contains plastoglobules) into 45% sucrose/TrE to reach a concentration of 2–6 mg of chlorophyll per mL (typically, ~10–20 mL 45% sucrose/TrE solution will be required) (see Notes 24 and 25). 21. Homogenize the resuspended membranes by 20 strokes in a 15-mL Potter homogenizer (see Note 26). 22. Pour 5 mL aliquots of the membranes homogenate into the required number of UltraClear SW28 tubes, and carefully overlay each aliquot with the sucrose/TrE solutions in the following order: 6 mL of 38% sucrose, 6 mL of 20% sucrose, 4 mL of 15% sucrose, and finally 8 mL of 5% sucrose to the top of the tube (see Note 27).
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Fig. 1. Separation of total membranes from isolated Arabidopsis chloroplasts by flotation on a discontinuous sucrose gradient. Plastoglobules (PG) are visible as a yellowish layer at the top of the gradient. Outer and inner envelope membranes (OM/IM) are yellow; thylakoid membranes (Thyl) are green.
23. Carefully balance the tubes and ultracentrifuge them overnight at 100,000 × g (see Notes 28 and 29). An example of the gradient that is obtained after the overnight centrifugation is given in Fig. 1. 24. For each gradient, collect 1-mL fractions with a micropipette, starting from the top of the gradient (fraction 1) and ending at the bottom (approximately 32 fractions), and store them at −20°C. An illustration of the content of plastoglobules, envelopes and thylakoid membrane fractions observed with a confocal microscope is given Fig. 2. Typically, plastoglobules are contained in fractions 1–6, envelopes in fractions 14–18, and thylakoid membrane in fractions 25–32. However, the exact plastid membrane distribution varies from one experiment to another and has to be checked by immunoblotting as described below. 3.3. Protein Precipitation and SDS-PAGE
Here, we describe the procedure that we routinely follow to verify the plastid membrane distribution among the sucrose gradient fractions, and the purity of the plastoglobule fraction before subsequent analysis. However, the procedure needs to be modified depending on the goal of the experiment. For example, for mass spectrometry (MS) analysis, plastid membranes containing 25–30 mg of chlorophyll are loaded on one sucrose gradient and total proteins contained in 1 mL of the plastoglobule fraction are precipitated and separated on SDS-PAGE. Diverse staining methods (e.g., colloidal blue or silver staining) can then be used to visualize proteins before MS analysis.
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Fig. 2. Observation of the different plastid membrane fractions by confocal microscopy. Chloroplast membranes from Arabidopsis leaves expressing a plastoglobule marker (AtPGL35) fused to the green fluorescent protein (GFP) under the control of the constitutive 35S promoter were separated by flotation on a sucrose gradient. Fluorescence of fractions F1 (plastoglobules), F21 (envelopes), F33 (thylakoids), and of intact chloroplasts (pellet) is visualized by confocal laser scanning microscopy. GFP and chlorophyll autofluorescence were monitored with a Leica TCS SP5 microscope (Leica Microsystems, Germany) using the 488 nm laser line and detection windows of 490–540 and 650–800 nm, respectively. Scale bars: 5 mm.
3.3.1. P rotein Precipitation
1. Different volumes of each even (or odd) fraction are transferred to microtubes for protein precipitation: 400 mL from each of fractions 1–18 (split between two tubes of 200 mL for each fraction), and 200 mL of the upper fractions, stroma (St) and intact chloroplasts (see Note 30).
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2. To 200 mL of sample, add 160 mL of chloroform and 480 mL of methanol. 3. Vortex the tubes for 1 min and then add 640 mL of deionized H2O. 4. Centrifuge the tubes for 1 min at maximum speed in a microcentrifuge. 5. Remove the upper phase, taking care not to disturb the protein band at the interphase (see Note 31). 6. Add 480 mL of methanol to the lower phase and vortex. 7. Centrifuge for 5 min at maximum speed in a microcentrifuge, remove the supernatant, and dry the protein pellet with a speed vacuum concentrator (around 10 min at 30°C) (see Note 32). 8. Resuspend the proteins in SB buffer: 20 mL each for fractions 1–18 (10 mL for each tube of each fraction, which are then pooled together), 15 mL each for fractions 19–24, 30 mL each for fractions 25–28 and stroma, and 60 mL for total chloroplasts and for each of the remaining fractions (see Note 33). 9. Heat the protein samples for 10 min at 65°C. Proteins are now ready to be loaded on an SDS-PAGE gel. 3.3.2. Separation of Proteins by SDS-PAGE
The following steps describe the preparation and the use of a 12% SDS-PAGE gel using the PerfectBlue Dual Gel Twin ExWS electrophoresis system (PeqLab). 1. Scrupulously clean with deionized water and technical ethanol the glass plates (20 × 10 cm) before assembling the electrophoresis system. 2. For one 12% acrylamide running gel, mix in a 50-mL Falcon tube: 5 mL of deionized water, 3 mL of 40% acryl/bisacrylamide (see Note 34), 2 mL of 2 M Tris–HCl, pH 8.8, and 50 mL of 20% SDS. When everything is ready, add 5 mL of TEMED and 80 mL of 10% APS (see Note 35) to the running gel solution, rapidly mix, and pour the gel leaving a space for the stacking gel (see Note 36). Overlay with isopropanol. The gel should polymerize in less than 30 min at room temperature. 3. Remove the isopropanol with absorbing paper and prepare stacking gel by mixing in a 50 mL Falcon tube: 3.9 mL of water, 500 mL of 40% acry/bisacrylamide, 600 mL of 0.5 M Tris–HCl, pH 6.8, and 25 mL of 20% of SDS. Finish by adding 5 mL of TEMED and 40 mL of 10% APS. Mix and pour the stacking gel on the running gel until reaching the top of the plate and carefully insert the comb. The stacking gel should polymerize within 20 min at room temperature. 4. Once polymerized, assemble the gel into the electrophoresis system and add the running buffer to the upper and lower
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chambers. Carefully remove the comb and wash the wells by expelling running buffer into the wells with a micropipette. 5. Load different volumes of each sample in the wells: 20 mL for fractions 1–18, and 15 mL for the remaining samples. One well is loaded with 5 mL of the molecular weight marker. 6. Complete the assembly of the electrophoresis system and connect to a power supply. The gel can be run at 15 mA until the front migration penetrates the running gel, and then at 30 mA until the blue dye front reaches the edge of the glass plates (around 45 min). 3.4. Verification of Plastoglobule Purity by Immunoblotting
1. Open a blotting transfer cassette in a tray filled with transfer buffer. Cut two pieces of 3MM paper and a sheet of nitrocellulose membrane at the dimensions of the cassette. Soak the 3MM papers, the nitrocellulose, and two sponges with transfer buffer. 2. Disassemble the SDS-PAGE electrophoresis system and remove the stacking part of the gel. Cut a small piece of the running gel at one corner to allow future orientation of the gel. Soak the running gel in transfer buffer for 2–3 s. 3. Assemble the blotting “sandwich” in the cassette with soaked elements as follows. On the cathode side, avoiding any air bubble imprisonment, lay out the following: a sponge, a sheet of 3MM paper, the running gel, the nitrocellulose membrane, a sheet of 3MM paper, and a sponge (see Note 37). 4. Close the cassette and introduce it into the transfer tank. It is crucial to pay attention to the orientation of the cassette in the tank with the gel on the cathode side and the membrane on the anode side. Fill the tank with transfer buffer. A magnetic stir bar is added in the tank and the tank is placed on top of a magnetic stirrer to ensure constant homogenization of the buffer. Transfer is performed at 4°C for 2 h at 250 mA (see Note 38). 5. Disassemble the sandwich and stain the nitrocellulose membrane in a bath of AmidoBlack staining solution for approximately 10 min. Destain the membrane by three successive washes with destaining solution until a good contrast is obtained. 6. Rinse the membrane with deionized water and take a picture of it with a scanner or a camera. Mark the position of the standards with a pen. 7. To remove any trace of destaining solution, wash the membrane three times for 2 min each time on a rocking platform with PBS or TBS buffer, depending on the buffer to be used for the following steps.
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8. Incubate the membrane in 20 mL of blocking buffer for 1 h at room temperature on a rocking platform. 9. Incubate the membrane with the desired primary antibody at an adequate dilution for 2 h at room temperature or overnight at 4°C on a rocking platform. 10. Remove the primary antibody solution and wash the membrane three times for 10 min with PBS or TBS (see Note 39). 11. Incubate the membrane with secondary antibody solution for 30 min at room temperature on a rocking platform. 12. Wash the membrane in PBS or TBS three times for 10 min each time on a rocking platform. 13. In a square Petri dish, mix 2 mL of ECL reagent and 6.6 mL of 3% H2O2 (see Note 40). Remove excess liquid from the membrane using absorbing paper and place it into the Petri dish. Incubate the membrane for 2 min with the ECL solution making sure the membrane is uniformly covered with the solution. Remove excess ECL solution from the membrane using absorbing paper and place it into an X-ray film cassette. Cover the membrane with a clear plastic material such as wrapping film, and, once in a dark room, place a film on it. Expose the film for 30 s to 10 min. The X-ray film is developed in the dark room with a tabletop processor or by successive baths in standard developer and fixative solutions. Alternatively, the luminescent signal could be monitored using a luminescence imaging device. An example of the results that are obtained is given Fig. 3. 14. It may be necessary to remove the previous antibody before hybridizing a new one. For this, incubate the membrane 30 min in stripping buffer and then wash it with three successive baths of PBS or TBS for 10 min each. After stripping, the membrane must be blocked and the protocol restarts at step 8 of Subheading 3.4.
Fig. 3. Immunoblot analysis of chloroplast membrane fractions isolated from 6-week-old plants. Chloroplast membranes were separated by ultracentrifugation on a sucrose gradient. Fractions of 1 mL were collected from the top (fraction 1 in 5% sucrose) to the bottom (fraction 30 in 45% sucrose) of the sucrose gradient. Proteins from 400 mL aliquots of the even fractions 2–18, from 200 mL aliquots of the even fractions 20–24, from 100 mL aliquots of fractions 26, 28 and stroma (St), and from 50 mL aliquots of fraction 30 and total chloroplasts (Chl) were precipitated, separated by SDSPAGE, and transferred to a nitrocellulose membrane. The membrane was sequentially probed with antibodies raised against membrane marker proteins: AtPGL35 (plastoglobule protein), atToc75 (outer envelope membrane protein), and LHCB2 (thylakoid protein).
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4. Notes 1. EDTA is not soluble below pH 8; thus, NaOH should be added when preparing EDTA solution to help dissolution and after to reach the desired pH. 2. HB buffer is an isotonic solution which will prevent osmotic shock and thus preserve intact chloroplasts. It is prepared with the following stock solutions that are autoclaved and can be stored for several months at room temperature: 1 M Tricine– KOH, pH 8.4; 0.5 M EDTA, pH 8.5; 0.5 M NaHCO3; and 1 M MnCl2. 3. PMSF is a protease inhibitor. It is insoluble in water and must be dissolved at 0.2 M in a solvent such as isopropanol. It is toxic: wear adequate protective clothes when handling the stock solution. 4. HB buffer can be prepared the day before and stored at 4°C. However, sodium ascorbate, BSA, and PMSF should be added just before use. 5. RB buffer is prepared with the following stock solutions that are autoclaved and can be stored several months at room temperature: 1 M Tricine–KOH, pH 7.6; 0.5 M EDTA, pH 8.5; and 1 M MgCl2. 6. Since acetone is highly volatile, the concentration of acetone in the bottle will decrease with time and the extraction of chlorophyll will then be less efficient. Prepare new 80% acetone solution periodically. 7. TrE buffer can be prepared with the following stock solutions: 1 M Tricine–KOH, pH 7.5; 0.5 M EDTA, pH 8; and 1 M DTT (stored at −20°C). 8. APS is unstable at 4°C. Thus, after 2–3 days at 4°C a new aliquot of APS should be opened. 9. AmidoBlack solution must be stored at room temperature in a bottle surrounded by aluminium paper to avoid light damages. It can be reused several times until loss of efficiency but it is recommended to filter it from time to time. 10. ECL solution is prepared in an opaque bottle to prevent light damage, and it may be conserved at 4°C for several weeks. 50 mL of this solution should be prepared by mixing 50 mL of water with the following stock aliquots stored at −20°C: 110 mL of 90 mM coumaric acid and 250 mL of 250 mM luminol. 11. b-Mercaptoethanol is toxic. Work under a fume hood and wear adequate protective clothes.
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12. In order to ensure homogeneous density, seeds are mixed with water in a saltshaker and then distributed on soil. Plant density is critical: high density is needed to obtain enough starting material; however, too high density will induce early flowering plants with very small leaves. For each 30 × 50 cm tray, an amount of Arabidopsis seeds corresponding to a volume of approximately 20 mL is used. 13. Starch accumulates as heavy granules in chloroplasts which might break the organelles during centrifugation. Therefore, it is critical to avoid starch formation before starting chloroplast preparation. 14. Leaves collected from six trays should weigh 150–300 g. 15. Plants are immersed in chilled water to allow decantation of the soil that might have been collected with leaves. 16. Leaves are wrung to eliminate most of the tap water and then introduced into the homogenizer. Enough HB buffer is added to the leaves to allow good grinding. The total leaf amount is generally too important to be introduced all at the same time into the homogenizer; thus, only half or a third of the leaf volume is ground at a time. While the first-round homogenate is filtering, the next batch of leaves is ground with new HB buffer. 17. Resuspension of the pellet should be done as carefully as possible so as not to break the chloroplasts. Gently shake bottles on ice and use plastic Pasteur pipettes with large openings to transfer the solution into a new tube. The complete resuspension of the whole pellet is frequently not achieved with the first 1–2 mL of RB buffer. In such cases, add a supplemental 1–2 mL of RB buffer onto the remaining pellet and carefully scrape the pellet with a plastic Pasteur pipette to help its resuspension. Nevertheless, pay attention not to resuspend the pellet in an excessive total volume of RB, to allow loading the Percoll gradients with no more than 3 mL of crude extract. If necessary, prepare extra Percoll gradients. 18. Disconnect the break so as not to disturb the gradient during deceleration. 19. Chlorophyll content is measured by diluting 5 ml of resuspended chloroplasts in 1 mL of 80% acetone. Mix well by vortexing and spin for 2 min at maximum speed in a microcentrifuge. Transfer the supernatant into a quartz cuvette (a plastic cuvette can be used if the plastic is resistant to acetone) and measure the absorbance at 652 nm (A652) against a blank of 80% acetone. Chlorophyll concentration is calculated as follows:
[Chlorophyll] (mg/mL) = A652 × dilution factor/36
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20. The pellet can be resuspended by vortexing, since at this step the integrity of chloroplasts need not be preserved. 21. The freezing and thawing cycle participates in the disruption of the chloroplasts. 22. Defrosting and mechanical grinding using a Potter homogenizer breaks the chloroplasts. 23. Dilution of the chloroplast suspension followed by ultracentrifugation at 100,000 × g allows the separation of stroma from other chloroplast components (i.e., total chloroplast membranes, including plastoglobules). For protein analysis, an aliquot of the stroma fraction is taken from the supernatant and stored at −80°C. 24. The volume of 45% sucrose solution to be used for membrane resuspension is determined to (1) reach 2–6 mg/mL of chlorophyll concentration and (2) allow the loading of 2–4 gradients (5 mL on each gradient). With this concentration, each gradient will be loaded with 10–30 mg of total membranes, which is the optimal chlorophyll amount needed for efficient plastoglobule preparation with a gradient of 30 mL. 25. At this step, the membrane solution can be stored at −80°C for later fractionation. 26. Efficient homogenization with 20 strokes of the Potter homogenizer is critical to detach plastoglobules from the thylakoid membranes. Possibly, the chloroplast suspension can be sonicated for 1–2 min to help dissociation of plastoglobules from the thylakoids. 27. Use plastic Pasteur pipettes to gently load each sucrose solution on top of the preceding phase, taking care not to disturb the gradient. 28. This ultracentrifugation allows the separation of the different plastid membranes (envelopes, thylakoids and plastoglobules) by flotation in the discontinuous sucrose gradient. 29. Disconnect the break so as not to disturb the gradient during deceleration. 30. The precipitated volume of the first fractions (1–18) is doubled because of the low protein abundance in these fractions. 31. The proteins are present at the interface as a white band. However, in the first fractions, this white band is not visible due to low protein abundance. This does not prejudge the success of the experiment. 32. The protein pellet can also be dried by leaving tubes open under a chemical hood for 1 h to overnight.
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33. The abundance of proteins in the last fractions makes it difficult to resuspend the pellet. This is why a higher volume of SB is added to these fractions. 34. Nonpolymerized acrylamide is toxic; therefore, wear appropriated protective clothes. 35. APS and TEMED catalyze the gel polymerization. High temperature results in faster polymerization. It may be useful to pour the gel in a cold room during summer time. 36. The space needed for the stacking gel is approximately 1 cm below the comb. 37. To remove any air bubbles which may have formed between the different layers of the sandwich and which will interfere with the blotting process, use a plastic spatula and gently roll air bubbles out. 38. Alternatively, transfer could be done overnight at 100 mA, in the cold room. 39. Primary antibody solution can be kept at −20°C and reused three to five times depending on the antibody. 40. Once H2O2 is added to the ECL reagent, the stability of the solution is approximately 15 min. Thus, the following steps need to be rapidly performed.
Acknowledgments We would like to thank Dr. S. Melser and Dr. C. Garcion for critical reading of the manuscript.
References 1. Bréhélin, C., and Kessler, F. (2008) The plastoglobule: a bag full of lipid biochemistry tricks. Photochem. Photobiol. 84, 1388–1394. 2. Austin, J. R., Frost, E., Vidi, P. A., Kessler, F., and Staehelin, L. A. (2006) Plastoglobules are lipoprotein subcompartments of the chloroplast that are permanently coupled to thylakoid membranes and contain biosynthetic enzymes. Plant Cell 18, 1693–1703. 3. Vidi, P. A., Kanwischer, M., Baginsky, S., Austin, J. R., Csucs, G., Dörmann, P., Kessler, F., and Bréhélin, C. (2006) Tocopherol Cyclase (VTE1) Localization and vitamin E Accumulation in
chloroplast plastoglobule lipoprotein particles. J. Biol. Chem. 281, 11225–11234. 4. Ytterberg, A. J., Peltier, J. B., and van Wijk, K. J. (2006) Protein profiling of plastoglobules in chloroplasts and chromoplasts. A surprising site for differential accumulation of metabolic enzymes. Plant Physiol. 140, 984–997. 5. Lohmann, A., Schottler, M. A., Bréhélin, C., Kessler, F., Bock, R., Cahoon, E. B., and Dörmann, P. (2006) Deficiency in phylloquinone (vitamin K(1)) methylation affects prenyl quinone distribution, photosystem I abundance, and anthocyanin accumulation in the
12 Preparation of Plastoglobules from Arabidopsis Plastids… Arabidopsis AtmenG mutant. J. Biol. Chem. 281, 40461–40472. 6. Zbierzak, A. M., Kanwischer, M., Wille, C., Vidi, P. A., Giavalisco, P., Lohmann, A., Briesen, I., Porfirova, S., Brehelin, C., Kessler, F., and Dörmann, P. (2009) Intersection of the tocopherol and plastoquinol metabolic pathways at the plastoglobule. Biochem. J. 425, 389–399. 7. Gaude, N., Bréhélin, C., Tischendorf, G., Kessler, F., and Dörmann, P. (2007) Nitrogen deficiency in Arabidopsis affects galactolipid
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composition and gene expression and results in accumulation of fatty acid phytyl esters. Plant J. 49, 729–739. 8. Kessler, F., Schnell, D., and Blobel, G. (1999) Identification of proteins associated with plastoglobules isolated from pea (Pisum sativum L.) chloroplasts. Planta 208, 107–113. 9. Vidi, P. A., Kessler, F., and Bréhélin, C. (2007) Plastoglobules: a new address for targeting recombinant proteins in the chloroplast. BMC Biotechnol. 7, 4.
Chapter 13 Preparation and Proteomic Analysis of Chloroplast Ribosomes Kenichi Yamaguchi Abstract Proteomics of chloroplast ribosomes in spinach and Chlamydomonas revealed unique protein composition and structures of plastid ribosomes. These studies have suggested the presence of some ribosomal proteins unique to plastid ribosomes which may be involved in plastid-unique translation regulation. Considering the strong background of genetic analysis and molecular biology in Arabidopsis, the in-depth proteomic characterization of Arabidopsis plastid ribosomes would facilitate further understanding of plastid translation in higher plants. Here, I describe simple and rapid methods for the preparation of plastid ribosomes from Chlamydomonas and Arabidopsis using sucrose gradients. I also describe purity criteria and methods for yield estimation of the purified plastid ribosomes and subunits, methods for the preparation of plastid ribosomal proteins, as well as the identification of some Arabidopsis plastid ribosomal proteins by matrixassisted laser desorption/ionization mass spectrometry. Key words: Chloroplast ribosome, Plastid ribosomal protein, Sucrose density-gradient ultracentrifugation, Proteomics, Chlamydomonas reinhardtii, Arabidopsis thaliana
1. Introduction Chloroplast ribosomes, also more generally termed plastid ribosomes, are structurally related to eubacterial 70S ribosomes that are distinct from cytoplasmic 80S and mitochondrial 55S–75S ribosomes (1). Plastid ribosomes are responsible for translation of genes encoded in the plastid genome. In green leaves, chloroplast ribosomes are present at about equimolar amounts relative to cytoplasmic ribosomes (2). With respect to abundance by weight, chloroplast ribosomes account for over 25% of the total leaf ribosomes (1). Although chloroplast ribosomes synthesize only ~80 polypeptides encoded in the plastid genome, about 50% of total protein mass in leaves comprises products of chloroplast ribosomes (3).
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_13, © Springer Science+Business Media, LLC 2011
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Unlike in eubacteria, gene expression in plastids is regulated primarily at the translation level: i.e., the translation rate is not correlated with transcript abundance (4), and translation of many plastid mRNAs is activated in response to light illumination (5, 6). A majority of the studies on plastid translation, especially lightactivated translation, has been carried out in the unicellular green alga, Chlamydomonas (7, 8). Since two landmark discoveries published in 1962, of chloroplast ribosomes from spinach by Lyttleton (9) and of chloroplast DNA from Chlamydomonas by Ris and Plaut (10), the translational apparatus in chloroplasts has been mainly studied in spinach and Chlamydomonas (reviewed in refs. 2, 11–13). Proteomic characterizations of plastid ribosomes from spinach (14–16) and Chlamydomonas (17, 18) have revealed that plastid ribosomes contain some plastid-specific ribosomal proteins (PSRPs) in addition to bacterial orthologs. These proteomic studies also revealed differences in protein composition and the primary structure of each ribosomal protein between higher plants and green algae. Recent cryo-electron microscopy of plastid ribosomes from spinach (19) and Chlamydomonas (20) has visualized the 3D-localization of PSRPs and plastid-specific domains in the ribosomes, suggesting their involvement in translation regulation. Although functional analyses of some PSRPs (PSRP-1 in spinach and PSRP-7 in Chlamydomonas) have been reported (21, 22), the physiological roles of another five PSRPs (PSRP-2 to PSRP-6) remain unclear. In addition, posttranslational modifications, which may also affect translational activity of plastid ribosomes, remain to be elucidated. Although advanced proteomic analyses of Arabidopsis cytoplasmic 80S ribosomes have been performed (23–25), proteomic characterization of Arabidopsis plastid ribosomes has not been reported so far. This may be due to one or more of the following reasons: (1) since plastid ribosomes have been well-characterised in spinach and Chlamydomonas as mentioned above, all the putative plastid ribosomal protein genes of Arabidopsis have already been annotated by sequence homology (17, 18); (2) an isolation method for plastid ribosomes from Arabidopsis has not been established; (3) compared with preparation methods for cytoplasmic ribosomes, those for plastid ribosomes are generally laborious and time consuming, and do not readily yield sufficient quantities or the purity required for proteomics. Even though purification of Arabidopsis plastid ribosomes may not be easy by comparison with Chlamydomonas or spinach, once a method is established it could be used for advanced proteomics: e.g., protein dynamics in translation regulation, posttranslational modifications in translation regulation, etc. Light-dependent phosphorylation of few plastid ribosomal proteins in spinach has been reported (26, 27). Recent large-scale Arabidopsis phosphoproteome profiling suggested that some plastid ribosomal proteins
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are phosphorylated by uncharacterised chloroplast kinases (28). Therefore, detailed phosphoproteomics of plastid ribosomes must be an interesting issue. Large-scale (over 10,000 absorbance at 260 nm [A260] units) preparation methods employing zonal rotors have been established for tobacco (29, 30) and spinach (31). Although these procedures could be also applicable to Arabidopsis, these methods may not be practical for such a tiny plant. For proteomics and molecular biology in Arabidopsis, small-scale (10–100 A260 units) isolation methods using general ultracentrifugation rotors would be preferable. Here, I describe small-scale methods for plastid ribosome preparation, which have been established with Chlamydomonas (17, 18) and Arabidopsis (unpublished results). Methods for purity checking and yield estimation of purified ribosomal particles, as well as preparation of plastid ribosomal proteins for proteomic analysis, are also described.
2. Materials 2.1. Preparation of Total Ribosomes 2.1.1. Biological Materials
1. Plant leaves. For one isolation procedure, 20–200 g (fresh weight) of leaves would be appropriate (see Note 1). For example, 20 g fresh weight of leaves from 40- to 50-day-old Arabidopsis thaliana Columbia-0 plants (see Note 2). 2. Algal cells. For example, Chlamydomonas reinhardtii (strain CC-3395) grown at 25°C under constant light in 2 L of liquid TAP medium (32) with 50 mg/mL L-Arg to a density of 5–8 × 106 cells/mL (mid-late log phase) (see Note 3).
2.1.2. Buffers
1. Buffer A (all-round extraction buffer): 25 mM Tris–HCl, pH 7.6, 25 mM KCl, 25 mM MgCl2, 5 mM dithiothreitol (see Note 4). 2. Buffer B (alternative extraction buffer): 25 mM Tris–HCl, pH 7.6, 25 mM KCl, 25 mM MgCl2, 14 mM 2-mercaptoethanol (see Note 5). 3. Buffer C (cushion buffer): 1 M sucrose in Buffer A.
2.1.3. Homogenization
1. Laboratory blender (e.g., Waring Blender 7011HS). 2. One ice bucket, containing ice, for chilling buffers and blender during cell disruption.
2.1.4. Centrifuges, Rotors, and Tubes
1. High-performance centrifuge (e.g., Avanti J-E Centrifuge, Beckman Coulter, CA, USA). 2. Beckman Coulter JA-10 fixed-angle rotor or equivalent. 3. Tubes compatible with the JA-10 rotor (e.g., 500-mL Nalgene centrifuge tubes).
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4. Beckman Coulter JA-17 fixed-angle rotor or equivalent. 5. Tubes compatible with the JA-17 rotor (e.g., 15-mL Nalgene centrifuge tubes). 6. Tabletop ultracentrifuge (e.g., Optima TLX Personal Benchtop Ultracentrifuge, Beckman Coulter). 7. Beckman Coulter TLA-100.3 fixed-angle rotor or equivalent. 8. Tubes compatible with the TLA-100.3 rotor (e.g., thickwall polycarbonate tubes, 3.5 mL, 13 × 51 mm, Beckman Coulter). 9. Preparative ultracentrifuge (e.g., Optima L-100K Preparative Ultracentrifuge, Beckman Coulter). 10. Beckman Coulter Ti70.1 fixed-angle rotor or equivalent. 11. Tubes compatible with the Ti70.1 rotor (e.g., open-top thickwall polycarbonate tubes, 10 mL, 16 × 76 mm, Beckman Coulter). 2.2. Separation of Plastid Ribosomes and Subunits
1. Buffer A (see Subheading 2.1.2) and stock solutions of each buffer component (see Note 4).
2.2.1. Buffers and Solutions
3. Buffer D (dissociation buffer): 25 mM Tris–HCl, pH 7.6, 100 mM KCl, 5 mM MgCl2, and 5 mM dithiothreitol.
2.2.2. Apparatus for Gradient Preparation/ Fractionation System
1. Gradient maker, 50 mL (e.g., SG 50 Gradient Maker, Hoefer, MA, USA).
2. Sucrose stock solution: 2 M (68.5% [w/v]) sucrose.
2. Magnetic stirrer and stirring bar. 3. Density-gradient fractionator (e.g., Auto Densi-Flow II, Buchler, NJ, USA). This item is used for depositing a preformed sucrose gradient into an ultracentrifuge tube (i.e., preparation of sucrose gradients before ultracentrifugation), and for gentle withdrawal of centrifuged sample layers (i.e., fractionation of sucrose gradients after ultracentrifugation). 4. Peristaltic pump (e.g., Econo Gradient Pump with 0.8-mm internal diameter PharMed tubing; Bio-Rad, CA, USA). 5. UV-monitor (e.g., Model EM-1 Econo UV Monitor, BioRad). 6. Tubing (e.g., 0.8-mm internal diameter Tygon tubing, BioRad). 7. Fraction collector (e.g., Model 2110 Fraction Collector, Bio-Rad). 8. Chart recorder (e.g., Model 1327 Chart Recorder, Bio-Rad).
2.2.3. Centrifuges, Rotors, and Tubes
1. Preparative ultracentrifuge (e.g., Optima L-100K Preparative Ultracentrifuge, Beckman Coulter). 2. Beckman Coulter SW28 Ti swinging-bucket rotor or equivalent. 3. Tubes compatible with the SW28 rotor (e.g., open-top thinwall polyallomer tubes, 38.5 mL, 25 × 89 mm, Beckman Coulter).
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4. Beckman Coulter Ti70.1 fixed-angle rotor or equivalent. 5. Tubes compatible with the Ti70.1 rotor (e.g., open-top thickwall polycarbonate tubes, 10 mL, 16 × 76 mm, Beckman Coulter). 2.3. Purity Criteria and Yield Estimation
1. Spectrophotometer (e.g., NanoDrop ND-1000, Thermo Scientific, MA, USA). 2. Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) system (e.g., Mini-Protean System, Bio-Rad).
2.4. Preparation of Plastid Ribosomal Proteins for Proteomics
1. Ribosomal protein extracting solution: 50 mM magnesium acetate in glacial acetic acid. To prepare 100 mL of this solution, dissolve 1.07 g of magnesium acetate tetrahydrate (nuclease and protease tested) in ~90 mL of acetic acid (glacial, ³99.8%) at room temperature (20–25°C), then fill up to 100 mL with acetic acid. This solution can be stored in a Pyrex screw-cap storage bottle at room temperature for at least 6 months. 2. One ice bucket, containing ice. 3. One glass test tube (e.g., Pyrex test tube, 13 × 100 mm). An acid-stable glass tube is needed for extraction of plastid ribosomal proteins. 4. Stirring bar compatible with test tube (13 × 100 mm) and magnetic stirrer. 5. Dialysis buffers. For high-performance liquid chromatography (HPLC) or SDS-PAGE, prepare 30% (v/v) acetic acid, 10% (v/v) acetic acid, and 5% (v/v) acetic acid. For twodimensional (2D)-PAGE, prepare 8 M urea containing 0.1% (v/v) 2-mercaptoethanol. Urea should be ultrapure grade (e.g., Bio-Rad). 6. Dialysis membrane, with a molecular weight cut-off of 3.5 kDa (e.g., Spectra/Por 3 Dialysis Membrane, Spectrum, Japan). 7. Dialysis membrane closures (e.g., Spectra/Por Closures, Spectrum, Japan). 8. Solvent-absorbent powder (e.g., Spectra/Gel Absorbent, Spectrum, Japan). This is a gel powder (dehydrated polyacrylate-polyalcohol), which is used to concentrate dialysis samples (see Note 6).
2.5. Rapid Preparation of Plastid RibosomeRich Fraction
1. Buffer A (see Subheading 2.1.2) and stock solutions of each buffer component (see Note 4).
2.5.1. Buffers, Solutions, Reagents, and Apparatus
3. Sucrose stock solution: 2 M (68.5% [w/v]) sucrose.
2. Buffer C (see Subheading 2.1.2). 4. Solid ammonium sulfate (for molecular biology, ³99%).
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5. The apparatus needed for gradient preparation and the fractionation system are the same as those listed in Subheading 2.2.2, except that the size of gradient maker is different (15 mL, instead of 50 mL). 2.5.2. Centrifuges, Rotors, and Tubes/Bottles
1. Refrigerated centrifuge (e.g., Compact Refrigerated Centrifuge 7780, Kubota, Japan). 2. Kubota AG-508CA (see Note 7).
fixed-angle
rotor
or
High-Speed equivalent
3. Tubes compatible with the AG-508CA rotor (e.g., 50-mL polypropylene conical centrifuge tubes, Greiner Bio-One, Germany). 4. Preparative ultracentrifuge (e.g., Himac CP 75 beta, Hitachi, Japan). 5. Hitachi P70AT2 fixed-angle rotor or equivalent. 6. Bottles compatible with the P70AT2 rotor (e.g., 8.4-mL 10PC bottle B, Hitachi). 7. Hitachi P40ST swinging-bucket rotor or equivalent. 8. Tubes compatible with the P40ST rotor (e.g., 10.9-mL 13PA tubes, Hitachi). 2.6. SDS-PAGE of Plastid Ribosomal Proteins from Sucrose Gradient-Separated Ribosome Fractions
1. 100% saturated ammonium sulfate (SAS) at 0°C. To 100 mL of RNase-free water, add 70.7 g of ammonium sulfate (for molecular biology, ³99%) and dissolve completely by stirring. This SAS solution is saturated at 0°C (for practical use on ice). Make aliquots in 15-mL conical tubes for one-time to fewtimes usage, and store at 4°C. 2. Running buffer (10×): 250 mM Tris, 1.92 M glycine, and 1% (w/v) sodium dodecyl sulfate (SDS). 3. SDS-sample buffer (1×): 62.5 mM Tris–HCl, pH 6.8, 2% (w/v) SDS, 10% (w/v) glycerol, 5% (v/v) 2-mercaptoethanol, and 0.001% (w/v) bromophenol blue; as reported by Laemmli (33). 4. Precast polyacrylamide gels (e.g., Mini-Protean TGX Gels, Bio-Rad). 5. Prestained molecular-weight markers (e.g., Precision Plus Protein Dual Xtra Standards, Bio-Rad). 6. CBB staining solution: 0.1% (w/v) Coomassie Brilliant Blue-R250 (CBB), 40% (v/v) methanol, and 10% (v/v) acetic acid (e.g., JIS special grade, 99.7%, Wako Pure Chemical Industries, Japan). Alternatively, use a commercial stain (e.g., Bio-Rad CBB-R250 staining solution). 7. Destaining solution: 40% (v/v) methanol and 10% (v/v) acetic acid.
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2.7. Identification of Plastid Ribosomal Proteins by MALDI Mass Spectrometry
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1. Utility knife for handcrafts (e.g., Art Knife AK-1/5B, Olfa, Japan) (see Note 8). 2. Overhead projector (OHP) sheets. 3. Destaining solution: 30% (v/v) acetonitrile in 25 mM ammonium bicarbonate. 4. Acetonitrile. 5. Vacuum centrifuge (e.g., SpeedVac, Thermo Scientific). 6. Trypsin solution. Prepare a 10 mg/mL solution of proteomicsgrade trypsin (e.g., Trypsin, Proteomics Grade, T6567, Sigma) in 10% (v/v) acetonitrile, 25 mM ammonium bicarbonate. 7. Microwave oven. 8. Extraction solution: 5% (v/v) formic acid and 50% (v/v) acetonitrile. 9. Cup horn sonicator (e.g., Astrason Ultrasonic Processor XL2020, Misonix, NY, USA). 10. Matrix solution. For example, 1× DHBA solution (5 mg/mL of 2, 5-dihydroxybenzoic acid [e.g., DHBA, Shimadzu, Japan]) in 33% (v/v) acetonitrile, 0.1% (v/v) trifluoroacetic acid (TFA). 11. Matrix-assisted laser desorption/ionization (MALDI) mass spectrometer capable of tandem mass spectrometry (MS/MS) analysis (e.g., AXIMA Resonance, Shimadzu, Japan).
3. Methods 3.1. Preparation of Total Ribosomes
Chloroplast ribosomes can be purified from either isolated chloroplasts (34) or from total cell homogenates (18, 29, 30). It has been suggested that isolation of chloroplast ribosomes from total cell homogenates results in a higher yield compared with that from isolated chloroplasts (30). 1. Harvesting plant leaves. Wear disposable examination gloves and cut the leaves with sterile scissors. After the harvest, rinse the leaves with chilled sterile water, drain off the water with a strainer, and keep in the cold room (4°C, in the dark) for at least 1 h. This step is required to induce polysomal runoff, yielding an accumulation of free ribosomes (35, 36). The leaves should be frozen in liquid nitrogen before homogenization (see Note 9). 2. Harvesting algal cells. Pour the Chlamydomonas cell culture into six 500-mL Nalgene centrifuge tubes and harvest the cells by centrifugation at 2,800 × gmax in a high-performance centrifuge (e.g., Beckman Coulter Avanti J-E) with a fixed-angle rotor (e.g., Beckman Coulter JA-10) for 5 min at 4°C. Put the tubes on ice, suspend the cells with 40 mL of chilled liquid TAP
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medium, and then transfer the suspension to four 15-mL Nalgene centrifuge tubes. Pellet the cells by centrifugation at 4,000 × gmax in a fixed-angle rotor (e.g., Beckman Coulter JA-17). Cells in the tube should be frozen in liquid nitrogen before homogenization (see Note 10). 3. Homogenization. When working with plant leaves, homogenize the frozen leaves (e.g., 20 g with 40 mL of Buffer A) in a chilled Waring Blender for 1 min (three 20-s bursts). Alternatively, when working with algal cells, homogenize the frozen cells (e.g., 1.0–1.6 × 1010 cells from 2 L culture of Chlamydomonas reinhardtii CC-3395 with 40 mL of Buffer A) in a chilled Waring Blender for 1 min. 4. Centrifuge the homogenates at 10,000 × gmax for 10 min at 4°C (e.g., at 8,500 rpm in a Beckman Coulter JA-17 rotor). 5. Centrifuge the supernatant (the S-10 fraction) at 40,000 × gmax for 30 min at 4°C (e.g., at 27,000 rpm in a Beckman Coulter TLA-100.3 fixed-angle rotor) and collect the supernatant (the S-40 fraction). 6. Layer 5 mL of the S-40 fraction over 2 mL of Buffer C (cushion buffer containing 1 M sucrose) and centrifuge at 330,000 × gmax for 12 h at 4°C (e.g., at 60,000 rpm in a Beckman Coulter Ti70.1 fixed-angle rotor). 7. Discard the supernatant and dissolve the ribosomal pellet in a minimal volume of Buffer A. 8. Clarify the sample by centrifugation at 15,000 × gmax in a microfuge for 10 min at 4°C to remove insoluble materials, and measure absorbance at 260 nm (A260). The concentration of the total ribosomes should be 200–1,000 A260 units/mL. 9. Make aliquots of the total ribosome preparation, freeze them in liquid nitrogen, and store at −80°C. 3.2. Separation of Plastid Ribosomes and Subunits 3.2.1. Preparation of Sucrose Gradients
Use the following procedure to prepare a sucrose gradient (e.g., a 10–40% (w/v) linear sucrose gradient; 36 mL) that will be used to separate plastid (70S) and cytoplasmic (80S) ribosomes. 1. Assemble a gradient preparation system. Put the gradient maker on a magnetic stirrer. Add a magnetic stirring bar to the mixing chamber. Connect the gradient maker to the inlet of a peristaltic pump and a density-gradient fractionator to the outlet of the pump with tubing. Before starting sucrose-gradient preparation, clean the gradient maker and tubing with RNase-free water at the maximum flow rate at room temperature for ~30 min (e.g., the maximum rate is 1.74 mL/min using the Econo Gradient Pump with 0.8-mm internal diameter PharMed tubing). 2. Prepare 10% sucrose in Buffer A and 40% sucrose in Buffer A by diluting 2 M sucrose stock and buffer component stocks (see Note 4).
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3. Close the stopcocks and add 18 mL of 10% sucrose in Buffer A to the reservoir chamber of the gradient maker. Slowly open the connector stopcock and remove air in the connector channel, then close the connector stopcock. 4. Slowly add 18 mL of 40% sucrose in Buffer A to the mixing chamber. Open the connector stopcock and start stirring. 5. Place a centrifuge tube (e.g., Beckman Coulter thinwall polyallomer 38.5-mL tube) on the tube holder of the density-gradient fractionator. Open the delivery stopcock and make the gradient by pumping at a flow rate of 1.0–1.3 mL/min for ~30 min. 3.2.2. Plastid Ribosome Separation
1. Carefully load 1–2 mL of total ribosome preparation (diluted to 25–50 A260 units/mL with Buffer A) onto the top of a 36 mL 10–40% (w/v) linear sucrose gradient made up in Buffer A. 2. Centrifuge at 91,000 × gmax for 12 h at 4°C (e.g., at 22,500 rpm in a Beckman Coulter SW28 rotor). 3. Assemble a gradient fractionation system. Disconnect tubing from the gradient maker of the gradient preparation system (see Subheading 3.2.1, step 1) and connect the tubing to the inlet of a UV-monitor. Connect the outlet of the UV-monitor to a fraction collector with tubing (e.g., 0.8mm internal diameter Tygon tubing). Connect the UV-monitor to a chart recorder with an appropriate cable. Set the pumping direction to reverse (i.e., from the densitygradient fractionator to the UV-monitor). Before fractionation, wash the tubing and probe of the density-gradient fractionator, withdrawing 30–35 mL of RNase-free water from a clean ultracentrifuge tube. 4. Fractionate the gradient into ~40 microfuge tubes (1.0 mL/ tube) from the top surface of the gradient using the gradient fractionation system at a flow rate of 1.0–1.3 mL/min for ~30 min at room temperature. Monitor the absorbance at 254 nm using the UV-monitor and chart recorder (chart speed set at 5–10 mm/min). After the fractionation, the tubes should be immediately placed on ice. An example sucrose-gradient profile of total ribosomes from Chlamydomonas is shown in Fig. 1a. 5. Recover the 70S plastid ribosomes as pellets from appropriately pooled fractions (see Fig. 1a) by centrifugation at 330,000 × gmax for 12 h at 4°C (e.g., at 60,000 rpm in a Beckman Coulter Ti70.1 fixed-angle rotor) (see Note 11). 6. Repurify the ribosomes by using a second sucrose gradient, if highly pure 70S ribosomes are needed (i.e., repeat steps 1–5 above) (see Note 12).
3.2.3. Subunit Preparation
1. Add a small volume of Buffer D to the 70S pellet and dissolve the pellet by pipetting to dissociate the ribosomes into subunits.
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Fig. 1. Isolation of chloroplast 70S ribosomes and their subunits from the green alga Chlamydomonas reinhardtii. (a) Sucrose-gradient (10–40%) analysis of a total ribosome preparation (50 A260 units) from C. reinhardtii cells in Buffer A. Peaks corresponding to plastid ribosomes (70S) and cytoplasmic ribosomes (80S) are indicated by arrowheads. The underlined 70S fractions were pooled and subjected to a second sucrose-gradient purification. (b) Separation of chloroplast 30S and 50S subunits in a sucrose density-gradient (10–30%) after dissociation of the 70S ribosomes with Buffer D; the underlined fractions were collected for the making of total protein preparations (TP30 and TP50, respectively).
Adjust the subunit concentration to 20–40 A260 units/mL with Buffer D. 2. Carefully load 1–2 mL of subunits (20–40 A260 units) onto a 36 mL 10–30% sucrose gradient made up in Buffer D. Centrifuge at 91,000 × gmax for 20 h at 4°C (e.g., at 22,500 rpm in a Beckman Coulter SW28 rotor). 3. Fractionate the gradients as described in Subheading 3.2.2 (steps 3–4). An example sucrose-gradient profile of Chlamydomonas plastid ribosome subunits is shown in Fig. 1b. 4. Recover the 30S and 50S subunits as pellets from appropriately pooled fractions (see Fig. 1b) by centrifugation at 330,000 × gmax for 14 h at 4°C (e.g., at 60,000 rpm in a Beckman Coulter Ti70.1 fixed angle rotor) (see Note 13). 3.3. Purity Criteria and Yield Estimation
Fine proteomic analysis will require highly pure ribosomes. The following section describes easy and rapid methods for purity checking and yield estimation of plastid ribosomes. 1. Measure absorbance at 260 and 280 nm of appropriately diluted solutions of ribosomal particles using spectrophotometer. 2. Purity criteria. Pure plastid ribosomes and subunits must show an A260/280 ratio greater than 1.9. Although ratios in the range 1.7–1.9 for A260/280 have been an accepted criterion for a pure plastid ribosome preparation (31), significant contamination
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Table 1 Conversion table for 1 A260 unit of plastid ribosomal particles of a higher plant (spinach) Particle amountd
Ribosomal particle
Protein mass a (kDa)
RNA mass b (kDa)
Particle mass c (kDa)
(mg)
(pmol)
Protein amount e (mg)
70S
1,022
1,558
2,580
66
26
26 (TP70)
50S
571
1,040
1,611
62
38
22 (TP50)
30S
429
518
947
73
77
33 (TP30)
The sum of the masses of individual ribosomal proteins (14, 15), taking into account that L12 protein is present in four copies per 50S subunit and that a 22-kDa protein (pRRF) is present in 70S but absent from either 30S or 50S subunits (15, 49) b As the sodium salt, calculated from the nucleotide sequences (50) of 16S rRNA (1491 nucleotides [nt]) of the 30S subunit, and of 23S rRNA (2811 nt), 5S rRNA (121 nt), and 4.5S rRNA (103 nt) of the 50S subunit c The sum of the total protein mass and the total RNA mass, not taking into account protein/RNA modifications, metal ions (e.g., Mg2+) required for rRNA folding/stabilizing, and other ribosomal components such as polyamines d Calculated from the RNA amount, using the relationship that one A260 unit of RNA corresponds to 40 mg e Calculated from the following equation: (total protein of the ribosomal particle) = (particle weight) − (RNA weight, i.e., 40 mg). TP70, TP50, and TP30 stand for total proteins present in 70S, 50S, and 30S ribosomal particles, respectively a
of RuBisCO is usually suspected even in samples with a high A260/280 ratio (>1.9). SDS-PAGE analysis of sucrose-gradient fractions (see Subheading 3.6) can be utilized to examine the extent of RuBisCO contamination (50-kDa and 15-kDa bands correspond to the large and small subunits of RuBisCO, respectively). 3. Yield estimation. The yields of plastid ribosomes and their subunits can be estimated spectrophotometrically using the table of amount conversion (see Table 1). For example, 1 A260 unit of plastid 70S ribosomes corresponds to approximately 66 mg (26 pmol) of particles. Since protein recovery with the procedure of acetic acid extraction (37) is close to 100% (over 98%), the amount of extracted ribosomal proteins can be also estimated with the conversion table, without the need for a protein assay; for example, acetic acid-extracted total protein (TP70) from 1 A260 unit of plastid 70S ribosomes corresponds to 26 mg (26 pmol) of protein. 4. RNA integrity. Intactness of rRNA is not a requirement for the maintaining of a full complement of ribosomal proteins, of sedimentation coefficients of the subunits, or of translation activity (30, 38), although highly degraded rRNA does cause anomalous sedimentation peaks on sucrose gradients (30). Intactness of rRNA can be easily assessed by agarose gel
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e lectrophoresis after RNA extraction by the acid guanidiniumthiocyanate–phenol–chloroform (AGPC) method (39) using an RNA isolation reagent (e.g., TRIzol reagent, Invitrogen). 3.4. Preparation of Plastid Ribosomal Proteins for Proteomics
The following procedure is a slightly modified method of acetic acid extraction method described by Hardy et al. (37). Total proteins of plastid ribosomal particles prepared using this procedure can be used for various separation techniques in proteomics: e.g., SDS-PAGE (17, 18), HPLC, liquid chromatography-mass spectrometry (LCMS), MALDI mass spectrometry, and 2D-PAGE (14, 15). 1. Dilute ribosomes or subunits with Buffer A and adjust the concentration to 20–100 A260 units/mL. 2. Transfer to a glass test tube with a stirring bar and chill the ribosome solution in an ice-filled bucket. Place the bucket on a magnetic stirrer and start stirring. 3. Slowly add 2 volumes of ribosomal protein extracting solution (50 mM magnesium acetate in glacial acetic acid). Cloudy insoluble rRNA will appear immediately. 4. Stir on ice for at least 1 h. 5. Transfer the cloudy suspension to 1.5-mL centrifuge tubes. Remove the insoluble rRNA by centrifugation at 15,000 × gmax for 10 min at 4°C (e.g., at 14,800 rpm in a Hitachi Himac RT15A3 rotor). Retain the supernatant. 6. For SDS-PAGE, HPLC, and MALDI mass spectrometry, take the supernatant (from step 5) and dialyze it against the following, in the following order: (1) 100 volumes of 30% acetic acid for 4 h, (2) 100 volumes of 10% acetic acid for 4 h, and (3) 200 volumes of 5% acetic acid for 16 h; all dialysis steps should be conducted in a cold room (4°C). Make aliquots of the desalted ribosomal proteins, then lyophilize or dry using a SpeedVac. Store the protein samples at −20°C or −80°C. An example SDS-PAGE gel resolving total proteins from 30S, 50S, 70S, and 80S particles from Chlamydomonas is shown in Fig. 2. 7. Alternatively, for 2D-PAGE (40), take the supernatant (from step 5 above) and dialyze it against the following: (1) 100 volumes of 8 M urea, 0.1% 2-mercaptoethanol for 4 h and (2) 200 volumes of 8 M urea, 0.1% 2-mercaptoethanol for 16 h; both dialysis steps to be conducted in a cold room (4°C). Concentrate the ribosomal proteins with solvent-absorbent powder to 1–2 nmol/mL. Store the protein samples at −20°C or −80°C. 8. Protein resolution for mass spectrometry. Because many plastid ribosomal proteins possess highly basic isoelectric points (pI > 11), commercially available IPG (Immobilized pH Gradient)-based 2D-PAGE systems, enabling separation with a
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Fig. 2. SDS-PAGE profiles of total proteins from 30S, 50S, 70S, and 80S ribosomal particles prepared from Chlamydomonas reinhardtii. Total proteins (TP) extracted from 10 pmol of each of the following fractions were resolved by SDS-PAGE using a 1.5-mm thick, 12% acrylamide gel: chloroplast small subunits (TP30), chloroplast large subunits (TP50), chloroplast ribosomes (TP70), and cytoplasmic ribosomes (TP80). Proteins were stained with Coomassie Brilliant Blue R-250. All the protein bands from TP30, TP50, TP70, and TP80 were analyzed by LC-ESI-MS/MS (17, 18, 47).
pI range of 3–10, are not suitable for proteomics of plastid ribosomal proteins. The hand-crafted acrylic plastic 2D-PAGE apparatus designed by Mets and Bogorad (41) and the buffer/ gel system improved by Subramanian (40) are recommended. The RFHR (radical-free and highly reducing) method (42) would also be preferable to resolve plastid ribosomal proteins. Although 2D-protein mapping is a powerful approach for visualizing individual ribosomal proteins, high-throughput mass spectrometry (e.g., liquid chromatography-electrospray ionization-tandem mass spectrometry [LC-ESI-MS/MS]) allows identification of an almost complete set of plastid ribosomal proteins from protein bands separated by 1D-SDS-PAGE
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K. Yamaguchi
a
10
80S
Abs 254 nm
8
70S 6
4
2
0
1
2
3
4
5
6
7
8
9 10 11 12
Fraction (0.8 ml/tube) b (kDa) 100 75 50
M 1
2
3
4
5
6
7
8
1a 6a
37 5a
25
6b
5b
20 15
1b
10 5 Fig. 3. Sucrose density-gradient separation of 70S chloroplast ribosomes and 80S cytoplasmic ribosomes from the leaves of Arabidopsis thaliana, and SDS-PAGE profiles of the sucrose-gradient fractions. (a) Ten A260 units of total ribosomes (A260/280 = 1.98) were loaded on a 10–40% sucrose gradient in Buffer A (10 mL) and centrifuged as described in Subheading 3.5. (b) Proteins (present in a half volume of each fraction) in fractions 1–8 of panel a were resolved on a Mini-Protean TGX precast gel (Any kD, 15-well comb, Bio-Rad) and stained with Coomassie Brilliant Blue R-250. Protein band sections used for mass spectrometric analysis are indicated by dotted boxes. Proteins in sections 1a (50-kDa band) and 1b (15-kDa band) were identified as RuBisCO large subunit and small subunit, respectively, by peptide mass fingerprinting. Almost invisible 50-kDa (corresponding to gel section 1a: RuBisCO large subunit) and 45-kDa (corresponding to gel section 6a: cytoplasmic 60S ribosomal subunit proteins L3 and L4; see Table 2) bands in fraction 5 indicate that fraction 5 is rich in plastid 70S ribosomes with a slight contamination of RuBisCO and 80S ribosomes. This was also supported by distinct MALDI MS spectra (see Fig. 4) of tryptic fragments from gel sections 5a (30-kDa band of fraction 5) and 6b (30-kDa band of fraction 6).
(e.g., Fig. 2) or from the whole ribosomal protein complex (17, 18). MALDI-quadrupole ion trap-time of flight (QITTOF) MS also allows identification of plastid ribosomal proteins (see Figs. 3 and 4), posttranslational processing, and
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Fig. 4. MALDI MS spectra of tryptic fragments obtained from Arabidopsis ribosome preparations. Section 5a (30-kDa band in fraction number 5, i.e., 70S-rich fraction; upper panel ) and section 6b (30-kDa band in fraction number 6, i.e., 80S-rich fraction; lower panel ) of the gel shown in Fig. 3 were excised and analysed. White arrowheads indicate ion peaks assigned as plastid ribosomal proteins by MS/MS analysis; black arrowheads indicate those for cytoplasmic ribosomal proteins. Regions of relatively low signal in both spectra (m/z 1,750–2,300) were amplified fivefold (×5). Peptide ions annotated by numeral (m/z ), and protein ID in parentheses, indicate that positive protein identification was performed by MS/MS ion searching with significant probability-based Mascot scores (p < 0.05). The scores, peptide sequences, and other para meters obtained by Mascot searches are listed in Table 2.
modifications (data not shown) without 2D-PAGE separation. Thus, 2D-PAGE is not indispensable to conduct the proteomic characterization of plastid ribosomes. 3.5. Rapid Preparation of Plastid RibosomeRich Fraction
This procedure employs ammonium sulfate precipitation to concentrate total ribosomes, which allows efficient purification of intact 70S ribosomes from E. coli (43).
