Membrane structural biology: with biochemical and biophysical foundations

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STRUCTURAL BIOLOGY WI H BlOCH M CAL AND BIOPHYSICAL FOUNDATIONS

MARY LUCKEY San Francisco State University

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CAMBRIDGE UNIVERSITY PRE S

CAMBRIDGE UNIVERSITY PRESS

Cambl'idge, Ne\\' York, Melbourne, MilclT-iu, Cape Town, Singapol'e, Sao Palllo, Delhi Cnnlbridgc University Pres~

32 Avenue or the America" New York, NY 10013-2473, USA \\·ww.calnbridge.org Jn~ormalion on

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o Man' LuckeI' 2008 This publication is in copvrighl. Subjeci to statuton exception and to the provisions of relevant collecrive licensing agl'cemenls, no reproduction or anv pan mil\' take place without the wrillen permission of Cambridge Universilv Press. First published 2008 PI'inted in Canada bv Friesen, rl cala/o,~ ,.cco,.d {()/' Ihis puhlicrllioll is nvai!a!JIc {iOiIl Ihe B,./Ii,/, Lihron·.

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Luckev, Mal'." Membl'ane structural biology: \\'itb biochemical anel bioflh~'sical foundations I Marv Luckev. p,; cm. Includes bibliographical rderences and index. ISBN 978-0-521-85655-3 (hardback) I. Membranes (Biology) 2. Membrane lipids. 3. IVlembrane proteins. 1. Title. [DNLM: l. Cell Membrane - phvsiologv. 2. iV1cmbrane Lipids - phvsiologv. 3. Membl'ane Proteins - phvsiologv. QU 350 1..941 m 2008] QH60l.L75 2008 571.6'4-dc22 2007031145 ISBN

978-0-521-85655-3 hardback

Cambl'idge University Press has no responsibilitv for the persistence or accuracy of URLs for external or third-partv Internet Web sile, refern::d to in thi, publication and doe~ not gu(\rantee that al1,v content on .such

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The title page shows high-resolution stl1.JClures of membrane proteins incorporated into a simulated lipid bilavel'. The proteins are, fl'Om left to r'ight: vitamin B '2 transponel's BlLrCD with BtuF, the light harvester LH2 with some chlorophylls, the mechanosensilive channel MscS, lactose permease, BtuB from the outer membrane, rhe pore domain of Kv1.2, aquilporin. ilnd Cal, -ATPase. Substr
Contents in Brief

Preface

Xl

1

Introduction

2

The Diversity of Membrane Lipids

13

3

Tools for Studying Membrane Components: Detergents and Model Systems

42

4

Proteins in or at the Bilayer

68

5

Bundles and Barrels

102

6

Functions and Families

127

7

Protein Folding and Biogenesis

160

8

Diffraction and Simulation

191

9

Membrane Enzymes and Transducers

213

10

Transporters and Channels

241

11

Membrane Protein Assemblies

271

12

Themes and Future Directions

309

Appendix I: Abbreviations

315

Appendix II: Single-Letter Codes for Amino Acids

318

Index

319

Contents

Preface

page xi

1 Introduction General Features of Membranes ParacUgm 1: The AmphiphiJic Molecules in Membranes Assemble Spontaneously due to the Hydrophobic Effect Paradigm 2: The Fluid Mosaic Model Describes the Membrane Structure A Shift in the Paradigm: Biomembranes Have Lateral Domains that Form "Rafts" A View for the Future: Dynamic Protein Complexes Crowd the Membrane Interior and Extend Its Borders

4 5

8

9

2 The Diversity of Membrane Lipids

13

The Acyl Chains Complex Lipids Phospho) i pids Sphingolipids Sterols and Linear Isoprenoids The Lipid Bilayer Matrix Structure of Bilayer Lipids Diffusion of Bilayer Lipids BOX 2.1 Fluorescence techniques Lipid Asymmetry and Membrane Thickness Lipid Polymorphism Lamellar Phase Hexagonal Phase and the Amphiphile Shape Hypothesis Cubic Phase Miscibility of Bilayer Lipids BOX 2.2 Phase diagrams Lateral Domains and Lipid Rafts Detergent-Resistant Membranes Diversity of Lipids BOX 2.3 Nonlamellar phase lipids and growth of E. coli

13 17 17

20 20 23 24 24 26 26 28 28 29 31 31 31 33 36 37 40

3 Tools for Studying Membrane Components: Detergents and Model Systems

42

Detergents Types of Detergen ts BOX 3.1 Surfactants and surface tension Mechanism of Detergent Action Membrane Solubilization Lipid Removal Model Membranes Monolayers Planar Bilayers BOX 3.2 Electrophysiology Patch Clamps Supported Bilayers Liposomes from SUVs to G Vs Mixed Micelles and Bicelles Blebs and Blisters Nanodiscs

43 43

4 Proteins in or at the Bilayer

68

Classes of Proteins that Interact with the Membrane Proteins at the Bilayer Surface Extrinsic/Peripheral Membrane Proteins Reversible Interactions of Peripheral Proteins with the Lipid Bilayer BOX 4.1 Binding of ligands to surfaces Proteins and Pep tides that Insert into the Membrane Toxins Colicins Peptides SecA: Protein Acrobatics Proteins Embedded in the Membrane Monotopic Proteins Integral Membrane Proteins Protein-Lipid Interactions BOX 4.2 Electron paramagnetic resonance Hydrophobic Mismatch

43 45 48 50 50 50 53 53 55 57 60 63 63 66

68 69 69 76 79 84 84 85 87 88 90 90 90 94 97 98

5 Bundles and Barrels

102

Helical Bundles Bacteriorhodopsin Photosynthetic Reaction Center I3-Barrels BOX 5.1 NMR determination of membrane protein structure Par'ins Specific Porins fron Receptors

102 102 107 113 116 118 120 123

6 Functions and Families

127

Membrane Enzymes BOX 6.1 Surface dilution effects Diacylglycerol Kinase P450 Cytochromes Transport Proteins Transport Classification System Superfamilies o( ATPases ABC Transporter Superfamily Group Translocation Symporlers Antiporters Ion Channels Membrane Receptors Nicotinic Acetylcholine ReceptOlG-Protein Coupled Receptors Bioinformatics Tools for Membrane Protein Families Predicting TM Segments BOX 6.2 Bioinformatics basics Hydrophobicity Plots Orientation 01" Membrane Proteins The Positive-Inside Rule BOX 6.3 Making and testing hydrophobicity plots Genomic Analysis of Membrane Proteins BOX 6.4 Statistical methods for TM prediction Helix-Helix Interactions Proteomics of Membrane Proteins I3-Barrels

127 128 129 130 131 132 134 134 135 137 138 138 139 \39 140 141 141 142 143 143 144 144 145 148 154 156 157

7 Protein Folding and Biogenesis

160

Protein Folding Folding ex-Helical Membrane Proteins BOX 7.1 Energetics of folding and insertion a hydrophobic a-helix into the bilayer Folding Studies of f3-BarrcllVlembrane Proteins Other Folding Studies

161 162

164 J67 169

Biogenesis of Memb.·ane Proteins Export from the Cytoplasm BOX 7.2 Evidence for cleavable signal sequences involved in protein translocation BOX 7.3 Import of mitochonddal proteins Integration of Nascent Proteins into the Membrane BOX 7.4 Cross-linking traces nascent peptides through the translocon into the bilayer Topogenisis in Membrane Pmteins Misfolding Diseases

170 170

171 177 178

179 184 187

8 Diffraction and Simulation

191

Back to the Bilayer Liquid Crystallography BOX 8.1 X-ray and neutron scattering Liquid Crystal Theory Joint Refinement of X-Ray and Neutron Di f[i'action Data Modeling the Bilayer Simulations of Lipid BiJayers Molecular Dynamics BOX 8.2 Molecular dynamics calculations Monte Carlo Lipids Observed in X-Ray Structures of Membrane Proteins The Crystallographer's Art Membrane Simulations

19\ 192 193 193

9 Membrane Enzymes and Transducers

213

Enzymes OMPLA Prostaglandin Hz Synthase BOX 9.1 Mechanism of action of prostaglandin Hz synthase Formate Dehydrogenase Transducers Rhodopsin. a GPCR BOX 9.2 Efficiency of light-induced signal transduction BOX 9.3 Numbering TM helices Mechanosensitive Ion Channels

214 214 216

10 Transporters and Channels Transporters LacY and GlpT Mitochondrial ADP/ATP Carrier Channels Aquaporins and Glyceroaquaporins Potassium Channels Calcium ATPase

194 196 196 197 J98 202 203 207 210

218 222 226 227 229 231 235 241 241 242 249 253 253 258 264

Contents

ix

11 Membrane Protein Assemblies

271

F1Fo-ATPase/ATP Synthase Subunit Structure and Function Regulation of the FIFo-ATPase Catalytic Mechanism of a Rotary Motor Complexes of the Respiratory Chain Cytochrome bCI Cytochrome-c Oxidase The Translocon The M. jannaschii Translocon Structure The Translocon-Ribosome Complex ABC Transporters and Beyond The Vitamin B 12 Uptake System Transport across the Inner Membrane Transport across the Outer Membrane Drug Efflux Systems Sav1866, an ABC Multidrug Transporter EmrE, Small but Powerful

272 273 276 277 279 279 284 286 286 288 290 291 291 294 296 297 299

Tripartite Drug EFnux via a Membrane Vacuum Cleaner AcrB, a Peristaltic Pump AnA, a Membrane Fusion Protein TolC, the Channel-Tunnel

300 300 302 304

12 Themes and Future Directions

309

OJigomeriza tion Conformational Changes Motifs and Patterns Conclusions

309 310 3I I 312

Appendix I: Abbreviations

315

Appendix II: Single Letter Codes for Amino Acids

318

Index

319

Preface The tremendous progress made over the last decade in our understanding of biomembranes calls for a new gestalt in a book about their structure and function. The need for such a book was apparent as I labored to capture the explosion of information about the structure and organization of biological membranes for my course on membrane biochemistry. Applications of new techniques and whole new methodologies have changed both how we acquire knowledge of the membrane and how we view it. For many years, the difficulties in crystallization of membrane proteins caused a scarcity of structural detail. Now sophisticated diffraction analysis allows description of fluid lipid bilayers, and highresolution structures have been determined for a variety of membrane proteins. Each new high-resolution stmcture of a membrane protein that graces a journal cover offers new insights into membrane functions. And yet, a full understanding of each new structure and its lipid environment is built on foundations of membrane biochemistry that derive from basic physical and life sciences. This book combines a physicochemical analysis of the membrane milieu with the latest structural biology on membrane lipids and proteins to give an exciting portrayal of biomembranes. The book's title, Membrane Siruciural Biology, emphasizes the successes of structural biology in revealing exciting details of many membrane components. To see the impact of structural biology on biochemistry textbooks, one need only compare a biochemistry book from 25 years ago with a current textbook, in which colorful and detailed molecular structures illustrate the functions of biomolecules and mechanisms of complex biochemical processes. A textbook on membrane biochemistry can only now approach that transformation to molecular detail, drawing from x-ray crystal structures, lipid diffraction and liquid crystallography, and computational modeling. To cover these advances and their foundations in one comprehensive volume, this book moves from basic membrane biochemistry to detailed examples of membrane structural biology. It includes numerous new topics, such as phase diagrams of lipid raft mixtures, reconstitu tion using bicelles and nanodiscs, binding domains of amphitropic proteins, effects of elasticity on folding of membrane proteins, a biological scale for identification of transmembrane helices, bioinfonnal ics and proteomics of membrane proteins, and joint refinement of x-ray and neutron diffraction data for lipid bilayers. It offers explana tions of techniques as varied as statistical methods for prediction of transmembrane sequences, surface effects in binding and kinetics, protein rolding

studies, liquid crystal theory, and molecular dynamics simulations. Written at a level appropl-iate for advanced students and scientists new to the field, Membrane Structural Biology assumes a background familiarity with the concepts covered in an undergraduate biochemistry course. Although il includes a wide range of material in a broad and rapidlv moving field, the book is not encyclopedic. Nor does it provide the thousands of references to the scientific papers on which it is based. Thal literature is vast, but fortunately it is readily accessible with the search engines now available. Readers who want to Jearn more can get started with the key references to seminal papers and reviews provided for each chapter. They can study the papers cited in the figure captions and ca n easily search for other contributions h'om those authors. A thorough familiarity with the examples described in the book will provide the reader with a solid foundation for f~urther studies, including the exploration of other important topics, such as membrane fusion, chemotaxis, endocytosis, and membrane recycling. It is a challenge to cover the full scope of this burgeoning field, including new methodologies and latest developments. Though it is mv hope that the numberof typographical and/or factual errors in this text will be small, I welcome the readers of this first edition to send me their corrections so that they may be incorporated into the next edition. I want to acknowledge assistance when I started writing the book [Tom my student, Aram Krauson, along with early feedback I received [TOm Professor JUI-g Rosenbusch. For their comments on specific topics I thank Professors Scott Feller, Steve White, Sam Hess, Rosemary Cornell, Ehud Landau, David Hackney, Paula Booth, Bill Plachy, and Hiroshi Nikaido. I am especially grateful to those who reviewed the entire manuscript: Professors Lin Randall and Stanlev Parsons, and former studenls Shyam Basharam, Marla Melnick, and Jared Matt Greenberg. I thank Andrea Dose for library help, J. C. Gumbart for the model membrane figures and graphic artist Diane Fenster of SFSU for the design of the publicity postcard, which led to the cover design. I appreciate how diligently Mary Paden and her staff worked on the production. For her unflagging enthusiasm and wise editorial help, I thank Dr. Katrina Halliday. Thanks as well to my colleagues and friends who supported my progress writing the book. Finally, I deeply appreciate the patience and encouragement 1 received from my family, Paul, Ariel, Amanda, SAM, and Ryan. Mary Luckey I [email protected]

1

Introduction

Essential for the compartmentalization that defines cells and organisms, biomembranes are fu ndamentalto life. Early membranes played a crucial role in the origin of life as the structures that defined what stayed in and what was kept out of primordial cells. In addition to their compartmentalization function, membranes provide modern cells with energy derived from chemical and charge gradients, organize and regulate enzyme activities, facilitate the transduction of information, and even supply substrates for biosynthesis and for signaling molecules. Some membranes have specialized functions; for example, the brush border membrane lining the intestines absorbs nutrients, the myelin surrounding nerves functions as insulation, and the rod cell membrane of the eye captures light. Wh ile prokaryotes either have one cell membrane (Gram positive) or have inner and outer membranes in the cell envelope (Gram negative), eukaryotic cells have many membranes (Figure 1.1). In addition to the plasma membrane, eukaryotes have membranes surrounding the nucleus, organelles such as mitochondria, chloroplasts in plants, Iysosomes, and of course the membranebased endoplasmic reticulum (ER), Golgi apparatus, and other vesicles involved in intracellular transport. Even some viruses have membrane envelopes. In spite of this variety, much can be genera lized abou t the structure and function of biomembranes.

GENERAL FEATURES OF MEMBRANES

Biological membranes consist of lipids, proteins, and carbohydrates (Figure 1.2). The lipid components include glycerophospholipids (also called phospholipids), sphingolipids, and sterols. The basic unit of the membrane is a bilayer formed by phospholipids and sphingolipids organized in two layers with their pola.· headgwups along the two surfaces and their acyl chains forming the nonpolar domain in between. Embedded in the lipid bilayer are integral membrane proteins, which cannot be removed without disrupting the membrane. Most of these proteins have one or more transmembrane (TM) segments, and they interact closely with nearby lipids as well as other proteins. In addition, there are peripheral membrane proteins that associate at the surface of the membrane and lipidanchored proteins that are held into the membrane by covalenL1v allached fatty acids or lipids. Although carbohydrate membrane constituents serve important h.lnctions, these hydrophilic moieties are always on the portions of glycoproteins and glycolipids external to the membrane bilayer. Such glycoconjugates deserve detailed consideration on their own and are not covered in this book. Membranes are responsible for the selective permeability of cell envelopes that enables cells to take

Endoplasmic reticulum

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Cell membrane

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Basal bodv Ribosomes ~ c.

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Nucleus

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Cell wall Vacuole

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Endoplasmic reticulum Nuclear membrane Milochondrion

(a)

B.

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(b)

Ribosomes

Outer membrane Peptidoglycan layer / Inner memarane

I Ribosomes

Gram-negative bacteria Outer membrane; peptidoglycan layer (b)

Cell envelope

Gram-positive bacteria No outer membrane; thicker peptidoglycan layer (a)

(e)

1.1. A variety of types of membranes. A. Plasma membrane and intracellular membranes in eukaryotic cells are shown in a diagram based on thin section electron micrographs of generalized animal (a) and plant (b) cells illustrating the plasma membrane and membrane-bound organelles. Redrawn from Jain, M. K., and R. C Wagner, Introduction to Biological Membranes, 2nd ed., Wiley, 1988, p. 2. © 1998. Reprinted with permission from John Wiley & Sons, Inc. B. Bacterial membranes are shown in a diagram of a bacterial cell (a) and in thin section electron micrographs of the cell envelope of Gram-negative (b) and Gram-positive (c) bacteria. Redrawn from Nelson, D. L, and M. M. Cox (eds.J, Lehninger Principles of Biochemistry, 4th ed., W H. Freeman, 2005, p. 6. (Q 2005 by W H. Freeman and Company. Used with permission.

General Features of Membranes

3

Oligosaccharide chains of glycoprotein

Outside

Peripheral protein

Integral protein (single transmembrane helix)

Peri pheral protei n covalently linked to lipid

Integral protein (multiple transmem brane helices)

1.2. Membrane components. Membranes contain lipids, proteins, and carbohydrates as glycolipids and glycoproteins. Nelson, D. L., and M. M. Cox (edsl. Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 372. © 2005 by W. H. Freeman and Company. Used with permission.

up many nutrients and exclude most harmful agents. The permeability properties are determined by both lipid and protein components of membranes. In general, the lipid bilayer is readily penetrated by nonpolar substances while proteins in the membrane make channels and transporters for ions and hydrophilic substances. This permeability barrier enables the membrane to maintain charge and concentration gradients that are critical to the cell's metabolism. The permeability barrier is maintained during activities such as cell division and exocytosis because the membrane is flexible and self-sealing. Membranes are also very dynamic structures, with constant activity on their surfaces as well as constant movements in the bilayer, both in the transverse direction across the bilayer and the latera] direction in the plane of this two-dimensional matrix. The latter movements give rise to the fluid nature of the membrane and enable interactions among proteins and between proteins and lipids to provide temporal associations that are important to membrane functions. Thanks to many, many scientists who have contributed to the enormous progress of the past decades, knowledge of the membrane goes beyond its basic architecture and properties to a multitude of details describing specific elements and functions. While the

particular tools and approaches used by biochemists, biophysicists, geneticists, and ceJI biologists \vho study the membrane vary greatly, two paradigms~ provide the framewOl-k for understanding their work. The starting point for understanding membrane structure is the hydrophobic effect. A far-reaching paradigm for many areas of chemistry, this principle governs the behavior of membrane components. The specific paradigm for membranes is the Fluid Mosaic Model, a description of membrane properties and organization that has endured for more than thl-ee decades. A description of these paradigms and the classic work on which they are based will lay the groundwork for the rest of this book. Yet today the current paradigm is shifting because of new aspects of membrane organization that have risen to the forefTont in the past few years. The importance of transient, specialized regions called membrane rafts affects the contemporary model of cell membl-anes. Finally. the organization of many membrane proteins into large assemblies that often involve molecules at the bilayer periphery and beyond Purudigms ure scientifiC models. According to science philosopher Thomas Kuhn, the pamdigllls of u held of study shupe it so thol'Oughly thut thev Illav be unacknowledged and even unobserved bv its practitioners. Yel, they determine the assumptions and the tools with which those scientists operate dail.".

indicates that a more complex and comprehensive view is needed for fclture work.

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PARADIGM 1: THE AMPHIPHILIC MOLECULES IN MEMBRANES ASSEMBLE SPONTANEOUSLY DUE TO THE HYDROPHOBIC EFFECT

All biomembranes contain amphiphilic lipid and protein constituents that have both polar and nonpolar parts, and this dual nature of its components is essential to membrane structure. Because proteins are simply polymers of amino acids, their polarity is a [unction of their amino acid composition; thus they have hydrophobic domains rich in residues with nonpolar side chains and hydrophilic domains generally Jacking them. On the other hand, by classification a lipid is quite nonpolar because the definition of lipids is empirical: a lipid is a biological substance that is soluble in organic solvents and has poor solubility in water. Yet all lipids have hydrophilic domains, called their headgroups, even when the headgroup is simply a hydroxyl group, as in cholesterol. The structures o[ lipids vary considerably (as described in Chapter 2) but all provide the amphiphilicity" that leads to the formation of distinct phases in aqueous systems, in which the lipids aggregate spontaneously to form polar and nonpolar domains. Mixing a pure lipid with water can result in formation of monolayers, micelles, bilayel's, hexagonal arrays, or cubic phases, depending on the nature o[ the lipid and the method of preparation. The spontaneous formation of each type of lipidic aggregate depends on the structure and hydrophobicity of the lipid, but it is always driven by the structure of watel~ In ice each water molecule has four hydrogen bonds worth ~S kca\l111ol each (Figure 1.3). When ice melts ~8S% of these hydrogen bonds arc preserved, but of course in Iiquid water they are dynam ic, with lOll/sec positional changes. The extensive hydrogen bonding of ,vater accounts for its special properties, such as its high boiling point and high dielectric constant (a measure of the extent to which it shields dissolved ions). It also provides the basis for the hydrophobic effect. Insertion of a nonpolar molecule, such as a falty acid with a long acyl chain, into liquid water reorders the water molecules closest to the hydrocal-bon chain to form a hydrogen-bonded cage around the nonpolar moiety. Depending on the size of the nonpolar domain, there may be no net loss or hydrogen bonds so enthal py does not necessarily have a strong effect. However, as the water molecules rearrange to form the cage around the nonpolar chains, their mobility is drastically reduced, resulting in a large loss of entropy. The best way to lower this entropic cost is to sequester the Aml'hi/Jhilicitv means "having po 1<11' and nonpolal' domains"; (1IlIphiphilic
1.3. Importance of hydrogen bonding in the structure of water In ice, each water molecule forms four hydrogen bonds with its nearest neighbors. In liquid water at room temperature and atmospheric pressure, each water molecule has on average 3.4 hydrogen bonds. Redrawn from Nelson, D. L., and M. M. COX (eds.), Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman,

2005, p. 49.

nonpolar moieties into large aggregates, thus reducing the total surface area of nonpolar material exposed to the aqueous layer and hence decreasing the number of immobilized water molecules. (This is possible because as a sphere increases in size, the volume increases as the cube of the radius while the surface area increases as only the square of the radius, with the resull that a larger l-adius gives a smaller surface area-to-volume ratio.) The end result of this entropic driving force is the separation of the aqueous and lipid molecules into two phases or domains. The nonpolar domain may then be further stabilized by van der Waals forces between the close-packed acyl chains. The hydrophobicity of a substance is traditionally measured by a partitioning experiment using two solvents, such as heptane and water. From the partition coefficient is calculated the .0.G"nn,rer for the solute of interest: Kp

= Kc q = [solute]H2o/[soluteJhepwne .0.G".

=

-RT In Kcq ,

where K p is the partition coefficient, K eq is the equilibrium constant, and .0.G lI is the free energy change for the transfer from heptane to water. When the solutes are fatty acids with varying chain lengths, the energy cost is proportional to the chain length: a cost per CH 2 unil of 0.8 kcal/mol is derived

Paradigm 2: The Fluid Mosaic Model Describes the Membrane Structure

In 1935 Davson and Danielli used thermodynamic arguments along wi th measurements of surface tension and permeabilitv to postulate a membrane structure that placed globular proteins on the outer surfaces of a membrane bilayer (Figure 1.5A). This model dominated

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0$ ~

I

5

Davson-Danielli model

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u

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.... <.J

~

-6


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Lipid

4

8 12 16 2022 Number of Carbon Atoms

1.4. The free energy of transfer of fatty acids from water to heptane is a function of the chain length. Fatty acids of varying lengths in n-heptane at 23°C to 25°C are equilibrated with dilute aqueous buffer and their activities (lJO) in each phase determined. The x-axis gives the number of carbon atoms, and the y-axis gives the free energies for transfer. Redrawn from Tanford, c., The Hydrophobic Effect: Formation of Micelles and Biological Membranes, 2nd ed., Wiley, 1979, p. 16. © 1979. Reprinted with permission from John Wiley & Sons, Inc.

B Robertson's unit membrane Protein

from the plot of 6G", versus the chain length (Figure 1.4). Like other structures in biology, the aggregate structures of lipids are stabilized by the cooperative su m of many weak interactions. Thus the thermodynamic stability of the membrane bilayer maximizes waterwater interactions outside and acyl chain interactions inside the nonpolal- interior while minimizing wateracyl chain interactions that are en tropically expensive. The hydrophobic effect explains the energetics of membrane formation but does not address the basic structure of the biological membrane.

Lipid

Protein C.

Benson-Green subunit model

PARADIGM 2: THE FLUID MOSAIC MODEL DESCRIBES THE MEMBRANE STRUCTURE

While the Fluid Mosaic Model for the structure o[ membranes is now familiar to all life science students, the amazing unity it brought to a divided field is not apparent without an appreciation of its historical development. The earliest evidence for a lipid bilayer is attributed to Ben Franklin's calculation of the thickness of an olive oil film on pond water as 25 A (2.5 nm). Then in 1925 Gorter and Grendel made surface area measurements for a compressed monolayer formed by acetoneextracted lipid from erythrocytes. Theirconclusion that the monolayer area covered twice the surface area of the erythrocytes was correct, in spite of ex peri mental errors that offset each othel- I

1.5. Early models for the structure of biological membranes.

A. The Davson-Danielli membrane model with layers of globular proteins outside the lipid bilayer. © 1935. Reprinted with permission from Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc. B. The unit membrane proposed by Robertson had the protein as l3-sheets, still outside the lipid bilayer. '(;1 1966. Reprinted with permission of Blackwell Publishing. C. In contrast, the BensonGreen model for the mitochondrial inner membrane showed protein particles that are solvated by lipids and are readily fractionated into complexes. (c;~ 1983 by Academic Press. Reprinted with permission from Elsevier. A and B redrawn from Gennis, R. B., Biomembranes: Molecular Structure And Function, SpringerVerlag, 1989, p. 8. C redrawn from Aloia, R. c., Membrane Fluidity in Biology, vol. I, Academic Press, 1983, p. 119.

B.

A.

proteins ---- Split lipid bilayer / 1.6. Visualization of the distribution of proteins in membranes. A. The freeze-fracture technique reveals the interior of a biological membrane by splitting a frozen membrane sample with a cold microtome knife. Redrawn from Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, p. 405. © 2004. Reprinted with permission from John Wiley & Sons, Inc. B. Electron microscopy of an erythrocyte plasma membrane split by freeze fracture shows the inner surface of the membrane is studded with embedded proteins. From Voet, D., J Voet, and C. W. Pratt, Fundamentals of Biochemistry Upgrade Ed., John Wiley, p. 247. © 2002. Reprinted with permission of John Wiley and Sons and Vincent Marchesi, Yale University.

thinking about membrane structure for the next three decades, with modifications such as changing the protein conformation to extended f)-sheets, and led to the concept of a "unit membrane" with a width of 6 to 8 nm, corresponding to the width of myelin sheath in x-ray diffTaction measurements (Figure 1.SB). In 19S9 Robertson argued that this unit mem brane was common to all biological membranes, citing "railroad track" images from thin section electron microscopy (EM) 01' tissues stained with osmium tetroxide, which stained the phosphates of phospholipid headgroups and washed proteins out (see Frontispiece). Other staining techniques in use atthe time, such as prior cross-linking with glutaraldehyde, produced images in which the full membrane was electron dense. A challenge to the Davson-Danielli-Robertson model came with the application of ft-eeze-fractuIT techniques: bumps visible by EM when the membrane was cleaved within the plane of the bilayer were attributed to embedded proteins (Figure 1.6). Suppor"t for the interpretation that the bumps were proteins came from their absence in membranes treated with proteases and in samples of myelin sheath, which has

very little protein. In studies of the respiratory chain of mitochondria by Benson and later Green, mitochondrial inner membrane could be separated into lipoprotein subunits and reconstituted to regain activity. These results supported a model in which the lipid is solvent for embedded, globular proteins, consistent with EM images obtained after negative staining with heavy metals that showed subunits (not "railroad tracks") that were unaffected by lipid extraction prior to staining. Thus the Benson-Green subunit model was the antithesis 01' the Davson-Danielli-Robertson model (Figure 1.Se). Today it is hard to realize the extent of controversy that occurred. As Singer and Nicolson wrote in J 972, "Some investigators who, impressed with the great diversity of membrane compositions and functions, do not think there are any useful generalizations to be made even about the gross structure of cell membranes ..." Of course, their now-classic paper on membrane structure did present a general model for the structure of biomembranes - the Fluid Mosaic Model which is included in every modern biochemistry and biology textbook (Figure 1.7). Their paper should be

Paradigm 2: The Fluid Mosaic Model Describes the Membrane Structure

7

1.7. The Fluid Mosaic Model proposed by Singer and Nicolson. The basic structure of the membrane is a lipid bilayer, with the fatty acyl chains from each leaflet forming a nonpolar interior. Intrinsic proteins are integral to the bilayer while extrinsic proteins are on its periphery. Redrawn from Singer, S J., and G. L. Nicolson, Science. 1972, 175:720-731.

read in full, for it provides a beautiful example of examining all the biomembrane's properties conducive to testing with available techniques and summarizing the results in a consistent model. In addition to the thermodynamic principles and EM results discussed above, Singer and Nicolson emphasized the lateral mobility of membrane components. Significant lateral diffusion of membrane proteins had been demonstrated in the elegant Frye-Edidin experiment that followed the mixing of surface antigens in cell fusion experiments (Figure 1.8), and the rates of diffusion of lipids in the plane of the membrane were being measured by fluorescence techniques (discussed in Chapter 2). Singer and Nicolson also described the limited transverse mobility of lipids and the lack of it for proteins; the permeability barrier provided by the membrane; the structure of membrane proteins based on circular dichroism, x-ray diffraction, and labeling experiments (revealing them to be a-helical, globular, and membrane spanning); the assays of certain enzymes that require lipids for activity; and the phase transitions detected \-vith differential calorimetry. Based on these results, their Fluid Mosaic Model puts forth simple principles: the bulk or the lipid forms the bi layer, wh ic h provides the solven t for em bedded proteins; most of the proteins are embedded and globular, termed intrinsic or integral membrane proteins. Some proteins are extrinsic (peripheral) as they can be removed by washes that change the pH or ionic strength. The bilayer, composed of two lipid layers, or leaflets, is fluid; in fact, it has the viscosity of olive oil,

which allows lateral mobility of lipids and some protein components. It is mosaic in that proteins are scattered across it or on its su rrace. Both lipids and integral memb.-ane proteins al-e amphipathic, allowing the nonpolar portions of proteins and lipids to interact and the polar portions of proteins and lipids to interact. Th is widely accepted model for membrane structu re is often a bbrevia ted as a pictu re of integra I proteins floating as icebergs in a sea of lipids, an oversimplification that denigrates the role of the lipids, whose diversity and polymorphic phases provide particular chemical activities as well as structural domains in that "sea," as the next section asserts. Furthermore, this simple picture obscures the wide variation in membrane composition (not overlooked in the original paper by Singer and Nicolson l ). As Table 1.1 shows, the proportion of membrane components varies from ~80% lipid and ~20% protein (myelin) to ~75% protein and ~25% lipid (mitochondrial inner membrane). A rough calculation for the mitochondrial inner membrane suggests that these membranes have on the order of 100 lipid molecules per protein. Because it requires at least 40 to 50 lipid molecules to form a single belt of lipid around a protein, clearly this is not enough lipid to solvate individual proteins and provide a "sea" in which they float. So how does the mitochondrial inner membrane fit the model? First, the total protein given in Table 1.1 includes peripheral proteins. In the mitochondrial inner membrane over half the proteins are peripheral, leaving much less embedded in the lipid bilayer. Second, the protein-protein interactions between integral proteins

exclude bulk lipid; thus the lipid solvates the respiratory complexes, not each individual protein. No wonder scientists who concentrated on this membrane argued strongly for the subunit model! While much additional work has contributed support for the Fluid Mosaic Membrane, the uniform mixing of bilayer lipids has been challenged by experimental observations of lipid heterogeneity based on the physical measurements of phase separations, as well as the detection of membrane domains wi th separate functions. Today there is wide acceptance of a shift in the paradigm that allO\.\Is membranes to have specialized microdomains called lipid rafis.

Mouse cell

Human cell

(a)

Sendai virus

I

TABLE 1.1. Composition of membrane preparations by percent dry weight" Source

Lipid

Protein

Cholesterol

30-50 15-30 60 20-25 30-40 15-40 60 20-25

5()....70 6()....80 40 7()....80 60-70 60-80 40 7()....80

20 6 10 <3 <5 10 8 14

6()....70 50 40 50

20-30 50 60 40

22 20 24 <3

Escherichia coli Bacillus subtilis

2()""3 0 2()""3 0

Chloroplast

35-50

70 70 5()....65

0 0 0

Rat liver Plasma Rough ER Smooth ER Inner mitochondria Outer mitochondria Nuclear Golgi Lysosomes Rat brain Myelin Synaptosome Rat erythrocyte Rat rod outer segment

• The percentages by weight of membrane preparations from various eukaryotic and prokaryotic sources are given. ER, endoplasmic reticulum. Source: Based on Jain, M. K., and R. C. Wagner, Introduction to Biological Membranes, 2nd ed. New York: Wiley, 1988, p. 34.

A SHIFT IN THE PARADIGM: BIOMEMBRANES HAVE LATERAL DOMAINS THAT FORM "RAFTS"

Fusion (b)

t

I

-40 min

t

(c) 1.8. Diffusion of membrane components after cell fusion. Human and mouse antigens are labeled with red and green fluorescent markers, respectively. Virus-stimulated fusion of the mouse cell and human cell (a) produces a heterokaryon with both types of antigen on its surface (b). After 40 minutes, the red and green markers have fully intermingled (c). From Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, p. 4-5. ~) 2004. Reprinted with permission from John Wiley & Sons, Inc, and the Company of Biologists.

In addition to the wide variation in composition shown in Table 1.1, many biomembranes have protein-rich domains and othel- domains. In fact, some membranes are so rich in a pal-ticular protein, they contain quasicrystalline arrays of that protein, such as bacteriorhodopsin in the purple membrane of halo bacteria and porins in the outer membrane of Gram-negative bacteria (see "Bacteriorhodopsin" and "POI-ins" in Chapter 5). Furthermore, protein-rich domains often need particulal- lipid species, because some proteins require specific lipids in their boundary layer. The boundary Jayel- of lipids, also called the annulus, is an old concept that is supported by much data from activity assays and electron spin resonance studies and more recently by x-ray structures (see "Protei n-Lipid Interactions" in Chapter4 and "Lipids Observed in X-ray Structures of Membrane Proteins" in Chapter 8). As Singer and Nicolson pointed out, specific lipid-protein interactions play important roles in the annulus. They did not anticipate that such interactions could extend the mosaic nature of the membrane to include functionally important lateral domains selective in terms of both protein and lipid components, which was unexpected in view of their emphasis on the fluidity of the bilayer. Since 1972 ·a number of new techniques have been developed to measure the fluidity of model membranes.

A View for the Future: Dynamic Protein Complexes Crowd the Membrane

9

Raft, emiched in sphingolipids, cholesterol

Outside

Prenylated protein

Acyl groups (palmitoyl, myristoy\)

1.9. Lipid rafts. Membranes have stable but transient microdomains that are enriched in cholesterol and sphingolipids, along with glycosylphosphatidylinositol (GPI)-linked proteins and proteins anchored by acyl groups. From Nelson, D. L., and M. M. Cox (eds.). Lehninger Principles of Biochemistry, 4th ed, W H. Freeman, 2005, p. 385. © 2005 by W H. Freeman and Company. Used with permission.

The physical definition of fluidity is the inverse of viscosity in an isotropic fluid, a liquid in which movement in all directions is equivalent. This definition does not directly apply to the membrane, which is highly anisotropic with a two-dimensional lipid bilayer as its base. Furthermore, the variation along the membrane normal (perpendicular to the bilayer) means the center is nearly isotropic, but a few angstroms away it is highly ordered, so position-dependent parameters are required. Therefore, measurements of membrane fluidity give results that depend on the method used, the probe for fluidity, and the conditions. More recently, considerable lateral heterogeneity in lipid bilayers has been detected employing newer techniques such as fluorescence recovery after photobleaching, single-particle tracking, and now mass spectrometry imaging. Characterization of "liquid-ordered" microdomains in biological membranes indicates thel-e are lateral domains with less fluidity, which form transient membrane "rafts" apart from the rest of the fluid bilayer (see "Organization of Bilayer Lipids" and "Lateral Domains and Lipid Rafts" in Chapter 2). Rafts are formed in the plasma membrane of many cell types as well as in many intracellular membranes. Although their composition varies, in general they are enriched in cholesterol and sphingolipids, which makes them thicker than the bulk membrane (Figure 1.9). They are also enriched with certain lipid-anchored proteins. Because many rah proteins are involved in signaling and trafficking, their

transient associations have profound biological implications.

A VIEW FOR THE FUTURE: DYNAMIC PROTEIN COMPLEXES CROWD THE MEMBRANE INTERIOR AND EXTEND ITS BORDERS

Even with the addition of microdomains of different sizes, lifetimes, and functions, the model of the Iluid mosaic membrane is incomplete. While the emphasis on lipid rahs focused attention on the lateral organization of the membrane, a variety of both old and new findings indicate the transverse organization across lhe plane of the mem brane is complex as well. The new view of the membrane acknowledges variation in this transverse dir'ection, encompasses layers outside the bilayer iIsel f, and recognizes the activities going on at its borders. The important activities OCCUlTing at the surfaces, along with striking differences across the bilayer, emphasize the significance of I he lhil-d dimension of the membrane. Thus the membrane is more than a layer of proteins embedded in a lipid bilayer. Crucial functions are carried oul by complexes involving interactions between integral and peripheral proteins at the interfaces. Many of the proteins are oligomers that operate in large assemblies in the membrane. Many large protein complexes operate in very close quarters in nOI-mally crowded biomembranes.

1.10. Peripheral proteins and complexes. Membranes encompass not only the bilayer but also peripheral proteins and, in the case of plasma membranes, the cytoskeleton. Typically crowded with proteins, membranes contain many complexes that extend beyond the bilayer. Engelman, D., Nature. 2005. 438:578-580 © 2005 Reprinted with permission of Macmillan Publishers Ltd.

To start to describe this complexity, researchers are mapping the microenvironments found along a line extending perpendicular to the plane of the bilayer at different sites along biological membranes. The asymmetry in lipid compositions of the inner and outer leaflets was detected long ago, yet new results show it is associated with complex patterns of lipid trafficking that can turn over components of the plasma membrane each hour. Also familiar for years has been the complex cytoskeleton on the internal surface of the eukaryotic plasma membrane that provides structural support and limits the mobility of some membrane proteins. In Gram-negative bacteria lipoproteins from the outer membrane provide a similar structural SuppOrlthrough their hnks to the underlying peptidoglycan. These alT stable structures at the borders of membranes. Gaining attention today are important transient associations with peripheral proteins along the SUI-face of many biomembl-anes. Even the picture of the lipid bilayer itself has been revised from the "lollipop" depiction of lipids in most drawings. Sophisticated analyses of diffraction data and computational modeling (described in Chapter 8) present a new picture of the bilayer in which the nonpolar domain, defined as the center that is free of water, is only about half of its thickness. Each interfacial region, made up of lipid headgroups and amphiphilic domains of proteins and containing some \vater molecules, contributes another quarter. Furthermore, these regions

are the dynamic playgrounds for lipid-metabolizing enzymes and other proteins that insert into the bilayer (described in Chapter 4.) The exterior surface of many cell membranes is crowded with peripheral proteins that interact at the interface of the bilayer. Many of these proteins have activities that are regulated by binding the membrane in dynamic cycles (see "Amphitropic Proteins" in Chapter 4). Their specific binding is mediated by highly conserved motifs and often by divalent cations. Thus, the focus of membrane research has expanded to include the membrane periphery as an additional important layer of the membrane. Large complexes made up of integral membrane proteins and associated peripheral proteins carry out many functions of the membrane (see Chapter 11). The typical biomembrane is crowded with protein assemblies, many of which are tightly associated heterooligomers (Figure 1.10). Furthermore, some complexes are large enough to span two membranes, either from the same cell or organelle, seen in the double-membrane systems of Gram-negative bacteria and mitochondria; fyom different cells, such as at gap junctions; or from a cell and a virus, as observed in the fusion events enabling viral penetration. Future challenges in understanding membrane functions include characterizing these larger functional membl-ane complexes and must not neglect the essential activities at the bilayer interface and on its surface (see Chapter 12).

A View for the Future: Dynamic Protein Complexes Crowd the Membrane In short, the biomembrane consists of amphiphilic molecules that respond to the hydrophobic effect by spontaneously assembling into the bilayer. The bilayer is fluid and mosaic with respect to both lipids and proteins and contains dynamic lateral domains, or rafts, which appear to be critical to many biological functions. The central nonpolar region that excludes virtually all water molecules spans about half the bilayer, sandwiched between the two interfacial layers where considerable activity takes place. The surface of the bilayer is often crowded with peripheral proteins, some coming and going and others providing mechanical support. Many functions of the membrane are carried out by molecular assemblies that are large multicomponent complexes, some of which even span two membranes. To study this marvelous, multifaceted biological structure in some detail and to appreciate the stunning molecular structures of membrane constituents now emerging requires an initial understanding of the physical and chemical properties of the membrane bilayer. Thus, this book starts with the fundamental properties of membrane components and ends with examples of membrane proteins whose high-resolution structures give insights into their functions. Chapter 2 takes a close look at the structure and function of membrane lipids, moving from the diversity and properties of membrane lipids to the properties of domains and the formation of lipid rafts. It concludes with the examination of the role of non-bilayer-formi ng lipids in biological membranes. Chapter 3 describes the tools needed for in vitro characterization of membrane constituents. It opens with the properties of the detergents that are so important in purification of membrane components and then surveys the old and new model systems available for studying lipid aggregates and for reconstituting membrane proteins. Chapter 4 portrays the different types of proteins at or in the bilayer. It describes amphitropic proteins, including peripheral proteins and lipid-anchored proteins, and the dynamics of their binding to the interface. It considers pep tides and proteins that can insert into bilayers, a group that includes some ionophores and toxins. Finally, it examines the constraints placed by the bilayer on the general Features of integral membrane proteins. Then Chapter 5 gives an in-depth view of the two major classes of integral protein structuresbundled a-helices and f3-barrels - and presents the proteins that are paradigmatic examples of each class. It ends with a reminder that the knovvn atomic structures represented in these classes at present account For only a small proportion of the membrane proteins in the genomes. With this foundation of the structural principles that determine the characteristics of membrane constituents, the book turns to biochemical, biophysical,

11

and proteomic studies of membrane proteins. Chapter 6 describes the three major (and overlapping) classes of membrane protein functions - enzymes, transport proteins, and receptors - with examples of each. Then it covers the bioinformatics tools that are used to predict structures and families of membrane proteins for genomic analysis and describes proteomics techniques designed to reveal inter-subunit and other proteinprotein interactions. After a major focus on a-helical membrane proteins, it ends with prediction methods for f3-barrel proteins. Chapter 7 Focuses on Folding and biogenesis of membrane proteins, starting with studies on the Folding of purified membrane proteins andmoving to the complex systems involved in their biogenesis in cells. It concludes with a look at human diseases that result from misfolding membrane proteins. Chapter 8 presents diffraction and simulation techniques that give representations of the fluid membrane. It moves from structures of lipids in crystalline arrays to liquid crystallography and other techniques that describe the fluid bilayer based on diffraction data. It shows how computational modeling using molecular dynamics gives simulations of increasingly complex lipid bilayers. Then it describes lipids observed in highresolution x-ray structures of membrane proteins and ends with a brief discussion of techniques used to obta in suitable crystals of membrane proteins. The last part of the book tours the new and growing field of membrane structural biology with a sample of representative membrane proteins of known structures. Chapters 9 and J 0 showcase examples of membrane enzymes and transducers, transporters and channels. At this initial stage of the development of membrane structural biology, each atomic structure of a membrane protein that is solved provides fresh insights into its functions and relationships. Yet many functions of the membrane are carried out by large protein complexes, so Chapter 11 describes the high-resolution structures of several important multicomponent membrane systems. Then Chapter 12 identifies some of the common issues that have arisen during work on these first structures and that are likely to provide tbe themes and directions for h.Jture work. Molecular cbaracterization of the membrane bas lagged behind otber fields of biochemistry, in part because of tbe difficulties in obtaining high-resolution structural inFormation on its components. Today, structural insights into the membrane are taking off, hleled by the recenl growth in the number of membrane proteins whose structures have been solved. As always in biochemistry, these first structures provide models that suggest possibilities For structures and mechanisms still unknown, those involved in similar as well as more complicated membrane functions. From the fl.Jndamental principles covel-ed in the first chapters to the

descri ptions of the high-resolution structures in the last part, this book gives a solid gr8sp of the field needed for an appreci8tion of the progress to come. Membrane research is leading to significant understanding or- topics such as membrane fusion and viral infection, sign8ling 8nd roles of v8rious oncogenes in cancer development, uptake and efflux of drugs affecting drug resistance, drug targeting and drug design, and many other·s. Components of the membrane are the focus of basic research into differentiation and development, immunology, neurochemistry, and nutrition. Finally, defects in membrane components are the cause of many human diseases, 8nd membrane components are the targets of the m8jority of newly designed/discovered drugs. It is an exciting time to study the biomembrane.

FOR FURTHER READING

Edidin, M., Lipids on the frontier: a century of cell-membrane bilavers. Nat Rev Mol Cell Bioi. 2003. 4:414-418. Engelman, D. M., Membranes are more mosaic than fluid. N(llure. 2005, 438:578-580. Green, D. E., and A. Tzagoloff, Role of lipids in the structure and function of biological membranes. ] Lipid Res. 1966, 7(5):587-602. Kuhn, T. S., The StrucLUre of'Scientific Revolutions, 3rd ed. University of Chicago Press, 1996. Simons, K., and E. lkonen, Functional rafts in cell membranes. Nature. 1997,387:569-572. Singer, S. J., and G. L. Nicolson, The Fluid Mosaic Model of the stnlcture of cell membranes. Science. 1972, 175:720731. Tan ford C, The Hydrophobic Eli'ecl: Formation oj'MiceJles and Biological Mernbrarles, 2nd ed. New YOl-k: Wiley, 1980.

2

The Diversity of Membrane Lipids

Crystal

Gel

Fluid

To understand biological membranes, and especially to predict their behavior, requires a detailed knowledge of their components. it is appropriate to start with the lipids that make up the bilayer, because they are not just a solvent providing the "sea" in which membrane proteins float. if this were the case, a few lipidic species would suffice to provide the amphiphilic base of the bilayer and some variation of shapes for packing it. instead, the diversity of membrane lipids is amazing. A typical biomembrane contains more than a hundred species of lipids, which vary in general structure and in the length and degree of saturation of their fatty acyl chains. This chapter begins \vith the properties and modulation of the acyl chains and then reviews the structural features of the major complex

lipids. it describes the properties of lipid aggregates, including their polymorphism and phase separations, which are the basis of their ability to form lateral microdomains. lL addresses the characteristics of lipid rafts in membranes. The chapter ends with studies or lipid metabolism in Escherichia coli that address the biological need ror diversity of lipids and support a role for non-bilayer-forming lipids in maintaining the elasticity of the membrane.

THE ACYL CHAINS

When lipids are extracted from a cell with organic solvent, such as a 2: I mixture of chloroform and

TABLE 2.1. Some naturally occurring fatty acids: structure, pt'operties, and nomenclature a

Carbon skeleton

Structure b

Systematic name c

Common name (derivation)

12:0

CH3(CHz)lOCOOH

n-Dodecanoic acid

Lauric acid (Latin laurus," laurel plant")

14:0

CH3(CHz)lZCOOH

n-Tetradecanoic acid

Myristic acid (Latin

Melting point (OC)

Solubility at 30°C (mg/g solvent) Water

Benzene

44.2

0.063

2600

53,9

0.024

874

myristica, nutmeg genus)

16:0

CH3(CHzh4COOH

n-Hexadecanoic acid

Palmitic acid (Latin palma, "palm tree")

63.1

0.0083

348

18:0

CH3(CHz)16COOH

n-Octadecanoic acid

5tearic acid (Greek stear, "hard fat")

69.6

0.0034

124

20:0

CH3(CHZ)lSCOOH

n-Eicosanoic acid

Arachidic acid (Latin Arachis, legume genus)

76.5

24:0

CH3(CHz12zCOOH

n-Tetracosanoic acid

Lignoceric acid (Latin lignum, "wood" + cera, "wax")

86.0

16:1(69)

CH3(CHz)s CH=CH(CHzhCOOH

cis-9-

Palmitoleic acid

0.5

cis-9Octadecenoic acid

Oleic acid (Latin oleum, "oil")

13.4

cis-,cis-9,12-

Linoleic acid (Greek finon, "flax")

-5

Hexadecenoic acid

18:1(69)

CH3(CHzh CH=CH(CHzhCOOH

18:2(69,12}

CH3(CHZ)4 CH=CHCHz CH=CH(CHzhCOOH

Octadecadienoic acid

18:3(69,12,15)

CH3CHZ CH=CHCH z CH=CHCH z CH=CH(CHzhCOOH

cis-,cis-,cis9,12,15Octadecatrienoic acid

ex-Linolenic acid

-11

20:4(65,8,11,14)

CH3(CHz)4 CH=CHCH z CH=CHCH z CH=CHCHz CH=CH(CH zl3COOH

cis-,cis-, cis-,cis5,8,11,14Eicosatetraenoic acid

Arachidonic acid

-49.5

• The symbol for fatty acids gives the number of carbon atoms, followed by the number of carbon-carbon double bonds. For unsaturated fatty acids, the notations in parentheses denote the positions of their double bonds. For example, t;9 denotes a double bond between C9 and Cl0. All the double bonds in these fatty acids have cis configuration. b All acids are shown in their nonionized form. At pH 7, all free fatty acids have an ionized carboxylate. Note that numbering of carbon atoms begins at the carboxyl carbon. e The prefix n indicates the normal unbranched structure. For instance, dodecanoic simply indicates 12 carbon atoms, which could be arranged in a variety of branched forms; n-dodecanoic specifies the linear, unbranched form. Source: Data from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed. New York: W. H. Freeman, 2005.

methanol, free fatty acids make up only about 1% of the total; most fatty acids are bound covalenLly in COI11pJex lipids. There are more than 500 different species of fatty acids according to the searchable database at http://sofa.bfel.de/. Table 2.1 shows the fatty acids normally occurring in the lipids of biological membranes, grouped as Saturated or unsaturated (those with at least one carboncarbon double bond). The acyl chains are typically 10 to

24 carbons long, with an even number of carbons resulting h'om their synthesis from the precursor acetylcoenzyme A. The systematic names of fatty acids are rarely used. Instead they may be referred to by common names or by sym boIs indicating their chain length and degree of saturation. For example, stearic acid is CI8:0, a saturated 18-carbon chain, drawn in Figure 2.IA with the acyl chain Fully extended. If it is part of a complex lipid, the

The Acyl Chains

15

A. Stearate

B. Oleate

C. Elaidate

o

o

o

~ /

0-

~/

C

0-

C

~/

0-

C

2.1. Saturated and unsaturated fatty acids with l8-carbon chains. With the carboxyl group deprotonated at neutral pH, stearic acid becomes stearate (C18; A), oleic acid becomes oleate (C18:l, /),9 cis; B), and elaidic acid becomes elaidate (C18:l /),9 trans; C). While the trans double bond does not affect the conformation of the acyl chain, the cis double bond introduces a kink in the chain, as illustrated by the space-filling model for oleic acid. Redrawn from Nelson, D. L., and M. M. Cox (eds.l, lehninger Principles of Biochemistry, 4th ed, W. H. Freeman, 2005, p. 345. © 2005 by W. H. Freeman and Company. Used with permission.

18-carbon saturated acyl chain is stearoy!. If the chain is monounsaturated (with one double bond), it is oleoyl, because oleic acid is C18:1 (69), an 18-carbon chain with a double bond between carbons 9 and 10 (Figure 2.1 B). A single cis double bond makes a 30° bend. or kink, in the acyl chain, as sho\.vn in the figure. Naturally occurring unsaturated fatty acids have cis double bonds. Partial dehydrogenation of dietary fats prod uces trans fatty acids such as elaidic acid. C18:1 (69) trans (Figure 2.IC), which are found in processed foods and enter the human body (to the apparent detriment of health). The fully extended chain in the illustration of stearic acid (Figure 2.1 A) results when all the dihedral angles along the hydrocarbon chain are 180', which is the favored angle of rotation around the single C-C bonds in fatty acids. This angle, called the torsion angle, is evident when bonds between four neighboring C atoms are considered (Figure 2.2). The most favored value of the torsion angle of a hydrocarbon chain is 180" and is called trans or anti; a second favored value is around 60° and is called gal/che. In the liquid state. hydrocarbon chains sample different torsion angles with rotation around their C-C

bonds; rotation is more restricted around C=C double bonds. Phospholipids in biological membranes ohen have a saturated acyl chain on CI of the glyceml moiety and an unsaturated chain on C2 (see Figure 2.5). Polyunsaturated fatty acids. including omega-3 fatty acids such as ex-linolenic acid (CI8:3. 69. 12, IS). whose last C=C is three carbons fTom the end of the chain. are important nutritional precursors to molecules such as prostaglandins and isoprenes. As minor constituents of membrane lipids. polyunsaturated acyl chains are bulky yet highly flexible and affect membrane elasticity (see below); they are usually absent in bacteria. Unusual fatty acids are found in the membranes of some organisms. Some bacteria have branched. hydroxylated, or iso-acyl chain fatty acids. E. coli membranes can have ~25% cyclopropane-containing fatty acids. In some marine organisms, an odd number of carbons is common. Pure fatty acids undergo a sharp phase transition when they are melted. This transition is detected by the inCl'eased heat uptake measured by differential scanning calorimetl-y (DSC) as illustrated in Figure 2.3. For the acyl chains of a lipid. such as phosphatidylcholine.

:x0~> CH 3

anti conformation (j = 180°

][3:$: (CH 3

~

CH 3

H

HOH CH7\ line-and-lvedge formulas:

H

-<:if

. H,\

-------FCH 3

H

A. viewing a model of butane from one cnd of the central carbon-carbon bond

/

CH 3

CH 3

/ C\"" H

H O C H3

HVH H,~/H ~

CH 3 HVH H gauche conformations (j =:!:60°

"'\'!'//"',. H

H ..

CH 3

B. end-on view

C. Newman

D. energetically favored torsion angles

projection

2.2. Bond torsion angles in hydrocarbon chains. Four atoms of a hydrocarbon chain, labeled ABCD, may be represented by butane, shown in A. The torsion angle is the dihedral angle defined by the planes between atoms ABC and atoms BCD. This angle may be visualized from the end-on view (B) and is diagrammed in the Newman projection (C). The most stable torsion angles for hydrocarbon chains are called anti and gauche, shown for butane in D. A long hydrocarbon has considerable flexibility with varying torsion angles along the chain. Redrawn from Loudon, G. M., Organic Chemistry, 4th ed., Oxford University Press, 2002. © 2002 by Oxford University Press, Inc. Reproduced by permission of Oxford University Press, Inc.

this transition reflects the change from the gel state ("solid") along the chains to the liquid crystalline state ("fluid") as the temperature is increased. In the gel state, the acyl chains are fairly ordered, with a high Irans/gauche ratio. Tn the liquid crystalline state, the rotational freedom decreases the lrans/gauche ratio and allows many more configurations of the hydrocarbon chains (see chapter Frontispiece). From the trend in Figure 2.3 as well as the melting temperatures (T,n) of the fatty acids listed in Table 2.1, it is evident that Till increases with increasing chain length and decreases with increasing unsaturation. In the gel phase, the extended saturated acyl chains pack closely, stabilized by van der Waals forces. Fatty acids with longer chains have higher Till' because with more extensive contact areas, more heat is required to disrupt this structure. The kinks introduced by double bonds disrupt this order, destabilizing the gel phase so less heat is required to melt unsaturated fatty acids.

Many examples of the adaptation of unicellular organisms to their environments illustrate the functional importance of these phase transitions. In E. coli growing at 37 C, the ratio of satUl-ated to unsaturated fatty acids is about 1:1; when growing at 17 c C, it changes to about 1:2 (see Table 2.2). When gro\.ving at high temperatures, bacterial membranes are enriched in saturated and longer acyl chains, as is especially evident in thermophilic bacteria such as those found in the hot springs at Yellowstone National Park, with ambient temperatures around 85 C. Marine organisms both bacteria and deep-sea fish - have adapted to high pl-essures by increasing the proportion of unsaturated fatty acids to enable them to maintain the fluidity of their membranes. In gener'al, organisms vary the composition of acyl chains in their membranes to achieve a state that is fluid but not too fluid, indicating that the acyl chains may be important determinants of phase polymorphism (see below), even in a complex u

v

Complex Lipids

17

TABLE 2.2. Fatty acid composition of E. coil cells

cultured at different temperatures· Percentage of total fatty acids b

Myristic acid (14 :0) Palmitic acid (16:0) Palmitoleic acid (16: 1) Oleic acid (18:1) Hydroxymyristic acid Ratio of unsaturated to saturated C

lO-C

20°C

30°C

40'C

4 18 26 38 13 29

4 25 24 34 10 2.0

4 29 23 30

8 48 9 12 8 038

10 16

The values are given in weight percent of total lipid. The exact fatty acid composition depends not only on growth temperature but on growth stage and growth medium composition. C Ratios calculated as the total percentage of 16: 1 plus 18: 1 divided by the total percentage of 14:0 plus 16:0. Hydroxymyristic acid was omitted from this calculation. Source: Data from Marr, A. G., and J L Ingraham, Effect of temperature on the composition of fatty acids in Escherichia coli J Bocteriof. 1962, 84: 1260-1267 Reprinted with permission from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed. New York: W H. Freeman, 2005. a

b

With fally acyl chains in ester linkage on carbons 1 and2 it becomes phosphatidic acid (PA). Esterification of PA with another alcohol creates the following PLs: phosphalidvlcholine (PC), phosphalidylethanolamine (PE), phosphatidvlserine (PS), phosphalidylglycerol (PG), and phosphatidylinositol (PI) (Figure 2.5). In addition, PG can link through its glycerol headgroup to PA to Form diphosphatidylglycerol (CL for its common name, cardiolipin). These abbreviations are coupled with lhe abbreviated common names for the acyl chains: DOPC is Lhus dioleovlphosphaLidylcholine and MPoPS is I-myristyl 2-palmitoleoylphosphatidylsel'ine. (See Appendix I for a list of abbreviLltions.) The phosphate groups and headgroups are the polar portions, and the acyl chains are the nonpolar parts of these amphiphilic molecules. Jn many PLs of biologicLl] membranes, the acvl chain on Cl is saturated and 16 or 18 Main transition

t

J-

mixture that would not exhibit a simple "melting" point.

DMPC

PMPC COMPLEX LIPIDS

Lipids found in biomembranes fall into three main classes of lipids (Figure 2.4):

u

MPPC

E '-< v

• glycerophospholipids (ohen called phospholipids) sphingolipids (including sphingophospholipids) sterols and linear isoprenoids

-S 3 0

c

0

~

v c

lJ-l

C
In addition, glycolipids may be considered a separate class, although they consist of either glycerophospholipids or sphingolipids with oligosaccharide headgroups. Their importance in human health and disease is evident: the blood groups A, B, and 0 are determined by the glycosphingolipids on cell surfaces, and several hereditary diseases, such as Tay-Sachs disease, result from abnormal accumulalion of glycosphingolipids. The ganglioside GM 2 that accumulates in patients with Tay-Sachs disease is a sphingolipid with galactose (Gal), glucose (GIc), N-acetylneuraminic acid (NeuNAc), and N-acetylglucosamine (GJcNAc) moieties. Phospholipids

A glycerophospholipid, commonly known simplv as a phospholipid (PL), is buill on a glycerol molecule, which becomes chiral when derivatized to glycerol-3phosphate. The backbone of membrane PLs is the Lisomer, called 511-glyceroI3-phosphate (511, for stereochemical numbering, is used instead of D and Lor Rand S).

v

..c

DPPC

--L ~SPC

c I1J

SPPC

~ IX

DSPC I

0

I

I

JO 20

30 40 50 60 Temperature (OC)

2.3. Detection of the gel-to-liquid crystal tronsition in different phosphatidylcholine (PC) molecules with acyl choins varying from 14 carbons (dimyristoyl PC) to 18 carbons (disteoroyl PC). Differential scanning calorimetry is used to measure the heat consumption oS the temperature is increased. The peaks correspond to the enthalpic "melting" events, which occur ot higher temperatures as the chain lengths increase. When the PC contains two acyl chains of different lengths, its melting temperature is midway between that of the two PC molecules having identical choins of the two types. Redrawn from Keough, K. M., and P. J. Davis, Biochemistry. 1979, 18: 1453. © 1979 by American Chemical Society. Reprinted with permission from American Chemical Society.

A

0,

(~H2 o CH

0-

o

o

I + 'CH2-0-P-0~N___ II ---

0/

Glycerophospholipid

0,

(~H2

o

HO

0-

/CH I + 'CH2-0-P-0~N:::::::

oII

Lysophospholipid

---

B

OH

~"CH I

0-

I

/CH + NH 'CH2-0-P-0~N:::::::

r o

II

Sphingomyelin

---

0 /OH

~"CH I

/CH NH 'CH 2-O-Glc-Gal-GloNAc-Gal

I

o

NeuNAc

I

Ganglioside

NeuNAc

c. OH

Sterol

Farnesyl Linear isoprenoids Geranylgeranyl .;:?' 2.4. Three major classes of membrane lipids. The structures of representative glycerophospholipids (Al, sphingolipids (Bl, and sterols and linear isoprenoids (C) are shown. An additional class is the glycolipids, which have sugar headgroups typically on a sphingomyelin base. A and B redrawn from Gennis, R. B., Biomembranes, Springer-Verlag, 1989, p. 24; C redrawn from Nelson, D. L., and M. M. Cox (eds.l, Lehninger Principles of Biochemistry, 4th ed., W H. Freeman, 2005, p. 355.

carbons long while that on C2 is frequently unsaturated and often longer. Although phospholipids do act as solvent for membrane proteins and define the polar and nonpolar domains of the bilayec they also have important chemical, biological. and physical properties. The anionic PLs (PS, PI, PG, and CL) have a net negative charge at physiological pH, while the zwitterionic PLs (PE and PC) are neutral. PE and PS contain reactive amines that can participate in hydrogen bonding. PI, PC, and cardiolipin (CL) are relatively bulky. \vhich affects their packing in bilayers. When a phospholipid loses one acyl chain through the action of a phosphoJi-

pase, it becomes a lysophospholipid with increased water solubility that gives it surfactant (detergent) activity. Phospholipids provide sources of second messengers for signaling across the membrane and enhance the activity of membrane enzymes and transport proteins (see Chapter 4). Their degree of unsaturation contributes to the elasticity of the membrane, which influences insertion and sequestering of proteins (see Figure 2.18 and the discussions of folding studies in Chapter 7). Archaebacteria have a unique set of phospholipids that have ether linkages to their phytanyl chains (instead of ester bonds to acyl chains) as they are

Glycerophospholipids

o

"CH I

o

o

0-

2

I

/CH

"CH 2 -O- P -OH

II

o

Phosphatidic acid (PA) +

-0-CH 2 -CH 2 -N(CH 3h Phosphatidylcholine(PC)

0

+

- 0 - CH 2 - CH 2 - NH 3

Phosphatidylethanolamine (PE)

+

-0-CH 2 -CH-NH 3 I CO;

Phosphatidylserine (PS)

OH

I - 0 - CH 2 - CH - CH 20H

-0

~ 6

HO 2

Phosphatidylglycerol (PG)

OH

~H H~

;

Phosphatidylinositol (PI)

OH

0"

o

o

~H2 /CH

0-

I

"CH)-O-P =0

-

o

I

0

I

CH)

I -

CHOH

Cardiolipin (CL)

I

o

CH 2 "CH I

o

o

o

I 0

2

/CH

I I

"CH -O-P =0 2

0-

2.5. Structures of glycerophospholipids. The common glycerophospholipids in biological membranes contain one of the polar headgroups shown. In addition, they vary greatly in the length and saturation of their acyl chains, although in general the acyl chain on C1 is saturated and the acyl chain on C2 is unsaturated. Redrawn from Gennis, R. B., Biomembranes, Springer-Verlag, 1989, p. 25.

,

,

HalO o Archaeol

OH

CH 3

I

o OH

~ /0

0

P -0/ \

o

o HO

HO 0/ "

~ ..0'0

sY"

-0/ \

i-

~-OH

~OH

_O~

o

'OH

Archaeol SuI fated triglycosylarchaeol

-0

/

o

~P / "Archaeol

o

PhosphatidylgJycero phosphate methylester

O~ /

/ -0

OH

o

P" Archaeol

Phosphatidylglycero sulfate

2.6. Structures of the three major polar lipids found in the purple membrane of Ha/obacterium sa/inarium, which are derived from archaeol (2,3-di-O-phytanyl-sn-glycerol, shown above). Redrawn from Lee, A G., Biochim Biophys Acta. 2003,1612:1-40.

derived fTom archaeol. Archaeol is 2,3-di-0-phytanylsl1-glycerol, and its phytanyl groups are 20-carbon branched chain isoprenoids (3,7, \1,1 5-tetramethyl hexadecanoic acid) (Figure 2.6). They also have unusual headgroups and differ in stereochemistry, esterified to the phosphate head group or sulfated glycolipid on the SI1 I instead of the S/1.3 carbon. Some archeallipids even have the two archaeol groups fused with headgroups on both ends (not shown in the figure). Sphingolipids

Sphingolipids are built not on a glycerol backbone but on sphingosine, a long-chain amino alcohol, to which a fatty acyl chain is attached in amide linkage (Figure 2.7A). The most common sphingoJipids are sphingomyelins, which are sphingophospholipids with either phosphocholine or phosphoethanolamine headgroups, giving them overall shapes much like those of PC and PE (Figure 2.7B). Other sphingolipids have headgroups made up of sugars or oligosaccharides, providing the great diversity of structure of cerebrosides (with monosaccharide headgroups) and gangliosides (with oligosaccharides). The ability of the amide bond and the hydroxyl group of sphingolipids to hydrogen bond at the

membrane/water interface allows specific interactions \vith PL headgroups, the hydroxyl group of cholesterol, or other polar groups. In mammalian cell membranes, the fatty acid is generally saturated, with 16 to 24 carbons, making both "chains" o[ the sphingolipids fully saturated. A small fraction of the 24-carbon chains have a single C=C far along the chain (C24:1 6.15), so they still pack tightly together. Sphingolipids are important components of nerve membr'anes, and their carbohydrate moieties are vital in cell recognition and differentiation. Sterols and Linear Isoprenoids

A third major class of biological lipids includes compounds derived [Tom five-carbon units called isoprene (2-methyJ-l ,3-butadiene). The linear isoprenyl groups farnesyl (CI5) and geranylgeranyl (C20) are used to anchor certain proteins to the bilayer (see Chapter 4). Dolichols are long (C90) poly isoprenoid lipids needed tc attach sugars to membrane proteins in the ER of animal cells. Moreover; all steroids are derived from cyclized polyisoprene precursors that have 30 carbon atoms. The dominant sterol in animal membranes is cholesterol (FigUl"e 2.8). Other eukaryotes have different

Complex Lipids

21

B.

A.

Phosphocholine headgroup

A sphingomyelin 2.7. Structure of a sphingomyelin. A. Sphingosine is shown with a palmitoyl chain in amide linkage and a phosphocholine headgroup. B. Comparison of space-filling models of this sphingolipid (a) with SOPC, the glycerophospholipid 1-stearoyl-2-0Ieoyl phosphatidyl choline (b), reveals how very similar they are in spite of the lack of an unsaturated chain in the sphingolipid. Redrawn from Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, pp. 386-387. © 2004. Reprinted with permission from John Wiley & Sons, Inc.

B.

A.

CH J C2 H 5

CH 3

HO

Stigmnsterol 1-10

Cholesterol

C.I-I 3 C2 H,

CH J

.1-10

I3-Si tosterol CH 3 CH J .1-10

Ergosterol 2.8. Sterols found in biological membranes. A. The structure and space-filling model of cholesterol, a major component of animal membranes. Redrawn from Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, p. 389. © 2004. Reprinted with permission from John Wiley & Sons, Inc. B. Different sterols occur in membranes of other organisms: plants have stigmasterol and l3-sitosterol, whereas yeast and fungi have ergosterol.

CH 3

2.9. Close packing of cholesterol with phospholipids Molecular dynamics portrays the interactions between cholesterol and phospholipids and reveals a close association between saturated acyl chains and the rigid tetra cycle of the sterol, whereas the bulky PL headgroup can cover the small (OH) headgroup of cholesterol in the manner of an umbrella. A. Side view of a simulated cholesterol-PL complex. The PL in the model is 1-stearoyl-2-docosahexaenoyl-PC, which has a polyunsaturated omega-3 fatty acid on C2. The saturated acyl chains of the PL extend past the sterol. Saturated acyl chains (blue) pack along the smooth faces of the cholesterol, whereas polyunsaturated acyl chains (red) have a much lower affinity for it. Even with oleoyl on C2 (not shown), the kink from the double bond at C9-10 interferes with this close packing. On the other hand, sphingomyelin usually has saturated chains of 16 to 24 carbons, and when unsaturated the double bond occurs at C 1S, well below the rigid portion of cholesterol, which explains the preference of cholesterol for sphingomyelin. B. Side and top views showing probability functions for the headgroup of PC in dynamic complex with cholesterol, illustrating the umbrella effect. The probability density for phosphate is gray, and that for choline is orange. As the PC orientation varies, the headgroup completely covers the sterol. From Pittman, M. c., F. Suits, A D. MacKerell, Jr., and S E. Feller, Biochemistry. 2004, 43:1S318-1S328. © 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

sterols in their membranes (ergosterol in yeast and fungi, sitosterol and stigmasterol in plants), while prokaryotes have essentially none. The cholesterol content of various ceJlmembranes varies from 0% to ~25%

(Table 1. 1). Eukaryotic cells have as much as 90% or their- cholesterol in the plasma membrane, maintained by a dynamic cholesterol supply route from the ER. Both the plasm membrane and intracellular

The Lipid Bilayer Matrix

23

Bilayer normal

Acylglycerol part

i

Polar region d p thickness of the polar region

Headgroup

~

1 Bilayer interface S, molecular area

2.10. The structure of a PC molecule determined with x-ray crystallography. The acyl chains are fully extended (all anti dihedral angles) and are tilted from the bilayer normal. The polar region (circled) includes the headgroup and the glycerol moiety. Other PL molecules give a different chain tilt and thickness of the polar region, depending on the way their polar headgroups pack in the crystal. Redrawn from Gennis, R. B., Biomembranes, Springer-Verlag, 1989, p 38. It) 1981 by Elsevier. Reprinted with permission from Elsevier.

membranes have cholesterol-rich domains (see below). Experimental results following the effects of depletion of cellular cholesterol by treatment with cyclodexll-in suggests that many different functions in eukaryotic cells involve cholesterol. Pure cholesterol cannot form a bilayer, and excess cholesterol (beyond 50-60 mol 'Yo') precipitates out of PL biJayers. X-ray diffraction detection of the crystalline precipitate establishes precise cholesterol solubility limits of 66 mol % in PC and 51 mol % in PE bilayers. In calorimetric studies of mixtures of cholesterol and pure phospholipid, the melting transition of the lipid broadens and is eventually eliminated as the percent of cholesterol is increased. This phenomenon has been attributed to the endothermic dissolution of condensed cholesterol-phospholipid complexes of defined stoichiometries. These cholesterol-PL complexes form cooperatively and are described by [CqPp]n,

in which q molecules of cholesterol complex with p molecules of PL, with n denoting the cooperativity. The interactions between cholesterol and othet, lipids have been cha1'3cterized by nuclear magnetic resonance (NMR) and simulated using molecular dynamics (see "Molecular Dynamics" in Chapter 8). The rigid portion of the sterol imposes conformational order on neighboring lipids, while the larger headgroups of the phospholipids form "umbrell£ls" over their cholesterol neighbm-s (Figure 2.9). Because its rigid tetracycle can align better next to saturated acyl chains, cholesterol exhibits a strong preference for saturated sphingomyelin over unsaturated PLs. The presence of cholesterol increases bilayer thickness, tight packing of acyl chains, and compressibility, while it decreases the translational diffusion rates of PLs.

THE LIPID BILAYER MATRIX , Surface concentration terms are either mole fraction ([specific Iipid]/[totallipid]) or mol % (mole fraction x 100). Thus 60 mol % cholesterol would be 60 moles of cholesterol per 100 total moles; typically this would be in a mixture with 40 moles of other lipids.

These diverse lipid molecules form bilayers vvi th properties tha t renect their individual structures and collective

interactions. A basic description of the bilayer as a matrix starts with the well-characterized structure of phospholipid molecules in a bilayer. It then considers the diffusion of phospholipids within the two dimensions of the bilayer: the rapid lateral movements that generate the fluidity of the bilayer and the relative lack of rapid transverse movements that allows it to be asymmetric. Structure of Bilayer Lipids

Details of the structure of PL bilayers have been obtained with x-ray crystallography, NMR, and molecular modeling. X-ray crystallography of pure PLs crystallized from aqueous acid shows structures in which the headgroups are bent in a position almost parallel to the plane of the bilayer, with the acyl chains aligned with each other as a result of a bend in the acyl chain at C2 (Figure 2.10). The position of the headgroup and the angle of chain Lilt vary for different PL species. Precise measurements of their cross-sectional areas are obtained. Of course, these structures do not represent the structure of lipids in the fluid membrane, particularly because in the crystalline state the chains are rigid and fully extended in all trans conformations. NMR experiments show that in the fluid membrane the acyl chains are not rigid. Motion along the acyl chain can be probed by NMR using 2H labels at different positions along the acyl chains. The deuterium NMR spectra of different DMPC preparations labeled in different positions clearly show a gradation of mobility along the chain (Figure 2.11). The spectrum for DMPC with 2H at the terminal methyl residue is narrow, reflecting the disorder near the center of the bilayer, in contrast to the broadened peaks obser'ved when the 2 H is closer to the interface and less mobile. Similar results are obtained using probes suitable for fluorescence depolari7.ation and recovery (see Box 2.1) and electmn paramagnetic resonance (see Box 4.2). Molecular modeling of aqueous lipid systems illustrates the mobility along the acyl chains in bilayers. In these simulations, the freedom of Irans/gauche rotations of the carbon-carbon bonds of long chains makes the chains highly flexible, with the result that they are clearly not aligned in a snapshot of a simulated bilayer' (Figure 2.12). The mobility of the acyl chains that is r'evealed by a molecular dynamics approach can be obser'ved at sites such as hllp://courses.washington. edu/physeng/membrane/bilayer.htm or http://www. umass.edu/micmbio/rasmol/bilayers.htm. (Molecular modeling methods and their results are described in Chapter 8.) The angular rotations along hydrocarbon chains of individual molecules contribute to the fluid ity of the membrane that is so essential to life it is main-

3,3

6,6

10,10~

12,12 I

14,14, 14 1 40,000

/

,---

20,000

o

-20,000

-40,000

Hz 2.11. Deuterium NMR of DMPC with 2H at different positions on the acyl chains as indicated on the left. The deuterium located closer to the center of the bilayer experiences greater disorder, giving a sharper peak, than deuterium located near the interface. Gennis, R. B, Biomembranes, Springer-Verlag, 1989, p. 53. 11") 1989. Reprinted with permission from E. Oldfield.

tained by organisms living under extreme conditions (as discussed above). Another important source of fluidity is the movement of whole molecules within the two-dimensional matrix of the bilayer Diffusion of Bilayer Lipids

Lipid molecules move within the bilayer in three different modes: mtational, lateral, and transverse. Rotational diffusion, the spinning of a single molecule around its axis, affects a lipid's interactions with its nearest neighbors but does not alter its position. Lateral diffusion occurs when neighboring molecules exchange places via Brownian motion; it enables lipids to travel within a monolayer. Transverse diffusion is the exchange of lipid molecules between leaflets and is commonly called "flip-flop." The fluid aspect of the Fluid Mosaic Model is primarily due to lateral difr'usion of membrane components. One technique used to measure the rates of lateral diffusion of lipids is called FRAP, fluorescence recovery after photobleaching (see Box 2.1). The diffusion rates obtained for freely diffusing lipid molecules

The Lipid Bilayer Matrix

25

2.12. A snapshot from a simulated model of a fully hydrated DMPC bilayer. This molecular dynamics simulation shows clearly the disorder among the acyl chains (green). The starting point for the simulation was the lipid configuration from the x-ray crystal structure, which was then "heated" to a constant temperature and pressure. Note the penetration of water molecules (blue and white) into the extensive interfacial regions, while water is absent from the nonpolar center Phospholipid headgroups are orange, water hydrogens are white, water oxygens are blue, and phospholipid hydrocarbon chains are green. From Chiu, S. W, et al., Biophys. J 1995,69:1230-1245. :1=; 1995 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

are very fast, typically around 10- 7 to 10- 8 cm 2 /sec. This is fast enough for a single lipid molecule in the erythrocyte plasma membrane to go around the whole cell in seconds. Clearly, such a fast rate of diffusion can be expected to randomize the positions of lipids in the bilayer, leading to the concept that most of the bilayer consists of undifferentiated bulk lipids. However; rates of .lateral diffusion of PLs in membranes differ widely in various cell types and are significantly retarded (by factors up to 100) compared with rates in model membranes. Newer techniques observe lateral diffusion by single-particle tracking using very sensitive video fluorescence microscopy to follow the motion of a single fluorescent lipid molecule on the surface of a cell. These methods reveal an irregular path for lipid motion in the plane of the membrane, called "hop diffusion" because the tagged molecule St
diffusion are observed in plasma membranes from many cell types and are likely to be formed by interactions with the cytoskeleton (see Figure 4.3). While latera] diffusion in the membrane is fast, transverse diff'usion is very slow: lipid molecules do not readily flip from one leaflet to the other because it is energetically unfavorable fOI- their polar headgroups to pass through the nonpolar center of the bilayer. When the rate of transverse exchange (flip-flop) of lipid mo.lecules between the two leaflets of the bilayer is measured in model membranes called liposomes (see Chapler 3), the half-times are several hours or more in the absence of proteins or olher defects. A pH gradient can stimulate some lipids, such as PG and PA, lO cross the bilayer. Some biological processes, such as incor'poration 01' newly synthesized .lipids into membranes, require a much fasler rale of Iransbilayer movement. For Ihis purpose, lipid Iransfer is carried oul by flippases, enzymes Ihal calalvze flip-flop al Ihe cost of adenosi ne tri phospha le (ATP) hydrolvsis.

BOX 2.1. Fluorescence techniques A number of spectroscopic methods make use of probes or derivatized biomolecules that fluoresce; that is, they absorb energy to reach an excited electronic state and then emit radiation (photons) when they return to the ground state. The excitation wavelength, which is the wavelength of incident light required to excite the fluorescent molecule, depends on the nature of the fluorophore, the energy-absorbing group. The emission spectrum is the variation of fluorescence intensity with the wavelength of the emitted light, and is always at a lower frequency (higher wavelength) than the excitation spectrum. This difference between the excitation wavelength and the emission wavelength increases the sensitivity of fluorescence 1 DO-fold over absorption spectroscopy simply because the detector on the instrument is set for the emission wavelength and «>00000OO does not pick up background from the light source. Because Cell the excited f1uorophore can interact with surrounding solvent molecules before emission, both the intensity of the emission and its maximum wavelength (A max ) are sensitive to the environment of the molecule. In general, movement of the fluorescent group from a nonpolar environment to an aqueous milieu will decrease the intensity and shift the A max to a higher wavelength. ~ A variety of techniques employ fluorescence in membrane research. Fluorescence depolarization measures rotational diffusion and thus quantitates viscosity (the inverse of fluoOooooeo<> Fluorescent idity). It requires excitation with plane-polarized light and probe on observation through analyzing polarizers to resolve the flulipids orescence intensity into parallel and perpendicular compoView surface with nents. Fluorescence recovery after photobleaching (FRAP) Auorescence uses fluorescence microscopy to follow fluorescence intensity microscope over time after a laser beam destroys the fluorophores in a small observation area. The rate of recovery of fluorescence is a measure of the rate of diffusion of unbleached molecules into the bleached area. FRAP was used to investigate the latIntense laser eral diffusion of lipids in cell membranes using fluorescent beam bleaches probes attached to PL headgroups. Within milliseconds, the bleached patch of membrane recovered its fluorescence as unbleached lipid molecules diffused into it and bleached lipid molecules diffused away (see Figure 2.1.1). Many experimental designs make use of specific quenching processes to provide additional information on the location of the fluorophore. Effective quenching decreases the fluorescence intensity. Quenchers such as oxygen; heavy ions, including iodide and bromide; or other paramagnetic molecules remove the excited state energy upon collision With time, unbleached with the fluorescent molecule. An example is the use of phosphospholipids pholipids with bromide bound to carbon atoms at differdiffuse into ent positions of the acyl chains to probe the depth of the bleached area • '" """ I.e. bilayer occupied by a fluorophore (as described in Chapter ~ .-.,1 7). Fluorescence resonance energy transfer (FRET) involves a second type of quenching process that does not require Measure rate collisions. In this process, the fluorophore interacts through of fluorescence a short distance (1-50 A [0.1-5.0 nm]) with an acceptor return molecule with similar electronic properties. As the excited donor molecule passes its energy to the acceptor, it does not emit a photon. Because the acceptor is now excited, it can 2.1.1. Measurement of lateral diffusion of lipids by FRAP. From decay to the ground state, emitting a photon with a longer Nelson, D. L., and M. M. Cox, Lehninger Principles of BiochemA max . If the fluorescent probe is not close enough to an accepistry, 4th ed. New York: W H. Freeman, 2005. © 2005 by W H. tor, its emission will not change. Freeman and Company. Used with permission.

-~

--

j

_,m~]

j

E]

~

Lipid Asymmetry and Membrane Thickness

Because the transverse exchange of lipids between monolayers is slow, the lipids do not readily equilibrate between them. In addition many cells use ATP-driven

transpon of lipids catalyzed by flippases to maintain different lipid compositions in the inner and outer leaflets of their membranes. Analysis of this asymmetry has used phospholipases that cannot permeate the membrane and therefore only hydrolyze lipid substrates

The Lipid Bilayer Matrix

\-----1 ~ 'I.~r~

27

33 ms Resolution (lOs Observation)

~~

500nm

DF-mlsh

B.------------11 0 Resolution 500 nm

25 ms

~s

,Finish

40 ms

(50 -100 ms Observation) 6 ms 20 ms

.

2.13. Hop diffusion of individual lipid molecules. Computerized time-resolved single-particle tracking was used to follow the diffusion path of a single gold-labeled DOPE molecule on the surface of the cell. A. At a resolution of 33 msec, the path appears to be simple brownian diffusion. Each color represents 60 step periods or 2 seconds. B. At a resolution of 110 microseconds (J-lS), the pattern of movement reveals the phenomenon of hop diffusion as the lipid hopped from one region to the next. Each color indicates confinement within a compartment, with black for intercompartmental hops. The residency time for each compartment is indicated. From Murase, K., et al., Biophys J. 2004, 86:4075-4093. © 2005 by W H. Freeman and Company. Used with permission.

from the outer leaflet, as well as chemical labeling with nonpenetrating agents such as trinitrobenzenesulfonic acid (TNBS). A good example of lipid asymmetry (and the first observed) is the erythrocyte membrane, whose outer leaflet is enriched in sphingomyelin and PC, while the inner leaflet contains most of the PE and nearlv all of the PS found in the membrane (Figure 2.14). Furthermore, the same phospholipid species may have acyl chains in the outer leaflet different from those in the inner leaflet. The concentration of sphingolipids is typically six-fold higher in the outer leaflet of membranes than in the inner leaflet; in contrast, cholesterol is commonly distributed in both leaflets of eukaryotic membranes. The consequences of lipid asymmetry are being explored in vitro with new techniques that produce supported bilayers with asymmetric lipid composition (see Chapter 3). Numerous in vitro studies have shown that the two leaflets of a lipid bilayer can be coupled together by interdigitation produced when some of the acyl chains extend past the bilayer midplane, pushing their termi-

nal methyl groups into the opposing leaflet. This may result from chain length asymmetry within individual lipid molecules (when a lipid bears one acyl chain that is much longer than the other), as frequently occurs in sphingolipids. Because the tlVO monolayers become physically coupled, interdigi tation may be observed as a distinct phase in calorimetric studies and as line broadening in the 31 P-NIVIR spectrum (see belo'.v). An important consequence of interdigitation is a decrease in the bilayer thickness, because it allows the two monolayers to approach each other more closelv. The thickness of the lipid bilayer is strongly influenced by the number of carbons and degree of saturation of acyl chains. In addition, much experimental evidence indicates that cholesterol increases the thickness of a Ji pic! bi layer. Th icken ing by cholesterol is attributed to stabilizing the neighboring acyl chains in their most extended conformations (with all anti dihedral angles). thus increasing their effective length (see Figure 2.9A). However. a challenge to this vie\-v is presented by experiments that measure the thickness of various cell

Percent 01 lOtal Membrane

membrane

Distribution in

phospholipid

phospholipid

membrane Inner

Outer

monolayer

monolayer

100 Phosphatidylethanolamine

30

Phosphatidylcholine

27

Sphingomyelin

23

Phosphatidylserine

IS

a

100

Phosphatidylinositol Phosphatidyli nositol 4-phosphate 5 Phosphatidylinositol 4.5-bisphosphate Phosphatidic acid 2.14. The asymmetric distribution of lipids in erythrocyte membranes. The graph shows the content of each lipid type expressed as mol % in the inner and outer leaflets. Redrawn from Nelson, D. L., and M. M. Cox (eds.l. lehninger Principles of Biochemistry, 4th ed., W H. Freeman, 2005, p. 373. © 2005 by W H. Freeman and Company. Used with permission.

Hexagonal

Lamellar

Hu

La

HI

c.

B.

A.

Hexagonal

Cubic Q

( yo'/f!!!! •••••••17

D.

E.

2.15. Structures of lamellar, hexagonal, and cubic phases, the most common polymorphic states observed with membrane phospholipids. Lamellar phase (B) is La. Hexagonal phase is either normalHI (A), with nonpolar regions inside the tubes - or inverted - H II (C), with polar groups and water inside. Cubic phases are three-dimensional systems of lipid channels or networks interpenetrated by water channels, represented by the bicontinuous type (0) and the micellar type (E) that occur in excess water. Redrawn from Gruner,S., Nonlamellar lipid phases, in P. L. Yeagle (ed.), The Structure of Biological Membranes, 2nd ed., CRC Press, 2005, p. 174. © 1997 by Academic Press. Reprinted with permission from Elsevier.

membranes by x-ray scattering and find thickness is not correlated with their cholesterol content but rather seems to be strongly inOuenced by the protein content (see "Hydrophobic Mismatch" in Chapter 4).

Lamellar Phase

The commonly observed lamellar states are called La and L(\: La = lamellar liquid CJystalJine, also called Ld (or ld, liquid-disordered)

LIPID POLYMORPHISM

The bilayer is only one of the possible lipid aggregates that form spontaneously when amphiphilic lipids are mixed with water: Different compositions and/or changes in conditions bring about polymorphic (from polv+mo/71h. "many shapes") phase changes. The three general categories of lipid phases are lamellal~ hexagonal, and cubic (Figure 2.15). The familiar bilayer of the membrane is lamellar, yet the presence of lipid components that prefer hexagonal phase has turned out to be crucial for many of its functions (see below). The cubic phase has received much interest as a likely intermediate in membrane fusion and a medium for crystallization of membrane proteins.

Lp = lamellar- gel, also called So (ordered solid) The highly ordered, dehydrated PL arrays characterized by x-ray crystaJJography (Figure 2.1 0) are in a third lamellar phase: Lc = lamellar crystalline. (See the chapter Frontispiece for a compal-ison of the La. L p , and L c states.) The melting temperatures that were described earlier for transitions from "solid" (gel) to "fluid" (liquid crystalline) states characterize the L p to La transition (Figure 2.16). Additionally. some pure PLs exhibit minor transitions between Lp and L", with phases called L~', in which the chains are tilted, and P r3 and P r,', which are ripple phases seen with pure PC. Figure 2.16B shows why the term ripple is appropriate.

Lipid Polymorphism

29

A.

B. a

I

I

b

1/ililililililililililiitU/ii It!ililili/i/i/i/t/filt/t/ilililililtt,

I

IT 1J7J7J7J7J7J7J7J7J7J7J7J7J7J7J7~ 17;1;1;7;7;7;7;7;7m;1;1;1;1;7;7;7;7;

~~u~~u~~~~u~~~~

~~A~~~,~n~~~~~~

T

I-Je «JtltiLltil/tltltltltltltltlt-w t/t!filill/I/I/I/i/I/I/I/i/i/I/i/Uuu ~uuu~~~~~~~~~~ ~ L 13 '

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2.16. Differential scanning calorimetry of an aqueous dispersion of DPPC. A. The excess enthalpy (heat taken up) of the sample compared with a reference is measured as the temperature is raised. DPPC exhibits two phase transitions. The first is a pretransition called Tml, which produces the ripple phase P~', followed by the transition T m 2 to Ln , as diagrammed schematically above the graph. © 1976 by American Chemical Society. Reprinted with permission from American Chemical Society. Main image: © 1972 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center. B. Electron density map for the ripple phase of DMPC with 25% water at 18"C at high resolution achieved by x-ray diffraction. From the dimensions of the unit cell (drawn in white), the rippling repeat period is 142 A and the lamellar repeat is 58 A. However, the thickness of the hydrocarbon varies, being different at A and B; the thickness of the water layer between bilayers is shown at C. Redrawn from Nagle, J. F., and S. Tristram-Nagle, Curl' Opin Struct Bioi. 2000, 10:474-480. (C; 2000 by Elsevier. Reprinted with permission from Elsevier.

Hexagonal Phase and the Amphiphile Shape Hypothesis Hexagonal phases consist of hexagonally packed arrays of lipids in long cylindrical tubes. They have t,vo topologies (see Figure 2.15A and C): HI = cylinders with nonpolar centers and polar groups and water outside H I1 = cylinders with polar groups and water inside, nonpolar groups outside

Because its orientation is opposite the usual bilayer orientation, H I[ phase is called inverted. Lipids that prefer hexagonal phases are important constituents of biological membranes, as described below. What determines whether lipids form lamellar or hexagonal phases under certain conditions? The amphiphile shape hypothesis suggests the lipid aggregates formed by aqueous dispersions of pure PLs and other amphiphilic compounds reflect the general shape of the individual molecules (Figure 2.17).

in. C.vlindel·

Bilayer

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2.17. The amphiphile shape hypothesis: the general relationship between lipid shape and aggregates, which influences lipid polymorphism. The shape parameter, 5, is the ratio of the volume to the area of the polar headgroup times the length. When 5 ~ 1, the lipid is roughly cylindrical and in aqueous dispersion can form stable bilayers (lamellar phase). When 5 > 1, the lipid tends to organize into micelles or HI; when 5 < 1, the lipid forms inverted 'nicelles or H II . L, lipid; W, water. Redrawn from Jain, M. K., and R. C. Wagner, Introduction to Biological Membranes, 2nd ed. New York: Wiley, 1988, p. 53.

Bilayer-forming lipids such as PCs that favor lamellar aggregates are roughly cylindrical in shape, with similar cross-sectional areas for the heaetgroup and the acyl chains, This can be described by a shape par'ameter, S, calculated as lipid volume/(cross-sectional area of polar headgroup x lipid length). FN a fairly cylindrical lipid, S equals ~J. A lipid that is conical (S > I) or wedge shaped (S < 1) introduces curvature, leading to the fonnation of non lamellar phases. For example, DOPE, with its small headgroup and unsaturated acyl chains, is conical and forms the H I [ phase at temperatures at which the mOl'e cylindrical DOPC is lamellar. The physical situation in a bilayer is more dynamic than that implied by the amphiphile shape hypothesis because of the mobility of acyl chains above the temperature of their L~ to La transition. Bilayer-forming lipid molecules in the fluid L" phase are not confined to cylindrical spaces, as illustrated by the bilayer snapshot in Figure 2.12. Thermodynamics provides a more complete account of the shape concept. The free energ~f per lipid molecule differs in lamellar and hexagonal phases, in which the molecule occupies different volumes. This shape-dependent Free energy has Four components: hydrocarbon-packing energies, the elastic bending of the lipid monolayel-s, hydration, and electrostatic potentials: The hydrocarbon-packing energies, due to . See Gruner (1985) ror a quantitative treatment.

the hydrophobic effect, depend on the extent of hydration. In the H I1 phase with water molecules sequestered inside the cylindrical tubes, inueasing the water-tolipid ratio increases the hydrocarbon-packing free energy. The elastic bending of lipid molecules describes their tendencies to form curved monolayers. A lipid monolayer in lamellar phase is essentially flat, whereas in hexagonal phase it is tightly rolled into cylinders, When the decrease in elastic energy that results from curling the layers competes favorably with the increase due to packing the hydrocarbon chains, the system undergoes the La to H II transition to lower the total free energy. The elastic bending of a monolayer is described in terms of R, the radius of curvature of the lipid/water interFace (Figure 2.18A and B). R o is the intrinsic value of R for each lipid species - that is, the radius of CUfvature it would reach at equilibriulll in the absence of other Forces. When other forces drive the lipid A Negative curvature

Positive curvature

~

~

Zero curvature

mmmnt B

C

Interfacial

•

Tension

t

Fh

-- "Y--

Fe ..

)i

t~

2.18. Curvature of a lipid monolayer. A. R is the radius of curvature of the lipid/water interface, which is defined as positive for HI phase (right) and negative for H II phase (left). B. A larger value

of R produces less curvature than a smaller value of R. C. The zero curvature of one leaflet of a lamellar phase is the result of a balance of forces: Fe is the lateral pressure pushing the chains apart due to motions of bond rotation, and Fh is the lateral pressure in the headgroup region that consists of steric, hydrational, and electrostatic effects, in addition to some positive contributions from hydrogen bonds. Fy is the result of the hydrophobic effect at the interface, where the interfacial tension minimizes the hydrocarbon-water contacts. Redrawn from Lee, A. G., Biochim Biophys Acta. 2004, 1666:62-87. © 2004 by Elsevier. Reprinted with permission from Elsevier.

miscibility of Bilayer Lipids molecules into a phase with a different R value, R-R o is an indication of how far they lie from their intrinsic curvature. Factors that widen the splay of the lipid tails (e.g., temperature or unsaturation) decrease R o , while factors that increase the effective headgroup area (e.g., size and charge of the headgroup) increase Ro. In general, PC species have larger Ro values and remain in the L", phase at higher temperatures while PE species have lower R o values and go into H II phase at those temperatures. A mixture of the two has an intermediate Ro value. However, an aqueous mixture of DOPE and DOPC will adopt a lamellar phase with just over 20 mol % of the bilayer-forming lipid. When a bilayer restricts the tendency of the lipids of each monolayer to curve, the forces exerted on the lipid create a state of "CUl-vature frustration." The frustration is the result of the lateral pressures pushing apart the acyl chains and/or the polar headgroups within the bilayer, countered by the hydrophobic effect stabilizing it (Figure 2.18C). Their intrinsic curvature explains a role for nonbilayer-forming lipids in the membrane. The average R o of a biological membrane made up of a variety of lipids should not be too small- to avoid destabilizing its lamellar structure - but also should not be too large - to allow the transient local perturbations of lamellar structure needed for fusion, endocytosis, or a similar event. Thus membranes of many organisms contain a significant fraction of non-bilayer lipids, which the organism varies homeostatically to control the curvature of its membranes (see below). Cubic Phase In addition to lamellar and hexagonal phases, Iipids can fonn cubic phases (Figure 2.ISD and E). Like hexagonal phases. cubic phases are type I (positive curvature, acyl chains inside) and type II (negative curvature, acyl chains outside). Cubic phases have a much greater voriety of three-dimensional structures as they are formed from cubic packing of rod-like elements, resulting in discontinuous phases. Various geometries of periodic minimal surfaces are fanned by different lipids in aqueous solvents. Only two of the cubic phases - 0 224 and 0 227 - can exist in excess water. 0 224 is bicontinuous, containing t,vo networks of rods, each with tetrahedral joints providing connections (Figure 2.1SD). The walls of the rods are curved bilayers, with water on either side. 0 227 has quasi-spherical micelles packed into cubes (Figure 2.1SE). Because the frustration of bilayer curvature in cubic phases is less than that in lamellar and hexagonal phases, cubic phases can form in the transition between L", and HI! phases. Cubic phases have received much attention as putative intermediates in membrane fusion and lamellar to hexagonal (H II ) phase transitions, as well as providing novel environments for crystallizing membrane proteins.

31

MISCIBILITY OF BILAYER LIPIDS

The polymorphism of mixtures of pure lipids is summarized in diagrams that show the phases of a twocomponent system as a fLlnction of temperature and the mole fraction of one component (see Box 2.2). Such BOX 2.2. Phase diagrams Simple phase diagrams show the phases of a pure substance as a function of temperature and pressure. To learn more about phases in biological systems, which are more complex but are normally at a constant pressure, phase diagrams can be applied to simple mixtures, showing their phases with temperature on one axis and the composition of the mixture - in terms of the mole fraction X of one component - on the other. When the two substances in the mixture are miscible in both solid and fluid phases, as are many combinations of two similar phospholipids, the diagram looks like the example in Figure 2.2.1. The x-axis gives the mole fraction of the component, N, which has the higher melting point. At higher temperatures, the mixture is liquid at all mole fractions of N (L region, above both curves); at lower temperatures, it is solid (S region, below both curves). Between the curves is a region where both solid and liquid coexist (L + S). As the temperature is lowered for a particular composition - for example, starting at position x - the liquid pool is depleted of the higher melting point component. At point Q between t1 and t2, both phases are present, with the overall mole fraction of x. However, the tie line from P to R gives the mole fraction in each phase: Xp is the mole fraction of N in the liquid, and XR is the mole fraction of N in the solid. This type of phase diagram is observed in parts A and B of Figure 2.19. If the two phospholipids are sufficiently different, they may be immiscible in the liquid phase or in both liquid and solid phases, as observed in parts C and D of Figure 2.19.

2.2.1. Phase diagram for a mixture of two substances, designated M and N, that are miscible in both liquid (L) and solid (5) phases. From Lee, A. G, Biochim Biophys Acta. 1977, 472:237-281. © 1977 by Elsevier. Reprinted with permission from Elsevier.

B.

A.

DEPC-DSPC

60 DEPC-DPPC

50

50

40

40

30

30

iJ0- 20

20

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2-.

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00

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2.19. Phase diagrams of aqueous binary PC and PE mixtures that were determined from the mobility of the lipid-soluble nitroxide spin probe TEMPO (inset). A and B. The DEPC-DPPC and DEPC-DSPC mixtures show regions of fluid (f), gel (g), and liquid-solid (f+ g) miscible phases as described in Box 2.2. C. The DOPC-DPPE mixture shows two different liquid phases coexisting with a solid (fl + g and f2 + g). D. The DE PC-DPPE mixture shows these regions and an additional one consisting of immiscible liquid phases (f, + f2). Redrawn from Wu, S. H., and H. M. McConnell, Biochemistry. 1975, 14:847-854. © 1975 by American Chemical Society. Reprinted with permission from American Chemical Society.

diagrams are constructed using data obtained with a variety of probes that can detect the immiscibil ity of two phases, such as electron par'amagnetic resonance (EPR, see Box 4.2), which detects the mobility of spin probes, compounds carrying unpaired electrons. Early examples, constructed using EPR data on the mobility of TEMPO, a smaJi lipid-soluble spin probe whose nitroxide group has an unpaired electron, are shown [or different aqueous binary dispersions of two PLs (Figure 2. J 9). The phase diagrams for mixtures of DEPC and DPPC (Figure 2.19A) and for OEPC and OSPC (Figure 2.19B) indicate they are completely miscible, with ideal mixing in both fluid and gel states and with a solid state coexisting with a fluid state at intermediate temper'atUI'es and compositions. This ideal mixing suggests that the two lipid molecules are completely interchangeable and is limited to PLs with acyl chains that differ by less than [our methylene residues. The phase diagrams for DOPC

plus DPPE (Figure 2.19C) and for DEPC plus DPPE (Figure 2.19D) show immiscibility in fluid plus gel states and even immiscible fluid states (in Figure 2.19D). An intermediate situation occurs when the two lipids are miscible in the fluid state and immiscible in the gel state, as seen in mixtures of DMPC and DEPC probed with FRAP. The fluorescence probe NBD-DLPE, a PE labeled with the nitrobenzoxadiazolyl group, partitions almost exclusively in the liquid crystalline phase. When added to bilayers consisting of varying proportions of DMPC and DSPC, the extent of recovery after photobleaching has revealed the discontinuity between the gel and liquid crystalline phases. Similardiscontinuities have been observed with the techniques of singleparticle tracking and optical tweezers. Ternary phase diagrams for aqueous mixtures o[ cholesterol and two other lipids have been constructed using several techniques, including confocal

Lateral Domains and Lipid Rafts

33

Cholesterol

0.66

/----c---------'\

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0.30 IIIIIIIIIIIIIII!llllllllI/gllllllllllllllllll!!fl.111111111111 0.25

=

E

D

~

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A (La) DLPC

~

0.25

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2.20. Ternary phase diagram of DLPC, DPPC, and cholesterol at constant temperature (24 C). Each vertex represents a pure component in excess buffer, and each axis corresponds to the mole fraction of one component: X chol for both PC: cholesterol axes and X DPPC i~ P for the bottom axis. Seven regions are identified, labeled A to G, based on experiments carried out along the trajectories of the four dotted lines Clnd the bottom axis. RedrClwn from ChiClng, Y W, et al., Biophys J. 2004, 872483-2496. © 2004 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

fluorescence microscopy, fluorescence resonance energy transfer (FRET, see Box 2.1), and EPR using a spin probe at the end of a 16-carbon acyl chain. In addition to the phases La and L~\ (13mellar liquid crystalline and lamellar gel) and crystalline lipid, these techniques detect a "liquid-ordered" state, called L o , which occurs as a result of the close interactions between cholesterol and PLs (see Figure 2.9), 3S well as between cholesterol and sphingolipids. In Lo the acyl chains of the lipid are extended and tightly packed as in the L1\ state, but they exhibit rates of lateral diffusion close to that of lipids in L". To contrast with the Ln state, La is also called the liquid-disordered state (LJ). The diagram for the mixture of DLPC, DPPC, and cholesterol at 24 C describes seven regions, labeled A to G (Figure 2.20). Atlo\ov concentrations of cholesterol, the transition from La (A), which is rich in DLPC, to Lf3 (C), rich in DPPC, goes through an intermediate area (B) where both La and Lf3 are present. At higher concentrations of cholesterol (mole fTactions from 0.25 to 0.66, area F), the Lo phase occurs: Even higher concentrations of cholesterol produce crystalline cholesterol in a cholesterol-saturated lipid lamellar phase (G). At intermediate mole fractions of cholesterol (between 0.16 Q

" At these concentrations, cholesterol can interact with phospholipids to produce cooperative condensed complexes, as described above. Because the cholesterol in these complexes confers rigidity upon the lipids, it is difficuiL to distinguish between a phase that has microscopic phases containing condensed complexes and two immiscible liquid phases, such as Lo and Ld.

and 0.25) are two areas (D and E) where more than one phase coexists. The area labeled E has both ordered and disordered phases (L" or Ld and Lo ), while D appears to have L rJ and another unidentified phase. Coexistence of Ld and C, phases is also observed in the philse diagram constructed for cholesterol, sphingomyelin, and PO PC at compositions that mimic lipid rafts (see below). Physical studies of lipid mixtures cleilrly indicate that nonideal mixing occurs 3S 3 function of composition and temperature and demonstrate the possibility that diffel-entlipid domains coexist in bilayers composed of much more complex lipid mixtures. Recently a great deal of atlention has focused on the properties and roles of segregil ted lipiJ domains in biomembranes, generally called lipidmfts.

LATERAL DOMAINS AND LIPID RAFTS

While the Fluid Mosaic Model lor membrane structure emphasizes the fluidity of the bulk lipid phase of the membrane, allowing random diffusion of its components not bound by the cytoskeleton, Singer and Nicolson did ilcknowledge the possibility of small membrane domains. Particular cases or lateral organiz3tion in the membrane have long been recognized. For example, buckling of membrane-enclosed viruses occurs in select regions 01' the hosl membranes, and epithel ia I cells have diCferent lipid (and protein) compositions in their apical and basolateral domains, that is, in the portions of

~ ~

I I

61A ~ II

~ I

lo-phase lipid

= Lo

Ie-phase lipid

= L"

Raft protein (transmembrane or lipid-anchored)

I I I

t

I

I

Cholesterol

r

Nonraft transmembrane protein

2.21. Model depicting lipid rafts as distinct domains of the plasma membrane, domains in the Lo phase that are enriched in cholesterol, and sphingolipids. Certain proteins are excluded from rafts, while others are enriched in them, conferring upon them special functions. Larger rafts may form from coalescing smaller rafts (A) or by recruitment of additional proteins and lipids (B). Redrawn from Brown, D., and E. London, J BioI Chem. 2000, 275: 17221-17224. © 2000 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

their cytoplasmic membranes that provide theil- exter~ nal (exposed) and internal surfaces on opposite ends of the cells. The question of how differenllipids are delivered to the two distinct surfaces of epithelial cells led to the concept of lipid rafts, lateral regions distinguished from the bulk lipid of the bilayer that are involved in lipid trafficking as well as protein targeting and other important biological functions (Figure 2.21). The surface of a plasma membrane on a eukaryotic cell is studded by close to a million lipid rafts, and evidence is emerging for rafts in intracellular membranes as well. Rafts may only reside in the outer leaflet of the membrane, as mixtures of lipids imitating the composition of the inner leaflet do not form rafts ill vi/roo Although there is no direct evidence I'm- co-localized raf1 formation in the inner and outer Jeaflets of the bilayer; the two leaflets may be sufficiently coupled by interdigitation of the longer acyl chains to sustain bilayer raft organization. Certain proteins concentrate in rafts, whereas others are excluded from them (see below); in fact, the clustering o[ some proteins seems

to induce raft formation or at least increase the size of ra[1s. The lipids in ra[ts have physical properties different from those of the bulk lipids. Domains enriched in glycosphingolipids and cholesterol are several angstroms thicker than the rest of the bilayec Indeed they look like rafts on the surface of the fluid bilayer when their increased thickness is detected by atomic force microscopy, producing images that validate the name lipid rafis (Figure 2.22). The raft lipids, with a preponderance of saturated acyl chains, are in the L tl state, and are thus are more ordered than the bulk lipids. Coexistence oflo and Lei phases has now been demonstrated in raft-imitating model membranes consisting of cholesterol, palmitoyl-sphingomyelin, and PO PC in 1:1:1 molar ratios (Figure 2.23). Segregated lipid domains are also observed in monolayers and bilayers made \-vith the lipids extracted from epithelial brush border membranes [yom the apical microvilli of the absorptive cells lining the intestine. Treatment of these membranes with methyl-13-cyclodextrin, which removes cholesterol, abolishes the domains. I3-Cyclodextrin appears to disrupt rafts on the surface of whole cells as well, but these experiments must be interpreted with caution as cholesterol depletion has many other effects on cells and l3-cyclodextrin has been shown to retard di[fusion of membrane proteins independent of its effect on cholesterol. Lipid rafts are diverse in terms of their composition (and therefore [·unctions), their lifetimes, and their sizes. Because membrane microdomains of different sizes form dynamically, methods to detect them employ different time scales and different length scales. The results must be integrated into a consistent model, combining observations from model membranes and from cell membranes. Depending on the method of observation used, raft sizes vary from ~50 nm to up to 700 nm. The larger rafts can be detected by light microscopy or fluorescence microscopy. However~ when immunolabeling techniques are used, antibody interactions can stimulate fusion of microdomains to create the larger sizes. Very small microdomains (containing as few as 25-50 lipid molecules) have been detected in model membranes using fluorescence quenching techniques, in which incorporation of lipid-linked bromines identifies quencher-rich and quencher-poor domains that form in response to addition of cholesterol or sphingomyelin. Rafts of intermediate sizes can now be detected with a very new application of mass spectrometry that provides quantitative images of lipid bilayer components with spatial resolution of less than 100 nm. The size of small rah domains is limited by bilayer curvature and by the domain boundary at the interface between the domain and the surrounding bulk lipid. The energy per unit length of this interface is referred to as the line /ension, and it physically determines the

Lateral Domains and Lipid Rafts

35

2.22. Observation of rafts by atomiC force microscopy, which detects the Increased thickness of the Lo domains. From Nelson, D. L., and M. M. Cox (eds.), Lehnmger PrinCIples of BIochemIstry, 4th ed., H. Freeman, 2005.

w.

-I-~~----r-~~~=t=:;=~==~==5S0~~0,0 0,2

0,6

0,8

1,0

So + Lei Mole fraction PSpM 2.23. Phase diagram for the aqueous ternary mixture of raft lipids at 23 C. The lipids, POPC, palmitoylsphingomyelin (PSpMl, and cholesterol, are vaned In concentration along each of the three axes The regions of the phase diagram are labeled Le, Ld , and So (solid, ordered) and show coexistence of Le with both Ld and So phases as a function of composition. The "raft mixture" of a 1:1'1 molar ratio is indicated by the tie line (dotted) in the Lo + Ld region Above 66 mol % cholesterol, excess cholesterol separates out in crystalline phase (dashed line). Redrawn from Simons, K., and W L. Vaz, Annu Rev B/ophys Biomof Struct. 2004, 33:269-295. © 2004 by Annual Reviews. Repnnted with permission from the Annual ReView of BiophysIcs and Blomolecular Structure, www.annualreviews.org.

Preexisting organization

Induced "rafts"

2.24. A model for the clustering of small, preexisting membrane domains into larger rafts. The clustering of small domains into relatively large rafts is actively organized in both space and time. The affinities between some lipids and proteins form preexisting structures (left) that can coalesce into rafts (right). The red and pink circles represent different nonraft lipid species, the yellow circles represent raft lipids, the green circles represent cholesterol, and the larger black circles represent GPI-linked raft proteins. The scale bar is ~5 nm. Redrawn from Mayor, $., Traffic. 2004, 5:231-240. © 2004. Reprinted with permission of Blackwell Publishing

sizes and shapes of the domains. If a large portion of the membrane is occupied by small rafts, then raft boundaries must be extensive. In this case, the line tension is small, entropy dominates, and the domains are small. If the line tension is large, it favors fusion into larger domains because when many small rafls merge into a large one, the total length of the raft boundary is reduced, thus t-educing the boundary's energy. Large rafts appear 10 encompass smaller heterogeneous domains within them and likely form by coalescence of preexisting structures (Figure 2.24). In cells, cluslering of smaller rafts into larger domains is probably stimulated by particular proteins and may have important functional or regulatory consequences. These clusters are probably transient, resulting from very dynamic formation and growth. High-speed video microscopy has recorded the formation and dissolution of small rafts (with diameters ~SO nm, containing ~ 3000 Ii pid molecules and probably only 10-20 protein molecules) with lifetimes or less than 1 msec. It is thought that certain raft proteins stimulate raft fOl~ mation, as suggested by models that posit that "lipid shells" around such proteins come together to make r·afts. In this case. raft heterogeneity is a consequence of the particular lipid-protein and lipid-lipid interactions that trigger their formation. Detergent-Resistant Membranes

The proposition that I-afts exist in cell membranes \NaS given a huge boost by methods for extraction of an L" fraction of biologicaJ membranes. Treatment of mammalian cells with the non ionic detergent Triton X100 (see Chapter 3) produces a Triton-insoluble lowdensity membrane fTaction. These detergent-resistant membranes (DRMs) are rich in cholesterol and sphingolipids, observed to be in the L" state. and enriched in

fatty acid- or GPI-linked proteins, as are rafts. According to x-ray diffraction measurements, they are 9 Athicker than nom'aft lipid bilayers. Because the DRMs are isolated at low temperatures, it is difficult to assess how much of this membrane fraction would have been in the L" phase at growth temperatures, and this adds uncertainty regarding the specificity of the procedure for raft extraction. Raft proteins are used as markers, most successfully in the case of caveolae (see below). Other questions focus on the type and amount of detergent used. Titrated addition of Triton X-IOO can disrupt preexisting L u domains, as well as induce formation of L" domains. Several different detergents produce variations in the insoluble fraction, compared with the "raft selectivity" of Triton X-IOO; the heterogeneity of the resulting DRMs may not reveal anything more than the selectivity of the detergents for subsets of lipids and proteins. Non-detergent methods to isolate rafts employ sonication, which may cause undesirable mixing of membrane components and, in fact, give a DRM fraction enriched in ar-achidonic acid-containing plasmalogens, which are polyunsaturated lipids that would be excluded from typical rafts. DRMs contain specific types of proteins, most of which are also detected in lipid rafts by other techniques. These raft proteins include GPI-linked proteins; doubly acylated proteins, such as tyrosine kinases of the Src family and GiX subunits of heteromeric G proteins; and certain TM proteins. (See Chapter 3 for a description of the proteins.) The proteins enriched in lipid rafts suggest they have special biological functions, such as signal t1-ansduction and protein trafficking. Logically, the segregation of signaling proteins in rafts could speed the rates of their interactions \Nith other raft proteins and slow their interactions with non raft proteins. Furthermore, treatment with f3-cyclodextrin to disrupt rafts by removing cholesterol causes defects in some

Diversity of Lipids

A.

37

DSM

DRM

1

48 A

j B. 0.15 •

P-23

o P-29

:g 20.10 (5

5 u

;g, (j)

] 0.05 0, (j)

c..

0.00

-'--li=-=-_-'--.............._....IDSM DRM

....._

L-----L~_

DSM

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DRM

\.~--~ ---~)

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4°C

37°C

Y

Y

2.25. Sorting peptides based on their TM lengths. Insertion of synthetic peptides of different lengths suggests their TM lengths could direct their segregation into rafts, or into DRMs in vitro. A. Two peptides, P-23 and P-29, with 17 and 23 nonpolar amino acids, respectively, were designed to match the transbilayer thickness of DRMs and DSMs (detergent-soluble membranes) obtained from x-ray diffraction analysis. B. The ratio of peptide to total lipid for each peptide in DRMs and DSMs at 4"( and 37'( shows more P-29 than P-23 inserts into DRMs at both temperatures. Redrawn from Allende, D., et al., Trends Biochem Sci. 2004, 29:325-330. © 2004 by Elsevier. Reprinted with permission from Elsevier.

signaling pathways. Among the tyrosine kinases associated with rafts are the receptors for epidermal growth factor (EGF), and treatment of fibroblasts in culture with EGF causes its receptor to leave the raft. The dynamic process of signal transduction could thus take advantage of constant change in raft constituents as well as size. Protein trafficking through the Golgi apparatus appears to use rafts to sort membrane proteins, because domains rich in cholesterol and sphingomyelin form in the Golgi and become vesicles targeted for the plasma membrane. One of the factors in protein localization could be the thickness of the TM domains (see "Hydrophobic Mismatch" in Chapter 4), because the increased thickness of these domains could help "raft proteins" partition into them (Figure 2.25). Thus proteins with shorter TM domains could be excluded from rafts and retained in the Golgi, while those \-vith longer

TM domains partition into rafts that move to the plasma membrane. A minor fraction of the DRMs contain caveolae, small invaginations in the membrane associated with the protein caveolin in addition to other raft proteins. Caveolae may function in both protein trafflcking and signal transduction. Although caveolae are considered lipid rafts, they are a special case, because caveolin inserts from the cytoplasmic side, which is not enriched in raft lipids, and then oligomerizes to force an inward curvatLIl-e of both leaflets of the membrane (Figure 2.26). A similar process may occur when membrane domains segregate for viral envelope formation. The envelope of influeni'a virus is enriched in cholesterol and sphingomyelin compared with the host membrane from which it is formed, and its formation involves the clustering of specific glycoproteins before budding. It is believed that a matrix protein, M I, docks at the inner leaflet, selectively binds the glycopmteins, and polymerizes to induce the cun'ature that triggers the budding process. Whether lipid rafts are defined as domains of L" phase floating in the Lei membrane, Triton X-IOOinsoluble membrane f.-actions of low density, or heterogeneous microdomains of membrane containing proteins involved in signaling and trafficking, they clearly have profound effects on the nature of the membrane. The presence of rafts may also contribute to the need for lipid diversity, to support the dynamic formation of such stable but iluidlipid domains segregated from the bulk bilayer lipids and to maintain permeability barriers at their boundaries.

DIVERSITY OF LIPIDS

The typical biological membrLllle contains hundreds of lipid species when their particular acyl chains are considered, and the tvpes of complex lipids predominant in memb.-anes from different sources vary significantly. An obvious example is the absence of sterols and sphi ngoJipids from prokaryotic mem branes. Among phospholipid species, PG and CL are found in bacterial membranes but not in eukalyotic membranes other than mitochondria. E. coli normally lacks PC but grows well when carrying the Rhodobacter sphaeroides gene for the PE methylase, producing PC to levels as high as 20% of its total PL. Given these variations, what are the requirements for diverse PLsJ This question has been addressed using mutations affecting the PL biosynthetic pathway (see FigLIl-e 2.27) to manipulate the PL composition of E. coli membranes. In E. coli, the normal PL composition is 70% to 80% PE, 20% to 25% PG, and _5% CL, all of which are produced from phosphatidic acid in a few enLymatic steps. The anionic lipids are essential, as psgA- null

Plasma membrane

Outside

Caveolin dimer (six fatty acyl moieties) 2.26. Formation of caveolae by insertion of the specific protein caveolin. Caveolin monomers linked to three acyl chains insert into the membrane from the cytoplasm. When they dimerize, they force an inward curvature that leads to budding of the membrane. The inset shows a thin section electron micrograph of caveolae from fibroblasts (with arrows pointing to the ER). Main figure: Redrawn from Nelson, D. L., and M. M. Cox (eds.), Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005. '1':) 2005 by W. H. Freeman and Company. Used with permission. Inset: From Anderson, R. G., Ann Rev Biochem. 1998,67:199-225. (1;. 1998 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, wwwannualreviewsorg.

mutants that cannot make PG and CL alT not viable. The negaLively charged membrane is required for initiation of DNA synthesis, attributable to the binding of PG by the DnaA protein. Other examples of proteins that have affinity for anionic PLs include type I topoisomerases, some DNA polymerases, SecA protein (which is involved in protein translocation, see "Biogenesis of Membrane Proteins" in Chapter?), and some signaling proteins. On the other hand, PE does not seem to be essential in spite of its normal abundance: pssA - null mutants, with <0.1 % PE in their membranes, are able to grow in rich medium supplemented with divalent cations, under which conditions the membrane is 90%

PG and CL. However, these mutants grow poorly on defined minimal media and show defects (transport and motility problems, filament formation, and early entry into stationary phase) that indicate zwitterionic lipids are needed for normal membrane functions. Finally, mutants deficient in PE regulate their CL content, allowing them to maintain the proportion of nonlamellar lipids. indeed, mutants lacking both PE and CL are not viable, indicating that the membrane requires nonbilayer-forming lipid. Lipid diversity appears to be essential to reach the desired R, value and maintain the general state of the bilayer in a narrow range between the lamellar gel state and the inverted phase (L~ < L" < H Il ).