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1. Homogenize frozen leaves (e.g., 20 g) as described in Subheading 3.1. 2. Transfer the homogenate to two 50-mL conical tubes and centrifuge at 15,000 × gmax for 10 min at 4°C (e.g., at 10,600 rpm in a Kubota AG-508CA rotor). 3. Transfer the supernatant (25 mL of supernatant from each tube: total 50 mL) to a 200-mL glass beaker using a disposable pipette. Put a stirring bar in the beaker and put the beaker in an ice bucket. Place the bucket on a magnetic stirrer and start stirring. 4. Add 24 g of solid ammonium sulfate per 50 mL of supernatant under stirring on ice. 5. Stir for 5–10 min (until solid ammonium sulfate is completely dissolved). 6. Transfer the cloudy solution to two 50-mL conical tubes and then centrifuge at 15,000 × gmax for 10 min at 4°C (e.g., at 10,600 rpm in a Kubota AG-508CA rotor). 7. Discard the supernatant and add 5 mL of Buffer A to each tube. 8. Suspend the pellet by pipetting up and down (most of the pellet will be dissolved). 9. Transfer the suspension to two ultracentrifuge bottles (e.g., Hitachi 8.4-mL 10PC bottle B) and centrifuge at 40,000 × gmax for 30 min at 4°C (e.g., at 20,800 rpm in a Hitachi P70AT2 fixed-angle rotor). 10. Layer 3–5 mL of the supernatant (the S-40 fraction) over 2 mL of Buffer C and centrifuge at 230,000 × gmax for 16 h at 4°C (e.g., at 50,000 rpm in a Hitachi P70AT2 fixed-angle rotor). 11. Discard the supernatant and dissolve the ribosomal pellet in a minimal volume of Buffer A. 12. Clarify by centrifugation at 15,000 × gmax for 10 min at 4°C to remove insoluble material, and measure absorbance at 260 nm. The concentration of the total ribosomes should be 200–1,000 A260 units/mL. 13. Prepare one or more 10 mL 10–34% (w/v) linear sucrose gradients made up in Buffer A (e.g., in a Hitachi 10.9-mL 13PA tube) as described in Subheading 3.2.1. 14. Load 0.1–0.2 mL of total ribosome preparation (equivalent to 10–20 A260 units; the concentration should be adjusted with Buffer A if necessary) onto each sucrose gradient. Centrifuge at 111,000 × gmax for 5 h at 4°C (e.g., at 25,000 rpm in a Hitachi P40ST rotor). 15. Fractionate the gradients into 12 microfuge tubes (0.8 mL/tube) from the top surface of the gradient using the density-gradient
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fractionator by pumping at a flow rate of 0.8 mL/min through a UV-monitor and fraction collector (as described in Subheading 3.2.2, steps 3–4). Monitor the absorbance at 254 nm using a UV-monitor. 3.6. SDS-PAGE of Plastid Ribosomal Proteins from Sucrose Gradient-Separated Ribosome Fractions
This procedure is a rapid and reproducible method using a precast gel system (Bio-Rad) to separate plastid ribosomal proteins immediately after fractionation on a sucrose gradient. This method is also useful to evaluate the purity of sucrose gradient-separated ribosome fractions (see Fig. 3). However, attention should be paid to data interpretation in the case of proteomic analysis, since some plastid ribosomal proteins may not appear on gels depending on the source of ribosomal particles. For example, some plastid ribosomal proteins of Chlamydomonas (e.g., S2, S3 and S5) are hardly extracted by the following procedure (i.e., SDS-treatment alone). whereas most of the other proteins are extracted. In such cases, an acetic acid extraction method (see Subheading 3.4) should be employed. 1. Ammonium sulfate precipitation. Transfer 400 mL of each sucrose-gradient fraction to a 1.5-mL centrifuge tube and add 1 mL of SAS to each tube. Shake vigorously by hand for few seconds and keep them on ice for 10 min. Spin down the precipitate by centrifugation at 15,000 × gmax for 10 min in a microfuge. Withdraw and discard the clear supernatant using a 1-mL Gilson pipette tip. Centrifuge again at 15,000 × gmax for 1 min and remove as much residual liquid as possible using a 200-mL Gilson pipette tip. 2. SDS-PAGE sample preparation. Add 20 mL of 1× SDS sample buffer to the pellet (from step 1) and incubate the tube at 95°C for 1 min. Suspend the precipitated sample by pipetting at room temperature and incubate the tube again at 95°C for 5 min. Centrifuge at 15,000 × gmax for 10 min in a microfuge to remove insoluble material. 3. Assemble the gel unit and connect to a power supply. 4. Load prestained molecular-weight markers and the samples. Run the SDS-PAGE at 200 V (~28 V/cm) until the front protein marker (2 kDa) is 1 cm from the bottom of the gel. 5. Stain with CBB staining solution for 15–30 min with constant shaking and then destain with destaining solution until a clear background is obtained. 6. Sandwich the gel between clear plastic sheets (e.g., clean OHP sheets) to protect the gel from physical damage and to avoid contamination (e.g., by keratin) and then place it in a plastic zip-seal bag and store at 4–10°C until use (e.g., the following section).
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3.7. Identification of Plastid Ribosomal Proteins by MALDI Mass Spectrometry
Here described is an example procedure for conducting MALDI mass spectrometry analysis of Arabidopsis plastid ribosomal proteins using MALDI-QIT-TOF MS (see Figs. 3 and 4). Microwaveassisted protein enzymatic digestion (44, 45) allows rapid and accurate identification of the plastid ribosomal proteins (see Table 2). 1. Place the CBB-stained gel on a clean OHP sheet. 2. Excise the protein band of interest using a sterile cutting blade (e.g., half of a band; see Fig. 3b) (see Notes 8 and 14). 3. Cut the excised gel slice into several 1-mm2 pieces and transfer two pieces to a 1.5-mL tube. 4. To the tube, add 500 mL of destaining solution and agitate for 10 min. 5. Discard the solution and repeat step 4 (usually two to three times) until the gels look clear. 6. Dehydration. Add 500 mL of acetonitrile to the tubes and agitate for 10 min (during agitation, the clear gel pieces will gradually shrink and turn white). Discard the solution. 7. Dry the gel pieces by using a vacuum centrifuge (e.g., SpeedVac) for 10 min at 45°C. 8. Microwave-assisted protein enzymatic digestion. Add 10 mL of trypsin solution per one tube and put on ice for 20–30 min (until the gel pieces look clear). Remove any excess amount of trypsin solution using a 10-mL Gilson pipette tip and close the caps of the tubes. Put the tubes in a plastic tube stand. In a microwave oven, place a 300-mL beaker containing 200 mL of water (see Note 15) and put the tube stand on top of the beaker. Microwave the samples for 5 min at 200–300 W. 9. Peptide extraction. Add 20 mL of extraction solution to each tube and sonicate in a cup horn sonicator (e.g., Astrason Ultrasonic Processor XL2020, Misonix, at output level 4) for 1 min. Transfer the solutions to new 0.5-mL centrifuge tubes. 10. Dry the peptide fragments by using a vacuum centrifuge (e.g., SpeedVac) for 15 min at 45°C. 11. Add 2 mL of matrix solution (e.g., DHBA solution) and dissolve the peptides by pipetting. 12. Spot 1 mL of the sample solution onto a MALDI target plate. 13. Conduct mass spectrometry and analyze the MS/MS spectra (e.g., by MS/MS ion searching using Mascot (46), Matrix Science) (see Note 16).
RL131
60S RPL6 60S RPL7a
60S RPL13
2,066.2 1,783.1 1,783.1 1,661.1 1,350.9 1,355.9 1,019.6 1,147.7 1,244.8
983.6 1,320.7 1,149.7 1,444.7 1,572.8 1,862.1
1,499.8 1,838.2 1,198.7 1,456.9
1,817.1 2,112.1 1,257.8 1,570.0 1,726.1 2,209.3 1,159.7 1,180.7 1,635.1
m/z
0.26 0.16 0.16 0.21 0.14 0.10 0.09 0.10 0.14
0.06 0.03 0.08 0.04 0.05 0.03
0.13 0.27 0.11 0.13
0.12 0.05 0.08 0.11 0.11 0.20 0.04 0.09 0.15
Delta
0 1 1 1 1 1 0 1 0
0 0 1 0 1 0
0 0 0 1
1 1 1 1 2 1 2 1 0
Missc
47 73 73 41 36 42 24 16 7
43 50 25 71 75 84
58 69 34 30
65 78 49 40 22 48 38 53 51
Score
R.HTPGTFTNQMQTSFSEPR.L K.KVLQFAGIDDVFTSSR.G K.KVLQFAGIDDVFTSSR.G R.FSLKQGMKPHELVF.K.TLDKNLATSLFK.V R.LKVPPALNQFTK.T K.TWFNQPAR.K K.TWFNQPARK.T R.SLEGLQTNVQR.L
R.HGSLGFLPR.K K.FIDTASIFGHGR.F R.MFAPTKIWR.R K.AGHQTSAESWGTGR.A K.KAGHQTSAESWGTGR.A R.HAIVSAIAATAVPALVMAR.G
R.GFEGGQTALYR.R R.LGTTQSHHSLWFAQPK.K R.GFEGGQTALYR.R R.FRLDNLGPQPGSR.K
K.AGTVTANIPQAIEEFKK.G K.GTGQTVIVAVLAQGEKVDEAK.S K.SATLKLPSGEVR.L R.KKPVTPWGYPALGR.R R.KKPVTPWGYPALGRR.T R.GVVMNPVDHPHGGGEGRAPIGR.K K.DWSIKINRK.E K.TKDFLAAMQR.W R.TPLRPGGGVVFGPRPK.D
Sequence
b
a
PRP, plastid ribosomal protein. Nomenclature of PRPs is in accordance with Yamaguchi and Subramanian (15) Abbreviated Swiss-Prot accession numbers omitting “_ARATH” (e.g., RK1_ ARATH is abbreviated as RK1) c Number of missed cleavage sites in the tryptic fragment
RSSA1 RS21, RS22 RS23, RS24 RL62 RL7A1, RL7A2
40S RPSa 40S RPS2
RL4A, RL4B
60S RPL4
6b
RL31
60S RPL3
6a
RR2 RR3 RK15
RK4
50S PRPL4
30S PRPS2 30S PRPS3 50S PRPL15
RK2
50S PRPL2
5b
RK1
50S PRPL1
5a
Accession No.b
Protein IDa
Section
Table 2 Identification of Arabidopsis ribosomal proteins from the gel sections 5a, 5b, 6a, and 6b in Fig. 3
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4. Notes 1. For example, 20 g of Arabidopsis leaves yields at least 50 A260 units of total ribosomes and ~5 units of 70S-rich fraction at the first sucrose gradient. 2. Alternatively, leaves of Brassicaceae plants (closely related to Arabidopsis thaliana), such as Brassica rapa subsp. chinensis (bok-choy) and Brassica rapa var. peruviridis (komatsuna), are also applicable for plastid ribosome preparation and proteomics. These edible Brassicaceae plant leaves can easily be obtained from a fresh market (one batch is enough for one preparation). I have found that sucrose gradients of total ribosomes from bok-choy and komatsuna show similar patterns to that of Arabidopsis (see Fig. 3a), and Mascot MS/MS ion searches of in-gel digests obtained from the bok-choy 70S-rich fraction allowed efficient identification of Arabidopsis orthologs (unpublished observation). 3. The recipe for TAP medium is also available at the Web site of the Chlamydomonas Center (http://www.chlamy.org/TAP. html). Compared with cells of wild-type strains, those of strain CC-3395 (an arginine-requiring cell-wall-deficient mutant) can be readily disrupted by the homogenization protocol described here, or by the N2-bomb method (17). Steady-state cells (over 107 cells/mL) contain a lesser amount of plastid ribosomes and a greater amount of cytoplasmic ribosomes than the cells in mid-late log phase (5–8 × 106 cells/mL). 4. Unless otherwise stated, all solutions for ribosome extraction and purification should be prepared with RNase-free water (either diethyl pyrocarbonate [DEPC]-treated water or RNase/ pyrogen-free Milli-Q water purified through an ultrafiltration cartridge [e.g., BioPak, Millipore]) and RNase-free reagents (i.e., sucrose, ammonium sulfate, other salts, etc.). For convenient and reproducible preparation, I use stock solutions of each buffer component: 1 M Tris–HCl, pH 7.6; 1 M KCl; 1 M MgCl2 (store at 4°C, and can be used at least for 6 months); and, 1 M dithiothreitol (make aliquots in 0.5-mL centrifuge tubes for one-time usage and store at −20°C or −80°C). These stock solutions are also used to make other buffers (i.e., Buffer C, sucrose gradients, etc.). All the buffers should be prepared just before ribosome preparation, diluting the stock solutions with RNase-free water. Addition of polyamines (final concentrations, 0.05 mM spermine and 2 mM spermidine) to ribosome preparation buffers is helpful to prolong intactness of rRNAs (unpublished observation) and conformational stability (47, 48), though I have not tested addition of polyamines for the ribosome preparation toward proteomics.
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5. Dithiothreitol (final concentration, 5 mM) in Buffer A can be used in the place of 2-mercaptoethanol (final concentration, 7–14 mM). I prefer to use Buffer B for mid- to large-scale preparations (>200 g leaves) instead of Buffer A. 6. Alternatively, dry powder of Sephadex (e.g., Sephadex G-50, GE Healthcare, Chalfont St. Giles, UK) can be used as a solvent-absorbent powder. 7. This fixed-angle rotor is convenient, accepting 8 × 50-mL conical tubes. 8. I prefer to use a utility knife (e.g., Art Knife AK-1/5B, Olfa, Japan) rather than a generally employed scalpel or razor blade. It works nicely for precise gel excision and splitting gels into small pieces, as well as for picking up and transferring gel pieces to microfuge tubes. The knife blade must be cleaned by wiping with a 70% ethanol-sprayed KimWipe tissue (Kimberly-Clark, TX, USA). 9. Liquid nitrogen-frozen leaves can be stored at −80°C until ribosome preparation. 10. Liquid nitrogen-frozen cells can be stored at −80°C until ribosome preparation. 11. 70S ribosomes can be stored at −80°C after dissolving the ribosomal pellets in a small volume of Buffer A containing 10% (v/v) glycerol. 12. Multistep sucrose-gradient ultracentrifugation is a very effective method for obtaining highly pure chloroplast ribosomes (18). 13. Subunits can be stored at −80°C after dissolving the ribosomal pellets in a small volume of Buffer A containing 10% (v/v) glycerol. 14. Half of a band is enough for tryptic peptide preparation. The residual half can be used for other peptide preparations (e.g., other enzymatic digestions with Lys-C, Arg-C, Asp-N, etc., or chemical cleavage) if required. 15. The beaker with water is to absorb excess microwave energy. 16. Identification of small (<10 kDa) and highly basic ribosomal proteins by mass spectrometry (peptide mass fingerprinting or MS/MS ion searching) is difficult or impossible (17, 18). In such cases, N-terminal sequencing of individual ribosomal proteins using a protein sequencer (e.g., PPSQ-33A, Shimadzu, Japan) would be effective, since most proteins in plastid 70S ribosomes possess free N-termini (14, 15), unlike those of cytoplasmic 80S ribosomes (25).
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Acknowledgments I thank Dr. Alap R. Subramanian, Dr. Stephen P. Mayfield, Dr. Don P. Bourque, and Dr. Tatsuya Oda for their support and advice. I am grateful to Dr. Paul Jarvis for his encouragement and helpful comments on the manuscript. I also thank Ms. Sawako Iwanaga, Ms. Moemi Yamawaki, and Ms. Akiko Myouse for technical assistance with plant/algal culture and ribosome preparation. This work was supported by funds from the Grant-in-Aid from the Japan Society for the Promotion of Science (16770036), and in part by a Grant-in-Aid for Scientific Research from Nagasaki University, Japan. References 1. Subramanian, A. R., Stahl, D., and Prombona, A. (1991) Ribosomal proteins, ribosomes, and translation in plastids. In, The Molecular Biology of Plastids (Bogorad, L. and Vasil, I. K., eds.) Academic Press, New York, USA, pp. 191–215. 2. Bourque, D. P., Hagiladi, A., and Naylor, A. W. (1973) A method for extracting intact chloroplast and cytoplasmic ribosomal RNA from leaves. Biochem. Biophys. Res. Commun. 51, 993–999. 3. Subramanian, A. R. (1993) Molecular genetics of chloroplast ribosomal proteins. Trends Biochem. Sci. 18, 177–181. 4. Eberhard, S., Drapier, D., and Wollman, F. A. (2002) Searching limiting steps in the expression of chloroplast-encoded proteins: relations between gene copy number, transcription, transcript abundance and translation rate in the chloroplast of Chlamydomonas reinhardtii. Plant J. 31, 149–160. 5. Barkan, A., and Goldschmidt-Clermont, M. (2000) Participation of nuclear genes in chloroplast gene expression. Biochimie 82, 559–572. 6. Malnoë, P., Mayfield, S. P., and Rochaix, J. D. (1988) Comparative analysis of the biogenesis of photosystem II in the wild-type and Y-1 mutant of Chlamydomonas reinhardtii. J. Cell Biol. 106, 609–616. 7. Bruick, R. K., and Mayfield, S. P. (1999) Lightactivated translation of chloroplast mRNAs. Trends Plant Sci. 4, 190–195. 8. Somanchi, A., and Mayfield, S. P. (2001) Regulation of chloroplast translation. In, Advances in Photosynthesis and Respiration (Aro, E.-M. and Andersson, B., eds.) Kluwer Academic Publishers, Dordrecht, Netherlands, pp. 137–151.
9. Lyttleton, J. W. (1962) Isolation of ribosomes from spinach chloroplasts. Exp. Cell Res. 26, 312–317. 10. Ris, H., and Plaut, W. (1962) Ultrastructure of DNA-containing areas in the chloroplast of Chlamydomonas. J. Cell Biol. 13, 383–391. 11. Harris, E. H., Boynton, J. E., and Gillham, N. W. (1994) Chloroplast ribosomes and protein synthesis. Microbiol. Rev. 58, 700–754. 12. Beligni, M. V., Yamaguchi, K., and Mayfield, S. P. (2004) The translational apparatus of Chlamydomonas reinhardtii chloroplast. Photosynth. Res. 82, 315–325. 13. Zerges, W., and Hauser, C. (2009) Protein synthesis in the chloroplast. In, The Chlamydomonas Sourcebook, 2nd edn. (Stern D. B., ed.) Academic Press, New York, USA, pp. 967–1025. 14. Yamaguchi, K., von Knoblauch, K., and Subramanian, A. R. (2000) The plastid ribosomal proteins: Identification of all the proteins in the 30S subunit of an organelle ribosome (chloroplast). J. Biol. Chem. 275, 28455–28465. 15. Yamaguchi, K., and Subramanian, A. R. (2000) The plastid ribosomal proteins: Identification of all the proteins in the 50S subunit of an organelle ribosome (chloroplast). J. Biol. Chem. 275, 28466–28482. 16. Yamaguchi, K., and Subramanian, A. R. (2003) Proteomic identification of all plastid-specific ribosomal proteins in higher plant chloroplast 30S ribosomal subunit. Eur. J. Biochem. 270, 190–205. 17. Yamaguchi, K., Prieto, S., Beligni, M. V, Haynes, P. A, McDonald, W. H, Yates, J. R 3rd, and Mayfield, S. P. (2002) Proteomic characterization of the small subunit of Chlamydomonas
13 Plastid Ribosome Preparation for Proteomics reinhardtii chloroplast ribosome: identification of a novel S1 domain-containing protein and unusually large orthologs of bacterial S2, S3, and S5. Plant Cell 14, 2957–2974. 18. Yamaguchi, K., Beligni, M. V., Prieto, S., Haynes, P. A., McDonald, W.H., Yates, J. R. 3rd, and Mayfield, S. P. (2003) Proteomic characterization of the Chlamydomonas reinhardtii chloroplast ribosome: identification of proteins unique to the 70S ribosome. J. Biol. Chem. 278, 33774–33785. 19. Sharma, M. R., Wilson, D. N., Datta, P. P., Barat, C., Schluenzen, F., Fucini, P., and Agrawal, R. K. (2007) Cryo-EM study of the spinach chloroplast ribosome reveals the structural and functional roles of plastid-specific ribosomal proteins. Proc. Natl. Acad. Sci. USA 104, 19315–19320. 20. Manuell, A. L., Quispe, J., and Mayfield, S. P. (2007) Structure of the chloroplast ribosome: novel domains for translation regulation. PLoS Biol. 5, e209. 21. Sharma, M. R, Dönhöfer, A., Barat, C., Marquez, V., Datta, P. P, Fucini, P., Wilson, D. N., and Agrawal, R. K. (2010) PSRP1 is not a ribosomal protein, but a ribosome-binding factor that is recycled by the ribosome-recycling factor (RRF) and elongation factor G (EF-G). J. Biol. Chem. 285, 4006–4014. 22. Beligni, M. V., Yamaguchi, K., and Mayfield, S. P. (2004) Chloroplast elongation factor Ts proprotein is an evolutionarily conserved fusion with the S1 domain-containing plastid-specific ribosomal protein-7. Plant Cell 16, 3357–3369. 23. Giavalisco, P., Wilson, D., Kreitler, T., Lehrach, H., Klose, J., Gobom, J., and Fucini, P. (2005) High heterogeneity within the ribosomal proteins of the Arabidopsis thaliana 80S ribosome. Plant Mol. Biol. 57, 577–591. 24. Chang, I. F., Szick-Miranda, K., Pan, S., and Bailey-Serres, J. (2005) Proteomic characterization of evolutionarily conserved and variable proteins of Arabidopsis cytosolic ribosomes. Plant Physiol. 137, 848–862. 25. Carroll, A. J., Heazlewood, J. L., Ito, J., and Millar, A. H. (2008) Analysis of the Arabidopsis cytosolic ribosome proteome provides detailed insights into its components and their posttranslational modification. Mol. Cell. Proteomics 7, 347–369. 26. Posno, M., van Noot, M., Debise, R., and Groot, G. (1989) Isolation, characterization, phosphorylation and site of synthesis of Spinacia chloroplast ribosomal proteins. Curr. Genet. 8, 147–154. 27. Guitton, C., Dorne, A. M, and Mache, R. (1984) In organello and in vitro phosphoryla-
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40. Subramanian, A. R. (1974) Sensitive separation procedure for Escherichia coli ribosomal proteins and the resolution of high-molecularweight components. Eur. J. Biochem. 45, 541–546. 41. Mets, L. J., and Bogorad, L. (1974) Twodimensional polyacrylamide gel electrophoresis: an improved method for ribosomal proteins. Anal. Biochem. 57, 200–210. 42. Wada, A. (1986) Analysis of Escherichia coli ribosomal proteins by an improved two dimensional gel electrophoresis. I. Detection of four new proteins. J. Biochem. 100, 1583–1594. 43. Kurland, C. G. (1966) The requirements for specific sRNA binding by ribosomes. J. Mol. Biol. 18, 90–108. 44. Juan, H. F., Chang, S. C., Huang, H. C., and Chen, S. T. (2005) A new application of microwave technology to proteomics. Proteomics 5, 840–842. 45. Sun, W., Gao, S., Wang, L., Chen, Y., Wu, S., Wang, X., Zheng, D., and Gao, Y. (2006) Microwave-assisted protein preparation and enzymatic digestion in proteomics. Mol. Cell. Proteomics 5, 769–776. 46. Perkins, D. N., Pappin, D. J., Creasy, D. M., and Cottrell, J. S. (1999) Probability-based protein identification by searching sequence
databases using mass spectrometry data. Electrophoresis 20, 3551–3567. 47. Manuell, A. L., Yamaguchi, K., Haynes, P. A., Milligan, R. A., and Mayfield, S. P. (2005) Composition and structure of the 80S ribosome from the green alga Chlamydomonas reinhardtii: 80S ribosomes are conserved in plants and animals. J. Mol. Biol. 351, 266–279. 48. Agrawal, R. K., Penczek, P., Grassucci, R. A., Burkhardt, N., Nierhaus, K. H., and Frank, J. (1999) Effect of buffer conditions on the position of tRNA on the 70S ribosome as visualized by cryoelectron microscopy. J. Biol. Chem. 274, 8723–8729. 49. Rolland, N., Janosi, L., Block, M. A., Shuda, M., Teyssier, E., Miège, C., Chéniclet, C., Carde, J. P., Kaji, A., and Joyard, J. (1999) Plant ribosome recycling factor homologue is a chloroplastic protein and is bactericidal in Escherichia coli carrying temperature-sensitive ribosome recycling factor. Proc. Natl. Acad. Sci. USA 96, 5464–5469. 50. Schmitz-Linneweber, C., Maier, R. M., Alcaraz, J. P., Cottet, A., Herrmann, R.G., and Mache, R. (2001) The plastid chromosome of spinach (Spinacia oleracea): complete nucleotide sequence and gene organization. Plant Mol. Biol. 45, 307–315.
Chapter 14 The Workflow for Quantitative Proteome Analysis of Chloroplast Development and Differentiation, Chloroplast Mutants, and Protein Interactions by Spectral Counting Giulia Friso, Paul Dominic B. Olinares, and Klaas J. van Wijk Abstract This chapter outlines a quantitative proteomics workflow using a label-free spectral counting technique. The workflow has been tested on different aspects of chloroplast biology in maize and Arabidopsis, including chloroplast mutant analysis, cell-type specific chloroplast differentiation, and the proplastid-to- chloroplast transition. The workflow involves one-dimensional SDS-PAGE of the proteomes of leaves or chloroplast subfractions, tryptic digestions, online LC-MS/MS using a mass spectrometer with high mass accuracy and duty cycle, followed by semiautomatic data processing. The bioinformatics analysis can effectively select best gene models and deals with quantification of closely related proteins; the workflow avoids overidentification of proteins and results in more accurate protein quantification. The final output includes pairwise comparative quantitative analysis, as well as hierarchical clustering for discovery of temporal and spatial patterns of protein accumulation. A brief discussion about potential pitfalls, as well as the advantages and disadvantages of spectral counting, is provided. Key words: Spectral counting, Label-free quantitative proteomics, Chloroplast mutants, Chloroplast development
1. Introduction Recent improvements in sensitivity, mass accuracy, and speed of mass spectrometers (1–3) have enabled large-scale proteome quantifications at wider coverage and higher sensitivity than twodimensional gel-image-based quantification methods. We took advantage of these new developments, and using a fast and accurate mass spectrometry (MS) instrument (LTQ-Orbitrap) (4, 5), we optimized a workflow for label-free MS/MS-based quantification, also known as spectral counting (6–10) (Fig. 1). The spectral
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One LC-MS/MS LTQ-Orbitrap run for each gel band. Optional: repeat the MS analysis (technical replicate). Use ≥ 2 blank runs between each sample
a. MASCOT search-1 at 30 ppm against relevant nuclear genome + chloroplast & mitochondrial genomes b. Recalibrate precursor spectra for each individual sample (offset by ∆ppm) c. MASCOT search-2 at 6 ppm (full tryptic) and 3 ppm (semi-tryptic; not for quantification) d. Post-MASCOT cleanup: - no protein identification based on subsets of peptides nor based on semi-tryptic peptides only - minimum ion score 33 ppm (for ATH) and 6 ppm for protein identifications with ≥ 2 SPC (FDR peptides <1%) - minimum ion score 35 ppm (for ATH) and 3 ppm for protein identifications with 1 SPC (FDR peptides <1%) e. Calculate SPC, unique SPC and adjusted SPC per accession for each sample f. Calculate total SPC, total unique SPC and total adjusted SPC for each identified protein across all samples Identified proteins (including different protein models) Select best scoring protein models for each gene Identified proteins (best protein model); calculate NadjSPC and NSAF for each protein per gel lane Remove proteins with only 1 SPC Across all samples
Group proteins by similarity matrix based on shared MS/MS spectra Remove protein groups with total adjSPC <2; manual correction groups
Proteins and protein groups Annotate for function and location
Hierarchical clustering for temporal and spatial analysis (based on NadjSPC or NSAF)
Statistical analysis for determination of significant changes across treatments, genotypes and/or cell types (based on NadjSPC)
Fig. 1. Experimental and bioinformatics workflow of the proteome analysis as described step by step in this chapter. The procedure starts with SDS-PAGE separation of extracted proteomes and is completed with hierarchical clustering for spatial and/or temporal analysis, and with statistical analysis for differential accumulation. Explanation of abbreviations used: MS mass spectrometry, ATH Arabidopsis thaliana, SPC spectral counts, FDR false discovery rate, NadjSPC normalized adjusted spectral counts, NSAF normalized spectral abundance factor.
counting technique is based on the observation that the number of successful MS/MS acquisitions of peptides coming from a protein shows a positive and linear correlation to the relative concentration of this protein in the studied sample (6–8, 11). Spectral counting requires a high-resolution and fast mass spectrometer, such as the LTQ-Orbitrap (4, 12), the newer LTQ-Velos instrument (13), or the LTQ-FTICR (14). Two orthogonal separations are usually required for sufficient proteome coverage and quantification. The multidimensional separation described in this chapter is a combination of SDS-PAGE, in-gel digestion, and online reverse-phase nano liquid chromatography (nano-LC) interfaced with an LTQOrbitrap mass spectrometer. We applied this workflow on pairwise comparative proteome analysis of Arabidopsis mutants with reduced levels of chloroplast Clp protease subunits (15, 16) and comparative proteome analysis of isolated bundle sheath and mesophyll cell chloroplasts (17–19). Furthermore, we used the workflow to determine the kinetics of maize leaf development and C4 differentiation; developmental
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protein accumulation profiles and hierarchical cluster analysis established the kinetics of organelle biogenesis, formation of cellular structures, metabolism, and coexpression patterns (20). We also used this workflow to characterise MDa-sized macromolecular chloroplast stromal protein assemblies. Proteins migrating above 0.8 MDa were fractionated under nondenaturing conditions by gel filtration, followed by 1-dimensional (1D) SDS-PAGE of each of the fractions. After in-gel digestion and MS/MS analysis, normalized protein spectral counts were calculated for each of the fractions and then hierarchical clustering and protein heat maps were used to assign proteins to complexes (21). The main advantage of the label-free spectral counting technique, as compared to mass spectrometry-based quantification methods that employ stable isotope labeling techniques such as iTRAQ and ICAT, is the relative simplicity and speed, as well as the capacity to detect large fold changes. Spectral counts (SPC) have been shown to correlate well with the abundance of the corresponding protein extending over a linear dynamic range of at least two orders of magnitude for complex protein mixtures (6, 8). In addition, since SPC can be readily extracted from the MS search result files, spectral counting can be relatively straightforward. As such, due to its relative simplicity and speed, spectral counting offers a practical alternative to label-based quantification methods and to peak intensity analysis, which relies heavily on computational efforts for chromatogram alignment and peak processing (1). The disadvantage of the spectral counting technique is that small fold changes (i.e., less than ~2-fold) are harder to detect, in particular for proteins of lower abundance. We have demonstrated that it is possible to carry out quantitative comparative chloroplast analysis from total leaf extracts without actually isolating chloroplasts. This was possible because more than 1,200 of the most abundant chloroplast proteins have now been identified in Arabidopsis from a large number of (independent) chloroplast proteomics studies supplemented by many detailed experimental papers for individual plastid-localized proteins (10). Importantly, this set of chloroplast proteins was established after manual curation of all available data and updated versions can be found in the Plant Proteome Database, PPDB (http://ppdb.tc.cornell.edu). The advantages of characterizing quantitative effects on the chloroplast proteome through analysis of total leaf extracts, rather than analysis of isolated chloroplasts, are as follows: (1) mutants with strong growth defects can be analyzed; isolation of chloroplast from such mutants can be very hard or even practically impossible; (2) more accurate results are obtained for chloroplast mutants with heterogeneity in their leaf phenotype (often with strongest phenotypes in the youngest leaves), which is most pronounced in the variegated mutants (22); isolation of chloroplasts from
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such leaves could result in selection of a subset of chloroplast phenotypes, not representing the overall chloroplast population. Our workflow has sufficient sensitivity and throughput rate to detect and quantify a high number of chloroplast proteins. Thus, it is far more practical if one can assess the chloroplast proteome without actual isolation of organelles, with the added benefit that the extrachloroplastic responses can also be determined.
2. Materials 1. All chemicals should be of the highest degree of purity available, and be freshly prepared. 2. Extraction buffer (EB) for total leaf proteomes: 50 mM Tris– HCl, pH 8.0, 2% (w/v) sodium dodecyl sulfate (SDS), and protease inhibitor cocktail (50 mg/mL antipain, 40 mg/mL bestatin, 20 mg/mL chymostatin, 10 mg/mL E64, 10 mg/mL phosphoramidon, 50 mg/mL pefabloc SC, 2 mg/mL aprotinin) (see Note 1). 3. 0.8-mL centrifuge columns (pore size 30 mm) with microcentrifuge receiver tubes (Thermo Scientific, part numbers 89868 or 89869). These are used for efficient removal of unsolubilized leaf material during proteome extraction. 4. BCA assay kit (Thermo Scientific) for protein quantification compatible with SDS (in EB) but not with reductants. Instead of purchasing a commercial kit, the components can be made quite easily from individual chemicals as detailed in ref. 23. 5. 2× Laemmli solubilization buffer: 125 mM Tris–HCl, pH 6.8, 10% (v/v) b-mercaptoethanol, 20% (v/v) glycerol, 4% (w/v) SDS, plus a few grains of bromophenol blue. 6. Coomassie (de)staining and gel storage. Fixing solution: 50% (v/v) ethanol, 5% (v/v) acetic acid; use this for fixation of the proteins in gels, and removal of SDS to avoid interference with the MS analysis. Coomassie staining solution: 50% (v/v) ethanol, 5% (v/v) acetic acid, and 0.04% (w/v) Coomassie R-250. Destaining solution: 30% (v/v) ethanol, 5% (v/v) acetic acid. Gel storage solution: 1% (v/v) acetic acid. 7. Peptide extraction solutions and materials. 500-mL or 1.5-mL Eppendorf microtubes should be washed before use: soak in an excess volume of 50% (v/v) acetonitrile, 0.1% (v/v) trifluoroacetic acid (TFA) for 30–60 min. After the wash, the tubes
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should be dried in a laminar flow hood. The wash removes polymers that can interfere with MS analyses. 8. Vacuum concentrator. This instrument must be kept free of keratins. 9. Trypsin stock solution and reagents for reduction and alkylation. Trypsin digestion solution: dissolve 20 mg trypsin in 10 mM NH4HCO3, 10% (v/v) acetonitrile to a final concentration of 12.5 ng/mL. We use Promega Trypsin (product code V5111; 5 vials, 20 mg per vial); one vial is typically sufficient for 20–40 gel slices. Alternatively, if less trypsin is needed, dissolve the trypsin in 1 mM HCl at 100 ng/mL and store aliquots at −20°C; dilute the trypsin stock eightfold to 12.5 ng/mL using 10 mM NH4HCO3, 10% (v/v) acetonitrile shortly before use (check that the pH is close to 7.8). Reducing solution: 10 mM dithiothreitol (DTT), 0.1 M NH4HCO3; make this shortly before use. Alkylating solution: 55 mM iodoacetamide, 0.1 M NH4HCO3; make this shortly before use and keep it in the dark. 10. C18 microcolumns for sample cleanup. If needed, clean and concentrate the peptide extracts using C18 ZipTips (Millipore, product code ZTC18S096, micro 0.6 mL) or hand-made microcolumns using gel loading tips with Poros R2 (50-mm bead diameter) which can be purchased in bulk (Applied Biosystems) (24). 11. A high-throughput, high accuracy, high resolution reversephase LC nano electrospray ionization (RP-LC-nanoESI) tandem mass spectrometer. The workflow presented in this chapter was developed and tested with an LTQ-Orbitrap mass spectrometer (Thermo Scientific), interfaced with a nano-LC system (Surveyor MS Pump, Thermo Scientific) with a precolumn splitter, C18 guard column, and C18 analytical column. Samples are loaded using a Micro AS Autosampler (Thermo Scientific). 12. In-house search engine for matching of mass spectral data to predicted protein sequences. For the workflow developed in this chapter, we used a licensed copy of Mascot (Matrix Science; http://www.matrixscience.com/). 13. Hardware and software for postprocessing. Any PC or Macintosh computer with Microsoft Excel software. Hierachical cluster analysis (correlation metric) as described in our workflow is done in MatLab (http://www.mathworks. com/products/matlab/), but alternative software solutions do exist. The recalibration script (in Perl) and the Perl script for filtering results after Mascot searches are available upon request.
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3. Methods 3.1. Total Leaf Proteome Extraction
The workflow described in this chapter is summarized in Fig. 1. This is a quantitative extraction method that extracts hydrophobic and soluble proteins. 1. Grinding. Grind the frozen leaf tissue in liquid nitrogen into a very fine powder using a precooled pestle and mortar. We typically use between 150 and 300 mg of fresh leaf weight for Arabidopsis, providing a yield of ~10 mg protein per 1 mg fresh-weight (as determined in the assay in step 3 below) (see Note 2). 2. Extraction. Transfer the powder into an Eppendorf microtube, and add ice-cold EB. Use 2 mL of EB per ~1 mg of freshweight leaf material; do not use more than 150–300 mg of fresh leaf material, corresponding to 300–600 mL of EB. Vortex the sample for 20 s, leave it on ice for 5 min, transfer it into the 0.8-mL centrifuge columns, and then spin in a microcentrifuge (1 min, 10,000 × g at 4°C) (see Note 3). 3. Collection. Collect the flow-through (the extracted leaf proteome) and transfer it to a new microtube. Avoid the white pellet (starch). Determine the protein concentration of the sample using the BCA assay (see Note 4).
3.2. Gel Electrophoresis
1. Select optimal gel. We typically use 10.5–14% acrylamide gradient gels (13.3-cm wide × 8.7-cm long; 1-mm thick) with 12 sample wells, either made in the laboratory (see Note 5) or precast by a commercial supplier (e.g., Bio-Rad Criterion Tris–HCl gels, 12 + 2 wells, 45 mL well volume; product code 345–9949). For optimal resolution, protein identification, and quantification, load 50–75 mg proteins per lane, corresponding to 0.1–0.2 mg protein/mm3 of gel lane volume. Higher proteins loads are not beneficial, since they result in decreased gel resolution and likely decrease the number of protein identifications due to oversampling of the most abundant proteins (see Note 5). 2. Solubilization. Mix the sample (in EB) with 2× to 4× (as needed) Laemmli solubilization buffer (including fresh reductant), vortex, and then keep at room temperature for 10 min. Flash-freeze any unused sample in liquid nitrogen and store in −80°C. Heat the sample to be loaded at 75°C using a heat block or water bath for 5 min (see Note 6). Spin down the sample in a microcentrifuge (10 min, ~14,000 × g at 4°C) to pellet starch, unsolubililized material, and aggregates prior to gel loading.
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3. Gel loading. The proteome samples that will be compared should be run on the same gel in neighboring lanes to minimize variations between quantitative proteomics results. 4. Electrophoresis. For complex proteomes such as the leaf, run the sample the whole length of the gel for maximum resolution and best proteome coverage and quantification. When running the gel, do not allow it to overheat but run it fast enough to avoid diffusion that will reduce gel resolution. In the case of the suggested commercial gels, run at ~30 mA/ gel (constant current) for about 2 h at room temperature or in the cold room. 5. Staining. After the electrophoresis run, incubate the gel in fixing solution at room temperature with gentle mixing (3 × 30 min incubations, with fresh solution each time). Stain the gel in Coomassie staining solution for at least 3 h at room temperature (for best results, stain for >12 h) and then destain accordingly (see Note 7). Store the gel in gel storage solution at 4°C. 3.3. Gel Processing, Peptide Extraction, and Cleanup
1. Gel band excision. After staining and destaining, wash the entire gel once or twice with water for 15 min at room temperature, and then put a clear plastic tray containing the gel onto a light box and excise each lane in ~10–20 slices with a clean scalpel (be careful to avoid keratin contamination; see Note 8). Cut the gel bands into smaller pieces (ca. 1 × 1 mm) but do not macerate them. Note that smaller pieces can clog pipette tips. Transfer the gel pieces into prewashed microcentrifuge tubes (500 mL) and spin them down briefly. For complex samples, cut each gel lane into at least ten slices (12–16 slices per lane is typical). When excising gel lanes, try to (1) keep each highly abundant protein (e.g., Rubisco large subunit, RBCL) within one gel slice, (2) separate the highly crowded areas into more bands than areas with fewer visible bands, (3) cut the low molecular mass region (below ~20 kDa) into wider gel slices (see Note 5). 2. Washing the gel pieces. Wash the gel pieces once with water (100 mL) for 5 min at room temperature. Remove all liquid and shrink the gel pieces by adding ca. 400 mL of 100% acetonitrile (use 3–4 times the volume of the gel pieces) and incubating at room temperature for 10–15 min. Collect the gel pieces by briefly (~5 s) spinning them in a microcentrifuge, and then remove all the liquid; you can, but do not have to, dry the gel pieces in the vacuum centrifuge (15 min is typically enough, since acetonitrile is volatile and the volumes are small). 3. Reduction and alkylation. Add a sufficient volume of reducing solution (containing DTT) to completely cover the gel pieces (i.e., 30–50 mL, depending on gel volume), and incubate for
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30 min at 56°C to reduce the proteins. Cool the tubes to room temperature, add ca. 400 mL of acetonitrile, incubate at room temperature for 10 min, and then spin down briefly in a microcentrifuge before removing all liquid. Add enough alkylating solution (containing iodoacetamide) to cover all the gel pieces and incubate in the dark at room temperature for 20 min. Spin down briefly, remove the liquid, and then shrink the gel pieces with ca. 400 mL of acetonitrile; spin down again and remove all the liquid. Wash the gel pieces at least twice with ~100 mL of a 0.1 M NH4HCO3–100% acetonitrile 1:1 (v/v) mixture, to remove residual iodoacetamide; incubate each change for 15 min on the bench with occasional vortexing. Finally, spin down again, remove all the liquid, and dry the gel pieces in the vacuum centrifuge (15 min is typically enough, since acetonitrile is volatile and the volumes are small). 4. In-gel digestion. Rehydrate the dry gel pieces in about 50 mL of freshly prepared and chilled trypsin digestion solution at ice. Make sure that the buffer covers the gel pieces, and then incubate on ice for 30 min. Check if all solution is absorbed and add more trypsin digestion solution if the gel pieces are not completely covered with solution (in the case of silver-stained gels and/or lower protein loads per lane, top-up with digestion solution lacking trypsin to avoid overly high trypsin concentrations). Leave the gel pieces for another 90 min and then add 10–20 mL of chilled 10 mM NH4HCO3. Incubate overnight at 37°C (use of an air circulation thermostat is recommended). During the digestion, it is important to avoid a temperature gradient between the bottom and the lid of the tube, to prevent premature dehydration of the gel pieces (because of water condensation on the lid). The completed digest with gel pieces can be stored at −20°C if needed. 5. Extraction of peptides. Cool the tube(s) to room temperature and spin down the gel pieces using a microcentrifuge. (Optional: withdraw 0.5–1 mL aliquots directly from the digest for matrixassisted laser desorption/ionization time-of-flight peptide mass finger printing [MALDI-TOF PMF] to verify digestion and estimate peptide concentration). Transfer the digest solutions to clean 500-mL prewashed tubes. To the gel pieces, add 50–100 mL (enough to cover the gel pieces) of a 100% acetonitrile–5% formic acid 1:1 (v/v) mixture; vortex briefly, incubate, and spin down, and then transfer the supernatant to a fresh prewashed microtube. Repeat twice more and pool the supernatants together. Then, add 50–100 mL of 100% acetonitrile to the gel pieces, spin down, and add the supernatant to the previously combined supernatants. Dry the pooled supernatants (typically between 225 and 300 mL) down in the vacuum concentrator (this step can take several hours depend-
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ing on the efficacy of the vacuum concentrator; see Note 9). For storage, stop here and keep the dry pellets at −20°C; dried extract can be safely stored at −20°C for a few months. For immediate MS/MS analysis, redissolve each pellet in 15–20 mL of 2% formic acid, vortex two times for 60 s, and then spin down for ~5 s (desalting and concentration may optionally be conducted at this stage; see step 6 below). Transfer the supernatant liquid (containing the peptides) to a 96-well plate that fits the autosampler, using fine gel loading tips, to avoid transferring small gel pieces which could clog the needle of the autosampler injector (see Note 10). 6. Desalting and concentration of peptides prior to LC-MS/MS (Optional). For total leaf proteome analysis, we recommend cleanup of the peptide extracts using C18 ZipTips (Millipore) or hand-made microcolumns filled with Poros R2. The cleanup procedure should be conducted at room temperature and is as follows (employing a 20 mL pipettor). Step 1: (equilibration): rinse the tip four times with 10 mL of 100% acetonitrile, and then wash it two times with 10 mL of 2% formic acid. Step 2: load the sample in 2% formic acid (from the penultimate step in the extraction procedure above; point 5) by aspirating (pipetting up and down) 7–10 times. Step 3: wash the column twice with 10 mL of 2% formic acid. Step 4: elute the peptides, first with 2 × 10 mL of 50% acetonitrile, 0.1% formic acid, and second with 2 × 10 mL of 90% acetonitrile, 0.1% formic acid (for peptides with higher affinity to the matrix). In addition to concentrating the peptides and removing salts, this off-line purification step also helps to (1) remove very hydrophilic peptides with low affinity for the C18 matrix of Poros R2 and (2) capture gel fragments and detergents, as well as very hydrophobic and/or long peptides that would not elute from the analytical C18 column. We also use ZipTips for hydrophobic proteomes, i.e., total thylakoid or cellular membranes, and for samples from first-dimension native gels in which detergents were used (1D blue native [BN]-PAGE) (17). 3.4. Nano-Capillary Reverse-Phase LC MS/MS Using an LTQ-Orbitrap Mass Spectrometer
Peptide extracts resuspended in ~15 mL of 2% formic acid are subjected to MS/MS analysis by reverse phase nano-LC-ESI-LTQOrbitrap (Thermo Scientific). We typically inject 5–6.4 mL for each run; if the peptides are in ³15 mL, this allows duplicate MS/MS analyses (technical replicates) to be run. Analysis by nano-LCLTQ-Orbitrap is done as follows (see Note 11). 1. Using automated sample pickups, load 5–6.4 mL of peptide extracts at 20 mL/min for 6 min (this can be shorter or longer depending on the dead volume) on a guard column (LC Packings; MGU-30-C18PM), followed by elution and separation on a PepMap C18 reverse-phase nano column (LC
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Packings; nan75-15-03-C18PM), using gradients with 95% water, 5% acetonitrile, 0.1% formic acid (solvent A), and 95% acetonitrile, 5% water, 0.1% formic acid (solvent B) at a column flow rate of 200 nL/min. Use a precolumn splitter to reduce the flow rate of ~80 mL/min (this flow rate depends on dead volumes and back pressures) to 200 nL/min (see Note 12). 2. Each sample injection and analysis should be followed by two blank injections using 60 min gradients to prevent carry-over from sample to sample. 3. The duration of the gradients should depend on the sample complexity. For highly complex samples (i.e., leaf extract), we typically perform peptide separation using a 120 min gradient, which contains an internal blank, followed by two 60-min blanks. For lower complexity samples or less abundant samples (i.e., silver-stained bands, coimmunoprecipitates), we typically use a shorter gradient (i.e., 85 min with internal blank) followed by two shorter blanks (i.e., 45 min). 4. The acquisition cycle should consist of a survey scan in the Orbitrap analyzer at the highest resolving power (100,000), followed by five data-dependent MS/MS scans in the LTQ analyzer. Use dynamic exclusion with the following parameters: exclusion size 500, repeat count 2, repeat duration 30 s, exclusion time 180 s, and exclusion window ±6 ppm. Target values should be set at 5 × 105 and 3 × 104 for the survey and tandem MS scans, respectively; set the maximum ion accumulation time at 200 ms for both analyzers (LTQ and Orbitrap). 5. When separating peptides extracted from a 1D gel, analyze them sequentially from top to bottom (according to the gel lane) and add extra blanks between different gel lanes. 3.5. Processing of the MS Data, Database Searches, and Additional Data Processing
1. Generation of peak lists. From the RAW files, generate peak lists (*.mgf format) for database searching using DTASuperCharge (version 1.19) software (http://msquant.sourceforge.net/). 2. Create searchable database. Create the maize database (currently version 4a.53 with 53,764 entries; http://www.maizesequence.org/index.html) or the Arabidopsis database (version 8 with 33,013 entries; http://www.arabidopsis.org/). These databases must include the plastid and mitochondria-encoded proteins and sequences for known contaminants (e.g., keratin and trypsin). In the case of maize, download the organellar database from http://megasun.bch.umontreal.ca/ogmp/projects/other/cp_list.html. In the case of Arabidopsis, these organellar genomes are already part of the public ATH database (proteins encoded by the plastid and mitochondrial genomes can be recognized as their accession numbers start with ATCG and ATMG, respectively). For determination of
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false discovery rates (FDRs) during peptide and protein identification in step 5 below, concatenate a decoy database where all the sequences are in reverse orientation using a script that can be downloaded from the Matrix Science Web site ( h t t p : // w w w. m a t r i x s c i e n c e . c o m / h e l p / d e c o y _ h e l p . html#MANUAL). The resulting final database (target plus decoy) should include 108,452 and 66,025 entries for maize and Arabidopsis, respectively. 3. Recalibration. Perform a preliminary database search with Mascot (version 2.2.0) using a broad precursor tolerance window set at ±30 ppm. Recalibration should be performed using a Perl script, which adjusts precursor masses in the peak lists (see Subheading 2). 4. Database searches. The recalibrated peak lists should be searched once more against the database of interest. For each of the peak lists, one should typically perform two searches: (1) tryptic search with the precursor ion tolerance window set at ±6 ppm; and (2) semitryptic search with N-terminal acetylation (this is an enzymatic reaction that occurs in situ) or N-terminal glutamine to pyroglutamate conversion (N-terminal glutamine residues can spontaneously cyclize to become pyroglutamate) set as variable modifications. For both types of search, the first and second 13C peaks should be considered as precursors without widening of the precursor ion tolerance, using the corresponding Mascot 2.2.0 feature. Additionally, in both searches set methionine oxidation as a variable modification and allow two missed cleavages. One may optionally carry out a third, error-tolerant search with the precursor ion tolerance window set at ±3 ppm (25). For quantification, use only the results from the tryptic search (1). The semitryptic searches (2) help to increase sequence coverage, in particular for proteins that are N- or C-terminally processed, such as proteins with N-terminal chloroplast or mitochondrial transit peptides, or signal peptides for delivery to the endoplasmic reticulum, or those with C-terminal peptides for delivery to the peroxisomes. 5. Postsearch filtering. Using an in-house written filter, combine the results of the two or three types of Mascot searches in an Excel spreadsheet. For identification with two or more peptides, the minimum ion score threshold should be set to 33 for the Arabidopsis database, and to 30 for the maize database. For protein identification based on a single peptide, the minimum ion score threshold should be set to 35 for the Arabidopsis database, and to 40 for the maize database, and the mass accuracy of the precursor ion is required to be within ±3 ppm. Calculate the peptide false discovery rate (FDR) as: 2 × (decoy_ hits)/(target + decoy hits). We obtain a final peptide FDR below 1% at the aforementioned ion score thresholds and post-
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Mascot filtering. Only matches from the tryptic search should be considered in protein identifications and quantifications. The FDR of proteins identified with two or more peptides is zero. Peptides with less than seven amino acids should be discarded. Submit mass spectral data (the *.mgf files reformatted as PRIDE XML files) to the PRIDE database at http://www.ebi.ac.uk/pride/. 3.6. Determination of Relative Protein Abundance and Selection of the Best Gene Models
1. Quantification of protein abundance by spectral counts. For each identified protein, calculate the total number of spectral counts (SPC) and the number of unique (not shared with other proteins) SPC. In cases where two or more homologous proteins are identified with shared and unique peptides, the number of spectra from shared peptides assigned to each protein should be determined based on the ratios of spectra derived from the unique peptides that identified each protein, using the following formula: AFi = uSPC*i + Shared _ SPC ×
uSPC*i , ∑ uSPC*j j
where uSPC*i is the initial number of unique SPC for ith protein in the group, AFi is the abundance factor used for quantification, Shared_SPC is the number of total shared SPC within the group, and ∑ j uSPC*j is the sum of all unique SPC within the group. In cases of ambiguous groups (with no unique SPC), as well as in the cases of protein subsets, AF was set as equal to Shared_SPC, with the whole ambiguous group treated as one protein. 2. Selection of highest scoring gene models. Many Arabidopsis or maize genes have multiple predicted protein models. Only the protein models with the highest total adjSPC across all samples in a certain experiment are analyzed. If the protein models do not differ in total adjSPC, select protein model 1 (a “model 1” is present for each protein-encoding gene). 3. Normalized relative mass (NadjSPC) or relative concentration (NSAF). To calculate the relative abundance for each protein per gel lane, divide the total adjSPC by the number of observable tryptic peptides within the mass range 700–3,500 Da (with the predicted transit peptide removed), yielding the spectral abundance factor (SAF). Then, normalize the SAF values to the total SAF of proteins identified in the gel lane, yielding normalized spectral abundance factors (NSAFs). The normalized adjSPC (NadjSPC) for each protein is calculated through division of adjSPC by the sum of all adjSPC values for the proteins from that gel lane. NadjSPC provides a relative
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protein abundance measure by mass, whereas NSAF estimates relative protein concentration within a particular sample. 4. Grouping of proteins. Proteins that share more than about 80% of their matched peptides with other proteins across the complete dataset should be grouped into clusters; this is done by generating a similarity matrix through calculation of the dice coefficient between each pair of identified proteins. The dice coefficient is defined as s = 2 |X∩Y|/(|x| + |Y |), where x and y represent the SPC number for each protein. The similarity cut-off should be set at 0.80 (based on our experience). The MCL software (26) is used to cluster the proteins into groups, with the inflation value set at 5. All groups are manually verified and ungrouped if needed. Additional proteins are grouped manually as needed, in particular if they had a low adjSPC number (e.g., less than 20). The linkage of homologous proteins that are identified by the same set of MS/MS spectra should be recorded (by marking their identifications as “ambiguous”), and the matched MS/MS spectra should be marked as “shared” spectra (or “not unique”). For proteins identified with unique spectra, as well as shared spectra, a linkage with the related identified protein should be recorded (by marking them as “related” proteins). 3.7. Correction for the Reduced Amount of Chloroplast Proteins, When Specifically Assessing the Chloroplast Proteome in Mutants with Strong Phenotypes
In case of chloroplast mutants with strong phenotypes, the chloroplast proteome in the mutant can represent a smaller percentage of the total leaf proteome mass. This reduced chloroplast protein mass is often due to loss of abundant thylakoid and Calvin cycle proteins, and possibly also due to an overall lower chloroplast volume per cell, as we observed for the clpr4-1 mutant with complete loss of the ClpR4 protein (15). To better understand the chloroplast proteome, it will be necessary to correct for the overall decrease in chloroplast proteome in the mutant. To that end, first calculate the SAF for each chloroplast-localized protein in the wild type (or other genotypes that are part of the comparison) and in the mutant. Then, recalculate NSAF using only the SAFs from the chloroplast-localized proteins. In this way, one can compare the wild-type and mutant chloroplast proteins directly without any influence of an overall change in the amount of chloroplast protein per leaf proteome.
3.8. Statistical Analyses
We have used the G-test (10, 15, 16) for significance analyses of pairwise comparisons based on SPC data using the workflow presented in this chapter. To determine significance (95% confidence) of upregulated and downregulated proteins, first the G-test (27) should be used for each biological replicate. For each protein across m conditions tested, calculate the G-statistic and then compare it to the Chi-square distribution with m − 1 degrees of freedom,
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resulting in a p-value. The p-values determined by G-testing should be corrected for multiple hypotheses testing using the Benjamini– Hochberg method (28). Briefly, p-values are ranked from 1 to N, where 1 corresponds to the lowest p-value and N to the highest p-value; a protein with p-value of rank k is said to pass the G-test at 95% significance, if inequality p-value < (k/N) × 0.05 holds. Subsequently, accessions are deemed significantly upregulated or downregulated only if (1) they pass the G-test in both biological replicates at 95% confidence, and (2) they are consistently either up or down in all biological replicates. Whereas the G-test is useful, and one of only a few tools currently available for significance tests of pairwise differences, the G-test is rather limited and better statistical tools need, therefore, to be developed. 3.9. Hierarchical Clustering Analyses
To group proteins with similar accumulation patterns across a certain experiment (i.e., when comparing two different cell types), we employ hierarchical clustering using the Statistics toolbox of MATLAB version 7 (Mathworks, Inc.). We derive the linear correlation (r) between every pair of proteins with NAdjSPC distribution across the experiment, X1, …, Xn and Y1, …, Yn where n is the number of sample types (e.g., cell types) as follows: rXY =
∑ ∑
n j =1
n j =1
(X j − X )(Y j − Y )
(X j − X )2 ∑ j =1 (Y j − Y )2 n
This is then converted into a distance measure, DXY = 1 – rXY. Protein pairs with higher correlations have smaller distance values. A linkage map based on the average distance among protein pairs can be constructed to yield a hierarchical cluster tree (dendrogram). An example of a simple dendrogram is shown in Fig. 2, which depicts the distribution of 30S and 50S chloroplast ribosomal proteins, (co-) translational factors, and ribosome biogenesis factors separated by size-exclusion chromatography; this is adapted from ref. 21.
4. Notes 1. No reductant is added, since it will interfere with protein quantification. 2. You can stop at this step and store the dry powder at −80°C for future extractions.
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Fig. 2. Dendrogram of 30S and 50S chloroplast ribosomal proteins, (co-) translational factors, and ribosome biogenesis factors separated by size-exclusion chromatography. Hierachical clustering of 53 assigned ribosomal proteins, (co-) translational factors, and ribosome biogenesis factors with 20 or more adjSPC. Three clusters were distinguished, with cluster A representing the 30S particle, cluster B the 50S particle, and cluster C the translating 70S ribosome. Native mass ranges of the five selected size exclusion fractions 1–5 are, respectively, >5, 3–5, 2–3, 1–2, and 0.8–1 MDa. The colors reflect the relative abundance distribution of each protein across the fractions, with red the most enriched and green the least enriched. For more details, see ref. 21.