Diversity of Lipids

39

Glycerol 3-phosphate

PbB y - - AoyJ-"yl

,",,'ee pco,"'c

o II

CH 2 -O-C-R 1

I

HC-OH

I

0

II

CH2-0-P-OH

I OH PlsC

rI ~

Acyl-acyl carrier protein

o

o II

II

CH 2 -O-C-R 1

I

R 2 -C-O-C-H

I

0

II

CH 2-O-P-OH

I

OH

Phosphatidic acid

Cd'Ay--CTP

L-Serine ~ / /

CDP-Diglyceride "" PssA

r---

Glycerol-3-P

PgsA""

Phosphatidylserine

P'd~CO' Phosphatidylethanolamine

Phosphatidylglycerol-3-P

'~P; Phosphatidylglycerol Cis

r~ I

.

Phosphatidylglvcerol ~"

Cardiolipin + Glycerol 2.27. The pathway for phospholipid biosynthesis in E. coli. All PLs are formed from the activated precursor cytidine 5' -diphosphate (CDP)-diacylglycerol, from which PE is synthesized via P5, and PG and CL are synthesized on a second branch of the pathway. Note that the pssA- mutant cannot make P5 or PE. CTP, cytidine 5' -triphosphate. Redrawn from Cronan, J. E., Annu Rev Microbiol. 2003, 57:203224. © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Microbiology, www.annualreviews.org.

The role of PE in maintaInIng the polymorphism of E. coli membrane lipids is illustrated by NMR studies of 31 P-labeled phospholipids carried out in whole cells, with purified mem brane vesicles or with extracted lipids, because the shift in the 31 P-NMR powder spectra can distinguish JameJlar and HJI phases. When the total E. coli lipids are first extracted, they are lamellar (Figure 2.28). After incubation at 42°C, they shift to hexagonal phase, giving an NMR spectrum that resem-

bles that of isolated PE from the same strain. If 5% lysophosphoJipid, whose inverted cone shape complements the shape of PE, is added to the total lipid extract, the shift to H I1 at higher temperature is abolished. In other words, E. coli lipids dominated by PE are lamellar at growth temperatures and convert to H II at higher temperatures. Because PE isolated from other sources prefers HI! even at the growth temperature, E. coli must control the L~-HII transition temperature (T LH ) by the

tial for the cur"Vature of the membrane; the resistance to curvature creates pressure gradients that may be triggers for mechanosensitive channels and membraneinserting peptides such as gramicidin and melittin (see Chapters 4 and 9).

A.

BOX 2.3. Nonlamellar phase lipids and growth of E. coli Different cations affect the T LH transitions of isolated CL. Experiments using 31 P-NMR reveal specific effects of different mono- and divalent cations on CL isolated from a pss- mutant strain that has increased CL content (see Figure 2.3.1). In the 31 P powder spectra, the hexagonal-phase lipid gives a peak at around 0 ppm whereas the lamellarphase lipid gives a peak around -20 ppm. The effects of five cations are examined at three temperatures. The differences show that the cation effects involve more than just providing counter-ions for anionic lipids. The most physiological cations have the strongest effect in promoting the nonlamellar phase of the isolated CL, with the same ranking as their ability to support growth.

B.

c.

35°C

50

25

0 -25 -50 PPM

2.28. Detection of lamellar and hexagonal phases in E. colj lipids by 31 P-NMR. The lipids were extracted fro ,n a fatty acid auxotroph grown at on oleic acid, in which over half the acyl chains are C18:1 The initial NMR signal indicates the lipid is lamellar (A). After incubation at 42 C for 15 minutes (B) and 90 minutes (C), the signal shifts to that of an isotropic lipid and now resembles the NMR spectrum for HII-phase PE. Redrawn from deKruijff, B., et ai, in R. M. Epand, (ed), Lipid Polymorphism and Membrane Properties, vol 44 of Current Topics in Membranes, Academic Press, 1997, pp. 477-515. (~: 1997 by Academic Press. Reprinted with permission from Elsevier.

3rc

degree of saturation of its acyl chains. In this way E. coli membrane lipids, which al'e up to 80% PE, stay between Till (L1\-L,,) and T LH (L,,-H i1 ) to achieve the desired fluidity. Further evidence for the importance of nonJamelJar phase lipids in E. coli is provided by the cation specificity of phase transitions of CL isolated from a pss- strain (see Box 2.3). In fact, the zone from Till to TI.H transitions is imponant [or many organisms that keep their membranes close 10 T lI;. Clearly if the whole membrane passes beyond T lIi , the loss of lamellar lipids would be deleterious because of the loss of the permeability barrier. However, this does occur in cer"lain sites in mammals, such as tight junctions in poJari7.ed cells and networks of myelin in lung tissue. The possible roles for non lamellar membrane structures in processes such as membrane fusion, cell division, and gene lI-ansfer keep interest in these transitions high. As discussed above, nonbilayer lipids are essen-

50°C

75°C

NO~~~ BoJL ~ JL

s"JL ~ JL Mg~JL~ Ca 2 + ~ r.",I",,,,,,,III!!"II,I,'III

50

0 -50

PPM

JL~ I.

I"

50

"1",,111,,1,,,,1

!""I" '

0 -50

50

PPM

!

1",,1

0 -50

PPM

2.3.1. Cation dependence of cardiolipin phase transitions observed with 31 P-NMR. From deKruijff, B., et ai, in R. M. Epand (ed.), Lipid Polymorphism and Membrane Properties, vol 44 of Current Topics in Membranes, Academic Press, 1997, pp. 477515 © 1997 by Academic Press. Reprinted with permission from Elsevier.

Conclusion CONCLUSION This chapter describes the diversity of lipid stnJCtures and the polymorphic phase behavior it enables. The properties of these remarkable amphiphiles allow them to assemble spontaneously and to form the fluid bilayer that characterizes the structure of all biomembranes. From the need for nonlamellar lipids in effecting membrane elasticity to the specialized functions of lipid rafts involved in cellular communication and intracellular trafficking, membrane lipids do much more than provide the structure of the bilayer and the solvent for membrane proteins. Applications of sophisticated biophysical and biochemical techniques will undoubtedly continue to reveal their complex and crucial roles. The portrayal of the nature and diversity of lipids is essential for understanding membrane structure and fl.Jnction. Of course, the other major contributors to membrane properties are the proteins of the membrane, which have to be isolated from the membrane to characlerize them. Chapter 3 describes Ihe detergents used for this process and Ihe model systems used to reconstitute membrane mimetics. The importance of lipids will reemerge when their interactions wilh membrane proteins are described in Chapter 4 and the detailed structures of lipid bilayers are examined in Chapter 8.

FOR FURTHER READING

Reviews Anderson, R. G. w., and K. Jacobson, A role for lipid shells in targeting proteins to caveolae, raft· and other lipid domains. Science. 2002, 296: 1821-1825. Daleke, D. L., Regulation of transbilayer plasmil membrane phospholipid asymmetry. J Lipid Res. 2003,44:233-242.

41

Dowh'ln. W., Molecular basis for membrane phospholipid diversilv: why al-e there so many lipids) Am7u ReI' Bioche17l. 1997,66: 199-232. Edidin, M., The state of lipid rafts: from model membranes 10 cells. Mil/II Rev Biophvs Biol7lo! Siruci. 2003, 32:257283. McConnell, H. M., and A. Radhakrishnan, Condensed complexes 01' cholesterol and phospholipids. Biochi17l Biophl's Acla. 2003,1610:159-173. McConnell, H. M., and M. Vdjic, Liquid-liquid immiscibility in membranes. AWIII Rev Biophvs Biomo! SII"IICI. 2003, 32:469-492. Ohvo-Rekila, H., el aI., Cholesterol interactions wilh phospholipids in membranes. Prog Lipid Res. 2002, 41: 66-97. Simons, K., and W. L. C. Vaz, Model s.vstems, lipid rafts and cell membranes. AI7I711 Rev Biophvs Biomo! Sirt/CI. 2004, 33:269-295. Seminal Papers Anderson, D. M., S. M. Gruner, and S. Leibler, Geometrical aspects of the frustration in the cubic phases of lyotropic liquid crystals. Proc Nal! Acad Sci USA. 1988,85:53645368. Baumgart, T., et aI., Imaging coexisting Iluid domains in biomembrane models coupling curvature and line lension. Nnillre. 2003,425:821-824. Chapman, D., Phase transilions and Iluidit.v characteristics of lipids and cell membranes. 0 Rev Biophvs. 1975,8: 185235. Feigenson, G. W., and J. T. Buboltz, Ternaf\' phase diagram of dipalmiloyl-PC/dilaumvl-PC/cholesteml: nanoscopic domain I'ormat ion driven by cholesterol. Biophvs 1. 2001, 80:2775-2788. Gruner, S. M., Intrinsic curvature hypothesis for biol11embrane lipid composition: a mle for nonbilaver lipids. Proc NaI! Acad Sci USA. 1')85, 82:3665-3669. Jain, M. K.. and H. B. White, 3rd, Long range order in biomembranes. Adl' Lipid Res. 1977, 15: 1-60.

3

I Tools for Studying Membrane

Components: Detergents and Model Systems

One classic<J1 model membrane system consIsts of mulrilamellar veSicles, which are layered spheres of phosphohpids suspended 111 water, From Nagle, J. r., and 5 Tristram-Nagle. Bioch;rn B,ophys Acta. 2000. 1469159 195. :: 2000 by Elsevier. Rcpnnled w,lh permIssion from Elsevier.

While progress in biochemistry, biophysics, and structural biology relies on studies of purified biological components, the purification of membrane components is complicated by their amphipathic nature. First, their removal from the membrane usually requires disruption of the lipid bilayer. Once removed, they tend to aggregate in aqueous buffers due to their low solubility in water. And finally, study of the functions of many membrane components requires their insertion back into a reconstituted membrane. The critical tools that allow i/1 vitro characterization of membrane components are detergents and model membranes. Detergents are used to solubilize membrane components, removing them from the lipid bilayer and preventing their aggregation. This chapter begins with an overview of detergents, emphasizing their mechanisms of action in solubilizing membrane components. The goal of reconstitution is to insert the purified membrane component into a good mimic of the biological membrane, usually a lipid bilayer. Because 42

many aqueous lipid mixtures spontaneously assemble in lamellar phase (see Chapter 2), model bilayers tend to form spherical vesicles called liposomes. Liposomes are just one type of model system used for;'1 vilro characterization of membrane components. Even for that one type, the nature of the lipid vesicles depends on how they are made and determines their suitabil ity for different experimental techniques. By necessity, these models are simpler than biological membranes, which contain hundreds of lipid species; depending on the objective, a single lipid species can often suffice. Model membranes used to study lipid properties offer control over the stoichiometry in a mixture of t\.vo or three types of lipids. The availability of widely varying model membrane systems is critical in the characterization of purified membrane components because they allow the LIse of different experimental tools. Some membrane mimetics allow the application of the powerful techniques that have provided a wealth of information about soluble proteins. while others enable use of methods based

Detergents

'r\ ci\~ ; Oil" \

>(rl

43

{~""',

on select properties of the membrane, such as electrical conductance or pressure effects. The latter part of this chapter surveys the characteristics of the different model membrane systems, from classic systems such as black films to new technologies such as nanodiscs. Examples are provided to iJJustrate the uses of these systems in membrane research.

DETERGENTS

Detergents are defined as water-soluble surfactants, which makes them amphiphiles that are effective in the solubilization of membrane components. Their solubility in water is much greater than that of most lipids; for example, sodium dodecylsulfate (SDS) has a monomer solubility of 10- 2 M, compared with the solubilities of DPPC and cholesterol of 10- 10 M and 10- 8 M, respectively. Occasionally, detergents are considered synonymous with surfactants, because they reduce the surface tension of a liquid when dissolved in it (see Box 3.1); however; the water solubility of detergents is essential for their role in disrupting membranes. Purification of a membrane protein typically begins with solubilization of the membrane with a detergent, after which one or more types of chromatography in detergent separates the desired protein from the others. The purpose of the detergent is to prevent aggregation of the membrane protein as it is removed from its lipid environment. During these procedures, one detergent can replace another, often to maintain mild (nondenaturing) conditions. Thus it is essential to be familiar with the different detergents, their properties, and their modes of action. Types of Detergents

A variety of detergents are in common use for membrane biochemistry. While most detergents are synthetic, there are natural compounds with detergent activity, such as the bile salts that solubilize dietary fats in the intestine, and saponins, a varying group that includes the heart stimulant digitonin. Dozens of detergents are commercially available (examples are shown in Figure 3.1). They may be ionic (e.g., SDS, cetyltrimethylammonium bromide [CTABJ), zwitterionic (e.g., lauryldimethylamine oxide [LDAOJ, which varies from SDS in its headgroup), or nonionic (e.g., Triton X-IOO, octylphenol linked to a hydrophilic polyoxyethylene headgroup with an average of 9 to J 0 repeats). Most synthetic detergents have a polar or ionic headgroup and a nonpolar hydrocarbon tail, while some, like sodium cholate, sodium deoxycholate, and 3-[3-(cholamidopropyl) dimethyl-ammonio]-lpropanesulfonate (CHAPS) are derivatives of the bile salts, with a tetracyclic structure similar to that of

BOX 3.1. Surfactants and surface tension By definition, surfactants are substances that reduce surface tension. Surface tension results from the cohesive forces between liquid molecules that are unopposed by other molecules at an air/liquid interface. These unbalanced forces produce the tendency for a liquid to minimize its surface area, which is why a drop of liquid is spherical. Surface tension is measured as the work required to break a liquid film, and has the units dynes per centimeter (1 dyne = 10- 5 newtons). With its extensive hydrogen bonding, water at 20°C has a relatively high surface tension, 72.8 dynes/em compared with 22.3 dynes/em for ethyl alcohol.

cholesterol. Some commonly used nonionic detergents, such as octyl l3-o-glucoside (OG) and dodecyl l3-o-maltoside (DDM), have sugar headgroups. The wide number of detergents available allows researchers to choose from many options, often testing several for best results. While non ionic detergents like Triton X-IOO are relatively mild and can stabilize membrane proteins, some ionic detergents, notably SDS, bind strongly to proteins and usually denature them. The bile salt derivatives are much less denaturing than ionic detergents having linear nonpolar chains with the same headgroups. Short-chain nonionic detergents, including OG, denature some proteins, in which case DDM, with its longer chains, is a good alternative. Because it allows retention of biological activities and is generally very effective at solubilization, Triton X-I 00 has been widely used in studies of membrane proteins. However, its aromatic ring interferes with spectroscopic studies such as determination of ultraviolet (UV) absorbance, fluorescence, and circular dichroism, so a reduced form of Triton X-IOO is available. Several of the nonionic detergents, including Triton X-IOO, are synthesized by condensation of ethylene oxide with the parent alcohols, resulting in polydisperse mixtures containing variable chain lengths. Although the manufacturers give the average chain lengths, they do not indicate the variability, which fits a nearly Poisson distribution of oxyethylene chain lengths from one to >20, and the resulting heterogeneity can be quite deleterious, especially in crystal formation. For homogeneous alternatives, a series of alkyl polyoxyethylenes of defined chain length (CxE N , where X is the number of C atoms in the alkyl group and N is the number of oxyethylene monomers in the headgroup) is used. Commercially available detergents may have problems of impurities; for example, SDS often contains n-dodecanol and polyoxyethylene-based detergents may contain peroxide and aldehyde, which necessitates additional purification steps or the purchase of "protein-grade" or "especially purified" quality. A few detergents, including sodium cholate and

ANIONIC Sodium dodecy\ sulFale (Sodium lauryl sulFale)

Sodium dodecyl-N-sarcosinale (SodiuIl11auryl-N-sarcosinale) (Sarkosyl L) CH 3

o

0

I

II

II

~C /N 'CH /CO-Na+ 3

~O-S-O-Na+

II

II

o

o CH 3

CATIONIC Celyl lrimelhylammonium bromide (Hexadecvl lrimelhylammonium bromide) (CTAB)

I

~N+-CH3Br­

I CH 3

ZWITTERIONIC Lauryldimelhylamine oxide (LDAO) (Dodecvlamine N-oxide)

CHAP~S 0 HO

CH 3

,

I

~N+-O-

I

HO

CH 3

~H3

~

N~N+~S-O-

I

I

II

H

CH 3

0

OH

SulFobetaines (Zwiuergent bl'and)

0

CH 3

II

I

~N+~S-O-

I

II

0

CH 3 UNCHARGED

~

Digilonln

Polyoxyelhylene alcohols (denoted CxENl (I) Brij series (2) Lubrol (vVX,PX)

HoroL...J'OB Glc-GIc-Gal-Gal-Xyl-O

~(O

: H

I3-D-oclylglucoside

~~

~O

~OHO

OB

CH OH

I3-D-Dodecylmalloside (laury! maltoside)

CH 20H

OH

Fatly acid ester~ of polyoxyelhylene sorbitan (denoled C,-SOI bltan-E n ) .

CH 3 CH 3 )n -OH

CH OH 2

~2 ~ HO

0

HO

OH 0

OH

(0 CH CH ), - OH 2 2 J~

I

II

TlVe~ C -

OH

(0 CH 3 CB 3 )" - 0 - CH 2 - CH

r

(0 CH 2 CH 2)y - OH

(n = w+ x + Y + z) Alkyl-N-melhylglucamides (MEGA"" brand) CH 3 I OH OH OB

~rN;YY o

OH OH OH

(0 CH CH2)z - OH 2 Polyoxyelhylene p leu octylphenols (denoted IeI'I - C80 E,,) (I) Trion X-lOO, n = 9.6 (2) Trion X-114, n = 7.8 (3) Nonidet PAD, n= 9

~ ( O CH 2 CH 2)n - OH

BILE SALTS Sodium cholale

0

&C'ON,'

Sodium deoxycholate

HO

'OH

0

§C'ON"

II

II

HO

3.1. Structures of some detergents used to solubilize membrane components. From Gennis, R. B., Biomembranes, New York: Springer-Verlag, 1989, pp. 90-91.

Detergents

45

c.

B.

~ zCO

Na+

~O r-0 / HN HN

Y

FFFFFF

S~H F

F F

F F

F

OA~H

Ht00H OH

3.2. Three new alternatives to detergents. A. The tripod amphiphile that extracts bacteriorhodopsin. Redrawn with permission from Yu, S. M., et aI., Prot Sci. 2000,9:2518-2527. B. An example of an amphipol called A8-35 with variation in the polymeric backbone giving x = 35%, y = 25%, and z = 40%. Redrawn with permission from Gohon, Y, et al. Anal Biochem. 2004, 334:318-334. C. A hemifluorinated surfactant called HF-TAC [Cz Hs C6 H1ZCZ H 4 -S-poly-Tris-(hydroxymethyl)aminomethane] with a single hemifluorinated hydrocarbon chain. Redrawn with permission from Breyton, c., et al., FEBS Lett. 2004, 564:312-318.

l3-octylglucoside, can be purified by crystallization prior to use. Because no detergent satisfies all criteria desired by researchers, alternatives to the traditional detergents are being designed and tested (Figure 3.2). One of the tripod amphiphiles (Figure 3.2A), which have three hydrophobic tails and a hydrophilic headgroup, has been used to solubilize bacteriorhodopsin. The amphipols (Figure 3.2B) are small amphiphilic polymers of various structures that form water-soluble complexes with membrane proteins, presumably by wrapping around their nonpolar domains. Hemifluorinated surfactants (Figure 3.2C) are unlike detergents because the perfluorinated regions of their chains are hydrophobic but stilllipophobic. The nonfluorinated tails enable them to interact with membrane proteins while presumably allowing the interactions between the proteins and lipids to remain intact in aqueous solutions.

The action of most detergents involves micelle fonnation. Micelles are roughly spherical assemblies of surfactant molecules, in which most of the nonpolar tails are sequestered from the aqueous environment in a disorganized (liquid-like) hydrophobic interior. Thus the chains are not fully extended like the spokes of a wheel, and the radius of the micelle is 10% to 30% smaller than the fully extended monomer (Figure 3.3A). Furthermore, the surface is rough and heterogeneous rather than smoothly covered by polar headgroups: NMR studies of SDS micelles revealed that only onethird of the surface was covered by hydrophilic headgroups (Figure 3.3B). At high concentrations of detergent, micelles change shape to become elliptical or rod-like; this occurs at lower concentrations for surfactants with weakJy polar headgroups. Micelles of small B.

A.

(a)

Mechanism of Detergent Action

(b)

3.3. A. Cross-sectional views of detergent micelles. The old view (a) incorrectly portrays the chains as ordered like spokes, whereas they are actually disordered and fluid (b), resulting in an uneven surface. Redrawn with permission from Menger, F. M., R. Zana, and B. Lindman, J Chem Educ. 1998,75:93 and 115. B. Model of the surface of a micelle, showing the uneven surface at the detergent/water interface. Redrawn from Lindman, B. et al., in J.-J. Delpuech (ed.), Dynamics of Solutions and Fluid Mixtures by NMR, Wiley & Sons, 1995, p. 249. © 1995 by John Wiley & Sons Limited. Reprinted with permission from John Wiley & Sons Limited.

TABLE 3.1. Properties of micelles derived from some commonly used detergents Monomeric MW

Detergent Octyl-(3-D-glucoside Dodecyl-maltoside Lauryldimethylamine oxide (LDAO) Lauramido-N,N-dimethyl-3-n-propylamineoxide (LAPAO) Dodecyl-N-betaine (zwittergent 3-12) Tetradecyl-N-betaine (zwittergent 3-14) Myristoylphosphoglycerocholine Palmitoylphosphoglycerocholine 3-[[3-cholamidopropyl)-dimethylammonio]-1propanesulfonate (CHAPS) Deoxycholic acid Cholic acid Taurodeoxycholic acid Glycocholic acid Sodium dodecylsulfate (SDS) in 50 mM NaCI Dodecylammonium ClGanglioside GM , PEG-dodecanol Polyoxyethylene glycol detergents CsE 6

ClO E6 C'2 E6 C12 ES C'2 & 14 E9.5{Lubrol PX) C'2 E12 C,2E23 (Brij 35) C,6&1SE17 (Lubrol WX) tert-p-Cs0E9.5(Triton X-1 00) tert-p-Cs0R7.S (Triton X-114) C12 sorbitan E20 (Tween 20) C1S:1 sorbitan E20 (Tween 80) Cetyltrimethylammonium bromide (CTAB)

Critical micelle concentration (M)

Aggregation number

x 10- 2 x 10-4

292 528 229 302 336 350 486 500 615

2.5 1.7 2.2 3.3

393 409 500 466

3 x 10- 3 1 x 10- 2 1.3 x 10- 3

27 140 75

x 10- 3 x 10- 3

8 x 10- 2 6 9 1 5

x x x x

87 130

10- 3 10- 5 10- 5 10- 3

22 4 20 6 62

8 x 10- 3 15 x 10- 3

55

10-9

150 130

x 10- 4

394 422 450 538 620 710 1200

1 x 10- 2 9 x 10- 4 8.2 x 10- 5 8.7 x 10- 5

32 73 105 120 100 80 40 90 140

9 x 10- 5 9 x 10- 5

1000 1625 540 1240 1320 364

3 x 10- 4 2 x 10- 4 6 x 10- 5 1.2 x 10- 5 9.2 x 10- 4

60 169

MW, molecular weight. Source: Jain, M. K., and R. C. Wagner, Introduction to Biological Membranes, 2nd ed. New York: Wiley, 1988, p. 71.

TABLE 3.2. Effect of ionic strength on micelle formation by ionic surfactants in aqueous solutions at 25: C Surfactant Anionic Sodium n-octylsulfate Sodium n-decylsulfate Sodium n-dodecylsulfate (SDS) Sodium n-dodecylsulfate Sodium n-dodecylsulfate Sodium n-dodecylsulfate Cationic n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide n-Dodecyltrimethylammonium bromide

Medium

CMC (mM)

N

piN

H2 O H2O H2 O 0.1 M NaCI 0.2 M NaCI 0.4 M NaCI

130 33 8.1 1.4 0.83 0.52

58 91 105 129

018 0.12 014 013

14.8 10.4 7.0 4.65

43 71 76 78

0.17 0.17 0.16 016

H2O 0.0175 M NaBr 0.05 M NaBr 0.10 M NaBr

N, aggregation number; p, micellar charge. Source: Jones, M. N., and D. Chapman, Micelles, Mono/ayers, and Biomembranes. New York: Wiley-Liss, 1995, p. 68

Detergents

47

I I I I I I I I I I I I I /

~

2

(/)

V

..c

c: c:

.:2 0:1

'c:"" Q)

U

Micelles >,

t: V

0. 0 0. '""

c:

Monomers

.g :l

(3

------~-

C/)

/

/

/.

c:

0

u CMC

Total concentration 3.4. The critical micellar concentration. As detergent (or surfactant) is added to an aqueous solvent, the concentration of dissolved monomers increases until the critical micellar concentration (CMC) is reached. At that concentration, micelles form. Further addition of detergent increases the concentration of micelles without appreciably affecting the concentration of monomers. Redrawn with permission from Helenius, A., and K. Simons, Biochim Biophys Acta. 1975, 415:38.

detergents exhibit even more fluctuations in shape as they can deform, split, and fuse over time. Micelle formation is a direct consequence of the degree of amphiphilicity of surfactants. The surfactant molecules that form micelles are more water soluble than most lipids but still contain nonpolar groups with a propensity to form hydrophobic domains. They also tend to have conical shapes with bulky headgroups relative to their nonpolar groups (see Figure 2.17). In addition to detergents, Iysophospholipids (phosphol ipids lacking one acyl chain) form micelles, as do PLs with very short acyl chains (e.g., PC with four to nine carbon chains) under certain conditions. Self-association of detergents into micelles is strongly cooperative and occurs at a defined concentration called the critical micellar concentration, or CMC (Table 3.1). Below the CMC, the amphipath dissolves as monomers; as its concentration increases beyond the CMC, ideally the monomer concentration is unchanged while the concentration of micelles increases (Figure 3.4). The CMC can be detected by measuring surface tension or other aqueous properties, such as conductivity or turbidity (Figure 3.5). Micelle formation is dynamic, allowing constant interchange between constituents of aggregates and soluble monomers. For ionic surfactants, it is strongly affected by ionic strength (see Table 3.2). Micelle formation is also a function of temperature. The critical micellar temperature (CMT) is defined as the temperature above which micelles form (Figure 3.6). The Krafft point, also called the cloud point, is the temperature at which a turbid solution of surfactant becomes clear due to the formation of micelles.

Concentration

3.5. Variation in surface tension (y), specific conductivity (K), and turbidity (T) as a function of detergent concentration. The schematic plots show the dependence on concentration of detergent (surfactant) in solution of properties commonly used to find the CMC. (Note that conductivity only applies to ionic surfactants.) At the CMC, denoted by the dashed line, there is a break in the line for each property. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 65. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

Thus the Krafft point falls at the intersection of the lines for the CMT and the CMC, and at the Krafft point the temperature dependence of solubili ty rises steeply as the result of micelle formation. At the Krafft point, insoluble crystalline detergent is in equilibrium ,\lith monomers and micelles, so if the temperature is lowered, the detergent crystallizes out of solution. A familiar illustration is the precipitation of SDS in aqueous solutions below 4°C (its Krafft point). The CMT CMT ~

E

d

Detergent crystals

Detergent miceJles

.g 0:1

l::::

c:Q)

u

c: ou

L-----rCMIC Detergent monomers Temperature,OC

3.6. Detergent phase diagram. At temperatures below the Krafft point, the detergent exists as monomers at very low concentrations and insoluble crystals at higher concentrations. Raising the temperature increases the monomer concentration until the critical micellar temperature (CMT) is reached, when micelles form. At (and above) that temperature, the solution clears at temperatures because the only two phases present are micelles and monomers. The Krafft point falls at the intersection of the lines for the CMT and the CMC, where the temperature dependence of solubility rises steeply due to micelle formation, Redrawn from Helenius, A., and K. Simons, Biochim Biophys Acta. 1975,415:37. © 1975 by Elsevier. Reprinted with permission from Elsevier.

Membrane

strongly dependent on the ionic strength of the aqueous medium (see Table 3.2), as well as the kind of counterions available to shield the charged headgroups. Membrane Solubilization

+Detergent

Membrane "vith bound detergent

+More detergent

Mixed micelles: Detergent-Ii pid-protei n complexes

+More detergent

+

Mixed micelles: Detergent-protein complexes and detergent-lipid complexes

3.7. The stages in membrane solubilization. This schematic illustration follows the addition of increasing amounts of detergent to a membrane. Initially, integral membrane proteins are embedded in the lipid bilayer. At low concentrations of detergent, some detergent molecules penetrate the bilayer but do not disrupt it. As more detergent is added, disruption of the bilayer results in mixed micelles containing detergent, lipid and protein. At even higher detergent concentrations, most of the lipid is removed from the protein, prodUCing detergent-protein complexes, along with detergent-lipid complexes.

for nonionic surfactants and the common bile salts is below G°c. The size of detergent micelles is usually described by the aggregation number (N), the average number of surfactant moJecu les per micelle, although for some situations the molecular weight or hydrodynamic radius is reported (Table 3.J). The aggregation numbers given in the literature are averages, and the size distribution may be quite large. Micelle size can be determined by light scattering, ultracentrifugation, viscometry, and gel filtration. It varies widely, reflecting the size of the nonpolar domain: N increases with increasing tail length for a series of surfactants in which only the hydrocarbon chain length is varied. For ionic surfactants, N is

Detergents are used to extract membrane lipids and proteins into an aqueous suspension. When a low concentration of detergent is added to a membrane. the detergent molecules intercalate into the bilayer. When a higher concentration is added, the detergent disrupts the bilayer and forms mixed micelles containing lipid, protein, and detergent (Figure 3.7). Mixed micelles vary considerably in structure and size. The detergent concentration must be kept above its CMC to maintain the mixed micelles. Sometimes adding still higher concentrations displaces the lipid completely, producing detergent-protein complexes Free of lipid. Thus both the detergent concentration and the detergentto-protein ratio are important variables that influence how a particular membrane protein will be extracted from the membrane. The behavior of the membrane protein in further purification and characterization steps will depend on detergent-protein and detergentdetergent interactions, along with detergent-lipid and lipid-protein interactions if lipid remains. The amount of a particular detergent that solubilizes the membrane is roughly propoltional to its CMC. Bile-type detergents solubilize segments or

PC

Logiludinal

vie\v

Cmss-seclional

vie\."

3.8. Schematic illustration of the structure of mixed micelles of bile salts and phospholipid sandwiches of bile salt detergent with lipids. Redrawn from Jones, M. N, and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 97. © 1980 by American Chemical Society. Reprinted with permission from American Chemical Society.

Detergents

49 vQ) V

Q)

N

5

13

(a)

-

1i 4 ::I 1i "0 ::I VJ v 3 "0 VJ N

C

9

0-

"0 'v .£ 0 0- 2 .... p...

~

'"

5

VJ

0

.£

p...

~

a

0.0

0.2

0.4

0.6

0.8

1.0 a 2 4

Triton X-lOa concentration

6 8 10 12 14 16

Detergent concentration

3.9. Ratio of protein to phospholipid solubilized from epithelial cells by four different detergents. A spike at the low detergent concentrations indicates protein leaking out before the lipid is solubilized. 0, Triton X-1 00; _, sodium dodecylsulfate; 6, dodecyltrimethylammonium bromide; .... , sodium cholate. Redrawn from Jones, M. N., and D. Chapman, Micelles, Monofayers and Biomembranes, Wiley-Liss, 1995, p. 148. © 1991 by Elsevier. Reprinted with permission from Elsevier.

the membrane as detergent/bilayer sandwiches (Figure 3.8). The success of an extraction procedure is determined by checking the amount and composition of the desired component (usually protein) in the supernatant following sedimentation of the membrane. The

ratio of phospholipid to protein solubilized can indicate whether proteins leak from the membrane before it is completely disrupted, revealing whether a detergent concentration is suf-ficient to disrupt the membrane or only to solubilize segments of it (Figure 3.9).

3.10. Belts of detergents around purified membrane proteins. The positions of detergent molecules in solutions of detergent-solubilized proteins are revealed in neutron diffraction density maps obtained at different H20/D20 ratios to provide contrast variation. This image obtained with OmpF porin in ClODAO also shows C<x traces for protein obtained from the x-ray structure (protein is pink and detergent is green). From Pebay-Peyroula, E., et al., Structure. 1995,3:1053. © 1995 by Elsevier. Reprinted with permission from Elsevier.

Fixed barrier

Moveable balTier

I

'Y

~_!--~.-l'\'.l.~.l.l ...

.. .l.,L./..

'Yo

I

Aqueous phase

I~~~~

,

Trough

!

3.11. Formation of a monolayer in the Langmuir trough. The moveable barrier allows the area covered by the monolayer to vary. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995 p. 26. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

For reconstitution experiments, it is often desirable to replace the solubilizing detergent with lipids to better imitate biological conditions. Methods for detergent removal include dialysis, gel filtration, adsorption to polystyrene beads, and pH changes. Detergents that have a low CMC, like Triton X-I 00 (CMC = 0.24 mM), are much more difficult to remove by dialysis than detergents like OG (CMC = 25 mM) because so little of the detergent is present as monomers. Gel filtration is most effective when there is a large difference in the sizes of the detergent micelle and the detergent-protein mixed micelle, and thus works best with detergents with small values of N (and high CMCs). Adsol-ption to polystyrene beads (e.g., SM 2 Bio-Beads) is effective for most detergents, including Triton X-I 00, OG, DDM, cholate, CHAPS, and C I2 E g . Of course it is a problem if the protein of interest also adsorbs to the beads, in which case the beads can be placed outside a dialysis chamber. Some ionic detergents, such as cholate and deoxycholate, precipitate at about one pH unit above their pKas (5.2 and 6.2, respectively), which greatly simplifies their removal provided the mildly acidic conditions do not harm the protein being studied. SDS can be precipitated after exchange of sodium for potassium, as potassium dodecylsulfate is insoluble at room temperature. For some hardy proteins (such as bacterial porins), complete removal of detergent is effected by precipitating the protein with organic solvents. Although detergents have been widely used in pUlifying proteins for crystallization studies, their disorder prevents their resolution in the resulting highresolution structures obtained by x-ray diffraction. To visualize the detergent in protein-detergent complexes, low-resolution images of the structure of detergent domains in crystals can be obtained by neutron diffraction with H 2 0ID 2 0 contrast variation (see Box 8.1 on neutron diffraction). In this procedure, several crystals are prepared that vary in their H 2 0ID 2 0 content, giving relative contrasts to the protein and detergent with respect to the solvent to allow visualization of individual components. Discrete belts of detergent around the nonpolar portion of the protein are clearly detected by neutron diffraction studies of membrane proteins

such as the l3-barrel OmpF porin (Figure 3.10). In the case of OmpF protein, the "hardness" of the detergent torus affected the observed shape: with softer detergents, such as OG, the belts of detergents fuse with those of their nearest neighbors. Clearly the size and shape of the detergent molecules - along with smaller additives, such as heptane - are crucial to the success of the crystallization process (see Chapter 8), which has led to much interest in new detergents. Lipid Removal

Although it is rare for detergent extraction to completely remove bound lipid from membrane proteins, lipid removal may lead to loss of biological activity. There are dozens of examples of proteins (including succinate dehydrogenase and other components of the electron transport chain, several ATPases, numerous transferases, and receptors) that are inactivated when stripped of lipid by detergent or organic solvent and are reactivated by addition of lipid (see Table 4.2 for more examples). The lipid requirement may be quite specific, such as the absolute requirement of l3-hydroxybutyrate dehydrogenase extracted from mitochondrial inner membrane for Pc. Even for the cases of a general lipid requirement, it is clear that a portion of the lipids in a biomembrane associates dynamically with membrane proteins. This boundary lipid, which differs in mobility from the bulk lipid of the bilayer, may be functionally important and can be studied in model membranes.

MODEL MEMBRANES

The functions of the membrane and of many of its components are lost upon its disruption, necessitating reconstitution of the membrane in an in vitro model system for studies to elucidate mechanisms of transport and energy transduction, to measure enzyme kinetics and ion flows, and to explore phase changes and microdomains. The availability of a wide variety of model membrane systems is fortunate as no single system is suitable for all the membrane components being characterized or for all the techniques used to study them. Hundreds of papers describe the applications of each classical system, while the promise of the newest membrane mimetics has yet to be realized. Monolayers

Amphipathic lipid molecules with sufficiently large hydrophobic portions will line up at an air-water interface with their hydrophobic tails in the air. Such monolayers are commonly formed in a Langmuir trough, a container with a movable barrier on one side that allows control of the area and measurement of the pressure of the monolayer (Figure 3.11). The surface pressure (n)

Model Membranes

51

expanded

gaseous

____ solid ZOO

400

A2/molecule

I

E

z

E l=

expanded

20

30

40

50

70

60

80

AZ/molecule 3.12. A surface pressure-versus-area isotherm for a monolayer of a fatty acid at the air-water interface. A diagram of the surface pressure (TI) versus the area per molecule shows the relationship between TI and the area of the molecules forming the monolayer. The isotherm reveals phase changes: the monolayer approaches a solid state at high pressure, changes to liquid states at lower pressures, and changes to gaseous states at very low pressures, as shown in the inset. Redrawn from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 27. © 1990. Reprinted with permission from John Wiley & Sons, Inc.

A.

is created by the dilference between the surface tension of the monolayer (y) and that of the air-water interface (Yo): 7t = 1'0 - y. The high surface tension of water means that it takes work to cover the area of the air-water interface. To decrease that work, the monolayer spreads over the surface, putting pressure on the movable barrier. Inward movement of the barrier to decrease the area increases the surface pressure of the monolayer. In forming monolayers, the composition is controlled and the amount of lipid is known. A surface pressure-versus-area isotherm, showing the relationship between 7t and the area of the molecules forming the monolayer, reflects phase changes (Figure 3.12). Highly compressed molecules are so condensed they approach a solid state, whereas at very low pressures, the molecules are so spread out they do not interact and are considered to be in a two-dimensional gaseous state. Between these extremes, the monolayer is in a Iluid phase described as liquid. The effect of chain length on the phase of the monolayer is revealed in the pressurearea isotherms for a series of monolayers composed of PC with varying acyl chains (Figure 3.13A). Two fluid phases, called L E (liquid expanded) and L c (liquid condensed), are evident in the curve for DPPC, but only B.

50

50 r x - - - - - , - - - - - - , . - - - - - , - - - - - ,

x

~

x

40

40

E 30

E 30 E '--'

Z

Z

E

(j)

....

(j)

'-< :::l

:::l if) if)

(j)

20

if) if)

(j)

'-<

20

'-<

0...

0...

10

60

80

100

120

Area (AZ/molecule) 3.13. Phospholipid phase changes in monolayers. Surface pressure-versus-area isotherms reveal the effects of chain length and temperature on lipid monolayers. A. When monolayers are made of phosphatidylcholine with saturated acyl chains of varying lengths (0, dibehenoyl [C22); 0, distearoyl [C18]; x, dipalmitoyl [C16); 6., dimyristoyl [C14]: and '\7, dicapryloyl [Cl0)) phospholipid behavior in the monolayers correlates with their melting temperatures. The PLS with longer chains and higher melting temperatures form liquid condensed (Lc) monolayers at 22°C. while the PLs with shorter chains and lower melting temperatures form liquid extended (LE) monolayers at 22°C B. For a particular PL, increased temperature changes the monolayer state from Lc to LE, shown in surface pressure-versusarea isotherms for DPPC in 0.1 M NaCI at varying temperatures. (e, 34.6°C; 6., 29.5°C; _, 26.0°C; x, 21.1 cC; 0, 16.8°C; &, 12.4°C; 0, 6.2°C) Redrawn from Phillips, M. C, and D. Chapman, Biochim Biophys Acta. 1968, 163:301. © 1990. Reprinted with permission from John Wiley & Sons, Inc.

120

3.14. Alveoli with and without pulmonary surfactant. The presence of natural surfactants enables the alveoli to withstand changes in pressure (P) in the lungs. From Johns Hopkins School of Medicine Interactive Respi ratory Physiology, http://oac.med.jh m i.edu/res_phys/Encyclopedia/Surfactant/ Surfactant.HTML. © 1995 by Daphne Orlando. Reprinted by permission of Daphne Orlando.

when the temperature is held at values bet\veen 15"C and 30°C (Figure 3.13B) indicating the LEIL e transition is a function of the state of the acyl chains. Monolayers provide a means to study the effects on lipids of factors such as pH, ionic strength, and addition of multivalent versus monovalent ions. Monolayel's have also been used to examine the intercalation of membrane-active peptides, such as gramicidins (see "Proteins and Pep tides That Insert into the Membrane" in Chapter 4) and signal peptides. In spite of their obvious limitations as membrane mimics, monoJayers can give useful physiological information, such as an understand i ng of respira tory distress syndrome j n newborns. Pulmonary surfactant is a complex of proteins and phospholipids that coats the surface of alveoli and small bronchioles to reduce surface tension and confer compressibility to lungs (Figure 3.14). The lipid is predominantly DPPC with a significant fraction of PC. Monolayer studies indicated that compression squeezes out the PC and the remaining PC-rich surfactant in the L c state is less resilient in responding to pressure, Thus the lack of PC in the surfactant of premature babies can trigger alveoli collapse. resulting in respiratory failure.

Example (Briggs, M. S., and L. M. Gierasch, Exploring the conformational roles of signal sequences: synthesis and conformational analysis oflambda receptor protein wildtype and mutant signal peptides. Biochemistry. 1984, 23:31 I 1-31 14; Briggs, M. 50, et al., Confo17nations ofsignal peptides induced by lipids suggest initial steps in protein export. Science. 1986, 233:206-208) Monolayers were used to dete17nine the conformation of signal peptides as they interacted with membrane

Lipids. Signal pep/ides are the N-terminal extensions of newly synthesized proteins targeled (or secrelion from the cytoplasm (see Chapter 7). Their intel71al sequence of aboul 10 to 15 nonpolar amino acids suggested they could insert into the membrane. Synthelic signal peptides were found to lay on the sUijace of a monolayer 9

I

I

I

I

6

bil v

3

-0

5

0

-3

-6 1 190

1

210

1

230 A. (nm)

1

1

250

3.15. Circular dichroism spectra for synthetic signal peptide interacting with a monolayer of POPE:POPG in a 2:1 ratio at high pressure (dashed line) or low pressure (solid line). e, molar ellipticity; A, wavelength. Redrawn from Briggs, M. S., et al., Science. 1986, 233:206-208. © 1986. Reprinted with permission from AAAS.

Model Membranes

53

I-

r-/"//"7 _ ' / .,-/,--- -

I I I I I I I I I I I I I I

1

I I I I I I I I I I I I I I

3.16. Schematic diagram of a black film separating two aqueous compartments with electrodes Inset shows the planar bilayer region of the film. Redrawn from Jain, M. K., Introduction to Biological Membranes, Wiley, 1988, p. 95. © 1998. Reprinted with permission from John Wiley & Sons, Inc.

with high surface pressure and to insert into th.e monolayer when the pressure was lowered. Circular dichroism revealed the inserted peptides have an a-helical structure (Figure 3. /5). This early evidence for inseriiol1 of a-helices into the membrane was given functional significance by the much 10'wer tendency of mutant signal peptides to form a-helices in monolayers. Planar Bilayers

While a monolayer reveals pressure effects of membrane constituents, a planar bilayer separating two aqueous compartments is used to study electrical properties because it offers access for electrodes on both sides of the membrane. Pure lipid bilayers are not permeable to ions (which is why the myelin membrane is a good insulator), so the introduction of molecules that form ion channels can be closely monitored. These systems mimic the electrophysiological aspects of the cell membrane with its electrochemical potential and numerous ion channels, and they have been the technique of choice for biophysical studies of ion channels (see Box 3.2). Historically, planar bilayers were called "black films" because they appear black when made on a Teflon sheet. The lipid is dissolved in organic solvent (such as hexane, decane, or hexadecane) and is painted on a tiny orifice (about 1 mm in diameter) in the Teflon sheet, which is then inserted between two aqueous compartments. As the solvent dissipates and the lipid gradually drains or "thins," a region of single bilayer stretches across part of the orifice and can last for a few hours (Figure 3.16). Peptides, small proteins, and other lipids will diffuse into the bilayer when added to one of the aqueous compartments, although the incorporation of larger proteins may be problematic. By insert-

ing electrodes in the two compartments. a voltage may be applied across the bilayer and current may be measured. This system permits precise measurements of ion flows, such as those observed \vhen OmpF porin inserts into the black film, and even detects the closing of single channels (Figure 3.17). However, the electrical capacity of a black film is considerably lower than that of cell membranes, implying there are structural differences. which have been attributed to the presence of excess solvent. Montal and Mueller introduced a significant new method for the formation of a planar lipid bilayer by apposing two monolayers derived from air-water

BOX 3.2. Electrophysiology Current (I) measures the flow of charge from one place to another in the units amperes (amps). The flow of charge responds to a potential difference (V, in volts), requires a conducting path, and obeys Ohm's law: V = IR, where R is the resistance of the path with the units ohms. The conductance is the reciprocal of the resistance, giving the ease of the flow of current, and is measured in siemens. An electrochemical gradient exists across any membrane that separates two compartments containing different amounts and kinds of ions. The potential difference between the compartments drives the movement of ions, provided there is a pathway such as a pore or ion channel. Many excitable cells (i.e., nerve, sensory, and muscle cells) have a potential across the plasma membrane in the resting state of around -60 mV, which creates an enormous electric field across the membrane (calculated to be 200,000 V cm- 1 for a membrane that is 3 nm thick). In E. coli, the electric potential measured across the cytoplasmic membrane is around -95 mV. Many in vitro measurements utilize application of a "voltage clamp" in which the experimenter applies a voltage to control the potential across the membrane and then measures the current.

A.

1500 ~


p,

';:; 1000 >::


.... .... ;:l u

500

4

8

12

16

Time (sec)

B.

100 pA

L I_ _

5s

-03 -02 -01

!

·-c

-:.,.

c.

AL ~ J.....~...,IItII_.~••t1.'.' ,••"......,. ~ ••~ - 03 20 P

2

~ ~

26~

20PAL

------

-02

I

40~PS

3.17. Conductance data obtained with pure OmpF porin added to the cis side of black films. OmpF is a channel-forming protein described in detail in Chapter 5. A. When OmpF inserts into a planar lipid bilayer at a membrane potential of +50 mV, a stepwise current increase results from sequential insertion events of open porin trimers (pA, picoamperes). B. After insertion of a single trimer is complete, channel closures occur in response to a membrane potential of + 140 mV. These closures provide evidence of the voltage gating of OmpF channels and typically occur in three steps believed to correspond to the three channels in OmpF trimers. C. Small fluctuations occur at -70 mV, indicating fast flickering events involving subconductance states (monomeric conductance of 380 picosiemens IpSI marked by tick marks). From Basle, A., et aI., Biochim Biophys Acta. 2004, 1664:100-107. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Model Membranes

ss

A.

I Direction of Teflon film -

__~

B.

, displacement Septum

!

Direction of displacement Teflon film Air

Aperture ___

Water Trough

3.18. Technique introduced by Montal and Mueller for making a planar lipid bilayer by apposing two monolayers. A. The apparatus has two compartments containing solutions into which electrodes can be inserted. The compartments are separated by a movable partition called a septum, which has a Teflon film with a tiny aperture. After monolayers are formed at the air/water interfaces of the two compartments, the septum is moved down, allowing the two monolayers to come together to form a bilayer across the aperture. Redrawn from Montal, M., and P. Mueller, Proc Natl Acad Sci USA. 1972, 69:3561-3566. B. A dose-up view of the bilayer formed across the aperture in the Montal-Mueller apparatus, with polar headgroups from one monolayer shaded to indicate the asymmetry achieved in the bilayer. Redrawn with permission from Jain, M. K., Introduction to Biological Membranes, Wiley, 1988, p. 95. © 1998. Reprinted with permission from John Wiley & Sons, Inc.

interfaces (Figure 3.18). The organic solvent introduced with the lipid is evaporated from each monolayer prior to forming the bilayer, and the electrical capacity of the bilayer thus formed matches that of biomembranes, supporting the claim that the bilayer is solvent-free. One clear advantage of the Montal-Mueller system is that it allows independent manipulation of the lipid contents of the two leallets. Schindler introduced a further modification by spreading the monolayers from preformed lipid vesicles, making it easier to incorporate proteins and to direct their introduction from one leaflet specifically. Recent advances in the design of the compartment to allow both optical and electrical measurements have enabled the determination of electrical properties to be correlated with other physical characteristics of the bilayer, such as fluidity.

Example (Armah, C. N., et aI., The membrane-penneabilizing effect ofavenacin A-I involves the reorganiz.ation of bilayer cholesterol. Biophys J. /999, 76:281-290) To investigate hemolytic pore {onnation by saponins such as digitonin and Ihe avenacins (a group of fi.mgicidal sleroid glycoside planl natural products), MontalMueller planar lipid bi/ayers were fonned in an oplical chamber thaI allowed simultaneous measurements of both conductance and fluorescence. The data sho'w that an increase in conductivily of bilayers coincides will? a decrease in the lateral mobilily ofNBD-cholesterol

determined by FRAP (Figure 3.19). Monolayer studies showed thaI il1sertion of the saponin into the bilayer does not require cholesterol (not shown), pore forma.tion occurs only in the presel1ce of cholesterol. The results indicate that saponin-cholesterol complexes {onn pores in the bilayer, consistent with their biological aClivity.

Patch Clamps Although technological improvements have enhanced their sensitivity, traditional planar bilayers have exhibited a high level of noise, making it difficult to detect currents flowing through single channels whose amplitude is very small (typicaJly I or 2 picoamperes [pA]). A crucial advance that aJJowed detection of single ion channels in cell membranes was recognized by the award of the Nobel Prize in Physiology or Medicine to Erwin Neher and Bert Sakmann in 1991. Their patch clamp technique accesses a tiny portion of a membrane in a way that aJJows electrical measurements to be made and has now been used with a variety of membrane systems. To clamp a patch of membrane bilayer, a portion is sucked into a polished tip of a glass pipette, sealing off an area of 1 to 6 ~m2. The pipette can remain attached or, if the seal is sufficiently stable, the pipette may be withdrawn to excise the patch of membrane

A.

B.

12

N

I: ll)

2 10

u

u

:;:c ::i. '--' -'

.:.c ........ ~ Vl 60

8

<.J';;-

o <.JE 50

I:

ll)

~ <.J u

.:.c

I:

6

'iii'i' 40 ..20 ~ X 30 ",--,

4

'u ll)

0, if)

90 80 70

-'

~

"@

2

'-<

20

ell

10

~

-1

0 -20

0

20

0 -20

40

Time (minutes)

0

20

40

Time (minutes)

3.19. Effect of saponin treatment on conductivity and on lateral diffusion in Montal-Mueller bilayers consisting of POPC:DOPE:cholesterol (7:3: 10) containing 1 mol % NBD-cholesterol. Avenacin A-1 (1.0 11M) was added at time zero. A. Conductivity increases after ~20 minutes, which is not observed without addition of a saponin (data not shown). B. The decrease of the rate of lateral diffusion of NBDcholesterol observed by FRAP corresponds with the increase of conductivity. Redrawn from Armah, C. N., et aI., Biophys J. 1999, 76:281-290. © 1999 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

A.

. . .-.1. B.

Preamplifier

Inside-out

.:. . P.: .ul:. :.-I_~. ~ Suction ~ Ou tside-ou l Pull

~

3.20. The patch clamp method. A. Excision of a portion of bilayer by a patch clamp. A clean firepolished pipette is pressed against the cell membrane to form a gigaohm seal. As it is withdrawn, the membrane reseals. Conductance can be measured across the patch on the cell surface or across the excised patch. B. Methods for patch recording with cell-attached patch or excised patches in the insideout and outside-out configurations. The cell membrane is represented by a solid line with a dotted line, and the dotted line indicates the inner surface of the membrane in the two patch configurations. Redrawn with permission from Jain, M. K., Introduction to Biological Membranes, Wiley, 1988. © 1998. Reprinted with permission from John Wiley & Sons, Inc.

Model Membranes

51

lj~: '~.lJ1fT'I'~: 20 ms

ACh

end-plate channel

=iJ~~~j~ Closed

3.21. Early single-channel recording by Neher and Sakmann showing the current through rat muscle activated by acetylcholine (ACh). The diagram under the tracing illustrates how the channel opens reversibly in response to ACh binding. Redrawn from the Nobel lecture by Bert Sakmann, December 9, 1992. © 1991 by The Nobel Foundation. Reprinted by permission of the Nobel Foundation.

Open

(Figure 3.20). With a gigaohm seal (having an electrical resistance higher than 10 9 ohms), the patch clamp allows detection of current down to the level of picoamperes 00- 12 amperes). Because of its sensitivity and small size, the conductance behavior of a small number of channels may be monitored through the patch, revealing single-channel opening and closing (Figure 32\).

Example (Nehel; E., and B. Sakmann, Single-channel currents recorded fl'om membrane of denervated frog /"nuscle fibres. Nature. /976, 260:799-802, and Nobel lectures) The first patch clamp studies were peljonned on denervated frog and rat muscle and measured ion flow through individual acetylcholine receptors (AChRs) ofthe neuromuscularjunctio11. The recorded currents indicated that the acetylcholine-activated channels exist in only two COl1ductance states, open and closed, and showed that bursts ofcurrent through open channels result when acetylcholine binds. The obselvation oftwo classes ofcurrents, which differ in amplitude and duration, enabled detection of two isofonns of the receptor, now knOl·\m to be fetal and adult AChRs (Figure 3.22). Supported Bilayers

Planar lipid bilayers that sit on glass, quartz (mica), or gold supports allow direct observation of their surFace using atomic force microscopy (AFM), immunolabeling, fluorescence, and other spectroscopic techniques. They can be made by fusion of lipid vesicles of the desired composition on the surFace of the support in an aqueous environment or by sequential deposition of monolayers, which allows asymmetric bilayers to be rormed. Incorporation of specific celJ surface receptors enables the supported bilayers to be used as biosensors

for cell populations or ligands. However, artiFacts due to the presence of the solid support arise when incorporating components with soluble portions that can adhere to the support. Also, the distance between the support surFace and the supported lipid bilayer is typically about 2 nm (and water filled). Thus membranespanning proteins incorporated in supported bilayers can contact the support, losing their lateral mobility and possibly affecting their runctional properties as well. A variety of new strategies address this problem by employing polymerized lipids or tethered polymers to alter the distance to the support. A cushion provided by polyethylene glycol linked to a lipid on one end and a reactive silane to attach to the glass support on the other end increased the distance to around 4 nm (Figure 3.23), which allowed successful reconstitution of green fluorescent protein-labeled membrane proteins.

Example (Crane, 1. M., and L. K. Tamm, Role ofcholesterol in the formation and nature of lipid rafts in planar and spherical model membranes. Biophys J. 2004, 86:2965-2979) Addition of cholesterol to binary lipid mixtures (PC + sphingomyelin) stimulates formation of an Lo phase, which has been of great interest as the basis of domain {onnation in lipid rafts (see Chapter 2). The use of val'ious lipid-linked dyes that show differen t preferences for Lo and LJ phases (and are excluded from gel phase) allovved obseniation of domain formation in response to cholesterol in planar lipid bilayers supported on quartz slides. /n the absence of cholesterol, the PC/sphingomyelin bilayers exhibit coexistence ofgel and fluid phases; with addition 0[' cholesterol, the gel phase disappears and the rounded domains of La phase appear (Figure 3.24). The extent of La phase correlated with the mobility of the (lyes revealed by FRAP measurements.

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3.22. Characterization of two forms of the acetylcholine receptor. A. Patch clamps observed in postnatal rat muscle fiber treated with 0.5 pM ACh revealed two classes of currents through the acetylcholine receptor (AChR; marked with' and #) that differ in amplitude and duration. B. The two types of AChR responsible for different classes of currents have been shown to be fetal and adult isoforms, which differ in subunit composition as shown. Redrawn from the Nobel lecture by Bert Sakmann, December 9, 1992. © 1991 by The Nobel Foundation. Reprinted by permission of the Nobel Foundation.

closed

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Model Membranes

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3.23. Diagram of a tethered polymer-supported planar bilayer, representing a 3400-Da polyethylene glycol covalently attached to a silicon oxide surface and at the other end to a phospholipid, which doubles the distance from the bilayer to the support. From Kiessling, v., and L. K. Tamm, Biophys J. 2003, 84:408-418. © 2003 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

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3.24. Fluorescence micrographs of PC/sphingomyelin (SMl supported planar bilayers stained with rhodamine-DOPE at different ratios of PC/SM and different concentrations of cholesterol. The top row has different ratios of PC/SM, as labeled, in the absence of cholesterol. The middle row has PC/SM of 1:1 with different cholesterol concentrations as labeled, and the bottom row is the same, with PClSM of 2:1. The irregularly shaped domains in the absence of cholesterol correspond to regions of gel phase (top row: C, D) As the cholesterol concentration increases, the lighter regions corresponding to Lo domains increase. At 20% all three domains (gel, fluid, and Lol coexist. The white bar represents 10 ~lm. From Crane, J. M., and L. K. Tamm, Biophys J. 2004, 86:2965-2979. © 2004 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

Multilamellar vesicle (MLV)

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In contrast to planar lipid bilayers, liposomes are closed bilayer' vesicles that do not allow access to the inner compartment, although they can be preloaded with diverse compounds From the dispersion medium in which they are Formed. Liposomes ['esult when bilayerForming lipids are mechanically dispersed in aqueous suspensions due to the tendency [or bilayer edges to seal so the acyl chains are not exposed to water. Depending on the method used, they may be uniJamelJar (encapsulated by a single bilayer) or mulLilamellar. They are also classified by size, and the large variation in size effects large differences in curvature (Figure 3.25). The lipids that form vesicles are roughly cylindrical in shape and generally have more than 11 carbons in their acyl chains. Liposomes are used to study effects of particular lipids, such as cholesterol, in mixtures of lipid components. In addition, liposomes are used in many studies of membrane protein function, folding, and assembly. When proteins are reconstituted into them, the vesicles are called proleoliposomes. Proteoliposomes are judged by numerous criteria, such as homogeneity regarding size and number of lamellae when visualized \-vith thin section EM. Proteins should be distributed fairly evenly and oriented nonrandom]y to mimic their incorporation in biomembranes. Like other Jiposomes, proteoliposomes should have low permeability to ions, giving them the ability to maintain a charge gradient. Such characteristics may depend on variation of the lipid-toprotein ratio. Multilamellar Vesicles

50->10,000 nm.

Aqueous compartment 3.25. The structures and dimensions of three types of liposomes. Multilamellar liposomes (MLVs) have many more layers than indicated. Comparison of small unilamellar liposomes (SUVs) and large unilamellar liposomes (LUVs) reveals the difference in curvature that results in more loosely packed acyl chains in SUVs. Redrawn with permission from Jones, M. N., and D. Chapman, Micelles, Mono/ayers and Biomembranes, Wiley-Liss, 1995, p. 119. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

Multilamellar vesicles (MLVs) contain concentric spheres of lamellae and may be made by simply shaking a thoroughly dried lipid film into an aqueous solution. They are usually polydisperse, with diameters from 0.2 to 50 IJ.m, and have as many as 20 concentric layers of membranes. Their internal volume is unknown but quite small. To increase the internal volume, they can be converted to vesicles with one to four lamellae by extrusion through polycarbonate filters. They have been used in studies of lipid phase transitions by DSC and in many studies of enzyme and peptide binding. Their osmotic sensitivity allows quantitative measurements of solute uptake rates from turbidity changes.

Example (Luckey, M., and H Nikaido, Specificity of diffusion channels produced by lambda phage receptor protein o(Escherichia coli. Proc Nat] Acad Sci USA. 1980, 77:167-171) The liposome swelling assay {ollows absorbance changes as the MLVs respond to osmotic pressure; when the liposomes are suspended in hypotonic solutions,

Model Membranes

swelling results li'omwater entry that pushes the concentric bilayers further apart, causing a decrease in light scattering. In isotonic solution, swelling does not occur without solute uptake, so MLVs can be used to measure the rate of uptake, which is more sensitive than methods that measure the extent of uptake after reaching equilibrium. This liposome swelling assay was crucial to discovering the specificity of maltoporin (LamB protein) because it could distinguish among uptake rate::; of disaccharides (Figure 3.26).

61

0.7

Small Unilamellar Vesicles

Example (Beschiaschvili, G., and 1. Seelig, Peptide binding to lipid bilayers. Nonclassical hydrophobic effect and membrane-induced pK shifis. Biochemistry. 1992, 31: 10044-1 0053) This study elnployed SUVs made using PO PC with and witlwllt POPG, which had diameters of around 30 nm, to compare the binding of a small al11phiphilic peptide to that in larger vesicles. Since the acyl chains are less tightly packed in the small vesicles than in the larger vesicles, they have less lateral tension, which is related to membrane elasticity. High-::;ensitivity titrati011 calorimetry was used to measure the heats of reaction {or binding the cyclic peptide, an analog of somatostatin called SMS that is an amphiphilic peptide with a positive charge. l\!lonolayer studies had already indicated that SMS intercalated into lipid with little change in its conformation according to circular dichroism, and 2H NMR studies had shown it could diffuse rapidly on the surface when bound. The binding enthalpy for SMS was -7.3 kca 1111101 for SUVs, independmt of pH or lipid composition, in contrast to the enthalpy of binding to larger vesicles of -1.4 kcallmol. The ditfermce in t::.G for binding to the two classes of vesicles was less than 1 kcal/mol, indicating there is a large enthalpy-entropy compensation, which means el1tropy is not important in SUV binding but is the driving force for binding the larger vesicles. The entropy effect is explained in terms of the increased area of the bilayer that accompanies peptide binding, which lessens

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o Small unilamellar vesicles (SUVs) with diameters of 20 to 50 nm result from extensive sonication of MLVs. They are also made by extrusion through polycarbonate filters of defined pore size and can be further sized by gel filtration or gradient centrifugation. Another method to make SUVs is by injection of lipids in organic solvent into aqueous media, followed by removal of the organic solvent. SUVs are vel)' asymmetric due to their extreme curvature. For example, SUVs of PC are 22 nm in diameter and have 1900 and 1100 molecules in the outer and inner leaflet, respectively. Although the acyl chains are less tightly packed than in larger liposomes, the extreme curvature of SUVs makes it difficult to incorporate proteins.

+ LamB protein

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Time, min 3.26. The liposome swelling assay follows a decrease in optical density (OD) at 500 nm upon mixing MLVs with permeant solutes. The optical density tracings in the control panel show very little change observed in the absence of maltoporin (LamB protein). When MLVs have incorporated purified maltoporin (3 IJ.g/mg PL). the rate of uptake of maltose is significantly higher than that of lactose and sucrose. Redrawn with permission from Luckey, M .. and H. Nikaido. Proc Natl Acad Sci. USA. 1980. 77167-171.

the intemal bilayer tension of the more tightly packed acyl chains in larger vesicles. Large Unilamellar Vesicles

Large unilamellar vesicles (LUVs) with diameters hom 100 nm to 5 ~m can be made by freeze-thaw methods that induce fusion of SUVs. Since the introduction of commercial extruders that force the liposomes under nitrogen pressure through polycarbonate filters of defined pore size, more uniform LUV sizes have been obtained, especially with repeated extrusions. Other procedures make proteoliposomes in this size range by mixing the protein in detergent with an excess of lipid and then gently removing most of the detergent by dialysis or dilution. Dialysis is slow (often requiring days) and is successful for detergents with high CMCs and small aggregation numbers, such as OG, sodium cholate, and CHAPS. Dilution to well below the detel-gent CMC is rapid; micelles break up and proteins (along with detergent monomers) incorporate into the lipid vesicles, which are collected by centrifugation. The rate of detergent removal affects how well the proteins are distributed. Gel fiJtration is used to size the vesicles, as well as to separate proteoliposomes from excess detergent. LUVs have the advantage of large encapsulated volumes, up to 50 Llmol of lipid, but their disadvantages include heterogeneous size distributions and fragility of larger vesicles.

Example (Costello, M. 1., et aI., Morphology of proteoliposomes reconstituted with purified lac carrier protein li'om Escherichia coli. J Bioi Chem. 1984, 259: 1557915586,' Costello, M. 1., et al., Purified lac permease and cytochrome-o oxidase are functional as monomers. J Bioi Chem. 1987, 262:17072-17082)

3.27. A series of pictures showing the effect of Ca 2 + on the elastic compressibility of GUVs manipulated with micropipettes. The suction pressure is sufficient to hold the vesicles firmly on the pipette tips (A). When the suction pressure is reduced as they are brought together, the vesicles in 3 mM Ca 2 + (B) do not adhere to each other, while in 0.12 M sucrose (e), they do. From Akasyi, K., et aI., Biophys J. 1998, 74:2973. © 1998 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

Lactose permease (also called the LacY protein; see Chapter 10) was originally reconstituted by DC dilution followed by fi~eez.e-thaw/sonication to make proteoliposomes that were examined by fi'eeze-fracture EM. At a molar protein-to-lipid ratio o[ 1:2500, the majority of the proteoliposomes had diameters of 30 to 150 n111 and exhibited fairly even distributions of protein particles. Quantitation ofthe siz.e and distribution ofthe lactose permease with variation of protein/lipid ratios led to the conclusion that the protein was incorporated as a monDma To answer the question o[ the quaternary state of lactose permease during active transport, proteoliposomes were reconstituted with both lactose pennease and cytochrome o (a tenninal oxidase of the E. coli respiratoly chain) and energized by providing ubiquinol, generating an elecli'ical gradient ofaboU! -130 mV Alternatively, proteoliposomes containing the lactose pennease were suspended in bu[fers containing high levels ofK+ Q/1Cllater treated with valinomycin, which carries K+ across the 111embranes to dissipate the K+ gradient. Changes in energiz.ed states did not lead to dimerizat ion o[ the lactose permease, proving the reconstituted lactose permease was capable o[ both passive and active lactose transport as a I1101wmer. For many subsequent experimel1ts, LUVs were used to compare the activities o[ lactose pennease variants made when evelY residue of Lac Y protein was altered by mutation. Short-Chain/Long-Chain Unilamellar Vesicles Short-chain/long-chain uniJamellar vesicles (SLUVs) form spontaneously from aqueous suspensions of longchain phospholipid (saturated PC, PE, and sphingomyelin with acyl chain lengths of at least 14 carbon atoms) mixed with small amounts of short-chain lecithin (acyl chain lengths of six to eight carbons). They range in diameter from 10 nm to > 100 nm, depending on the ratio of short-chain to long-chain compo-

nents (increasing short-chain PL produces smaller vesicles). Inclusion of cholesterol can increase the size of the SLUV. While they have not been of general importance, SLUVs have been employed in functional studies of lipolytic enzymes because they are superior as substrates for the water-soluble phospholipases (phospholipase C and phospholipase A 2 ; see Chapter 4). Giant Unilamellar Vesicles Giant unilamellar vesicles (GUVs) are 5 to 300 !J.m in diameter. These giant liposomes are cell-size vesicles that are large enough to insert a microelectrode or to visualize surface sections by optical microscopy. They can be manipulated by micropipettes to test their elastic compressibility by their adherence to other vesicles (Figure 3.27). While they are generally viewed as excellent membrane mimetics, their large internal volume may be a disadvantage. Also, proteoliposomes of this size are very fragile. GUVs can be made by slowly hydrating lipid at low ionic strength and high lipid concentration, followed by sedimentation through sucrose to eliminate MLVs and amorphous material. Alternatively, a homogeneous population of < 100 !J.m diameter can be prepared by electroswelling, applying a voltage to the solution of lipids in 100 mM sucrose at 60 v C. Preparation of GUVs at high ionic strengths (comparable to physiological salt concentrations) requires 10% to 20% of a charged PL and millimolar concentrations of Mg 2+ or Ca 2+. GUVs may also be made from native membrane with addition of lipid: for E. coLi membrane, the optimal lipid concentration is 90 mg/ml. To better incorporate membrane proteins into GUVs, a fusion technique has been devised. First LUVs are prepared with the proteins and then coupled to fusion-inducing peptides. One such fusogenic peptide is a short ex-helix called WAE; since it is negatively charged, a positively charged target is

incorporated in the GUV to facilitate docking of the LUVs. Thousands of LUVs dock onto the surface of a GUV, and after a few minutes they fuse, as demonstrated by free diffusion of the lipids between them.

Example (Korlach, I, et aI., Characterization of lipid bilayer phases by confocal microscopy and fluorescence correlation spectroscopy. Proc atl Acad Sci US A.1999, 96:8461-8466) Two fluorescent probes were incorporated into CUVs of DLPClDPPClcholesterol. The probe Dil-C2o (1,1'dieicosanoyl-3,3,3',3'-tetramethylindolcarbocyanine perchlorate) partitions preferentially (3:1) in the La phase and the probe Bodipy-PC (2-(4,4-difluoro-5,7dimethyl-4-bora-3a, 4a-diaza-s-indacene-3-pentanoyl)-Ihexadeca11.0yl-sl1-glycero-3-phosphocholine) partitions preferentially (4:1) in the Ld phase. Their appearance in complementary regions of the images obtained by confocal microscopy facilitated phase assignments for a set of CUVs of varying compositions and gave conclusive evidence for the coexistence of separate lipid phases (see Chapter 2).

Mixed Micelles and Bicelles

Because micelles have little resemblance to bilayers, they are not generally considered to be good membrane mime tics. Even so, mixed micelles of phospholipid and detergent have been used in a multitude of studies of membrane proteins, especially when either detergent removal led to denaturation or detergent inclusion gave better activity. Micelles have been used to determine quaternary structure of membrane proteins by gel filtration and electrophoresis. Mixed Triton X- LOO micelles are especially useful for kinetic analysis of enzymes with phospholipid substrates, because variations in PL concentration (up to 15 mol %) have little effect on the micelle structure. Finally, due to their small size, micelles form isotropic solu tions that are advantageous for NMR studies of membrane-associated peptides and small proteins. An exciting advance for NMR studies that provided a better imitation of the membrane and avoided the severe curvature of micelles was the development of bilayered micelles, or bicelles. Bicelles are discoidal lipid aggregates composed of long-chain phospholipid and either detergent or short-chain phospholipid. The center of the disc is a lipid bilayer with its edges stabilized by the detergent or short-chain lipid (Figure 3.28). Bicelles made with detergent have a much lower detergent content than mixed micelles. Varying the longchain lipid can alter features of the bilayer, changing both headgroups (typically DMPC doped with DMPS or OMPG) and length of acyl chains (OM PC, OPPC, and OLPC) in ways that allow investigation of these features of membranes. The size of the bicelles is dependent on both the ratio of long-chain to short-chain PL (q) and

A.

4nm

20-40 nm 3.28. Schematic cross sections of bicelles. Bicelles contain a mixture of long-chain phospholipids, such as DMPC, and either shortchain PLs, such as DHPC (Al. or bile salt detergents, such as CHAPSO (B). The planar region is composed mainly of long-chain PLs, while the rim is formed by a monolayer of the short-chain PL (in A) or the bile salt detergent (in B). A polytopic membrane protein is shown incorporated in the bicelles in B. Redrawn from Sanders, C. R., and R. S. Prosser, Structure. 1998,6:1227-1234. © 1998 by Elsevier. Reprinted with permission from Elsevier.

the total concentration of PL (Cl). When q > 3 and CL is 15% to 20%, the bicelle diameter is 500 A and the bicelles orient in strong magnetic fields, allowing them to be used for solid-state NMR. When q < 1 and CL is 5% to 15%, the bicelle diameter is only 80 Aand these form isotropic suspensions suitable for high-resolution MR. Recently, bicelles have been used for crystallization of bacteriorhodopsin.

Example (Czerski, L., and C. R. Sa l1ders, Functionality ofa membrane protein in bicelles. Anal Biochem. 2000, 284:327-333) Kinetic analysis of the integral membrane protein diacylglycerol kinase (DCK, which functions to phosph01ylate the lipid diacylglycerol using Mg-ATP; see "Membrane Enzymes" in Chapter 6) shows its activity is optimal in mixed micelles containing decyl maltoside and cardiolipin and decreases in bicelles. The DCK activity in bicelles is dependent 011 lipid composition, demonstrating a preference for DMPC or DPPC with 3-([3-cholamidopropyl]dimethylammonio)2-hydroxy-l-propanesulfonale (CHAPSO). The kinetic data show a reduced V",ax rather than changes in KI1I , suggesting lillie perturbation at the subst rate-binding site. The enzyme activity exhibited by DCK in bicelles validates the use of this system for NMR studies. Blebs and Blisters

The goal of most reconstitution systems is to reproduce the membrane environment in a model system. A different approach is to use protrusions from the membrane, which are called blebs or blisters. Because they allow experimentalists to examine portions of membranes still attached to living cells, blebs are model membranes with a different set of advantages. Blebs are composed of the physiological mixture of lipids,

3.29. Bleb on Xenopus laevus oocyte surface viewed by confocal microscope. After using hypertonic stress to induce blebbing, the sample was treated with NBD-phallacidin, a fluorescent dye that stains actin. The bleb is readily visualized in the light image (A) and is absent in the fluorescence image (BJ, where the plasma membrane from which it derived is clearly stained. Similar results are obtained with a stain for tubulin, indicating the bleb lacks an associated cytoskeleton. Redravvn with permission from Xhang, Y, et aI., J Physio/. 2000, 523: 117-130. © 2000. Reprinted with permission from Blackwell Publishing.

providing a native environment for other constituents free of detergents. They preserve the asymmetric orientation of membrane proteins and may also preserve the distribution of lipids in inner and outer leaflets. Because they are still connected to the cell, they may be reached by diffusible intracellular compounds that modulate important membrane properties. Finally, they allow comparisons of different cells to test the effect of specific mutations or of up-regulation or down-regulation of membrane components. The formation of such protrusions of the plasma membrane is one of the many changes induced by eukaryotic cell injury. When a bleb bursts, the loss of the permeability barrier triggers the onset of cell death, but the events leading up to rupture are reversible. Cell surface blebbing has been observed '.\lith numerous cell types and may be caused by mechanical or chemical (e.g., depletion of ATP with potassium cyanide, injection of polar organic solvents, addition of iodoacetic acid) treatments, as well as bacterial infection of macro phages. The absence of actin and tubulin from blebs formed on oocytes of Xenopus laevus clea r1y indicates the bleb membrane is detached from the cell cytoskeleton (Figure 3.29). For this reason, the lateral mobility of certain integral membrane proteins measured by FRAP reveals faster diffusion rates in membrane blebs than in intact cells. Mass spectrometry shows a large number of (if not most) cell membrane lipids are found in the lipid composition of vesicles derived from blebs. Blebs have also been studied by confocal fluorescence microscopy, immunocytochemistry, EM, and patch clamp techniques. Example (Baumgart, T., et aI., Large-scale fluid/fluid phase separation of proteins and lipids i/1 giam plasma membrane vesicles. Proc Natl Acad Sci USA. 2007, 104:3/65-3170)

Membrane microdo111ains were observed in blebs induced by treat ing cultured fibroblast cells and leukemia cells with polar organic solvents. Imaging of two fluorescent dyes, napthopyrene that partitions preferentially in La phase and rhodamine-B-DOPE that partitions preferentially into Ld phase, shows that Lo-Ld fluid-fluid phase coexistence occurs in blebs at room tempemture as well as at 4°C (Figure 3.30). The variation in shape of the observed regions provides evidence for phase boundmy line tension. Selective partitioning into L o domains of the blebs by lipid-anchored proteins associated with rafis, obtained using antibodies against specific membrane proteins and green fluorescem protein/membrane protein chimeras, supports the hypothesis that the ordered regions ofthe blebs are rafis, which makes itthetirst physical demonstration of rafis in biological membranes on the micrometer scale.

In pathogenic bacteria, such as Neisseria gonorrhoeae, Pseudomonas aeruginosa, and Borrelia burgdorferi (Lyme disease), membrane blebs occur as part of the growth cycle and lead to the shedding of' membrane vesicles that probably have a role in the spread of infection. These blebs have been visualized by various microscopic techniques bu t have not been extensively used as model membranes. Bleb-like stnlctures called blisters have been made on giant liposomes containing reconstituted E. coli membrane fractions for patch clamp studies. Treatment with 20 mM Mg 2+ induces collapse of the liposomes, followed by the emergence of blisters of 50 to 100 ~lm diameters that are stable for several hours. Example (/yer, R., and A. H. Delcow; Complex inhibition of011lpF and OmpC bacterial porins by polyamines. J Bioi Chem. 1997,272:18595-/860/; Samartzidou, H., and A. H. Delcow; Distinct sensitivities of OmpF and PhoE poril1s to charged modulators. FEBS Lett. /999, 444:65-70)

3.30. Evidence for coexisting Ld and lo domains in blebs derived from mammalian plasma membranes. Direct visualization of large lateral domains in membrane blebs of cultured mammalian cells at 25"C is achieved by labeling with specific dyes, napthopyrene in A and rhodamine-DOPE in B. Previous experiments have shown that naphthopyrene preferentially labels the Lo phase and rhodamine-DOPE preferentially labels the lei (L,,) phase. Blebs are induced with injection of 4% (VN) ethanol. Scale bars 5 >J,m. From Baumgart, T., et ai, Proc Natl Acad Sci. USA. 2007, 104:36153170. Reprinted with permission of PNAS.

Model Membranes

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3.31. Electrophysiological tracings obtained by patch clamps on blisters on the surface of E. coli show the effects of physiological modulators of porin activities. A. OmpF porin with and without 0.1 mM spermine (a, b) and PhoE protein with and without 3 mM ATP (c, d). The current level corresponding to a large number of open pores is indicated by the baseline (BL) on the right. Redrawn from Samartzidou, H., and A. H. Delcour, FEBS Lett. 1999,444:65-70. © 1999 by the Federation of European Biochemical Societies. Reprinted "vith permission from Elsevier. B. Opposite voltage dependence was observed for OmpF (e) and PhoE (0). Redrawn from Samartzidou, H., and A. H. Delcour, EMBO J. 1998,17:93-100. © 1998. Reprinted by permission of Macmillan Publishers Ltd.

Application of the patch clamp lechnique to E. coli ouler membrane blisters gives much higher resolution lhan electrophysiological studies of purified outer membrane proleins in planar lipid bilayers and reveals velY fast and cooperative galing belween open and closed slales lhal is modulaled by physiological compounds. The opening of the cation-selective channels lhrough OmpF porin can be inhibited by polyamines, while the anion-seleclive PhoE channel can be blocked by ATP (Figure 3.31A). Because the ouler membrane fractions seem to incOl7Jorale in a preferred orienlalion in the blislers, they reveal lhe opposite voltage deperldence ofOmpF and PhoE channels (Figure 3.31B).

Nanodiscs The newest model bilayer system resulted from the study of high-density lipoproteins (HDLs), the "good" lipoproteins that circulate in plasma to transport cholesterol esters from the periphery of the body to the liver. Apolipoprotein (apo) A-I, one of the most common proteins found in these lipid-protein complexes, was shown to form amphipathic ex-helical belts that wrap around the lipid (Figure 3.32). The HDL complex can be reconstituted using purified apo A-I with different PLs, such as PC, PS, PA, PE, and/or cholesterol, as well as different acyl chain compositions. With controlled amounts of lipid, the reconstituted complex is a disc rather than a sphere (Figure 3.33). It contains approximately 160 PL molecules in a circular bilayer approximately 10 nm in diameter stabilized within the apo A-I protein, hence the name nanodiscs. Insertions or deletions in apo A-I were used to create modified apo A-I proteins called membrane scaffold proteins (MSPs) that give different controlled sizes of the nanod iscs. To incorporate membrane proteins, nanodiscs are assembled in the presence of the detel-gent-solubilized membrane protein of interest. Removal of the detergent produces a population of nanodiscs, many of which contain individual proteins for characterization. The first proteins to be reconstituted into nanodiscs were bacteriorhodopsin and cytochrome P450 reductase. Example (Shaw, A. W, el ai., Phospholipid phase transitions in homogeneous nanometer scale bilayer discs. FEBS Lett. 2004, 556:260-264) Nanodiscs prepared with eilher DPPC or DMPC were analyzed for phase tral1sitiol1s using DSC as well as parlitioning of a fluorescent probe sensilive lo the gel-loliquid clyswlline phase lransilion. When compared with the same sludies in lipid vesicles, the phase lransilions were broader and shifled 3° to 4" higher il1 nanodiscs thal1 il1 vesicles (Figure 3.34). The small size oflhe nanodisc bilayers bOlh decreases the coopera tivity ofthe lipids and increases the proporlion of lipids that inleraCl wilh

3.32. Structure of apo A-I lipoprotein (apo A-I) determined by x-ray crystallography. Deletion of the first 43 residues of this 243residue protein gives a structure in aqueous solution that is consistent with the structure of lipid-bound apo A-I, allowing solution of the crystal structure. The crystal reveals a dimer formed by two molecules of apo A-I tightly associated in an anti parallel orientation and curved into an elliptical shape with a hydrophobic strip along the interior. The amphilic nature is shown in this spacefilling model by the coloration of the amino acid side chains (Arg and Lys, blue; Asp and Glu, red; Phe, Tyr, Trp, Leu, Met, and Val, yellow; Pro, green; all others and main chain atoms, white.) From Brouillette, C. G., etal., Biochim BiophysActa. 2001,1531:4-46. Reprinted with permission from Elsevier.

lhe protein (boundary lipids; see Chapter 4). The broader transition o[ the l1anodisc lipids makes il more similar lhanlipid vesicles to a biomembrane whose complex composition eliminates a shal7J phase transition. In conclusion, while no model membrane is perfect, the an-ay of systems available enables researchers to apply a wide variety of techniques to studies of membrane components. Most of the information on lipidlipid interactions described in the last chapter was obtained using such systems. Furthermore, they are

A.

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,-

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3.33. Formation of nanodiscs from protein derived from apo A-I and detergent-solubilized phospholipids. The detergentsolubilized PLs are added to the protein derived from apo A·I (A). followed by removal of detergent (B) to produce nanodiscs (e). Redrawn from Shaw, A. W, et aI., FEBS Lett. 2004, 556:260. © 2004 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Model Membranes

67

B.

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Temperature (0C) 3.34. Comparison of the fluidity of phospholipid vesicles and nanodiscs revealed by phospholipid phase transitions. Two techniques were used to measure phospholipid phase transitions. A and B. Fluorescence data obtained with the dye laurdan (6-dodecanoyl-2-dimethylaminonaphthalenel, which incorporates readily into lipid bilayers with its hydrophobic tail embedded in the acyl chains. Its maximum emission wavelength shifts from 440 to 490 nm during the gel-to-liquid crystalline transition. Its generalized polarization (GP) is calculated from (1440 - 1490)/(1440 + 1490) and plotted as a function of temperature. For both DMPC (A) and DPPC (Bl, the transitions observed in vesicles (_) and nanodiscs (A) are quite similar. C. The phase transitions were compared by DSC, with the curves shown for DMPC nanodiscs (solid linel, DMPC vesicles (dashed linel, and DMPC nanodiscs after scanning to 90°C (dotted line). The scale on the left is for nanodiscs, while the scale on the right is for vesicles. Redrawn from Shaw, A. w., et aI., FEBS Lett. 2004, 556:260. © 2004 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

indispensable for the exploration of protein-lipid interactions as covered in the next chapter. FOR FURTHER READING

Jones, M. N., and D. Chapman, Micelles, MO/10layers and Biomembra.11es. New York: Wiley-Liss, Inc., 1995. Reviews

Garavito, R. M., and S. Ferguson-Miller, Detergents as tools in membrane biochemistry. J Bioi Chem. 200 1,276:3240332406. Helenius, A., and K. Simons, Solubilization of membranes by detergents. Biochim Biophys Acta. 1975,415:29-79. Rigaud, J.-L., B. Pitard, and D. Levy, Reconstitution of membrane proteins into liposomes: application to energytransducing membrane proteins. Biochim Biophys Acta. 1995,1231:223-246.

Seminal Papers

Denisov, 1. G., Y. V. Grinkova, A. A. Lazarides, and S. G. Sligar, Directed self-assembly of monodisperse phospholipid bilayer nanodiscs with controlled size. J Am Chem Soc. 2004,26:3477-3487. Montal, M., and P. Mueller, Formation of bimolecular membranes from lipid monolayers and study of their electrical properties. Proc Natl Acad Sci USA. 1972,69:3561-3566. Neher, E., and B. Sakmann, Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature. 1976, 260:779-802. Sanders, C. R., and G. C. Landis, Reconstitution of membrane proteins into lipid-rich bilayered mixed micelles for NMR studies. Biochemistry. 1995, 34:4030-4040. Sanders, C. R., and R. S. Prosser, BiceJles: a model membrane system for all seasons? Structure. 1998,6: 1227-1234. Other references given in examples and figures in text.

4

Proteins in or at the Bilayer


02 '-

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>\ u
- Phosphate

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.}

- Choline

.

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- Lys

The chemical and physical properties of the lipids described in Chapter 2 make it clear that the lipid bilayer provides a special milieu for proteins. Its constraints affect the structure, function, and regulation of proteins that dock on it or assemble in it to perform jobs such as energy transduction, nutrient transport, and signaling. The variety among proteins that interact in some \Nay with the membrane is quite astonishing and provides fascinating examples of how these proteins are structured to work in their environment. This chapter first defines the classes of proteins that are found in or at the bilayer. It describes many examples of peripheral proteins, as well as some lipidanchored proteins, before looking at what modulates the interactions of these proteins with membl-ane lipids. Next it focuses on molecules that insert into the membrane, including examples of toxins, colicins, and ioncarrying peptides. Then a look at the special qualities of the nonpolar milieu of the membrane explains many characteristics of integral membrane proteins. The chapter ends with studies of protein-lipid interactions in membranes.

68

Protein Insertion into lipid bilayers involves polar ilnrl nonpolar interactions. [Jxcillplified by the interaction of ,11118residue ~-hellx from coliCin A with the membrane interfacial re9lOr' and nonpolar i"terior. Three lysine residues (yellow) extend toward the aqueous milieu, while a tryptophan residue (green) reaches toward the cenler of the bililyer From Zakharov, S. D., and W A. Cri,mer, Biochimlc. 2002. 84:472. 2002 by Socete Fran~a,se de B,ochllll,e et Biolo· gie MoleccJlaire Rep, inled with permissior Irom Elsevier

CLASSES OF PROTEINS THAT INTERACT WITH THE MEMBRANE

A typical biomembrane contains many species of proteins, some embedded in the lipid bilayer and others on its surface. The Fluid Mosaic Model described in Chapter 1 distinguished between extrinsic and intrinsic membrane proteins by how easily they could be isolated from the membrane: extrinsic (peripheral) proteins can be removed by washes of the membrane, while the extraction of intrinsic (integral) proteins requires disruption of the membrane. Most integral membrane proteins have at least one TM peptide segment: bilOpic proteins have only one TM span, while polytopic proteins have more than one. In addition, there are 17'/onotopic proteins that insert but do not span the membrane and lipid-anchored proteins held in the membrane by covalently linked lipid groups, whether or not the polypeptide is membrane spanning (Figure 4.1). Those peripheral proteins that bind the membrane weakly and reversibly and are regulated by that binding are called amphitropic proteins and are involved in many

Proteins at the Bilayer Surface

69

Peripheral protein

Integral protein

GPI-linked protein 4.1. Schematic diagram showing classes of membrane and membrane-associated proteins: peripheral proteins, monotopic proteins, and TM proteins, including bitopic and polytopic proteins. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., w. H. Freeman, 2005, p. 374.

important biological functions. A few soluble proteins and many peptides insert into membrane bilayers to accomplish their functions, often coming from the outside and causing deleterious consequences to the cell or organism. Finally, some proteins blur the boundaries between classes since they are found in the cytoplasm, on the membrane periphery, and even inserted into membrane structures and can be fully characterized only when the dynamics of their localization are understood. To discuss the characteristics of proteins capable of existing in or interacting with the membrane, each of these classes will be described, giving attention to how the proteins and lipids interact.

ble proteins. In addition to electrostatic interactions, hydrophobic interactions with the acyl chains often contribute to the association of peripheral proteins \-\lith the membrane, as described below. The classic example of a peripheral protein is cytochrome e, whose cluster of lysine residues enables it to bind to anionic lipids of the bilayer as well as to acidic residues on the surfaces of cytochrome bel and cytochrome-e oxidase. The first use of ion exchange chromatography (in Uppsala, Sweden) was in the purification of this highly basic protein: 20 amino acids of the 104-residue human cytochrome e are lysine and arginine residues. Nine highly conserved lysine residues form a ring around the only exposed edge of its heme group (Figure 4.2). Differential labeling experiments have shown that the complex between yeast cytochrome e and cytochrome-e oxidase shields these lysine residues of cytochrome e and also shields three acidic residues of cytochrome-e oxidase. Further evidence that this is the interaction site between cytochrome e and the other cytochromes with which it reacts derives from the x-ray structure of a complex of cytochrome e and cytochrome-e peroxidase (which together reduce organic hydroperoxides) thaI reveals specific ion pairs between Lys73 and Lys87 of cytochrome e and Glu290 and Asp34 of the peroxidase. In the absence of other proteins, cytochrome c binds to bilayers fonned with anionic lipid and has been used in many studies of the interactions between peripheral proteins and bilayer lipids (see below). Electrostatic interactions are also paramount in the interactions of myelin basic protein with the myelin

PROTEINS AT THE BILAYER SURFACE

Extrinsic/Peripheral Membrane Proteins

The surface of some biomembranes is almost crowded with peripheral proteins in dynamic interplay with the lipid bilayer, with integral membrane proteins, and with each other. Typically, extrinsic membrane proteins bind to the surface of the membrane via electrostatic interactions, interacting with anionic lipids or with charged groups on other proteins, or both. Thus they sediment with the membrane and then can be isolated from it by treatment with buffers that vary the pH or increase the ionic strength; after sedimentation of the membrane under the new conditions, the extrinsic proteins remain in the supernatant and can be purified as solu-

73

4.2. Cluster of basic residues on surface of cytochrome c. Some lysine residues (dark blue balls) are strongly protected against acetylation when complexed with cytochrome-c oxidase or reductase, while others (light blue balls) are less strongly protected. Redrawn from Voet, D., and J. Voet, Biochemistry, 2nd ed., John Wiley, 1995, p. 579. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

IU

I

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4.3. Role of ankyrin in attaching various components of the cytoskeleton. Ankyrin (green) binds spectrin (pink, helical filaments) and numerous TM proteins (orange). Redrawn from Voet, D., and J. Voet, Biochemistry, 2nd ed., John Wiley, 1995, p. 302. © 1995. Reprinted with permission from John Wiley & Sons, Inc.

sheath. Myelin basic protein is a 21-kDa peripheral membrane protein that exists in isomers of different net charge. It is involved in the electrical activity of the myelin sheath and is implicated in the pathology of the autoimmune disease multiple sclerosis. Like cytochrome C, myelin basic protein requires anionic lipids for membrane binding and has been used to explore the energetics of the association between peripheral proteins and the bilayer (described below). Binding of one myelin basic protein immobilizes 18 lipid molecules of the bilayer as probed with EPR (see Box 4.2). Another familiar example of a peripheral protein is ankyrin, a 200-kDa globular protein that mediates the linkage of the cytoskeleton to the plasma membrane by binding to spectrin and numerous integral membrane proteins, including ion channels and cell adhesion molecules (Figure 4.3). The membrane-binding domain of ankyrin contains 24 ANK repeats, organized into four six-repeat folding domains. Each domain consists of a bundle of stacked antiparallel ex-helices connected by f)-turns, whose tips make contacts with other proteins (Figure 4.4). While ANK repeats have been found with assorted other domains in over 1500 proteins, ankyrin is composed almost entirely of them. Different isofonns of ankyrin have different combinations of the ANK repeat domains to make up distinct, highaffinity sites for protein binding. A hereditary ankyrin deficiency severely weakens erythrocytes, leading to anemia, jaundice, and eventually hemolysis.

An important family of peripheral proteins is the diverse group of phospholipases, water-soluble

4.4. Structure of ANK repeats, each composed of 33 amino acids that make up a pair of lX-helices connected by a tight f3-turn. ANK repeats bind to target proteins with the aligned f3-turns and the surface of the helical bundle. Three repeats from the yeast transcription factor Swi6 are shown. From Cell Signaling Technology Catalog, 2000, with permission from Pawson lab, (http://pawsonlab.mshri.on.ca/index.php?option=com_content &task=view&id=142<emid=64). © 2000. Reprinted with permission from Cell Signalling Technology.

Proteins at the Bilayer Surface

71

verted to arachidonic acid and then to the eicosanoidsprostaglandins, thromboxanes, and leukotrienes which trigger inflammation and other physiological reactions. For these peripheral enzymes, binding the membrane bilayer is mediated in part by binding their substrates (Figure 4.5A). The x-ray crystal structure of PLA 2 binding to DMPE illustrates its "lipid clamp," a highly specific binding pocket for lipid (Figure 4.5B). Crystal structures of PLA 2 with and without a phosphonate transition state analog have revealed the movement of a "flap" that appears to uncover the substrate binding site as well as a hydrophobic patch that seals the enzyme to the bilayer surface once the PL substrate is inside (Figure 4.6). A similar flap B.

A.

~ "Lid"

,

.-.'

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"Lid" 4.5. Membrane binding mediated by substrates. A. Model for PLA2 binding to bilayer surface via substrate binding. PLA2 binds to the membrane periphery (1) and then binds a phospholipid molecule in its active site (2). It cleaves the acyl chain (3) and then diffuses from the bilayer (4). Redrawn from Seaton, B. A., and M. F. Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, p. 363. © 1996 by W. Cho. B. X-ray crystal structure of PLA2 from cobra venom binding to DMPE. The bound Ca 2+ is magenta and the substrate, DMPE, is represented with a space-filling model in the active site. The PL substrates in a biological membrane would typically be four to six carbons longer and would be integrated into a leaflet as diagrammed in A. From Dennis, E. A., J Bioi Chem. 1994,269:13057-13064. @ 1994 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

enzymes that cleave the phospholipids of the bilayer. Some phospholipases are key regulatory enzymes, producing second messengers. For example, phospholipase C cleaves the glycerophosphate ester bond to liberate a phosphorylated alcohol and diacylglycerol, which activates protein kinase C (PKC). Phospholipase A2 (PLA2) releases the 511-2 fatty acid, which can be con-

Phospholid headgroup analog

4.6. Model for flap movement in PLA2. A. In the unliganded state, the left side of the substrate binding site forms the flap, or lid region, which can close over the bound Ca 2+ B. With the phosphonate transition state analog bound, the lid is held in place over it, which would seal the enzyme to the membrane if the substrate were a lipid from the bilayer. From Seaton, B. A., and M. F. Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, pp. 361-362. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

rTolelllS In or al lne ollayer

72

Membrane

A.

C.

Pl'otein core domain

N-terminal domain B.

Membr'ane

4.7. General structure of annexins. A. Schematic drawing of an annexin attached to a membrane surface through bound Ca 2 + ions (blue). Four ANK domains are shown, with the N,terminal domain on their surface away from the membrane. B. Structural model of annexin core based on alignment of over 200 annexin sequences. Each ANK repeat is colored differently, with Nand C termini black. Oxygens involved in binding Ca 2 + are red, and nitrogen atoms of highly conserved basic residues are blue. C. Annexin A6, with eight ANK repeats in two halves connected by a flexible linker (green), is capable of binding two membranes (at arrows). From Gerke, v., C. E. Creutz, and S. E. Moss, Nat Rev Mol Cell Bioi. 2005, 6:449-461. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

mechanism is observed in many other lipases and in some annexins. Annexins al'e a large family of proteins that require Ca2+ to bind to membranes and carry out a variety of functions. Their affinity for Ca2+, which enables them to mediate Ca 2+ signaling, is low in the absence of lipids and high (~M Kd) in the presence of anionic lipids. Members of the annexin family are structurally related proteins with four or eight repeats of a right-handed superhelical binding site for Ca2+, which together form a curved disc that binds the surface of the membrane (Figure 4.7). The dissimilar N-terminal domains of different annexins form an outward-facing concave side of the disc. They are "interaction domains" that enable annexins to form complexes with other proteins. Some annexins form complexes in a way that allows them to bring two membl'anes into close contact (although not to fuse without the help of fusogenic proteins; Figure 4.7C). Some participate in actin cytoskeleton attachment and others may playa role in cholesterol-dependent raft formation, while others have

been obser'ved to form two-dimensional ordered alTays on model membranes. Clearly their dynamic roles in lipid organization al'e important to many of their intracellular functions, such as endocytosis and lipid trafficking. Some annexins also have extracellular roles, such as suppression of inflammation and coagulation. Interest in annexins is high because of their roles in immune functions, their use as early markers for apoptosis, and their increased expression in certain tumors. Amphitropic Proteins

Many of these peripheral proteins are considered amphitropic proteins because their activity is regu, lated by the change from a water-soluble form to a membrane-bound form. This regulation may affect their catalytic function and/or their access to substrates, as well as their assembly into complexes (sometimes linking them to the cytoskeleton). While most amphitropic proteins, such as phospholipase C and PKC, come from inside the cell and al'e vital in signal transduction, some are extracellular proteins such as

Proteins at the Bilayer Surface

73

Membrane C2

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4.8. Model showing interaction of PKC with the membrane in response to elevated Ca 2 + levels. Ca 2+ binds to the C2 domain to tether PKC to the membrane with a low affinity, allowing the C1 domain to find and bind DAG, giving PKC a high affinity for the membrane and expelling its pseudosubstrate (N-terminal portion of the peptide chain), at the same time exposing the C terminus for autophosphorylation. Signaling activity is further controlled by phosphoryation and dephosphorylation, as well as association with Hsp70 and the cytoskeleton. For structural details of the C1 and C2 domains, see Figure 4.16. Here the C2 domain is yellow and the C1 domain is orange on the blue PKC; the pseudosubstrate is green. Redrawn from Newton, A C, et aI., Biochem J. 2003, 370:361-371. (C) 2003 by the Biochemical Society. Reprinted with permission from Portland Press Ltd.

blood clotting factors and apolipoproteins involved in transport of lipids through the blood. Activation of amphitropic proteins upon binding the membrane can be due to the proximity of effector molecules or of substrate, as in the case of the phospholipases. When restricted to the two-dimensional plane of the membrane, the effective concentration of the substrate (or effector) goes up approximately 1000-fold, calculated by comparing the volume of a sphere to the volume of surface phase: (4/3)nr 3 /4nr 2 d, assuming a spherical cell radius r of 10 j..lm and a surface thickness d of I to 10 nm. This concentration effect is also very important in facilitating interactions between membrane-bound proteins. Alternatively, activation can be due to structural changes induced in the proteins that relieve autoinhibition, as observed in PKC. PKC has two well-consel\led membrane-binding domains - CI, which binds diacylglycerol (DAC) or phorbol esters, and C2, which binds Ca 2 + - each triggering confonna tional changes. It appears that the initial interaction between Ca2+ -bound C2 and anionic lipids brings PKC close to the membrane to allow C 1 to penetrate and bind DAC (Figure 4.8; see also Figure 4.16A and B). Once both regulatory domains have bound, PKC undergoes a structural rearrangement that releases a pseudosubstrate N-terminaJ group from its active site and undergoes phosphorylation to produce the catalytically active enzyme. The CI and C2 domains are two of a small set of membranebinding domains that are used for signaling and subcellular targeting by many other proteins (described below).

Lipid-Anchored Proteins Lipid modification gives an opportunity to target proteins to the membrane (and even to specific intracellular membranes in eukaryotes) as well as to stabilize their interactions at the bilayer. The lipids can be fatty acids, terpenes, or glycosylphosphatidylinosi tol (CPI; Figure 4.9). The acyl chains are typically myristoyl groups in amide linkage to N-terminal glycine residues and palmitoyl groups covalently linked to seline or cysteine residues. Covalent modification with myristate occurs on nearly a hundred different proteins, not all of \vhich bind membranes. Peripheral membl-ane proteins with mutations at their myristoylation site no longer bind membranes, indicating the myristoyl group is required for membrane binding. Often these proteins have more than one lipid molecule bound because binding a single lipid anchor is not sufficient to maintain a stable attachment to the bilayer. The second acyl chain is frequently palmitate. Detergent-resistant membranes (thought to correspond to lipid rafts, see Chapter 2) are enriched with proteins modified with two or more acyl chains, and loss of one of these abolishes raft targeting. Terpenes, derived from isoprene, are either farnesyl or geranylgeranyl groups in thioether linkages to cysteine residues and are generally observed on proteins at intracellular membranes. Many members of the Ras-guanine triphosphatase (CTPase) superfamily have farnesyl adducts and are localized to the plasma membrane. Modification with isoprenyl groups tends to exclude a protein from lipid rafts, presumably since these groups would disfilpt the packing of acyl chains in the La phase.

t-'rotetns in or at the I:3llayer

74

+

NH)

Palmiloyl group on inlernal Cys (or Ser')

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Inside

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4,9. Lipid anchored proteins. Proteins may be anchored to the bilayer by covalent modification with acyl chains (typically myristate and palmitate), with terpenes (such as farnesyl or geranylgeranyl groups) and with GPI. In the plasma membrane the proteins covalently attached to acyl chains and terpenes are found on the cytoplasmic surface, while proteins attached to GPI are on the extracellular surface. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 379. © 2005 by W. H. Freeman and Company. Used with permission.

Proteins at the Bilayer Surface

75

GPI core structure Sugar modifications Attachment of sugars or phosphoethanolamine

o

0

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Lipid modifications • Acylation of inositol dng • Fatty-acid exchange or modification (changes in saturation, hydroxylation, linkage) • Lipid-backbone exchange (DAG -7 ceramide)

4.10. Core structure of G PI anchors, showing locations of sugar and lipid modifications. This structure has three mannose residues and a glucosamine on the phosphatidylinositol, but it can be varied by addition of extra sugars or ethanolamine phosphates to the mannose residues, acylation of the inositol, changes in the fatty acids (length, saturation, hydroxylation) or their linkages to the glycerol backbone, or remodeling of the entire DAG to ceramide. Note also the amide linkage between the ethanolamine moiety of GPI and the carboxyl group created by cleavage at the w-site of the precursor protein. Redrawn from Mayor, S., and H. Riezman, Nat Rev Mol Cell BioI. 2004, 5:110-119. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Examples of lipid-anchored proteins are found in bacteria, such as lipoproteins in E. coli and penicillinase in Bacillus lichel1ifonnis, as well as in eukaryotes, such as the catalytic subunit of cyclic adenosine monophosphate (cAMP) protein kinase, the G protein complex (GO<. is acylated and Gy is isoprenylated), the Src family of tyrosine kinases, and the Ras superfamily of small GTPases. While some lipid-modified proteins such as rhodopsin and the transfer.-in receptor are membrane spanning (and will be covered in other sections of this book), many are soluble except for the lipid anchor and thus are held on the membrane surface. Proteins linked by GPI are common in animal cell membranes, and in fact account for ~0.5% of all eukaryotic proteins. Normally components of plasma membranes, they can also be found in different internal membranes following endocytosis. They play many important roles, including essential functions in embryonic development in animals, and they are also essential for viability in lower eukaryotes such as fungi. Proteins destined for attachment to GPI are synthesized as precursors with GPI-anchor sites, called w-sites, near their carboxyl termini. The GPI moiety is synthesized by a number of glycosyl transferases. On the luminal side of the ER membrane, a GPI transamidase cleaves the peptide bond in the precursor and forms an amide bond to the ethanolamine of the GPI moiety. When these proteins do not get their GPI anchor, they remain soluble. While GPI anchors have a common core structure, they differ in their sugar and fatty acid composition, including both saturated and unsaturated acyl chains

(Figure 4.10). Furthermore, some lipid moieties are modified after attachment to the protein; for example, in many yeast GPI-anchored proteins DAG is replaced with ceramide. The GPI anchor of prion is modified with sialic acid, which could increase its lateral mobility in the membrane. Most likely these differences provide additional information for targeting GPI-linked proteins. The role of the GPI anchor as a sorting tag was suggested by early studies of the membranes of epithelial cells whose apical and basolateral surfaces differ in composition. GPI-anchored proteins were found concentrated at apical surfaces. Furthermore, if typically basolateral membrane proteins were modified with a GPI tag, they also localized in the apical surface. GPIanchored proteins are found in DRMs (see Chapter 2), along with sphingolipids and cholesterol. The presence of GPI-anchored proteins in DRMs depends on the presence of cholesterol, since they are solubilized by detergent treatment if the cells are first treated with saponin, which complexes cholesterol. These results have led to the concept that lipid rafts are enriched in GPIanchored proteins, where they interact with tyrosine kinases and other signaling complexes. Severa] techniques have been used to probe the association of GPI-anchored proteins with rafts. Results from FRET measurements carried out with varying protein concentrations indicate that some portion of GPI-anchored proteins are not randomly distributed, although the proportion depends on the study. Dynamics were revealed by single-particle tracking of a

rrOlelnS III

76

A~

u~

B(]J -K on ~

Koff

C.

-~

4.11. Different modes of binding amphitropic proteins to membranes. A. Electrostatic interactions between a polybasic motif on the protein and the anionic lipids, which can be prevented by polyphosphorylation. shown together with insertion of a lipid anchor, in this case an acyl chain that was sequestered in the protein interior. B. Membrane clamp, a binding pocket with an affinity for a specific lipid headgroup, which in some cases is an inhibitory domain to activate the enzyme. C. Insertion of an amphipathic <x-helix into the bilayer. Redrawn from Johnson, J. E., and R. B. Cornell, Mol Membr BioI. 1999, 16:217235. (~J 1999. Reprinted by permission of Taylor & Francis Ltd, http://www.informaworld.com.

gold-linked GPI-anchored protein that recorded "transient confinement zones" of 200 to 300 nm diameter in which the panicles were trapped for 10% to 15% of the time. It is likely that such large zones consist of smaller domains, each of which contains a few GPI-anchored proteins. Whether and how the proteins are targeted to these domains is still unclear, but it is possible that the nature of the GPT anchor is one determinant of raft localization since GPI anchors containing unsaturated acyl chains are expected to be much less compatible in Lo phase than those containing saturated chains (see "Lateral Domains and Lipid Rafts" in Chapter 2). Reversible Interactions of Peripheral Proteins with the Lipid Bilayer

The reversible associations between peripheral proteins and membranes are accomplished by a number of mechanisms, singly or in combination (Figure 4.11).

or

iH lne Dllayer

An important mechanism is electrostat.ic attract.ion bet.ween charged groups on the peripheral protein and chal-ged groups on either lipid 01- protein component.s of the membrane, exemplified by cytochrome c and myelin basic protein described above. As will be seen below, changes Lhat affect the net charge on t.hese peripheral prot.ei ns alter their affinity for the membrane, setting up an "electrostatic s\vitch" that can control their binding. A second mechanism is the insertion of a lipid anchor, which occurs when proteins with covalently bound acyl chains can markedly shift the position of the chains in the aqueous and membrane-bound forms. Tn the water-soluble form of such a protein, the acyl chain is sequestered int.o the protein interior; then at the membrane, it projects from t.he protein to embed into t.he closer leaflet of the membrane. In the third mechanism, the protein has a binding site for a particular lipid that is often specific for the headgroup. In this case, association of the protein with the membrane depends on it.s affinity [or the lipid as well as the concentration of t.he lipid. To modulate the binding to a high-affinit.y site, the protein may have a "flap" that covers the binding site in the aqueous form, as described for PLA 2 above. The fourt.h mechanism is insertion of an amphipathic helix in the interfacial region of the bilayer, requiring a large conformational change in the protein. While the peptide usually occupies the polar headgroup region of t.he membrane, in some cases it may penetrate the nonpolar region, in which case it generally does not extend more than halfway into the leaflet. to which it binds (see Frontispiece). Before describing these interact.ions in more detail, the effects on membr'ane lipids of binding peripheral protein will be considered.

Effects of Peripheral Protein Binding on Membrane Lipids Thermodynamic st.udies reveal the impact of binding a peripheral protein on the general properties of lipids in the bilayer~ exemplified by DSC measurements of lipid phase transi Lions in the presence of increasing amounts of protein. These calorimetry experiments measure the heat capacity of t.he system, which is the heat input divided by the temperature change, as described in Chapter 2. Both cytochrome c binding to DMPG bilayers and myelin basic protein binding to DMPS bilayers produce shifts in Ton, although t.he shifts are in the opposit.e directions (Figure 4.12). Binding increasing amounts of cytochrome c results in increased broadening of the phase transition as well as a shift of Too up to +5'°C, chieny due to the effect of protein binding on the surface electrostatics of the bilayer. When t.he chains melt, the charge density is less since the area per lipid is larger, which means there is a lower electrostatic binding energy between cytochrome c and the membrane. (In addition when cytochrome c binds,

Proteins at the Bilayer Surface

77

3,=-----------------,

Myelin basic protein + DMPS

cytc + DMPG

biJ

2

Q)

""? "0

biJ

-2

Q)

-0

~ ~

c.

u

P/L:

,2:,

'u (\l

0..

(\l

u

(\l Q)

::c

0 0 0

-.,r-

o

:

"0 E

LlT m ""'5°: :

--

~

:

=. U

1~

U

1/20

P/L

= 1/25

-)

1I1.i!0_--.i.-----:-

32

27

=0

0

1/~

22

P/L

Temperature [OC]

20

25

30 35 T [OC]

40

45

4.12. Effect of peripheral proteins on heat capacity of lipid bilayers. Changes in heat capacity detected by DSC illustrate the influence of binding peripheral proteins on the chain melting of the lipid. A. Binding cytochrome c to DMPG bilayers was measured at five different degrees of surface saturation indicated by the protein: lipid ratio (P/L). Increasing surface saturation broadens the transition and shifts the heat capacity maximum (Tm), until at ~ 100% saturation it shifts by 5°, corresponding to a change of -4.3 kcallmol. From Ramsay, G, et aI., Biochemistry. 1986,25:2265-2270. © 1986 by the Biophysical Society. Adapted with permission from the Biophysical Society. B. Binding myelin basic protein to DMPS bilayers broadens and lowers (by 1.6 kcal/mol) the heat capacity of the phase transition. Redrawn from Heimburg, T., and R. L. Biltonen, Biophys J. 1996, 70:84-96. © 1980 by American Chemical Society. Adapted with permission from American Chemical Society.

Homogeneous mixture

peripheral

Demixing according to mass action law

'-/'-./'-"'-'-../'-"''-,-'-.-./-P~i~ ~1ijm peripheral protein

Total demixing

4.13. Schematic representation of the different effects of peripheral proteins on lipid organization. Two different species of PL are indicated by the light and dark headgroups. Binding of a peripheral protein can cause (1) no lipid rearrangement (top); (2) redistribution of lipids with no preferential interaction between them, according to different affinities for protein (middle); and (3) concentration of certain lipids in a domain that binds protein, resulting in total demixing (bottom). Redrawn from Heimburg, T., and D. Marsh, in K. Merz and B. Roux (eds.), Biological Membranes, Birkhauser, 1996, p. 410. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

the strong ionic attractions to anionic headgroups can cause those phospholipids to cluster, as described in the next paragraph.) Titration calorimetry indicates the binding of cytochrome c is endothem1ic; therefore, it is driven by entropy. The opposite shift is observed with myelin basic protein, where the binding is driven more by enthalpy, most likely because in addition to the electrostatic interaction with anionic lipid headgroups, myelin basic protein has hydrophobic segments that intercalate into the nonpolal- domain of the bilayer. In addition to the eFFect on heat capacity, protein binding to the bilayer can affect the organization and state of the lipids. HO\,\f much the lipids rearrange when the protein binds depends on the relative affinities of the different lipid species for the protein as well as the mixing properties ofthe lipids, that is, whether the intel-actions between lipids are favorable or unfavorable (Figure 4.J 3). This subject is of great relevance to the mechanism of formation of lipid rafts. Measurements of cytochrome c binding to the surface of DOPG bilayers at different ionic strengths fit quite well to predicted isotherms for nonlocalized binding with little interaction between proteins, with the effective charge determined to be +3.8 for cytochrome c (Figure 4.14; see Box 4.1). The calculations give a lipid stoichiometry of 12 lipids per protein, like the stoichiometry determined by crystallography. Interactions between Peripheral Proteins and Lipids

Binding of peripheral proteins typically involves electrostatic interactions. If the binding shows an absolute requirement for anionic lipid and is abolished in high ionic strength, it can be attributed solely to electrostatic interactions. Studies of peptides whose binding to the membrane is strictly electrostatic indicate that these peptides actually remain about 3 A from its surface. When basic peptides bind to acidic lipid vesicles, the contlibution from each positive charge on the peptide is ~ - J.4 kcal. The strength of the association is not dependent on the chemical nature of the acidic lipids (e.g., PG, PI, or PS) or of the amino acids (Lys or Arg), but obviously it is diminished by deletion of one or more of the basic amino acids. Reversible reactions that decrease the net charge provide opportunities for modifying the force of the eleco-ostatic interaction (see below). Quite often, binding of peripheral proteins to the membrane surface involves hydrophobic interactions in addition to electrostatic interactions. When a peripheral protein inserts one or more acyl chains to anchor into the bilayel~ the hydrophobic interaction makes a significant conttibution to the binding. Studies with model acylated peptides show the binding energy is proportional to the chain length, with -0.8 kcallmol per CH z group, mirroring Tanford's partitioning data for

30 28 26 24 22 20 18

2: ::i ;::; 16

a = J 1.9

, .. - - -

-_ ... -- ...... -_ ...... - ...... -- _... -- - --- ... -_ ...... -- _... - - --

v

<:

6 14

...

J:)

0...

'-' 12 10 8 6 4 2 20

40

60

[Pw1aiJ (iJ.M) 4.14. Binding isotherms for cytochrome c binding to DOPG at neutral pH, 200C, which plot the concentration of protein bound (y-axis) as a function of the total protein (x-axis) at ionic strengths of 41.9,54.4,79.4, and 104.4 mM (from top to bottom). Redrawn from Heimburg, 1., and D. Marsh, in K. Merz and B. Roux (eds.), Biological Membranes, Birkhauser, 1996, p. 415. © 1995 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

fatty acids (see Figure 1.4). Myristoylated peptides bind electrically neutralliposomes with ~G = -8 kcal/mol, which indicates that 10 methylenes of the 14 in the chain are embedded in the nonpolar domain. The corresponding binding constant (K n ) of 10 4 M- 1 is insufficient to maintain stable binding of the protein to the membrane. Additional binding strength is provided by a second acyl chain or prenyl group, or by another mechanism, such as a group of basic residues that participate in electrostatic interactions. The latter mechanism is used by the signaling kinase Src, which is myristoylated on its N-terminal glycine and has three lysine and three arginine residues in its membrane-binding N-terminal domain (Figure 4.15). Measurements of the binding of the J5-residue peptide corresponding to this domain (myristate-GSSKSKPKDPSQRRR') to vesicles containing 2:1 PCPS show the myristoylated peptide binds with K" = 10 7 M- 1 , while the non myristoylated Single-leLter amino acid codes. often used for motif names and for peptide sequences, are given in Appendix II.

Proteins at the Bilayer Surface

79

BOX 4.1. Binding of ligands to surfaces The Langmuir isotherm gives the simplest mathematical model to describe the binding of a ligand to defined binding sites on a surface and can approximate the binding of peripheral proteins to the surface of a membrane. It characterizes the binding of the surface, S, by the ligand, L, as S + L --+ SL by describing the binding constant, K, in terms of bound and free sites:

K = [Sbl I ([Sfl [Lj), where Sf are free binding sites and Sb are sites with ligand bound, and [L] is the concentration of free ligand (protein). Solving this equation for [LI gives

Then if 8 is the fraction of occupied sites, 8 = [Sbl I ([SbJ [Sil and substituting that into the equation derived for [L] gives the Langmuir isotherm,

Langmuir-type isothenn:

+ [SIl). Solving this equation for [Sb] I

Localized binding sites

8

1

[Ll =

Continuous surface:

In-plane interactions: Distributional entropy, Protein-protein interactions 4.1.1. Different models for binding of peripheral proteins to a surface. Redrawn from Heimburg, T., and D. Marsh, in K. MerL and B. Roux (eds.), Biological Membranes. Birkhauser, 1996, p. 408. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

K . (1 -8).

In the Langmuir-type isotherm the proteins bind to a surface with fixed, independent binding sites (Figure 4.1.1, top). However, the Langmuir isotherm is a fair approximation for protein binding to a bilayer only in the condition of very little protein binding (8 « 1) as it does not account for steric interactions between proteins that become likely at higher amounts of binding. Furthermore, the binding sites for peripheral proteins on a membrane are dynamic and delocalized (Figure 4.1.1, bottom). When they bind to a continuous surface, the proteins can rearrange on the surface and interact with neighboring proteins. The mathematics to describe such delocalized binding utilizes the Gibbs absorption isotherm, along with Gouy-Chapman and Debye-Huckel theories to model the electrostatic energy of binding. The Gibbs absorption isotherm treats the lateral interactions between proteins on the surface as a two-dimensional pressure, II(i), where i is the number of proteins bound. Then d

n

(i) I d In [L]

= i kT I

n AA,

where n is the number of proteins that saturate the surface and AA is the excluded area per protein. [LI is still the free protein concentration. In the simplest case,

.J-

n

(i) = ikTI (n-i) AA

( IJ,)

-1-

I ~D

(the Volmer equation of state). After substituting and integrating, the isotherm becomes

1

[Ll

= K.

8

8

(1 -8) exp (1 -8)

with e = j / n. This equation fits the binding to continuous surfaces with delocalized sites much better than the Langmuir isotherm and allows for a higher binding constant. To calculate the free energy of binding of a charged ligand to a membrane with consideration of surface electrostatics, both the Gouy-Chapman and Debye-Huckel theories are used. The Gouy-Chapman (and later Gouy-Chapman-Stern) theory describes quantitatively the electrical potential energy as a function of the distance from the bilayer surface, assuming the charges on the surface are spread out rather than localized. An important result is the existence of an electrostatic double layer created by the balance between the entropic drive for ions to randomize and their electrical attraction to the surface. The Debye-Huckel theory gives the distribution of electric potential around an ion in solution, describing the thermal and electrical forces around the ion. Using Gouy-Chapman to calculate the electrostatic free energy of the bilayer surface and Debye-Huckel for the electrostatic energy of the free peripheral protein (the ion) gives a complex mathematical expression for the change in total electrostatic free energy when the charged protein binds to the surface. From the fit of empirical data to the theoretical curves, it appears that most peripheral proteins bind to membranes with some demixing (nonrandom rearrangement) according to the law of mass action, as shown in the intermediate diagram of Figure 4.13.

A.

C

+++ +++

:-

minus and the cluster of basic residues. In other peripheral proteins the distance is greater, yet the anchorc ing greatly increases the probability that the oppositely charged groups will come in close proximity. A good example of combining both hydrophobic and electrostatic interactions to bind the membrane is provided by the Cl and C2 domains of PKC described above. The Cl domain has a well-characterized binding site for DAG and phorbol esters (analogs of DAG that promote tumor formation), and the C2 domain has a Ca 2+-binding site that interacts with anionic lipids. The Cl domains do not interact with membranes lacking DAG and show strict specificity for the physiological stereoisomer. The binding has been characterized by fluorescence (using phorbol ester analogs), NMR, and x-I'ay crystallography. Lipid binding displaces water from the pocket and increases the hydrophobicity of the protein surface, thus increasing its affinity for the membrane. The C2 domain binds two to three calcium ions coordinated by aspartate residues and carbonyl groups, which stabilizes a loop region of its structure and increases the affinity for the membrane. Domains Involved in Binding the Membrane

4.15. Models of the protein-lipid interactions involved in binding the N-terminal Src peptide to lipid vesicles. A. A cartoon of the domain structure of c-Src shows the myristate chain on the Nterminus, near a cluster of basic residues that interact with anionic lipids in the bilayer. B. A space-filling representation of the portion surrounded by a dashed line in A illustrates the docking of the myristoylated eighteen-residue peptide in a PCPS (2:1) bilayer. The model shows the insertion of the myristoyl group (green) and the interactions of the six basic residues (blue) of the N-terminal peptide with phosphat idyl serine headgroups (acidic residues red and the nitrogen atoms blue). Insertion of the acyl chain confines the peptide, increasing the chance of forming electrostatic interactions between the basic residues with anionic lipids. From Murray. D., et al., Structure. 1997, 5: 985-989. © 1997 by Elsevier. Reprinted with permission from Elsevier.