3. Avoid longer incubations, as this may result in protein loss due to proteolysis. 4. Include EB in the matrix during protein quantification, since protease inhibitors contribute to the protein signal. Typical concentrations for protein concentrations range from 2 to 10 mg/mL. 5. Several considerations should be made when choosing the type and size of gel, including the width of the gel lane (comb), how far to run the proteins into the gel, and the number of cut
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slices. This is clearly a compromise between protein amount and sensitivity, gel processing time, and MS analysis. Different gel dimensions are possible but aim to obtain about 0.1–0.2 mg protein/mm3 of gel lane volume. Minimize the amount of gel per amount of protein while still having effective protein separation and reduction of proteome complexity. We have noted that washing and digestion of large gel slices or pooled gel slices in large 1.5-mL Eppendorf tubes is not effective. It is better to cut small gel slices and to pool some of the final peptide extracts at the end of the digestion and extraction process. If using homemade gels, an extra caution has to be taken to let the gel polymerize completely as failure to do so might lead to creation of covalent peptide acrylamide adducts (25). 6. Boiling the samples is not recommended, as it may lead to aggregation of membrane proteins. 7. For less abundant proteins, use mass spectrometry-compatible silver stain (which uses formalin solution instead of glutaraldehyde) (29) or SYPRO Ruby (a fluorescent stain from Molecular Probes) as an alternative to Coomassie staining. However, for large-scale spectral count analysis we strongly recommend using sufficient protein amounts (50–75 mg). 8. To avoid contamination by human keratin (very abundant proteins in skin and hair), take several precautions, as follows: (1) never touch the gel, gel boxes, or other materials that come in direct contact with the gel with bare hands; instead, always use clean, powder-free gloves; (2) wash and rinse the gel equipment and gel boxes with water and soap, and then rinse with ethanol; (3) use a lab coat; (4) cover your hair with a hair net; and (5) keep a clean pipette just for protein digests. These precautions are not important after you have completed the peptide extraction, but they are very important in the immediate steps prior to adding the trypsin solution. 9. It is beneficial to carry out the drying step as quickly as possible without heating the samples. A powerful pump resulting in high vacuum, and a cold trap that is as cold as possible, all contribute to shorter drying times. Shorter drying times and absence of heating will help to reduce nonphysiological posttranslational modifications (PTMs) (25). 10. For maximum protein/peptide recovery, minimize losses from adsorption to plastics including pipettes and walls of the tubes. Keep the volumes as low as possible (i.e., during extraction) and choose the smaller 500-mL tubes instead of the bigger 1.5-mL tubes. In this way, the peptides will be exposed to a smaller surface area. Ideally no PTMs are introduced during protein/peptide storage and digestion. Also, for final resuspension of the peptides prior to MS/MS analysis, aim to keep the volume small, for example between 15 and 20 mL.
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11. The setup described here has proven to be very robust and allows for fully automated acquisition for up to 4 × 24 h. Alternative setups, e.g., without a guard column but with an autosampler, or which preload samples using a “pressure bomb” are possible, but require much more manual intervention. 12. The analytical column (C18) is protected by a guard column (C18). Therefore, minor amounts of salts do not pose a problem for the LC system. However, gel fragments are a huge problem for the autosampler and LC system, as they clog the in-line filters and capillaries/tubes/connections. Detergents reduce the resolution of the chromatography, suppress ionization, and give background signals in the mass spectrometer. References 1. Bantscheff, M., Schirle, M., Sweetman, G., Rick, J., and Kuster, B. (2007) Quantitative mass spectrometry in proteomics: a critical review. Anal. Bioanal. Chem. 389, 1017–1031. 2. Mann, M., and Kelleher, N. L. (2008) Precision proteomics: the case for high resolution and high mass accuracy (Special Feature). Proc. Natl. Acad. Sci. USA 105, 18132–18138. 3. Domon, B., and Aebersold, R. (2010) Options and considerations when selecting a quantitative proteomics strategy. Nat. Biotechnol. 28, 710–721. 4. Hu, Q., Noll, R. J., Li, H., Makarov, A., Hardman, M., and Cooks, G. R. (2005) The Orbitrap: a new mass spectrometer. J. Mass Spectrom. 40, 430–443. 5. Makarov, A., Denisov, E., Lange, O., and Horning, S. (2006) Dynamic range of mass accuracy in LTQ Orbitrap hybrid mass spectrometer. J. Am. Soc. Mass. Spectrom. 17, 977–982. 6. Liu, H., Sadygov, R. G., and Yates, J. R., 3rd (2004) A model for random sampling and estimation of relative protein abundance in shotgun proteomics. Anal. Chem. 76, 4193–4201. 7. Zybailov, B., Coleman, M. K., Florens, L., and Washburn, M. P. (2005) Correlation of relative abundance ratios derived from peptide ion chromatograms and spectrum counting for quantitative proteomic analysis using stable isotope labeling. Anal. Chem. 77, 6218–6224. 8. Old, W. M., Meyer-Arendt, K., Aveline-Wolf, L., Pierce, K. G., Mendoza, A., Sevinsky, J. R., Resing, K. A., and Ahn, N. G. (2005) Comparison of label-free methods for quantifying human proteins by shotgun proteomics. Mol. Cell. Proteomics 4, 1487–1502.
9. Lu, P., Vogel, C., Wang, R., Yao, X., and Marcotte, E. M. (2007) Absolute protein expression profiling estimates the relative contributions of transcriptional and translational regulation. Nat. Biotechnol. 25, 117–124. 10. Zybailov, B., Rutschow, H., Friso, G., Rudella, A., Emanuelsson, O., Sun, Q., and van Wijk, K. J. (2008) Sorting signals, N-terminal modifications and abundance of the chloroplast proteome. PLoS ONE 3, e1994. 11. Sandhu, C., Hewel, J. A., Badis, G., Talukder, S., Liu, J., Hughes, T. R., and Emili, A. (2008) Evaluation of data-dependent versus targeted shotgun proteomic approaches for monitoring transcription factor expression in breast cancer. J. Proteome Res. 7, 1529–1541. 12. Scigelova, M. and Makarov, A. (2006) Orbitrap mass analyzer - overview and applications in proteomics. Proteomics 6 (Suppl. 2), 16–21. 13. Olsen, J. V., Schwartz, J. C., Griep-Raming, J., Nielsen, M. L., Damoc, E., Denisov, E., Lange, O., Remes, P., Taylor, D., Splendore, M., Wouters, E. R., Senko, M., Makarov, A., Mann, M., and Horning, S. (2009) A dual pressure linear ion trap Orbitrap instrument with very high sequencing speed. Mol. Cell. Proteomics 8, 2759–2769. 14. Syka, J. E., Marto, J. A., Bai, D. L., Horning, S., Senko, M. W., Schwartz, J. C., Ueberheide, B., Garcia, B., Busby, S., Muratore, T., Shabanowitz, J., and Hunt, D. F. (2004) Novel linear quadrupole ion trap/FT mass spectrometer: performance characterization and use in the comparative analysis of histone H3 post-translational modifications. J. Proteome Res. 3, 621–626. 15. Kim, J., Rudella, A., Ramirez Rodriguez, V., Zybailov, B., Olinares, P. D., and van Wijk, K. J.
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(2009) Subunits of the plastid ClpPR protease complex have differential contributions to embryogenesis, plastid biogenesis, and plant development in Arabidopsis. Plant Cell. 21, 1669–1692. 16. Zybailov, B., Friso, G., Kim, J., Rudella, A., Rodriguez, V. R., Asakura, Y., Sun, Q., and van Wijk, K. J. (2009) Large scale comparative proteomics of a chloroplast Clp protease mutant reveals folding stress, altered protein homeostasis, and feedback regulation of metabolism. Mol. Cell. Proteomics 8, 1789–1810. 17. Majeran, W., Zybailov, B., Ytterberg, A. J., Dunsmore, J., Sun, Q., and van Wijk, K. J. (2008) Consequences of C4 differentiation for chloroplast membrane proteomes in maize mesophyll and bundle sheath cells. Mol. Cell. Proteomics 7, 1609–1638. 18. Majeran, W., Cai, Y., Sun, Q., and van Wijk, K. J. (2005) Functional differentiation of bundle sheath and mesophyll maize chloroplasts determined by comparative proteomics. Plant Cell 17, 3111–3140. 19. Friso, G., Majeran, W., Huang, M., Sun, Q., and van Wijk, K. J. (2010) Reconstruction of metabolic pathways, protein expression, and homeostasis machineries across maize bundle sheath and mesophyll chloroplasts: large-scale quantitative proteomics using the first maize genome assembly. Plant Physiol. 152, 1219–1250. 20. Majeran, M., Friso, G., Ponnala, L., Connolly, B., Huang, M., Reidel, E., Zhang, C., Asakura, Y., Bhuiyan, N. H., Sun, Q., Turgeon, R., and van Wijk, K. J. (2010) Structural and metabolic transitions of C4 leaf development and differentiation defined by microscopy and quantitative proteomics. Plant Cell, in press. 21. Olinares, P. D., Ponnola, L., and van Wijk, K. J. (2010) Megadalton complexes in the chloroplast stroma of Arabidopsis thaliana characterized by size exclusion chromatography, mass
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Chapter 15 Use of Phosphoproteomics to Study Posttranslational Protein Modifications in Arabidopsis Chloroplasts Anne Endler and Sacha Baginsky Abstract The posttranslational modification of proteins is important for the regulation of enzymatic activity, protein half-life, and interaction with other molecules. One of the best understood posttranslational modifications is the reversible phosphorylation of proteins at serine, threonine, or tyrosine residues. These phosphoamino acids are relatively stable in acidic solutions, and their comprehensive identification by mass spectrometry is, therefore, feasible. Phosphoproteomics-type experiments require some modifications in the sample preparation, mass spectrometry setup, and software-based data interpretation compared to standard proteomics workflows. Furthermore, phosphoproteome analyses are incompatible with long organelle isolation procedures prior to analysis, because of the highly dynamic nature of regulatory phosphorylations. In this chapter, we provide a detailed step-by-step overview of the complex experimental setup required for successful chloroplast phosphoproteome analysis, report our experience with existing methods, and comment on their application in the field. Key words: Proteomics, Protein phosphorylation, Affinity chromatography, Mass spectrometry
1. Introduction The systematic analysis of posttranslational modifications (PTMs) requires a specialized experimental setup particularly when the modification interferes with peptide ionization properties (1, 2). In the case of phosphorylation, affinity enrichment of phosphopeptides is required to increase their relative concentration compared to their nonphosphorylated counterparts (1, 3). This is necessary because phosphopeptides are often substoichiometric and the negative charge of the phosphate group has an unfavorable influence on their detection in positive ion mode. Commonly employed phosphopeptide enrichment strategies combine strong cation exchange chromatography (SCX) with immobilized metal R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_15, © Springer Science+Business Media, LLC 2011
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affinity chromatography (IMAC) or titanium dioxide affinity chromatography (TiO2). The two affinity methods deliver partially complementary results and their combined use allows more comprehensive analyses to be performed (4). In addition to peptide enrichment, large-scale phosphorylation analyses require a specialized database search that uses dramatically increased search spaces because several additional amino-acid masses can be used for matching the measured mass with peptides in the database. Thus, more potential solutions to the peptide and protein inference problem exist that will inevitably lead to higher numbers of false assignments, i.e., an increased false discovery rate (FDR). This must be taken into account for the setup of the database search and the interpretation of the results (5). Phosphoproteome analyses provide insights into the regulatory role of protein phosphorylation and its dynamics in the cell. To this end, the analysis should be quantitative and the quantification results must reflect the biochemistry. One of the most important sources for quantification errors is the preparation of the biological material, which is especially critical for organellar phosphoproteins. Considering that changes in phosphorylation state occur within minutes, lengthy organelle isolation procedures should be avoided. To circumvent this problem, we propose an approach that acquires the phosphorylation data at the level of the entire cell and makes use of organelle proteome reference lists to assign proteins to organelles a posteriori (6). Even with excellent biological material, the robust quantification of phosphorylation dynamics is still critical despite established isotope labeling techniques. This is a consequence of undersampling of highly complex peptide mixtures. Therefore, targeted analyses that specifically screen for precursor masses and that allow determining the area under curve for phosphopeptide peaks are the methods of choice for quantification. Such methods, which are known as accurate inclusion mass scanning (AIMS), require detailed prior knowledge about the phosphoproteome that can be obtained with the phosphoproteome mapping method described here (7). In the following, we describe in detail procedures for the extraction of proteins from Arabidopsis shoots, the enrichment of phosphopeptides by SCX combined with IMAC or TiO2 affinity chromatography, and the acquisition of mass spectrometry (MS) data along with their interpretation.
2. Materials 2.1. Plant Material, Protein Extraction, and In-Solution Tryptic Digest
1. Osmo buffer : 40 mM Tris–HCl, pH 8, 5 mM MgCl, 1 mM dithiothreitol (DTT), inhibitor cocktail for proteases (e.g., EDTA free, Roche, Basel, Switzerland), inhibitor cocktail for phosphatases (e.g., phosSTOP, Roche). Add inhibitor cocktails
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freshly. Alternatively, store buffers including the inhibitor cocktails at −20°C. 2. SDS buffer: 40 mM Tris, 4% (w/v) sodium dodecyl sulfate (SDS), 40 mM DTT. 3. Resolution buffer: 20 mM Tris–HCl, pH 8.3, 3 mM ethylenediaminetetraacetic acid (EDTA), 8 M urea. 4. Digestion buffer: 20 mM Tris–HCl, pH 8.3, 3 mM EDTA. 5. Trypsin (sequencing grade, modified). 6. 10% (v/v) trifluoroacetic acid (TFA). 7. Other chemicals required: 100% acetone, 100% methanol, 100% chloroform, DTT, and iodoacetamide. 8. Standard protein assay (e.g., Bradford assay). 9. Mortar and pestle. 10. Vortex mixer. 11. Benchtop centrifuge, with rotor to accommodate 50-mL Falcon tubes. 12. Falcon tubes, 50 mL. 13. SpeedVac sample concentrator. 14. Heating block (for incubations at 50°C and 37°C). 15. pH test strips. 2.2. Desalting of Peptides
1. Sep-Pak reverse phase columns (C18) (see Note 1). 2. 3% (v/v) acetonitrile (ACN), 0.1% (v/v) TFA. 3. 80% (v/v) ACN, 0.1% (v/v) TFA. 4. 0.1% (v/v) TFA. 5. 60% (v/v) ACN, 0.1% (v/v) TFA. 6. Other chemicals required: 100% methanol. 7. SpeedVac sample concentrator.
2.3. Fractionation of Peptides by SCX Chromatography
1. 4.6 × 200-mm polySULFOETHYL aspartamide A column (PolyLC Inc., Columbia, MD, USA). 2. High-performance liquid chromatography (HPLC) system including an autosampler and variable wavelength detector (e.g., Agilent HP1100 binary HPLC system, Santa Clara, CA, USA). 3. Buffer A: 10 mM KH2PO4, pH 2.6 in 25% (v/v) ACN (see Note 2). 4. Buffer B: 10 mM KH2PO4, pH 2.6, 350 mM KCl in 25% (v/v) ACN (see Note 2). 5. SpeedVac sample concentrator.
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2.4. IMAC
1. Chelating Sepharose Fast Flow beads in ethanol (GE Healthcare, Piscataway, NJ, USA). 2. 25% (v/v) ACN, 0.1% (v/v) acetic acid. 3. 0.1 M FeCl3 (prepare freshly). 4. Washing buffer: 74:25:1 H2O:ACN:acetic acid. 5. 100 mM Na-phosphate buffer, pH 8.9. 6. 10% (v/v) TFA. 7. pH test strips. 8. SpeedVac sample concentrator. 9. Fine gel loading tips.
2.5. TiO2 Chromatography
1. TiO2 (GL Sciences, Tokyo, Japan) (see Note 3). 2. Small columns for affinity chromatography (e.g., Mobicols with screw caps and small filters, MoBiTec, Göttingen, Germany). 3. Phthalic acid solution: 80% (v/v) ACN, 2.5% (v/v) TFA, 0.13 M phthalic acid. 4. 80% (v/v) ACN, 0.1% (v/v) TFA. 5. 0.1% (v/v) TFA. 6. 0.3 M NH4OH. 7. 3% (v/v) ACN, 0.1% (v/v) TFA. 8. Other chemicals required: 100% ACN; 100% TFA; phthalic acid; 100% methanol. 9. End-over-end rotator. 10. SpeedVac sample concentrator.
2.6. ZipTip
1. ZipTips (mC18; Millipore, Billerica, MA, USA) (see Note 4). 2. 80% (v/v) ACN, 0.1% (v/v) TFA. 3. 0.1% (v/v) TFA. 4. 60% (v/v) ACN, 0.1% (v/v) TFA. 5. Other chemicals required: 100% methanol. 6. SpeedVac sample concentrator.
2.7. Reverse Phase Chromatography
1. Reverse phase nanocolumn: 11-cm length with 75 mm inner diameter packed with Magic C18 AQ 3 mm resin (Michrom BioResources, Auburn, CA, USA). 2. Chromatography buffer A: 5% (v/v) ACN, 0.1% (v/v) formic acid. 3. Chromatography buffer B: 80% (v/v) ACN, 0.1% (v/v) formic acid.
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1. High-accuracy mass spectrometer capable of tandem mass spectrometry (MS/MS) (e.g., LTQ-Orbitrap, FTICR, Q-TOF), coupled with a nano-LC system. 2. Analysis software for MS-data interpretation (e.g., SEQUEST or Mascot).
3. Methods 3.1. Plant Material and Protein Extraction
1. Grow Arabidopsis seedlings for 22 days under short-day conditions on soil. 2. Harvest shoots and immediately freeze samples in liquid nitrogen. Store samples at −80°C until ready to proceed (see Note 5). 3. Grind 2 g of Arabidopsis shoots in liquid nitrogen to a fine powder with mortar and pestle, and then aliquot 4 × 0.5 g in 2-mL Eppendorf tubes.
3.2. Extraction of Soluble Proteins
1. Add to each Eppendorf tube 900 mL of osmo buffer (including protease and phosphatase inhibitors) and vortex. 2. Centrifuge for 10 min at 16,000 × g (4°C) to remove cell debris and pool all four supernatants into a 50-mL Falcon tube. 3. Repeat steps 1–2 and add the supernatants to the first collection. Save the sedimented plant material for membrane protein extraction on ice. 4. Estimate the total volume of the pooled supernatants and then add 5 volumes of 100% acetone (precooled at −20°C) and vortex. 5. Precipitate proteins for 3 h at −20°C (see Note 6). 6. Centrifuge at 4,000 × g for 10 min. Discard the supernatant and let the pellet air-dry at room temperature. 7. Dissolve the dry protein pellet in 100 mL of resolution buffer and transfer to a 2-mL Eppendorf tube. Rinse the Falcon tube with 700 mL of digestion buffer and add to the protein suspension in the 2-mL Eppendorf tube (see Note 7). 8. Determine the protein concentration using a standard method (e.g., Bradford assay).
3.3. Extraction of Membrane Proteins
1. Add to each plant sediment (from step 3 of Subheading 3.2) 500 mL of SDS buffer and pipette up and down. Try to avoid the formation of foam. 2. Centrifuge for 10 min at 16,000 × g and pool the supernatants in a 50-mL Falcon tube. 3. Add 1 mL of SDS buffer to each of the four sediments and mix by pipetting up and down. Centrifuge for 10 min at 16,000 × g.
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Pool all supernatants with the first collection in the 50-mL Falcon tube, mix, and then split into two 50-mL Falcon tubes. 4. Estimate the volume and then add 4 volumes of 100% methanol and vortex for 30 s. 5. Add 1 volume (in relation to the original volume determined in step 4 above) of 100% chloroform and vortex for 30 s. 6. Add 3 volumes (original volume, see above) of water and vortex for 30 s, then incubate for 1 min at room temperature before centrifuging for 5 min at 4,000 × g. 7. During centrifugation, the sample separates into three phases: the lower phase, the interphase, and the upper phase. The proteins reside in the lower phase and the interphase. Discard the upper phase. 8. Estimate the combined volume of the lower phase and interphase, and then add 3 volumes of 100% methanol and vortex. 9. Centrifuge for 3 min at 4,000 × g to precipitate the proteins (see Note 8). Discard the supernatant. 10. Do not dry the pellet; directly dissolve the proteins in 100 mL of resolution buffer and transfer to a 2-mL Eppendorf tube (see Note 9). 11. Rinse the Falcon tube with 700 mL of digestion buffer and add to the protein suspension (see Note 10). 12. Briefly run the samples in a SpeedVac, for 5 min, to remove the remaining solvents. 13. Determine the protein concentration using a standard assay (e.g., Bradford assay). 3.4. In-Solution Tryptic Digest
1. Use as much protein as possible for the tryptic digest, but use at least 1 mg of soluble proteins and 500 mg of membrane proteins. 2. Add DTT to a final concentration of 10 mM and reduce cysteine residues for 45 min at 50°C. Subsequently, alkylate reduced cysteine residues by adding iodoacetamide to a final concentration of 50 mM. Incubate in the dark at room temperature for 1 h (see Note 11). 3. Add trypsin in a ratio of 1:20 based on mass (trypsin: protein) and digest overnight at 37°C. 4. After overnight digestion, centrifuge the mixture for 10 min at 16,000 × g to remove insoluble material. Transfer the supernatant, which includes the tryptic peptides, to an Eppendorf tube. 5. Adjust the sample to pH <3 by adding 10% TFA. Use 0.5 mL of the solution to check the pH with pH test strips. Centrifuge to precipitate and remove the sediments that may appear due to the change of pH.
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6. Concentrate the peptides in the supernatant in a SpeedVac to a final volume of approximately 10 mL, and then desalt the two samples (the peptides of membrane and soluble proteins) with Sep-Pak columns (see below) (see Note 12). 3.5. Desalting of Peptides
1. Dilute the peptides (see Subheading 3.4, step 6) in 1 mL of 3% ACN, 0.1% TFA. 2. Wet the Sep-Pak columns with 3 × 1 mL of 100% methanol (see Note 13). 3. Wash the Sep-Pak columns with 1 mL of 80% ACN, 0.1% TFA. 4. Equilibrate the columns with 2 × 1 mL of 0.1% TFA. 5. Load the diluted peptides onto the columns (see Note 14). 6. Wash the columns with 2 mL of 0.1% TFA. 7. Elute the peptides into 1.5-mL Eppendorf tubes with 500 mL of 60% ACN, 0.1% TFA. 8. Before continuing with SCX, concentrate the desalted peptides to a few microliters in a SpeedVac.
3.6. Fractionation of Peptides by SCX Chromatography
1. Dissolve the peptides in 200 mL of Buffer A and load them on a polySULFOETHYL aspartamide A column on an Agilent HP1100 binary (or equivalent) HPLC system. 2. Elute the peptides with the following increasing KCl gradient: 10–40 min, 0–30% Buffer B; 40–60 min, 30–100% Buffer B. 3. Collect the fractions (e.g., 27 fractions) with the auto sampler. 4. Concentrate the peptides in a SpeedVac to approximately 50 mL. Never dry them completely (see Note 15). 5. Pool the collected fractions according to their readout at 214 and 280 nm. The fractions eluting first from the column do not possess absorbance at 214 and 280 nm but usually contain primarily the phosphopeptides (see Note 16); pool them into fraction 1. Pool the subsequent fractions with the first absorbance peaks at 214 and 280 nm into fraction 2. Pool the remaining samples (mainly nonphosphorylated peptides) into the last three fractions (3–5). 6. Desalt all five fractions using Sep-Pak columns as described in Subheading 3.5. 7. Each desalted fraction will be used individually for both IMAC and TiO2 chromatography. Thus, split the 500 mL eluates in two equal aliquots (2 × 250 mL). Store one aliquot at −20°C to use it later on for TiO2 chromatography. SpeedVac the second aliquot down to approximately 100 mL volume to remove ACN, and then use it for IMAC.
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3.7. IMAC
1. Prepare the samples for IMAC by diluting peptides in 25% ACN, 0.1% acetic acid. 2. To charge the beads, take 20 mL of Fast Flow beads per sample and pipette into an Eppendorf tube. Use a cut pipette tip (lacking ~3–5 mm from the end) for pipetting to avoid damaging of the beads. 3. Spin for 30 s at 400 × g at room temperature in a microcentrifuge, and then discard the supernatant. Avoid removing the beads. 4. Estimate the volume of the beads and add the same volume of 0.1 M FeCl3. Mix gently by finger tapping. Spin for 30 s at 400 × g and then remove the supernatant. Repeat the charging at least four times with 0.1 M FeCl3 until the beads are yellow to brown. 5. Wash the beads with washing buffer 4–8 times until the supernatant is clear. Check the volume of the beads before the last wash. 6. Add 3 volumes of washing buffer to obtain 25% bead slurry. Aliquot 40 mL of 25% slurry per sample into Eppendorf tubes. 7. Start the phosphopeptide enrichment by applying the prepared peptides to the 25% slurry, and then mix gently by finger tapping. 8. Incubate at room temperature for 45 min with occasional finger tapping. 9. Spin at 400 × g for 30 s and collect the supernatant (flowthrough) in a fresh Eppendorf tube. 10. Wash four times with 100 mL of washing buffer and add the first two supernatants obtained from the washing to the flowthrough. 11. Wash once briefly with 70 mL of water. 12. Add 30 mL of 100 mM Na-phosphate buffer, pH 8.9, to elute the phosphopeptides (see Note 17). Mix gently by finger tapping and spin at 400 × g of 30 s. Collect the eluted phosphopeptides in a fresh Eppendorf tube. Use fine gel loading tips to remove as much of the supernatant as possible. 13. For ZipTip (described in Subheading 3.9), adjust the eluted samples to pH 2–3 by adding 10% TFA. Check sample pH by applying 0.5 mL of the sample solution to pH test strips (see Note 18). 14. Reduce the volume of the flow-through by 25% in a SpeedVac, to remove ACN. ZipTip the samples before proceeding to MS (see Subheading 3.10).
3.8. Enrichment of Phosphopeptides Using TiO2
1. Thaw the peptides (from step 7, Subheading 3.6). Adjust the peptide solution to 80% ACN, 2.5% TFA, 0.13 M phthalic acid. To do so, add 250 mL of 100% ACN and 12.3 mL of 100% TFA. Add 10 mg of phthalic acid to each sample (see Note 19).
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2. Apply the small filters into blocked Mobicol columns and close them with the small caps. 3. Weigh 0.3 mg of TiO2 per sample into an Eppendorf tube. Wash with 80 mL of water per 1.5 mg TiO2. Spin at 2,000 × g for 1 min and discard the supernatant. 4. Wash the TiO2 with 80 mL of 100% methanol per 1.5 mg TiO2. 5. Resuspend the TiO2 in 120 mL of phthalic acid solution, mix, and aliquot 24 mL per sample into separate Mobicol columns. 6. Load the samples and close the Mobicols with the screw caps. Incubate for 30 min at room temperature on an end-over-end rotator. 7. Remove the small cap and let the flow-through drop through the filters (see Note 20). 8. Wash the resin with 200 mL of phthalic acid solution. Repeat the washing with 150 mL of phthalic acid solution. 9. Wash with 300 mL of 80% ACN, 0.1% TFA. Perform a quick additional wash with 150 mL of 80% ACN, 0.1% TFA. 10. Wash once with 300 mL of 0.1% TFA. 11. Wash once with 300 mL of 80% ACN, 0.1% TFA. 12. Elute the phosphopeptides into fresh Eppendorf tubes with 150 mL of 0.3 M NH4OH per column. Carry out a second elution with 100 mL of 0.3 M NH4OH, and then pool both eluates. Fully dry the peptides in the SpeedVac. Afterward, dissolve each peptide sample in 10 mL of 3% ACN, 0.1% TFA for ZipTip. 3.9. ZipTip
1. Prepare one Eppendorf tube with 1 mL of 100% methanol, one with 1 mL of 80% ACN, 0.1% TFA, and two with 1 mL of 0.1% TFA (one for equilibration, one for washing). 2. For elution, prepare for each sample an Eppendorf tube containing 10 mL of 60% ACN, 0.1% TFA. 3. During the ZipTip procedure described below, set your pipette to a volume of 10 mL and pipette up and down with the appropriate solution. 4. Wet the ZipTips by pipetting three times up and down with 100% methanol. 5. Wash the ZipTips three times with 80% ACN, 0.1% TFA. 6. Equilibrate the ZipTips three times with 0.1% TFA. 7. Bind the peptides by carefully pipetting the peptide solution ten times up and down. 8. Wash three times with 0.1% TFA. 9. Elute the peptides into the prepared Eppendorf tubes of step 2 by pipetting five times up and down with 60% ACN, 0.1% TFA.
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10. Dry the eluted peptides in a SpeedVac and dissolve the peptides for MS analysis in 10 mL of chromatography buffer A (see Subheading 2.7). 3.10. Acquisition of MS Data for Phosphopeptide Identification 3.10.1. Reverse Phase Chromatography
3.10.2. Mass Spectrometry
1. Equilibrate a reverse phase nanocolumn coupled to a mass spectrometer with at least 10 column volumes of chromato graphy buffer A at a flow rate of 200 nL/min. 2. Load 5 mL of the peptide solution (see Subheading 3.9, step 10) onto the reverse phase column; keep the flow rate at 200 nL/min for column loading and peptide elution. 3. Separate the peptides by gradient elution from 3 to 35% ACN in 90 min (see Note 21). 1. Acquire MS data over the entire chromatography run with a calibrated high-accuracy mass spectrometer (see Note 22). 2. Acquire up to five data dependent MS/MS spectra in the linear ion trap for each Orbitrap-Fourier transform mass spectrum (FTMS) (see Note 23). 3. Acquire the latter at 60,000 full width at half maximum (FWHM) nominal resolution settings with an overall cycle time of approximately 1 s. 4. Enable dynamic exclusion with a setting that up to 500 m/z ± 20 ppm values are excluded from tandem MS for 120 s. Set the automatic gain control to 5e5 for full Orbitrap (FTMS) trap and to 1e4 for linear ion trap MS/MS. 5. It is advisable to acquire peptide masses using the internal lock mass calibration option with the masses m/z 429.088735 and 445.120025 (see Note 24).
3.10.3. Interpretation of the Results
1. Search the raw file obtained from the analysis in a confined database, e.g., the Arabidopsis thaliana database containing common contaminants such as keratin (see Note 25). 2. Use Mascot 2.1.04 (Matrix Science, London, UK) and allow as fixed modification carbamylation of cysteines and as variable modifications oxidation of methionine and phosphorylation of serine, threonine and tyrosine. 3. Restrict the search space to tryptic peptides with a maximum of two missed cleavages and protein C- and N-terminal peptides. 4. Allow for 2+ and 3+ charged peptides at a parent mass error tolerance of 5–10 ppm and a daughter ion error tolerance of 0.6 Da (see Note 26). 5. Assess the false discovery rate (FDR) in decoy database searches and adjust the FDR to the desired value (an FDR of 0.1–0.2% is acceptable) by choosing the respective Mascot score as a threshold for acceptance (a Mascot ion score >30 and a Mascot
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expect value <0.015 usually delivers reasonable results) (see Note 27). 6. Determine the exact assignment of the phosphorylation site by calculating a normalized delta ion score (DI) for those phosphopeptides for which the only difference between the rank 1 and the rank 2 hit is the position of the phosphate group. DI is calculated by taking the difference of the two top ranking ion scores and dividing that difference by the ion score of the first ranking phosphopeptide. Phosphorylation site assignments with DI ³ 0.4 are usually considered acceptable. DI scores lower than 0.4 indicate an ambiguous assignment of the phosphorylation site (see Note 28). 7. In critical cases when biological information is based on the detection of a phosphopeptide or a certain phosphorylation site, we strongly encourage performing a manual spectrum interpretation. Even nonexperts can easily identify the dominant neutral loss of phosphoric acid from the precursor mass, which indicates the presence of phospho-Ser or phospho-Thr in the phosphopeptide. Phospho-Tyr is maintained in the MS/ MS spectrum sequence derived from collision induced dissociation (CID) experiments. In order to validate the phosphorylation site assignment, use the manual de novo sequencing rules (e.g., provided in http://www.ionsource.com). 8. To extract the chloroplast phosphoproteins, we suggest using chloroplast proteome reference tables that are available in public databases based on previously published proteomics data (e.g., from http://suba.plantenergy.uwa.edu.au/).
4. Notes 1. Sep-Pak columns contain a silica-based stationary phase of strong hydrophobicity and are used to absorb analytes of even weak hydrophobicity from aqueous solutions. Here, Sep-Pak reverse phase columns (C18) are used to desalt peptides. 2. Precisely adjust to pH 2.6. This is important for a proper separation of phosphopeptides and nonphosphorylated peptides during SCX. 3. TiO2 is delivered in HPLC columns. Cut the columns to recover the TiO2 powder and transfer the powder to Eppendorf tubes. 4. ZipTips are 10 mL tips with 0.6 mL mC18 material fixed to the end of the tip. These tips are used to concentrate and purify peptides prior to MS. 5. During the whole procedure of harvesting, protein extraction, and phosphopeptide enrichment, wear nonpowdered gloves to
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minimize human keratin contamination. Try to work as cleanly as possible since contamination could prevent the identification of relevant proteins. 6. During acetone precipitation, continue with the extraction of membrane proteins. Time of precipitation can be extended up to overnight to increase protein yield. 7. Increase the volume of the resolution buffer if the pellet cannot be dissolved in 100 mL. In this case, also add more digestion buffer to a final urea concentration of approximately 1 M. Urea concentrations higher than 1 M inhibit trypsin during digestion. 8. Do not centrifuge longer since the pellet might become too dense to dissolve the membrane proteins. 9. Dissolving membrane proteins always takes time. So keep calm and take your time. Start with a small volume of resolution buffer (e.g., 50 mL). Give the pellet time to dissolve. The suspension might be incubated for 1 h in the fridge at 10°C, but not colder; otherwise, urea will precipitate. Rinse the tube with 50 mL of resolution buffer. If you are unable to dissolve the pellet in 100 mL of resolution buffer, increase the volume of the resolution buffer. 10. If you used in step 10 more than 100 mL of resolution buffer you have to add more digestion buffer. The final urea concentration must be diluted to £1 M for the tryptic digest. 11. Cysteine residues tend to oxidize to give rise to disulfide bonds, which disturbs mass spectrometry analysis. To prevent the formation of disulfide bonds, cysteine residues of peptides are first reduced with DTT and further alkylated with iodoacetamide resulting in a covalent addition of a carbamidomethyl group (57.07 Da) to the cysteine residues. If iodoacetamide is present in limiting quantities and at a slightly alkaline pH, cysteine modification will be the exclusive reaction. Excess iodoacetamide or nonbuffered iodoacetamide reagent can also alkylate amines (lysine, N-termini), thioethers (methionine), imidazoles (histidine), and carboxylates (aspartate, glutamate). The sulfhydryl-reactive alkylating reagent iodoacetamide is unstable and light sensitive. Therefore, iodoacetamide must be freshly prepared and incubation has to take place in the dark. 12. From this step on, always use low-binding Eppendorf tubes to minimize peptide loss by adsorption to the tube. This is especially important when lyophilizing peptides in the SpeedVac. Avoid complete drying during lyophilization to reduce the losses by peptide adsorption to the tube. 13. To be faster you can carefully apply pressure with a pipette and cut blue tips.
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14. Do not apply pressure with a pipette to guarantee an optimal binding of the peptides to the columns. 15. The very last eluted SCX fractions contain a high concentration of KCl that may result in precipitation during lyophilization. Do not dismay. After pooling fractions in step 5 and dissolving peptides in 3% ACN, 0.1% TFA for Sep-Pak desalting, the precipitate will disappear. 16. At pH 2.6, most tryptic peptides have a charge state of 2+, while phosphopeptides have a lower net charge state due to the negatively charged phosphate group. Thus, phosphopeptides will elute earlier than nonphosphorylated peptides during SCX. Keep in mind that in comparison to nonphosphorylated peptides the amount of phosphopeptides is very low. Owing to this low abundance, the earlier eluted phosphopeptides will not be detectable at 214 and 280 nm. The detectable absorbance peaks at 214 and 280 nm mainly correspond to nonphosphorylated peptides. 17. Phosphopeptides bind to Fe-ions only under acidic conditions and will be eluted by increasing the pH. Under acidic conditions the Fe-loaded beads are yellow to brown. When adding Na-phosphate buffer, the color of the beads should change to white due to the increased pH. If the beads remain yellow, add more Na-phosphate buffer. 18. Phosphopeptides are unstable at basic pH; therefore, it is important to adjust the pH to 2–3 as soon as possible after elution of the phosphopeptides. 19. At 0.13 M, phthalic acid is saturated; therefore, it takes some time till the salt is completely dissolved. 20. In the following steps, pressure can be applied with a pipette and a capped blue tip. 21. The gradient shape depends on the sample complexity and the goal of the analysis. For comprehensive analyses of the chloroplast phosphoproteome, we use 90 min to increase the ACN concentration in the mobile phase to 35% ACN; at this concentration the majority of the phosphopeptides are eluted from the column. We usually use such a gradient for the analysis of complex mixtures with the goal to comprehensively analyze a complex phosphoproteome. 22. As described in the Introduction, the analysis of PTMs with mass spectrometric data allows for additional combinatorial solutions to the mass matching problem; thus, a high mass accuracy is desirable to decrease the search space and reduce the risk of false positive phosphopeptide assignments. 23. It is advisable to restrict the MS/MS data acquisition to a minimum threshold, which depends on the chemical noise in the MS run. In general, a reasonable setting for data-dependent
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MS/MS acquisition (DDA) is to define the four most abundant peaks eluting from the column at a given time as targets. 24. The settings of the mass spectrometer were optimized for an LTQ-Orbitrap and can deviate considerably for other instruments and laboratories. Thus, these settings should only provide a rough guidance for phosphoproteome analysis. Consult an MS expert for the optimal setting for your specific analytical questions. 25. The risk of false positive assignments increases with larger search spaces; i.e., the FDR of a search result is higher when it is obtained from searches in large databases. We, therefore, strongly encourage using a database that is tailored for the needs of the analysis. 26. The error-tolerance depends on the mass accuracy of the instrument. The higher the mass accuracy, the smaller is the necessary error tolerance. A small error tolerance improves the accuracy of the database search. 27. A decoy database has the same size and elemental composition as the target database, but the sequences are scrambled. At any given score threshold, a number of assignments will be in the target database and another number in the decoy database. The number of decoy hits must be divided by the sum of the number of target hits and decoy hits, and then multiplied by 2 to obtain the FDR at the chosen score threshold. 28. This is only necessary if several Ser, Thr, or Tyr residues are present in the identified phosphopeptide. References 1. Preisinger, C., von Kriegsheim, A., Matallanas, D., and Kolch, W. (2008) Proteomics and phosphoproteomics for the mapping of cellular signalling networks. Proteomics 8, 4402–4415. 2. Lemeer, S., and Heck, A. J. (2009) The phosphoproteomics data explosion. Curr. Opin. Chem. Biol. 13, 414–420. 3. Baginsky, S. (2009) Plant proteomics: concepts, applications, and novel strategies for data interpretation. Mass Spectrom. Rev. 28, 93–120. 4. Bodenmiller, B., Mueller, L. N., Mueller, M., Domon, B., and Aebersold, R. (2007) Reproducible isolation of distinct, overlapping segments of the phosphoproteome. Nat. Methods 4, 231–237.
5. Nesvizhskii, A. I., and Aebersold, R. (2005) Interpretation of shotgun proteomic data: the protein inference problem. Mol. Cell. Proteomics 4, 1419–1440. 6. Reiland, S., Messerli, G., Baerenfaller, K., Gerrits, B., Endler, A., Grossmann, J., Gruissem, W., and Baginsky, S. (2009) Largescale Arabidopsis phosphoproteome profiling reveals novel chloroplast kinase substrates and phosphorylation networks. Plant Physiol. 150, 889–903. 7. Jaffe, J. D., Keshishian, H., Chang, B., Addona, T. A., Gillette, M. A., and Carr, S. A. (2008) Accurate inclusion mass screening: a bridge from unbiased discovery to targeted assay development for biomarker verification. Mol. Cell. Proteomics 7, 1952–1962.
Part IV Photosynthesis and Biochemical Analysis
Chapter 16 Use of a Pulse-Amplitude Modulated Chlorophyll Fluorometer to Study the Efficiency of Photosynthesis in Arabidopsis Plants Matthew D. Brooks and Krishna K. Niyogi Abstract Chlorophyll a fluorescence has long been used as a noninvasive means to assess photosynthetic performance in plants. Pulse-amplitude modulated (PAM) fluorometry is one of the most common techniques used to study the induction and quenching of chlorophyll fluorescence in physiological studies. In this chapter, we briefly describe the basics of PAM fluorometry and how to configure the instrument before moving on to some examples of common measurements that can be made using a PAM fluorometer. Photosynthetic performance and energy dissipation are compared between wild type and the npq4-1 mutant by examining the maximum photochemical efficiency during high-light stress, the induction and relaxation of non-photochemical quenching, and by plotting light curves. Key words: Pulse-amplitude modulated fluorometry, Arabidopsis thaliana, Chlorophyll a, Fluorescence, Photosystem II, Photosynthesis
1. Introduction Light energy that is absorbed by the light-harvesting pigment- protein complexes of plants is quickly transferred to chlorophyll a, which then relaxes by one of several different pathways: photochemistry, non-photochemical processes, or fluorescence (1). In the mid-twentieth century, Hans Kautsky demonstrated that the chlorophyll fluorescence signal carries information on the light reactions of photosynthesis (2). In the 50 years since Kautsky’s characterization of induction curves, chlorophyll fluorescence has become one of the primary methods that researchers use to assess photosynthesis in vivo (3–5). With the proper instrumentation, the chlorophyll fluorescence signal can be determined in an accurate, rapid, and noninvasive manner.
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Fig. 1. Example of a typical PAM trace from a wild-type Arabidopsis plant. Fo is the dark-adapted fluorescence yield; Fm is the maximum fluorescence yield in the dark; Fv is the difference between Fm and Fo; Fs is the fluorescence yield in the light; F ´o is the light-adapted minimal fluorescence yield; and F ´m is the maximum fluorescence yield in the light. Note that not all of the saturating pulses are labeled. All of the other parameters used in this chapter can be calculated from these basic measurements (see Table 1).
One of the most common methods used is pulse-amplitude modulated (PAM) fluorometry (6, 7). PAM fluorometers use three distinct light sources: a weak (<0.1 mmol photons/m2/s) pulsed measuring light, an actinic light capable of moderate intensities (0–3,000 mmol photons/m2/s) used to drive photosynthesis, and a saturating light of high intensity (up to 18,000 mmol photons/m2/s). The advantage of the PAM system is that changes in the fluorescence yield excited by the pulsed measuring light can be isolated and quantified in a background of fluorescence intensity changes originating from the actinic and saturating lights that are orders of magnitude larger. An example of a typical PAM trace is given in Fig. 1 and descriptions of the parameters used in this paper are given in Table 1. Under ambient conditions, fluorescence originates primarily from the chlorophyll a molecules associated with photosystem II (PSII), and this fluorescence can be quenched photochemically by electron transfer to the oxidized primary electron acceptor of PSII, Q A (8). In a dark-adapted leaf, Q A is fully oxidized and the reaction centers are said to be in the open state. The fluorescence yield when Q A is maximally oxidized, Fo, is determined by exposure to the weak measuring light that has a negligible effect on photochemistry. A short, high-intensity pulse can then be given to the sample to reduce the Q A acceptor completely, closing all of the PSII
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Table 1 Fluorescence parameters used in the protocols described in this chapter Parameter
Formula
Definition
Fo
–
Minimal fluorescence in the dark (fluorescence yield when all reaction centers are open)
Fm
–
Maximum fluorescence in the dark (fluorescence yield when all reaction centers are closed)
Fo′
–
Minimal fluorescence in a light-adapted leaf when all reaction centers are open
Fm′
–
Maximum fluorescence in a light-adapted leaf when all reaction centers are closed
Fs
–
Steady-state fluorescence level
Fv/Fm
(Fm−Fo)/Fm
Maximum photochemical efficiency of PSII (efficiency at which light absorbed by PSII is used for photochemistry when all reaction centers are open)
NPQ
(Fm − Fm′ )/Fm′
Non-photochemical quenching of PSII fluorescence, directly proportional to the level of energy dissipation
FPSII
(Fm′ − Fs ) / Fm′
PSII operating efficiency (efficiency at which light absorbed by PSII is used for photochemistry in a light-adapted plant)
ETR
PPFD × 0.84 × 0.5 × FPSII
Electron transport rate through PSII. The constants 0.84 and 0.5 are estimates of the fraction of light absorbed by the leaf and the fraction of absorbed light received by PSII, respectively
qP
(Fm′ − Fs )/(Fm′ − Fo′)
Coefficient of photochemical quenching (fraction of PSII maximum efficiency that is actually realized). Also used to estimate the fraction of PSII centers that are in the open state according to the puddle model. Also referred to as Fq′ /Fv′
qL
q P × (Fo′/Fs )
Fraction of PSII centers that are in the open state according to the lake model of the PSII photosynthetic unit
reaction centers. Because fluorescence emission competes with photochemistry, the fluorescence yield will rise to its maximum value, Fm, when the reaction centers are closed. The difference between the Fo and Fm is called variable fluorescence, Fv, and the ratio Fv/Fm is equivalent to the maximum quantum efficiency of PSII (9). The Fv/Fm values of unstressed leaves from different species of plants are consistently around 0.83 (10), and lower values are often used as an indicator of stress (11) or altered function of the photosynthetic apparatus (4). When a dark-adapted leaf is exposed to an actinic light, the initial rise in the fluorescence induction curve is followed by a
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slower decrease that is due to photochemistry and non- photochemical quenching (NPQ). PAM fluorometry has been the most common technique used in studying and separating these quenching processes. To distinguish between photochemistry and NPQ in a leaf exposed to an actinic light, all of the PSII reaction centers are closed with a saturating light pulse, eliminating the contribution of photochemical quenching, and the light-adapted maximal fluorescence, Fm′, is determined. The difference in fluorescence yield, between Fm and Fm′, is due to non-photochemical processes, whereas the difference between Fm′ and Fs is due to photochemical processes. NPQ has traditionally been divided into three components: feedback de-excitation, state transitions, and photoinhibition. The contribution of each component can be determined by plotting the relaxation of NPQ versus recovery time and performing linear regression analysis (12). Another common measurement made using a PAM fluorometer is a light curve (9). In this method, an actinic light is applied to the sample in a series of increasing intensities. At each light intensity, once the fluorescence reaches a steady state, Fs, a saturating pulse is applied to close the reaction centers and measure Fm′. This is immediately followed by a few seconds when the actinic light is turned off and a short pulse of far-red light is applied to oxidize QA and determine Fo′. From these measurements several different informative indicators of photosynthetic performance can be determined and plotted versus light intensity. There is a wellestablished correlation between PSII photochemical efficiency, as measured by fluorescence, and the efficiency of CO2 assimilation (9) and oxygen evolution (13). The following sections describe how to configure the settings on a PAM fluorometer and how to record the basic measurements needed to calculate photochemical efficiency, follow NPQ induction and relaxation, and finally how to plot a light curve. Wild-type Arabidopsis thaliana plants are compared to the npq4-1 mutant, which lacks the feedback de-excitation component of NPQ (14), in order to demonstrate the usefulness of the PAM system. The instrument used in the following protocols is the commercially available Hansatech FMS 2. However, because the basis of the parameters and the experimental design do not depend on the specifics of this instrument, the following methods can be performed on any other PAM system.
2. Materials 1. Arabidopsis thaliana plants (Columbia-0 wild type and npq4-1) are grown at 21°C for 4–6 weeks with a 12-h light/12-h dark cycle and measured before bolting. Plants are grown under
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low-light conditions of 100–200 mmol photons/m2/s. Leaves should be larger than the diameter of the PAM fluorometer fiber optic when measured. High-light treatments are done at 1,700 mmol photons/m2/s. Plants should be dark adapted for at least 20 min prior to measurements (see Note 1). 2. Hansatech FMS 2 (Hansatech Instruments Ltd., King’s Lynn, UK) or similar pulse modulated fluorometer with leaf clip and light/temperature sensor, connected to a suitable computer loaded with the FMS software.
3. Methods 3.1. Setting Up the Instrument
1. Attach the fiber optic to the leaf clip by sliding it into the groove, aligning the marked graduations, and tightening the screw. The following steps need to be performed whenever the position of the fiber optic is adjusted relative to the sample. 2. It is necessary to make sure that the measuring light does not significantly impact the physiology of the leaf. Take a darkadapted plant and gently place a leaf in the clip while the leaf is still attached to the plant (see Note 2). Turn on the measuring light and start recording the fluorescence. In order to get a reliable measurement, the fluorescence from the leaf should be high enough (between 100 and 600 units for the FMS 2) so that the noise is negligible. If it is too low, adjust the gain and/ or move the fiber optic closer to the leaf. Follow the trace for a couple minutes to make sure that the fluorescence does not increase. If it does change, lower the modulation frequency and/or move the fiber optic further away from the sample. 3. Measure the intensity of the actinic light source for the light levels that are to be used. To do this, place the light/temperature sensor in the front mount and then slide the upper part of the clip forward while gently pressing down so that the fiber optic is at the same position and angle over the sensor as it would be over the leaf (see Note 3). Set the actinic light to a low intensity (a value of 5 for the FMS 2, approximately 10 mmol photons/m2/s), turn on the lamp, and note the photosynthetically active radiation (PAR) value (see Note 4) displayed at the bottom of the window. Turn off the light, increase the light intensity by 5, turn the lamp back on and note the PAR value again. Continue until you reach the maximum actinic light intensity for the system (in the case of the FMS 2, a value of 50, approximately 3,000 mmol photons/m2/s). 4. Check that the saturating light pulse is strong enough to record the maximum fluorescence. Place a new dark-adapted leaf in the clip. Set the duration of the pulse to 0.8 s and the intensity
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to a value of 50. Start the measurement and give the sample a pulse of saturating light. Wait 30 s, increase the pulse intensity by 10 and give the leaf another saturating pulse. Repeat until a pulse intensity of 100 (approximately 18,000 mmol photons/ m2/s) is reached and then stop the recording. The pulse intensity at which no further increase in maximal fluorescence (Fm) is seen when the intensity is increased further should be used for further measurements (see Note 5). If the fluorescence value is greater than the maximum detection range for the instrument (4,095 for the FMS 2) at any point during the experiment, the gain and/or orientation of the fiber optic will need to be adjusted. 5. Measure the saturating pulse intensity with the light sensor. The length of the pulse need to be set to ~3 s in order to get an accurate measurement. Turn on the saturating light source and note the PAR value. Slide the clip back to the measurement position and reset the pulse length to 0.8 s. 3.2. Measurement of Maximum Photochemical Efficiency (Fv /Fm) in High-Light-Treated Leaves
1. Transfer a subset of wild type and npq4 plants to high light. 2. To measure the Fv/Fm, insert a dark-adapted, low-light-grown leaf into the clip. Turn on the measuring light and start recording fluorescence, wait 30 s to confirm that the Fo is constant, and then give a saturating pulse using the Fv/Fm function (see Note 6). After the measurement is complete, stop recording and note the Fv/Fm value calculated by the instrument. 3. Repeat step 2 with a low-light-grown npq4-1 plant and then with wild-type and npq4-1 plants exposed to 30, 60 and 90 min of high light. Be sure to dark adapt each plant for 20 min prior to measurement. An example of the expected results is shown in Fig. 2. The decrease in Fv/Fm is due to photoinhibition of the PSII reaction centers. Relative to wild type, the npq4-1 mutant is more susceptible to photoinhibition due to the lack of feedback de-excitation and over reduction of PSII electron acceptors (15). Recovery of Fv/Fm during a subsequent lowlight period can also be measured, and different stresses or treatments can be substituted for high light, depending on the experiment.
3.3. Induction and Relaxation of Non-photochemical Quenching
1. Create a simple script using the FMS software as follows in order to measure several plants using the same routine. Use the configuration established in Subheading 3.1 for the position of the fiber optic, modulation frequency, and saturating pulse intensity (for the Fv/Fm and FPS2 functions) (see Note 7). (a) 30 s of dark followed by an Fv/Fm measurement (b) Actinic light on at an intensity of 1,200 mmol photons/ m2/s (as set up in Subheading 3.1)
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Fig. 2. Changes in the maximum photochemical efficiency of wild-type (gray bars) and npq4-1 (white bars) Arabidopsis plants as the result of high-light stress treatment. Plants grown in low light (100–200 mmol photons/m2/s) were exposed to high light (1,700 mmol photons/m2/s) for 0, 30, 60 and 90 min. Fv/Fm was measured after 20 min dark acclimation. Data are shown as the means ± SD (n = 3). Asterisks indicate significant differences (P < 0.05, Student’s t-test).
(c) Take a measurement using the FPS2 function after 30 s and then every minute for 5 min (d) Actinic light off (e) Take a measurement using the FPS2 function after 30 s and then every 2 min for 6 min (see Note 8) 2. Place a dark-adapted wild-type leaf in the leaf clip and start the script. Once the script has finished, save the data to a spreadsheet. 3. Repeat step 2 with a dark-adapted npq4-1 leaf. 4. Plot NPQ versus time for each sample to obtain a graph as seen in Fig. 3. The npq4-1 mutant lacks the feedback de-excitation component of NPQ that is rapidly turned on in high light and off in darkness in wild-type plants (14). The remaining slowly inducing and relaxing NPQ is mainly due to photoinhibition. 3.4. Light Curve
1. Create a script using the FMS software as follows. Use the configuration established in Subheading 3.1 for the position of the fiber optic, modulation frequency, and saturating pulse intensity (for the Fv/Fm and FPS2R functions) (see Note 7). (a) 30 s of dark followed by Fv/Fm measurement (b) Actinic light on at an intensity setting of 5 (c) Wait 5 min (see Note 9)
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Fig. 3. Non-photochemical quenching (NPQ) induction curves in wild-type (solid line) and npq4-1 (dotted line) Arabidopsis plants. Plants grown in low light (100–200 mmol photons/m2/s) were dark-adapted for 20 min before exposure to actinic light of 1,200 mmol photons/m2/s. The white bar at the top of the figure indicates when the actinic light was on, and the black bar when the actinic light was turned off. Data are shown as the means ± SD (n = 3).
(d) Take a measurement using the FPS2R function (e) Increase actinic light intensity by 5 (f) Wait 5 min and take another measurement with the FPS2R function (g) Repeat steps e and f until the maximum intensity is reached (an actinic light setting of 50 on the FMS 2) 2. Place a dark-adapted wild-type leaf in the leaf clip and start the script. Once the script has finished, save the data to a spreadsheet. 3. Repeat with a dark-adapted npq4-1 leaf. 4. Export the data to a spreadsheet. Plot 1 − qL (see Note 10), FPSII, the electron transport rate through PSII (ETR), and NPQ versus light intensity. An example is shown in Fig. 4 (see Note 11). Due to the absence of the feedback de-excitation component of NPQ in the npq4-1 mutant, the excess light energy at higher PPFDs, which is dissipated as heat in wildtype plants, results in the over reduction of PSII electron acceptors and an increased excitation pressure (15). The FPSII and ETR parameters, changes in which can be indicative of problems with light harvesting or photochemistry, are the same in npq4-1 and wild-type plants (14).
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Fig. 4. Light curves of wild-type (solid lines) and npq4-1 (dotted lines) Arabidopsis plants. Plants grown in low light (100–200 mmol photons/m2/s) were dark-adapted for 20 min and then exposed to increasing light intensities. Data are shown as the means ± SD (n = 3). (a) Electron transport rate through PSII, (b) Non-photochemical quenching, (c) PSII operating efficiency, (d) Fraction of closed PSII reaction centers (excitation pressure).
4. Notes 1. Adequate dark adaptation (by simply placing the plant in a box or drawer) is required to relax NPQ and to fully oxidize the primary electron acceptor QA and get an accurate measure of Fo and Fm. This can be checked by giving the leaf a pulse of weak far-red light, which will preferentially excite photosystem I and oxidize QA. If a change in Fo occurs due to the far-red pulse, increase the amount of time the plant is dark adapted. 2. It is possible to detach a leaf or punch out a leaf disc (larger than the area of the fiber optic) and use that sample for the measurement. This is often necessary if one wishes to infiltrate the sample with a photosynthetic inhibitor. Water loss from these samples occurs more quickly than in attached leaves and can cause errors in the estimation of various parameters. Therefore, measurements that take more than 10–15 min should be done on attached leaves.