Src peptide binds with K. = 10 3 M- l . Recalling that the K, for insertion of the myristate is 10 4 M- l , it is clear that the hydrophobic and electrostatic energies add (i.e., the binding constants multiply). In addition to its contlibution to the binding energy. the insertion of a lipid chain can help localize the basic residues of the peripheral protein to the appropriate sites on the membrane. In the case of the Src peptide, there is only I to 3 nm between the lipid-anchored N ter-

C I and C2 domains of PKC have now been' observed in many proteins involved in cell signaling. In addition there are two other dominant types of domains utilized for membrane binding by peripheral proteins: FYVE zinc-binding domains, named for their amino acid motif, bind to polyphosphorylated inositol, and pleckstrin homology (PH) domains bind phosphoinositides and are essential in P13-kinase signaling pathways. These Cl, C2, FYVE, and PH domains have now been identified in hundreds of proteins involved in signal transduction and membrane trafficking, enabling them to bind the membrane and thereby regulating their localization and activity. Each domain type consists of a specific binding site (usually for a lipid ligand) that is often flanked by additional, less specific membrane-binding sites. The nonspecific binding is important in increasing the overall affinity for the membrane andlor making an additional point of contact to define the stereoselectivity of the specific binding site. The weak interactions can also be enhanced by the combination of two (or more) domains in the protein or by the oligomerization of protein subunits, which allows the temporal variation needed in regulatory systems. Examples of each of the four types of domains have been characterized by x-ray crystallography (Figure 4. 16), NMR, fluorescence, mutagenesis, and localization of green tluorescent protein fusions. The Cl domain, now identified in > 200 different proteins, has ~50 amino acids making t\VO smalll3-sheets 'with a short ex-helix built around two 3-Cys-l-His clusters that bind

Proteins at the Bilayer Surface

81

A.Cl

B. C2

Interface Hydrocarbon core

C. FYVE

D.PH

Interface Hydrocarbon core

4.16. Four types of membrane-binding domains found in hundreds of peripheral proteins involved in signal transduction. The x-ray structures reveal major features of four membrane-binding domains: A. C1 domain from protein kinase C; B. C2 domain from PLA2; C. FYVE domain from Vps27p (a yeast protein for endosomal maturation); and D. PH domain of phospholipase C. Selective residues that make contact with the surface are labeled, and specifically recognized lipids are modeled. PI3P is phosphatidylinositol 3-phosphate and PI(4,S)P2 is phosphatidylinositol 4,S-bisphosphate. The membrane leaflet is divided into an interfacial zone and the hydrocarbon core. Hydrophobic residues are colored green and basic residues are blue. From Hurley, J. H., and S. Misra, Annu Rev Biophys Biomol Struct 2000,29:49-79. © 2000 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

Zn 2+ very tightly. As described for PKC (above), this domain binds DAG and phorbol esters. The binding occurs at the tip of the domain, unzipping the two 13strands to expose the binding site. This groove is located in a hydrophobic end of the domain that is adjacent to a ring of basic residues posi tioned so that membrane penetration by the hydrophobic tip allows the basic ring to contact the membrane surface. Most Cl domains are not targeted to the membrane if they lack specific binding sites for DAG, although there are a few atypical Cl

domains that do not require DAG. Most PKCs and DAG kinases have pairs of Cl domains, and in some, DAG is an aHosteric activator. C2 domains, identified by a conserved sequence motif that binds Ca2+ reversibly, have been found in >400 proteins, including many involved in signal transduction, inflammation, synaptic vesicle trafficking, and membrane fusion. The C2 domain is a f3-sandwich like the immunoglobulin fold, with the Ca 2+ -binding sites formed by three loops at a tip analogous to the

C=O

C=O

I

C=O

I

0-

I

C=O 0

I

0-

C=O

I

I

OH

OH H+ H+ H+I-{+

interfacial pH == bulk pH Zwitterionic surface

C=O OH

H+

~ interfacial pH < bulk pH Anionic surface

4.17. Helical wheel of the amphipathic helix of cytidyltransferase. Since the pH at the surface of anionic (but not zwitterionic) membranes is lower than the bulk pH due to the attraction of protons to the negative surface, the probability of protonation of three Glu residues increases, which effectively increases the hydrophobicity of the surface of the peptide. Redrawn from Johnson, J. E. et aI., J BioI Chem. 2003, 278:514-522. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

antigen-binding site. It is not easy to generalize about this domain. There are five different C2 domain structures that Fall into two permutations of this fold. Some are hydrophobic enough to penetrate the membrane, while others are not; most require acidic PLs, while the C2 domain of the c isoform of PLA 2 prefers neutral lipids, especially PC. There are even C2 domains that do not bind calcium! The FYVE domain, identified in ~60 proteins, is noted For its specificity in binding phosphatidylinositol 3-phosphate (P1P3), which enables it to target proteins to endosomal membranes that are enriched in PIP 3 . This domain consists of 70 to 80 residues, forming two small double-stranded l3-sheets and an <x-helix, with a conserved RKHHCR motif that binds PIP 3 . It also contains eight Cys or seven Cys with one His that coordinates two Zn 2+ ions, two Leu residues at one end that protrude into the membrane, and a few less-conserved Lys residues that probably contact the membrane surface. The PH domains bind different phosphoinositides with varying degrees of specificity and thus respond to signaling that interconverts phosphoinositides having different phosphorylation patterns. Found in >500 proteins, this domain consists of two curved l3-sheets of three or four strands capped by an <x-helix. It has different subsites that bind phosphate groups to make up the substrate-binding site of varying a ffinity and specifici ty. It has a positively charged face that interacts with acidic lipids in the membrane; mutations that strengthen this

interaction can result in constitutive activation, while mutations that decrease it can lead to loss of function. Some PH domains also participate in proteinprotein interactions and some provide allosteric regulation. Like the other domains, adjacent portions of the PH domain contribute nonspecific interactions to give variable interplay with the membrane. A less common mechanism for binding to the membrane is the insertion of an amphipathic <x-helix parallel to the plane of the bilayer, observed in a miscellaneous group of proteins including DnaA, adenosine diphosphate (ADP)-ribosylation factOl~ vinculin, epsin, several regulators of G protein signaling, and cytidyltransferase (CT). The hydrophobic interactions between the hydrophobic face of the helix and the nonpolar core of the bilayer provide the driving force for insertion, which is opposed by the resulting perturbation of lipid packing. Formation of the <x-helix concomitant with insertion can provide a significant additional driving force (see below). CT is a well-characterized representative of this group of amphitropic proteins. It carries out the transfer of a cytidyl group fTom cytidine triphosphate (CTP) to phosphocholine in the key regulatory step for the synthesis of Pc. Studies employing circular dichroism indicate that its ~60-residue membrane-binding domain changes from a mix of l3-strand, l3-turn, and disordered conformations to <x-helix upon binding the membrane. The amphipathic helix has basic residues on one face and a mixture of acidic and basic residues on another,

Proteins at the Bilayer Surface

83

with a center strip dominated by acidic residues. A nonpolar face contains a total of 18 hydrophobic residues giving a hydrophobic surface area of ~2500 A2 .lnterestingly, three glutamate residues positioned at the interfacial region contribute to the selectivity of CT for anionic lipids because they become protonated in the low pH milieu at the surface of anionic, but not zwitterionic, membranes (Figure 4.17). Studies of CT bindi ng to multilamellar vesicles (MLVs) suggest that the first step of membrane association is electrostatic: CT binds when the MLVs are in gel phase, but can only insert its amphipathic helix upon raising temperature above the transition to liquid crystalline phase. Modulation of Binding Since membrane binding regulates the actIvIty of amphitropic proteins and thus controls many key processes of the cell, modulation of reversible binding to the membrane is crucial. It is accomplished by one or more of several mechanisms that respond to signaling kinases, altered levels of ions or effector molecules, or changes in local compositions of the membrane that are sometimes linked to trafficking, which is the targeted movement of specific molecules to particular regions or organelles. A simple mechanism is the disruption of the electrostatic interactions between basic groups of the protein and anionic lipids by the addition of negative charges when the protein is phosphorylated on serine or tyrosine residues in the membrane-binding region. In the example of the Nterminal region of Src, the phosphorylation site is a serine located between the clusters of lysine and arginine residues. Because the phosphate can be cleaved by phosphatases, the reversible change in charge provides an "electrostatic switch" that affects membrane binding. Another mode of regulation is by the enzymatic addition ctnd removal of an acyl chain. In these cases, the first lipid anchor is permanently added in a posttranslational modification and barely provides enough energy for the protein to bind to the bilayer. Modification with a second acyl chain (or prenyl group) doubles the hydrophobic interactions with the bilayer, converting a weak association into a strong one; thus its addition and removal regulate localization to the membrane. For example, after the ex subunit of the G, protein complex is myristoylated in a posttranslationaJ modification that remains for the lifetime of the protein, specific addition and removal of a palmitate chain by acyltransferase and thioesterase activities control its affinity for the membrane. A third major type of regulation involves binding ligands such as nucleotides or calcium. The binding of guanosine triphosphate (GTP) to the G pl-otein ADPribosylation factor causes a conformational change that exposes its amphipathic helix, which then inserts

4.18. Model for the Ca 2 + -triggered extrusion of a Trp residue in annexin V. The ribbon diagrams show annexin V viewed from the side with the membrane-binding surface face up and the mobile Trp residue at the right. Ca 2 + ions are blue spheres. A. If Ca 2 + is absent from domain 3, the Trp side chain is buried. B. When Ca2+ binds domain 3, the Trp side chain emerges from its buried position to interact with the lipid bilayer. From Seaton, B. A., and M. F. Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, p. 38S. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

into the bilayer. Calcium binding can trigger structural changes in the protein that affect its membrane-binding surface, as seen in annexin V. The crystal structure of annexin V shows a flattened molecule with opposing convex and concave faces. EM studies suggest the convex side flattens on the surface of the membrane, aJlowing multiple Ca2+ -binding loops to contact the surface. Ca2+ binding triggers the rotation of a single tryptophan side chain, moving it from a buried position to a protrusion that intercalates into the bilayer (Figure 4.18). Binding calcium has a very different effect on the retinal protein recoverin. When recoverin binds Ca 2+, its conformational change induces the extrusion of a bound myristoyl group that becomes a membrane anchor (Figure 4. J 9). This "myristoyl switch" has been observed in other peripheral membrane proteins that are involved in signaltTansduction. Finally, those amphitropic proteins that bind a specific lipid component of the membrane, such as DAG or polyphosphol)dated inositols, are subject to temporal changes in the concentrations of their ligands in the bilayer, and these concentrations are controlled by the phospholipases in response to signaling. Furthermore, the different concentrations of the specific lipid ligands in various membranes ofeukaryotic cells can contribute

Calcium-myristoyl switch

2 Ca 2 + ~

•

T

R

4.19. Diagram of the calcium-myristoyl switch. A conformational change in recoverin enables it to extrude its bound myristoyl group upon binding calcium. The protruding acyl chain interacts with the lipid bilayer and activates the protein to prolong the photoresponse of rod and cone cells. From Seaton, B. A., and M. F Roberts, in K. Merz and B. Roux, Biological Membranes, Birkhauser, 1996, p. 393. © 1996. Reprinted with permission from J. Ames, M. Ikura, and L. Stryer.

to membrane-selective targeting, as seen in the enrichment of endosomal membranes for PIP,.

carries out its translocation. DT has three domains, a catalytic C domain at the N terminus, a receptorbinding R domain at the C terminus, and between them a T domain involved in translocation (Figure 4.20). After binding to a receptor on the cell surface, DT is cleaved into the A fragment consisting of the C domain and the B fragment containing both T and R domains; A and B are linked by two disulfide bridges. Because DT is internalized by endocytosis, it enters the cell in acidic membrane-bound compartments called endosomes. The low pH of the endosomes triggers an acid-induced conformational change that enables the T domain to insert into the endosomal membrane and translocate the A domain into the cytosol where it carries out ADP-ribosylation of elongation factor 2, inhibiting protein synthesis and leading to cell death. The T domain of DT is a bundle of 10 ex-helices, two of which are hydrophobic and insert as a helical hairpin that has been shown to span the membrane. In planar lipid bilayers at low pH (::;6), DT as well as its B fragment and isolated T domain have all been observed to form cation-selective channels, but the structure of the pore and how it translocates the A fragment is not known.

PROTEINS AND PEPTIDES THAT INSERT INTO THE MEMBRANE

While a fe\-\' amphitropic proteins have small segments that insen into the nonpolar domain of the bilayer, there are soluble proteins and peptides that insert more extensive portions to cross the bilayer, often via major conformational changes. Insertion of these TM domains is generally governed by the same forces that drive the insertion o[ TM segments of integral membrane proteins, described in detail below. Besides the hydrophobic and electrostatic forces, insertion of a peptide across the bilayer involves the perturbation of the acyl chains in the membrane, immobilization of the peptide, and possibly its unfolding or refolding. Clearly the state and lateral tension of the lipid are important: the enthalpy of peptide insertion into small unilamellar vesicles was greater for insertion into tightly packed vesicles than loosely packed ones. How soluble proteins and peptides such as toxins, colicins, and ionophores move into the membrane milieu to accomplish their functions is fascinating.

Toxins Some protein toxins that come from outside the cell and affect cytosol ic targets provide their own mechanism of translocation across the membrane. Decades of research on diphtheria toxin (DT) established the AB model, in which part A of the toxin carries out its catalytic attack on an intracellular target and part B

4.20. High-resolution structure of diphtheria toxin. Diphtheria toxin has three domains: the C (catalytic), R (receptor-binding), and T (translocation) domains. The active site cleft of the C domain contains the endogenous dinucleotide ApUp. Two helices of the T domain insert into the membrane as a helical hairpin. From Collier, R. J., Toxicon. 2001,39:1793-1801. © 2001 by The International Society on Toxinology. Reprinted with permission from Elsevier.

Proteins and Peptides that Insert into the Membrane

85

PA

ATRJ CMG2

't

/

'\

::~+\

calmodulin)

I

cAMP

EF

--_.~ LF

4.21. Steps in the internalization of anthrax toxin. Anthrax toxin is made up of three proteins, PA (protective antigen), EF (edema factor), and LF (lethal factor). (1) PA binds to a receptor, either ATR or CMG2 (2) Cleavage by a furin protease removes PA20. (3) PA 6 3 self-associates to form the heptameric prepare. (4) Up to three molecules of EF and/or LF bind to the prepore. (5) Endocytosis brings the complex to an acidic intracellular compartment. (6) The low pH triggers conversion of the prepore to a pore, and EF and LF are translocated to the cytosol, where EF catalyzes the formation of cAMP and LF proteolytically inactivates MAPKKs. Redrawn from Collier, J. R., and J. A. T. Young, Annu Rev Cell Dev Bioi. 2003, 19:45-70. © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Cell and Developmental Biology, www.annualreviews.org.

The family of AB toxins has grown to include cholera toxin, pertussis toxin, tetanus and botulinum neurotoxins, and recently, anthrax toxin, although it is structurally more complicated than the others. Anthrax toxin is made up or three different proteins, PA (protective antigen), EF (edema factor), and LF (lethal factor). PA is the antigen for the anthrax vaccine in current use. PA binds to a receptor on the cell surface and then gets cleaved, releasing a 20-kDa fragment. The change in conformation of the remaining 63-kDa PA fragment allows it to form heptamers that can bind up to three molecules of EF and/or LF (Figure 4.2\). The low pH of the endosome triggers a conformational change in PA, enabling each protomer to insert t,vo f3-strands into the membrane, forming a pore that is a 14-stranded 13barrel. The other two proteins, EF and LF, unfold at least partially to translocate through this pore into the cytosol, where they exert their lethal inhibitory actions. The structure of the pore formed by anthrax toxin PA is very similar to that formed by (X-hemolysin ((XHL), a hemolytic toxin secreted by Sraphy/ococcus aureus in a soluble form. A major difference is that (XHL requires no catalytic domain or other inhibitors; it simply lyses human erythrocytes and other cells by pore formation, causing leakage. Another difference is that each 33.4-kDa polypeptide inserts into the membrane

before forming the heptameric prepore at the surface of the bilayer. Once the heptamer forms, each monomer extends a f3-hairpin, forming a f3-barrel pore (see Chapter 5) that is 52 A long and 26 A in diameter, with an inner diameter that is only ~ 15 Ain the narrowest part (Figure 4.22). The rest of the l3-sheet structure fonns a much wider cap, with hydrophobic residues at the rim that contacts the membrane bilayer. This mushroomlike structure was first observed for aerolysin, another l3-barrel channel-forming toxin, which is a virulence factor of Aeromonas bacteria that cause gastrointestinal disease and wound infections. Calicins

Colicins are bacteriocins, a class of toxins synthesized and released by bacteria to kill competing microorganisms. More than a third of E. coli strains produce colicins. These bacteria harbor plasm ids that encode specific colicins, along with specific immunity proteins, which are membrane proteins that ensure their own protection from the lethal action of the colicins. Colicins enter their target cells utilizing outer membrane receptors and either the Tol or Ton intermembrane translocation systems (see Chapter I J on TolC). Those in the channel-forming subfamily then insert into the

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4.22. The a-hemolysin heptameric pore, viewed from the side (A) and the top (8). In (C), one protomer is shown in the open-pore configuration; before insertion, the extended [3-strands of the stem are folded into the rest of the [3-sandwich. From Gouaux, E., j Struct Bioi. 1998, 121: 110-122. © 1998 by Elsevier. Reprinted with permission from Elsevier.

A:coJicin Ja

'l

B:colicin B

C:colicin N

E:colicin A (C-domain)

F:colicin E1 (C-domain)

4.23. Structures of four colicins and the C domains from two others. Pore-forming domains are shown in blue and brown, connecting helices in red, translocation domains in green, and one immunity protein in purple. From Zakharov, S. D., et ai, Biochim Biophys Acta. 2004, 1666:239-249. © 2004 by Elsevier. Reprinted with permission from Elsevier.

I

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S

Proteins and Peptides that Insert into the Membrane cytoplasmic membrane to form voltage-gated channels that leak ions at a very great rate (> 10 6 ions channel-I seC I), depolarizing the membrane and eventually killing the cell. Colicins are proteins of around 60 kDa organized into three functional domains: the N-terminal T domain, which mediates translocation across the cell envelope; the R domain for receptor binding; and the C-terminal domain called C for cytotoxicity, or more specifically, P for pore formation. The x-ray structures of four colicins reveal a central helix or pair of helices usually connecting two structural domains in a dumbbell fashion and giving a very elongated shape to the molecules (Figure 4.23). The lengths o( the coiled coils of colicins 1a and E3 are 160 'A and 100 'A, respectively! In most of the channel-forming colicins, the C domain is a 10-helix globule, which characteristically contains a pair of hydrophobic helices and is very similar to the T domain of DT (compare E and F in Figure 4.23 to T in Figure 4.20). Although the average length of the C domain exhelices is ~ 13 amino acids - clearly not enough to span the bilayer - isolated C domains from many different channel-forming colicins have been shown to form voltage-gated pores in planar bilayers. Experiments with biotin-labeled single cysteine mu tants of the C domains of both colicin Ia and colicin A indicate that a large portion of the peptide chain (115 and 70 amino acids, respectively) moves to the opposite side of the membrane duling insertion and voltage-gated channel opening. Therefore, the sequence of events in channel formation is postulated to be binding to the membrane, unfolding to an extended flexible helical network in the interfacial layer of the membrane, helix elongation, and then insertion (Figure 4.24). It is possible that lipid curvature is involved in pore forma tion, as suggested for some peptides that insert into membranes.

Peptides Peptides that insert into membranes include many antimicrobial pep tides and peptide toxins - some well characterized, like melittin of bee venom, and others newly identified, such as the peptide toxins of spiders and sea anemones. Numerous antimicrobial peptides are under study not only for their role in the natural immune defense of mammals but also their potential as valuable new therapeutics either by themselves or as transporters, as in the case of the family called 'Trojan pep tides," which can deliver other agents into cells. Other pep tides capable of membrane insertion are the ionophores, named for their affinity for ions, which can be highly specific. While some ionophores simply chelate the ion, surrounding it with a lipid-soluble coat, others insert into the bilayer to make ion channels. Alamethicin and gramicidin are two widely stud-

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4.24. Model of two steps in pore formation of channel-forming colicins. A. After binding the membrane surface, the C domains unfold into a helical network that extends in the ~ 15 A-thick interfacial layer of the lipid headgroups, anchored by insertion of the hydrophobic helical hairpin. B. Insertion is predicted to involve helix elongation as well as toroidal configuration of lipids to make the open pore. In A, the membrane potential is trans-positive or is absent, and in B it is trans-negative. Anionic lipids that interact with basic side chains in the pore lining of the toroidal pore structure are red. From Zakharov, S. D., et aI., Biochim Biophys Acta. 2004, 1666:239-249. © 2004 by Elsevier. Reprinted with permission from Elsevier.

ied examples of channel-forming ionophores that have provided many insights for the understanding of protein ion channels (see Chapter 10). In addition, many synthetic peptides have been designed to insert into membrane bilayers. The wealth of data on peptide insertion into model membranes [Tom studies using conductance, Fourier transform infrared (FT1R) spectroscopy and oriented circular dichroism, solid-state NMR, neutron diffraction, and other techniques emphasizes the importance of the lipid composition, temperature, extent of hydration, and the peptide-to-lipid ratio. In general, the peptides can insert in one of two orientations, parallel to the plane of membrane or perpendicular to it, and then permeabilize the membrane by one of four possible mechanisms. In the Carpet mechanism, pep tides bind as ex-helices to the membrane surface and embed in the headgroup region oriented parallel to the bilayer plane. Although they remain in this orientation, at high concentrations they disrupt the membrane integrity

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the plane of the membrane even at high peptide-to-lipid ratios. Alamethicin is a 20-r-esidue peptide with eight helix-stabilizing amino isobutyric acid residues, only one charged amino acid (GluI8), an internal proline (Pro14l. an acetylated N terminus, and an alcohol (phenylaJaninol) at the C terminus. An amphipathic (Xhelix that can bend at Pro14, it forms voltage-gated ion channels of the barrel-stave configuration, as indicated by much experimental evidence. Its single-channel conductance is characterized by multiple discrete states, suggesting the channel is oligomeric and changes its conductance state when a single alamethicin molecule joins or leaves the aggregate. The pore dimensions determined by neutron scattering give it a thickness that matches the helix diameter. Solid-state NMR results indicate that in the nonconductive state, the helices are tilted by 10° to 20 from the bilayer normal, suggesting a possible "preaggregate" state that leads to oligomerization and an open channel. Gramicidin A, which forms channels that are specific for monovalent cations, is composed of 15 nonpolar amino acids of alternating Land D configurations. Gramicidin A can form ~-helices, which are helical structures made up of 13-sheets twisted into cylinders, with hydrogen bonding of the backbone N-H and car~ bonyl groups roughly parallel to the axis of the helix. The ~-helices have hydrophobic exteriors since with alternating L- and D-amino acids, all the side chains are on the outside. The Nand C termini are blocked, so with the lack of polar side chains, the most polar part of the molecule is the peptide backbone; indeed the ion path in gramicidin Ainvolves the carbonyl oxygens. Different gramicidin structures are observed depending on the solvent (lipid, organic solvent, or detergent) and ions present. Detailed structures obtained by x-ray crystallography and NMR have been classified as either a double ~-helical pore that can span the bi layer or a ~-helical dimel~ both of which can have open and closed states (Figure 4.26). Since conditions under which the double helix forms a conducting pore are velY limited, the helical dimer is probably the major conducting form and exemplifies the fourth mechanism for peptide channel formation. There are actually several species of natural gramicidins with slight differences in amino acid composition, as well as numerous synthetic analogs. Because of its self-associating dimer, gramicidin A has been found to be a suitable nanodevice for membrane biochips. 0

4.25. Two mechanisms for pore formation by inserted peptides: the barrel-stave model (A) and the toroidal model (B). See text for details. Redrawn from Yang, L., et al., Biophys J. 2001,81 :14751485. © 2001 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

(without forming pores). In two of the insertion mechanisms, the peptides bind in parallel orientation but oligomerize when a critical concentration is reached, changing their orientation to approximately perpendicular to the bilayer and resulting in pore formation. There are two mechanisms of oligomeric pore formation (Figure 4.25). In the barrel-stave model, the peptides span the two leaflets of the bilayer to line the pore like the staves of a barrel. In the other mechanism, the peptides form a toroidal pore when the insertion of helices stimulates the lipid monolayer to bend back on itself like the inside of a torus (a mathematical term for a surface containing a hole). In contrast to the barrelstave model, the toroidal pore has a continuous bending of a PL monolayer, stabilized by the peptides. In the fourth mechanism, the peptides insert to span the two leaflets of the membrane and then form an open pore when they align as dimer-s. Melillin is an amphipathic ex-helix of 26 amino acids, with five basic residues along its polar side and a bend of ~I20° at its internal proline. Because of its large pore size (4.2 nm inner diameter and 7.7 nm outer diameter), the hydration and temperature dependence of pore formation, and the lack of discrete conductance steps, melittin is considered to form a toroidal hole. In contrast, a solid-state NMR study of the antimicrobial peptide ovispirin supports the Carpet mechanism for its membrane disruption by showing it remains parallel to

SecA: Protein Acrobatics

Perhaps the best example of a protein that is so versatile it can move readily between the cytosol, the membrane surface, and the membrane interior is SecA, the motor for the translocon that exports proteins across the

Proteins and Peptides that Insert into the Membrane

89

A. Channel (helical dimer)

Closed

Open

B. Pore (double helix)

.... Closed

Open

4.26. Channel and pore structures of gramicidin A. A. Gramicidin A forms a helical dimer when two f)-helices form a channel by associating end to end, as diagrammed schematically on the left. The hydrogen bonds are intramolecular except at the interaction between the two N-terminal groups. On the right is the high-resolution structure of the open helical dimer in SDS as determined by NMR. The polypeptides are portrayed with ball-and-stick models (one molecule is white and the other yellow), and the interior lumen of the channel is indicated by overlapping green spheres. B. In the pore form, gramicidin A forms a double helix with intermolecular hydrogen bonding along two antiparallel f)-strands, which opens in response to environmental factors (such as cations), as diagrammed on the left. The high-resolution structure of the cesium-containing open-pore double helix shown to the right was determined by x-ray crystallography. From Wallace, B. A., J Struct BioI. 1998, 121: 123-141. © 1998 by Elsevier. Reprinted with permission from Elsevier.

bacterial plasma membrane (described in Chapters 7 and 12), SecA is a remarkably flexible protein composed of an N-terminal ATPase domain and a C-terminal domain that binds membrane lipids as well as the preprotein to be exported and the cytoplasmic chaperone SecB (Figure 4.27). SecA is quite concentrated in the cytoplasm (5 ~M inside E. coli). Its membrane bind-

ing is dependent on the presence of anionic PLs, except at high temperatures, and is enhanced by nonlamellar lipids. It binds the SecYEG translocon with high affinity (20-40 nM KJ) and inserts repeatedly into the translocon in cycles that couple ATP hydrolysis to movements of the exported polypeptide chain (see Chapter 7). The resulting insertion is so extensive that several regions of

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4.27. The domain structure of SecA. The major SecA domains are the ATPase motor, the C-domain that binds lipids and SecB, and the substrate specificity domain (SSD) that binds to the preprotein. Three views are shown of the space-filling model of the x-ray structure of SecA from B. subtilis. While SecA is shown here as a monomer, two different x-ray structures show SecA dimers of quite different organization. From Vrontou, E., and A. Economou, Biochim Biophys Acta. 2004, 1694:67-80. © 2004 by Elsevier. Reprinted with permission from Elsevier.

the SecA protein can be chemically modified from the outside (periplasmic side). SecA can be cross-linked to portions of both SecY and SecE, although not to membrane lipids.

PROTEINS EMBEDDED IN THE MEMBRANE The operational definition of integral (or intrinsic) membrane proteins implies that they are embedded in the membrane, since disruption of the membrane is required to solubilize them. The exceptions are those peripheral proteins that are held in the membrane by two or more lipid anchors, binding them to the membrane with enough strength to require its disruption for their release (see above). Embedded membrane proteins include monotopic proteins, which insert into the membrane but do not span it, and proteins with one or more TM segments. Monotopic Proteins

There are only a few well-characterized examples of monotopic proteins. Some enzymes involved in lipid metabolism access their substrates by integrating into one leanet of the membrane. Structures have been solved for three of these: prostaglandin H 2 synthase (described in Chapter 9), squalene-hopene cyclase, and fatty acid amide hydrolase, which are all important pharmaceutical drug targets. A high-resolution structure is available for another monotopic protein with clinical importance: monoamine oxidase, which binds to the outer membrane of mitochondria. It is important for inactivation of several neurotransmitters, such as serotonin and dopamine, as well as catabolism of monoamines ingested in foods.

Another example of monotopic proteins is the caveolins, which associate with rafts to form caveolae (see Chapter 2). Caveolin is inserted into the plasma membrane from vesicles derived from the Golgi and remains in the inner leatlet, strongly immobilized by associations with the cytoskeleton. It forms dimers that bind cholesterol and form a striated coat as the membrane invaginates for endocytosis (see Figure 2.26). Integral Membrane Proteins

Most integral membrane proteins have one or more TM segments. Bitopic proteins span the bilayer one time and are classified by their topology as type I, \vith their N terminus outside, or type II, with their C terminus outside. Poly topic proteins are called type III integral membrane proteins and have multiple spans connected by loops. When several bitopic integral membrane proteins oligomerize with interacting TM segments, they are called type IV (Figure 4.28). Chapters 9, J 0, and J J present examples of detailed structures of integral membrane proteins. Up-todate lists of the membrane proteins with structures solved at or near atom ic levels can be found at http:// blanco. biomol. uci .ed u/Mem brane_Protei ns_xtal. h tml and http://wwwmpibp-frankfurt.mpg.de/m ichel/pu bl ic/ memprotstruct.html. The possible folds of integral mem brane proteins are dictated by the process of export to and assembly in the bilayer as well as by the stability of the embedded protein. These constraints on their folds may explain why all of the type III integral membrane proteins whose detailed structures have been solved are of two structural types, bundles of ex-helices and ~-barrels, described in detail in the next chapter. Since most of these structures were solved by x-ray crystallography, it is also possible that crystallization

Proteins Embedded in the Membrane

91

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TABLE 4.1. Properties of the cytosolic and membrane

environments that affect proteins

Bitopic

Polytopic TypenI

Oligomeric Type IV

Inside

Outside

4.28. Classification of integral membrane proteins by topology. Both type I and type II are bitopic proteins with only one TM helix. Type I has the N terminus outside while type II has it inside. Type III proteins have multiple TM segments in a single polypeptide. In contrast, type IV proteins are oligomers assembled from several polypeptides, each having one TM helix. From Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 374. © 2005 by W. H. Freeman and Company. Used with permission.

methods favor these two structural types. With known atomic structures for fewer than 10% of all predicted integral membrane proteins, a more varied menu of structures, such as the I)-helix, will likely be revealed in [l.lture work. As many of the following chapters will illustrate, the TM ex-helix is currently the focal point for most researchers dedicated to understanding the nature of these intriguing proteins. The structures of membrane-spanning proteins must cope with many chemical and physical differences from soluble proteins, as indicated in a list of important environmental factors (Table 4.1). Notable among these are the lack of homogeneity and isotropy; the presence in most membranes of gradients of pH, electric field, pressure, dielectric constant, and redox potential; the paucity of solvent; and the very low dielectric constant. Some properties vary as a function of the depth in the membrane (see Chapter 8). Together these factors significantly alter the ~GO of functions associated with folding processes. Overall, it is much harder to break a main-chain hydrogen bond, ionize a side chain, or break a salt bridge of a protein domain in the membrane interior than in the cytosol; of course it is easier to expose a hydrophobic group, as well as to bring subunits in close proximity.

Property

Cytosol

Plasma membrane'

Solvent chemical homogeneity Chemical groups available

Yes

No

HOH, ions, -SH

-CH3, -CHz-, = CHNo Yes b ~2 x 106b Yes Yes Yes b

~Yes Isotropy pH gradient No ~O Electric field (V.m- 1 ) Pressure gradient No No Dielectric constant gradient Redox potential gradient No ~17 Volume or surface occupancy [protein/solvent (%)JC Separation between two proteins: Distance (A) ~50 Intervening solvent ~ 15-20 molecules ~1O-11 Exchange time between solvent molecules (s)d Viscosity at 20°C 0.001 (Tl; N.s.m -2) Dimensions 3 Translational diffusion: e ~lO-'O Dla, (m 2 .s -') ~250 Average range explored in 1 ~lS (x; A) Dielectric constant (c) 80 t.Go (kcal.mole- 1 ) for: ~O Breaking a main-chain H-bond Deprotonating a Glu side -4 chain (pH7) Opening a salt bridge <1 Exposing one A2 of +0.025 hydrophobic surface Exposing a Leu side chain +2.8 to the solvent Associating two 50-kDa 8 proteins (T t. S at 20°C)

~35

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a For properties that vary as a function of the depth in the membrane, the data correspond to those at the membrane center. b In most but not all membranes. C Estimated from data for the cytosol and plasma membrane of an E. coli cell. Calculations for the cytosol assume a 1:2.5 w/w ratio of RNA to protein; calculations for the plasma membrane assume the average integral protein (ohen an oligomer) to comprise ~ 12 TM helices and to have about half of its volume buried into the membrane. Estimates published in the literature vary from 17% to 50%. d In pure solvent. e For a middle-sized protein (~50 kDa) in either pure water or pure lipids; in the cytosol and in real membrances, diffusion coefficients vary with the distance range considered. Source: Popot, J. L., and D. M. Engelman, Annu Rev Biochem. 2000, 69:881-922. © 2000 by Annual Reviews. Reprinted with permission from the Annual Review a Biochemistry, www.annualreviews.org.

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The membrane milieu strongly favors the formation of secondary structure in TM segments. The low dielectric constant of the nonpolar domain of the membrane and the scarcity of water molecules favor formation of hydrogen-bonded secondary structure. This is readily shown by computing the change in free energy of transfer (~Glr) for peptide bonds with and without H-bonds:

Non-H-bonded-NH-C=O H-bonded-NH-C=O

~Glt- from water to alkane +6.4 kcal/mol +2.1 kcallmol

Therefore the per-residue cost of disrupting Hbonds in the membrane is ~4 kcaJlmol. For a TM segment of 20 amino acids, this is 80 kcaJlmol driving the formation of an (X-helix! Analysis of the solved integral membrane protein structures and mutagenesis of particular residues in them have led to some generalizations about the locations of amino acids in the nonpolar membrane domain: I. Nonpolar amino acids are typically found in membrane-spanning (X-helices with their side chains pointing into the hydrophobic interiOl- of the bilayer. This is expected from thermodynamic arguments and was tested when the small bitopic protein phospholamban from sarcoplasmic reticulum was engineered to replace nonpolar residues in its TM helix with polar ones, resulting in a water-soluble analog. 2. Acidic and basic amino acids either (i) remain uncharged due to the effect of the low dielectric environment on their pKas, (ii) form ion pairs that neutralize their charges, or (iii) playa special role, for example, in transport of protons or electrons or in binding a cofactor such as heme or retinal. Polar residues are not completely excluded from the nonpolar region, since the partitioning of many hydrophobic side chains into the interior is so favorable it can overcome the cost of including a few less-favorable groups. In addition, the charged residues found in TM segments can move their polar groups toward the interface by snorkeling, adopting configurations that orient their polar atoms to partially escape from the hydrophobic membrane core toward the interface. Snorkeling can be quantified by measuring the displacement of the polar atom(s) from the f)-carbon of the amino acid, which shows that the largest snorkeling distances are achieved by lysine residues. However, side chains of Arg, Tyr, Asp, Glu, Asn, and GIn also snorkel, in addition to Trp residues at the in terfaces. 3. Hydrogen-bond formers often use H-bonds to link their side chains to backbone carbonyl groups. These can provide "caps" for the ends of helices, as well as stabilizing the interactions between helices in a heli-

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cal bundle or oligomer. Even the hydrogens on the main-chain (X-carbons can form hydrogen bonds with backbone or side chain oxygen atoms. Several of these bonds, each wi th about half the strength of a conventional hydrogen bond, can stabilize a helix in the nonpolar bilayer interior and are often found in glycine-, alanine-, serine-, and threonine-rich packing interfaces. 4. Of the amino acids considered to be helix breakers, glycine and proline are found frequently in TM helices, often at conserved positions. Glycine ,-esidues are important for allowing the close packing that minimizes interhelical distances in bundles of (X-helices (see below). While proline residues are rare in (X-helices of soluble proteins, they are common in TM (X-helices and are often located near the center of the bilayer. The restricted backbone angles of the peptide chain at proline causes a kink in the helix. Inclusion of proline also leaves one carbonyl of the helix without an intrachain H-bond. The kink is a bend in the chain of ~120' in the direction away from the missing backbone Hbond. Interestingly, mutagenesis of bacteriorhodopsin showed that substitution of alanine for proline does not remove the kink, indicating that the tertiary structure of the integral membrane proteins maintains the helix distortion. The conservation of prolines at kinks in many integral membrane proteins led to the suggestion that the TM segments with kinks that do not have prolines evolved from TM segments with proline, since homologous proteins do have proline in the position of the kinks. 5. Aromatic amino acids, especiallyTrp and Tyr, play a special role at the interface of the hydrophilic and nonpolar domains in both (X-helical and f)-baITel integral membrane proteins. NMR studies with model indole compounds reveal that this is not due to their dipole moment or H-bonding ability but rather to their flat rigid ring and aromaticity that lead to complex electrostatic interactions with the hydrocarbon core. The well-characterized integral membrane proteins that are bundles of (X-helices typically have an even number of helices, with the notable exception of the family of seven-helix bundles that are involved in signal transduction, such as bacteriorhodopsin (see Chapters 5 and 6). Analysis of predicted membrane proteins from 26 genomes (see Chapter 6) shows the number of predicted TM helices is distributed over all integers from two to 13, with the occurrence decreasing as the number increases except for spikes at foul~ seven, and 12. When all the inner membrane proteins of E. coli are similarly analyzed, by far the highest incidence is for bundles of 12 predicted helices, with the next-largest groups havi ng two and six predicted TM helices and significant numbers with four, five, and 10 predicted TM helices.

Proteins Embedded in the Membrane

93

c.

D.

4.29. Distortions of a-helices in TM segments of integral membrane proteins. A. n-Bulge at Ala215 in helix G of bacteriorhodopsin, which causes the peptide plane to tilt away from the helix axis locally. B. Unwinding of helix M4 in the calcium ATPase of the sarcoplasmic reticulum exposes backbone carbonyl groups that participate in coordinating Ca 2+. C. Proline kink in helix C of bacteriorhodopsin, resulting in a lack of hydrogen bond to the carbonyl of Leu87. D. Half-helices in the glycerol facilitator, numbered 3 and 7. From Ubarretxena-Belandia, I., and D. M. Engelman, Curr Opin Struct Bioi. 2001, 11 :370-376. © 2001 by Elsevier. Reprinted with permission from Elsevier.

Distortions from the classical ex-helix are not uncommon in the TM segments seen in the x-ray structures (Figure 4.29). Many of the distortions probably have a functional role; alternatively they may serve to facilitate folding by preventing off-pathway intermediates (see Chapter 7). The proline-induced kinks described above are one class of distortions seen often in TM helices. Anothertype is a 7f bulge, where one backbone carbonyl is not H-bonded, such as a site involved in retinal binding in bacteriorhodopsin (see Chapter 5). Helix unwinding is a third kind of distortion, seen in the Ca 2 + pump from sarcoplasmic reticula (see Chapter 10), vvhere the unwinding frees up carbonyls to coordinate Ca 2+. Hal f-helices, where two short helices that do not individually span the bilayer stack end-toend to span the bilayer, have been observed, for example, in the glycerol facilitator and the aquaporins (see Chapter 10). A number of approaches have been taken to study helix-helix interactions. In model peptides that form TM helices, inserting glutamine in the middle of the TM sequence drives formation of oligomers. Indeed, any amino acid capable of acting simultaneously as both donor and acceptor of hydrogen bonds (Asp, Glu, Asn, and His) promotes oligomerization, while serine, threo-

nine, or tyrosine does nol.ln type III integral membrane proteins, H-bonds play an important role in the tertiary structure. For example, there is at least one hydrogen bond between each pair of helices in bacteriorhodopsin (see Chapter 5). Glycophorin A, the primary sialoglycoprotein of human erythrocyte membranes, has a single TM helix with a critical Gxx.x.G amino acid sequence that is needed for the helix-helix interaction of dimer formation (Figure 4.30). Finding this sequence in numerous other TM peptides has defined a TM-oligomerization motif of GxxxG, along with the less common Gxx.'\.A, which clearly reflects the importance of small amino acids at positions buried between the helices. In glycophorin A the critical sequence is LlxxGVxxGVxxT. Two helices cross at an angle of 40°, making a righthanded coiled coil in which the helices mesh closely by "knob-into-hole" interactions. The knobs are formed by isoleucine and valine residues and the holes by glycine residues. These interactions bring the helices close enough for important van del' Waals interactions along the coiled coil (see Figures 4.30 and 7.5). Even though its [unction is unknown, the detailed analysis of glycophorin A by saturation mutagenesis, NMR, and computational approaches provides a good model for

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type IV proteins as well as for type III integral membrane proteins with multiple TM helices.

PROTEIN-LIPID INTERACTIONS

The properties of membrane proteins are best understood in the context of their lipid surroundings. Indeed, the contacts between integral membrane proteins and lipids must be very tight to maintain the seal of tbe membrane as a permeability barrier. The presence of a protein has essentially no effect on distant lipids but has a large effect on the shell or ring (annulus) of lipids that surround it, forming the interface between it and the rest of the membrane. These lipids are called annular or boundary lipids and can be distinguished experimentally from the bulk lipids of the bilayer. In addition to bulk lipids and annular lipids, there is a third class of lipids comprising those which are tightly bound in crevices or between subunits of the proteins. These lipids are called nonannular lipids (not to be confused with bulk lipids) or lipid cofactors, as they are frequently required for activity. Experiments with purified proteins have shown that many membrane proteins require specific lipids or classes of lipids to stably bind or insert into bilayers, while numerous enzymes require specific lipids for activity (Table 4.2). The activity of Ca2+ -ATPase, for example, increases as lipid is added up

to 30 moles of I ipid per mole of ATPase. Both the nature of interactions with annular lipids and the influences of the physical state of the lipid bilayer on the functions of membrane proteins have been extensively studied using EPR, fluorescence quenching and energy transfel~ and molecular dynamics simulations. Additional information can be gleaned from those lipids detected in highresolution structures obtained by x-ray crystallography described in Chapter 8. EPR is especially suited for studying boundary lipids because it can readily detect two populations of membrane lipids (Figure 4.31; Box 4.2). Since the mobility of the acyl chains is greatest near the center of the membrane (see Figure 2.11), inCOllJOration of a nitroxyl spin label close to the terminal methyl of the chain gives an EPR spectrum with quite narrow line widths in a pure lipid bilayel~ The rotation around the C-C bonds (trans-gauche isomerism, see Chapter 2) is fast (~1O-'O seconds) and averages out, so the spectrum results [Tom the axial rotation of the lipid molecule as a whole (10- 8 -10- 9 seconds). With proteins present, the axial rotation of a spin-labeled lipid molecule in the annular layer is hindered. Because it is relatively immobilized, it produces broader line widths, resulting in a second component most easily seen in the "outer wings" of the EPR spectrum. The selectivity of a protein for annular lipids can be determined From the relative intensity of the peaks in the outer wings, as obse,'ved for

Protein-Lipid Interactions

95

TABLE 4.2. Specific lipid requirements of membrane proteins and enzymes assessed by various techniques A. Lipid specificity for reactivation of delipidated enzymes Enzyme

Source

Cytochrome-c oxidase

Bovine heart mitochondria

j3-Hydroxybutyrate dehydrogenase

Bovine heart mitochondria

Sarcoplasmic reticulum Ca z+ -ATPase Monoamine oxidase

Delipidation by

Reactivation by

Reference

Cardiolipin, not PE, not PC

Sedlak and Robinson, Biochemistry 1999, 3814966

Phospholipase A

Only PCs

Sandermann et al., J Bioi Chem 1986, 261 :6201

Rabbit skeletal muscle

Cholate extraction

Phosphatidylinositol-4phosphate

Starling et aI., J Bioi Chem 1995,270:14467

Rat brain mitochondria

PLAz

PI, negatively charged PLs

J Bioi Chem 1981,

Huang and Faulkner, 256:9211

B. Lipid specificity in reconstitution of membrane proteins Protein

Source

Reconstituted in

Specific requirements

Reference

Acetylcholine receptor

Torpedo californica

DOPC vesicles

cholesterol, PA

Fong and McNamee, Biochemistry 1986, 25:830

Rhodopsin

Bovine retinal rod

Egg PC or DOPC/DOPE supported bilayers

PE (favors activated M2 state)

Alves et aI., Biophys J 2005,88:198

C. Lipid requirement of amphitropic proteins for binding and activation Protein

Source

Binding to

Specific requirements

Reference

Protein kinase C

Rat brain

PClPS LUVs

DAG and PS, anionic lipids

Slater et aI., J Bioi Chem 1994,269:4866

MARCKS

Mouse

PC/PS monolayer

Phosphocholinecytidylyltra nsferase

Rat

PC LUVs or SUVs

a series of spin-labeled lipids reconstituted with myelin proteolipid protein (Figure 4.32). The selectivity for differenl lipids is a reOection of different exchange rates, since there is constant exchange of lipids between the bulk and the annular layer (Figure 4.33). For the exchange equilibrium, LNP + L* ++ L N-! L'P + L, the lipid association constant, K" is ([L'P] [L])/([LP] [L'J). (Typically the concentration of the spin-labeled lipid, [L'"J, is less than 1 mole % of [L].) The reference lipid in these studies is PC, the most abundant PL in most animal cell membranes; thus the K, for PC in this example is 1.0. When there is no selectivity, K, = 1; with fairly high selectivity, K, approaches 10. The restriction of the bilayer to two dimensions produces a high effective concentration of lipids that can further enhance the selectivity. Since K, gives an average of affinities of a particular lipid, which may be due to several sites of quite different binding

Wang et aI., J Bioi Chem 2001,276:5013 Anionic lipids, unsaturated PE, DAG

Arnold and Cornell, Biochemistry 1996, 35:9917; Davies et aI., Biochemistry 2001, 40:10522

affinities, it may mask the presence of a tightly binding site for the lipid. A comparison of the lipid selectivity of different proteins shows that other proteins do not have the large variation in K r seen with myelin proteolipid protein and some, like rhodopsin, do not discriminate at all (Figure 4.34). Thus in general, the composition of lipids in the boundary layer appears to be quite similar to the composition in the bulk lipid bilayer. The exchange into the annulus (on-rate) is diffusion limited (~108 sec!) and thus is the same for different lipids, while the off-rate reflects the specificity of interaction with the protein and can be slowed to 10 7 or even 10 6 sec l . By performing experiments at different lipid-to-protein ratios, both K, and the fraction of spin-labeled lipid associated with protein can be determined. The stoichiometry of annular lipids for a number of different proteins has been found to correlate well

noterns In or at me tlilayer

96

Time scale 10- 9 sec

10- 8 sec

10- 7 sec 10- 6 sec Fluid lipids

•

•

Restricted lipids

-----==-==-- - - -

--:;;;.---------1\ l\

ij /.

_,-h

II

4.31. EPR detection of two populations of lipids: bulk and annular. A. Diagram of components of the membrane with time scales for the rotational mobility of each. The mobility of the annular lipid is about 1OO-fold slower than that of the bulk lipid. B. EPR spectrum of a lipid spin labeled on (14 (see Box 4.2). The solid line is the spectrum arising from contributions of two components, annular and bulk lipids, shown by dashed lines. The spectral ranges of the two components are identified over the spectrum, with the black indicating the more fluid (bulk) lipid and the gray indicating the restricted (annular) lipid. From Marsh, D, and L. I. Horvath, Biochim Biophys Acta. 1998. 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

with the shape of the protein. Given a helix diameter of ~I nm and a lipid diameter of ~0.5 nm, a bitopic protein with one TM helix has 10 lipids in the shell around the helix. For poly topic proteins, the number of annular lipids is proportional to the number of TM helices and dependent on whether the geometric arrangement of the helices is sandwiches or polygons (Figure 4.35). An example is the Ca 2+ -ATPase with an estimated circumference of 14 nm, which was calculated to require a lipid shell of ~30 lipid molecules and was found by EPR

measurements to have 32 annular lipids. Similar calculations may be done for oligomeric proteins by treating the geometry of the arrangement of subunits instead of the TM helices. In integral membrane proteins that are /3-barrels, the TM segments are more extended and thinner, with diameters of ~0.5 nm. The predicted number of annular lipids works out to be the same as the number of /3strands if there is no tilt as they cross the bilayer; if there is a 60 tilt, there are about twice as many annular 0

Protein-Lipid Interactions

97

BOX 4.2. Electron paramagnetic resonance Electron paramagnetic resonance (EPR), also called electron spin resonance (ESRl, detects the orientation of unpaired electrons on paramagnetic molecules placed in a strong magnetic field. EPR can be used for biochemical substances containing paramagnetic transition metals or free radicals. To broaden its applicability, researchers employ free radical probes that are spin-labeled analogs of the molecules of A. interest, such as DPPC with a nitroxyl group. Such a compound, phosphatidylcholine carrying a spin probe on carbon 14 of the acyl chain (14PCSL), is shown in part A of Figure 4.2.1 o The absorption spectrum created by irradiation of the sample in the magnetic 4PCSL field is displayed as a first derivative, characterized by its intensity, line width, g6PCSL value (for positions), and multiplet structure. The nitroxide group gives three lines, 8PCSL whose line widths (related to the spin relaxJOPCSL ation times, T1 and T2) indicate the mobility of the molecule carrying the unpaired 12PCSL electron, which can vary from freely tumbling to strongly immobilized. In a series 14PCSL of PCSL probes, the position of the probe on the acyl chain determines its mobility. Placing the spin probe near the terminal methyl group gives relatively narrow lines due to the angular fluctuations from rotations around C-C bonds all along the chain, as shown in part A. The sharp lines that result are typical of isotropic mobilDescription of Approx rotational B. ity. Of course in a lipid bilayer, the spin spectra tumbling times (ns) probes do not rotate isotropically, but their large fluctuations average the orientational Freely tumbling O.J anisotropy when they are labeled near the end of the acyl chains. Cooling the lipid sample will slow the movement of the spin Weakly probe, broadening the lines, until it even0.6 immobilized tually produces a rigid glass or powder, as shown in part B of Figure 4.2.1. When the spin label is not free to tumble in all directions, it has anisotropic motion. The Moderately 2.5 frequent trans-gauche isomerizations along immobilized the acyl chains give motional averaging, while the axial rotation of the lipid is slower and is characterized by the rotational correlation times TRII and TR.L as shown in 5.0 part A. In addition to its application in studies of lipid-protein interactions described in this chapter, EPR is also used to probe protein Strongly -300 conformations with a procedure called siteimmobilized directed spin labeling (SDSL). The first step in these studies is the use of site-directed Rigid glass mutagenesis to create a single reactive site, -100°C >300 or powder typically by replacing an individual residue in I a protein with cysteine. Then reaction with a sulfhydryl-reactive spin label positions a 4.2.1. A. Dependence of the EPR spectra of nitroxide-Iabeled spin-labeled side chain at that site to proDPPC on the location of the spin probe. B. Effect of temperature on vide information about structure, orientathe mobility of a spin-labeled Pc. Both redrawn from Campbell, I. tion, and conformational changes in memD., and R. A. Dwek, Biofogica/Spectroscopy, Benjamin Cummings, 1984, pp. 197 and 192. © 1984 by lain D. Campbell and Raymond brane proteins. A. Dwek. Reprinted with permission from the authors.

- ~V-O:-;:T

rrOH~InS

98

14-SASL

14-PASL

14-PSSL

14-PGSL

14-PCSL

In or dl Ule Dlldyer

EPR experiments have addressed the detailed nature of the selectivity for annular lipids by comparing phospholipids with different headgroups and varying the ionic strength and pH, as well as by examining the importance of the glycerol backbone and the length of the acyl chains. In general, most proteins are found to prefer negatively charged lipids. Tn some cases, this is simply an electrostatic effect that is overcome by high ionic strengths, and in other cases the selectivity for the headgroup holds even in high ionic strength. (The few examples of headgroups resolved in highresolution structures reveal that extensive electrostatic and hydrogen-bonding interactions stabilize them in binding pockets, described in Chapter 8.) There is little or no difference when sphingomyelin and gangliosides are compared to PC, indicating the glycerol backbone is not a factor in selectivity. On the other hand, the acyl chain length is important: the free energy of association shows a linear dependence on chain length from 13 to 17 carbons. Hydrophobic Mismatch

4.32. EPR spectra of different lipids reconstituted with myelin proteolipid protein in DMPC. The protein/DMPC ratio is 23:1 and the temperature is 30°C. All the lipids contain a spin label on C14. They are stearic acid (14-SASLl, phosphatidic acid (14PASLj, phosphatidylserine (14-PSSU, phosphatidylglycerol (14PGSL), and phosphatidylcholine (14-PCSLl, where SL stands for the spin label in each case. The increasing relative intensity of the outer peaks arising from motion ally restricted lipids indicates increasing selectivity for the protein. Redrawn from Marsh, D., and L. I. Horvath, Biochim Biophys Acta. 1998, 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

lipids as the number of I)-strands. The exchange rate was found to be slo\ver for annular lipids of I)-strands, presumably because lipids aligned along the I)-strands are more extended and less flexible than annular lipids around (X-helices. This result was determined with M13 coat protein, whose TM domains can be either (X-helices or I)-strands, and the exchange mte was four to five times slower for lipids associating with M 13 coat in 1)strand conformation than those associating with M13 coat in (X-helix.

© tt

~

4.33. Lipid exchange between bulk and annular lipids. Two lipids exchange at one "site" on the surface of a membrane protein, as L leaves and L' takes its place. Redrawn from Lee, A. G., Biochim BiophysActa. 2003, 1612:1-40.

The importance of acyl chain length on lipid-protein interactions produced the concept of hydrophobic mismatch, which results when the nonpolar region of the bilayer is thinner or thicker than the hydrophobic thickness l-equired by an integral membrane protein. The thickness of a bilayer is strongly influenced by its lipid composition: for example, a PC bilayer with saturated chains is 2.5 A wider than a bilayer with unsaturated chains of the same number of carbon atoms (see Chapter 2). If the hydrophobic regions of the protein and lipid do not match, either the lipid bilayer must stretch or compress to match the hydrophobic thickness of the protein (Figure 4.36), or' the protein must change by tilting helices or rotating side chains to fit to the bilayer to avoid exposing nonpolar groups to the aqueous environment. Since proteins are more rigid than lipids, the bilayer might be expected to deform to accommodate the dimensions of their TM segments, contributing to the lateral tension of the bilayer. This is observed when the thickness of lipid bilayers changes to accommodate gramicidin channels: insertion of gramicidin causes a DMPC bilayer to become 2.6 A thinner and a DLPC bilayer to thicken by 1.3 A. The perturbation of the membrane due to the mismatch creates a tension that contributes to its free energy: the t.GC' for bilayer deformation has been calculated to be ~ 1.2 kcallmol for a large hydrophobic mismatch of loA. While lipid bilayers adjust to accommodate gramicidin, they do not similarly accommodate single TM helices. Synthetic peptides designed to be TM helices of different lengths have no effect on the thickness of model bilayers. Rather, NMR measurements revealed

Protein-Lipid Interactions

99

PA

/' /' /'

/'

/'

/'

/' /'

/'

/'

/'

/'

/'

/'

/' /'

Kr

4.34. Patterns of lipid selectivity of different proteins. Kr , the relative association constant between each protein and each lipid, varies from 1 (shown in light gray) to > 6 (shown in dark gray). The data show the Kr for each protein {listed along the right edge}, with each of the lipids identified at the top. Lipid selectivity increases from front (with rhodopsin exhibiting almost no lipid selectivity) to the back (highest selectivity with PLP, the myelin proteolipid protein). Redrawn from Marsh, D., and L. I. Horvath, Biochim Biophys Acta. 1998, 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

that TM helical peptides tilt with respect to the bilayer normal to match the hydrophobic thickness of the lipids. The peptides have sufficient flexibility of orientation to accommodate to the bilayer and not deform it. These findings suggest that proteins that cross the membrane with a small number of lX-helices are likely to accommodate the bilayer thickness by helix tilting. However, larger proteins or proteins with less flexibility impact the lipid enough for hydrophobic mismatch to induce changes in bilayer thickness. The latter includes proteins that cross the bilayer as l3-barrels (see Chapter 5), which have structural constraints thai prevent them from adapting to the lipid bilayer and thus are

more likely to select for lipids that provide hydrophobic matching (see Chapter 7). A comparison of relative binding constants for PCs with acyl chains of different lengths indicates that some integral membrane proteins bind more strongly to lipid that requires no change in bilayer thickness than to

A

dp

\,J

B.

Helical sandwich

Polygon

4.35. Geometries considered for determining the lipid-toprotein stoichiometry. Two geometries suffice to describe the stoichiometry of lipid to protein for integral membrane proteins with up to six TM helices. (With seven or more, there may be centrally located helices that do not contact the lipid.) On the left is a helical sandwich and on the right is a regular polygon. From Dc<, the diameter of a TM helix, and dch, the diameter of a lipid chain, the number of lipid molecules in the first shell around the protein (represented by the dotted line) can be calculated. Redrawn from Marsh, D., and L. I. Horvath, Biochim Biophys Acta. 1998, 1376:267-296. © 1998 by Elsevier. Reprinted with permission from Elsevier.

4.36. Distortion of lipid bilayers due to hydrophobic mismatch. Lipids in a bilayer can be distorted to match the hydrophobic thickness of the protein, as viewed from the side (Al and top (B). In both diagrams on the left, the hydrophobic thickness of the protein (d p ) is greater than that of the lipid bilayer {dll, whereas in the diagrams on the right the reverse is the case. When the acyl chains stretch (d p > dl), the surface area they occupy decreases; when they compress (d l > d p ), it increases. Redrawn from Lee, A. G., Biochim Biophys Acta. 2004, 1666:62-87. © 2004 by Elsevier. Reprinted with permission from Elsevier.

IIUlt::lIl;:) III UI

lUU

Cll

1I1~

Ulldy'-=l

T ~

4.37. The consequence of hydrophobic mismatch in biological membranes may be a high-energy state as lipids and proteins try to compensate by extension of acyl chains (E), compression of acyl chains (C), and/or tilting of TM helices (T). Redrawn from Mitra, K., et aI., Proc Natl Acad Sci USA. 2004, 101 :4083-4088. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

lipid that requires such a change. Preference for chain length has been demonstrated with both rhodopsin and the photosynthetic reaction center. When covalently spin-labeled rhodopsin was reconstituted in PC with different-length saturated chains, it was active in DMPC (CJ4), segregated into protein-rich domains in DLPC (CI2), and aggregated in DSPC (CI8). Similarly, \-vhen the incorpora tion of photosynthetic reaction cen tel' into lipid bilayers was monitored by DSC, the T m for DLPC (CI2) increased by goC whereas that for DPPC (CI6) decreased by 3°e. These differences suggested that the protein partitioned into the gel phase with the shorter acyl chains and into the liquid crystalline phase with the longer acyl chains, since the bilayer is thicker in gel phase than in fluid phase. Such findings su pport the idea that hydrophobic mismatch could drive integral membrane proteins to regions of the bilayel- of appropriate thickness, which could be important in raft formation. Hydrophobic mismatch could also be involved in sorting membrane proteins to different membrane compartments. For example, along the secretory pathway that carries proteins to the plasma membrane in eukaryotic cells, some proteins remain in the Golgi apparatus, where they glycosylate secreted proteins. These Golgi-resident proteins have TM domains that are typically five amino acid residues shorter than the TM domains of proteins of the plasma membrane. This length difference was shown to be critical to sorting when constructs were engineered with shorter and longer TM segments. The bilayer thicknesses of the membranes of the secretory pathway have been determined by x-ray scattering. The ER, Golgi, basolateral, and apical plasma membranes from rat hepatocytes were treated with proteases (and puromycin and ribonuclease [RNase] as appropriate to remove ribosomes) prior to measuring their distances from P atom to P atom to determine bilayer thickness. The thickness of these membranes was expected to increase along the pathway, proportional to their cholesterol contenl. While the thickness does increase from the ER to the Golgi to the apical

plasma membrane, the basolateral plasma membrane is significantly thinner than the others. Since proteins targeted to the apical plasma membrane of the rat hepatocyte pass through the basolateral plasma membrane, hydrophobic mismatch must occur along the pathway. It is possible that the strain of hydrophobic mismatch puts the membrane in a high-energy state useful for vital functions such as fusion or protein insertion (Figure 4.37). In addition to protein sorting, hydrophobic mismatch is involved in membrane protein folding (see Chapter 7). The stress induced by mismatch is likely to affect the environment in which integral membrane proteins fold and assemble, which may at least partially account for the need for specific lipids in folding certain proteins. For example, the E. coLi transporter lactose permease (see Chapter 10) requires PE for correct folding but does not require PE for function. Thus it is proposed that the lipids have the role of chaperone in the folding process. With this understanding of how the special environment of integral membrane proteins constrains their structure and how they interact closely and dynamically with their boundary lipids, Chaptet- 5 focuses on the properties of some very well-characterized proteins. The following chapters describe the kinds of functions membrane proteins carry out, the structural principles used to predict their structures, and their folding and biogenesis.

FOR FURTHER READING

Reviews Peripheral Proteins

Gerke, v., C. E. Creutz, and S. E. Moss, Annexins: linking Ca 2+ signaJJing to membrane dynamics. Nal Rev Mol Cell Bioi. 2005, 6:449-461. Heimburg, T., and D. Marsh, Thermodynamics of the interaction of proleins with lipid membranes, in K. Men and B. Roux (eds.), Biological Membra l1es. Cambridge, Mass.: Birkhauser, 1996, pp. 405-462.

For Further Reading Hurley, J. H., and S. Misra, Signaling and subcellular targeting by membrane-binding domains. Annu Rev Biophys Biomol Struct. 2000, 29:49-70. Johnson, J. E., and R. B. Cornell, Amphoteric proteins: regulation by reversible membrane interactions. Biochim Biophys Acta. 1999,16:217-235. Mayor, S., and H. Riezman, Sorting GPI-anchored proteins. Nat Rev Mol Cell BioI. 2004,5:110-119. McLaughlin, S., and A. Aderem. The myristoyl-electrostatic switch: a modulator of r'eversible protein-membrane interactions. Ii-ends Biochem Sci. 1995,20:272-276. Seaton, B. A., and M. F. Roberts, Peripheral membrane proteins, in K. Merz and B. Roux (eds.), Biological Me1'l'lbrClnes. Cambridge, Mass.: Birkhauser, 1996, pp. 355-403.

101

Zakharov, S. D., et aI., On the role of lipid in colicin pore Formation. Biochim Biophys Acta. 2004,1666:239-249. General Features of Integral Membrane Proteins Curran. A. R., and D. M. Engelman, Sequence motifs. polar interactions and conformational changes in helical membrane proteins. CWT Opin Struct Bioi. 2003.13:412-417. Popot. J. L., and D. M. Engelman, Helical membrane protein Folding, stability and evolution. Annu Rev Biochem. 2000, 69:881-922. White, S. H., and G. von Heijne, Transmembrane helices before, during and after insertion. Curl' Opin Struct Bioi. 2005, 15:378-386. White, S. H., et aI., How membranes shape protein structure . .1 Bioi Chern. 2001,276:32395-32398.

Toxins and Colicins Collier, J. R., and J. A. T. Young, Anthrax toxin. Amw Rev Cell Dev Bioi. 2003, 19:45-70. Falnes, P.O., and K. Sandvig, Penetration of protein toxins into cells. Curr Opin Cell Bioi. 2000, 12:407-413. Gouaux, E., ex-Hemolysin from Staphylococcus aureus: an archetype of l3-barrel, channel-forming toxins. J Struct Bioi. 1998, 121: 110-122. Zakharov, S. D., and W. A. Cramer, Colicin crystal structures: pathways and mechanisms for colicin insertion into membranes. Biochim Biophvs Acta. 2002, 1565:333-346. Zakharov, S. D., and W. A. Cramer, Insertion intermediates of pore-forming colicins in membrane two-dimensional space. Biochimie. 2002,84:465-475.

Protein-Lipid Interactions Lee, A. G., How lipids affect the activities of integral membrane proteins. Biochim Biophys Acta. 2004, 1666:62-87. Lee, A. G., Lipid-protein interactions in biological membranes: a structural perspective. Biochim Biophys Acta. 2003,1612:1-40. Marsh, D., and L. 1. Horvath, Structure, dynamics and composition of the lipid-protein interface. Perspectives from spin-labelling. Biochim Biophys Acta. 1998, 1376:267296. Marsh, D., and T. Pali, The protein-lipid interface: perspectives ITom magnetic resonance and crystal structures. Biochim Biophys Acta. 2004, 1666: 118-141.

5

Bundles and Barrels

B. Structures of helical bundle and j3-barrelmembrilne proteins d Her in many respects, seen ,n the nbbon diagrams of the photosynthetic reaclion center from Rb. sphaeroldes (A) and themallopo.inlrimer frol11 E. coli outer membrane (B) A redrawn fro III Jones, M. R., et aI., Biochirn Biophys Acta. 2002. 1565:206-214 ,i 2002 by ElseVier. Reprinted Witt, permiSSIon from ElseVier. B redrawn 2003 by Elsevier. Reprinted wltl permiSSion from Elsevier from Wrmley, W. C, CurT Opin Struci Bioi. 2003, 13:404-411

The thermodynamic arguments discussed in the previous chapter make it clear that the TM segments of proteins will utilize secondary structure to satisfy the hydrogen bond needs of the peptide backbone. While a variety of combinations of secondary structures might be imagined in type ITl membrane proteins, all known protein structures cross the bi layer wi th ei ther ex-hel ices or l3-strands, producing either helical bundles or 13barrels. This chapter looks at how understanding structure and fu nction for a few proteins has provided the paradigms for these two known classes of integral membrane proteins.

x-ray structu re solved for mem brane proteins, that of the photosynthetic reaction center (RC). The majodty of integral membrane proteins whose high-resolution structures have been solved by x-I-ay crystallography exhibit the helical bundle motif (see examples in Chapters 9, 10, and 11). Helix-helix interactions have been analyzed in many of these, providing details of both tertiary and quaternary interactions. Identification of new integral membrane proteins in the proteome relies heavily on prediction of TM helices, as described in Chapter 6. Bacteriorhodopsin

HELICAL BUNDLES

Transmembrane (TM) ex-helices have dominated the picture of membrane proteins, guided by early stn.1Ctural information on bacteriorhodopsin and by the first

If a single protein dominated the thinking about structure, dynamics, and assembly of membrane proteins in the decades following 1970, that protein was bacteriorhodopsin (BR) from the purple membranes of the salt-loving bacterium Halobacler salinarum. From early

electrochemical proton gradient that supports the synthesis of AT? The ability of reconstituted vesicles containing BR and beef heart mitochondrial AT? synthase to synthesize AT? in response to light provided crucial early support for Mitchell's chemiosmotic hypothesis that the energy of an electrochemical gradient across the membrane could be lIsed to do work (Figure 5.2). Like rhodopsin. the light-absorbing protein in the rod outer segments of the eye's retina (see Chapter 9), BR has seven helices labeled A to G that span the membrane, and a retinal that is bound to a lysine residue in helix G via a protonated Schiff base (Figure 5.3). Whereas BR undergoes a light-induced photocycle involving conversion of the retinal from all-trans to J3-cis accompanied by conformational changes in the Light

+.+.• •

Bacteriorhodopsin

•

Cell~

wall

•

+

·.41~-~-~-[f.;-~ +

FJageJla 5.1. Schematic showing the different membrane domains of a halobacterium cell. The patches of purple membrane containing bacteriorhodopsin (BR) are separate from the regions of membrane containing the respiratory chain and the ATP synthase. Protons are pumped out of the cell in response either to light absorption by BR in the purple membrane or to cytosolic substrates for the electron transport chain. The ATP synthase normally uses the uptake of protons to drive the synthesis of ATP, although it can act as an ATPase and eject protons at the expense of ATP. Redrawn from Stoeckenius, w., Sci Am. 1976,234:38-46. © 1976 by Scientific American. Reprinted with permission from Scientific American.

structural images and spectroscopic characterizations, BR became the paradigm for ion transport proteins and indeed for ex·helical TM proteins in general. From the wealth of studies of its structure and function, a truly detailed understanding of this membrane protein has emerged. BR is the only protein species in the discrete membrane domains called purple membranes, the lightsensitive regions of the plasma membranes of H. salinarum (Figure 5. J). Together with specialized lipids, this protein forms functional trimers that pack as ordered two-dimensional arrays on the bacterial cell. In photophosphorylation BR functions to pump protons out of the cell in response to the absorption of light by its chromophore, retinal, converting light energy into an

Lipid vesicle

AD? +

Y

Mitochondrial FIFo-AT? synthase

~TP H

5.2. Schematic of reconstituted vesicles containing BR and ATP synthase. When the light is turned on, ATP is synthesized from ADP + P;. When the light is turned off, ATP synthesis stops. These vesicles gave important evidence to support the chemiosmotic theory of Peter Mitchell. Redrawn from Garrett, R. H., and C. M. Grisham, Biochemistry, 2nd ed., Brooks/Cole, 1999, p. 897. © 1999. Reprinted with permission from Brooks/Cole, a division of Thomson Learning: www.thomsonrights.com. Fax 800 730221S.

Bundles and Barrels

104

C4

Cl8

Cl9

C20

I

I

I

rCS~ /C7~ . . . . . C9.:::::::. /clr~ . . . . .C13..::::::....C6 C8 CIO Cl2 Cl4

I

C.- Lys216

I

C3 CI 'C2/ \~CI7 Cl6

~ hv

CIS

CI9

I C4

I

C20

I

/C5.:::::::.

C6

/C7~

I

. . . . . C9.:::::::.

C8

CIO

. . . . . Cll~

CI2

. . . . .CJ3~

CI4

I

C3 CI ' 0 / \~CI7 CI6

5.3. Retinal bound to lysine 216 in bacteriorhodopsin. The Schiff base linkage (shaded) between the aldehyde of retinal and the Eamino group of lysine 216 is protonated, as shown, before light stimulation. The all-trans retinal converts to 13-cis retinal upon absorption of a photon. Redrawn from Neutze, R., et aI., Biochim BiophysActa. 2002,1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier.

protein that result in proton transfer across the membrane, visual rhodopsin's activation cascade involves an II-cis to all-trans conversion of the retinal, followed by its dissociation from the protein. In addition to BR, H. salinaru111 contains three related rhodopsins, and similar molecules are found in eubacteria and unicellular eu karyotes. A.

B.

The purple membrane is composed of 75% protein and 25% lipid by weight, with 10 halobacteriallipids per BR monomer. These native lipids are based on archeol (see Figure 2.6), so they differ in headgroups but not in length of acyl chains. Delipidation by treatment with a mild detergent affects the kinetics of the BR reaction; addition of halobacterial lipids but not phospholipids restores activity. When BR is crystallized (see below) the bound lipids that are retained from the membrane fit well into the grooves along the protein surface (see Figure 8.9). BR was the first integral membrane protein whose topological organization in the membrane was elucidated. Electron diffraction of the native twodimensional crystalline alTays of purple membrane provided early images of BR trimers, revealing the monomer structure to be seven TM ex-helices arranged in an arc-like double crescent in the plane of the bilayer (Figure 5.4A and B). Models fitting the primary structure of the protein, with 70% of its 248 residues being hydrophobic, to the observed images were aided by the sensitivity of exposed loop residues to partial proteolysis in situ (carried out on BR in the membrane), although the precise beginning and end of each helix were uncertain for years. Such studies also showed that a few N-terminal amino acids are exposed to the exterior and the last 17 to 24 amino acids of the C terminus

c.

B

A

c

o

E

F

Gr I MOAO I

5.4. EM structure of BR. A. The electron density profile of the 2D-crystalline purple membrane shows arrays of BR trimers. Each trimer is arc-shaped with three well-resolved peaks in the inner layer and four less resolved peaks in the outer layer. By Unwin and Henderson. Redrawn from Garrett, R. H., and C. M. Grisham, Biochemistry, 2nd ed., Brooks/Cole, 1999, p. 273. C9 1999. Reprinted with permission from Brooks/Cole, a division of Thomson Learning: www.thomsonrights.com. Fax 800 730-2215. B. Seven helices are modeled to correspond to the seven peaks of a BR monomer. Based on neutron diffraction data, a retinal has been placed in the center of the protein. From Subramaniam, S., and R. Henderson, Biochim Biophys Acta. 2000, 1450:157-165. © 2000 by Elsevier. Reprinted with permission from Elsevier. C. A topology model of BR shows the predicted sequence composition of the seven helices and their connecting loops. The model was adjusted periodically based on genetic mutations of targeted residues (colored boxes) until the x-ray structure was solved. From Khorana, H. G., j Bioi Chem. 1988,263:7439-7442. © 1988 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

G

Helical Bundles

105

= 5 ms

T

BR ~4 ps A l11ax = 570 nm ~

o

K

Amax = 640 nm An/ax = 590 nm \ T

= 5 ms

7

(

N~Amax

=

T

560 nm

f-

1 "'

L

Ama.x = 550 nm

= 5 ms

T

= 40

J1.s

AlIlax = 410 nm

M z _ Mt T

= 350

J1.S

5.5. The photocycle of bacteriorhodopsin. In response to light,

BR undergoes a series of transitions through intermediates K, L, M1, M2, N, and 0, which have different lifetimes (T) and different absorbance maxima as shown. The photocycle is initiated by isomerization of the retinal from all trans to 13-cis (BR --+ K, L), followed by transfer of a proton (L --+ M), the conformational change that switches the accessibility of the Schiff base from the extracellular side to the cytoplasmic side (M1 --+ M2), another proton transfer (M --+ N), and conversion of the retinal back to all trans (N --+ 0). From Neutze, R., et aI., Biochim Biophys Acta. 2002, 1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier.

are accessible in the cytoplasm. The location of retinal in the center of the protein (Figure 5.4B) was determined by neutron diffraction, and the lysine to which it binds was identified by reduction of the Schiff base with NaBH 4 . Once it was clear that the retinal binds to Lys2 J6, nearby residues were investigated by sitedirected mutagenesis, which identified residues that interact with the retinal, such as Asp212 and Arg82, as well as residues crucial to proton pumping, such as Asp85 and Asp96. These results led to a model for the topology of BR that was further modified as genetic studies defined the positions of many residues (Figure SAC). Extensive mutagenesis of the gene for BR provided a large collection of mutant proteins that could be studied in cell suspensions or reconstituted in lipid vesicles, with changes of pH, temperature, and salt conditions used to further characterize the protein function. In addition, a variety of retinal analogs were incorporated to observe their effects on the absorption spectrum and activity. Researchers used a number of pH-sensitive dyes to investigate the stoichiometry of proton pumping. And over many years, increasingly sophisticated instrumentation for visible and ultraviolet absorbance, fluorescence, circular dichroism. Raman, and infTared spectroscopy have been employed to follow the response of BR to light. The primary event when BR absorbs a photon is the isomerization of retinal. This event triggers subse-

quent structural changes and pKa shifts in the protein that allow deprotonation of the Schiff base, vectorial transfer of the proton to the extracellular side of the membrane, and uptake of a proton from the cytosol. These processes are accompanied by differences in the absorbance spectru mol' BR, allowing detection of intermediates with lifetimes varying from a few picoseconds (ps, 10- 12 sec) to a few milliseconds (msec, 10- 3 sec). The light-induced changes in BR are summarized in a photoreaction cycle, or photocycJe (Figure 5.5). BR in its resting state has a )'ma, of 570 nm (purple); when it absorbs a photon, it rapidly isomerizes to the K intermediate ()'max 590 nm) and then converts to the L intermediate (),.max 550 nm). The transition from L to M ()'m", 410 nm) occurs when the proton fTom the Schiff base is transferTed to Asp85, the primary acceptor. At this point a structural rearrangement occurs to switch the accessibili ty of the Schi ff base, described as M I -> M2, which is essential for vectorial proton transport by preventing reprotonation from the extracellular side that

Cytoplasmic

A

Extracellular 5.6. The first high-resolution structure of BR. This overview of the structure shows the seven TM helices labeled A to G, and the residues involved in proton translocation as well as the retinal. From Pebay-Peyroula, E., et al., Biochim Biophys Acta. 2000, 1460: 119-132. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Bundles and Barrels

106

B.

A.

Cytoplasmic side

) Helix G

Q7

ca

~

82

3.29

W408

0

3 • 11

_

2.36

3.25

.J

Extracellular side -../

5.7. Proton path in BR. A. The ribbon diagram for the seven TM helices, labeled A to G, is marked to show proton transfer steps indicated by arrows numbered in chronological order from 1 to 5. Step 1 is release of a proton from the Schiff base to Asp85. In step 2 a proton is released to the extracellular medium, possibly via Glu204 or Glu194. In step 3 the Schiff base is reprotonated by Asp96. Step 4 is the reprotonation of Asp96 from the cytoplasmic medium. Step 6 is the final proton transfer step from Asp85 to the group involved in proton release at the extracellular side, either Glu204 or Glu194. From Neutze, R., et al., Biochim Biophys Acta. 2002, 1565:144-167. © 2002 by Elsevier. Reprinted with permission from Elsevier. B. Details of the proton path on the extracellular side of the Schiff base reveal a network of hydrogen bonds between Asp85, Asp212, Arg82, Glu194, and Glu204 and discrete water molecules (red, labeled W). Interatomic distances are given in angstroms. From Pebay-Peyroula, E., el aI., Biochim Biophys Acta. 2000, 1460: 119-132. © 2000 by Elsevier. Reprinted with permission from Elsevier.

would result in a zero net effect. The next three steps are s!owel', each taking around 5 msec. Transfer of a proton to the Schiff base from Asp96 creates the N intermediate U.max 560 nm). With the return from 13-cis to alltrans retinal, the 0 intel-mediate o..max 640 nm) is formed and the release of a proton from Asp85 completes the cycle. The first high-resolution structure of BR was achieved by x-ray crystallography of microcrystals prepared in bicontinuous cubic phase lipids, either monoolein or monopalmitolein. (These are not phospholipids but rather racemic mixtures of glycerol esterified to one acyl chain, either oleoyl or palmitoleoyl, on Cl.) In cubic phase-grown crystals, BR trimers are stacked in layers that have the same ori-

entation and lipid content as that observed in purple membrane. Furthermore, spectroscopic studies show that BR in cubo undergoes the main steps of the photocycle. As expected from the electron density images, the seven TM hel ices cross the mem brane nearly perpend icular to the plane of the bilayer and are packed closely together and connected by short loops (Figure 5.6). The structure has now been refined to better than 2 A resolution and provides sufficient detail to trace the proton channel, including several important water molecules (Figure 5.7A). The central cavity that contains retinal in Schiff base linkage to Lys21 6 is quite rigid, with the n-bulge in helix G stabilized by an H-bond From Ala215 to a water molecule. In the resting state, the positive charge on the protonated Schiff base

Helical Bundles (pKa ~13.5) is stabilized by the nearby deprotonated carboxylate groups of Asp85 and Asp212. Polar side chains and 'vvater molecules make a clear proton path in the extracellular half of the molecule from Asp85, the proton acceptor, to the extracellular surface where the proton is released (Figure 5.7B). At the cytoplasmic surface are several acidic groups that may be involved in transferring protons from the cytoplasm, but no clear proton path connects the central cavity to them. The pKo of Asp96, the proton donor during reprotonation of the retinal from the cytoplasmic side, is very high due to its nonpolar environment and to its side chain H-bond to the side chain ofThr46. This part of the protein is more flexible than the extracellular half and must undergo a conformational change to open a proton path between the cytoplasmic surface and Asp96. To relate the elegant structure of BR in its resting state to the dynamic events of its photocycle, structural information has been obtained for different intermediates in the photocycle by crystallization of mutants prevented from completing the photocycle (such as D96N, which stops at the late M state) and by "kinetic crystallography" of wild-type crystals, which uses low temperatures and different wavelengths of light to Irap a significant population of the molecules in the crystals in one state. The structures show that the geometric and electrostatic effects of photoisomerization of retinal produce tensions in the protein molecule, which responds with small motions of residues and movements of discrete water molecules, as well as movements of helices G, F, and B (Figure 5.8). These detailed structures reveal a high-resolution "movie" that complements the spectroscopic data to present an exciting view of the dynamic mechanism of this light-driven proton pump, which is nature's simplest photosynthetic machine.

107

A.

Trp182

0/

.W407

B.

Photosynthetic Reaction Center

Nearly a decade before the first high-resolution x-ray structure for BR was published, the 1988 Nobel Prize in Chemistry was awarded to Hartmut Michel, Johann Deisenhofer, and Robert Huber for the elucidation of the x-ray structure of the photosynthetic reaction center (RC) from Rhodopseudomol1as viridis, Ihe first highresolution structure achieved for integral membrane proteins. When Michel and coworkers first crystallized the RC, the gene sequences encoding its protein constituents were not even available' The beautiful struclure of this multicomponent complex provided specific descriptors of its protein domains including TM helices, along with the locations of cofactors involved in light absorption and electron transfer. In photosynthesis light energy is converted into chemical energy when the absorption of a photon drives an electron transfer thai is otherwise

5.8. Examples of the structural shifts that occur during the photocycle in SR. Small differences are revealed when the high resolution structures of the active sites of the K and L intermediates are overlain on the structure of the ground state. A. The K intermediate, obtained for wild-type SR illuminated with green light at 11Oo K, shows disordering of a water (W402) and slight movements of Asp85 and Lys216. The circle indicates where another water molecule may appear. Positive and negative difference electron densities are shown in blue and yellow, respectively. B. Structural models for two different intermediates, K (blue) and L (red), are overlain with the ground state (green backbone with colored residues). The larger shifts in L include reorientation of the guanidinium group of Arg82, flexing of the backbone of helix C, and movement of the side chain of Trp182 toward the cytoplasm. From Neutze, R., et al., Biochim Biophys Acta. 2002, 1565: 144167. © 2002 by Elsevier Reprinted with permission from Elsevier.

Bundles and Barrels

108

5.9. The structure of the photosynthetic reaction center from Blastochloris viridis, a group II reaction center. The complex contains four protein subunits, L, M, H, and a cytochrome, and 14 cofactors (red). The TM helices are highlighted in yellow. Compare it with the group I reaction center shown in the chapter frontispiece. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., w. H. Freeman, 2005, p. 376. © 2005 by W. H. Freeman and Company. Used with permission.

thermodynamically unfavorable. The subsequent passage of electrons through spatially arranged carriers is coupled wi th the expulsion of protons, just as it is in oxidative phosphorylation, thus providing the proton gradient that drives the ATP synthase. Plants have two types of photosystems, called PSI and PSII, which di ffer in the electron acceptors used. The much simpler photosynthetic RCs of purple bacteria are considered the ancestors of PSII. Recent x-ray structures (at somewhat lower resolution) of both PSI and PSII reveal common structural features with the RCs in spite of their enormous size and complexity. RCs were discovered in photosynthetic purple bacteria and characterized by biophysical techniques such as EPR (see Box 4.2) and optical spectroscopy before they proved amenable to crystallization. Soon aher elucidation of the structure from R. viridis (renamed BlaslOchloris viridis), another high-resolution structure was obtained for the RC [yom Rhodopseudomonas sphaeroides (renamed Rhodobac/er sphaeroides). More recently, the x-ray structure for the RC from Ther-

l11ochromariul11 repidul11 was solved at 2.2 A resolution. While the cofactor-protein interactions are nearly the same in all three complexes, they represent two types of RCs. The Rb. sphaeroides RC is a member of group I and contains three protein subunits called L, M, and H, along with 10 cofactors (see Frontispiece). The other two RCs are members of group II, and they contain an additional subunit, a c-type cytochrome with its four heme cofactors (Figure 5.9). The three protein subunits, L (light), M (medium), and H (heavy), were named for their apparent molecular weights determ ined wi th SDS gel electrophoresis.

The Proteins The B. viridis RC has a size of ~ 130 A by ~ 70 A; its TM domain consists of five <x-helices from L, five C(helices from M, and one <x-helix from H. The remainder of the H subunit caps the structure on the cytoplasmic side, and the c-type cytochrome lies on the peripJasmic side (Figure 5.9). The TM helices composed of 21 to 28 amino acids cross nearly perpendicular to the plane of

Helical Bundles

109

L

H

M

cyt

weo

5.10. The peptide backbones of L, M, H, and cytochrome subunits of the photosynthetic reaction center from B. viridis. From Deisenhofer, J, et aI., Nature. 1985, 318:618-624. © 1985. Reprinted by permission of Macmillan Publishers Ltd.

the membrane. Three helices from each Land M subunit are nearly straight, while one is curved and one is bent more than 30 at a proline residue and ends with a 3 10 helix (a slightly narrower helix with three amino acids per turn). The structural similarity of Land M (Figure 5.10) gives a high degree of twofold symmetry in the TM domain in spite of only 26% sequence identity. The cytochrome with its two pairs of hemes also has twofold symmetry, unrelated to that ofL and M.lts com· pact structure consists of five segments: the N-terminal segment (residues Cl to C66), the first heme-binding segment (C67 to CI42), a connecting segment (C143 to C225), the second heme-binding segment (C226 to C315), and the C-terminal segment (C316to C336). As the first available detailed structure of integral membrane proteins, the RC confirmed expectations about distribution of amino acids on its surface, yet it revealed a new concept of interior polarity. The surface of the RC complex is polar in the peripheral subunits, with a net negative charge on the peri plasmic side and a net positive charge on the cytoplasmic side. The mem brane-spanning surface is very hydrophobic. There are no charged amino acids in this 30 A-wide band around the center of the RC, and very few water molecules associate with it. Like other membrane proteins, it has tyrosine and tryptophan residues distributed at the interfacial borders. Surprisingly, the polarity of the interior of the membrane-spanning G

domain is like that of the interior of soluble proteins, intermediate between the polarities of amino acids exposed to water and the hydrophobic interiOl- of the bilayer. The 10 TM helices of Land M together have 74 polar side chains. Furthermore, most of these polar residues do not appear to be involved in forming hydrogen bonds, as there are at most two hydrogen bonds between any pair of TM helices. Their predominant roles seem to be interactions with cofactors and protein subunits. Comparison of related RC sequences indicates that residues buried in the interior of the structure are conserved more than are residues on the surface. The M, L, and H subunits of the Rh. sphaeroides RC have 59%,49%, and 39% homology to subunits in B. viridis, respectively, and are very similar in structure. While some differences in amino acid sequence lead to differences in the interactions of the cofactors with the peptide chains, the complex has the same approximate twofold symmetry. The site-specific mutants first available in Rb. sphaeroides revealed the roles of GluL212 and SerL223 for reduction and protonation of the quinone and TyrM21 0 for efficient electron transfer. Lipids Based on its size and shape, the RC is predicted by EPR to have 30 to 35 annular lipids (see Chapter 4). However, most of the lipids are replaced by detergent during purification. For crystallization, the RC is solubilized with LDAO (N,N-c1imethyl-dodecylamine-N-oxide), and a few detergent molecules are included in the crystal structure. While it is often difficult to ascertain whether an acyl chain detected in the structure is from a lipid or a detergent, the x-ray structures give evidence for specific lipids (see Chapter 8), including a cardiolipin and a PE, which fit closely into hydrophobic grooves at the surface of the protein exposed to the nonpolar membrane domain.

The Cofactors The proteins provide a scaffold for the cofactors, holding them in the same spatial arrangement in all three RCs crystallized. The RC core has 10 cofactors: Four bacteriochlorophyJls (BChl), which resemble heme except for the replacement of iron by Mg2+ ions, a cyclopentenone ring fused to one pyrrole ring, and different substituents off two of the pyrrole lings Two bacteriopheophytins (BPh), which are BChl with two protons in place of the Mg 2+ Two qui nones (one ubiquinone and one menaquinone in B. viridis) A nonheme ferrous ion A carotenoid, which is a largely linear C40 polyene such as l3-carotene

110

There are two types of both BChl and BPh, a and b. The a type is found in the RC from Rh. sphaeroides and has either a phytyl or geranylgeranyl side chain, whereas the b type, found in the RC from B. viridis and T tepidum, has only a phytyl side chain and has one more C=C in the side chain of ring II. The cofactors form two symmetrically related branches within the hydrophobic environment of the closely packed TM helices (Figure 5.11). At the top, two molecules of BChl are positioned so close together that the edges of their tetrapyrrole rings overlap. Called the special pair, they receive the photon of light and release an electron in the primary event of photosynthesis. Each branch has another BChl (called the accessory BChl), a BPh, and a quinone. The nonheme iron is between the two quinones. The two branches follow the same local symmetry displayed by the Land M chains. The symmetry is not perfect, and the two branches have different electron transfer properties - in fact, electron transfer uses only the branch that associates with the L subunit. The cofactors are differentiated by groups of the protein that alter their environments. For example, the two quinones play different roles (see below) and the primary quinone is in a more hydrophobic environment than the secondary quinone. After being reduced and protonated, the secondary quinone (now quinol) diffuses from the RC, its leaving facilitated by nearby carboxylate groups. The similarities in the arrangement of cofactors in all photosystems, including the ironsulfur type PSI complexes, allow definition of a common motif: a dimer of tetrapYITole molecules nanked

5.11. The arrangement of the RC cofactors. The tetrapyrrole rings of BChl-b, BPh-b, and the quinone headgroups follow the same local symmetry displayed by the Land M chains. The figure shows the special pair, PA and PB (coral); the accessory BChl molecules, BA and BB (rose); the two BPh, HA and HB (cyan); the ubiquinones, OA and OB (yellow); carotenoid, Crt (purple); and non-heme iron atom (gray). The electron path utilizes only the A half, indicated by the arrows. From Jones, M. R., et al., Biochim Biophys Acta. 2002, 1565:206-214. (<;) 2002 by Elsevier. Reprinted with permission from Elsevier.

Bundles and Barrels

5.12. Structure of the tetraheme cytochrome subunit of the B. viridis reaction center. Location of the hemes is apparent when the polypeptide backbone of the cytochrome subunit is represented by a thread, while the heme groups are represented by space·filling models. The x's mark the visible positions of the sulfur atoms in the thioether side chains for the heme bridges, as well as free cysteines. From Knaff, D. B., et aI., Biochemistry. 1991, 30: 1303-131 O. © 1991 by American Chemical Society. Reprinted with permission from American Chemical Society.

by four monomeric tetrapyrrole molecules 'with an iron at the center. The cytochrome of group n RCs has four hemes, located in pairs on the two heme-binding segments of the polypeptide, each having a helix follo'wed by a turn and the Cys-X- Y-Cys-His sequence typical of c-type cytochromes. The hemes lie parallel to the axis of the helix with thioether bonds bet'ween each heme and the two cysteines (Figure 5.12). The fifth ligand to the iron is His, and for three of the four hemes the sixth ligand is a methionine residue within the helix. The reduction potentials for the hemes vary, and paradoxically the arrangement of hemes by potential does not follow the internal symmetry of the cytochrome. The closest heme to the special pair is heme-3, with the highest reduction potential (370 mY). However, in order of increasing distance from the special pail~ the heme reduction potentials follow the sequence high, low, high, low.' Spectroscopic measurements indicate that electrons are • In B. viridis, the reduction potentials are heme-I, -60 mV; heme2, +300 mV; heme-3, +370 mV; and heme-4, + 10 mV.

Helical Bundles

5.13. Model of the antenna system in purple photosynthetic bacteria. Viewed from above the plane of the membrane, each photosynthetic reaction center is surrounded by a light-harvesting complex called LH1. In addition, several LH2 complexes associate with it and with each other to rapidly transfer the excitation generated by absorption of light. From Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, p. 878. © 2004. Reprinted with permission from John Wiley & Sons, Inc.

transferred through heme-2 and heme-3 to the special pair, perhaps through heme-4, which is located between them but has a much lower reduction potential. Antennae

To maximize the absorption of light, the membrane contains about 100 times more bacteriochlorophyll A.

111

molecules than RC complexes. These other chlorophylls function as antennae to collect photons and pass the energy on to the special pair, greatly increasing the efficiency of each RC. These chromophores are organized by light-harvesting proteins into complexes called LH 1 and LH2. LHI is the core complex, found in a fixed stoichiometry with the RC. LH2 is peripheral, and its synthesis depends on factors such as light intensity (Figure 5.13). LH2 absorbs light at shorter wavelengths than LH I, so it rapidly passes the energy to LH I, which passes it to the RC. Both LH I and LH2 contain many copies of two short protein subunits called ex and 13 (in LH2, ex has 56 amino acids and 13 has 45 amino acids), each containing a single TM helix. In low-resolution EM images of native B. viridis membrane, the RC appears to be surrounded by a ring of 15 to 17 LHI molecules. A crystal structure of LH2 shows a double cylinder formed of an inner ring of its ex subunits and an outer ring of its 13 subunits, with eight- or ninefold symmetry depending on the organism of origin. A ring of ninefold symmetry has 18 BChl molecules between the rings, an-angecl like a water wheel, and nine more BChl molecules between the helices of its outer wall, along with accessory pigments such as carotenoids (Figure 5. I4A). However, both theoretical considerations and spectroscopic data suggest that the ring is not intact but is C shaped. Dimers of LH I observed at low resolution indicate that two Cshaped complexes face each other to form an S shape B.

5.14. Organization of the light-harvesting complexes. A. Crystal structure of LH2 from Rhodopseudomonas acidophila, viewed from the peri plasmic side. The complex has ninefold symmetry, shown with (X subunits in yellow, [3 subunits in green, BChl in gray, and carotenoids in orange. B. Projection map of membranes from Rb. sphaeroides grown under photosynthetic conditions in the presence of nitrate, in which two C-shaped LH1-RC complexes dimerize to an S shape. From Vermeglio, A., and P. Joliot, Trends Microbiol. 1999, 7:435-440. © 1999 by Elsevier. Reprinted with permission from Elsevier.

Bundles and Barrels

112

Soluble electron canier proteins (Cyt c2 or HiPIP) H+ -ATPase HI

,0

Lighl energy -1.0

~/

BChl:; BChl BPhe

~

"0

2: -05

] c:

QA

'"00 0.0

, OB

c:

H+ RC

LHI & LHII

8-

~

~'

0.5

BChl 2

1.0

5.15. Illustration of the photosynthetic electron transfer reactions in purple bacteria. The RC (pink) accepts energy from the antenna complexes LH 1 and LH2 (purple). After charge separation in the RC complex reduces the secondary quinone (OB) to quinol, it diffuses through the membrane to the cytochrome-bc, complex (green). where it is oxidized back to quinone. Cytochrome bCl transfers the electrons to soluble electron carrier proteins (blue). either cytochrome C2 or HiPIP, which carry them back to the RC Cytochrome bCl also generates a proton gradient used by the ATP synthase (H+ATPase, yellow) to drive the synthesis of ATP. Redrawn from Nogi, T., and K. Miki, J Biochem (Tokyo). 2001, 130:319-329. © 2001. Reprinted with permission from the Japanese Biochemical Society. Inset: Potentials of cofactors involved in electron transfer in purple bacteria. The dotted line represents the absorption of light by the special pair of BChl, and the solid line represents the energy transfer steps in the RC to the quinones, OA and 0B (shown in Figure 5.11). The dashed line represents steps that occur outside the RC Redrawn from Allen, J. P., and J. C Williams, FEBS Lett. 1998,438:5-9. © 1998 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

(Figure 5.14B). Dimers of LHI could be isolated only in the presence of another membrane protein caIJed PufX, named for photosynthetic unit Formation. This protein, which is requit-ed for anaerobic photosynthetic growth, associates closely in a 1:1 ratio with RC-LHI complexes. The Reaction Cycle The overall reaction carried out by the photosynthetic RC is the reduction of ubiquinone by cytochrome C2, using the energy fyom light to initiate the electron transfer. It occurs through the following steps (Figure 5.15): 1. The light energy absorbed via antenna complexes LHl and LH2 is transmitted to the RC, where it promotes a charge separation with the oxidation of the special pair of bacteriochlorophyll and the two-electron reduction of quinone to quino!. When the special pair absorbs light, it gives a distinct optical absorption band in the near infrared. The cofactors in one branch are close enough to delocalize the electrons in a conjugated system, so electron transFel- is fast - \vithin 200 ps the electron reaches the primary quinone. The primary quinone accepts one electron and releases it to the secondary quinone, which then accepts a second electron from a second photon event. Next the secondary

quinone picks up two protons from the cytoplasm and enters the membrane's quinone pool as quinol (Q B H 2 ). 2. The quino! diffuses in the membrane to the cytochrome-bcl complex, where it is reoxidized, with concomitant release of electrons to periplasmic electron carriers, either cytochrome C2 or in some species (including T. lepidum) the abundant high-potential iron-sulfur protein (HiPlP). The cytochrome-bcl complex is similar to complex III in the mitochondrial membrane. It releases protons into the periplasmic space, From which they reenter the cytosol via the FIFo-ATP synthase (see Chapter 11) to drive the synthesis of ATP. 3. The periplasmic electron carriers bring electrons to the RC to reduce the special pair. Cytochrome C2 is very similal- to mitochondrial cytochrome c and diffuses along the surface of the membrane to the RC. The Rb. sphaeroides RC has been co-crystallized with cytochrome C2. which appears to bind quite near the special pair and to reduce it directly. In group II RCs, the soluble electron carriers dock on the cytochrome subunit of the RC and reduce its hemes. The binding sites envisioned on crystal structures of the separate redox components indicate that the cytochrome C2 interaction with the RC cytochrome is electrostatic, while that between HiPlP and the T. lepidum RC is hydrophobic (Figure 5.16).

I3-Barrels

113

5.16. Recognition of electron carriers by the cytochrome subunit of the RC. In the case of HiPIP, the interacting surfaces on both proteins are hydrophobic (A and B) while the recognition of cytochrome C2 is electrostatic (C and 0), as the B. viridis cytochrome C2 has a large basic surface and the cytochrome subunit of RC has a large acidic surface. Negatively charged surfaces are red, and positively charged surfaces are blue. The green dotted circles indicate the interacting surfaces involved in recognition. From Nogi, T., and K. Miki, J Biochem (Tokyo). 2001, 130:319-329. © 2001. Reprinted with permission from the Japanese Biochemical Society.

Completion of the cycle takes less than 100 ~s and gives a quantum yield close to one, meaning that for almost every photon absorbed by the RC, one electron is transferred. There is much more to learn about how the RC regulates the electrochemical properties of its cofactors and precisely how it takes up protons from the cytosol (when it reduces quinone) and electrons from reduced periplasmic carriers (to reduce the special pair at the end of the cycle). To address these and other questions, researchers are now explori ng structural differences between RCs in the light and those adapted to darkness, between wild type and mutants, and between isolated RC and the complex consisting of RC and its partners of the photosynthetic membrane.

I3-BARRELS

While the structures of bacteriorhodopsin and photosynthetic reaction center came to represent the image

of membrane proteins, a wholly different picture of membrane protein structure was emerging in studies of bacterial porins. Early data from circular dichroism and infrared spectroscopy indicated that these poreforming proteins contain l3-structure and little or no helical content; EM and x-ray diffTaction studies suggested they span the membrane as l3-barrels. The first x-ray crystal structure of a TM f)-barrel was that of the porin from Rhodobacter capsuLatus (Figure 5.17) and was quickly followed by the high-resolution structures of other bacterial porins. The cell envelope of Gram-negative bacteria consists of two membranes separated by an agueous compartment called the periplasm and a thin layer of peptidoglycan, a net-like polymer of amino acid and sugar residues that confers structural stability (see Figure l.IB). The f)-ban-el proteins are found in the outer membrane, while most proteins in the inner (plasmal membrane are bundles of ex-helices. This difference is attributed to mechanisms of biogenesis, since the more hydrophilic

Bundles and Barrels

114

A.

B.

5.17. Tracing of porin from Rhodobacter capsulatus, the first porin with an x-ray crystal structure. A. View from the side of the monomer, with chain ends marked by dots. B. View of the trimer from the top. From Weiss, M.S., et al., FEBS Lett. 1990,267:379-382. © 1990 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

f)-barrel proteins are secreted into the periplasm before incorporation into the outer membrane, while the inner membrane proteins are incOl-porated directly into the bilayer from the translocon (see Chapter 7). f)-Barrel proteins are predicted to make up 2% to 3% of the proteome of Gram-negative bacteria, in which they carry out diverse functions. They also are found in mitochondria and chloroplasts of eukaryotes but not in Archaea or in Gram-positive bacteria, except Mycobacteria. Due to its extended peptide backbone, a f)-strand needs only seven amino acid residues to cross the nonpolar domain of the membrane. Typically TM f)-strands have nine to J 1 residues, usually tilted 45 fTom perpendicular to the plane of the bilayer, although observed tilts vary from 20' to 45°. To satisfy the H-bonding of the carbonyl and amino groups along their peptide backbones within the nonpolar domain, f)-strands must partner with other f)-strands. By formi ng a circular pattern in a barrel, none of the strands are left without a partner. The interstrand H-bonding makes the structure rigid, as well as very stable: for a small f)-barrel of eight TM strands, the H-bonding contributes a stabilization of around 40 kcal/mol (eight strands of ~ 10 amino acids, each forming 80 H-bonds x 0.5 kcallmol H-bond). Among known f)-ban-el structures of membrane proteins, the number of f)-strands varies from eight to 22. In the smallest of these proteins, the center is filled with polar residues, while barrels of intermediate sizes (16-18 strands) contain aqueous pores. The larger barrels with 22 strands have "hatch" or plug domains that 6

close their channels. An even number of strands allows the Nand C termini to meet since the barrels are made from meandering antiparallel strands, which means all of them are hydrogen-bonded to their next neighbors along the peptide chain. With strands connected by tight turns or loops, the twisted f)-sheet rolls into a cylindel~ In the known structures, the peri plasmic ends of the f)-strands form tight turns and the other ends are loops of varying lengths, either pointing into the extracellular space or folding back inside the barrel. [n general, the f)-strands are amphipathic, with polar side chains pointing to the aqueous interior of the protein alternating with nonpolar side chains contacting the hydrophobic bilayer. (See the sequence of OmpF in Figure 5.19D for an example.) This means the composition of the lipid-exposed surfaces of f)-barTels, like those of £x-helical integral membrane proteins, is rich in aromatic and nonpolar amino acids (Phe, Tyr, Trp, Val, Leu, lie; Figure 5.18). At the two bilayer interfaces, aromatic residues are ~40% of the lipid-exposed residues. These aromatic side chains form gird les whose intermediate polarity helps define the two nonpolarpolar interfaces of the membrane, also observed in £x-helical membrane proteins. The exposed loops at the extracellular surface provide binding sites for colicins and bacteriophage, as well as recognition sites for antibodies. Interestingly, half of the strands of the small f)-barrel called OmpX protrude into the external medium and nonspeciRcally bind proteins with exposed f)-strands (see Figure 5.18A and Box 5.1). Because of this unusual feature, OmpX, which is not a porin, is called an adhesion protein. It is induced by stress and

(3-Barrels

115

B.

PRO

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o 5.18. The structure of OmpX, a small, closed l3-barrel protein, Because OmpX is monomeric, all of its exposed side chains go into the bilayer. Additionally, four of its eight l3-strands protrude on the outside of the cell. The barrel interior is filled with polar residues that form a hydrogen-bonding network. A Ribbon drawing shows nonpolar side chains (yellow) on the surface of OmpX. From Schulz, G. E., Curr Opin Struct Bioi. 2000, 10:443-447. © 2000 by Elsevier. Reprinted with permission from Elsevier. B, Topology model shows the primary structure and distribution of amino acids in OmpX, with lipidexposed (red), interior (black), and external loops (green). The y-axis gives the transbilayer location with the center of the bilayer as 0, To show the interstrand hydrogen bonds between each pair of strands, the eighth strand is repeated on the left (gray). C. Bar graphs show the distribution of amino acid residues on the external surface (left) and in the interior (right). Band C from Wimley, W. C, Curr Opin Struct Bioi. 2003, 13:404-411. © 2003 by Elsevier. Reprinted with permission from Elsevier.

is thought to be a part of the virulence mechanism of E. coli. There is much variation in quaternary structure among the known (3-barrels. Several of them are monomers, while all the porins are homo-oligomers (usuaJJy trimers) with very tight interactions between the subunits. The enzyme activity of outer membrane

phospholipase A (OMPLA; see Chapter 9) is regulated by its quaternary structure: it is only active as a homodimer. The (3-barrel formed by ct-hemolysin (see Chapter 4) results from a completely different motif, in that each subunit of the heptamer contributes a (3hairpin to make the 14-stranded barrel that inserts into target membranes. The rich diversity of structure and

116

Bundles and Barrels

BOX 5.1. NMR determination of membrane protein structure Structure determination by nuclear magnetic resonance (NMR) spectroscopy is limited to fairly small proteins and is even more difficult for membrane proteins in detergent micelles with their increased particle sizes. Complete structures have been solved for only a few small membrane proteins (~20 kDa or less) in detergent micelles. While there has long been hope for application of solid-state NMR to solve the structures of membrane proteins, the structures obtained to date are from solution NMR. These successes were made possible by the development of TROSY (transverse relaxation optimized spectroscopy) by Kurt Wuthrich, for which he was awarded the Nobel Prize in Chemistry in 2002. NMR

has the potential to measure dynamics within the protein structure, as well as to give information about protein-lipid interactions.• Structures of f)-barrels are easier to solve by NMR than those of helical bundles because (1) the 1 H chemical shift

A detailed explanation of these NMR methods is beyond the scope of this book. See Fernandez, C., and G. Wider, TROSY in NMR studies of the structure and function of large biological macromolecules, Curr Opin Struct BioI. 2003, 13:57Q.-580. For a general introduction, read MagnetiC Resonance in Chemistry and Medicine by Ray Freeman, published by Oxford University Press, 2003.

5,1,1. OmpX structure solved by solution NMR. A. NMR spectra of OmpX in DHPC micelles obtained using 'H lsN COSY (a) and 'H 1sN TROSY (b). B. Structure of OmpX determined by NMR. When the NMR assignments are mapped onto the x-ray structure of OmpX (a), the barrel portion is well-structured (red) and three loops are disordered (yellow). The flexibility of the loops is clear in the NMR structure represented by the superposition of 20 conformers (b). From Fernandez c., et aI., FEBS Lett. 2001, 504: 173-178. © 2001 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

I)-Barrels

117

BOX 5.1. (continued)

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(a) 5.1.2. OmpA structure solved by NMR. A. TROSY-based 1 H_ 15 N spectrum of the TM domain of OmpA in DPC micelles. B. NMR structure of OmpA. The NMR solution structure of OmpA is represented by the superposition of the ten confomers having lowest energy (a). Again the l3-barrel (red) is well-defined. while the loops are disordered. Four flexible loops are resolved in the ribbon diagram based on the NMR results (b). C. Comparison of the NMR (red) and x-ray (blue) structures for OmpA show good agreement on the l3-strands and considerable differences in the loops. From Arora, A., et aI., Nat Struct Bioi. 2001, 8:334-338. © 2001. Reprinted by permission of Macmillan Publishers Ltd.

(continued)

Bundles and Barrels

118

BOX 5.1. (continuec/)

c.

~C(X77

Outer membrane

(Figure 5.1.1 A) shows the enhancement obtained with TROSY. which allowed the eight-stranded fold of the polypeptide backbone of OmpX to be determined from 107 nuclear Overhauser effect (NOE)-derived distance constraints and 140 dihedral angle constraints. The structure was further refined with selective protonation of the methyl groups of Val. Leu, and lie on a perdeuterated background. which enabled assignment of 526 NOE distance constraints. When 20 NMR conformers are superimposed. the result clearly reveals the f)-barrel with its flexible loops (part b of Figure 5.1.1 B). The ribbon diagram in part a of the figure maps the NMR results on the x-ray structure and distinguishes the well-structured regions (red) from the disordered loops (yellow). The NMR result is strikingly similar to the structure from x-ray crystallography and in particular confirmed the extension of f)-strands on the exterior of the protein.

OmpA TM Domain from E. coli

Periplasmic end

C N

5.1.2. (continued)

dispersion is larger for f)-sheets than for lX-helices. especially with alternating hydrophilic/hydrophobic residues. and (2) f)-barrels have higher thermal stability. so they can withstand higher temperatures for the longer times needed to get well-resolved NMR spectra. Furthermore. the two f)-barrel proteins described here could be overexpressed in E. coli and purified in denatured form from inclusion bodies before refolding in detergent micelles (see Chapter 7). resulting in a mixed micelle particle size of 60 to 80 kOa. Uniform labeling of the proteins with 2H. 13C. and 15N allows detection of the protein NMR signals with little or no interference from the signals of the unlabeled detergent molecules. OmpX from E. coli Triply labeled OmpX. a 148-residue protein. was solubilized from inclusion bodies with guanidinium chloride and reconstituted in dihexanoyl-PC (OHPC) micelles. Comparison of the 20 COSY (Correlation Spectroscopy) and TROSY spectra

function among 13-barrels continues to be uncovered, yet the paradigm for this group of membrane proteins is the porins.

Porins The outer membranes of Gram-negative bacteria protect the cells from harmful agents while allowing nutrient uptake via porins and other transport proteins. Porins are pore-forming TM proteins that function as passive diffusion channels and thus allow rapid diffusion of their solutes. even at O°c' Porins are grouped in two categories: general porins. like OmpF and OmpC, and specific pOl-ins, like PhoE (phosphoporin), LamB (maltoporin). ScrY (sucrose porin). and Tsx (the nude-

The f)-barrel domain of OmpA (residues 172-325) was purified after denaturation in urea and refolding into dodecylphosphocholine (OPC) micelles. The protein was labeled with 15N, 13C. and 2H, and TROSY experiments were carried out at 600 and 750 MHz. The protein in a large excess of OPC gave the best NMR spectra, shown in Figure 5.1.2A with several of the assigned resonances labeled. TROSY experiments were carried out with several specific amino acid-labeled samples to aid in assignments. The backbone fold of the OmpA TM domain was initially calculated from 91 NOE distance constraints and 142 torsional angle constraints. It was refined by introducing 116 H-bond constraints between adjacent f)strands that were identified in the initial fold calculations. The structure of the eight-stranded f)-barrel is well defined. Figure 5.1.2B shows 10 superimposed conformers (a) and the corresponding ribbon diagram of the solution structure (b) of OmpA. Like the OmpX structure. the NMR structure of OmpA shows much agreement with its x-ray structure, illustrated in Figure 5.1.2C, where the solution structure determined by NMR (red) is overlain with the x-ray crystal structure (blue). As NMR gives information about protein dynamics. some of the poorly defined loops are thought to have intrinsic high mobility. A study of the dynamics of the backbone from 15N relaxation times indicates the H-bonded core is not completely rigid but moves on the microsecond-millisecond time scale.

oside channel). The channels of general par'ins do not discriminate among solutes thal are hydrophilic. under ~600 Da. and not highly charged. which allows them to take up many nutrients such as mono- and disaccharides. In contrast. the specific porins have channels that are selective for their solutes. although the line of demarcation is blurred as described below for PhoE. Note that aquaporin, which is a TM channel for water molecules in plasma membranes (see Chapter 10). is not a 13-barrel. Porins were first detected by the permeability of the outer membrane of Gram-negative bacteria to hydrophilic antibiotics. Thus the first assay was based on hydrolysis of 13-lactams (penicillin and its derivatives) in intact cells. since their rate of permeation

~-Barrels

through the outer membrane determines their availability for hydrolysis by the enzyme ~-Iactamase in the periplasm. Channel activity of pOl'ins is evident when the purified proteins are reconstituted with lipids: porins make lipid vesicles permeable to small, hydrophilic solutes and make voltage-gated conductance channels in black films. OmpF and Om pC Typically Gram-negative bacteria have ~lOs copies of general porins per cell, with the number of species of porins varying in different strains. The outer membrane of E. coli K12 is dominated by OmpF and OmpC pOl'ins, both homotrimers of around lIS kDa whose levels are controlled by osmolarity of the growth medium. This regulation is carried out by a two-component system made up of EnvZ, the protein sensor of osmolarity, and OmpR, the transcriptional activator for their two genes. OmpC has been called osmoporin, because its expression is induced by high osmolarity as well as high pH and high temperature. On the other hand, higher levels of OmpF are expressed in medium of low osmolarity. OmpF has a larger channel diameter than does OmpC, and although the size difference is only ~IO%, the flux of larger solutes through the OmpF channel is significantly faster than that through OmpC. This difference allows the bacteria to respond to two very different habitats: in mammalian hosts, high osmolal-ity and high temperature induce OmpC, whose narrower channel keeps out some of the body's inhibitory substances, \-vhereas outside the host the wider channel of OmpF speeds the uptake of nutrients h'om very dilute environments, such as ponds and rivers. Purification of porins is facilitated by their strong, noncovalent association with peptidoglycan, as this complex can be isolated by extraction of the cell envelope with SDS at 60°C: while most of the membrane proteins are solubilized, the porin remains in homogeneous two-dimensional crystalline aggregates. These aggregates have porins in hexagonal arrays (with patches of phospholipids between porin trimers) and give typical conductance properties when incorporated into planar bilayers. To remove the peptidoglycan, further purification involves salt treatment before gel filtration or ion exchange chromatography. POI-ins can also be solubilized [Tom cell envelope by extraction with non ionic detergents such as octyl-POE (see Chapter 3). Detergent-solubilized porin binds 0.6 g detergent per gram of protein, which amounts to around 200 detergent monomers per protein trimer. The protein is remarkably stable: it is resistant to proteases, chaotropic agents and most organic solvents, in addition to SDS and other detergents. It is even functional after lyophilization and storage at -20°C.

119

Crystals of OmpF from E. coli were described as early as 1980 but due to their unusually symmetric packing, the phase problem could not be solved. lL was another decade before a crystal structure of a porin \,vas solved, that of the analogous porin from Rb. capsulaIus. The high-resolution structure of OmpF, as well as that of PhoE, was then determined. The highly homologous OmpC resists crystallization but is expected to share many of the structural features of Rb. capsulalus porin and OmpF. Even a porin with weak homology can have very similar architecture, as demonstrated in the x-ray structure of the porin h'om Rhodopseudomonas blaslica. These porins are homotrimers, in which each subunit forms a ~-barrel (Figure 5.19). The barrel consists of 16 antiparallel ~-strands tilted by 4SO, with a salt bridge between the amino and carboxyl termini to complete the cyclic structure. At the extracellular end one loop latches onto the next subunit, and one or more of the loops fold back. in to the channel. Loop 3 in OmpF has the highly conserved sequence motif PEFGG and folds into the channel to constrict the pore at the middle of the barrel to IS x 22 'A. This eyelet of the pore has clusters of acidic residues on one side and basic residues on the other, which create a local transverse electric field that accounts for the slight cation specificity observed in conductance studies (Figure 5.20A). As discussed in Chapter 3, conductance measurements can detect Single-channel openings, as well as channel closures at voltages above 100 mV (see Figure 3.16), although the mechanism of channel closing and its significance under physiological conditions are unknown. Characterization of structure and function of porins with mutations affecting the constriction site of the channel has enabled properties such as single-channel conductance and transport rates to be correlated with precise changes in the crystal structures. None of the mutations altered the barrel framework of the protein. Single amino acid substitutions for each of the charged residues of the eyelet alter the ion selectivity but produce little change in pore size and conductance (Table 5.1). If, however, both acidic residues are mutated (DI13N/E 1170), the conductance drops by 50% and the cation selectivity is removed. Simulations of ion flow by Brownian dynamics illustrate the role of the acidic residues in drawing cations through the constriction site. The change in pore size does not always COlTelate with changes in conductance but is well reflected in uptake rates for disaccharides measured by the liposome-swelling assay, as seen in the effects of deletion of six residues of Loop 3 (6109-114) and substitution of neutral amino acids for five charged residues (R42A1R82A/RI32A1DI13N/EI170). Clearly both pore dimensions and the constellation of charged residues

A.

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5.19. Structure of OmpF porin. A. Ribbon diagram of the OmpF monomer showing the long loops facing the cell exterior and the l3 loop that folds inside the barrel, as well as the proximity of the Nand C-terminal residues. B. View of the OmpF trimer from the top (outside) showing the constriction of each pore by the l3 loop. C. Slice through the center of the OmpF trimer using a stick model enhanced with the molecular surfaces, showing the dimensions of the pores. A through C from Schirmer, T., j Struct Bioi. 1998, 121: 101-1 09. © 1998 by Elsevier Reprinted with permission from Elsevier. D. The topology of OmpF as an unrolled barrel with the last two l3-strands repeated to emphaSize the cyclic nature of the structure. Residues in l3-strands are in diamonds, residues in !X-helices are in rectangles and residues in turns and loops are in circles. From Cowan, S. W., et al., Nature. 1992,358:727-733. © 1992. Reprinted by permission of Macmillan Publishers Ltd.

at its narrowest site determine the transport properties of the general pm-ins. Specific Porins

Sharing many characteristics of general pOI-ins, the specific porins add to them the ability to discriminate

among solutes. Since they carry out passive diffusion, they cannot rely on energized conformational changes to release substrates from high-affinity binding sites (see below and Chapter 10). Rather, the specific porins have low affinities for their substrates and have achieved selectivity through features of their channel architecture. Their specificity is apparent only with

~-Barrels

121

B.

A.

5.20. Key amino acid residues in the constriction zones of porins. A. Viewed in cross-section, the channel of the OmpF porin monomer is constricted by Loop 3, forming an eyelet lined by clusters of basic residues (K16, R42, R82, and R132) and acidic residues (D113 and E117). B. The channel constriction in PhoE, also formed by Loop 3, has two additional basic residues, here numbered K18 and K131. Note the residue numbers in PhoE have been adjusted to match those in OmpF. From Karshikoff, A., et aI., j Mol BioI. 1994, 240:372-384. © 1994 by Elsevier. Reprinted with permission from Elsevier.

TABLE 5.1. Channel properties of OmpF, PhoE, and OmpF mutants

Protein OmpF PhoE D111G E117Qe D113N/E117Q(NQ) D113N/E117Q/R42A/R82A/R132A (NQAAA) R42C R82C R132P V18K1G131K(KK)

Mimimal cross-section a (%)

Conductance b (nS)

SelectivityC PNa/PC!

Disaccharide permeation d Relative swelling rates

100 70 129 100 94 177

0.S4 ±0.06 0.63±0.06 0.SO±0.06 0.64±0.02 0.40±0.02 0.64 ±0.04

4.5 ± O.S 0.44 ±0.05 1.4±0.1 2.9 ±0.2 1.0±003 12.3 ±0.9

5 2 2 2 12±1.5 7±3 NT 37 ± 1.5

127 114 109 75

07S±0.07 0.76±0.06 o 77±0.04 0.75±0.02

9.7 ±2.0 2.1 05 nO±1.9 2.1 ±O.OS

NT NT 1O±3 <1

D Minimal cross-section describes the accessible area of the pore available to a probe with a radius of 1.4 A; values are normalized to 31 A2, that of OmpF b Mean of > 100 single-conductance step measurements. nS, nanosiemen. C PNa/P CI (the relative rates of permeation by Na+ and (1-) was determined from at least four different preparations. d Disaccharide uptake is measured by the liposome-swelling assay in MLVs and given as the mean swelling rates for sucrose, lactose, melibiose. and maltose. NT, not tested. e The liposome-swelling assay utilized E117C instead of E117Q. Source: Phale, P. S., et aI., Biochemistry. 2001, 40:6319-6325. © 2001 by American Chemical Society. Reprinted with permission from American Chemical Society.