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3. It is important that the position of the leaf relative to the fiber optic remains the same for each sample. If the distance or angle changes, the amount of incident light on the sample will also change and differences between samples will not be informative. 4. PAR is light in the spectral range from 400 to 700 nm which can be used in photosynthesis. It is often quantified as photosynthetic photon flux density (PPFD) with the units mmol photons/m2/s. 5. If the pulse intensity is too low, not all of reaction centers will be closed, and there will be an underestimation of Fm. If the pulse intensity is too high and/or the pulse length too long, there is a risk of the saturating pulse causing changes in the physiology of the sample, for example inducing NPQ. 6. Commercial systems, such as the FMS 2, have specific functions that can be carried out to perform a series of actions to calculate many common parameters automatically. For example, the Fv/Fm function will determine the Fo, give a saturating pulse to measure Fm, and then calculate the Fv and Fv/Fm. 7. The FPS2 function on the FMS 2 will give a saturating pulse and record several light-adapted parameters, including FPSII, Fs, Fm′, NPQ, and ETR. The FPS2R function does the same as the FPS2 function with an additional far-red pulse to oxidize QA and measure Fo′, allowing the Fv′, Fv′ / Fm′, and qP parameters to be calculated as well. 8. It is possible to decrease the time interval between saturating pulses in order to get better resolution of the rapid components of NPQ induction and relaxation. However, a potential problem with doing so is that the saturating pulses themselves may induce NPQ or prevent relaxation if given too frequently. 9. When determining how long the leaves are exposed to each light intensity, there is a trade-off between reaching a steady state and how long the experiment will take. While it may take up to 30 min to reach a true steady state at each light level, measuring a single plant in this manner could take several hours. Often, it is necessary to take measurements on many plants in a single day, and this approach would be impractical. Adapting plants to each light level for 5 min will bring the plants to a near steady-state value and also allows the experimenter to process several plants in a single day with just one fluorometer. 10. qP has often been used to measure the fraction of PSII reaction centers in the open state. This relationship is only accurate for the “puddle” model of the photosynthetic unit and has been shown experimentally to be nonlinear (16, 17). Kramer et al. (18)
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therefore introduced the term qL to represent the fraction of open PSII centers using the “lake” model, which can easily be calculated from available fluorescence parameters as q L = q P (Fo′ / Fs ). 1 − qL then represents the fraction of closed PSII centers. 11. The definitions of several photosynthetic parameters (for example, Fo, Fm, and ETR) include terms for the absorption of radiation by the leaf and the fraction of absorbed light that is received by PSII. These terms are assumed to be constant, and likely are for a single sample, but can deviate between samples, especially in different mutants or under stress conditions (3). Therefore, interpreting changes in photosynthetic parameters that depend on these terms may not be possible. On the other hand, when comparing ratios of fluorescence parameters (e.g., Fv/Fm), the assumed terms cancel out and comparison between samples becomes informative.
Acknowledgment This work was supported by the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, US, Department of Energy (FWP number 449A449B). References 1. Butler, W.L. (1978) Energy distribution in the photochemical apparatus of photosynthesis. Annu. Rev. Plant Physiol. 29, 345–378. 2. Kautsky H, Apel W, Amann H. (1960) Chlorophyllfluoreszenz und kohlensäureassimilation. XIII. Die fluoreszenkurve und die photochemie der pflanze. Biochem. Zeit. 322, 277–292 3. Baker, N.R. (2008) Chlorophyll fluorescence: a probe of photosynthesis in vivo. Annu. Rev. Plant Biol. 59, 89–113. 4. Maxwell, K. and Johnson, G.N. (2000) Chlorophyll fluorescence - a practical guide. J. Exp. Bot. 51, 659–668. 5. Govindjee (2004) Chlorophyll a fluorescence: a bit of basics and history. In, Chlorophyll a Fluorescence: A Signature of Photosynthesis (Papageorgiou, G.C. and Govindjee, eds.), Springer, Dordrecht, Netherlands, pp. 1–42. 6. Schreiber, U., Schliwa, U. and Bilger, W. (1986) Continuous recording of photochemical and non-photochemical chlorophyll fluorescence quenching with a new type of modulation fluorometer. Photosynth. Res. 10, 51–62.
7. Schreiber, U. (2004) Pulse-AmplitudeModulation (PAM) fluorometry and saturation pulse method: an overview. In, Chlorophyll a Fluorescence: A Signature of Photosynthesis (Papageorgiou, G.C. and Govindjee, eds.), Springer, Dordrecht, Netherlands, pp. 279–319. 8. Duysens, L. N. M., Sweers, H. E. (1963) Mechanism of two photochemical reactions in algae as studied by means of fluorescence. In, Studies on Microalgae and Photosynthetic Bacteria (Tamiya, H. ed.), Univ. Tokyo Press, Tokyo, Japan, pp. 353–372. 9. Genty, B., Briantais, J. and Baker, N.R. (1989) The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence. Biochim. Biophys. Acta 990, 87–92. 10. Björkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77 K among vascular plants of diverse origins. Planta 170, 489–504. 11. Adams, W.W. III, Demmig-Adams, B. (2004) Chlorophyll fluorescence as a tool to monitor
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plant response to the environment. In, Chlorophyll a Fluorescence: A Signature of Photosynthesis (Papageorgiou, G.C. and Govindjee, eds.), Springer, Dordrecht, Netherlands, pp. 583–604. 12. Guadagno, C., Virzo De Santo, A. and D’Ambrosio, N. (2010) A revised energy partitioning approach to assess the yields of non-photochemical quenching components. Biochim. Biophys. Acta 1797, 525–530. 13. Genty, B., Goulas, Y., Dimon, B., Peltier, J. M., Moya, I. (1992) Modulation of efficiency of primary conversion in leaves, mechanisms involved at PSII. In, Research in Photosynthesis, Volume 4 (Murata, N. ed.), Kluwer Academic Publishers, Dordrecht, Netherlands, pp. 603–610. 14. Li, X.-P., Björkman, O., Shih, C., Grossman, A. R., Rosenquist, M., Jansson, S., and Niyogi, K. K. (2000) A pigment-binding protein essential for regulation of photosynthetic light harvesting. Nature 403, 391–395.
15. Li, X.-P., Müller-Moulé, P., Gilmore, A. M., and Niyogi, K. K. (2002) PsbS-dependent enhancement of feedback de-excitation protects photosystem II from photoinhibition. Proc. Natl. Acad. Sci. USA 99, 15222–15227. 16. Schreiber, U., Bilger, W. and Neubauer, C. (1994) Chlorophyll fluorescence as a nonintrusive indicator for rapid assessment of in vivo photosynthesis. In, Ecophysiology of Photosynthesis (Schulze, E.D., Caldwell, M., eds.), Springer, Berlin, Germany, pp. 49–70. 17. Baker, N. R., Oxborough, K., Lawson, T., and Morison, J. I. (2001) High resolution imaging of photosynthetic activities of tissues, cells and chloroplasts in leaves. J. Exp. Bot. 52, 615–621. 18. Kramer, D., Johnson, G., Kiirats, O., and Edwards, G. (2004) New fluorescence parameters for the determination of QA redox state and excitation energy fluxes. Photosynth. Res. 79, 209–218.
Chapter 17 Gas Exchange Measurements for the Determination of Photosynthetic Efficiency in Arabidopsis Leaves Giles Johnson and Erik Murchie Abstract Photosynthesis is one of the most readily measured metabolic processes in a plant, with fluxes being measurable non-invasively even under field conditions. In this chapter, two principal approaches are described to measure photosynthesis – O2 evolution as determined using an O2 electrode, and CO2 fixation which can be quantified using an infrared gas analyser. The advantages and disadvantages of these different methods, as applied to Arabidopsis, are discussed, and some major forms of analysis are described. Key words: Photosynthesis, Assimilation, Gas exchange, Light response curve, CO2 response curve
1. Introduction Photosynthesis is one of only two metabolic processes (the other being respiration) that it is possible to routinely monitor in real time in intact plant material. Photosynthesis involves the production of oxygen, from the splitting of water, and the fixation of CO2, to form carbohydrate, and both O2 evolution and CO2 fixation are readily measured. Commercially available systems allow either of these parameters to be measured in laboratory or field conditions, and it is possible to assemble systems that measure both simultaneously (see Note 1). In addition, measurements of gas exchange can be combined with chlorophyll fluorescence analysis (see Chapter 15, Vol. 2) or other forms of optical spectroscopy to obtain detailed information about a range of individual steps in the photosynthetic pathway. The decision about whether to measure using CO2 uptake or O2 evolution (or both) may depend on the resources available and the nature of the material to be studied (see Note 1). Measurements
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of CO2 uptake using infrared gas analysers (IRGAs) (often sold commercially as single portable units) allow a high degree of control of the leaf environment, including CO2 concentration, and can be performed on intact plants (leaves still attached to the plant), including under field or greenhouse conditions. Measurements can even be made on whole plants of Arabidopsis. Instruments required are, however, expensive. If it is less important to maintain leaves under physiological conditions, then detached leaves can be studied using an oxygen electrode system, where saturating levels of CO2 are applied. These can be bought for a fraction of the cost and provide robust, reproducible, and straightforward measurements.
2. Materials 2.1. Plant Material
A major limitation in making measurements of photosynthetic gas exchange in Arabidopsis has been the size of the plants grown. Typical experimental systems are set up to measure leaves having an area of 2.5 cm2 or more. Leaves of this size can be obtained by growing Arabidopsis plants in short days (8–10 h) for periods of up to 8 weeks, although different accessions vary substantially. Plants can be grown in 3″ diameter pots in peat-based compost at irradiances between 100 and 500 mmol/m2/s white light, and their rosettes will reach a diameter of approx. 7 cm, with a typical leaf length of 3 cm, depending on the accession used. Hydroponic systems, where root confinement is not an issue, can result in plants of a larger size. Where plants do not produce such large rosettes (e.g., certain mutants), it is possible to make measurements on detached leaves using oxygen electrode chambers (see below). Alternatively, various manufacturers produce small area chambers for CO2 analysers, often with long reach arms, allowing access to smaller plants, or chambers in which a pot can be inserted and gas exchange from the whole plant measured. In all measurements of photosynthesis, it is essential that the physiological state of the material is controlled rigorously. Expanding or senescing leaves are liable to have different photosynthetic performances and substantial changes in photosynthesis can be induced after plants begin to bolt. As far as possible, one should only make measurements on the youngest fully expanded leaf. There can also be diurnal variations in photosynthesis, which may need to be accounted for in experimental design. Where many measurements are being performed over several hours, it is important to randomise the timing of replicate measurements on different samples.
2.2. O2 Evolution Measurements
The most widely used method for measuring O2 evolution by intact leaves is the Clark-type oxygen electrode (Fig. 1). Instruments developed especially for photosynthesis research are based on systems
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Fig. 1. Diagram showing the assembly of an oxygen electrode. A drop of saturated KCl solution is placed on the Pt cathode and filling the trough above the Ag anode. A piece of cigarette paper and a piece of PTFE membrane are placed over the Pt cathode. An O-ring is then pushed over the membrane and paper, holding them in place over the cathode. Care should be taken to ensure that the paper reaches into the KCl solution over the anode, to make a salt bridge between the two electrodes. Ensure that the PTFE membrane is placed on the electron smoothly and is clean of fingerprints. Excess membrane and any overflowing electrolyte should be removed before placing the electrode into the chamber, to discourage crystallisation of KCl which can damage the electrode. When placing the electrode into the electrode chamber, care should be taken to ensure that all O-rings are present. The electrode should be placed tightly in the chamber but do not over tighten, as this can damage the membrane.
developed by David Walker and manufactured by Hansatech Instruments (King’s Lynn, UK) (1). Similar systems for measuring liquid samples can be applied to isolated chloroplasts or thylakoid membranes; however, such measurements lie outside the scope of this review. For measurements on intact leaves, changes in O2 in the gas phase are measured in a sealed chamber. In a sealed system, it is necessary to supply a source of CO2 that is sufficient to be saturating for photosynthesis throughout the measurement. This is supplied in the form of a 1 M NaHCO3 (pH 9, adjusted using 1 M NaCO3). A piece of felt matting soaked in this buffer is placed at the bottom of the leaf chamber. The leaf is placed on matting soaked in water and protected from contact with the NaHCO3 buffer using a piece of foam as a spacer. Note that the efficacy of the buffer declines over time and so it should be made fresh, approximately weekly. To set up the oxygen electrode, a drop of saturated solution of KCl is placed on the Pt cathode and covering the Ag anode (Fig. 1). A piece of cigarette paper (Rizla blue or similar) and then a piece of PTFE membrane (supplied by Hansatech) are placed over the cathode and fixed to the electrode using the O-ring provided with the electrode. The electrode then needs to be inserted into the gas chamber, attached to the Hansatech control unit, switched on and left to equilibrate for at least 20 min prior to measurements. Once assembled, the electrode can usually be used for up to a week; however, if it has been left for longer periods, it should be disassembled and cleaned. If allowed to dry, KCl crystals
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form and can damage the electrode. Electrodes also need regular cleaning to remove oxide deposits on the silver. This can be performed using commercial silver polish or aluminium oxide paste; however, these can damage the electrode and shorten its life. Hansatech now recommend using their specialised cleaning solution. When cleaning and assembling the electrode, care should be taken to prevent the electrical connections from coming into contact with water or solutions. Wet contacts can lead to electrode malfunction and contact with chemicals may cause corrosion (see Note 2 for trouble shooting). Prior to measuring with an oxygen electrode, it is necessary to perform a calibration. Using a 1 mL disposable syringe, inject 1 mL of air into the chamber through the gas tap and then close the tap to seal in the gas. The change in voltage across the electrode is recorded using a chart recorder or computer software. Following injection of air, the signal from the electrode should reach a new stable value after approx. 30–60 s. The increase in the voltage output from the O2 electrode control unit induced by injecting 1 mL of air into the chamber is equivalent to 2,558/T mmol O2, where T is the chamber temperature in degrees Kelvin. The rate of O2 evolution or uptake under any given condition is measured as the change in the oxygen electrode signal over time. This is normalised to the amount of leaf material in the chamber, usually expressed either as leaf area (mmol O2/m2/s) or the amount of chlorophyll in the leaf (mmol O2/mg Chl/h). Other sensors for measuring O2 are usually limited in their ability to measure small changes in O2 concentration against a background high concentration of atmospheric O2. Sensors do exist that claim to be able to measure ±1 mL/L against background atmospheric concentrations of O2 (e.g., Qubit Systems Differential O2 Analyzer, Qubit Systems, Kingston, ON, Canada), allowing measurements of O2 evolution in flow-through systems; however, such systems need to be combined with a custom made leaf chamber. Simultaneous measurements of O2 and CO2 exchange can be measured in such systems and can provide useful information about leaf physiology, for example to estimate the rate of electron flow to alternative electron acceptors such as nitrate, but these methods lie beyond the scope of this review. 2.3. CO2 Uptake Measurements
A number of different commercially available systems can be used for measurements of CO2 exchange, and users should carefully refer to information provided by manufacturers (see Note 1). The basic principles are, however, the same, regardless of the system used. An intact leaf, still attached to the plant, is clamped into an airtight chamber (or cuvette) and a gas with known concentrations of CO2 and H2O is passed through the chamber at a constant flow rate. The amount of CO2 and H2O produced or consumed in the chamber is measured as differences in absorbance in IR analyser
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cells of gas that has passed though the chamber, relative to gas that bypasses the chamber. Knowing the rate at which gas is flowing through the chamber, it is possible to calculate the rate of gas exchange occurring between the leaf and air. Most systems are referred to as ‘open’ systems because they do not recycle gas but rather take in ambient supply, adjust the CO2 and H2O content, and then eject the gas after it has passed through the chamber and analysers. Gas exchange analysis using an IRGA can provide a large amount of information on the photosynthetic rate, photosynthetic efficiency, and the biochemical/physiological components that are limiting photosynthesis of the leaf contained within the chamber. Much of this is based on the work of von Caemmerer and Farquhar (2) which has been expanded and developed since its publication. Calculated parameter values are usually provided instantly and automatically from on-board software. Curve fitting and modelling is usually performed on a separate computer. The most commonly used single-point measurements generated by IRGAs are CO2 assimilation (A), stomatal conductance to water vapour (Gs), transpiration rate (E or T), internal leaf CO2 concentration (Ci), and vapour pressure deficit (VPD). Analysers measure the difference in gas concentrations between a reference line and the leaf cuvette (sample) and, with flow rate, this is used to calculate the flux in CO2 and H2O per unit leaf area. The total conductance of the leaf to water vapour can be calculated from the following values: the humidity in the internal leaf air spaces (assumed to be saturated), the leaf temperature (this is measured), and the humidity external to the leaf. The conductance to H2O is calculated from Fick’s law of diffusion (2). Stomatal conductance is calculated by removing the boundary layer contribution. This is directly used to give the CO2 conductance (correcting for the physical properties of the two molecules). The calculation for the exchange of H2O (transpiration) and the assimilation of CO2 (photosynthesis) are linked because the former can be substantial and dilutes the CO2 concentration in the small chamber, requiring a correction. The CO2 conductance can then be used to calculate Ci, which is called the sub-stomatal or the mesophyll cell wall CO2 concentration. This is often assumed to be the in vivo substrate concentration for Rubisco, the enzyme that directly fixes CO2, allowing the generation of A versus Ci curves. However, calculations for mesophyll conductance itself are becoming more common. IRGAs are generally more stable than O2 electrodes, and newer models are much more stable and forgiving than older ones. Nevertheless, they are delicate instruments and require handling with care and treating with respect (see Note 3). It is strongly advisable to consult the manufacturer’s literature and to search the troubleshooting guides. The guide here provides some basic
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recommendations for the user to enhance awareness and below are listed some common errors, based on the authors’ experience (see Note 3). One also needs to be sure that it is the machine giving the error and not the leaf! Unreasonably high expectations of photosynthetic performance in stressed plants are common. 2.3.1. Calibration Issues
It is normally necessary to allow a short time for analysers to warm up after switch-on, and the manufacturer’s advice should be followed. Similarly, IRGAs require frequent zero-setting to ensure that a common ‘base-line’ is used for all measurements. There are actually different types of ‘zeroing’ and the terminology differs between models. Again, refer to the manufacturer’s instructions. The usual advice is to perform a ‘full’ zero calibration after switching on the machine and before starting the measurements. This requires an empty cuvette and is done by removing all of the CO2 and water from the system. Other types of calibration involve the rapid short-term equilibration of reference and analysis lines to remove small differences in humidity and temperature. This can be done with a leaf in the chamber and is frequently used when there are small differentials. It is particularly important, where the CO2 concentration and light intensity are being changed frequently. Some machines perform these operations automatically. Note that ‘spanning’ of the analysers against gas of a precise CO2 concentration is a much more complex procedure and performed much less frequently with modern IRGAs (in older models, it was necessary on a daily basis). This is, however, part of the essential maintenance.
2.3.2. Cuvette Type and Conditions
Ideally, measurements of photosynthesis should be made on the largest leaf area possible, to maximise the gas differentials and so the accuracy of the measurement. If only small leaves are formed, it may be necessary to measure on whole plants and some manufacturers (e.g., Li-Cor, http://www.licor.com) produce chambers in which whole plant pots can be placed. It should be remembered in such cases that measurements are being made not only on the plant (including leaves of different ages), but also on any soil microbes. Most manufacturers now produce specialised Arabidopsis chambers with long reach arms. These, however, have the drawback that environmental conditions and gas mixing may not be as good as in standard chambers. Thus, if plants can be grown that can be measured in standard chambers, this is preferable. Where small leaves are being measured, which do not fill the leaf chamber, this can give problems with estimating the leaf area. For some IRGA models, this has to be entered before making measurements and is difficult to estimate without damaging the plant. In such cases, it may be necessary to enter an arbitrary value and recalculate data post hoc. Other models allow correction after measurement.
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Where leaf area is small or photosynthesis low, accuracy can be somewhat increased by reducing gas flow (and so increasing the absolute differential in gas composition). On the other hand, it is common to use as high a gas flow rate as possible (e.g., 200–500 mmol/s) to speed up the response time of the measurements. It must be noted that the high flow rate typically used in IRGAs reduces boundary layer conductance substantially (i.e., they increase air mixing in the chamber). Boundary layer conductance is generally a characteristic of the chamber but variation needs to be taken into account. The user must decide on the temperature, humidity, CO2, and light intensity settings according to the problem in hand. Other conditions should be set according to advice given below. Some devices, especially older ones, use a gas source which is simply drawn in from ambient. If this is the case, the inlet tube must be positioned away from enclosed spaces and sources of combustion where CO2 will accumulate. 2.4. Light Sources
To measure a rate of photosynthesis, it is necessary to supply energy in the form of light. In IRGA systems used in the greenhouse or field, this can be in the form of sunlight. However, natural light intensities can vary rapidly over orders of magnitude and so, unless there are specific reasons to use natural light, it is normal to supply light from an artificial source. In the laboratory, where ambient light levels are low, this is essential. Many commercial light sources are available, having different advantages and disadvantages. Importantly, the spectrum of different lights may be important. Traditionally, measurements have been made using halogen light sources. These produce white light with a relatively even distribution of wavelengths but with, however, a significant amount of light in the far-red part of the spectrum (>700 nm). It is necessary to filter out longer wavelength light, using heat reflecting glass filters, to avoid heating of leaves. Increasingly, systems are becoming available that use LED lights. Red LEDs produce light that is optimal for photosynthesis, corresponding to the red absorption maximum of chlorophyll; however, it needs to be borne in mind that red light is less efficient at controlling stomatal opening and so stomatal behaviour will not be the same as in white light. This is not a problem in O2 electrode systems, where stomatal resistance is overcome with high CO2 concentrations. However, in IRGA systems, especially where measurements are made at ambient or varying CO2 concentrations, this may be a problem. Most commercial systems, therefore, use a mixture of red and blue LEDs. White LEDs, although appearing most like sunlight to our eyes, actually either mix red, blue, and green lights or produce a mixture of blue and green-yellow light. These are, therefore, rather inefficient at driving photosynthesis. Whatever the source chosen, it should be remembered that experiments using different light sources cannot be directly compared and details of the light source, including any filters used, should always be reported.
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In measurements of photosynthesis, light levels should preferably always be expressed as a quantum flux density, in mmol/m2/s estimated using a photosynthetically active radiation (PAR) metre (available from, e.g., Li-Cor [http://www.licor.com] or Skye [http:// www.skyeinstruments.com]).
3. Methods Measurements of photosynthesis can be performed in a variety of ways depending on the scientific questions being asked. In the following, we first outline the general principles of making a simple measurement (Subheading 3.1), and then describe a series of procedures that use this approach in more complex measurements (Table 1). Simple measurements of steady-state photosynthesis can be made under growth conditions, to estimate actual photosynthesis (Subheading 3.1), or under saturating conditions, to estimate maximal photosynthesis and assay the overall photosynthetic capacity (Subheading 3.2). Where differences are seen in these basic measurements, a more detailed understanding of what gives rise to these differences can be obtained by measurements of light (Subheading 3.3) or CO2 (Subheading 3.4) response curves.
Table 1 A simple decision chart showing the types of photosynthesis measurement described in this guide and the possible motivation for performing them Do I just need a single rate of photosynthesis, transpiration, and stomatal conductance?
Yes
For in situ rates of photosynthesis, perform the measurements under conditions used for growth (see Subheading 3.1) For photosynthetic capacity, use a light-saturated rate (Amax, Pmax) (see Subheading 3.2)
Do I need quantum yield, light compensation point, or overall leaf radiation use efficiency?
Yes
Light response curve (see Subheading 3.3)
Do I need information on Rubisco activity, conductance, and electron transport?
Yes
A vs. Ci curve (ACi curve) (see Subheading 3.4)
To prevent over-heating of the leaf cuvette by IR radiation
a
In a growth chamber, measure with no environmental control and ambient light In laboratory or outsidea, simulate conditions Mostly Rubisco-limited: use ambient CO2 Mostly RuBP limited: use saturating CO2
Do I need to know about photorespiration? If yes, combine ACi and chlorophyll fluorescence with O2 control
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Measurements of photosynthesis under any given set of conditions can be made using either an oxygen electrode or an IRGA. To measure photosynthetic rate, place a leaf (or leaf piece) of known area into the appropriate gas exchange chamber. Leave to equilibrate in the dark for 5 min. Measure the rate of gas exchange over a period of 1–2 min. Illuminate the leaf for 20–30 min until a stable rate of gas exchange is attained. Measure the rate of O2 evolution (see Subheading 2.2) or CO2 uptake (see Subheading 2.3). The net rate of photosynthesis is equivalent to the rate of gas exchange in the light. The gross rate of photosynthesis is taken to be the difference between the light and dark rates of gas exchange; however, it should be remembered that the rate of respiration may be different in the light compared to that recorded in the dark. Data recorded are commonly expressed relative either to the area of leaf (mmol/m2/s), or to the chlorophyll content of the leaf (mmol/mg Chl/h). The chlorophyll content of the leaf piece can be estimated at the end of the measurement. Extract chlorophyll by homogenising the leaf piece of known area in a total volume of 10 mL of 80% (v/v) acetone in a pestle and mortar. Centrifuge the extract for 5 min at 3,000 × g. The absorbance of the resulting solution is then measured in a glass cuvette at 663.6, 645.6, and 750 nm. The concentration of chlorophyll in the extract in mg/mL is given as: [Total chlorophyll] = 17.76 A(646.6 - 750) + 7.34 A(663.6 - 750) (1) where A(x−750) is the difference in absorbance between wavelength x and 750 nm (3). Expressing gas exchange data in these different ways can give different information, and it can be informative to compare the two. For example, plants with reduced chlorophyll content may have the same rate of photosynthesis per unit area, with there being an increased rate per unit chlorophyll. This would indicate that the leaf content of photosynthetic reaction centres is constant but that the amount of chlorophyll per reaction centre (the size of the light harvesting antenna) is reduced. Data can also be expressed relative to leaf (fresh or dry) weight; however, this is less common.
3.2. Measurements of Photosynthetic Capacity (Amax ; Pmax )
The rate of photosynthesis achieved by a leaf will vary depending on the light absorbed by that leaf and on the CO2 concentration inside the leaf (which is in turn a function of the external CO2 concentration and the stomatal conductance). To estimate the maximum possible rate of photosynthesis, measurements should be made under conditions that remove these limitations. In an O2 electrode chamber, the concentration of CO2 provided by the NaHCO3 buffer will be high (~5%) and sufficient to give a saturating Ci. In IRGA systems, the gas concentration can typically be set to 2,000 mL/L (0.2%; often expressed as mmol/mol or ppm).
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This is usually sufficient to be saturating for CO2 fixation, but in leaves with very low stomatal conductance (e.g., drought stressed plants) it may still be limiting. The irradiance required to saturate photosynthesis varies depending on the conditions in which plants have been grown. For Arabidopsis grown in a growth cabinet, 1,000 mmol/m2/s light is usually sufficient to saturate photosynthesis; however, this may need to be confirmed by performing measurements at different irradiances. Measurement at high irradiances (>2,000 mmol/m2/s) may give rise to progressive photoinhibition (light-induced damage to the photosynthetic apparatus), resulting in a declining rate of gas exchange over time. Mutants may saturate photosynthesis at lower irradiances and may be more vulnerable to photoinhibition. The rate of photosynthesis can be monitored over time, following the onset of illumination, and should increase to a stable level. If it increases and then declines, this is a sign of photoinhibition. Further indications of photoinhibition can be obtained by making simultaneous measurements of chlorophyll fluorescence (see Chapter 15, Vol. 2). 3.3. Light Response Curves
A fuller understanding of the functioning of the photosynthetic apparatus is obtained by measuring rates of photosynthesis as a function of the substrate concentration. Measurements at different light intensities can provide a measure not only of the maximum rate of photosynthesis achieved by a leaf, but also of the maximum quantum yield and light compensation point (see Fig. 2). The most robust light response curves are measured using different leaves for each irradiance. Leaves are measured at a given irradiance for a fixed time period, sufficient to allow photosynthesis to reach a steady state (20–30 min). By using different leaves for each measurement, individual leaves are less sensitive to damage accumulated during long measurements. This is especially true if measurements of gas exchange are to be correlated with chlorophyll fluorescence – measurements of non-photochemical quenching in particular are sensitive to photoinhibition. Rapid light response curves can be measured using a single leaf for all irradiances. Leaves are firstly pre-illuminated at a saturating irradiance for a period of 20–30 min, until a steady rate of photosynthesis is attained. This ensures that stomata are open and that enzymes and metabolites involved in CO2 fixation are fully active. The irradiance is then lowered in steps, with each irradiance being maintained for 2 or 3 min. The rate of photosynthesis is estimated at each irradiance. To determine whether there are any timedependent changes in photosynthesis, the leaf can be returned to the original irradiance either directly or step-wise and any hysteresis in the response identified. Modern instruments usually come with programs to allow automation of such measurements. Light response curves can be analysed in various ways. At the most simple, it is possible to estimate the maximum quantum yield
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Fig. 2. Light response curves from leaves of Arabidopsis thaliana (accession Ws-2) grown at 100 mmol/m2/s (circles) or then transferred to 400 mmol/m2/s (triangles) for 9 days. Growth light was provided by high-frequency fluorescent lamps. Net CO2 fixation was measured using a CIRAS-1 IRGA (PP Systems) at an external CO2 concentration of 2,000 mL/L with light provided by a quartz halogen lamp (Schott KL-1500). Data were fitted with a non-rectangular hyperbola using OriginPro 8 software (Origin Lab, Northampton, MA, USA). Values for quantum yield (F) Q and Pmax resulting from curve fitting are given for each curve. Note how these values change depending on the conditions in which the plants were grown. See ref. 10 for experimental details.
of photosynthesis by measuring the gradient of a straight line fitted through the lowest light intensities. This will, however, tend to underestimate the quantum yield. Alternatively, data can be mathematically fitted using a non-rectangular hyperbola:
P = (FI + Pmax - ((FI + Pmax )2 - 4QFIPmax )0.5 / 2QZ
(2)
where P is the rate of photosynthesis at any given irradiance (I) (4–6). This allows estimation of the maximum rate of photosynthesis, where this may not be attained with the irradiances used and of F, the quantum yield of photosynthesis. The function Q describes the curvature of the light response and is a complex function, depending on the leaf thickness and absorption as well as on the composition of the photosynthetic apparatus and the physiological status of the leaf. A full discussion of this is beyond the scope of this review (see refs. 4–6 for further discussion). 3.4. ACi Curves
Plotting assimilation (A) against Ci is commonly used to separate the various limiting steps in photosynthesis, notably Rubisco activity, ribulose bisphosphate (RuBP) regeneration, and triose phosphate utilisation. Stomatal limitation can also be estimated. Once in the leaf chamber, the leaf is exposed to a series of CO2 concentrations. A typical procedure would be to equilibrate the
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leaf at ambient CO2 levels (currently 385 mL/L CO2) and then expose the leaf to sequentially lower values down to a typical minimum of between 10 and 40 mL/L. ACi curves should be performed quite quickly (no more than a few minutes at each CO2 level) since it relies on instantaneous responses of Rubisco activity, and it is desirable to avoid any CO2-induced stomatal responses. Higher values of CO2 are then used. It is useful to take advantage of rapid zeroing procedures between CO2 levels as gas analyser sensitivity changes with CO2 concentration (see above). An example of what an ACi curve may look like is shown in Fig. 3. The slope of the lower linear phase of the ACi response should denote the amount of active Rubisco in the leaf (Vc,max). At higher CO2 levels, A is increasingly limited by the regeneration of RuBP, the substrate for the CO2 fixing enzyme Rubisco, and this is commonly given as representing a measure of the maximum rate of linear electron transport, Jmax. A further limitation is provided by the utilisation of triose phosphate in the leaf (VTPU). Mesophyll conductance can also be estimated directly from ACi curves or in combination with chlorophyll fluorescence (7). It is possible to fit curves according to the equations given in the literature (7) and derive these values for one’s self. However, software that is capable of fitting curves to ACi data is available. Software packages may differ in terms of assumptions made. For the purposes of this guide, the reader is referred to Sharkey et al. (7) and Long and Bernacchi (8). A useful software utility, described in Sharkey et al. (7), is available for download as an Excel spreadsheet from Plant Cell and Environment (http://www.blackwellpublishing.com/PlantSci/pcecalculation/).
Fig. 3. Typical ACi curve showing the separation of phases in which Rubisco, RuBP, or triose phosphate are limiting for photosynthesis. The three phases are fitted to the data to determine the variables Vc,max, Jmax, and VTPU as described in the text.
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In addition, a simple method for using ACi responses to separate mesophyll and stomatal limitations has been described (9). The assimilation rate (A) at ambient CO2 is subtracted from the rate that would occur if stomatal limitation was zero, which is the value at which the Ci equals the ambient CO2 value (A0). This gives the proportional drop in A that should be caused by stomata, i.e., the relative limitation on photosynthesis caused by stomata (L):
L = (A0 - A) / A0
(3)
The rate of leaf photorespiration can also be measured if the user has access to combined IRGA and chlorophyll fluorescence device (most IRGA models come with optional fluorimeter modules; see Chapter 15, Vol. 2 for technical details on chlorophyll fluorescence) that is connected to two interchangeable supplies of air with 2 and 20% O2. This can be achieved by switching between gas cylinders of air and 2% O2. The proportion of electrons that are diverted to photorespiratory and non-photorespiratory sinks can be calculated. Readers are referred to Long and Bernacchi (8) for details of this technique.
4. Notes 1. Systems available for measuring photosynthetic gas exchange. It is not our intention, or appropriate, to provide an exhaustive buyer’s guide. When purchasing, we consider these to be the most important considerations. In all cases, the user must define the scientific objectives and decide what type of measurement is appropriate. (a) The type of measurement to be made: An O2 electrode allows for budget measurements of photosynthesis, but with those being under non-physiological conditions; this can be useful for, e.g., comparative measurements but limits the range of conditions that can be explored. IRGAs allow for more refined measurements but at a considerably higher level of investment. This decision may be influenced by the type of data required. Table 1 gives advice on the type of measurement required depending on the information sought. (b) The level of environmental control over the leaf: This should be as high as possible but often raises the price and may not be required. Photosynthetic capacity measurements absolutely require saturating irradiance and often this means the addition of a light source. Some devices have the ability to raise the cuvette humidity as well as reduce it.
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(c) The rapidity of measurements: Some machines have the analysers positioned next to the cuvette; this speeds up the response time. Higher flow rates can be advantageous if many rapid measurements are required. (d) Software control and programing: The ability to program the device to perform automatic ACi and light response curves and automatic logging over long time periods. (e) Inbuilt capacity to perform (modulated) chlorophyll fluorescence measurements and in some cases imaging: This is useful for advanced measurements of Photosystem II activity and efficiency (see Chapter 15, Vol. 2). (f) Ability to connect the analyser to alternative cuvettes: This is normally provided as an extra. For example, some manufacturers provide whole-plant Arabidopsis cuvettes. (g) Size, weight, portability, and battery life: This varies greatly. A few devices are bulky and users may have difficulty if travelling. The only manufacturer of O2 electrodes for measurements on leaves of which we are aware is Hansatech Instruments, King’s Lynn, UK (http://www.hansatech-instruments.com/). Qubit Systems, Kingston, ON, Canada (http://www.qubitsystems.com/) supply a differential oxygen sensor which can be incorporated in a flow-through gas exchange system; however, this is not currently available in a complete system. A number of different manufacturers produce complete portable IRGA systems, with varying degrees of sophistication (and price). The following lists the main suppliers (limited largely to those familiar to the authors): ADC Bioscientific, Great Amwell, Herts, UK (http://www.adc. co.uk) Heinz Walz, Effeltrich, Germany (http://www.walz.com) Li-Cor Biosciences, Lincoln, NE, USA (http://www.licor. com) Photon Systems Instruments, Brno, Czech Republic (http:// www.psi.cz) PP Systems, Amesbury, MA, USA (http://www.ppsystems. com) Qubit (teaching system) Kingston, ON, Canada (http://www. qubitsystems.com) 2. Common problems in using an oxygen electrode. Oxygen electrodes are notoriously fickle instruments and making them work consistently is something of an art. The following is a list of the most common problems we have experienced and solutions available:
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(a) Signal is noisy: This is usually caused by a damaged PTFE membrane. This can commonly occur during the assembly of the O2 electrode chamber or may be a result of the electrode drying with age. Disassemble and reassemble the electrode. (b) Electrode signal drifting, or slow to respond when air injected during calibration: This is usually a result of a dirty or poorly assembled electrode. Clean the silver electrode with recommended cleaning solution, dry thoroughly, and reassemble. Slow response can be caused if the PTFE membrane is dirty. Take care to avoid touching the membrane surface during assembly. (c) During calibration the signal rises and then falls again: This is because the chamber is not gas tight. Check that taps are sealed and that all O-rings are present and intact. O-rings can be greased lightly with vacuum grease. If the source of the leak cannot be identified, it may be necessary to instal new O-rings. (d) Signal fluctuates or oscillates: This type of behaviour can be caused by electrical interference or temperature fluctuations. Move the instrument away from possible sources of electrical interference, ensure that all connections are wired correctly and that connectors are dry and clean. We have in the past observed regular signal oscillations that were found to be occurring in time with the heating and cooling of an air conditioner. The solution in that case was to place the O2 electrode chamber in a polystyrene box. 3. Common problems in using an IRGA. (a) Before you start: It is essential to check that the IRGA is running satisfactorily before placing a leaf in the chamber (it is surprising how often this is not done!). Check that the gaskets around the leaf give a good seal but are not placing undue physical pressure on the leaf; when not in use, chambers with spring closing should be stored open to avoid pressure on the gaskets, and these should be replaced periodically, when they become compressed. Confirm that conditions in the cuvette are as set and that the photosynthesis and transpiration is zero (i.e., no differential). If this is not the case, it could indicate a leak. (b) Leaks are perhaps the most notorious source of error although are less common with newer equipment. The best way of checking for leaks is to ensure that the set values of CO2 and H2O are attained with no leaf in the chamber. The most common position for a leak is around the foam gaskets that hold the leaf itself. You can check for a leak in this region by breathing out heavily around the gaskets.
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(c) Calibration: Errors will result when the analysers are not fully calibrated (see above). Make sure that baseline and differential zeroing is performed as recommended by the manufacturer after changes in conditions. (d) Leaf temperature measurements should be checked. If a thermocouple is used, this should be touching the under surface of the leaf. (e) Leaf area: Check that leaf area has been measured accurately and entered into the software calculation. References 1. Delieu, T. J., and Walker, D. A. (1983) Simultaneous measurement of oxygen evolution and chlorophyll fluorescence from leaf pieces. Plant Physiol. 73, 534–541. 2. von Caemmerer, S., and Farquhar, G. D. (1981) Some relationships between the biochemistry of photosynthesis and the gas exchange of leaves. Planta 153, 376–387. 3. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) determination of accurate extinction coefficients and simultaneous-equations for assaying chlorophyll-a and chlorophyll-b extracted with 4 different solvents - verification of the concentration of chlorophyll standards by atomic-absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394. 4. Ögren, E., and Evans, J. R. (1983) Photosynthetic light response curves 1. The influence of CO2 partial pressure. Planta 189, 182–190. 5. Ögren, E., Jakobsen, I., and Evans, J. R. (1983) Photosynthetic light response curves 2. Gradients of light absorption and photosynthetic capacity. Planta 189, 191–200. 6. Akhkha, A., Reid, I., Clarke, D. D., and Dominy, P. J. (2001) Photosynthetic light
response curves determined with the leaf oxygen electrode: minimization of errors and significance of convexity term. Planta 214, 135–141. 7. Sharkey, T. D., Bernacchi, C. J., Farquhar, G. D., and Singsaas, E. L. (2007) Fitting photosynthetic carbon dioxide response curves for C-3 leaves. Plant Cell Environ. 30, 1035–1040. 8. Long, S. P., and Bernacchi, C. J. (2003) Gas exchange measurements, what can they tell us about the underlying limitations to photosynthesis? Procedures and sources of error. J. Exp. Bot. 54, 2393–2401. 9. Hall, D. O., Scurlock, J. M. O., BolharNordenkampf, H. R., Leegood, R. C., and Long, S. P. (1993) Photosynthesis and Production in a Changing Environment; a Field and Laboratory Manual. Chapman and Hall, London, UK. 10. Athanasiou, K., Dyson, B. C., Webster, R. E., and Johnson, G. N. (2010) Dynamic acclimation of photosynthesis increases plant fitness in changing environments. Plant Physiol. 152, 366–373.
Chapter 18 Measurement of the DpH and Electric Field Developed Across Arabidopsis Thylakoids in the Light Steven M. Theg and Curtis Tom Abstract Measurement of the different components of the proton motive force (pmf) gives information about the coupling of proton movement within thylakoids to chemiosmotic processes such as photophosphorylation and protein transport, as well as that relating to the overall quality of a thylakoid preparation. The techniques to assess the pmf have been known for many years, as they have been applied to the most popular model plants for photosynthetic research. The emergence of Arabidopsis thaliana as the prominent model plant in developmental and genetics research prompted us to apply these techniques to thylakoids isolated from Arabidopsis chloroplasts. We describe here two spectroscopic techniques to measure the transmembrane pH gradient and electric field developed in the light in Arabidopsis thylakoids. Key words: Chloroplast, Thylakoid, Proton gradient, Proton motive force, Delta pH, Electrical potential, 9-Aminoacridine, Electrochromic shift, Arabidopsis, Pea
1. Introduction Chloroplasts are the organelles in green plants responsible for the conversion of sunlight into chemical energy, upon which all the biosphere depends. The first steps in this process are embodied in the so-called light reactions taking place in the chloroplast’s internal thylakoid system. The photosynthetic reaction centers transduce the energy of absorbed photons into redox energy subsequently utilized to transfer electrons from water to NADPH. Proton pumping during electron transfer reactions leads to a proton gradient across the thylakoid membrane which, in turn, powers the synthesis of ATP. The NADPH and ATP thus produced are then utilized in the “dark reactions” in the fixation of carbon from CO2 to carbohydrates. A central function of the light reactions is the temporary storage of energy as an electrochemical gradient of protons across
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_18, © Springer Science+Business Media, LLC 2011
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the thylakoid membrane, an intermediary on the path of the transduction of redox energy into the chemical energy contained in ATP and NADPH. The energy manifested in the proton electrochemical gradient ( ∆m H+ ), termed the proton motive force (pmf) by Mitchell (1), contains a chemical component described by the proton concentration difference in the aqueous compartments on the different sides of the membrane (DpH), and an electrical component described by the transmembrane electrical potential (Dy). When divided by the Faraday constant (F), the pmf is expressed in volts (R is the gas constant and T is temperature in Kelvin), thus:
pmf =
∆m H+ F
=
2.3RT ∆pH + ∆y . F
At room temperature the constants 2.3RT/F = 59.1 mV, so that pmf (mV) ≈ 60 ∆pH + ∆y ,
from which it is evident that a DpH of 1 unit across a membrane is energetically equivalent to a membrane potential of approximately 60 mV. It is useful for many purposes to evaluate the two components of the pmf developed in thylakoids in the light (see for instance, refs. (2–4)), and numerous methods have been elaborated to do so. These methods have generally been applied to thylakoids derived from spinach or pea, two plant species popular with photosynthesis researchers. The emergence of Arabidopsis thaliana as a model plant, and in particular, the relatively recent development of techniques to isolate its chloroplasts retaining physiological activity, has led us to extend the application of two methods for evaluating components of the pmf to Arabidopsis thylakoids. These methods, one to monitor the DpH with 9-aminoacridine (9-AA) fluorescence and the other to measure the onset and decay of the Dy using the electrochromic shift of carotenoids (ECS), are described here.
2. Materials Unless otherwise noted, all chemicals are prepared in distilled water. 2.1. Plant Growth
1. Seeds from Arabidopsis thaliana (ecotype Columbia).
2.1.1. Arabidopsis
2. Murashige and Skoog Basal Salt Mixture (MS) (Sigma–Aldrich, St. Louis, MO, USA). 3. Phytagar. 4. Sucrose.
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5. 50% (v/v) Clorox bleach (~3% sodium hypochlorite) with three drops of Tween 20 in 50 mL of solution. 6. Petri dishes. 7. 95% (v/v) ethanol. 8. Sterile distilled water. 9. Surgical tape or equivalent. 10. Laminar flow hood. 11. Plant tissue culture chamber. 2.1.2. Peas
1. Seeds from Pisum sativum var. Little Marvel (Seedway LLC, Hall, NY, USA). 2. Vermiculite.
2.2. Chloroplast Isolation
1. 2× Grinding buffer (2× GB): 0.1 M 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES)–KOH, pH 7.3, 0.66 M sorbitol, 2 mM MgCl2, 2 mM MnCl2, 4 mM Na2EDTA (ethylenediaminetetraacetic acid), 0.2% (w/v) bovine serum albumin (BSA). 2. 2× Storage buffer (2× SB): 0.1 M Tricine–KOH, pH 8.0, 0.66 M sorbitol, 6 mM MgCl2. 3. Percoll (GE Healthcare Sciences). 4. Miracloth. 5. Ascorbic acid. 6. Isoascorbic acid. 7. Glutathione. 8. 80% (v/v) acetone. 9. Waring blender with sharpened blades. 10. Swinging-bucket rotor (e.g., Sorvall HB-4 or equivalent) and suitable centrifuge (e.g., Sorvall RC 6). 11. Fixed-angle rotor (e.g., Sorvall JA-20 or equivalent) and suitable centrifuge (e.g., Sorvall RC 6). 12. 50-mL centrifuge tubes. 13. Large funnel to filter 200 mL; small funnel to filter 5 mL. 14. 500-mL beaker. 15. Whatman #1 filter paper, 70-mm diameter circles.
2.3. Thylakoid Isolation
1. Chloroplast lysis buffer: 10 mM 2-(N-morpholino) ethanesulfonic acid (MES)–KOH, pH 6.5, 5 mM MgCl2. 2. 80% (v/v) acetone. 3. 50-mL centrifuge tubes.
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2.4. Measurement of DpH with 9-Aminoacridine
1. 9-Aminoacridine (1 mM stock made up in ethanol). 2. Methyl viologen. 3. 1× Storage buffer (see Subheading 2.2). 4. Fluorimeter equipped for actinic illumination (see Note 1).
2.5. Measurement of Dy via the Carotenoid Electrochromic Shift
1. Methyl viologen. 2. 1× Storage buffer (see Subheading 2.2). 3. Sensitive spectrophotometer equipped for actinic illumination (see Note 2).
3. Methods 3.1. Plant Growth 3.1.1. Arabidopsis Growth Media and Plate Preparation
1. Add 4.4 g of MS and 10 g of sucrose to approximately 800 mL of distilled water. Stir until completely dissolved. 2. Set the pH to 5.7 with KOH. 3. Bring the final volume up to 1.0 L. 4. Add 8 g of Phytagar and autoclave for 30 min. 5. Allow the media to cool to touch and then pour plates in a laminar flow hood. 6. Allow the plates to dry and solidify in the hood. 7. Store plates upside down in a sealed bag at 4°C.
3.1.2. Arabidopsis Seed Sterilization and Growth
All of the following procedures are to be performed under sterile conditions in a laminar flow hood. Materials and instruments are sterilized by ethanol treatment and illuminated with germicidal light for 15 min prior to seed handling. 1. Place approximately 10 mg of seeds into a microcentrifuge tube. 2. Fill the tube with 1 mL of 95% ethanol and mix repeatedly by inverting for approximately 10 min. Allow the seeds to settle to the bottom of the tube or alternatively, spin briefly in a tabletop microcentrifuge. Remove the ethanol using a pipette. 3. Fill the tube with 1 mL of the bleach/Tween 20 solution. Shake or vortex for 20 min and then remove the supernatant using a pipette. 4. Fill tube with 1 mL of sterile water. Shake for 1 min and then remove the water using a pipette. Repeat this process four more times. 5. Resuspend the seeds in 1 mL of sterile water. Place the seeds on the plates by drawing them into a pipette tip and gently releasing them onto the surface of the plates.
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6. Remove excess water with a pipette and seal the plates with surgical tape. 7. Place the plates in dark at 4°C for 2 days. 8. After 2 days, allow the seeds to germinate by moving them to 20°C under 100 mmol/m2/s white light with a long-day cycle (16-h-light–8-h-dark). 9. Seedlings are harvested at 2–4 weeks. 3.1.3. Peas
1. Soak approximately 100 g seeds in distilled water for a few hours. 2. Sow the seeds on a wet bed of vermiculite in a flat plastic pan (35 cm× 20 cm× 6 cm). Cover the seeds with a thin layer of wet vermiculite. 3. Plants are grown in a controlled environment chamber at 20°C in a 12-h-light–12-h-dark cycle. 4. Seedlings are harvested at 10–14 days.
3.2. Chloroplast Isolation 3.2.1. Chloroplasts from Arabidopsis
Chloroplast isolation should be performed on ice and in dim light. All solutions are diluted to 1× as needed using distilled water. 1. Place 15 mL of Percoll and 15 mL of 2× GB into a 50-mL centrifuge tube. Add 10 mg isoascorbic acid and 10 mg glutathione. 2. Centrifuge the tube at 37,000 × g for 30 min (with the brake on) in a fixed-angle rotor (16,000 rpm in a JA-20 rotor) to create a linear gradient. Be sure to handle the tube gently after centrifugation so as not to disturb the gradient. 3. Place 5–10 g of leaves into a Waring blender in which the blades have been sharpened and add 150 mL of 1× GB containing 50 mM ascorbate. Grind briefly. 4. Pour the blended slurry into a funnel draped with Miracloth and gently squeeze the cloth to push all the liquid through into a collection beaker. 5. Divide the filtrate between four 50-mL centrifuge tubes and centrifuge at 3,000 × g with the brake on in a swinging-bucket rotor (4,000 rpm in a HB-4 rotor) for 5 min. 6. Decant the centrifuge tubes and gently resuspend each pellet in approximately 1 mL of 1× GB (without ascorbate). 7. Gently overlay the chloroplast-containing suspensions onto the preformed Percoll gradient. 8. Centrifuge the gradient at 8,000 × g for 10 min in a swingingbucket rotor with the brake off (6,500 rpm in an HB-4 rotor). 9. After centrifugation, two Chlorophyll (Chl)-containing bands will have separated in the gradient; the intact chloroplast will
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collect in the lower of the two bands, whereas the broken chloroplast will collect at the upper band. Aspirate the upper portion of the gradient down to the lower band and then transfer the lower band to an empty prechilled 50-mL centrifuge tube. 10. Bring the tube to full volume with ice-cold 1× SB and centrifuge at 1,500 × g for 5 min (3,000 rpm in an HB-4 rotor). 11. Remove the supernatant and gently resuspend the pellet in 1× SB, then bring to full volume and repeat the centrifugation at 1,500 × g for 5 min. 12. Resuspend the pellet containing intact chloroplasts in a small volume (~1 mL) of 1× SB and place on ice in the dark. 13. Measure the Chl content of the intact chloroplast suspension (see Note 3) by adding 10 mL of chloroplasts to 5 mL of 80% acetone in a glass test tube. Mix well, filter through Whatman filter paper, then read the absorbance (A) of the filtrate at 645, 663, and 720 nm. [Chl] (in mg/mL) is calculated as 4.02 × (A663 – A720) + 10.14 × (A645 – A720). 3.2.2. Chloroplasts from Peas
3.3. Isolation of Thylakoids from Intact Chloroplasts (see Notes 4 and 5)
The procedure for the isolation of chloroplasts from peas is identical to that for the Arabidopsis chloroplast isolation described in Subheading 3.2.1, except that ascorbate is omitted from the grinding buffer in step 3. See also ref. 5. 1. Centrifuge the intact chloroplasts at 1,500 × g for 5 min. Decant and discard the supernatant. 2. Resuspend the pellet in chloroplast lysis buffer to a Chl concentration of 1 mg/mL (see Note 6). Incubate on ice in the dark for 5 min. 3. Add an equal volume of 2× SB to the chloroplast lysis reaction and centrifuge at 1,500 × g for 5 min. 4. Decant the supernatant and resuspend the pellet in a small volume (~1 mL) of 1× SB. Place on ice in the dark. 5. Measure the Chl concentration of the thylakoids as in Subheading 3.2, step 13.
3.4. Measurement of DpH Across the Thylakoid Membrane with 9-Aminoacridine
9-Aminoacridine is a fluorescent amine that can be used to monitor the DpH established across the thylakoid membrane in response to illumination (6). This amine is membrane permeable in its uncharged, unprotonated state, but upon protonation becomes charged and thus membrane impermeable. Its fluorescence is quenched when it enters the thylakoid lumen, and hence, the extent of 9-AA fluorescence quenching upon illumination is a reflection of the extent to which the amine is drawn into and trapped in the lumen in response to the DpH.
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The kinetics of fluorescence quenching and the maximal quenching observed are functions of the rates of proton entry into and leakage from the thylakoid lumen, whereas the kinetics of fluorescence recovery is a function of the leakage rate only (see Note 7). The kinetic parameters (see Note 8), while not directly convertible to rates of proton movement (see Note 9), are, nonetheless, indicators of the general physiological well-being of the thylakoid preparation. Faster establishment of the maximal quenching indicates high rates of electron transport and/or good membrane integrity, and slower fluorescence recovery also indicates good membrane integrity. High-integrity membranes are expected to provide good coupling of electron transport to photophosphorylation or protein transport via the pmf. 1. 9-AA measurements in our laboratory are performed in a fluorimeter equipped for red (>650 nm) actinic illumination delivered 180° from the weak 420 nm excitation beam and 90° from the 520 nm emission direction (see Note 1). Reactions are typically performed in a stirred cuvette with a 500-mL reaction volume containing 1× SB, 20 mM methyl viologen, 20 mM 9-AA, and thylakoids at 40 mg Chl/mL. 2. After a baseline is established, the maximum, unquenched fluorescence signal is recorded by admitting the excitation beam. Fluorescence quenching is recorded with the onset of actinic illumination, which is allowed to proceed until a maximum quenching is observed. With the cessation of actinic illumination, the fluorescence returns to its original unquenched level, and we usually end the measurement by extinguishing the excitation beam and recording the original baseline again. Figure 1 shows representative traces recorded with Arabidopsis thylakoids, and for comparison, thylakoids isolated from peas. The timing sequence is typical for us, with intervals of 5, 15, 80, 145, and 5 s
Fig. 1. DpH-indicating quenching of 9-AA fluorescence using Arabidopsis and pea thylakoids. Traces were recorded as described in the text with thylakoids isolated from intact chloroplasts by osmotic lysis. Data (points) were recorded at 500 ms per point; the solid lines represent fits to the data using Eqs. 1 and 2 in Note 8, with the kinetic parameters given in Note 11. Upward and downward arrowheads, measuring beam on and off; upward and downward arrows, actinic light on and off. Fluorescence levels (arbitrary units, a.u.) corresponding to Fo, Fm, and Fq are indicated by dashed lines.