Bundles and Barrels

122

Cell exterior

-PeripJasm 5.21. X-ray structure of a monomer of LamB, the maltoporin. Ribbon diagram of the LamB monomer viewed from the membrane-exposed surface, with the external chains given in ball representation. (Oxygen atoms are in black.) The arrows point to the aromatic girdles at the boundaries between the nonpolar and interfacial regions of the bilayer, and the vertical bar (~25 A) denotes the nonpolar region. The charged side chains are mainly in the outer region of the barrel, where they could interact with lipopolysaccharide. From Schirmer, T., J Struct Bioi. 1998, 121: 101-1 09. © 1998 by Elsevier. Reprinted with permission from Elsevier.

larger molecules because small solutes like arsenate and glucose easily permeate the nonspecific channel interiors. Clues to their specific functions come from their regulation: for example, in E. coli, PhoE protein is induced under phosphate starvation and LamB protein is induced by growth on the carbon source maltose. PhoE, the Phosphoporin

In spite of the very high homology between PhoE, OmpF, and OmpC, PhoE has a specific transport function. When their functions are compared in whole cells, mutants constructed to have only PhoE take up phosphate and phosphorylated compounds much more efficiently (,vith a ninefold decrease in K m of tl-ansport) than mutants with only OmpF or OmpC. Conductance studies with purified PhoE protein demonstrate a strong anion selectivity. Furthermore, polyphosphates such as ATP inhibit the flux of small ions through reconstituted PhoE channels. Like OmpF protein, the PhoE protein is a 16stranded j3-barrel; in fact, the barrel structures of the two proteins are superimposable, and the differences in their folds are confined to the loop regions and a single short turn. The constriction zone of the PhoE pore has two additional basic groups (Figure 5.20B),

and the calculated electrostatic potential of the pore is more strongly positive than that for the pore of OmpF. A genetic approach was taken to identify critical residues of PhoE that might form a phosphatebinding site, targeting basic residues that replace neutral or acidic amino acids in OmpC or OmpF. Indeed the single mutation K125E changes the ion selectivity of the PhoE pore. In the crystal structure of PhoE, this lysine residue is in Loop 3 at the constriction site (as KI25 in PhoE corresponds to amino acid 131 of OmpF; it is labeled Lys131 in Figure 5.20B). Another unique lysine is at the mouth of the pore. Howevel~ introduction of lysine residues at these locations in OmpF (OmpF mutant V18KJG131K) did not completely convert it to a PhoE-like pOl-e (Table 5.1). Other residues in PhoE must contribute to its selectivity; for example, Serl15 contributes by changing the position of the backbone in Loop 3 to make room for the side chain of Lys 131. Electrostatic contributions from other residues are probably important, as in the pore in OmpF. Given the lack of a specific binding site, PhoE might be viewed as a general porin. [ts selectivity is achieved with nonsaturable binding sites of low affinity. Howevel~ it clearly facilitates the transport of phosphorylated compounds into cells, and its blockage by polyphosphates is velY similar to the inhibition of the LamB pore by maltodextrins. LamB, the Maltoporin

LamB protein is named for its role as the receptor for phage A.It is required for growth of E. coli in chemos tats with limiting maltose as the sole carbon source, which provided early evidence for its role in maltose transport. The specificity of the LamB channel for maltose and maltodextrins can be detected by comparing rates of sugar uptake in the liposome-swelling assay (see Figure 3.26). The reconstituted LamB pore shows little discrimination among monosacchal-ides, but among disaccharides uptake of maltose is > 10 times faster than that of lactose and 40 times faster than that of sucrose. Other than maltodextrins, sugars larger than d isaccharides do not permeate the channel. The affinity for maltodextrins can be quantitated by their inhibition of glucose uptake as well as by binding to immobilized starch; both show a weak affinity (K,s in the low mM range). Maltodextrins also block conductance through the LamB channel when reconstituted in black films. Overall, the high-resolution structure of LamB protein shows similar architecture to the other porins: it is a homotrimer in which each monomer has 18 strands in the f)-barrel (Frontispiece and Figure 5.21). Three of the loops fold in to constrict the channel to a minimum diameter of 5 A. Six aromatic residues line one side of the channel, forming the "greasy slide," a smooth hydrophobic path through the otherwise aqueous pore

0-Barrels

123

(Figure 5.22). Soaking the maltoporin crystals in maltodextrin and different disaccharide substrates showed the substrates threading lengthwise through the channel with the hydrophobic sides of their pyranose rings along the greasy slide. The specificity for the maltose configuration is determined by numerous hydrogen bonds to charged side chains on the other side of the channel. When sucrose diffuses into the crystals, it gets stuck above the channel constriction, which explains its very low rate of uptake into liposomes. Interestingly, a sucrose-specific porin, called SrcY, is homologous to LamB protein, with a few strategic differences in the residues revealed in its high-resolution structure. Thus, like PhoE, the selectivity of these pores for their sugar substrates is achieved by the specificity built into their channels.

Iron Receptors 5.22.

Side view of the LamB monomer showing the "greasy slide". Maltodextrins pass through the pore down a slide made up of six aromatic residues: Trp74 from the adjacent monomer, Tyr41 , Tyr6, Trp420, Trp358 and Phe227 (in purple, with oxygen atoms red and nitrogen atoms blue). Tyrl18 (green) constricts the channel from the other side. From Koebnik, R., et ai, Mol Microbiol. 2000, 37:239-253 © 2000. Reprinted with permission from Blackwell Publishing.

Receptors involved in iron transport are 0-barrel proteins with an entirely different transport mechanism. Iron is abundant but unavailable in the environment due to its insolubility as ferric hydroxides. To solubilize this essential mineral, microorganisms synthesize and secrete siderophores, iron cheJators of 500 to 1500 Da with extremely high affinities for ferric ions. The

5.23. Structures of FepA and FhuA proteins involved in iron transport. A. Side views with the extracellular surface on top and the periplasmic end on the bottom. B. Views from the external solvent show the blockage of the interior. The FhuA l3-barrel is blue and plug domain is yellow; the FepA l3-barrel is green and plug domain is orange. From Ferguson, A. D., and J. Deisenhofer, Biochim Biophys Acta. 2002, 1565:318-381. © 2002 by Elsevier. Reprinted with permission from Elsevier.

124

enteric bacteria, including E. coli, synthesize and transport an iron chelator called enterobactin and also have transport systems for siderophores secreted by other microorganisms, such as ferrichrome. These transport systems consist of outer membrane receptors, periplasmic binding proteins, and inner membrane transport proteins. Unlike the porins, the outer membrane iron receptors bind their substrates with high affinities (K! ~O.l I-lM) and pump them into the periplasm at the expense of energy. The energy is provided via a complex of three proteins, TonB, ExbB, and ExbD, anchored in the inner membrane. In E. coli the iron receptors, along with another TonB-dependent protein, BtuB, the receptor for vitamin B 12 , interact with TonB at a conserved sequence of five amino acids near the N terminus called the ''TonB box" (TXXV[S/T], where X is a hydrophobic residue; see Chapter 11). While these interactions are well characterized, the mechanism of energy delivery is unknown. ExbB and ExbD are candidates for a proton translocation apparatus that couples chemiosmotic potential with conformational changes in TonB. In this scenal-io, the energized TonB then binds FhuA or FepA to open a high-conductance channel either by moving the plug domain into the peri plasm as postulated for the BtuB protein (see Figure 11.41) or by conformational changes more like those of other transporters, such as the sugar transpOl-ters LacY and GlpT described in Chapter 10. High-resolution structures show many similarities between the E. coli receptors for enterobactin and for ferrichrome, the FepA protein and the FhuA protein, respectively. They each have t\,vo domains, a C-terminal domain that makes a 22-stranded f)-barrel, and a globular N-terminal domain of ~ 150 residues that fills the interior; making a plug (Figure 5.23). The barrel has a diameter of ~40 Aand extends beyond the bilayeron the outside. Some of the 11 loops on the external membrane surface are unusually long: they range from seven to 37 l-esidues in FepA. The plug domain has a four-stranded I)-sheet and interspersed LX-helices and loops, including two loops that extend 20 A beyond the outer membrane interface to frame a pocket with the binding site for the siderophores. The largest difference between the two \-eceptors is the nature of this site, which is tailored for siderophores that differ in composition and charge. Comparison of the x-ray structures of FhuA in the presence and absence of substrates indicates that a conformational change takes place in the N-terminal domain upon ligand binding. Small movements of the loops in the binding pocket trigger unwinding of an LXhelix and a large movement of the polypeptide chain to the opposite side of the barrel wall (Figure 5.24). These changes do not open a transport channel, but

Bundles and Barrels

5.24. Conformational change in FhuA when it binds its siderophore. The (3-barrels of unliganded and liganded FhuA are shown as wire frame models. The plug domain of unliganded FhuA is blue and that of liganded FhuA is red. From Ferguson, A. D., and J Deisenhofer, Biochim BiophysActa. 2002, 1565:318332. © 2002 by Elsevier. Reprinted with permission from Elsevier.

could enable the binding of TonB to do so (see Chapter 11). The structural information from studies of FepA and FhuA sharpens the understanding of iron transport in E. coli and also provides models for its investigation in other microorganisms. Certainly iron receptors in pathogenic bacteria are imponant from a medical standpoint. Acquisition of iron is critical for invading microbes, which extract iron from host proteins such as transferrin and hemoglobin. In addition, the iron receptors are used by an ti biotics as well as col ici ns to enter the cell. Finally, siderophore-drug conjugates show potential for targeted drug delivery. From the porins to the iron receptors, tremendous progress has produced elegant structures of I)-barrel membrane proteins along with enhanced understanding of their transport mechanisms. Even so, it is likely they represent a small h-action of the proteins in this class of membrane proteins. If the genomic analyses are correct, there are many more I)-barrel proteins to be characterized. The next chapter looks at bioinformatics techniques used to predict and analyze membrane proteins and shows how they al-e grouped in families.

For Further Reading FOR FURTHER READING

Bacteriorhodopsin Reviews Haupts, U., J. Tittor, and D. Oesterhelt, Closing in on bacteriorhodopsin. Ann Rev Biophys BioI/wi Strltcl. 1999,28:367399. Lanyi, J. K., and H. Luecke, Bacteriorhodopsin. CUlT Opil1 Strltct Bioi. 2001,11:415-419. Lanyi, J. K., X-ray diffraction of bacteriorhodopsin photocyde intermediates. l\!lol NJembr Bioi. 2004, 21:143-150. Neutze, R., E. Pebay-Peyroula, K. Edman, A. Royant, J. Navarro, and E. M. Landau, Bacteriorhodopsin: a highresolution structural view of vectorial proton transport. Biochim Biophys Acta. 2002,1565:144-167. Seminal Papers Henderson, R., and P. N. T Unwin, Three-dimensional model of purple membrane obtained by electron microscopy. Natllre. 1975,257:28-32. Oesterhelt, D., and W. Stoeckenius, Functions of a new photoreceptor membrane. Proc Natl Acad Sci USA. 1973, 70:2853-2857. Pebay-Peyroula, E., G. Rummel, J. P. Rosenbusch, and E. M. Landau, X-ray structure of bacteriorhodopsin at 2.5 angstroms h'om microcrystals grown in lipidic cubic phases. Science. 1997,277:1676-1681. Winget, G. D., N. Kanner, and E. Racker, Formation of ATP bv the adenosine triphosphatase complex from spinach chloroplasts reconstituted together with bacteriorhodopsin. Biochiln Biophys Acta. 1977,460:490-499. Photosynthetic Reaction Centers Reviews Deisenhofer, J., and H. Michel, Structures of bacterial photosynthetic reaction centers. Al/lnt Rev Cell Bioi. 1991, 7: 1-23. Nogi, T, and K. Mild, Structural basis of bacterial photosynthetic reaction centers. ] Biochem (Tokyo). 200 I, 130:319329. Vermeglio, A., and P. Joliot, The photosynthetic apparatus of Rhodobacter sphaeroides. Trends NJicrobiol. 1999,7:435440. Seminal Papers Deisenhofer, J., O. Epp, 1. Sinning, and H. Michel, Crystallographic refinement at 2.3 A. resolution and refined model of the photosynthetic reaction centre from RhodopseudOlnonas viridis. ] Iv101 Bioi. 1995,246:429-457. Deisenhofer, J., O. Epp, K. Miki, R. Huber, and H. Michel, Structure of the protein subunits in the photosynthetic reaction centre of Rhodopseudol1lcmas viridis at 3A. resolution. Nature. 1985,318:618-624. I3-Barrels Buchanan, S. K., [3-Barrel proteins from bacterial outer membranes: stnJCture, function and refolding. ClilT Opin Struct Bioi. 1999,9:455-461.

125

Schulz, G. E., [3-Barrel membrane proteins. Curl' Opin Struct Bioi. 2000, 10:443-447. Wimley, W. c., The versatile [3-barrel membrane protein. Curr Opin Struct Bioi. 2003, 13:404-411. Porins Achouak, W, T Heulin, and J.-M. Pages, Multiple facets of bacterial porins, FEMS Microbiol Lett. 200 I, 199: 1-7. Delcour, A. H., Solute uptake through general porins. Frontiers Biosci. 2003, 8:1055-1071. Dutzler, R., Y-F. Wang, P. J. Rizkallah, J. P. Rosenbusch, and T Schirmer, Crystal structures of various maltooligosaccharides bound to maltoporin reveal a specific sugar translocation pathway. Structure. 1996, 4: 127-134. Nikaido, H. Porins and specific channels of bacterial outer membranes. Mol Microbiol. 1992,6:435-442. Schirmer, T, General and specific porins h'om bacterial outer membranes. ] Struct Bioi. 1998, 12\: I 0 I-I 09. Iron Receptors Cao, Z., and P. E. Klebba, Mechanisms of colicin binding and transport through outer membrane proteins. Biochimie. 2002, 84:399-412. Clarke, T E., L. W Tari, and H. J. Vogel, Structural biology of bacterial iron uptake systems. CUlT Top Med Chem. 2001, 1:7-30. Ferguson, A. D., and J. Deisenhofer, TonB-dependent receptors - structural perspectives. Biochim Biophys Acta. 2002, 1565:318-332. Locher, K. P., B. Rees, R. Koepnik, A. Mitschler, L. Moulinier, J. Rosenbusch, and D. Moras, Transmembrane signaling across the ligand-gated FhuA receptor: crystal structures of fTee and felTichrome-bound states reveal allosteric changes. Cell. 1998,95:771-776. First Crystal Structures of Proteins Discussed in Chapter 5 Buchanan, S. K., B. S. Smith, L. Venkatramani, D. Xia, L. Esser, M. Palnitkar, R. Chakraborty, D. van del' Helm, and J. Deisenhofer, Crystal structure of the outer membrane active transporter FepA from Escherichia coli. Nat Struct Bioi. 1999,6:56-63. Cowan S. W., T Schirmer, G. Rummel, M. Steiert, R. Ghosh, R. A. Pauptit, J. N. Jansonius, and J. P. Rosenbusch, Crvstal structures explain functional properties of two E. coli pOl'ins. Nature. 1992,358:727-733. Deisenhofer, J., O. Epp, K. Miki, R. Huber, and H. Michel, Structure of the protein subunits in the photosynthetic reaction centre of Rhodopseudomonas viridis at 3A. resolution. Nature. 1985,318:618-624. Ferguson, A. D., E. Hofmann, J. W. Coulton, K. Diederiche, and W. Welte. Siderophore-mediated iron transport: crystal structure of FhuA with bound lipopolysaccharide. Science. 1998,282:2215-2220. Forst, D., W. Welte, T Wacker, and K. Diederichs, Structure of the sucrose-specific porin ScrY from Salmonella tvphinntriu111 and its complex with sucrose. Nat Struct Bioi. 1998, 5:37-45.

126

Kreusch, A., A. Neubuser, E. Schiltz, J. Weckesser, and G. E. Schulz, Structure of the membrane channel porin from Rhodopseudol11on£ls blastica at2.0 Aresolution. Protein Sci. 1994, 3:58-63. Pebay-Peyroula, E., G. Rummel, J. P. Rosenbusch, and E. M. Landau, X-ray structure of bacteriorhodopsin at 2.5 angstroms from microcrystaJs grown in lipidic cubic phases. Science. \997,277:1676-1681.

Bundles and Barrels Schirmer, T., T. A. Kellel-, Y-F. Wang, and J. P. Rosenbusch, Structural basis [01' sugar translocation through maltopOlin channels at 3.\ A resolution. Science. \995,267:512514. Weiss, M. S., A. Kreusch, E. Schiltz, U. Nestel, W. Welte, J. Weckesser, and G. E. Schulz, The stn.1Cture of porin from Rhodobactercapsulalus all.8 Aresolution. FEBS Lell. 199\, 280:379-382.

6 x

Functions and Families

10 3

50

100

150

200

250

50

100

150

200

250

Amino terminus

Q)

-0

.:: ;>.,

-5 ell

00 H -0

a

;>.,

::r:: -3

10

Residue number A hyclrcpil'ny plot pred,-ts . day r-sF r nit g leg'ons u! m "bru _ proteir s lik bactcriorhooopsln, s own I ere w th Its TM helices colored to maId he corr
Useu With permiSSion.

The functions of most membrane proteins are traditionally described as enzymes. transporters and channels, and receptors. although the demarcations of these groups are blurred. For example. ATPases that are ion channels and other active tr-ansport protei ns ("permeases") are studied as enzymes as well as transporters. Some receptors involved in signaling are also ty.-osine kinases; other receptors are gated ion channels. Membrane proteins can also be classified using data from bioinformatics, \vhich describe families of proteins, f'unctions of homologous domains. and evolutionary relationships. This chapter utilizes both approaches to look at the roles of membrane proteins before turning to tools for prediction of membrane protein structure and genomic analysis of membrane proteins. It starts by describing the general characteristics of membrane enzymes. transporters. and receptors, giving specific examples and briefly identifying their protein families.

MEMBRANE ENZYMES The enzymes found in membranes carry out diverse covalent catalytic functions. In addition to solute transport and signaling. membrane-bound enzymes partici-

pate in electron transpor-t chains and other redox reactions. as well as the metabolism of membrane components such as phospholipids and sterols. Many membrane enzymes require specific lipids or particular types of lipids for activity (see "Protein-Lipid Interactions" in Chapter 4). In addition. soluble enzymes may bind to the membrane periphery for catalysis involving substrates in the membrane. for efficient access to substrates that are passed along a series of bound enzymes to avoid dilution in the cytoplasm. or for regulation by modulation of their activities as described in Chapter 4. In all these cases. the heterogeneity and dimensionality of the mem brane affect the activi ty of the enzymes. Whether mem brane enzymes require lipids for their activity or catalyze reactions involving membranebound substrates. they are restricted to the available lipid or substrate near them in the membrane. This means the rate of enzyme activity (velocity) depends 00 the concentration of available lipid or substrate in the bilayer in the vicinity of the enzyme. not the concentration in the bulk solution. For kinetic treatment of these cases. the 'lmount of substrate available to the enzyme is not determined by its bulk concentration but by its concentration in the lipid bilayer, so it is described with surface concentration terms. either mole fraction 127

Functions and Families

128

BOX 6.1. Surface dilution effects brane. Thus the first step is the binding of the soluble enzyme to the micelles, followed by binding the substrate in the bilayer:

A description of the actions of lipid-dependent enzymes must consider both three-dimensional bulk interactions that occur in solution and two-dimensional surface interactions in the bilayer. Thus a kinetic model for surface dilution effects takes into account the concentrations of the required lipid in both phases, whether the enzyme is reconstituted into micelles or liposomes. The detergent-lipid mixed micelle is particularly amenable for study of surface dilution kinetics because it allows both the bulk concentration and the surface concentration of a lipid substrate to be varied. Then the surface concentration of the lipid is expressed in terms of its mole fraction, [lipid)l([lipid] + [detergent]). With a fixed number of lipids at several different micelle concentrations, the bulk concentration of lipids does not vary, but the lipid/micelle ratio changes. This is illustrated by the lipid-dependent binding of a phorbol ester by PKC in mixed detergent-lipid micelles (see Figure 6.1.1). PKC requires PS, as described in Chapter 4. When the level of Triton X-100 is increased as the amount of PS is held constant, the activity drops: the required PS is not as available to activate the enzyme. If the ratio of PS to Triton X-100 is held constant as the concentration of Triton X-100 increases, the activity stays the same because the mole percent of PS is maintained. Thus PKC shows a loss of activity at high detergent concentration (without addition of PS) because the surface concentration of lipid has been diluted. The surface dilution kinetic model was developed for cobra venom PLA2 (also described Chapter 4) in Triton X-100/PC mixed micelles. In the case of such peripheral proteins, the analysis includes the binding step that takes place in the three-dimensional bulk solution and results in restricting the interactions to the two-dimensional surface of the mem-

"'0

~~

o

l:>.

0

k,

k2

E+A .... EA

L,

Surface step

k3

EA + B .... EAB .... EA + Q,

L

2

L3

where E is the enzyme, A is the mixed micelle, EA is the enzyme-mixed micelle complex, B is the substrate, EAB is the catalytic complex, and Q is the product. (Note that the equation for the first step applies whether the enzyme binds nonspecifically, in which case A is the sum of the molar concentrations of the lipid and the detergent, or specifically to a phospholipid species, in which case A is the molar concentration of that lipid.) Once bound, the association between EA and B is a function of their surface concentrations, expressed in units of mole fraction or mole percent. For a water-insoluble integral membrane enzyme, the protein is delivered to the assay as a detergent-protein mixed micelle, which is likely to fuse with lipid micelles. In this case, E represents the concentration of the enzyme-detergent complex. The kinetic equation becomes v

=

Vmax[AJ[B]

-

Ks A Km B + Km B [AJ + [AJ[BI

}

where the dissociation constant, KsA = k.., /k" and the interfacial Michaelis constant, KmB = (k_ 2 + k3)/k 2 , are expressed in surface concentration units.

Constant mol percent phosphatidylserine

1.2

:J-

Bulk step

~ 1.0

l:>.

l:>.

.D ~~

eU

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~ 0.8

CIl

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:.a;Q -

s:;

2.... 'v

0.6

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o

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2 0.4

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Constant bulk concentration phosphatidylserine

""""'0

'-'

0.2 0

0

0.25

0.5

0.75 1.0 1.25 1.5 Triton X-IOO (WN %)

1.75

2.0

6.1.1. Surface dilution effect on the lipid-dependent enzyme, PKC. Redrawn from Gennis, R. B., Biomembranes: Molecular Structure and Function, Springer-Verlag, 1989, p. 228. © 1989 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Membrane Enzymes

129

([substrate]/{[substrate] + [total lipid]}) or mole percent (mole fraction x 100). Then the classical MichaelisMenten equation is applicable, with surface concentration units used for the substrate concentration (see Box 6.1). When an enzyme that loses activity upon dilution of the lipid is solubilized and reconstituted into micelles or liposomes, the amount of lipid remaining will affect its activity. For this reason, the extent of separation of lipid and protein components during solubilization of the membrane components (see Chapter 3) is critical in studies of membrane enzymes. Only a few high-resolution structures of integral membrane proteins that are classical enzymes are available (see Chapter 9) in addition to those involved in energy transduction and transport. HO'vvever, many membrane enzymes have been extensively characterized biochemically. In addition, some enzymes that are integral membrane proteins have extensive soluble portions that can be removed by proteolytic cleavage and crystallized. When the soluble portion of the enzyme carries out the catalysis, its structure reveals the binding site and catalytic groups to give a picture of the enzyme Function, even though it is missing the portion that anchors the enzyme in the membrane and perhaps plays a regulatory role. Diacylglycerol kinase (DGK) is an example of a well-characterized mem brane enzyme lacking a complete high-resolution structure. Some of the P450 cytochromes provide examples of membrane enzymes whose soluble portions have been crystallized and their structure solved. Both of these examples are enzymes that occur in mammals in numerous isoforms, different Forms of the enzymes that are encoded by difFerent genes. IsoForms, also called isozymes, are catalytically and structurally similar and are typically located in different tissues oFthe organism, where they respond to different regulators. Diacylglycerol Kinase

Diacylglycerol kinase carries out the reaction Diacylglycerol

+ MgATP

---+

Phosphatidic acid

+ MgADP with Michaelis-Menten kinetics and rates limited by substrate diffusion. Both the substrate and product of the DGK reaction are allosteric effectors and second messengers in signal transduction in mammals, which have 10 isoForms of DGK. Localized to the cytosol or the nucleus, the mammalian DGKs are peripheral proteins that dock on the membrane to access their substrate. Two of the isozymes are activated by both PE and cholesterol and inhibited by sphingomyelin when reconstituted in large unilamellar vesicles. All the mammalian isozymes appear to have specialized roles in signaling based on their diFFerent sites and pat-

terns of expression. Since lower organisms such as the nematode worm Cael10rhabditis elegans and the fruit fly Drosophila melal10gaster have only a few isozymes of DGK, and none has been detected in yeast, the mammalian isoforms appear to be involved in processes of development, neural networking, and immune functions that are essential in higher vertebrates. The E. coli DGK provides an example of a very well-characterized integral membrane enzyme whose stnlcture has not been determined at high resolution. Located in the inner membrane, DGK functions to replenish phosphatidic acid. The phosphatidic acid is needed in a surprising turnover of membrane phospholipid that provides the cell with osmoprotectants called membrane-derived oligosaccharides (MDOs). MDOs are made in the peri plasm under conditions of low osmolarity, when they can account For up to 5% of cell dry weight. Because they are water soluble and too large to diffuse through the porins, MDOs stay in the peri plasm and keep it from shrinking too much. MDOs contain six to 12 glucose units joined by ~-1,2 and ~-I ,6 linkages that are variously substituted with sl1-1-phosphoglycerol, phosphoethanolamine, and 0succinyl ester residues. The phosphoglycerol and phosphoethanolamine are enzymatically added from PG and PE, respectively, leaving diacylglycerol. It is the job of DGK to phosphorylate the diacylglycerol to return it to the phospholipid pool in the membrane. The activity of DGK in E. coli is determined by the rate of TM flip-flop supplying diacylglycerols from the outer to the inner leaflet of the plasma membrane. The smallest known kinase, E. coli DGK is a homotrimer of 13-kDa subunits, with three active sites at the subunit-subunit interfaces. It has been purified and reconstituted in detergent micelles and in phospholipid vesicles; in the latter, the enzyme activity depends on the structure of the surrounding lipids. Spectroscopic studies using FTIR spectroscopy and circular dichroism indicate that DGK is ~90% <x-helical, and topology predictions using fusions with f3-1actamase and ~-galactosidase(see below) indicate it has three TM helices (Figure 6.1). Many mutants of DGK have been made, including some that exhibit a remarkable increase in stability. Patterns of disulfide bond formation between singlecysteine mutants indicate the second TM segment from each subunit mediates trimerization by forming a bundle within the trimer. Indeed, addition of a free peptide corresponding to this TM segment interferes with trimer formation. In contrast, the first TM helix appears to be a passive membrane anchor since its replacement with polyaJanine produced an active enzyme. with the polyalanine (influenced by the flanking residues of the protein) spanning the membrane. Several highly conserved residues that are critical for activity « 10% activity when mutated) cluster near the N terminus in a

Functions and Families

130

A.

Periplasl11

Cytoplasln

B. TM-l: 31

50

KKKK-WINEAAFRQEGVAVLLAWIACWLDV-KKKK TM-2: 50

75

KKK-VDAITRVLLISSVMLVMIVEILNSAI-KKK TM-3: 95

121

KKKK-DMGSAAVLIAIIVAVITWAILLWSHFG-KKKK 6.1. A and B. Model of the predicted structure of DGK. The protein has two amphipathic helices at the membrane surface and three TM helices, whose sequences are given in (8). The encircled numbers on the model show where each given sequence begins and ends. From Partridge, A. W., et aI., J BioI Chem. 2003, 278:22056-22060. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

region (residues 9-28) shown by NMR to form a helix at the interface just before TMJ. NMR studies also indicate that TM J and TM2 are connected by a shon loop (residues 48-52) that is exposed to the aqueous environment. In spite of years of effort, no highly diffracting crystals of DGK suitable for x-ray crystallography have been obtained. However, nanocrystals of the enzyme give promising results with solid-state NMR. P450 Cytochromes

P450 cytochromes are a ubiquitous superfamily of heme-containing monooxygenases that are named for their absorption band at 450 nm. They are involved in metabolism of an unusually wide range of endogenous and exogenous substances. They panicipate in the metabolism of steroids, bile acids, fatty acids, eicosanoids, and fat-soluble vitamins, and they convert lipophilic xenobiotics (foreign compounds) to more polar compounds for funher metabolism and excretion. P450 cytochromes catalyze hydmxylation of an organic substrate, RH, to R-OH with the incorporation of one oxygen a tom of O 2 , while reducing the other oxygen atom to H 2 0. Their source of reducing power is NAD(P)H, with electrons either donated directly fmm a flavin-containing reductase (class II) or shuttled to the P450 by smaJI soluble electron carrier proteins (class I; Figul-e 6.2). Some self-sufficient P450s in bacteria have

been found to contain heme, flavin, and iron-sul fur centers in one polypeptide. In eukaryotes, most P450 cytochromes are bound to the mitochondrial inner membrane or to the ER. The P450s in the ER are integral membrane pmteins, each bound by a single N-terminal TM domain. Truncation of the N-terminal domain is not sufficient to express soluble protein for crystallization and must be accompanied by several point mutations to disrupt a pel-ipheral membrane-binding site. Even so, detergent is needed to prevent aggregation. High-resolution structures of such constructs show the large soluble domain is a triangular prism shape with ~-sheets along part of one side and ex-helices forming the rest (see Figure 6.2). Structures of these P450 catalytic domains in the presence and absence of substrates reveal that dramatic conformational flexibility must be needed for binding such a variety of compounds. About half of the 57 P450 cytochmmes in the human genome metabolize endogenous compounds, while many of the rest metabolize drugs and other xenobiotics. Some plants have around 300 or more P450s. The genes are classified into families based on sequence identity: to the root symbol CYP is added a number for the family (one of more than 200 gmups with >40% sequence identity), followed by a letter for subfamilies (having> 55% identity), followed by a number for the gene. For example, sterol 27-hydmxylase

Class I

Class II

Heme

e

Fe-S

Heme

e

FAD/NAD(P)H

FMN/FAD/NADPH

6.2. Examples of the two classes of P450 cytochromes. The classes are distinguished by their redox partners. Class I is represented by the P450 system from Pseudomonas putida with P450cam (with heme), putidaredoxin (with Fe-Sl, and putidaredoxin reductase (with FAD/NAD(P)H). Class II is represented by P450BM3 from Bacillus megaterium (with heme) and cytochrome P450 reductase from rat liver (with FMN/FAD/NADPH). The high-resolution structures for P450cam and P450BM3 have been solved for proteins lacking their TM segments. From Li, H., and 1. L. Poulos, Curr Top Med Chern. 2004, 4: 1789-1802. ~, 2004. Reprinted by permission of Bentham Science Publishers Ltd.

is CYP27A and vitamin D 24-hydroxylase is CYP27B, because their amino acid sequences are >40% identical (placing them both in CYP27) but <55% identical. If there were another P450 cytochrome in the CYP27A class, then it vvould be CYP27A2. The presence of many gene clusters, each containing up to 15 CYP genes, in most genomes suggests that the diversity of P450 cytochromes arose from many gene duplications in addition to likely gene amplifications and lateral transfers.

TRANSPORT PROTEINS

Transport of molecules across the bilayer is obviously an important function of membrane proteins and utilizes a variety of mechanisms. These mechanisms are defined and classified by both the stoichiometry and the energetics of the transport process. The definitions are essential for understanding the systematic classification of all membrane transport proteins. Uniport is the movement of one molecule at a time across the membrane, and cotransport is the tightly coupled movement of more than one substrate. SympOrl is the tightly cou-

pled transport of two different molecules in the same direction, ancl antiport is the tightly coupled transpOl-t of two different molecules in opposite directions. When symport and antiport involve transport of ions, they may be electroneutral - resulting in no net transfer of charge - or electrogenic - creating (\ charge separation across the membrane. Proteins involved in transport carry out either active or passive transport. Active transporters enable a cell to accumulate solutes against electrical and/or concentration gr«dients by making use of energy sources to "pump" them thermodynamically "uphill," whereas passive transporters allow the "downhill" flow of solutes across the membrane until their electrical and/or concentration gradients are dissipated. Like the specific porins described in the last chapter, passive transport proteins provide saturable pathways that are susceptible to competitive inhibition. The classic example is the glucose transporter in erythrocytes, a glycoprotei n predicted to have 12 TM ex-helices (Figure 6.3). Unlike the porins, the channel of the glucose transporter does not leak small molecules or ions, so it is described as a "gated pore" that opens alternatively to one side of the membrane or the other depending on its conformation

Functions and Families

132

'"A. I I

o 1.0

1 max = 1.0

10-6

X

m

Transport Classification System

M

om ,eo-'

The transport classification (TC) system endorsed by the International Union for Biochemistry and Molecular Biology describes five broad classes: channels and pores, carriers or porters, primal)' active transporters, group translocators, and TM electron carriers, in addition to lists of accessory factors and incompletely char'acterized transport systems (Table 6.1 and http://www.tcdb.org). About 400 families of transport proteins are now included in the TC system, divided as follows:

/

x I

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Vl

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0.5

::E

5 ~

o

()

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....,eo 2

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6

8

10

[Glucose] mM B.

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Class 1: The channels and pores are proteins (and peptides) that allow the relatively free flow of solute through the membrane. This class is divided into five subclasses: cx.-helical protein channels; (3-barreJ

Glucose + 10 mM 6-0-benzylD-galactose

.,,..,..,.

,.,.,.

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'

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-l/KM

1/[Glucose]

6.3. Glucose transport into erythrocytes. A. A plot of the flux for glucose uptake by erythrocytes at 5°C as a function of the external concentration of glucose shows the passive uptake of glucose follows Michaelis-Menten kinetics, with the flux (J. in units of mM·cm·sec') in place of the velocity of a reaction. The saturable curve indicates that the flux utilizes a carrier. B. The flux of glucose can be inhibited by 6-0-benzyl-o-galactose, and the kinetic analysis indicates it is competitive inhibition, providing evidence for a binding site on the carrier. Redrawn from Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, pp. 728729. © 2004. Reprinted with permission from John Wiley & Sons, Inc.

(Figure 6.4). This type of conformational change is also seen in active transporters, such as LacY, the lactose permease (see Chapter 10). Active transport is further divided into primary processes, which are directly driven by hydrolysis of ATP (or another exergonic chemical reaction), and secondary processes, which carry out symport 01- antiport coupled to ion gradients made by primary active transporters like ATPases. These distinctions are part of the functional basis for c1assi fication of all transporters into hierarchies of families and superfamilies.

Inside

6.4. The model for conformational changes involved in transporting glucose via a gated pore. Step 1, glucose binds from the outside; step 2, the protein undergoes a conformational change, changing the exposure of the binding site to the inner surface; step 3, glucose is released to the cytoplasm; step 4, the protein returns to the original conformation. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed, w. H. Freeman, 2005, p 394. © 2005 by W. H. Freeman and Company. Used with permission.

Transport Proteins

133

I.A. a-Helical protein channels I.B. ~-Barrel protein porins

I

I. Channels! Pores

~

I.e. Toxin channels

~

1.0. Nonribosomally synthesized channels I.E. Holins

2.A. Protein porters

I

2. Electrochemical Potential-driven Transporters

~

2.B. Nonribosomally synthesizcd porters 2.e. Ion gradient-driven energizers

3.A. P-P-bond hydrolysis-driven systems 3.B. Decarboxylation-driven systems

I

3. Primary Active Transporters

~

3.e. Methyl transfer-driven systems 3.0. Oxidoreduction-driven systems 3.E. Light absorption-driven systems

4. Group Translocators

4.A. Phosphotransfer-driven systems

5.A. Two-electron transfer calTiers 5. Transmembrane Electron Carriers 5.B. One-electron transfer carriers The five main classes are listed on the left, with subclasses provided on the right. Source: Busch, W., and M. H. Saier, Jr., Cril Rev Biochem Mol Bioi. 2002, 287-227. © 2002. Reproduced by permission of Taylor & Francis Group, LLC., http://www.taylorandfrancis.com.

a

porins (see Chapter 5); channel-forming toxins, including colicins, diphtheria toxin, and others (see Chapter 4); nonribosomally synthesized channels, such as gramicidin and alamethicin (see Chapter 4); and holins, vvhich function in expor1 of enzymes that digest bacterial cell walls in an early step of cell lysis. Class 2: The carriers, also called porters, form complexes by binding their solll tes before transporting them across the bilayer by the secondary processes of symport or antiport. For this reason, class 2 is also called electrical potential-driven transporters. A very large family of porters in this class is called the major facilitator superfamily (MFS) and

is exemplified by the lactose permease (see Chapter 10). Class 2 also contains the ionophores like valinomycin and nigericin, which are nonribosomally synthesized ion carriers, as well as the TonB family of proteins involved in transferring energy to the outer membrane of Gram-negative bacteria (see Chapter 11). Class 3: The class of primary active transporters includes the ATPases and the ATP-binding cassette transporters described next. Other examples of primary active transporters are those driven by oxidoredllction reactions and those driven by light, including the photosynthetic reaction center (see Chapter 5).

Functions and Families

134

Class 4: The group translocators provide a special mechanism for the phosphorylation of sugars as they are transported into bacteria. described below. Class 5: The TM electron transfer carriers in the membrane include two-electron carriel-s. such as the disulfide bond oxidoreductases (DsbB and DsbD in E. coli) as well as one-electron carriers such as NADPH oxidase. Often these redox proteins are not considered transport proteins.

CD

Cells have multiple transport systems to take up many nutrients. and the families of uptake systems employed for any nutrient type are not limited to one class. For example. the uptake of sugars is carried out by nine families of ABC transporters. 20 families of secondary caITiel-s. seven families of porins. and six families of group translocation systems. In addition to descriptions of transport functions. the TC system derives evolu tionary relatedness from sequence inrormation obtained using the tools of bioinformatics described below that allow in.vestigators to compare primary sequences. predict topologies. and an.alyze genomes. Superfamilies of ATPases

The TC system has t"vo superfamilies or ATPases. one for the P-type ATPases that 3r'e found in plasma membranes and include the Na+ -K+ ATPase (see below) and the Ca 2+ pump (see Chapter 10). The other superfamily consists of three other types of ATPases: F-type. such as the mitochondrial and bacterial ATP synthases (see Chapter 11); A-type, which transports anions such as arsenate and is mainly found in Archaea; and Vtype, which maintains the low pH of vacuoles in plant cells and lysosomes, endosomes. the Golgi. and secretory vesicles of animal cells. Both the subunit compositions and their protein sequences indicate that the Atype is more similar to the V-type ATPases than to the F-type. The Na+-K+ ATPase, or the Na+ -K+ pump. transports two K+ in and three Na+ out for every ATP hydrolyzed, establishing gradients that are critical to osmotic balance in all animal cells and to the electrical excitabi lity of nerve cells. as well as driving the uptake of glucose and amino acids. It consists of a and 13 subunits, with the ion-binding sites and catalytic site on the a subunit. which is predicted to have eight TM a-helices. ATP phosphorylates an Asp residue in the highly conserved sequence DKTG only in the presence of Na+; the aspartyl-phosphate thus produced is hydrolyzed only in the presence of K+. giving strong evidence for two conformational states. Thus the mechanism of coupling active transport with ATP hydrolysis involves shifting from the phosphorylated form with high affinity for K+ and low affinity for Na+ to the dephosphorylated rorm with high affinity for Na+ and low affinity for K+

Inside

Outside

6.5. The proposed mechanism of Na+ and K+ transport by the Na+ -K+ ATPase. Phosphorylation and dephosphorylation of the ATPase trigger conformational changes that determine the direction the channel opens. The dephosphorylated ATPase (EI) has an inward-facing high affinity binding site for Na+ . and the phosphorylated ATPase (EII-P) has an outward-facing high affinity binding site for K+. Thus EI binds 3 Na+ and ATP from inside(l); allowing phosphorylation to EI-P . 3 Na+(2). This is followed by the conformational change to EII-P, releasing Na+ to outside(3). Then EII-P binds 2 K+ from outside(4). and hydrolysis of phosphate yields Ell· 2K+(5}; the conformational change releases K+ inside and the ATPase reverts back to EI(6) ready for step (1). Redrawn from Nelson, D. L.• and M. M. Cox. Lehninger Principles of Biochemistry, 4th ed .. W. H. Freeman, 2005, p. 399. ,E) 2005 by W. H. Freeman and Company. Used with permission.

(Figure 6.5). This electrogenic process helps create the typical TM poten tial of -50 to -70 m V (inside negative) across the plasma membrane of most cells. The Na+K+ ATPase is inhibited by digitalis in the well-known treatment for congestive heart failure. ABC Transporter Superfamily

An important class of ATP-dependent transporters is the supel-family of ABC transporters. named for their ~TP-2.inding (:assettes. which are nucleotide-binding

Transport Proteins

135

Pcriplasm Inner membrane

(3)

(b)

_~""",\..,..-~P~e_ri plasm Inner membrane

(
(e)

(f)

6.6. Organization of the structures of ABC transporters. The subunit composition of the ABC transporter system varies in different organisms. The two nucleotide-binding domains (NBDs) (white) and two TM domains (shaded) may be four separate subunits or fused into one, two, or three proteins as shown. Redrawn from Higgins, C. F, Res Microbial. 2001, 152:205-210 © 2001 by Elsevier Reprinted with permission from Elsevier.

domains. ABC transporters are found in large numbers in animals, plants, and microorganisms: the genome of E. coli has 80, while the human genome has 49. (See http://nutrigeneAt.com/humanabc.htm.) Some ABC transporters function in uptake and others, in efflux. Some are very specific for their substrates while others are quite promiscuous. The first ones to be identified functioned in uptake of amino acids (histidine), peptides, sugars (maltose), and vitamin B 12 • ABC transporters also export cell wall polysaccharides, participate in cytochrome-c biogenesis, and secrete cellulases, proteinases, and toxins. All have two nucleotidebinding domains and two membrane domains, which are separate subunits in Archaea and Eubacteria and are usually fused together in higher organisms (Figure 6.6).ln addition, the ABC transporters that function in uptake have a substrate-specific domain or protein on the external side (the periplasmic binding proteins in Gram-negative bacteria). The membrane domains typically have 12 TM helices in either a single peptide or two subunits. The nucleotide-binding domains of ABC transporters are highly conserved, whether they are separate subunits or are fused to the TM domains. Several of them have been crystallized either as aqueous proteins or water-soluble fragments of transporters, and x-ray structures have been solved for two of the intact transporters (see Chapter 11). As expected, the various NBD domains show strong structural similar-ities. Each NBD domain contains an ATPase subdomain with similarities to the F 1 ATPase (see Chapter 11) and a helical subdomain specific to the ABC transporters (Figure 6.7A). The ATPase subdomain contains typical Walker A and Walker B motifs found in proteins that hydrolyze ATP and has a f3-sheet region that positions the base and ribose moieties of the nucleotide. The helical subdomain contains the signature motif of ABC transporters,

LSGGQ. In the dimel~ the Walker A motif from one subunit binds the same ATP as the LSGGQ from the other subunit, thus the two NBD domains make an "ATP sandwich." Each round of ATP hydrolysis involves three conformations of the NBD domains: a nucleotidefree conformation, an ATP-binding conformation, and a more open conformation after the hydrolysis reaction has produced ADP plus P j • Recently the MalK protein, the NBD of the maltose transport system, was crystallized in all three conformations (Figure 6.7B). The movements of the NBD domain are transmitted to the TM regions of the membrane transporter to drive active transport. The substrate-specific components of ABC transporter systems in the periplasm of Gram-negative bacteria are obligatory to the transport of their substrates. High-resolution structures of several of the periplasmic binding proteins with and without bound substrate indicate large conformational changes typically occur upon binding. Each protein has two lobes that close over the substrate when it binds, giving a very high substrate affinity (micromolar K:is). For some transport systems, mutants of the inner membrane transporters have been character-ized in which delivery of substrate by these soluble binding proteins is no longer obligatory. Now high-resolution structures of intact ABC transporters have been obtained for the vitamin B n transporter (BtuBC) from E. coli and the drug efflux protein Sav 1866 [Tom Staphylococcus aureus (see Chapter 11). Many of the ABC transporters in humans have medical significance. The CFTR protein, the cystic fibrosis transmembrane conductance regulator, is a CI- channel that is defective in most cases of cystic fibrosis (see Chapter 7). Both Savl866 and CFTR are closely related to the multidrug resistance protein I (MDRl, also called p-glycoprotein) that confers resistance of tumor cells to chemotherapy. Group Translocation

A different mechanism of coupling to an exergonic reaction is utilized for group translocation of sugars in bacteria, in which the sugar substrates are phosphorylated during the transport process. This unusual group of transport systems constitutes class 4 in the TC system. The best-characterized example is the uptake of glucose, fructose, mannose, and other sugars by the phosphoenolpyn.lvate-dependent phosphotransferase system (PEP PTS) of E. coli (Figure 6.8). The two cytoplasmic proteins involved in transfer of the phosphoryl group from PEP, E1 (enzyme 0, and HPr (histidine-containing phosphocarrier protein) are used in transport of all the sugar substrates of this system. Uptake of each sugar uses a specific TM component called Ell, which in some cases has one or more

B.

Resting state

Pr'e-hydrolysis state

+ATP

,J

ADA

Post-hydrolysis state

Hydrolysis

Pi

~

6.7. The MalK dimer, an example of an NBD domain of an ABC transporter. A. The x-ray structure of the NBD domains of the MalK dimer shows the ATP-binding site. Each NBD domain has two subdomains, an ATPase subdomain (green) and a helical subdomain (cyan). The conserved segments shown are the Walker A motif iyVA, red), Walker B motif (WB, blue), LSGGO motif (magenta), and the 0 loop (yellow) that connects the ATP-binding region to the helical subdomain, as well as to the TM portions of the ABC transporter. The bound ATP is shown in ball-and-stick model. The regulatory domain of MalK has been omitted for clarity. From Davidson, A. L., and J. Chen, Annu Rev Biochem. 2004, 73:241268. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org. B. The conformation of the MalK dimer varies from the resting state (with the two NBDs separated from each other), the ATP-bound state (closed), and the post-hydrolysis state (open). Each subunit has an NBD domain (green and blue or cyan) and a regulatory domain (yellow). The Walker A motif (red) shows where the nucleotide (ball-and-stick model) binds. From Lu, G., et aI., Proc Natf Acad Sci USA. 2005, 102:17969-17974. © 2005 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

Transport Proteins

137 Outside Lactose Glycerol

+ ATP

Glvcerol-3-phosphate

+ ADP

PEP Py,u," "

X

EIXHPr-py ..J\--..

El - P

HP,

+ H+

Glucose

EllA''' - P

cAMP + PP; 6.8. The PEP PTS for glucose in E. coli. Group translocation utilizes a combination of shared and specialized proteins, as exemplified by the PTS for glucose. El and HPr are the shared cytoplasmic proteins utilized for transfer of phosphoryl groups. EIIAg lc is a specialized cytoplasmic protein that phosphorylates the glucose as it enters the cytoplasm and is sensitive to catabolite repression. EIIBCg1c is the TM glucose transporter. In the cytoplasm EI is phorphorylated by PEP; the phosphoryl group is transferred to HPr, and then to a specific EllA. Then EllA phosphorylates the membrane protein(s), in this case EIIBC, which then phosphorylate the substrate as it enters. Variations in the number of Ell proteins are seen with different substrates. LacY is shown transporting lactose, which plays an inhibitory role in uptake of glucose by the PEP PTS. Redrawn from Voet, D., and J. Voet, Biochemistry, 3rd ed., John Wiley, 2004, p. 745. © 2004. Reprinted with permission from John Wiley & Sons, Inc.

separate cytoplasmic subunits, called EllA and EIlB. For example, the TM component of the PTS for glucose is EIIBC gk , which transports and phosphorylates glucose and transfers it to the cytosolic component. The cyLosolic component, ElIAg k , is sensitive to catabolite repression. Thus transfer of the high-energy phosphoryl group in PEP to glucose drives its uptake as glucose6-phosphate. Many components of the PEP PTS have been charactelized thoroughly, and several of the soluble proteins have high-resolution structures. Although the EI and HPr components are common elements that are shared by transport systems for different substrates, different forms of them are usually present, encoded by more than one gene. For example, the PTS f
acids, while Na+ symport is used in E. coli for uptake of melibiose, glutamate, and other amino acids, as weJJ as in the intestinal epithelium for uptake of glucose and some amino acids (Figure 6.10).

Symporters

6.9. Symport of lactose and protons in E. coli. The proton gradient made by the respiratory chain or other proton pumps is used to drive the uptake of lactose by the lactose permease. When cells are treated with CN- to inhibit the energy-yielding oxidation pathways that support the proton pump, active transport of lactose is abolished. Under this condition, the lactose permease carries out passive transport. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W H. Freeman, 2005, p. 404. © 2005 byW. H. Freeman and Company. Used with permission.

Secondary active transport that uses ion gradients typically involves cotr
Lactose transporter

Proton pump (inhibited by CN-)

Lactose (outside)

Fuel

CO 2

Lactose (inside)

Functions and Families

138 Basal surface

Apical surface

Intestinal lumen Micmvilli •

0

o

Glu

~'~

Na ;- -glucose sympol-tcl-

(driven by high extracellular [Na+]) 6.10. Symport of glucose and Na+ ions in the intestinal epithelium The Nat gradient made by the Na+ -K+ ATPase provides the driving force for glucose uptake from the intestinal lumen A passive glucose uniporter exports glucose to the bloodstream. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W H. Freeman, 2005, p. 405. © 2005 by W. H. Freeman and Company. Used with permission.

\J\lhen symport uses Na+, it is driven by the Na+ gradient created by [he Na +-K+ ATPase. The energy used can be calculated from the chemical potential and electrical potential of the ion gradient. For example, for the Na+ gradient, 6G = l1RT In ([Na+]in/[Na+]OLll)

+ 11:'16£.

In intestinal Na+-glucose symport, n = 2 for both the concentration and electrical terms, as two Na+ ions enter per glucose. Using typical values for 6£ (-50 mV), [Na+]in (12 mM) and [Na+]nUI (145 mM), the 6G for glucose transport is -22.5 kJ, enough to pump glucose inside until its concentration is 9000 times the external concen u-ation. Antiporters

Antiport is important for ion exchange across many biological membranes. Forexample, the Na+/H+ antiporter in the plasma membrane exchanges extracellular Na+ for intracellular H+ and is activated in response to mitogenic agents such as epidermal growth factor. The slight increase in cytoplasmic pH produced by this exchanger may activate other enzymes in the mitogenic response. A prominent anti porter in the erythrocyte membrane is the anion exchanger called Band 3, \vhich is about 25% of the total protein in that membrane. It carries out a one-to-one exchange of CI- and bicarbonate (HC03l to facilitate the removal of CO 2 from respiring cells. (Carbonic anhydrase in the erythrocyte converts waste CO 2 to HCOj', which then circulates to the lungs where it reenters erythrocytes to be converted back to CO 2 and exhaled.) Since the exchange of anions is oblig-

atory, the net flow of transport depends on the sum of the chloride and bicarbonate concentration gradients. The membrane portion of the Band 3 protein is predicted to have 12 or 14 TM hel ices, and the cytoplasmic portion interacts with cytoskeletal components while the extracytoplasmic portion carries the carbohydrate antigenic determinants for several blood groups . Besides the anion exchanger in the erythrocyte, humans have two other anion exchangers that are closely related to Band 3. Similar exchangers are also found in plants and microorganisms. Not all antiporters are electroneutral; for example, the ADP/ATP carrier, the most abundant protein in the mitochondrial inner membrane, exchanges one ATp 4 - (exiting) for one ADp 3 - (entering). This electrogenic exchange is driven by the inside-negative potential across the mitochondrial membrane and utilizes one third of the energy stored in the electrochemical proton gradient generated by the respiratory chain! To meet the energy needs of the cell, it facilitates the exchange of adenine nucleotides with a turnover number of 500/min. Two natural inhibitors bind to alternate sides of the transporter, indicating that it has two conformations that allow its adenine nucleotide binding site to face in or out. The high-resolution structure of the ADP/ATP translocator in the presence of one of these inhibitors shows a bundle of six TM ex-helices forming a basket with the inhibitor inside (see Chapter 10). The ADP/ATP can-ier belongs to the mitochondrial carrier family (MCF), whose functions ensure that metabolites (such as pyruvate, tricarboxylates and dicarboxylates of the Krebs cycle, carnitine, and citrulline) as well as nucJeotides and reducing power are exchanged between the cytosol and the mitochondrial matrix. The family also includes uncoupling protein of brown adipose tissue, which carries protons back into the matrix, allowing respiration to generate heat, and is involved in type II diabetes. The transporters in this family are in the inner membrane of mitochondria and share 20% sequence identity and a common threefold symmetry in their structure. New members of the family are readily identified by the MCF signature, which consists of three copies of the sequence PX[D,E]XX[K,R], along with the threefold sequence repeats that generate the overall fold. Mitochondria in different eukaryotes contain from 35 to 55 different mitochondrial carriers; the human genome encodes 48. Ion Channels

Most eukaryotic membranes contain selective ion channels that are unlike the ion pumps driven by the hydrolysis of ATP. They are characterized by ion specificity, high fluxes - up to 10 8 ions/sec - and gating (opening and closing) in response to ligands or voltage detected by patch clamp measurements (see Chapter 3).

Membrane Receptors

TABLE 6.2. Families of some representative receptors in eukaryotes Immunoglobulin superfamily - many diverse functions T cell receptor (a, 13 subunits) Major histocompatibility complex II MHC (a, 13 subunits) Lymphocyte function-associated antigen-3 (LFA-3) CD2 (T cell LFA-2) Immunoglobulin A (lgA)/lgM receptor IgG Fe receptor High-affinity IgE receptor (ex subunit) Surface immunoglobulins (heavy, light chains) N-CAMs (neuronal cell adhesion molecules) Myelin-associated glycoprotein Integrins - bind to extracellular matrix and adhesion proteins Fibronectin receptors Vitronectin receptors Platelet glycoprotein complex (lib/lila) Leukocyte adhesion proteins (LFA-1, Mac 1) T cell very late antigens (VLA family) Mitogen/growth factor receptors with tyrosine kinase activity - stimulate cell growth Epidermal growth factor (EGF) receptor Platelet-derived growth factor (PDGF) receptor Insulin receptor Insulin-like growth factor-1 (IGF-1) receptor Colony stimulating factor-1 (CSF-1) receptor Neurotransmitter receptors/ion channels receptor-operated channels Nicotinic acetylcholine receptor (nAChR) y-Aminobutyric acid (GABA) receptor Glycine receptor Receptors that activate G proteins I3- Adrenergic receptors (131, (32) <x-Adrenergic receptors (al, az) Opsins (rhodopsin) Muscarinic acetylcholine receptors (M 1, M2) Miscellaneous AsiClloglycoprotein receptors Low-affinity (lymphocyte) IgE receptor (unknovvn function) Insulin-like growth factor-2 (IGF-2) receptor Cation-independent mannose-6 phosphate receptor Cation-dependent mannose-6-phoshate receptor (extracytoplasmic domain) Source: Gennis, R. B., Biomembranes: Molecular Structure and Function, Springer-Verlag, 1989, p. 324. ~) 1989 by Rober1 B. Gennis. Reprinted by permission of the author.

High-resolution structures for a number of ion channels reveal their basis o[ selectivity and transport mechanisms, exemplified by the K+ channels described in Chapter 10. Ion channels that are involved in the passage of signals in nerves and muscle cells are also receptors for agonists and antagonists that control their activity (see below).

MEMBRANE RECEPTORS

Membrane receptors are integral proteins that function in information transfer by triggering a response to their

139

binding of ligands. Receptors have many diverse [unctions, listed in Table 6.2. Many receptors are involved in cell surface interactions, while some trigger endocytosis or recycling of membrane components. Among the receptors "vith enzyme activity are many receptors involved in signaling, such as the insulin receptor. The insulin receptor is an ex2 ~2 tetramer that responds to insu lin by au tophosphorylation of three Tyr residues on each ~ subunit, activating its protein kinase activity to start a signal cascade. Nicotinic Acetylcholine Receptor (Neurotransmitter Receptor Superfamily)

The most thoroughly studied receptor is the nicotinic acetylcholine receptor of muscle fiber. It is one of two structurally different classes of acetylcholine receptors; the other is the muscalinic acetylcholine receptor, a rhodopsin-like bundle of seven TM ex-helices. When the nicotinic acetylcholine receptor binds acetylchol ine, a neurotransmitter released by the motor neuron, it opens channels for Na+, Ca 2 +, and K+, triggering a depolarization of the membrane that causes the muscle fiber to contract. After several milliseconds the channel closes. The quater'nary structure of the muscle receptor is ex213y6, and each highly homologous subunit has four TM ex-helices (M J -M4). The channel is formed from the M2 helices from each of the five subunits, which are thought to twist in response to acetylcholine binding, turning bulky leucine side chains out of the pore interior to open the channel. The t\NO ex subunits have binding sites for acetylcholine, which are evident as cavities located in the extracellular domains in the EM image of the receptor from the electric organ from the Torpedo ray (Figure 6.11), Cations enter the central vesti bule of the tunnel and exit at narTOW openings close to the cytoplasmic membrane surface. The nicotinic acetylcholine receptor is in a superfa mily with receptors for other neurotransmitters, such as y-aminoblltyric acid and serotonin, \",hich are also pentameric ligand-gatee! ion channels (LGICs). Each subunit has a large extracelJular N-terminal domain, four TM segments, and an extracellular C terminus. A related superfamily inclue!es the excitatory glutamate receptors and ionotropic ATP receptors, \",hich are also LGICs, but their protein fold differs in that they have three TM segments and an intracellular C terminus. Together these superfamilies have over 200 entries in the LGIC database (http://www.ebi.ac.ukJ compneur-srv/LGICdb/LGICdb.php). While there is not yet a crystal structure for any of these neurotransmitter receptors, the crystal structure of the acetylcholine binding protein from the snail LYl11naea stagnalis provides insigh t into the strllctu re and fu nction of the extracellular domain of nicotinic acetylcholine receptor. Although the overall homology is weak «25% identity),

Functions and Families

140

B.

A.

C.

Ci-y

Cia

////

'/////);

/////

~I I

"caB

. ?Jj.

D.

Rapsyn 6.11. A through C. Images of the nicotinic acetylcholine receptor from Torpedo ray postsynaptic membranes at 4.6 A resolution achieved vvith electron microscopy. The cross-sections run through the pentameric structure as shown by the dotted line in the insets. The asterisk denotes the cavities for acetylcholine binding, and the arrow points to an opening for the ion channel to the cytoplasm. D. Cartoon showing the opening of the channels in response to acetylcholine binding. From Miyazawa, A., et al., J Mol Bioi. 1999,288:765-786. © 1999 by Elsevier. Reprinted with permission from Elsevier.

the homology in the regions of the ligand-binding sites is higher and the overall fold is consistent with EM data on the Torpedo acetylcholine receptor. G-Protein Coupled Receptors

The largest and most diverse group of receptors involved in signaling is the G-protein coupled receptors (GPCRs). The majority of these receptors respond to ligand binding by activating heterotrimeric guanine nucleotide binding proteins (G proteins) on the cytoplasmic surface of the membrane, while a few open ion channels directly. The G proteins transmit and ampl ify the signals by triggering changes in the concentrations of second messengers, such as cAMP (Figure 6.12). GPCRs respond to many intercellular messenger molecules as well as sensory messages. The more than

1000 GPCRs in humans include receptors for nucleotides, neurotransmitters (acetylcholine, dopamine, histamine, and serotonin), prostaglandins and other eicosanoids, and many hormones, as well as olfactory and gustatory receptors. With a common protein fold, the GPCRs are caJled serpentine receptors because their seven TM ex-helices "snake" across the membrane. The first member of the GPCRs to have its structure solved at high resolution is rhodopsin (described in Chapter 9). Additional structures are highly desirable to aid in drug design, as nearly three quarters of modern pharmaceuticals are targeted at GPCRs. The GPCRs form a superfamily that is divided into six families with no statistically significant sequence similarity between them (see www.gpcr.org/7tm/). Their seven-TM topology was attributed to convergent evolution until some of the bioinformatics tools described next enabled searches

Bioinformatics Tools for Membrane Protein Families

141

Hormone

Adenylate cyclase

G-protein (a)

GDP

P1

A

C)

(,

GTP Cellular responses -

cAMP + PPi

6.12. Interaction of a receptor with a G protein to stimulate the production of cAMP. The activation/deactivation cycle for hormonally stimulated adenylate cyclase (AC) starts with inactive AC and guanosine diphosphate (GDP) bound to the G protein. When hormone binds to its receptor (a). the hormone-receptor complex stimulates the G protein to exchange GDP for guanosine triphosphate (GTP; b). The GTP-G protein complex binds to AC, activating it to produce cAMP (c). When the G protein catalyzes hydrolysis of GTP to GDP. it dissociates from AC. inactivating it. Redrawn from Voet. D .. and J. Voet. Biochemistry, 3rd ed. John Wiley, 2004, p. 674. © 2004. Reprinted with permission from John Wiley & Sons, Inc.

for distant homology and new members of the superfamilies.

BIOINFORMATICS TOOLS FOR MEMBRANE PROTEIN FAMILIES

Because purification and crystallization of membrane proteins are complicated by the presence of lipids and detergent (see Chapters 3 and 8), the number of high-resolution structures of membrane proteins, while growing rapidly, is still relatively small. With under 200 structures of membrane proteins in the Protein Data Bank (see Box 6.2), analysis of the wealth of genomic information available relies on methods for interpretation and comparison of the approximately 285,000 identified protein sequences in Swiss-Prot, a large portion of which are likely membrane proteins. The planar dimensionality and the hydrophobicity of the bilayer simplify the application of bioinformatics to membrane proteins to give topology models describing the number and orientation of TM segments. Classification of membrane proteins into families has relied on both sequence homologies and common topologies or folds. An early advance was the construction of hydropho-

bicity plots to identify potential TM ex-helices based on occurrences of ~18 or more contiguous nonpolar amino acids. The problem then became how best to assign values for the hydrophobicity of the different residues, with numerous scales sometimes offering contrasting results. Application of various statistical tools brought large improvements in prediction methods, and recently methods were added to identify f)-barrels by their amphipathic sequences. The rest of this chapter looks at these developments in some detail. Structural predictions of integral membrane proteins have evolved, from fairly simple computations using physical data to the increasingly sophisticated algorithms that are the tools of bioinformatics today. At the heart of the searches for structural similarities that are the basis for determining families and superfamilies is the ability to predict TM segments fTOm the primary structure data. Predicting "rM Segments

With the vast majority of membrane proteins expected to be bundles of ex-helical TMs, the prediction of integral membrane protein structure has focused on

Functions and Families

142

BOX 6.2. Bioinformatics basics Bioinformatics develops statistical methods and computational models for analysis of biological data to identify and predict the composition and structure of biological molecules. The rapid growth of sequence data and the huge numbers of structures solved by x-ray crystallography or NMR created the need for a central repository for structure information, the Protein Data Bank (PDB) (http://www.pdb.org/). In use since 1971, the PDB provides atomic coordinates, chemical and biochemical features, details of structure determination, and features derived from the structures of over 45,000 proteins, with thousands more added yearly. Many of the PDB entries are redundant, because the same structure was determined by different groups or resulted when the sequence contained a point mutation, so the Astral compendium (http://astral.berkeley.edu) provides tools to provide nonredundant data based on unique protein domains. Swiss-Prot (http://expasy.ch/) is a widely used annotated protein sequence databank that contains nearly 300,000 entries for sequences assigned to proteins based on homologies or known identities. It provides a description of the proteins' features, such as disulfide bonds, binding sites, and secondary structure elements (including TM a-helices). along with references and links to other databases. It also points out conflicts between data provided by different references. Many bioinformatics tools use the structural information of the PDB to display images that allow visualization of the structures, to simulate their dynamic interactions, to classify proteins into families and superfamilies, and to predict

three-dimensional structures based on sequence homologies. Two programs in use for pairwise sequence alignments are BLAST, a Basic Local Alignment Search Tool (http://www. ncbi.nlm.nih.gov/BLAST), and FASTA (http://www.ebi.ac.uk/ fasta33/), which search a chosen database for sequences that align with the query sequence. A more powerful search can be carried out with the new program PSI-BLAST (same URL as BLAST), an iterative process that extends the search based on the profile first generated by BLAST. Created to probe evolutionary relationships, SCOP, the Structural Classification of Proteins (http://scop.mrc-Imb. cam.ac.uk/scop/), is a database that describes the threedimensional structure of proteins. SCOP organizes proteins by family, superfamily, fold, and class, based on common structural domains (see Figure 6.2.1). A similar database, CATH, for Class, Architecture, Topology, and Homologous superfam iIy (http://cathwww.biochem .ucl.ac. uk/latest/i ndex. html). takes a slightly different approach to classify folds into 30 major protein architectures, such as a-bundle, f.l-barrel, af.l-propeller, and so on. There are over a thousand superfamilies of proteins identified in CATH and SCOP. Other tools probe the interactions between proteins (see for example, DIP, the Database of Interacting Proteins; http://dip.doembi.ucla.edu/) and between proteins and their ligands or their solvent. These tools were developed for soluble proteins; bioinformatics of membrane proteins relies more on specialized algorithms described later in the chapter. For details see Structural Bioinformatics, edited by P. E. Bourne and H. Weissig, published by Wiley-Liss in 2003.

scop Classes March 200 I Alia

All (3

138 Folds

93 Folds

Other Proteins

0:/(3

0'+(3

97 Folds

184 Folds

• Multiple domain • Membrane and cell : surface Small S-S stabilized: • Coiled coil • Low resolution • Small peptides • Designed proteins

seop classifies proteins into four classes based on common folds. From Reddy B. V. B., and P. E. Bourne. Protein structure evolution and the seop database, in P. E. Bourne and H Weissig (eds.). Structural Bioinformatics. Wiley-Liss, 2003. pp. 239-248. © 2003. Reprinted with permission from John Wiley & Sons, Inc. 6.2.1.

recognizing those portions of polypeptides with favorable free energy changes for transfer from the aqueous solvent to the nonpolal- milieu of the memb.-ane. The transfer free energy change (6G"., see Chapter 1) for partitioning of each amino acid between ethanol and water was the basis of the first scale for hydrophobicity used to predict which sequences of amino acids wel-e likely to form a TM helix. To emphasize the polarity of the side chain, each 6G" was normalized to that

of glycine to subtract the contributions of the amino group, the carboxyl group, and the ex carbon: 6G tr = RT In (Xw/xo) - RT In

(XglyW/XglyO),

where X is the mole fraction of the amino acid in the aqueous phase (\-v) or the organic phase (0) at equilibrium. Numerous other hydrophobicity scales have been developed to model partitioning into the membrane

Bioinformatics Tools for Membrane Protein Families

143

interior based on partitioning into other TABLE 6.3. Hydrophobicity scales for partitioning of the amino organic solvents, vapor pressures of side aCids into nonpolar phases, as determined by Tanford; Goldman, Engelman, and Steitz (GES); and Wimley and White 0/If\N)a chain analogs, or distributions of amino acids in the interior of soluble proteins. GoldmanWimley-White Today the most frequently used are the Amino Nozaki and EngelmanWimleywith charged Goldman-Engelman-Steitz (GES) and the side chains acid Tanford Steitz White Wimley-White (WW) scales (Table 6.3). Phe -2.65 -3.70 -1.71 The GES scale is based on calculated esti-2.00 -0.02 Cys mates for the 6G" of amino acids in (X~1.12 lie -3.11 -2.96 helical peptides arrived at by combining -1.30 -3.39 -0.67 Met -241 -2.80 -1.25 Leu terms for the appropriate hydrophobic and -2.61 Val -046 -1.68 hydrophilic contributions. To the data from -301 -1.90 -209 Trp partitioning experiments the hydrophobic 011 b -0.67 b 3.01 233 His component adds a function for the sur~2.53 Tyr -0.71 0.70 face area of the amino acid side chain in Ala -0.73 -1.60 0.50 -044 -1.20 Thr 0.25 an (X-helix, computed based on geometric -1.00 1.15 Gly considerations. The hydrophilic component 0.01 4.80 Asn 0.85 considers the pKas for charged groups and -004 -0.60 Ser 046 the energy required to produce an uncharged Pro 0.20 0.14 species by protonation or deprotonation. Gin 4.11 0.77 0.10 0.11 b Glu -055 b 8.20 3.63 In contrast, the WW scale is an empirical -0.54 b 9.20 043 b 3.64 Asp scale based on water/octanol partitioning of Arg -073 123 1.81 small random-coil peptides, so it provides an -1.50 Lys 8.80 2.80 empirical measure that includes the contria The D.G l , IS given for the transfer of single amino acids from water to ethanol bution from the peptide backbone. It is aug(Tan ford), from water to an (X-helix in the membrane (GES). and for the transfer mented by varying the charged states of Asp, from water to octanol of the amino acid in a peptide 0NWJ. Note that WimGlu, and His residues to allow for formaley and White compare the un-ionized and ionized states of Asp, Glu, and His. They also determined the D.G" for residues linked in salt bridges, such as tion of salt bridges. For example, neutralizArg+ Asp- (not shown). ing Asp 115 in BR improved detection of helix b Values for the un-ionized state. D as a TM segment (see below). These bioSources: Jones, M. N., and D. Chapman, Micelles, Monolayers, and Biomembranes, Wiley-Liss, 1995, p. 16 (Tanford and GES). Jayasinghe, 5., et ai., J Mol physical hydrophobicity scales can now be Bioi. 2001, 312:927-934 (WW). compared with a biological hydrophobicity scale that describes quantitatively the tendency for each amino acid to be inserted in the membrane as part of a TM helix during biogenesis satisfying picture of the predicted fold of many inteof membrane proteins in cells (see Chapter 7). gral membrane proteins (see Box 6.3), yet they are quite imprecise at the end points of the TM helices and they are unable to establish the orientation of the TM helices. Hydrophobicity Plots A simple yet powerful advance was the creation of hydrophobicity plots, also called hydropathy plots, to display the distribution of hydrophobic segments along the linear peptide chain and thus predict its twodimensional topology. The original Kyte-Doolittle algorithm moves a sliding window along the sequence of amino acids and calculates the mean 6G" at each position (initially using the values of 6G,r from water to vapor phase). To correspond to the length of biJayerspanning helices, the size of the window should be around 18 amino acids. When plotted with the primary structure along the x-axis and hydrophobicity along the y-axis, with positive values (for hydrophobic residues) above the line, the peaks correspond to predicted TM segments. This method became widely used, with results depending heavily on the hydrophobicity scale used (Figure 6.13). Hydrophobicity plots give a

Orientation of Membrane Proteins

Establishing the correct orientation of an integral membrane protein has long been a challenge for membrane biochemists, and many biochemical techniques have been used to label the external portions of membrane proteins, with varying degrees of success. Chemical modification with impermeant reagents is often invalidated by findings that the reagent can permeate the membrane after all. Protease sensitivity is limited by the extensive protease resistance of many membrane proteins, once they are fully folded and inserted into the bilayer. Epitopes recognized by antibodies do identify external binding sites as long as the antibody-binding data give unambiguous results. A successful, though laborious, genetic approach to determining orientation was developed in E. coli using

Functions and Families

144 A. ~

1.76 1.25

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:.0 0

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0..

...0 -0

>,

:I:

0

-0.76 B.

1.96

0

'0 .D 0

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0

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i

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I

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because its substrate is ampiciJJin, which inhibits peptidoglycan formation; thus only when this reporter is exported to the peri plasm does it confer ampicillin resistance to the cell. The third reporter enzyme, ()galactosidase (LacZ), is active only in the cytoplasm because attempts to translocate it across the inner membrane leave it embedded in the membrane, which renders it inactive. When normalized to their expression levels, the enzyme activities obtained for different inserts can be correlated with the sites of localization and used to map the results as a topology model (Figure6.14). While the gene fusion approach works well 1'01- bacterial proteins, only a few eukaryotic membrane proteins have been cloned into E. coLi, where their expression can take advantage of these markers. Identification of glycosylation sites has long been used to indicate which loops or domains are exported from the cytoplasm in eukaryotes, since the sugar chains are always extracellular. A new strategy to localize proteins in eukaryotes uses fusions to green fluorescent protein, \'vhich can reveal the fusion pmtein's compartmentalization if the resolution of the fluorescence microscope is high enough.

20


Wimley-White

30

0

25

50 75 100 125 150175200225250 Center amino acid in window

6.13. Hydrophobicity plots for bacteriorhodopsin using different hydrophobicity scales. A. Kyte-Doolittle; B. Goldman-EngelmanSteitz (GES); C. Wimley-White 0NW) with varying charge on Asp115. The agreement between the WW plot and the structure of bacteriorhodopsin is indicated by the lines for identified (red) and known (blue) TM helices in C, and is illustrated in the frontispiece to this chapter. A and B redrawn from Gennis, R. B., Biomembranes: Molecular Structure and Function, Springer-Verlag. 1989. p. 125. © 1989 by Robert B. Gennis. Reprinted by permission of the author; C from Jayasinghe. S.• et aI., J Mol Bioi 2001, 312:927-934. © 2001 by Elsevier. Reprinted with permission from Elsevier.

gene fusions with reporter enzymes inserted into predicted loop regions of the membrane protein. Three reporter enzymes that each requires a particular location in the cell are commonly used. Alkaline phosphatase (PhoA) is normally exported to the peri plasm, where essential disulfide bonds are formed. so if it is expressed in the cytoplasm (on a cytoplasmic domain of a membrane protein), it gives little or no activity. ()Lactamase (Bla) is also effective only in the periplasm.

Using the topological data on E. coli inner membrane proteins plus data from the pmteins in the photosynthetic reaction center, a statistical analysis of the amino acids of internal and external loops revealed a prevalence of basic residues in the cytoplasmic loops. In general the non-translocated loops of membrane proteins have two to four times more Lys and Arg residues than found in translocated domains. This characteristic

BOX 6.3. Making and testing hydrophobicity plots MPTopo (http://blanco.biomol.uci.edu/mpex) is a tool for the simple generation of hydrophobicity plots using the WW scale and a window size of 19. It allows the user to choose a protein sequence from the PDB database or to choose a protein of known topology. The site divides the latter proteins into three groups: "3D-helix" contains helical membrane proteins whose structures are known from x-ray crystallography or NMR; "1 D-helix" contains proteins with evidence from gene fusions and other techniques that supports their predicted topology; and "3D-other" contains f3-barrels and monotopic proteins. It is easy and fun to generate and compare hydrophobicity plots for known integral membrane proteins. However, MPTopo predicts TM segments in a large fraction (up to 43%) of soluble proteins chosen from the PDB because it picks up segments that are simply buried in the hydrophobic interior of a globular protein, as well as cleavable signal sequences on secreted proteins (see Chapter 7). As a research tool, it must be combined with other methods to improve the success of topology predictions (see below).

Bioinformatics Tools for Membrane Protein Families A.

I

t --+-+----+--+--+----+- Periplasm --+-+---+--+-+----+- Cvtoplas rn N

t

C

2

N

~

=

Alkaline phosphatase

B.

PERI MEM CYT

36 6.14. Analysis of membrane protein topology using alkaline phosphatase fusions. A. Diagram that demonstrates the method: when the fusion puts alkaline phosphatase in the peri plasm, as at site 1, it is active, and when it puts it in the cytoplasm, as at site 2, it is inactive. From Manoil, c., and J. Beckwith, Trends Genet. 1988,4:223-226. © 1998 by Elsevier. Reprinted with permission from Elsevier. B. Application of the method to the LacY protein. The sites of fusions are labeled by the level of alkaline phosphatase activity they produced: high (dark blue arrows), > 190 units; medium (hatched green arrows), 46 to 99 units; and low (light blue arrows), <35 units. The ends of some TM segments are adjusted to maximize their hydrophobicity. Redrawn from Calamia J., and C. Manoil, Proc Natl Acad Sci USA. 1990,

87:4937-4941.

is the basis of the von Heijne positive-inside rule and makes it possible to predict the orientation of the TM helices from the amino acid sequences. Specifically, a loop with several basic residues within 30 residues [Tom the end of a TM helix will be an internal loop, probablv due to restricted insertion of helical hairpins carrying highly positively charged sequences. The positive-inside rule \vas tested by engineering the charge distribution in the E. coli inner membrane

145

protein leader peptidase. Leader peptidase (Lep) has two TM helices (HI and H2) separated by a cytoplasmic domain (PI), with a short N terminus and a large C terminus (P2), both in the peri plasm. The small PI domain has nine positively charged residues. When basic residues were redistributed to put more in the N terminus than the PI domain, the orientation of Lep flipped to the inverted topology (Figure 6.15A). More tests of engineered proteins with at least four TM segments show that altering the charge distribution can invert or "frustrate" the topology of the protein (Figure 6.15B). An example of the positive inside rule at work during evolution is the pair of highly homologous proteins called RnfA and RnfE, membrane proteins involved in nitrogen fixation in Rhodobactercapsulatus that differ in orientation across the membrane and exhibit opposite distributions of Lys and Arg residues (Figure 6.16). Analysis of tbe amino acid distributions in integral membrane proteins [,-om over a hundred genomes (see below) indicates that the positive-inside rule is universal. Combining the positive-inside rule with hydrophobicity plots significantly improves the power of topology predictions, as carried out by TopPred. an algorithm that calculates a standard hydrophobicity profile and ranks the possible topologies according to the positiveinside rule. Structural predictions also benefit from knowledge of which side of the membrane the N or C terminus is on. A new approach is the use of gene fusions to rapidly identify the localization of C termini using PhoA as a periplasmic marker and green fluorescent protein as a cytoplasmic marker, since it does not fold correctly in the E. coli peri plasm. This approach enabled a global topology analysis of over 700 inner membrane proteins in E. coli. Genomic Analysis of Membrane Proteins

Identification of membrane proteins in genomes requires fast, accurate methods designed to start with the primary structure of a protein derived h'om a genetic sequence and predict the locations of TM segments (Figure 6.17). Of the algorithms that carry out more sophisticated statistical analyses (Table 6.4), the most widely used are TMHMM and HMMTOP, which use hidden Markov models; MEMSAT, which uses dynamic programming to optimally "thread" a polypeptide chain through a set of topology models; and PHD, which builds a neural network for predicting secondary structure from multiple sequence alignments (see Box 6.4). Various investigations have examined the accuracy of these predictions, which range up to 85% for predicting individual TM helical regions in a new dataset. In a comparison with a test set of 60 bacterial integral membrane proteins with known topology, the

A.

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6.15. Demonstration of the positive-inside rule with leader peptidase (Lep). A, Mutations that affect charge distribution in the loops can change the topology of Lep. At left is wild-type Lep in its normal orientation with Pl in the cytoplasm. The middle construct is a mutant with a shortened Pl, still in normal orientation. The right illustrates that the addition of four basic residues to the N-terminal end gives an inverted orientation. From von Heijne, G., Annu Rev Biophys Biomof 5truct. 1994, 23: 167192. © 1994 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org. B, A variety of Lep constructs have been engineered from duplicates of the lep gene to have four TM segments and a varying number of lysine residues in soluble portions as indicated. Some of them, such as the 3K10K/OKl3K construct, are "frustrated," meaning that regions that would normally span the membrane cannot do so. Redrawn from Gafvelin, G., et aI., J Bioi Chem. 1997,272:6119-6127. © 1997 by American Society for Biochemistry & Molecular Biology, Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

C '0'

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1+ ORF 193 (RnfA homolog)

Cyloplasm

Pet'iplasm

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I

, YdgQ (RnfE homolog)

Cyloplasm

N

6.16. Topology of the homologous proteins RnfA and RnfE from R. capsu/atus. The E. coli homologs, ORF193 (RnfA) and YdgO (RnfE), were tested by PhoA fusion analysis. The opposite orientations of the two proteins are consistent with the positive-inside rule, indicated by the number of Lys + Arg residues in each loop and tail (with the number of + charges shown). Redrawn from von Heijne, G., Q Rev Biophys. 1999,32:285-307. © 1999. Reprinted with permission from Cambridge University Press.

Bioinformatics Tools for Membrane Protein Families

147

TABLE 6.4. Some of the most commonly used methods for predicting topology of helix bundles, along with their

web URLs Name (reference)

URL

Method

HMMTOP rrusnady & Simon, 1998) MEMSAT (Jones et al., 1994)

http://www.enzim.hu/hmmtop/ http://www.psipred.net/

PHDhtm (Rost et al., 1996) PRED-TMR (Pasquier et al. 1999)

http://www.predictprotein.org/ http://biophysics.biol.uoa.gr/PRED-TMR2/

TMAP (Persson & Argos, 1996)

http://www. psc. ed ul gene ra I/softwa re/packageslem bossl tmap.html http://www. bioinformatics-ca nada.orglTMI http://www.cbs.dtu.dk/servicesITMHMM/ http://bioweb.pasteur.frlseqanol/interfaces/toppred .htm I

Model-based, HMM Model-based, dynamic programming Neural network Sliding-window + edge detection Multiple sequence alignments Sliding-window Model-based, HMM Sliding-window + positive-inside rule

TMFinder (Deber et aI., 2001) TMHMM (Krong et aI., 2001) TopPred (C1aros & von Heijne, 1994)

Source: Lehnert, U, et 01,0 Rev Biophys. 2004, 37:121-146.

~

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Protein sequence (lD) FHEPIVM VTIAG IILGG LALVG LITYFGKWTYLWK EWLTS VDHKRLGIMYIIVAIVMLLRGFADAIMMRSOQALASAGEA GFLPPHHYDQIFTAHGVIMIFFVAMPFVIGLMNLVVPLQI

Input

t

Prediction I algorithms

I

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Predicted helical transmembrane segments (lD) 9999999864442001112346689999999999999999 9999863202567788888887765214678899999999 9999887622567788888888888888777665544442

numbers Predicted helical transmembrane segment

Observed helical transmembrane segment

Reliability indices

6.17. Strategy for predicting topology from protein sequence. Algorithms such as TMHMM, PHD, and MEMSAT all derive topology information from the primary structure of proteins. The example shows a partial sequence of cytochrome-o ubiquinol oxidase subunit 1 (cyob), diagrammed to show the three TM helices determined experimentally (top) and the analysis by PHDhtm (bottom). The bars in the last panel allow comparison between predicted (white) and observed (shaded) TM segments, and the numbers below the line give the reliability of the prediction for each residue on a scale of 0 to 9. The low reliability values for the top segment illustrate how TM segments can be underpredicted. Redrawn from Rost, B, et aI., Protein Sci. 1995, 4:521-533. © 1995. Reprinted with permission from Protein Science.

Functions and Families

148

BOX 6.4. Statistical methods for TM prediction While most of the methods listed in Table 6.4 employ hydrophobicity analysis and the positive-inside rule to find TM domains, some are greatly enhanced by the computational power of the statistical methods they employ. These methods give improved accuracy over the simple sliding-window approach that is limited to analysis of the linear sequence of the protein of interest. In particular, they can be based on multiple sequence alignment since there is now a sufficient databank of membrane proteins of known topology to be used in the search for other membrane proteins. Two successful approaches are neural networks and hidden Markov models.

input for the second. For the first level, the local input is a weighted term including the frequency of occurrence of each amino acid at that position in the multiple alignments plus the number of insertions and deletions in the alignment for that residue. The output of the first level indicates whether or not the segment is predicted to be a TM helix. This is the local input for the second level. The global input for both levels describes characteristics of the protein outside the window of 13 residues and includes its amino acid composition and length, plus the distance (number of residues) from the first residue in the window of 13 adjacent residues to the N terminus and the distance from the last residue in the window to the C terminus. The output is expressed in units that signify TM helix and not-TM helix. As shown in Figure 6.4.2, the first level of PHD analyzes the protein sequence in windows of 13 amino acids (only seven are shown) and produces the output code for HTM (helix TM) or NotHTM, which becomes the input for the second level. The box on the left shows the data from sequence information from the protein family used for both local and global factors. The center box shows how at each position in the alignment the amino acid frequencies are compiled, the number of insertions and deletions are counted, and a conservation weight is computed for the local input. Global information gives the amino acid composition, length of the protein, and position of the current window from the Nand C termini. All of this was codified and fed into the neural network input for the first level and then to the second level, shown in the box on the right.

Neural Networks The program PHD uses neural networks to score "preferences" for TM helices generated from multiple sequence alignments. Neural networks compute relationships between units (amino acid residues) in layers, described as the input layer, one or more hidden layers, and the output layer, that together define the network architecture (Figure 6.4.1). For multiple protein sequences, the input is a sequence profile of a protein family, with each sequence position represented by the amino acid residue frequencies derived from multiple sequence alignments in the database. The hidden layer is set up to determine the state of each unit as it depends on the states of the units in the previous layer to which it is connected and the weights of the connections. The output layer has the results of the analysis. PHD analyzes protein sequences with two levels of neural network analysis, so that the output from the first level is the

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Functions and Families

150

BOX 6.4 (continued)

Amino acid seq:

MGDVCDTEFGILVA ... SVALRPRKHGRWIV ... FWVDNGTEQ ... PEHMTKLHMM ...

State seq:

o oooooooohhhhh ... hhhhi iii iii hhh ... hhhooooOO ... OOOoooohh h ...

Topology:

Tail Out

Tail-Tail

Hetix

SlWl't loop

Helix

Tail - Loop - Tail Long loop

6.4.3. Structural states defined for HMMTOP, where thick lines represent tails while thin lines are loops. Redrawn from Tusnady, G. E., and I. Simon, J Mol Bioi. 1998,283:489-506. © 1998 by Elsevier. Reprinted with permission from Elsevier.

When PHD was tested with an initial dataset of 69 membrane proteins with experimentally determined locations of TM segments, its accuracy was >95%. Assessments of its accuracy when tested on a dataset not used in setting up the analysis are considerably lower. One weakness of the system is the step it uses to filter out TM helices that are too long (>35) or too short «17). If too long, they are split in the middle into two helices; if too short, they are deleted or elongated.

Hidden Markov Models Hidden Markov models (HMMs) apply statistical profiles to describe a series of states connected by transition probabilities. For proteins, each state corresponds to the columns of a multiple sequence alignment, which are intermediate steps in the algorithm that the user does not see. A matrix describes the possible states and the transitions between the states, and an algorithm is employed for a "random walk" through the states, that is, one that derives each possibility from the previous state. For sequence alignment, the HMM generates profiles compiled of high scores (if sequence is highly conserved), low scores (if sequence is weakly conserved), and negative scores (if sequence is unconserved) and identifies sequences with the highest scores. In HMMs for membrane protein topology, structural states are defined to describe portions of the protein. HMMTOP

uses five structural states: inside loop (I), inside tail (i, the region of a loop that is close to the TM helix), TM helix (h), outside tail (0), and outside loop (0) (Figure 6.4.3). TMHMM further divides the residues in TM helices according to whether they are in the center of a helix (helix core) or on one end of a helix (helix cap). Thus seven states are considered in TMHMM: helix core, helix caps, short loop on inside, short and long loop on outside, and globular regions. Each state has a probability distribution over the 20 amino acids, based on the dataset with known topologies. The overall layout of the HMM is a function of the different structural states (Figure 6.4.4). Arrows show the possible connections between the different states, which are limited by the constraints of the protein structure as depicted in (A), with boxes corresponding to one or more states in the model. The connectivity of the different states varies. For the inside and outside loops and helix caps the connectivity is shown in (8), and for the helix core it is shown in (C). This model allows the helix core to be between five and 25 residues, which makes the entire length of TM helices (including caps) between 15 and 35 residues. The program follows rules of "grammar" that state a helix must be followed by a loop, and inside and outside loops must alternate. Then it calculates probabilities for the sequences of these states. This is depicted in the architecture of HMMTOP (Figure 6.4.5), which shows states within the same transition matrices (gray = helix states, yellow = tail states, red = loop states). The rectangular areas represent fixed-length states,

Bioinformatics Tools for Membrane Protein Families

151

BOX 6.4 (continued)

A.

Cytoplasmic side

Me.mbrane

non-cytoplasmic side

Loop

C.

cap

Helix core 22

2J

24

2S

6.4.4. The layout of the hidden Markov model. Redrawn from Krogh, A, et aI., J Mol Bioi. 2001,305:567-580.

"f MA,'(L, ()

MINL,

tail L, MAXL"

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6.4.5. The architecture of HMMTOP. Redrawn from Tusnady, G. E., and I. Simon, J Mol BioI. 1998, 283:489-506.

© 1998

by Elsevier.

Reprinted with permission from Elsevier.

(continued)

Functions and Families

152

BOX 6.4 (continued) which include the helices whose lengths need to be sufficient to cross the nonpolar domain of the bilayer and the tails that are defined to be the short segments of loops adjacent to ends of helices. The hexagonal areas represent the nonfixed-length states, which are the loops that are allowed to be any length. By considering fixed-length states and non-fixed-

length states, the methods allow realistic length constraints on TM helices. While the current programs using these methods (listed in Table 6.4) are very powerful, new computational approaches can be expected to make them even more useful in the near future.

highest fraction (around three fourths) were correctly predicted with two algorithms using hidden Markov model, TMHMM, and HMMTOP, while MEMSAT and TOPPRED had less success and the lowest fraction (around one half) were correctly predicted with a neural network predictor, PHD. The best results were achieved by carrying out analyses by all five and searching for consensus: a very high rate of correct predictions occurs when all five or four of the five agree. A new algorithm that compares the results of nine different methods is called CONPRED. The success of the predictions is still consistently higher with prokaryotic genomes than with eu kalyotic genomes. Analysis of 26 genomes for membrane proteins produced a total of 637 families of poly topic TM domains, based on a combination of TM helices predicted using

TMHMM and those annotated in Swiss-Prot. The domains were classified based on homologies with known families or characterized using sequence alignment, hydrophobicity plots with the GES scale, and identification of consensus sequences for TM segments (Figure 6.18). When the families are sorted by the number ofTM helices, the number of families with domains of a given number of TM helices decreases as the number of helices increases (Figure 6.19). Within this trend, the plot shows the highest occurrences of two- and four-TM helices, a slight excess of seven-TM helices, and a major spike at twelve-TM helices due to the prevalence of this topology among transporters and channels. Interestingly, the total number of membrane protein domains in the genome is roughly proportional to the number of open reading fTames in all of the genomes

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6.18. Classification of a family of polytopic membrane domains. The example shown is family PF01618. The steps involved are (top to bottom) sequence alignment, hydrophobicity plot based on GES scale, consensus sequence displayed by sequence logo, and consensus sequences of TM helices, where the nonconserved amino acids are represented by "x." From Liu Y, et al., Genome Biology 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

Bioinformatics Tools for Membrane Protein Families

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7 8 9 10 11 12 13 Number of TM helices 6.19. The number of TM helices in families of polytopic membrane domains. The number of families of polytopic membrane domains is plotted on the y-axis as a function of the number of TM helices, plotted on the x-axis. The green bars are the numbers from all studied families from the Pfam-A database. The yellow bars are the numbers from families from the Pfam-A database that are annotated as transporters and channels. Redrawn from Liu, Y, et al., Genome Biology, 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

D 1% K

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except that of the nematode worm C. elegans, which has a huge number (754) of seven-TM chemoreceptors. In fact, the genome of C. elegan5 conforms to the general picture when its three large families of chemoreceptors are removed from the total. Clearly this organism that lacks sight and hearing has finely developed chemosensation to find its food l Approximately half of the membrane proteins identified by genome analysis have an even number of TM helices with the Nin-C in topology. The other three combinations of N- and C-terminal locations are about equally represented in the other half. The high proportion of proteins with both Nand C termini inside is attribu ted to the mechanism of biogenesis, because this topology results from insertion of helical hairpins (see Chapter 7). Amino acid distributions in the TM segments of the putative membrane proteins in the 26 genomes show the expected high amounts of nonpolar amino acids, along with the polar residues Ser and Thr that participate in hydrogen bonding (see below; Figure 6.20A).

B. Positionally conserved residues K

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6.20. Amino acid distributions in TM helices. Amino acid distributions were determined for the TM segments from the 168 families from the Pfam-A database that have more than 20 members. A. A pie diagram of the amino acid compositions of TM helices shows their high content of nonpolar residues, along with glycine, serine, and threonine. B. Consensus sequences identified as shown in Figure 6.18 allow comparison of the amino acid residues in conserved positions of the TM helices. The diagram of the compositions of these positionally conserved residues shows that the prevalence of three amino acids (Gly, Pro, and Tyr) has increased significantly, indicating they are the ones whose positions are highly conserved. Redrawn from Liu, Y, et al., Genome Biology, 2002, 3:research0054.1-0054.12. © 2002 by Yang Lui, Donald M. Engelman, and Mark Gerstein. Reprinted with permission from the authors.

,. .'

Functions and Families

154

Similarly, the composItion of nearly 50,000 TM segments annotated in the Swiss-Prot database (see Box 6.2) reveals that the six amino acids Leu, Ile, Val, Phe, AJa, and Gly make up two thirds of TM residues. When the sequences are aligned to determine conserved residues (see Figure 6.18), the nonpolar amino acids are not prevalent in conserved positions, presumably because they al-e quite interchangeable. Interestingly, the amino acids that are prevalent at highly conserved positions in the helices are Gly, Pro, and Tyr (Figure 6.20B). The proline residues form kinks in the helices, which are conserved even after mutation of the Pro residues, while tyrosine residues playa special role near the interfaces due to their electronic propel1ies (see Chapter 4). The positions of glycine residues are often highly conserved in soluble proteins because they occur at positions where the proteins do not accommodate larger side chains. This is also the case in TM helices, where Gly is commonly observed where two helices are in close contact. The GxxxG motif, llrst observed in glycophorin dimers, places both Gly l-esidues on the same side of the <X-hel ix (see Figure 4.30). Genomic analysis for pair motifs shows a very high presence of GxxxG and GxxxxxxG pairs, as well as similar motifs with Ala or Ser replacing Gly residues. The resulting location of small side chains is expected to be important fOI' helix-helix interactions in a broad range of membrane proteins. Helix-Helix Interactions

The prevalence of the GxxxG motif in poly topic membrane proteins in the genome reflects both the close packing of TM helices observed in many helix-bundle proteins and the importance of protein-protein interactions in oligomeric membrane proteins and complexes. To analyze helical packing patterns within proteins, a comparison of pairs of helices in membrane proteins and pairs in soluble proteins shows that most helixhelix pairs in membrane proteins have homologs in soluble proteins. The exceptions to this correlation are the irregular helices of some membrane proteins (described in Chapter 4). The high occurrence of GxxxG and GxxxA in helices of membrane proteins allows the helices to approach more closely than those in soluble proteins, forming "knob-into-hole" interactions, as well as to interact over longer distances, that is, over the width of the bilayer. The information on helix pairs within proteins probably also applies to helixhelix interactions between subunits because the TM helices from different subunits often align as much as TM helices within one subunit. This can be seen with oligomeric proteins, where a cross-section through the middle of the membrane shows the extensive interaction of helices from different subunits (Figure 6.21): the

Photosynthetic react ion center

(Rhodopseudomonas virdis) Deisenhofer et al. (1995)

I

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6.21. Helix interactions viewed from the membrane midplane. Positions of five-residue sections at the middle of TM helices are shown for photosynthetic reaction center, cytochrome-c oxidase, and cytochrome-bcl complex, as labeled. The subunits are colored differently. The gray-scale image of cytochrome-c oxidase shows that the subunit composition cannot be inferred from the relationships of the helices. Redrawn from Liu, Y, et aI., Proc Natl Acad Sci USA 2004. 101:3495-3497. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

gray-scale image of cytochrome-c oxidase in the figure shows how difficult it is to assign subunits from the relationships of the helices! Additional information about helix-helix intemctions comes from the analysis of interhelical three-body interactions to find "triplets" where three atoms from at least two different helices are closely packed. A program called INTERFACE-3, which computes interhelical atomic triplets based on algorithms for geometric shapes and distances, detects six diffel-ent types of triplets in the glycophOlin dimer: GGV, GTV, GVV, ILL, nT, and ITT (Figure 6.22A). Likewise, bacteriorhodopsin and its homologs halorhodopsin and sensory rhodopsin II have three triplets that are conserved in sequence and in structure (Figure 6.22B). Furthermore, an additional 13 triplets are conserved structul-ally but not fOl-med by identical residues, allowing conservative mutations that suggest that the triplet interaction has been conserved to maintain the orientation of the helices in the three highly homologous proteins. Comparison with a set of soluble <x-helical proteins identified triplets that are unique to membrane proteins, such as AGF, AGG, GLL, and GFF. Such triplets are often found in regions of the closest contact between helices, and are therefore often correia ted with GxxxG-type motifs. Close contact between TM helices is also stabilized by two types of hydrogen bonding. The hydrogen bonds

Bioinformatics Tools for Membrane Protein Families

B.

A.

a

b G79:C V80:Ca

V80:Ca

6.22. Three-body contacts, or triplets, in helix-helix interactions. A. The TM helices (residues 73-91) of a glycophorin dimer with one of the triplets illustrated in space-filling representation. The atoms involved, C from Gly79 and Ccx from Val80 on both chains, are shown in space-filling representation (a), and viewed from the top (b). (Orange, C from G79, chain A; green, Ccx from V80, chain A; blue, Ccx from V80, chain S.) B. A conserved triplet in the archaei rhodopsins is shown by superposition of helices C, E, and F from bacteria rhodopsin, halorhodopsin, and sensory rhodopsin II. The residues in the triplet are two Leu and a Thr, corresponding to L97, L152, and T178 in SR, again in space-filling representation. The retinal is drawn in green From Adamian, L., et aI., J Mol Bioi. 2003, 327:251-272. © 2003 by Elsevier. Reprinted with permission from Elsevier.

observed in glycophorin dimers are relatively weak bonds because they use the CO( proton as the donor. Each of these hydrogen bonds contributes less than 1 kcal/mol to the stability of the dimer. The second type B.

A.

155

of hydrogen bond makes use of polar residues in TM segments; such bonds are detected in high-resolution structures of poly topic membrane proteins such as bacteriorhodopsi n. Interactions between polar residues account for around 4% of all atomic interhelical contacts in membrane proteins, as ,.veil as in soluble proteins. In contrast to soluble proteins, where interacting ionized residues typically form salt bridges, the types of polar interactions are more varied in TM segments of membrane proteins, which also have H-bonds bet,veen ionizable and polar residues (the most common are DY and Y-R) and between polar nonionizable residues (the most common are Q-S and S-S). How prevalent are these interhelical hydrogen bonds? In a dataset based on the high-resolution structures of 13 membrane proteins, 134 unique TM helices form nearly 300 helical pairs, 53% of which are connected by at least one H-bond. In the 299 interhelical H-bonds identified, almost half involve Ser, Tyr, Thr, and His. Hydrogen bonds between two side chains and hydrogen bonds between a side chain and a backbone carbonyl oxygen occur at the same frequency. The majority of these helical pairs have one H-bond between two amino acids from two neighboring helices. However, two other types ofH-bonding patterns emerge. One type is a "seline zipper" motif, named for its similarity to a leucine zipper (Figure 6.23). A search for homologous serine zippers by PSI-BLAST identified more than 100 sequences with highly conseJ\led Ser residues in positions 7, 14, and 21 of one helix and 1,8, and 15 of the other. A three-body analysis also found triplets of two serine residues with a leucine. The other type of H-bonding pattern is called a polar clamp because the

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6.23. The serine zipper motif in helix-helix interactions. A. Schematic representation of hydrogen bonding between two serine residues on two adjacent helices. B. An example of a serine zipper in bovine cytochrome-c oxidase. The two helices shown are helices III and IV, and the hydrogenbonded serine residues are 5101-5156, 5108-5149, and 5115-5142. C. The pairs of serines can be predicted from helical wheels of helices III and IV from subunit 1 of bovine cytochrome-c oxidase. A helical wheel designates residues i and i + 3 as a and d and shows how they fall on the same side of the helix. Nonpolar residues are shaded pink. Three 5er-5er pairs and two Leu-Leu pairs form a zipper at different a and d positions in this example. A and B redrawn from Adamian, L., and J. Liang, Proteins. 2002, 47:209-218. © 2002. Reprinted with permission from John Wiley & Sons, Inc.; C from Adamian, L., et aI., J Mol Bioi. 2003, 327:251-272. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Functions and Families

156

A.

B.

S159

6.24. Polar clamp hydrogen bonding in helix-helix interactions. A. Three residues in rhodopsin, W161, T160, and N78, form a clamp between helix IV and helix II. The side chain of N78 (on helix II) makes two hydrogen bonds: its 0 atom is hydrogen bonded to the N atom from W161, and one of its amide hydrogen atoms is hydrogen bonded to the oxygen of T160. B. A polar clamp in subunit I of cytochrome-c oxidase from Thermus thermophifus involves residues S155 and S159 from helix 0:4 and 086 from helix 0:2. From Adamian, L., and J. Liang, Proteins. 2002, 47:209-218. © 2002. Reprinted with permission from John Wiley & Sons, Inc.

side chain of an amino acid at a given posItIOn I that is capable of forming at least two hydrogen bonds (i.e., E, K N, Q, R, S, T) is "clamped" by H-bonding to two other residues, one at position i + 1 or i + 4 and one on the othel- helix (Figure 6.24). The polar clamp shown in rhodopsin, involving l-esidues T160 and WI61 (both on helix IV), and N78 (on helix U), is highly conserved in the GPCR family. When the numbers of intermolecular and intramolecular helix-helix interactions are estimated for membrane proteins in whole genomes, the totals are in the millions. Clearly an understanding of them will deepen as the number of high-resolution stnlctures increases and will provide in turn a more complete insight for predicting membrane protein structure. Furthermore, helix-helix interactions are a critical step in the assembly of polytopic membrane proteins, as desclibed in the next chapter. Since the nonpolar residues are quite nonspecific in their interactions, the often conserved hydrogen bonds between polar residues in TM segments must be crucial in helix alignments. Proteomics of Membrane Proteins

With thousands of membrane proteins predicted by genomic analysis, many questions are raised about their

identities and interactions. Using TMHMM predictions combined with PhoA/GFP fusions to localize the C termini, a global topology analysis of E. coli membrane proteins detected 601 inner membrane proteins. When these proteins are sorted by their known or predicted functions, 40% of them are involved in transport, while nearly 20% are involved in metabolism, biogenesis, and signaling (Figure 6.25). Over one third are "orphans" with unknown functions. The functions of some orphan memb.-ane proteins can be ascertained from their association with known proteins in complexes. (The yeast two-hybrid analvsis, an important proteomic tool for identifying proteinprotein interactions in complexes, does not work with membrane proteins because loss of function in the fusion proteins generated can result from loss of correct compartmentalization, topology, or orientation in the membrane.) Complexes of membrane proteins can be detected by two-dimensional gel electrophoresis using native gel electrophoresis for the first dimension, combined with mass spectrometry to identify the proteins. To carry out this procedure with Gram-negative bacteria such as E. coli, the inner and outer membrane fractions were first separated by gradient centrifugation and solubilized in a mild detergent, such as 0.5% 11dodecyl-0-D-lllaltoside. When native gel electrophoresis was carried out in Coomassie blue, complexes involving 44 inner membrane proteins and 12 oute.- membrane proteins were isolated. The results enabled roles to be assigned to six Ol-phan proteins of unknown function, in addition to identifying a number of Imown proteins. A majority of the inner membrane proteins in complexes are members of .-espiratory chains (e.g., succinate dehydrogenase, F I Fo-ATP synthase, and cytochrome b0 3 ubiquinol oxidase). Others are involved in biogenesis (including the SecYEG translocon; see Chapter 7) and

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6.25. Functional categorization of the E. coli inner membrane proteome. The 737 proteins identified in the inner membrane are assigned to different functional categories. Redrawn from Daley, D.O., et al., Science. 2005, 308: 1321-1323. © 2005. Reprinted with permission from AAAS.

Bioinformatics Tools for Membrane Protein Families

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4,-------------------------------------, OmpF 3

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6.26. Prediction of ~-barrel proteins by BBF, the ~-barrel finder program. BBF uses both hydropathy plots (solid lines) and amphipathicity plots (dotted lines) since the predicted ~-strands (indicated along the bottom) are based on the overlap of hydrophobic and amphipathic segments. As an example, the plot for OmpF (described in Chapter 5) is shown, Since it takes only seven to nine residues to span the membrane as a ~-strand, the window size is set for seven. However, this introduces noise in the baseline (compare with Figure 6.13). Redrawn from Zhai, Y, and M. H. Saier, Protein Sci. 2002, 11 :2196-2207. © 2002. Reprinted with permission from Protein Science.

some are transporters (including the intact PTS systems for mannose and galactitol, as well as ABC transporters for maltose and glutamine). The trimeric porins in the outer membrane were also identified, as the method also recognizes homo-oligomers. 13-Barrels The prediction methods described above for genomic analysis utilize algorithms that identify TM ex-helices and are not useful for recognition of (3-barrel mem brane proleins. One way to illustrate this is to apply MPTopo to the (3-barrel proteins in group 3 of the program (See Box 6.3). To identify 13-barrels, algorithms combine secondary stnJcture prediction and hydrophobicity analysis. For example, the Beta-Barrel Finder (BBF; http://ww\'v. biology.ucsd .edu/~msaier/transport/ software.html) carries out four steps. First, because these proteins are exported to outer membranes, protein sequences are screened for an N-terminal signal sequence, which has a hydrophobic segment that is within the first 50 residues from the N terminus (see Chapter 7). The second step is prediction of secondary structure and the selection of predicted (3strands for [Llrther analysis. The third step is analysis

of the (3-structure regions for hydrophobicity and also For amphiphilicity, using a window size of seven. Since the TM (3-strands of the porins contain alternating polar and nonpolar amino acids, they are more hydrophobic than the loops. Therefore, BBF looks for peaks of hydrophobicity and amphiphilicity that coincide (Figure 6.26). The last step takes the sequences of these regions for known (3-barrel proteins and uses BLAST to look for homologous sequences. An alternative approach uses a neural network to predict (3-barrels. These methods can effectively identify the TM (3strands of known (3-barrel proteins, exemplified with BtuB (Figure 6.27). Application of these methods to genomes of different Gram-negative bacteria indicates that 2% to 3% of the genomes encode (3-barrel proteins, which corresponds to <10% of integral membrane proteins, In addition, the methods signify how many (3-barreJ proteins remain to be characterized: one third to one half of the approximately 100 (3-barrels they identify in the E. coli genome are unknown proteins. There are now 35 families of (3-barrels, including 29 in Gram-negative bacteria. The others are in mitochondria, chloroplasts, and the acid-fast Grampositive bacteria, such as mycobacteria. There is no significant sequence similarity between members of

Functions and Families

158

6.27. A and B. Correlation between predicted and actual I:'-strands in the E. coli outer membrane protein BtuB. The arrows show the peaks that are color coded to correspond to the [3-strands in the ribbon diagram of the structure. From Wimley, W. C, Curr Opin Struct Bioi. 2003, 13:404-411. © 2003 by Elsevier. Reprinted with permission from Elsevier.

different families, in spite of frequent resemblances in the fold. Although a relatively new field, bioinformatics of membrane proteins has already provided a wealth of information about families of membrane proteins and their characteristics, especially for the large class of helical bundle proteins. It provides patterns and frameworks that clarify much of what is known about membrane proteins, and by viewing data on hundreds of predicted membrane proteins with unknown structures and functions, it stimulates the desire to know more about the many important roles these proteins play. FURTHER READING Enzymes

Li, H., and T. L. Poulos, Crystallization of cytochromes P450 and substrate-enzyme interactions. Curl' Top Med Chel11. 2004,4:1789-1802. Carman, G. M., R. A. Deems, and E. A. Dennis, Lipid signaling enzymes and surface dilution kinetics. J Bioi Chell1. 1995, 270:1871 J-187J4. Transporters

Busch, W., and M. H. Saier, 11'., The transporter classification (TCl system, 2002. Cril Rev Biochem Mol Bioi. 2002, 37:287-337. Saier, M. H., Jr., Families of transmembrane sugar transport proteins. Mol Microbial. 2000, 35:699-710. Gruber, G., et aI., Structure-function relationships of A-, Fand V-ATPases. J Exp BioI. 2001, 204:2597-2605.

Locher, K. P., Structure and mechanism of ABC transporters. Curl' Opin Slrucl Bioi. 2004, 14:426-431. Tchieu,1. H., et aI., The complete phosphotransferase system in Escherichia coli. J Mol Microbial Biolechnol. 2001,3:329346. Kunji, E. R. S., The role and structul-e of mitochondrial carriers. FEBS Lell. 2004, 564:239-244. Pebay-Peyroula, E., and G. Brando)in, Nucleotide exchange in mitochrondria: insight at a molecular level. Curl' Opil1 Slrucl Bioi. 2004, 14:420-425. Receptors

Miyazaya, A., et aI., Nicotinic acetylcholine receptor' at 4.6A resolution. J Mol Bioi. 1999,288:765-786. Sixma, T. K., and A. B. Smit, Acetylcholine binding protein (AchBP) ... pentameric ligand-gated ion channels. AI1I1lt Rev Biophys Biomol Slrucl. 2003, 32:311-334. Bockaen, 1., and J. P. Pin, Molecular tinkering of G proteincoupled receptors: an evolutionary success. EM 80 J. 1999, 18:1723-1729. Predictions and Genomic Analysis

von Heijne, G., Membrane proteins from sequence to structure. A/1/1U Rev Biophys BiOI/wi Siruci. 1994, 23: 167-192. von Heijne, G., Recent advances in the understanding of membrane protein assembly and structure. Q Rev Biophys. 1999,4: 285-307. Tusnady, G. E., and 1. Simon, Principles governing amino acid composition of integral membrane proteins: application to topology prediction. J Mol Bioi. 1998,283:489-506. Lehnert, U., et aI., Computational analysis of membrane proteins: genomic occurrence, stnJCture prediction and helix interactions. Q Rev Biophys. 2004, 37:121-146.

Further Reading Daley, D.O., et a!., Global topology analysis of the Escherichia coli inner membrane proteome. Science. 2005, 308:13211323. Zhai, Y, and M. H. Saier, Jr., The l3-barrel finder (BBF) program. Protein Sci. 2002, I 1:2169-2207. Wimley, W. c., Toward genomic identification of l3-barrel membrane proteins, Protein Sci, 2002,11:301-312.

159

Protein-Protein Interactions

Adamian, L., et a!., Higher-order interhelical spatial interactions in membrane proteins . .1 Mol Bioi. 2003, 327:251272.

Stenberg, E, et a!., Protein complexes of the Escherichia coli cell envelope. .1 Bioi Chem. 2005, 280:3440934419.

7

Protein Folding and Biogenesis

Folding ~

Ftn

'~I~n~1fn~~~lI~~

!'. . . . .~

A populated unfolded state in the membrane

ll'n"",on

~V~nnnnn

U~~~U~~~~U1~,

11

Cou~led

folding and IIlserlJon

Folding ilnd Insertion of helical bundle and j3-barrel membrane proteins utilize different mechanisms, according to results from In VItro folding studies From Bowie, J, U., Pmc Natl Acad SCI USA 2003, 1013995-3996

With an appreciation of the structural characteristics and functional diversity of membrane proteins, the question of their biogenesis arises, How does a nascent peptide, a chain of amino acids emerging from the ribosome, fold into a three-dimensional structure and insert into the membrane bilayer? Evidence suggests that folding and insertion are coupled processes in cells, However, given the na tu re and complexi ty of these processes, different approaches are taken to study folding and insertion in vitro, Protein folding studies give information about the thermodynamic forces that drive folding and about its kinetic pathways, identifying transient but detectable intermediates, Recently these techniques have been applied to purified membrane proteins of both the (Xhelical and B-barrel classes, While the folding mechanisms determined by in vitro studies of membrane 160

proteins may differ significantly from mechanisms of their biogenesis in the cell, these studies do provide insights into the necessary steps, as well as their stability and their lipid requirements, Thermodynamic analysis of the in vitro folding process gives insights into the evolution of the complex machinery used to assemble membrane proteins in cells, Finally, the in vitro studies provide valuable practical information on refolding techniques that are applicable to other membrane proteins that have been denatured during purification, Insertion of nascent proteins into the membrane involves their translocation out of the cytoplasm by the same export machinery used to secrete proteins, There are strong similarities between prokaryotic and eukaryotic protein translocation systems, including recognition of the signal for export, structure of the main translocation apparatus, and mechanisms for insertion

Protein Folding into the bilayer. Progress in defining and characterizing the components of these systems is leading to a detailed picture of the processes involved in export and integration of proteins. This chapter views how membrane proteins fold in vitro before turning to the complex cellular process of translocation and integration of nascent proteins into the membrane. It ends with a look at how misfolding of membrane proteins can lead to diseases in humans.

161

A.

o 70

I

•

x

Q 60

~

E c: o

V)

c

~

c

Il)

u C Il) u V) '-

Before discussing folding studies of membrane proteins, it is useful to review some principles established with protein folding studies using purified small soluble proteins, with which both theory and methodology for in vitro folding studies were developed. Typically the protein of interest is unfolded with high concentrations of a denaturant such as urea or guanidinium hydrochloride, \-vhose removal by dilution or dialysis triggers refolding. Temperature and extremes of pH can also be used to cause denaturation, although fTequently it is not reversible in these cases. Of course, solution conditions including temperature, pH, and ionic strength are critical in all folding studies. A standard assay follows the changes in intensity and wavelength of the intrinsic fluorescence of the protein: as it unfolds, the aromatic side chains move to more polar environments and their fluorescence emissions shift to longer wavelengths. A complementary method monitors the loss of secondary structure in the protein using circular dichroism, a method of spectroscopy that detects secondary structure in a peptide backbone by following the differential absorption of circularly polarized light. Other techniques include monitoring changes with ultraviolet and visible absorption and FTlR spectroscopy. Both kinetic and equilibrium measurements are informative: kinetic experiments reveal intermediates in the folding pathway, and equilibrium measurements give thermodynamic constants if the process is reversible. An intact peptide chain can fold to a vast set of different configurational isomers, whose relative free energies are determined by the interactions within the fold and the surfaces they present to the environment. A few configurations have a relatively low free energy, corresponding to multiple minima in the energy landscape that depicts how the energy of the system (comprising all configurations of the protein) depends on the positions and orientations of all its atoms during the folding process. The native protein samples this conformational space by dynamic motions, with small, fast fluctuations between configurations. This dynamical complexity is overlooked in the elementary fonn of the folding reaction, which views it as

/"".

""o

Il)

PROTEIN FOLDING

o

N N

50

-;;; ~,

40

0 :l

u:

30

o

0.5

1.0

1.5

2.0

GdmCI (M) B.

-0

Il)

-0

]

C :l

c

o

0.5

u
[..L.

00-<>00---------------1

o

0.5

1.0 GdmCI (iv!)

1.5

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7.1. Folding studies of the soluble enzyme phosphoglycerate kinase. The data for unfolding phosphoglycerate kinase show a two-state mechanism. A. Unfolding as a function of guanidinium chloride (GdmCI) concentration is monitored by fluorescence (closed circles) and circular dichroism (open circles). Samples are allowed to go to equilibrium. The steepness of both curves indicates that unfolding is a highly cooperative process. B. When the fraction unfolded is plotted as a function of the concentration of GdmCI, the two methods give the same unfolding curve, consistent with a two-state mechanism. Redrawn from Creighton, T. E., Proteins Structures and Molecular Properties, 2nd ed., Freeman, 1992, p. 288. © 1992 by W. H. Freeman and Company Used with permission.

a two-state process: U B N, where U is the unfolded protein and N is the native (folded) protein. For small, one-domain globular proteins, the unfolding process appears to be a first-order transi tion, which typically exhibits cooperativity (Figure 7.1). When this reaction

Protein Folding and Biogenesis

162

C

N

~

JL\G~ =

C

C

°- 0

L\G I

N

DcY,or1a"n

•

JL\G~ = -60

+46

N L\G 4O = -106

..

----------------------

Helix

Helix Polar headgroups

~

-------------------

Helical hairpin 7.2. The helical hairpin hypothesis describes the insertion of a hairpin structure composed of two helices into the nonpolar interior of the bilayer. It is driven by the free energy arising from burying hydrophobic helical surfaces, estimated to be -60 kcal/mol for a pair of membrane-spanning helices. The alternate pathway of inserting a random coil prior to folding in the bilayer, with a free energy change of +46 kcal/mol, is so unfavorable that the insertion of an unfolded peptide effectively cannot occur. Redrawn from Engelman, D. M., and 1. A. Steitz, Cell. 1981, 23:411-422. © 1981 by Elsevier. Reprinted with permission from Elsevier.

is fully reversible, it can go to equilibrium, at which point L'.Gfolding

= GN

- GU

= -RT In Ke q ,

where K eq = [N]/[Ul

Keq is also equal to (l

~ Oi.)/Oi., with Oi. defined as the average fraction of unfolding, so it is directly obtained from equilibrium folding or unfolding data. In most unfolding studies, the environment of the protein is simply aqueous solvent; for membrane proteins, it is complicated by the presence of either detergent or lipids, due to the need for an amphipathic system that provides a nonpolar domain to solubilize the native protein. However, once such a system has been defined. it can be exploited to give additional information, such as demonstrating the importance of bilayer elasticity by obtaining thermodynamic parameters in the presence of different lipids, as shown below.

Folding ex-Helical Membrane Proteins

Because integral membrane proteins require the special environment of the lipid bilayer (as described in Chapter 4), theil- in vi/ro folding process includes their insertion into a lipid bilayer, micelle, or other model membrane. An important early proposal for the mechanism of spontaneous membrane insertion was the helical hairpin hypothesis. It described how two hydrophobic (X-helices connected by a hairpin turn can more Favorably penetrate the nonpolar region of the bilayer as

a pair of helices than as unfolded peptides (Figure 7.2). The next important development built on the recognition that TM helices of a polytopic membrane protein could independently insert into the membrane as a first step in assembling the helical bundle, the premise ofthe widely accepted two-stage model for folding membrane proteins (Figure 7.3). In stage I, the hydrophobic (Xhelical TM segments partition into the bilayer, and then in stage n, they assemble by packing together. For some proteins, there is a third stage that involves the binding of prosthetic groups, the Folding of loops. the assembly of oligomers, or even the movement of other segments into the bilayer region of the protein, all of which can occur once the bundle of helices creates an environment that is both more organized and m01-e polar than the bilayer itself. Stage I of the two-stage model describes the insertion of each individual Oi.-helix as driven by the hydrophobic effect and stabilized by hydrogen bonding along the backbone. The prediction that TM segments can insert independently is supported by the partitioning behavior of peptides that result from either gene splicing that cuts polytopic membrane proteins into separate TM pieces or synthesis of peptides that correspond to single TM domains of membrane proteins. Furthermore, the TM segments observed in known structures of membrane proteins fit quite well with those predicted based on hydrophobicity algorithms. supporting the idea that they each could interact with the lipid nonpolar domain before clustering together to decrease protein-lipid contacts.

Protein Folding

163

A.

STAGE I

STAGE II

7.3. The two-stage model for folding helical membrane proteins. A. The two-stage model for folding helical membrane proteins consists of (I) helix insertion and (II) helix interaction. In stage I, the stability of individual helices is postulated to allow them to insert as stable domains in the lipid bilayer. In stage II, side-to-side helix association results in a folded and functional protein. From Popot, J.-L., and D. M. Engelman, Annu Rev Biochem. 2000, 69:881-922. © 2000 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org. B. A third stage for some proteins involves additional folding steps, such as loop rearrangement (top), or addition of prosthetic groups (P, bottom). From Engelman, D. M., et al., FEB5 Lett. 2003, 555: 122125. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

solvation of the protein (or peptide), the perturbation of the lipid by the protein, and the loss of degrees of fyeedom of the protein. Folding to <x-helix is likely to be induced by partitioning, when the L':.G of partitioning the folded peptide is more negative than the L':.G of

For thermodynamic analysis, stage I can be further separated into three stages: partitioning, folding, and insertion (Figure 7.4). As described in Chapter 4, the hydrophobic side chains provide more than enough free energy for parti tioning, even considering the changes in

/

10""·'"1 path

1~

tJGif

dGW tJG WII

~ tJ G wiu

Water path

~

~

Unfolded (u)

Partitioning

...