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for recording of baseline, maximal fluorescence, fluorescence quenching, fluorescence recovery, and final baseline, respectively. 3. As shown in Fig. 1, we can define Fo, Fm, and Fq as the baseline fluorescence, maximal fluorescence, and level of the fluorescence after maximal quenching, respectively. Then, the extent of fluorescence quenching in the light can be used to estimate (see Note 10) the DpH according to the equation:
Q ∆pH = log − log(V ), 1 − Q
(1)
where Q = (Fm − Fq ) / (Fm − Fo ) and V = ratio of the volume of the thylakoid lumen to the total volume (in this experiment estimated as[20 µL/mg Chl × 0.02 mg Chl] / 500 µL = 8 × 10 −4 ) (7). Using Eq. 1, the trace obtained from Arabidopsis thylakoids shown in Fig. 1 indicates that the DpH developed was 3.04 pH units. This trace displays the characteristics of physiologically healthy thylakoids, with a membrane not inordinately leaky to ions (protons) (see Note 11), and which would then be expected to be well coupled to ATP synthesis and protein transport. 4. A typical 9-AA fluorescence quenching trace for pea thylakoids is shown for comparison on the right side of Fig. 1. As mentioned above, thylakoids from pea have been characterised extensively with respect to bioenergetics parameters. These thylakoids display a faster and deeper fluorescence quenching compared to the Arabidopsis thylakoids, reaching a DpH of 3.36 pH units (see Note 12). The reason for the difference between pea and Arabidopsis thylakoids is not at present known, but may be due to many years of optimization of the pea thylakoid preparations that have not been applied to Arabidopsis, or simply to daily variation in the preparations. 3.5. Measurement of the Dy Across the Thylakoid Membrane Using the Carotenoid Electrochromic Shift
As first described by Junge and Witt (8), carotenoids in the pigment bed surrounding photosystem II undergo an electrochromic red shift in response to a transmembrane electric field. This ECS signal can be followed noninvasively as an absorbance change at 520 nm, which thus becomes an indicator of the electric field developed across the thylakoid membrane in response to short flashes of light (see Note 13). The absorbance signal is quite small, usually DA <0.01, and so it is usually measured in a spectrophotometer with sensitivity beyond that found in typical laboratory spectrophotometers (see Note 2). The initial height of the flash-induced ECS is proportional to, among other things, the magnitude of the electric field established by the flash (see Note 14). The decay of the ECS with time after the flash reports on the ability of the membrane to maintain a
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charge, which is a function of the permeability of the membrane to ions, i.e., membrane integrity. One can easily spot the effects of membrane damaging conditions or an increase in ion permeability as an increase in the rate of decay of the ECS. 1. ECS signals are recorded in our laboratory in one of two homebuilt spectrophotometers set up for actinic illumination at 90° to the measuring and detection beam (4, 9, 10). 2. Reactions are typically carried out in unstirred four-sided fluorescence cuvettes in a total volume of 2 mL (see Note 15), and contain 20 mM methyl viologen and thylakoids at 40 mg Chl/mL. 3. A baseline is recorded at 520 nm before an actinic flash is fired, and then the flash is fired and the absorbance is monitored subsequently, typically for a few seconds until the absorbance returns to the initial baseline level (see Note 16). 4. It is usually the case that the signal-to-noise ratio in this measurement is so small so as to require signal averaging to achieve sufficient resolution. If that is necessary, step 3 is repeated at approximately 10-s intervals (see Note 17) until an acceptable resolution is achieved; typically, 4–16 repetitions will suffice (see Note 18). 5. An ECS measurement performed with Arabidopsis thylakoids is shown on the left in Fig. 2, with the better characterised signal obtained with pea thylakoids on the right. The pea thylakoids show a higher initial ECS signal (see Note 19), perhaps indicating, as suggested from Fig. 1, that they are more robust than the Arabidopsis thylakoids. However, also as with the 9-AA measurements, the Arabidopsis samples display an ECS signal with characteristics of a well-coupled, healthy thylakoid preparation.
Fig. 2. Electric field-indicating absorbance change at 520 nm in Arabidopsis and pea thylakoids. Traces were recorded as described in the text and Note 16 with thylakoids isolated from intact chloroplasts by osmotic lysis. Data are represented by points, with solid lines derived from the fits using Eq. 4 in Note 19 and the kinetic parameters listed therein. Upward arrows mark the application of a 10-ms actinic pulse.
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4. Notes 1. Most of our 9-AA measurements have been made using a home-built instrument that can be configured for visible absorbance, fluorescence, or luminescence measurements. However, many commercial fluorimeters can be set up for illumination of the samples at right angles to the emission direction. Indeed, with the proper light filtering one can provide actinic illumination at 0° and 180° to the emission direction, and we (and many others) have done so with good results. An important point to consider is that neither the excitation beam nor the actinic light can be allowed to scatter into the detector. This is generally only a problem for the red actinic beam, which is intended to drive photosynthetic electron transport and so must be fairly bright. This light scattering problem can be handled by placing appropriate optical filters in the different light paths. We have found that narrow-band-pass interference filters are not sufficient on their own for this purpose, as they transmit enough light in their cut-off wavelengths to spoil the measurement. Accordingly, we use a combination of colored glass and interference filters. A good rule of thumb for selecting filters is to stack together those that are to select and block a particular color (in this case, you want to transmit the green fluorescence and block the red actinic light), hold them up to your eye, and then look directly at a bright light source. If any light can be seen through the combination, however dim, then the filters will not perform adequately. A working filter set up for us has a Corion S-10 520 narrow-band-pass filter (Newport Corp., Franklin, MA, USA) with a Corning CS4-96 colored glass filter (Kopp Glass, Pittsburgh, PA, USA) in front of the detector, and a Corion 650 high-pass filter with a Corning CS3-67 colored glass filter in line with the actinic light. 2. As with the 9-AA measurement, we often perform ESC measurements with the homemade instrument described set up in the spectrophotometer configuration; cf. (4). The selection of appropriate color blocking filters applies to this measurement as well (see Note 1), but additional difficulties are manifested because the signal is smaller than can be measured with typical commercial laboratory spectrophotometers. Appropriate flash kinetic spectrophotometers are found in many laboratories engaged in the study of the photosynthetic light reactions. A full description of the two instruments used in our laboratory to measure ECS signals (the flash kinetic spectrophotometer and the nonfocusing optics spectrophotometer [NoFOSpec]) is beyond the scope of this chapter, but can be found in refs. 4, 9, 10.
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3. In our laboratory, we often perform various experiments using intact chloroplasts, although they are not needed for the measurements described herein. Accordingly, we often prepare both intact and broken chloroplasts (which are thylakoids) at the same time. When doing so, we measure the Chl content of the intact chloroplasts prepared as described before going on to isolate thylakoids. Should only thylakoids be required, one could postpone the Chl measurement until the final step in the thylakoid preparation, i.e., Subheading 3.3, step 5. 4. Thylakoids are prepared by breaking the rather fragile chloroplast envelope membranes. This is achieved by placing the intact chloroplasts in hypotonic buffer, causing the osmotically active inner envelope membrane to swell to the point of bursting. The heavier and more robust thylakoid membranes are then separated from the broken envelope membranes by centrifugation. 5. As mentioned in Note 3, one may wish to prepare only thylakoids and not have need for intact chloroplasts in a particular day’s experiments. If so, steps 12 and 13 in Subheading 3.2.1 and step 1 in Subheading 3.3 can be skipped. That is, the pellet from the second wash after the Percoll gradient, or perhaps even after the first wash, can be resuspended directly in chloroplast lysis buffer as in Subheading 3.3, step 2. 6. We do not have experience with lysing intact Arabidopsis chloroplasts at different Chl concentrations, and we have settled on 1 mg/mL purely for historical reasons. What appears to be important is to expose the intact chloroplasts to a medium of sufficiently low osmotic strength so as to cause them to swell and break. 7. The decline in fluorescence is due to the acidification of the thylakoid lumen. The rate of this decline is determined by the balanced rates of proton deposition into the lumen and escape back into the external medium. The extent of quenching is obviously dependent on these rates as well, which match each other in the steady state when the signal plateaus. The rate of return of fluorescence after the actinic light is extinguished reflects only the rate of proton return from the lumen to the external medium (11). In the measurement described, the proton gradient is not coupled to other chemiosmotic processes (i.e., ATP synthesis or protein transport), and so release of the gradient will take place through proton leakage. 8. The time course of fluorescence decline and recovery can be fitted to kinetic equations to allow a more quantitative description of the traces. We have found that the declining and recovery phases of the signal can be fit well as first-order reactions, with the fit to the recovery phase improved considerably if it
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is treated as the sum of two rising exponentials. The falling phase is fitted to
F (t ) = Fq + (1 − Fq )e − kf t ,
(2)
where F(t) and Fq are the levels of fluorescence at time t and the maximal quenched level, respectively. The rising phase is fitted to either:
F (t ) = Fq + (1 − Fq )(1 − e − krt )
(3)
for a single exponential fit, or
(
fast
F (t ) = Fq + (1 − Fq ) 1 − f fast e − kr
t
slow
− f slow e − kr
t
)
(4)
for the sum of two rising exponentials. In Eqs. 2 and 3, kr is the rising first-order rate constant and f fast and fslow are the fractions of the signal going with the fast and slow rate constants so superscripted, respectively. Whereas Eq. 3 gives the better fit, Eq. 2 allows the use of halftimes for comparing traces, as in Fig. 1. In Eqs. 1–3 the fitted parameters are Fq, f fast (fslow = 1 – f fast), and the rate constants. 9. Quenching of 9-AA reports the DpH developed across thylakoid membranes. This can be related to the flux of protons through the lumen only (a) through a detailed knowledge of the buffering capacity of the thylakoid lumen, which is difficult to assess (12, 13), or (b) empirically and indirectly through measurements of proton pumping with a pH-indicating dye (2). 10. While 9-AA fluorescence quenching can theoretically be used to calculate the value of the light-dependent transmembrane DpH (6), problems associated with its binding to charged membranes make this calculation less than completely reliable, especially at low DpH values (2). Accordingly, we prefer to use it as a tool to estimate the DpH, and in particular, to compare the DpH values developed under different conditions. 11. The kinetic parameters for the fits (solid lines) of Eqs. 1 and 2 to the data (points) in Fig. 1 are as follows: Arabidopsis: Fq = 0.54, kf = 0.085 s−1 (or t½ = 8.15 s); ffast = 0.33, krfast = 0.045s −1 , krslow = 0.0096s −1 ; a slightly worse, but still acceptable fit to Eq. 3 gives t½ = 52 s for the recovery phase. Pea: Fq = 0.35, kf = 0.17 s−1 (or t½ = 4.01 s); f fast = 0.69, krfast = 0.050s −1 , krslow = 0.014s −1 ; a slightly less favorable, but still acceptable fit to Eq. 3 gives t½ = 24 s for the recovery phase. 12. It is noteworthy that a seemingly large difference in the amount of fluorescence quenching observed between the traces in Fig. 1 for Arabidopsis and pea thylakoids translates into a rather modest change in the calculated DpH in the two samples. It is
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Fig. 3. DpH as a function of the extent of quenching of 9-AA fluorescence. DpH was calculated from Eq. 1 with V = 8 × 10−4, Fo = 0, and Fm = 1; Q = 1 – Fq.
instructive to examine a plot DpH as a function of Q to get a feel for the limits of detection and responsiveness of 9-AA fluorescence to changes in DpH. As seen in Fig. 3, fluorescence quenching of just 5% (Q = 5%, Fq = 95%) already translates to a DpH of 1.8 pH units so that this technique is not responsive to small DpH values. In addition, the slope through the middle of the curve is shallow, such that relatively large changes in quenching result in rather small changes in DpH, as observed for the two traces in Fig. 1. 13. The absorbance change measured in the ECS signal in response to a single turnover flash is typically 0.001 absorbance units. Since thylakoids undergo slow volume changes in response to illumination, which are manifested as scattering changes in the visible region of the spectrum, it is challenging to record this and many other small absorbance changes in thylakoids under steady-state light. Accordingly, the ECS signal has most often been measured in response to a short-duration flash which is so brief so as to prevent the volume-associated light-scattering changes. 14. The magnitude of the electric field across the thylakoid membrane is difficult to quantitate (14), and the ECS signal alone does not contain this information. The height of the flashinduced absorbance change is a function of many parameters in addition to the electric field, including the number of active photosystem II reaction centers in the sample, the number of photons absorbed per reaction center per flash, the ion conductivity of the membrane, and the sampling time of the spectrophotometer. In contrast to determination of the magnitude of the electric field, comparison of the electrical properties of the membranes between samples is much more straightforward, where, for instance, changes in the conductivity of ions in different samples can be readily observed.
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15. Because thylakoid suspensions are optically inhomogeneous, stirring introduces considerable noise in small absorbance signals. Additionally, the volume and cuvette chosen allow us to use a measuring beam covering a relatively large area without the undesirable result of having photons bypass the sample before encountering the detector. 16. The timing sequence that we have used in the traces in Fig. 2 is typical for our measurements made with the NoFOSpec (9): 500 points are collected at 10-ms intervals, with a 10-ms flash delivered from a set of four red light-emitting diodes after 100 points have been collected. The remaining 400 points carry the ECS signal as it decays back to the original absorbance level, with the rate of decay reflecting the ability of the membrane to maintain the electric field. 17. Sufficient time should be allowed for the electric field to decay to zero between measuring sequences; 10 s is usually enough when measuring the ECS in isolated thylakoids. 18. The signal-to-noise ratio describes the resolution of the measurement and varies widely depending on the instrument used. In the NoFOSpec, the signal-to-noise ratio can be sufficiently high after one actinic pulse, whereas in the laboratory’s other flash kinetic spectrophotometer, averaging is required to achieve the desired resolution. The signal-to-noise ratio improves as the square root of the number of signals averaged so that a twofold improvement of the signal over a single trace requires four traces to be averaged, and improving the signal fourfold requires the averaging of 16 traces. 19. The time course of the decay of the ESC signal to a small offset value can often be fitted by a single falling exponential:
A (t ) = Aoffset + (A0 − Aoffset )e − kt
(5)
where A(t), A0 and Aoffset are the absorbance at 520 nm at time t, at t = 0 (immediately after the flash), and at 4 s, respectively, and k is a first-order rate constant. The fitted parameters are A0, Aoffset, and k, which in the traces in Fig. 2 are as follows: Arabidopsis: A0 = 0.0048, Aoffset = 0.0005, and k = 2.20 s−1 (t½ = 0.32 s); pea: A0 = 0.0075, Aoffset = 0.0007, and k = 2.72 s−1 (t½ = 0.25 s).
Acknowledgments This work was supported by US Department of Energy Grant DE-FG02-03ER15405.
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tions on a molecular level. Z. Naturforsch. 23, 244–254. Sacksteder, C. A., Jacoby, M. E., and Kramer, D. M. (2001) A portable, non-focusing optics spectrophotometer (NoFOSpec) for measurements of steady-state absorbance changes in intact plants. Photosynth Res. 70, 231–240. Junge, W. (1976) Flash kinetic spectroscopy in the study of plant pigments. In, Biochemistry of Plant Pigments, Vol. 11 (Goodwin, T., ed.) Academic Press, New York, USA, pp. 233–333. Berry, S., and Rumberg, B. (1999) Proton to electron stoichiometry in electron transport of spinach thylakoids. Biochim. Biophys. Acta 1410, 248–61. Walz, D., Goldstein, L., and Avron, M. (1974) Determination and analysis of the buffer capacity of isolated chloroplasts in the light and in the dark. Eur. J. Biochem. 47, 403–407. Junge, W., Auslander, W., McGeer, A. J., and Runge, T. (1979) The buffering capacity of the internal phase of thylakoids and the magnitude of the pH changes inside under flashing light. Biochim. Biophys. Acta 546, 121–141. Junge, W. (1982) Electrogenic reactions and proton pumping in green plant photosynthesis. In, Current Topics in Membranes and Transport, Vol. 16 (Slayman, C. L., ed.) Academic Press, New York, USA, pp. 431–464.
Chapter 19 Measurement of Chloroplast ATP Synthesis Activity in Arabidopsis Aleel K. Grennan and Donald R. Ort Abstract There are numerous options for monitoring ATP synthesis in chloroplasts using isolated thylakoid membranes, intact chloroplasts, and even whole leaves. Currently, the most commonly used method employs isolated thylakoids coupling the synthesis of ATP to light emission from luciferin in a reaction catalyzed by luciferase. The luciferin–luciferase assay can be highly sensitive and is a direct measure of ATP. Another direct measurement of ATP is the incorporation of 32P into ATP, which, while more technically difficult, has the advantage over the luciferin–luciferase assay of being able to distinguish newly synthesized from total ATP. The phosphorylation of ADP results in a net decrease in pKa (acid disassociation constant) between the reactants and the product ATP, resulting in an increase in the pH of the assay media, which can be used as a convenient, continuous measurement of ATP synthesis. The formation of DmH+ across the thylakoid membrane and its concomitant dissipation as ATP is synthesized can be measured by an electrochromic absorption band shift (ECS) of thylakoid pigments measured at 518 nm (Witt, Biochim. Biophys. Acta 505:355–427, 1979; Petty and Jackson, Biochim. Biophys. Acta: Bioenergetics 547:463–473, 1979). The first-order decay time of the ESC can be used to estimate the rate of ATP synthesis providing a noninvasive, indirect method for measuring ATP synthase activity that can be used with intact leaves. Key words: ATP synthase, Thylakoid, pH change, Proton motive force, Electrochromic shift
1. Introduction The chloroplast ATP synthase complex, located in the thylakoid membrane, catalyzes the synthesis of ATP from ADP and inorganic phosphate. Energetically, this is achieved by coupling the anhydride bond formation with the proton transmembrane electrochemical potential (DmH+) across the thylakoid membrane formed by light-driven proton-coupled photosynthetic electron transport. Thus, unlike its counterparts in the mitochondria or bacteria, the
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chloroplast ATP synthase does not have a constant driving force for ATP synthesis and, therefore, has the potential to catalyze net hydrolysis of ATP when the driving force is absent or too small to drive net synthesis. Net hydrolysis of ATP is prevented by regulation of chloroplast ATP synthase, which has at least three components, the primary one being DmH+ formation across the thylakoid membrane, but also nucleotide binding and release to the catalytic and regulatory sites located on the a- and b-subunits, and thiol modulation of disulfide bridge-forming cysteine residues on the g-subunit (1, 2). The DmH+ has two components, both created by light-driven electron transport coupled to the movement of protons from the stroma across the thylakoid membrane to the lumen, causing the formation of both a DpH as well as an electric potential (DY) across the thylakoid vesicle membrane. In chloroplasts, much of the DY is dissipated by counter ion movement, leaving the DpH, which can be as large as 3 pH units, as the major driving force for ATP synthesis. All of the techniques described in this chapter have been successfully used to study ATP synthase activity in Arabidopsis and can be readily adapted to a wide range of plant species. 1.1. Luciferin– Luciferase Bioluminescence
Firefly luciferin is a bioluminescent pigment found in many species of the Lampyridae (firefly) family. In this two-step reaction that requires oxygen as well as ATP, the oxidation of luciferin catalyzed by luciferase results in light emission with a high quantum yield. 1. Luciferin + ATP → luciferyl adenylate + pyrophosphate (PPi) 2. Luciferyl adenylate + O2 → oxyluciferin + AMP + light The stoichiometric requirement for ATP forms the basis for an excellent method for quantifying ATP (2–6). Luciferins isolated from some other luminescent organisms (i.e., marine invertebrates and bacteria) do not require ATP and thus cannot be used for monitoring ATP synthesis reactions. The luciferin–luciferase assay is a sensitive reaction; less than a femtomole of ATP can be detected using 0.2 mg of luciferase (Sigma). This assay is sensitive enough to measure ATP formed in a thylakoid suspension driven by a single-turnover flash (ref. (6) and references within). A limitation to this method for some applications is that it measures all of the ATP present in the sample, so any ATP present prior to the initiation of the ATP-synthesizing reaction will also be included in the final measurements. The potential for contamination of ADP with ATP also exists. In most cases, background ATP levels are manageable and can be determined in appropriate controls. Measurement of the luminescence requires a luminometer or spectrofluorometer. Many plate readers have luminescence modules available, which provides a convenient platform for high-throughput ATP synthesis measurements. Numerous luciferin–luciferase assay
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kits are commercially available. Most of these are used for monitoring industrial samples for microbial contamination and thus need to be carefully evaluated to ensure that they are appropriate for detecting ATP at levels required. Although convenient, kits are not necessary, as all of the reagents necessary to quantify ATP by this method can be purchased from chemical supply companies. If one chooses to use a kit, it is recommended that one be selected with separate, individual components to allow the user to “customize” concentrations of the reagents. 1.2. 32 P Incorporation
The incorporation of 32P into ATP is a very sensitive measure of ATP synthesis and has the advantage over the luciferin–luciferase method of having the capacity to distinguish between newly synthesized ATP, which will be radiolabeled, from preexisting ATP, which will not. Before the amount of 32P-labeled ATP can be quantified, it must be separated from unreacted 32P and any radioactive polyphosphates that might be present (7–9). For laboratories already set up to work with radioactive compounds, this methodology is straightforward, very sensitive, and, as already mentioned, can distinguish newly synthesized ATP from preexisting pools that might exist in the sample.
1.3. Monitoring pH Change Due to ATP Formation
The phosphorylation of ADP with inorganic phosphate (Pi)
ADP + Pi + nH+ → ATP + H2O results in a net decrease in pKa of the phosphate hydroxyls of the reactants (ADP + Pi) compared to ATP (here n = DH+/DPi or number of hydrogen ions lost/number of ATP ions formed). Thus, ATP synthesis results in a rise in pH of the assay medium. Nishimura et al. (10) long ago worked out the underlying chemistry that allowed the quantification of ATP formation by this method in a physiologically relevant pH range (pH 7–9). The change in hydrogen ion concentrations as well as the buffering capacity of the reaction mixture is determined by titration, allowing the amount of ATP formed to be calculated for a specific pH range (pH 7–9) by assuming the following for a weak acid (HA) pH = pK a + log(γA − /γHA) + log([A − ]/[HA]) = pK a′ + log([A − ] / [HA]) where gA− and gHA are the activity coefficients for the conjugate basic and acidic forms. These small changes in pH can be measured in a weakly buffered solution with a sensitive pH electrode or appropriate pH-sensitive dye.
1.4. Electrochromic Shift
Measuring the relaxation kinetics of the electrochromic shift (ECS) is a spectroscopic technique that can be used to estimate the rate of ATP synthesis in isolated thylakoids, intact chloroplasts, and even
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attached leaves. The ECS in thylakoids is a bathochromic shift (i.e., red shift) that results from an electric field effect (i.e., DY) on the absorption spectrum of a subset of thylakoid pigments, mostly carotenoids. The difference spectrum is largest at 518 nm (11), and the absorption change is most often monitored at this wavelength. Because the rapid phase of the relaxation of the electric field that gives rise to the ECS is due primarily to protons exiting the thylakoid lumen through the ATP synthase, these kinetics can be used to estimate the rate of ATP formation (11, 12). While, based on a set of assumptions, it is possible to estimate absolute rates of ATP formation (13), this method is most frequently used as a comparative technique. For example, ECS methodology has been used to isolate mutants of the g-subunit of ATP synthase in Arabidopsis (14). By examining the amplitude of the ECS, this technique can also be used to examine the effects of environmental stress on proton-coupled electron transport (15, 16). ECS has the advantage of being a noninvasive technique for measuring ATP synthase activity; these measurements can be performed on whole leaves (14, 15, 17), intact plastids, or on isolated thylakoid membranes. For our purposes here, we focus on using attached leaves, as this is essentially the only technique available to monitor ATP synthesis in whole leaves. Access to a kinetic spectrophotometer is necessary for these measurements. While these have been previously custom built (e.g., refs. 16–18), there are now specialty commercial instruments available (JTS-10 from BioLogic; see Subheading 2.4).
2. Materials 2.1. Luciferin– Luciferase Bioluminescence
1. Thylakoid membranes (see Chapter 19, Vol. 1; alternatively, see ref. 18) (see Note 1). 2. Assay buffer I: 100 mM Tricine–NaOH, pH 8.0, 500 mM NaCl, 5 mM MgCl2, 0.05 mM phenazine methosulfate (PMS), 2 mM potassium phosphate, pH 7.0, 1 mM ADP (ATP-free; Sigma, St. Louis, MO, USA), and 0.1 mM diadenosine pentaphosphate. 3. 100% (w/v) trichloroacetic acid (to denature proteins). 4. Assay buffer II: 25 mM Tris–acetate, pH 7.75, 2 mM ethylenediaminetetraacetic acid (EDTA), 50 mM dithiothreitol (DTT), 0.02 mM d-luciferin, 1.5 mg/mL bovine serum albumin (BSA), 20 mM magnesium acetate, and 0.3 mg/mL luciferase (see Notes 2 and 3). 5. ATP standards (10−11 to 10−5 M in tenfold increments) dissolved in 25 mM Tris–acetate, pH 7.75, freshly prepared.
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6. Light source. Actinic light can be provided by a 24-V/250-W tungsten–halogen lamp with a sharp red cutoff filter (Corning 2–64), and the duration of the light pulse can be controlled by an electronic shutter (e.g., Uniblitz, Rochester, NY, USA). 7. Luminometer (e.g., Sirius SmartLine TL luminometer). 8. Microcentrifuge and microcentrifuge tubes (1.5 mL). 2.2. 32 P Incorporation
1. Acid-washed charcoal, for the preparation of AMP-washed charcoal (see Subheading 3.2.1). 2. Charcoal wash buffer I: 0.3 M perchloric acid, 0.05 M PPi, 0.1 M Pi, and 2 mM AMP. 3. Charcoal wash buffer II: 4 volumes of 95% (v/v) ethanol, mixed with 6 volumes of 1 N NH4OH. 4. Thylakoid membranes (see Chapter 19, Vol. 1; alternatively, see ref. 18). 5. Reaction buffer: 50 mM sorbitol, 50 mM Tricine–KOH, pH 8.0, 5 mM MgCI2, 0.1 mM methyl viologen, and 1.0 mM P1, P5-diadenosine-5¢-pentaphosphate. 6. 2 M trichloroacetic acid in 10 mM EDTA (to stop reaction). 7. Unlabeled ATP. 8. Phosphate solution: 0.5 M orthophosphate (NaH2PO4) and 0.125 M pyrophosphate (Na4P2O7). 9. Charcoal wash buffer III: 0.025 M Na4P2O7, 0.10 M H3PO4, and 0.3 M HClO4. 10. Ammonia–ethanol solution: 0.6 M NH4OH in 40% (v/v) ethanol. 11. 0.2 M Tris–HCl, pH 8.0. 12. Light source (see Subheading 2.1, item 6). 13. 0.22-mm membrane filter, for charcoal removal. 14. Quaternary amine resin (Dowex AG1-X4, 200–400 mesh). 15. Small column (2 mL) or syringe barrel (5 cc). 16. Scintillation counter, vials, and fluid (fluid is optional; see Note 4). 17. Buchner funnel and filter paper. 18. Microcentrifuge and microcentrifuge tubes (1.5 mL). 19. pH paper.
2.3. Monitoring pH Change Due to Photophosphorylation
1. Thylakoid membranes (see Chapter 19, Vol. 1; alternatively, see ref. 18). 2. Monitoring buffer: 10 mM KH2PO4, 30 mM MgCl2, 1 mM Na succinate, and 1 mM ADP.
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3. 50 mN HCl for calibration of ATP synthesis rate. 4. pH meter. 5. Calomel glass micro combination pH electrode. 6. Cylindrical cuvette (22 mm outer diameter, 15 mm depth). 7. Light source (see Subheading 2.1, item 6). 2.4. Electrochromic Shift
1. Attached leaf or detached leaf with water source (see Note 5). 2. Kinetic spectrophotometer; built in-lab or purchased from BioLogic (JTS-10; BioLogic SAS, Claix, France; or, BioLogic USA, LCC, Knoxville, TN, USA). 3. Data analysis software (e.g., OriginLab Corp., Northampton, MA, USA).
3. Methods 3.1. Luciferin– Luciferase Bioluminescence
3.1.1. Luciferin–Luciferase and ATP Measurement (2)
The amount of ATP produced is determined by the amount of light emitted from the luciferin–luciferase assay, calibrated by standards using known amounts of ATP. Although this assay has an advantage in that it does not use radionuclides, it can indicate higher formation of ATP when compared to other methodologies (i.e., 32Pi incorporation). The luciferin– luciferase assay is a measurement of all the ATP present in the sample, so any ATP present prior to the reaction will also be included in the final measurements. Commercially available ADP typically has some fraction of ATP present leading to nonzero background readings. For these two reasons, it is very important to obtain an accurate background measurement before beginning the assay. Luciferase activity is suppressed by Cl−, so acetates are used instead of chlorides in the buffers for this assay. 1. Resuspend thylakoid membranes equivalent to 10 mM of chlorophyll (Chl) in 1 mL of assay buffer I. Allow the reaction to proceed at room temperature (22°C) at desired light intensity. 2. Following illumination, stop the reaction by adding trichloroacetic acid to the reaction mixture to a final concentration of 0.5% (w/v). Centrifuge the mixture at 30,000 × g for 5 min at room temperature in a microcentrifuge to remove denatured proteins. Keep the supernatant. 3. Often, it is desirable to dilute the sample (up to 20-fold) before assaying for ATP concentration with a luminometer or spectrofluorometer. Mix 100 mL of the supernatant solution in a recommended tube for the detection device to be used, together
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Fig. 1. Measurement of ATP synthesis by bioluminescence as a function of the number of light flashes. Thylakoid samples were denatured 1 s after the actinic light flash and bioluminescence catalyzed by luciferase was determined. Figure adapted from Graan and Ort (3).
with 100 mL of rapidly injected assay buffer II. Measure the luminescence by integrating the signal for 5 s after injection. A typical curve of nmol ATP produced per mg Chl as a function of the number of light flashes can be seen in Fig. 1. 4. The same reaction mix can be used to determine concentrations of freshly prepared ATP standards (10−11 to 10−5 M in tenfold increments) (see Note 6). 3.2. 32 P Incorporation
ATP synthesis can be determined by measuring the incorporation of radiolabeled Pi into ATP (7). The newly synthesized radiolabeled ATP is separated from the unincorporated nucleotide on an anion-exchange column (Dowex AG1-X4) after initial absorption and deabsorption of the nucleotides on activated charcoal (3, 8).
3.2.1. Preparation of AMP-Washed Charcoal
Treating the charcoal with AMP aids in the recovery of the nucleotides (8). 1. Suspend 10 g of acid-washed charcoal in charcoal wash buffer I and mix for 15 min. 2. Filter the suspension through filter paper in a Buchner funnel and rinse with water. 3. Resuspend the charcoal in 200 mL of charcoal wash buffer II and mix for 15 min. Filter the suspension through filter paper and resuspend the washed charcoal in water at 100 mg charcoal/mL.
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3.2.2. ATP Synthesis Reaction
Intense actinic light can be provided using a tungsten–halogen lamp or other suitable source. A heat filter and temperature control should be used to maintain constant conditions in the sample. This method is sensitive enough to measure ATP generated from short illumination periods or even flashes of light. Longer illumination periods can be used and the amount of radioactivity can be lowered accordingly. 1. One minute prior to the illumination, suspend thylakoid preparations containing 10 mmol Chl in 2 mL of reaction buffer at 25°C, to allow the thylakoids to temperature equilibrate. Fifteen seconds before the flash, add 0.5 mM 32Pi (15 mCi) and 0.1 mM ADP to the suspension (see Note 7). 2. Illuminate the reaction for 1 s with actinic light. 3. Terminate the reaction 1 s after the end of illumination by the addition of 1 mL of a 2 M trichloroacetic acid in 10 mM EDTA.
3.2.3. Determining Radioactive Orthophosphate Incorporation
The incorporation of radioactive orthophosphate into ATP is measured from the acid-quenched samples after purification. 1. Centrifuge the samples at 23,000 × g for 10 min in a microcentrifuge to remove insoluble proteins. Carefully remove the supernatant and neutralize with 0.2 M Tris–HCl to pH 5.4. 2. Combine the supernatant with 1.5 mmol of unlabeled ATP (to calculate % recovery), 0.3 mL of AMP-washed charcoal, and 0.4 mL of phosphate solution, then mix well and allow to stand for 5 min (to remove unincorporated radiolabeled orthophosphate). Collect the charcoal with the adsorbed nucleotide on filter paper (in a Buchner funnel). Wash the charcoal once with charcoal wash buffer III followed by three washes with distilled water. Elute the absorbed nucleotides by placing the charcoal and filter paper in a 125-mL flask with 14 mL of ammonia–ethanol solution. Shake for 20 min. Filter the solution through a 0.22-mm filter to remove the charcoal. Wash the collected charcoal again in the same flask with the ammonia–ethanol solution and filter. 3. Prepare a 2 mL column; alternatively, a 5 cc syringe barrel can be used. Pack the column with a quaternary amine resin (e.g., Dowex AG1-X4, 200–400 mesh). Wash the resin with 1.0 M HCl and follow with distilled water until the eluate reaches neutral pH, by testing with pH paper. Apply the filtrate to the Dowex column. Once the filtrate has passed through under gravity, rinse the column with 2 mL of distilled water followed by 5 mL of 0.2 M Tris–HCl, pH 8.0. Elute the nucleotide fractions (typically, 2.5 mL each) with a stepwise gradient of increasing
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Fig. 2. Incorporation of 32P into ATP as a function of the number of light flashes. The ADP and 32P were added 15 s prior to the actinic light flash. The thylakoid samples were denatured 1 s after the flash and assayed for 32P incorporation into ATP. Figure adapted from Graan and Ort (3).
concentrations of HCl as follows: 30 mM HCl (AMP), 60 mM HCl (ADP), and 1 M HCl (ATP). Remove a 1.0-mL aliquot from each to determine the recovery of nucleotides by measuring the absorbance peak at 257.5 nm (A260 or A257.5). 4. To determine the incorporation of labeled orthophosphate, remove another 1 mL of sample, combine with scintillation fluid and measure counts in a scintillation counter. An example of typical results can be seen in Fig. 2. 3.3. Determination of pH Change Due to ADP Phosphorylation (10)
1. Resuspend thylakoid membranes in monitoring buffer to ~10 mM Chl and adjust pH to 7.4 using dilute HCl. 2. Add the suspension to the measuring cuvette and insert pH electrode and start monitoring (see Note 8). 3. Illuminate the mixture at the required intensity. The intensity used will be dependent upon the experiment as well as the growth light of the plants. A good starting intensity is 500 mmol/m2/s. 4. Record the rate of pH change due to illumination. 5. Determine the buffering capacity of the solution by adding a known amount of 50 mN HCl to the thylakoid suspension after each measurement (5 mL of 50 mN HCl is 250 ng ions H+). Record the pH change. 6. The amount of P incorporated into ATP is calculated from the n-value (=DH+/DPi) and the buffering capacity determined in
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Fig. 3. Change in pH due to ADP phosphorylation as a function of the number of light flashes. Isolated thylakoid preparations were exposed to different numbers of actinic light flashes each lasting 6 ms while the pH was monitored. Figure adapted from Graan and Ort (18).
step 5 (=added H+/DpH; this value was calculated to be 0.891 in Rhodospirillum rubrum chromatophores (10)). Figure 3 shows typical results for pH change due to ADP phosphorylation plotted against the number of light flashes the sample received. 3.4. Electrochromic Shift
The details of how these measurements are to be made will be idiosyncratic with the design and features of the kinetic spectrophotometer used. As such, the methodology given below is intended to outline general steps involved and the factors to be considered (see Note 9). In general, plants are dark-adapted for 12 h to ensure that the g-subunit regulatory dithiols of ATP synthase within the leaves are fully oxidized (16). The procedure outlined below is for the laboratory-built kinetic spectrophotometer described in Wise and Ort (16). All measurements can be conducted at room temperature. 1. Position an intact leaf in the leaf cuvette clamp in which a dim (~6 × 10−2 J/m2/s) measuring beam of light is delivered to a spot on the adaxial leaf surface via a fiber-optic light guide, collected from the abaxial surface by one arm of a bifurcated light guide, and routed to a photomultiplier tube or sensitive photodiode. 2. A bright actinic light pulse is delivered to the abaxial surface through the second arm of the bifurcated light guide. Comple mentary blocking filters are needed to prevent scattered light from the actinic light from entering the photomultiplier.
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Fig. 4. DA518 relaxation kinetics in wild-type Arabidopsis leaves. DA518 relaxation kinetics was measured in 20 leaves from 45-day-old plants using a flash kinetic spectrophotometer (14). Plants were dark-adapted for 12 h and then illuminated with 65 mmol/m2/s red light for 7 s.
3. The relaxation kinetics of the flash-induced DA518–540 absorption transient is biphasic. Since only the initial, faster component of the thylakoid membrane depolarization is associated with ATP synthesis, the slow component is digitally subtracted. The remaining fast decay can be expressed as the relaxation time constant, t, according to the equation
∆A518−540 = ∆A518−540 max e −t / t where both t (time) and t are in ms. These relaxation time constants can be calculated by fitting the initial 600 ms of the decay to a first-order exponential using an iterative nonlinear least-squares program (Fig. 4).
4. Notes 1. Not all thylakoid preparations are capable of performing photophosphorylation. If the results are not as expected, incremental improvements to the preparation are recommended. 2. Depending on the sensitivity requirements, luciferase and luciferin concentrations can be altered significantly. The higher the luciferase concentration, the higher is the sensitivity. 3. The pH optimum of the luciferase reaction is about 7.7. However, it is possible to measure from pH 6.5–8.5 (although it might be necessary to increase the luciferase concentration).
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4. Owing to the relatively high energy of the beta-particle emission, [32P]ATP can be conveniently measured based on Cherenkov radiation in a scintillation counter avoiding the expense and complications of scintillation cocktails. 5. If using detached leaves, recut the petioles while they are under water to ensure that the xylem stream is intact and ensure a water supply for the duration of the experiment. Plants that produce latex should not be used for detached leaf experiments, as latex will block xylem flow. 6. Luciferin and luciferase can vary significantly with each preparation and lot. A standard should be prepared and run each time. 7. 33P can be substituted for 32P. It has the advantage of having a longer half-life and has a weaker beta-particle emission making it somewhat easier to use. 8. For the most sensitive determinations, stray light and electrostatic disturbances can be a problem. To overcome this, the cuvette assembly can be shielded in a metal box. 9. The physiological state of the plants will influence the measurements; this must be taken into account when comparing between plants. References 1. Ort, D. R., and Oxborough, K. (1992) In situ regulation of chloroplast coupling factor activity. Annu. Rev. Plant Physiol. Plant Mol. Biol. 43, 269–291. 2. He, F., Samra, H. S., Johnson, E. A., Degner, N. R., McCarty, R. E., and Richter, M. L. (2008) C-Terminal mutations in the chloroplast ATP synthase gamma subunit impair ATP synthesis and stimulate ATP hydrolysis. Biochemistry 47, 836–844. 3. Graan, T., and Ort, D. R. (1981) Factors affecting the development of the capacity for ATP formation in isolated-chloroplasts. Biochim. Biophys. Acta 637, 447–456. 4. Steigmiller, S., Turina, P., and Gräber, P. (2008) The thermodynamic H+/ATP ratios of the H+-ATP synthases from chloroplasts and Escherichia coli. Proc. Nat. Acad. Sci. USA 105, 3745–3750. 5. Turina, P., Samoray, D., and Graber, P. (2003) H+/ATP ratio of proton transportcoupled ATP synthesis and hydrolysis catalysed by CF0F1-liposomes. EMBO J. 22, 418–426. 6. Beard, W. A., and Dilley, R. A. (1988) ATP formation onset lag and post-illumination phosphorylation initiated with single-turnover flashes. 1. An assay using luciferin-luciferase
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luminescence. J. Bioenerg. Biomembr. 20, 85–106. Hangarter, R. P., and Good, N. E. (1982) Energy thresholds for ATP synthesis in chloroplasts. Biochim. Biophys. Acta: Bioenergetics 681, 397–404. Smith, D. J., Stokes, B. O., and Boyer, P. D. (1976) Probes of Initial phosphorylation events in ATP synthesis by chloroplasts. J. Biol. Chem. 251, 4165–4171. Flores, S., and Ort, D. R. (1984) Investigation of the apparent inefficiency of the coupling between photosystem II electron transfer and ATP formation. Biochim. Biophys. Acta: Bioenergetics 766, 289–302. Nishimura, M., Takeru, I., and Chance, B. (1962) Studies on bacterial photophosphorylation III. A sensitive and rapid method of determination of photophosphorylation. Biochim. Biophys. Acta 59, 177–182. Witt, H. (1979) Energy-conversion in the functional membrane of photosynthesis: analysis by light-pulse and electric pulse methods: central role of the electric-field. Biochim. Biophys. Acta 505, 355–427. Petty, K. M., and Jackson, J. B. (1979) Correlation between ATP Synthesis and the decay of the carotenoid band shift after single
19 Measurement of Chloroplast ATP Synthesis Activity in Arabidopsis flash activation of chromatophores from Rhodopseudomonas capsulata. Biochim. Biophys. Acta: Bioenergetics 547, 463–473. 13. Junge, W., and Jackson, J.B. (1982) The development of electrochemical potential gradients across photosynthetic membranes. In, Photosynthesis in Green Plants and Bacteria, Vol. 1 (Govindjee, ed.) Academic Press, New York, USA, pp. 589–646. 14. Wu, G., Ortiz-Flores, G., Ortiz-Lopez, A., and Ort, D. R. (2007) A point mutation in atpC1 raises the redox potential of the Arabidopsis chloroplast ATP synthase g-subunit regulatory disulfide above the range of thioredoxin modulation. J. Biol. Chem. 282, 36782–36789. 15. Ortiz-Lopez, A., Ort, D. R., and Boyer, J. S. (1991) Photophosphorylation in attached
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leaves of Helianthus annuus at low water potentials. Plant Physiol. 96, 1018–1025. 16. Wise, R. R., and Ort, D. R. (1989) Photophosphorylation after chilling in the light: Effects on membrane energization and coupling factor activity. Plant Physiol. 90, 657–664. 17. Sacksteder, C. A., Jacoby, M. E., and Kramer, D. M. (2001) A portable, non-focusing optics spectrophotometer (NoFOSpec) for measurements of steady-state absorbance changes in intact plants. Photosynth. Res. 70, 231–240. 18. Graan, T., and Ort, D. R. (1983) Initial events in the regulation of electron transfer in chloroplasts. The role of the membrane potential. J. Biol. Chem. 258, 2831–2836.
Chapter 20 Methods for Analysis of Photosynthetic Pigments and Steady-State Levels of Intermediates of Tetrapyrrole Biosynthesis Olaf Czarnecki, Enrico Peter, and Bernhard Grimm Abstract Tetrapyrroles and carotenoids are required for many indispensable functions in photosynthesis. Tetrapyrroles are essential metabolites for photosynthesis, redox reaction, and detoxification of reactive oxygen species and xenobiotics, while carotenoids function as accessory pigments, in photoprotection and in attraction to animals. Their branched metabolic pathways of synthesis and degradation are tightly controlled to provide adequate amounts of each metabolite (carotenoids/tetrapyrroles) and to prevent accumulation of photoreactive intermediates (tetrapyrroles). Many Arabidopsis mutants and transgenic plants have been reported to show variations in steady-state levels of tetrapyrrole intermediates and contents of different carotenoid species. It is a challenging task to determine the minute amounts of these metabolites to assess the metabolic flow and the activities of both pigment-synthesising and degrading pathways, to unravel limiting enzymatic steps of these biosynthetic pathways, and to characterise mutants with accumulating intermediates. In this chapter, we present a series of methods to qualify and quantify anabolic and catabolic intermediates of Arabidopsis tetrapyrrole metabolism, and describe a common method for quantification of different plant carotenoid species. Additionally, we introduce two methods for quantification of non-covalently bound haem. The approach of analysing steady-state levels of tetrapyrrole intermediates in plants, when applied in combination with analyses of transcripts, proteins, and enzyme activities, enables the biochemical and genetic elucidation of the tetrapyrrole pathway in wild-type plants, varieties, and mutants. Steadystate levels of tetrapyrrole intermediates are only up to 1/1,000 of the amounts of the accumulating end-products, chlorophyll, and haem. Although present in very low amounts, the accumulation and availability of tetrapyrrole intermediates have major consequences on the physiology and activity of chloroplasts due to their additional photoreactive and possible signalling functions. Although adjusted for Arabidopsis tetrapyrrole metabolites, the presented methods can also be applied for analysis of cyanobacterial and other plant tetrapyrroles. Key words: 5-Aminolevulinic acid, Carotenoids, Chlorophyll, Coproporphyrin III, Haem, Mg protoporphyrin IX, Mg protoporphyrin IX monomethylester, Porphobilinogen, Porphyrins, Protoporphyrin IX, Tetrapyrroles, Uroporphyrin III
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1. Introduction In plants, the end-products of the branched pathways of tetrapyrrole and carotenoid biosynthesis serve various important biological functions. As components of light-harvesting complexes, chlorophyll (Chl) and carotenoids absorb light, dissipate excessive excitation energy (carotenoids) or transfer excitation energy ultimately to the reaction centres of the two photosystems (carotenoids, Chl), and perform charge separation (Chl), which eventually leads to the generation of ATP and reducing power. Haem-containing proteins are involved in respiration, detoxification of reactive oxygen species (ROS), and many other redox reactions. Bilins are products of haem breakdown reactions and act as chromophores of phytochromes, which light-dependently control the expression of various genes. Last but not least, sirohemes serve as cofactors of nitrite or sulphite reductases. This chapter describes different extraction procedures and methods for the determination of photosynthetic pigments as well as different intermediates and end-products of the branched tetrapyrrole biosynthetic pathway; the latter range from the common precursor of all tetrapyrroles (5-aminolevulinic acid, ALA) to the end-products, Chl and haem, and also Chl degradation intermediates. The presented methods are optimised for the analysis of Arabidopsis thaliana, but they can also be applied in studies of cyanobacteria or other plant species with minimal modification. Although Arabidopsis is a small and unpretentious weed, scientists have taken advantage of its short life cycle; its small, fully sequenced genome; and the availability of well-established transformation techniques. For these reasons and others, Arabidopsis is now a widely used model plant for the analysis of genetic and physiological processes, and as such is an excellent system for studying photosynthetic pigments and other tetrapyrroles. 1.1. General Remarks
Quantifying pigments and tetrapyrroles in plant tissues or isolated chloroplasts is always connected with the question “which reference value should be chosen?” When plant tissues are analysed, common references are dry or fresh weight. Also, the Chl content is often used. When isolated chloroplasts are used, Chl and protein contents are appropriate references. The volume of isolated chloroplasts is also a parameter that can easily be determined (using special tubes for biomass determination, e.g., Sartorius-Stedim VoluPAC tubes), and this can serve as a reference value. However, the reference value can strongly be modified upon deregulation of Chl biosynthesis, and leaf chlorosis and necrosis can occur. Additionally, reduced Chl synthesis and content are often accompanied by reduced photosynthesis and reduced protein levels. This can easily be monitored by reduced levels of
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Fig. 1. Serial extraction of different tetrapyrroles from one sample.
the two most abundant proteins in chloroplasts: light-harvesting chlorophyll-binding proteins (LHCPs) and the ribulose-1, 5-bisphosphate carboxylase (RuBisCO). Thus, a careful decision is recommended before steady-state levels of tetrapyrrole intermediates are determined. It is often worth performing a serial extraction of different tetrapyrrole intermediates and end-products from a single sample. Porphyrins and Mg porphyrins (including Chl) are extracted in a basic solvent, and haem can be extracted subsequently using acidic acetone. Figure 1 presents a possible strategy to analyse different tetrapyrroles from a single sample based on protocols given in Subheading 3.
2. Materials Plant tissue needs to be thoroughly homogenised for pigment extraction. We recommend using a precooled pestle and mortar, because some remaining activity of tetrapyrrole processing enzymes may change individual steady-state levels during extraction. Addition of quartz sand for better cell disruption is optional. However, when many samples have to be analysed, the usage of tissue homogenisers, grinders, or rotating cell-mills is recommended. For almost all protocols presented under Subheading 3, a spectrophotometer (e.g., Ultrospec 3300 Pro, GE Healthcare) and a refrigerated tabletop microcentrifuge (e.g., Biofuge Fresco, Heraeus)
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are mandatory. The quantification of the extracted tetrapyrroles can be performed by measuring absorption and fluorescence emission spectra using a spectrophotometer or a fluorescence spectrometer, or after separation by HPLC using a photodiode array detector or a fluorescence detector, depending on the complexity of the pigment and precursor mixture, their spectral properties, and steady-state levels of particular intermediates. Due to the photoreactivity of all pigments, it is generally recommended to extract pigments under dim or green light. 2.1. Determination of the ALA-Synthesis Rate
1. Incubation buffer: 50 mM Tris–HCl, pH 7.2, and 40 mM levulinic acid (see Note 1). For 200 mL buffer, dissolve 1.21 g Tris base in 100 mL of H2O. Add 0.93 g of levulinic acid (4-oxopentanoic acid) or a corresponding amount of stock solution (see Note 1). Adjust the pH to 7.2 with HCl and fill up with H2O to 200 mL. 2. Extraction buffer: 20 mM potassium phosphate, pH 6.8. Dissolve 0.27 g KH2PO4 in 100 mL of H2O. Dissolve 0.35 g K2HPO4 in another 100 mL of H2O. Adjust the pH of the K2HPO4 solution to 6.8 by stepwise addition of the KH2PO4 solution. 3. Modified Ehrlich’s reagent: 75% (v/v) glacial acetic acid, 12.5% (v/v) perchloric acid, 11.5 mM HgCl2, and 122 mM p-dimethyl aminobenzaldehyde. For 500 mL, mix 373 mL of glacial acetic acid, 90 mL of 70% (v/v) perchloric acid, and 1.55 g of HgCl2, and adjust cautiously to a final volume of 500 mL with H2O. HgCl2 may require some time to dissolve completely. The solution can be stored at 4°C in darkness for up to 2 months. p-Dimethyl aminobenzaldehyde should be added fresh. Add 2.0 g per 110 mL (9.10 g per 500 mL) prior to use and mix well. Handle the solution with care. HgCl2 is toxic and spilled solution stains almost every surface with nasty blotches. 4. Ethyl acetoacetate. 5. 1 mM ALA stock solution: Prepare at first a 1 M stock by dissolving 1.68 g 5-aminolevulinic acid hydrochloride in exactly 5 mL extraction buffer. Fill exactly to 10 mL with extraction buffer. Dilute to 1,000 by mixing 10 mL of the 1 M stock with 9.99 mL of extraction buffer. Both stock solutions (1 M and 1 mM) can be stored at −20°C.
2.2. Determination of Porphobilinogen Steady-State Levels
1. Extraction buffer: 200 mM sodium phosphate, pH 8.0, and 600 mM ethylenediaminetetraacetic acid (EDTA). Dissolve 3.12 g NaH2PO4 ⋅ 2H2O in 100 mL of H2O. Dissolve 2.84 g Na2HPO4 in another 100 mL of H2O. Adjust the pH of the Na2HPO4 solution to 8.0 by stepwise addition of
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the NaH2PO4 solution. Add 120 mL of a 0.5 M disodium EDTA solution (pH 8.0) per 100 mL of the buffer. 2. Modified Ehrlich’s reagent (see Subheading 2.1). 2.3. Extraction of (Mg) Porphyrins According to Mock and Grimm (1)
1. Extraction solution 1: 50 mM potassium phosphate, pH 7.8. Dissolve 0.68 g KH2PO4 in 100 mL of H2O. Dissolve 0.87 g K2HPO4 in another 100 mL of H2O. Adjust the pH of the K2HPO4 solution to 7.8 by stepwise addition of the KH2PO4 solution. 2. Extraction solution 2: 90% (v/v) methanol and 10% (v/v) 0.1 M NH4OH. To produce a 0.1 M NH4OH solution, add 14 mL of a 28% NH3 solution to 986 mL of H2O. For 100 mL of extraction solution 2, mix 90 mL of methanol with 10 mL of 0.1 M NH4OH. 3. Extraction solution 3: 90% (v/v) acetone and 10% (v/v) 0.1 M NH4OH. For 100 mL of extraction solution 3, mix 90 mL of acetone with 10 mL of 0.1 M NH4OH. 4. 1 M acetic acid. For 100 mL, dissolve 6 mL of glacial acetic acid in 80 mL of H2O and fill up with H2O to 100 mL. 5. 2-Butanone peroxide.
2.4. Extraction of (Mg) Porphyrins According to Moulin and Smith (2)
1. Extraction solution 1: 90% (v/v) methanol and 10% (v/v) 0.1 M NH4OH. To produce a 0.1 M NH4OH solution, add 14 mL of a 28% NH3 solution to 986 mL of H2O. For 100 mL of extraction solution 1, mix 90 mL of methanol with 10 mL of 0.1 M NH4OH. 2. Extraction solution 2: 80% (v/v) acetone and 20% (v/v) 0.1 M NH4OH. For 100 mL of extraction solution 2, mix 80 mL of acetone with 20 mL of 0.1 M NH4OH. 3. N2 or air to dry the pigment solution. A SpeedVac is also recommended (see Note 2).
2.5. Preparation of (Mg) Porphyrin HPLC Standards
1. Dimethyl sulfoxide (DMSO). 2. HCl at different concentrations: 0.1, 0.5, and 2.7 M. Conc. HCl (37%) is around 12 M. Dilute 835 mL, 4.2 mL, or 22.5 mL with H2O in a final volume of 100 mL. 3. Ethanol. 4. Diethyl ether.
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5. Uroporphyrinogen III (Uro III), coproporphyrinogen III (Copro III), protoporphyrin IX (Proto IX), Mg protoporphyrin IX (Mg Proto IX), and Mg protoporphyrin IX monomethylester (Mg Proto IX MME). These compounds can be purchased from Frontier Scientific, Inc., USA. There are also more porphyrin standards available. The preparation of standard solutions is described in detail in Subheading 3.5. 2.6. HPLC Analysis of (Mg) Porphyrins According to Mock and Grimm (1)
1. HPLC device with fluorescence detector and RP 18 column (Nova-Pak C18, 3.9 × 150 mm, 4-mm particle size). 2. Solvent A: 10% (v/v) methanol and 10% (v/v) 1 M ammonium acetate, pH 5.2. Dissolve 77.08 g ammonium acetate in 1,000 mL of H2O to make a 1 M stock; adjust the pH to 5.2 using acetic acid. Mix 100 mL of methanol, 100 mL of 1 M ammonium acetate, and 800 mL of H2O. Filter the solution through a 0.22-mm mesh filter. 3. Solvent B: 90% (v/v) methanol and 10% (v/v) 1 M ammonium acetate, pH 5.2. Mix 900 mL of methanol and 100 mL of 1 M ammonium acetate (see item 2 above). Filter the solution through a 0.22-mm mesh filter.
2.7. HPLC Analysis of (Mg) Porphyrins According to Moulin and Smith (2)
1. HPLC device with fluorescence or photodiode array (PDA) detector and RP 18 column (Nova-Pak C18, 2.1 × 150 mm, 4-mm particle size). 2. Solvent A: 20 mM ammonium acetate, pH 5.16. Dissolve 1.54 g ammonium acetate in 1,000 mL of H2O. Adjust the pH to 5.16 using acetic acid. Filter the solution through a 0.22-mm mesh filter. 3. Solvent B: 100% methanol. 4. Solvent C: 100% acetonitrile. 5. Solvent D: 100% acetone.
2.8. Extraction of Protochlorophyllide and Chlorophyllide
1. Boiling water bath (e.g., kettle). 2. Mesh to expose the leaf sample to steam. 3. 15-mL centrifuge tubes, closable (e.g., Falcon™ type). A centrifuge and rotor to accommodate these tubes are also required. 4. Extraction solution: 90% (v/v) acetone and 10% (v/v) 0.1 M NH4OH (see Subheading 2.4, item 1). 5. n-Hexane. 6. Fluorescence spectrometer (e.g., FP-6500, Jasco Inc.). 7. HPLC device with fluorescence detector and RP 18 column (Waters RP-18 ODS Hypersil, 4.0 × 125 mm, 3-mm particle size).
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This item is optional and only required when HPLC is used for quantification (see Subheading 3.8.2). 8. Solvent A: 80% (v/v) methanol and 20% (v/v) 1 M ammonium acetate, pH 7.0. Dissolve 77.08 g ammonium acetate in 1,000 mL of H2O. Adjust the pH to 7.0 using acetic acid. Mix 800 mL of methanol and 200 mL of 1 M ammonium acetate. Filter the solution through a 0.22-mm mesh filter. This item is optional and only required when HPLC is used for quantification (see Subheading 3.8.2). 9. Solvent B: 80% (v/v) methanol and 20% (v/v) acetone. Mix 800 mL of methanol and 200 mL of acetone. Filter the solution through a 0.22-mm mesh filter. This item is optional and only required when HPLC is used for quantification (see Subheading 3.8.2). 2.9. Preparation of Protochlorophyllide and Chlorophyllide Standards
1. The same equipment and solutions needed for protochlorophyllide (Pchlide) extraction (see Subheading 2.8). 2a. For Pchlide standard extraction, etiolated barley seedlings are needed. Germinate barley (Hordeum vulgare L.) seedlings on wet vermiculite for 5 days at around 23°C in complete darkness (N.B. In contrast to small etiolated Arabidopsis seedlings, Pchlide can easily be extracted from etiolated barley leaves). 2b. For Chlide standard extraction, the etiolated barley seedlings should be illuminated for 30 min at 100 mmol photons/m2/s. 3. Diethyl ether. 4. Saturated NaCl solution. Heat 50 mL of water to approximately 50°C and add stepwise around 25 g NaCl up to the solubility limit under continuous stirring. Let the solution cool down. NaCl crystals will precipitate; the supernatant is a saturated NaCl solution. 5. Saturated KH2PO4 solution. Heat 50 mL of water to approximately 50°C and add stepwise around 15 g KH2PO4 up to the solubility limit under continuous stirring. Let the solution cool down. Crystals will precipitate; the supernatant is a saturated KH2PO4 solution.
2.10. Spectrophoto metrical Determination of Chlorophylls and Carotenoids
1. Extraction solution: 80% (v/v) acetone and 10 mM KOH; or 90% (v/v) acetone and 10% (v/v) 0.1 M NH4OH (see Subheading 2.3, item 3); or 90% (v/v) methanol and 10% (v/v) 0.1 M NH4OH (see Subheading 2.3, item 2); or 90% (v/v) acetone and 10% (v/v) 0.2 M Tris–HCl, pH 8.0 (see Subheading 2.16, item 1).