~

~

tJG w {

Folding

Folded (f)

Insertion

1

dG,.

~ tJ Gwhf

+tJG""ha

r I ............ .........

~

tJG wa

Association (a)

7.4. The four-step model for the thermodynamics of folding helical membrane proteins breaks stage I into three steps: partitioning, folding, and insertion. All four steps are represented occurring in the aqueous phase as well as at the bilayer, with the possibility of partitioning at each step. The process can follow either the f'.Gw,u or f'.Gw.F step, or a combination of both. The f'.G symbols indicate the standard transfer free energies, with subscripts indicating the steps: w, water: i, interface; h, hydrocarbon core of the bilayer; u, unfolded; f, folded; and a, association. For example, f'.G wi ( is the standard free energy of transfer from water to interface of a folded peptide. This allows the overall standard transfer free energy to be described by a sum of contributions from the various steps, which may be obtained from different types of experiments. From White, S. H., and W. C. Wimley, Annu Rev Biophys Biomol 5truct. 1999, 28:319-365. © 1999 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

164

BOX 7.1. Energetics of folding and inserting a hydrophobic ex-helix into the bilayer

While numerous peptide studies have been carried out, it is not feasible to obtain values of all the free energy changes depicted in Figure 7.4 from experiments for several reasons. First, most peptides that are sufficiently hydrophobic to partition into the membrane will aggregate in water and possibly in the interfacial region of the lipid bilayer as well. Without changing experimental conditions, most such peptides will either partition to the interface or perhaps insert across the bilayer, and it is difficult to control the insertion step. Finally, peptides that form a helix in the bilayer will not unfold in the bilayer. The last of these reasons is illustrated by considering the energetics associated with folding and insertion of a peptide of 20 hydrophobic amino acids. When the peptide forms an a-helix in aqueous solution, the net number of hydrogen bonds does not change significantly, since there is an exchange of some of the hydrogen bonds with the water for hydrogen bonds along the helical backbone. When the peptide partitions into the bilayer, the favorable partitioning of its nonpolar side chains into the bilayer gives an estimated free energy change on the order of -30 kcal/mol, mainly due to the hydrophobic effect discussed in Chapter 1. This is the favorable ~G for inserting the hydrophobic helix into the bilayer. In contrast, if the unfolded peptide transfers from water to the nonpolar milieu of the bilayer, it will dehydrate and the loss of hydrogen bonds is estimated to cost about 40 kcal/mol. Inserting the backbone of the helix into the nonpolar region of the membrane is unfavorable, since the hydrogen-bonded peptide bonds are hydrophilic. The cost of moving each peptide bond from the water to the bilayer, based on the ~Gtr obtained for partitioning an internally hydrogen.bonded peptide bond from water into an alkane, is +2.1 kcal/mol. Including the partition. ing of the aC of each residue brings that cost down to + 1.15 kcal/mol, which can be viewed as the per residue cost of transferring a polyglycine a·helix into the bilayer. It means the full cost of dehydrating the backbone of a 20-residue a-helix is 23 kcal/mol. What is the cost if the peptide is not folded into a helix? The ~Gtr for a peptide bond that is not internally hydrogen bonded to partition into alkane is +6.4 kcal/mol. Again, partitioning of the aC is favorable, -0.95 kcal/mol, so the cost of partitioning a residue of an unfolded peptide into the nonpolar milieu of the bilayer is about +5.4 kcal/mol. For a helix .in the bilayer to unfold, the per residue cost is therefore +5.4 kcal/mol - 1.15 kcal/mol (because the cost of dehydration has already been paid), which comes to ~4.2 kcallmol. This means the cost to unfold a 20-residue helix in the bilayer is 84 kcallmol! Clearly the energetics depend on the amino acid composition of the TM helix. In the example of glycophorin, the membrane-spanning helix has 19 hydrophobic residues. The ~G for its insertion is estimated to be -36 kcallmol. Since the cost of dehydrating the helix backbone is +26 kcal/mol, the net energy favoring insertion is -10 kcal/mol.

Protein Folding and Biogenesis

partitioning the unfolded peptide (see Box 7.1). Insertion of the helix to span the bilayer restricts both its rotational degrees of freedom and its space, in addition to ordering the nearby acyl chains. These entropic costs are also compensated by the hydrophobic effect. Since stage I is determined by interactions relating to the hydrophobic effect, other factors must drive the assembly of helices in stage n. These factors include intrinsic forces such as packing, electrostatic effects, and interactions among the loops between helices, as well as interactions with prosthetic groups or with components at the surface of the membrane. Hydrogenbonding side chains (Asp, Asn, Glu, GIn, Sec Thl; Tyr; His) are often important in helix-helix interactions (see Chapters 4 and 6). As helix packing increases helix-helix interactions, it also increases lipid-lipid interactions while decreasing helix-lipid interactions. This effect lessens the the en tropic cost of helix-lipid interactions due to the packing of relatively "soft" lipids against the rigid, r'elatively "hard" helices. Helix packing is sufficiently close for helix-helix interactions to be dominated by van der Waals forces. The tight packing between two helices typically involves "knobs" formed by branched residues such as valine and isoleucine fitting into "holes" formed by glycine residues. This was originally seen in the dimerization of glycophorin A, a protein with only one TM helix, as described in Chapters 4 and 6 (Figure 7.5A). Similar knob-into-hole packing has been observed in type HI membrane proteins having multiple TM helices, such as the glycerol facilitator and the calcium ATPase (Figure 7.5B). While insertion and packing of separate TM helices are emphasized in the two-stage model, the loops between helices can also be critical. This was demonstrated in experiments that tested the ability of bacteriorhodopsin (BR, described in Chapter 5) expressed as two fragments to assemble into stable, functional molecules in lipid bilayers. When its seven helices (A to G, see Figure 5.6) ",,:ere combined as two fragments, such as AB + CDEFG, BR usually assembled as a stable protein of the correct structure. Ho\vever, some fragment combinations, such as ABC + DEFG, produced less stable proteins. TherefOI-e the connecting loops between helices C and D, and also between E and F, while not indispensable for assembly, are essential for the stability of the native protein. A different approach tested the contribution of each loop by genetically l-eplacing each with a linker of Gly-Gly-Ser of the same length, and showed that four loops (AB, CD, EF, and FG) contribute Lo the resistance of BR to SDS. Furthermore, only helices A, B, D, E, and also C when its aspartate residues are protonated are able to insert independently into phospholipid vesicles, as expected by the two-stage model. A great deal has now been learned about BR folding in vitro.

Protein Folding

165

A.

B.

Calcium ATPase

GIV(l'l'ol f"acililalor (ClpF)

GlpF

GlpF

Ca ATPase

Hdi.\ I-Heli\ 4

Helix 5-Ht.'li:\ 8

Helix 5-1-ldi'\ 7

7.5 "Knobs-in to-holes" packing is observed at helix-helix interfaces in many helical membrane proteins. A. Glycophorin A dimerizes when the TM helices from two molecules associate, bringing the "knobs" of branched amino acid side chains into "holes" due to glycine residues. Viewed from the top, the close contacts between pairs of valine and glycine residues are evident. Viewed from the side, the proximity of such pairs allows the two helices to interact tightly. The third drawing shows the close contacts between the two TM strands, with interatomic distances given in A. From Smith, S. 0., et al., Biochemistry. 2001,40:6553-6558. © 2001 by American Chemical Society. Reprinted with permission from American Chemical Society. B. Other examples of helix-helix packing are seen in the glycerol facilitator and the calcium ATPase, with the overall structures above and examples of specific pairs of TM helices below. B and the third drawing in A from Senes, A., et aI., Proc Natf Acad Sci USA. 2001, 98:90S6-9061. © 2001 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

Protein Folding and Biogenesis

166

SDS-denatured bacterio-opsi n

12 intermediates, dependent on lipid la teral pressure

1I

intermediate

---

---

Central regions of most helices folded, but ends of helices have to form and helix D lies in lipid headgroup regionlin contact with SDS

t!

:~

Partially folded with native secondary structure, with helix D inserted, bu t different helix packing in the two intermediates

Functional bacteriorhodopsi n

IR

intermediates

Schiff base formation possibly involving another retinal-protei/1 intermediate retinal

~

retinal-protein - - intermediates

_?)R~ ~

Two intermediates, and I R440 , with retinal noncovalently bound formed in parallel

IR380

7.6. Intermediates in the folding pathway of bacteriorhodopsin. Kinetic techniques have distinguished three intermediates, 11,12, and IR, in the process of refolding BR from its SDS-unfolded state, which has roughly half its helical content. The two forms of IR that are distinguished by their absorption maxima are affected by pH. Redrawn from Allen, S. J., et aI., J Mol BioI. 2004, 342:1279-1291. © 2004 by Elsevier. Reprinted with permission from Elsevier.

BR Folding Studies BR was the first integral membrane protein to be fully unfolded and then refolded to regain its activity. Early studies demonstrated its complete unfolding requires formic acid or anhydrous trifluoroacetic acid; when transferred into SDS, it regains about half its helical content, and then when added to retinal and mixed micelles (e.g., cholate or CHAPS and lipid), it refolds to the native structure. Since BR sufficiently unfolds in SDS to lose its native structure and its chromophore, kinetic studies can follow the folding and insertion of SDS-unfolded BR into micelles (or bilayers) using circular dichroism, fluorescence, or absorption spectroscopy. The protein in the absence of retinal is called bacterioopsin (BO). Binding of retinal to BO is marked by the return of the absorbance maximum at ~560 nm, as well as by its quenching of intrinsic fluorescence. Stopped-flow absorption spectroscopy and circular dichroism reveal a rate-limiting, pH-dependent, and lipid-dependent step that produces an intermediate with most of the normal helical content. This could be

attributed to the slow folding of some of the helices. In this case BR folding would not strictly follow the two-stage model. Instead it would follow a pathway that involves formation of an intermediate composed of a five-helix bundle, after which the last two helices (F and G) are inserted as a helical hairpin before retinal can bind (Figure 7.6). An alternative explanation is that all seven TM helices form incompletely at this intermediate stage. Additionally there is now evidence that a middle helix, D, inserts to become TM during this step. The simplest pathway consistent with the kinetic data from folding SDS-denatured BR is:

R

BO~ II~ h~ IR

-+

BR

(Figure 7.6). The first intermediate, II, is accompanied by changes in the sUIToLtnding lipid/detergent solvent. The second intermediate, 12 , has native-like secondary structure but lacks some tertiary contacts of the native protein. Formation of 12 is rate-limiting in apoprotein formation, and this folding event must occur before

Protein Folding

167

retinal can bind. It has a binding pocket that enables il to bind retinal noncovalently, forming the intermediate IR . Formation of the Schiff base between retinal and Lys216 converts I R lo BR. The intermediate I R has at leasltwo forms distinguished by their absorption maxima at 380 and 440 nm, which are populated differently at different values of pH.

100

" Ly o-PP 90

'"DMPC

80 70 ..

kla

IRJ80

/'

~

/' I R440

"0

60

BR klb

'"

k2a

Iz+R

~

" "

k2b

o

DPoPE

50 ...,....~-r-~--.-~-r-~--.-~-r-~--.-~ 0,0 0.1 0.2 0,3 0,4 0,5 0.6

Mole fraction of lipid added to DPoPC It is likely that the two forms of I R arise from different

conformational states of 12 having different protonation states of groups near the retinal-binding site. Thus there are parallel folding pathways to the same native structure, consistent with the view of the folding process as a funnel of different pathways leading from an ensemble of unfolded states. Refolding BR into lipid bilayers in a detergentfTee environment provides the ability to vary the lipid composition and investigate the effect of the state of the bilayer lipids on membrane protein folding. Experiments have been designed to determine the effect of lateral pressures in the bilayer on the folding and insertion of BR. Lateral pressure derives from the tendency of each leaflet to curve away from the plane of the bilayer (see Figure 2.18). This curvature stress confers bending rigidity to the bilayer, decreasing its elasticity, Such stress can be introduced into reconstituted bilayers by mixing bilayer-forming lipids with a nonbilayer former (e.g., PC and PE) and can be altered by varying acyl chain composition; for example, it is reduced with a shorter chain length that decreases crowding (pressure) near the middle of the bilayer. The yields of BR formed from BO added to retinal and different lipid vesicles are clearly affected by curvature stress of the bilayer (Figure 7.7). The vesicles are made of DPoPC (dipalmitoleoyl PC) under conditions that give 70% refolding. The yield increases with addition of DMPC, and even more with addition of a lysophospholipid having one acyl chain removed to reduce the lateral pressure. The yield decreases with the addition of PE with unsaturated acyl chains, which increases the la teral pressure in the center of the bilayer. In bicelles consisting of different proportions of DHPC (dihexanoyl PC, chain length of six) and DMPC (chain length of 14), the bending rigidity of the bilayer is calculated to increase twofold as the mole fraction of DMPC increases from 0.3 to 0.7. In refolding experiments in this system the rate of BR folding decreased 10-fold over the same range, revealing a clear effect of lateral pressure in the fatty acyl chain region of the bilayer on

7.7. Effect of bilayer curvature stress on folding of bacteriorhodopsin, Three different lipids are added to DPoPC: DpoPE (red circles). which increases curvature stress; DMPC (black triangles). which decreases curvature stress; and Iyso-PPC (blue squares), which relaxes the bilayer still further The folding yields are determined from the area under the purple absorption band of folded functional BR, From Allen, S. j" et aI., J Mol BioI, 2004. 342: 1293-1304, © 2004 by Elsevier Reprinted with permission from Elsevier,

kinetics of BR folding. These results suggest that BO has more difficulty inserting into stressed, rigid bilayers as it refolds to BR. From its assembly as combined fTagments to its refolding in different conditions of bilayer stress, in vitro folding studies with BR have started to produce a sophisticated understanding of the interplay of the protein with the membrane during the folding process. Even with expected differences in the process of folding and insertion in the cell, these studies give valuable insight, especially since in the BR photocycle the retinal becomes detached and rebinds. The elegant folding studies ofBR provide a paradigm for other helical membrane proteins. Folding Studies of I3·Barrel Membrane Proteins

How do f)-barrels fit the model for folding and insertion of membrane proteins? They are fundamentally unlike bundles of a-helices, because each helix can be formed independently with hydrogen bonds along the helix axis while f)-barrels have hydrogen bonds between neighboring strands, even between the strands closest to the Nand C termini. A single f)-strand is nOl a stable structure; rather, formation of at least three to five strands is required for stability in all f)-structures, including domains of soluble proteins. Furthermore, geometric considerations make it difficult to envision a portion of a barrel folding first, so all the strands of a f)-barrel can be expected to form at roughly the same time. Since the f)-barrel proteins are found in outer membranes, they fold and insert from an aqueous

Protein Folding and Biogenesis

168

Structure

A.

Topology

Loops

Extracellular space ,\11, ~

,.

"

,

'.

r

R

t t

DOPC,

hydo-ophobic thickness

-30'\

30.'\

~

I Mem brane

Turns

Periplasm

B. ~

1

/

II

III

N

~~~~ -qj

\

---------~~~Encounter Complex

Molten Disk

Molten Globule

J , 7i , ,(\ , \ '."

' , , '

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Membrane

Native

Location of Tryptophan in Folding Intermediates Identified by TDFQ Distance from Center Tryptophan

(J) (15.57,102,143)

I

I

I 14-16

II 10

III

A-

ID

lOA A

0-5

N

lOA lOA

7.8. Folding studies of the TM barrel domain of OmpA. A. The structure of the (3-barrel domain has been determined by NMR. The topology of OmpA shows the location of its five Trp residues, four of which cross the bilayer in the process of insertion. The position of OmpA in the membrane is indicated schematically by the PL and LPS molecules represented on the left of the figures. B. Schematic drawing of four stages in the insertion of OmpA, with the locations of the Trp residues at each stage as determined with the "spectroscopic rulers." Each circle represents a Trp residue, with the Trp that remains on the inner side marked "7" (see text). From Tamm, L. K., et aI., J Bioi Chem. 2001,276:3239932402. © 2001 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

environment, such as the periplasm of Gram-negative bacteria. For this reason, in vitro folding studies of l3-barrels may be more directly applicable to the in vivo situation than to those of <x-helical proteins that are inserted via a translocation apparatus in cells (see below). Extensive in vitro foldi ng studies of a l3-barrel membrane protein have been done with the l3-barrel domain of OmpA, an eight-stranded monomeric protein that can be unfolded in urea and refolded upon dilution into a suspension of PL vesicles. From the structure solved with NMR (see Box 5.1), the positions of its five tryp-

tophan residues are oriented such that four of them are near one end of the barrel while the fifth is close to the opposite end. During assembly of the l3-ban-eI, four Trp residues must translocate to the ou ter leaflet of the bilayel~ crossing its midsection, while the fifth l-emains in the inner leaflet (Figme 7.8A). In an ingenious experiment, the locations of the Trp residues during the folding process were followed with a set of "spectroscopic rulers," lipids carrying a quenching bromine atom at different positions on their acyl chains. The four positions of BR reported on locations at the interface, 14 to 16 A from the center of the bilayer, around 10 A

Protein Folding

169

from the center, or at the center of the bilayer (without distinguishing between the two leaflets). Using I)-barrel domains from OmpA proteins that had all but one of the Trp residues deleted, the spectmscopic rulers could detect the position of each remaining tryptophan during folding. As predicted, four Trp residues cross the bilayer while the fifth stays at the J0 Aposition during insertion. The time course of insertion suggests a model for l)-balTel assembly from a flattened disc composed of I)-strands at the interface to an inserted "molten globule" as the I)-strands penetrate the bilayer in a relatively slow temperature-dependent step (Figure 7.8B). Like folding of BR, folding of the OmpA I)-barrel domain is affected by stresses in the bilayer. Bilayer stresses were created by refolding in lipid compositions with varying chain lengths, with different headgroups, and with doubly unsaturated chains to intmduce curvature stress. Refolding was observed both by fluorescence and electrophoretic mobility. The reference bilayer was PO PC with 7.5 mol % POPG, which gives two-state folding with a free energy of unfolding of 3.4 kcallmol. The 6GO for unfolding was linearly proportional to chain lengths when saturated and monounsaturated acyl chains were used, and inversely proportional to chain lengths when doubleunsaturated acyl chains with more curvature stress were used (Figure 7.9). It also increased with increased lateral pressure due to addition of PE. The studies with both BR and OmpA make it clear that folding and stability of membrane proteins are affected by the 8

di-C I4 : IPC

di-C I8I PC

6

di-C I8I PC di-C 20I PC

E 4

CI8:0CISIPC

,......, I

A.

Individual leaflets prefer to curve

~~~~j\

~U~V~

I B.

Curvature frustration when forced into bilayer

~~IT~~~~~~~~~

UVU~V~~V~~~~

I c. CUI-vature frustration relieved by hourglassshaped protein

~ Cylindrical lipid

?l

(bilayer)

ACone-shaped lipid 1~ (nonbilayer)

7.10. Effect of protein insertion on bilayer curvature stresss. Bilayer stress resulting from incorporation of a non bilayer lipid (cone shaped) can be relieved by insertion of a protein with an hourglass shape. Redrawn from Bowie, J., Nature. 2005, 438:581589. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

0

C;; u

~

CI8:0CISIPC di-C I4 PC

a 2

:£' ~

G
material characteristics of the bilayer, in particular the bending rigidity resulting h-om curvature stress (Figure 7.10). Other Folding Studies

0 di-CloPC -2 15

20

25

30

35

dhydrophobic (A) 7.9. Folding studies of the TM barrel domain of OmpA reveal the effects of bilayer stresses. The graph shows the free energy of unfolding (L'>Go u, in kcal/mol) as a function of the hydrophobic thickness of the PC bilayer. For saturated and monounsaturated chains (filled circles), the L'>Go u increases linearly with chain length (thickness), while for cis double-unsaturated acyl chains (open circles), it decreases linearly with chain length. From Hong, H., and L. K. Tamm, Proc Natl Acad Sci USA. 2004, 101 :4065-4070. © 2004 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

Classical il1 vitro folding studies in lipids or detergents have now been carried out with several other membrane proteins, including the enzyme DGK, the lactose permease, and the potassium channel, KcsA. In addition, some of these studies employed different strategies to gain different types of information about folding membrane proteins. For example. the stability of various mutants of DGK was assessed with thermodynamic analysis of refolding the partially denatured pmtein. When DGK is reversibly denatured in SDS, it retains most of its a-helical content (as is typical of soluble proteins in SDS). The refolding that takes place upon the removal of SDS is sufficient to differentiate the stability of different mutants.

Protein Folding and Biogenesis

170

C

(Y4

7.11. Ribbon diagram showing the lowest energy conformer of Mistic determined by NMR. From Roosild, 1. P., et aI., Science. 2005, 301: 1317-1321. © 2005. Reprinted with permission from

MAS

To study the TM portion of the KcsA potassium channel (see Chapter 9), a novel approach combined pallia] de novo synthesis with folding. The semisynthetic TM protein was constructed by fusion of a recombinant peptide (residues 1-73) with a synthetic peptide (residues 74-125). The refolding of this construct was then used to define the lipid requirement for folding. Folding of membrane proteins can be a very important step of theil- purification after they are harvested in denatured form from insoluble inclusion bodies, which are lipid-bounded storage sites within the cytoplasm of bacterial cells that are overproducing the proteins from high-expression vectors. An example of a membrane protein that was refolded after it was obtained in inclusion bodies is the enzyme OMPLA (described in Chapter 10). The denatured OMPLA was solubilized in 8 M urea and diluted into Triton X-lOa, which produced a mixtlll'e of folded and unfolded OMPLA that was resolved by ion-exchange chromatography to recover the native enzyme. The choice of detergent is critical, as refolding studies under different conditions revealed a strong preference for Triton X-lOa in the case of OMPLA. Often, however, refolding from inclusion bodies is not effective for membrane proteins. An alternative procedure is now available that makes use of Mistic (the acronym for Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs), a l3-kDa protein from Bacillus subtilis that spontaneously inserts into membranes in vivo. The structure of Mistic \.vas determined using sophisticated NMR methods along with site-directed spin labeling, which revealed a four-helix bundle with an exposed periplasmic C terminus (Figure 7.11). In spite of its hydrophilic character, Mistic associates tightly with the membrane. When it is expressed as a fusion with an integral membrane protein, it has a unique ability to insert the TM domains from the membrane proteins. Over 20 eukaryotic membrane proteins have now been overexpressed

in E. coli as Mistic fusions, and those tested had functionality. The use of Mistic might overcome the usual problems caused by expressing high amounts of eukaryotic membrane proteins in bacteria: (l) their targeting signals may not be recognized by the host, and (2) their overexpression is toxic because it clogs the machinery for inserting the bacterium's own membrane proteins or causes other problems like proton leaks. Finally, the use of in vitro transcription/translation systems to synthesize mem brane proteins in the presence of lipid vesicles or micelles has revealed conditions needed for folding and insertion. This approach was pioneered in studies of the E. coli outer membrane protein PhoE (see Chaptel- 5) and demonstrated that folding and insertion of PhoE require lipopolysaccharide. In similar studies of the E. coli lactose permease (see Chapter 10), protein folding took place only with inside-out vesicles, mimicking the vectorial insertion of the lactose permease into the inner membrane.

BIOGENESIS OF MEMBRANE PROTEINS

Folding and insertion of membrane proteins are just two elements of the very complex process needed for the assembly of proteins into cell membranes. The first steps of that process are shared by membrane and secreted proteins and utilize a complex apparatus called a translocon that has been conserved through eukaryotic, bacterial, and archaei kingdoms. In later steps integral membrane proteins are differentiated from secreted proteins by a process that allows their lateral integration into the lipid bilayer and detel-mines their topology. The process of integration and topogenesis (the genesis of topology) will be examined after describing the process for export and the steps involved in translocation. Export from the Cytoplasm

Since membrane proteins are not synthesized by a special population of ribosomes, they must be targeted for their destinations as they exit the ribosomes. In eukalYotes, they may enter the ER and end up in the plasma membrane or endocytic organelles (the ER pathway) or they may follow other pathways for mitochondrial, chloroplast, and nuclear membranes. The destinations in prokaryotes are the plasma membrane and, in Gramnegative bacteria, the outer membrane. Once targeted for export, the processes of translation and translocation may be coupled (cotranslational translocation) or sequential (posttranslational translocation). Three types of signals initiate the export of membrane proteins: cleavable signal sequences, signal. anchors, and reverse signal-anchors. (Since reverse signal-anchors differ only in the resulting orientation

Biogenesis of Membrane Proteins

171

BOX 7.2. EVidence for cleavable signal sequences Involved in protein translocation The protease protection assay distinguishes between translocated membrane proteins and secreted proteins and provides evidence that both can have cleavable signal sequences. The experiment uses SDS polyacrylamide gel electrophoresis (PAGE) to distinguish products of in vitro protein synthesis in the presence or absence of membrane vesicles, with and without added protease. When translation of a protein with a classical N-terminal cleavable signal sequence is carried out in vitro in the absence of membranes, a precursor containing the signal is produced. The precursor is not protected by membranes, so it is digested by added protease (sample 1 in Figure 7.2.1). When it is carried out in the presence of membrane vesicles, the signal insert into the membrane, along with any TM segments of the protein (samples 2

and 3). Signal peptidase on the membrane cleaves the signal from both secreted and inserted proteins, resulting in mature forms that migrate faster on SDS-PAGE as shown. To determine whether the protein is integrated into the membrane or secreted across it, a nonspecific protease such as proteinase K is added. If the protein is soluble and outside the vesicles, it is fully digested (sample 1); if it is integrated into the membrane, the external portion is digested, resulting in a smaller protein or fragments (sample 2); and if it is fully transported into the vesicle (secreted), it is protected from the protease and retains its size (unaltered mobility on the gel, sample 3). Many applications of this basic protocol have revealed the nature and role of signal sequences on hundreds of proteins.

CD ...... .....;;;:i{L Q)

~ .C

....<}:

Simple membrane protein

,

Signal peptidase Q)Full translocatio~ of secreted proteIn SDS·PAGE

I

Before external protease 2

3

After external protease 1

2

3

Precursor_ Mature form _

7.2.1. In vitro assay for protein insertion into membrane vesicles, utilizing protease sensitivity (top) and analysis by 50S-PAGE. Based on unpublished figure from Professor A. Driessen.

Protein Folding and Biogenesis

172

NH

r

~t.

++ 1-5 aa

3-7 aa mature chain

7-15 aa Hydrophobic

7.12. Signal sequences for the ER pathway and bacterial export systems follow the same fundamental design. They have a basic N terminus followed by a stretch of seven to 15 nonpolar residues; then three to seven hydrophilic and charged residues precede the cleavage site. Redrawn from von Heijne, G., in W. Neupert and R. Lill (eds.), Membrane Biogenesis and Protein Targeting, Elsevier, 1992, p. 77. © 1992 by Elsevier Reprinted with permission from Elsevier.

of the membrane proteins, they are discussed further under Topogenesis in Membrane Proteins.) Cleavable signal sequences at the N termini of nascent peptides are present in many proteins targeted to the ER

3'

pathway in eukaryotes and the export pathway in bacteria. Their presence is readily recognized experimentally by the larger size, hence slower electrophoretic mobility on SDS polyacrylamide electrophoresis (SDS-PAGE), of the precursor protein that emerges from the ribosome compared to the mature, functional protein (see Box 7.2). The precursor proteins have an N-terminal signal sequence of ~20 amino acids, consisting of a few polar and basic residues followed by a stretch of seven to 15 primarily hydrophobic residues before the polar cleavage site at the beginning of the mature protein (Figure 7.12). While not all N-terminal signals are cleaved, the cleavable signal is the first type of signal sequence that was recognized by researchers and is sometimes called the classical signal sequence. In those integral membrane proteins that lack a signal sequence, their first TM segment functions to

5'

I

Eubacterial SRP system E. coli

Periplasm

3'

5'

,c

_ ,

~

l

AlII; S

Eukaryotic SRP system human

Endoplasmic reticulum 7.13. The core components of the E. coli and human signal recognition particle (SRPj and its receptor are highly homologous. Both SRP54 in eukaryotes and the Ffh protein of the E. coli SRP have three domains. The M domain is methionine rich and binds RNA (lavender). The N domain is the ex-helical domain, and the G domain is a nucleotide-binding site with GTPase activity (drawn together in purple). The receptor, which is FtsY in E. coli and SRex in eukaryotes, also has similar domains, homologous N and G (light pink) and A, the N-terminal domain (dark pink). The remaining proteins of the eukaryotic SRP along with SR(3 do not have homologs in E. coli, and the larger eukaryotic RNA has an Alu domain that, along with two of the proteins, functions in translational arrest. From Luirink, J., and I. Sinning, Biochim Biophys Acta. 2004, 1694: 17-35. © 2004 by Elsevier Reprinted with permission from Elsevier.

Biogenesis of Membrane Proteins

173

Periplasm

G

G

Cyloplasmic membrane

--

Cytoplasm

I

COO-

7.14. Cartoon representation of the role of SecA in the translocation of nascent peptides. The SecS chaperone binds the nascent chain (1) and brings it to SecA (2). This complex binds the translocon composed of SecY, SecE, and SecG (3). Then SecA binds and inserts into the translocon, carrying the bound precursor (4). When SecA hydrolyzes the ATP, it withdraws from the translocon, while the peptide remains engaged within the channel (5). SecA binds ATP and another segment of the peptide chain to repeat the cycle (6). The changes in the line for SecG (gold) indicate the topology inversion that appears to accompany SecA insertion. From Mori, H., and K. Ito, Trends Microbial. 2001,9:494-500. © 2001 by Elsevier. Reprinted with permission from Elsevier.

initiate topogenesis (see below) and is called a signalanchor. Since they must span the bilayer, the signalanchor sequences are longer, with 18 to 25 mostly nonpolar residues. Responsible for both insertion and anchoring, they are not necessarily at the N termini of the proteins. Other types of sorting sequences occur in the middle or even at the C termini, some of 'which are involved in the topogenesis of polytopic membrane proteins, as described below. In addition, there are signal sequences for proteins destined to be secreted beyond the periplasm or even beyond the ou tel' membrane, such as the TolC-HlyABC system (see Chapter 11). Signal sequences are highly degenerate. Their primary structures vary greatly and can be altered significantly 'without affecting their roles, as long as the nonpolar region is sufficiently hydrophobic and long enough (but not so long as to become a TM segment). This is because their requisite featuIT is simply a helical stretch of hydrophobic amino acids to bind to a hydrophobic groove on a chaperone or piloting protein, which recognizes the nascent signal as it emerges from the ribosome, protects the nascent chain from aggregation, and targets il to the membrane. Tbis role is often carried out by a ribonucleoprotein complex caJled the signal recognition particle (SRP). In eukaryotes the SRP is composed of six proteins and a 7S RNA. It slows the rate of elongation and brings the nascent chainribosome complex to Ibe SRP receptor of tbe ER membrane. Botb SRP and its receptor are GTPases, and when the SRP is docked, the catalytic sites on both SRP and the SRP receptor come together to form a unique catalytic chamber that binds two GTPs. Once

bound, they stimulate each other's GTPase activity. The hydrolysis of GTP by the complex releases SRP, which enables the ribosome to dock on the translocon and resume translation coupled to translocation, producing cotranslational translocation. The E. coli SRP is composed of a 4.5S RNA and the Ffll protein and carries out the same functions with the exception of translational arrest (Figure 7.13). The E. coli SRP and iIs receptor in the inner membrane, FtsY, are required for integration of very hydrophobic proteins into the inner membrane. The SRP pathway is not utilized by alJ membrane proteins in E. coli. Those that are less hydrophobic, including those destined for the outer membrane, are piloted by SecB and SecA. SecB is a tetrameric cbaperone that prevents newly synthesized proteins from folding or aggregating in the cytosol by binding to exposed hydrophobic surfaces without an expenditure of ATP. (Other chaperones, e.g., CsaA, play tbis role in other bacteria.) SecB has well-defined binding sites for SecA and pilots the nascent peptide to the SecA protein. Transfer of the nascenl peptide chains from SecB to SecA releases SecS and allows Ihe initiation of translocation. SecA provides the energy for translocation. It is highly conserved in bacteria, functions as a dimeI', and undergoes conformational changes, allowing it to be in the cytoplasm, peripheral to the membrane or inserting into the translocon (see Chapler4). SecA hydrolyzes ATP to drive the insertion of 30- to 40-residue segments of the nascenl peptide chain through the translocon in a repetitive cycle (Figure 7.14). The activity of SecA is modulated by SecG, a component of the translocon that

Protein Folding and Biogenesis

174

~

1 ~(~

\ GDP

+ Pi

t ~ ATP Cytoplasm

'Inner

~embrane Periplasm

~~ 7.15. The two pathways, the SRP pathway and the Sec pathway, that target proteins for export from the cytoplasm in E. coli. Both pathways start with the ribosome, where the emerging nascent peptides interact with ribosomal protein L23 in the exit tunnel. SRP with its FtsY protein binds to signals that are especially hydrophobic and bring the nascent chain-ribosome complex to an unidentified site on the membrane for translocation (Ieh side). Alternatively (right side), SecB binds the nascent chain and brings it to SecA. The complex binds the SecYEG translocon and SecA hydrolyzes ATP as it catalyzes insertion of the peptide. As the peptide reaches the periplasm, the signal is hydrolyzed by leader peptidase (Lep). Then either the protein is released with the help of SecDF and YajC, or it inserts into the bilayer with the help of Vide. TF is trigger factor, a cytoplasmic protein that interacts with the peptide as it emerges from the ribosome. From Luirink, J., and I. Sinning, Biochim Biophys Acta. 2004, 1694: 17-35. © 2004 by Elsevier. Reprinted with permission from Elsevier.

appears to invert its topology concomitant with SecA insertion. A nascent peptide destined for the membrane (or secretion) may be recognized in the exit tunnel of the ribosome. Fluorescence and cross-bnking experiments indicate that two proteins of the large subunit make contact with helical TM segments of nascent membrane proteins and not with signal sequence segments of nascent secretory proteins. This contact may also

involve contact between the helical TM segments and the translocon (the translocation apparatus described below). One of the ribosomal proteins, L23 at the exit of E. coli ribosomes. makes contact with SRP and a chaperone called trigger factor (TF), and then \,\Iith the translocon when the ribosome docks at the membrane (Figure 7.15). Thus TF may playa role in sorting proteins between the SecB/A pathway and the SRP pathway.

Biogenesis of Membrane Proteins

175

TABLE 7.1. Components of the protein transloction apparatus In bacteria and mammalian cells·

Function Translocon core Homologs

Piloting complex Pilot receptor Driving force Chaperones In Out Insertion aid Other components

Bacterial system: E.

coli

SecYEG SecY SecE SecG Ffh + 4.5 S RNA FtsY SecA

+ ATP;

pmf

SecS Skp, SurA YidC

Mammalian system: ER Sec61cx(3y Sec61cx Sec61y Sec61 (3 SRP 6 proteins + 7S RNA SRP receptor Ribosome + GTP

=

Hsp70 Sip TRAM

SecD, SecF, YajC

The bacterial system is represented by that of E. coli, and the mammalian system is that found in the ER. These may be compared with the translocon from the archaebacteria Methanococcus jannaschii described in Chapter 11. Based on Wickner, W., and R. Schekman, Science. 2005, 310: 1452-1456, and earlier papers.

a

The Translocon

Remarkably, the insertion of nascent membrane proteins into either ER membranes or bacterial plasma membranes generally utilizes the same translocation apparatus that exports most proteins bound for extracellular destinations. Therefore the translocon is a pore that functions in two dimensions and recognizes which proteins go into the membrane and which go across it into the lumen of the ER or the periplasm of Gramnegative bacteria. It does this while maintaining the permeability barrier of the membrane, even though its channel is large enough to accommodate at least one ex-helix. The translocon is highly conserved: the mammalian translocon has subunits Sec61 ex, 13, and y, with homo logs in yeast and bacteria (see Table 7.1). A unified picture of this extraordinary molecular apparatus is now complemented by the high-resolution structure of the translocon from Methanococcus janl1aschii (see Chapter 11). In the crystal structure of the iVl. jwmaschii translocon, the Sec heterotrimer has a visible pore and therefore could function as a

unit. This is not consistent with higher-order assemblies that were postulated after larger structures were observed with cryo-EM. However, the translocon may oligomerize in different complexes to carry out different functional roles, as it appears to be quite promiscuous in its dynamic association with different partner proteins. Additional Proteins Involved in Translocation

Many of the proteins involved in translocation are named Sec, for secretion. The core of the E. coli translocon consists of SecY and SecE proteins, which are essential for viability. The nonessential SecG protein stimulates translocation by facilitating the insertion-deinsertion cycle of SecA that moves peptide segments through the channel. In addition to the SecYEG pore, a number of other components play roles in the translocation process in the E. coli membrane (Table 7.2 and Figure 7.15). SecD and SecF, along with the small protein YajC, assemble at the translocon on the periplasmic side to help in protein release. Their precise function is unclear, but they appear to stabilize the inserted form of SecA, to help release translocated proteins from the membrane, and to mediate the interaction between SecYEG and YidC in the biogenesis of membrane proteins. YidC, an integral membrane protein that binds to the Sec OF component of the translocon, is clearly involved in the insertion of membrane proteins, although its mechanism of action is unknown. It could function either as a membrane-bound chaperone for TM segments or as a catalyst for membrane integration. YidC is essential for insertion of Sec-independent and some Sec-dependent poly topic membrane proteins. The Sec-independent coat proteins from the bacteriophages M 13 and Pf3 require YidC to adopt their TM conformation. YidC is found in both Gram-positive and Gram-negative bacteria; it has homologs in mitochondria (Oxa I) and in chloroplasts (Alb3). An additional participant that works with the translocon to integrate membrane proteins in eukaryotic cells is called TRAM (translocating chainassociated membrane protein). TRAM is either stimulatory or required for protein translocation, depending

TABLE 7.2. Proteins involved in protein translocation in E. colr" Protein

SecY

SecE

SecG

SecA

SecD

SecF

YajC

YidC

Relative mass (kDa)

48

14

12

102

67

35

12

62

# TM spans Partners

10 SecEG

3

2

6

6

1

6

SecYG

SecYE

b

SecS

SecFYajC

SecDYajC

SecDF

Numerous proteins are involved in protein translocation in E. coli in addition to those that make up the translocon. Not listed are proteins involved in the TAT pathway and in tripartite systems that export hemolysin and other toxins. bSecA inserts into the membrane as part of its catalytic cycle. See text and "SecA: Protein Acrobatics" in Chapter 4. Based on Veenendaal, A. K. J., et al., Biochim Biophys Acta. 2004. 1694:81-95.

a

Protein Folding and Biogenesis

176

· 11I Cf ,

Endoplasmic reticulum

<

~;:

l.

,

---

\

/!

Ribosome

'---~

TM

~

~,

I,

'.

\

-

Cytoplasm

Membrane Lumen 7.16. Insertion of proteins into the ER membrane utilizes the Sec61 translocon aided by TRAM. After the nascent chain binds to SRP as it emerges from the ribosome, the complex docks on the SRP receptor at the membrane. Hydrolysis of GTP releases SRP and initiates translocation. TRAM facilitates the lateral movement of TM segments into the bilayer. From Dalbey, R. E., et al., Curr Opin Struct BioI. 2000, 12:435--442 © 2000 by Elsevier Reprinted with permission from Elsevier.

on the protein. Data from cross-linking experiments of the sari described in Box 7 A show that TRAM specifically interacts with signal sequences during their passage through the Sec61 channel and associates with TM segments as they leave the translocon and integrate into the lipid bilayel- (Figure 7.16). TRAM recognizes some sequences based on their length and charges and may use this information to guide the integration of TM segments to provide the correct topology. TRAM, along wi th the Sec61 ex, (3, and y subunits and the SRP, was sufficient to reconstitute in vitro protein translation and integration into liposomes. Other proteins that play imparlant roles as the nascent polypeptide gets across the membrane include the signal peptidase and chaperones. When exported proteins have cleavable N-terminal signal sequences, they are cleaved by signal peptidase (also called leader peptidase, or Lep), which is anchored with its active site on the noncytoplasmic side of the membrane. In the E coli peri plasm, chaper'ones such as Skp and SurA prevent aggregation of outer membrane proteins secreted by the translocon and may help them fold (although in vitro studies of OmpA folding suggest that folding and inser"tion into the outer membrane of the (3-batTel proteins occur spontaneously and are highly synchronized), Also in the periplasm, the Dsb system oxidizes cysteine residues to form disulfide bonds, and other enzymes make bonds to lipid or lipopolysaccharide. Similarly, covalent modification of the nascent proteins to bind sugars and/or lipid is a crit-

ical part of the maturation process in the ER lumen of eukaryotes. Before examining the detailed mechanism of translocation, it is helpful to review the molecular picture of the process of protein export in E coli or protein translocation in eukal)'otes based on the characterization of the many proteins involved in the two pathways. Cytosolic chaperones and targeting complexes, either SecB/SecA or the SRP and its receptOI- (Ffh/FtsY), guide the ribosome-nascent peptide complex to the translocation apparatus in the membrane. In cotranslational translocation, the growing peptide chain threads from the ribosome into the Sec translocon and ei ther crosses to the periplasm or the ER lumen or inserts into the lipid bilayer. For protein secretion, the signal is cleaved by signal peptidase as it emerges from the translocon; when translation and translocation are complete the protein is released. For integration of membrane proteins, the translocon recognizes hydrophobic TM segments as signal-anchors or stop transfer signals (defined below) and alJows them to integrate laterally into the lipid milieu of the membrane, as discussed below and in Chapter II. Besides the Sec translocon, other translocation mechanisms exist in the cytoplasmic membrane of E coli, including the YidC pore for integration of membrane proteins and a poorly understood system called the twin arginine translocation (TAT) path\vay that transports folded proteins to the periplasm. Also, eukaryotic cells have specialized mechanisms for protein import into chloroplasts and mitochondria (see Box 7.3).

Biogenesis of Membrane Proteins

177 -

- - -

BOX 7.3. Import of mitochondrial proteins Nearly all (98%) of the ~1 000 mitochondrial proteins are synthesized in the cytosol and imported into the mitochondria, destined either for the matrix, the intermembrane space, the inner membrane, or the outer membrane. The imported proteins are classified in two major groups: precursor proteins

~

'''''''' "'1110

r""

with cleavable amino-terminal signals, called presequences, and hydrophobic membrane proteins with targeting signals dispersed throughout their primary structures. In addition, the proteins of the outer membrane and the inter membrane space constitute special classes.

pn:~tlrwrrmtdn!

TOM

~UI~I"rlI1T""I.·on.

, Ill!

""I.

11l["nul! .Iiln.,·,

II,

II11er",,1 ',gnub

nn( /J

)

~

+f\Tr ATP ~

.. TOO1Zl .... lhm5 Cytosol

!i)]l120

•

OM

Tom70

,...,.,

~

Tom7

TOln6

T011140· 1011140:

, IMS

c

r,."lllnl'_'·'!''')1 ,"Inl·l,·~

,\l;llilt

llInn

t'lCrtlbr,'I'l:t,rol"ltl

TIM23

P"cpr01cin \\'illl

~

•C

\\. , ,

'"

.

OUh'!" mCfllhr~uw

Inl"""cr'!h";"l.~ ~r;K"

{jjJ

M:lllor,

maldx protein

7.3.1. Model of the mitochrondrial import pathway showing the components of the translocation across the outer membrane (OM) in the TOM complex and the two translocases for the inner membrane (1M). TIM22, and TIM23 complexes, in addition to chaperones and other helpers in the intermembrane space (IMS) and matrix. The insets show the protein components and the functions of the three complexes. From Pfanner, N., and N. Wiedemann, Curr Opin Cell BioI. 2002, 14:400-411. © 2002 by Elsevier. Reprinted with permission from Elsevier. (continued)

Protein Folding and Biogenesis

178

BOX 7.3 (continued) All imported proteins are transported across the outer membrane of the mitochondria via the translocase of the outer membrane (TOM). Two more translocases are located in the mitochondrial inner membrane: the TIM23 complex transports cleavable preproteins, while the TIM22 complex inserts hydrophobic proteins into the inner membrane (see Figure 7.3.1). The components of these complexes are proteins that act as receptors, form pores, and/or exhibit chaperone functions and are well characterized in yeast. These complexes appear to accumulate at adhesion sites where the inner and outer membranes are 18 to 20 nm apart, which could allow protein complexes in the two membranes to cooperate during translocation. The TOM Complex

Translocation across the outer membrane is a passive process mediated by receptors (Tom20, Tom22, and Tom70) and a general import pore (GIP). The presequences of cleavable precursor proteins are positively charged and can form amphipathic helices that fit into a hydrophobic groove of Tom20. They are transferred to Tom22 by its recognition of the opposite, charged surface of the helices. Tom70 is a receptor for hydrophobic precursors and forms oligomers upon binding them, which could help prevent aggregation. The main component of the GIP is Tom40, which forms a pore with a diameter of around 20 A, large enough to accommodate two ex-helical polypeptide segments but not a folded protein. In the membrane, Tom40 functions with the smaller proteins TomS, Tom6, and Tom7, which may help stabilize several Tom40 dimers to form a large, dynamic complex containing several pores. Interestingly, in biogenesis the import of precursors to the Tom proteins utilizes the TOM appara-

Integration of Nascent Proteins into the Membrane

Biogenesis of membrane proteins may start to be differentiated as early as their passage through the ribosomal exit tunnel, yet the point of demarcation from other exported nascent proteins is their transit from the aqueous channel of the translocon to insert into the lipid bilayer: The mechanism of insel-tion determines the alTangement of the TM segments, and thus the signals controlling insertion determine the topology of the protein. These topics will be discussed starting with methods that reveal the steps in the process of insertion of TM segments, before describing more fully the types of signals and their topogenic consequences. Surprising dual pathways through the translocon into the lipid bilayer and to the other side of the membrane have been discovered using sophisticated photoactivated cross-linking methods. These experiments use knowledge of the location of the crosslinking residue in the translocating peptide to follow its progress by irradiation at different times (see Box 7.4). The nascent peptides carrying TM segments were observed to cross-I ink initially to cytosolic components,

tus, and the import of each Tom protein requires several other Tom proteins. TIM Complexes

The pathways for the two main classes of imported proteins diverge after tanslocation via TOM. Translocation of presequence-containing proteins across the inner membrane uses the T1M23 complex, which consists of Tim23, Tim17, and Tim50 in the inner membrane, plus a peripheral membrane protein, Tim44, and two soluble proteins in the matrix, the mitochondrial Hsp70 (mHsp70 or Ssc1 p) and its helper (Grep or Yge1 pl. When reconstituted in liposomes, purified T1M23 forms a voltage-activated cation-selective channel that is activated by presequence peptides and a membrane potential (~IJI, negative inside). while inhibited by presequence peptides alone. While this suggests that ~ IJI is reo quired for channel opening, translocation across the membrane also uses hydrolysis of ATP. The mitochondrial Hsp70, Ssc1 p, hydrolyzes ATP either to drive the uptake of imported peptides or to unfold and bind them to prevent their return (or both). The mitochondrial processing peptidase (MPP) cleaves the amino-terminal presequence in the matrix. The T1M22 complex is a carrier translocase responsible for insertion of inner membrane proteins such as the ATP-ADP carrier (described in Chapter 5) as well as the Tim proteins themselves. It utilizes the Tim9-Tim1 0 complex in the inter· membrane space, the peripheral protein Tim12, and three integral membrane proteins, Tim22, Tim54, and Tim18, of which only Tim22 is essential for viability. Tim22 uses the membrane potential to drive insertion; when purified and reconstituted, it forms a gated channel with two pore sizes, the larger of which allows insertion of tightly packed loops.

then to subunits of the translocon and then to lipid (Figure 7.17). The data showing that nascent TM segments move from the transloeon into the lipid suggest that conformational changes in the subunits of the translocon create a sideways opening to allow lateral movement of the peptide out of the translocon. At this opening the nascent TM segments are exposed to the lipid as well as the aqueous channel of the translocon, and they partition into the bilayer due to their hydrophobicity. At the end of a TM segment, the side of the translocon closes to allow transport of the hydrophilic portion of the nascent polypeptide into the cytosol or periplasm/lumen of the ER. Thus integration of polytopic membrane proteins involves a dynamic cycling between two conformations of the translocon, one that exports the polypeptide across the membrane and one that allows it to insert into the lipid bilayer. The regulation of this cycling depends on interplay between the translocon, the lipid, and the signals contained in the sequence of the protein substrate. These signals are segments of ~20 nonpolar residues called stop transfer sequences because they stop the export of the polypeptide chain and insert into the lipid bilayer to become TM segments. The nonpolar character of these segments allows their prediction

Biogenesis of Membrane Proteins

179

BOX 7.4. Cross-linking traces nascent peptides through the translocon Into the bilayer To investigate the path of nascent membrane proteins through the translocon, full-length or truncated proteins are engineered with sites for incorporation of chemical crosslinking agents. Translation intermediates of various lengths are generated using polymerase chain reaction (PCR) with different downstream primers to amplify portions of the gene before transcription to produce a set of truncated mRNAs. In vitro translation is carried out in the presence of 35S_ methionine and is stopped with cycloheximide, which inhibits the peptidyl transferase activity of the ribosome. Many different experiments have employed cross-linking to study the fate of translocated proteins. For the protocol described here, the truncated mRNAs are designed to allow the incorporation of a photoactivatable derivative at a unique position in the TM segment of the protein to be studied by incorporating a UAG stop codon at the position of interest. A suppressor tRNA carrying the anticodon CUA will bind to the UAG stop codon and incorporate its amino acid into the growing chain, instead of letting translation terminate. When the suppressor tRNA carries trifluoromethyl-diazinyl phenylalanine, it incorporates this UV-sensitive cross-linking agent in that position (Figure 704.1). Only when irradiated will it react, forming a covalent bond with other molecules in its immediate environment. In addition, chemical cross-linking was carried out with bis-maleimidohexane, a homobifunctiona I sulfhydryl-reactive agent (Figure 704.1). It can react with two cysteines, linking one in the protein substrate to another within ~15 A. In this case, the targeted cysteine is in the Sec61 13 subunit. The substrate protein for these experiments is a signalanchor protein engineered for glycosylation at the N terminal as well as for cross-linking as described (Figure 7.4.2A). The TM domain from residues 19 to 41 (shaded gray) contains a site for incorporation of the photoreactive phenylalanine derivative (*, residue 28 labeled "stop codon") as well as the cysteine residue (C) residue at position 35 for the chemical cross-linking. Below it the positions marked with (++) indicate where two arginine residues were inserted in experiments designed to test whether charges in the TM segment disrupted translocation and lipid partitioning (and they do). In vitro translation is carried out in the presence or absence of rough (i.e., not highly purified) microsomal membranes, and proteinase K is added to see at what point the translation intermediates are no longer protected from digestion across the membrane (see Box 7.3). The extent of glycosylation indicates whether the N terminus extends into the lumen of the microsomes. The results in Figure 7A.2B are autora-

NHt 0

I

~

N II N

7"1 ~

II

CH2-CH-C-tRNA

diographs from experiments in which radiolabeled chains are synthesized from different length messages in the presence or absence of rough microsomes and in the presence and absence of proteinase K, as indicated. Samples are analyzed by SDS-PAGE prior to autoradiography. The 35S-labeled products show that at least 61 amino acids (the 61mer) are needed to see some protection from proteolysis by the addition of microsomal membranes (bands marked with arrows), indicating the ribosome-nascent chain complex has reached the translocon at that length. By the length of 85 amino acids (85mer), glycosylation can take place, producing slowermigrating bands (marked with stars), which indicates that the chain is emerging on the other side of the membrane. (Open arrows and open stars mark nonproteolyzed nonglycosylated and glycosylated chains, respectively; closed arrows and closed stars mark membrane-protected nonglycosylated and glycosylated chains, respectively, in the 85mer to the 154mer. When samples are irradiated after translation and then sedimented at either neutral or alkaline pH, cross-links to the translocon (Sec61 a and Sec61(3) and to lipid are observed (Figure 704.3). Sedimentation at neutral pH that does not occur at alkaline pH indicates that a nascent peptide chain is associated at the membrane periphery and not yet integrated across it, as observed with the 61 mer and 68mer in Figure 7A.3A. These two nascent chains are also able to cross-link to the Sec61 a of the translocon (marked with closed diamonds, lanes 14 and 22; see also Figure 7A.3B). Chemical cross-linking to Sec61 13 appears with the 71 mer, the 78mer, and the 85mer (closed circles). Membrane integration begins with the 71 mer, with clear cross-linking to lipid (open diamonds; see also Figure 7A.3C), while the 71mer does not cross-link to Sec61 a. As the length increases, the lipid crosslinks remain, indicating the TM segment remains in the lipid bilayer. Figure 7A.3B shows the results when the samples are immunoprecipitated with antibodies to Sec61 a and verifies that the peptides of 61 and 68 residues are the only ones that remain in its close vicinity. Figure 7A.3C shows what happens when the cross-linked 71 mer is treated with phospholipase A and verifies that the faster-migrating band is the result of cross-linking to lipid. When all the data are plotted according to the length of the translation intermediate, they give a sequential picture of contacts, indicating that the nascent TM segment is first exposed to the cytosol, and then enters the translocon and inserts into the lipid bilayer; finally, the N terminus becomes glycosylated as the C terminus diffuses away from the translocon into the cytosol (see Figure 7.17).

y~w~ J-J o

o

CF 3

7.4.1. Chemical structures of the cross-linking agents used: trifluoromethyl-diazinyl phenylalanyl-tRNA (left), which is added during translation to incorporate a photoreactive derivative of Phe in the TM segment, and bis-maleimidohexane (right), a reagent for homobifunctional sulfhydryl-reactive cross-linking with a spacer arm of 13 A). (continued)

Protein Folding and Biogenesis

180 BOX 7.4 (continued)

glycosylation site

Y

A. N.(

3

stop codon

54-154mers

19

28

41

I

I

I

(

*II

'1

c _.)

I-C 297

/;

++

54 mer

61mer

lumen

- + +

RM 2

4

3

5

78mer

6

8

7

85mer Q-

RM

>-..,J

71mer

protK

protK

cytosol

N

SA I membrane protein B.

'I

9 10 11 12

95mer

..

.

~

....

-'I

- + - ++ - + - +

- + - + - I +

-

I

- I +

1314 15 16

I 134mer I

IIlmer

protK RM

--

---?II -. j

-

"'Q

154mer

-

t1.

"'-

'-

~

.-..

~-

*-

.

- + - + - + - +1 • + - + • I + - I + I I+

.

25 26 27 28 29 30 3\ 32 33 34 35 36 7.4.2. A. The signal-anchor membrane-protein used in photocross-linking experiments. B. Protection assay of radiolabeled chains of different lengths. Synthesis of the signal-anchor nascent chains of different lengths in the presence and absence of rough microsomes (RM) was followed with and without treatment with proteinase K (protK) by SDS-PAGE and autoradiography. From Heinrich, S., et aI., Cell. 2000, 102:233-244. ~) 2000 by Elsevier. Reprinted with permission from Elsevier.

Biogenesis of Membrane Proteins

181

BOX 7.4 (continued)

A.

neutral pH P

UV

P

S

S

P

P

S

+

- + -+

alkaline pH

neutral pH

alkaline pH

S

P

P S

+

P

S

S

P

S

+ - +

+

•

54mer

9 3

2

5

4

7

6

11 12

8

13 14 15 16

•

•

-

71mer

68mer

17 18 19 20 21 22

25 26 27 28

23 24

•

29 30 31

32

• 78mer

•

--<> ~

-

~

•

<>

+.

85mer

~

+•

~

41 42 43 44 45 46 47

33 34 35 36 37 38 39 40

Sec61 a-IPs

B.

c.

48

71mer

154mer chain length

49 50 51 52 53 54 55 56

uv 54 61 6ll 71 85 95 134

1234567

PLA2

+

+ + 2

7.4.3. Chain length requirement for membrane integration. A. Chains of different lengths were cross-linked by exposure to ultraviolet irradiation and the membranes were sedimented at alkaline or neutral pH. Pellets (P) and supernatant (5) were analyzed. B. Immunoprecipitation with antibodies to Sec61 c< showed which chains interacted with the translocon. C. Treatment with phospholipase A z showed which chains interacted with lipids. From Heinrich,S., et al., Cell. 2000,102:233-244. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Protein Folding and Biogenesis

182

40mer

~ IO~I <.i

1

targeting

75J

V'~r,.v _..... _

_'1- - - - - - -

sedimentation (neutral pH)

'7-<1

~

S

'0

25

" •

0.0

$3

=

~

~

25


....<.i

:

01 30

-

-

q

r

j

40

50

.-.

60

~

••

/ '

S"61~-,ro,,lin.,

~;~.-

70

I

I

80

90

i

-;

I

I

I

0.0 ~

-=

•

Q.,

o

•

30

III

=

100

•

I i-----'-

I

I

40

50

60

diffusion &

70

l

-=

I

90

.

I

I

<.i

~ -8

i

I

•

..

!

;ntegn"on

71mer

---I

I i i

.. ..

=~ N

ITI

diffusion & glycosylation

1

l34mer


0.0

.

$3

=
Q.,

targeting

61mer

n

'0 1::
~

~

•

100 110 120 130 140 150 160

exposure or cytosolic domain

9IYCOSYlati(~

';

I

80

I

• lipid crosslinks

~..

..-----..----'


I

sedimentation (alkaline pH)

~


~o

..... I

£

'0

I

100 110 120 130 140 150 160

------.~

<.i

=D 0==

Sec61a-crosslinks

.S IOOll integration I

1::
N

50


~

50

-v - - - - - - ~

0-1

•

•

,~

i i i

30

40

50

60

I

70

80

90 100 110 120 130 140 150 160

Chain length

~ N

7.17. Cross-linking results indicate three stages in the biosynthesis of the SAl protein. a signal-anchor protein Data were obtained as described in Box 7.4 and were quantitated to show the percentage of total chains that had each characteristic plotted (y-axis) as a function of the length of the truncated chain (x-axis). (Note: the top plot also gives a scale for the percentage sedimented at neutral pH. right axis.) The top graph shows cross-linking to Sec61, which requires a minimum of about 60 amino acids. The middle graph shows that over 70 residues are required before the nascent chain sediments at alkaline pH. at which length it is integrated into the membrane. At the same length, it can be cross-linked to lipid. The bottom graph shows that glycosylation occurs on chains longer than 85 residues, indicating they have reached the outside, while the proportion that is exposed to protease after translocation continues to grow. These stages are diagrammed along the side of the graphs. From Heinrich,S .. et al., Cel/. 2000, 102:233-244. © 2000 by Elsevier. Reprinted with permission from Elsevier.

h-om the primary sequence based on hydrophobicity scales. As shown in Table 6.3, the biophysical scales for hydrophobicity give values of the free energy for partitioning of each amino acid into the lipid phase. Based on knowledge of the role of the transJocon, an ingenious set of experiments established a biological scale For the apparent l]-ee energy (6G app ) of inserting each amino acid, as a part of a TM helix, into the bilayer.

The amino acid is placed in a test segment engineered to follow the t\",O TM segments of leader peptidase (see Figure 6.15) so that when synthesized in the cell, it is either inserted into the membrane or exported past it; in the latter case, it is glycosylated, which allows quantification (Figure 7.18A). The test segment consists of leucine and alanine residues nanked by GGPG linkers, with the amino acid of interest in the center position.

Biogenesis of Membrane Proteins

183

PI

4.0,----------------------------,

3.5 3.0 2.5

I

(3

E

2.0

(ii u

1.5

6

1.0

0.5

....__._.___,---r_,_e,___..,....,

O. 0 r--,..rr-.......--n.--r_____,---r_,_e,___~ -0.5 -1.0 '-----

---.J

I L F V C MAW T Y G S N H P 0 R E K D Amino acid 7.18. Biological hydrophobicity scale for insertion of TM segments. A. A test segment is engineered in leader peptidase (Lep) constructs that will be either inserted into the membrane or translocated across it. The test segment, H, consists of a flanking sequence GGPG, followed by 19 leucine residues, then the flanking sequence GPGG, with each amino acid to be tested inserted in its center. Two sites for N-glycosylation are also inserted, as indicated by G1 and G2. Each construct was expressed in BHK cells that were immunoprecipitated with Lep antiserum after labeling with 35S-methionine. The bands corresponding to singly and doubly glycosylated proteins were quantified on a phosphorimager after separation by SDS-PAGE. B. The 6G app for membrane insertion of each amino acid placed in the center of a TM segment is calculated from the apparent equilibrium constant of the singly glycosylated and doubly glycosyated Lep molecules obtained with the contructs described in A. The bar indicates the standard deviation in the 6G app for lie; the standard deviation for each of the others is similar. Redrawn from Hessa, T., et al., Nature. 200S, 433:377-381. © 200S. Reprinted by permission of Macmillan Publishers Ltd.

As expected, the nonpolar amino acids have ~Gapp < 0, promoting membrane insertion, while the polar and charged residues have ~Gapp > 0 (Figure 7.I8B). Thus the biological scale for membrane insertion correlates very well with the biophysical scales derived earlier, supporting the postulate that during translocation the

segment is directly exposed to the bilayer. Interestingly, some amino acids give different values when they are positioned away from the center of the segment: notably, positions closer to the ends of the segment are more favorable than the center for Trp and Tyr, as well as Lys and Asn. Structural data show a prevalence

Protein Folding and Biogenesis

184

A.

(C N

B.

N~VVV~V\.C aa ~.!W -=-~lVVV-JtV\.C .~tltl ~.~tl _

N

C

C 7.19. Functional topogenic determinants throughout the polypeptide. Downstream portions of polytopic membrane proteins may influence topology, depending on loop size. A. Insertion of the bitopic ASGP receptor follows the positive-inside rule. However, in an engineered protein consisting of the four copies of its signal-anchor sequence with flanking charges, separated from each other by ~100 residues, the TM segments insert sequentially in spite of their flanking charges. It appears that the long loops between TM segments allow insertion to override the positive-inside rule. B. A natural protein with 12 TM segments with short loops between them, Glut1, inserts according to the positive-inside rule (wild type, in center). Perturbing its topogenic determinants by introduction of different charges disturbs the local topology but not the other TM segments (right and left). From Goder, V, and W. Spiess, FEBS Lett. 2001, 504:87-93. © 2001 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

ofTrp and Tyr in aromatic girdles of integral membrane proteins at the interfacial regions of the bilayer, while "snorkeling" of Lys and Arg residues enables their basic groups to move to the more polar interface (see "Integral Membrane Proteins" in Chapter 4). The presence of more than one stop transfer sequence in polytopic proteins allows each hydrophobic TM segment to be inserted sequentially as each one reaches the translocon and acts as a stop transfer sequence. An engineered protein that contains four copies of a stop transfer sequence inserts each as a TM segment, provided the loops between them are of sufficient length (Figure 7.19). However, some polytopic proteins appear to exhibit cooperation between adjacent TM segments, which implies more than one can be in the translocon simultaneously. Although the channel observed in the high-resolution structure of the translocon from 111.. jannaschii is only wide enough for the passage of an unfolded polypeptide or a single a-helix, it is possible that alternate conformations of the translocon involving dimers or higher oligomers allow more than one helical peptide segment to fit in the channel. This would explain the evidence that some TM segments can influence the insertion of neighboring TM domains in some poly topic proteins (see below). Topogenesis in Membrane Proteins

In discussing the signals that determine the topology of integral membrane proteins, '.vhich have been studied extensively in the mammalian ER and also in E. coli, it is useful to refer to the outside compartment (lumen or peri plasm) as the exocytoplasmic side of the membrane. Thus for bitopic membrane proteins, type 1 are oriented Nexa/C m and type II are Nevl/Cexo. Poly topic proteins, called type III membrane proteins, may have an even or an odd number of TM segments and thus have their N and C termini either on the same side of the membrane or on opposite sides with either orientation. The first

signal occurring in the sequence of a poJytopic protein is treated much as the signal of a bitopic protein, and insertion of the remaining TM segments can be assumed to follow in sequence, even though there are other factors influencing insertion, as wiJJ be discussed. Signals that govern topogenesis are classified as (l) signal sequences, (2) signal-anchor sequences, and (3) reverse signal-anchor sequences. The presence of one of these sequences determines the orientation of the first TM segment of the protein (Figure 7.20). Insertion of the later TM segments of polytopic membrane proteins is initiated by stop transfer sequences, which have sufficient length to span the membrane and can insert stably into the bilayer. Signal sequences have already been discussed ror their role in targeting the nascent chain to the memo brane. Not all signal sequences are cleaved by signal peptidase. Those that are cleaved are released and degl-aded by signal peptide peptidase. After cleavage, \vhich makes a new N terminus outside the membrane, the rest of the peptide is fully exported as a secreted protein unless it is followed by a stop transfer sequence to create a TM segment (see a and b of Figure 7.20). A stop transfer sequence that can insert stably in the membrane is called a signal-anchor sequence. Signalanchor sequences remain un cleaved and have enough nonpolar residues to span the hydrophobic domain of the bilayer. When it is the only TM segment and is not preceded by a cleavable signal sequence, a signalanchor inserts to create a type 11 membrane protein (c in Figure 7.20). Reverse signal-anchors are like signal-anchors except that they create type I membrane proteins (d in Figure 7.20). They are differentiated from signalanchors not by obvious physical characteristics but by more subtle factors that affect their function, allowing them to insert with an Nc\o/Cc.\·1 orientation. The factors that influence the orientation of the signal-anchor

Biogenesis of Membrane Proteins

185

-----------{c::::J}-----

1-------c:r-----c=J---------100 aa

.~ N

c

c .~

~

N

exo N'

cyt

a

b

--:

c

cleavable signal

d

f

e

---

signal-anchor

g ~

reverse signal-anchor

7.20. Three types of signals initiate topogenesis. Cleavable signal sequences (green, with arrowheads for cheavage sites) are translocated, leading the N-terminus to the outside before they are cleaved. After cleavage by signal peptidase, they are digested by signal peptide peptidase outside the membrane (not shown). The protein is released (a) unless the cleaved signal is followed by another TM segment (b). Uncleaved signal-anchor sequences (red) induce translocation of the rest of the peptide while they remain TM (c). Reverse signal-anchors (blue) insert to become TM as the N terminus is translocated (d). Thus for bitopic proteins, both a cleaved signal and a reverse signal-anchor produce the NexolCcyt orientation (b and d), while a signal-anchor produces the NcytlCexo orientation (c). For polytopic proteins (e, f and g), additional TM segments insert in alternating orientations (light red for NcytlCexo and light blue for NexoICcyt). The sequences shown represent a secreted protein (a, prolactin), two type I membrane proteins (b, asialoglycoprotein receptor and d, cation-dependent mannose-6phosphate receptor). a type II membrane protein (c, synaptotagmin I) and three type III proteins (e, gap junction protein; f, vasopressin receptor V2; g, glucagons receptor). From Higy, M., et aI., Biochemistry 2004, 43:12716-12722. © 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

sequence as it emerges from th.e translocon are just being elucidated. The topologies that result from the different signals indicate that a TM segment can insert into the bilayer in either orientation and can thus translocate its C terminus or its N terminus (Figure 7.21). Current evidence points to five factors that influence the orientation of signals within the translocon. 1. The charges flanking the hydrophobic sections of signals greatly influence the resulting topology. As described in the last chapter, the positive-inside rule, discovered in bacteria where positive residues are more abundant in cytoplasmic loops than in peri plasmic loops, holds for all organisms. In eukaryotes the charge difference between the flanking segments of a signal's hydrophobic core. described as L'.(N-C), results in the more positive flan king sequence preferri ng the cytoplas-

mic side. Therefore if the signal can change directions while in the translocon, it will orient to aJlow the more positive flanking residues to stay on the more negative side of the membrane and perhaps to interact with residues of the translocon on the cytoplasmic side. 2. Because a large globular folded domain will not get through the translocon, an N-terminal domain that folds into a large globular domain will not be translocated, even if the flanking charges suggest it should. Evidence for this comes h'om many studies with fusions and truncations that attach or eliminate rapidly folding globular domains. However, natural proteins that have extensive globular domains are probably protected from folding in the cytoplasm by chaperones. 3. Another factor that can override the positiveinside rule is hydrophobicity, specifically the length and composition of the nonpolar sequence of the signal. When artificial signals are made up of leucine residues,

Protein Folding and Biogenesis

186 ~

Positive charge difference

~

~(N-C)

-

Folding of N-terminal domain - -

.

Hydrophobicity

N

C

•

r.

?

•

..

f'\

.

\~(U~

N

N~)

"y~~ ')

\

~. N

(./~ )

7.21. Several factors affect the orientation of signals in the translocon. Signals translocate either their C or their N terminus once they enter the translocon, depending on factors such as the difference in positive charges, the hydrophobicity, and the folding of the N-terminal domain. These factors are thought to affect the orientation of signal-anchor and reverse signal-anchor sequences as they engage with the translocon. The resulting orientation determines whether the C-terminus or the N terminus emerges. From Goder, V, and W Spiess, FEBS Lett 2001, S04:87-93. © 2001 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

orientation is a function of length: Leu7 drives mostly Cterminal u-anslocation, resulting in Nin/Coul' while Leu22 and Leu25 produce N-terminal translocation, resulting in Ncxo/Ccyt. When amino acids are ranked for their ability to promote N-terminal translocation, the best are the most hydrophobic, which also have the highest propensity to form ex-helices in nonpolar environments. 4. An additional factor for multispanning membrane proteins derives from cooperation with the rest of the molecule. Charge mutations designed to invert the topology of polytopic proteins can instead affect a region of the protein without inverting it, as obse[\led with the human glucose transporter, Glutl, a 12-TM helix bundle (Figure 7.18). One or two hydrophobic segments can be "frustrated" enough by the mutation to remain outside the membrane, rather than perturbing the rest of the topology. The short loops between many TM segments suggest they insert as helical hairpins, preserving the expected topology, if the translocon accommodates two helices. 5. Finally, any glycosylated proteins have sites for glycosylation on the outside. In the ER, an oligosaccharyl transferase associates with the translocon and glycosylates nascent chains as they emerge. An engineered si te that becomes glycosylated will fmce that portion of the polypeptide to end up on the outside and prevent reorientation of that segment. A model to explain these effects on topology postulates that all signals enter the translocon with their N termini first, as they come off the ribosome, and therefore the signals that \-esult in a type II orientation must

somehow turn around in the translocon (Figure 7.22). There does seem to be a time limit fOl- the reorientation, based on results obtained with a collection of chimeric model proteins ranging in length from lOO to 580 residues. The proportion of proteins with translocated C termini increased linearly from about 20% for a length of 100 residues to ~55% for lengths over 300 residues, corresponding to a period of 40 to 50 seconds, after which the topology is fixed. This discussion of topogenesis has assumed that once the first TM segment of a poly topic protein has integrated into the membrane, insertion of the remaining TM segments follows in a linear manner. However, the topology of some polytopic proteins is dependent on the order of TM segments, suggesting that they contain "weak" TM domains that are not sufficiently hydrophobic to insert on their own. That strong TM domains can help integrate adjacent "weak" TM domains is observed in MDR1 and the erythrocyte Band 3. Presumably these interactions take place in the transJocon, when the two helices are close enough in the sequence to enter it together; in experiments that prevent the interaction by engineering a longer distance between them, the "weak" helix no longer inserts into the bilayer. A number of polytopic proteins can insert into the membrane with alternative topologies, including the prion protein, PrP, and the ion channel involved in cystic fibrosis, CFTR (see below). Regulation of insertion is hypothesized to involve the recognition of sequences called stop-transfer effector (STE) sequences, domains flanking the hydrophobic membrane-spanning domain

Misfolding Diseases

187

N

SRP (

\---

\

)

7.22. A model for the topogenic effect of the signal as it emerges from the translocon. Since translocation is initiated at the N terminus of the signal as it comes off the ribosome, that end of the signal must enter the translocon first. The only way both possible orientations can result is if the signal is able to reorient while in the translocon in response to various factors (see previous figure). From Goder, and W. Spiess, FEBS Lett. 2001, 504:87-93. © 2001 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

v.,

that appear Lo affecL whether it becomes a TM segment. Receptors for STE sequences have been identified by cross-linking and are proteins associated with the translocon. So few of these have been identified that their characteristics are not well defined. The STE sequences in IgM have negatively charged residues that are importanL in their function, while the STE sequences in PrP have positively charged residues, suggesting Lhey bind different STE receptors. These mechanisms are importanL subjects for research for proLeins whose altered topology results in disease.

of the disorders caused by missense mutations. Over a hundred different rhodopsin mutanLS produce retinitis pigmentosa. CysLic fibrosis is a relatively common hereditary disease that results in severe obstruction of the respiratory and gastrointestinal tracts of people with two defective copies of the gene for CFTR. The CFTR protein (Figure 7.24A) functions as a CI- ion channel thaL is

MISFOLDING DISEASES

Since correct insertion of membrane proteins determines their topology (and therefore their fold) and mistakes cannot be remedied simply by refolding due Lo the barrier to reorienting TM segments across the bilayer, errors in this process can be costly. In an intriguing case, when the prion protein, PrP, is inserted with its N terminus in the ER lumen, it does not result in disease; however, when iL is inserted with its N Lerminus in the cytoplasm, iL can lead to neurodegeneration, apparently by a mechanism involving aggregation (Figure 7.23). In general, damage can resulL from eiLher accumulation of misfolded proteins or from loss of needed hlllctions of the affected proteins. In humans the misfolding of membrane proteins is known to cause several heritable diseases, such as cystic fibrosis and retinitis pigmentosa. Many of the thousand point mutations causing cystic fibrosis resu IL in the misfolding of the cystic fibrosis TM conductance regulator (CFTR), and even the wild-type CFTR assembles with a low efficiency under some conditions. Almost a third of the 55 genes linked to disorders of the human retina encode integral membrane proteins, with many

Localization of the N terminus b Integration of / TM domain

unLrans]ocaLed PrP ER lumen N

NtmprP

elmprP SecPrP

7.23. The three topologies of the prion protein, PrP. The TM segment of PrP can insert in the bilayer in both orientations, Next/Ccy, (NtmprP) and NcyrlCext emprP). In addition, it can be secreted into the lumen in a soluble form (seCprP), which can then become attached to a GPI-anchor. There is evidence for a role of CtmprP in the neurodegenerative scrapie diseases. Redrawn from Ott, C. M., and R. Lingappa, Biochemistry. 2004,43: 11973-11982. c 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

v.

Protein Folding and Biogenesis

188

Oilgosaccharide chains of CFTR A.

R domain B.

CFTR (cystic fibrosis)

.....

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-.

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.... ~

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it .. NBD2

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*-*

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rZZ7ZZZLi

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*-

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tZZZZZZZ)

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7.24. The CFTR protein and cystic fibrosis disease. A. The topology of the cystic fibrosis TM conductance regulator (CFTR) is shown with 12 TM helices and three cytoplasmic domains. Two of the cytoplasmic domains are nucleotide-binding domains, NBD1 and NBD2, and the third is a regulatory domain (R domain) in which the protein is phosphorylated by cAMP-dependent protein kinase. The position of the most common mutation in cystic fibrosis patients, Phe508, is indicated. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 403. © 2005 by W. H. Freeman and Company. Used with permission. B. The locations (starred sites) of other mutations in the CFTR sequence that result in cystic fibrosis disease. The hatched bars below the sequence correspond to TM segments of the protein. From Sanders, C. R., and J. K. Myers, Annu Rev Biophys Biomol Struct. 2004, 33:25-51. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

*-

*-

1400

Misfolding Diseases

189

A.

p B.

PMP 22 (CharcotMarie-Tooth) Aquaporin (diabetes insipid is)

Vasopressin receptor (diabetes insipid is)

*

****,

* *

*

,,77777711

I ti

it

.,...* **

*i*

LZ7LlllIlA

=

20\

MIt

t:Wr***

*oM:

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*** .. t*

\ * .............. 'fMH***

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***t

*

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**

it ........

*

*

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,,77777J

t

* .... IZZZJ

*

200

.* *

I

**

*

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I

t ,,"

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• .....

I

200

378

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1....

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\llili/ZIA

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,,777777777l

IlZI

t

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'l'IIZII'A

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Rhodopsin (reti n i tis pigmentosa)

Connexin 32 (CharcotMarie-Tooth)

*.,

1l17777777J

200

349

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**=1**

*

11"1" 1,,1'11 .... 200

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284

7.25. Sites in membrane proteins of mutations linked to diseases. The sites for which mutations have been linked to disease are well distributed throughout the sequences of numerous membrane proteins. A. Ribbon diagram of the structure of rhodopsin showing the locations of missense mutations implicated in retinitis pigmentosa. The affected side chains (shown in black) are distributed all over the structure. B. The sequences of rhodopsin and other membrane proteins linked to diseases, with the positions of mutations (asterisks) and of TM segments (hatched bars) shown. From Sanders, C. R., and J. K. Myers, Annu Rev Biophys Biomol Struct. 2004, 33:25-51. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

=

Protein Folding and Biogenesis

190

activated by cAMP-protein kinase. Loss of the channel activity in epithelial cells alJo'ws the lungs to be clogged with debris and become subject to frequent bacterial infections that damage the lungs, reduce respiratory efnciency, and shorten the afflicted person's lifetime. In 70% of cystic fibrosis cases, a deletion of PheS08 from CFTR results in misfolding of the protein, which results in its degradation. In addition, many other diseaserelated mu tations are distribu ted throughout the protein sequence (Figure 7.24B). Like CFTR, the mutations in rhodopsin that lead to retinal disease are located in many different parts of the protein structure (Figure 7.2SA). From the variety of domains affected by mutations, it is evident that the mutations do not target an active site, or even a couple of localized regions. In fact, this pattern of widely distributed disease-causing mutations is true in five other membrane proteins associated with diseases (Figure 7.2SB). It strongly suggests that these are misfolding mutations, because a Jat-ge variety of single changes in virtually any part of a protein are not expected to critically impair function. They could, instead be crucial to folding by tipping the delicate energetic balance between correct folding and misassembly. The high frequency of misfolding of these membrane proteins provides new targets for drug development, as ligands that stabilize the native state could act as chemical chaperones that rescue the misfolded membrane proteins.

FOR FURTHER READING

Seminal Papers Engelman, D. M., and T A. Steitz. The spontaneous insertion of proteins into and across membranes: the helical hairpin hypothesis. Cell. 198 I, 23:411-422. Gruner, S. M., Instrinsic curvature hypothesis for biomembl-ane lipid composition. Proc Nat! Acad Sci USA. 1985, 82:3665-3669. Randall, L. L., Translocation of domains of nascent peripIasmic proteins across the cytoplasmic membrane is independent of elongation. Cell. 33:231-240. Hartl, F. D., S. Lecker, E. Schiebel. J. P. Hendrick, and W. Wickner, The binding cascade of SecB to SecA to SecY/E mediates preprotein targeting to the E. coli plasma membrane. Cell. 1990, 63:269-279. Simon, S. M., and G. Blobel, A protein-conducting channel in the endoplasmic reticulum. Cell. 1991. 65:371380.

Heinrich, S. U., W. Mothes, J. Brunner, and T A. Rapoport, The Sec61p complex mediates the integration of a membrane protein by allowing lipid partitioning of the transmembrane domain. Cell. 2000. 102:233-244. Selected Reviews Folding Booth, P. J., and A. R. CU1Tan, Memb,-ane protein folding. Curl' Opin Sirucl Bioi. 1999, 9: J 15- 121. Popot, J.-L., and D. M. Engelman, Helical membrane protein folding, stability and evolution. Annu Rev Biochem. 2000, 69:881-922. White, S. H., and W. C. Wimley, Membrane protein folding and stability: physical principles. Armu Rev Biophys Bion-wl Siruct. 1999, 28:319-365. Bowie, J.. Solving the membrane protein folding problem. Nall.lre. 2005, 438:581-589. Sanders, C. R., and J. K. Myers, Disease-related misassembly of membrane proteins. AI1I1U Rev Biaphys Biomal SLrUCI. 2004, 33:25-51. Biogenesis Veenendaal, A. K. J., C. van del' Does, and A. J. M. Driessen, The protei n-cond ucti ng cha nnel SecYEG. Biochim Biophys Acta. 2004, 1694:81-95. Luirink, J., G. von Heijne, E. Houben, and 1.-W. deGier, Biogenesis of inner membrane proteins in Escherichia coli. AI1J7U Rev Microbial. 2005. 59:329-355. Luirink, J., and 1. Sinning, SRP-mediated protein targeting: structure and function revealed. Biochim Biophys Acta. 2004,1694:17-35. Dalbey, R. E., and M. Chen, Sec-lJ-anslocase mediated membrane protein biogenesis. Biochim Biophys Acta. 2004, 1694:37-53. White, S. H., and G. von Heijne, The machinery of membrane protein assembly. Curr Opil1 StruCl Bioi. 2004, 14:397-404. \"'hite, S. H., and G. von Heijne, Transmembrane helices before, during and after insertion. CUlT Opin Struct Bioi. 2005,15:378-386. Ott, C. M., and V. R. Lingappa, Integral membrane protein biosynthesis. J Cell Science. 2002, 115:2003-2009. Higy, M., T Junne, and M. Spiess, Topogenesis of membrane proteins at the endoplasmic reticulum. Biochemistr\'. 2004, 43: 12716-12722. Mitochondrial Biogenesis Pfanner, N., and N. Wiedemann, Mitochondrial protein import: two membranes. three translocatases. Cllrr Opil1 Cell Bioi. 2002, 14:400-411. Endo. T, H. Yamamoto, and M. Esaki, Functional cooperation and separation of translocators in protein impol"t into mitochondria, the double-membrane bounded organelles. J Cell Science. 2003, 116:3259-3267.

8

Diffraction and Simulation

Experimental sample

Diffra Ion studies oeus a be~1Tl of x-rays or neutrons 0 mullililmellar s.acks containIng thousanos ot lipid bdayers and hus differ greatly in seal from s'rnulations of a bilayer contallling only a ew hundred I p'd molecu es. From Benz, R. W, e aI., Blophys j. 2005, 88:805-8 7. 2005 by tl e Blop 'yslcal Society Reprinted With permissl n from he Biophysical SOCie y.

Multilamellar stack of 1000's of bilayers

Important tools for structural determination of membrane components include x-ray and neutron diffTaction techniques. While the most familiar use of x-ray diffraction is the solution of crystal structures providing high-resolution structures of proteins and lipids in crystalline arrays, other diffraction techniques can provide structural information on membranes or reconstituted systems with lipids in the fluid phase. Such membrane diffraction studies give information that is onedimensional, normal to the bilayer plane, because of the liquid nature of the acyl chains. The constant motions of lipids in the L~ phase (see Chapter 2) introduce several types of disorders (Figure 8.1) that prevent precise delineation of their structure at atomic resolution and invite description of their dynamic properties by sophisticated computer modeling. Today the interplay between diffraction techniques and simulation methods contributes even more to understanding the structure of the fluid membrane. This chapter describes diffraction and simulation methods as tbey apply to the lipid bilayer and then looks at the lipids that are resolved in crystal structures of membrane proteins. It will close with a few comments on the art of crystallog-

raphy of membrane proteins, which allows solution of their high-resolution structures. The following chapters illustrate how x-ray crystallography of membrane proteins is providing insights toward detailed understanding of their structure and functions.

BACK TO THE BILAYER

A starting point to depict the lipids in a bilayer is a view of the static structures obtained from x-ray crystallography of several pure lipids in crystalline phase (Figure 8.2). Analysis of such single-crystal structures describes the conformations of lipids in Lc phase, giving their dimensions, torsion angles, and tilt angles relative to the bilayer normal, along with the positions of their headgroups. In addition, several high-resolution structures of membrane proteins have a few well-delineated lipids (see below), although these must be viewed as boundary lipids with constraints on their mobility from the close interactions with the proteins. Because the biologically relevant fluid lipid bilayer is a two-dimensional liquid crystal with lateral motions

191

Diffraction and Simulation

192

Rotational diffusion wobble 10- 8 sec

O~~1I' Protrusion , ~ 10- 9 sec

Gauche-trans

~ Flip-flop

I

(J 10-

~

3

-

4

10 sec

~

Lateral diffusion 10- 7 sec

~ Undulations 10- 6 sec-I sec 8.1. Disorder in lipid bilayer is due to several kinds of lipid motions, shown here with their approximate correlation times. Redrawn from Gawrisch, K., in P. Yeagle (ed." The Structure of Biological Membranes, 2nd ed .. CRC Press, 2005, p. 149. © 2005. Reproduced by permission of Taylor & Francis. a division of Informa pic.

~~t ~~

- ~~

:~; .<~

')-<

3~

,),

DMPC'j<

~~ 'i;

DLPEM 2

L~;"""h. ):.

,...{

'J..

'}~=

,1

)~) /\~ ,

:;~~~~~~ "~: ';

t.

AD = 2(VL

+ nwVw)

since the thickness is for two lipid layers. For fully hydrated DPPC at SO?C, these methods give o = 67.2,.\. VL = 1230,.\3. and A = 71.2,.\2.

LIQUID CRYSTALLOGRAPHY

'S: DLPA>

of its molecular components and constant fluctuations of its acyl chains. its structure is much less amenable to analysis than that of less dynamic lipid phases. A simple parameter can be difficult to pin do\.vn: for example, experimental measurements of the interfacial area of DPPC in La phase from diffraction and NMR studies give results varying from 56 ,.\2 to 72 ,.\2 per monomer. Recent simulations employing different values suggest that the appropriate value for the area is 69 ,.\2, which exemplifies how contributions [Tom both experimental and computational approaches can refine knowledge of fundamental membrane characteristics such as bilayer thickness and distributions of components. A number of methods have been used to determine such general structural characteristics for a wellhydrated lipid bilayer. X-ray diffraction studies give a measure of D. the thickness of the bilayer, as described below. Flotation experiments can determine VL • the volume of a lipid, by finding what mixture of H 2 0 and D2 0 matches its density. Using data that correlate relative humidity with the activity of water determined either gravimetrically or isotopically, the number of water molecules (n,,) is calculated from the % humidity, and the volume of a water molecule (Vw ) is known. Thus the area can be calculated from the volume and the thickness:

DMPG, 4

jr U-c. f

8.2. Single-crystal structures of four phospholipids. The structures of DMPC, DLPA. DMPG, and DLPEM2 (dilauroyl-glycero1-phosphate-N,N-dimethylethanolamine. or DLPC lacking one methyl group) are shown as labeled. The crystal structures were all determined with DL lipids. The four differ in both chain stacking and the orientation of the glycerol group. Redrawn from Hauser, H. et al., Biochemistry. 1988.27:1966.

The techniques called liquid crystallography use data derived by diffraction h'om oriented muJtilamellar arrays of lipids (see Frontispiece and Box 8.l). Since the disorder inherent in the fluid bilayer defies structural delineation at atomic resolution. the positions of atoms in bilayer lipids are properly described by broad statistical distribution functions. Typically presented as one-dimensional projections along the axis normal to the bilayer surface. the structure derived from diffraction data gives the time-averaged probability distributions of the groups that make up the lipid (Figure 8.3). Examples of profiles of DOPC biJayer-s obtai ned with xray electron density and neutron scattering show how simple the data appear (Figure 8.4); however. they contain the information needed for liquid crystallography to describe a fully resolved bilayer structure profile. Two distinct approaches are taken for liquid crystallography: direct analysis with liquid crystal theory and the application of the refinement methods of crystallography to these lamellar diffraction patterns.

Liquid Crystallography

193

BOX 8.1. X-ray and neutron scattering which has the reversible property

Incident rays

J ox;

f(xl =

F(Sle -i21fsX dx

-00

•

•

•

•

•

•

•

•

•

•

•

•

8.1.1. Reflection of x-rays from different layers of atoms, with d the distance between layers and El the tilt angle of reflection. From Chang, R., Physical Chemistry for the Chemical and Biological Sciences, University Science Books, 2000, p. 835. In diffraction experiments beams of x-rays or neutrons (or electrons or light) are directed at ordered samples and reflect off components in the sample. X-rays are scattered by electrons, while neutrons are scattered by nuclei. The diffracted waves interfere with each other and many cancel each other out, while some are added together to produce a diffraction pattern that can be analyzed mathematically. The radiation from an x-ray beam penetrates enough to act with atoms in many layers of the sample. The distance between layers is represented by the variable d, and 8 is the tilt angle of reflection, as shown in Figure 8.1.1. Thus d sin 8 is the difference in path length between two waves. The mathematical relationship between the diffraction pattern and the object that produced the scattering is called a Fourier transform. It is a computation that converts an intensity signal (from the electron density) into a series of numbers that characterize the relative amplitude and phase components of the signal as a function of frequency (the diffraction pattern): The Fourier transform of a function f(x)' is defined as

J 00

F(S) =

f(xle -ibm dx

(see Figure 8.1.2). Many results of Fourier transforms can be seen at http://www.ysbl.york.ac.uk/~cowtan/fourier/fourier. html. These principles apply to the more familiar technique, x-ray crystallography, in which the object is a crystal with the particles in a very ordered array, as well as to diffraction studies of liquid-crystalline lipid bilayers. Since the observed diffraction is the result of time-averaged positions of the diffracting particles, it is affected by thermal motion, which is much larger in the bilayers than in crystalline materials. The diffraction pattern may be analyzed by Bragg's law, which is based on the fact that constructive interference occurs only when the difference in distance of the x-rays reflected from adjacent planes is equal to the wavelength of the x-ray beam. Therefore 2d sin 8 = A, where 8 is the reflection angle, d is the distance between the layers, and A is the wavelength. A stack of fluid lipid bilayers can produce five to 10 sharp Bragg reflections, analyzed as structure functions whose Fourier transform yields the one-dimensional structure profile across the bilayer. When x-rays are scattered by electrons, the profile is an electron density distribution; when neutrons are scattered by nuclei, the profile is called a scattering length distribution. Both can be generalized as probability distributions. The energy of a neutron beam is very weak, compared with x-rays, but neutron diffraction has been utilized very productively with selectively deuterated molecules incorporated into the bilayer. Since protons and deuterons scatter neutrons very differently, it is easy to pinpoint the locations of the deuterated groups. In addition, nondeuterated lipid can be suspended in D20 to locate the water molecules in the bilayer with neutron diffraction. There are excellent books on diffraction, as well as on-line courses such as http://www.uni-wuerzburg.de/mineralogie/ crystal/teaching/teaching.html and http://www-structmed. cimr.cam.ac.uk/Course/Overview/Overview.html.

-00

o I

I I I

FT

<~

~>

x __ [bl

I I I I I _____ L

o

__

--s

8.1.2. Fourier transform (FT) pair that relates a step function of width xp to a sin y/y function when xp s/2 = n1r, as an example of the Fourier transform, F(Sl. of a function f(x). Redrawn from Campbell, I. D., and R. A. Dwek, Biological Spectroscopy, Benjamin Cummings, 1984, pp. 359-360.

Liquid Crystal Theory

L"

smectic liquid crystals (Figure 8.5). The constituents of liquid crystals are ordered with respect to their long-

phase of the fluid bilayer denoles a liquid crys-

range positions (orientation) bUl still undergo rapid

talline material that is intermediate bel\-veen the liq-

rota tiona I and translationa I di ffusion. Thei r orien ta tion

uid and solid states of matter and can be treated like

makes them anisotropic, meaning their properties are

The

Diffraction and Simulation Probability

+z

Bilayer normal (z) 8.3. Illustration of the correspondence between a peak in the time-averaged probability distribution in the z-axis and the group that it represents. The ball-and-stick model shows the structure of DMPC with shading to illustrate its space-filling dimensions lacking the hydrogen atoms. The peak represents the phosphate group. From Wiener, M. c., and S. H. White, Biophys J. 1991, 59:162-173 and 174-185. © 1991 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

not distributed equally in x, y, and z directions, and establishes directionality, described with x and y in the plane of the layer and z normal to it. Their tilt angle off the z-axis, denoted e, can vary significantly. Their degree of orientational order is described by the function (3cos 2 e-1 )/2, which varies from 0 (totally random, i.e., isotropic) to t (totally aligned, so e = 0° and cos e = 1). The order parameter, S, of a liquid crystal is the average of this function: S = ((3cos 2 e-t)/2). X-ray diffraction of a smectic liquid crystal often produces strong first- and fourth-order scattering peaks, but the peaks are very diffuse in x-ray scattering data from fluid lipid bilayers (Figure 8.6). The peaks decrease in intensity as the order number increases until diffuse scattering overcomes the ability to detect the peak. Liquid cl)'stal theory provides a mathematical treatment of the scattering that can give electron density profiles for samples with at least four orders of diffraction.

ture, both sets of data are combined, after they are scaled mathematicaJly to establish agreement on the positions of the groups. With this joint refinement of xray and neutron diffraction data, the positions of eight groups have been defined for a phospholipid molecule plus associated waters (Figure 8.7). The result is the structure of a DOpe bilayer in La phase (Figure 8.8). Each peak represents the time-ave.-aged distribution of a principal structural group of the lipid projected onto the z-axis normal to the bilayer plane. Each peak is a Gaussian distribution whose area represents the number of structural groups the peak represents. The apex of each peak gives the position of the group in the bilayer normal, and the half-width of the peak gives the thermal motion. A numberoffeatures ofthe bilayer are evident in the structure profile obtained for DOPe. For example, there A.

.S' "D

..c:

~ 0.5

~ OIl ~

'C

~ 0.0 C1l U


C 0

b

::l

Z -0.5 -30

Since the number of observable diffraction orders is limited when x-ray or neutron diffraction data are collected on lipid bilayers, a powerful advance in determining bilayer structure combines the information from both. The joint refinement of x-ray and neutron diffraction data is effective because the two kinds of data ,-eport on different regions of the bilayer. X-rays interact with electrons, so they scatter most strongly from the phosphate group, while neutrons interact with nuclei and sca tter most strongly [Tom the carbonyl groups (see Figure 8.4). Thus to more fully describe bilayer struc-

-10

10

30

-to

10

30

B.

.q
cQ)

"D

9

8

..c: bJJ

7

~

6

~

O!l ~

'C

Joint Refinement of X-Ray and Neutron Diffraction Data

10


~

C1l

u

5 4


;>, C1l

....

X

3 2

-30

Distance from bilayel- center (A.) 8.4. Examples of diffraction data for DOPC bilayers at 23°C and

66% relative humidity presented as eight-order scattering-length density profiles. A. Neutron diffraction profile, whose peaks correspond to the carbonyl groups. B. X-ray diffraction profile, whose peaks correspond to the phosphate groups. From from Wiener, M. c., and S. H. White, Biophys J. 1992, 61 :434-447. © 1992 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

Liquid Crystallography

195

.~=

rffl

Crystal

Liquid

Smectic A

Smectic C

8.5. Crystals, smectic liquid crystals, and liquids. Smectic liquid crystals are ordered in layers consisting of rod-like molecules oriented with their long axes perpendicular to the plane of the layers. They may also be at an angle from the normal, as shown. The layers are free to slide over one another, giving the structural properties of a two-dimensional solid. Redrawn from Chang, R., Physical Chemistry {or the Chemical and Biological Sciences, University Science Books, 2000, p. 884. © 2000. Reprinted with permission from University Science Books.

2

4

5

---T--------------------~-----r-------l

I

A.

Ir1I

Gel phase

&f-------------_

~

I I

I

IH

,

Mica peaks from substrate

I I

:

:I

I:

:H

H:

HI

I

I

I I

I

: II

:

'0

:

:I

I:

I I I I

I I .. - - - _...r

:0

:

I I I I

I I 1 I

I

:

: : : CH 2 ----~--------------------T-----~-r-----~ I

qr

8.6. Bragg peaks for x-ray diffraction of phospholipid bilayers. A. X-ray scattering data from a sample of DMPC at 10°C (gel phase) show peaks (h) up to the seventh order. The variable q represents the scattering in two dimensions, z and r. B. X-ray scattering data from fully hydrated DOPC in fluid phase is much more diffuse, with overlap between Bragg orders 1 and 2. From Tristram-Nagle, 5., and J. F Nagle, Chem Phys Lipids. 2004, 127:3-14. © 2004 by Elsevier. Reprinted with permission from Elsevier.

: I

: II

:

°

I

CH 3 -{-(CH 2l7 -;C =C T-(CH2)r~--C-0 -{-CH

I I I I

B.

t

J

CH 3 -;- (CH 2)r-; C =C ~(CH2l7 +C -0-;-CH 2

I

,

3

I I

:

h ....0 _1_2_3__ 4_5_

1-----1

1 2

3 4 5 6 7 8

CH 3 CH 2 C=C COO GLYC P0 4 CHOL. WATER

0

I

I

:

6 :

O-P=O:

I

I

I

:

0 : ~ -- -- -- -- -- -- --1-- -- -- -- -- -- < I H-C-H

: I

7 : : I

: 1

I

H-C-H

I H 3C-N-CH 3

I

CH 3

--------

8 :----~=--H2-0--------: ...

J

8.7. A phospholipid such as DOPC is divided into submolecular groups to be used in the structure determination. Redrawn from White, S. H., and M. C. Wiener, in K. M. Merz, Jr., and B. Roux (eds,), Biological Membranes, A Molecular Perspective {rom Computation and Experiment, Birkhauser, Boston, 1996.

Diffraction and Simulation

196

Inte'rface

Hydrocarbon core

Intelface

o :0

'2"

.0

0..

r;h l'l'!'O!

holine Phosphate

8.8. The structure of a DOPC bilayer determined by the joint refinement of x-ray and neutron diffraction data. The peaks correspond to the methyl groups (CH31. methylene groups (CH21. double bonds (C=Cl. carbonyl groups, glycerol, phosphate, choline, and water The bilayer is divided into two interfacial regions (defined by the penetration of water) and a hydrocarbon core, as labeled at the top. From White, S. H., et aI., Biochim Biophys Acta. 2003, 555: 116-121. © 2003 by Elsevier. Reprinted with permission from Elsevier.

is no water in the internal hydrocarbon core. Also the two interfacial layers (each about 15 A thick) together make up about hall' the total thickness of the bilayer. Furthermore, this method of presenting the structure of the bilayer can convey other features of membranes. For example, when a peptide consisting of ] 8 alanine residues with neutralized N- and C-terminal groups (Ac18A-NH 2 ) inserts into a DOPC bilayer, it is detected 12 .".

-

0

X

10

-0

'6. 1-0

~

b)

A-DOPC + 18A

8

v

0.

0 if)

6IDOPC~\

c v

-0 OJ)

c

'C ~

4

2~' ..... /

'" o

u

.

I

(/)

I

-20

\

' , - - - Densily difference I '\ (approx. /SA dislribuliorli

,

).-.-.-:.-.-

.... -

I

\

,

,

I

,-"

-10 0 10 20 Distance from bilayer center (A)

8.9. The scattering density profiles of a DOPC bilayer with and without 5 mol % Ac-18A-NH2 (18Al. expressed on a per lipid basis. The dashed curve gives DOPC alone, the solid curve gives DOPC + Ac-18A-NH2, and the dot-dash curve gives the difference, showing the localization of the peptide. From Hristova, K., et al., J Mol Bioi. 1999,290:99-117. © 1999 by Elsevier Reprinted with permission from Elsevier

entirely in the headgroup region, indicating it is not hydrophobic enough to insert as a TM helix (Figure 8.9).

MODELING THE BILAYER

Simulations of Lipid Bilayers

For over t\NO decades. advances in computer hardware and software, including computer graphics, have made simulations of lipid bilayers more realistic and more predictive. To look at the resulting molecular models as more than pretty pictures requires some appreciation of the theoretical steps that go into generating a simulation. A static model requires specification of all the atoms in the system, which can be done with the familiar Cartesian coordinates x, y, and z. Therefore a system of N particles (atoms) can be described by 3N coordinates, which becomes formidable when the system contains more than a fevv hundred atoms. The system is also dynamic. so each atom has a velocity and direction and therefore is constantly changing positions. Furthermore, the atoms are part of molecules, so they have interactions due to bonds specified by lengths, angles, and rotations as we]] as non bonded interactions, such as electrostatic and van derWaals interactions. By specifying all these factors in a model of the system, welJestablished algorithms calculate the potential energy of the system as a function of its atomic coordinates. The changes in potential energy of all the atoms in the system are considel-ed as movements on a multidimensional surface called the energy surface

Modeling the Bilayer

197

A.

Metastable Stable

B.

Potential energy (kcal mo]-\)

ensemble generated by the simulation. The ensemble is defined by the variables held constant as the replications are generated: the canonical ensemble has a constant number of particles, volume, and temperature, and is therefore referred to as NVT. [t can be useful to work with other ensembles. such as NPT (holding pressure constant instead of volume) or NVE (holding energy constant instead of temperature). Simulations allow calculation of thermodynamic properties, such as heat capacity, and kinetic properties, such as order parameters. When possible, comparison of the calculated results with results from experiments is used to evaluate the accuracy of the simulation. Computer simulations in frequent use for lipid bilayers employ molecular dynamics (MD) and, to a lesser extent, Monte Carlo (MC) methods. Molecular Dynamics

HD +H

RHO

(bohrs) D + H2 8.10. Energy surfaces depict potential energy with analogy to forces on a ball. A. In one dimension, a ball will roll downhill until it reaches equilibrium at the bottom. A minimum corresponds to a stable or metastable state. In the metastable state, it is at a local minimum but not at the global minimum. B. In three dimensions, the energy surface can depict the transition states of a reaction, as shown here for the reaction D + H2 ---> HD + D, or it can depict the various conformational states of a molecule or macromolecular system. Redrawn from Dill, K. A, and S. Bromberg, Molecular Driving Forces, Garland Science, 2003, pp. 29 and 349. © 2003 by Ken A Dill, Sarina Bromberg, Dirk Stigter. Reproduced by permission of Taylor & Francis, a division of Informa pic.

(Figure 8.10). Stable structures correspond to minima on the energy surface. Since any movement away from a minimum describes a configuration with a higher energy, algorithms make small changes in the coordinates and determine whether the energy increases or decreases. In a complex energy surface with more than one minimum, the one with the lowest energy is caUed the global minimum, which usually (but not always) corresponds to the state observed in a biological system. When experimentalists measure some property of a system, they usually detect an average of that property over a large (often macroscopic) number of molecules and over the time it takes to make their measurement. If the property is called A, the experiment produces the time average, Aavc . However, computers can rapidly examine a large number of replications of the system and produce an ensemble average, denoted (A), the average value of property A over all replications of the

The first MD simulation of a lipid bilayer by van del' Ploerg and Berendsen in 1982 consisted of two leaflets of J 6 decanoate molecules each and lacked waters or lipid headgroups. Advances in computational abilities greatly increased the size of bilayer simulations. By 1996, a simulation of 17.000 atoms, representing 72 phospholipid molecules and 2511 water molecules, could be run for a duration of ~10 ns. A major quantitative result from MD simulations is the very rapid rate (20 ns- 1 ) of isomerization of dihedral angles along the acyl chains in Lex phase phospholipids (Figure 8.11), which increases the mobility of acyl chains compared with the standard picture of the fluid lipid bilayer. Today longer (sometimes ~ 100 ns) and more complex simulations model more than one kind of bilayer constituent, such as PL plus cholesterol, PL plus detergent, or PL plus a peptide or protein. Molecular dynamics calculates time averages of properties based on the dynamics of the system. Sequential determination of sets of atomic positions at very short time steps 0-10 femtoseconds) are derived using Newton's equations of motions (see Box 8.2). From the initial coordinates all intramolecular and intermolecular interactions of the atoms are computed to determine where the atoms will move and, with many repetitions, to generate their trajectories and thus predict the future state of the system. Because in a fluid bilayer the force on one atom depends on its position relative to many other atoms, solution of Newton's equations for all the atoms becomes very computationally expensive. The steps in setting up an MD simulation include the following: 1. Specify initial conditions, giving the positions and velocities of the particles and the interparticle forces. Impose boundary conditions.

Diffraction and Simulation

198

t

=0

t = 10

Starting from the initial set of coordinates and velocities, the forces on each atom are calculated from the derivatives of the potential energy function. The potential energy function, U, is differentiated into a number of components or parameters, as shown in Box 8.2. They include intramolecular parameters, such as bond length, bond angle, vibrational modes, and torsional potential (for rotation along the C-C axis), and intermolecular forces such as van der Waals interactions and electrostatic interactions. Each parametel- is described as the mathematical sum of the differences between instantaneous and equilibrium values. For example, the harmonic (meaning symmetrical) function for bond length (1), is

L

=

U(l)

k/;(l -lO)2.

BONDS

BOX 8.2. Molecular dynamics calculations

t = 20

8.11. Mobility of PL molecules in a fluid bilayer. The rapid rotation around C-C bonds of the acyl chains in L~ phase phospholipids results in striking chain mobility, illustrated in snapshots of three individual lipid molecules from the molecular dynamics simulation of a DPPC bilayer shown at 0, 10, and 20 ns. Kindly provided by S. E. Feller and R. W. Pastor.

2. Describe the potential energy function and algorithm to be used. 3. Determine the simulation time period, which depends on computer power. To get starting parameters for a simulation, the phospholipid molecule has been divided into portions that are similar to groups on other macromolecules or to small molecules (Figure 8.12). Some of these portions correspond to small model compounds whose geometries and interaction energies have been determined with ab initio calcula tions. Alternatively, the starting point can be derived from x-ray stll1ctures of lipids in the Lc phase, or the simulation can start with the lipids in L p phase and "melt" the structure to L~ phase, although the latter approach is rarely successful. Initial velocities can be assigned to the atoms using MaxwellBoltzmann distributions at the temperature of interest. Now that several MD simulations are available, they provide the starting point for others. To avoid the unrealistic effect of atoms hitting the wall at the boundary of the simulation, the boundary is considered permeable. Since the number of particles in the system is held constant, when a particle leaves the system an identical particle enters [Tom the opposite side.

Newton's laws of motion state that (1) a body in motion continues to move in a straight line at constant velocity unless a force acts on it; (2) force equals mass times acceleration (F = mal, which is the rate of change of momentum; and (3) to every action, there is an equal and opposite reaction. The trajectory for a motion of a particle is obtained by solving the differential equation d2Xi/dt2

=

Fx;/mj,

where mj is the mass of the ith particle, Xi is the coordinate along which it moves, and FXi is the force on the particle in the X direction. Similarly, the trajectories in the other two coordinates are Fy;/m; and Fzj/mj. The forces on each atom are calculated from the derivatives of the potential energy function: Fx

=

dU/dx, Fy

=

dU/dy, and Fz

=

dU/dz,

with Fx as the X component of force, Fy as the ycomponent of force, and Fz as the z component of force. U is the potential energy of the system. For a simulation U is equal to (E), which is the ensemble average of the energies of states generated during the course of the simulation. The total energy is calculated as: Etot

=

L

kb(r - ro)2

+

bonds

+L

L

k~(lX - lXQ)2

angle

K1 _3(r'-3 -

r6- 3)2

UB

+

L

ky(Y - YO)2

improper

+

L nonbonded

+

L

V[cos(m - TO)

+ 1J

dihedrals qjqj 12 ( 6] } {4TI€0 rij + € [ (r;j ) - r;j ) (J

(J

This form of the energy function is used in a program called CHARMM. The algorithm performs calculations using CHARMM or other available programs, such as AMBER and GROMOS. A more detailed introduction to the mathematics of computer simulations may be found in Molecular Modelling: Principles and Applications, by Andrew R. Leach, second edition, Prentice Hall, 2001.

Modeling the Bilayer

199

0

, f

5

o II

IC-C-c-ct c - c -

P

0

3

C-O"-- "O-C

O"C ..--C

c-c-c-cEEJ-c-o II

X:

o 4

X

C '.,,,'C C-N' "C H C_!'J-,,,,H H 2

8,12, The portions of a phospholipid molecule (either PC or PEl that are used to determine model compounds for the parameters in an MD simulation: 1. trimethanolammonium moiety of the PC headgroup, modeled by tetramethylammonium or choline; 2, an ammonium ion from the PE headgroup, for which parameters are taken from the protonated €-amino group of lysine or ab initio calculations for ethanolammonium; 3, phosphate group. like those in the nucleic acid parameter set; 4, ester bond to the acyl chains, modeled by methyl acetate, methyl propionate and ethyl acetate; 5, aliphatic chain, for which parameters are taken from aliphatic amino acids in proteins. Redrawn from Schlenkrich, M., et aI., in K. M. Merz, Jr., and B. Roux (eds.), Biological Membranes, A Molecular Perspective (rom Computation and Experiment, Birkhauser, 1996, p. 36. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

wherelo is an equilibrium bond length taken [Tom a simple model compound of known geometry - for example, for an alkyl chain - and k" is the vibrational frequency for that same compound. The bond angle energy and vibrational energy of the system can each be represented by a similar quadratic [unction, while the equation for torsional potentia] can describe two local minima for gauche conformations and a global minimum for the trans position, or it can describe a double bond having cis and trans states. The van der Waals interaction is modeled to account for the attractions between atoms and the sizes of atoms and is limi ted by the short-range nature of these interactions. Its value is based on experimental data such as heats of vaporization and densities. For electrostatic interactions the partial charge of each atom is needed to include the charge component of headgroup-headgroup, headgroup-solvent, and solvent-solvent interactions. These assignments may be based on ab initio calculations or may be calculated with both short-range and long-range summations. Each simulation has two phases, equilibration phase and production phase. For a simulation of a lipid bilayer, equilibration can take a relatively long time. When little or no change occurs, the system is assumed to have reached equilibrium. The longer the production phase, the more likely macroscopic properties

8.13. MD simulation of a DPPC bilayer. This view of a DPPC bilayer is from an 800-ps trajectory with A = 62.9 P/lipid. The atoms and atom groups are colored as follows: yellow, chain terminal methyl; gray, chain methylene; red, carbonyl and ester oxygen; brown, glycerol carbon; green, phosphate; pink, choline; dark blue, water oxygen; and light blue, water hydrogen. From Feller, S. E., R. M. Venable, and R. W. Pastor, Langmuir. 1997, 13:6555-6561. © 1997 by American Chemical Society. Reprinted with permission from American Chemical Society.

Diffraction and Simulation

200

ing stearoyl (CI8:0), oleoyl (CI8:169cis), and elaidoyl (CI8:169Irans), in both same-chain lipids (e.g., DOPC,

Stearic - - Oleic - - _. Elaidic ::;

~ ii (\l ~

e

0..

" " " .--~-----

-I

o

-0.5

0.5

Cos 8 8.14. Probability distribution of a kink in the acyl chain. The presence of a kink at the carbon-carbon double bond is indicated by the probability distribution function for the angle made between vectors representing the upper and lower halves of the acyl chain = 1, the two halves are parallel, as is on C1 of pe. When cos dominant for both stearic (no double bond) and elaidic (with a trans double bond) but not oleic (with a cis double bond), which has a fairly broad distribution of angles. Redrawn from Roach, e., et al., Biochemistry. 2004, 43:6344-6351. © 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

e

will emerge. For bilayers most simulations are ~ 10 ns, while some have reached ~ 100 ns. While this is still too short to detect macroscopic properties such as phase changes, MD can approach simulating domain formation. By the late 1990s an MD simulation of a DPPC bilayer demonstrated just how mobile the acyl chains could be in the fluid phase (Figure 8.13). Four simulations with constant particle number, pressure, temperature, and area (an NPAT ensemble) were can'ied out to determine the best value for the surface area per DPPC molecule, which turned out to be 62.9 A2. That simulation has provided a starting point for numerous other simulations, as \Nell as a baseline for comparisons when the type of lipid is changed. Both the acyl chains and the headgroup of the lipid have now been varied, with several interesting conclusions. The extreme mobility of the acyl chains in L",-phase DPPC could make it surprising that kinks made by cis double bonds in unsaturated chains (Figure 2.1 B) increase the disorder of the acyl chains enough to significantly lower mel ting transitions of unsaturated fatly acids compared with saturated fatty acids (see Chapter 2). The dynamic effects of unsaturated acyl chains can be appreciated in MD simulations of bilayers containing acyl chains with cis and lrans double bonds. The effects of frans double bonds are interesting in view of the link between consumption of frans fatty acids in foods and hean disease. In a systematic study, a number of J 8-carbon hydrocarbons were compal-ed, includ-

DEPC) and mixed-chain lipids (e.g., SOPC). Since the double bond is between C9 and CIO, chain packing of lipids was analyz.ed From the MD simulations by representing each chain as a pair of vectors stretching from C2 to C9 and from C10 to C 17. The ensemble-averaged measure of the relative orientations of the chain segmen ts, shown as probability distributions (Figure 8.14), indicate that for both steamyl and elaidoyl chains the most likely state is a parallel orientation (maximum probability at I), while a much higher distribution of bent conformations (kinks) is observed for oleoyl chains. Experimental measurements of fluidity and lateral mobility also reveal similarities between the saturated lipids and lipids with lrans double bonds, suggesting that replacement of cis unsaturated fatty acids by frans fatty acids depletes the lipids of the disorder that is needed in the membrane. Using the POPC bilayer as a template, an MD simulation of 72 SOPE molecules (with ~ 1500 waters) was equilibrated with NPT until a volume could be determined and then used for an NVE production phase, allowing both structural and dynamic analyses. The striking structural difference between PE and PC is the extensive hydrogen bond network between the primary amines of PE and phosphate groups (Figure 8.15). This hydrogen bonding dominates the dynamics of the interfacial region, where clusters of eight to 12 headgroups become locked together by hydrogen bonds for short times (on a nanosecond time scale). The segmental motion of the chains near the headgroup is slower in PE than in pc, while the dynamics of the acyl chains show diffel-ences between saturated and unsaturated chains.

....."

"\ ,

\

\

.....

...../- ........

~#~ - .....\.~~-.t \ .,.'. l" ~ 4-' -. ,~, .... '" ~ \ .,~ .%~, ~,w.~" ~ '( , ~ #.,;,~~~ "- U' :\

...

,

!\_,

I

,,"'.':--'.,

I

. , . . \. , h 'tiM 1...~II\II\\.... I~'" ~ 'r~'~~ .~ ~t '~~1( ,

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I

~~ '~-

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,"

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~~

.

•

~1 ~ .:1'11 11. ....' ; Jt.!'~'" (~ (), '\V'\;J!!l't- ,'\" '\'~ • .W ~\ l~ ~ ~ \ ,,.. ~ \' I~ ~ . 't.... ", , '7::.9t!' v·....., "" ~\ ~. r::-f~

.... '""4Wo

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-

....

\

I

~\'''., J~~' '. '111'. -

8.15. Snapshot of a 14-second NVE MD simulation of a fully hydrated SOPE bilayer showing the extensive hydrogen bond networks on the surface. Hydrogen bonds (blue and white stripes) between amine and phosphate groups make single, double and even triple linkages in rich patterns. From Suits, F., M. e. Pitman, and 5. E. Feller, J Chem Phys. 2005, 122:244714. © 2005 by American Institute of Physics. Reprinted with permission of the Office of the Publisher, American Institute of Physics.

Modeling the Bilayer

8.16. Snapshot of rhodopsin interacting with two molecules of 1stearoyl-2-docosahexaenoyl-PC in a simulated bilayer. Note how the polyunsaturated chains (spheres) bend in conforming to the surface of the protein (ribbon diagram). From Feller, S. E., and K. Gawrisch, Curr Opin Struct Bio/. 2005, 15:416-422. © 2005 by Elsevier. Reprinted with permission from Elsevier.

201

MD simulations have also examined the role of polyunsaturated fatty acids such as the beneficial omega-3 fatty acids (see Chapter 2). In contrast to early views that suggested polyunsaturated fatty acids increased the rigidity of the bilayer, MD simulations along with NMR studies indicate that the extra flexibility and rapid conformational fluctuations of these hydrocarbon chains increase the sohness of the bilayer. The most prevalent omega-3 fatty acid, docosahexaenoic acid (DHA, C22:6), is round in high concentrations (up to 50 mol %) in membranes of the nervous system. It is the dominant fatty acid in the rod cell membrane, where it is required for rhodopsin activation (see Chapter 9). DHA could be affecting rhodopsin indirectly by modulating membrane elasticity or activating it directly, since MD simulation indicates it interacts closely with helices on the surface of rhodopsin (Figure 816). Simulations of DPPC or DMPC and cholesterol have been done in several labs and at several different concentra tions of cholesterol. The condensing effect of cholesterol is evident in the decreased lipid surface area above approximately 10% cholesterol and can be attributed to close contact with acyl chains that "wets" the cholesterol, allowing it to come cJoser together (Figure 8.17). The Lo phase appears when cholesterol

8.17. Snapshots of DPPC-cholesterol interactions. The cholesterol molecules are depicted as spacefilling molecules, while the DPPC molecules are depicted as stick molecules. A. A lipid-cholesterol cluster seen in a 1: 1 DPPCcholesterol system. B. Two closely associated cholesterol molecules in a system of DPPCcholesterol at a 7:1 ratio. The hydroxyl groups of the cholesterol molecules are hydrogen bonded to different lipids, but are also quite close to each other. From Chiu, S. W., et aI., Biophys J. 2002, 83: 1842-1853. © 2002 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

Diffraction and Simulation

202

A.

B.

0.35 0.3

Co

Co 0.25

'(J)

c

'(J)

cQ)

-0 C

g

Q)

0.2

-0

0.15

j3

Co
<.)

..0

Q)

'iiJ

o

Stearic (w/chol)

0...

DHA (w/chol)

'-<

0.1

Stearic (neat)

0.05

DHA (neat)

0 -32 -24 -16

-8 0 8 z/Angstroms

16

24

32

-20

-10

o

10

20

z/Angstroms

8.18. Location of cholesterol in a lipid bilayer containing polyunsaturated fatty acids. A. Distribution of molecular groups along the z-coordinate (bilayer normal) for a lipid bilayer consisting of cholesterol and 1-stearoyl-2-docosahexaenoyl-PC in a 1:3 ratio. Because its hydroxyl group hydrogen bonds with water and polar headgroups in the interfacial region. cholesterol is located in the upper acyl chain region. as it is in DPPC. B. Distribution of acyl chains along the z-coordinate for stearic acid and docosahexaenoic acid (DHA) in the presence and absence of cholesterol. The cholesterol has a negligible effect on the location of stearoyl chains. but a significant effect on the location of DHA chains. Redrawn from Pittman, M. c., et al., Biochemistry. 2004, 43:1531815328. © 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

is ~ 12% to 50%; the lower concentration suggests that one cholesterol molecule can affect eight to nine PC molecules. In mixtures of PC, PE, PS, and cholesterol, cholesterol interacts preferentially with mixed-chain polyunsaturated PC, for which it prefers the saturated chain on Cl. MD simulations, along with NMR and xray data, indicate that DHA has a low affinity forcholestero!. While the loca tion of the cholesterol is not affected by the degree of unsaturation of the acyl chains (Figure 8.18A), the DHA chain is affected by it much more than a saturated chain is (Figure 8.18B). Monte Carlo

Because they solve equations of motion for all atoms in the system, MD simulations are limited on both time and length scales: currently the longest time is ~100 ns and the largest bilayer consists of fewer than 2000 lipid molecules, with most having fewer than 100 lipid molecules. MC methods, on the other hand, can be applied to model cooperative, large-scale behavior. Alternatively, for simple models they can describe molecular interactions on the atomic level, for example, simulating phase changes. In contrast to MD simulations where the successive configurations of the system are connected in time, in MC simulations small random changes in conformation are generated and each is compared only with its

predecessor to determine whether it represents a state of lower potential energy. By calculating the potential energy after making random small changes, such as in the rotation of a bond, and assigning a higher acceptance probability for the new configuration if the potential energy has decreased and a lower probability if it has increased, the simulation progresses toward configurations of lowest energy (greatest stability) without a kinetic input. Traditional MC simulations use NVT ensembles, although like MD, it can use others, such as NTP. For an MC simulation of a PL molecule, the hydrogen atoms may be omitted, reducing the lipid to the equivalent of three chains, the two acyl chains plus a chain corresponding to the phosphate and headgroup (Figure 8.19). The flexibility of each of these chains gives an enormous number of degrees of freedom that contribute to the conformation of the molecule. For example, DMPC in L", phase has around 15 to 20 degrees of freedom per molecule, mainly associated with the acyl chains. Initial MC simulations focused on the conformations of the acyl chains and applied lattice or polymer methodology. In the mid-1990s, a standard MC method allowed calculation of equilibrium properties of a bilayer consisting of 36 DPPC molecules and no water molecules. Once MC methods got up to ~100 PL molecules, variation of acyl chain length and incorporation of cholesterol were performed.