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2.11. Quantification of Carotenoids and Chlorophylls by HPLC
1. Extraction solution (see Subheading 2.10). 2. HPLC device with PDA detector and a Sperisorb ODS2 column (Waters 4.6 × 250 mm, 5-mm particle size). 3. Solvent A: 90% (v/v) acetonitrile, 9.9% (v/v) H2O, and 0.1% (v/v) triethylamine. For 1,000 mL, mix 900 mL of acetonitrile with 99 mL of H2O and 1 mL of triethylamine. 4. Solvent B: 100% ethylacetate. 5. Ethanol. 6. Acetone. 7. N2 or air to dry the pigment solution. A SpeedVac is also recommended (see Note 2).
2.12. Extraction of Haem from Green Tissues
1. 15- and 50-mL centrifuge tubes, resistant against diethyl ether; closable (e.g., Falcon™ type). A centrifuge and rotor to accommodate these tubes are also required. 2. Extraction solution 1 (ice cold): 90% (v/v) acetone and 10% (v/v) 0.1 M NH4OH (see Subheading 2.4). 3. Extraction solution 2: 80% (v/v) acetone, 16% (v/v) DMSO, and 4% (v/v) conc. HCl. Mix 200 mL of acetone, 40 mL of DMSO, and 10 mL of conc. HCl. 4. Diethyl ether. 5. Saturated NaCl solution (see Subheading 2.9). 6. Ethanol. 7. Pasteur pipette.
2.13. Quantification of Haem Using Ion Exchange Chromatography
1. DEAE Sepharose CL-6B (GE Healthcare). 2. 1 M sodium acetate, pH 7.0. Dissolve 82.03 g sodium acetate (anhydrous) in 900 mL of H2O. Adjust the pH to 7.0 using acetic acid. Fill up with water to 1,000 mL. 3. Acetone. 4. Wash solution 1: 75% (v/v) diethyl ether and 25% (v/v) ethanol. For 200 mL, mix 150 mL of diethyl ether with 50 mL of ethanol. 5. Wash solution 2: 50% (v/v) diethyl ether and 50% (v/v) ethanol. For 200 mL, mix 100 mL of diethyl ether with 100 mL of ethanol.
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6. Elution solution: 81% (v/v) ethanol, 10% (v/v) H2O, and 9% (v/v) glacial acetic acid. Mix 90 mL of ethanol, 11 mL of H2O, and 10 mL of glacial acetic acid. 7. 5-mL polypropylene columns (Qiagen). 2.14. Quantification of Haem in HighThroughput Assays
1. 2× Reaction buffer: 200 mM Tris–HCl, pH 8.4. For 100 mL, dissolve 2.42 g Tris base in 80 mL of H2O. Adjust the pH to 8.4 using HCl. Fill up with water to 100 mL. 2. 1 mM hemin stock solution. Prepare a 100 mM stock by dissolving 652 mg hemin (Sigma) in exactly 10 mL DMSO. Mix 100 mL of the 100 mM hemin stock with 9.9 mL DMSO. 3. 10 mM KOH. Dissolve 561 mg KOH in 1,000 mL of H2O. 4. 2.5 mM horseradish peroxidase apoenzyme (HRP) (Sigma) in H2O (1,000-fold concentrated). The molecular weight of the apoenzyme is approximately 40 kDa. Dissolve 100 mg in 1 mL of water, or prepare homemade HRP apoenzyme (see Subheading 3.15). 5. Very sensitive commercial Western blot detection reagent (e.g., ECL, GE Healthcare). 6. For detection of the luminescence signal, use a CCD camerabased gel-doc apparatus including software to quantify spots densitometrically (e.g., Stella 3200 with AIDA Image Analyzer, Raytest), or a 96-well plate reader with luminescence mode (e.g., Spectra Max M2, Molecular Devices). 7. White 96-well plates. Transparent or black plates are not recommended to detect luminescence signals. 8. 8-Channel pipette, 20–200 mL. This item is optional.
2.15. Homemade Horseradish Peroxidase Apoenzyme
1. HRP holoenzyme (Sigma). 2. 0.05 M HCl. Conc. HCl (37%) is around 12 M. Therefore, dilute 420 mL of this in 100 mL of H2O. 3. 2-Butanone. 4. 600 mM NaHCO3. Dissolve 50 mg NaHCO3 in 1,000 mL of H2O. 5. Devices to concentrate protein solutions or to perform buffer exchange (e.g., concentrator tubes or dialysis tubes).
2.16. Quantification of Chlorophyll Catabolites
1. Extraction solution: 90% (v/v) acetone and 10% (v/v) 0.2 M Tris–HCl, pH 8.0. Dissolve 2.42 g Tris base in 80 mL of H2O. Adjust the pH to 8.0 using HCl and fill with H2O to 100 mL. Mix 90 mL of acetone with 10 mL of 0.2 M Tris–HCl.
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2. 100 nM pheophorbide a standard. Dissolve pheophorbide a (Wako Chemicals) in extraction solution. Quantify the concentration spectrophotometrically using the molar absorption coefficient: e Pheophorbide in 80% acetone, 663 nm = 4.4 × 104 L/mol/cm (3). Dilute the standard solution to 100 nM. 3. 100 nM pheophytine a standard. Dissolve pheophytine a (Wako Chemicals) in extraction solution. Quantify the concentration spectrophotometrically using the molar absorption coefficient: e Pheophytine in 80% acetone, 663 nm = 4.4 × 104 L/mol/cm (3). Dilute the standard solution to 100 nM.
3. Methods 3.1. Determination of the ALA Synthesis Rate
The first intermediates of Chl and haem biosynthesis are water soluble and can be detected by a colorimetric assay based on the reaction of p-dimethyl aminobenzaldehyde with pyrroles (4). Steady-state levels of 5-aminolevulinic acid (ALA) are usually not detectable, unless radio-labelled substrates are used. The method of choice for non-radioactive quantification is the determination of an ALA synthesis rate instead of steady-state levels. This requires the inhibition of enzymatic ALA conversion by levulinic acid, a competitive inhibitor of ALA dehydratase. Additionally, ALA must be condensed with ethyl acetoacetate to form a pyrrole which can react with Ehrlich’s reagent prior to the colorimetric determination. 1. Cut leaves from Arabidopsis plantlets and incubate the leaf material in light for 2 h in a sufficient volume of incubation buffer (see Note 3). 2. Remove leaves from the buffer and dry them on paper towels; determine the fresh weight and then freeze the samples in liquid nitrogen. 3. Thoroughly homogenise the samples in liquid nitrogen using a pestle and mortar. Store the samples in liquid nitrogen. 4. Resuspend the powder in 1 mL of the extraction buffer. Centrifuge at 4°C in a precooled microcentrifuge at 16,000 × g for 10 min. Transfer the cleared supernatant to a new tube and then keep it on ice. 5. Transfer 400 mL of the supernatant into a new reaction tube and add 100 mL of ethyl acetoacetate. Mix carefully and incubate at 100°C for 10 min (see Note 4). 6. Cool the samples on ice and subsequently add 500 mL of modified Ehrlich’s reagent. Centrifuge the sample for 5 min at 16,000 × g at room temperature to obtain a clear supernatant.
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7. Measure absorption at 526, 553, and 720 nm and calculate the ALA concentration according to a calibration curve (see Note 5). 8. Generate a calibration curve by spiking extraction buffer with known amounts of an ALA stock solution. Final concentrations of the calibration samples should be between 0 and 50 mM ALA. To record the calibration curve, treat the standard solutions like samples as described in steps 5–7 above. 3.2. Determination of Porphobilinogen Steady-State Levels
1. Harvest leaf material, determine the fresh weight, and freeze samples in liquid nitrogen. Grind approximately 250 mg of plant tissue in liquid nitrogen using a pestle and mortar, and extract porphobilinogen (PBG) with 1 mL of extraction buffer. 2. Centrifuge samples in a precooled microcentrifuge at 16,000 × g for 5 min. 3. Mix 400 mL of the supernatant with 500 mL of modified Ehrlich’s reagent and incubate for 10 min. 4. Read absorption at 555 nm and calculate the PBG content using a molar absorption coefficient of e 555 nm = 6.2 × 104 L/ mol/cm (4).
3.3. Extraction of (Mg) Porphyrins According to Mock and Grimm (1)
Since tetrapyrroles are generally hydrophobic, they are extracted from plant tissues or isolated chloroplasts with organic solvents, such as ethanol, methanol, or acetone. An extraction method was initially developed for analysis of porphyrins ranging from uroporphyrinogen III to Mg protoporphyrin IX monomethylester (Mg Proto MME) employing leaf samples from tobacco plants (5). However, this method is not always applicable for Arabidopsis leaves. Firstly, in contrast to tobacco, steady-state levels of Chl precursors are lower in Arabidopsis leaves (see Note 6). Secondly, detection and quantification of tetrapyrroles from Arabidopsis samples are hampered by chlorophyllase activity converting Chl to chlorophyllide (Chlide) in organic solvents of up to 80% acetone or methanol. The accumulation of non-physiological amounts of Chlide interferes with detection of intermediates of Chl biosynthesis (see Note 7). Two commonly used methods to extract and quantify (Mg) porphyrins are presented in this chapter (1, 2). 1. Harvest leaf material, determine the fresh weight, freeze samples in liquid nitrogen, and grind approximately 100 mg of plant tissue in liquid nitrogen using a pestle and mortar (see Note 8). 2. Resuspend the samples in 250 ml of extraction solution 1 and incubate for 5–10 min on ice. 3. Centrifuge the cell extract at 10,000 × g for 10 min at 4°C. Transfer the supernatant into a dark 2.0-mL reaction tube and keep on ice.
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4. Resuspend the pellet in 500 mL of extraction solution 2 and centrifuge at 10,000 × g for 10 min at 4°C. Combine the supernatant with the supernatant from step 3. 5. Resuspend the pellet in 500 mL of extraction solution 3 and keep the solution for 15–20 min at −20°C. Centrifuge the last extract at 16,000 × g for 10 min and combine the supernatant with the supernatants from steps 3 and 4. 6. Centrifuge the extracts at 16,000 × g for 30 min at 4°C (see Note 9). 7. Optional step: For quantification of Uro(gen) III, Copro(gen) III, and Proto(gen) IX, take a 200 mL aliquot of the extract and mix with 5 mL of 1 M acetic acid and 5 mL of 2-butanone peroxide prior to injection into the HPLC (see Note 10). 3.4. Extraction of (Mg) Porphyrins According to Moulin and Smith (2)
1. Harvest leaf material, determine the fresh weight, freeze samples in liquid nitrogen, and grind approximately 50 mg of plant tissue in liquid nitrogen using a pestle and mortar (see Note 8). 2. Add 500 mL of extraction solution 1, mix vigorously, and then centrifuge the extract at 10,000 × g for 10 min. Transfer supernatant to a new dark 2-mL tube. 3. Resuspend the pellet in 500 mL of extraction solution 2. Centrifuge at 10,000 × g for 10 min at 4°C. Combine the supernatant with the supernatant from step 2. 4. Repeat step 3 two times. 5. Centrifuge the extracts at 16,000 × g for 10 min at 4°C and transfer the pooled supernatants to a fresh tube. 6. Dry the pooled supernatants under a stream of N2. Try to keep the pigments in solution as long as possible, since pigments sticking to the upper part of the tube can hardly be redissolved (see Note 2). 7. Redissolve the pigments in 200 mL of extraction solution 2 (see Note 11). 8. Centrifuge the extracts at 16,000 × g for 30 min at 4°C (see Note 9).
3.5. Preparation of (Mg) Porphyrin HPLC Standards
All porphyrin solutions can be prepared according to the method described by Jensen et al. (6) and de Rooij et al. (7). Table 1 summarises the preparation of different HPLC standard stock solutions for porphyrin quantification. A small amount of the porphyrin is dissolved in DMSO and centrifuged (step 1). For determination of the concentration, dilute the stock solution initially 100× (step 2). Increase dilution until there is a linear correlation between absorption (at wavelengths indicated in Table 1, step 4) and dilution factor. Calculate the porphyrin concentration in the stock solution using the molar absorption coefficients given in Table 1. Dilute the
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Table 1 Preparation of porphyrin standards for HPLC Step
Uro III
Copro III
Proto IX
Mg Proto IX Mg Proto IX MME
1
Stock solution
2
Dilute 10 mL 990 mL 990 mL 990 mL 990 mL stock in: 0.5 M HCl 0.1 M HCl 2.7 M HCl ethanol
3 4
Dissolve a small amount in a 1 mL DMSO and centrifuge at 16,000 × g for 5 min at room temperature 990 mL diethyl ether
Read absorption spectra from 300 to 700 nm Absorption maximum
405 nm
400 nm
554 nm
419 nm
419 nm
e [L/mmol/ cm]
541
489
13.5
308
100
5
Dilute stock in 80% (v/v) acetone and 10% (v/v) 0.1 M NH4OH to a final concentration of 100 nM
Table 2 Solvent elution programme for HPLC separation of tetrapyrroles according to Mock and Grimm (1) Time (min)
Solvent A (%)
Solvent B (%)
0
100
0
7
0
100
20
0
100
23
100
0
29
100
0
stock solution stepwise to a final concentration of 100 nM (step 5). Inject 5 mL (0.5 pmol), 10 mL (1.0 pmol), and 20 mL (2.0 pmol) to calibrate the HPLC (see Subheading 3.6 or 3.7). 3.6. HPLC Analysis of (Mg) Porphyrins According to Mock and Grimm (1)
1. Set the solvent flow rate to 1.0 mL/min. Set the solvent elution programme as given in Table 2. 2. Inject 10–100 mL of the porphyrin extracts or 0.5, 1, and 2 pmol of the standards. 3. Elution of tetrapyrroles can be monitored by fluorescence (see Note 12). For detection of Uro III, Copro III, and Proto IX, set the excitation wavelength to 405 nm and monitor fluorescence at 625 nm. For detection of Mg Proto IX and Mg Proto IX MME, set the excitation wavelength to 420 nm and monitor fluorescence at 595 nm (see Note 13).
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Table 3 Solvent elution programme for HPLC separation of tetrapyrroles according to Moulin and Smith (2) Time (min)
Solvent A (%)
Solvent B (%)
Solvent C (%)
Solvent D (%)
0
40
60
0
0
2
40
60
0
0
25
10
0
70
20
35
10
0
70
20
40
40
60
0
0
4. Identify the (Mg) porphyrins by their retention time and fluorescence spectra. Calculate the amounts of (Mg) porphyrin in the samples according to the standard peak areas. 3.7. HPLC Analysis of (Mg) Porphyrins According to Moulin and Smith (2)
1. Set the solvent flow rate to 0.2 mL/min. Set the solvent elution programme as given in Table 3. 2. Inject 10–50 mL of the porphyrin extracts or 0.5, 1, and 2 pmol of the standards. 3. Elution of tetrapyrroles can be monitored either by a photodiode array (350–800 nm) or by fluorescence (see Subheading 3.6 and Notes 12 and 13). 4. Identify the (Mg) porphyrins by their retention time and fluorescence spectra. Calculate the amounts of (Mg) porphyrin in the samples according to the standard peak areas.
3.8. Extraction of Protochlorophyllide and Chlorophyllide
Pchlide accumulates during darkness and in etiolated plant seedlings, but steady-state levels are also detectable in light-exposed samples. In all angiosperms, NADPH:Pchlide oxidoreductase (POR) light-dependently converts Pchlide to Chlide. To prevent reduction of Pchlide to Chlide during extraction, POR needs to be inactivated, which is achieved by steam fixation of leaf material. When Pchlide levels are analysed from green leaf material, Chl has to be removed from extracts by hexane washing. 1a. For Pchlide determination, harvest etiolated or dark-incubated leaf samples under green safety light, determine the fresh weight, and then transfer the material into 1.5-mL reaction tubes (see Note 8). Continue with step 2.
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1b. When Chlide contents should be determined, harvest green plant material, and determine the fresh weight. Freeze samples in liquid nitrogen. Continue with step 3 (see Note 14). 2. Fix the opened tubes containing plant material on a mesh and put them into steam over boiling water for 2 min. Subsequently freeze the samples in liquid nitrogen. All following steps should be carried out under dim light. 3. Grind approximately 100 mg leaf material in 1 mL of extraction solution using a pestle and mortar. Centrifuge for 10 min at 10,000 × g and transfer the supernatant to a fresh test tube. 4. Re-extract the pellet with 1 mL of fresh extraction solution. Centrifuge again and then combine the supernatants. When the extracts are free of Chl, Pchlide can be detected directly (continue with Subheading 3.8.1 or 3.8.2; see Note 9); otherwise, proceed with steps 5–8 below (note that steps 5–8 below are optional, as they are only required when Chl-containing samples are analysed). 5. Add 1 volume of n-hexane to the combined acetone extracts in a 15-mL tube and mix thoroughly. Centrifuge for 1 min at 10,000 × g. Remove and discard the upper hexane phase (containing Chl). 6. Repeat the washing with 1 volume of hexane. 7. Repeat the washing with 0.3 volume of hexane. 8. The remaining acetone phase is a Chl-free Pchlide and Chlide extract. Adjust the volumes of all samples by adding extraction solution. 3.8.1. Quantification of Pchlide and Chlide Using a Spectrofluorometer
Monitor the fluorescence emission spectra of the samples between 600 and 700 nm using an excitation wavelength of 433 nm. Dilute the samples with extraction solution until there is a linear correlation between dilution factor and fluorescence intensity. Determine maximum fluorescence at 633 nm (Pchlide) and 675 nm (Chlide). When samples contain Chlide, its fluorescence will be subtracted to obtain the value of Pchlide fluorescence. Calculate Pchlide fluorescence intensity using the following equation: Fmax Pchlide = Fmax 633 nm − 0.032 Fmax 675 nm. Determine the Pchlide and Chlide content of the samples by comparing the fluorescence intensities of the samples with Pchlide and Chlide standards (see Subheading 3.9).
3.8.2. Quantification of Pchlide and Chlide via HPLC
Quantification of Pchlide and Chlide can be performed using an HPLC system with a fluorescence detector (8). 1. Set the solvent flow rate to 1.0 mL/min. Set the solvent elution programme as given in Table 4.
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Table 4 Solvent elution programme for HPLC separation of Pchlide and Chlide Time (min)
Solvent A (%)
Solvent B (%)
0
100
0
15
0
100
25
0
100
28
100
0
30
100
0
2. Inject 10–50 mL of the extracts or 0.5, 1.0, and 2.0 pmol of the standards (see Subheading 3.9). 3. Elution of Pchlide and Chlide can be monitored by fluorescence (see Note 12). Set the excitation wavelength to 435 nm and monitor fluorescence at 635 nm or 680 nm for Pchlide or Chlide detection, respectively (see Note 13). 4. Calculate the Pchlide and Chlide amounts in the samples according to the standard peak areas. 3.9. Preparation of Protochlorophyllide and Chlorophyllide Standards 3.9.1. Preparation of a Pchlide Standard
Pchlide extraction from etiolated barley seedlings follows the protocol of Koski and Smith (9).
1. Fix 5 g of etiolated barley leaf material in steam for 2 min (see Subheading 3.8). 2. Grind the leaf material in 20 mL of extraction solution. Centrifuge and then transfer the supernatant to a fresh tube. 3. Remove carotenoids by washing with 1 volume of n-hexane. Centrifuge briefly for better phase separation. Remove and discard hexane phase. 4. Repeat step 3 two times. 5. Add 5 mL of diethyl ether, 2 mL of saturated KH2PO4, and 2 mL of saturated NaCl to transfer Pchlide from the acetone to the ether phase. Transfer the ether phase to a fresh tube. This step improves stability of the extracted Pchlide. 6. Wash the diethyl ether with 1 volume of water to remove remaining acetone. Transfer the ether phase to a fresh tube. The ether extract can be stored in darkness and at −20°C for approximately 2 weeks.
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7. Determine the concentration of Pchlide in the ether extract spectrophotometrically at 623 nm using a molar absorption coefficient of e Pchlide in diethyl ether, 623 nm = 3.56 × 104 L/mol/cm (10). 8. Prepare a 100 nM standard solution by diluting the ether extract in extraction solution. 3.9.2. Preparation of Chlide Standard
1. Perform steps 1–6 above using illuminated barley seedlings as described for Pchlide standard preparation (see Subheading 3.9.1); steam fixation is not necessary. Transfer of Chlide to ether is not mandatory. 2. Determine the concentration of Chlide in the acetone or ether extract spectrophotometrically at 663 nm using a molar absorption coefficient of eChlide in acetone, 663 nm = 7.51 × 104 L/mol/cm (10) or eChlide in diethyl ether, 663 nm = 9.12 × 104 L/mol/cm (10). 3. Prepare a 100 nM standard solution by diluting the extract in extraction solution.
3.10. Spectrophoto metrical Determination of Chlorophylls and Carotenoids
Chl and carotenoids (Car) can be quantitatively and qualitatively extracted with organic solvents. Higher plants, ferns, mosses, and algae contain Chl a and Chl b: Chl a is the more abundant pigment in photosynthetic protein complexes and forms the reaction centre pigment (special pair), while Chl b functions as an accessory pigment. Carotenoids serve as additional pigments of the photosynthetic apparatus and broaden the range of wavelengths that can be absorbed. The spectrophotometrical determination of the pigment content of a leaf sample is the simplest and fastest method. There are two commonly used references for equations to calculate the pigment contents in organic solvents (11, 12). 1. Harvest leaf material, determine the fresh weight, freeze samples in liquid nitrogen, and then grind approximately 50 mg of plant tissue in liquid nitrogen using a pestle and mortar (see Note 8). 2. Add 500 mL of extraction solution, mix vigorously, and then centrifuge the extract at 10,000 × g for 10 min. Transfer the supernatant to a new tube. 3. Repeat step 2 until the pellet obtained by centrifugation is almost white. Combine the supernatants. 4. Read absorption of the extracts at wavelengths given in Tables 5–7. Dilute samples in extraction solution until there is a linear correlation between dilution factor and absorption. Calculate the Chl concentrations in the extracts according to the equations of Lichtenthaler (11) given in Tables 5 and 6 or of Porra et al. (12) given in Table 7. Calculate the total Car concentration in the extracts according to the equations of Lichtenthaler (11) given in Tables 5 and 6.
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Table 5 Equations for the determination of the concentration of Chl and Car in leaf extracts in commonly used acetone-containing solvents according to Lichtenthaler (11)* 100% acetonea
80% acetone a
Chl a
11.24 A662 − 2.04 A645
12.25 A663 − 2.79 A647
Chl b
20.13 A645 − 4.19 A662
21.50 A647 − 5.10 A663
Chl (a + b)
7.05 A662 + 18.09 A645
7.15 A663 + 18.71 A647
Car
[1,000 A470 − 1.90 (Chl a) − 63.14 (Chl b)]/214
[1,000 A470 − 1.82 (Chl a) − 85.02 (Chl b)]/198
Axxx denotes absorption at xxx nm * Concentrations are calculated as mg/mL and can be converted to molar units by the molar weights given in Note 15
a
Table 6 Equations for the determination of the concentration of Chl and Car in leaf extracts in commonly used methanol-containing solvents according to Lichtenthaler (11)* 100% methanola
90% methanola
Chl a
16.72 A665 − 9.16 A652
16.82 A665 − 9.28 A652
Chl b
34.09 A652 − 15.28 A665
36.92 A652 − 16.54 A665
Chl (a + b)
1.44 A665 + 24.93 A652
0.28 A665 + 27.64 A652
Car
[1,000 A470 − 1.63 (Chl a) − 104.96 (Chl b)]/221
[1,000 A470 − 1.91 (Chl a) − 95.15 (Chl b)]/225
Axxx denotes absorption at xxx nm * Concentrations are calculated as mg/mL and can be converted to molar units by the molar weights given in Note 15
a
Table 7 Equations for the determination of the concentration of Chl in leaf extracts according to Porra et al. (12)* 100% methanola
in 80% acetonea
Chl a
16.29 A665 − 8.54 A652
12.25 A664 − 2.55 A647
Chl b
30.66 A652 − 13.58 A665
20.31 A647 − 4.91 A664
Chl (a + b)
2.71 A665 + 22.12 A652
7.34 A664 + 17.76 A647
Axxx denotes absorption at xxx nm * Concentrations are calculated as mg/mL and can be converted to molar units by the molar weights given in Note 15
a
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3.11. Quantification of Carotenoids and Chlorophylls by HPLC
In general, information on the total carotenoid content of a leaf sample (as determined in Subheading 3.10) is not sufficient. For more detailed information, for example on the composition and quantities of different carotenoid species, separation and detection of individual carotenoid species can be achieved by means of HPLC. Using this method, Chl a, Chl b, and chlorophylide can be quantified in parallel. In the first step of this procedure, carotenoids are extracted together with Chl (see Subheading 3.10, steps 1–3).
3.11.1. Preparation of Chl and Carotenoid HPLC Standards
Carotenoid standards can presently not be purchased but they can easily be prepared together with Chl from Arabidopsis pigment extracts. In the course of an HPLC-driven separation of pigment extracts (see Subheading 3.11.2), carotenoid and Chl fractions can be directly collected at the outlet of the PDA detector. Carotenoid or Chl species can be identified according to their absorption spectra, and standard concentrations can be determined spectrophotometrically using known absorption coefficients (11, 13, 14). 1. Prepare a pigment solution (see Subheadings 3.10 and 3.11). Use up to 1 g of plant material and linearly increase the extraction volume. 2. Perform an HPLC run and inject a small volume (e.g., 10 mL, see Subheading 3.11.2) of the pigment extract. 3. Monitor the pigment elution and identify the carotenoids and Chls according to the retention times indicated in Table 8 and also by their typical absorption spectra (13).
Table 8 Retention time and molar extinction coefficients of Chl a, Chl b, and different carotenoids* Retention time (min)
Wavelength (nm)
Neoxanthin
9
Violaxanthin
Molar and specific absorption coefficient (L/mol/cm)
(mL/mg/cm)
438
1.35 × 105
0.224
11
441
1.50 × 105
0.250
Antheraxanthin
14
444
1.37 × 105
0.235
Lutein
16
445
1.45 × 10
0.254
Zeaxanthin
17
450
5
1.45 × 10
0.254
Chlorophyll b
18
645
4.69 × 104
0.052
Chlorophyll a
20
663
8.26 × 104
0.092
b-Carotene
26
451
1.41 × 10
0.262
5
5
*Specific absorption coefficients at indicated wavelengths are given according to Rowan (14) and Lichtenthaler (11) and molar absorption coefficients were calculated
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4. Perform another HPLC run and inject as much of the pigment extract as possible (e.g., 200 mL). Collect chromatographically separated carotenoids and Chl directly on the outlet of the PDA detector using a 1.5-mL reaction tube. Decide which fraction contains the desired pigments based in the HPLC run in step 3 and the retention times given in Table 8. 5. Repeat step 4 several times (e.g., five times) and combine identical fractions. 6. Dry the collected carotenoids and Chl standards under a stream of N2 or by using a SpeedVac (see Note 2). 7. Dissolve the carotenoid standards in a small volume (e.g., 100 mL) of ethanol. Dissolve the Chl a and b standards in a small volume (e.g., 100 mL) of acetone. 8. Determine the concentration of the carotenoid and Chl standards spectrophotometrically at the wavelengths indicated in Table 8. Dilute the standards until there is a linear correlation between absorption and dilution factor. Calculate the concentration of the standards using the absorption coefficients given in Table 8. We provide molar (L/mol/cm) and specific (mL/mg/ cm) absorption coefficients because both are commonly used. 9. Dilute the standard solutions to a final concentration of 2 mM or 1 ng/mL. Inject 10 mL (20 pmol or 10 ng), 30 mL (60 pmol or 30 ng), 50 mL (100 pmol or 50 ng), and 75 mL (150 pmol or 75 ng) to calibrate the HPLC (see Subheading 3.11.2). 3.11.2. Quantification of Chl and Different Carotenoid Species by HPLC
1. Set the solvent flow rate to 1.0 mL/min. Set the solvent elution programme as given in Table 9.
Table 9 Solvent elution programme for HPLC separation of Chl and carotenoids Time (min)
Solvent A (%)
Solvent B (%)
0
100
0
1
100
0
16
66.7
33.3
22
59.7
40.3
34
33.3
66.7
34.2
0
100
36.7
0
100
36.9
100
0
41
100
0
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2. Inject 10–100 mL of the pigment extracts or standards (see Subheading 3.11.1). 3. Elution of the pigments can be monitored by measuring absorption at a wavelength of 440 nm. 4. Identify the Chl and carotenoids by their retention times and absorption spectra. Calculate the amounts of pigments in the samples according to the standard peak areas. 3.12. Extraction of Haem from Green Tissues
Non-covalently bound haem (mainly the b-type haem) is extracted from plant tissues or chloroplasts using acidic acetone after removal of pigments and lipids by using cold alkaline acetone (15). There are different methods to quantify extracted haem. The pyridine hemochrome method (16) is based on the complex formation of haem and pyridine. Different absorption spectra of hemi- and hemochromes (central Fe2+ or Fe3+ ion) can be used to calculate the haem content. Here, we present two other quantification methods. Firstly, haem can be purified by ion exchange chromatography and quantified by absorption at 398 nm (see Subheading 3.13) (17). The second and more sensitive approach was developed recently (18, 19) and uses the reconstitution of horseradish peroxidase (HRP) activity by application of haemcontaining extracts to the HRP apoenzyme. The latter method allows haem quantification in a pM range or from less than 5 mg plant material, and is very useful to determine haem contents in high-throughput assays or when plant material is limited (see Subheading 3.14). 1. Grind up to 1 g (depending on the subsequent quantification method; see preamble above) of frozen leaf material using a precooled pestle and mortar. 2. Dissolve the powder in 5 mL of ice-cold extraction solution 1 per gram fresh weight and transfer the suspension to 15-mL centrifuge tubes. Scale up or down linearly all volumes when more or less plant material is used. 3. Centrifuge at 10,000 × g for 10 min and discard the supernatant (see Note 16). 4. Repeat the pigment extraction with ice-cold extraction solution 1 until all Chl is removed from the pellet (see Note 16). 5. Extract haem from the pellet by adding 5 mL of extraction solution 2 per gram fresh weight and mix thoroughly. Incubate for 10 min at room temperature and centrifuge at 10,000 × g for 10 min. Transfer the haem-containing supernatant to a fresh 50-mL centrifuge tube. 6. Repeat the extraction step (step 5 above) and combine the supernatants. Continue with step 7 when ion exchange chromatography is to be used to quantify the haem content of the
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sample (see Subheading 3.13). For haem quantification by reconstitution of HRP activity, continue directly to Subheading 3.14. 7. Add 3 mL of diethyl ether, 2 mL of saturated NaCl solution, and 10 mL of water (in this order) per 10 mL of haem extract (from step 6); mix well; and then centrifuge for 1 min at 1,000 × g for better phase separation. Transfer the upper (mostly yellowish) ether phase to a fresh 15-mL centrifuge tube (see Note 17). 8. Add another 3 mL of diethyl ether to the remaining acetone extract, mix, and then centrifuge for 1 min at 1,000 × g. Combine both ether phases. 9. Centrifuge the ether extracts for 1 min at 1,000 × g and remove any remaining water or acetone from the bottom of the tube using a Pasteur pipette. 10. Reduce the volume of the haem-containing ether extract to 2 mL by applying a continuous air flow or by incubating in a water bath at approximately 30°C (see Note 18). 11. Add ethanol (approximately 0.7 mL) to have a final solution of 75% (v/v) diethyl ether and 25% (v/v) ethanol. 3.13. Quantification of Haem Using Ion Exchange Chromatography
1. Mix sepharose suspension (approximately 4–5 mL per sample) with 1 volume of water in a beaker and decant the supernatant from sedimented sepharose. 2. Resuspend the sepharose in 1 volume of 1 M Na acetate, pH 7.0, and then discard the Na acetate after allowing the sepharose to settle. 3. Wash the sepharose with 1 volume of water and discard the supernatant after allowing the sepharose to settle. 4. Resuspend the sepharose in 1 volume of acetone. Fill polypropylene columns with 4–5 mL of sepharose suspension; the final volume after sedimentation will be around 2–3 mL of sepharose. The sepharose must be sedimented before the acetone is removed by gravity flow. 5. Equilibrate the column matrix with 2 column volumes of wash solution 1 (see Note 19). 6. Carefully apply the sample to the column (see Note 19). Allow the solvent to pass through the column by gravity flow. 7. Wash with 2 column volumes of wash solution 1 (see Note 19). 8. Wash with 1 column volume of wash solution 2 (see Note 19). 9. Wash with 1 column volume of ethanol (see Note 19). 10. Elute haem with 2 column volumes of elution solution. Collect the flow-through. Determine the exact elution volume.
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11. Quantify the haem concentration in the eluate spectro photometrically. Monitor the absorption spectrum between 300 and 720 nm. Absorbance at 720 nm should be zero. Calculate the haem content using the molar absorption coefficient e 398 nm = 144 L/mmol/cm (17). 3.14. Quantification of Haem in HighThroughput Assays
HRP apoenzyme spontaneously reconstitutes with haem, resulting in an active enzyme (20). The peroxidase activity of HRP is used to oxidise luminol in the presence of H2O2. The resulting luminescence signal is proportional to the amount of haem that is present in the sample. Luminescence signals can be detected in two ways: (A) by directly measuring the luminescence using a luminometer or a 96-well plate reader for luminescence at 430 nm; or (B) by using a sensitive CCD camera followed by densitometric analysis of the spots. In both cases, it is important to calibrate with hemin standards and to adjust the signal range of the calibration standards and the samples. Although this step is time consuming, precise and cautious calibration ensures high reproducibility of results. Because the standard homemade HRP substrate typically used for Western blotting is not sensitive enough for this assay, commercial Western blot detection kits containing signal enhancers should be used.
3.14.1. Calibration Solutions
Prepare at least two different sets of calibration solutions by serial dilution of the 1 mM hemin stock in 10 mM KOH. The following concentrations are initially recommended: Set 1: 0, 5, 25, 50, 100, 200, 300, and 400 pM. Set 2: 0, 250, 500, 1,000, 2,000, 2,500, 3,000, and 4,000 pM (see Note 20).
3.14.2. Sample Dilution
Haem extracts (see step 6, Subheading 3.12) need to be diluted in 10 mM KOH before they can be applied to the enzyme assay because the acetone interferes with HRP. The minimum dilution of the acetone extract is 1,000-fold. Prepare different dilutions of the sample (e.g., 1,000-, 2,000-, and 4,000-fold) to ensure that the luminescence signal linearly correlates with the haem concentration in the sample (see Note 20).
3.14.3. Method
Table 10 illustrates how the 96-well plate can be used. The haem content of each standard (S1–S8) and sample should be measured in triplicate. The samples A–X in the scheme consist of different samples or different dilutions of identical samples. 1. Prepare a fresh assay master mix: per reaction, 50 mL of 2× reaction buffer and 30 mL of H2O. Add HRP stock solution to a final concentration of 2.5 nM. 2. Add 80 mL of the master mix in each well of a white 96-well plate; use an 8-channel pipette if available.
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Table 10 Pipetting scheme of a 96-well plate 1
2
3
4
5
6
7
8
9
10
11
12
A
S1
S1
S1
A
A
A
I
I
I
Q
Q
Q
B
S2
S2
S2
B
B
B
J
J
J
R
R
R
C
S3
S3
S3
C
C
C
K
K
K
S
S
S
D
S4
S4
S4
D
D
D
L
L
L
T
T
T
E
S5
S5
S5
E
E
E
M
M
M
U
U
U
F
S6
S6
S6
F
F
F
N
N
N
V
V
V
G
S7
S7
S7
G
G
G
O
O
O
W
W
W
H
S8
S8
S8
H
H
H
P
P
P
X
X
X
3. Add 20 mL of standard solution or the diluted sample. Mix well by pipetting up and down. 4. Incubate for 30 min at room temperature. 5. Add 100 mL of Western blot detection reagent prepared according to the manufacturer’s instructions. 6. Mix well by pipetting up and down. Incubate the assay mix according to the manufacturer’s instructions. 7a. Measure luminescence at 430 nm using a 96-well plate reader in the luminescence mode. 7b. Alternatively, monitor luminescence using a CCD camera. Quantify the intensity of the spots densitometrically. 8. Calculate the haem content of the samples by comparing with calibration standards. 9. If the luminescence signal differs between standard solutions and sample dilutions, prepare new sets of standard solutions or use different sample dilutions. 3.15. Homemade Horseradish Peroxidase Apoenzyme
If commercial HRP apoenzyme is not available, it can be prepared using the holoenzyme as follows (20–22). 1. Dissolve the holoenzyme in 0.05 M HCl. The concentration of the enzyme should be around 1% (w/v) (e.g., 50 mg in 5 mL). Cool down the sample on ice. 2. Wash with 1 volume of ice-cold 2-butanone. Centrifuge for 1 min at 1,000 × g for better phase separation and then discard the upper, brown haem-containing phase. 3. Wash with 0.5 volume of ice-cold 2-butanone and discard the upper phase after centrifugation.
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4. Repeat the washing step (step 3). 5. Exchange the buffer against 600 mM NaHCO3 until all butanone is removed (the characteristic smell disappears) using concentrator tubes or dialysis tubes. 6. Exchange the NaHCO3 against H2O using concentrator tubes or dialysis tubes. 7. Determine the concentration of the apoprotein solution spectrophotometrically using a molar absorption coefficient of e HPR in H2O, 278 nm = 20 L/mmol/cm (20). Dilute or concentrate the protein solution to 2.5 mM. Store it at −80°C. 3.16. Quantification of Chlorophyll Catabolites
Chl a breakdown includes the removal of the central Mg ion to form pheophytin a. Subsequently, the phytol chain is removed and pheophorbide a is released. Pheophorbide a is further processed to red Chl catabolites (RCC), fluorescent Chl catabolites (FCC), and finally non-fluorescent Chl catabolites (NCC) that are stored in the vacuole (23, 24). Usually, Chl catabolites can only be analysed in senescent leaves. Since there is a huge number of chemical variations among FCCs and NCCs, we cannot provide detailed protocols to identify and quantify all these Chl catabolites. Please refer to (24) and references therein for more detailed information. 1. Harvest leaf material, determine the fresh weight, freeze the samples in liquid nitrogen, and then grind up to 1 g plant tissue in liquid nitrogen using a pestle and mortar. 2. Add 20 mL of extraction solution per gram fresh weight, mix vigorously, and then incubate for 2 h at −20°C in darkness. Centrifuge the extract at 10,000 × g for 10 min. Transfer the supernatant to a new tube. 3. Use the HPLC system and setup as described for Chlide detection (see Subheading 3.8.2). 4. Inject 10–100 mL of the extracts or 0.5, 1.0, and 2.0 pmol of the pheophytin a or pheophorbide a standards. 5. Calculate the pheophytin a and pheophorbide a amounts in the samples according to the standard peak areas.
4. Notes 1. Solid levulinic acid is a hygroscopic compound. More convenient is the use of commercially available levulinic acid solution. Alternatively, prepare a concentrated levulinic acid stock solution directly after purchasing the powder. 2. Dry the samples with care. It is very important to keep the pigments in solution as long as possible by repeated vigorous
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vortexing during the drying process. This makes subsequent resuspension easier. Instead of applying a stream of N2, a SpeedVac can be used to evaporate the solvent. 3. The activity of enzymes of ALA synthesis is high in young and greening tissues and decreases in older leaves. Furthermore, the enzyme activity changes during daytime and follows diurnal oscillation with a maximum activity directly after the darkto-light transition. The activity decreases in the second half of the light period and is hardly detectable in darkness. These varying activities of ALA synthesis have to be considered when the incubation time of leaf discs with levulinic acid is assessed. A minimum of 30-min incubation with levulinic acid is recommended (25). ALA accumulation in levulinic acid-treated young leaves occurs at a constant rate for approximately 3 h. A longer incubation may lead to incorrect results, and should only be used in experiments with samples of older leaves or when ALA synthesis should be determined at unfavourable times of the day. 4. The samples should be vigorously shaken to mix ethyl acetoacetate and the ALA-containing buffer. Repeated vortexing of the sample may help to prevent phase separation during the incubation at 100°C. When mixing or handling the samples, ensure that the tubes are closed safely to avoid injuries due to hot spraying. Prepare a blank for the spectrophotometric measurement. Use 400 mL of extraction buffer and proceed as described for the samples (steps 5–7). 5. The measurement of the absorption should be performed within 30 min after addition of modified Ehrlich’s reagent, since prolonged incubation may cause changes in the absorption of the coloured complex. The absorbance ratio between the two wavelengths (553–526 nm) should be 1.3–1.5. If the absorption of the sample is too high, dilute an appropriate volume of the cleared extract (step 4) with extraction buffer and repeat steps 5–7. 6. The first reliable and reproducible detections of intermediates of Chl biosynthesis were possible upon ALA feeding. Usually, concentrations between 0.1 mM ALA (for long-term experiments) and 5 mM ALA (for short-term experiments) were applied to (Arabidopsis) plantlets or leaf samples. Nevertheless, this approach causes accumulation of intermediates and is suitable to determine limiting or impaired enzymatic steps in mutants, but does not necessarily allow one to draw conclusions about steady-state levels of (Mg) porphyrins in unfed plants. Nowadays, technological progress in combination with improved methods for extraction and detection enables measurement of these steady-state levels.
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7. Besides the low steady-state porphyrin levels in Arabidopsis leaves, the extraction and detection of (Mg) porphyrins suffer from two other main problems. Firstly, disruption of plant tissues and sample homogenisation release the acidic content of the vacuole, which can provoke the loss of the central Mg ion of Mg porphyrins and Chls. Use of alkaline or buffered extraction solutions avoids this problem. Secondly, Arabidopsis seedlings possess a very rugged chlorophyllase activity, which is still active in 80% (v/v) acetone or methanol. Therefore, the measurement of Mg porphyrins can be disturbed by accumulating Chlide. Certainly, the easiest way to avoid this problem is to use sufficient volumes of the extraction solutions and to minimise the aqueous fraction of the extract. To avoid hydrolysis of chlorophyll, take heed that the first extraction of the leaf sample should occur in a large buffer volume relative to the fresh weight of the sample. 8. Please note that all steps for extraction should be carried out under green safety light or dim light, and samples as well as extracts should be kept on ice, since many porphyrins are very labile. Therefore, it is also recommended that extracts be analysed directly after extraction, since prolonged storage may lead to degradation of porphyrins. 9. Any remaining particles (e.g., cell debris) in the extract should be removed by centrifugation prior to HPLC analysis to avoid clogging of the column. 10. Protogen IX is non-enzymatically converted to Proto IX in the presence of light and oxygen. To avoid quantification errors caused by partial conversion of Protogen IX to Proto IX, a complete oxidation of Protogen IX by butanone peroxide in the presence of acetic acid is recommended. Hence, the measured “Proto IX pool” cumulatively represents both intermediates. 11. Resuspending the dried pigments may take some time. Again, repeated vortexing can aid the dissolving of the precipitated pigments. 12. Even though porphyrins can be analysed using a PDA detector, the fluorescence-based analysis is more sensitive and allows the detection of femtomoles of tetrapyrroles. 13. However, the fluorescence-based detection of tetrapyrroles has one essential drawback. Due to the different excitation and emission wavelengths, the detection of Proto IX and Mg Proto IX (MME) or Pchlide and Chlide requires separate sample runs, unless a second fluorescence detector can be used in parallel. 14. For the extraction of Pchlide and Chlide from green, lightexposed tissues, the steam fixation (step 2) is optional. The heat treatment of etiolated or dark-incubated plants should
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inactivate the POR, which light-dependently converts Pchlide to Chlide. If the enzyme is not inactivated, Pchlide that accumulates in dark grown plants is rapidly decomposed upon light exposure. Nevertheless, the detection of steadystate levels of light-grown plants does not require POR inactivation. 15. The molar weights of Chl a and b are 893.49 and 907.47 g/mol respectively. 16. If the determination of Chl from the same leaf sample is required, the supernatants from steps 3 and 4 should be collected and combined in a 50-mL tube. After dilution (e.g., in 80% acetone), the Chl content can be determined as described in Subheading 3.10. When processing non-green tissues (e.g., roots or etiolated plants), the use of alkaline acetone for Chl removal (steps 2–4) is not required. After homogenising the sample (step 1), proceed directly to step 5. 17. Because of the high vapour pressure, saturate the piston volume of the pipette with diethyl ether by pipetting a fresh volume diethyl ether up and down. Working under a fume hood is recommended when handling diethyl ether. 18. To avoid boiling of ether, do not increase the temperature. The boiling point of ether is at approximately 35°C. 19. Pipette carefully when adding new wash or elution solvent to the column. Do not disturb the settled sepharose, as this may interfere with binding and elution of haem. Do not allow the column to run dry. 20. The decision, which dilution fits best, can only be made from case to case. Ensure that the intensity of the luminescence signal of the standard dilution series and that of the sample are in the same range. For the sake of faster and easier sample processing, fill diluted samples and standards in a 96-well plate. This enables the use of an 8-channel pipette for further steps and makes handling of large sets of samples more convenient. References 1. Mock, H. P., and Grimm, B. (1997) Reduction of uroporphyrinogen decarboxylase by antisense RNA expression affects activities of other enzymes involved in tetrapyrrole biosynthesis and leads to light-dependent necrosis. Plant Physiol. 113, 1101–1112. 2. Moulin, M., and Smith, A. G. (2008) A robust method for determination of chlorophyll intermediates by tandem mass spectrometry. In, Photosynthesis. Energy from the Sun (Allen, J.F., Gantt, E., Golbeck, J.H. and Osmond, B., eds.) Springer, Dordrecht, Netherlands, pp. 1215–1222.
3. Lorenzen, C. J., and Downs, J. N. (1986) The specific absorption-coefficients of chlorophyllide-a and pheophorbide-a in 90-percent acetone, and comments on the fluorometricdetermination of chlorophyll and pheopigments. Limnol. Oceanogr. 31, 449–452. 4. Mauzerall, D., and Granick, S. (1956) The occurrence and determination of delta-aminolevulinic acid and porphobilinogen in urine. J. Biol. Chem. 219, 435–446. 5. Kruse, E., Mock, H. P., and Grimm, B. (1995) Reduction of coproporphyrinogen oxidase level by antisense RNA synthesis leads to
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11. 12.
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deregulated gene expression of plastid proteins and affects the oxidative defense system. EMBO J. 14, 3712–3720. Jensen, P. E., Gibson, L. C. D., Shephard, F., Smith, V., and Hunter, C. N. (1999) Introduction of a new branchpoint in tetrapyrrole biosynthesis in Escherichia coli by co-expression of genes encoding the chlorophyll-specific enzymes magnesium chelatase and magnesium protoporphyrin methyltransferase. FEBS Lett. 455, 349–354. de Rooij, F. W. M., Edixhoven, A., and Wilson, J. H. P. (2003) Porphyria: A diagnostic approach. In, The Porphyrin Handbook, Vol. 14 (Kadish, K.M., Smith, K.M. and Guilard, R., eds.) Academic Press, Amsterdam, Netherlands, pp. 211–245. Langmeier, M., Ginsburg, S., and Matile, P. (1993) Chlorophyll breakdown in senescent leaves: demonstration of Mg-dechelatase activity. Physiol. Plant. 89, 347–353. Koski, V. M., and Smith, J. H. (1948) The isolation and spectral absorption properties of protochlorophyll from etiolated barley seedlings. J. Am. Chem. Soc. 70, 3558–3562. Dawson, R. C. M., Elliott, D. C., Elliot, W. H., and Jones, K. M. (1986) Data for Biochemical Research, 3rd edn. Oxford University Press, Oxford, UK. Lichtenthaler, H. K. (1987) Chlorophylls and carotenoids: pigments of photosynthetic membranes. Meth. Enzymol. 148, 350–383. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Determination of accurate extinction coefficients and simultaneous-equations for assaying chlorophyll-a and chlorophyll-b extracted with 4 different solvents - verification of the concentration of chlorophyll standards by atomic-absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394. Britton, G., Liaaen-Jensen, S., and Pfander, H. (2004) Carotenoids, Handbook, 1st edn. Birkhäuser Verlag, Basel, Switzerland. Rowan, K. S. (1989) Photosynthetic Pigments of Algae. Cambridge University Press, Cambridge, UK. Stillman, L. C., and Gassman, M. L. (1978) Protoheme extraction from plant-tissue. Anal. Biochem. 91, 166–172.
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16. Berry, E. A., and Trumpower, B. L. (1987) Simultaneous determination of hemes-a, hemes-b, and hemes-c from pyridine hemochrome spectra. Anal. Biochem. 161, 1–15. 17. Weinstein, J. D., and Beale, S. I. (1983) Separate physiological roles and subcellular compartments for two tetrapyrrole biosynthetic pathways in Euglena gracilis. J. Biol. Chem. 258, 6799–6807. 18. Masuda, T., and Takahashi, S. (2006) Chemiluminescent-based method for heme determination by reconstitution with horseradish peroxidase apo-enzyme. Anal. Biochem. 355, 307–309. 19. Takahashi, S., and Masuda, T. (2009) High throughput heme assay by detection of chemiluminescence of reconstituted horseradish peroxidase. Comb. Chem. High. Throughput Screen. 12, 532–535. 20. Tamura, M., Yonetani, T., and Asakura, T. (1972) Heme-modification studies on horseradish-peroxidase. Biochim. Biophys. Acta 268, 292–304. 21. Teale, F. W. J. (1959) Cleavage of the haemprotein link by acid methylethylketone. Biochim. Biophys. Acta 35, 543–543. 22. Breslow, E. (1964) Changes in side chain reactivity accompanying binding of heme to sperm whale apomyoglobin. J. Biol. Chem. 239, 486–496. 23. Schelbert, S., Aubry, S., Burla, B., Agne, B., Kessler, F., Krupinska, K., and Hörtensteiner, S. (2009) Pheophytin pheophorbide hydrolase (pheophytinase) is involved in chlorophyll breakdown during leaf senescence in Arabidopsis. Plant Cell 21, 767–785. 24. Pruzinska, A., Tanner, G., Aubry, S., Anders, I., Moser, S., Muller, T., Ongania, K. H., Kräutler, B., Youn, J. Y., Liljegren, S. J., and Hortensteiner, S. (2005) Chlorophyll breakdown in senescent Arabidopsis leaves. Characterization of chlorophyll catabolites and of chlorophyll catabolic enzymes involved in the degreening reaction. Plant Physiol. 139, 52–63. 25. Richter, A., Peter, E., Pörs, Y., Lorenzen, S., Grimm, B., and Czarnecki, O. (2010) Rapid dark repression of 5-aminolevulinic acid synthesis in green barley leaves. Plant Cell Physiol. 51, 670–681.
Chapter 21 Analysis of Starch Metabolism in Chloroplasts Carmen Hostettler, Katharina Kölling, Diana Santelia, Sebastian Streb, Oliver Kötting, and Samuel C. Zeeman Abstract Starch is a primary product of photosynthesis in the chloroplasts of many higher plants. It plays an important role in the day-to-day carbohydrate metabolism of the leaf, and its biosynthesis and degradation represent major fluxes in plant metabolism. Starch serves as a transient reserve of carbohydrate which is used to support respiration, metabolism, and growth at night when there is no production of energy and reducing power through photosynthesis, and no net assimilation of carbon. The chapter includes techniques to measure starch amount and its rate of biosynthesis, to determine its structure and composition, and to monitor its turnover. These methods can be used to investigate transitory starch metabolism in Arabidopsis, where they can be applied in combination with genetics and systems-level approaches to yield new insight into the control of carbon allocation generally, and starch metabolism specifically. The methods can also be applied to the leaves of other plants with minimal modifications. Key words: Arabidopsis, Photosynthesis, Partitioning, Amylose, Amylopectin
1. Introduction Starch is the major storage carbohydrate in higher plants. It is composed of polymers of glucose (glucans) that adopt an insoluble, semicrystalline structure, resulting in massive, osmotically inert granules that can reach up to 200 mm in diameter in the storage tissues of some species. The major glucan in starch is amylopectin, accounting for 70–90% of the granule weight. Amylopectin molecules contain between 100,000 and 1,000,000 glucosyl units, which are linked by a-1,4-bonds to form linear chains with a degree of polymerization from 6 to over 100. These chains are linked to each other by a-1,6-bonds, resulting in a tree-like, or ‘racemose’ structure. Approximately 4–5% of the bonds are branch points. Amylopectin is responsible for the semi-crystalline nature of starch. R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_21, © Springer Science+Business Media, LLC 2011
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In contrast, amylose is a smaller molecule (typically around 1,000 glucosyl units) and contains far fewer or no branch points. Amylose is synthesized within the matrix formed by amylopectin. Further information on starch structure can be found elsewhere (1). Starch biosynthesis is typically associated with long-term storage of carbon in specialized tissues such as tubers or the starchy endosperm of seeds. In these cells, starch is made in specialized plastids – the amyloplasts – using carbon imported from the leaves. However, most plants also accumulate starch in their chloroplasts as a direct product of photosynthetic carbon assimilation. The rate of starch synthesis in the leaves can be very high (comparable to that in a developing sink organ), but the period of starch accumulation is limited to the day and is generally followed by a period of net starch degradation at night. Thus, leaves generally contain much less starch on a gram-for-gram basis than storage organs. When considering chloroplast function, photosynthetic carbon assimilation, and the allocation of resources into biomass, it is important to consider the role of the starch pool. The amount of starch made and the rate at which it is remobilized are carefully controlled processes that respond to changes in environmental conditions (1). In Arabidopsis, there is a tight inverse correlation between the complete utilization of stored starch and the rate of plant growth (2). Not surprisingly, plants deficient in the ability to make starch or to mobilize it effectively show severe growth retardation when cultivated in a diurnal cycle (3, 4). In continuous illumination, this growth retardation is usually not observed, illustrating the primary requirement of starch to support metabolism at night (3). In this chapter, we present methods to visualize starch by iodine staining, and quantify it by enzymatic hydrolysis to glucose (see Subheadings 3.1 and 3.2). In Subheading 3.3, we present a radiological method for the quantification of starch biosynthesis rate, and the investigation of starch turnover. This can help place the simple quantification of starch into an appropriate context. For example, an increase in starch content in chloroplasts in a given set of experimental conditions, or in a given plant accession, could reflect an increase in biosynthesis, a decrease in degradation, or a combination of both factors. Furthermore, these effects may be integrated over time. Our earlier studies of the sex4 mutant (for starch excess 4) illustrate this (5). The sex4 mutant is deficient in a phosphoglucan phosphatase, an enzyme necessary for starch degradation, without which the degradation rate is reduced (4, 6). Newly emerged leaves have a starch content that is similar to or slightly above that of the wild type. After repeated diurnal cycles in which more starch is synthesized during the day than is degraded at night, mature leaves have accumulated much higher levels of starch than the equivalent wild-type leaves, where synthesis and degradation are balanced (5). Paradoxically, however, the rate of starch biosynthesis in the mutant is also lower than that in the wild
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type, due to a compensatory shift in assimilate partitioning away from starch biosynthesis (4, 5). This was revealed both by a timecourse analysis of starch amount and by the analysis of the percentage of photo-assimilated 14CO2 partitioned into starch. It is also possible that the rate of starch biosynthesis is not equal to the rate of starch accumulation if there is simultaneous synthesis and degradation. This can be revealed by ‘pulse-chase’ experiments where first, the amount of 14C label incorporated into starch via photosynthesis in 14CO2 in the light is determined (the pulse). Then, the amount of 14C remaining in starch during a chase period in 12CO2 (also in the light) can be monitored. A drop in the amount of 14C in starch indicates that starch made during the pulse is being degraded, despite the continuation of photosynthesis (7). Starch composition and structure are key determinants of starch functionality, which is important in numerous ways in the food and nonfood industries. Therefore, we also present methods for the extraction of starch (see Subheading 3.4) and investigation of three key aspects of starch structure: the amylose to amylopectin ratio (see Subheading 3.5), the chain length distribution of amylopectin (see Subheading 3.6), and the amount of phosphate covalently bound to starch (see Subheading 3.7). The measurement of the amylose to amylopectin ratio depends on the different iodinebinding characteristics of the two polymers. Amylopectin binds less iodine than amylose, and the glucan–iodine complexes have different absorption spectra. With appropriate standard curves, the proportion of amylose can be easily calculated after simple spectrophotometric measurements. The fine structure of amylopectin is a complex trait determined by the co-operative activities of at least three classes of enzymes. The most frequently used and informative method for investigation of amylopectin structure is the analysis of the distribution of chain lengths. This is achieved by the selective enzymatic hydrolysis of a-1,6-branch points, followed by the separation and detection of the liberated a-1,4-linked chains. The method we present uses high performance anionexchange chromatography coupled with pulsed amperometric detection (HPAEC-PAD). Amylopectin, but not amylose, has covalently linked phosphate groups. The presence of phosphate on amylopectin enhances its functionality, and extracted starches are often treated to increase the number of charged groups. Phosphate groups are predominantly located at the C6 position of glucosyl residues, with a smaller fraction at the C3 position. Starches from different botanical sources have quite different amounts of phosphate, with potato tuber starch having high levels (1 in every 200– 300 glucosyl residues), Arabidopsis leaf starch having intermediate levels (1 in every 2,000 glucosyl residues), and cereal endosperm starches have very low phosphate (1 in every 10,000 glucosyl residues or less). The phosphate plays an important biological function; a cycle of glucan phosphorylation and dephosphorylation is an
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intimate part of the starch degradation process in chloroplasts. Phosphorylation is mediated by enzymes of the glucan, water dikinase class (GWD), and serves to disrupt the crystalline packing of amylopectin, rendering the glucan chains susceptible to hydrolysis and subsequent dephosphorylation (by SEX4, see above). Differences in the phosphate content of starch in chloroplasts could, therefore, reflect a change in this glucan phosphorylation cycle (see Note 1).