Lipids Observed in x-Ray Structures of Membrane Proteins

203

A configurational-bias MC method is now used in combination with MD to give more complete equilibration and sampling. For this method, the MD simulation stops at random times. and around 100 MC configurations are generated - for example, for each position of a randomly chosen acyl chain - to push the system closer to equilibration or energy minima.

LIPIDS OBSERVED IN X-RAY STRUCTURES OF MEMBRANE PROTEINS

8.19. Stick model of DMPC used in Monte Carlo simulations. The model shows three flexible chains: the upper chain is the polar headgroup. and the two lower chains are the hydrophobic fatty acyl chains. All have large degrees of freedom providing large numbers of random conformations during the simulation. From Scott, H. L., in K. M. Merz, Jr., and B. Roux (eds.l. Biological Membranes, A Molecular Perspective from Computation and Experiment, Birkhauser, 1996. © 1996 by Springer Verlag. With kind permission of Springer Science and Business Media.

Both membrane diffraction and bilayer simulations enhance the appreciation of the environment of membrane proteins by providing insight into the nature of bulk lipids in the membrane. In contrast, observations of lipids in the x-ray structures of membrane proteins show how specific lipid-protein interactions affect the structure of individual lipid molecules. Overall, these lipids exhibit much more varied confolmations than the bulk lipids of the bilayer (Figure 8.20), sometimes even displaying energetically disfavored eclipsed angles. The unusual configurations of these lipids are most likely stabilized by strong electrostatic and van der Waals interactions with specific groups or regions of the proteins, which probably also accounts for their adherence to the protein during the purification and crystallization procedures. It is possible, however, that they are forced into those configurations during crystallization, as discussed below.

Cytoplasmic 8.20. Lipid molecules in the high-resolution structure of bacteriorhodopsin. The identified lipids are varied in conformation. Lipids on the outer surface of the BR trimer are modeled as 2,3-diphytanyl-sn propane. Oxygen atoms are red. The green helix is helix Awith Arg7, Glu9, and Trp1 0 at the extracellular end and Lys30 at the cytoplasmic end in space-filling representation. From Lee, A. G., Biochim Biophys Acta. 2003, 1612: 1-40. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Diffraction and Simulation

204

TABLE 8.1. Lipids associated with integral membrane proteins in the protein database Lipid

Protein

Resolution (nm)

PDB file

DSPC DSPE PLPC SAPE PVPG Acy l4CL Acy l2PE Acy'2 PC Acyl2PI Acyl4CL Acyl2PE Acyl2PC Acyl4CL Acy l2PC (Glc Gal) acyl2Gro Acy l4CL Acy l4CL Acy l4CL DPPE DPPG Galactosyl S2GrO DOPC Acy l4CL Acy l4CL Acy l2PE Acy l2Gro PhY2 Gro Triglycosyl PhY2Gro PhY2 Gro PhY2 Gro PhY2 Ptd (Triglycosyl) CL PhY2 PG - P PhY2PG LPS

Paracoceus denitrificans CO Rb. sphaeroides CO

0.30 0.23 018 0.18 018 0.18 0.23 0.23 0.23 0.23 0.316 0.22 0.22 0.255 0.255 0.255 0.21 027 0.22 0.25 0.25 0.30 0.16 026 0.26 0.20 0.27 0.27 0.155 0.19 0.25 025 0.25 025 0.25

1QLE 1M56 1V54 1V54 1V54 1V54 1KB9 1KB9 1KB9 1KB9 1BCC 10KC 10KC 1M3X 1M3X 1M3X 100V 1E14 1EYS 1JBO 1JBO 1UM3 1KOF 1NEK 1NEK 1K4C 1BRR 1BRR 1C3W 10HJ 10M8 10M8 10M8 10M8 10FG

Bovine Bovine Bovine Bovine

CO CO CO CO

Saccharomyces cerevisiae CR S. cerevisiae CR S. cerevisiae CR S. cerevisiae CR Chicken CR Bovine AAC Bovine AAC Rb. sphaeroides RC Rb. sphaeroides RC Rb. sphaeroides RC Rb. sphaeroides RC Rb. sphaeroides RC

Thermochromatium tepidum RC Synechococeus efongatus PS I S. efongatus PS I Mastigoc!adus /aminosus cyt b 6 f E. coli Fdh-N E. coli Sdh E. coli Sdh Streptomyces lividans KcsA Halobacterium salinarum BR H. salina rum BR H. salina rum BR H. salina rum BR H. salinarum BR H. salinarum BR H. sa/inarum BR H. safinarum BR E. coli FhuA

PL abbreviations as in Appendix II with acyl chains S, stearoyl; P, palmitoyl; Phy, phytanyl; L, linoleoyl; V, vaccenoyl; A, arachidonoyl; and unidentified acyl chains where indicated Fragments of PLs abbreviated Ptd, phosphatidyl; Gro, glycerol. Other abbreviations: Glc, glucose; Gal, galactose; LPS, lipopolysaccharide; CO, cytochrome-c oxidase; CR, cytochrome-c reductase (cytochrome-bcl complex); AAC, ADP/ATP carrier; RC, photosynthetic reaction center; PS, photosystem; cyt b6 f, cytochrome-b6 f complex; Fdh-N, nitrate-induced formate reductase; Sdh, succinate dehydrogenase; KcsA, pH-gated potassium channel; SR, bacteriorhodopsin; FhuA, Fe-siderophore transporter. From Marsh, D., and T. Pali, Biochim Biophys Acta. 2004, 1666:118-141

In the analysis of the x-ray data, lipids are modeled to fit electron-dense regions observed in clefts or at the edges of the crystallized proteins. Unless the resolution is very high, better than ~2 A, the lipid is probably not well defined. Often only portions of the lipid are clearly resolved; for example, the relatively fixed glycerol and top part of the acyl chains of a phospholipid may be clear while the ends of the chains are lost. Headgroups are fTequently not well resolved (as in Figure 8.20), which suggests they are bound wi th lower affinity or less specificity than the rest of the molecule. In the cases where the headgroups are complete in the x-I-ay structure, they usually devia te from the conformation observed in Lc phase lipids.

As the number of high-resolution structures of membrane proteins ino-eases, the data on lipid configuration grow (Table 8.1). These lipids fall into three categories: annular lipids in the first shell of lipid surrounding the TM regions of the protein, nonannular lipids in crevices bet\.veen subunits, and integral lipids in unusual positions within the proteins (Table 8.2). Annular lipids mediate between integral membrane proteins and the bilayer, as they fit tightly in the grooves and clefts on the protein to cover its rough surface. The crystal structure of bacteriorhodopsin, which is unusual in showing a nearly complete shell of annular lipids, clearly demonstrates the complementarity between the protein surface and bound lipid molecules

Lipids Observed in x-Ray Structures of Membrane Proteins

205

TABLE 8.2. Numbers of annular and nonannular lipids observed in x-ray structures of integral

membrane proteins Nonannular lipids' Protein

PDB code

Annular lipids

Bacteriorhodopsin Rhodopsin Bacterial photosynthetic reaction centers

10HJ 1GZM

6 1

Rb. sphaeroides

100V

Between helices

Between subunits

2

lOGV

Tch. tepidum Photosystem 1 from S. elongatus Light-harvesting complex from spinach Cytochrome-c oxidase from P denitrificans Cytochrome bCI from S. cerevisiae Cytochrome b 6 (from Chlamydomonas reinhardtii Succinate dehydrogenase from E. coli Nitrate reductase ADP/ATP carrier from mitochondria Potassium channel KcsA

1M3X 1EYS lJBO lRWT 10LE lKB9 1090 lNEK 1016 10KC lK4C

1?

1? 2

1 1 1 2 1 4 2 1 1

7

"Nonannular lipids are classified as being located either between transmembrane (X-helices within a monomer or between subunits in a multimeric complex. From Lee, A. G., Biochim Biophys Acta. 2004, 1666:62-87

(Figures 8.20 and 8.21). Another example is the close fit of cardiolipin and phosphatidylcholine molecules into grooves on the surface of the mitochondrial ADP-ATP carrier (Figure 8.22). Some tightly bound lipids on the surface are essential to the function of the proteins, such as the cardiolipin in the photosynthetic reaction center (Figure 8.23).

In contrast to annular lipids, which are readily exchanged with bulk lipids in the bilayer, nonannular lipids are more tightly bound in compartments defined by protein molecules, between TM helices or betvveen subunits of either oligomeric membrane proteins or

PC

R151

8.21. Annular lipid molecules on the surface of bacteriorhodopsin. The high-resolution structure of BR is shown with portions of lipids bound to the surface. Lipids are green, and a squalene molecule is red. From Lee, A. G., Biochim Biophys Acta. 2003, 1612: 1-40. © 2003 by Elsevier. Reprinted with permission from Elsevier.

8.22. Annular lipid molecules on the surface of the mitochondrial ADP-ATP carrier (MC). Two molecules of cardiolipin and two of phosphatidylcholine are shown on the surface of the mitochondrial Me. In this view Arg151 and Trp70 are prominent in forming the grooves to bind lipids. The protein surface is color coded by electrostatic potential (blue, positive; red, negative; gray, neutral). From Lee, A. G., Biochim Biophys Acta. 2004,166662-87. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Diffraction and Simulation

206

A.

(j~

EI~6_

B.

M R267 M R267

M

WI48

HY30 MHI45 10

A

8.23. A molecule of cardiolipin binds to the hydrophobic surface of the photosynthetic reaction center of Rb. sphaeroides. A. View of the entire complex shows the location of the cardiolipin (in spacefilling representation), with Trp residues also in space-filling representation to define the interfacial region. The position of Glu1 06 is indicated to help define the cytoplasmic surface. B. Close-up view of the cardiolipin-binding site from the side and from the cytoplasm. The cardiolipin is in a depression between the TM a-helix of the H subunit and TM helices 3 and 5 of the M subunit. Residues involved in interactions with the cardiolipin molecule are shown in ball-and-stick representation. From Lee. A. G .. Biochim Biophys Acta. 2003, 1612: 1-40. © 2003 by Elsevier. Reprinted with permission from Elsevier.

lar'ge membrane complexes. A nonannular lipid in BR is the haloarchaeal glycolipid STGA (3-HS0 3 -Galp(3 16Manpal-2Glcpcd -archaeol), which binds in the central compartment of the trimer and therefore is separated from the bulk lipid (Figure 8.24). Integral lipids reside within a membrane protein complex and may be involved in its function and/or assembly. In contrast to both other groups, integral lipids may not even be aligned normal to the bilayer as they fit into internal sites. For example, the yeast cytochrome-bcl complex has an internal lipid, assigned as PI, whose acyl chains extend into a hydrophobic cleft that is parallel to the membrane plane and whose phosphate group is submerged lOA below the interfacial

zone of phosphodiester groups (Figure 8.25). Of the six PE molecules identified in the structure of cytochromec oxidase fyom Rh. sphaeroides, four are between subunits and are responsible for binding subunit IV in the complex. An important caveat is the reminder that lipids observed in static crystals of membrane proteins may be very dissimilar from the lipids in the native membrane. Problems result from the use of detel-gents during protein purification, which removes many native lipids. As a result, the protein is crystallized in the presence of less lipid than the membrane environment provides, and the remaining lipids could be atypical. Furthermore, detergent micelles cover the nonpolar regions of purified

The Crystallographer's Art

10

A

8.24. A lipid in the central cavity of the bacteriorhodopsin trimer. A nonannular lipid molecule, STGA (3-HS03-GalpI31-6Manpa12Glcpa1-archaeol) is observed in the central cavity of the BR trimer. Viewed from the trimer interior, this STGA molecule lies among helices from two neighboring BR monomers, with hydrogen bonds to Tyr64 and Thr67 (in ball-and-stick models) on one monomer. From Lee, A. G., Biochim Biophys Acta. 2003, 1612: 140. © 2003 by Elsevier. Reprinted with permission from Elsevier.

membrane proteins (see Figure 3.9). New techniques that avoid the presence of detergent, such as crystallization in cubic phase lipids (see below), can circumvent these problems. Indeed, the number of lipids bound to BR when crystall ized in cuba matches the stoichiometry of the purple membrane (see Chapter 5). Secondly, to be detected by crystallography, the lipids must be "frozen" in place, having lost both their dynamic fluctuations and fast exchange rates (~108 sec-I for annular lipids according to EPR studies; see Chapter 4). The unusual lipid configurations observed in annular lipids could result from crystal packing, for example, when flexible acyl chains are forced to adapt to the irregular protein surface during the dehydration step. Finally, important elastic properties of the membrane (such as lateral pressure) are obviously lost in crystals and could be essential to understanding key roles of the lipids. Nonetheless, the information provided by high-resolution structures has enriched the view of membrane lipids by expanding notions of their possible configurations.

THE CRYSTALLOGRAPHER'S ART

Crystallography of membrane components is a thorny challenge, given the inherent disorder of the fluid

207

mosaic membrane. Even obtaining material of the required high degree of pur;ty can be elusive for both lipids and membrane proteins. Purified lipidic materials are usually crystallized by a drop in temperature to achieve the Lc phase. It can take years, on the other hand, to find conditions that promote crystallization of integral membrane proteins, especially the very hydrophobic ones like the lactose permease (LacY). For decades, low-resolution structures from diffraction studies of proteins such as BR and porin that occur naturally in two-dimensional arrays provided the only images of integral membrane proteins. That situation changed dramatically with recent successes in obtaining high-resolution structures of membrane proteins. While the number of available atomic structures of membrane proteins is still relatively small, the structures showcased in the rest of this book exemplify their powerful impact on the field of membrane biochemistry. Many of these structures were obtained only after years of effort, and successes have been heralded with much attention and excitement. The first solution of a crystal structure of a membrane protein garnered the 1988 Nobel Prize for Johann Deisenhofer, Harmut Michel, and Robert Huber for their structure of the photosynthetic reaction center (see Chapter 5). The 2003 Nobel Prize in Chemistry for stnlctures of channel proteins was awarded to Roderick MacKinnon for the stmcture of the KcsA potassium channel and to Peter Agre for the discovery of aquaporins (see Chapter 10). During the J 5 intervening years, studies of the porins, bacteriorhodopsin, and many other membrane proteins contributed much to a fundamental understanding of membrane structure and function, but only recently have these and other integral membrane proteins yielded to analysis by x-ray crystallography. The excitement of seeing a new x-ray structure, usually visualized as a complex ribbon diagram, can eclipse the uncertainties inherent in its determination. One obvious issue is the limit of resolution in the observed electron density. When the resolution is greater than ~3.0 A, the structure may provide important new information on the fold of the peptide backbone, but it gives no knowledge of side chain positions and water molecules. Another issue is how complete the structure is. Sometimes portions of the protein are removed to facilitate its crystallization. Even when present in the molecule, the ends of polypeptide chains as well as some internal loops may be disordered enough in the crystal to go undetected by x-ray diffraction. The degree of disorder is indicated by the B factor, also called the temperature factor or the Debye-Waller factor. The B factor describes the degree to which the electron density is spread out. A higher B factor indicates a higher degree of uncertainty in fitting a model of the structure to the electron density, which can be due to higher mobility of the molecule in the region or can result from an error

Diffraction and Simulation

208

A.

B.

8.25. Internal phospholipids in the yeast cytochrome-bcl complex. A. The tightly bound phospholipids of the cytochrome-bcl complex from yeast include lipids in two internal cavities, circled on the structure (black circle and brown broken circle). The lipids, including those in the center numbered L2 and L7, are shown in space-filling representation (yellow, except for cardiolipin, which is cyan). Cofactors, including ubiquinone 6 (Uq6) are shown as ball-and-stick models (black), and the helices are colored according to subunits: cytochrome b (red), cytochrome c, (black). Rieske protein (green). Ocr6p (cyan), Ocr7p (mid-gray). Ocr8p (white), and Ocr9p (magenta). B. For context, the homodimeric complex is viewed from the side, with the intermembrane space at the top and the matrix at the bottom (same color scheme), and with yellow and red bars at the sides delineating the plane of the membrane. For a thorough discussion of this complex, see Chapter 11. From H. Palsdottir and Hunte, Biochim Biophys Acta 2004, 1666: 2-18. © 2004. Reprinted with permission of Elsivier.

in the model. lL is typical for membrane proteins to be quite ordered in the plane of the membrane and to have more disordered loops (with higher B factors) in the solvent-exposed loops (see Figure 8.28 and Chapter 9 Frontispiece). Examples of all these issues will be seen in the nexl three chapters. II is notoriously difficult to get well-diffracting crystals of some membrane proteins. The OmpF porin was crystallized for more than a decade before its structure was solved. More than 1 g of purified LacY protein was required for crystallization attempts, allowing more than J 000 crystals to be examined at a synchrotron over a similar period before its structure was determined. Success was finally achieved using a mutant of LacY that strongly favors one of its conformations (see Chapter 10). An alternative to this brute force approach involves screening multiple homologs of the protein of interest to select the one most suitable for crystallization, as was done for numerous proteins described in the next two chapters, including GlpT, MscL, MscS, and KcsA. Optimization of conditions employed multiple trials carried out in different detergents. Typically, integral membrane proteins are purified in detergents, with the choice of detergent being largely empirical (see Chapter 3). Often the detergent used for

purification is not appropriate for crystallization, so trials are carried out to test detergent combinations along with other additives, typically small molecules such as heptane. Crystallization of a protein-detergent complex requires judicious choice of detergents to avoid such undesirable characteristics as polydispersity and large micellar size, which interfere with packing in an ordered array If the polar head group on the detergent is too small, the detergent is likely to denature the protein. On the other hand, if it is too large, it will interfere with protein-protein contacts needed to form a crystal lattice. Since the detergent micelle covers the hydrophobic region, crystal formation depends on contacts between the exposed polar ends of the protein. Membrane proteins with small hydrophilic portions may lack sufficient exposed regions for protein-protein contacts to form a crystal lattice. To enlarge the hydrophilic regions and create space for detergent micelles, the protein can be bound to a Fab or Fv fragment of an antibody. These portions have the antigen-binding site of the antibody and are therefore extremely specific for the protein of interest. This method was first used to crystallize cytochrome oxidase and has been used successfully for other components of the respiratolY chain (Figure 8.26). An advantage of

The Crystallographer's Art

209

8.26. Crystallization of cytochrome oxidase from Paracoccus denitrificans mediated by Fv antibody fragments. Crystal lattices of the four-subunit cytochrome oxidase (COX) in A and the two-subunit COX in B both show crystal contacts involving the antibody fragments. Colors indicate the pairs, so a blue COX is bound to a red Fv and a green COX is bound to a magenta Fv. From Hunte, C, and H. Michel, Curr Opin Struct BioI. 2002, 12:503-508. © 2002 by Elsevier. Reprinted with permission from Elsevier.

this approach is that it enables the protein to be purified employing affinity chromatography with a tag Fused to the antibody fragment. However, it does raise the possibility that the relatively large antibody fragment (56 kDa or 28 kDa) alters portions of the membrane protein structure, especially when a domain of the protein is inherently flexible. A novel crystallization method that avoids the presence of conventional detergents utilizes lipids in cubic phases (see Chapter 2), usually created with racemic monooleoyl or monopalmitoleoyl glycerol. The cubic phase has viscoelastic properties that mimic the native membrane and allows protein diffusion throughout the sample to Feed the crystal nuclei. The crystalline array is formed of stacked two-dimensional sheets. Pioneered for crystallization of BR, in cuba crystallization has also been used For halorhodopsin, sensory rhodopsin II, and photosynthetic reaction center, all of which have colored pigments that ease the difficulty of harvesting the tiny crystals that result. Another new approach is the use of bicelles to promote crystallization of membrane proteins. Bicelles, small bilayer discs containing both lipid and detergent (see Chapter 3), become gel-like (promoting crystal growth) at 37"C but have low viscosity at room temperature, facilitating retrieval of crystals from the bicelles. When this relatively simple method was applied to BR, the crystal packing was completely different from that of previous crystallizations and generated a monomeric structure (Figure 8.27). Finally, there are membrane proteins whose structures have been solved only after deleting their TM

portions to increase their solubility. One example is cytochrome P450, which crystalJized in detergent only after it was genetically engineered to both delete the single TM sequence and substitute amino acids at several other sites involved in binding to the membrane (see Chapter 6). Of course the resulting structure, while informative, is not actually depicting a TM protein' While making very significant advances, the field of crystallography of membrane proteins has much Further to progress in some notable areas. First of all, Few x-ray structures of membrane proteins have achieved better than 2 A resolution. For those proteins with structures solved at 3 A or more, improving the resolution can be expected to provide many details in the positions of amino acid side chains and small molecules such as lipids ,md water. Thus achieving greater resolutions will lead to more certainty, as well as at least minor alterations in the structures. Also, more than one crystal form can be needed to eliminate artifacts arising from the crystallization contacts. For example, the J 5 C-terminal residues of bovine rhodopsin are resolved in only one of the two high-resolution structures, in which they are apparently immobilized in an artificial position due to their involvement in the crystal contacts (Figure 8.28). The only other difference in the two rhodopsin structures is the positions of two cytoplasmic loops, which have the most flexibility (highest B factors) as is typical of loops outside the membrane region. In addition, few structures are available for membrane protein conFormations in the presence and absence of bound substrate or ligand to indicate what

Diffraction and Simulation

210

A.

Lipid cubic phase crystals B.

""

C.

BicelJe crystals 8.27. Comparison of bacteriorhodopsin packing in bicelles and lipid cubic phase crystals. A. Top view of a layer of lipid cubic phase crystals. B. Top view of a layer of the bicelle crystals. The box delineates the ac face of the unit cell of dimensions a = 45.0 A. b = 108.9 A, and c = 55.9 A, and antiparallel molecules are shaded differently. C. Side view of the monomeric BR molecules showing the vertical displacement of the molecules. From Faham, 5., and J. U Bowie, J Mol Bioi. 2002. 316: 1-6. © 2002 by Elsevier. Reprinted with permission from Elsevier.

conformational changes might be involved in the function of the protein. The Ca 2+ ATPase is one example of a membrane protein whose structure has been solved with different ligands bound (Chapter 10). And the xray structures of several intermediates in the photocycle carried out by BR capture short-lived conformations involved in its mechanism (Chapter 5). While neither has yet portrayed an entire mechanism, these examples illustrate what structures of additional conformations can be expected to reveal.

Finally, the ability to compare the cl)'staJ structures with structural information obtained by NMR, which was crucial to the acceptance of x-ray crystallography of macromolecules, is not possible for the vast majority of membrane proteins. The solution of membrane protein structures by NMR has been rare because their size usually exceeds current limits for NMR. One exception is the very small Eml-E protein involved in multidrug efflux (see Chapter 11), which was characterized by high-resolution heteronuclear NMR (see Box 5.1) as welJ as by solid-state NMR before its x-ray structure was available. At present a small number of (3-barrel membrane proteins also have structures solved by both xray cl)'stallography and NMR. The solution structure of the small barrel protein OmpX in DHPC micelles determined by NMR is very close to its crystal structure in detergent and, in particular, confirmed the protrusion of its extracellular domain, where four of the (3-strands extend beyond the bilayer (see Figure 5.8 and Box 5.1). The structures of another eight-stranded (3-barrel called PagP, an enzyme in the outer membrane that transfers a palmitate chain from a phospholipid to lipid A, show excelJent agreement along the barrel, although as expected, the NMR structure gives more flexibility to the loops (Figure 8.29). In the case of OmpA protein (see Chapter 7), whose channel-forming activity has been controversial, the x-ray structure of its TM domain does not show a pore; however, NMR spectroscopy is able to detect dynamic gradients along the barrel axis that may contribute to pore formation. NMR of membrane proteins is a frontier that promises to make significant contributions as its future capabilities develop, which will alJow more comparisons with x-ray crystal structures. Given the small percentage of membrane proteins encoded in the genome whose structures have been solved, it is possible that the methods that have been successful in crystaJlizing membrane proteins to date have been selective. This may even account for the apparent existence of only two classes of polytopic membrane protein structures, as bundles of ex-helices and (3-barrels could be the types of mem brane proteins most amenable to crystallization. Future work could reveal much more variation in membrane protein structure.

MEMBRANE SIMULATIONS

Now that some structures of membrane proteins have been determined at the atomic level, they can be inserted into simulated lipid bilayers in systems that consist of proteins, lipids, waters, and ions and require 50,000 to 300,000 atoms to describe (Figure 8.30). This use of data [Tom x-ray crystallography with molecular dynamics can approach a description of integral membrane proteins in their physiological environment, the

Membrane Simulations

211

C-terminal fragment

C2loop

30

50 70

90 110 130 150

B-factor scale (I~)

A and a trigonal crystal (P3Jl at 2.65 A resolution. The position of the C terminus in P41 is likely an artifact due to its stabilization by intermolecular contacts in the crystal packing. In addition, the flexibility of the loops (indicated by their B factors; see coloring scale at bottom) results in their different orientations in the two structures. (See also the Frontispiece for Chapter 9.) From Li, J., et al., J Mol Bioi. 2004, 343: 1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

8.28. Variation in two crystal structures of bovine rhodopsin. A tetragonal crystal (P4,) at 2.2

8.29. PagP structures determined by NMR and x-ray crystallog-

,. J

,

'. ,~

raphy. When the PagP crystal structure (red) is superimposed on the 20 lowest-energy NMR structures in diacylPC micelles (black), the differences are in the loop regions and the position of the amphipathic helix, which is expected to lie at the membrane interface. From Ahn, V. E., et al., EMBO J. 2004, 23:2931-2941. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Diffraction and Simulation

212

8.30. Representation of proteins studied by MD in lipid bilayers. The proteins shown are BR, the KcsA potassium channel, the GlpF aquaporin, the OmpF porin, and the mechanosensitive channel MscS. From Gumbart, J, Y. Wang, A. Aksimentiev, E. Tajkhorshid, and K. Schulten, Curl' Opin Struct Bio/. 2005, 15:423-431. © 2005 by Elsevier. Reprinted with permission from Elsevier.

lipid milieu of the fluid membrane, which is usually so different from the crystal lattice. The interplay between simulations and crystallography will continue to enhance our understanding of membrane proteins and serves as a reminder that the x-ray structures of membrane proteins, including those described in the next chapters, should be viewed in the rich complexity of their dynamic lipid environment, the fluid mosaic membl'ane.

Schlenkrich, M., el aI., An empirical potential energy func, tion For phospholipids, in K. M. Merz, Jr., and B. Roux (eds.), Biological Membranes, A Molecular Perspeclive (rom Computation and Experimenl, Boston: Birkhauser, 1996, pp.31-81. Gumbarl, J., Y. Wang, A. Aksimentiev, E. Tajkhorshid, and K. Schullen, Molecular dynamics simulations of proteins in lipid bilayer·s. Curl' Opin Strucl Bioi. 2005, 15:423-431. Lipids Viewed in Membrane Protein Structures

White, S. H., and M. C Wiener, The liquid-cryslallographic structure of fluid lipid bilayer membranes, in K. M. Merz, Jr., and B. Roux (eds.), Biological Membranes, A Molecular Perspective /i'011"1 Compulation and Experimem. BoSlon: Birkhauser, 1996. ''''iener, M. C, and S. H. White, Fluid bilayer structure delel~ mined by the combined use of x-ray and neutron diffraclion. Biophys 1. J 991,59: 162-173. Nagle, J. F, and S. Tristl'am-Nagle, Structure of lipid bilayers. Biochim Biophys AC/Q. 2000,1469:159-195. Trislram,Nagle, S., and J. F Nagle, Lipid bilayers: thermodynamics, structure, fluctuations and interaclions. Chem Phys Lipids. 2004,127:3-14.

Marsh, D., and 1. Pali, The protein-lipid interface: perspectives from magnetic resonance and crystal structures. Biuchim Biophys Acla. 2004, 1666: 118-141. Palsdotlir, H., and C Hunte, Lipids in membrane protein sllUctures. Biochim Biophys Acla. 2004, 1666:2-18. Lee, A. G., Lipid-prolein interaclions in biological membranes: a stlUctural perspective. Biochim Biophys Acta. 2003,1612:1-40. Cal1:ailler, J.-P., and H. Luecke, x-ray crystallographic analysis of lipid-protein interactions in the bacteriorhodopsin purple membrane. A11IW Rev Biophys Biol11ol Struct. 2003, 32:285-310. Fyfe, P. K., et aI., Probing the interface between membrane proteins and membrane lipids by x-ray crystallography. Trends Biochem Sci. 2001, 26: I 06-112. Pebay-Peyroula, E., and J. P. Rosenbusch, High-resolulion structures and dynamics of memb.'ane protein-lipid complexes: a critique. Curl' Opin Scrucl Bioi. 2001, 11 :427-432.

Molecular Modeling

Crystallography of Membrane Proteins

Pastor, R. W., and S. E. Feller, Time scales of lipid dynamics and molecular dynamics, in K. M. Merz, JI~, and B. Raux (eds.), Biological Membranes, A Molecular Perspeclive from Computali011 and Experinlenl, BOSlon: Birkhauser, 1996. Scott, H. L., Statistical mechanics and Monte Carlo sludies of lipid membranes, in K. M. Merz, Jr., and B. Roux (eds.), Biological Membranes, A Molecular Perspeclive li'om Com, putation and Experiment, Boston: Birkhauser, 1996. SCOlt, H. L., Modeling the lipid component of membranes. CUlT O]1in Struct Bioi. 2002, 12:495-502.

Hunte, C, and H. Michel, Crystallisalion of membrane proLeins mediated by antibody ft-agments. CUlT Opin Struc[ Bioi. 2002, 12:503-508. Landau, E. M., and J. P. Rosenbusch, Lipidic cubic phases: a novel concept for the crystallization of membrane proleins. Proc Natl Acad Sci USA. 1996,93: 14532-14535. Faham, S., and J. U. Bowie, Bicelle crystallization: a new method for cryslallizing membrane proleins yields a monomeric bacteriorhodopsin structure . .J Mol Bioi. 2002, 316:1-6.

FOR FURTHER READING

Membrane Diffraction

9

Membrane Enzymes and Transducers

Uncertainties in filling structure to electron densities limit the resolution of structures produced by x ray crystallogrilphy, as indicated by the B factors represented on this rib on diagram of rhodo Sin. The 1M helices have a low B factor (green), while the external loops and some of the regions tow
An understanding of their lipid environment, structural constraints and predictions, types of fu nctions, and biogenesis lays the foundation for a survey of membrane protein structures. The remaining chapters showcase a gallery of high-resolution structures selected to be representative of the different well-characterized membrane proteins. The fact that the structures of the vast majority of the proteins predicted to be transmembrane (see "Predicting TM Segments" in Chapter 6) are unknown means these first-obtained structures will not likely portray all types of integral membrane proteins. The class of f)-barrel membrane proteins is overrepresented in the structure database, with over half of the approximately ] 00 unique structures of integral membrane proteins solved as of 2005. They undoubtedly have less stnlctural variation than the class of helical bundle proteins. The progress of determining the structures of helical membrane proteins has increased tremendously with 38 new structures in the last 5 years (Figure 9.1). Even more variety can be

expected as new structures are obtained for helical bundles and for membrane proteins of mixed secondary structures. The availability of structures has provided great insight into how membrane proteins function, igniting much interest and fresh excitement in the field. Often these structures are representative of many others in their families; occasionally they seem unique. This chapter looks at examples of membrane enzymes and transducers (including receptors) that are not part of extensive macromolecular machines; therefore, their structures tell much about how they carry out their functions. Chapter ] 0 describes structures of transporter-s and channels that are beginning to illustrate the conformational changes involved in mechanisms for passage of small molecules and ions across mem branes. Chapter 11 views examples of the complex assemblies of membrane protei ns that interact to do jobs such as electron transport, protein export, drug efflux, and vitamin uptake. 213

Membrane Enzymes and Transducers

214

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9.1. Acquisition of structures for helix bundle membrane proteins since the first crystal structure was solved in 1985. Data include only unique structures and were obtained from http:// blanco.biomol.uci .edu!Membrane _Proteins...xtal.html, Red rawn from Bowie, J. U" Nature, 2005, 438:581-589. © 2005, Reprinted by permission of Macmillan Publishers Ltd,

ENZYMES

membrane proteins, this 31-kOa protein has a f3-barrel structure. Most unusual is the fact that its active site is on the external surface of the f3-baITei. Finally, OMPLA is inactive as a monomer but active as a dimer. The enzyme activity of OMPLA is dependent on calcium. It has phospholipase AI and A2 activities, which means it can cleave the acyl chain from either carbon 1 or carbon 2 of phospholipids. In addition, it can cleave the acyl chain from a lysophosp.holipid. Its broad substrate specificity shows a minimum requirement of a polar headgroup esterified to an acyl chain of at least 14 carbon atoms. With such broad specificity, OMPLA must be strictly regulated to avoid significant losses of membrane phospholipids. The monomer of OMPLA is inactive, while the dimer is active. Activation or the enzyme is triggered by perturbations of the integrity of the outer membrane from events such as heat shock, EDTA treatment and spheroplast formation, phage-induced lysis, and colicin release. It is likely that such disruptions affect the normal lipid asymmetry of the outer membrane, which consists of an outer leaflet of lipopolysaccharide and an inner leaflet of phospholipids. The resulting nonbilayer structures allow phospholipids access to the active site of OMPLA, which promotes dimerization (see below). What then is its physiological role? OMPLA is constitutively expressed. Tn E. coli OMPLA is involved in release of bacteriocins, and in pathogenic bacteria it is involved in virulence, most likely because an increase in

As discussed in Chapter 6, very few structures of membrane enzymes are solved in their entirety, Three that are available at high resolution portray very different modes for carrying out catalysis in the membrane environment. The outer membrane phospholipase A (OMPLA) is a f3-barrel that functions as a dimer when it traps its phospholipid substrate, Prostaglandin H 2 synthase (PGHS) is a monotopic membrane protein that sits in one leaflet of the membrane to have ready access to its substrates, And fumarate reductase creales a "redox loop" for the generation of a proton motive force under anaerobic conditions. In addition, there are stnJCtures for enzymes that function in large complexes, such as cytochrome-c oxidase, that are described in Chapter 11.

OMPLA OMPLA is an unusual membrane enzyme for a number of reasons, It is one of the few integral membrane proteins in the outer membrane of Gram-negative bacteria that has enzyme activity. (The other known enzyme in the E. coli outer membrane is OmpT, a protease.) It belongs to a large family of lipolytic enzymes (enzymes that catalyze the hydrolysis of lipids and phospholipids), but it has no sequence homology with the water-soluble members of the family. Like other outer

9.2. The catalytic triad of OMPLA. Serine, histidine, and asparagine residues at the active site of OMPLA, with a hexadecylsulfonyl inhibitor covalently attached to Ser144, are shown from the crystal structure of OMPLA of E. coli, The dashed lines represent hydrogen bonds. Note the proximity of the calcium ion. Redrawn from Kingma, R, L" et ai" Biochemistry. 2000, 39: 1001710022, © 2004 by American Chemical Society. Reprinted with permission from American Chemical Society.

Enzymes

r T2

9.3. Structure of a monomer of OMPLA. The monomer has 12 l3-strands (blue), and two small ex-helices (red). The active site residues on the external surface are shown as ball-and-stick models. From Snijder, H. J., et al., Nature. 1999,401 :717-721. © 1999. Reprinted by permission of Macmillan Publishers Ltd.

Iysophospholipid content of the outer membrane gives increased invasive capacity. In addition, it has been proposed to enable cells to tolerate organic solvents by increasing phospholipid turnover to allow incorporation of trans fatty acids produced by a peri plasmic cistrans isomerase, which makes the membrane less permeable. The E. coli gene for OMPLA, called pldA, codes for 289 amino acids, the first 20 of which are a typical signal sequence that is removed by leader peptidase during export from the cytosol (see Chapter 7). Comparison of the E. coli gene sequence with the OMPLA genes (Tom 15 other Gram-negative bacteria reveals 20 strictly conserved residues and an additional 18 that are highly conserved. One conserved region contains the catalytic triad, which consists of histidine, serine, and asparagine residues (AsnI56-His142-Serl44 in E. coli; Figure 9.2). Thus OMPLA is a serine hydrolase, and site-directed mutagenesis of Serl44 abolishes its activity, except in the case of S144C, which has 1% the activity of the wild type. Similarly, HI42G has an activity four orders of magnitude lower than \-vild type. Asn156 is less essential and is less strictly conserved (Asp in one and GIn in another); its role is to orient the histidine residue, in the same manner of an Asp residue in classical serine protenses. In the mechanism, Ser144 performs a nucleophilic attack on the carbonyl carbon of the ester bond in the substrate, forming a tetrahedral intermediate. Cleavage of the bond generates a free lysophos-

215

pholipid, which can diffuse away, and an acyl-enzyme intermediate. which is subsequently cleaved by hydrolysis, releasing a fatty acid. Early results from circular dichroism and EM images suggested that OMPLA is a l3-barrel composed of 12 l3-strands. and this has been confirmed by x-ray crystallography when the structure was solved with a resolution of 2.6 A(Figure 9.3). Like other membrane 13barrels. the TM l3-strands of OMPLA are amphipathic. giving a hydrophobic surface with interfacial regions rich in aromatic residues, while the in terior is polar. It has polar loops on the outside end of the barrel and short turns on the inside end. The active site residues His 142 and Serl44 are located at the exterior of the barrel in the outer leaflet side of the membrane. The OMPLA monomer has a convex side and a flat side, and dimers form by the association of their flat sides (Figure 9.4). The dimer interface is hydrophobic, with a patch of four leucine residues from each monomer forming knobs that fit into holes on the opposite subunit (see Figure 9.5B). However, embedded in the hydrophobic region is a strictly conser'ved glutamine residue, Gln94, which makes a double hydrogen bond with the same residue in the other monomer. When cysteine residues are inserted into the flat side with the mutation H26C, the disulfide bridge that forms cross-links OMPLA to produce a covalent dimer. Reversible dimerization is triggered by addition of Ca 2+ i11 vitro, and binding of the inhibitor hexadecanesulfonyl Huoride stabilizes the dimeI'. The highresolution structure of the dimer binding the inhibitory hexadecanesulfonyJ chain shows very little difference

9.4. OMPLA dimer formation. One side of the OMPLA monomer is flattened, and this side interacts with another subunit to form the dimer, as evident in a view of the crystal structure of the OMPLA dimer from the top, looking down on the l3-barrels. The arrows point to the active sites, with the active site residues and the hexadecanesulfonyl inhibitor shown in ball and stick and the calcium ions represented by large spheres. From Snijder, H. J, and B. W. Dykstra, Biochim Biophys Acta. 2000, 1488:91-101. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

216

A.

9.5. Structure of the OMPLA dimer inhibited with hexadecanesulfonyl fluoride. A. Each subunit is colored as in Figure 9.3; in addition, two inhibitor molecules (ball-and-stick models) occupy the active sites between the subunits. The black lines indicate the polar-nonpolar interface of the membrane. B. The surface representation of the subunit interface on one monomer shows the clefts for substrate binding by indicating the surface area within 1.5 A of the van der Waals surface of the inhibitors (blue). Residues involved in the dimer interaction are labeled, with asterisks marking the knob-andhole pattern and a dashed triangle to label the hydrophilic cavity. From Snijder, H. J., et al., Nature. 1999, 401 :717-721. © 1999. Reprinted by permission of Macmillan Publishers Ltd.

from the structure of the monomer. The dimer has two deep clefts that extend 25 A from the active sites along the subunit intel-face (FigUl-e 9.5). The 16 carbon atoms of the inhibitor fit into these clefts and interact with both monomers. Thus the substrate binding pocket is only formed in the dimeric enzyme. Calcium is abundant in the bacterial outer membrane, and each monomer has two Ca2+ -binding sites, one with 1a-fold higher affinity than the other. However, in the dimer, both have high affinity (Kd ~5a I-tM). The Ca 2+ -binding site seen in the structure of the monomer is around I a Afrom the active si te, between loops L3 and L4, with two aspartate side chains as ligands (Asp149 and AspI84). The second Ca 2+ -binding site in the dimer is located at the active site and is the catalytic calcium site (Figure 9.6). The binding involves the side chain of Ser152 and one main-chain carbonyl oxygen atom from each monomer, with three water molecules in the binding site. The effect of the catalytic calcium ion is to polarize the two water molecules, forming an oxyanion hole that stabilizes the negatively charged intermediates during the reaction. Thus three factors contribute to the inactivity of the monomer: 1) the absence of a substrate-binding pocket, 2) the lack of the oxyanion stabilization that results from Ca 2+ binding the catalytic site, and 3) the physical separation of the active site (outer leaflet) from the substrate (PL in the inner leaflet). Perturbations of the outer membrane can make substrate available; then substrate binding triggers dimer formation and generates an active complex. Such a perturbation might be

triggered by the colicin release protein that presents phospholipids in the outer membrane, allowing dimerization to activate OMPLA. In this case, the job of OMPLA is to hydrolyze PLs to increase the permeability of the outer membrane so the colicins may be secreted. Clearly the structure of this unusual dimer of f3-ban'els held the key to understanding its regulation as well as its function. Prostaglandin Hz Synthase

In vertebrates from humans to fish, two isoforms of PGHS, also called cyclooxygenase (COX): carry out the committed step in prostaglandin synthesis. Their main catalytic function is the conversion of arachidonic acid to prostaglandin H 2 (PGH 2 ), using two molecules of oxygen (Figure 9.7). These enzymes are monotopic integral membrane proteins: they bind to the luminal leaflet of the ER membrane, as well as to nuclear membranes, and they require detergent solubilization to release them from the membrane. Implicated in thrombosis, inflammation, neurological disOl-ders, and cancer, they receive a great deal of attention as the targets of nonsteroidal anti-inflammatory drugs (NSA1Ds) such as aspirin, acetaminophen, and ibuprofen, as well as newer drugs such as rofecoxib (Vioxx) and celecoxib (Celebrex) that specifically target the second iso[orm.

, The enzyme is called PGHS here, rather than the more familiar name COX, because it contains t\Vo active sites, COX and peroxidase (POX), Ihat are discussed in more detail below.

Enzymes

217

A.

B.

9.6. View of the active site of the dimeric complex inhibited with hexadecanesulfonyl. A. The binding site at the active site of OMPLA is formed by residues on the outsides of both l3-barrel monomers (yellow and green). The calcium (white sphere) is liganded by the carbonyl groups from Ser106 and Arg147 along with four waters; none of the calcium ligands is charged. Below it, the inhibitor hexadecanesulfonyl (purple) occupies the substrate-binding pocket formed between the two monomers. The catalytic residues from the subunit on the right, Asn 156-His283-Ser144, are clearly visible, with an arrow indicating the sulfonyl oxygen that occupies the oxyanion hole. B. A close-up view of the polar end of the site shows electron densities of the active site residues, water molecules, and Ca 2+. From Snijder, H. J., et aI., Nature. 1999,401:717-721. © 1999. Reprinted by permission of Macmillan Publishers Ltd.

AH 2

0""'"

-I OOH

OH

AA 9.7. The main reaction carried out by prostaglandin H2 synthase. The enzymes accept a number of other fatty acid substrates, and the second isoform accepts neutral derivatives of arachidonate. AA, arachidonic acid; PGG2, prostaglandin G2; PGH2, prostaglandin H2. Redrawn from Rouzer, C. A., and L. J. Marnett, Biochem Biophys Res Commun. 2005, 338:34-44.

Membrane Enzymes and Transducers

218

BOX 9.1. Mechanism of action of prostaglandin H2 synthase Y38S'

~~.RHS

~C4H9

Y 38S

U

HR

~ ~-

~

•

C4 H 9

0,

O,,~

.

~

CH 2CH 2 COOH

1CH'CH2COOH

5-exo trig

C4 H 9

O~

O~C4H9

~ 0,

2

CH2CH'COOH

S-exo trig

CH 2CH 2COOH

O-~

Prostaglandin G 2

o---~-

0

o 1~

Y'"i

l";

C4H9~

CH,CH,COOH

o~

Reductam ~

(peroxidase site)


1

C,H,

~H'CH'COOH

Prostaglandin H 2

9.1.1. Reaction mechanism for the conversion of arachidonic acid to PGH2. Redrawn from Furse, K. E., et aI., Biochemistry 2006, 45:31893205 The crystal structures, along with data from mutant studies, spectroscopy, and EPR, provide insight into the mechanism of the reaction. The mechanism at the COX site is a controlled free radical chain reaction that can be divided into four stages (see Figure 9.1.1). Entry of arachidonic acid into the substrate channel where it interacts with Arg 120 positions the proS hydrogen on carbon 13 next to a free radical on Tyr385. Abstraction of this hydrogen creates an arachidonyl radical centered at carbon 13. A rearrangement of the radical to carbon 11 is followed by attack by 02 to form an 11-peroxy radical. The 11-peroxy radical attacks carbon 9 to form the endoperoxide with isomerization of the radical to carbon 8, and ring closure occurs between carbon 8 and carbon 12, producing the bicyclic peroxide. This is hypothesized to change the configuration of the substrate, repositioning the acyl tail to allow attack by the second oxygen on carbon 15 to generate a 15-hydroperoxyl radical. Abstraction of a hydrogen atom from Tyr385 produces PGG2 and regenerates the tyrosyl radical for the next round of catalysis.

The active enzymes are homodimers with subunit molecular weights of 70 kDa. The sequences of PGHS-I and PGHS-2 have 60% to 65% identity and only minor difFerences in structure, but they differ in expression and function. PGHS-l is expressed constitutively in a wide range of tissues, while expression of PGHS-2 is induced by inflammatory and proliferative stimuli and it is mainly found in nervous, immune, and renal cells. Due to catalytic controls, PGHS-2 is active in cells where PGHS-l is latent (see Box 9.1). PGHS-2 also has accepts a wider range of substrates, which aJlows for design of inhibitors that specifically target this isoform.

Initial generation of the free radical at Tyr385 is the result of a redox reaction at the POX site. The ferric heme reacts with a hydroperoxide activator (proposed to result from the reaction between nitric oxide and superoxide anion) to produce a ferryl-oxo derivative of the heme. Intramolecular migration of the electron transfers the radical from the heme to Tyr385. Once the COX reaction occurs, its product, PGG 2, serves as the substrate for the peroxidase at the POX site, generating PGH2 and alleviating the need for the peroxide. The initial need for peroxide to generate the free radical provides a way to differentiate between the PGHS isoforms catalytically. Activation of PGHS-1 by peroxide is much less efficient than activation of PGHS-2. Under many conditions, glutathione levels and cellular glutathione peroxidase keep the peroxide concentrations below those needed for PGHS-1. Therefore even though its expression is constitutive, PGHS-1 is catalytically latent in cells under many conditions.

Ovine PGHS-l (from rams) was the first monotopic protein for which a high-resolution structure was obtained. When the crystal structure for the murine PGHS-2 (from mice) was solved, it showed the backbones of the two structures to be superimposable. In this discussion common Features will be described in PGHS-l and residue numbers will refer to its sequence. The structures show three domains: a domain similar to epidermal growth Factor (EGF, residues 33-72), a membrane-binding domain (MBD, residues 73-116), and a catalytic domain (residues 117-586; Figure 9.8). As portrayed, the protein lacks its signal sequence that has been cleaved from the N terminus as is typical for

Enzymes

219

i

Flurbiprofen

EGF-like domain

9.8. The structure of the PGHS dimer viewed from the side and colored to indicate the different domains: EGF domains (red), membrane-binding domains (yellow), and catalytic domains (blue and gray). In the monomer on the left, the heme is pink and a bound flurbiprofen (an NSAID) is green. The plane of the membrane is indicated by the line at the bottom. From Fowler, P. W., and P V. Coveney, Biophys J. 2006, 91 :401-41 O. 11;:) 2006 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

proteins targeted to the ER, and it lacks the C-terminal end, which is not resolved in the x-ray structure (Figure 9.9). Three of the sites for glycosylation are shown on the monomer. Glycosylation is necessary for folding but not for enzyme activity once folded. The interface in

the climer involves the EGF-like and catalytic domains, with patches of polar, electrostatic, and hydrophobic contacts between the subunits (Figure 9.10). The role of the EGF-like domain is unclear, although it is a feature of many cell surface proteins. It contains three highly conserved, interlocking disulfide bonds, with a fourth disulfide linking it to the catalytic domain. The membrane-binding domain consists of

N

9.9. The x-ray structure of the PGHS monomer. Each monomer has three domains. The catalytic domain (blue) has two active sites, the POX site (top, at the heme) and the COX site (bottom), where arachidonic acid (yellow space-filling model) is bound. The membrane-binding domain (orange) is below the arachidonic acid, and the epidermal growth factor domain (green) is on the side that becomes the subunit interface in the dimer. From Garavito, R. M., and A. M. Mulichak, Annu Rev Biophys Biomol Struct. 2003, 32: 183-206. © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

9.10. View of dimer interface on one subunit, with space-filling representation of atoms within 5.0 A of the other subunit. The membrane anchor helices are at the bottom. Patches of polar and nonpolar interactions are evident in the colored side chains showing acidic residues (red), basic residues (blue), and hydrophobic residues (purple). The labeled positions indicate conserved residues. From Kulmacz, R. A., et al., Prog Lipid Res. 2003, 42:377-404. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

220

9.11. The side of the PGHS dimer that faces the membrane interior. The PGHS dimer (colored as in Figure 9.8) is viewed from the bottom. The four <x-helices of the MBD are labeled A, B, C, and D in the monomer on the left. From Fowler, P. W., and P. V. Coveney, Biophys J 2006, 91 :401. © 2006 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

four amphipathic <x-helices, three of which lie roughly in the same plane \vhile the fourth angles away from them into the catalytic domain (Figure 9.11). The MBD has hydrophobic and aromatic residues along the surface that interacts with the lipid bilayer. These are predicted to contribute to a strong interaction with the

membrane (with calculations of ll.G for binding as large as -37 kcal/mol). In a molecular dynamics simulation of a PGHS monomer inserting into POPC bilayers (Figure 9.12), hydrogen bonds between the phosphate groups of the lipids and basic residues on the enzyme are shown to further stabilize their interaction. The large catalytic domain has two active sites for the two steps of the reaction: the COX site that converts arachidonic acid to prostaglandin G 2 (PGG 1 ), a hydroperoxy endoperoxide, and a heme-containing peroxidase (POX) site that reduces PGG 2 to the hydroxyl endoperoxide, PGH 2 (see Figure 9.9). The catalytic domain has direct homology to members of the myeloperoxidase family. Thus PGHS-I and -2 are in the superfamily of heme-dependent peroxidases, but they are the only membrane proteins in the family. At the POX site, the heme is unusually open to sol· vent. It lies in a shallow c1eh on the surface opposite the MBD, coordinated by proximal (His388) and distal (His207) axial ligands (Figure 9.13). Although the groups around the heme are not highly conserved, the ferric/felTous midpoint potential is quite similar in the two isofom1s of PGHS. Below the heme is a long channel for binding long-chain fatty acid substrates (Figure 9.14). The COX catalytic center encompasses half the channel. from Argl20 to Tyr385. Arachidonic acid binds in an extended L-shape, with its carboxylate liganded by the guanidinium group of Argl20 (Figure 9.15). Carbons 7 through 14 weave around the

z

1 "

t =

t

0 ns

= 5 ns

t =

1 ns

t = 2 ns

t = 10 ns

9.12. Snapshots of a molecular dynamics simulation of the integration of a monomer of PGHS with a POPC bilayer, with 17 lipid molecules removed to accommodate the inserted protein. The nitrogen atoms of the choline in POPC are blue spheres to aid visualization of the lipid-water interface. The enzyme MBD is stably inserted by 5 ns in a repeatable manner. Simulation of the insertion of a dimer is expected to give similar results. From Fowler, P. W., and P. V. Coveney, Biophys J 2006, 91 :401 © 2006 by the Biophysical Society. Reprinted with permission from the Biophysical Society.

A.

B.

9.14. Substrate access channel of the COX site of the catalytic domain of PGHS. The ex-C tracing of the PGHS-1 monomer (blue) is shown with the substrate access channel highlighted (pink). Active site residues are Arg120, Tyr385, and Ser530 (yellow). and the heme is also indicated (red). From Rouzer, C. A, and L. J. Marnett, Biochem Biophys Res Commun. 2005, 338:34-44. © 2005 by Elsevier. Reprinted with permission from Elsevier.

9.13. Heme in the POX site of the catalytic domain of PGHS. A. The heme is in a cleft that is quite exposed on the surface of the catalytic domain. B. A close-up of the heme-binding site of PGHS-1 shows the proximal (His33) and distal (His207) ligands, along with other amino acids that define the heme-binding pocket. From Garavito, R. M., and A M. Mulichak, Annu Rev Biophys Biomol Struct. 2003, 32:183-206. © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure, www.annualreviews.org.

side chain of Ser530 (the residue acetylated by aspirin) and carbon 13 is pointed toward the phenolic oxygen of Tyr385. The rest of the acyl chain (carbons 14-20) binds in a hydrophobic groove above Ser530 but is not resolved in crystal structures. There is room for movement in the channel that enables a few minor products to be synthesized from arachidonic acid and allows several other fatty acids to bind as substrates, forming other bioactive products. In PGHS-2 substitution of a valine for lle523 creates an additional pocket that allows fatty acyl derivatives to bind, such as 2-arachidonylglycerol and arachidonyl-ethanolamine. The products of these two reactions are both endogenous ligands for the cannabinoid receptors that mediate the effects of the

9.15. A portion of arachidonic acid in the substrate channel of a mutant PGHS-1. The first 12 carbon atoms of arachidonic acid are modeled (pink) into the electron density (light green) and compared with a simulated structure (blue). The structure shows that the mutations, V349A and W387F, do not significantly affect the structure of the active site. The side chains that contact the substrate are identified and shown with stick representations (gray carbon atoms, red oxygens, dark blue nitrogens, and yellow sulfur). From Harman, C. A, et al., J Bioi Chem. 2004, 279:4292942935. © 2004 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Membrane Enzymes and Transducers

222

A.

B.

Flurbiprofen binding PGHS-l

SC-588, a COX-2 inhibitor

9.16. Binding of NSAIDs to PGHS as revealed by crystal structures. The amino acid side chains that define the binding sites are shown, with those critical to the differences between the two isoforms in space-filling models: isoleucine residues at positions 434 and 523 in PGHS-1 are replaced with valine residues in PGHS-2, allowing a shift in Phe518 (all three copper-colored) that gives access to a more polar side pocket between Ser530 and Tyr385 for specific COX-2 inhibitors. A. Flurbiprofen (yellowgreen) binds in the substrate channel of PGHS-1 B. SC-588 binds in the substrate channel of PGHS-2, filling the side pocket with the phenylsulfonamide group that prevents its binding to the active site of PGHS-1. From Garavito, R. M., and A. M. Mulichak, Annu Rev Biophys Biomo! Struct. 2003, 32: 183206. Q 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure. www.annualreviews.org.

active component of marijuana. Clearly the Oexibility at the active site has important biological consequences that are of interest. Manv NSAIDs act as competitive inhibitors. Highresolution structures solved with NSA1Ds bound to the enzyme sho"v how they prevent su bstJ'ate binding by occupying the upper part of the COX channel between Arg120 and Tyr385 (Figure 9.16). The interactions between the drugs and the enz.yme are hydrophobic, with the exception of inter-actions of the acidic NSAIDs with Arg120 and the potential of forming a hydrogen bond with Ser530. The extra binding pocket in the active site ofPGHS-2 has been exploited to design specific inhibitors that do not fit the channel ofPGHS-1 (commonly known as COX-2 inhibitors). For example, nurbiprofen interacts with Arg120 and fills a portion of the substrate channel in PGHS-l, while the phenylsulFonamide group of SC-588 fits into the pocket of the channel in PGHS-2. Other NSAlDs follow binding with covalent mod ification of the enzyme. When aspirin acetylates Ser530 it blocks the binding of arachidonic acid to PGHS-l, thereby inhibiting the enzyme completely. Acetylation has a diFferent effect on PGHS-2, in which it does not inhibit substrate binding but results in formation of a different product, presumably because it aFFects the alignment of the substrate in the channel. As these differences are likely to have significant biological consequences, developing new inhibitors to help minimize side eFfects ensures continuing interest in these enzymes.

Formate Dehydrogenase

A very different membrane enzyme is Formate dehydrogenase. Many prokaryotes can live anaerobically and carry out oxidative phosphorylation with a variety

Formate dehydrogenase-N HCOO-

CO 2

+ H+

\J 2 H+

2 e-: P-side

bp ;'

b,.: N-side

.' .' ..-rviQH~·

r.. · .

Mg.

~/

.

2H+

N0 2 - + H 2 0

N0 3 - + 2 H+

Nitrate reductase 9.17. Diagram of the nitrate respiratory pathway. Formate dehydrogenase-N and nitrate reductase form a redox loop, in which formate is oxidized and nitrate reduced with electron transfers that are accompanied by the transfer of protons across the membrane. From Jormakka, M, et al., FEBS Lett. 2003. 545:2530. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Enzymes

223

-1-20 m\

Mo

bis-\1<'0 [~Fe-~

J-O

(~Fc-~Sl-I I

',obooi, {

.

",.

",.

1~ll'-~SH

2B

(~ ,:-~Sl-2

L~r . . ~~SJ.1

..,... . . MQH 2

hI' -y subunit {

,

~ bc..-MQ • ·

-75 mV

MQ ........

.,""'-"

MQ-~

hI'

~

-75

mv}

•

-y subunit

be

t

(3Fc-4SJ-3 V N

IAF<,-451-2

,

L4Fe-4SH

} ~

subunit

L4Fc-4SJ-1

~

his-MGD Mo

~

} ex subunit

400 mV 9.18. Redox potentials of the electron carriers in the nitrate respiratory pathway. Electron carriers of FDH-N are given in red, and those of nitrate reductase are given in blue The highly exergonic reaction allows the electrons to be transferred against the membrane potential. From Jormakka, M, et aI., FEBS Lett. 2003, 54525-30. '10 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

ex su bunit

13 subunit

-y ~ubl1llit

9.19. High-resolution structure of FDH-N. The mushroom-shaped heterotrimer is viewed from the side with the catalytic <X subunit (dark blue), the 13 subunit (red), and the y subunit (light blue) Electrons are transferred from the active site in the periplasm with its Mo atom (green) to five [4Fe-4S] clusters (gray/yellow) to two heme b groups (black) and then to the quinone (purple). A single cardiolipin (yellow-green) is also visible in the trimer interface. The P-side is the peri plasmic side of the membrane and the N-side is the cytoplasmic side of the membrane. From Jormakka, M., et aI., Curr Opin Struct BioI 2003, 13:418-423. ~: 2003 by Elsevier. Reprinted with permission from Elsevier.

224

Membrane Enzymes and Transducers site on the cytoplasmic side. There nitrate is reduced to nitrite, consuming two more protons from the cytosol. The transfer of electrons from the outside to the inside of the membrane, followed by their transport back to the outside accompanied by protons was called a "redox loop" in Mitchell's original chemiosmotic theory. The high-resolution structure of formate dehydrogenase-N (FDH-N) from E. coli provides insight into the steps involved in generating a proton motive force from the electron transfers of the nitrate respiratory pathway. This enzyme is a 51 O-kDa dimer of a heterotrimer that consists of three subunits called lX, {3, and 'y (Figure 9.19). The catalytic lX subunit is a large peri plasmic polypeptide composed of 982 amino acids, including an intrinsic selenium-cysteine (SeCys) residue, a [4Fe4SJ cluster, and a molybdopterin guanine dinucleotide (MGD) cofactor. The {3 subunit (289 amino acids) has one TM helix near its C terminus, which is on the cytoplasmic side of the membrane, while its N terminus is on the periplasmic side. It coordinates four [4Fe-4SJ clusters positioned in two symmetrical domains (Figure 9.20). The y subunit (216 amino acids) is a polytopic protein with four TM helices and both Nand C termini on the cytoplasmic side (Figure 9.21). This

9.20. Structure of the [3 subunit of FDH-N. Each of the two symmetric domains outside the membrane contains two 14Fe-4S] clusters (colored as in Figure 9.19). From Jormakka, M., et aI., Curr Opin Struct BioI. 2003, 13:418-423. © 2003 by Elsevier. Reprinted with permission from Elsevier.

of terminal electron acceptors, including nitrogen. The presence of nitrate under anaerobic conditions induces the nitrate respiratory pathway, which consists of two membrane enzymes, formate dehydrogenase and nitrate reductase (Figure 9.17). Both enzymes are heterotrimers with numerous cofactors involved in electron transfers that enable proton transfer across the membrane to generate a proton motive force that can be used for the synthesis of AT? by F1-F o AT? synthase (see Chapter II). During anaerobic respiration, bacteria can produce formate from acetyl-coenzyme Aderived from pyruvate. In the reaction carried out by formate dehydrogenase, formate is oxidized to CO 2 and H+ on the peri plasmic side of the cytoplasmic membrane, releasing two electrons that are transferred across the membrane to a lipid-soluble quinone called menaquinone (MQ). The reduced MQ picks up two protons from the cytosol to form menaquinol (MQH 2 ). The redox potentials for the electron carriers indicate this is a highly exergonic reaction (Figure 9.18). The MQH 2 then diffuses through the membrane to nitrate reductase, which oxidizes it, releasing its two protons to the periplasm and transferring its two electrons via a series of co factors to the nitrate reduction

Heme b p

Heme be

N ("I subunit)

9.21. Structure of the integral membrane domain of FDH-N. The y subunit has fourTM helices, numbered tmyl through tmylV, and the TM helix of the [3 subunit (tm[3; red) is also shown. Histidine residues that coordinate heme b and residues involved in binding MQ are labeled. From Jormakka, M., et al., Curr Opin Struct Bio/. 2003, 13:418-423. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Enzymes

225

60

A

ex subunit

:\'10

MGDI MGD2

I]3.8 (6.0)

Fe, -0 3.6 FeS-] 111.9 FeS-4 113.5 FeS-2 I ] 2.0 Fe -3

I]

f3 subunit

'Y subunit

(10.6) (9.0) (10.3) (9.0)

114.9 (10.2) Heme b p 1 20.5 (10.7) Heme be

I 8.0 (0.0)

HOQ 0

~l

9.22. The electron path in an ex(3y protomer of FDH-N. The linear electron transfer pathway from Mo to the quinone-binding site is shown with center-to-center and edge-to-edge (parentheses) distances between the redox centers. From Jormakka, M., et ai, FEBS Lett. 2003, 545:25-30. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier. Inset: The structure of the molybdenum guanine dinucleotide (MGD) cofactor in the ex subunit of FDH-N. Redrawn from Khangulov, S. v., et ai, Biochemistry. 1998, 37:3518-3528.

Membrane Enzymes and Transducers

Heme be

•...•....

.', -. ... -:

~_ ... ,. w1990 _ w1729 w1370 .

or

Cytoplasm 9.23. Menaquinone-binding site in FDH-N. The inhibitor HOONO (dark blue) binds very close to the heme b (red) at the cytoplasmic side. A possible proton pathway from the cytoplasm to the bound MO is indicated by the location of several waters From Jormakka, M., et aI., FEBS Lett. 2003, 545:25-30. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

subunit is a cytochrome b that contains two heme b groups, one coordinated by His residues of TMs II and IV near the per'iplasmic side and the other coordinated by His residues of TMs I and IV near the cytopl8.smic side of the membrane. It also has the site for MQ reduction. Based on the linear path of the 11 redox centers in an ex~y heterotrimei~ as well as the close distances between them, it appears that the functional unit of FDH-N is the heterotrimer (Figure 9.22). The electron path spans almost 90 Aacross the enzyme. The sequence is initiated by the oxidation of formate by the ex subunit, in a reaction like that of the soluble formate dehydrogenases, whose catalytic subunits are supei-imposable with the FDH-N ex subunit. The Mo at the active site is at the end of a funnel lined with positively charged residues. The Mo is coordinated by cis-thiolate groups of the two MGD cofactors (Figure 9.22 inset), the SeCys, and a hydroxide ion and directly receives electrons from the substrate. The position of the SeCys is very close to the ex proton of the substrate, indicating it may be involved in proton removal and transfer to a nearby His side chain. The location of the quinone-binding site \-vas determined by soaking the crystals with the inhibitor 2heptyl-4-hydroxyquinoline-N-oxide (HOQNO) and was

found to be near the heme b on the cytoplasmic side of the membrane. This establishes a mechanism for proton translocation, since earlier EPR studies showed that the MQHz-binding site on nitrate reductase is close to its heme b on the periplasmic side. A group of waters near the FDH-N quinone-binding site suggests a proton path from the cytosol to the bound MQ (Figure 9.23). Thus when two electrons are transferred fl-om the formate oxidation site in the periplasm, two protons are taken up h'om the cytoplasm. When they are translocated across the membrane (as MQH 2 ), they are released to the peri plasm from the quinone-binding site in nitrate reductase . The overall structure of FDH-N is like that of other membrane-bound respiratory enzymes, which generally have two membrane-associated subunits and one integral membrane subunit, and unlike the many formate dehydrogenases that use nicotinamide adenine dinucleotide (NAD+) in aerobic organisms. The other NAD+ -independent formate dehydrogenases differ in the wide variety of redox centers they utilize. However, in E. coli, FDH-N is similar to formate dehydrogenase-O, another membrane-bound heterotrimer, and its ex subunit is highly homologous to the catalytic subunit of the soluble FDH-H described above. The high-resolution structure has given insight into the group of formate dehydrogenases with SeCys at their active sites, as well as a model for how a redox loop generates a proton gradient.

TRANSDUCERS Some membrane proteins transduce signals across the membrane in response to various triggers. G-protein coupled receptors (GPCRs) make up a huge superfamily of membrane receptors that respond to chemicals, light, or odor and activate G proteins to initiate signal cascades in many cells. Humans have more than a thousand different types of GPCRs, including many receptors fOJ- hormones and neurotransmitters (see Chapter 6). A prototype for this superfamily is rhodopsin, the GPCR that responds to light in ways that are thought to be shared by many o[ the other receptors. A much different trigger is involved when mechanosensitive (MS) channels transduce physical perturbations of the membl-ane into chemical and electrical signals. In response to pressure changes, they change the membrane permeability, leading to depolarization or hyperpolarization, entry of calcium, or release o[ osmolytes. Physiologically important pressures in humans range [Tom the effect of a faint sound on the auditory hair cell (\0-4 N/m L ) to the large tensions

Transducers

227

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9.24. The generalized function of G-protein coupled receptors. GPCRs respond to a variety of stimulants, including light, odorants, calcium ions, small molecules, and proteins. They trigger activation of a G-protein cxl3y complex, which stimulates the release of second messengers or affects channels in the membrane. FSH, follicle-stimulating hormone; LH, luteinizing hormone; TSH, thyrotropin. From Bockaert, J., and J. P. Pin, EMBO J 1999, 18:1723-1729. © 1999. Reprinted by permission of Macmillan Publishers Ltd.

in skeletal muscles and the heart 00" N/m 2 ). With such a broad range, it is not surprising that MS channels of very different structures have evolved. The best-characterized MS channels are those that enable bacteria to respond to sudden drops in osmotic pressure, two of which have high-resolu tion structures. Although they do not provide a prototype forthis diverse group of membrane proteins, they do give insight into how this kind of signal transduction can occur. Rhodopsin, a GPCR GPCRs respond to external signals such as light, odorants, hormones, and neurotransmitters and activate G proteins in signal transduction pathways that alter levels of second messengers and/or control ion channels (Figure 9.24). As key elements in sensory physiology, neurobiology, and pharmacology, they make up ~6% of the human genome and are targets of over half the drugs currently on the market. GPCRs share a common structure of seven TM helices, the serpentine structure already described for bacteriorhodopsin (see Chapter 5). Also like BR, they have been thoroughly studied with EM, genetics, and many spectroscopic techniques. Rhodopsin is the only GPCR

whose structure has been solved at high resolution; therefore, it provides a prototype for this important superfamily. Rhodopsin consists of an apoprotein called opsin and a chromophore, II-cis retinal. The bovine rhodopsin that has been crystallized has 348 amino acids (40 kDa) and spans the membrane with seven ['(helices, with its C terminus in the cytosol and N terminus on the exo-cytoplasmic surface (Figure 9.25). The seven helices of serlJentine GPCRs have highly conserved residues at key positions that form the basis for a generic system for numbering residues. For comparison of different GPCR structures, each residue is given a number for the helix it is in, followed by a number for its position relative to the most conserved residue in the helix, which is assigned an index of 50. For example, all residues in helix H7 are 7.xx, wher-e xx refers to its position relative to the most conserved residue in helix H7. Thus Lys296 in bovine rhodopsin, which makes the Schiff base to II-cis retinal, is Lvs7.43 because it is in helix H7 and is seven residues before its most conserved residue. GPCRs also have in common many aspects of their [-unction, which can be illustrated by the role of rhodopsin in signal transduction.

Membrane Enzymes and Transducers

228

C-II

C-III

Cytoplasmic side

Extracellular side

9.25. The topology of bovine rhodopsin. The protein is a bundle of seven TM ex-helices (yellow region) with a glycosylated exo-cytoplasmic N-terminal tail and a cytoplasmic C terminus. Lys296 (purple) binds to retinal, acyl chains are on Met1 (orange) and Cys322 and Cys 323 (light green). and the disulfide between Cys110 and Cys187 (dark yellow) stabilizes the helices. The predominant phosphorylation sites are Ser334, Ser338, and Ser343 (green). From Palczewski, K., Annu Rev Biochem. 2006, 75:743767. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Rhodopsin Function Rhodopsin is a photoreceptor in the rod cell of the vertebrate eye. The eye has two types of light-sensitive neurons, rod cells and cone cells (Figure 9.26). Rod cells are very sensitive to light and responsible for high-resolution and night vision, while cone cells are responsible for discerning color. Cone cells are of three types, each of which has one type of photoreceptor that detects red, green, or blue color. All three cone photoreceptors are very similar to rhodopsin, with slight differences in the opsin structure that changes the absorption spectrum of the chromophore II-cis retinal. Color blindness results from mutations in one of the genes for these opsins. The rod cell has two parts, an outer segment loaded with membrane discs containing rhodopsin and an inner segment containing the nucleus and mitochondria. The electrical potential of the cell is produced by the Na+K+ -ATPase in the inner segment. The outer segment has an ion channel for Na+ and Ca 2+ that is opened in response to cyclic guanosine monophosphate (cGMP). When light hits the membranes in the outer segment of the rod cell, the concentration of

cGMP decreases, closing this channel and changing the electrical potential from -45 mV to -75 mY. This change, called hyperpolat-ization, sends a signal to the optic nerve (Figure 9.27). The main events of hyperpolarization are catalyzed by rhodopsin and the G protein called transducin (Figure 9.28). When a photon hits rhodopsin it triggers conversion of the II-cis retinal to all-lral1s retinal, leading to conformational changes that stimulate rhodopsin to interact with transducin. Transducin is the member of the family of heterotrimeric GTP-binding proteins that is involved in visual transduction. In the dark, the transducin Ct, 0, and y subunits form a complex at the cytoplasmic surface of the membrane, with GDP bound to the Ct subunit. Excited rhodopsin stimulates the replacement of GDP with GTP, allowing the Ct subunit (with GTP) to dissociate and bind an inhibitory subunit of cGMP phosphodiesterase and thus activating this enzyme to convert cGMP to 5'-GMP. The rapid drop in cGMP levels causes the cGMP-gated ion channels to close, blocking reentry of Na+, which hyperpolarizes the cell. A typical signaling cascade amplifies the signal at different steps (see Box 9.2).

Transducers

229

BOX 9.2. Efficiency of light-induced signal transduction

Light -

Rhodopsin is an example of a protein that is highly enriched in the tissue from which it is extracted. In the mammalian eye, each rod's outer segment contains 1000 to 2000 stacked discs that originated from the plasma membrane. Greater than 90% of the protein in the disc membranes is rhodopsin. The mouse retina, which has ~5 million discs, contains ~80,000 molecules of rhodopsin per disc. A single photon is sufficient to activate rhodopsin. Once activated, phosphorylation can prolong the excited state. Each excited rhodopsin can activate >500 molecules of transducin, each of which can activate a molecule of the cGMP phosphodiesterase by binding its inhibitory subunit. Now activated, phosphodiesterase can hydrolyze >4000 cGMP molecules per second. Three cGMPs are required to open one ion channel. so a small change in the concentration of cGMP makes a large change in ion conductance. Thus each photon absorbed by rhodopsin can close 1000 or more channels and change the potential by about 1 mY.

/Rod

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To optic nerve Ganglion Interconnecting neurons neurons 9.26. Rod cells and cone cells in the retina of the eye. In vertebrates, the lens of the eye focuses light on the photosensory neurons of the retina, which are the rod cells and cone cells that respond to the light by sending signals to the ganglion neurons and from there to the brain via the optic nerve. Redrawn from Nelson, D. L., and M. M. Cox, lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 456. © 2005 by W. H. Freeman and Company. Used with permission.

Light

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.5 9.27. Hyperpolarization of rod cells in response to light. In the absence of light, the cGMP-dependent ion channels in the rod's outer segment are open, decreasing the gradient of Na+ that is being pumped out by the Na+K+-ATPase. When light absorption by rhodopsin triggers degradation of cGMP, the channels close and the cell becomes hyperpolarized. Redrawn from Nelson, D. L., and M. M. Cox, lehninger Principles of Biochemistry, 4th ed., w. H. Freeman, 2005, p. 457. © 2005 by W. H. Freeman and Company. Used with permission.

Membrane Enzymes and Transducers

230

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9.28. The phototransduction cycle in the rod cell. When rhodopsin (Rh) is excited by absorption of a photon of light (1). it catalyzes the exchange of GTP for GDP on the (X subunit of transducin (T",) (2). allowing it to dissociate from transducin 0 and y (T ~y) and bind the inhibitory subunit (I) (3). Removing I turns on cGMP phosphodiesterase (PDE) so that it hydrolyzes cGMP (4), and the ion channels in the plasma membrane close (5). Ca 2+ efflux through the Na+ -Ca 2 + exchanger reduces cytoplasmic [Ca2+) (6). activating guanylyl cyclase (GCl to replenish cGMP (7), which enables channels to reopen. Adaptation occurs, slowing the hydrolysis of GTP on T<x, when "bleached" rhodopsin binds recoverin (Recov) (8) and is phosphorylated by rhodopsin kinase (RK) (9). Binding of arrestin (Arr) to phosphorylated rhodopsin prevents further interaction between rhodopsin and T <x. Dissociation of arrestin allows rhodopsin to be dephosphorylated and to replace the all-trans retinal with 11-cis retinal for a new cycle (10). Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles or Biochemistry, 4th ed., W. H. Freeman, 2005, p. 459.:g 2005 by W. H. Freeman and Company. Used with permission.

If the light is turned off, the rod cell recovers quickly due to the GTPase activity of the transducin ex subunit, which allows GDP-ex to rebind the transducin l3y complex. Release of the inhibitol)' subunit allows it to rebind the cGMP phosphodiesterase and block the hydrolysis of cGMP (see Figure 9.28). Guanylyl cyclase replenishes the level of cGMP, enabling the ion channel to reopen. Prolonged exposure to Ijght triggers adaptation through the phosphorylation of rhodopsin by rhodopsin kinase. Phosphorylated rhodopsin is bound by a protein called an-estin to prevent its interaction with transducin. As an-estin slowly dissociates, rhodopsin is dephosphorylated. Hydrolysis of the Schiff base aJlows the opsin to lose its all-trans retinal and bind a new II-cis retinal and initiate the cycle again. Rhodopsin Structure In addition to its serpentine fold, a number of other features of the rhodopsin structure -were known before the x-ray structure was solved. Several residues of rhodopsin are subject to posttranslational modifications, including N-acetylation of MetI, N-glycosyJation of Asn2 and Asn 15, palmitoylation of Cys322 and Cys323, and the Schiff base linkage of II-cis retinal to

Lys296 (as numbered in the bovine sequence; see Figure 9.25). In addition, a disulfide forms between CysiiO and Cysl87 that stabilizes the interaction between helix 3 and the second extracellular loop. Residues from two TM helices, Glu122 and His211, coordinate a Zn 2+ ion, whose physiological significance is unknown. In the three-dimensional Sl!-ucture, most of the 74 highly conserved residues are close to the retinal in the center of the TM domain, although several are found in the cytoplasmic domain and t\,vo in the N-tcrminal domain (Figure 9.29). There are two highly consen/ed motifs in the primary structures of over 100 opsins (shaded in Figure 9.25): the motif (D/E)R(Y/W) corresponds to Glu I 34-ArgI35-Tyrl 36 in the cytoplasmic end of TM helix 3, and NPXXY corresponds to the sequence NPVIY that starts with Asn302 near the cytoplasmic end of the seventh TM helix, a region thought to be involved in G-protein coupling (see below). Two crystal structures have been solved for the inactive state of bovine rhodopsin, at 2.8 Aresolution, later impl"Oved to 2.2 A, and at 2.65 Al-esolution, and they are in good agreement except for the cytoplasmic domain discussed below (see Figure 8.28). The structure could be positioned in the membrane by docking it onto a

Transducers

231

c

BOX 9.3. Numbering TM helices Several conventions are used to denote specific helices of integral membrane proteins. For the well-known helical bundles of Chapter 5, there is consensus on the designations for the helices: in bacteriorhodopsin, the helices are given letters A to G, while in the photosynthetic reaction center they are given letters to indicate the subunit, followed by numbers; for example, L1 is the first TM helix of the Light subunit. For some of the examples of helical proteins that follow, different authors use different conventions. Most formally, TM helices are given Roman numerals, while other helices are given Arabic numbers, numbered sequentially according to the primary structure. This convention results in naming the helices of rhodopsin as helices I through VII and helix 8. Frequently, the helices are just numbered sequentially with Arabic numbers, often with "H" to distinguish them from loops, which are numbered after "L". Both Arabic and Roman numerals are found in the literature on rhodopsin, lactose permease, and others. For helices in hetero-oligomers, a letter to designate the subunit typically precedes the number. These conventions make comparison of homologous structures straightforward.

9.29. Conserved residues among over 100 opsins, mapped onto the structure of bovine rhodopsin The 74 conserved residues cluster around the retinal (shown in red), with their color indicating successive levels of contact, defined in the shell on the right. From Teller, D. C, et ai, FEBS Lett. 2003, 555: 151-159. ~. 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

cryo-EM map of two-dimensional crystals. It resolves the retinal embedded near the center, the acyl chains near the C terminal that go into the plane of the bilayer, and portions of the exo-cytoplasmic oligosaccharide chains near the N terminal (Figure 9.30). In addition to the seven TM a-helices, numbered HI to H7 (or HI to HVII; see Box 9.3), there is an eighth a-helix lying in B

H4

H8

9.30. High-resolution structure of bovine rhodopsin. A. Ribbon diagram of rhodopsin showing two views from the side, rotated 180'-. The eight helices are numbered (different colors), with retinal (purple) near the center and the f)-hairpins below it. The resolved portions of the glycosyl chains are shown at the bottom, and the acyl chains near the amphipathic helix (H8) are clearly entering the TM region. B. The view from the exo-cytoplasmic (intradiscal) side (same coloring scheme) shows how the 13hairpins plug the access to the retinal C. The view from the cytoplasm shows this end has greater surface area than the intradiscal end. From Palczewski, K, Annu Rev Biochem. 2006, 75:743-767. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

232

the interfacial region on the cytoplasmic side. Four f)-strands at the exo-cytoplasmic end make two hairpins W1-f)2 and f)3-f)4) that "plug" the chromophorebinding pocket, as seen in the structure viewed from the exterior (Figure 9.30B). The cytoplasmic end of the molecule has a greater surface area than the intradiscal (exo-cytoplasmic) end (Figure 9.30C). The seven TM helices vary in length from 20 to 33 residues and do not line up normal to the bilayer; rather, their arrangement is varied by tilt angles and bends in the helices. Kinks in HI, H4, H5, H6, and H7 are made by proline residues, which are highly conserved in rhodopsin-like GPCRs. H2 bends at a pair of glycine residues, Gly89-Gly90, enabling the side chain ofThr92 to hydrogen bond to the main chain at residue 88. H3 is bent at Glu113, which is the counter-ion for the protonated Schiff base to the retinal. Two proline residues near the retinal-binding site in H7 (Pro291 and Pro303) serve to elongate the helix. In short, the kinks in the helices of rhodopsin help enable them to pack more closely and to make the pocket tha t binds the retinal. The helices are linked by hydrogen-bonding networks involving some very highly conserved residues and several key water molecules. One network extends from the kink in HI at Asn55 to H2 to H3 to H6 to H7 in the crystal structure and involves some residues shown by mutation to be important in the activation of rhodopsin (Figure 9.31). A hydrogen bond from Ser298 to the main chain 0 at residue 295 stabilizes the begin-

Membrane Enzymes and Transducers

9.32. Interface between the exo-cytoplasmic and TM domains of rhodopsin. The ends of the helices and the (I-strands are labeled, and loops are colored to distinguish between TM segments (yellow) and loops at the interface (blue). Hydrogen bonds between the two (I-hairpins are mediated by waters 3, 5, and 13. Critical residue Pr023 (see text) is between stacked residues Pr012Pr0285 on one side and Pr027-Tyr102 on the other. From Li, J., et aI., J Mol BioI. 2004, 343:1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

9.31. Interactions between TM helices in rhodopsin. The view

ning of a 3\0 helix' (residues 296-299). The conformational changes that occur aher absorption of a photon can be expected to crea te different H-bonding networks. Van del' Waals contacts are also important, as seen in a highly conserved group of hydrophobic residues, L76, L79, Ll28, M257, Ll31, and M253. Genetic experiments show that a change of Met257 to any residue but Leu gives constitutively active rhodopsin. A close look at the interface between the exocytoplasmic and TM domains shows waters mediating hydrogen bonds between the two f) hairpins with hydrogen bonds connecting them (FigUl-e 9.32). Interaction between these regions is also stabilized by hydrophobic stacking of Pro12 with Pro285 and of Pro27 with Tyr107. Right between them is a critical residue, Pro23. P23H is the most Frequent mutation in North American patients with retinitis pigmentosa and results in misfolcled rhodopsin that is degraded by the ubiquitinproteosome pathway, impairing that pathway and leading to neurodegeneration. The high-resolution structure gives good defini tion to the environment of the retinal in the inactive protein (Figure 9.33). It binds strongly in a pocket formed by the kinks in H3 and H5 and stabilized by hydrogen bonds. The protonated Schiff base to Lys296 in helix 7 is near its counter-ion, Glu 113 in helix 3. The ionone ring of

of a TM slice between the retinal-binding pocket and the cytoplasmic surface shows both a hydrogen-bonded network and a cluster of hydrophobic residues linking helices 1,2,3, 6, and 7. From Li, J .. et al., J Mol BioI. 2004, 3431409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

A 3'0 helix has three residues per tLirn and 10 atoms in the ring enclosed by hydrogen bonds. Compared with the a-helix with 3.6 amino acids per turn and 13 atoms in the ring. the 3'0 helix is thinner and moo'e elongated.

Transducers

233

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9.33. The environment of 11-ci5 retinal in bovine rhodopsin. A. The amino acid residues in the vicinity of the 11-c;5 retinal (pink) include Lys296 on H7, which forms the Schiff base linkage to retinal, and Glu 113 on H3, its counter-ion that is H-bonded to the peptide N of Cys187 in f)4 near the plug to the extracellular surface. From Palczewski, K., Annu Rev Biochem. 2006, 75:743-767. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org. B. The retinal binding pocket is defined by the electron density map in bovine rhodopsin. C. A sketch of the residues that surround the retinal indicates ligand bonds, hydrogen bonds, and polar and hydrophobic contacts. From Li, J., et ai, j Mol BioI.; 2004, 343: 1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

234

C3 Loop 9.34. The cytoplasmic domain that interacts with transducin. The cytoplasmic region consists of the four segments that project above the lipid phosphate groups: three cytoplasmic loops, C1, C2, and C3; the peripheral helix H8 (purple); and the (truncated) C-terminal tail. Side chains of individual residues that interact with transducin are shown, including those that mutate to give constitutive activation (green). those that have been cross-linked to the transducin ex subunit (Gtex; red). those that inhibit stabilization of Metall by Gtex (pink). those that reduce activation (yellow), and those that regain function upon reverse mutation from Ala (greenish yellow). The footprint of retinal (light pink) from the TM region of rhodopsin illustrates its alignment with key residues at the interface. Interhelical cross-links that inhibit activation are indicated by chains from H6 to nearby helices. From Li, J, et aI., J Mol BioI. 2004, 343: 1409-1438. © 2004 by Elsevier. Reprinted with permission from Elsevier.

retinal fits at the second kink in helix 2, bordered by hydrophobic groups, except for Glu 122. Some of the polar residues in the binding pocket are those that vary in the color pigments, alJowing the pigments to absorb light of different wavelengths. After its isomerization is triggered by absorption of a photon, retinal no longer fits the pocket, and conformational changes occur in the active intermediate that enable it to leave by hydrolysis of the Schiff base. These structuml changes are the subject of much current investigation employing various NMR, EPR, and mutagenic approaches since there is no high-resolution structure of the active intermediate called MetaU. Mutagenesis to introduce Cys residues For disulfide formation and spin-labeling studies has allowed comparisons between distances across the activated and ground-state structures. The results suggest that activation results in an expansion and opening of the cytoplasmic end, which could open a crevice For Gprotein binding. This activation mechanism is thought to be a general Feature of GPCR activation. As the portion of rhodopsin that interacts with transducin, the cytoplasmic domain receives much interest. This region includes the three cytoplasmic loops and adjoining ends of the TM helices, along with the peripheral helix and the C-terminal tail (Figure 9.34). It contains many of the residues that are highly conserved among different GPCRs, including the (D/E)R(YIW) and NPXXY motifs described above, and

is the site of many mutations that affect rhodopsi n activity. It is also the site of the main differences between the two crystal structures of bovine rhodopsin, and its loops have the highest crystallographic B factol~ suggesting it is the most flexible portion of the structure (see Figure 8.28 and Frontispiece.) A Fairly new question about the interaction of rhodopsin with transducin is the oligomerization state of rhodopsin in native membranes. Long considered a monomel~ rhodopsin has been shown by SDS-PAGE, EM, and AFM to be dimeric. The dimer contact is thought to involve helices 4 and 5 (Figure 9.35). The data suggest higher oligomers are likely in disc membranes and these may form and dissociate dynamicaJJy and may have important functional roles. If rhodopsin dimers bind transducin, rhodopsin is an even better prototype F01- a general signal transduction mechanism since many other GPCRs appear to function as dimers. Genomic and proteomic studies of the GPCRs indicate they have strong evolutionary relationships in spite of their diverse functions. Since they respond to different ligands or stimulants, the portion of the molecules that diverges the most is the ligand-binding domain. This domain is usually on the extracellula.- surface, unlike the buried [-etinal of rhodopsin. The binding sites of many GPCRs have been characterized by Cys scanning to probe accessibility of different residues to sulfhydryl reagents in the absence and presence of lig· ands. With many GPCRs, the binding site is the most studied domain, as it is valuable for designing new

Rho'"

Rho

9.35. Model of a rhodopsin dimer. The dimer is viewed from the cytoplasm and includes an inactive rhodopsin subunit (pink) and an activated subunit (yellow, marked with an asterisk). (Acidic residues are red and basic residues are blue.) The model has the constraints from the x-ray structure fitted to the AFM data from native membranes. Cross-linking data were applied to model the activated form. From Palczewski, K., Annu Rev Biochem. 2006, 75:743-767 11;) 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Transducers

9.36. Model of the structure of another GPCR, the atl angiotensin receptor. based on information from the rhodopsin structure. The functional regions are shown in space-filling models, with ligand binding (red), signal propagation (green), and G-protein binding (blue) from the crystal structure of rhodopsin. The positions of these regions are based on locations of important residues in the sequence of the atl receptor. From Filipek, S, et al., Annu Rev Biophys Biomol Struct. 2003, 32:375-397 © 2003 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular Structure. www.annualreviews.org.

drugs that interFere with binding. Since details about the TM domain are lacking, the structure of other GPCRs are modeled based on the crystal structure of rhodopsin, as exemplified in a model [or the angiotensin receptor (Figure 9.36). Thus. as the first high-resolution structure of a GPCR, the structure of rhodopsin is a highly significant prototype.

235

Bacteria maintain a slight outward turgor pressure as they respond to varying osmotic conditions. When osmotic pressure increases, the cells accumulate solutes such as betaine, proline, and potassium ions to balance the pressure and minimize water efflux. When osmotic pressure drops, they avoid rupture by opening MS channels For solute efflux. MS channels were discovered in 1987 in patch clamp experiments on giant bacterial spheroplasts (see Chapter 3). E. coli has three types of nonspecific MS channels, named for their different single-channel properties: MscL (large), MscS (small), and MscM (mini; Figure 9.38). The smallest channels, MscM, open at lo'vv tensions, are not essential, and are poorly characterized. Two species of the small MS channels that open next have been identified, MscK, which opens a K+ channel, and MscS, which is anion specific. At high tension, near that which would rupture the cell, the large nonselective channel MscL opens. Double mutants thal lack both MscL and MscS do nOl survive osmotic downs hocks. Furthermore, introducing MscL from E. coli into marine bacteria gives them increased resistance to drops in osmotic pressure. MscL MscL, the large MS channel of the E. coli inner membrane, Forms a nonselective ion channel that is activated by quite high levels of membrane tension. The MscL pore is large, giving a conductance of 2.5 nS in vitro. AFter crystallization attempts with a dozen MscL homologs from different bacteria, the MscL Crom Mycobacterium tuberculosis was crystal Iized fTom dodecylmaltoside and its structure solved at 3.5 A resolution. This protein, called Tb-MscL, has 37% sequence identity with the E. coli MscL, Eco-MscL, and its structure is consistent with cross-linking and EM data on Eco-MscL. Tb-MscL is a homopentamer consisting of two domains, a TM domain and a cytoplasmic domain

Mechanosensitive Ion Channels

Mechanosensitive (MS) channels respond to mechanical stresses applied to the membrane or to membraneattached elements of the cytoskeleton, enabling organisms to respond to touch, sound, pressure, and gravity. They fall into two broad classes, depending on whether or not cytoskeletal elements are involved (Figure 9.37). MS transduction in vertebrate auditory hair cells provicks examples of fast and sensitive responses involving the cytoskeleton. The prokaryotic MS channels that respond to deCl-eased osmotic pressure exemplify the class that does not involve the cytoskeleton. Crystal structures of two of the prokaryotic MS channels illustrate the best understood models for how membrane tension can drive conformational change and lead to channel opening.

,-;:

.... .,....,.---

,

I

I

I

L--..+-

I \ ~'lo._

,.------'\ (

\ \.~

__

~ ~

__

.J

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r---

, \

9.37. Two classes of MS channels. In many vertebrate cells. the channel has a swinging gate that is unlocked by tension from the cytoskeleton (left). In contrast, some channels respond directly to pressures in the bilayer and can be envisioned as an expandable barrel (right). Redrawn from Sukharev, S., and A. Anishkin, Trends Neurosci. 2004, 27:345-351. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

236

Membrane breakdown

A.

in

MscL

/

~

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ftL

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0'

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I I

o

I

o

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11111

nn 111111111111

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Tension

o

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9.38. Three types of MS channels are detected in E. coli. The channels listed on the left give different single-channel conductance characteristics (center) and open at different threshold pressures (right). From Perazo, E., and D. C. Rees, Curr Opin Struct Bioi. 2003, 13432-442. © 2003 by Elsevier. Reprinted with permission from Elsevier.

(Figure 9.39). Each monomer has two TM helices that tilt about 30° from the bilayer normal and a cytoplasmic helix that tilts about 1SO, making two bundles with fivefold symmetry. The first TM helix, TM1, lines the pore and is quite buried, as it is in contact with two TM 1 helices from other subunits, TM2 from the same subunit, and another TM2 [yom another subunit. The TM2 helices from neighboring subunits are separated by ~20 A. The peri plasmic loops between TM 1 and TM2 form a flap that makes the extracellular surface, and part of

each loop folds into the pore. Overall the TM domain has a funnel shape with the typical location of aromatic residues at the polar/nonpolar interfaces. The TM I helix that forms the pore of the channel is one of the most highly conserved regions in the sequences of MscL proteins. One conserved motif in this helix is a GxxxG motif for close helix packing and consists of AoxGXxxGAAxG at residues 20 to 30 (where X is a residue at the pore constriction). Several mutations in TM1 alter the channel gating (see below). In

A.

B.

c

9.39. X-ray structure of MscL from Mycobacterium tuberculosis. A. The side view shows the separate cytoplasmic and TM domains, with shading to indicate the membrane. B. Each monomer contributes two TM helices to the TM domain, as seen in the view of the TM domain from the top. In the homopentamer the pore is lined with five TM1 helices. From Perozo, E., and D. C. Rees, Curr Opin Struct BioI. 2003, 13:432-442. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Transducers

237

is important to channel function. The cytoplasmic domain is a compact left-handed five-helix bundle that might act to filter the solutes permeating an open channel upon osmotic downs hock. The C termini beyond the bundle are disordered in the crystal and are of varying length in the different MscL homologs.

9.40. Pore structure in Tb-MscL. In the MscL homopentamer, the pore lining has many polar and charged residues from TM 1. A cutaway side view of the molecular surface of the pore shows areas with charged residues (blue, basic; red, acidic). The green arrows show where the channel is closed. From Chang, G., et al., Science. 1998, 282:2220-2226. © 1998. Reprinted with permission from AAAS.

the structure, the pore of Tb-MscL is lined with a series of hydrophilic residues and is only 2 A at its most constricted point; hence this is considered the structure of the closed channel (Figure 9.40). Genetic experiments indicate the cluster of charged groups at the cytoplasmic end of the MscL pentamer

MscS The family of small MS channels found in some yeasts and plants as well as bacteria and archaea is highly diverse: it includes proteins that vary from 286 residues (in E. coli) to over a thousand residues in length. /n vitro the small MS channel from E. coli shows a slight preference for anions and gives a conductance of ~ LnS, which is still several orders of magnitude larger than that of voltage-gated K+ channels (see Chapter LO). MscS is gated by vol tage as well as pressure, as it requires less tension to open as the membrane is depolarized. The 3.9 A-resolution structure of E. coli MscS also reveals a TM domain and a cytoplasmic domain, but is otherwise very distinct from the structure of MscL. MscS is a homoheptamer, with each subunit containing three TM helices and a much more extensive cytoplasmic region (Figure 9.41). Each subunit corkscrews around the pore axis. In the membrane-spanning region, TM Land TM2 pack closely, while TM3, which lines the pore, has a kink that positions L4 residues parallel to the bilayer, making it similar to the paddle of the K VAP K+ channel (see Figure 10.38). The cytoplasmic domain has a large

Membrane domain

}

}

~ domaio

o~

domain

} l3-barrel 9.41. X-ray structure of MscS from E. coli. A. The fold of a single MscS monomer shows the three TM helices (labeled) and the mixed odf3 structures of the cytoplasmic domain. The N terminus is at the top and the C terminus at the bottom of the peptide, as drawn. Two arginine residues are shown in the TM region as space-filling models. B. The MscS heptamer viewed from the side, with each subunit in a different color. C. Space-filling representation of the MscS heptamer. With each subunit a different color, this view emphasizes how they corkscrew around the central axis. In addition, the cytoplasmic domain is divided into a f3 domain near the membrane, an wf3 domain, and a f3-barrel at the end. A and B from Bass, R. B., et al., FEBS Lett. 2003, 555:111-115. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier. C from Edwards, M. D., et al., Curr Opin Microbio/. 2004, 7: 163-167. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Membrane Enzymes and Transducers

238

A.

B.

lipid headgroups, which suggests that sensor regions are located at the interfacial regions of the membrane, where the lateral pressure is higher than it is in the centel~

9.42. Structural determinants in gain-of-function mutants in MscL (A) and MscS (B). Mutants were produced by random mutagenesis and screening of growth rates, potassium ion retention, or overall survival. Affected residues are thought to playa critical role in permeation and/or gating. From Peraza, E., and D. CRees, Curr Opin Struct Bioi 2003, 13:432-442. © 2003 by Elsevier. Reprinted with permission from Elsevier.

interior chamber formed by a ~ subdomain and an (X~ subdomain. This large vestibule (40 A in diameter) has seven side openings of J 4 A diameter and ends at the C termini with a seven-stranded ~-barrel with an opening of 8 Adiameter, \.vhich may pl-efilter the solutes that reach the open channel. In the TM domain of MscS the seven TM3 helices are tightly packed to form a pore of ~ 11 A diameter, which suggests that this is the open conformation of the channel. The pore is lined with hydrophobic residues, with the narrowest par1. at Leu j 05 and Leu) 09 near the cytoplasmic surface. TM3 has a conserved sequence starting at Ala97 in E. coli MscS and consisting ofAxxGAAGXAxGXAxyG (where x is a hydrophobic residue, X is a hydrophobic residue at the pore constriction, and y is a hydrophilic residue). The interhelical A-G pairs that result not only allow close packing in the crystal form but would also enable helix rotations to reach another conformation, as discussed below. The loop between TM2 and TM3 also extends into the channel and contains polar residues that make the pore more favorable for watet: Mutations that introduce charges close to the pore do not alter its ionic selectivity, so its slight preference for anions can be attributed to rings of positive charge within the vestibule. MS Channel Gating Portions of the MS channels involved in opening and dosing the pores have been identified by the characterization of mutants that affect channel gating. Loss-offunction (LOF) mu tants either require a higher pressure threshold to open the channel or cannot open it at all, in which case the bacteria do not survive osmotic downshock. Gain-of-function (GOF) mutants have a lower threshold for opening or stay open, generally resulting in slow growth rates. When mapped onto the crystal structures of MscL and MscS, the residues affected by GOF mutations cluster in the permeation pathway and on the outer helices (Figure 9.42). LOF mutations affect loops and regions of the protein that interact with

The direct effect of later'al pressure on MS proteins has been tested with a combination of conductance and EPR techniques that investigated how MS proteins respond to changes in their lipid environment. Spin probes incorporated into TM 1 of MscL give information on its subunit interactions as well as its mobility. The conductance of the spin-labeled channels reconstituted into vesicles of varying lipid compositions was measured in patch clamp experiments (see Chaptet- 3). The results indicate that varying the acyl chain length from J 0 to 20 carbon a toms is not su fficien t to open the MscL channel, although the threshold for activation is lower in vesicles made with short-chain phospholipids. However, introduction of lysophosphatidylcholine into the external leaflet to modify the lateral pressure tr'iggel'S channel opening. A comparison of the crystal structures of MscS and MscL gives a model for channel opening, since they represent the open and closed channels, respectively. When viewed from the top, their TM domains look very different (Figure 9.43). Although the TM helices of MscL are more intimately associated, both MscS and MscL have a central set of helices flanked by sets of outer helices that are less tightly packed. The outer helices are likely to be the sensors when the channels respond to tension in the bilayer. When the membrane is stretched, the MS channels will experience a pulling force, which concentrates at the interfacial regions according to analyses of the lateral pressure profile of the lipid bilayer. For the closed MscL structure, the increased tension at the inner and outer rims of the barrel could tilt TM helices to allow the channel to expand like tbe opening of the iris of a camera. This would involve TM 1 helices moving

A.

B.

9.43. TM helices from MscL and MscS. The TM regions of the MS channels are viewed from the top. The 10 helices from MscL (A) make an interlocking pattern, while of the 21 helices from MscS (B), only the pore helices are connected and the others splay out from the center. From Peraza, E., and D. CRees, Curr Opin Struct Bioi. 2003, 13:432-442. © 2003 by Elsevier. Reprinted with permission from Elsevier.

For Further Reading

239

Open (TM tilted)

Closed

Thus the challenges that make studying these membrane proteins difficult also provide a wealth of information in the ou tcomes.

FOR FURTHER READING

9.44. One model for channel opening in MscL. The MscL chan· nel is expected to open by a cooperative tilting of all TM helices by ~40°, limited by the periplasmic loops on the peri plasmic end and held by the helix bundle on the cytoplasmic side of the membrane. From Perozo, E., and D. C. Rees, Curr Opin Struct Bioi. 2003, 13432-442. (9 2003 by Elsevier. Reprinted with permission from Elsevier. away from the center and sliding over adjacent helices to unlock the pattern. All the TM helices would tilt by an additional 40' in a cooperative manner, producing a channel opening of ~ I 00;\2 (Figure 9.44). This model is supported by conductance studies that predict the open channel to have a diametel- of 35 A. It is also consistent with data indicating that TM I is exposed to water \-vhile TM2 remains in an apolar environment, as well as with properties of mutants anel cysteine cross-I inki ng studies. Finally, since mutagenesis and proteolysis indicate the normal gating response requires the periplasmic loops between the TM helices, the structure can be viewed as pairs of stiff helices connected by springlike loops that nllow the barrel to expand or contract in response to pressure. The helix tilt mechanism of MscL is only one mechanism for gating of channels formed by ex-helices. The nicotinic acetylcholine receptor, a ligand-gated ion channel, opens by rotation of the channel-lining helices to displace bulky residues from the pore (see Chapter 6). Finally, some potassium channels open by bending the inner helices at specific hinge points (see Chapter 10). Given the wide diversity among MS channels, it is possible that they utilize different mechanisms to open and close in response to membrane tension.

OMPLA 'Snijder, H. J., et aI., Structural evidence for dimerizationregulated activation of an integral membrane phospholipase. Nature. 1999,401:717-721. Snijder, H. J., and B. W. Kijkstra, Bacterial phospholipase A: structure and function of an integral membrane phospholipase. Biochim Biophys Acta. 2000, 1288:91-101 (review). Prostaglandin H2 Synthase Garavito, R. M., and A. M. Mulichak, The structure of mammalian cyclooxygenases. AI11'IU Rev Biophys Biomol Struct. 2003,32: 183-206.

Kulmacz, R. J., et aI., Comparison of the properties of prostaglandin H synthase-l and -2. Prog Lipid Res. 2003, 42:377-404.

'Loll, P. J., et al., The structural basis of aspirin activity inFerred From the crystal structure 01' inactivated prostaglandin H2 synthase. Nat Struct Bioi. 1995, 2:637643.

"Malkowski, M. G., et aI., The x-ray stnlcture 01' prostaglandin endoroperoxide H synthase 1 compJexed with arachidonic acid. Science. 2000, 289:1933-1937. Rouzer, C. A., and L. J. Marnell, Structural and functional differences between cyclooxygenases: [atty acid oxygenases with a critical role in cell signaling. Biochelll Biophys Res Comnnm. 2005, 338:34-44.

FORMATE DEHYDROGENASE

Jormakka M., B. Byrne, and S. Iwata, Formate dehydrogenase - a versatile enzyme in changing environments. CUlT Opin Struci Bioi. 2003, 13:418-423. Jorrnak.ka M., B. Byrne, and S. Iwata, Proton motive Force generation by a redox loop mechanism. FEBS Lelt. 2003, 545:25-30.

'Jormakka, M., S. Tornroth, B. Byrne, and S. Iwata, Molecular basis of proton Illotive force generation: structure of formate dehydrogenase-No Scie1'tce. 2002, 295:1863-1868. Rhodopsin

CONCLUSION

The examples of enzymes and transducers in this chapter show a wide variation in how proteins carry out catalysis and signal transduction in the membrane milieu. Each has a fascin8ting story that will be more completely revealed when higher-resolution structures and structures of mutants or of activated states become available. From the dimerization of OMPLA that enables it to trap its lipid substrate to the gating of MS channels in response to bilayer tension, the role of membrane lipids in protein function is clear.

Filipek, S., D. C. Teller, K. Palczewski, and R. Stenkamp, The crystallographic model of rhodopsin and its use in studies of other G protein-coupled receptors. AI1I1U Rev Biophys Biomo! Stmct. 2003, 32:375-397. 'Li, J., et aI., Structure 01' bovine rhodopsin in a trigonal crystal form. J /vIol Bioi. 2004, 343: 1409-1438. 'PaJczewski, K., et aI., Crystal structure of rhodopsin: a G protein-coupled receptOl~ Science. 2000,289:739-745. Palczewski, K., G protein-coupled receptor rhodopsin. Am'/U Rev Biochem. 2006, 75:743-767.

,. Paper presents original structure.

240

Schertler, G., Structure of rhodopsin and the meta rhodopsin [photointermediate. CUlT Opin Struct Bioi. 2005,15:408415.

Mechanosensitive Channels

"Bass, R. B., et aI., Crystal structure of Escherichia coli MscS, a voltage-modulated and mechanosensitive channel. Science. 2002, 298:1582-1587.

Membrane Enzymes and Transducers

'Chang, G., et al., Structure of the MscL homolog [Tom Mycobacterium tuberculosis: a gated mechanosensitive ion channel. Science. 1998, 282:2220-2226. Peerozo, E., and D. C. Rees, Structure and mechanism in prokaryotic mechanosensitive channels. Curl' Opil1 SlruC[ Bioi. 2003,13:432-442. Sukharev, S., and A. Anishkin, Mechanosensitive channels: what can we learn from "simple" model systems? Trends Neuyosci. 2004, 27:345-351.

10 Transporters and Channels

H,gh resolution structures can r1lustrate mcchanrs[l details, such as the Ion conduction pore of the KcsA potassium c laon ·1 exposed In " cutaway vIew 0 two of Its four subumts to reveal he pOSItions 0 Ih pore- orming heli (re ) and the s lectivi y filter (gold). The extracellular SIde is" he top Fronl MacKinnon, R., FEBS le t. 2003. 555 62 65. ,:' 2003 by the Fe eration of European Biochemical Societies Reprinted With permission from Elsevier.

For most of the past century, transporters and ion channels have been the object of many physiological, genetic, biochemical, biophysical, and bioinformatic investigations that have provided a wealth of data about how molecules and ions cross biological membranes (see Chapter 6). In recent years they have also become the subject of structural biology, as a few highresolution structures have added details from a rich vein of structure-function relationships. The award of the 2003 Nobel Prize to Peter Agre and Rod McKinnon for the x-ray structures of aquaporin (AQP) and the potassium channel brought wide attention to the progress in this field. This chapter presents examples that have varied histories, [Tom the long-studied lactose permease encoded by the lac Y gene in the lac operon to a relative newcomer, the water channel AQP.

TRANSPORTERS

Transport proteins are required for all cells to take up nutrients and to dispose of waste materials, as well as to mediate flux of metabolites between intracellular compartments of eukaryotes. In the cell envelope of Gramnegative bacteria, transport across the outer membrane is facilitated by porins that contain water-filled channels with varying specificity for solutes (see "Porins" in Chapter 5). In contrast, transport proteins of the inner membrane tend to be highly specific for their substrates, and consequently the cell has many diverse transporters in this membrane. Lactose permease and the glucose-3phosphate transporter of E. coli are exa m pIes of specific inner membrane transport proteins, both of whose xray crystal structures were reported in 2003. That same 241

Transporters and Channels

242

LacY and GlpT

Cytoplasm (inside)

A.

B.

10.1. Alternating access model of transport proposed for LacY (A) and GlpT (B). The proposed mechanism alternates between a conformation facing out (to the periplasm) and a conformation facing in (to the cytoplasm), with a rocker-type switch between them. Not shown is a possible intermediate state that is closed to both. From Abramson, J., et aI., Curr Opin Struct Bioi. 2004, 14:413-419. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Lactose peromease (LacY) and the glycerol-3-phosphate transporter (GlpT) of E. coli aloe members of the major facilitator superfamily (MFS) of secondary active transporters (see "Transport Proteins" in Chapter 6). As the largest of all transporter families, this group has 70 members identified in E. coli and over 3000 members identified in genomes of organisms from all three kingdoms of life. In spite of low-sequence homology, the proteins in the family appear to have a common fold, as predicted by hydrophobicity plots and supported by gene fusion data. For this reason, the high-resolution structures of LacY and GlpT provide information rdevant to a vast number of transporters. In addition, they provide clear support for a transport mechanism proposed over 50 years ago (Figure 10.1).

year the structure of the mitochondrial ADP/ATP carrier was solved, giving a first example from the mitochondrial carrier family (MCF). The calcium pump became the first P-type ATPase to have a high-resolution structure (described at the end of this chapter). Chapter 1 I will return to transport to examine structures of tr·ansporters that are part of multicomponent complexes, including the uptake system for vitamin B 12 , the bacterial drug efflux system, the mitochondrial F 1 Fa-ATPase, and the translocon involved in export of nascent proteins.

LacY The lactose permease, which is the galactoside/H+ symporter of E. coli simply called the LacY protein, utilizes the proton gradient of the inner membrane to drive the lOa-fold accumulation of lactose inside the cell. The lacY gene is part of the lac operon induced when lactose is in the medium. It was the first transporter gene to be cloned and sequenced, ushering in the use of molecular biology techniques to investigate membrane transport. Many of the features of the LacY protein subsequently determined by extensive genetic, biochemical, and biophysical analyses have been confirmed with the

Lactose H+

Lactose H+

G3P

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10.2. X-ray structure of LacY. The high-resolution structure of the LacY C154G mutant is shown in ribbon representations viewed from the membrane plane (A) and from the cytoplasm (B), with the 12 helices colored from dark blue at the N terminus to red at the C terminus. The sugar-binding site in the cavity is occupied by the inhibitor f3-D-galactopyranosyl 1-thio-f3-D-galactopyranoside (TDG), represented with black spheres. In B, the loops are removed to view the helices normal to the membrane, revealing their curves and kinks. Helices are labeled with Roman numerals (see Box 9.3). From Abramson, J., et aI., Science. 2003, 301 :61 0-615. © 2003. Reprinted with permission from AAAS.

Transporters

243

A,

B,

VII

V ........_" • ...,

10.3. Substrate-binding site of LacY with TDG present. Interactions between the galactosyl moiety and the N-terminal domain produce the binding specificity, while the interactions with the C-terminal domain are less specific and give room for more bulky adducts, (See text for details,) A. The residues involved in TDG binding, viewed along the membrane normal from the cytoplasmic side, B, A closeup view of the N-terminal domain interactions with TDG From Abramson, J" et ai" Science, 2003, 301 :61 0-615, © 2003. Reprinted with permission from AAAS,

x-ray structure, which took 10 years to achieve, The first structure vvas determined at 3,5 A resolution for a mutant of LacY (CI54G) that can bind substrate with high affinity but not transport it. This phenotype suggested the mutant protein is stabilized in one conformation, which facilitated its crystallization, Recently, crystals of wild-type LacY protein have been obtZlined thaI provide general confirmation to the overZiIl fold seen in the mutant structure described in detail here, A1l417 amino acids of LZicY have now been mutated to study its structure and function, which allows precise identi fication of critical residues in the crystal structure. As predicted by spectroscopic measurements, LacY is mainly ex-helical (86%), and 80% of the protein is in the membrane. Its twofold pseudo-symmetry, which suggests a gene duplication event occurred in its evolution, gives it a heart shape when vie\'ved from the plane of the membrane. The structure consists of two bundles of six ex-helices forming N-terminal and C-terminal domains that are linked by a long loop between helices 6 and 7 (Figure 10.2), The irregularity of the helices due to curves and kinks may be important in the conformational change involved in the mechanism of transport (see below). The two domains are separated by a large hydrophilic cavity that opens to the cytoplasm and contains a single sugar-binding site. LacY is highly specific for the galactose moiety of lactose: it will not bind or transport glucose or 0glucopyanosides, but it will transport galactose and many derivatives of galactose with hydrophobic groups on the anomeric carbon, including the familiar ones used as inducers and dyes, The substrate-binding site in the crystal structure is very well defined because in the crystal LacY is binding a substrate analog, [3-0galactopyranosyl l-thio-(3-o-galactopyranoside (TDG)

(Figure 10.3). The galactose moiety interacts with severZll residues or the N-terminal domain and only one from the C-terminal domain. The galactopyranosyl ring sits over the indole ring or Trp151 of helix 5 (expbining why substrate binding shifts its fluorescence mZlXimum), and this interZlction positions it to make specific hydrogen bonds, Another hydrophobic interaction involves the C6 atom of the galactose ring with Met23 in helix 1. An essential arginine, Arg144 of helix 5, makes Zl criticZlI bidentate hydrogen bond with the 03 and 04 atoms of the galactose ring, The irreplaceable residue Glul26 of helix 4 is positioned to interact with the galactose 04, OS, or 06 atoms via water molecules; biochemical data indicZlte it interacts with Arg144 in the absence of a bound substrate. Another essential residue, Glu269 in helix 7 appeal-s to Form a salt bridge with Argl44 close to Trpl51. As this residue is in the C-terminal domZlin and is also involved in proton translocation (see below), it may be a critical energetic link between the two domZlins. Other residues of helix 7 and helix 10 in the C-terminZlI domain that are near the substrate appear to have less specific interactions and probably provide the additional affinity LacY has for disaccharide and galactosyl adducts compared with galactose. The structure of LacY also gives information about proton translocation and coupling of proton and lactose upt8ke for symport. Interestingly, LacY catalyzes passive exchange of sugar (down its concentration gradient) without translocation of protons but does not translocale protons in the absence of a sugar substrate, suggesting that proton binding is the initial step, followed by substrate binding, which is supported by quantitative determinations of pH changes with substrate addition. The x-ray structure represents the protonated form, with the proton on Glu325 in a

Transporters and Channels

244

B.

A.

10.4. Residues of LacY involved in proton translocation and coupling. The labeled residues form a network of salt bridges and hydrogen bonds (black broken lines) as described in the text. A. View from the side. B. View from the cytoplasm. From Abramson, J., et al., Science. 2003, 301 :61 0-615. © 2003. Reprinted with permission from AAAS.

hydrophobic pocket made by residues from helices 7, 9, and 10 of the C-terminal domain (Figure 10.4). Remarkably, this residue is part of a salt bridge/hydrogen bond network that lies parallel to the plane of the membrane, instead of crossing it as in BR and other proton pumps. This strongly suggests that the proton is bound and released in the same way as the sugar substrate, via the conformational rearrangement that alternates their access from one side of the membrane to the other. The steps of coupled translocation can be summarized as follows. Initially LacY is open to the peri plasm (outward facing) and may be protonated spontaneously. Then substrate can bind. When binding of substrate disrupts the salt bridge between Arg144 and Glu126, the proton from His322 (from helix 10) will transfer to GJu325. Glu269 is recruited to form a salt bridge with Arg144, triggering the proposed conformational change that results in the inward-facing conformation observed in the crystal. While the conformational change can be

modeled for the helical segments (Figure 10.5), further understanding of the mechanism depends on achieving higher-resolution structures, complemented with dynamic studies combining genetic and biophysical approaches. GlpT E. coli uses the GlpT transporter to take up glycerol-3phosphate (G3P) for use as an energy source (via glycolysis) and as a precursor for phospholipids. The presence of G3P in the medium induces the glpT gene for uptake; when G3P is lacking, the glpT gene is l-epressed and G3P is made f\'Om glycerol by the action of glycerol kinase or from dihydroxyacetone phosphate by G3P dehydrogenase. GlpT is an antiporter that carries out the exchange of G3P for inorganic phosphate, which is generally at millimolar concentrations inside the cell, so its efflux is energetically favored. Because GlpT recognizes the phosphate moiety of G3P, it \-vill not transport

B.

A.

~ffi~m~

~oo~~

jMl1

mlllllll1 PeripJasm

(X-ray structure)

(Simulation)

10.5. Model for structural changes between the two conformations of LacY. A. The x-ray structure reveals the inward-facing conformation with the N- and C-terminal TM domains shown in blue and red, respectively. Residues colored yellow are those that have been replaced with cysteine and found to increase their reactivity to N-ethylmaleimide when substrate binds. B. Suggested model for outwardfacing conformation, obtained by applying a rotation of ~60° to the N- and C-terminal domains. From Abramson, J., et aI., Science. 2003, 301 :61 0-615. © 2003. Reprinted with permission from AAAS.

Transporters

245

Membrane

10.6. Topology of GlpT showing the amino acid sequence. The N- and C-terminal domains, each with six TM segments, have weak sequence homology to each other. The essential arginine residues (Arg45 and Arg269) are in bold. From Lemiuex, M. J., Y. Huang, and D.-N. Wang, Curr Opin Struct Bioi. 2004, 14:405-412. © 2004 by Elsevier. Reprinted with permission from Elsevier.

glycerol. It will transport glycerol-2-phosphate, as well as a phosphate-based antibiotic called fosfomycin. Fosfomycin has been used to select for mutations in glpT in laboratory strains and has been used clinically to treat bacterial uri nary tract infections. GlpT consists of 452 ami no acids, most of which are embedded in the membrane as it has very short connecting loops except between TMs 6 and 7, which connect the N- and C-terminal domains (Figure 10.6). Overall, the architecture of GlpT is a trapezoid made up of these two domains, which appear to be fairly rigid helical bundles, with the substrate-binding site between them. The crystaJJized form of GlpT has a wide opening at the cytoplasmic side and is constricted at the periplasmic side (Figure 10.7). Therefore, the GlpT structure, like the structure of LacY, has the inward-facing conformation. The overall surface along the translocation pore is neutral except for two arginine residues (seen in Figure 10.7C). There are extensive van der Waals interactions between the two domains where they come together at the peri plasmic side, while they are connected by the long cytoplasmic loop at the open side. The two sides have complementary geometries in that the N-terminal side has a numbel- of bulky side chains opposite pockets on the C-terminal side. The helices vary in length and tilt angles; they have many glycine residues, allowing them to pack closely. Most are straight, but the four at the interface between the Nand C domains are curved (Figure 10.8). This curvature makes it easy to envision a rocker-switch type of movement that would change the conformation to the outward-facing form. GlpT was crystallized with no substrate bound. Since it must bind both G3P and inorganic phosphate at the hydrophilic cavity between the two domains, the substrate-binding site is identified by the two essential

arginine residues, Arg45 from helix] and Arg269 from helix 7. The distance between the two arginine residues is a little too far for both guanidino groups to form optimal hydrogen bonds with a bound phosphate (Figure 10.7C). Thus it appears that binding phosphate or G3P pulls the arginine residues closer together, resulting in a more compact conformation and a narrower cytoplasmic pore. Evidence for this change comes from tryptophan fluorescence studies, as weJJ as measurement of the Stokes radius of GlpT in detergent. To close the pore on the cytoplasmic side requires a ] 0° to 15° rotation of each domain and results in exposing the substrate-bindi ng site to the periplasm. In other words, like LacY, the transport mechanism operates via a single binding site, alternating access with the rocker-switch type of movement of the N- and C-terminal domains (Figure 10.9). Within the major facilitator superfamily, GlpT is in the organophosphate: phosphate antiporter family, whose members do share significant sequence homology. In particular, there are conserved residues on both sides of the essential arginine residues in both the N- and C-terminal domains. The conserved residues include both hydrophilic and hydrophobic amino acids and may be involved in substrate binding in the two conformations (Figure 10.10). Interestingly, this family contains human transporters for G3P and for glucose-6phosphate that are implicated in a variety of conditions, including deafness, hyperglycerolemia, respiratory distress, and seizures. Comparison of LacY and GlpT The simultaneous publication of high-resolution structures for two MFS transporters naturally invited their detailed comparison. The overall folds of LacY and GlpT are nearly supetimposable, and the fit is even better when the separate N- and C-terminal domains are

Transporters and Channels

246

B.