2. Materials 2.1. Visualizing Starch Using Iodine Staining
1. Lugol (I2–KI) solution: 0.34% (w/v) I2 and 0.68% (w/v) KI (Sigma–Aldrich, Buchs, Switzerland). 2. 80% (v/v) ethanol. 3. Orbital shaker. 4. Light table and/or optical microscope.
2.2. Quantitative Starch Measurement
1. 1.12 M perchloric acid. 2. 80% (v/v) ethanol. 3. Digestion mix: 200 mM sodium acetate–acetic acid, pH 4.8, a-amyloglucosidase (6.3 U/mL) EC 3.2.1.3 from Aspergillus nidulans (Roche, Rotkreuz, Switzerland), and a-amylase (50 U/mL) E.C. 3.2.1.1 from Bacillus subtilis (Roche). 4. 200 mM sodium acetate–acetic acid, pH 4.8. 5. Glucose assay mix: 100 mM 2-(4-[2-hydroxyethyl]-1piperazinyl)-ethanesulfonic acid-KOH (HEPES–KOH), pH 7.5, 1 mM MgCl2, 1 mM adenosine triphosphate (ATP), 1 mM nicotinamide adenine dinucleotide (NAD+), and hexokinase (6 U/mL) E.C. 2.7.1.1 from Saccharomyces cerevisiae (Roche). 6. Glucose 6-phosphate dehydrogenase E.C. 1.1.1.49 from Leuconostoc mesenteroide (Roche). 7. Liquid nitrogen. 8. All-glass homogenizer (e.g., Sartorius Potter S homogenizer with 2- or 5-mL homogenizer vessel and ground-in glass plunger). 9. Bench-top centrifuge with swing-out rotor suitable for 10-mL tubes. 10. Water baths/heating blocks set to 37 and 95°C. 11. Spectrophotometer or microtiter plate reader for absorbance measurements at 340 nm. 12. Plastic semi-micro cuvettes (1.5-mL, 10-mm path length) for a spectrophotometer.
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1. Sodium 14C-bicarbonate, 50 mCi/mmol (Hartmann Analytic, Braunschweig, Germany). 2. 50% (w/v) lactic acid. 3. 20% (v/v), 50% (v/v), and 80% (v/v) ethanol. 4. Tissue solubilizer (NCS II, GE Healthcare, Glattbrugg, Switzerland). 5. Digestion mix: 200 mM sodium acetate–acetic acid, pH 4.8, a-amyloglucosidase (6.3 U/mL) EC 3.2.1.3 from A. nidulans (Roche), and a-amylase (50 U/mL) E.C. 3.2.1.1 from B. subtilis (Roche). 6. Two-part plexiglas labeling chamber sealed with rubber foam as shown in Fig. 1 (custom-made, ETH Zurich, Switzerland). 7. Light source providing uniform white light (150 mmol photons/m2/s) to the labeling chamber. 8. Adjustable flat-topped laboratory stand (e.g., Laborboy; Bochem Instrumente, Weilburg, Germany). 9. Rubber stopper with turn-up lip for the Plexiglas chamber injection port (e.g., Subaseal, Sigma–Aldrich). 10. Syringe (5 mL) with long needle. 11. Small glass Petri dish (~5-cm diameter). 12. Water baths/heating blocks set to 80°C and 95°C. 13. All-glass homogenizer (see Subheading 2.2).
Fig. 1. Custom-made chamber system for labeling Arabidopsis rosettes. (a) Photograph of assembled chamber. (b) Schematic presentation of the chamber system. (1) Upper part of the plexiglas chamber; (2) rubber foam for sealing; (3) lower part of the plexiglas chamber; (4) height-adjustable laboratory table; (5) glass Petri dish with sodium 14C-bicarbonate; and (6) plants for labeling.
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14. Bench-top centrifuge with swing-out rotor suitable for 6.5-mL vials. 15. Rotary evaporator (e.g., Büchi Rotavapor® R-210) with vacuum supply. 16. Thermomixer (Eppendorf, Hamburg, Germany), set to 20°C and to shake at 1,400 rpm. 17. Scintillation counter, fluid, and vials. 2.4. Leaf Starch Granule Preparation
1. Starch extraction medium: 20 mM HEPES–KOH, pH 8.0, 0.2 mM ethylenediaminetetraacetic acid (EDTA), and 0.5% (v/v) Triton X-100. 2. Percoll (Sigma–Aldrich). 3. 500 mM HEPES–KOH, pH 7.0. 4. 0.5% (w/v) sodium dodecyl sulphate (SDS). 5. 50 mM HEPES–KOH, pH 7.2. 6. Liquid nitrogen. 7. Mortar and pestle. 8. Waring blender. 9. Nylon nets with 100-, 60-, 30-, and 20-mm mesh widths (Sefar Inc. Mesh+ Technology, Heiden, Switzerland). 10. Wide-stem (~25 mm) funnels. 11. 30-mL Corex centrifuge tubes. 12. Bench-top centrifuge with swing-out rotor and rubber sleeves suitable for 30-mL Corex tubes. 13. Vertical rotary wheel mixer. 14. SpeedVac sample concentrator system.
2.5. Determination of the Amylose: Amylopectin Ratio
1. Fresh Lugol (I2–KI) solution (see Subheading 2.1). 2. Water bath/heating block set to 95°C. 3. 1-mL quartz cuvettes (10-mm path length). 4. Spectrophotometer for absorbance measurements at 525–700 nm.
2.6. Chain Length Distribution
1. Debranching mixture: 50 mM sodium acetate–acetic acid, pH 4.8 (filter sterilized), isoamylase (40 U/mL) from Pseudomonas amyloderamosa (Sigma–Aldrich), and pullulanase (40 U/mL) from Klebsiella planticola (Megazyme, Bray, Ireland). 2. 1 M sodium acetate. 3. 2 M HCl. 4. Water baths/heating blocks set to 37°C and 95°C. 5. Thermomixers set to 95°C and 37°C, shaking at 400 rpm.
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6. Dowex 50 × 4 – 200-mesh resin and Dowex 1 × 8 – 200- to 400-mesh resin (Sigma–Aldrich). 7. 2-mL syringe barrels. 8. Miracloth tissue. 9. Freeze dryer (e.g., Martin Christ Freeze dryer ALPHA 1-2/ LDplus). 10. HPAEC-PAD system with a Dionex CarboPac PA200 column. The method uses NaOH as the eluant, so the HPLC system needs to be constructed from inert materials (e.g., Dionex ICS 5000 or earlier models, with PEEK plastic fittings). Sugars are quantified using a gold electrode performing pulsed amperometric detection. 2.7. Quantification of the Covalently Bound Phosphate of Starch
1. 1 M HCl. 2. 1 M NaOH. 3. Antarctic phosphatase (5 U/mL) and 10 times Antarctic phosphatase reaction buffer (New England Biolabs, Bioconcept, Allschwil, Switzerland). 4. Phosphate solution: 100 mM NaH2PO4. 5. 3.75 and 1 M NaCl solutions. 6. Malachite green solution: 1.5 mM Malachite green hydrochloride and 0.35% (w/v) polyvinyl alcohol (Mowiol 40–88, Sigma–Aldrich). Stir preferably overnight to dissolve the polyvinyl alcohol completely and then add the malachite green afterward. 7. Molybdate solution: 28 mM ammonium heptamolybdate tetrahydrate and 2.1 M sulfuric acid. 8. Vertical rotary wheel mixer. 9. Water baths/heating blocks set to 37 and 95°C. 10. Plastic semi-micro cuvettes, suitable for 600 mL volume (10mm path length) for a spectrophotometer. 11. Spectrophotometer for absorbance measurements at 620 nm.
3. Methods The following protocols are optimized for Arabidopsis rosettes. For other tissues or plant species, the protocols may need to be adapted according the tissue type, tissue weight, reaction volumes, etc. 3.1. Visualizing Starch Using Iodine Staining
Starch can be visualized by iodine staining (using Lugol solution). Iodine is incorporated into the secondary structures formed by glucans and thus staining is highly dependent on their molecular
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architecture. The staining is dark-blue for amylose, red-brownish for amylopectin, or pale orange-red for soluble polyglucans such as glycogen. Iodine staining can be used for qualitative screens, for microscopy, and for visualizing starch distribution in different cell types. 1. Harvest plant material (single leaves, rosettes, etc.) into a suitably sized tube and cover with 80% (v/v) ethanol. 2. Incubate for 3 min at 80°C in a water bath. Discard the supernatant. 3. Repeat step 2 until the plant material is free of chlorophyll and pigments. 4. Add Lugol solution and shake for 5 min on an orbital shaker (see Note 2). 5. Discard Lugol solution and wash twice for 2 min with water. 6. Observe the stained sample on a light table or, for cellular resolution, with an optical microscope. 3.2. Quantitative Starch Measurement
1. Harvest 100–300 mg plant material (record the exact weight). Transfer the samples immediately into 6.5-mL vials and freeze in liquid nitrogen. Store at −80°C. 2. Add the frozen tissue to an ice-cold all-glass homogenizer. Add 3 mL of ice-cold 1.12 M perchloric acid. Grind the plant material until it is completely and uniformly homogenized (see Note 3). 3. Transfer a defined volume (e.g., 2.7 mL) of the homogenate to a round-bottomed 10-mL centrifuge tube and spin down the insoluble material, including the starch (10 min, 3,000 × g, swing-out rotor, 4°C). 4. Discard the supernatant and resuspend the pellet (see Note 4) by vortexing in 4 mL sterile water to wash away contaminating soluble compounds. Spin down again (10 min, 3,000 × g, swing-out rotor, 4°C) and discard the supernatant. 5. Wash the pellet in 10 mL of 80% (v/v) ethanol to remove remaining sugars and pigments which could interfere with the subsequent analytical steps. Repeat three times, or until the pellet is whitish or light brown (8). Between each wash, spin down the insoluble material (10 min, 3,000 × g, swing-out rotor, 4°C) and discard the supernatant. 6. After the last wash step, carefully invert the reaction tubes and let the pellet air-dry on the bench (see Note 5). 7. Resuspend the pellet in 0.5 mL of sterile water using a widebore pipette tip and transfer the suspension to a 1.5-mL screwcapped microcentrifuge tube. Fill up with sterile water to give a final, defined volume (e.g., 1 mL; record the final volume for
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later calculations). The resuspended starch pellet can be stored at −20°C (see Note 6). To measure the starch in the insoluble fraction obtained as above, a glucose assay is performed after hydrolyzing the starch to glucose using a-amyloglucosidase and a-amylase (see Note 7). Glucose is determined via an enzyme-coupled assay. Hexokinase uses ATP to phosphorylate glucose to glucose 6-phosphate (Glc6P). Subsequently, glucose 6-phosphate dehydrogenase (G6PDH) converts Glc6P to 6-phophogluconate and simultaneously reduces an equimolar amount of NAD+ to NADH. NADH but not NAD+ absorbs light with a wavelength of 340 nm. Hence, NADH production can be monitored spectrophotometrically. 8. After resuspending the pellet, transfer 500 mL to a 1.5-mL screw-capped microcentrifuge tube (see Note 4). 9. Heat the samples in a water bath to 95°C for 15 min (see Note 8). 10. Cool the samples to room temperature and spin briefly in a microcentrifuge to remove condensed water and residual pellet particles from the tube cap. Mix well to obtain a homogenous suspension and pipette 200 mL to each of two 1.5-mL microcentrifuge tubes. To the first, add 200 mL of digestion mix. To the second (a control), add 200 mL of 200 mM sodium acetate–acetic acid, pH 4.8. 11. Incubate the samples for 4–6 h at 37°C in a water bath (see Note 9). 12. Spin samples in a microcentrifuge (10 min, 14,000 × g, 20°C) to remove undigested material. The supernatant contains the glucose released from starch. 13. Mix between 20 and 200 mL of the digested samples (record the volume used) with 250 mL of glucose assay mix in a 1.5mL semi-micro cuvette. Make up to a final volume of 1 mL with sterile water. After recording the absorption at 340 nm (OD340), add a minimal volume (e.g., 1 mL) containing 0.25 U G6PDH onto a plastic microspatula and mix rapidly with the cuvette contents without removal from the spectrophotometer. Record the OD340 again after a constant value is reached (this should take no more than 10 min; see Notes 10 and 11) and calculate the difference (DOD340). The mmol glucose in the cuvette is calculated as follows:
∆OD340 = A, 6.22 where 6.22 is the extinction coefficient of NADH at 340 nm (see Note 12). To obtain the net value of A (Ae), subtract the value of A for control samples from the value of A for digested samples.
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Calculate the mmol of glucose in starch in the original plant sample as follows: Ae ×
×
Vol. of starch digest Vol. of insoluble fraction × Vol. added to glucose assay Vol. added to digest Vol. of total homogenate 1 × . Vol. of analyzed homogenate Fresh weight
If you used the exact volumes suggested in this protocol, then the calculation is as follows (see Note 13): Ae ×
×
400 µL 1, 000 µL × Value between 20 and 200 µL 200 µL 3000 µL + (0.9 × Fresh weight [mg]) 1 × . 2, 700 µL Fresh weight (g)
To express the value as mg starch per gram of fresh weight, multiply by 162 (the mass of a glucosyl residue in starch). Alternatively, multiply by 180 to give mg ‘glucose equivalents’ per gram of fresh weight. 3.3. Investigation of Starch Biosynthesis Rate and Starch Turnover
We recommend using screw-cap tubes when working with radioactively labeled material, as the risk of contamination is reduced. The labeling chamber is assembled as shown in Fig. 1. 1. Place the plants in the labeling chamber under the light source within a fume cupboard and adjust the table and/or lights to give the desired light intensity (see Note 14). 2. Place the small glass Petri dish containing sodium 14 C-bicarbonate with a specific activity of 50 mCi/mmol in the holder and seal the labeling chamber. 3. Using a syringe, inject an excess of lactic acid through the rubber stopper into the glass Petri dish to release 14CO2 from sodium 14C-bicarbonate. 4. Start the fan to distribute the 14CO2 evenly within the chamber. 5. Leave the plants in the labeling chamber for a pulse period of up to 1 h (see Note 14). 6. After the pulse period, open the chamber, remove the plants, and harvest the ‘pulse’ samples, while replacing the ‘chase’ samples back under the lights for the desired chase periods (see Note 15). 7. Prepare a 50-mL centrifuge tube containing 20 mL of 80% (v/v) ethanol for each plant sample and heat to 80°C. 8. Harvest the fresh plant material after the pulse and at different time points within the chase period. Quench the tissue in the preheated 80% (v/v) ethanol.
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9. Close the tubes and incubate for 10 min at 80°C in a water bath, after which the samples can be cooled and stored at 4°C until subsequent steps. 10. Take out the plant tissue and homogenize it in an all-glass homogenizer with 1 mL of 80% (v/v) ethanol. Keep the 20 mL 80% (v/v) ethanol as the soluble fraction. 11. Collect the homogenized material in a 6.5-mL vial. Rinse the all-glass homogenizer twice with 1 mL of 80% (v/v) ethanol, add it to the 6.5-mL vial, and spin down the insoluble material (12 min, 2,400 × g, swing-out rotor, 20°C). Add the supernatant to the soluble fraction. 12. Wash the pellet in 5 mL of 50% (v/v) ethanol and spin down as before. Repeat washing with 5 mL of 20% (v/v) ethanol, with 5 mL of sterile water, and finally with 5 mL of 80% (v/v) ethanol, each time adding the supernatant to the soluble fraction. 13. Dry down the soluble fraction (~48 mL) by rotary evaporation under a vacuum to approximately 1 mL. 14. Add sterile water and record the final volume (e.g., 2 mL; see Note 16). 15. Dry the pellet (step 12) briefly at room temperature to evaporate remaining ethanol and resuspend in 2 mL of sterile water. This is defined as the insoluble fraction. Both fractions can be stored at −20°C. 16. Incubate three aliquots of 100 mL insoluble fraction in 900 mL of tissue solubilizer at 20°C overnight in a Thermomixer, shaking at 1,400 rpm. 17. Determine the radioactivity in the incubated insoluble fractions and, additionally, in three 200-mL aliquots of the soluble fraction by liquid scintillation counting. Transfer both fractions to a 6.5-mL scintillation vial and add up to 5 mL of scintillation cocktail. Vortex the vials and incubate in the dark for 30 min before scintillation counting. These data give an overview of the partitioning into the soluble and insoluble compounds (see Note 17). 18. To determine the radioactive label in starch, heat three 500-mL aliquots of the insoluble fraction for 15 min at 95°C in a water bath (see Note 8). 19. Let the samples cool to room temperature and spin briefly to remove condensed water and residual pellet particles from the tube cap. Add 500 mL of digestion mix and vortex briefly. 20. Incubate for 6 h at 37°C in a water bath. 21. Spin down the samples in a microcentrifuge (5 min, 12,000 × g, 20°C) and transfer each supernatant, containing glucose units
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released from starch, into a 6.5-mL scintillation vial (see Note 18). Wash the pellets once with 200 mL of sterile water, spin down again (5 min, 12,000 × g, 20°C), and add the supernatants to the corresponding scintillation vials. 22. Determine the radioactivity in the supernatant by liquid scintillation counting as in step 17. 23. Obtain the mean radioactivity (disintegration per minute) in the three replicates per fraction and calculate the total label in each fraction, taking into account that the label in the different fractions was determined only in aliquots of the fraction. 24. Express the label in starch as a proportion of the total label in the plant by dividing the amount of label in starch by the overall label in the insoluble and soluble fractions (step 17 and see Note 19). 3.4. Leaf Starch Granule Preparation
For some analytical techniques, such as the determination of amylose:amylopectin ratio (see Subheading 3.5), or starch phosphate content (see Subheading 3.7), it can be an advantage to work with purified starch granules. Inevitably some starch is lost during purification, so the following method is not suitable for determining total starch content. 1. Harvest whole rosettes of 4- to 5-week-old Arabidopsis plants (20–50 g) at the end of the light period (see Note 20), freeze them in liquid nitrogen, and store at −80°C. 2. Precool a mortar and pestle with liquid nitrogen and grind the frozen leaves to a fine powder. 3. Mix the pulverized plant material with 20–50 mL of ice-cold starch extraction medium and homogenize for 20–40 s using a Waring blender (see Note 21). 4. Filter the homogenate through a 100-mm nylon mesh (see Note 22). 5. Transfer the resulting filtrate to 30-mL Corex centrifuge tubes and spin down (5 min, 1,500 × g, swing-out rotor, 20°C). A whitish starch pellet should be clearly visible. 6. Discard the supernatant and resuspend the starch pellet in 10 mL of starch extraction medium. 7. Filter the suspension sequentially through nylon meshes with 100-, 30-, and 20-mm pore sizes. 8. Overlay the filtrate on a 10-mL cushion of 95% (v/v) Percoll and 5% (v/v) 500 mM HEPES–KOH, pH 7 (see Note 23), in a 30-mL Corex centrifuge tube and spin down the starch (15 min, 2,000 × g, swing-out rotor, 20°C). 9. Carefully remove the supernatant (see Note 24). Resuspend the pellet in 1 mL of 0.5% (w/v) SDS and transfer to a 2-mL
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microcentrifuge tube. Spin down the starch in a microcentrifuge (1 min, 14,000 × g, 20°C). 10. Wash the resulting starch pellet twice more with 1–2 mL of 0.5% (w/v) SDS (see Note 25) and then spin down the suspended starch granules (1 min, 14,000 × g, 20°C). Discard the supernatants (see Note 26). During the last washing step, mix the starch granule suspension for 10–15 min on a vertical rotary wheel mixer (15 rpm). 11. Wash the starch pellet four times with 50 mM HEPES–KOH, pH 7.2, and once with sterile water to remove SDS from the samples. 12. Transfer the starch to pre-weighed 1.5-mL microcentrifuge tubes and dry the pellet in a SpeedVac concentrator system. Continue until the tube weights remain constant. The difference between final weight and the initial tube weight is equal to the amount of extracted starch. 13. Use starch directly or store it at −20°C. 3.5. Determination of the Amylose: Amylopectin Ratio
This method can be used for the determination of the starch composition. It is based on the observation that amylose and amylopectin have different iodine-binding capacities. After staining with Lugol solution, amylose and amylopectin show different absorption spectra, depending on the amount of iodine bound and the type of complex formed between iodine and glucan secondary structures. When complexed with iodine, amylopectin has a wavelength of maximal absorbance (lmax) of 550 nm, whereas amylose has a lmax of 620 nm (9). Standard curves of the absorbance of the iodine–polymer complexes for purified amylose and amylopectin from Arabidopsis are shown in Fig. 2. These were used to generate specific absorbance coefficients such that the amylose:amylopectin ratios of starch samples can be calculated from absorbencies at two wavelengths using the following formula: Percentage amylose =
3.039 − (7.154 × OD700 / OD525 ) . (3.048 × OD700 / OD525 ) − 19.192
The wavelengths 700 and 525 nm are not the lmax values, but are used to give the maximal sensitivity (Fig. 2). The procedure for Arabidopsis starch may be used for starches from other species as well, provided that amylose and amylopectin from such starches have the same iodine-binding capacities. However, it will always be preferable to establish new standard curves for the starch in question. Furthermore, iodine staining may be affected if starch structure as well as composition is altered (see Subheading 3.6). In such cases, this method is unsuitable for comparisons between starch types.
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1. Resuspend purified starch granules in sterile water to a concentration of 5 mg/mL (see Note 27). 2. Incubate the samples for 15 min at 95°C in a water bath (see Note 8). 3. Add 5–20 mL of gelatinized starch to one of two 1-mL quartz cuvettes (see Note 28) and an equal volume of water to the second (as a control). Add freshly prepared 10% (v/v) Lugol solution to give a final volume of 1 mL. Mix by inverting gently a few times. 4. Measure the absorbance immediately after mixing in a spectrophotometer equipped with a program for measuring at 700 and 525 nm. Correct the values for the control cuvette and calculate the ratio of the absorbencies (OD700/OD525). 5. Determine the amylose percentage by applying the formula above (see Note 29). 3.6. Analysis of the Chain Length Distribution of Starch
The chain length distribution (CLD) of amylopectin or related glucose polymers yields valuable structural information. It is the most commonly used method to give insight into the molecular architecture of the polymers. Alterations in CLD between genotypes (e.g., between wild-type and mutant plants) are frequently presented as ‘difference plots.’ However, it is important to realize that the method delivers an ‘average’ result. It cannot yield information about heterogeneity in chain lengths within polymer molecules, or distinguish between polymer types that might accumulate in a single tissue.
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The method also does not yield information about the positioning of branch points. Thus, even if there are no differences in CLD, it is still possible (albeit unlikely) that the polymers differ (e.g., in branch point distribution). We recommend starting with at least 100 mg of starch, either in the insoluble fraction of a perchloric acid extract (see Subheading 3.2) or as pure starch (see Subheading 3.4). This method is also suitable to analyze the architecture of similar amounts of other polymers (glycogen/phytoglycogen). Larger amounts can also be used, with corresponding increases in the reaction volumes (see Note 30). 1. Resuspend your sample, containing the starch, vigorously by vortexing to obtain a homogenous suspension (see Note 4) and transfer the required volume to obtain 100 mg immediately to a 1.5-mL screw-capped microcentrifuge tube using wide-bore pipette tips. Make up to a final volume of 100 mL with sterile water (see Note 6). 2. Heat the sample for 15 min at 95°C. Use either a thermomixer set at 400 rpm or a water bath (in the latter case, vortex once after 5 min) (see Note 8). 3. Cool the samples to 20°C and spin briefly to remove the condensed water and residual insoluble material from the tube cap. 4. Add 25 mL of debranching mixture to each sample to hydrolyze the a-1,6-linkages specifically. Mix well by vortexing. 5. Incubate for 3 h at 37°C in a thermomixer set at 400 rpm or in a water bath (in the latter case, vortex briefly every hour). Stop the reaction by heating the sample for 10 min at 95°C. 6. Spin down the cell debris and denatured enzymes in a microcentrifuge (10 min, 10,000 × g, 20°C). The samples can be stored at this stage at −20°C. If necessary, clean-up the samples by using Dowex 1 and Dowex 50 ion-exchange columns (see Note 31). 7. Weigh out each Dowex resin (1 g per sample) into separate glass beakers (such that the resin occupies no more than 20% of the volume). Wash the resins in an excess of sterile water. Let the resin settle and pour off the water. Repeat until the supernatant is colorless. Charge the Dowex 1 and Dowex 50 resins with 1 M sodium acetate and 2 M HCl, respectively (use twice the resin volume). Stir for 30 min and let the resin settle. Remove the supernatant without losing the resin. Wash the resin with excess sterile water four times or until the pH of the supernatant is neutral. 8. Prepare the Dowex columns. We use the barrels of 2-mL syringes (see Note 32). Cut circles of Miracloth tissue to fit the outflow of the syringe and rack them in vertical orientation.
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Fig. 3. Sample preparation with self-made Dowex columns. 2-mL syringes are fitted with Miracloth to retain Dowex 50 (orange, top) and Dowex 1 (white, bottom) resins. Dowex 50 columns are racked on top of the Dowex 1 columns. The Dowex 1 column outlet fits into a 6.5-mL collection vial (the stacking plexiglas support is made in-house).
9. Resuspend the charged resin in an equal volume of sterile water, giving a slurry. Mix the resin on a magnetic stirrer while pipetting it into the columns. After the water has drained from the column, the resin bed volume should be 1 mL. Wash the columns twice with 1 mL of sterile water. Avoid disturbing the resin and do not let the columns dry out. 10. Stack the Dowex 50 columns above the Dowex 1 columns in a suitable rack (see Fig. 3), so that the flow drops directly from one column to the next. Place a 6.5-mL vial below the columns. 11. Apply 100 mL of the supernatant of the centrifuged samples from step 6 to the Dowex 50 column. Let the sample run into the resin. 12. Add 500 mL of sterile water and let it run into the resin. Repeat this step every 5 min nine times to give a final elution volume of 5 mL (if a different size of column is used, the final elution volume should be 5 times the bed volume of the column). The glucan chains will not bind to the resins and are collected in the outflow. Freeze the samples in liquid nitrogen and freeze dry. 13. Add 150 mL sterile water to the freeze dried samples and dissolve well by vortexing. Transfer the dissolved samples to a
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new 1.5-mL microcentrifuge tube and spin down any resin remnants in a microcentrifuge (5 min, 10,000 × g, 4°C). 14. Transfer 100 mL of the supernatant carefully to a vial suitable for the autosampler of the HPAEC-PAD system. We normally inject 10 mL for analysis (see Note 33). 15. For HPAEC-PAD, we use an ICS 5000 HPLC (Dionex) system equipped with an electrochemical detector and gold electrode. We recommend using a CarboPac® PA200 column (Dionex) which is designed for separation of neutral oligosaccharides. Separation is achieved at a high pH, such that the hydroxyl groups are ionized. Longer glucans bind more tightly to the column. The chains are eluted with a NaOH/sodium acetate gradient. Standard program: Eluent A (see Note 34): (100 mM NaOH) Eluent B (see Note 34): (150 mM NaOH and 500 mM Na-acetate) Starting conditions: 95% Eluent A and 5% Eluent B Gradient 0–13 min: linear to 60% Eluent A and 40% Eluent B Gradient 13–50 min: linear to 15% Eluent A and 85% Eluent B Re-equilibrating 50–70 min: 95% Eluent A and 5% Eluent B Individual peak areas are determined and summed. Each peak area is then expressed as a percentage of the total area of all peaks (Fig. 4) and plotted accordingly. Subtracting one CLD from another, results in a difference plot to visualize easily the differences in the relative amounts of each chain length (Fig. 4). 3.7. Quantification of the Covalently Bound Phosphate of Starch
This method for measuring starch-bound phosphate levels consists of three steps. First, starch is hydrolyzed to glucose and glucose phosphates. Second, orthophosphate is released from glucose phosphates by treatment with a phosphatase. Third, malachite green binds orthophosphate in the presence of molybdate solution, giving rise to a complex that can be measured in a colorimetric assay (10). 1. Weigh out at least 15 mg purified starch granules (see Subheading 3.4) in a clean 1.5-mL microcentrifuge tube (see Note 35). Note the exact weight. 2. Resuspend the starch in 0.5 mL of sterile water and incubate on a vertical rotary wheel mixer (5 min, 20°C, 25 rpm) to wash and rehydrate the starch. Spin down the starch (5 min, 10,000 × g, 20°C). 3. Repeat step 2 twice. 4. Resuspend the pellet in sterile water to a final concentration of 25 mg starch per mL.
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Fig. 4. Analysis of the chain length distribution of amylopectin. (a) The racemose model of amylopectin. Circles represent glucose molecules linked via a-1,4-linkages (closed circles) or a-1,6-linkages (open circles). (b) Amylopectin treated with a debranching mixture containing isoamylase and pullulanase to hydrolyze the a-1,6-linkages, generates linear chains. (c) Typical CLD chromatograms obtained by HPAEC-PAD of glucans from wild-type Arabidopsis (WT, amylopectin) and from a double mutant (isa1/2, phytoglycogen) with altered starch structure (8). (d) CLD calculated from the chromatograms. Peak areas of chains containing between 2 and 40 glucose units are summed and the relative percentage of the total area plotted against the chain length. (e) Difference plots for better visualization of changes in CLD calculated by subtracting the relative percentage of the wild-type from the isa1/2 mutant.
5. Transfer 200 mL (i.e., 5 mg starch) into each of three 1.5-mL screw-capped tubes (two replicates and one control). Spin down the starch (5 min, 10,000 × g, 20°C). 6. Resuspend each of the starch pellets in 100 mL of 1 M HCl. Tightly seal the tubes and incubate for 2 h at 95°C in a water bath to hydrolyze the starch (see Note 8). 7. Cool the samples to room temperature and neutralize with 100 mL of 1 M NaOH. Note the exact neutralization volume (see Note 36). 8. Phosphatase treatment: transfer 175 mL from each sample into clean 1.5-mL microcentrifuge tubes. Add 20 mL of 10 times Antarctic phosphatase reaction buffer to each, and 5 mL of Antarctic phosphatase to the two replicates. Add 5 mL of sterile
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water to the control. Incubate for 2 h at 37°C in a water bath (see Note 37). 9. Prepare a series of 10 orthophosphate standards (at least 100 mL of each) in the range of 0–200 mM (equivalent to 0–20 nmol in the assay) by diluting the phosphate solution with sterile water. 10. Transfer up to 150 mL of each phosphatase-treated sample or control into semi-micro cuvettes. 11. Transfer 100 mL of each phosphate standard into semi-micro cuvettes. Adjust to the same buffer and salt conditions as the starch samples by adding 35 mL of 3.75 M NaCl and 15 mL of 10 times Antarctic phosphatase reaction buffer to each phosphate standard. 12. Prepare the phosphate reagent by mixing 1 volume of malachite green solution and 1.34 volumes of molybdate solution (see Note 38). 13. Add 450 mL of the phosphate reagent to each semi-micro cuvette. Mix thoroughly and incubate at 20°C for 10–15 min. 14. Measure the absorbance in a spectrophotometer at 620 nm wavelength (see Note 39). 15. Plot the absorbance of the standards against the amount of phosphate. Determine the linear range (usually between 2 and 20 nmol phosphate) and calculate the slope of the standard curve. Calculate the phosphate content of the starch samples, taking into account dilutions made during the procedure (see Note 40).
4. Notes 1. The mechanism of starch breakdown in the endosperm differs from that in chloroplasts and amyloplasts. It probably does not require transient glucan phosphorylation (1). 2. Ethanol destroys the membranes and solubilizes pigments such as chlorophyll. The tissue needs to be thoroughly decolourized for effective iodine staining. The staining is reversible and the iodine complexed with the glucan structures is washed out over time. Work in a fume hood, as iodine vapor is toxic. 3. Perchloric acid inactivates proteins by denaturation and precipitation. Keep the samples ice cold and do not leave the extract in perchloric acid for an extended time, as this could cause acid hydrolysis of metabolites. 4. The insoluble pellet contains starch granules, cell debris, and denatured proteins. It is very important that the pellet is uniformly
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resuspended for washing and for further processing. Large pellet fragments impede accurate pipetting, leading to imprecise results. 5. It is important that excess ethanol is removed, since it could prevent the subsequent enzyme reactions. However, make sure that the pellet does not dry out completely as it becomes very difficult to resuspend uniformly (see Note 4). When inverting tubes, take care that material is not dislodged from the pellet. 6. The volume for resuspension should depend on the starch content – use larger volumes for starch-rich samples. The final resuspension volume should be determined as accurately as possible. When defrosting frozen samples, ensure that the sample is well mixed before removing aliquots. 7. Use a-amylase and a-amyloglucosidase free of contaminating enzymes that could degrade glucose-containing polymers other than starch (e.g., cellulose). We use enzymes from Roche (Rotkreuz, Switzerland). 8. This step gelatinizes (solubilizes) starch, making it more accessible for enzymatic degradation. Use tubes with a tightly sealing cap to prevent water loss during heating. Avoid cooling below room temperature as secondary structures can reform, causing precipitation (retrogradation) of gelatinized starch. 9. It is critical that starch is completely digested. You can check digestion efficiency by pipetting aliquots of the digest and control suspensions into microcentrifuge tubes containing Lugol solution. The digested sample should not stain for starch, whereas the control sample will. 10. For maximal accuracy, ensure that your assay yields a DOD340 between 0.2 and 1. For large-scale analyses, the glucose assay can also be performed in a 96-well format using a microtiter plate reader. In this case, we recommend a final reaction volume of 200 mL, four technical replicates per sample, and a calibration curve between 0 and 200 nmol glucose. Again, after monitoring the initial OD340 values, add 0.25 units G6PDH in a minimal volume (e.g., 1 mL) to each well and record the OD340 after a constant value is reached in all wells. Use a microtiter plate reader with an integrated shaking/mixing function. 11. If the kinetic curve of the reaction is slow or does not occur, check the assay for the following: (a) verify that the pH of the assay is 7–7.5; (b) confirm that the assay is working by adding a defined amount of glucose and see if there is the corresponding OD340 change; and (c) add more coupling enzymes or reaction substrates to see if one component is limiting the reaction. 12. If the glucose assay is performed in a 96-well format, determine the mmol glucose in each well by comparing the obtained sample values with the calibration curve for known amounts of glucose, rather than by using the NADH extinction coefficient.
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13. The amount of plant tissue affects the total homogenate volume. We designate the total volume as the volume of perchloric acid added plus 0.9 times the fresh weight of the sample. 14. We typically use 4- to 5-week-old Arabidopsis plants illuminated with a light intensity of 150 mmol photons/m2/s. We release 50 mCi 14CO2, which does not significantly alter the CO2 concentration in the 18-L chamber. The length of the pulse can be varied. For investigating starch turnover, we recommend pulse periods of between 10 and 60 min and chase periods between 60 and 180 min. We recommend five replicate samples per time point. 15. Avoid exposure to 14CO2. The chamber should be opened in a fume hood. Additional precautions can be taken; adding an excess of potassium hydroxide to the acidified sodium 14 C-bicarbonate prevents the release of residual 14CO2 and will absorb the CO2 in the chamber. The 14CO2 can also be pumped out through a 10% potassium hydroxide solution if suitable inlet and outlet ports are integrated in the chamber design. However, such steps may compromise the efficiency of the experiment. 16. Further separation into neutral, acidic, and basic fractions is possible by running the sample over an ion-exchange column (11). 17. Ensure your scintillation cocktail is suitable for measurements of both aqueous and nonaqueous samples. Ensure complete mixing of the sample and the scintillation cocktail. 18. The pellet can be further digested with protease to determine the amount of 14C in protein (12). 19. During the chase period, radioactive label is exported from the Arabidopsis shoot to the root system. It must be taken into account that after longer chase periods, a considerable part of the overall label assimilated by the leaves can be found in this fraction (5). 20. Arabidopsis leaves generally accumulate 10 mg starch per gram of fresh weight during a 12-h photoperiod (4). Depending on the amount of starch required for subsequent applications, calculate how much plant material is needed. Starch-excess mutants contain up to eight times more starch than the wild type (6, 13). However, be aware that 20% or more of the starch may be lost during the extraction and purification procedures. 21. The extraction medium:plant material ratio is generally 1:1 (v/w). However, it should be adjusted as necessary to obtain a fluid homogenate. 22. It is recommended to use funnels with a wide stem to facilitate the filtration of the homogenate. Fix the net at the bottom of the funnel stem securely with a rubber band. Help filtration by gently stirring the homogenate with a spatula to prevent cell debris clogging the mesh.
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23. Percoll consists of colloidal silica particles of 15–30 nm diameter (23% [w/w] in water) coated with polyvinylpyrrolidone (PVP). It is ideal for the isolation of cells, organelles, and other molecular components by density centrifugation. In this case, dense starch granules pass through the Percoll layer and sediment upon centrifugation, while cell debris and other impurities remain on top of the Percoll layer. 24. Remove the supernatant with a pipette to avoid dislodging the starch pellet. 25. SDS effectively removes proteins bound to the starch granule surface. 26. We recommend removing the residual impurities from the surface of the starch pellet manually using a pipette tip. Gently scratch the exposed surface to remove any brown residue from the white pellet. Some of the extracted starch will inevitably be lost during this step. 27. With this ratiometric method the concentration of solubilized starch is not critical for the analysis, as long as the absorbance values change linearly with the amount added. In our experience, the suggested concentrations yield absorbance values between 0.2 and 0.5, and are within the linear range. However, we recommend that the linear range is always confirmed by measuring different amounts of each sample. 28. It is possible to use plastic cuvettes, but then it is essential to change cuvettes for every measurement since they quickly stain red when exposed to iodine and this may affect the absorbance values. 29. The amylose content of wild-type Arabidopsis starch extracted from plants grown in a growth chamber is approximately 6–9%. Under other growth conditions and in starch-excess mutants, the amylose content can increase up to 33% (9), comparable to the amounts seen in storage starches from seeds and tubers. 30. The comparison between different samples is based on relative percentage differences of peaks (see Fig. 4). Thus, differences in the amount used lead to the same results if the sample is within the linear range of the detection limits of the HPAEC-PAD. 31. If working with purified starch, samples can be analyzed by HPAEC-PAD (or alternative detection methods, see Note 33) directly after filtering the samples through a 0.2-mm spin filter to remove larger particles and precipitated proteins. Go to step 14. 32. The column volume can be adapted depending on the volume of sample. Larger syringe barrels or empty purchased columns can be used. Alternatively, 1-mL pipette tips can be used for smaller volumes.
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33. An alternative method for separation and detection involves labeling the chains at the reducing end with a charged fluorophore. Chains are separated either via gel or capillary electrophoresis. Detection and quantification rely on established methods for DNA sequencing using the same equipment (14). This results in similar chromatograms as obtained by HPAEC-PAD. As each chain carries one label, the fluorescence intensity correlates with the number of chains. Subsequent data analyses are essentially the same as described in Fig. 4. 34. When preparing Eluents A and B, degass with helium for 10 min to displace dissolved CO2 prior to adding NaOH. This prevents the formation of Na2CO3 which interferes with HPAEC-PAD analysis. 35. The amount of phosphate covalently bound to starch varies considerably between species and tissues. Therefore, the amount of starch needed for the analysis may vary as well. We recommend including a reference starch with known phosphate content when first analyzing starches with unknown phosphate contents. 36. The amount of NaOH needed for neutralization might vary slightly. Always test neutralization with tubes containing just 100 mL of 1 M HCl, incubated at 95°C together with the samples, before neutralizing your samples. 37. In this step, Antarctic phosphatase dephosphorylates glucose phosphates to glucose and orthophosphate (Pi). It is possible to use pure glucose 6-phosphate standards as a control for both the acid hydrolysis step (ensuring no acid hydrolysis of glucose 6-phosphate) and the subsequent dephosphorylation step (ensuring complete enzymatic hydrolysis of glucose 6-phosphate). 38. Malachite green and molybdate solutions can be stored at 4°C for several weeks or even months. The phosphate reagent must be prepared freshly (use up the same day). It is crucial to use the same preparation of phosphate reagent for all samples, controls, and standards as the mixture is never exactly the same. 39. The measurement can alternatively be performed in a microtiter plate reader. However, the volumes will have to be adjusted accordingly. The absorption maximum of the complex is at 620 nm, but reliable measurements can be achieved in the range of 590–660 nm. 40. The sample volume needed varies as starches from different origins have different levels of starch-bound phosphate. Adjust the sample volume individually to a phosphate amount lying within the linear range of the standard curve.
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References 1. Zeeman, S. C., Kossmann, J., and Smith, A. M. (2010) Starch: its metabolism, evolution, and biotechnological modification in plants. Ann. Rev. Plant Biol. 61, 209–234. 2. Sulpice, R., Pyl, E. T., Ishihara, H., Trenkamp, S., Steinfath, M., Witucka-Wall, H., Gibon, Y., Usadel, B., Poree, F., Piques, M. C., Von Korff, M., Steinhauser, M. C., Keurentjes, J. J. B., Guenther, M., Hoehne, M., Selbig, J., Fernie, A. R., Altmann, T., and Stitt, M. (2009) Starch as a major integrator in the regulation of plant growth. Proc. Natl. Acad. Sci. USA 106, 10348–10353. 3. Caspar, T., Huber, S. C., and Somerville, C. (1985) Alterations in growth, photosynthesis, and respiration in a starchless mutant of Arabidopsis thaliana (L) deficient in chloroplast phosphoglucomutase activity. Plant Physiol. 79, 11–17. 4. Zeeman, S. C., Northrop, F., Smith, A. M., and ap Rees, T. (1998) A starch-accumulating mutant of Arabidopsis thaliana deficient in a chloroplastic starch-hydrolysing enzyme. Plant J. 15, 357–365. 5. Zeeman, S. C., and Ap Rees, T. (1999) Changes in carbohydrate metabolism and assimilate export in starch-excess mutants of Arabidopsis. Plant Cell Environ. 22, 1445–1453. 6. Koetting, O., Santelia, D., Edner, C., Eicke, S., Marthaler, T., Gentry, M. S., Comparot-Moss, S., Chen, J., Smith, A. M., Steup, M., Ritte, G., and Zeeman, S. C. (2009) STARCH-EXCESS4 is a laforin-like phosphoglucan phosphatase required for starch degradation in Arabidopsis thaliana. Plant Cell 21, 334–346. 7. Schneider, A., Hausler, R. E., Kolukisaoglu, U., Kunze, R., van der Graaff, E., Schwacke, R., Catoni, E., Desimone, M., and Flugge, U. I. (2002) An Arabidopsis thaliana knock-out mutant of the chloroplast triose phosphate/ phosphate translocator is severely compromised
only when starch synthesis, but not starch mobilisation is abolished. Plant J. 32, 685–699. 8. Delatte, T., Trevisan, M., Parker, M. L., and Zeeman, S. C. (2005) Arabidopsis mutants Atisa1 and Atisa2 have identical phenotypes and lack the same multimeric isoamylase, which influences the branch point distribution of amylopectin during starch synthesis. Plant J. 41, 815–830. 9. Zeeman, S. C., Tiessen, A., Pilling, E., Kato, K. L., Donald, A. M., and Smith, A. M. (2002) Starch synthesis in Arabidopsis. Granule synthesis, composition, and structure. Plant Physiol. 129, 516–529. 10. Cogan, E. B., Birrell, G. B., and Griffith, O. H. (1999) A robotics-based automated assay for inorganic and organic phosphates. Anal. Biochem. 271, 29–35. 11. Canvin, D. T., and Beevers, H. (1961) Sucrose synthesis from acetate in germinating castor bean - kinetics and pathway. J. Biol. Chem. 236, 988–995. 12. Runquist, M., and Kruger, N. J. (1999) Control of gluconeogenesis by isocitrate lyase in endosperm of germinating castor bean seedlings. Plant J. 19, 423–431. 13. Yu, T. S., Kofler, H., Hausler, R. E., Hille, D., Flugge, U. I., Zeeman, S. C., Smith, A. M., Kossmann, J., Lloyd, J., Ritte, G., Steup, M., Lue, W. L., Chen, J. C., and Weber, A. (2001) The Arabidopsis sex1 mutant is defective in the R1 protein, a general regulator of starch degradation in plants, and not in the chloroplast hexose transporter. Plant Cell 13, 1907–1918. 14. O’Shea, M. G., Samuel, M. S., Konik, C. M., and Morell, M. K. (1998) Fluorophore-assisted carbohydrate electrophoresis (FACE) of oligosaccharides: efficiency of labelling and high-resolution separation. Carbohydr. Res. 307, 1–12.
Chapter 22 Analysis of Lipid Content and Quality in Arabidopsis Plastids Anna Maria Zbierzak, Peter Dörmann, and Georg Hölzl Abstract Chloroplasts of plants contain an intricate membrane system, the thylakoids, which harbor the complexes of the photosynthetic machinery. Chloroplasts are confined by two membranes, the inner and outer envelope. The major glycerolipids of chloroplasts are the glycolipids monogalactosyl diacylglycerol (MGD), digalactosyl diacylglycerol (DGD), and sulfoquinovosyl diacylglycerol (SQD). Furthermore, two phospholipids, phosphatidyl glycerol (PG) and phosphatidyl choline (PC), are found in chloroplast membranes. The photosystems and light-harvesting complexes in the thylakoids are rich in photosynthetic pigments (chlorophyll, carotenoids, and xanthophylls) and contain a unique set of prenylquinol lipids (tocochromanol/vitamin E, plastoquinol, and phylloquinol/vitamin K1). In this chapter, methods for the isolation and quantification of chloroplast and leaf glycerolipids and prenylquinol lipids are presented. Glycerolipids are separated by thin-layer chromatography prior to conversion of the fatty acids into methyl esters. Fatty acid methyl esters are subsequently quantified by gas chromatography. Prenylquinol lipids are separated by HPLC and quantified by UV absorption (plastoquinol) or fluorescence (tocochromanol, phylloquinol). Key words: Prenylquinol, Membrane, Lipid, Galactolipids, Phospholipids, Glycolipid, Tocopherol, Phylloquinol, Plastoquinol
1. Introduction Glycerolipids are the major building blocks of all biological membranes. Chloroplast membranes are characterised by the prevalence of glycolipids, i.e., monogalactosyl diacylglycerol (MGD) and digalactosyl diacylglycerol (DGD). A further glycolipid, sulfoquinovosyl diacylglycerol (SQD), and the phospholipid phosphatidyl glycerol (PG) are found in the thylakoids in lower amounts. The latter two lipids are negatively charged under physiological pH and, therefore, are anionic. In addition, the outer chloroplast envelope contains another phospholipid, phosphatidyl choline (PC) (1). The synthesis of chloroplast lipids in Arabidopsis involves two pathways (2–4). Fatty acids are synthesized in the R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3_22, © Springer Science+Business Media, LLC 2011
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chloroplast stroma. In many plants including Arabidopsis, a certain proportion of fatty acids are employed for the synthesis of plastidial glycerolipids (prokaryotic pathway). These lipids are characterised by the presence of hexadecatrienoic acid (16:3) at the sn-2 position of the glycerol backbone, in particular of MGD. Plants such as Arabidopsis containing considerable amounts of 16:3 in MGD are designated “16:3 plants”. The other fraction of plastidderived fatty acids is exported to the endoplasmic reticulum for the production of extraplastidial glycerolipids. Some of the extraplastidial lipid precursors are retransported into the chloroplast leading to the synthesis of chloroplast lipids carrying mostly a-linolenic acid (18:3) at the sn-2 position of glycerol (eukaryotic pathway). A number of plant species such as pea use the prokaryotic pathway for the synthesis of plastidial PG, while their glycolipids are derived from the eukaryotic pathway (“18:3 plants”). Under normal growth conditions, the occurrence of MGD, DGD, and SQD is restricted to the chloroplasts. During phosphate starvation, however, the synthesis of DGD and SQD is increased. While SQD remains in the chloroplasts, DGD accumulates in plastidial and extraplastidial membranes during phosphate deprivation (5, 6). DGD and SQD serve as surrogate lipids for plastidial and extraplastidial phospholipids. This mechanism provides the means to save phosphate during growth in phosphate-deficient environments. Chloroplast glycerolipids constitute ca. 75% of total leaf lipids (Table 1). Therefore, glycerolipids are most often extracted from whole leaves rather than from isolated chloroplasts. This strategy
Table 1 Lipid composition (mol% of total) of Arabidopsis leaves (45) and spinach chloroplast membranes (46) MGD
DGD
Leaves (+P)a
40
17
PI
PG
PC
PE
9
d
10
17
8
29
11
d
Leaves (−P)
40
7
9
4
17
29
6
5
10
32
0
Inner envelopec
55
30
5
1
9
0
0
Thylakoidsc
57
27
7
1
7
0
0
b
Outer envelope
c
SQD
Arabidopsis plants grown on synthetic medium with phosphate Arabidopsis plants grown on synthetic medium under phosphate deprivation c Isolated from spinach leaves d PI and SQD were not separated in these experiments a
b
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avoids the need for tedious chloroplast isolation, which can result in undesired lipid breakdown. In contrast to chloroplasts, extraplastidial membranes are enriched in phospholipids, i.e., PC, PG, phosphatidyl ethanolamine (PE), phosphatidyl inositol (PI), and phosphatidyl serine (PS). Separation and analysis of glycerolipids can be achieved by HPLC (7), a combination of thin-layer chromato graphy (TLC) and gas chromatography (GC) (8, 9), or by electron spray ionization-mass spectrometry (ESI-MS) (10). In addition to glycerolipids, chloroplasts contain a unique set of prenylquinol lipids, i.e., tocochromanols, phylloquinol, and plastoquinol. Numerous studies were devoted to the elucidation of the function and localization of prenylquinol lipids in chloroplasts (11–16). Tocochromanols serve as antioxidants (17–19), while phylloquinol (vitamin K1) and plastoquinol are the electron carriers of PSI and PSII, respectively (20–23). To this day, there is no good evidence for the occurrence of these prenylquinol lipids outside of chloroplasts. Similar to glycero lipids, prenylquinol lipids are commonly measured in wholeleaf extracts rather than isolated chloroplasts. Tocochromanols encompass a group of four forms of tocopherols (a-tocopherol, b-tocopherol, g-tocopherol, d-tocopherol) and four forms of tocotrienols, which differ in number and positions of methyl groups and in the degree of side-chain desaturation. Tocopherols carry a phytyl side chain, while tocotrienols harbor a geranylgeranyl moiety. Furthermore, plants including Arabidopsis contain another form of tocochromanol with a solanesyl side chain (“plastochromanol”) (11, 14, 24). In Arabidopsis, as in many other dicot plants, a-tocopherol is the most abundant form in leaves, while g-tocopherol is predominant in seeds (25). Tocotrienols are found in monocot plants such as wheat, rice, barley, and palm oil (26) and only in a limited number of dicot species (Carum carvi) (27), but are absent from Arabidopsis. The standard protocol for tocochromanol quantification involves HPLC separation with fluorescence detection in the presence of an internal standard. Furthermore, tocochromanols can be identified by thin-layer chromatography (TLC) (28) or by gas chromatography-mass spectrometry (GC-MS) (29). Phylloquinol (vitamin K1) and plastoquinol each harbor a redox-active quinone ring which is the basis for their function as electron carriers in photosynthesis. Therefore, the two compounds can exist in a reduced, quinol (hydroquinone) or an oxidized, quinone state. The semiquinone state, which is a radical, is not stable over time. Phylloquinol is measured after reduction by fluorescence HPLC. Plastoquinone, however, is quantified in its oxidized state by HPLC with UV detection.
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2. Materials 2.1. Glycerolipid Extraction
1. Lipid extraction solutions: chloroform–methanol–formic acid (1:1:0.1); and, 1 M KCl, 0.2 M H3PO4 in H2O. 2. Glass tubes with screw thread (e.g., 12 mm × 100 mm) and screw caps/teflon septa. 3. Glass test tubes (e.g., 12 mm × 100 mm). 4. Pasteur pipettes.
2.2. Glycerolipid Separation by Thin-Layer Chromatography
1. TLC plates with concentration zone (SI250, J.T. Baker, code 7003-04). 2. Ammonium sulfate solution: 0.15 M (NH4)2SO4; TLC plates are impregnated by briefly submerging in ammonium sulfate solution. The plates should be dried and stored in a dust-free place for at least 2 days. 3. Glass tanks with lid for TLC. 4. TLC developing solution: acetone–toluene–H2O (91:30:8). 5. ANS reagent: 0.2% aniline naphthalene sulfonic acid (ANS) in methanol. 6. Glass sprayers and compressed air for the application of staining reagents onto TLC plates. 7. UV light table or lamp (wavelength: 366 nm). 8. Glass tank filled with ca. 5 g crystalline iodine for lipid staining. Iodine will sublimate into the gas phase after ca. 24 h. 9. Micropipettes to handle volume ranges of 10–100 mL.
2.3. Synthesis of Fatty Acid Methyl Esters and Analysis by Gas Chromatography
1. Glass tubes with screw thread (e.g., 12 mm × 100 mm) and screw caps/teflon septa. 2. Water bath set at 80°C. 3. Glass test tubes (12 mm × 100 mm). 4. Methanolic hydrochloric acid: 1 N HCl in methanol; diluted from 3 N methanolic HCl (Sigma) with methanol; store at 4°C. 5. Pentadecanoic acid (15:0) as internal standard; working stock: 50 mg/mL 15:0 in methanol; store at −20°C. 6. Hexane for fatty acid methyl ester (FAME) extraction. 7. 0.9% NaCl solution for washing and phase separation of lipid extracts. 8. Pasteur pipettes. 9. Sample concentrator with nitrogen gas stream (e.g., SC-4 Hood and Stand from Techne). 10. Glass vials for GC autosampler.
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11. Rapeseed FAME standard reference mix (Supelco). 12. Gas chromatograph (e.g., Agilent HP6890) with autoinjector (injector temperature: 220°C) and flame ionization detector (detector temperature: 250°C; detector gasses: hydrogen, flow rate 30 mL/min; synthetic air, flow rate 400 mL/min; helium makeup flow, 19 mL/min). 13. GC column: SP 2380, fused silica capillary column, 30 m × 0.53 mm, 0.20-mm film (Supelco), carrier gas flow rate: 7 mL/min of helium; splitless injection of 2 mL. 2.4. Quantification of Tocochromanols
1. Glass tubes with screw thread (e.g., 12 mm × 100 mm) and screw caps/teflon septa. 2. Tocochromanol isolation solution: diethyl ether; and 1 M KCl, 0.2 M H3PO4. 3. Tocol as internal standard: 10 ng/mL tocol in ethanol; store at −20°C. 4. Sample concentrator with nitrogen gas stream. 5. Isocratic HPLC solvent: hexane–tertiary butylmethyl ether (96:4). 6. Glass vials for HPLC autosampler. 7. Tocopherol standards: a-tocopherol, b-tocopherol, g-tocopherol, d-tocopherol (Calbiochem); a linseed oil extract can be used as standard for plastochromanol. 8. HPLC (e.g., Agilent series 1200 HPLC) with fluorescence detector. 9. HPLC column: LiChrospher 100-Diol 250 mm × 3 mm, 5-mm particle size (Knauer, Berlin).
2.5. Quantification of Phylloquinol
1. Glass tubes with screw thread (e.g., 12 mm × 100 mm) and screw caps/teflon septa. 2. Phylloquinol isolation solvents: hexane; methanol–water (9:1); and methanol–dichloromethane (9:1). 3. Menaquinone-4 (from Sigma vitamin K2) as internal standard: 5 ng/mL menaquinone-4 in ethanol; store at −20°C. 4. Sample concentrator with nitrogen gas stream. 5. Isocratic HPLC solvent: mix 900 mL of methanol, 100 mL of dichloromethane, and 5 mL of a methanolic solution of 1.37 g of ZnCl2, 0.41 g of sodium acetate, and 0.30 g of acetic acid. 6. Glass vials for HPLC autosampler. 7. HPLC with fluorescence detector. 8. Reversed-phase (RP18) column: Eurosphos-100, 250 mm × 4.6 mm, 5-mm particle size (Knauer, Berlin). 9. Postcolumn reduction cartridge: 30 mm × 4 mm, filled with zinc powder, ca. 63-mm particle size.
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2.6. Quantification of Plastoquinol
1. Glass tubes with screw thread (e.g., 12 mm × 100 mm) and screw caps/teflon septa. 2. Ubiquinone-4 from Sigma as internal standard: 10 ng/mL ubiquinone-4 in ethanol; store at −20°C. 3. Plastoquinol isolation solutions: hexane; and 1 M KCl, 0.2 M H3PO4. 4. Sample concentrator with nitrogen gas stream. 5. Isocratic HPLC solvent: hexane–tertiary butylmethyl ether (90:10). 6. Glass vials for HPLC autosampler. 7. HPLC with UV or diode array detector. 8. HPLC column: Lichrospher 100 Diol, 250 mm × 3 mm, 5-mm particle size (Knauer, Berlin).
3. Methods 3.1. Glycerolipid Extraction (3, 9)
1. Grind plant material (one Arabidopsis leaf, ca. 150 mg fresh weight) in liquid nitrogen with mortar and pestle (see Note 1). 2. Add 1 mL of chloroform–methanol–formic acid (1:1:0.1) and transfer slurry into screw cap glass tube (see Notes 2–4). 3. Add 0.5 mL of 1 M KCl, 0.2 M H3PO4 and mix vigorously. The acids inactivate lipid-degrading enzymes. 4. Centrifuge for 3 min at 4,000 × g for phase separation. 5. Go through the upper aqueous phase with a Pasteur pipette and transfer the lower chloroform phase containing the lipids to a fresh test tube. 6. Evaporate the solvent in sample concentrator with nitrogen gas stream (see Note 5). Dissolve lipids in 50–100 mL chloroform–methanol (2:1) and store at −20°C.
3.2. Glycerolipid Separation by ThinLayer Chromatography (3, 9, 30)
1. Use ammonium-sulfate impregnated TLC plates. Activate the plate at 120°C for 2.5 h prior to chromatography (see Note 6). 2. Apply the lipid samples onto the concentration zone of the TLC plates using a micropipette or a yellow (200 mL) pipette tip. Reference lipids (phospholipids, galactolipids) can be applied in extra lanes. 3. Develop the plate in a TLC tank containing acetone–toluene– H2O (91:30:8; volumes in mL). This takes about 50 min. 4. Take out and dry the plate in a fume hood for 15 min. 5. Visualize the lipids after spraying with ANS reagent by fluorescence under UV light (366 nm). Lipid bands are marked with a pencil (see Notes 7–9).
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6. Scrape the bands of the silica material containing the lipids (MGD, PG, DGD, SQD, PE, PC) from the plate with a razor blade and individually collected in screw cap glass vials. Use a small glass funnel if necessary. 3.3. Synthesis and Analysis of Fatty Acid Methyl Esters by Gas Chromatography (8)
1. Fill plant material or silica gel with lipid isolated by TLC into a screw-cap glass vial. 2. Add 1 mL of methanolic hydrochloric acid and 100 mL of internal standard (15:0, 5 mg) to the sample (pipette with yellow tips may be used) (see Note 10). 3. Close the vials tightly, vortex, and incubate the samples in water bath at 80°C for 20 min. 4. After cooling, add 1 ml of 0.9% NaCl solution and 1 mL of hexane; mix vigorously by vortexing and inverting tubes several times. 5. Centrifuge (3,000 × g, 3 min) for phase separation. 6. Transfer the upper hexane phase containing the FAMEs to a test tube with a Pasteur pipette. 7. Evaporate the solvent in sample concentrator with nitrogen gas stream; dissolve sample in 50–100 mL of hexane and transfer to a GC vial. 8. FAMEs are separated by GC with a temperature gradient of: initial, 100°C; to 160°C with 25°C/min; to 220°C with 10°C; to 100°C with 25°C. 9. The fatty acids are identified after comparison to the retention times of the rapeseed FAME standard mix. Table 2 shows the fatty acid composition of Arabidopsis leaves, and Fig. 1b shows a typical GC chromatogram. Detector response factors are calculated from the signals of the rapeseed FAME standard mix. The amounts of fatty acids in mg are obtained from the report table. The mg values are converted into nmol and used to calculate the amounts of glycerolipids after division by two (two fatty acids per glycerol backbone). The composition of lipid classes (mol%) and the fatty acid composition of each individual lipid class (mol%) can then be calculated. Perform at least three independent measurements of different leaf lipid extracts to obtain reliable data for mean and standard deviation (see Note 11).
3.4. Quantification of Tocochromanols (11, 31)
1. Harvest one Arabidopsis leaf, determine leaf fresh weight (ca. 100–150 mg) and freeze in liquid nitrogen. Grind samples with mortar and pestle in liquid nitrogen to obtain fine powder (see Note 1). 2. Extract lipids on ice in a fume hood. To each sample, add 50 mL (500 ng) of tocol, 0.5 mL of diethyl ether, and 0.25 mL of 1 M KCl, 0.2 M H3PO4. Vortex and centrifuge for 5 min at 10,000 × g at 4°C (see Notes 2 and 12).
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Table 2 Fatty acid composition of an Arabidopsis leaf* Fatty acid
mol%
16:0
15.0
16:1-cis
1.1
16:1-trans
2.7
16:2
1.1
16:3
13.8
18:0
1.0
18:1
3.5
18:2
15.7
18:3
46.0
a
16:1-cis and 16:1-trans coelute on many GC columns *Data were taken from ref. 47 a
Fig. 1. Glycerolipid and fatty acid composition of an Arabidopsis leaf. (a) Separation of a leaf lipid extract by TLC. Lipids were stained with iodine vapor. (b) GC analysis of fatty acid methyl esters from an Arabidopsis leaf. Pentadecanoic acid (15:0) was used as internal standard.
3. Collect the upper phase containing tocochromanols in a glass test tube. To the residual plant material, add again 0.5 ml of diethyl ether and vortex vigorously. Centrifuge and combine the organic phases. Repeat extraction until the leaf tissue is white. 4. Evaporate solvent under nitrogen gas stream (see Note 5).
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5. Dissolve samples in 50 mL of hexane, vortex, and transfer to an HPLC vial. 6. The best separation of tocochromanols yielding individual peaks for a-tocopherol, b-tocopherol, g-tocopherol, d-tocopherol and tocochromanol is obtained by HPLC on a Diol column (11, 31) (Fig. 2a). Lipids are separated by isocratic chromatography with hexane–tertiary butylmethyl ether (96:4) (flow rate: 0.75 mL/min, time of run: 30–35 min) with fluorescence detection
Fig. 2. HPLC analysis of chloroplast prenylquinol lipids from Arabidopsis leaves. (a) Separation of tocochromanols by HPLC using a Diol column. Tocopherols and plastochromanol were detected by fluorescence. (b) HPLC quantification of phylloquinol using menaquinol-4 as internal standard. Prenylquinol lipids were subjected to postcolumn reduction prior to detection by fluorescence. (c) Plastoquinone measurement by HPLC and UV detection. IS, internal standards (tocol, menaquinol-4, and ubiquinone-4).
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(excitation of 290 nm, emission of 330 nm). Peaks are identified by cochromatography with the respective tocochromanol standards. The standards are used to calculate detector response factors. Tocopherol and plastochromanol concentrations are adjusted using published absorption coefficients (E, 1 cm, 1% in ethanol: a-tocopherol, 75.8 at 292 nm; b-tocopherol, 296 at 89.4 nm; g-tocopherol, 91.4 at 298 nm; d-tocopherol, 87.3 at 298 nm; plastochromanol, 55.5 at 296 nm) (32, 33). The amounts of tocochromanols in mg are converted into nmol per g fresh weight. Mean and standard deviation should be calculated from a minimum of three measurements (see Notes 13 and 14). 3.5. Quantification of Phylloquinol (16, 34)
1. Harvest one leaf and determine fresh weight (ca. 100 mg). Freeze in liquid nitrogen. 2. Grind the leaf with mortar and pestle in liquid nitrogen (see Note 1). 3. Extract lipids on ice under the fume hood. Add 0.8 mL of 2-propanol–hexane (3:1) and 50 mL of menaquinone-4 as internal standard (250 ng) (see Notes 2 and 12). 4. Transfer slurry to microfuge tubes. Vortex. 5. Centrifuge samples at 14,000 × g for 5 min. Transfer the upper lipid phase to a glass test tube. 6. Extract the remaining pellet again with 0.6 mL of hexane. To the combined organic phases, add 0.6 mL of methanol–water (9:1), and after vortexing and centrifugation, collect the organic phases. The solvent is evaporated with nitrogen gas and the residue dissolved in 100 mL of methanol–dichloromethane (9:1) for HPLC measurement (see Note 5). 7. Quantify phylloquinol by HPLC separation with fluorescence detection (16, 34) (Fig. 2b). Lipids are separated by isocratic chromatography (flow rate: 1 mL/min, time of run: 15 min) on a reversed-phase column equipped with a postcolumn reduction cartridge filled with zinc. Phylloquinol is measured by fluorescence (excitation, 243 nm; emission, 430 nm) (see Note 15). 8. Use phylloquinone and menaquinone-4 standards to determine the retention times and the detector response factors. The absorption coefficients at 248 nm in petroleum ether are: phylloquinone, e = 18,880 L/mol/cm; menaquinone-4 (e = 18,870 L/mol/cm) (35). The final amount of phylloquinol is determined in mg or nmol per g fresh weight. Calculate mean and standard deviation after measuring three separate replicas.
3.6. Quantification of Plastoquinol (11)
1. Harvest one leaf (ca. 100 mg), determine fresh weight and freeze in liquid nitrogen. 2. Grind the leaf material with mortar and pestle in liquid nitrogen (see Note 1).
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3. Extract lipids on ice under the fume hood. To each sample, add 50 mL of ubiquinone-4 solution (500 ng) as internal standard, 0.5 mL of hexane and 0.25 mL of 1 M KCl, 0.2 M H3PO4 (see Notes 2 and 12). 4. Vortex and centrifuge for 5 min at 10,000 × g at 4°C. 5. Collect the upper organic phase into a glass test tube. Extract again with 0.5 mL of hexane. Centrifuge and combine organic phases. Repeat extraction until the leaf is white. 6. Evaporate samples under nitrogen gas stream (see Note 5). 7. Dissolve samples in 50 mL of hexane, vortex, and transfer to HPLC vials. 8. Lipids are separated by isocratic HPLC (flow rate: 0.75 mL/ min, time of run: 15 min) on a Diol column by recording the absorption at 255 nm (Fig. 2c) (11). 9. The content of plastoquinone is determined based on the amount of the internal standard (ubiquinone-4) after calculation of response factors. Concentrations are adjusted using the absorption coefficients of plastoquinone (e = 15,200 L/mol/cm at 255 nm, in ethanol) and ubiquinone-4 (e = 14,900 L/mol/cm at 275 nm, in ethanol) (35). Determine mean and standard deviation of plastoquinol in nmol per g fresh weight after measuring three separate replicas (see Notes 16 and 17).
4. Notes 4.1. Glycerolipid Extraction
1. The protocols given above are based on the isolation of chloroplast lipids from whole leaves frozen in liquid nitrogen immediately after harvest. It is absolutely essential to ensure that the plant material does not thaw prior to getting in contact with the extraction buffer to prevent lipid degradation. 2. When working with lipids, use glassware rather than plasticware whenever possible (glass tubes, Pasteur pipettes). Do not use rubber septa but teflon ones. Organic solvents (chloroform, hexane) can dissolve certain components of plastic or rubber materials (e.g., plasticizers) that give rise to background peaks during GC or HPLC separation. 3. An alternative method for glycerolipid extraction from leaves includes the deactivation of lipid-degrading enzymes by boiling of the plant tissue followed by extraction with chloroform– methanol–NaCl solution (36, 37): (a) Boil the plant material (one Arabidopsis leaf of ca. 100 mg) with 1.5 mL of water for about 10 min in water bath. (b) Pour off water and after cooling, add 1.6 mL of chloroform–methanol (1:2) and mix gently for 30 min.
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(c) Collect chloroform phase in new vial. (d) Repeat extraction of the plant material with another aliquot (1.6 mL) of chloroform–methanol (2:1). (e) Combine chloroform extracts and add 1.6 mL of chloroform and 1.2 mL of 0.9% NaCl; the final ratio of chloroform–methanol–0.9% NaCl should be 2:1:0.75; mix vigorously. (f) Centrifuge 3 min at 4,000 × g, go through the upper aqueous phase with a Pasteur pipette and transfer the lower organic phase to a new glass vial. (g) Evaporate solvent under nitrogen gas stream. Dissolve lipids in 100 mL of chloroform–methanol (2:1) and store at −20°C. 4. Glycerolipids can also be extracted from aqueous membrane fractions, e.g., isolated chloroplasts, thylakoids, or envelope membranes (5): (a) Add 2 volumes of chloroform–methanol–formic acid (1:1:0.1) to the aqueous membrane fraction and mix vigorously. (b) Centrifuge 3 min at 4,000 × g and transfer the lower organic phase to a new glass vial. (c) Evaporate solvent under nitrogen stream. Dissolve lipids in 100 mL of chloroform–methanol (2:1) and store at −20°C. 5. Reduce the volume of organic solvents in test tubes by placing under nitrogen gas stream in a sample concentrator. If nitrogen gas is not available, compressed air might be used provided that it is absolutely free of oil. 4.2. Glycerolipid Separation by Thin-Layer Chromatography
6. Activation of ammonium sulfate-impregnated TLC plates at 120°C leads to the conversion of (NH4)2SO4 into NH3 and H2SO4. While ammonia is volatile, the sulfuric acid results in acidification of the silica material. Thus, the migration of anionic lipids is altered, leading to an improved separation of phospholipids and glycolipids (3) (Fig. 1a). 7. Lipids can be stained by placing the TLC plate in an air-tight glass tank containing iodine vapor. Lipids are stained in a yellow-brownish color. Staining is partially reversible, but polyunsaturated fatty acids can be covalently modified. Therefore, take care to protect lipids destined for quantification against overexposure to iodine. Figure 1a shows a TLC plate with Arabidopsis leaf glycerolipids. 8. Alternatively, the lanes containing the reference lipids can be selectively stained with iodine. Protect the lanes containing lipids
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destined for quantification with a sheet of paper or a glass plate. Stain the reference lipids with a Pasteur pipette, filled with some iodine crystals fixed between two layers of glass wool. Direct a gentle iodine stream onto the lipid spots by attaching the Pasteur pipette to a tubing connected to compressed air. 9. Spraying with a-naphthol reagent represents an alternative, irreversible method to stain glycolipids on TLC plates: (a) Produce a-Naphthol–H2SO4 reagent by dissolving 8 g of a-naphthol in 250 mL of methanol. Add concentrated H2SO4 dropwise (total of 30 mL) to the a-naphthol solution under stirring on ice. (b) Spray the TLC plate with a-naphthol reagent and heat to 137°C for 5–15 min. Glycolipids (MGD, DGD, SQD) are stained in a red color. 4.3. Synthesis and Analysis of Fatty Acid Methyl Esters by Gas Chromatography
10. Fatty acid methyl esters are synthesized by acidic transesterification of lipid-bound and free fatty acids to methanol. It is not necessary to isolate the lipids prior to derivatization. Instead, whole plant material (e.g., a single Arabidopsis leaf) or silica gel from the TLC plate containing the lipids can be directly used. 11. Because of the complexity of the molecular composition of the glycerolipids, different methods have been developed for their identification and quantification. In addition to the TLC–GCbased method described here, photodensitometric scanning of TLC plates after lipid staining has been used for lipid quantification (38–40). Recent developments have resulted in the establishment of liquid chromatography (LC) and electronspray ionization (ESI) MS-based methods (see: Analytical Laboratory at the Kansas Lipidomics Research Center, Kansas State University) (10).
4.4. Quantification of Tocochromanols
12. For the extraction of prenylquinol lipids (e.g., tocochromanols, phylloquinol, or plastoquinol) from isolated chloroplasts, add 2 volumes of chloroform–methanol (2:1) and 1 volume of 1 M KCl, 0.2 M H3PO4. 13. An alternative HPLC method is based on normal-phase column chromatography (41). Tocochromanols are separated on a LiChrosorb Si60 column (250 mm × 4.6 mm, 5-mm particle size, Knauer, Berlin) with isocratic elution (hexane–diethyl ether [95:5] at a flow rate of 0.75 mL/min). Tocochromanols are detected by fluorescence (see Subheading 3.4). However, in normal-phase chromatography, g-tocopherol tends to coelute with plastochromanol (11). 14. HPLC separation on a reverse-phase column represents yet another, robust method for tocochromanol separation.
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Lipids are injected onto an RP18 column (Eurosphere-100, 250 mm × 4.6 mm, 5-mm particle size, Knauer, Berlin) with gradient elution: from 0 to 10 min: 100% methanol; from 10 to 30 min linear gradient to 3% methanol/97% 2-propanol; hold until 40 min; flow rate: 0.5 mL/min (42). Alternatively, an isocratic solvent of 5% (v/v) 2-propanol in methanol can be used at a flow rate of 2 mL/min (43). Tocochromanols are detected by fluorescence (see Subheading 3.4). During reversephase chromatography, the order of elution of tocopherols is reversed as compared to normal-phase HPLC. Furthermore, b-tocopherol and g-tocopherol cannot be separated because they coelute on reverse-phase HPLC. 4.5. Quantification of Phylloquinol
15. During the isolation from plant tissue, the predominant fraction of phylloquinol is oxidized by contact with oxygen from the air. Only the reduced form, phylloquinol, but not the oxidized form, phylloquinone, can be measured by fluorescence detection. Therefore, the oxidized form of phylloquinone is reduced during HPLC in a postcolumn derivatization cartridge filled with zinc powder prior to fluorescence detection. Because zinc is oxidized to Zn2+, the zinc in the cartridge should be replaced on a regular basis.
4.6. Quantification of Plastoquinol
16. Take care to correctly identify the plastoquinone peak in the HPLC chromatogram between the large carotenoid peak and the phylloquinone peak. 17. A recent publication describes the separation of the oxidized and reduced form of ubiquinol and plastoquinol by HPLC with UV and fluorescence detection (44). This method is useful to determine the redox state of the plastoquinol pool.
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5. Härtel, H., Dörmann, P., and Benning, C. (2000) DGD1-independent biosynthesis of extraplastidic galactolipids after phosphate deprivation in Arabidopsis. Proc. Natl. Acad. Sci. USA 97, 10649–10654. 6. Kelly, A. A., Froehlich, J. E., and Dörmann, P. (2003) Disruption of the two digalactosyldiacylglycerol synthase genes DGD1 and DGD2 in Arabidopsis reveals the existence of an additional enzyme of galactolipid synthesis. Plant Cell 15, 2694–2706. 7. Christie, W. W., Gill, S., Nordbäck, J., Itabash, Y., Sanda, S., and Slabas, A. R. (1998) New procedures for rapid screening of leaf lipid components from Arabidopsis. Phytochem. Anal. 9, 53–57.
22 Analysis of Lipid Content and Quality in Arabidopsis Plastids 8. Browse, J., McCourt, P. J., and Somerville, C. R. (1986) Fatty acid composition of leaf lipids determined after combined digestion and fatty acid methyl ester formation from fresh tissue. Anal. Biochem. 152, 141–145. 9. Benning, C., and Somerville, C. R. (1992) Isolation and genetic complementation of a sulfolipid-deficient mutant of Rhodobacter sphaeroides. J. Bacteriol. 174, 2352–2360. 10. Welti, R., Li, W., Li, M., Sang, Y., Biesiada, H., Zhou, H.-E., Rajashekar, C., Williams, T., and Wang, X. (2002) Profiling membrane lipids in plant stress responses. Role of phospholipase Da in freezing-induced lipid changes in Arabidopsis. J. Biol. Chem. 277, 31994–32002. 11. Zbierzak, A., Kanwischer, M., Wille, C., Vidi, P.-A., Giavalisco, P., Lohmann, A., Briesen, I., Porfirova, S., Bréhélin, C., Kessler, F., and Dörmann, P. (2010) Intersection of the tocopherol and plastoquinol metabolic pathways at the plastoglobule. Biochem. J. 425, 389–399. 12. Lichtenthaler, H., Prenzel, U., Douce, R., and Joyard, J. (1981) Localization of prenylquinones in the envelope of spinach chloroplasts. Biochim. Biophys. Acta 641, 99–105. 13. Soll, J., Schultz, G., Joyard, J., Douce, R., and Block, M. A. (1985) Localization and synthesis of prenylquinones in isolated outer and inner envelope membranes from spinach chloroplasts. Arch. Biochem. Biophys. 238, 290–299. 14. Food and Nutrition Board, Institute of Medicine (2003) Dietary Reference Intakes for Vitamin C, Vitamin E, Selenium, and Carotenoids. Natl. Acad. Press, Washington, D.C., USA, pp. 162–196. 15. Vidi, P. A., Kanwischer, M., Baginsky, S., Austin, J. R., Csucs, G., Dörmann, P., Kessler, F., and Brehelin, C. (2006) Tocopherol cyclase (VTE1) localization and vitamin E accumulation in chloroplast plastoglobule lipoprotein particles. J. Biol. Chem. 281, 11225–11234. 16. Lohmann, A., Schöttler, M. A., Bréhélin, C., Kessler, F., Bock, R., Cahoon, E. B., and Dörmann, P. (2006) Deficiency in phylloquinone (vitamin K1) methylation affects prenyl quinone distribution, photosystem I abundance and anthocyanin accumulation in the Arabidopsis AtmenG mutant. J. Biol. Chem. 281, 40461–40472. 17. Fryer, M. J. (1992) The antioxidant effects of thylakoid vitamin E (a-tocopherol). Plant Cell Environ. 15, 381–392. 18. Sattler, S. E., Gilliland, L. U., MagallanesLundback, M., Pollard, M., and DellaPenna, D. (2004) Vitamin E is essential for seed longevity and for preventing lipid peroxidation during germination. Plant Cell 16, 1419–1432.
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19. Maeda, H., Sakuragi, Y., Bryant, D. A., and DellaPenna, D. (2005) Tocopherols protect Synechocystis sp. strain PCC 6803 from lipid peroxidation. Plant Physiol. 138, 1422–1435. 20. Jordan, P., Fromme, P., Witt, H. T., Klukas, O., Saenger, W., and Krauss, N. (2001) Threedimensional structure of cyanobacterial photosystem I at 2.5 Å resolution. Nature 411, 909–917. 21. Ben-Shem, A., Frolow, F., and Nelson, N. (2003) Crystal structure of plant photosystem I. Nature 426, 630–635. 22. Fyfe, P. K., Hughes, A. V., Heathcote, P., and Jones, M. R. (2005) Proteins, chlorophylls and lipids: X-ray analysis of a three-way relationship. Trends Plant Sci. 10, 275–282. 23. Knaff, D. B., Malkin, R., Myron, J. C., and Stoller, M. (1977) The role of plastoquinone and b-carotene in the primary reaction of plant photosystem II. Biochim. Biophys. Acta 11, 402–411. 24. Whittle, K. J., Dunphy, P. J., and Pennock J. E. (1965) Plastochromanol in the leaves of Hevea brasiliensis. Biochem. J. 96, 17c–19c. 25. DellaPenna, D., and Pogson, B. J. (2006) Vitamin synthesis in plants: tocopherols and carotenoids. Annu. Rev. Plant Biol. 57, 711–738. 26. Cahoon, E. B., Hall, S. E., Ripp, K. G., Ganzke, T. S., Hitz, W. D., and Coughlan, S. J. (2003) Metabolic redesign of vitamin E biosynthesis in plants for tocotrienol production and increased antioxidant content. Nature Biotechnol. 21, 1082–1087. 27. Aitzetmüller, K. (1997) Antioxidative effects of Carum seeds. J. Am. Oil Chem. Soc. 74, 185. 28. Porfirova, S., Bergmüller, E., Tropf, S., Lemke, R., and Dörmann, P. (2002) Isolation of an Arabidopsis mutant lacking vitamin E and identification of a cyclase essential for all tocopherol biosynthesis. Proc. Natl. Acad. Sci. USA 99, 12495–12500. 29. Fiehn, O., Kopka, J., Trethewey, R. N., and Willmitzer, L. (2000) Identification of uncommon plant metabolites based on calculation of elemental compositions using gas chromatography and quadrupole mass spectrometry. Anal. Chem. 72, 3573–3580. 30. Dörmann, P., Hoffmann-Benning, S., Balbo, I., and Benning, C. (1995) Isolation and characterization of an Arabidopsis mutant deficient in the thylakoid lipid digalactosyl diacylglycerol. Plant Cell 7, 1801–1810. 31. Balz, M., Schulte, E., and Thier, H.-P. (1992) Trennung von Tocopherolen and Tocotrienolen durch HPLC. Fat Sci. Technol. 94, 209–213. 32. Schüep, W., and Rettenmeier, R. (1994) Analysis of vitamin E homologs in plasma and
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tissue: high-performance liquid chromatography. Meth. Enzymol. 234, 294–302. 33. Leerbeck, E., Søndergaard, E., and Dam, H. (1967) Occurrence of a plastochromanol in linseed oil. Acta Chem. Scand. 21, 2582. 34. Jakob, E., and Elmadfa, I. (1996) Application of a simplified HPLC assay for the determination of phylloquinone (vitamin K1) in animal and plant food items. Food Chem. 56, 87–91. 35. Dawson, R. M. C., Elliott, D. C., Elliott, W. H., and Jones, K. M. (1985) Data for Biochemical Research. Oxford University Press, Oxford, UK. 36. Bligh, E. G., and Dyer, W. J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. 37. Hölzl, G., Leipelt, M., Ott, C., Zähringer, U., Lindner, B., Warnecke, D., and Heinz, E. (2005) Processive lipid galactosyl/glucosyltransferases from Agrobacterium tumefaciens and Mesorhizobium loti display multiple specificities. Glycobiol. 15, 874–886. 38. Grether-Beck, S., Bonizzi, G., SchmittBrenden, H., Felsner, I., Timmer, A., Sies, H., Johnson, J. P., Piette, J., and Krutmann, J. (2000) Non-enzymatic triggering of the ceramide signalling cascade by solar UVA radiation. EMBO J. 19, 5793–5800. 39. Weerheim, A. M., Kolb, A. M., Sturk, A., and Nieuwland, R. (2002) Phospholipid composition of cell-derived microparticles determined by onedimensional high-performance thin-layer chromatography. Anal. Biochem. 302, 191–198. 40. Domergue, F., Abbadi, A., Ott, C., Zank, T. K., Zähringer, U., and Heinz, E. (2003) Acyl carriers used as substrates by the desaturases and elongases involved in very long-chain polyunsaturated fatty acids biosynthesis recon-
stituted in yeast. J. Biol. Chem. 278, 35115–35126. 41. Thompson, J. N. a. H., G. (1979) Determination of tocopherols and tocotrienols in foods and tissues by high performance lipid chromatography. J. Liquid Chromatog. 2, 327–344. 42. Johnson, T. W., Shen G., Zybailov, B., Kolling, D., Reategui, R., Beauparlant, S., Vassiliev, I. R., Bryant, D. A., Jones, A. D., Golbeck, J. H., and Chitnis, P. R. (2000) Recruitment of a foreign quinone into the A1 site of photosystem I. I. Genetic and physiological characterization of phylloquinone biosynthetic pathway mutants in Synechocystis sp. PCC 6803. J. Biol. Chem. 275, 8523–8530. 43. Collakova, E., and DellaPenna, D. (2003) Homogentisate phytyltransferase activity is limiting for tocopherol biosynthesis in Arabidopsis. Plant Physiol. 131, 632–642. 44. Yoshida, K., Shibata, M., Terashima, I., and Noguchi, K. (2010) Simultaneous determination of in vivo plastoquinone and ubiquinone redox states by HPLC-based analysis. Plant Cell Physiol. 51, 836–841. 45. Hölzl, G., Witt, S., Gaude, N., Melzer, M., Schöttler, M. A., and Dörmann, P. (2009) The role of diglycosyl lipids in photosynthesis and membrane lipid homeostasis in Arabidopsis. Plant Physiol. 150, 1147–1159. 46. Moreau, P., Bessoule, J. J., Mongrand, S., Testet, E., Vincent, P., and Cassagne, C. (1998) Lipid trafficking in plant cells. Prog. Lipid Res. 37, 371–391. 47. Miquel, M., and Browse, J. (1992) Arabidopsis mutants deficient in polyunsaturated fatty acid synthesis. Biochemical and genetic characterization of a plant oleoyl-phosphatidylcholine desaturase. J. Biol. Chem. 267, 1502–1509.
Index A ABRC. See Arabidopsis Biological Resource Center ACi curves............................................................. 318, 321–324 Affymetrix........................................................... 123, 124, 132 ALA. See 5-Aminolevulinic acid Alkaline extraction.............................................................96 Amino acid assay.............. 162, 163, 165–166, 168, 170, 173–174, 179 biosynthesis........................................................162, 163 9-Aminoacridine (9-AA)..........328, 330, 332–336, 338, 339 5-Aminolevulinic acid (ALA)...........358, 360, 366–367, 382 Amylopectin......................387–390, 392, 394, 398–400, 404 Amyloplast............................................... 145, 150, 388, 405 Amylose............................. 388, 389, 392, 394, 398–400, 408 Analytical ultracentrifugation (AU)................. 68, 69, 80–81 Antibody (-ies).................................7, 14, 15, 17, 33, 71–74, 76, 84, 194, 198, 200, 203–205, 227, 234, 238 Arabidopsis Biological Resource Center (ABRC)......98, 169 ARAMEMNON.......................................................96, 106 AT_CHLORO....................................................95–98, 110 AtProteome.............................................. 101, 104–106, 110 ATP synthase.......................... 4, 28, 210, 343, 344, 346, 352 ATP synthesis........................................... 334, 337, 343–354 Autofluorescence.................................................. 54, 60, 231 Autoradiography.................................................................76
B Basic local alignment search tool (BLAST).............. 99, 103, 108–111 Bimolecular fluorescence complementation (BiFC)..................................................51–64, 71 Bioinformatics............................................................94, 266 Biolistic (gun, particle delivery system, particle bombardment)................................ 55, 58–59, 63 Blot dot.............................................................. 68, 69, 74–77 Northern...............................................................12, 123 Southern.......................................................................12 Western or immunoblot..............................4, 5, 7, 10, 11, 14–17, 27, 33, 47, 48, 71, 72, 193–195, 197–200, 203, 204, 224, 226–227, 230, 233–234, 365, 379, 380
Blue native polyacrylamide gel electrophoresis (BN-PAGE).........................3–17, 19, 20, 23, 25, 27–29, 67, 273
C Calmodulin......................................32–33, 35, 36, 38, 43, 44 Calmodulin binding peptide (CBP).............................32, 33 Carotenoids (Car) carotenoid........................... 118, 224, 328, 330, 334, 346, 357–384, 424 ECS......................................328, 330, 334–336, 339, 340, 345–346, 348, 352–353 quantification by HPLC............................. 364, 375–377 spectrophotometric(al) determination.......................363, 373–374 cDNA.......................56, 62, 68, 104, 120, 126, 128, 129, 132 cDNA synthesis................................................ 120, 128–129 CFP. See Cyan fluorescent protein Chemical cross-linking..............37, 40–42, 45–47, 67, 71–73 Chemiluminescence (or ECL)...................... 7, 14, 195, 200, 204, 227, 234, 235, 238, 365 Chlorophyll (Chl) catabolites........................................... 358, 365–366, 381 extraction, quantification......................47, 152, 211, 214, 216, 220, 235, 236, 319, 359, 363–364, 375–377 fluorescence................................164, 169, 180–181, 231, 299–309, 311, 318, 320, 322–324 quantification by HPLC............................. 364, 375–377 spectrophotometric(al) determination................. 47, 220, 363, 373–374 Chlorophyllide (Chlide)...................362–363, 367, 370–373, 381, 383, 384 Chloroplast DNA (plastid genome)................. 112, 241, 242 Chloroplast Function Database............................98–99, 110 Chloroplast isolation......................................... 10, 191–192, 195–196, 198, 201–202, 210–211, 213–214, 219–220, 224–225, 228–230, 235–326, 329, 331–332, 413 Chromatography affinity............................................. 31–48, 209, 284, 286 gas (GC).....................................135–137, 141–142, 144, 145, 150, 152–155, 157, 163, 167, 175, 413–415, 417, 418, 421, 423
R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume II, Methods in Molecular Biology, vol. 775, DOI 10.1007/978-1-61779-237-3, © Springer Science+Business Media, LLC 2011
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Chromatography (Continued ) gel filtration. see Size exclusion immobilized metal affinity (IMAC)..................283–284, 286, 289, 290 reverse phase...............................100, 266, 273–274, 285, 286, 292, 293, 423–424 size exclusion (SEC).........................3, 68, 69, 75, 77–79, 278, 279 strong cation exchange (SCX)...........................283–285, 289, 293, 295 thin layer (TLC)..................413, 414, 416–418, 422–423 Chromoplast.....................................................................223 Clear native electrophoresis.................................... 20, 23, 27 CO2 fixation (CO2 uptake, CO2 assimilation).................162, 302, 311–326, 388–389, 407 Confocal microscopy.................................. 60, 202, 230, 231 Coomassie stain (Coomassie Brilliant Blue, CBB)..............4, 5, 10, 11, 14, 16, 20, 27, 193, 197, 198, 246, 253, 254, 257, 268, 271, 280 cpDNA. See Chloroplast DNA Cross-linking.............................37, 40–42, 45–47, 67, 71–73 Cyan fluorescent protein (CFP)................. 53–57, 60–62, 64
D Databases ARAMEMNON.................................................96, 106 AT_CHLORO..............................................95–98, 110 AtProteome........................................ 101, 104–106, 110 Chloroplast 2010................................................161–163 Chloroplast Function Database......................98–99, 110 insertional mutants.....................................................162 organelle....................................... 95, 101–103, 110–111 P3DB . ..............................................................108–110 PhosPhAt........................................... 101, 106, 107, 111 plprot............................................................... 99–100, 111 PODB2...................................................... 101–102, 111 PPDB..............................................96, 100–101, 111, 267 proteomics................................5, 93–112, 267, 274–276, 284, 292–293, 296 SUBA (SUBA II)................101–103, 106, 111, 112, 209 TAIR........................................ 96, 98, 101–103, 106, 112, 123, 125, 132 transcriptomic..................................... 102, 112, 117–133 Detergent..................................................4–7, 15, 16, 20, 25, 26, 29, 33, 35, 36, 46, 47, 72, 76, 81, 100, 210, 273, 281 Digitonin..............................4–7, 14, 15, 20, 22–23, 25–29, 47 DNA archiving..................................... 163, 167–171, 177–178 cDNA...............................................56, 62, 68, 104, 120, 126, 128, 129, 132 chloroplast (plastid genome)....................... 112, 241, 242 preparation, extraction................................................177 Dot blot.............................................................. 68, 69, 74–77
2D-PAGE. See Two-dimensional polyacrylamide gel electrophoresis 1D-PAGE or 1D SDS-PAGE............... 3–17, 197, 253, 267 DpH (delta pH)........................................ 327–340, 344, 352
E ECS. See Electrochromic shift Electric(al) field, thylakoid............................... 327–340, 346 Electric(al) potential, thylakoid................................328, 344 Electrochromic shift (ECS)..................................... 328, 330, 334–336, 339, 340, 345–346, 348, 352–353 Electron microscopy........................................... 33, 156, 242 Electrophoresis agarose..........................................6, 12, 16, 23, 27, 29, 120, 122, 123, 251–252 BN-PAGE........................................3–17, 19, 20, 22, 23, 25, 27–29, 67, 273 denaturing......................... 4, 6, 7, 11–13, 15–16, 19, 122 2D-PAGE....................................3–17, 19–29, 207, 208, 210, 218, 219, 245, 252, 253, 255 1D-PAGE or 1D SDS-PAGE......... 3–17, 197, 253, 267 gradient gel................ 4, 6–10, 15, 21, 23–26, 29, 46, 270 native.........................................3–17, 19–29, 67, 209, 273 polyacrylamide............................................. 3–17, 19–29, 34–36, 38, 44, 46, 67, 72, 74, 94, 96, 108, 192–193, 197–198, 204, 207–210, 216, 218–219, 226, 230–234, 245–246, 251–255, 257, 266–267, 270, 273 RNA............................ 119–120, 122–123, 131, 251–252 SDS-PAGE..................................4–7, 11–16, 19, 23–24, 26–29, 34–36, 46, 72, 74, 94, 96, 108, 192–193, 197, 198, 204, 216, 226, 230–234, 245, 246, 251–254, 257, 266, 267 urea........................................23–24, 26–27, 212–213, 252 Envelope inner...................................... 4, 70, 72, 73, 190, 194, 199, 207, 230, 337, 412 lipids......................................................190, 411–412, 422 outer........................................ 4, 70, 72, 75, 85, 190, 199, 207, 227, 230, 234, 411, 412 protein topology...................................10, 72, 75, 85, 87, 96, 194, 196, 197, 199, 203, 204, 234 proteomics...................................... 96–97, 189–205, 208 purification, fractionation................... 189–205, 224, 230 Etioplast........................................................................ 99, 100
F False discovery rate (FDR).............................. 105, 124, 131, 266, 274, 275, 284, 292, 296 Fatty acid assay.............................. 162–163, 166–167, 170, 174–175 biosynthesis........................................ 162–163, 411–412 other..............................................76, 118, 154, 218, 224, 414, 417, 418, 422, 423
Chloroplast Research in Arabidopsis 429 Index
Fluorescence microscopy.............................. 51–64, 202, 231 Fluorimeter, PAM.............................164, 169, 180, 299–309 Förster resonance energy transfer (FRET)..................51–64, 71, 86
G Galactolipids....................................................................416 Gas-chromatography (GC)............................. 135–137, 141, 142, 144, 150, 152–155, 157, 163, 167, 175, 413–415, 417, 418, 421, 423 Gas exchange............................................................311–326 Gel filtration (size exclusion chromatography)..............3, 68, 75, 77–79, 267, 278, 279 Genetics.............................................112, 136, 161–183, 358 Genotyping...................................... 163, 167–171, 177–178 GFP. See Green fluorescent protein Glycerolipids.....................190, 411–414, 416–418, 421–423 Gradient centrifugation non-aqueous fractionation (NAF)......................135–159 Percoll............................................ 191–192, 195–196, 198, 201–203, 209, 224–225, 228–229, 236, 329, 331–332, 337 sucrose..............................3, 192, 196, 198, 202, 224–225, 228–231, 234, 237, 244–246, 248–251, 254–257, 260–261 Gradient gel electrophoresis.......................... 4, 6–10, 15, 21, 23–26, 29, 46, 270 Green fluorescent protein (GFP)....................... 70, 202, 231
H Heme or haem......................................... 118, 125, 358–359, 364–366, 377–380, 384 Hierarchical clustering..............104, 182, 266, 267, 278, 279 High-performance liquid chromatography (HPLC)..................................100, 136, 150, 152, 154, 157, 162, 166, 173, 174, 179, 183, 245, 252, 285, 289, 293, 360–364, 368–372, 375–377, 381, 383, 393, 403, 413, 415, 416, 419–421, 423, 424 Homogenization......................... 24, 33, 35, 36, 47, 136–137, 145–147, 158, 195, 213–215, 218, 220, 228–229, 236–237, 243, 248, 256, 260, 359, 366, 381, 383, 394, 397, 398 Homogenizer glass...............................211, 214, 215, 390, 391, 394, 397 oscillating ball mill.............................. 137, 146, 147, 158 pestle (and mortar, or microtube).......................... 37, 41, 119, 121, 158, 167, 177, 211, 214, 215, 225, 270, 285, 287, 319, 359, 366–368, 371, 373, 377, 381, 392, 398, 416, 417, 420 Potter.............................211, 214, 215, 225, 229, 237, 390 rotor-stator (Polytron, Ultra-Turrax)............................21 Waring blender...................................191, 195, 225, 228, 243, 248, 329, 331, 392, 398 Yeda press....................................210, 212, 213, 217, 220
Horseradish peroxidase (HRP)....................... 205, 227, 365, 377–381 Hybridization..........................................6, 12, 123, 131, 234
I IgG beads (IgG sepharose)................... 33, 35–37, 39–47, 72 Image acquisition and/or analysis.................... 55, 59–62, 64, 175–177, 180–181 Immunoblot or Western blot................................. 4, 5, 7, 11, 14–17, 27, 33, 47, 48, 71, 72, 193–195, 197–200, 203, 204, 224, 226–227, 230, 233–234, 365, 379, 380 Immunoprecipitation (coimmunoprecipitation)................52, 67, 69, 71–74, 209, 274 Infrared gas analyser (IRGA).......................... 312, 315–317, 319, 321, 323–326 In-gel digestion (trypsin)........................... 35–36, 39, 44–45, 96, 108, 247, 258, 260, 266–267, 269, 272, 280 Insertion mutants............................................... 98, 162–163 Intermembrane space.................................................33, 190 In vitro translated proteins.....................................72, 74–76 Iodine.................................162, 164, 165, 168, 182, 388–390, 393–394, 399, 400, 405, 408, 414, 418, 422, 423 Isothermal titration calorimetry (ITC)...................68, 81–83
L Laemmli buffer.................................23, 26, 28, 29, 193, 246, 268, 270 Light harvesting chlorophyll protein, or complex (LHCP, LHC)............................4, 20, 27, 28, 83, 85, 127, 194, 198, 203, 208, 216, 227, 234, 299, 319, 358, 359 Lipids extraction and analysis............................... 146, 162–163, 166–167, 169, 174–175, 411–424 galactolipids................................................................416 glycerolipids.................190, 411–414, 416–418, 421–423 prenylquinols.............................................. 413, 419, 423 Luciferase (luciferin)................................ 344–349, 353, 354 Lugol...........................165, 172, 390, 392–394, 399, 400, 406 Lumen..................................33, 190, 204, 207–221, 332–334, 337, 338, 344, 346 Luminometer........................................... 344, 347, 348, 379 Lyophilisation of leaf material.................. 137, 145–147, 158
M Macrocarrier........................................................... 55, 58, 59 Mascot........................................... 98, 101, 103, 106, 110, 255, 258, 260, 266, 269, 275, 287, 292 Mass spectrometry..........................33–36, 45, 47, 71, 94–96, 98, 103, 105, 107–109, 135–137, 141–142, 144, 150, 152, 154–155, 162, 173–174, 179, 183, 191, 209–210, 230, 247, 252–255, 258–261, 265–269, 272–281, 284, 287, 292–296
Chloroplast Research in Arabidopsis 430 Index
Metabolite analysis...................135–159, 162–169, 172–175, 179–180 Metabolomics...........................................................135–159 Microarray.............................95, 112, 118–120, 123–126, 131 Microcarrier............................................................ 58, 59, 63 Microscopy BiFC���������������������������������������������������������������� 51–64, 71 confocal................................................ 60, 202, 230, 231 electron......................................................... 33, 156, 242 epifluorescence..............................................................55 FRET.......................................................... 51–64, 71, 86 Morphology chloroplast...........................163, 167, 170, 175–177, 182 seed..........................................................163, 168, 178–179 whole plant......................................... 162, 165, 170–172 Murashige and Skoog (Murashige–Skoog, MS medium)...................36, 39, 54, 58, 328, 330 Mutagenesis.......................................................................73 Mutant.............................. 4, 62, 68, 69, 76, 81, 98, 118–120, 124–126, 129, 130, 132, 136, 157, 161–183, 260, 265–281, 302, 304–306, 309, 312, 320, 346, 382, 388, 400, 404, 407, 408
N NASC. See Nottingham Arabidopsis Stock Centre National Center for Biotechnology Information (NCBI)............................................. 95, 103, 109 Native electrophoresis (native PAGE)............... 3–17, 19–29, 67, 209, 273 Non-aqueous fractionation (NAF)...........................135–159 Northern blot.............................................................12, 123 Nottingham Arabidopsis Stock Centre (NASC).............123
O Oxygen (O2) electrode...............312–315, 317, 319, 323–325
P PCR genotyping.................................. 163, 167–171, 177–178 quantitative (real-time)...................... 120, 123, 126–130, 132–133 RT-PCR............................................. 120, 126–130, 132 Percoll cushion............................................... 392, 398–399, 408 gradient.......................................191–192, 195–196, 198, 201–203, 209, 224–225, 228–229, 236, 329, 331–332, 337 pH change..........................288, 339, 345, 347–348, 351–352 DpH (delta pH).................................. 327–340, 344, 352 gradient.......................................................................252 Phenomics . ..............................................................161–183 Phosphoproteomics....................95, 108, 109, 111, 242–243, 283–296
Photosynthesis gas exchange.......................................................311–326 measurement......................164, 169, 180–181, 299–309, 311–326 photosynthetic assimilation....................... 162, 302, 315, 321–323, 388 photosynthetic efficiency................... 164, 169, 180–181, 299–309, 311–326 photosynthetic pigments............ 334–335, 346, 357–384 photosynthetic protein complexes..................... 4, 19–29, 51, 117, 210, 299, 343, 358, 373 Photosystem................................. 4, 13, 20, 27, 28, 117, 210, 300, 307, 324, 334, 339, 358 Phylloquinol..............................413, 415, 419, 420, 423, 424 32 P incorporation...................................... 345, 347–351, 354 Plant growth conditions................................. 36, 39, 54, 119, 120, 162, 180, 210, 224, 287, 318, 328–331, 408, 412 medium. see Murashige and Skoog Plant Organelle Database 2 (PODB2)....... 95, 101–102, 111 Plant Proteome Database (PPDB).................... 96, 100–101, 111, 267 Plastid Protein Database (plprot)....................... 99–100, 111 Plastoquinol............................................. 413, 416, 420–421, 423, 424 Porphobilinogen (PBG)................................... 360–361, 367 Porphyrins ................................................................357–384 Posttranslational modification (PTM)...................... 80, 100, 242, 280, 283–296 Predotar............................................................................... 101 Preparation (purification, isolation) of chloroplasts.................191–192, 195–196, 198, 201–203, 209–211, 213–214, 218–220, 224–225, 228–229, 236, 329, 331–332, 337 DNA.................... 120, 128–129, 163, 167–168, 177–178 envelope membranes............189–205, 224, 230–231, 237 plastoglobules.....................................................223–238 ribosomes............................................................241–262 RNA.....................................119–123, 131, 251–252, 260 stroma.................................192, 196, 198–199, 202–203, 207–221, 224–225, 229, 231–232, 237 thylakoid lumen..................................................207–221 thylakoid membranes..........................................207–221 thylakoids...........................207–221, 224–225, 229–231, 234, 237, 329, 332, 337, 346 preSSU....................................................................72, 84–85 Protease inhibitors......................................5, 6, 37, 192, 196, 202, 225, 235, 268, 278 Protease Clp................................................................ 14, 266, 277 TEV...................................................... 32–36, 38, 42–47 Protein complexes or multiprotein complexes......................3–17, 19–29, 31–48, 117, 209–210, 241–262, 267, 299, 343, 373
Chloroplast Research in Arabidopsis 431 Index
degradation.................................................................228 extraction................................ 37, 41–42, 47, 94, 96, 108, 139–141, 151–152, 251–255, 257, 268, 270, 278, 284–285, 287–288, 293–294 import ..................... 51, 62, 68, 70–71, 81, 125, 190, 209 interactions......................................... 51–64, 67–87, 112 phosphorylation..............................21, 28, 101, 106–111, 242–243, 283–296 precipitation..................................38, 43–44, 48, 96, 226, 230–232, 255–257, 287, 294 quantification......................................37, 39, 42, 47, 108, 203, 211, 215, 217, 221, 251, 268, 270, 278, 285, 287–288 targeting prediction.................................... 100–103, 111 topology........................................................................96 Protein A (ProtA)....................................... 32, 33, 47, 71–72 Protein phosphorylation databases P3DB ......................................................... 108–109, 111 PhosPhAt........................................... 101, 106, 107, 111 Proteomics cross-contamination.................................. 190, 197–199, 202–204, 213, 224, 250–251, 254 databases...............................93–112, 209, 267, 274–276, 284, 292–293, 296 envelope.................................................. 96–97, 189–205 lumen . ..............................................................207–221 plastoglobule.......................................................223–238 post-translation modification............................100–101, 106–111, 283–296 quantitiative...................93–112, 190, 197, 199, 265–281 ribosome.............................................................241–262 spectral counts (spectral counting).........................96–98, 110, 199, 203, 265–281 stroma..................................................96–97, 101, 207–221 thylakoid................................. 96–97, 100–101, 207–221 Protochlorophyllide (Pclide).................... 362–363, 370–373, 383–384 Proton electrochemical gradient....................... 327–328, 343 Protonmotive force (pmf )........................................328, 333 Protoplast......................................................... 57, 63, 70, 71 Pull-down assay................................................ 67, 69, 71–74 Pulse-chase (pulse labelling)............................. 389, 396, 407
R Radiolabeling................................4, 71–72, 74–76, 345, 347, 349–351, 366, 388, 396–398, 407 Redox.........................................125, 327, 328, 358, 413, 424 Reticulocyte lysate..............................................................71 Retrograde signalling................................................117, 118 Reverse genetics........................................................161, 163 Ribosome (ribosomal) other....................................................................117, 203 proteins............................................... 241–262, 278–279 proteomics.......................................... 241–262, 278–279
PSRP...........................................................................242 purification.........................................................241–262 RNA (rRNA)......................123, 126, 131, 251, 252, 260 RNA analysis...............................................................117–133 editing ........................................................................112 interference (RNAi)............................................112, 118 messenger (mRNA)....................................................242 preparation, extraction.........119–123, 131, 251–252, 260 ribosomal (rRNA)...............123, 126, 131, 251, 252, 260 Robin.........................................119, 120, 123–125, 131–132 Root(s).............................................. 104, 157, 312, 384, 407 Rubisco................................ 13, 72, 84–85, 96, 105, 157, 198, 208, 209, 216, 251, 254, 271, 315, 318, 321, 322, 359
S Salk (SALK) lines....................................................168, 178 SDS-PAGE............................ 4–7, 11–16, 19, 23–24, 26–29, 34–36, 46, 72, 74, 94, 96, 108, 192–193, 197, 198, 204, 216, 226, 230–234, 245, 246, 251– 254, 257, 266, 267 Sec......................................................................................... 70 Seed amino acids..........................163, 168–169, 179, 181–183 C and N...................................... 163, 168–169, 179–181 morphology................................ 163, 168–169, 178–179 proteome..............................103–104, 106–108, 110–111 sowing.................... 39, 169–170, 191, 227–228, 330–331 starch............................................................. 164, 168–180 sterilization.........................................................330–331 Solubilization (-isation)..............................4–5, 7, 10, 13–15, 19–20, 23, 25–29, 33, 36, 41, 46, 47, 72, 81, 193, 197, 210, 218, 221, 268, 270, 391, 397 Sonication.................. 136–138, 145, 147–148, 237, 247, 258 Southern blot......................................................................12 spectinomycin...............................................................54, 59 Spectral counts (spectral counting)...................... 96–98, 110, 199, 203, 265–281 Split-ubiquitin..............................................................68–70 SRP (spSRP)................................................................83–85 Starch amylopectin.................387–390, 392, 394, 398–400, 404 amylose........................ 388, 389, 392, 394, 398–400, 408 analysis...............................................................387–409 biosynthesis rate......................... 388, 391–392, 396–398 chain length................................ 389, 392–393, 400–404 degradation, turnover...162, 388, 390–392, 396–398, 407 grain (granule).... 182, 201, 236, 387, 392, 398–400, 403, 405–406, 408 granule preparation..................................... 392, 398–399 phosphate............................390, 393, 398, 403–405, 409 staining....................... 162, 164, 165, 168, 171–173, 180, 182, 388–390, 392–394, 399–400, 405, 406, 408
Chloroplast Research in Arabidopsis 432 Index
Stock centre ABRC...................................................................98, 169 NASC.........................................................................123 RIKEN...........................................................98–99, 110 Stroma............................ 29, 33, 70, 73, 96, 97, 101, 150, 157, 190, 194, 196–199, 201–204, 207–221, 229, 231, 232, 234, 237, 267, 344, 412 SUBA (SUBA II).....................................................102, 111 Sucrose cushion...............................................................243, 248 gradient...........................................3, 192, 196, 198, 202, 224–225, 228–231, 234, 237, 244–246, 248–251, 254–257, 260–261 Surface plasmon resonance (SPR) resonant mirror.............................................. 68, 69, 83–85 SYBR Green.................................................... 120, 129, 132 SYPRO Ruby............................................27, 38, 44, 48, 280 Systems biology........................................................161–183
T TAIR.......................96, 98, 101–103, 106, 112, 123, 125, 132 Tandem affinity purification (TAP)................. 31–48, 67, 72 TAP tags.............................................................................32 TargetP......................................................................100, 101 T-DNA....................................................... 98, 163, 168, 178 TEM....................................................................................... 156 Tetrapyrroles biosynthesis........................................ 118, 125, 357–384 quantification and analysis..................................357–384 Thylakoid DpH .................................................. 327–340, 344, 352 electric field, potential........................ 327–340, 344, 346 energetics............................................ 327–340, 343–354 isolation............................................. 207–221, 224–225, 229–231, 234, 237, 329, 332, 337, 346 lumen............................................33, 190, 204, 207–221, 332–334, 337, 338, 344, 346 membrane.......................................10, 19–29, 33, 70, 83, 100, 162–163, 189–190, 192, 194, 199, 203, 207–221, 224, 227, 230, 313, 327–340, 343–354 protein transport........................70, 83–85, 333–334, 337 Tocochromanols........................413, 415, 417–420, 423–424 TOC/TIC TIC................................................................... 4, 72, 190 Tic proteins................................................ 14, 70, 72–73 TOC..............................................4, 72, 74, 81, 190, 227 Toc proteins.............................. 14, 70–72, 74, 76, 79, 81, 84–85, 227, 234
Topology, protein................................................................96 Transcript..................................112, 118, 126, 132, 242, 357 Transcription.................................70, 75, 118, 127, 131, 190 Transcriptomics........................................ 102, 112, 117–133 Transfection........................................................................57 Transient expression......................................... 54–55, 57–60 Transit peptide...................................62, 72–73, 84–85, 101, 208, 275–276 Transit peptide prediction................................ 100–103, 111 Translation............................................75, 77, 190, 241, 242, 251, 278–279 Transmembrane helices/domains...................... 70, 74, 84,96 Transmission electron microscopy (TEM).......................156 Transposon.........................................................................98 Trizol........................................................................... 121, 252 Trypsin............................................. 15, 39, 96, 108, 247, 258, 266, 269, 272, 274, 280, 285, 288, 294 Trypsin in-gel digestion......................................... 35–36, 39, 44–45, 96, 108, 247, 258, 260, 266–267, 269, 272, 280 Two-dimensional polyacrylamide gel electrophoresis (2D-PAGE)............................... 3–17, 19–29, 94, 100, 103–104, 107–111, 204, 207–208, 210, 212–213, 217–219, 221, 245, 252–253, 255
U Ultracentrifugation (ultracentrifuge).............................6, 10, 13–14, 33, 37, 41, 47, 68–69, 80–81, 192, 211–212, 215, 217, 225, 229–230, 234, 237, 243–244, 246, 249, 256, 261 Ultrasonication..................................136–138, 147, 247, 258
W Western blot (or immunoblot)..........................4, 5, 7, 10, 11, 14–17, 27, 33, 47, 48, 71, 72, 193–195, 197–200, 203, 204, 224, 226–227, 230, 233–234, 365, 379, 380 Wheat germ.................................................................71, 75
Y Yeast two-hydrid....................................................52, 67–70 Yeda press........................................................... 210, 212, 213, 217, 220 Yellow fluorescent protein (YFP)............... 53–58, 60–62, 64 YFP. See Yellow fluorescent protein
Z ZipTip .............................................269, 273, 286, 290–293