10.7. X-ray structure of GlpT. A. A ribbon diagram of the two domains (green and purple) shows the trapezoid shape of GlpT and its orientation in the membrane. The directions for translocation of G3P and Pj are shown. B. The symmetry between the two domains of GlpT is highlighted when its TM segments are colored according to function. Four of the TM helices are not involved in pore formation (H3. H6, H9, and H12; green), four line the central pore (H2, H5, H8, and H11; yellow), and the remaining four are central in the structure (purple). The two arginine residues in the transport path are shown. C. At the substrate-binding site the distance between Arg45 (helix 1) to Arg269 (helix 7) is 9.9 A. From Lemiuex, M. J., Y. Huang, and D.-N. Wang, Curr Opin Struct Bioi. 2004, 14:405-412. © 2004 by Elsevier. Reprinted with permission from Elsevier.

A.

B.

c.

10.8. Curved helices in the structure of GlpT A. The ribbon diagram viewed from the cytoplasm shows the GlpT structure (colored as in Figure 10.7A) has many curved helices. The curves are evident in views of pairs of helices, H2 and H11 (B) and H5 and H8 (Cl, viewed from the periplasm (top). Viewed from within the plane of the membrane (bottom), each pair of helices has an hourglass shape, making contact in the membrane and opening toward the cytoplasm. From Lemiuex, M. J., Y. Huang, and D.-N. Wang, Curr Opin Struct Bioi. 2004, 14:405-412. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Transporters

247

B.

A. A.

Co-S

Co-Pi

B.

Co

C o-G3P Periplasm

t ....

'-'

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-0 CIl

t

+

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0.; Cytoplasm

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10.9. Rocker-switch mechanism proposed for GlpT. A. Model of conformational changes proposed to accompany substrate translocation. The protein alternates between an outward-facing conformation (Co) and an inward-facing conformation (C,). The substrate phosphate (red) is shown as it moves from the inside (C) to the outside (A). The crystal structure corresponds to the inward-facing conformation without substrate (D). The outward-facing conformation was generated by rotating the two halves of GlpT by 16° each in opposite directions (8), while the Co-Pi conformation (A) required a 10° rotation. B. Proposed mechanism for single binding site, alternating access with a rocker-switch type of movement. From Lemiuex, M. J., Y. Huang, and D.-N. Wang, Curr Opin Struct Bioi. 2004, 14:405-412. © 2004 by Elsevier. Reprinted with permission from Elsevier.

10.10. Conserved residues among members of the organophosphate: phosphate antiporter family, mapped onto the structure of GlpT. A. The conserved residues are shown in ball-and-stick representation on the ribbon diagram of GlpT. B. Surface representation of the N-terminal domain with conserved residues colored, viewed from the C-terminal domain within the membrane plane. C. Surface representation of the C-terminal domain with conserved residues colored, viewed from the N-terminal domain within the membrane plane. From Lemieux, M. J., Y. Huang, and D.-N. Wang, Res Microbio/. 2004, 155:623-629. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Transporters and Channels

248

B.

II IV . III

~ ,.\

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/1jJ ~,U

~~ VI

~~ I

V

XI

:?

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1

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10.11. Superimposition of structures of LacY and GlpT. A. Superimposition of the structures of LacY (yellow) and GlpT (blue and red, for the N- and C-terminal domains, respectively) viewed parallel to the membrane. B. Superimposition viewed along the membrane normal from the cytoplasmic side (same color scheme), with helices numbered and loops removed. From Abramson, J., et al., Curr Opin Struct BioI. 2004, 14:413-419. © 2004 by Elsevier. Reprinted with permission from Elsevier.

superimposed (Figure 10.11). A view of the helices from the top of the membrane shows close alignment of all except the helix 2/helix 7 pair along the hydrophilic cavity, which may be partly due to the fact that in the crystals, LacY has a substrate analog bound while GlpT does not. In addition the LacY core helices are more distorted by kinks, although helices in the GlpT structures bend at or near the same places. The connecting loop between the two domains also varies as LacY has two short helices on both the N- and C-terminal sides of the loop, while GlpT lacks a helical segment at the N terminus of the loop. The biggest differences are likely to reside in the substrate-binding sites, given the physical differences bet\veen lactose and G3P (compare Figure 10.3 with Figure 10.7C). The lactose-binding site of LacY is very well characterized (see above) since the crystal contained a substrate analog, whereas the binding site for G3P is inferred from the two arginine residues that bind to the phosphate. Interestingly, the site in LacY is in the center of the membrane, while the site in GlpT appears to be closer- to the peri plasm, shifted by approximately one turn of the helix. A Paradigm for MFS Transporters Prokaryotic MFS pmteins consist of 400 to 600 amino acids in a single polypeptide chain, with 12 TM helices that are likely to be organized in two domains, as seen in LacY and GlpT. These similarities suggest they share a common architecture and mechanism of transport, in spite of needing unique binding sites for their diverse substrates, The 6.5 A resolution cryo-EM structure of a third E. coli MFS transporter, the oxalate/formate anti porter OxlT, exhibits these featUl-es and is useful as a template for the positions of TM helices in the

outward-facing conformation since it is observed facing the periplasm. Overall the sequence homology among MFS members is weak. In fact, their only consel\led sequence is the motif RxxRR, which is found in connecting loops L2-3 and L8-9, HoweveI; many of these proteins transport sugars, and these do exhibit conservation at the sugar-binding sites. When bacterial MFS transporters for other sugars are compared with E. coli LacY, the sequence homology varies from 35% to 75%, 'which allows their sequences to be aligned with the LacY sequence by homology threading to generate models of str'uctures (Figure 10.12). Recently the sequence of a maltose permease from a deep sea bactel-ium, 'which has consenation of the most essential amino acid residues in LacY (17% identity), was employed in a BLAST search of pmkaryotic and eukaryotic genomes (see Chapter 6). Interestingly, the eukaryotic pmteins thus identified are predicted to have 12 TM segments and many of the essential residues at their substratebinding sites, even though in general they are about t"vice the size of the prokaryotic sugar transporters. The common architecture, and likely mechanism, of MFS transporters does not entirely reveal how these uniporter-s, symporters, and antiporters for widely varied substrates can)' out secondary active transport. Along with the detailed pictures of substrate-binding sites and transport paths starting to emerge, it will be important to learn what drives the conformational changes to couple the cotransport pmcesses. Deduction of further mechanistic details will require higher resolution in crystal structures, capture of other conformations by crystallography or' cryo-EM, and dynamic studies employing spectmscopic, biophysical, and computational approaches.

Transporters

249

The mitochondrial AAC was discovered over 40 years ago through investigations of the mechanism of poisoning by atractyloside (ATR), a diterpene heteroglucoside that is produced by a widespread Mediterranean thistle used in herbal medicine. ATR was very useful in early studies, especially because it binds to the AAC

Intermembrane space

*

Matrix + +

+

ATR

Adenine nucleotide ATp4 - ~!S:======-=lS:-_ lranslocase ADp3(anti porter)

H8:l-~~'::=~~=--.t--~""

ATP synthase

10.12. Models for MFS sugar transporters from homology threading. The predicted structures are low resolution, as accuracy depends on the extents of homology between the two proteins. All are viewed from the plane of the membrane with the cytoplasm on the top. TM helices are depicted as ribbons from helix 1 (blue) to helix 12 (red). Conserved residues that are shown include Gly residues on helices 1 and 5 (yellow spheres), Gly residues on the extracellular surface between the N- and Cterminal helical bundles (pink spheres), and aromatic residues facing the hydrophilic cavity (gray spheres). A. LacY. B. E. coli sucrose permease CscB. C. E. coli reaffinose permease RafB. D. Bacillus megaterium fructose permease FruP. From Abramson, J., et al. Science. 2003, 301:610-615. © 2003. Reprinted with permission from AAAS.

Phosphate translocase (symporter) + + + + CH,OH

-03S-0~0 -o)s-o

o=c I

CH,

I -

H)C

Mitochondrial ADP/ATP Carrier The extensive traffic of hydrophilic metabolites that occurs in eukaryotic cells requires specialized transporters in membranes of organeJJes. Like the Gramnegative bacteria from \vhich they evolved, mitochondria are surrounded by two membranes: an outer membrane with pores that allow fairly nonselective passage of molecules and ions up to around 500 Da, and an inner membrane with at least 20 specific transport functions. Prominent among the mitochondrial inner membrane transporters is the ADP/ATP carrier (AAC), whose job is to provide the ADP substrate needed inside the matrix for ATP synthase and to export the ATP it produces (Figure 10.13). The AAC is an electrogenic antiporter that exchanges one ADp3- for one ATp 4 - without bound Mg2+ ions, with the net export of one negative charge driven by the membrane potential. Indispensable to the generation of ATP by respiration, it is the most abundant of the mitochondrial carriers and makes up 10% of the protein extracted from the inner mitochondrial membrane.

0

R

CH /'. CH)

R = eOOH Carboxyatractyloside (CATR)

Bongkrekic acid (BA)

10.13. Schematic of transport process across the inner mitochondrial membrane. The mitochondrial ADP/ATP carrier (top) exchanges ATp4- produced inside the mitochondrial matrix for ADp3- from the outside via the intermembrane space. Transport is driven by the potential gradient across the membrane created by the ATP synthase (center), and inorganic phosphate is replenished by a symporter (bottom) The ADP/ATP carrier is inhibited from the matrix side by bongkrekic acid (BA) and from the intermembrane side by atractyloside (ATR), as indicated. Redrawn from Nelson, D. L., and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., W. H. Freeman, 2005, p. 714. ©2005 byW. H. Freeman and Company. Used with permission. Inset: Chemical structures of the AAC inhibitors BA and CATR.

Transporters and Channels

250

C2

Cl

C

Intermembrane space

H6 Matrix

h 3 .• 10.14. The topology of the mitochondrial AOP/ATP carriers. Schematic diagram of the secondary structure of the bovine AAC shows the six TM helices with their connecting loops, along with MCF motif residues (gray) and the RRRMMM signature (black). From Nury, H., et al., Annu Rev Biochem. 2006, 7S:713-741 © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

from the intermembrane side, while another inhibitOl~ bongkrekic acid (BA; a complex fatty acid derivative), was discovered that binds from the matrix side. (See inset of Figure 10.13 for their structures.) Characterization of the binding of these inhibitors and various derivatives of them demonstrated that the ADP/ATP carrier exists in two conformations. AAC requires cardiolipin for efficient transport.

membrane space (IMS) (Figure 10.14). The yeast carrier has been the object of studies employing site-directed mutagenesis to identify critical residues; it shares some of these residues, along with general features, with the bovine AAC that has been crystallized. Purified bovine AAC in detergent was bound to carboxyatractyloside (CATR) to stabilize it in one conformation for crystallization. In the crystals the carrier is packed into protein-lipid layers stacked into three dimensions, with endogenous lipids between protein molecules. Two crystal forms have been solved, one at 2.2 A resolution and a second allowing resolution of three cardiolipin molecules per protein. The crystal structure shows the 297 amino acids of bovine AAC form a bundle that is closed toward the matrix with a wide funnel-shaped opening toward the IMS (Figure 10.15). The backbone exhibi ts the threefold symmetry that is expected from the primary structure. Helices 1,3, and 5 are sharply kinked (from 20° to 35°) at proline residues, and all the helices are tilted. The three loops on the matrix side contain small amphipathic helices parallel to the membrane surface to form the bottom of a "basket." TM helices form a deep hydrophilic cavity into the basket, lined by the conserved arginine residues that give a very specific signature, RRRMMM, to all ACC sequences (Figure 10.16).

Overall Structure

Like other mitochondrial carriers (see below), the AACs from different species are organized into three homologous repeats of around 100 residues each. They form six TM helices with both N- and C-termini in the inter-

Binding in the

Me "Basket"

Located in the cavity of the basket is the bound CATR, where it fills the cavity and interacts with the carrier through electrostatic interactions, hydrogen bonds,

Intermembrane space A.

CDL802

c.

B.

CDL801

hi

2

CDL800 CDL801 Matrix 10.15. The overall structure of the bovine AOP/ATP carrier. The ribbon diagrams of AAC viewed from three perspectives are colored to reflect the primary structure (N terminus, blue, to C terminus, red). The TM helices (H1-H6l, cytoplasmic loops (C1 and C2l, and the short helices on the end facing the matrix (h 1-2, h3-4, and h s- 6 ) are labeled, along with three cardiolipins (COL, gray). A. View from the IMS; B. View from the membrane plane; and C. View from the matrix. From Nury, H., et al., Annu Rev Biochem. 2006, 7S:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

20

A

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A ~

..

J:' ~

!

... ~......

R279

30

lC022

R137

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~

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.

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.

e>......

R235 'f!/:>, .....

.

~

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C0221:

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C0220 10.16. Surface representation of a cross-section through the cavity in the AAC basket. The cavity is accessible to the IMS (top) and is 20 A deep. It contains three arginines of the AAC signature, one of which forms a salt bridge with E264 (yellow). From Nury, H., et al .. Annu Rev Biochem. 2006, 75:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

and van der Waals contacts involving many conserved residues (Figure 10.17). The cavity has several patches of basic residues located at positions ranging from the entrance Lo the bottom. It also has a ladder of tyrosine residues (Tyrl94, Tyr190, and Tyr186) entering the cavity along helix 4. At the third basic patch the cavity is constricted to 8 A. by Tyr186 and three basic residues (Lys22, Arg79, and Arg279). Many of the charged and polar residues are linked by extensive hydrogen-bond

-180 0

10.18. A large hydrogen bond network in the AAC at the binding site for the inhibitor CATR. Highly conserved polar, acidic, and basic residues, along with main-chain carbonyls (labeled CO) are hydrogen bonded, often via numerous water molecules. From Nury, H., et aI., Annu Rev Biochem. 2006, 75:713-741. «;; 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

networks (Figure 10.18). These networks link all the TM helices except helix 4, which deviates from the threefold symmetry and has the bulk of the buried aromatic residues. The inhibitor binds at or near the binding site for ADP, as ATR blocks uptake of ADP from the IMS, so the cavity can be analyzed as a nucleotide-binding site. The patches of positive charge \vill attract the anionic nucleotide to the entrance. The funnel-shaped opening

120

A211>

14

084

YRO

IMS 180 Top of the cavity

T --

, S212

0

0

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.

Y;'1904- _.... 197

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Bottom of the cavity 180

0

10.17. Two-dimensional projection of the residues present at the surface of the cavity in the AAC basket. Each circle represents an atom of a residue located within the cavity, with a size proportional to its solvent accessibility and a color indicative of the type of residue (basic, blue; acidic, red; aromatic, gray; hydrophobic, yellow; polar, green). The tyrosine ladder is marked by Y and four positive patches are circled and numbered. From Nury, H., et aI., Annu Rev Biochem. 2006, 75:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Transporters and Channels

252 A.

10.19. Conserved residues on external surfaces of Me. Sequence similarities within ADP/ATP carriers are mapped onto the structure of the bovine MC and colored to show extent of homology (0% to 100%, white to red). The highly homologous regions include residues lining the cavity and are well defined on both the end of MC that faces the IMS (A) and that facing the matrix (B). From Nury, H., et al., Annu Rev Biochem. 2006, 75:713741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Homology

IMS view

:.

Matrix view

of the cavity will orient an incoming nucleotide substrate with stacking interactions with the row of tyrosine residues along helix 4. Selectivity of the transport for adenosyl and not guanylyl nucleotides can be explained by specific interactions with conserved residues. The adenosyl amino groups and sugar hydroxyl groups will form hydrogen bonds with polar residues, while the phosphoryl groups of the nucleotides will interact with the basic residues. Hypothesized Transport Mechanism The exchange process catalyzed by AAC requires that it bind to and release substrates on opposite sides of the membrane. Binding ATP from the matrix is predicted to disrupt the salt bridge between Arg236 and Glu264 (shown in Figure 10.16), which will shi ft the extensive hydrogen-bond networks stabilizing this conformation. Thus binding of negatively charged substrates perturbs the salt bridges at the bottom of the cavity and shifts the network of polar interactions to trigger conformational changes in the carrier molecule. Interestingly, among AACs fyom different species the most conserved residues are inside the cavity and include several basic residues, along with aromatic residues to an unusual degree (Figure 10.19). It is clear from the high-resolution structure of the conformation that opens to the IMS that AAC must undergo large conformational changes during transport. Large movements of TM helices are needed to invert the "basket" when AAC opens to the matrix. Most likely the proline residues that form kinks in the odd-numbered helices are the hinges that permit these conformational changes (Figure 10.20). This may be a general mechanism for mitochondrial carriers, since these proline residues are part of a highly conserved sequence found in each of the three repeats of AAC and other members of the MCF. The MCF motif is PxD/ExxKlRxKlR-(20-30 residues)-D/EGxx.;xxaKlRG, where the letter x denotes any amino acid and the letter a denotes an aromatic residue. In the AAC structure this motif is found in the bottom of the cavity where matrix loops close off the basket. Specifically, in the third repeat

of the AAC. the MCF motif includes the Glu264-Arg236 salt bridge along with Arg271, which clamp together the matrix ends of helix 5 and helix 6 at the bottom of the basket. The role of the MCF motif in shaping the "basket" seen in the structure of AAC suggests a common architecture and mechanism for mitochondrial calTiers. Many mitochondrial carriers are proposed to hmction as dimers. and a variety of biochemical and biophysical data have indicated that AAC forms a dimeI'. Covalent dimers made by fusing two tandem copies of the yeast gene for AAC produced functional carriers. However, the crystal structures strongJy suggest that a monomer of AAC can perform its transport lntelmembrane

space

K32

Matrix 10.20. Proline-induced kink in helix 1 of the Me. This example shows the kink after Pr027 in helix 1, where the carbonyl of Thr23 hydrogen bonds with Trp70. which interacts with cardiolipin (green). Hydrogen bonds are indicated by gray lines. From Pebay-Peyroula, E., and G. Brandolin, CurrOpin Struct BioI. 2004, 14:420-425. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Channels

A.

N

253

N

B.

of the pore itself, in addition to special features like vestibules (both external and internal) that attract ions and selectivity filters that exclude other ions from the pores. Further, many channels have gating mechanisms that enable them to open in response to outside signals communicated via ligand binding, electric potentials, and/or pH. Channel gating involves the response mechanism itself and the conformational change it stimulates. All these characteristics are better understood thanks to high-resolution structures of several AQPs and potassium channels. Both families of channels can)' out passive transport: the AQPs transport water in response to an osmotic pressure gradient, while potassium channels are driven by the electrochemical K+ gradient created by the Na+K+-ATPase. The chapter ends with a description of the calcium pump, an ion channel that undergoes large conformational changes of its cytoplasmic domains driven by the hydrolysis of ATP. Aquaporins and Glyceroaquaporins

10.21. Protein-protein interactions mediated by lipid in crystalline arrays of AAC. The crystal packing of AAC monomers allow them to interact via cardiolipin molecules (gray), as seen both from the side (A) and from the plane of the membrane (B). From Nury, H., et al., Annu Rev Biochem. 2006, 75:713-741. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

function and also show that protein-protein interactions are mediated by endogenous lipid (Figure 10.21). The oligomeric state of AAC in its native environment is still unclear. If dimers are functional in vivo, they could face opposite orientations, allowing them to be loaded simultaneously with an internal and an external nucleotide. Trapped in open conformations that resemble a heart, a trapezoid, and a basket, the three transporters described appear to have the common general mechanism of alternating access for a single binding site. Thus their functions necessitate quite large conformational changes to open to the other side of the membrane. While this mechanism distinguishes them from channels, mutations can stabilize them in an open conformation that allows passive diffusion, giving them channel characteristics.

CHANNELS In contrast to the flexibility of transporters, channels do not require large conformational changes to allow their substrates to cross the membrane, since they contain conduction pores. The passage of ions and molecules through the pores is not uncomplicated, however. The pore characteristics depend on the nature of the walls

Although predicted in the 1950s, water channels in biological membranes were not discovered until 1992, when a protein purified from the er)'throcyte membrane was reported to greatly increase the water flux in response to a gradient of osmotic pressure. Since the discover)' of this first AQP, over 350 different AQPs have been identified in all forms of life. Mammals have 11 isoforms, designated AQPO to AQPI0, that faIJ into two classes, AQPs and glyceroaquaporins. The latter group aIJows entry of glycerol and a few other small molecules in addition to water (see below). Four of the human AQPs (AQP3, AQP7, AQP9, and AQP10) are glyceroaquaporins. The physiological roles of the different human AQPs val)' widely because they differ in cellular locations as well as in modes of regulation. For example, in secretory glands, AQP5 is specifically expressed in the apical membrane where water passes into secretions such as tears, saliva, and sweat, while in respiratol)' epithelia of the lungs, different cell types express different AQPs (Figure 10.22). The role of AQPs in the lungs was explored in the interesting case of a few asymptomatic AQP I-null individuals, whose pulmonar)' capillaries swelled normally in response to saline but their surrounding tissue did not accu mulate fluid (Figure 10.23). In addition to the basic need for all cells to keep water in balance, the AQPs are involved in several illnesses, including abnormalities of kidney function, loss of vision, onset of brain edema, and starvation. Water channels are typically bidirectional, allowing influx and efflux of water molecules in response to changing osmotic conditions. They are extremely fast: water Hovvs through a single AQPI molecule at a rate of three billion molecules per second. And they are very selective, allowing the passage of water molecules without protons (or other ions). AQPs transport only

Transporters and Channels

254

structure arose from a gene duplication event in evolution. The highly conserved sequence motif NPA is repeated near the center of each half of the primary structure, and several other conserved residues, such as GJu 14 and Glu 152, are repeated near the beginning of each segment.

A.

AQP3 AQP5 Basement/ membrane

AQP4 H 20

Respiratory airspace

B.

Structure of Aquaporins Early images of AQP I in lipid bilayers obtained by both AFM and EM gave evidence for pores. The structure of AQP 1 from hu man red blood cells was determined by EM at a resolution of 3.8 Ain the same year that the crystal structure of GlpF, the glyceroaquaporin from E. coli, was reported at 2.2 A resolution. These structures A.

130 Goblet cell AQPI-null control

v 120

AQP4

>:::

(\$':'::::

Surface layer

v v '-

CIl

'fl

CIl

-~ V ~ ~ >:::

110

v v

>

~ v

8 JOO

Basal layer

90

L '- - ' - - - - - - - ' - - - - - - - ' - - - - - - ' - - -

bsln

IL

2L

3L

Normal saline challenge B.

~':

150 AQPI

Capillary

10.22. Localizations of different human aquaporins. A. In secretory glands, AOP3 and AOP4 are found in the basement membrane, while AOP5 resides in the apical membrane where water passes into secretions. B. In epithelial tissues of the lungs, AOP4 is expressed in surface columnar cells, AOP3 in basal cells, and AOP1 in underlying fibroblasts and capillaries, while no AOPs are expressed in goblet cells. From Agre, P., Proc Am Thorac Soc. 2006, 3: 5-13. © 2006 by American Thoracic Society. Reprinted with permission from Proceedings of the American Thoracic Society.

CIl V

v

Control

140

~

.~

@~ 130 - ro c:3~ 120 :
>,

£'I

.-
0

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>:::

110

~ 100

'-

v

8

.......

90 80'

b~ln

I~

2~

• AQPI-null

3~

Normal saline challenge water, while the related glyceroaquaporins conduct small organic molecues like glycerol, urea, DL-glyceraldehyde and glycine, in addition to water. The stereoselectivity of glyceroaquaporins is indicated by the J O-fold higher rates of transport of ribitol than of D-arabitol, both of which are simple five-carbon reduced sugars. The high-sequence homology among the AQPs suggests that they use a common architecture to accomplish selective watel- transport. Although they are fairly small (AQPI is only 28 kDa), the high similarity between their N- and C-terminal segments indicates that the

10.23. Decreased pulmonary vascular permeability in AOP1-null humans. Computed tomography scans of the lung before and after intravenous infusion of saline revealed differences in water permeability of lung tissue. AOP1-null individuals and normal controls received infusions of up to 3 L of physiologic saline, and images of their bronchioles and adjacent venules were recorded and quantified. In both groups, the vessel wall of the bronchiole became thickened due to the accumulation of fluid (Al; however, the surrounding area did not accumulate fluid in the AP01-null individuals (B) since they lack the AOP to secrete water. Redrawn from Agre, P., Proc Am Thorac Soc. 2006, 3:5-13. © 2006 by American Thoracic Society. Reprinted with permission from Proceedings of the American Thoracic Society.

Channels

255

Extracellular

Membrane

Intracellular N terminus 10.24. X-ray structure of AOP1. Each TM helix (colored differently) tilts about 30" off the bilayer normal. Two half-helices (HB and HE) form one of the seven TM segments. The helices are twisted into a right-handed bundle. From Fujiyoshi, Y, et al., Curr Opin Struct Bioi. 2002, 12:509-515. (() 2002 by Elsevier. Reprinted with permission from Elsevier.

were quickly followed by that of the bovine AQPI at 2.2 A and AqpZ, the E. coli aquaporin, at 2.5 A ITSO]Ution. More recently, the structures of sheep and bovine AQPO have become available. The structures all share a unique AQP fold that consists of six TM (X-helices with two additional half-helices, so called because they each span half the bilayer. The TM helices cross i.n a righthanded twist at a 30° tilt to form an hourglass-shaped pore, with the two half-helices meeting in the center of the membrane (Figure 10.24). Both Nand C termini are cytoplasmic, with the loop connecting the N- and C-terminal segments on the extracellular side and varying in length. The NPA signature sequence for the AQPs is repeated in each of the half-helices and makes the interface between them. Purified AQPs assemble into

tetramers both in the crystal lattice (Figure 10.25) and when reconstituted into lipid bilayers, but clearly each protein has a pore. Since there is no evidence for cooperation between subunits, the functional unit of AQP is a monomer. New insights into channel selectivity have come from details of the structure of GlpF, the first highresolution AQP structure.

Glyceroaquaporins: GlpF The GlpF pmtein provides a channel for passive diffusion of glycerol into E. coli. In the cytosol glyceml is immediately converted to G3P, ensuring that the inside concentration of glycerol is low, which drives its uptake. The topology diagram for GlpF shows the symmetry between the N- and C-terminal segments (residues 6-108 and 144-254), with similar TM segments on each side of the two half-helices (M3 and M7) (Figure 10.26). In three dimensions the two segments form two inverted halves of the channel and are linked by a protruding periplasmic region (Figure 10.27). The two half-helices form an important junction in the center of the membrane, held by van der Waals interactions between the proline residues of the NPA signature motifs. The NPA motifs cup each other between the pmline and alanine side chains of the opposite helix, with their orientation stabilized by other very highly conserved residues. In addition, conserved glycines allow close contact between the (X-helices where they cmss near the center of the bilayer, stabilized by CH-O hydmgen bonds. The channel in GlpF starts on the outer surface with a wide vestibule and then constricts to form a selective channel that is 28 A long. The crystal structure sho\vs three glycerol molecules in transit (see Figure 10.27). The narmwest point in the channel defines the selectivity filter, with a very close fit for glycerol that involves hydmphobic interactions at the corner between Trp48 and Phe200 and hydrogen bonds between the hydmxyl

10.25. Tetramers of AOP1. Each monomer contains a pore, and hydrophobic interactions between monomers stabilize the tetrameric assembly, which is viewed from the top (A) and the side (8), with each monomer colored differently. From DeGroot, B. L., and H. Grubmuller, Curr Opin Struct BioI. 2005,15:176-183. © 2005 by Elsevier. Reprinted with permission from Elsevier.

Transporters and Channels

256

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10.26. Topology of GlpF. From the N terminus in the cytoplasm, the peptide chain makes three and a half helices in the membrane before reversing the pattern, ending with the C terminus in the cytoplasm. Two short helices contribute to the periplasmic portion of GlpF. The two segments of GlpF are colored to show their symmetry, with highlighted residues to show which interact with the glycerol molecules in the channel (black). contribute carbonyls to the central channel (red), and contribute hydrocarbon to the channel (purple). From Stroud, R. M., et aI., Curr Opin Struct BioI. 2003, 12:424-431. © 2003 by Elsevier. Reprinted with permission from Elsevier.

groups on C1 and C2 of the glycerol and Arg206, Gly19, and Phe200. The channel is lined by nonpolar residues except along one side (Figure 10.28). The polar side has a striking row of carbonyl oxygens along the channel, with four on the periplasmic side and four on the cytoplasmic side separated by a gap of ~3 A.. From the cytoplasm, these are the carbonyl groups from Gly64, Ala65, His66, and Leu67, and from the periplasm they are from Phe200, Ala201, Met202, and Asn203 (Figure 10.29). In each row the four residues are in an extended confomlation and have altemating right-handed and left-handed helical backbone configurations, making a ladder that is maintained by hydrogen bonds to the conserved bLllied glutamates, Glu 14 and Glu 152. These carbonyls provide hydrogen-bonding acceptors for a line of watet- molecules in the channel, which orient in opposite directions outward from the center. The only hydrogen bond donors in the water channel are Asn68 and Asn203 from the NPA signatures at the center. As each water molecule reaches the centel~ it forms hydrogen bonds to the two asparagine residues and then flips its dipole orientation to continue out the channel (Figure 10.30). In this way each water molecule is hydrogen bonded in the channel, while avoiding a continuous network of water molecules of the same orientation. This channel architecture explains how AQPs conduct water but do not leak protons. Normally protons move along a line of hydrogen-bonded waters in a

Peri plasm

35

A.

Cytoplasm 10.27. X-ray structure of GlpF. The ribbon diagram of the GlpF structure viewed from the membrane plane shows the tilted helices make a bundle, each half of which is derived from the N- or C-terminal segment of the molecule (gold and blue, respectively). Three glycerol molecules (red space-filling models) are observed transiting the channel. From Stroud, R. M., et al., Curr Opin Struct Bioi. 2003, 12:424-431. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Channels

257

Periplasm

Cytoplasm 10.28. View of the channel interior in GlpF. When the GlpF molecule is opened like a book, one side of the channel is nonpolar and the other side is polar (oxygen atoms, red; nitrogen atoms, blue; carbon and hydrogen atoms, white; and sulfur, yellow). From Stroud, R. M., et al., Curr Opin Struct Bioi. 2003, 12:424-431. © 2003 by Elsevier. Reprinted with permission from Elsevier.

10.29. Orientation of water molecules in the channel of GlpF. A. The carbonyl groups within the channel are arranged in close proximity to conduct a line of water molecules. B. A close view of the hydrogen-bond acceptors and hydrogen-bond donors in the center of the channel shows the region where the orientation of the water molecule flips. From Stroud, R. M., et al., Curr Opin Struct Bioi. 2003, 12:424-431. © 2003 by Elsevier. Reprinted with permission from Elsevier.

Transporters and Channels

258

the hydrogen bonds, making it too costly to strip waters from hydrated ions. Another factor may be electrostatic repulsion from the placement of some charged residues in the channel (Figure 10.31). These factors that delineate pore characteristics will be important in understanding other ion channels. The KcsA potassium channel (although also a tetra mer) is completely different in overall architecture, and yet its pore also features lines of carbonyl groups from residues \.\Iith alternating rightand left-handed ex-helical configurations. Potassium Channels

10.30. Snapshot of a simulation of water molecules in the channel. The central water molecule is constrained by the two conserved Asn side chains to become a hydrogen-bond donor to the neighboring waters, thus polarizing the line of waters outward from the center, which prevents proton transfer. From Stroud, R. M., et al., Curr Opin Struct Bioi. 2003, 12:424-431. © 2003 by Elsevier. Reprinted with permission from Elsevier.

concerted mannel~ such that a proton attaches to one end of a water molecule and another leaves from the other end, as long as the dipoles of all the waters "face" the same way. Called the Grotthuss mechanism, this proton transfer can leak protons across a membrane without gener'ating any charge in the channel. The change in polarization at the center of the line of water molecules in the AQP channel prevents proton conductance by the Grotthuss mechanism. Although water transport through the AQPs is too fast for experimental verification of the mechanism, many molecular dynamics (MD) simulations support this mechanism for blocking flux of protons (see Figures 10.30 and 10.31). This is of fundamental importance since cells need to be able to transpon a lot of water without losing their membrane potential. Given their functional differences, AQPs and glyceroaquaporins would be expected to differ at their selectivity filters. Indeed, the channel diameter at the selectivity filter of GlpF is larger than that of AQPl or AqpZ. However, the diameters of all three are small enough to exclude aJl ions and charged solutes, because they are roo small for the passage of hydrated ions. Since much of the surface of the channel walls is hydrophobic, residues in the channel do not have groups to replace

The family of potassium channels is found in bacteria, archaea, and eukaryotes. Like the AQPs, these channels are both selective and fast: their selectivi ty for K+ over Na" is over 1000-fold, and their conduction rates of ~108 ions/sec are close to diffusion limited. They al-e passive channels that typically allow K+ ions to flow out of a cell in response to the electrochemical gradient created by the Na+K+ -ATPase, which pumps three Na+ out and two K+ into the cell. One grou p of voltage-sensitive potassium channels, called inward-rectifying channels, opens in response to a drop in the membrane potential to allow Kt- to flow into the cell. The diversity of potassium channels surpasses that of any other channel family. In excitable cells, such as neurons, they set the resting potential and regulate the action potential. In

Si7,e restriction

Electrostatic repulsion Water dipole reorientation

8 10.31. Schematic depiction of factors determining selectivity of the AOP channel. The internal pore (blue) of the AOP allows passage of water molecules and not protons and other ions by reorienting the water molecules during passage and by size restriction and electrostatic repulsion. From Agre, P., and D. Kozono, FEBS Lett. 2003, 555:72-78. Ii;) 2003 by Elsevier. Reprinted with permission from Elsevier.

A.

10.32. X-ray structure of KcsA channel. Each subunit of the tetra mer contributes two TM helices in addition to a pore half-helix, making a cone shape when viewed from the side (A). The view from the top (8) shows the channel at the subunit interfaces. Redrawn from Nelson, D. L, and M. M. Cox, Lehninger Principles of Biochemistry, 4th ed., w. H. Freeman, 2005, p. 410. © 2005 by W. H. Freeman and Company. Used with permission.

the cardiovascular system they regulate the heartbeat. In epithelial cells they help balance the passage of salts and watec Many cells have several types of potassium channels, some that respond to changes in the membrane potential or in pH and others that are triggered by binding ions, such as calcium ions; ligands, such as neurotransmitters and cyclic nucleotides; or even other proteins, such as certain G proteins. While potassium channels differ in gating mechanisms, they share a common pore architecture. Their TM domains consist of four usually identical subunits with the ion pathway down the center at the subunit interface. Their signature sequence forms the selectivity filter of the channel, described below. Gating enables conformational changes to open and close the pore in response to different signals, requiring a sensor domain to transmit the information 10 the pore domain. One or two regulators of K+ conductance (RCK) are gating domains that typically are attached to the intracellular C terminus of the pore domain, although RCK domains are also expressed as independent proteins. Thus the basic channel design appears to have evolved by addition of modular gating domains conferring sensitivity to voltage or ligands to generate the diverse potassium channels. Over 40 years after experiments measuring the flux of 42K+ across the membrane from squid axon suggested the single-file movement of K+ ions occurs through a putative membrane channel, the first highresolution structure of a potassium channel verified that model. In the intervening decades, electrophysiological, biochemical, biophysical, and computational techniques amassed an impressive amount of data characterizing these channels, while in recent years structures of more potassium channels have offered addi-

tional insights into channel opening and channel gating. To date, more than one conformation of any single K+ channel has not been solved, so the conformational changes involved in opening and gating can only be inferred by comparing the structures of different channels. Potassiu m channels belong to a family of letrameric calion channels, in which each tetramer has a central pore. The first x-ray structure was obtained for KcsA, a potassium channel from Streptomyces lividal1s chosen for its simple topology. While many K+ channels have six TM helices, each KcsA subunit has only two TM helices; however, KcsA still binds charybdotoxin, a small protein from scorpion venom that inhibits some of the larger eukaryotic potassium channels. After removal of its flexible C-terminal domain by cleavage with chymotrypsin, KcsA formed crystals that gave 3.2 A resolution. Attached to monoclonal Fab fTagments, KcsA gave better crystals that allowed resolution of 2.0 A. As described below, the KcsA pore domain crystallized in its closed conformation. The next K+ channel to be solved, calcium-activated MthK from Melhanobacleriwl1 Ihennoaulolrophicus, crystallized in its open conformation. It was followed by structures of KvAP, a voltage-gated channel from the thermophilic archaea Aeropyrum pemi.x and Kvl.2, a voltage-gated mammalian Shakerchannel- both open - and KirBac 1.1 and KirBac1.3, prokaryotic inward-rectifying channels both closed. KcsA Structure and Selectivity The overall shape of the KcsA protein is a cone within a cone, constricted at the cytoplasmic side and opening at the extracellular side (Figure 10.32). The channel is lined by four pairs of helices: each subunit contributes

Transporters and Channels

260

.~

.I@

10.33. The selectivity filter and potassium-binding sites of KcsA. A close-up view of the selectivity filter in the KcsA ion channel with the extracellular surface at the top shows the electron density (blue mesh) observed for dehydrated K+ ions at positions 1 to 4, in addition to a hydrated K+ ion in the central cavity. The chapter Frontispiece shows the location of the selectivity filter in the ion channel. From MacKinnon, R., FEBS Lett. 2003, 555:6265. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

an outer helix and an inner helix that span the bilayer. In addition, a shorter tilted half-helix from each subunit fills in the top of the cone to make the pore. Each pore helix is oriented with its C-terminal end inside, putting the negative end of its dipole toward the center. On the extracellular side of the channel, a narrow pore 12 A long defines the selectivity filter and ends at a large water-filled cavity in the center (Figure 10.33 and Frontispiece). A fully hydrated K+ ion is observed inside the cavity. In addition, within the pore there are four binding sites for K+, designated SI to S4 from the outside to the inside. Both the water-filled cavity and the orientation of the pore helices help overcome the electrostatic repulsion for the positively charged ions entering the channel. Indeed, in a similal- an-angement in the CJ- channel, the dipole of the pore helices is reversed, which places their N termini inside to contribute positive charge that a ttracts the anion. In the selectivity filter of KcsA, each signature sequence of TVGYG provides four evenly spaced layers of carbonyl oxygen atoms and a single layer of threonine hydroxyl oxygen atoms to create the four K+ -binding sites (see Figure 10.33). Both glycine residues and the threonine have dihedral angles that are allowed for a left-handed ex-helix. Alternating between right-handed and left-handed hel ical configurations enables the back-

bone carbonyl groups to line up. This puts at each binding site eight oxygen atoms at the vertices of a cube (or twisted cube) that corresponds to the arrangement of the water molecules surrounding the hydrated K+ observed in the central cavity. Therefore, the role of the selectivity filter is to mimic the waters of hydration for a queue of K+ ions, which pays the energetic cost of their dehydration. Na+ ion is apparently too small to fit (the atomic radii of K+ and Na+ are 1.33 Aand 0.95 A, respectively), so its dehydration energy is not compensated and it stays outside, explaining the selectivity for potassium. While the lines of oxygen atoms provide good binding sites, a strong binding of potassium would inhibit the fast rate of flux through the channel. Binding is weakened by electrostatic repulsion fyom neighboring ca tions in the queue, as well as by a conformational change that takes place upon potassium binding. At very low (nonphysiological) potassium concentrations, with only one K+ bound per tetramel; the selectivity filter collapses inward; binding a second K+ causes the channel to straighten out. This conformational change takes up some of the binding energy, making the binding weaker. The mechanism of potassium conduction through the channel is suggested by the stoichiometry of binding that was measured experimentally with Tl+ (thallium ion, an excellent replacement for K+ in laboratory work). It appears that the channel binds two ions with two waters between them. Computational experiments indicate that it has two states of equal energy with S I/S3 occupied or S2/S4 occupied. The flux of K+ through the channel involves a concerted movement in response to another K"" approaching, either from the internal cavity or from the extracellular site called SO (Figure J 0.34). Gating and Conformational Changes KcsA is controlled by intracellular pH values, and only a few percent of channels are open during functional assays in vitro. The crystal structure of the KcsA pore domain shows a closed channel, sealed from the cytoplasm by the helical bundle below the central cavity. Portions of the molecule not seen in this structure, including the unresolved N terminus and the proteolytically l-emoved C-terminal domain, are likely to be involved in conformational changes upon opening the channel at low pH, according to results obtained with NMR, EPR, and a novel surface plasmon resonance technique. The Ca 2+-gated channel MthK was crystallized in the presence of Ca2+ and reveals an open pathway that is abou t loA wide from the cytoplasm up to the selectivity filter (Figure I 0.35A). The helices of the pore domain are hinged at a conserved glycine residue about halfway across the membrane. Superimposition of the MthK and KcsA channels reveals the motion of the inner helix

Channels

261

10.34. Potassium binding during conduction through the KcsA pore. The selectivity filter has four binding sites, 51 to 54 (from the outside), and there is an additional extracellular site called 50. In a concerted process, an ion coming from the internal cavity fills the 54 site and moves the ions in the 51/53 sites to the 52/50 sites (left-hand pathway) as it enters the channel. The energetic state of 51/53 binding and 52/54 binding is the same, and the process is fully reversible. From Roux, B., Annu Rev Biophys Biomof Struct. 2005, 34:153-171. © 2005 by Annual Reviews. Reprinted with permission from the Annual Review of Biophysics and Biomolecular 5tructure, wwwannualreviews.org.

required to allow the helix bundle to open like the aperture of a camera (Figure 10.35C and D). Structure-based sequence alignment of a wide range of K+ channels shows consen/ation of this glycine residue (Gly83 in MthK and Gly99 in KcsA), suggesting this gating hinge is conserved. In addition, a conserved glycine or alanine is found at a position five amino acids toward the C terminus, below the hinge where a larger side chain would block the open pore. Conservation at these two positions implies the proposed conformational change is conserved in the different K+ channels. Each subunit of MthK consists of two TM helices that contribute to the pore plus a single C-terminal RCK domain (Figure 10.36). When the gene is overexpressed, additional copies of the RCK domain are synthesized, four of which associate with the MthK channel to make a gating ring composed of eight RCK domains at the intracellular membrane surface of the tetrameric pore (Figure J 0.37). The connection between the pore domain and the gating ring (residues 99-115) is not resolved in the crystal structure. The RCK domains have an ex-13 fold similar to dehydrogenase enzymes. A cleft between two RCK domains binds two Ca 2 + ions with carboxylate groups ofGlu210, Glu212, and Asp 184. The importance of these residues in gating is illustrated by the decrease in the ability of Ca 2+ to open the channel in a DJ 84N mutant. There are two kinds of domain interfaces within the gating circle, one flexible and one rigid. The mechanism of gating is proposed to involve movement of a flexible hinge near the Ca 2 +-binding site,

A

B.

c.

D.

10.35. Comparison of TM domains of MthK (open) and KcsA (closed). A and B. Traces of the Co: of the two K+ channels (extracellular side up) are viewed with one subunit removed from within the membrane for MthK (A) and KcsA (B). C and D. 5uperposition of the KcsA (red) and MthK (black) pores viewed from within the membrane (C, two subunits) and from the intracellular solution (D, four subunits, showing only the selectivity filter and inner helices). Arrows indicate the direction of inner helix displacement in going from closed to open states. From Jiang, Y, et al., Nature. 2002, 417:523-526. © 2002. Reprinted by permission of Macmillan Publishers Ltd.

Transporters and Channels

262

Pore domain M 1 = outer helix MZ = inner helix P = pore helix

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10.36. Structure of a protomer of the calcium-gated MthK potassium channel. The MthK protomer has two copies of a regulatory domain (RCK, regulators of calcium conductance; dark blue and red) that forms a gating ring at the intracellular membrane surface and binds two Ca 2 + ions (yellow spheres). It is connected via a disordered segment (dashed line) to the pore domain (blue cylinders). From Jiang, Y, et aI., Nature. 2002, 417:515-522. © 2002. Reprinted by permission of Macmillan Publishers Ltd.

which pushes two adjacent rigid domains to rotate, opening or closing the pore (see Figure 10.37C). While the gating domains of ligand-dependent channels are typically in the aqueous solution outside the membrane, the gating domains of voltagedependent potassium channels are integral to the membrane. Like MthK, the voltage-dependent channel KvAP is a tetra mer with a pore domain similar to that of KcsA, but the pore is surrounded by its voltage sensors. KvAP has six hydrophobic segments, numbered SI to S6 fmm the N terminus. SS and S6 correspond to the outer and inner helices of KcsA, and although the pore appears to be open at the glycine-gating hinge, in the extraceJJular leaflet of the membrane the pore helices are superimposable. The other four segments, SI to S4, form the voltage sensors. The crystal structure of KvAP shows that S 1 and S2 make a concentric layer of helices outside the pore helices, and S3 and S4 form a voltage paddle at the outer perimeter (Figure 10.38). The voltage paddle of KvAP is a helix-tum-helix structure connected to the rest of the protein by a flexible loop in the middle of S3. In the first crystals, which were obtained with Fab fragments bound at the tip of each paddle, crystal contacts pulled the paddles toward the intracellular side of the membrane in a conformation

likely to be non-native since cryo-EM images reveal the Fab fragments (and therefore the paddles) are pointing to the extracellular side in the native structure. As the position of the paddles is not altered in subsequent clystals obtained without Fab fragments, these results suggest the voltage sensors could be loosely attached to the pore domain and located at the protein-lipid interface. Voltage gating requires sensors that respond to electrical signals with conformational changes. The voltagesensor paddle in KvAP is made up of hydrophobic residues along with several arginine .-esidues on S4. Four of the arginine residues are considered the gating charges of the protein because they can move in response to the membrane potential, as demonstrated in numemus studies employing electrophysiology, scanning mutagenesis, and fluorescence techniques. When the charged helix-turn-helix structure senses a change in the electric field, the movement of the paddles carries positive charge (on two or more arginine residues) across the membrane. In view of the C1ystal structure, the voltage sensor paddle was postulated to make a large

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10.37. Pore and gating domains of calcium-gated MthK. The structure of the MthK channel shows the pore domain on top and the gating ring below it, with subdomains removed for clarity. RCK domains that interact with each other are the same color. Disordered linkers are dashed lines and Ca 2 + ions are yellow spheres. A. Crystal structure of the open channel. B. Hypothetical model for a closed conformation of MthK. C. Model for the gating motions, showing how a single rigid unit (red) of the gating circle could rotate to close the pore. From Jiang, Y, et aI., Nature. 2002,417:515-522. © 2002. Reprinted by permission of Macmillan Publishers Ltd.

Channels

263

A.

B.

10.38. X-ray structure of voltage-gated KvAP. A. The KvAP tetramer is viewed from the intracellular side of the membrane, with each subunit a different color and helices of one subunit labeled 1 to 6 and P for the pore half-helix. B. A single subunit viewed from the side with dashed lines to indicate the membrane and the intracellular solution on the bottom. The voltage-sensor paddle is at the bottom, with conserved arginine residues indicated. From Jiang Y, et al., Nature. 2003, 423:33-41. © 2003. Reprinted by permission of Macmillan Publishers Ltd.

movement from one side of the membrane to the other This model is supported by ingenious experiments measuring avidin binding to KvAP channels biotinylated at residues 121 and 122 (Figure 10.39). When the channel is closed, biotin is accessible to avidin from the intracellular side, and when it is opened, it becomes

accessible from the extracellular side. However, optical experiments monitoring the movement of bound fluorophores during voltage gating indicate a smaller movement of S4 occurs, one that could be achieved by rotating the helices instead of flipping them. The discrepancy may be resolved by allowing considerable

Closed External + + +

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10.39. Schematic of avidin-biotin detection of voltage-sensor paddle movement. Cysteine mutagenesis at different locations allows an activated biotin to be tethered to different locations in KvAP. Addition of avidin (green) from the intracellular side or extracellular side can abolish the K+ current measured in planar lipid membranes if the biotin is available on that side of the membrane. From Jiang, Y, et al., Nature. 2003, 423:42-48. © 2003. Reprinted by permission of Macmillan Publishers Ltd.

Transporters and Channels

264

thinning of the lipid bilayer at the KvAP structure, as observed in an MD simulation of the S4 helix in lipids. The simulation suggests that phosphate groups of some lipid headgroups make salt bridges vvith the buried arginine residues as they snorkel toward a deformed lipid interface. The movement of the voltagesensor S4 may be communicated to the pore helices to open the pore via a mechanism similar to the response of the mechanosensitive protein MscS (see Chapter 9). Voltage-gated channels for Na+ and for Ca 2+ share many of the features of voltage-gated K+ channels, although they are much larger proteins (~2000 amino acids). They consist of four domains, each of which is homologous to the K+ channel TM domain, so they are considered pseudo-tetramers and are expected to have folds similar to that of the K+ channel tetramer. Like the potassium channel, these passive channels open to allow the ions to flow dovvn gradients created by active ion pumps, the Na+K+ -ATPase and the Ca 2 +-ATPase.

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10.40. The El-E2 reaction scheme for the Ca 2 + -ATPase. The El form of the enzyme binds two Ca 2 + ions from the cytoplasm, along with ATP bound to M g 2+ (not shown) before the phosphorylation reaction. Phosphorylated enzyme (El P.2Ca 2 +) then exchanges Ca 2 + ions for H+ on the outside as it converts to E2. E2 is dephosphorylated and then converts back to El. This scheme is simplified because both the phosphorylation step and the dephosphorylation step are reversible, and also either the ATP or the Ca 2 + can bind first. From Toyoshima, c., et al., Annu Rev Biochem. 2004, 73:269-292.

Calcium ATPase

The Ca 2 +-ATPase enables organisms to keep the intracellular concentration of calcium low (~1O-7 M) to avoid formation of insoluble calcium phosphates. Important as a signaling molecule itself (see examples in Chapter 4), calcium also plays major roles in electrical signaling. In muscle contraction a nerve impulse triggers the release of Ca2+ from the sarcoplasmic reticulum. The released Ca2+ binds troponin, triggering a conformational change that exposes myosinbinding sites to bind actin in thin filaments. In the nervous system, depolarization stimulates voltagegated Ca2+ channels at the tip ofaxons to allow entry of Ca 2 + that triggers the release of acetylcholine into the synaptic cleft. Following such events, the Ca z+ATPases in the plasma membrane and membranes of the endoplasmic reticulum and the sarcoplasmic reticulum pump Ca2+ out of the cytoplasm to restore its 10 4 fold concentration gradient. The calcium pump is a P-type ATPase (see Chapter 6) because it carries out active transport at the expense of ATP hydrolysis and is phosphorylated as part of the transport cycle. It is autophosphoryJated on an aspartate residue at the start of the signature motif of the P-type ATPase superfamily, DKTGT[LIVM][TlSl Early work indicated it has two conformations, EI and E2. EI has a high-affinity Ca2+-binding site exposed on the cytoplasmic side, and E2 has a Jow-affinity Ca 2 + -binding site exposed to the outside. To couple the hydrolysis of ATP to the transport of calcium, the loaded calcium site must become closed to the cytosol in the phosphorylated (activated) enzyme. Thus its proposed mechanism is similar to that of the Na+K+ -ATPase (shown in Figure 6.4).

For each ATP hydrolyzed, the Ca 2 +-ATPase exports two Ca 2 + ions and allows import of two or three H+. The events of the transport cycle are summarized as follows (Figure 10.40): EI binds two Ca2+ ions and ATP from the cytoplasm. Phosphorylation at Asp3S1 leads to formation of an EI P·2Ca 2+ high-energy intermediate with occlusion of the Ca 2+ ions. Conformational change to the low-energy E2P intermediate allows release of Ca 2+ to the outside, in exchange for two or three protons. Dephosphorylation of E2P to E2 allows the enzyme to convert back to EI and release protons into the cytoplasm, where it picks up two more Ca 2 + to repeat the cycle. Understanding of the transport mechanism has been greatly enhanced by the determination of structures of the Ca 2 +-ATPase from rabbit sarcoplasmic reticulum in at least five different states. Conformational Change in the Ca 2 +-ATPase The sarcoplasmic reticulum Ca2+ -ATPase (SERCAI) is a single polypeptide of 994 residues (Mr 110 kDa) with 10 TM helices and an extensive cytoplasmic headpiece (Figure 10.41). Four of the helices (M2-MS) extend past the membrane into the cytoplasmic region. The cytoplasmic headpiece consists of three domains: the A domain that actuates the gating mechanism, the N domain that binds nucleotides, and the P domain, where phosphorylation takes place. The structures of the EI·2Ca 2+ and E2 forms of the enzyme have been solved at 2.6 Aand 3.1 A, respectively (Figure 10.42). The Ca2+ -free form of the enzyme was crystallized bound to an inhibitol~ thapsigargin (TG). The position of the bilayer was determined from symmetry constraints of SERCAI in different crystal forms and puts a ring of

Channels

265

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djs~ase

10.41. Topology of the sarcoplasmic reticulum Ca 2 + -ATPase (SERCA 1). The functions of various residues are indicated by the shapes listed on the right. From Toyoshima, C, et al., Annu Rev Biochem. 2004, 73:269-292. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

tryptophan residues at the interface between the polar and nonpolar regions on the cytoplasmic side. In addition, amphipathic segments of loops between TM segments as well as a short amphipathic part of MI reside in the interfaces. The two stl-uctures show that when SERCAI binds Ca 2+, the cytoplasmic domains undergo large movements. In the Ca 2+ -bound form the A, N, and P domains are widely separated, while in the Ca 2+ -free form they are in a more compact structure. The N domain rotates toward the P domain, and the A domain rotates 45" upward to fit into a crevice between the Nand P domains (see Figure 10.42). These movements influence the TM helices connected to these domains: the A domain is connected to M I to M3, and the P domain is connected to M4 and M5 and is linked by hydrogen bonds to M3. While Ml to M6 move considerably during the reaction cycle, M7 to M 10 do not move (Figure 10.43). The very long (~60 A) M5 helix functions as the spine of the molecule

and bends at Gly770 during the conformational change. Close contacts between a GxxxG motif (Gly841 and Gly845) on M7 and Gly770 in M5 are probably important during the conformational change (Figu re 10.44). The structures of the cytoplasmic domain indicate residues involved in binding and hydrolysis of ATP. The P domain has a Rossmann fold for nucleotide binding. Asp351, the site of phosphorylation in the P domain, is surrounded by three critical aspartate residues (Asp267, Asp703, and Asp707), one of which is involved in binding the Mg2+ associated with the ATP. Nearby is the highly consened Lys684, which binds the )'-phosphate of ATP. The adenosine moiety binds to groups of the N domain, including Phe487 that stacks with the adenine ring and other hydrophobic residues, along with ~ys515 and Glu442. The highly mobile A domain contains a conserved sequence motif near the phosphorylation site that appears to shield the aspartyl phosphate fTom water in the E2P state.

Transporters and Channels

266

2(TG)

EICa2+

.~

......... ,.,...

~> A

Lumen 10.42. X-ray structures of SERCA1 in the presence and absence of calcium ions. The Ca 2+ -free form of the enzyme was crystallized bound to an inhibitor, thapsigargin (TG). In the structure of E12Ca 2 + (violet), the three cytoplasmic domains are splayed open, with the ATP-binding site available. The two cyan circles point out the location of bound Ca 2+ ions. The red arrows indicate the movements of the cytoplasmic domains that would produce the Ca 2+ -free form. In the absence of Ca 2+, the E2(TG) structure (green) shows the N, P, and A domains gathered into a compact headpiece. The black dashed line illustrates the angular movement of the N domain. The cyan arrow shows the proposed entry path for Ca 2+. The bilayer shown is an MD simulation of DOPe. From Toyoshima, e., et al., FEBS Lett. 2003, 555: 106-11 O. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

A.

B. 10.43. Rearrangement of TM helices when SERCA 1 binds calcium. The TM helices of SERCA 1 (numbered) are superimposed in positions observed in E1·2Ca 2+ (violet) and E2(TG) (green). Both A and B are viewed from the side, with a 90° rotation in the viewpoints (such that the view in B is the back of the view in Figure 10.42). Helices M8 and M9 are removed in B to show the movements of M5, M2, and M4. The double circles (red and white) show pivot positions of M2 and MS. The red arrows indicate the direction of movements during the change from E1·2Ca 2+ to E2(TG), and the cyan arrow shows the proposed pathway for entry of the first Ca 2 + ion. From Toyoshima, e., et aI., Annu Rev Biochem. 2004, 73:269-292. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Channels

267

B.

A.

10.44. Close contact between TM helices M5 and M7 in SERCA 1. The GxxxG motif (green space-filling models) in M5 interacts with Gly770 (blue space-filling model) at the pivot point in M7. The view in B is rotated 90" from the view in A. From Lee, A. G., Biochim Biophys Acta 2002, 1565:246-266. © 2002 by Elsevier. Reprinted with permission from Elsevier.

The two Ca 2 + ions bind in a cooperative manner to sites located side by side in the TM domain (Figure 10.45). Site I is surrounded by Ms, M6, and M8 helices at the center of the TM domain, while site II is closer to the cytoplasmic surface on the M4 helix 'where M4 is partly unwound. Asp800 on an unwound part of M6 contributes to both sites. Most mutations of the specific residues involved in binding Ca 2 + at sites 1 and rr abolish binding, as expected. An entrance to the Ca2+ -binding sites located just below the phospholipid interface is suggested in the EJ -Ca 2+ crystal stmcture (Figure 10.46). For vectorial transport, Ihis site should be closed off upon phosphorylation; however, the intermediates EIP·ADP·2Ca2+ and E1P·2Ca2+ are short-lived. To see the occlusion of this site in phosphorylated intermediates, SERCA has been crystallized

10.45. The calcium-binding sites in SERCA 1. There are two high-affinity sites that bind Ca 2 + (blue spheres), each with seven oxygens from amino acid residues along with two waters (red spheres). From Toyoshima, C, et aI., Annu Rev Biochem. 2004, 73:269-292. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

with analogs of Pi and ATP present. The structure of EI-AlF4"·ADP·2Ca2+ (in which AIF4" is a phosphate analog) reveals movements ofTM helices MI, which makes a kink, and M2, which is lifted up about 10 A. These changes close the proposed entrance gate to site TI. The structure of yet another form or SERCA, the E2AIF.j(TG) complex, gives information about the dephosphorylation step. The N domain is moved away from the P domain by a rotation due to the bending of MS. The interactions of AIF4" with Asp3S1 and the catalytic Glul83 are consistent with an SN2 mechanism for dephosphorylation. In the next step protons musl enter the structure, although they are not seen at this

A.

10.46. Entrance gateway for Ca 2 + in SERCA1. A. The region in E1·2Ca 2 + at M 1 (gold) and M2 (lavender) makes a gateway lined by Glu109, Glu55. Asp59, and Glu58 (behind Asp59 at this angle) leading directly to Glu309 and the Ca 2 +ion (green) at site II. B. The conformational change occurring with phosphorylation blocks this entrance pathway as indicated in this region of the structure of E1AIF4" ·ADP·2Ca 2 +. From M011er, J. et aI., CurrOpin Struct BioI. 2005, 15:387-393.~) 2005 by Elsevier. Reprinted with permission from Elsevier.

v.,

Transporters and Channels

268

E1·2Ca 2 +

.....

E1·AIF ol-' ADP

A N

~

A

... '.,:

p

c> Eb-' E1'2Ca'';'' E1' AlP

2H~ 2Ca"

iJ

ATP

E2 P,

+

D

E1p·ADP

~

2H'

E2'P, -E2P

-r

t

ADP

E1P

2Ca"

E2(TG)

,

......

A

Q

2Ca 2' Cytoplasm

M

M1 ' SR Membrane M1

Lumen

Lx 10.47. Structural transitions during the transport cycle in SERCA 1. The conformational changes during the Ca2+ -ATPase transport cycle are shown with structures from the major stages of the E1-E2 cycle (shown in center). Motions of the headgroup domains are indicated by dashed arrows. Color changes gradually from the N terminus (blue) to the C terminus (red). Key residues, including R560, F487, E183, D351, and D703, are shown in ball-and-stick representation. When present, ATP, ADP, AIF4, and TG are shown in space-filling molecules and Ca 2 + is circled. From Inesi, G., et aI., Biochemistry. 2006, 4513769-13788. © 2006 by American Chemical Society. Adapted with permission from American Chemical Society.

For Further Reading resolution. Overall, this form of the enzyme has few structural differences from the E2(TG) structure except for the mobile A domain. Thus four structures of SERCA can portray most of its conformational changes during the EI-E2 reaction cycle to give remarkable insight into the function of this complex ion pump (Figure 10.47). Binding of two Ca 2+ ions opens the headpiece su fficiently for ATP to bind, which leads to large movements as the headpiece domains close. Bending of the A and P domains bring the ATP close to Asp351, allO\.ving transfer of the phosphoryl group to generate the phosphoenzyme intermediate. Another conformational change allows ADP to leave and presumably opens a channel for extrusion of the calcium ions into the lumen. Structural biology has come a long way toward elucidating the transport mechanism of the Ca 2+-ATPase and, by extension, other members of the family of Ptype ATPases, including the Na+K+-ATPases in animal cells and the H+ -ATPases in plant cells, which are both essential for the formation of the membrane potential. While the mechanism is more complex than those of the other transporters and channels described in this chapter, it is accomplished by a relatively simple protein structure when compared with the F 1 Fa-ATPase described in Chapter 11.

FOR FURTHER READING

269

Mitochondrial ADP/ATP Carrier Nury, H., et aI., Relations between structure and function of the mitochondrial ADP/ATP carrier. Armu Rev Biochem. 2006,75:713-741. 'Pebay-Peyroula, E., et al.. Structure of the mitochond.·ial ADP/ATP carrier in complex with carboxyatractvloside. Nature. 2003,426:39-44. Pebay-Peyroula, E., and G. Brandolin, Nucleotide exchange in mitochondria: insight at a molecular level. Curr Opin Slruct Bioi. 2004, 14:420-425. Aquaporins Agre, P.. Aquaporin water channels (Nobel lecture). Angew Chem Int Ed. 2004,43:4278-4290. DeGroot. B. 1.., and H. Grubmuller, The dynamics and energetics of water permeation and proton exclusion in aquapar·ins. Curl' Opin Struct Bioi. 2005, 15: 176-183. 'Fu, D. X., et aI., Structure of a glycerol-conducting channel and the basis for its selectivitv. Science. 2000,407:599605. Fujiyoshi, Y, et aI., Structure and function ot water channels. Curl' Opin Struct Bioi. 2002,12:509-515. 'Murata. K., et aI., Structural determinants of water permeation through aquaporin-l. Nature. 2000, 407:599605. Stroud. R. M., et aI., Glycerol facilitator GlpF and the associated aquaporin family of channels. Curl' Opin Struct Bioi. 2003, 12:424-431. 'Sui, H., et aJ.., Structural basis of water-specific transport through the AQP 1 water channel. Nature. 200 I, 414:872878.

LacY and GlpT 'Abramson, J., 1. Smirnova, V. Kasha, G. Verner, H. R. Kaback, and S. Iwata, Structure and mechanism of the lactose permease of Escherichia coli. Science. 2003, 301 :61 0615. Abramson, J., H. R. Kaback, and S. Iwata, Structtmll comparison of lactose permease and the glycerol-3-phosphate antiporter: members of the major facilitator superfamily. CurrOpin Struct Bioi. 2004,14:413-419. Guan, L., and H. R. Kaback, Lessons from lactose permease. Annu Rev Biophys Bi011101 Struct. 2006, 35:67-91. "Huang, Y, M. J. Lemieux, J. Song, M. Auer, and D.-N. Wang, Structure and mechanism of the glycerol-3-phosphate transporter from Escherichia coli. Science. 2003,301:616620. Lemiuex, M. J., Y Huang, and D.-N. Wang, The structural basis of substrate translocation by the Escherichia coli glycerol-3-phosphate tranSpOJ1er: a member of the major facilitatorsuperfamily. CurrOpinStruct Bioi. 2004,14:405412. Lemieux, M. J., Y. Huang, and D.-N. Wang, Glycerol-3phosphate transporter of Eschen:chia coli: structure, function and regulation. Res i\llicrobiol. 2004, 155:623-629.

• Paper presents original structure.

Potassium Channels 'Doyle, D. A., el aI., The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science. 1998,280:69-77. Gouaux, E., and R. MacKinnon, Principles of selective ion transport in channels and pumps. Science. 2005, 310: 14611465. •Jiang, Y, et aI., Crystal structure and mechanism of a calcium-gated potassium channel. Nature. 2002, 417:523526. •Jiang, Y.. et aI., X-ray structure of a voltage-dependent K+ channel. Nature. 2003,423:33-41. MacKinnon, R., Potassium channels and the atomic basis of selective ion conduction (Nobel lecture). Angew Chern Int Ed. 2004,43:4265-4277. Roux, B., Ion conduction and selectivity in K+ channels. Annu Rev Biophys Biomol Struct. 2005, 34: 153-171. Tombola, F, et aI., How does voltage open an ion channeP A,1I7U ReI' Cell Dev Bioi. 2006, 22:23-52. 'Zhou, Y. et aI., Chemistry of ion coordination and hydration revealed by a K+ channel-Fab complex at 2.0 A resolution. NalUre. 200 I, 414:43-48 .

Transporters and Channels

270

Calcium ATPase 2

Lee, A. G., Ca +-ATPase stmcture in the EI and E2 conformations: mechanism, helix:helix and helix:lipid interactions. Biochim Biophys Acta. 2002, 1565:246266. ·'Toyoshima, c., et aI., Crystal structur·e of the calcium pump of sarcoplasmic reticulum at 2.6 Aresolution. Nature. 2000, 405:647-655.

'Toyoshima, c., et aI., Structural changes in the calcium pump accompanying the dissociation of calcium. Nature. 2002, 418:605-611. Toyoshima, C., and G. Inesi, Structural basis of ion pumping by Ca 2 + -ATPase Ol the sarcolasmic reticulum. Al1rlU Rev Biochem. 2004, 73:269-292. Toyoshima, c., [on pumping by calcium ATPase of sar·coplasmic reticulum. Adv Exp Med Bioi. 2007, 592:295-303.

11

Membrane Protein Assemblies

Extracellular

OM

Assemblies oj membrane proteins can span two membranes, exempli ied 111 the dynamic coupling netween inner membrane transporters and a hannel-tlli1n€e1 ,KrOSS the periplasm and outer membrane or rug efflux In Gram negative bacteria TI15 model of the AcrABfTolC systeln, which can export drugs S ch as novobiocin from insidt> rhe cell and ampicillin from he penplasm, is composed 01 the x-ray s ucture of TolC modeled to be the open s ate (red], he x-ray structure 01 AcrB (green), and a ,'epresenta Ion of AcrA based on the x ray struc LJre of ItS close homolog, MexA (blue). rrom Eswaran. J., el ill, Curr Opin Struct Bioi. 2004, 14:741 747 7004 by Elsevi r Reprinted With permission from Elsevier

Periplasm

1M

Cytosol

Most of the membrane proteins described in the previous chapters can carry out their tasks without partners, although some form homo-oligomers and others are involved in transient interactions, for example, with signaling proteins. Because they can function on their own, their high-resolution structures reveal a great deal about their mechanisms. In contrast, many membrane proteins function in large complexes and can be understood only when the other protein components in these multi protein assemblies are characterized as well. This chapter describes structures of some multi-

component complexes that can be viewed as molecular machines, or nanomachines, in the membrane. These vary from large enzymes composed of many subunits, such as ATP synthase, to dimers of protomers* that each have many subunits, such as cytochrome-bel oxidase, to structures formed when separate proteins interact, A protomer is the minimal structure from which a larger structure is buill. Although the term prolomer can refer to monomers. it is used here to denote subunits which themselves contain subunits. Similarly, mullimers are the larger units resulting from the assembly of protomers.

271

272

11.1. ATP synthase molecules observed on inside-out vesicles of bovine heart mitochondria. The knob-on-stalk appearance of the ATP synthase is seen in negative staining EM. From Walker, J. E, Angew Chern Int Ed. 1998, 37:2308-2319. © 1998 by Gesellschaft Deutscher Chemiker. Used by permission of WileyVCH Verlag GmbH.

Membrane Protein Assemblies was recognized by the award of the 1997 Nobel Prize in Chemistry to Paul Boyer and John E. Walker for "the elucidation of the enzymatic mechanism underlying the synthesis of ATP." Further experimentation and structural biology have contributed to the present understanding of this marvelous mechanism. Unlike the calcium pump described in Chapter 10, which as a single polypeptide couples ion flux to the hydrolysis of ATP, lhe ATP synthase is a molecular machine with many different polypeptide components. With eight (prokaryotic) to 18 (mammalian) different subunits, the complex has a molecular mass of 550 to 650 kDa. The F 1 domain is the catalytic domain that carl-ies out the synthesis and hydrolysis of ATP. Forming the knobs in Figure 11.1, F 1 is extrinsic to the membrane, so it can be removed by mild treatmenls and function as a soluble ATPase. E. coli has the simplest F 1 domain, with the composition cx3f3,yb£. The F o domain is lypically composed of ab 2c(lo_12J, although some complexes have two different b subunits instead of a homodimer, and the number of c subunits varies in differenl

sometimes across more than one membrane as seen in the proteins involved in drug efflux in Gram-negative bacteria (Frontispiece).

F,Fo-ATPASE/ATP SYNTHASE Familiar fOJ'decades for its knob-on-a-stalk appearance, the FIFo-ATPase is named for its two major structural domains, F 1 and Fo (Figure 11.1). This fascinating complex is also called the ATP synthase because it couples the flow of protons across the membrane to the synthesis of ATP as well as its hydrolysis, depending on the direction of proton flux. Found in the plasma membrane of bacteria, the mitochondrial inner membrane in eukaryotes, and the chloroplasllhylakoid membrane in plants, the ATP synthase is the major producer of ATP in cells using either oxidative phosphorylation or photosynthesis to generate a proton motive fOl-ce (pmf, the proton electrochemical gradient across the membrane that stores energy). The importance of ATP generation is clear to humans, who each use around 40 kg of ATP each day of a sedentary life (an athlete uses much more!). With around 100 mmol in the pool of adenosyl nucleotides in the body, this rate of consumption means thal each molecule of ADP must be phosphorylated about a thousand times a day to provide the needed ATP. The reverse reaction of the F 1 Fo-ATPase is also important under conditions when the utilization of ATP is needed to replenish the proton gradienl. How lhe flow of protons across the membrane is coupled to the catalylic reaction is a question thal is fundamental to life, Biochemical data on the catalytic mechanism of the F 1 Fo-ATPase gained beautiful support from the first high-resolution structures of its components. This

11.2. Overall structures of the F1 Fo-ATPase from E. coli. The arrangement of the subunits making up Fo and F, is diagrammed with one ex subunit removed to reveal the y subunit down the center. The three pairs of exf3 subunits of F, (magenta/pink, dark blue/light blue, and green; the third ex subunit is removed) surround the y (red) of the stalk, with [ (yellow) at its base and & (orange) along the back. The c subunit ring (black) of Fo is linked to the central stalk (yt:) as well as to the peripheral stalk composed of b2 (green) and b. The five predicted TM helices of the a subunit are represented (gold). Note: the bows show some of the crosslinks that either have little or no effect on (green) or inhibit (red) the activity, which demonstrated that c, y, and [ rotate together. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002, 27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

F, Fa-ATPase/ATP Synthase

sources and even under different growth conditions. When the F I domains are stripped off, Fa by itself forms a passive proton pore. The Fa domain is sensitive to the inhibitor oligomycin, for which it was originally named. In the complex, these two structural domains are connected by two stalks: a central stalk made up of y and £ subunits (yb£ in mitochondria) and a peripheral stalk made up of the band b subunits (Figure 11.2). The additional subunits in mammalian F I Fa-ATPases are mostly in the stalk regions (see Table I 1.1). The oligomycin-sensitivity conferring protein (OSCP subunit) in the mitochondrial peripheral stalk is equivalent to the bacterial D subunit. The b subunit in mitochondria has the role of the £ subunit in bacteria, and the small mitochondrial £ subunit (only 50 amino acids) at the foot of the central stalk has no counterpart in bacteria or chloroplasts. The overall shape of F I assembled with the c subunits of Fa is shown beautifully in the low-resolution images of the yeast mitochondrial FI-c complex published in 1999 (Figure 11.3). Subunit Structure and Function

F, Domain In all F I Fo-ATPases, the F I domain has alternating <X and j3 subunits forming a spherical hexamer, like sections of an orange, 'with a cavity in the center (Figure 11.4). The <X and j3 subunits of E. coli have 20% sequence identity and very similar folds. Both can bind nucleotides, but the catalytic sites are located on the j3 subunits at the <x/j3 interfaces. As described below, the three active sites are in three different conformations,

273

TABLE 11.1. Equivalent subunits in ATP synthases in bacteria, chloroplasts, and bovine mitochondria Type

Bacteria

Chloroplasts

Mitochondria a

F,

ex

ex

ex

(3

(3

(3

y

y

6

6

y OSCpb

€

€

6

a b c

a (or x) band b' (or I and II) c (or III)

a (or ATPase 6) b c

€

Fa

, Mitochondrial ATP synthase has additional subunits that have no equivalents in bacteria or chloroplasts. b Oligomycin-sensitivity conferring protein. From Walker, J. E, ATP synthesis by rotary catalysis (Nobel lecture). Angew Chem Int Ed. 1998,37:2308-2319.

depending on whether they bind ATP (the loosebinding TP conformation, partly open), or ADP (the tight-binding DP conformation, closed), or are empty (the E conformation, open). The y subunit has very long N- and C-terminal helices, 'which form an antiparallel <x-helical coiled coil that traverses the central cavity of the <Xj3 hexamer and continues as the central stalk to the Fa domain. The two long helical domains of yare connected by a globular domain that protrudes at the base of the stalk. The globular domain of y, visible in the higher-resolution structure of the complete bovine mitochondrial F,-ATPase obtained in 2000, has an <X!j3 structure that wraps around the other two helices

11.3. Low-resolution x-ray structure of the F, Fa-ATPase from yeast mitochondria. The electron density map of the F,-ClO complex from Saccharomyces cerevisiae at 3.9 A resolution shows the architecture of the complex from the side (A) and the cytoplasmic end (B), with insets identifying the subunits in A and numbering the c subunits in B. Note the asymmetry in B. From Stock, D., et al., Curr Opin Struct Bioi. 2000, 10:672-679. © 2000 by Elsevier. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

274

A.

B.

11.4. The high-resolution structures of the ex, 13, and y subunits of the F,-ATPase from bovine mitochondria. The ribbon diagrams show ex (red), 13 (yellow), and y (blue) subunits as identified in the schematic drawing accompanying each figure. A. The entire F, particle shows the coiled coil of the y subunit in the central cavity between the ex and 13 subunits, with bound nucleotides (black ball-and-stick representation). The exl3 pairs are labeled E for empty, TP for binding ATP, and DP for binding ADP, as described in the text. B. A cutaway view showing only three subunits: CXTP, y, and I3DP (shaded in the diagram above the structure). From Walker, J. E., Angew Chern tnt Ed. 1998,37:2308-2319. © 1998 by Gesellschah Deutscher Chemiker. Used by permission of Wiley-VCH Verlag GmbH.

and contacts 8 and [ subunits (Figure 11.5). The position of the y subunit is asymmetric with respect to the cxl3 hexamer, which is important in the mechanism. Rotation of y, which unwinds the lower part of the coiled coil (see Figure 11.6A), is driven in one direction by hydrolysis of ATP and is driven in the opposite direction by the pmE. At the base of the y subunit where it contacts Fa is the [ subunit (8 in mitochondria), which has two domains, an N-terminal l3-sandwich and a helix-turnhelix C terminal. The N-terminal domain of [ makes contact with Fa and is essential for coupling. The [ subunit appears in very different positions in the E. coli and mitochondrial structures (Figure 11.6), and it also exhibits different conformations when crystallized as a pure protein and in complex with a portion of the y subunit. The closed conformation observed in isolated [ is similar to the conformation seen in the mitochondrial structure, but the open, partially unfolded conformation observed in complex with y is unlike that seen previously and suggests considerable flexibility in the structure (Figure 11.7). Cross-linking studies pro-

vide evidence for interactions between [ and 13 subunits that differdependingon which nucleotides are bound to F I , suggesting the flexible [subunit might have a regulatory function (see below). Data from cross-linking studies indicate that [ rotates with y and c, because both ATP synthesis and ATP hydrolysis coupled to proton pumping occur \",hen all three are covalently linked (see Figure 1J .2). The 8 subunit is required to bind F, to Fa, hence its historic designation in mitochondria as the oligomycin sensitivity-conferring protein. With the b 2 dimer from Fa ,8 forms the peripheral stalk that appears to hold the CXJ I3J knob in place while y rotates. For this reason b 2 8 has been called a stator, the stationary part of a machine in which a rotor revolves. As will be seen below, the rotor includes the central stalk and the ring of c subunits in the Fa domain of the enzyme.

Fa Domain The Fa structural domain consists of a ring of c subunits connected via the a subunit to the b dimer that forms the peripheral stalk. NMR studies indicate that

F, Fo-ATPase/ATP Synthase

275

11.5. The complete structure of the F,-ATPase from bovine mitochondria. The structure of the central stalk is visible in the x-ray structure of the complete F, domain obtained at 2.4 A resolution. A. The ribbon diagram shows the C( and 13 subunits (red and yellow, respectively) alternating around the y subunit (blue) with the y (green) and £ (magenta) at the base of the central stalk. B. Model of the mitochondrial ATP synthase based on EM data for bovine and yeast structures. Mitochondrial subunit names are different from those in the E. coli structure (see Table 11.1). In particular, the IS subunit is equivalent to the £. subunit in E. coli, and d along with six copies of F interact with the b subunits. From Stock, D., et al., Curr Opin Struct BioI. 2000, 10:672-679. © 2000 by Elsevier. Reprinted with permission from Elsevier.

the sm"l1 (8000 Da) c subunit forms two hydrophobic TM helices that make a coiled coil connected by a short polar loop. The number of c subunits varies, with J 2 in E. coli, 14 in chloroplasts, and 10 in bovine mitochondria. The c subunits form two concentric rings, with their N-terminal helices forming the inner ring and their C-terminal helices forming the outer ring (see Figure 11.3). The ring of c subunits is contacted by [ and y at the central stalk from F I , and half the polar loops contact the b subunit. An essential residue involved in proton uptake through Fo is cAsp61 (E. coli numbering) in the center of the C-terminal helix. When Asp61 binds " proton, the C-termin,,1 helix undergoes a conformational change that is critical to rotation of the ring. The very hydrophobic a subunit, predicted to form five TM helices, connects the ring of c subunits to the b subunit of the st"tor. A critical ,-esidue in the a subunit is Arg21 0, which is essential for ATP-driven proton translocation but not for passive proton transport when F I has been removed. The interaction of aArg210 with cAsp61 indicates there is a direct connection between the a subunit and the ring of c subunits. Therefore sub-

unit a is thought to contribute to the proton channel, with protons entering the translocation pathway either at an interface between the a and c subunits or via the a subunit, foJJowed by transfer to the c subunit. The a subunit has other basic and acidic residues essential for proton translocation, which are His245, GJu196, and Glu219. The b subunit is usually a homodimer, although some bacterial species have a heterodimer of band b' instead. It is very elongated and anchored in the membrane by its hydrophobic N-terminal region. The rest of the protein is hydrophilic and highly charged, except for a short stretch of hydrophobic amino acids near the C terminus. The C terminus is required for assembly of the ATP synthase. According to its circular dichroism spectra, the b subunit is highly helical. In agreement with this result, NMR data indicate the N terminus is helic"l, with residues 4 to 22 predicted as a TM segment. The two b subunits are in such close proximity that they are linked by disulfide bonds when cysteine residues are engineered at many different positions in the b subunit.

Membrane Protein Assemblies

276

A.

C

B.

11.6. Different conformations observed in mitochondrial and bacterial structures of the central stalk. The arrangement of the y (red) N-terminal helix (thicker), the C-terminal helix (thinner), and the [ subunit (yellow) varies between the mitochondrial ATPase (A) and the E. coli enzyme (B). The blue arrow shows the large movement of c involving rotation of 810 and translation of 23 A. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002, 27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

A.

Regulation of the F 1 Fo-ATPase

The FIFo-ATPase is a highly efficient motol~ Attempts to determine its thermodynamic efficiency (the ratio of free energy gained by pumping protons to the free energy expended in the phosphorylation of ADP to make ATP) from the measured torque of the F 1 domain suggest an efficiency near 100%. What then is the purpose of regulation? Cells would benefit by alteration of the efficiency of F I Fa-ATPase because if ATP concentrations in respiring cells are high, decreasing the net rate of ATP synthesis is beneficial. whereas if ADP concentrations are high, the cell needs rapid ATP synthesis. Crosslinking studies suggest that regulation of the activity of FIFo-ATPase could (partly) depend on the flexibility of the £ subunit described above (see Figure 11.6). The rate of cross-linking between £ and f3 subunits is affected by ATP/ADP concentrations: in the presence of ATP this rate increases, whereas in the presence of ADP it decreases. Thus the position of the £ subunit varies depending on the needed net rate of ATP synthesis. Recent studies show that £ is able to sample alternate conformations on a timescale of seconds. Do the conformations themselves suggest a possible mechanism? The conformation of the £ subunit seen in the E. coli structure allows its C-terminal helices to wind around the y subunit and extend toward the <X3f33 hexamer, with the central axis of the f3-sandwich roughly parallel to the N- and C-terminal helices of the y subunit. The conformation observed in the mitochondrial

B.

11.7. Conformational changes in the £ subunit. A. The closed conformation observed for the isolated £ subunit of E. coli is docked onto a model of the c subunit ring (below) and the end of the y subunit above. B. The open conformation observed when [ is crystallized with a portion of the y subunit has the C-terminal helices separated from each other as well as from the ~-sandwich and wrapped around portions of y. From Bulygin, V. v., et al., J Bioi Chem. 2004, 279:35616-35621. © 2004 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

F, Fa-ATPase/ATP Synthase

277

ATP

l

ADP + Pi

T

1

11.8. Sequential stages in the binding-change mechanism. Each catalytic site of the ATP synthase can have one of three conformations, designated 0 for open (empty) state, T for tight (ATP-bound) state, and L for loose (ADP + P; occupied) state. The figure shows one iteration through the states, which is coupled to a 120" rotation of the yc central stalk. This is also called an alternating sites hypothesis as it involves cycling of each of the three catalytic sites through three states. From Capaldi, R. A., and R. Aggeler, Trends Biochem Sci. 2002,27: 154-160. © 2002 by Elsevier. Reprinted with permission from Elsevier.

structure has shifted the central axis of the l3-barrel to be roughly perpendicular to the N- and C-terminal helices of the y subunit, ,vith the C-terminal helices flattened against the side of the l3-sandwich and away from Fl. These two states have been proposed to work as a rachet to affect the catalytic efficiency of F I , and this is supported by cross-linking studies showing different results when nucleotides are bound to Fl. Cross-linking studies also show that diFferential effects on hydrolysis and synthesis activities are possible. For example, formation of a cross-link between the C terminus of subunit £ and they subunit inhibited ATP hydrolysis by 75% and ATP synthesis by 25%, indicating that in some conformations, ATP synthesis is allowed when ATP hydrolysis is not. Finally, cross-linking results indicate that subunit £ can span the region of the central stalk and interact simultaneously with a 13 subunit of F, and subunit c of Fa. Catalytic Mechanism of a Rotary Motor

Kinetic studies of the F, Fa-ATPase established a number of characteristics of its mechanism: (1) Isotope exchange data are consistent with the idea that the enzyme does not release ATP at one active site until substrate is available to bind at another active site. (2) The exchange of '80 showed that all three catalytic sites on the three 13 subunits are equally capable of carrying out the reaction. (3) Because release of products is rate Ii 01iting, the Pi and ADP that remain bound can reversibly resynthesize ATP, resulting in the incorporation of 18 0 in different positions when the reaction is carried out in H 2 '80. (4) The catalytic sites on each of the 13 subunits in F 1 differ in affinity for ATP (when measured in excess ATP so ATP binds to all three sites): Kd for the first is < 1 nM, for the second is ~ 1 ~lM, and for the third is 30 IJ.M. Clearly, substrate binding shows negative cooperativity. However, when more than one site is occupied, the rate of ATP hydrolysis goes up 104 _ to lOS-fold, so catalysis shows positive cooperativity. Whether this cooperativ-

ity requires nucleotide binding to two or three sites is still controversial. Alternating Site Mechanism The kinetic results are all consistent with the Boyer binding-change mechanism that states that each of the three binding sites in F, is in a different conformation, either closed (tight), partly open (loose), or open (and empty; Figure J 1.8). The site in the open conformation is ready to bind substrate. When it binds nucleotide it closes, triggering conformational changes in the other two so that the closed one becomes partly open and the partly open one becomes fully open. In the mechanism, each site alternates between the three states in a cycle for both forward and reverse reactions (see Figure 11.8). The reaction for ATP synthesis involves (J) binding of ADP and Pi to the partly open site; (2) conformational change at that site that converts it to the closed site, where catalysis occurs (while changing the other two sites to the open and loose sites); and (3) a second conformational change to convert that site to the open site that allows dissociation of the product. The rotating y subunit interacts with one of the 13 subunits to drive its conformational change from closed (tight) to open (empty), which in tum triggers the conformational changes in the other two subunits. In the full F I Fa-ATPase, these conformational changes in F 1 are coupled to proton translocation through Fa. For ATP synthesis, the rotation is generated by the passage of protons, driven by the pmf. through Fa to cross the membrane; in the reverse direction, energy released by hydrolysis of ATP drives the rotation in the opposite direction and reverses the flow of protons (Figure 11.9). In this rotary motor, the central rotor formed by the y and £ subunits of F I rotates by 120 for each ATP synthesized or hydrolyzed, bringing subunit y into contact with a different 13 subunit with each rotation. The central stalk connects to the ring of c subunits and rotates with them to couple catalysis to proton transport. 0

278

Membrane Protein Assemblies

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3'

4'

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11.9. Proposed substeps in the mechanisms of ATP synthesis and hydrolysis by F1 Fa-ATPase. The three catalytic sites of F1 (colored green, cyan, and light green) alternate between different conformations, which are labeled C, high-affinity catalytic site (previously called tight-binding site); D, ADP-binding site and D' for binding both ADP and Pi; T, ATP-binding site; and 0, low-affinity site. Steps for ATP synthesis are accompanied by clockwise (cw) rotation (top), while steps for ATP hydrolysis drive counterclockwise (ccw) rotation (bottom). Note that this model indicates binding to only two sites is sufficient, as supported by studies using beef heart mitochondrial F,; however, studies using prokaryotic F, suggest binding to all three sites is required for cooperativity. From Milgram, Y. M., and R. L. Cross, Proc Natl Acad Sci. 2005, 102:13831-13836. © 2005 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

A mechanism has been proposed for hmv proton translocation drives rotation. Protons enter a hydrophilic channel between TM segments of both a and c subunits. When each proton enters, it binds to residue Asp61 of a c subunit, triggering a conformational change in the C-terminal helix as described above. This also breaks interaction between Asp61 of the c subunit and Arg21 0 of the a subunit, releasing the c subunit to allow it to move. Fa rotates in smaller inCl-ements than the cx3133 domain, so coupling requires that several rotations of the c ring occur before cxl3 rotates. Also the number of c subunits in the ring varies, so proton flux through the ring that drives its rotation is not necessarily related by an integer to the 120 rotation of Flat each stage. (In other words, the number of protons per ATP is nonintegral.) This "symmetry mismatch" could be important to avoid a deeper energy minimum if the stages of both were closely matched. A tremendous amount of evidence supports this mechanism, including the aforementioned kinetic data, cross-linking results, and structures of increasing resolution and detail. Subunit structures and interactions have also been probed using epitope mapping with monoclonal antibodies, NMR, and FRET. Exciting support for the rotary mechanism came from video fluores0

cence microscopy using the attachment of fluorescently labeled actin filaments to the y subunits. With time, the fluorescent label rotated 360 in three steps of 120 (Figure 11.10). The rotation was quite slow, so more recent experiments used smaller gold particles to tag the y subunit, and a faster rate was observed. Careful analysis of these experiments has detected two substeps, one of ~90° attributed to ATP binding and one of ~30° for product release. Substeps are of interest because coupling can be broken down into several mechanochemical steps: ATP binding, transition state formation, bond cleavage, Pi release, and ADP release in the direction of hydrolysis. Recent computational studies of the ATP synthase have probed several aspects of this rotary machine. MD simulations designed to investigate the effect of the y subunit on the f3 subunits indicate that the effect of y is to open the binding sites, with a spontaneous closing motion. Calculations relate the different binding constants at the three sites to the rree energy for hydrolysis at the three si tes and model how the energy for rotation might be stored in confomlations of the subunits. The interplay between compu tational biology and structural biology \-vill certainly continue. A complete understanding of the mechanism of rotation and how it is coupled to catalysis in the F, Fa-ATPase needs higher-resolution 0

0

Complexes of the Respiratory Chain

279

A.

Actin filament

His-tag Coverslip coated with Ni-NTA

••••••• •• • B.

11.10. Demonstration of rotation coupled to ATP hydrolysIs by video fluorescence microscopy. A. Design of the experiment involved engineering 10 histidine residues onto the N terminus of each 13 subunit of F, to bind the F,-ATPase to a nickel plate and using biotin-streptavidin binding to attach to the y subunit an actin filament carrying a fluorescence label. B. The fluorescence pattern observed with the rotation axis in the middle of the filament when ATP is provided shows a continuous rotation of the actin in 120" increments, with a time interval between images of 33 msec. Scale bar is 5 ~lm. Redrawn from Noji, H, et aI., Nature. 1997,386:299-302. © 1997. Reprinted by permission of Macmillan Publishers Ltd.

structures of the entire complex in different conformational states.

COMPLEXES OF THE RESPIRATORY CHAIN

The ATP synthase carries out the last step of a multistep process that occurs in mitochondrial and bacterial membranes for the generation and utilization of an electrochemical gradient across the membrane, the proton motive force. Thus ATP synthase is complex V of the respiratory chain, whose other complexes utilize the electrons extracted from various reduced metabolites and transport them via coupled redox carriers to terminal acceptors, usually O 2 . The familiar electron transport chain consists of complex 1, which transfers electrons from NADH to ubiquinone; complex n, which transfers electrons from the FADH 2 of succinate dehydrogenase to ubiquinone; complex llI, which transfers electrons from ubiquinol to cytochrome c; and complex IV, which transfers electrons from cytochrome c to O 2 , fonning water (Figure 11.11). Additional enzymes feed electrons from other sources, such as G3P or acyl-CoA, into the chain, reducing ubiquinone to ubiquinol, so they all provide the substrate for complex In and in turn, complex IV. Three of the complexes (I, III, and IV) pump protons across the membrane to create an

-

electrochemical gradient. If the system is "uncoupled" by the action of compounds that allow protons to leak back across the membrane, electron transfer proceeds without creating or maintaining a proton gradient and the energy is dissipated as heat. When physically separated by membrane solubilization and ion exchange chromatography in detergents, the respiratory complexes retain their functions in vitro. This property allowed their extensive characterization by kinetic, spectroscopic, and electrochemical techniques over decades of research. By the end of the 1990s, when structures became available for respiratory com plexes isolated from both eukaryotic and prokaryotic organisms, the field moved toward a detailed understanding of the principles of electron-driven proton transfer. This section focuses on describing that process in light of the high-resolution structures of complexes III and IV. Cytochrome be1

The cytochrome-bcl complex is a dimeric muJtimer, in which each protomer contains up to J J protein subunits. Three of the subunits are ca talytic, as they cOlltain the redox centers and make up the functional unit (Figure 11.12). As the name implies, this complex contains both b- and c-type cytochromes: cytochrome b has two

Membrane Protein Assemblies

280

2H+

eyl.

C

2H+ 3-4H+

~

NADH + H+

NAD+

-L:

~

Succinate

1. NADH dehydrogenase

Fumarate + H+

II. Succinate dehydrogenase

4H+ 2H+

III. Cytochromebc) complex

IV. Cytochrome-c oxidase

V. ATP synthase

11.11. Respiratory chain complexes of the E. coli and mitochondrial inner membrane. The L shape of complex I, NADH dehydrogenase, is based on low-resolution x-ray and EM structures. The highresolution structures of the remaining complexes are based on numerous x-ray structures. From Hosler, J. P., et al. Annu Rev Biochem. 2006, 75: 165-168. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

B.

11.12. Structure of the yeast cytochrome-bcl complex. A. The homodimer contains the catalytic subunits cytochrome b (blue), cytochrome c, (yellow), and the Rieske Fe/S protein (green) in addition to six additional subunits. The cofactors are shown in ball-and-stick form. The position of the membrane is indicated by the orange bars, with the intermembrane space (IMS) and matrix (MA) indicated. B. One functional unit has the three catalytic subunits, shown with transparent surface representation in the same orientation as the complete structure in A. The b- and c-type hemes (labeled) are buried. The primary sites are shown with white arrows, with a P-specific inhibitor, stigma tell in, bound to 0 0 and a quinone bound to 0; at the N center. From Hunte, C, et al., FEBS Lett. 2003, 545:39-46. rg 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Complexes of the Respiratory Chain

281

QH z + 2 cyl c++ 2H N+

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11.13. The mechanism of the 0 cycle illustrated on a functional unit of the chicken cytochrome-bcl complex. The cytochromeb (light green) and cytochrome-cl (yellow) subunits are shown with transparent surface representation, while the structure of the Rieske Fe/5 protein is represented by ribbon diagrams showing the two conformations (blue and red). The position of the membrane is indicated by dashed lines, with the intermembrane space labeled P-phase (for positive) and the matrix labeled N-phase (for negative). The [2Fe-25] center is represented by space-filling models in four positions, two showing the two conformations of the Rieske protein (blue and red) with two intermediate positions (yellow) to show the trajectory of its movement. The b-type hemes (blue) are near the quinone-binding sites, and the c-type heme (black) is not far from the docking site for cytochrome c. A quinone (green) binds to 0" and a quinone has been modeled to replace stigmatellin at 00 at the inhibitor binding sites (dotted light blue arrows). Electron transfers (small green arrows) and proton release and uptake (curved blue arrows) are indicated. In the reaction summarized in the text, two quinol molecules are oxidized to quinone at the 00 site, with two electron pairs following the divergent pathways (green arrows) to the [2Fe25] center, cytochrome C1 and cytochrome c and to the b·type hemes to reduce the quinone at the OJ site. From Crofts, A. R., Rev Physiol. 2004, 66:689-733. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

b-type hemes, called b H and b L , and cytochrome CI has a c-type heme (a-, b-, and c-type hemes differ in their substituents off the tetrapyrrole ring and their linkages to the proteins). In addition to the two cytochromes, the catalytic unit has an iron-sulfur protein of the Rieske type, which is a [2Fe-2S] cluster in which one Fe is coordinated by two histidine residues and the other by two cysteine residues. The redox sites in these three subunits are buried in the membrane interior (see Figure 11.12B) because their quinone substrates are very hydrophobic, with isoprenoid side chains 30 to 50 carbons in length. In addi tion, redox reactions of quinones involve a semiquinone intermediate that needs to be shielded from the aqueous environment to avoid formation of damaging reactive oxygen species.

The Q Cycle Unlike other proteins that pump protons, such as bacteriorhodopsin (see Chapter 5) or cytochrome oxidase (see next section), the cytochrome-bcl complex does not provide a direct proton path across the membrane. In the overall reaction, two electrons from ubiquinol reduce two molecules of cytochrome C with uptake of two protons from the matrix (or cytosol in bacteria) and release of four protons to the intermembrane space (or outside in bacteria). This is accomplished by a special mechanism called the Q cycle, which uses two quinonebinding sites. The Qi site is closer to the inner surface and with heme b H forms the N center (for negative side), and the Qo site is closer to the outer surface and with heme b L forms the P center (for positive side). Quinone reduction takes place at Qj, quinol oxidation takes place at Qo , protons are taken up at Q i and exit from Qo , and electrons are cycled between quinoJ and the hemes before delivery to cytochrome c at the outer surface (Figure 11.13). In brief, the first ubiquinol binds to the P center and is oxidized, with its two electrons taking divergent paths. One electron is transferred to the Rieske Fe/S center, and [Tom there to cytochrome c\ and then to cytochrome c. The other is transferred to the b L heme, and from it to the b H heme and then to a ubiquinone that binds to the Qi site at the N center, making ubisemiquinone. A second ubiquinol binds to the P center, and again the two electrons follow these different paths, resulting in the reduction of a second cytochrome c and the reduction of the semiquinone to ubiquinol at the N center. Each oxidation of ubiquinol releases two protons, and two protons are taken up from the matrix for reduction of ubiquinone at the N center. Evidence for the Q cycle includes the ability of the inhibitor antimycin, which binds to the Q i site, to inhibit oxidation of heme b H and stop all electron transfer to cytochrome c. High-Resolution Structures The x-ray structures of bovine, chicken, and yeast cytochrome-bcl complexes all show a symmetric, pearshaped dimer that protrudes from the membrane in both directions, ~75 A into the matrix on the inside and ~35 A into the intramembrane space on the outside (see Figure 11.12). The position of the lipid bilayer is clear because of the bound phospholipids (see Chapter 8). More than half the mass of the complex is in the matrix portion, including the misnamed subunits Core I and Core 2 that function in mitochondrial peptide processing. Each protomer has eight TM helices from cytochrome b and one each from cytochrome CJ and the Fe/S protein, along with two or three additional TM helices from small subunits that surround the functiona I unit (Figure 11.14). The Fe/S protein has a hinge between an extrinsic domnin nnd the TM domain containing the metal center, allowing the [2Fe-2S] center

Membrane Protein Assemblies

282

p'

H"

QCRS·"

A.

~B~i

F 11.14. TM helices of cytochrome-bcl homodimer. Segments of one subunit are designated with asterisks. The TM helices from cytochrome b (red) are labeled A to H, those from cytochrome c, (yellow) are labeled CYT1, and those from the Rieske Fe/S protein (green) are labeled RIP1. The additional TM helices from small subunits OCR8 and 9 (blue and gray) are also shown. The dimer interface has two large hydrophobic clefts, labeled CFT. The hemes and the headgroups of the quinone (behind helix A) and stigmatellin (between B and C) are shown in ball-and-stick models. From Hunte, C, et aI., Structure. 2000, 8:669-684. @ 2000 by Elsevier. Reprinted with permission from Elsevier.

to be shuttled bet,,veen cytochrome b and cytochrome c (see Figure 11.13). The conformational change around this hinge rotates the extrinsic domain 60" and moves the [2Fe-2S] cluster 16 A. Mutations that limit the hinge movement lower the activity of the complex. The crystallization of the yeast cytochrome-bel complex in tbe presence of substrate and inhibitors has provided insight into the mechanisms of electron transfer and proton conduction. In the crystal structures one molecule of ubiquinone is bound to Q; at the N center. At the Q o site, the inhibitor stigmatellin binds in the same position that ubiquinone is expected to bind, while the inhibitor 5-n-heptyl-6-hydroxy-4,7-dioxobenzothiazole (HHDBT) is a hydroxyquinone anion that resembles an intelmediate step of ubiquinol oxidation (Figure 11.15). Critical residues at the P center include Hisl81 on the Rieske Fe/S protein and Glu272 of cytochrome b. In the first step of the Q cycle ubiquinol is hydrogen bonded to both of these residues to form an electron donor complex, which allows essentially simultaneous electron transfer to the Rieske cluster and the b L heme. Oxidation of ubiquinol to ubiquinone breaks this complex, allowing the ubiquinone to diffuse out to the medium or possibly to the N center of the opposite protomer. 1n addition, formation of the complex increases the pKa on the imidazole nitrogen of Hisl81, allowing it to take up a proton from ubiquinol; when the complex dissipates and the Rieske Fe/S center moves away, the pKa is lowered and the proton is released. The second proton [Tom the ubiquinol is transferred to Glu272 after electron transfer to the b L heme (Figure 11.16).

B.

11.15. Binding of different inhibitors to the 0 0 site of yeast cytochrome bc,. A. Electron density fitted to the structure of stig· matellin at the 0 0 binding site between Glu272 and the 12Fe-2SI center of the Rieske protein coordinated by His181 and His161. Hydrogen bonds are shown to the carbonyl oxygen (04) and hydroxyl group (08) of stigmatellin. From Hunte, C, et aI., Structure. 2000, 8:669-684. © 2000 by Elsevier. Reprinted with permission from Elsevier. B. Electron density fitted to the structure of the inhibitor 5-n-heptyl-6-hydroxy-4, 7-dioxobenzothiazole (HHDBT), whose structure is given in the inset. Residues that stabilize the binding are labeled, and hydrogen bonds to the carbonyl oxygen (04) and the deprotonated hydroxyl oxygen (06) are shown. From Palsdottir, H., et aI., j Bioi Chem. 2003, 278:31303-31311. © 2003 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Complexes of the Respiratory Chain

283

11.16. A model of electron and proton transfer at the P center of the yeast cytochrome-bcl complex viewed from the membrane interior A. Ubiquinol is hydrogen bonded to His 181 of the Rieske Fe/S protein (residues 160-175 and 178-182 are represented as yellow strands) and to Glu272 of cytochrome b (residues 75-85 and 265-275 are represented as cyan ribbons) with the lengths of the hydrogen bonds given in angstroms. The side chains of His181, Glu272, and Arg79 are shown as stick models (carbon, green; oxygen, red; nitrogen, blue), along with the ubiquinol and the b L heme. This conformation is observed in stigmatellin-bound cytochrome bCI, with ubiquinol modeled to replace stigmatellin. B. The structure in cytochrome bC1 complexed with the inhibitor HHDBT, a hydroxyquinone anion inhibitor that resembles an intermediate step of ubiquinol oxidation, shows movement of the Glu272 toward the b L heme. From Hunte, c., et aI., FEB$ Lett. 2003, 54539-46.

11.17. The suggested pathways for proton uptake at the N center of the yeast cytochrome-bcl complex. Two distinct pathways connect the solvent at the matrix side with the quinone-binding pocket at the N center. The E/R pathway (right side of figure) has an entrance at Glu52 of the Ocr7 subunit and is gated by Arg218 (yellow side chain with blue nitrogen atoms). The cardiolipin/K pathway has a cardiolipin (CL, green) positioned at its entrance and is gated by Lys228 (yellow side chain with blue nitrogen atom). Water molecules (red spheres) are associated with charged residues, and hydrogen bond interactions (dotted and dashed lines) are indicated. The arrows indicate the access sites from the bulk solvent, and double-headed arrows indicate proton transfer between the residues and the ubiquinone (U06, cyan). From Hunte, c., et al., FEBS Lett. 2003, 545:39-46. © 2003 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

284 cyt. c

N-side

4W

I I

4W

Iran~localion ~ub~tratc

11.18. Schematic diagram showing the paths of proton and electron transfer in cytochrome-c oxidase. The chemical reaction of 02 reduction to water (blue arrows) is coupled with the translocation of four protons (red arrows). Electrons flow from reduced cytochrome c in the intermembrane space (P-side) to the CUA center, and from there to heme a and then to the heme a3-CuB center, where they reduce oxygen. Protons from the matrix are either shuttled to the heme a3-CuB site and consumed in the production of water (substrate protons) or are translocated across the membrane. From Wikstrom, M., Biochim Biophys Acta. 2004, 1655:241-247. © 2004 by Elsevier. Reprinted with permission from Elsevier.

Rotation of protonated Glu272 allows it to hydrogen bond with a water molecule that is hydrogen bonded to a heme propionate side chain. From there the proton is released to the surface via a hydrogen-bonded water chain associated with four charged residues of cytochrome b, Arg79, Asn256, Glu66, and Arg70. While protons are released from the P center of cytochrome bCI, they are taken up at the N centel~ The x-ray structure of the yeast complex reveals two distinct pathways for proton uptake, called the E/R pathway and the cardiolipin/K pathway (Figure J 1.17). The two pathways are located at either end of the substrate-binding site, indicating a quinone can be reduced on both ends of the molecule without changing positions. The E/R path runs from Glu52 of one of the minor subunits and is gated by Arg218 of cytochrome b. A molecule of cardiolipin is at the entrance of the other path, which is gated by Lys228 of cytochrome b. Upon reduction of ubiquinone, a proton is abstracted from each of the gating residues (Arg218 and Lys228) and is replenished with a proton [Tom the malLix via hydrogen-bonded networks to those residues. The structures have provided an elegant explanation for the coupling of proton movements to electron transfer.

Cytochrome-c Oxidase Complex IV of the respiratory chain, cytochrome-c oxidase, carries out the reduction of O 2 to H 2 0 using four electrons coming from four molecules of reduced

B.

11.19. x-ray structure of cytochrome-c oxidase. The crystal structure of cytochrome-c oxidase from Rb. sphaeroides was determined at 2.3 A resolution. The subunits are closely packed: subunit I (green), subunit II (light gray), subunit III (dark gray), and subunit IV (magenta). The redox centers include heme a (light blue), heme a3 (red), and CUA and CUB (dark blue), and have Mg (light green), calcium (pink). lipids (orange). and water molecules (red spheres) A. Ribbon diagram shows the structure viewed from the side. B. The TM helices in cytochrome-c oxidase are viewed from the P-side. From Svensson-Ek, M., et aI., J Mol BioI. 2002, 321 ;329-339. © 2002 by Elsevier. Reprinted with permission from Elsevier.

Complexes of the Respiratory Chain

285

cytochrome c while pumping a total of eight protons. four "substrate" protons that combine with the oxygen atoms and four "pumped" or "vectorial" protons that contribute to the electrochemical gradient. Like complex III. complex IV is a dimeric multimer with the prokaryotic protomercontaining three to foursubunits, while in eukaryotes it has eight to 13 subunits. AJI the redox centers are contained in subunits I and II: two a-type hemes called a and a3. and two Cu-containing centers, CUA, which has two copper ions. and CUB' In addition, the complex has a Mg 2+ ion that is not involved in redox and a site for binding Ca 2+ or Na+. Spectroscopic studies revealed the path of electron transfer is from cytochrome c to CUA to heme a, then to a binuclear complex of heme a3 and CUB. and finally to O 2 (Figure 11.18). The four-electron reduction of O 2 must occur by accumulating reduced intermediates without their release as reactive oxygen species. High-Resolution Structures

Crystal structures have been obtained for cytochrome-c oxidase [Tom Paracoccus denitrifrcans, from Rhodobactersphaeroides, and from bovine mitochondria. In these stmctures. both bacterial complexes have four subunits and the bovine complex has 13 subunits, of which subunits I. If, and III are very similar to the corresponding subunits in the other two. The dimeric complex has an ellipsoid shape that protrudes beyond the lipid bilayer 32 A into the intermembrane space and 37 A into the matrix (Figure I \.19). The proteins cross the membrane with 21 to 28 TM helices: 12 [Tom subunit I. two h-om subunit II. and seven from subunit III. The seven additional subunits in the bovine complex are type U membrane proteins (each with a single TM helix and the N terminal inside) thaI together surround subunits I. II. and HT. Six phospholipid molecules are resolved in the yeast structure. Both heme a and the bimetallic heme a3/CuB center are buried in the membrane interior of subunit I, ~13 A from the outer surface. The CUA center has two cystei ne residues as ligands to the two copper ions (analogous to [2Fe-2SJ) and binds to a globular domain of subunit II on the outside surface. This globular domain meets a corner of subunit I on the surface where cytochrome c likely binds, as there are 10 acidic residues that can interact with the ring of lysine residues on cytochrome c (see Figure 4.2).

11.20. Proton uptake pathways in yeast cytochrome-c oxidase. The ribbon diagram shows subunits I, II, and III with heme a and a3 (green) as stick structures, with Ca and Mg metals (green spheres) and Cu metals (orange spheres). The two paths for water molecules are the D path (red) and the K path (blue). with water molecules colored accordingly. From Hosler, J. P., et al.. Annu Rev Biochem. 2006, 75:165-187. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

[Tom time-resolved spectroscopic studies. indicate that O 2 binds first to the Fe 2 + of heme a3 and quickly picks up two electrons from the Fe 2+ and the CUB 1+ to form a peroxide bridge between Fe 3+ and CUB 2+. The next intermediate is an unstable hydroperoxo compound - Fe 3+OOH - that is quickly cleaved to the oxoferro state Fe4+ = 0 with CUB 2+ -OH-. By now, four electrons have been transferred. three from the bimetal center and the fourth probably from Tyr288, creating a tyrosyl radical. The next two steps are slower, limited by the time it takes the protons to reach the binuclear center. A proton reacts with the OH- on CUB to release the first water; then two electrons and two protons react with the 0 2 -- on Fe 4 + of heme a to release the second water. The electrons are supplied by additional molecules of cytochrome c operating through CUA and heme a.

Oxygen Reduction

The reaction can be viewed as starting with a metal reduction phase, when each cytochrome c molecule docks on the surface of cytochrome-c oxidase and transfers its electron to CUA. The electrons are then transferred to heme a and then to heme a3/CuB, where the reduction of O 2 takes place. The characteristics of the O2 reduction site with heme a3 and CUB have been determined in crystals of both fully oxidized and fully reduced enzyme. These structures, along with results

Proton Pathways

Protons from the matrix utilize two uptake pathways to the bimetallic center that have been identified in the structures of cytochrome-c oxidase. As observed in bacteriorhodopsin (see Chapter 5), a proton pathway consists of a series of hydrogen-bonded water molecules linked to residues that are essential for pumping protons. The D pathway goes from Asp132 on the surface to Glu286 between heme a and heme a3, a distance of

Membrane Protein Assemblies

286

Mg.

11.21. Molecular dynamics simulation of a proton exit pathway in yeast cytochrome-c oxidase. The simulaton of the Rb. sphaeroides cytochrome-c oxidase structure in added water lasted over a nanosecond and revealed a chain of hydrogenbonded water molecules from Glu286 to the M g 2+ ion. From Hosler, J. P., et al., Annu Rev Biochem. 2006, 75: 165-187. © 2006 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

~26 A, and the K path\vay begins at GlulOI on subunit II and passes in sequence Ser299, Lys362, Thr359, a farnesyl side group of heme a3, and Tyr288 of subunitl (Figure 11.20). Either the D or K pathway is used to conduct "substrate" protons to the heme a3/CuS site. Since genetic alteration of the D pathway eliminates proton pumping, the "pumped" protons use only the D pathway. The role of subunit III appears to be a proton antenna, as it has many chal-ged residues on the surface. When subunit III is Jacking, the rate of proton uptake into the D pathway is I-educed 50%. While a great deal of evidence supports the roles of the D and K pathways for proton uptake, the exit pathway for "pumped" protons is not clear from GJu286 to the outside. However, MD simulations of a singlefile column of water molecules suggest a path from Glu286 through a hydrophobic cavity to the Mg ion (Figure 11.2 J ). Add itional experimental and theoretical work is needed to define the exit pathway. A number of othel- questions are being explored, such as what controls the directionality of proton pumping) Are there mechanisms to regulate the efficiency of cytochrome-c oxidase? Yet tremendous progress has been made in understanding the mechanism of this vital enzyme complex and in general in addressing the nature of electron transfer coupled with proton pumping.

THE TRANSLOCON

The translocon (also called the translocase and the protein-conducting channel [PCC]) is a complex of

membrane proteins that functions to transport proteins across membranes as well as to incorporate proteins into the membrane (see Chapter 7). It has a wellconserved core of integral membrane proteins that partner with various peripheral proteins and chaperones. Since many proteins are translocated as they are synthesized, it also partners with the ribosome. The structures of the translocons from canine ER, yeast, and E. coli have been imaged with cryo-EM of twodimensional crystals. After a great deal of work, the first high-resolution x-ray structure for a translocon, that from the archaea Methanococcus jal1l1.Qschii, was solved at 3.2 A resolution. With this detailed structure as a model, the lower-resolu tion structure of the E. coli translocon with an actively translating ribosome could be analyzed. The translocon in bacteria, eukaryotes, and archaea are heterotrimers of similar composition. Two of the three proteins (corresponding to SecYE in bacteria) are essential for viability, while the third (SecG) stimulates translocation as well as the ATPase activity of SecA but is not essential under normal conditions of cell growth. Additional protein partners in E. coli include SecA, the motor for posttranslational translocation; YidC, a protein involved in insertion of hydrophobic TM segments into the bilayer; and the SecDFYajC heterotrimer that facilitates protein translocation in an unknown mechanism (see Chapter 7). The M. jannaschii Translocon Structure

The crystal structure of M. jal111a.schii SecY (X(3y greatly advanced understanding of the structure of the heterotrimeric translocon and suggested how it could move proteins both across the membrane and lateraUy into the bilayer. The (X subunit has JO helices positioned in the membrane to form a sort of balTel with a (-ectanguJar shape when viewed from the cytosol and a pseudo-symmetry between two halves formed by TM IS and TM6-10 (Figure 11.22). The loop between TM5 and TM6 connecting the two halves is proposed to act as a hinge at the back side of the rectangle. Many of the helices al-e tilted up to 35° from the bilayer normal, conU-ibuting to the funnel shape of the central cavity. Not all the helices span the bilayer fully, and in particular TM2a extends only halfway through the membrane and is only partially hydrophobic. Since it is bordered on three sides by the (3 and y subunits, the only lateral opening in the rectangular SecY complex is on the side opposite the proposed hinge, gated by TM2 and TM7. The nonessential (3 subu nit, at the side near the N terminus of the (X subunit, makes only limited contact with the (X subunit; its single TM he.lix is close to the C terminus of the y subunit at the external side of the membrane. The y subunit makes a girdle, or band, along two sides of the rectangle.It consists of two helices: an amphipathic helix that

The Translocon

287

A.

11.22. Structure of the M. jannaschii translocon viewed from the cytosol. The "front" is on the left, across from the hinge at the back made by TMS and TM6 on the right. A. The ex subunit is colored blue to red from the N to the C terminus, with the TM segments numbered; the 13 subunit is shown in pink, and the y subunit in purple. B. The structure is now colored to highlight the symmetry in the ex subunit, with the N-terminal half in blue and the C-terminal half in red. From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

lies on the cytoplasmic surface and a long, curved helix that runs along the back side of the ex subunit. With the f3 subunit on the third side, the two halves of the ex subunit can open only on the front like a clam shell. The M. jannaschii structure can be superimposed on the EM structure of E. coli translocon (SecYE) at 8 Aresolution with a good match of the TM ex-helices (Figure 11.23). This first high-resolution structure provided a number of insights into the function of the translocon and suggested it could function as a monomer (see below) with a singJe channel down the center. The channel is shaped like an hourglass, with funnel shapes above and below its central constriction (Figure 11.24). The opening of the constriction is only 3 Ain diameter and is lined by a ring of hydrophobic and inflexible lIe side chains. When cysteine residues were engineered at 30 positions throughout SecY, only the cysteines in the region of the central constriction made a disulfide bridge with a cysteine on a translocating polypeptide, giving strong evidence that this is the pore. However, the smalJ diameter of the constriction means that other TM helices would have to shift to open the channel enough to alJow passage of an extended peptide having a maximal width of around 12 A or an (X-helical peptide having a width of 14 A; this may happen dynamically as a peptide is passing through the channel to accommodate the peptide without a leak. TM2a is proposed to be a plug that closes the channel. Indeed, when the TM2a segment in E. coli is locked by an engineered disulfide bond to the y (SecE) subunit, the channel stays open. This result implies that TM2a moves 22 A to open the channel (see Figure 11.248).

11.23. Superposition of the x-ray structure of the M. jannaschii translocon on the cryo-EM structure of the E. coli translocon. The M. jannaschii x-ray structure (numbered helices, colored as in previous figure) was visually docked onto the electron density map of the E. coli SecY complex from cryo-EM (light blue). The labeled gray cylinders represent TM helices in E. coli that have no correspondence in M. jannaschii. The diamond indicates the axis of twofold symmetry in the E. coli complex. From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Membrane Protein Assemblies

288

A.

11.24. The channel pore in the M. jannaschii translocon. A. The channel is in the center of the translocon when viewed from the top. as in Figure 11.22. Half-helix TM2a (green) acts as the channel plug. B. The side view shows the hourglass shape of the pore with constrictions at the pore ring formed by three lie residues (gold). The arrow shows the modeled movement of the plug (green) toward the y subunit (magenta). From van den Berg, B., et aI., Nature. 2004, 427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

TM Insertion

The x-ray structure also suggests how a TM segment of the nascent protein could be released into the lipid bilayer. The "front" of the lX subunit is the only possible lateral opening, since the other three sides are closed by the 13 and y subunits. A hinge movement between TM5 and TM6 at the back would open the [Tont to allow the peptide access to the bilayer (Figure 11.25). A dynamic fluctuation of this hinge movement would

allow the peptide inside the channel to partition into the lipid if it were sufficiently nonpolal~ In addition to overall hydrophobicity, the positions of some residues influence this lateral partitioning, according to studies with model peptides (see Chapters 4 and 7). While the x-ray stl-ucture shows a central pore in the translocon from M. jQIlI1Qschii, suggesting it is active as a single copy of the SecY heterotrimer, there is substantial evidence for higher-order mul timers of the Sec translocons. The SecYEG complex purified h'om E. coli exists as oligomers. The cryo-EM images from twodimensional crystals formed by slow detergent removal in the presence of phospholipids clearly show dimers of SecYEG (see Figure 11.23). Translocon assembly into dimers, or even oligomers, could be a dynamic process that is triggered by interactions with other partners, such as the ribosome or SecA in E. coli. Cryo-EM has now been used to investigate how the SecYEG translocon might interact with the ribosome.

The Translocon-Ribosome Complex

11.25. Proposed lateral gate of the M. jannaschii translocon. The TM segment of the nascent protein is depicted as a magenta cylinder, and its movement into the lipid bilayer is indicated by the arrow to the left. This is postulated to involve an opening between TM helices 7 and 8 on one side and TM2b and TM3 on the other, resulting from the hinge at TM5 and 6 as shown by the arrow on the right. From van den Berg, B., et aI., Nature. 2004,427:36-44. © 2004. Reprinted by permission of Macmillan Publishers Ltd.

Images of the canine translocon bound to a ribosome achieved with cryo-EM show a larger assembly of translocons with a diameter of ~ 100 'A, which fits a tetra mer of Sec61 heterotrimers. The ribosometranslocon assembly shows a central depression between the four heterotrimers that is believed to be filled with lipids, in contradiction to the proposed aqueous pore. The ribosome exit tunnel is nearly centered over the tetra mer, but the contacts between the ribosome and the translocon indicate that only two of the Sec61 heterotrimers have access to the tunnel.

The Translocon

289

3' mRNA entrance

11.26. Cryo-EM image of the complex of ribosome and translocons from E. coli. The small (30S, yellow) and large (SOS, blue) ribosomes are shown with the A, P, and E sites (magenta, green, and orange), and the L7/L12 stalk is labeled. The mRNA (cyan) is visible and so is the nascent chain (gold) in the exit tunnel. The translocating SecYEG dimer (dark blue) is at the exit tunnel, while the nontranslocating SecYEG (red) is at the S'mRNA exit. From Mitra, K., et aI., Nature. 2005. 438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

High-resolution EM studies of a translocating SecYEG complex from E. coli with a translating ribosome show a complex of two Sec heterotrimers in a front-to-front orientation and also suggest different conformations for translocating and nontranslocating complexes. These studies were performed with detergent-solubilized SecYEG added to a cell-free system of E. coli ribosomes translating an mRNA that encodes a chimeric protein with the signal-anchor of

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Contacts with the Ribosome The contacts between the translocating SecYEG dimer and the ribosome occur at three regions, called Cl, C2, and C3. They involve both ribosomal proteins and rRNA from the ribosome and two large cytoplasmic loops (between TM6 and 7 and between TM8 and 9) of SecY

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FtsQ fused to a stalling domain of SecM, which slowed it sufficiently to allow a snapshot of the translocation process by cryo-EM. Computer modeling was used to fit a homology-based atomic model of the E. coli translocon derived from the 1vl. jannaschii structure, along with the detailed structure of the ribosome, to the 11 A resolution EM images. This complex shows two assemblies of translocons (PCCs) associated with the ribosome (Figure 11.26). One is associated with the exit pore of the large subunit and is apparently involved with the movement of the nascent peptide. The other one is nonphysiologically bound to mRNA near its exit site in the small subunit and is thus considered to be in a nontranslocating state. The atomic coordinates of two SecYEG translocons in a front-to-front arrangement could be fitted to the nontranslocating structure. consistent with the inactive/closed state of the x-ray structure, and enabled analysis of the structural changes required to obtain the activeltranslocating state for the other one. The model of the translocating state shows sufficient opening of the SecY halves within the membrane plane to allow formation of a wider pore (Figure 11.27), However, the pore is not shared between the two monomers, and the two pores of the SecYEG heterotrimers seem to be in different states, one open to the bulk lipid and the other inaccessible to lipids,

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11.27. Nontranslocating and translocating SecYEG dimers in front-to-front orientation. The van der Waals surface representations obtained by fitting to the nontranslocating (A) and translocating (B) electron density observed by EM are shown with the SecY C-terminal halves transparent. The green arrow indicates the change in the heterotrimer interface at the front, and the yellow arrows point out the changes in the opening of SecY. One heterotrimer is blue/green and the other is in shades of red. The ribosomal side is behind the plane of the membrane. From Mitra. K., et aI., Nature. 2005, 438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd .

Membrane Protein Assemblies

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11.28. Path of the nascent chain through the ribosome and translocon. A. The ribosome bound to a front-to-front SecYEG dimer is viewed from the side with a nascent chain (yellow) in the exit channel. Only the molecules from the ribosome that contact the translocon are shown. B. A schematic version of the molecules involved, with the rest of the ribosome and the membrane added. The nascent chain is yellow in the ribosome exit tunnel and the translocon pore is green. From Mitra, K., et aI., Nature. 2005,438:318-324. © 2005. Reprinted by permission of Macmillan Publishers Ltd.

(Figure 11.28). At Cl the rRNA helix 59 contacts one SecY, and at C2 the rRNA helix 24 contacts the other SecY. At C3 the proteins L29 and L23 from the ribosome make contact with the cytoplasmic region of SecG and possibly the N-terminaJ part of SecE in a nonessential but stabilizing connection. Since C3 is at the back, there is a large opening at the front, providing space between the translocon and the ribosome that is accessible to the cytoplasm. This access to the cytoplasm means the ribosome does not plug the translocon pore to prevent leakage, as earlier envisioned; thus the translocon itself must be capable of providing a tight seal to maintain the permeability barrier. Structures of the translocon with and without the presence of a ribosome-nascent chain complex answer many questions about protein export and leave many others unanswered (see Chapter 7). It is still not clear how the N-terminal portion of the nascent chain inserts into the translocon, presumably as a hairpin in the initial step. How does a TM segment reorient inside the transJocon, as indicated in studies of topogenesis? The dynamic interaction with SecA, presumed to include its insertion into the translocon, is not understood. What is the relation to other proteins that assemble at the translocon, such as SeeD, SecF, and YajC? StilJ other proteins, such as signal or leader peptidase and oligosaccharide transferase (in eukaryotes), are present on the outside of the membrane as part of the export

process. Clearly, the translocon is at the center of an amazing molecular machine that carries out dynamic and complex processes.

ABC TRANSPORTERS AND BEYOND

ABC transporters carry out the uptake or efflux of a wide variety of substances at the expense of hydrolysis of ATP (see Chapter 6). All ABC transporters have four domains, two TM domains and two nudeotidebinding domains (NBDs) that are synthesized as one to four polypeptides (see Figure 6.6). A molecular understanding of this important class of transporters is provided by the x-ray structures of two ABC transporters, the Sav1866 protein and the BtuCD complex. Many ABC transporters work in tandem with otber proteins to facilitate uptake or efflux across the two membranes and the space between them in the cell envelope of Gram-negative bacteria. The other components working with BtuCD in E. coli vitamin B I2 uptake include two other specific proteins 'whose structu res have been solved (BtuB and BtuF) and three less understood proteins (TonB, ExbB, and ExbD) required for energy coupling from the inner to the outer membranes. After a description of this complex system, the chapter ends with the structure and function of the Sav 1866 protein as well as others involved in drug efflux.

ABC Transporters and Beyond

291

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lizes two kinds of energy coupling and involves seven proteins to span the two membranes of the cell envelope. Interestingly, the genes encoding specific B I2 transport proteins are not in an operon but are scattered as bluE, blueD, and bluF loci in the E. coli chromosome. The recent acquisi tion of structures of all four B,rspecific transport components provides a structural basis to begin to understand the complexities of this system, which provides a good model for other ABC transport systems. Transport across the Inner Membrane Delivery by BtuF

5'-Deoxyadenosylcobalamin (coenzyme B l2 ) 11.29. S'-Deoxyadenosylcobalamin, vitamin B12. The Co(1I1} ion is liganded by four pyrrole N atoms of the corrin ring and the N atom of S,6-dimethylbenzimidazole (DMB), which is covalently linked through its 3'-phosphate group to a side chain of the corrin ring. The sixth ligand is a S'-deoxyadenosine in most physiological conditions, as shown, that is replaced during purification by a cyano group to produce cyanocobalamin.

Vitamin B '2 is transported across the inner membrane by an ABC transport system consisting of BtuCD and BtuF. BtuF is the soluble substrate-binding protein thaI avidly binds the cofactor (Kt ~ 15 nM) as it enters the periplasm through the BtuB channel in the outer membrane (discussed below). The x-ray structure of BtuF with bound vitamin B I2 shows two lobes that each consist of a central five-stranded l3-sheet surrounded by helices, a Rossmann-like fold (Figure 11.31). Betvveen the two lobes is a deep cleft with the substrate-binding site. The CN-cbl is bound with its DMB ligand present, and it contacts six aromatic residues, three from each lobe of BtuF. Unlike most of the ABC substrate-binding proteins in E coli, BtuF has a backbone ex-helix spanning the two domains that makes it unlikely to undergo a large hinge motion to the unliganded state. That such a large movement is not required to release the su bstrate is indicated by a zinc-binding protein from Treponema BtuB

The Vitamin 8 12 Uptake System

Vitamin B ,2 , or cyanocobalamin (CN-cbl), is a cofactor produced by some bacteria and archaea and is required by a variety of enzymes in most cells. Specific transporl systems enable cells to import this large, inflexible molecule, which consists of a corrin ring (a tetrapyrrole with a cobalt metal) plus two axial ligands, cyanide and 2,3-dimethyl-benzimidazole (DMB), covalently linked to the ring via aminopropanol-phosphate-ribose (Figure J 1.29). Transport of vitamin B '2 can be fully induced in E coli by growth on ethanolamine (because it is needed for the ethanolamine ammonia lyase reaction) and monitored by uptake of radiolabeled [57Co]CN-cbJ. Extensive biochemical and genetic studies have characterized two energized phases of vitamin B '2 transport: uptake across the outer membrane utilizing the specific receptor BtuB coupled with the TonB/ExbBD system for energy input, and uptake across the inner membrane via the ABC transporters BtuCD and BtuF (Figure J 1.30). Thus this system uti-

BtuCD

TonB-ExbBD complex 11.30. Components of the transport system for vitamin B12. Structures for BtuCD, BtuF, and BtuB have been solved, along with the C terminus of TonB, while the structures of ExbB and ExbD are not known. The general porin is included in the outer membrane because it may allow passive diffusion of vitamin B12 into the periplasm. From Kadner, R. J., et aI., in R. Benz (ed.), Bacterial and Eukaryotic Porins, Wiley-VCH, 2004, pp. 237-2S8. © 2004 by Wiley-VCH. Used by permission of Wiley-VCH Verlag GmbH.

Membrane Protein Assemblies

292

N

with gates at each end. Therefore to transport vitamin BIz to the cytoplasm requires a conformational change, which is likely to change the tilts of the TM helices analogous to the opening of LacY and GlpT (see Chapter 10). This conformational change is triggered by the binding and/or hydrolysis of ATP at the BtuD NBDs, likely A.

11.31. X-ray structure of BtuF, the periplasmic vitamin B1Zbinding protein. The ribbon diagram shows l3-sheets (blue) at the substrate-binding lobes and <x-helices (green) in the lobes and the backbone, with an asterisk denoting the helices that form the backbone. The substrate, vitamin B1Z, is shown as a ball-and-stick model. From Borths, E. L., et aI., Proc Natl Acad Sci USA. 2002, 99: 16642-16647. © 2002 by National Academy of Sciences, USA. Reprinted by permission of PNAS.

pallidum with a similar backbone, which requires a 4° tilt of the C-terminal domain to collapse the substratebinding site. BtuF carrying vitamin B 12 docks on the peri plasmic side of the BtuC dimer. Conserved glutamate residues on the surface of BtuF can be aligned with arginine residues on BtuC to form the stable complex described below. BtuCD in the Inner Membrane The BtuC and BtuD proteins transport vitamin BIz across the inner membrane at the expense of ATP hydrolysis. They both dimerize to form a heterotetramer, BtuCzD z, thus providing the two TM domains and the two NBDs that are standard for ABC transporters (see Figure 6.6). Crystals of the heterotetramer in the absence of both vitamin BIz and ATP allowed the structure to be refined at 3.2 A resolution, giving definition of the entire structure except for 17 C-terminal residues of BtuD and a few residues of the periplasmic loops of BtuC (Figure 11.32). Each BtuC subunit has 10 tilted TM helices, giving the assembled transporter significantly more TM segments than observed in most ABC transporters (which typically have 12). Between the BtuC subunits is a wide hydrophobic cavity forming a translocation pathway that is large enough to accommodate vitamin BIz but lacks a specific binding site for it. In the crystal structure the channel opens to the peri plasm. Two loops at the ends of TM helices 4 and 5 form a gate that closes the channel to the cytoplasm. The channel is proposed to work like an airlock

B.

11.32. X-ray structure of the ABC transporter BtuCzDz. A. Viewed from the side, the two BtuC subunits (purple and red) span the membrane with their L1 and L2 helices (gold) at the cytoplasmic surface. The TM helices are numbered. The two BtuD subunits (green and blue) associate with BtuC at the cytoplasmic side and have cyclotetravanadate molecules (ball-and-stick models) at their ATP-binding sites. B. The BtuD NBD domains (blue and green) are viewed from the cytoplasmic side, with the P-looplWalker-A motif (red), Walker-B motif (pink), and ABC signature motif (yellow) labeled (compare with Figure 6.7). From Locher, K. P, et aI., Science. 2002, 296:1091-1098. © 2002. Reprinted with permission from AAAS.

ABC Transporters and Beyond A.

293

B.

C.

11.33. MD simulation of the effect of ATP binding to BtuC2D2. Computer simulations based on the crystal structure of the BtuC2D2 heterotetramer were run for 15 ns in the absence of nucleotide (A). Then ATP was positioned at each of the two binding sites. In B, ATP is bound to binding site I (red) and in C, it is bound to binding site II (green). The cavities are colored to highlight the shifts in the transport channel (yellow) and the gap separating the NBDs (gray). From 0100, E. 0., and D. P. Tielelman, J BioI Chem. 2004, 279:45013-45019. © 2004 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

mediated by two short helices, L1 and L2, that make an L-shape at the interface between BtuC and BtuD (see Figure 1 1.32A). Both the overall fold and the nucleotide-binding sites of BtuD resemble those of other ABC NBDs described in Chapter 6 (see Figure 6.7). BtuD contains a six-stranded l3-sheet surrounded by nine a-helices, with a peripheral three-stranded l3-sheet. The nucleotidebinding sites, which contain critical conserved residues in the P and Q loops and the Walker-B motif, are occupied by vanadate salts in the crystals. Because the BtuD subunits face opposite directions in the dimel~ the ABC signature from one subunit is opposite the P-Joop of the other, creating two ATP-binding sites at the interface between BtuD subunits (see Figure 11.32B). In the structure lacking ATP, this interface is not extensive, so it is likely that the BtuD dimer is stabilized by the interfaces between each BtuCD pair, where conformational changes due to ATP binding and hydrolysis must be transmitted to the TM domains. Key residues for this transmission have been identified by analogy to other ABC transporters; for example, Leu96 in BtuD COlTesponds to PheS08 of CFTR, the si te of mutations in 70% of the cases of cystic fibrosis (see Chapter 6). Binding of ATP appears to have a significant effect on the BtuCD conformation. MD simulations used to dynamically probe the ATP-binding process suggest that docking of ATP draws the two NBDs closer to each other, which then alters the translocation pathway of the TM domain by reorienting TM helix 5 (Figure 11.33). Other effects that are likely to trigger further

BtuF-B I2

Membrane

Cytoplasm

11.34. Model of the BtuC2D2F heteropentamer. Ribbon diagrams of the BtuC, BtuD, and BtuF proteins are shown in the heteropentamer that is suggested from complementary groups on BtuC and BtuF. Two cyclotetravanadate molecules occupy the ATP-binding sites in BtuD, and vitamin B12 occupies the binding site of BtuF. From Locher, K. P., et al. FEBS Lett. 2004, 564:264268. © 2004 by the Federation of European Biochemical Societies. Reprinted with permission from Elsevier.

Membrane Protein Assemblies

294

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11.35. Assay systems for the activities of the BtuC2D2F heteropentamer. The stable heteropentamer is reconstituted into proteoliposomes in random orientation, which allows hydrolysis of external ATP (A) as well as uptake of vitamin B12 with preloaded ATP (B). ATP hydrolysis can also be measured in detergent micelles (C). From Borths, E. L., et aI., Biochemistry. 2005, 44: 16301-16309. @ 2005 by American Chemical Society. Reprinted with permission from American Chemical Society.

confm'mational changes and allo\,v delivery of vitamin B 12 to the cytoplasm include the hydrolysis of ATP and the binding of substrate-loaded BtuF. The specificity of the inner membrane transport system resides in the high-affinity binding site of BtuF, since the channel in BtuCD does not appear to specifically recognize vitamin B 12 . The complementary groups of BtuF and Btue allow a model to be made of the BtuC 2 D2 F heteropentamer, which shows the substrate is positioned directly over the translocation channel (Figure 11.34). Mixing purified Btu F with the purified BtuCD proteins in a 5: 1 ratio, followed by removal of excess BtuF, produces a stable BtuC 2 D 2 F complex that hydrolyzes ATP in detergent micelles and in reconstituted proteoJiposomes and also transports vitamin B I2 (Figure 11.35). Given the protein-protein interactions in the complexes, models have been proposed for the

coupling of ATP to vitamin B 12 transport, with conformational changes occurring in both TM and NBD domains (Figure 11.36). Crystallization of the proteins at other stages in the transport process as well as achievement of higher-resol ution structures will further the understanding of this process. Transport across the Outer Membrane

BtuB in the Outer Membrane The specific outer membrane ('eceptor for vitamin B I2 is the BlUB protein, a f3-barreJ protein with striking similarities to the FepA and FhuA iron receptors described in Chapter 5, These specific outer membrane transporters carry out active lI-ansport only in the presence of the TonB protein (see below), so they are called TonB-dependent transporters (TBDTs). BtuB has a high

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11.36. A model for the transport of vitamin B,2. BtuF (green) brings vitamin B'2 (red) to the BtuC2D2 complex (black), sending a signal (blue dashed arrows) across the membrane to trigger ATP binding and hydrolysis, Hydrolysis of two ATP molecules (yellow) promotes conformational changes that release vitamin B 2 to the cytoplasm and also release BtuF. From Locher, K. P., Curl' Opin Struct Bioi. 2004, ' 14:426-431. © 2004 by Elsevier. Reprinted with permission from Elsevier.

ABC Transporters and Beyond

11.37. Ribbon diagram of BtuB, the outer membrane receptor for vitamin B12. The 22-stranded j3-barrel surrounds the globular hatch domain in the interior, similar to the FepA and FhuA proteins also in the outer membrane (see Figure 5.23). From Chimento, D. P., et aI., Proteins. 2005, 59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

affinity for vitamin B 12 , and its crystal structure has been solved in the presence and absence of bound substrate. The BtuB protein has two domains: a C-terminal domain that makes the f3-barrel, and an N-terminal globular domain thaI folds into the barrel to make a hatch (Figure 11.37). The barrel consists of 22 tilted antiparallel amphipathic f3-strands connected by short turns on the periplasmic end and somewhat longer loops of nine to 19 residues on the outside. In the structure without bound substrate, the loop joining strands 9 and lO is ordered while loops 2, 3, and 4 are disordered; in the structure with vitamin B I2 bound, the reverse is the case, suggesting that the loops are mobile and close to trap the bound substrate. The hatch domain is highly conserved among TBDTs. It has a polar exterior (compatible with the polar interior walls of the barrel) and a hydrophobic core. The core is a four-stranded f3-sheet, with connecting loops that have distinct roles (Figure 11.38). The two helices are amphipathic and interact with the core 13sheet through their hydrophobic sides. The three apical loops are involved in binding vitamin B 12 , and the loop between the third and fourth strand is called a "latch" because it interacts with f3-strands 13 through 15 of the barrel that fold slightly into the lumen. Near the N terminus, seven residues (Asp6-Thr-Leu-Val-Val-Thr-Ala) form the "Ton box," defined as the major site of interaction with the TonB protein by data from genetic and cross-linking studies. The Ton box is inside the barrel on the peri plasmic side ofBtuB, and its exposure to the periplasm increases somewhat when substrate binds.

295

The BtuB protein binds eN-cbl with its axial ligand, DMB. The substrate-binding site on BtuB is on one side of the external face of the molecule and involves residues from both the barrel and hatch domains. Binding of two calcium ions is prerequisite to binding the vitamin B I2 and helps to order barrel loops 2, 3, and 4 that clamp around the bound substrate. Two apical loops from the hatch domain pack against the DMB, and equatorial side chains of the corrin ring fit into small pockets formed by both domains. The tight binding (nM KI) is not surprising in view of all the polar and van der Waals interactions between the receptor and the substrate, including 11 hydrogen bonds. The high affinity of BtuB for vitamin B I2 suggests a conformational change is required for the receptor to carry out transport. Indeed, no channel is evident in the available structures of BtuB or any other TBDT. However, the hatch domain in the internal cavity does not have a close fit to the barrel walls, and the many water molecules in the interface between them mediate half of the hydrogen bonds between the two domains. Yet two thirds of the water molecules present are hydrogen bonded to one domain or the other, not both, which is characteristic of a transient protein-protein interaction (Figure 11.39). Thus it is likely that the hatch moves out of the barrel to allow passage of vitamin B 12 , as BtuB

Substrate binding Loop / Loop 3 \ Loop 2 \

r'~

iRG"r

I

f\,,--, TDG

11.38. Conserved features of the hatch domain of TBDTs. The four j3-strands (labeled hj31-4, arrows) are connected by loops and <x-helices (cylinders) with particular roles: the apical loops 1 through 3 are involved in substrate binding (red), the loop between hj33 and hj34 is the latch (black). The site of interaction with TonB, called the Ton box, is connected to a "switch helix" (gold) in two iron receptors but not in BtuB. The connection of the hatch domain to the barrel domain is the linker (black). Residues of conserved motifs whose functions are not known are labeled (blue) From Chimento, D. P., et al., Proteins. 2005,59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

296

11.39. Water molecules at the interface between the barrel and hatch domains of BtuB. The polar interfacial regions are filled with many waters. Bridging waters (green) make hydrogen bonds to both domains, while nonbridging waters (blue) make hydrogen bonds to a single domain or to other water molecules. From Chimento, D. P., et al., Proteins. 2005, 59:240-251. © 2005. Reprinted with permission from John Wiley & Sons, Inc.

undergoes a significant conformational change to make room for this very large and inflexible substrate to pass.

TonB and Energy Coupling Such a major conformational change requires energy, and surprisingly BtuB and the other TBDTs in the outer membrane utilize the energy of the pmI' (proton motive force) across the inner membrane. The energy coupling is carried out by TonB and its accessory proteins, ExbB and ExbD, which are anchored in the inner membrane. TonB and ExbD are both predicted to have single TM segments near their N termini, with large periplasmic domains. ExbB is predicted to have three TM segments with a large cytoplasmic loop between the first two. In the cell, these proteins form a complex of ~260 kDa with the stoichiometry 1 TonB: 2 ExbD: 7 ExbB, which may dimerize. TonB has 239 amino acids in three domains: the N-terminal TM domain, a proline-rich periplasmic domain, and a C-terminal domain of 48 residues that contacts outer membrane receptors. The uncleaved signal sequence is its TM domain, which contains a highly conserved sequence along one face of the ex-helix. The TM domain of TonB interacts with the TM segment of ExbD. Little is known about ExbB and ExbD, but they seem to be similar to MotA/B, 'which use the pmI' to drive the motion of bacterial flagella. A high-resolution structure of the C terminus (residues 155-239) of TonB reveals a highly cylindrical dimer, in which each monomer has three l3-strands. one ex-helix, and a short 3 10 helix (Figure 11.40). If this structure has not been perturbed significantly by the lack of the rest of the protein, it indicates that TonB forms a dimer that extends at least 65 A, halFway across the periplasm. Models proposed for the energy coupling between TonB/ExbB/D and the TBDTs suggest that an

Membrane Protein Assemblies energized state of TonB contacts a TBDT in the outer membrane and alters the conformation of the transporter. Evidence for such a mechanism is provided by a high-resolution structure of BtuB complexed with the C terminus of TonB, which shows the Ton box of BtuB has become a l3-strand recruited by the l3-sheet in TonB (Figure 11.41). The interaction between the l3-strands is predicted to be sufficient to allowTonB to pull the hatch domain out of the barrel, opening a transport channel for vitamin B 12 • The function ofBtuB protein is the outer membrane transport of vitamin B 12 ; however, like other outer membrane proteins. it is also used as a receptor for colicins and bacteriophage. specifically colicins E and A and phage BF23. Entry of these agents into the cell, which is also TonB-dependent. is poorly understood. DRUG EFFLUX SYSTEMS In contrast to the mysterious TonB/ExbB/D complex that couples the inner and outer membranes for uptake, much more is known about how molecules on their way out of the cell cross the periplasmic space due to the high-resolution structure for the amazing channeltunnel made by ToIC, along with structures of several other proteins involved in efflux. Some of these proteins serve to export lipids from the inner membrane, and some are related to proteins involved in the secretion of proteins such as hemolysin and colicins. But they are best known for their role in drug efflux because the problem of multidrug resistance (MDR) is now limiting

11.40. The C-terminal domain of TonB. The ribbon diagram of residues 155 to 239 of TonB shows two molecules (red and blue) are intertwined along the entire 65 A length of the structure, with [3-strands alternating from each monomer. From Chang, C, et al., J Bioi Chem. 2001,27535-27540. © 2001 by American Society for Biochemistry & Molecular Biology. Reproduced with permission of American Society for Biochemistry & Molecular Biology via Copyright Clearance Center.

Drug Efflux Systems

297

ondary transporter AcrB belongs to the RND (resistance nodulation cell division) superfamily. AcrB is part of a well-characterized tripartite drug efflux system that includes the membrane fusion protein, AcrA, and utilizes the remarkable TolC tunnel to export drugs directly to the extracellular space. Representing the diverse mechanisms for drug efflux, these five proteins provide important models for the mechanisms that rid cells of unwanted toxic compounds. Sav1866, an ABC Multidrug Transporter

11.41. The ribbon diagram of the complex of BtuB and the C-terminal domain (residues 147-239) of TonB. The front of the BtuB protein, shown with the outer l3-barrel (copper) and the inner hatch domain (green). has been removed to show the lumen and the BtuB-TonB interaction. The purified proteins were combined in a molar ratio of 1:5 in the presence of vitamin B 12 and calcium. In the x-ray structure. the Ton box from BtuB (blue) has been recruited by the l3-sheet of the TonB C terminus (magenta). The substrate, vitamin B12 (red spheres). is bound in the extracellular region above the hatch domain of BtuB. From Shultis, D. D., et aI., Science. 2006, 312:1396-1399.

treatment options for many cases of cholera, pneumonia, gonorrhea, and tuberculosis. Today, pathogens resistant to almost any antibiotic can arise due to multidrug efflux pumps. Genome analyses predict a role in drug efflux for 6% to 18% of all transporters in bacterial membranes. A few of the bacterial drug transporters are related to human MDR proteins that are making many tumors highly drug resistant. In Gram-negative bacteria, many MDR proteins are organized into tripartite systems, each consisting of an inner membrane transporter and a peri plasmic lipoprotein called a membrane fusion protein, in addition to the ou ter membrane channel. The transporters use either ATP hydrolysis or the proton motive force as the source of energy and belong to at least six families in the transporter classification scheme described in Chapter 6 (ww,...,.tcdb.org). X-ray structures are now available for prokaryotic transporters involved in drug efflux from three of these classes. The Sav 1866 protein is an ABC transporter related to mammalian MDR proteins, including P-glycoprotein (see Chapter 6). The EmrE protein is a member of the small MDR family that exports toxic hydrophobic compounds. The sec-

Sav 1866 is a multidrug transporter from Staphylococcusaureus with homology to several human MDR transporters, including P-glycoprotein. It uses the hydrolysis of ATP to drive efflux of numerous drugs, including the cancer drugs doxorubicin and vinblastine. lL is a homodimeric ABC transporter, with each dimer contributingoneTM domain and one NBD. While the structure of the BtuCD transporter for vitamin B I2 uptake has been solved in the absence of bound nucleotide, the structure of Sav 1866 with bound nucleotide gives additional insight into the coupling of ATP hydrolysis with transport. Each Sav1866 subunit has an N-terminal TM domain (residues 1-320) and a C-tenninal nucleotidebinding domain (residues 337-578) with a short linking peptide (residues 321-336). The x-ray structure of Sav1866, obtained at 3.0 A resolution, shows an elongated molecule (120 A long) that extends well into the cytoplasm (Figure 11.42). The NBD structure is similar to those of other well-characterized ABC transporters, with the nucleotide-binding sites at the interface between subunits where the P-loop of one NBD is across from the ABC signature motif of the other (see Figure 6.7). In the x-ray structure, Sav1866 has two molecules of ADP tightly sandwiched at this interface. Like many ABC transporters, the TM domain of the Say 1866 dimer has twelve TM a-helices; their symmetry in the structure suggests the occurrence of a gene duplication event as postulated for AQPs and major facilitators (see Chapter 9), but not previously observed in an ABC transporter. Helices from the two subunits are intertwined at the center of the membrane, where they bend outward to form two wings, with TM 1-TM2 from one subunit aligning with TM3-TM6from the other (see Figure 11.42). This means the two subunits do not form separate lobes of the transporter, as postulated for many ABC transporters. Constrained by these interactions, the two subunits are not likely to act independently and the NBDs probably do not completely separate during the reaction cycle. Like other multidrug transporters (see below), Sav1866 has a large internal cavity and lacks a welldenned drug-binding site. Rather, drugs appear to bind nonspecifically in the cavity, whose affinity for the drugs is altered by conformational changes. In the x-ray

Membrane Protein Assemblies

298 A.

u

·s CJJ

Ol

TMDs

!i

2 u>.

NBDs

_1 C-ter 11.42. The ribbon diagram of the x-ray structure of Sav1866, an ABC drug efflux transporter in S. aureus. The views in A and B are rotated by 90° The subunits (yellow and green) are intertwined, especially as they cross the membrane (gray shading). Two molecules of bound ADP are visible between the NBDs. The TM segments (numbered in B) are connected by short loops on the extracellular side (labeled ECll, 2, and 3 in B) and long loops on the cytoplasmic side (labeled ICl). Based on Dawson, R. J. P., and K. P. locher, Nature. 2006, 443: 180-185. © 2005. Adapted by permission of Macmillan Publishers Ltd.

structure, the translocation pathway is accessible to the outside and the outward-facing cavity is polar. It has charged residues along its surface in the inner leaflet primarily from TM2-TMS, while at the outer leaflet it is lined by TMl, TM3, and TM6. In this conformation, extrusion of hydrophobic drugs occurs with simple diffusion from the low-affinity cavity. Biochemical studies of similar MDR proteins suggest that when the cavity is open to the cytoplasm, it has a high affinity for the drugs. Thus the transport mechanism appears to utilize an alternating access mechanism, like that observed with the major facilitators (described in Chapter 9) except using the hydrolysis of ATP to drive the conformational change. At the interface between the NBD and the TM domain lie portions of the long intracellular loops, ICLI and ICL2. Each of these loops contains a short helix running nearly parallel to the plane of the membrane; they are called "coupling helices" since both genetic and structural data implicate them in communication between domains. Interestingly, only one of the coupling helices contacts the NBDs of both subunits; the

11.43. Model for the interaction of the TM domains and the NBDs in Sav1866. Due to the extensive interactions between TM domains at the "wings" as well as the contact between one intracellular loop with both NBDs, small conformational changes rather than large domain movements are expected to result from ATP-binding. From Schuldiner, S., Nature. 2006, 443: 156-157. © 2006. Adapted by permission of Macmillan Publishers Ltd.

Drug Efflux Systems

A.

299

B.

11.44. X-ray structure of the EmrE dimer. Ribbon diagrams of the EmrE structure viewed from the plane of the membrane (A) and perpendicular to the plane of the membrane (8, top view). Each monomer is shown in rainbow color (N terminus, blue; C terminus, red), with a space-filling model of TPP+ (magenta). From Tate, C. G., Curr Opin Struct Bioi. 2006, 16:457-464. © 2006 by Elsevier. Reprinted with permission from Elsevier.

other one contacts only the opposite NBD. The portion of the NBDs that contacts the TM domain is primarily residues around the Q-Ioop. In addition, a newly recognized conserved motif called the "x-loop" near the ABC signature motif of the NBD is proposed to form and break a cross-link to the ICLs and thereby to transmit conformational changes upon ATP binding and hydrolysis. In short, the coupling between the NBDs and the TM domains appears to utilize communication of moderate conformational rearrangements [Tom the NBDs uponATP binding (Figure 11.43), rather than their association and dissociation as described for BtuCD (see above). EmrE, Small but Powerful

EmrE from E. coli is only 12 kDa (110 amino acids), and yet when overexpressed it causes bacteria to be resistant to tetracycline, tetraphenylphosphonium (TPP+), ethidium bromide, and other antiseptics and intercalating dyes. Found in both Gram-positive and Gramnegative bacteria but not in eukaryotes, it is a proton! drug antiporter that uses the energy of the proton motive force to drive transport of drugs out of the cytoplasm. Predicted to have four <x-helices, EmrE is very hydrophobic and has only eight charged residues. The most important of these is the only buried charged residue, Glu14, which is involved in coupling substrate transport and proton translocation. The pKa of Glu 14 is elevated to 8.5 due to nearby hydrophobic residues (Trp63, Tyr40, and Tyr60). When aspartate is substituted for Glu 14, its pK\ is only around 6.5 and it releases protons without using them to pump out drugs. Cysteine replacements to study accessibility of residues to alkylating agents identified other hydrophobic residues likely to be in the substrate-binding site. These residues

(Leu7, AlalO, Ilell, Tyr40, and Trp63) are from TM helices 1 (near GluI4), 2, and 3. While there is agreement that EmrE functions as a dimeI', the nature of the dimer is controversial. The structure of EmrE solved by both cryo-EM and x-ray crystallography appears to be asymmetric dimers (Figure 11.44). Each monomer contributes fourTM helices, as expected, with the substrate bound in a cleft formed by two helices [Tom one monomer and one from the other. An axis of pseudo-symmetry relates the first three helices of one monomer to the first three helices of the other. This dual topology was detected in a global topology genomic analysis of E. coli membrane proteins (see Chapter 6) and is supported by data from mutational analysis of EmrE fusion proteins. However, it is inconsistent with biochemical and cross-linking data. In the most compelling cross-linking result, the n08C mutant of EmrE was purified after cross-linking with 0phenylenedimaleimide, a rigid cross-linker about loA in length. In asymmetric dimers, the positions of T1 08 in the two monomers would be over 35 Aapart, strongly suggesting the cross-linked species, which has full activity, is a symmetric dimeI'. Some EmrE mutants have a C-out topology, while others have a C-in topology. Furthermore, different topologies are observed with different constructs, for example, if a histidine tag and epitope are placed at the N or C terminus. If the EmrE sequence is posed such that it is readily pushed into one topology or another, it may represent an interesting point in evolution. Gene duplication events that produced larger transporters have produced some with domains in parallel orientation, such as LacY, GlpT, and AcrB, and others with domains in antiparallel orientation, such as AQPs. The controversy over the EmrE dimer has intensified with the recent acknowledgment of a software

Membrane Protein Assemblies

300

~

Drug

Medium

/ Cytoplasm

11.45. Schematic representation of the MDR system composed of AcrABffolC. Drugs can enter the AcrB (yellow) in the membrane for export from either the cytoplasm or the peri plasm. AcrA {green} is thought to stabilize the docking of TolC (blue) on AcrB and allow drugs to be transported outside the cell. From Murakami, S., and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

glitch that affected the solution of the EmrE crystal structure. Further structural and biochemical studies are needed to determine whether two EmrE monomers, identical in sequence, can insert into the membrane with opposite orientations. If this does occur in cells, it raises interesting questions regarding the determinants of membrane protein topogenesis. Tripartite Drug Efflux via a Membrane Vacuum Cleaner

Drug efflux in Gram-negative bacteria often employs three-component systems that span the cell envelope to extrude drugs directly into the outside medium. A well-characterized tripartite system in E. coli is composed of AcrB, an inner membrane transporter; AcrA, a periplasmic lipoprotein; and ToIC, the channel-tunnel protein (Figure 11.45 and Frontispiece). An analogous system is made up of the secondary transporter EmrB, the peri plasmic lipoprotein EmrA, and Tole. Similarly, the system for exporting hemolysin is made up of HlyB, HlyD, and Tole. In E. coli several systems for drug and protein export are known to use TolC, whereas drug efflux systems in other bacteria, such as Pseudomonas aeruginosa, have different tunnel components for the different efflux systems (see below). Assay of different transporters in vitro demonstrates that substrate specificity of these MDR systems resides in the inner membrane transporters. The AcrAB/ToIC system exports a broad variety of mostly amphiphilic compounds that

may be positively or negatively charged, zwitterionic, or neutral, including bile salts, erythromycin, and 13lactams such as ampicillin. Such MDR systems have been called "membrane vacuum cleaners" because they can remove a large number of unwanted substances h-om the inner leaflet of the cytoplasmic membrane and pump them out of the cell, preventing their accumulation in either the cytoplasm or the periplasm. AcrB, a Peristaltic Pump

The inner membrane transporter of a tripartite drug efflux system may belong to one of several different classes of transport proteins. As a member of the RND superfamily, AcrB is a proton/drug anti porter that utilizes the proton motive force to export many clinically important drugs. When purified and reconstituted into proteoliposomes, AcrB carries out proton-dependent transport of these drugs. Like numerous other transpNters, including other MDR proteins, the sequence of AcrB can be divided into two halves that are homologous, suggesting they evolved h-om gene duplication events. AcrB Structure

The first crystal structure of AcrB revealed a homotrimer in the shape of a jellyfish, with each 110-kDa monomer (1049 residues) providing 12 TM a-helices and also contributing to the large peri plasmic headpiece (Figure 11.46). In the bilayer-spanning domain the TM segments are fairly loosely organized around a central cavity of 35 A diameter, which must be filled wi th mem brane Iipids in vivo to avoid loss of the permeability barrier. The central helices, TM4 and TM 10, contain the only three charged residues in tbe hydrophobic TM segments of each monomer, Asp407, Asp40S, and Lys940, which can form salt bridges and are assumed to be involved in proton translocation. The headpiece is divided into two layers: the upper TolC-docking domain and the lower pore domain. The TolC-docking domain opens like a funnel to a diameter of ~30 A, matching the diameter of the TolC tunnel (see below). Below it, the pore domain has a large central pore lined by three helices, one £1-om each subunit. In the pore domain each subuni t has four l3-a-13 subdomains that together form a large substrate-binding pocket. The central cavity of AcrB is enormous, measuring in total around 5000 A3 Between the subunits are three openings to the peri plasm, called vestibules. Substrates can thel-efore access the central cavity either from the periplasm or the bilayer for transport out the central pore (Figure 11.47). An Alternating Site Mechanism

Because the first crystals of AcrB were of the trigonal form, they confined the trimer to exact threefold

Drug Efflux Systems

11.46. X-ray structure of the drug efflux pump AerB. A. View of the AerB trimer from the side, oriented with the TM helices on the bottom and the peri plasmic headpiece on the top. Two of the three subunits (colored purple, green, and blue) are seen clearly from this angle. B. The AcrB trimer, viewed from the periplasm, reveals a peptide strand from each subunit that reaches far into the neighboring subunit. C. The view from the cytoplasm of the AerB homotrimer shows the large central cavity. From Murakami, S., and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

301

Membrane Protein Assemblies

302

-}, MCIPC OCAC TC

_ 1'1' 11.47. Solvent-accessible surface of AcrB in a cutaway model. The front subunit has been removed, allowing a view into the central cavity and revealing pore helices (yellow) and TM helices that form grooves (green). Broken lines indicate the putative membrane boundaries and the framework of the funnel and cavity. Broken arrows indicate the postulated translocation pathways for substrates from the membrane, such as deoxycholate (DOC). acriflavine (AC), and tetracycline (TC). as well as substrates from the periplasm/outer leaflet, such as cloxacillin (MCIPC). From Murakami, S, and A. Yamaguchi, Curr Opin Struct Bioi. 2003, 13:443-452. © 2003 by Elsevier. Reprinted with permission from Elsevier.

to O. Drug extrusion is accomplished by the diffusion of substrate along a pathway that bulges and occludes as it migrates toward the funnel, much like the action of a peristaltic pump. The funnel at the top of AcrB fits the dimensions of the bottom of the TolC tunnel. and their interaction can be modeled to involve six hairpins at the top of AcrB docking with six helix-tum-helix structures at the bottom of ToIC. Although cross-links can be inserted between the two molecules, both with chemical crosslinking and with disulfide Formation between inserted Cys residues, unmodified TolC does not bind AcrB in vitro. In contrast, purified AcrA exhibits micromolar affinity for both AcrB and ToIC, suggesting that the docking between AcrB and TolC is reinforced by AcrA. The role of AcrA is likely to be quite dynamic given that the complex Formation is transient, aJlowing TolC to partner with other efflux transporters. AcrA. a Membrane Fusion Protein

In the tripartite drug efflux systems, membrane fusion proteins do not contribute part of the transport pores but are required to stabilize the interactions of the inner membrane transporter with the channel protein.

L Access

symmetry. Solution of the AcrB structure from crystals of different space groups that allowed asymmetry of the monomers revealed three different conformations affecting the access in the pore region. The three conformations are called loose (L, or "access"), tight (T, or "binding"), and open (0, or "extrusion"), and they vary in the binding pocket and pore regions (Figure 11.48). A variation in the position of the central pore helices is key to the conformational differences. The central helix From the 0 monomer is inclined nearly 15° to\-vard the T monomer, which results in opening the pore from the 0 monomer to the exit funnel while contracting the T monomer' to Form the drug-binding pocket (Figure 11.49). The lining of the pocket has eight phenylalanine residues (see Figure 11.48), aJlowing hydrophobic and aromatic interactions with drugs of diverse sizes and structures, as seen in the crystal structures of AcrB with several different drugs. The observation of three subunits in three conformations suggests an alLernating site, Functionalrotation mechanism analogous to that of the F1-ATPase (Figure 11.50). In the rotation, a monomer first binds substrate in the L conformation, binds it more tightly in the T conformation, and then extrudes it to the funnel in the 0 conformation. The subunits communicate via the central helices. The conformational changes are coupled to proton translocation in the TM domain: proton uptake is postulated to drive the change from 0 to L and proton release to accompany the change from T

T Binding 11.48. Illustration of the three conformations of subunits of the AcrB trimer. Each subunit is in a different conformation: the access state (L, green). extrusion state (0, red). and binding state (T, blue) shown in a cut view of the pore domain from the exterior of the trimer. The vestibules are clefts that are open in the Land T conformations and closed in the 0 conformation, which is open to the exit pore. Phenylalanine residues are shown in ball-andstick representation, and the binding pocket of the T conformation contains a molecule of the drug minocycline (orange contour labeled "Drug"). The arrows indicate the relative movements of the subdomains, which are labeled PC1, PC2, PN1, and PN2. From Murakami, S., et aI., Nature. 2006, 443: 173-179. © 2006. Reprinted by permission of Macmillan Publishers Ltd.

Drug Efflux Systems

303

11.49. Visualization of the tunnels in the pore domain of the AcrB peristaltic pump. In a ribbon diagram of the AcrB trimer viewed from the side, the tunnels are highlighted (green surfaces). A. The tunnel in the L monomer (blue) goes from the cleft halfway toward the central pore. B. The tunnel in the T monomer (yellow) extends across the pore domain toward the pore. C. The tunnel in the monomer (red) is closed at the lateral edge and opens into the exit funnel. From Seeger, M. A., et aI., Science. 2006,313:1295-1298. © 2006. Reprinted with permission from AAAS.

°

Members of the membrane fusion protein family attach to the inner membrane either via lipid acylation ofa cysteine residue or with an N-terminal TM segment. However, studies of mutants lacking these regions reveal that membrane attachment is not essential for the drug efflux function of these proteins. MexA, from the

B.

%

11.50. Schematic representation of the AcrB alternating site functional rotation transport mechanism. The three conformational states are loose (L, access; blue), tight (1, binding; yellow), and open (0, extrusion; red). They are shown as viewed from the side (A, with dotted lines denoting the membrane) and from the top (B). The side chains presumed to be involved in proton translocation (Asp407, Asp408, and Lys940) are indicated in the TM part of each monomer in A. An acridine molecule is depicted as substrate, which first binds to the L state, then binds tightly to the aromatic pocket in the T state, and then is extruded toward the funnel in the state. From Seeger, M. A., et aI., Science. 2006, 313:1295-1298. © 2006. Reprinted with permission from AAAS.

°

MexAB/OprM tripartite system of P. aeruginosa, was the first membrane fusion protein to have a high-resolution structure. AcrA is highly homologous to MexA, so it is not surprising that its structure proved to be very similar. AcrA Structure AcrA consists of 397 amino acids including a cleavable N-terminal signal, residues 1 to 24. Its C-terminal ~90 residues are very sensitive to proteolysis and correspond to the region of MexA that did not give clear electron density in its x-ray structure, indicating it is highly flexible. Therefore, this C-terminal domain, which is essential for interaction with AcrB and ToIC, was cleaved fTom the AcrA molecule for structural analysis of its 28-kDa stable core. To solve the x-ray structure, four additional methionines were substituted in the AcrA fragment, allowing incorporation ofselenometbionine for phase determination. The effect of the four methionine residues on the activity of intact AcrA is not entirely clear: mutants with two of the substitutions function to give drug resistance in a 6AcrA mutant, but with all four substitutions the intact protein is not sufficiently expressed to give the resistance phenotype. The 2.7 A resolution structure of the AcrA fragment shows residues 53 to 299 in an elongated sickle that consists of three domains: a I)-barrel domain, a central lipoyl domain, and a coiled coil (X-helical hairpin (Figure 11.51). Nearest the membrane, the I)-barrel domain includes both the Nand C termini of the fragment. It has six antiparalJeJ I)-strands, two of which close off the barrel near the C-terminal end (where the proteolyticaUy sensitive domain would follow in the wildtype protein). The lipoyl domain contains a I)-sandwich

Membrane Protein Assemblies

304

cd

a1 u-helil:<.tJ hairpin

4

9if

lipoyl domain

.~ a3 53

r:l-barrel domain 11.51. X-ray structure of a monomer of AcrA. A ribbon representation of a monomer of the stable core fragment of AcrA (residues 45-312) with four methionine substitutions is viewed from two angles, differing by 90°. The AcrA monomer has three domains: helical hairpin (red), lipoyl domain (green), and j3-barrel (cyan). From Mikolosko, J., et aI., Structure. 2006, 14:577-587. © 2006 by Elsevier. Reprinted with permission from Elsevier.

composed of two half-domains of four strands each. This type of domain occurs in pmteins that bind lipoic acid or biotin, with the binding site (a lysine residue) present on a loop between the two half-domains. The lipoyl domain in AcrA lacks the lysine loop, instead forming the long coiled coil. The AcrA peptide chain Forms one half of the (:'>-sandwich, then the coiled coil, and then the other half of the sandwich. The coiled coil contains five heptad repeats in each helix, making it 58 A long. Packing between the helices involves canonical knobs into holes at the first and Founh positions of the heptad repeats. Conformational Flexibility

While the mle of the membrane fusion protein requires its interaction with both AcrB and TolC, just how AcrA interacts with its partners is not yet known. The position of AcrA can be predicted since its lipid anchor is on Cys25, the first residue of the mature protein, and the portion of its N terminus that is missing in the x-ray structure is sufficiently long to bring the (:'>barrel domain to a position alongside the peri plasmic domain of AcrE. This puts the coiled coil well into the peri plasm, where it can interact with TolC (see Frontispiece). While both AcrE and TolC are trimers, the oligomeric state of AcrA is not clear. In vilro both AcrA

and MexA, lacking membl-ane associations, are soluble monomers. In crystals, MexA formed six- and sevenmember rings while the AcrA Fragment Formed a dimer of dimers, which clearly does not represent physiological association. However, the crystal structure of AcrA is very informative because each of the Four monomers in the crystal has a different conformation due to a hinge at the base of the helical hairpin. The largest difference among the four shows a rotation of 15° (Figure 11.52). Interestingly, this hinge movement is due to variation in unwinding of the helices. It can be correlated with the proposed iris-like opening of the TolC channel (see below), but it is not clear whether it actively opens TolC or passively maintains the intermolecular contacts. TolC, the Channel-Tunnel

Drug efflux via the tripartite AcrAB/ToIC results in expelling the unwanted compounds into the outside medium because TolC provides a pathway acmss both the periplasm and the outer membrane. This is evident in the remarkable high-resolu tion structure of TolC (2.1 A resolution) that shows a homotrimer with a (:'>-barrel channel domain in the outer membrane that connects to a unique a-barrel in the peri plasm (Figure 11.53). Since the oc-barrel is 100 A in length, the sum of the

Drug Efflux Systems

305

_ 15° ~ Mol DMol A M () I B Mol C

a2

a1

al

ep

90"

11.52. Four conformations of AcrA observed in the crystal structure. The crystal form captured each monomer (labeled MolA, MoIB, MoIC, and MolD) in a different conformation due to the flexible hinge between the helical hairpin and the lipoyl domain. Superposition of the four conformations reveals a 15° rotation between the most different structures. From Mikolosko, J., et aI., Structure. 2006, 14:577-587. © 2006 by Elsevier. Reprinted with permission from Elsevier.

peri plasmic length of TolC and AcrB is 170 A, which is long enough to span the periplasm. TolC provides an uninterrupted pathway with an inner diameter of at least 20 A fTom the inner membrane transporter to the medium. Passage through the TolC channel-tunnel is passive, as the inner membrane transport partners carry out the energized step.

Extracellular

-40

A

-100

A

A Unique Structure

The TalC channel is an anti parallel 12-stranded I)-barrel with a right-handed twist. Aromatic residues delineate the regions located in the lipid bilayer interface (see Chapter 5). As the case for other outer membrane proteins, the external loops of TalC are sites for colicin and bacteriophage attachment. However, TolC differs from other outer membrane I)-barrels in significant ways. Rather than having each polypeptide form a barrel, each TolC subunit conttibutes four I)-strands to its single pore. With neither a constriction formed by inward folded loops nor a plug formed by a separate domain, the pore is constitutively open. Finally, the I)-barrel is connected directly to the ex-barrel, 'which has a lefthanded twist, requiring interdomain linkers with conserved proline residues that make abrupt turns (Figure 11.54). The peri plasmic tunnel of TolC is also 12-stranded, with ex-helices packed uniformly in an anti parallel lefthanded superhelical twist. Each subunit contributes two long helices (H3 and H7) and two pairs of shorter helices (H2/H4 and H6/H8) stacking end-to-end to span the length of the barrel (Figure 11.54B). Three short

Periplasm

Entrance 11.53. High-resolution structure of the TolC channel-tunnel. TolC is a homotrimer (each subunit is a different color) that spans the outer membrane with a l3-barrel channel and extends into the peri plasm with an ex-barrel. See text for details. From Koronakis, v., et ai, Annu Rev Biochem. 2004, 73:467-489. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

Membrane Protein Assemblies

306

Closed

Open

---<~f~~Q~'!R(t/M

.~;'';ii ~~ '~ '{Yr,:-

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l~ !§:f ~~ .'" ~

j'

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11.54. Structure of a monomer of TolC. A. The TolC monomer contributes four strands to the l3-barrel (yellow) and uses six helices to span the length of the tunnel (green). It also has an equatorial domain (red) containing both ex and 13 structures. B. Topology diagram for a monomer of TolC with helices (blue) and l3-strands (red) illustrates how the domains are connected and reveals a structural repeat. From Koronakis, et aI., Nature. 2000,405:914-919. © 2000. Reprinted by permission of Macmillan Publishers Ltd.

11.55. Model for opening the tunnel entrance of the TolC homotrimer. The closed state (left) is observed in the x-ray structure. The open state (right) has been modeled by replacing the inner set of coiled coils with the outer set to represent the rotation of the inner coiled coils. Both space-filling (upper) and ribbon (lower) representations are shown. The coiled coils H3/H4 and H7/H8 of one subunit are colored to clarify their positions, and the constraining links both within and between subunits are shown. From Koronakis,V., et ai, Annu Rev Biochem. 2004, 73:467-489. © 2004 by Annual Reviews. Reprinted with permission from the Annual Review of Biochemistry, www.annualreviews.org.

helices (HI, HS, and H9) and two short f)-strands thicken the middle of the tunnel at its equator-ial domain. Helix-helix interactions are stabilized by knobs-in to-holes packing of the side chains (described in Chapter 4). The lower part of the tunnel is less uniform: an inner pair of helices (H7 and H8) forms a

conventional coiled coil, while in the other pair helix H4 coils around helix H3. The result in the trimer is a 3.9 Aconstriction thal closes the tunnel at the cytoplasmic end, presumably in a resting state that maintains the integrity of the outer membl-ane. The TalC tunnel must open when it interacts with a substrate-carrying inner

v.,

TABLE 11.2. TolC-dependent drug efflux systems of E. coli and P. aeruginosa Substrates

Efflux systems (inner membrane/ outer membrane)

E. coli AC, AZ, BL, BS, CH, CM, CV, CP, DOC, EB, ER FA, FQ, FU, NAL, NV, RF, TC, SDS, TX LC, CCCP, NAL ML

AcrAB (RND)/ToIC AcrEF (RND)/ToIC EmrAB (MFS)/ToIC MacAB (ABC)/ToIC

P aeruginosa AC, AC, AH, AG,

AH, BL, CL, CM, CV, EB, FQ, ER, NV, SM, SDS, TL, TP AH, CL, CM, CV, EB, ER, FQ, NV, SDS, TC, TP, TS CM, ER, FQ, TP, TS ER, FQ, TC

MexAB (RND)/OprM MexCD (RND}/OprJ MexEF (RND)/OprN MexXY (RND)/OprN

AC, acriflavin; AG, aminoglycoside; AH, aromatic hydrocarbons; AZ, azithromycin; BL, 13-laetams; BS, bile salts; CCCP, carbonyl cyanide m-chlorophenylhydrazone; CH, cholate; CL, cerulenin; CM, chloramphenicol; CP, ciprofloxacin; CV, crystal violet; DOC, deoxycholate; EB, ethidium bromide; ER, ethyromycin; FA, fatty acids; FQ, fluoroquinolones; FU, fusidic acid; LC, lipophilic cations; ML, macrolides; NAL, nalidixic acid; NV, novobiocin; RF, rifampicin; SDS, sodium dodecyl sulfate; SM, sulfonamides; TC, tetracycline; TL, thiolactomycin; TS, triclosan; TP, trimethoprim; TX, Triton X-1 00. From Koronakis, v., et aI., Annu Rev Biochem. 2004, 73:467-489.

'I

Conclusion membrane transporter, such as AcrAB. The simplest mechanism for opening involves rotating the inner pair of helices around its partner, effectively untwisting the inner pairs like opening the aperture of a camera lens (Figure 11.55). The flexible helical hairpin of AcrA may playa role in opening TolC by engaging its inner helices, H7 and H8. Partners of TolC TolCjoins different E. coli inner membrane transporters to form tripartite efflux systems in a dynamic manner. In some bacteria, such as P aerugil1osa, numerous TolC homologs work with specialized transport functions (Table 11.2). Recently, the structures of two TolC homologs, OprM from P aeruginosa and VceC [Tom Vibrio cholerae, have been found to be cylindrical channels very similar to TolC. Homologs of TolC seem to be ubiquitous among Gram-negative bacteria and are used not only for drug export but for efflux of cations and secretion of proteins. TolC is utilized by specialized systems for secretion of hemolysin and colicin V from E. coli, leukotoxins from Pasteurella haemolytica and Actinobacillus actil1omycelemcomilans, adenylate cyclase from Bordetella pertussis, and several proteases and lipases. Thus the molecular picture provided by the structures of AcrA, AcrB, and TolC furthers the understanding of many important processes related to pathogenesis and drug resistance. TolC-dependent systems are one example of membrzlDe nanomachines whose mechzlDisms are being revealed with structural detnils. They show how the progress in structural biology of drug efflux systems in Grnm-negative bacteJin greatly enhances understanding of the general problem of drug resistance. While different classes of transporters are involved in drug efflux, organisms differ 'with regard to which classes are more represented. In Gram-positive bacteria MFS multidrug exporters are important, while in mammalian cells ABC transporters nre the dominant group. For this reason, the role of Sav 1866 in drug efflux in S. aureus heightens its importance as a model for human MDR proteins such as P-glycoprotein.

CONCLUSION

The examples described in this chapter range from complexes in the respiratory chain - FoFI-ATPase, cytochrome bCI, and cytochrome-c oxidase - to elaborate assemblies involved in transport - the translocon, the vitamin B I2 uptake system, and systems for drug efflux. They provide detailed revelations of the workings of various molecular machines in the membrane. Given the general difficulty of achieving highresolution stllJctures of membrane proteins, it is fantastic to see structures of multicomponent membrane

307

complexes and to begin to understand hm,v such molecular machines might operate. For some complexes, notably those of the respiratory chain, crystallization with cofactors remaining intact simplified the structural analysis in the same manner as the work on photosynthetic reaction centers described in Chapter 5. In other cases, each component of the membrane assembly was crystallized, affording tantalizing glimpses into the assembly of the whole. These examples give reason to expect many other membrane complexes that are subjects for biochemical, genetic, and biophysical analysis will become future candidates for structural analysis.

FOR FURTHER READING

F, Fa-ATPase Boyer, P. D., Energy, life and ATP (Nobel lecture). Angell' Chem 1111 Ed. 1998, 37:2296-2307. KurpJus, M., and Y. O. Gao, BiomolecuJar motors: the F I ATPase paradigm. Cun Opin Siruct Bioi. 2004, 14:250-259. Kinosita, K., Jr., et aI., Rotation of FI-ATPase: how an ATPdliven molecular machine may work. Annu Rev Biophys Biomol SiruCI. 2004, 33:245-268. Noji, H., et aI., Direct obselvation of the rotation of F 1 ATPase. Nature. J997, 386:299-302. ·Stock, D., et aI., Molecular architecture of the rotZlry motor in ATP synthase. Science. 1999,286: t700-\705. Stock, D., et aI., The rotary mechanism of ATP synthase. Cun Opin Siruci Bioi. 2000, 10:672-679. Respiratory Complexes Hosler, J. P., et al., Energy transduction: proton transfer through the respiratory complexes. Armu Rev Biochem. 2006,75:J65-187. "Hunte, c., et aI., Structure at 2.3 A resolution of the cytochrome bCI complex h-om the yeast Saccharomvces cerevisiae with an antibody Fv fragment. Siructure. 2000, 8:669-684. Hunte, c., et aI., Protonmotive pZlthw<1ys and mechanisms in the cvtochrome bCI complex. FEBS Lell. 2003, 545:39-46. 'Iwata, S., et aI., Structure at 2.8 Aresolution of cytochrome c oxidase from ParacocclIs denilrificaNs. Nature. J 995, 376:660-669. 'Svensson-Ek, M., et aI., The x-ray crystal structures of wild type and EO (1286) mutant cytochrome C oxidases from Rhodobacter sphaeroides. J Mol Bioi. 2002, 32\ :329-339. 'Tsukihara, T., et aI., The whole structure of the thineensubunit oxidized cytochrome c oxidase at 2.8 A. Science. 1996, 272:1136-t 144. 'Xia, D, et aI., Crvstal structure of the cytochrome bCI complex fmm bovine heart mitochonchia. Science. 1997,277:60-66. Yoshikawa, S., et aI., X-ray structure and the reaction mechanism of bovine heart cytochrome c oxidase. J Inorg Biochem. 2000, 82: \-7.

, Paper presents original structure.

Membrane Protein Assemblies

308 The Translocon

Beckmann, R., et aI., Alignment of conduits for the nascent polypeptide chain in the ribosome-Sec61 complex. Science. 1997,278:2123-2126. Breyton, e., et aI., Three-dimensional structure of the bacterial protein-translocation complex SecYEG. Narure. 2002, 418:662-664. "'Mitra, K., et aI., Structure of the E. coli protein-conducting channel bound to a translating ribosome. Nalure. 2005, 438:318-324. 'van den Berg, B., el aI., X-ray structure of a proteinconducting channel. Nalure. 2004, 427:36-44. White, S. H., and G. von Heijne, Transmembrane helices before, during and after insertion. CUlT Opin Snuct Bioi. 2005, 15:378-386. White, S. H., and G. von Heijne, The machinery of membrane protein assembly. Cun Opin Struct Bioi. 2004. 14:397404. Vitamin B1 2 Transport

Borths, E. L., et aI., The structure of Escherichia coli BtuF and binding to its cognate ATP binding cassette transporter. Proc Nail Acad Sci V S A. 2002,99:16642-16647. Chang, e., et aI., Crystal structure of the dimeric C-terminal domain of TonB reveals a novel fold. J Bioi Chern. 2001, 276:27535-27540. 'Chimento, D. P., Structure-induced transmembrane signaling in the cobalamin transporter BtuB. Nature Struct Bioi. 2003,10:394-401. Chimento, D. P., et aI., Comparative structural analysis of TonB-dependent outer membt-ane transporters: implications for the transport cycle. Proleins. 2005, 59:240251. Davidson, A. L., and J. Chen, ATP-binding cassette transporters in bacteria. Amw ReF Biachem. 2004, 73:241268. 'Locher, K P., et aI., The E. cali BtuCD structure: a framework for ABC transporter architecture and mechanism. Science. 2002,296:1091-1098. Locher, K. P., Structure and mechanism of ABC transporters. CUlT Opin So'uer Bioi. 2004, 14:426-431.

Postle, K, and R. J. Kadner, Touch and go: tying TonB to transport. Mol Microbial. 2003, 49:869-882. Drug Efflux 'Dawson, R. J. P, and K P. Locher', Structure of a bacterial multidrug ABC transporter. NalUre. 2006, 443: 180-185. "Koronakis, \I., et aI., Crystal structure of the bacterial membrane protein TolC central to multidrug efflux and protein export. Nann·e. 2000,405:914-919. Koronakis, \I., TolC - the bacterial exit duct for proteins and drugs. FEBS LeI!. 2003, 55:66-71. Koronakis, \I., et aI., Structure and function of Tole. Amw Rev Biochem. 2004, 73:467-489. "Ma, e., and G. Chang, Structure of the multidrug resistance efflux transporter EmrE from Escherichia coli. Proc Nat! Acad Sci USA. 2004, 101 :2852-2857. *Mikolosko, J., et a1., Conformational Aexibility in the multid rug efflux system protein AcrA. SlntClure. 2006, 14:577587. 'Murakami, S., et aI., Crystal structure of bacterial mullidrug efflux transporter AcrB. Nalure. 2002,419:587-593. Murakami, S., and A. Yamaguchi, Multidrug-exporting secondary tl-anSpOrlers. Curl' Opin SlntCI Bioi. 2003, 13:443452. Murakami, S., et aI., Crystal structures of a multidrug transporter reveal a functionally J'Otating mechanism. Nalure. 2006,443:173-179. Schuldiner, S., When biochemistry meets structural biology: the cautionary tale of EmrE. Trends Biachem Sci. 2007, 32:252-258. Seeger, M. A., et a1., Structural asymmetry of AcrB trimer suggests a peristaltic pump mechanism. Science. 2006, 313:1295-1298. Tate, e. G., Comparison of three structures of the multidrug transporter EmrE. Curl' Opin Siruct Bioi. 2006, 16:457-464. Yu, E. W, et aI., Structural basis of multiple drug-binding capacity of the AcrB multidrug efflux pump. Science. 2003, 300:976-980. Zgurskaya, H. 1., and H. Nikaido, Multidrug resistance mechanisms: drug efflux across two membranes. Mal Microbial. 2000, 37:219-225.

12 Themes and Future Directions

The crystal lattice of the Kvl.2 potassium channel, a mammalian voltage-dependen Shaker F<Jmily K channel, in complex with the I' su un t of its regula ory oxidoredllctaSE' reveals protem protein interactIons between I pid btl ayers The plane a the btlayer IS established y he TM pore and voltage sensor domaIns (red). The T1 domain ilnd (' sLJbuni me outSide the membrane (blue) and mediate the proteinprotein contacts ill th crystal. The box delll1eates a sngle ur It cell. From Long S. B., E. B. Campbell, and R. MilcKinnon, Seier ce. 2005, 309:897-903. 2005. Reprinted with permlssial from AAAS

The 2003 Nobel Symposium on Membrane Proteins presented progress based on a vast amount of work characterizing membrane proteins, including many of the proteins described in preceding chapters. For decades the methods of biochemists, biophysicists, geneticists, and cell biologists provided a wide range of techniques to probe the structural basis of how membrane proteins function. When crystallographers turned their attention to membrane proteins, it took over a decade to solve some of the first crystal structures and provide the long-awaited high-resolution structures. These first glimpses of their beautiful structural details changed the level of comprehension of membrane proteins, providing new insights into their mechanisms of action. Now that dozens of membrane proteins can be viewed at atomic resolution, recurring themes can be identified that are likely to engage structural biologists focusing on membranes. This last chapter not only summarizes these themes, which occurred throughout earlier chapters, but also points out directions for future work as it considers what the

current high-resolution structures are revealing about oligomerization, different conformational states, and structural patterns of membrane proteins.

OLIGOMERIZATION

The majority of the proteins discussed in this book, and indeed most of the membrane proteins for which there is a high-resolution structure, form oligomers. Even the monotopic prostaglandin Hz synthase forms dimers that are the active state. It is important to determine the oligomeric state of membrane proteins, whether enzymes, channels, or receptors. While crystal structures do not necessarily reveal the physiological state of proteins, they often provide structural evidence that bears on the possible quaternary associations, sometimes by defining them and sometimes by excluding them. In the cases where the x-ray structure reveals that an active site or transport channel is formed by monomers or, in contrast, is formed by more 309

310

than one subunit, the relationship between quaternary structure and the function of the membrane protein can be clarified. For example, the x-ray structures of channel-forming proteins make clear distinctions between those which have channels in each subunit, such as the tlimeric porins and the family of aquaporins, and those in which different subunits contribute to one central channel, such as the potassium channels, the mechanosensitive channels, and the TolC channeltunnel. Some hetero-oligomers form stable complexes that do not dissociate in detergent and have been more amenable to crystallization than other membrane proteins. Examples include the photosynthetic reaction centers and some components of the electron transport chain. Yet in the membrane even these complexes are likely to be part of higher-order structures. The photosynthetic reaction centers are associated with antennae containing many light-har'vesting proteins. Evidence from EM and native gel electrophoresis suggests the respiratory complexes are themselves protomers of higherorderoligomers. There are even EM images that suggest the ATP synthase can form dimers. How the membrane proteins pack to form crystal lattices affects the ability of researchers to detect oligomers or multimers in x-ray structures. During the purification process in detergents, protein-protein interactions may be disrupted. Then during the crystallization process, non-native protein-protein arrangements may form to stabilize the crystal lattice. Loss of higher-order structures is likely when protein-protein interactions are mediated by lipids. For example, there is extensive biochemical evidence for dimerization of the mitochondrial ATP/ADP carrier. Even though the AAC has been crystallized in layers of lipid bilayers, only one of its two crystal forms allows resolution of a cardiolipin between subunits, supporting the notion that lipids could facilitate dimer formation in vivo. Lipids clearly mediate the dimerization of OMPLA, whose active site is formed at the interface of two f)-barrels where they capture a phospholipid substrate. The activity of this enzyme is thus regulated by dimer formation, which depends on the state of surrounding membrane lipids. Subunit interactions are also important for understanding the regulation of many hetero-oligomers. An obvious example is the ATP synthase because the F I-ATPase can function to synthesize ATP only when coupled to the Fa subunits in the membrane. High resolution structures of increasing portions of the ATP synthase, in combination ",vith a wealth of data from other studies, are now providing insight into how its components function together. X-ray structures are not always consistent with previously obtained biochemical and EM evidence. A few proteins that appear by biochemical criteria to form

Themes and Future Directions homo-oligomers now have structures showing that they can actually Function as monomers, such as the mitochondrial carriers and the aquaporins. However, x-ray structures that reveal complete functional units do not eli minate the possibility of dynamic orconditional associations to Form higher-order oligomers, as seems to be the case with the AAe and the translocon. Clearly x-ray structures do not always define the protein associations that exist in the cell, and may in fact introduce non-native protein-protein arrangements formed to stabilize the crystal lattice. Of course protein associations that are dynamic and transient or that require input of energy are lost with crystall ization. Hence it is necessary to use other methods (EM, NMR, EPR with site-directed spin labeling, MD simulations) to probe which interactions are required for function and which result from artifacts.

CONFORMATIONAL CHANGES

While protein function frequently involves conformational changes, in the crystal lattice protein structure is usually Frozen in a single conformation. A Fascinating exception is the x-ray structure of AcrA in which the molecule has crystallized in four different conFormations, giving snapshots of the t1exibility needed for its role as a fusion protein between the AcrB transporter and the TolC tunnel-channel. Moreovel~ crystal structures can capture discrete conformations of subunits required for the function of the protein, as seen in the well-known OC3f?>J domain of the F 1 Fa-ATPase. The recent high-resolution structures of AcrB in crystal forms that allowed asymmetry in the trimer also show the subunits in different conFormations, pointing to an alternating-site rotation model analogous to that of the F\-ATPase. A typical approach to obtain different conformations is to carry out crystallization in the presence of substrate analogs or inhibitors. This approach has been especially successful with the sarcoplasmic reticulum calcium ATPase. Comparison of the structures of Ca 2+ -ATPase with and without calcium ions shows large movements of its three domains. Additional conformations have been obtained with AIF 4 -ADP as an ATP analog and with AlF4" and the inhibitor thapsigargin, which stabilizes the calcium-Free forms. As described in Chapter J 0, a nearly complete reaction cycle can be envisioned For this complex ATP-dependent pump based on high-resolution structures capturing its different conformations. Detailed comparisons of structures in di [ferent conformations clarify the specific sources of the structural changes. For example, AcrA is hinged at varying angles between the lipoyl domain and the long coiled coil, and the hinge angle depends on the unwinding of an

Motifs and Patterns

ex-helix at the base of the coiled coil. For the helicalbundle membrane proteins in general, conformational changes arise from bending of helices, rotation of helices, unwinding of helices, and/or loss of secondary structure creating short loops. These can result in quite large structural changes, such as hinging between domains and opening and closing of channels. Some membrane proteins have been crystallized only in the presence of a substrate analog or inhibitor to push the equilibrium to favor one conformation. This was the case for the transporters LacY and mitochondrial AAC, which are expected to undergo large conformational changes to accomplish substrate translocation. In contrast, addition of substrate or inhibitor to other membrane proteins has little effect on their overall conformation. These proteins often lack ,",veil-defined substrate-binding sites, having instead large internal pockets or cavities. The large binding pocket of AcrB can accommodate a variety of drugs that interact with different subsets of residues without triggering large stnlctural changes. Another example is the large cavity of the BtuC transporter that lacks a specific binding site and relies on BtuF for the specific delivery of their substrate, vitamin 8 12 . Also binding of inhibitors need not always cause large conformational changes but can instead give important insight into mechanisms of inhibition, as observed with cytochrome-bel complex and prostaglandin H 2 synthase. Even when structures of different conformations are lacking, the information from x-ray structures can contribute to understanding conformational changes. The B values (temperature factors) of the crystal structures show regions with high Aexibility that implies conformational freedom. Simulations based on one highresolution structure can also pick out flexible regions and then predict other conformations. Often features of a high-resolution structure lead to predictions for how the conformation is likely to change. For example, the bends in the basket helices of mitochondrial MC suggest these are Aexible sites where alternate conformations enable the carrier to open to the other side of the membrane. Another example is the proposal that the loop in the backbone or the SecY translocon functions as a hinge between the two halves of the heterotrimer, suggesting how the opposite side might open and allow peptides to move laterally into the bilayer. A different basis for prediction of conformational changes is the comparison of structures of related proteins, such as the potassium channels. The KcsA, KirBacl.J, and KirBac1.3 structures show the closed channels, whereas MthK, KvAP, and KvJ.2 show the open channels. In other cases EM images can indicate what a different conformational state might look like. For example, both LacY and GlpT x-ray structures show the transporters open to the cytoplasm, while a lower-resolution EM structure shows the related car-

311

rier OxlT in a state open to the peri plasm. In addition, evidence for the structure of LacY in the conformation open to the peri plasm comes from studies of Cys replacement mutants that determined reactivity to N-ethylmaleimide as well as cross-linking. Future progress in membrane structural biology will certainly include elucidation of x-ray structures in different conformations from those available today. In some cases these will be key to understanding the mechanism of the proteins, such as the TonBdependent outer membrane transporters whose channels are blocked in the structures now available. A significant achievement will be determination of the specific conformations of activated states, for example, of visual rhodopsin whose available x-ray structure portrays the inactive ground state. The solved structures of intermediates in the bacteriorhodopsin photocycle, trapped by different conditions at very low temperatures and/or by mutations, provide a hint of what can be achieved in future work on rhodopsin and other GPCRs.

MOTIFS AND PATTERNS

Recurring patterns in the structures of membrane proteins can involve general features or fine details. Only a few membrane protein structures with resolution around 2 A are now available. For these structures the ability to determine positions of amino acid side chains and of water molecules reveals the importance of hydrogen-bonding networks to both structure and f-unction. Interhelical hydrogen-bonding networks in rhodopsin include many highly conserved residues. Since genetic data indicate that some of these residues are essential for activation of rhodopsin, it is likely that conformational changes that accompany activation will produce different hydrogen-bonding networks. In the mitochondrial MC the hydrogen bond networks link all but one of the TM helices and thus must be altered to allow the outward-facing conformation to change to the inward-facing conformation. The hypothesized trigger for this change is perturbation by the negative charges on ATP of a particular salt bridge at the base of the hydrogen-bonding network. Important networks of interhelical and even intersubunit hydrogen bonds can be expected to be revealed in other structures as they are obtained at higher resolutions. Some general structural features that can be observed at lower resolution show that many helical membrane proteins deviate [Tom the now-classical model of bacteriorhodopsin with its fairly uniform serpentine structure. Helix irregularity would seem to be more the rule than the exception, as almost every description of the structure of a helical-bundle membrane protein reports on variations in helix lengths, tilt

Themes and Future Directions

312

angles, and bends. For example, four of the TM helices of the sarcoplasmic reticulum Ca 2+ -ATPase extend well past the membrane bilayer into the cytoplasmic region of the protein. Even visuali-hodopsin has a 33-residuelong TM helix. In addition to some very long helices, the presence of half-helices, as seen in aquaporins, certainly makes hydropathy predictions inadequate. The recently solved structure of the CIC chloride channel has been called "a jumble of helices." In view of such irregularities, specific helix-helix interactions are important in stabilizing helix packing. These are typically knob-into-hole nonpolar interactions, which account for the frequent occurrence of the GxxxG motif in the helices. While the character of most TM helices is predominantly hydrophobic, a surprising number of charged residues occur in some membrane domains. Notably the KvAP potassium channel has four arginine residues in its otherwise nonpolar voltage-sensor paddle. Another deviation h-om the serpentine structure is the protrusion of some mem brane proteins well beyond the membrane. Examples h-om the respiratory membrane include the FIFo-ATPase, cytochrome-bcl complex and cytochrome-c oxidase, all of which protrude significantly (h-om 32 to 75 A) on both sides of the membrane. Formate dehydrogenase in the E. coli nitrate respit-atory pathway has peripheral subunits that extend 90 A past the bilayer. The peri plasmic head of the AcrB drug efflux transporter is 70 Along, and the tube of the TolC channel-tunnel protrudes 100 Ainto the peri plasm. These extensions beyond the bilayer enlarge the depiction of membrane proteins as described at the end of Chapter 1. Symmetry in helical membrane proteins has been detected both in their amino acid sequences and, in cases of lower homology at the primary level, in their overall folds. While some have three repeats and threefold symmetry, as seen in the mitochondrial carriers, it is more common for membrane proteins to exhibit twofold symmetry. The sequences of aquaporins contain two homologous halves, and the twofold symmetry is apparent in their structure, in which each half folds into three and a half TM helices with a junction between the two half-helices in the center of the membrane. Twofold pseudo-symmetry is .-evealed in the xray structures of LacY and GlpT, with each consisting of two linked bundles of six ex-helices. Although the two halves of these transporters have low-sequence homology, bioinfo.-matics programs detect this pattern in most other members of the Major Facilitator Superfamily. The first ABC transporter to have symmetry noted in the two halves of its structure is the Savl866 drug exporter. Yet another example is the Sec translocon, whose major channel is formed by two pseudosymmetric halves of the SecY subunit (Sec61p in yeast and SecYex in archaea). Apparently a common occur-

rence III the evolution of helical membrane proteins has been gene duplication followed by fusion of the two genes. Interestingly, in some cases (e.g., the aquaporins) the repeated domains fused in inverted orientations. A precursor to this evolutionary step would be the association of two monomers with opposite orientations, as observed in the crystal structure of the drug efflux protein EmrE. Even though the physiological relevance of the asymmetric dimer of EmrE is controversial today, the fact that mutations can encourage this asymmetry suggests that EmrE could represent an early stage in the evolution of the proteins that contain inverted repeats. Symmetric folds and structural motifs often contain characteristic signature sequences that al-e usually first identified as highly consented residues among related or homologous proteins. The powerful tools of genetics and bioinformatics can be expected to identify more motifs in the many membrane proteins about which very little is known, and this in turn will provide critical information about their functions.

CONCLUSIONS

In spite of the rich diversity of their functions, it is not surprising that membrane proteins exhibit some structural themes, given the const.-aints of their lipid bilayer environment and of their mechanisms of biogenesis. As the number of high-resolution structures grows, more shared traits will be recognized. In addition, more specific interactions with lipids will likely be revealed, serving as a reminder of the complexity of the molecules that surround membrane proteins. Much more than a simple hydrophobic barrier, the heterogeneity of the lipid bilayer means that even the relative position of an amino acid in a TM helix affects the ability of a peptide to be a stable TM segment. Thus much more will be learned from ['urther studies of those proteins able to insert into the lipid bilayer, such as diphtheria toxin. Progress in the characterization of assemblies of proteins that form complexes, even nanomachines, in the membrane allows even more elaborate processes to be tackled at the molecular level. Topics such as intracellular trafficking, chemotaxis and flagellar movement, virus infection, and membrane fusion have long provided fertile areas for research. Now it is feasible to target the membrane proteins mediating such phenomenon for study by the methods of structural biology. Of course while membrane structural biology is making great advances, it continues to benefit fTom advances in membrane biochemistry, biophysics, genetics, bioinformatics, and computational biology. These productive methodologies are working together to solve a major riddle of life, as the very definition of a cell depends on how the membrane

Conclusions determines what it takes in and sends out and how it responds to the outside environment. And while many of the membrane proteins that comprise up to one third of the human genome are not well characterized, already there are many examples of membrane proteins involved in the etiology of diseases and a large majority of current drugs target membrane proteins. Therefore the benefits of increased knowledge of the structural biology of membranes will undoubtedly reach beyond advances in basic research to have an impact on human medicine and health.

313

FOR FURTHER STUDY

(multi-author) Nobel symposium on membrane proteins: structure, hmction and assembly. FEES Leu. 2003, 555. (multi-author) Insight on membranes. Nature. 2005, 438:587. (multi-author) Special section on crossing the bilayer. Science. 2005, 310: 1451. Pornillos, 0., and G. Chang, Inverted repeat domains in membrane proteins. FEBS Leu. 2006. 580:358-362. White S., et aI., Lipid bilayers, translocons and the shaping of polypeptide structure, in L. K. Tamm (ed.), Prolein-Lipid Interactions. New York: Wiley-VCH, 2005, pp. 3-25.

Appendix I

ABBREVIATIONS

AAC: ABC LnJnSporters: AChR: AFM: ANK: AQP: ATR: BA: BCl: BO: BPh: BR: CATR: CFTR: CHAPS: CHAPSO: CL:

CMC: CMT: CN-cbl: CT: CTAB: DAG: DAGK: DDM: DEPC: DI-IA: DHPC: DLPC: DLPE: DMB:

ex-hemolysi n f'-ee energy change for the transfer of a substance From one solvent to another ADP/ATP carrier a large class of transporters named for their ~TP-~inding

i::assettes acetyl choline receptor atomic Force microscopy repeated domain of ankyrins aquaporin atl-actyloside, an inhibitor of the mitochondriol ATP/ADP carrielbongkrekic acid, an inhibitor of the mitochondrial ATP/ADP carrier bacteriochlorophyll bacterio-opsin. which locks the retinal cofactor bacteriopheopyt in bacteriodlodopsi n carboxyatractyloside, an inhibitor of the mitochondrial ATP/ADP carrier cystic fibrosis tronsmembrane conductance regulator, a chloride channel 3-[3-(chola midopropvl) dimethyl-ammonioJ-l-propanesulfonate 3-([3-cholamidopropyl]dimethylommonio)-2hvdroxy-I-propanesulfonate cardiolipin critical micelbr concentration critical micellar temperature cyanocobalamin, or vitamin B I 2 cytidyl transferase cetyltrimethylammonium bromide diocylglycerol diacylglycerol kinase - but DGK in chapter 6 dodecyl I3-D-maltoside dieleidoyl phosphatidyJcholine docosahexaenoic ocid dihexanoyl phosphatidycholine dilauroyl phosphotidylcholine dilauroyl phosphatidylethanolamine 2,3-d imethvl-benzim idazole

DMPC: DMPE: DMPG: DMPS: DOPC: DOPE: DPPC: DPPE: DRMs: DSC: DSPC: DT: EDTA: EF: EGF: EM: EPR: ER: FDI-1: FRAP: FRET: FTIR: G3P: GdmCl: GOF: GPCR: GPl: GUVs: HDL: HHDBT:

HiPIP: HMM: HPr: ICL:

IMS: Ko: KcI:

dimyristoyl dimy,-istoyl dimyristoyl dimyristoyl

phosphatidylcholine phosphatidylethanolamine phosphatidylglycerol phosphatidylserine

dioleoyl phosphatidylcholine dioleoyl phosphatidylethanolamine dipalmitoyl phosphatidylcholine dipalmitoyl phosphatidyJethanolamine detel-gent-resistant membranes differential scanning calorimeu-y d iSlearovl phospha tidylchol ine diphtheria toxin ethylenediamine tetraacetic acid edema Factor. a component of anthrax toxin epidermal growth factor electron microscopy electron pa,-amagnet ic resonance endoplasmic reticulum Formate dehydrogenase fluorescence recovery aFter photobleaching fluorescence resonance energy transfer Fourie,- t1-onsForm infTared spectroscopy glycerol-3-phosphate guanidinium chloride gain of Function G-protein coupled receptor glycosylphosphatidylinositol giant unilamellar vesicles high-density lipoprotein 5-/7- heptyl-6- hydroxy-4, 7-diozobenwth iowle, an inhibitor of the cytochrome-be I complex hexagonal phase with nonpolar centers and polar groups and water outside inverted hexagonal phose (with polar centers and nonpolar exteriors) high-potential iron-sulfur protein hidden Markov model histidine-containing phosphocarrier protein intracellular loop intermembrane space of mitochondria binding constant dissociation constant

315

Appendix I

316

Kr:

lipid association constant describing selective

PEP PTS:

phosphoenolpyruvate-dependent

PG:

phosphotransferase system phosphatidylglycerol

PG H 2: PGHS:

prostaglandin H 2 prostaglandin H 2 synthase (also called

retention of lipids by pmteins)

L13,: LI3:

lamellar gel in which the chains are tilted lamellm- gel, also called So (ordered solid)

L~:

lamellar liquid crystalline, also called Lei (or Id,

Lc : LC:

lamellar crystalline

PH:

liquid condensed, a condensed phase in a lipid monolayer

pleckstrin homology domain that binds phosphoi nosi tol

PI:

phosphatidylinositol

LDAO:

lauryldimethyJamine oxide, or

pKa :

pH at which an acidic or basic hmctional group

LE:

dodecyldimethylamineoxide liquid expanded, an expanded phase in a lipid

PKC:

is 50% protonated protein kinase C

monolayer

PL:

phospholipid, glycerophospholipid

Lep:

leader peptidase

phospholipase A2

LF: LHJ, LH2:

PL A2: POPC:

light-hal\lesting complexes

liquid-disordered)

Lo :

lethal factor, a component of anthrax toxin liquid-ordered lipid state that occurs when PLs, sterols, and/or sphingomylins are present

cyclooxygenase, COX)

POPG: POX: PrP:

I-palmitoyl-2-0Ieoyl phosphatidylchoJine I-palmitoyl-2-0Ieoyl phosphatidylglycerol peroxidase prion protein

loss of function

PS:

phosphatidylserine

R:

MBD:

large unilamellar vesicles membrane-binding domain

the radius of curvature of the lipid/water interface

MC:

Monte Carlo

RC:

reaction center (photosynthetic)

MCF:

mitochondrial carrier family

RCK:

regulators of conductance of K+

MD: MDOs:

molecular dynamics

RND:

Resistance Nodulation cell Division, a

MDR:

membrane-derived oligosacchal-ides multidrug resistance

Ro :

superfamily of drug efnux proteins the inu-insic value of R for each lipid species

MFS: MLVs:

major facilitatOl- superfamily

S:

multilamellar vesicles

LOF: LUVs:

MPoPS:

I-myristyl 2-palmitoleoyl phosphatidylserine menaquinone

shape parameter for lipids, lipid volume/(cross-sectional area of polar head-group x lipid length)

SDS:

sodium dodecylsul fate, also called sodium

membrane scaffold proteins

SOSL:

site-directed spin labeling

N:

Newton, a unit of force

SOS-PAGE: SDS polyacrylamide electrophoresis

NBD: NBD-

nucleotide-binding domain cholestemllabeled with the

SERCA: SLUVs:

sarcoplasmic reticulum Ca2+ -ATPase short-chain/long-chain unilamellar vesicles

SM: SMS:

sphingomyelin an analog of somatostatin that is an

sn:

amphiphilic peptide with a positive charge stereochemical numbering

MQ:

MS: MSPs:

mechanosensitive

cholesterol: nitrobenzoxadia7.0lyl group NBD-DLPE: DLPE (a PL) labeled with the

laurylsulfate

nitrobenzoxadiazolyl group NMR:

nuclear magnetic resonance nuclear Overhauser effect (in NMR)

SOPC: SOPE:

J-steaml-2-0Ieoyl phosphatidylcholine

octyl-polyoxyethylene octyl f)-D-glucoside (sometimes put as f)OG)

SRP:

signal recognition particle

STGA:

outer membrane phospholipase A

3-HSOrGal pf)I-6Manpal-2Glcpal-archaeol small unilamellar vesicles

oligomycin-sensitivity conferring protein, a

SUVs: TBDTs:

TonB-dependent transporters

subunit of ATP synthase

TC:

transport classification

PA:

phosphatidic acid

TOG:

f)-D-galactopyranosyl

PM:

protective antigen, a component of anthrax TEMPO:

PC:

toxin phosphatidylcholine

a small lipid-soluble spin probe whose nitroxide group has an unpaired electron

PCC:

protein-conducting channel, also called the

TIM:

transJocatase across the inner mitochondrial

TU-I:

transition temperature for lipid transitions

NOE: NSAlDs: octyl-POE: OG: OMPLA: OSCP:

nonsleroidal anti-inflammatory drugs

I-thio-f)-D-galactopyranoside

translocon PCR:

polymerase chain reaction

PDB: PE:

Protein Data Bank phosphatidylethanolamine

J -stearol-2-oleoyl phosphatidylethanolamine

membrane from lamellar to hexagonal phases, typically L~-HII

Appendix I TO':

TM: TNBS:

317

melting temperature for fatty acids and pure lipids; also lIsed as transition temperature for lipid transitions from Lp to L" transmembrane tr-initrobenzenesulfonic acid

TOM: TRAM: TROSY:

translocase across the outer mitochondrial membrane translocating chain-associated membrane protein transverse relaxation optimized spectroscopy (in NMR)

Appendix II

SINGLE LETTER CODES FOR AMINO ACIDS A, C. D. E. F, G,

alanine cvsteine asparLate glutamate phenylalanine glycine H, histidine I. isoleucine K. lysine L, leucine

318

M. methionine N, asparagine p. proline Q. glutamine R, arginine S. serine T. threonine V. valine W. tryptophan Y. tyrosine

Index

I-palm itoyl-2-oJeoyl phosphatidylcholine (POPC) in lipid rafts, 34, 35 MD simulation of, 200, 220 2-heptyIA-hydroxyquinoline-N-oxide (HOQNO), 226 AAe. See ADP/ATP carrier (AAC) AB toxin model, 84 ABC transporters, 133-135, 136, 290-291. See also specific conformational changes in, 292-293, 294 nucleotide-binding domains of, 135, 136,292,293,297 structure of, 290 symmetry in, 312 acetylcholine, 139 acetylcholine receptors (ACchRs) muscarinic, 139 nicotinic, 139-140 patch clamp recordings of. 57, 58 AcrA, 302-304, 307 conformational Oexibility, 304, 305 slruclure of. 300, 303-304, 310 AcrB, 297, 300-302 alternating site mechanism, 300-302, 303 protrusion beyond membrane, 312 struclure of. 300, 301, 302, 310 active transporlers, 131, 132 primary, 133-134 secondary, 137-138 acyl chains, 13-17 addition or removal of, protein binding regulated by, 83 composition of, 16 in MD simulations, 200, 202 length of, in lipid-protein interactions, 98 names for, IS, 17 oleoyl-, 15 polyunsaturated, 15 stearyl-, 15,200 adhesion protein, 114 ADP/ATP carrier (AAC), 138,242, 249-250

ADP/ATP carrier (AAC) basket, 250, 251 binding in, 250-251, 252 conformational changes in, 252, 311 dimer formation in, 252, 253, 310 hydrogen bonding in, 251, 311 MCF motif in, 250, 252 structure of, 250 transport mechanism, 249, 252-253 adsorption, detergent removal by, 50 aerolysin, 85 Aeromonas, 85 Aeropyrul11 pemix, 259 aggregation number (N), 46, 48 Agre, Peter, 207, 241 helical proteins channels, 132-133 folding studies of, 162-164 helices, 102 distortion of, in transmembrane segments, 92-93 in MS channels, 239 location of. 113 membrane-spanning, amino acids in, 92 hemolysin (aHL), 85, 86, US alamethicin, 87, 88, 132-133 alkaline phosphatase (PhoA), 144, 145, 146 AMBER,198 amino acid distribution genomic analysis of, 153 in integral proteins, 145 amino acid(s) codes for, 78 hydrophobicity scales for, 142, 143, 182 in transmembrane domains, 92 positions of, 154 snorkeling of, 184 amphipathic helix in fJ-barrels, 114 insertion into interfacial region, 76, 82 amphiphile shape hypothesis, 29-30, 31 amphiphile(s) spontaneous assembly of, 4-5 tripod, 45

amphiphilicity,4 micelle formation and, 47 of fJ-barrels, 157 amphipols,45 amphitropic proteins, 68, 72-73 membrane binding by, 76 anaerobic respirat'ion, 222, 224 angiotensin receptor, 235 anion exchangers, 138 ANK repeats, structure of, 70 ankyrin, 70 deficiency, 70 almexins, 72, 83 annulus, 8, 94. See also boundary (annular) lipids antennae, 111-112,310 anthrax toxin, 85 anti torsion angle, 15, 16 antibiotics microbial resistance to, 296 porins and, 118 antibody fragments, proteins bound to, for crystallization, 208, 209 antimicrobial peptides, 87 antimycin, 281 anti port, 138 antiporters, 138 ApoAI, nanodiscs, 66 apolipoproteins,73 apparent free energy (DGapp), 182 aquaglyceroporins, 255-258 selectivity of, 254, 258 aquaporin I-null humans, 253, 254 aquaporins (AQPs), 253-254 physiological roles of, 253, 254 proton transfer in, 256-258 selectivity of, 253-254, 258 structure of, 254-255 symmetry in, 312 arachidonic acid, 220, 221 archaebacteria phospholipids in, 18, 20 translocons in, 286 archaeol, 20, 104 area vs. surface pressure isotherm, in monolayer formation, 51 aromatic amino acids, 92, 114

Index

320

arrestin, 230 artifacts, in x-ray crystallography, 209 Astral compendium, 142 atomic force microscopy, of lipid rafts, 34,35 AT? generation, 272 catalytic mechanism, 277, 278, 279 regulation of, 276 ATP synthases, 272. See a/so F 1 Fa-ATPase/ATP synthase catalytic sites, 277 composition of, 271-272, 273 equivalent subunits, 273 MD simulations of, 278 role in respiratory chain, 279 ATPase subdomain, of ABC transporters, 135 ATPases, 133-134 A-type, 134 F-type, 134 superfamilies of, 134 V-type, 134 ATP-binding cassettes. See ABC transporters ATP-sandwich, 135 atractyloside (ATR), 249 avenacins, 55 avidin-biotin binding, in potassium channels, 263 AxxxA motif, 154 B values (temperature factor), 207, 311 Bacillus subti/is, 170 bacteria. See also specific species anaerobic respiration, 222, 224 cell envelope structure, 113 drug efflux, 300 group translocation, 135, 137 multidrug resistance, 297 protein export, 170, 172, 176 topogenesis, 185 translocation apparatus, 175,286 bacterial membranes, I, 2 blebs in, 64, 65 fatty acid composition of, 17 lipid diversity in, 37, 39 lipid-anchored proteins in, 75 phase transitions in, 16,40 phospholipids in, 18, 20, 39 protein-rich domains in, 8 stnlctural support in, 10 bacteriochlorophyll (BChl), 109, Ill, 112 bacteriocins, 85 bacterio-opsin (BO), 166 bacteriopheopytins (BPh), 109 bacteriorhodopsin (BR), 8,45, [03, 166

bacteriorhodopsin (BR) crystallization of, 209, 210 helical bundles in, 102-107 helix-helix interactions in, 154 hydmphobicity plots fo.·, 144 integral proteins in, 92, 93 mutant proteins from, 105 nanodiscs,66 photoreaction cycle in, 105, 107 protein-folding studies in, 164, 166-167 proton path in, 106, 285 structure of, 103, 104,105,106,203, 204,210 Band 3 protein, 138, 186 barrel-stave model, 88 Basic Local Alignment Search Tool (BLAST), 142, 157 fl-barrels, 102, 113-118,213,214 amino acid residues in, 114, 119, 121 composition of, 114 enzymes with, 214 families of, 157 genomic analysis of, 157-158 hydrogen bonding, 114 hydrophobic mismatching in, 99 in porins, 113, 119, 120, 121, 132-133,157 in toxins, 85 location of, 113,167-168 number of fl-strands in, 114 pmtein folding in, 167-169 structure of, 115, 116, 210 transmembrane segments in, 96 BBF (Beta-Barrel Finder), 157 BChl (bacteriochlorophyll), 109, Ill, 112 fl-cyclodextrin, 34, 36 fl-D-galactopyranosyl I-thio-fl-o-galactopyranoside (TDG),243 bee venom, 87, 88 Beerendsen, H. J. c., 197 Benson, A. A., 5, 6 Beta-Barrel Finder (BBF), 157 fl-galactosidase (LacZ), 144 fl-hydroxybutyrate dehydrogenase, 50 bicelles,63 crystallization and, 209, 210 bile salts, 43, 48 binding ligands, 79, 83, 234 binding sites ATP synthase, 277 BtuB,295 calcium ATPase, 264-265, 267 conformational changes and, 311 cytochrome-bcl complex, 281,282 GlpT, 245, 246

GPCRs, 234 KcsA,260 LacY, 243 reaction centers, 112 binding-change mechanism, 277 bioinformatics tools for protein families, 141, 142 motifs and patterns studied with, 312 biotin-avidin binding, in potassium channels, 263 bitopic proteins, 69, 90, 184 Bla (fl-lactamase), 144 black films. See planar bilayers fl-Iactamase (Bla), 144 BLAST (Basic Local Alignment Search Tool), 142, 157 B/aslOch/oris viridis antennae, 111 reaction centers, 107, 108, 109, 110 blebs, 63-64, 66 blisters, 63-65, 66 blood clotting factors, 73 fJ-octylglucoside, 45, 50 bond angle energy, 199 bond length, function for, 198 botulinum neurotoxin, 85 boundary (annular) lipids, 50, 94, 204 EPR of, 94, 95, 96, 98 exchange with bulk layer; 95, 98 numbers of, 205 structure of, 205 Boyer binding-change mechanism, 277 Boyer, Paul, 272 BPh (bacteriopheopytins), 109 BR. See bacteriorhodopsin (BR) Bragg's law, 193, 194, 195 BtuB conformational change, 295 function of, 296 in outer membrane, 294-296 structure of, 295, 296 BtuBC (vitamin Bil transporter), 135 BtuBCDF, 272 BtuC, 311 BtuC2D2F complex, 292, 293, 294 BtuCD complex, 290 conformational changes, 293-294 in inner membrane, 292-294 BtuD,293 BtuF, 291, 292 bulk lipids, 94, 96 exchange with annular· layer, 95, 98 CI domain, 80, 81 C2 domain, 80, 81

Index Caenorhabdilis elegans, J 53 calcium ATPase, 264 binding sites, 264-265, 267 conformational change in, 264-267, 268, 269 E 1-E2 reaction scheme, 264, 269 structure of, 3 J0 x-ray crystallography of, 209-210 calcium binding, 83, 84 by calcium ATPase, 264-265, 267 dephosphorylation step in, 267 to OMPlA, 214, 216, 217 calcium channels, voltage-gated, 264 calcium pump, 264 calcium, in signaling, 264 carboxyatractyloside (CATR), 250 cardiolipin (CL), 17,38,40 in reaction centers, 109 mixed micelles containing, 63 structure of, 205. 206 cardiolipin/K pathway, 283, 284-286 carotenoids. in reaction centers, 109 Carpet mechanism, 87, 88 carriers, 133 catalytic domain. of prostaglandin Hz synthase, 217, 220, 221 catalytic mechanism, FiFo-ATPase, 277-278. 279 catalytic triad. in OMPlA, 2J4, 215 CATH (Class, Architecture, Topology and Homologus superfamily), 142 CATR (carboxyatractyloside), 250 caveolae, 36 formation of. 37, 38,90 caveolins,90 cell envelope, bacterial, 113 cell surface receptors, in supported bilayers, 57 cerebrosides, 20 CF (cystic fibrosis). 187, 188 CFTR (cystic fibrosis transmembrane conductance regulator), 135, 186 misfoJding of, 187, 188 channel(s). 253. See also specific channel in translocons, 287. 288 channel-forming toxins, 132-133 chaperones, J73. 176 CHAPS (3-[3-(cholamidopropyl) dimethyl-ammonioJ-lpropanesulfonate), 43 charge mutations, 186 CHARMM,198

321

cholate. 50 cholera toxin, 85 cholesterol, 20, 21 in lipid raf1s, 34, 35 in supported bilayers. 57. 59 in ternary phase diagrams. 32, 33 interactions with otber lipids, 22, 23, 33 MD simulation of, 201,202 proportion of, 8 solubility of, 22-23 thickening by, 22, 27, 100 chromophores, III circular dichroism. 161, 166 cis double bonds, 15,200 CL. See cardiolipin (Cl) Class, Architecture, Topology and Homologus superfamily (CATH), 142 CIC chloride channel. 312 cleavable signal sequences. 171. 172, 184, 185 cloud point. See Krafu point CMC (critical micellar concentration), 46, 47 CMT (critical micellar temperature), 47 cobra venom phospholipase Az, 128 cofactors, in photosynthetic reaction centers, 109-110 colic ins, 85-86, 87, 132-133 conduction pores, 253 cone cells, 228, 229 conformational changes, 310-311. See a.lso specific protein CONPRED, 152 cotranslational translocation, 170, 176 coupling helices. in Sav 1866, 298-299 COX (cycJooxygenase), 216, 218, 220, 221 critical micellar concentration (CMC), 46,47 critical micellar temperature (CMT), 47 cross-linking studies of ATP synthase, 277 of protein insertion, J 78, J 79, 182 crystal packing, 207, 209 crystallography. See a.lso x-ray crystallography process of, 207-210 CT (cytidyltransferase), 82 cubic phase, 28, 31, 207 crystallization in, 207, 209, 210 types of, 31 curvature frustration, 30, 31,40 curvature stress, in protein folding, 167,169 cyanocobalamin. See vitamin BIZ

cyclooxygenase (COX), 216, 218, 220, 221 cystic fibrosis (CF), 187, 188 cystic fibrosis transmembrane conductance regulator (CFTR), 135,186 cystic fibrosis transmembrane conductance regulator (CFTR) misfolding of, 187,188 cytidyltransferase (CT), 82 cytochrome b, 279 cytochrome-bc) complex, 154, 279-284 integral lipids in, 206, 208 proton pathways. 283, 284-285, 286 protrusion beyond membrane, 312 Q cycle, 281, 282 stTl.lctLll-e of. 279, 280, 281-282, 284 cytochrome-bcl oxidase, 271 cytochrome-c oxidase, 154, 284-286 integral lipids in, 206 oxygen reduction, 285 proton pathways, 283, 284, 285-286 protrusion beyond membrane, 312 structure of, 284, 285 cytochrome c, binding by, 69, 76, 77, 78 cytochromeci,281 cytochrome cz, 112 cytochrome oxidase, crystallization of, 208, 209 cytochrome P450, crystallization of, 209 cytochrome P450 reductase, nanodiscs, 66 cytochrome subunit, of reaction center. 110 cytoplasm. protein export from, 170-172.174,175,177 cytoplasmic domain, in rhodopsin, 234 cytoskeleton, of plasma membrane, 10 DAG (diacylglycerol), 73 DGK. See diacylglycerol kinase (DGK) Danielli, James,S Database of Interacting Proteins (DIP), 142 Davson, Hugh,S DDM (dodecyl ,B-D-maltoside), 43 Debye-HuckeJ theory, 79 Debye-Waller factor. See B values (temperature factor) decyl maltoside, 63 Deisenhofer, Johann, 107, 207 denaturation of proteins, 161 deoxycholate. 50 DEPC, phase diagram for, 32 detergent(s), 42,43. See also specific detergent

Index detergent(s) alternatives to, 45 bicelle formation, 63, 209, 210 crystallization and, 206, 208 definition of, 43 impurities in, 43 in protein-folding studies, 170 ionic, 43 lipid removal by, 50,206 mechanism of action, 45-48 membrane solubilization by, 48-50, 129 concentration required for, 48, 49 nonionic,43 porins solubilized with, 119 removal of, 50 nanodiscs produced by, 66 solubility of, 43 types of, 43-44, 45, 46 zwitterionic,43 detergent-resistant membranes (ORMs), 36-37, 73 GPI-anchored proteins in, 75 proteins in, 36, 37 DHA (docosahexaenoic acid), 201, 202 DHPC (dihexanoyl PC), 167 diacylglycerol (DAG), 73 diacylglycerol kinase (DGK), 129-130 kinetic analysis of, 63 mutations, 129 protein folding in, 169 structure of, 129, 130 differential scanning calorimetry (OSC) of fatty acid phase transitions, 15, 17, 29 of lipid phase transitions, IS, 17,29, 76, 77 of nanodiscs, 66 diffraction pattern, 193 diffusion of lipid bilayer, 24-25 rate of, 25 of membrane components, 7, 8 digitalis, 134 digitonin, 55 dihexanoyl PC (DHPC), 167 dilauroyl phosphatidylcholine (DLPC), 100 dimer formation, 271,309-310 mitochondrial carriers, 252, 253, 310 OMPLA, 215, 216, 310 dlodopsin, 234 translocons, 288 dimyristoyl phosphatidylcholine. See DMPC (dimyristoyl phosphatidylcholine)

dioleoyl phosphatidylchoJine (DO PC), 192,194,195 dioleoyl phosphatidylchoJine (DOPC), 196 DIP (Database of Interacting Proteins), 142 dipalmitoleoyl PC (DPoPC), 167 dipalmitoyl phosphatidylcholine (DPPC) MD simulation of, 198, 199,200, 201 phase diagram for, 32 diphosphatidyl glycerol. See cardiolipin (Cl)

diphtheria toxin (DT), 84,132-133 distearoyl phosphatidylcholine (DSPC), 32,100 disulfide bond oxidoreductases, 134 DLPC (dilauroyl phosphatidylcholine), 100 DMPC (dimyristoyl phosphatidylchol ine) hydrophobic mismatching in, 100 in protein-folding studies, )67 molecular dynamic simulation of, 201 Monte Carlo simulation of, 202, 203 NMR spectra of, 24 docosahexaenoic acid (DHA), 20 I, 202 dodecyl tl-D-maltoside (DDM), 43 dodecylmaltoside, 235 dodecyl-phosphocol ine (DPC), I 18 dolichols, 20 domains, protein-binding, 80-83 OOPC (dioleoyl phosphatidylchoJine), 196 DOPG,78 DPC (dodecyl-phosphocoline), I J 8 OPoPC (dipalmitoleoyl PC), 167 DPPC (dipalmitoyl phosphatidylcholine) MD simulation of, 198, 199,200, 201 phase diagram for, 32 DRMs. See detergent-resistant membranes (DRMs) drug efflux systems, 296-297 ToIC-dependent, 306 tripartite, 300 drug targeting cyclooxygenase, 216, 222 G-protein coupled receptors, 140, 227 Dsb system, 176 OSc. See differential scanning calorimetl)' (DSC)

DSPC (distearoyl phosphatidylcholine), 32, 100 DT (diphtheria toxin), 84 E2-AIF4-(TG) complex, 267 EBF (epidermal growth factor) receptors, 37 edema factor (EF), 85 Edidin, M., 7 EGF-domain, of prostaglandin H2 synthase, 219, 220 EI (enzyme I), 135 Ell, 135 elaidic acid, 15 electrical potential-driven transporters, 133 electrical properties patch clamp recordings of, 55-56, 57, 58 planar biJayers used to study, 53-55 electrochemical gradient, across membranes, 53 electron carriers, 134 in nitrate respiratory pathway, 223, 224 in reaction cycle, 112, 113 electron density distribution, 193 electron paramagnetic resonance (EPR), 32, 33, 97 boundary lipids, 94, 95, 96, 98 MS channels, 238 myelin basic protein, 70 photosynthetic reaction centers, 108, 109 electron spin resonance (ESR). See electron paramagnetic resonance (EPR) electron transfer, in reaction cycle, 112 electrophysiology, 53 electrostatic interactions in MD simulations, 199 in reaction centers, I J 2 lipid selectivity and, 98 protein binding by, 69, 76, 78, 79, 80, 83 x-ray crystallography and, 203 electrostatic switch, 76, 83 eleidoyl, 200 emission spectrum, 26 EmrE, 297, 299-300 asymmetry in, 312 mutations, 299 structure of, 210, 299 endoplasmic reticulum (ER) P450 cytochromes in, 130 protein export to, 170, 172, 176 energy coupling, in vitamin 812 transport system, 296

Index energy surFaces, 196. 197 ensemble average. 197 enterobactin, 124 EnvZ, 119 enzymes, 127-129.214. See also specific enzyme lipid requirements of. 94, 95. 127 lipolytic. 214 epidermal growth factor (EBF) receptors. 37 epitopes. antibody recognition of. 143 EPR. See electl'on paramagnetic I-esonance (EPR) equilibration phase. 199 equilibrium measurements, of protein folding. 161 ER (endoplasmic reticulum) P450 cytochromes in, 130 protein export to, 170. 172, 176 ER pathway, 283. 284-286 ergosterol. 21, 22 eIJ,throcytes, glucose transporters in. 131.132 Escherichia coli acyl chains. 16 adhesion proteins. ] 14 ATP synthase, 272. 276, 309 chaperones. 176 colicins, 85 DGK.129 fatty acid composition, J 7 glyceroaquaporins. 254. 255-258 group translocators. 137 integral pl·oteins. 92 iron receptors. 124 mechanosensitive channels, 235. 236.237 OmpA, 118 OMPLA.214-2]5 OmpX, 118 phase transitions in. 40 phospholipid composition. 37, 39 porins. ]] 9,122 protein Folding. 100, 145 protein odentation. J 43, 146 SRP, 172.173.174 TolC-dependent drug efflux systems. 306 translocation, 175 translocon. 175,287 transporters, 241. 242, 248 tripartite drug efflux. 300 eukaryotes. See also humans; yeast; specific species aquaporins, 253 cell membranes. 1, 2 ion channels. 138-139 lipid-anchored proteins. 75 mechanosensitive channels. 235

323

P450 cylochromes, 130 plasma membrane cytoskeleton. 10 protein export, ]70.176 protein orientation. 144 receptors. 139 signaling enzymes, 129 sterols, 20-21. 22 topogenesis. 185 translocation, 160. 176 translocation apparatus. 175 translocons. 175, 286 tl"ansporters. 248 ExbB,296 ExbD.296 excitation wavelength. 26 excitatory glutamate receptors, 139 extracellular proteins. 72 eye, photoreceptors in, 228. 229

Fa domain. 274-275 F 1 Fa-ATPase catalytic mechanism, 277-278, 279 conFormational changes in. 276. 277 protrusion beyond membrane. 312 regulation of. 276-277 F 1 Fa-ATPase/ATP synthase, 272-273 structure of. 272, 273 subunit structure and fLlnction, 273-274.275,276.302 F 1 domain. 273-274, 275, 276, 302 Farnesyl, 20 FASTA,142 Fatty acid amide hydrolase, 90 fatty acid(s). 73 cis, 15,200 names of, 14. 15 phase transitions. 15, 17.29 polyunsaturated, 15 MD simulations of, 200-20 1.202 species of. 14 frans.15,200

FDH-N (formate dehydrogenase-N) functional unit. 225, 226 structure of. 223. 224 FepA protein, J 23, 124 ferdchrome. J 24 ferrous ion, in reaction centers, 109 FhuA protein, 123. 124 flap movement, in phosphoJipases, 71 flip-flop diffusion. See transverse diffLlsion flippases. 25. 26 flotation experiments. 192 Fluid Mosaic Model, 3. 7. 8, 68 lateral diffusion in, 24 lateral domains in, 33 fluidity. membrane. 24 measurement of. 8-9

fluorescence depoJarization, 26 fluorescence recovery after photobleaching (FRAP), 24, 26, 32,64 fluorescence resonance energy transfer (FRET). 26. 33 fluorescence techniques, 24, 26 F 1 Fa-ATPase studied with, 278. 279 giant unilamellar vesicles studied using, 63 lipid rafts studied with, 34 supported bilayers studied with, 59 formate dehydrogenase. 222-226. 312 formate dehydrogenase-N (FDH-N) ftlllctional unit, 225. 226 structure of, 223, 224 FosFomycin.245 Fourier transForm, 193 Fourier transform infrared (FTIR) spectroscopy. 161 FrankJin. Ben. 5 FRAP (fluorescence recovery after photobleaching), 24. 26, 32. 64 free fally acids. 14 freeze-fracture techniques, 6, 62 FRET (fluorescence resonance energy transfer). 26, 33 Frye. L. D.. 7 FTIR (Fourier transfOim infrared) spectroscopy. 161 fumarate reductase, 214 FYVE zinc-binding domains, 80. 81. 82 G protein (guanine nucleotide binding). 140 gain-of-function (GOF) mutants, 238 y-aminobutyric acid. 139 ganglioside GM z . 17, 18 ganglioside(s). 20. 98 gated pores, 131, 132. 288 gating mechanisms, 253 KcsA, 260-264 MS channels. 238-239 voltage, 262-263 gauche torsion angle. 15, 16 gel filtration, detergent removal by, 50 gene fusion. 143. 145, 146 general import pore (GIP). 178 genetic approaches to motifs and pa ttems, 312 to protein orientation, 143 genomic analysis of GPCRs, 234 of proteins, ]41. ]45-154 geranylgeranyJ, 20 GES (Goldman-Engelman-Steitz) scale. 139, 143 giant unilamellar vesicles (GUVs). 62 Gibbs absorption isotherm. 79

Index GIP (general import pore). 178 GlpF, 254, 255-258 channel interior, 256, 257, 258 hydrogen bonding, 256, 258 selectivity of, 255-256 structure oC 255, 256 GlpT. See glycerol-3-phosphate transporter (GlpT) glucose transporters in erythrocytes, 131, 132 topology of, 186 glucose-3-phosphate transporters. 241 glyceroaquaporins, 255-258 selectivity of, 254, 258 glycerol, 17 glycerol-3-phosphate (G3P). 17, 244, 255 glycerol-3-phosphate transporter (GlpT).242,244-245 glycerol-3-phosphate transporter (GlpT) binding sites, 245, 246 compared to LacY, 245-248 structure oC 245. 246, 247, 311 symmetl'y in, 312 translocation mechanisms. 242, 245, 247 glycerophospholipids. See phospholipids (PLs) glycolipids, 17, 18 glycophorin A, 93, 94.164,165 glycosylation sites, 186 giycosyJphosphatidylinositol (GPO anchors. 73, 74, 75 glyceroaquaporins, 253-254 GOF (gain-of-function) mutants, 238 Goldman-Engelman-Steitz (GES) scale, 139, 143 Gorter, E., 5 Gouy-Chapman theory, 79 GPCRs. See G-protein coupled receptors (GPCRs) GPI (glycosylphosphatidylinositol) anchors. 73, 74, 75 G-protein coupled receptors (GPCRs). 140-141,226 G-protein coupled receptors (GPCRs) activation mechanisms. 234 function of, 227 genomic and proteomic studies of. 234 gramicidins, 52, 87, 88, 89, 98, 132-133 greasy slide, 122, 123 green fluorescent protein, 144, 145 Green, David E .. 5,6 Grendel. F, 5 GROMOS.198 Gronhuss mechanism, 256-258

group translocators, 134, 135-137 GTPases, 173 guanine nucleotide binding proteins (G proteins), 140 GUVs (giant unilamellar vesicles), 62 GuxG motif, 154,265,267,312 half-helices. 93,312 Halobacter salinarum, 102, 103 halorhodopsin, 154 HDL (high-density lipoproteins), model bilayer system of, 66 head groups, 4 binding sites For, 76 heat capacity, of lipid bilayers, effects of peripheral proteins on, 76, 77 helical bundles, 102,213,214 conFormational changes in. 311 in bacteriorhodopsin, 102-107 in photosynthetic reaction center, 107-108 topology of, prediction of, 147 Helical Hairpin Hypothesis, 162 helical symmetry, 312 helix distortion, in integral proteins. 92-93,311-312 helix packing, 154, 164, 165,312 helix pair motifs, genomic analysis of, 154 helix tilting, 94, 99 in MS channels, 239 helix unwinding, 93 helix-helix interactions, 93,154,156 hydrogen bonding in, 154. ISS. 156. 164 in helical bundles, 102,312 in TolC tunnel. 306 triplets in, 154, 155 helix-tum-helix structures, in voltage paddles, 262 heme binding site, prostaglandin H 2 synthase. 218, 221 heme subunits cytochrome-bcl complex. 279-280. 281 reaction center, 110 hemifluorinated surfactants, 45 hetero-oligomers, 3\ 0 hexagonal phase. 28, 29-31 NMR spectrum of, 39, 40 types of, 28, 29 HHDBT (5-n-heptyl-6-hydroxy-4, 7-diozobenzothiazole),282 hidden Markov models (HMM), 150 high-sensitivity titration calorimetry, 61 high-density lipoproteins (HDL), model bilayer system of, 66 high-potential iron-sulfur protein (HiPIP), 1 \2, 113

histidine, 135 histidine-containing phosphocarrier protein (HPr), 135 HMM (hidden Markov models), 150 HMMTOP, 145, 150 holins, 132-133 homo-oligomers, 310 hop diffusion, 25, 27 HOQNO (2-heptyl-4-hydroxyquinolineN-oxide), 226, 229 HPr (histidine-containing phosphocan'ier protein), 135 Huber, Robert, 107,207 humans ABC transporters, 135 aquaporins, 253,254 ATP generation, 272 G-protein coupled receptors. 140, 226,227 mechanosensitive channels. 226-227 P450 cytochromes, 130 hydrocarbon chains, torsion angles in, 15,16 hydrocarbon-packing energies, 30 hydrogen bonding. See also specific protein importance of, 311 interhelical, 154. 155, 156, 164, 311 MD simulations, 200 protein Folding, 162, 164 transmembrane secondary structures. 92, 102 water, 4 hydrophobic effect, 3, 4-5 in lipid phase transitions, 30, 31 in protein folding. 162 hydrophobic interactions, protein binding by, 69, 78, 80, 82 hydrophobic mismatch, 98-99, 100 hydrophobicity measurement of, 4, 5 of ,ti-barrels, 157 positive-inside rule overriden by, 185 hydrophobicity plots, 141, 143, 144 combined with positive-inside rule, 145 generation of, 144 hydrophobicity scales, 142, 143, 144 for protein insertion studies, 182, 183 IMS (intermembrane space), 250 inclusion bodies, protein refolding from, 170 inhibitors. crystallization in presence of,310 insulin receptor, 139 integral lipids, 204, 206

Index integral proteins, 68, 90-94. See also specific protein amino acid distribution in, 145 classification of, 90, 91, 204 crystallization oF, 207 environmental factors affecting, 91 hydrophobicity plots of, 143 lipid interactions with, 94-98, 99 lipids associated with, 204 numbers of lipids in, 205 structure of, predicting, 141-143 interaction domains, in annexins, 72 interdigitation,27 INTERFACE-3, 154 interfacial region, 10 insertion of amphipathic helix into, 76, 82 inten:nembrane space (IMS), 250 inward-rectifying channels, 258 ion, 138-139 ion carriers, non-ribosomally synthesized, 133 ion channels, 138-139 patch clamp recordings of, 55-56, 57, 58 planar bilayers used to study, 53-55 ion exchange chromatography, 69 ionic strength, effect on micelle formation, 46, 47 ionophores, 87, 133 ionotropic ATP receptors, 139 iron receptors, 123-124 structure of, 123, 124 iron-sulfur proteins, in cytochrome-bel complex, 281 isoforms (isozymes), 129 isoprene, 20 KcsA,259 binding sites, 260 gating and conformational changes, 260-263,264,311 potassium conduction mechanism, 260,261 protein folding in, 169, 170 structure and selectivity, 259-260, 261 kinetic crystallography, 107 of F l Fa-ATPase, 277 of protein folding, 161 KirBacl.1, 259, 311 KirBac1.3, 259, 3J J knob-into-hole interactions, 93, 154, 164,165,306,312 Krafft point, 47 KvAP, 259, 3JJ, 3J2 voltage paddle, 262, 263 Kyte-Doolittle algorithm, 143

lactose permease (Lac carrier; LacY), 1-3,137,241,242-244 compared to GlpT, 245-248 conformational changes in, 244, 311 crystallization of, 207, 208 protein folding in, 100,169,170 structure of, 242, 243 symmetry in, 312 symport in, 242, 243, 244 transport mechanism, 132, 133 LacZ (tl-galactosidase), 144 LamB (maltoporin), 61, 118, 122-123 lamellar phases, 28 La (liquid crystalline), 28,191,193 Lb (gel), 28 Lc (crystalline), 23, 28, 191 NMR spectrum of, 39, 40 Langmuir isotherm, 79 Langmuir trough, 50 large unilamellar vesicles (LUVs), 60, 61 lateral diffusion, 24 measurement oF, 24, 26 rate of, 25 lateral domains, 33-34, 36 lateral mobility, of membrane components, 7, 8 lateral pressures effect on MS channels, 238 in protein folding, 167, 169 lauryldimethylamine oxide (LDAO), 43, 49,109 LCIC (ligand gated ion channel) da tabase, 139 LDAO (lauryldimethylamine oxide), 43, 49,109 leader peptidase (Lep), 145, 146. See also signal peptidase in topogenesis, 183, 184 length scales, in simulations, 202 lethal factor (LF), 85 LH1, 111, 112 LH2, 111, 112 ligand binding, 79, 83, 209-210, 234 ligand gated ion channel (LCIC) database, 139 light-harvesting complexes, in reaction centers, 111,310 light-induced signal transduction, efficiency of, 229 line tension, 34 linear isoprenoids, 17, 18,20-23 lipid anchor, 76 lipid bi/ayers, 7 components of, 13 diffusion of, 24-25 rate of, 25

heat capacity of, effects of per"ipheral proteins on, 76, 77 matrix, 23-24 models, 10, 196,212 molecular, 24, 25, 196 organization of, 31-33 planar, 53-55 formation of, 53-54, 55, 56 reversible interactions of peripheral proteins with, 76 simulations of, 210-212 stresses in, protein folding and, 167, 169 structure of, 24,191-192 supported, 57, 59 thickness of, 22, 26-28, 98 x-ray crystallography of, 23, 24, 28, 191 lipid chain, insertion of, protein binding by, 80 lipid clamp, 71 lipid cofactors. See nonannular lipids lipid mono layers, 50-53 curvature of, 30, 40 formation of, 50, 51 interaction with signal peptides, 52 phase changes in, 51 roleof,31,40 lipid phases, categories of, 28 lipid raf-ts, 8-9, 10,33-36 clustering of smaller domains in, 36 domain formation in, 57 effects of protein binding on, 78 GPI-anchored proteins associated with,75 location of, 34 models of, 34 physical properties of, 34, 35 proteins in, 9, 34, 90 used as markers, 36 size oF, 34 lipid shells, 36 lipid(s). See also specific lipid asymmetry, 26-27, 28 boundary layer of, 8 classes of, 17, 18,94,204 complex, 17 configuration of, 204, 205 crystallization of, 207 definition of, 4 diversity of, 13, 37-40 effects of peripheral protein binding on, 76-78 headgroups, 4 interactions between, 22, 23, 33 phase transitions, 28, 29,191,192 polymerized, 57 polymorphism, 28, 30

Index

326

lipid(s) (COI1/") proportion of, 7, 8, 17 removal of, 50, 206 requirements fOl~ 94, 95, 127 selectivity for, 95, 98, 99 stoichiometry of, 95, 99 structu res of, 4 surface concentration of, 128 trafficking patterns, J0 volume of, determination of, 192 lipid-anchored pt'oteins, 68, 73-74, 75, 76, 90 lipid-protein interactions, 127 integral, 94-98, 99 peripheral, 77, 78-80 x-ray crystallography and, 203-207 lipolytic enzymes, 214 liposome swelling assay, 60, 6J, 122 Iiposomes, 42, 60-63 types of, 60 uses of, 60 liquid crystal theory, 193-194, 195 liquid crystallography, 192 liquid-disordered state (Ld), 33 liquid-ordered state (L o ), 33, 34, 35 extraction of, 36 loss-of-function (LOF) mutants, 238 LSGGO motif, 135,136 lungs aquaporinsin,253,254 pulmonary surfactant, 52 LUVs (large unilamellar vesicles), 60, 61 lysine, 105 Iysophosphatidylcholine, 238 lysophospholipids,47 MJ 3 coat protein, 98 MacKinnon, Roderick, 207, 241 major facilitator superfamily (MFS), 133,242 paradigm fOI~ 248, 249 symmetry in, 3 J 2 MalK dimer, 136 maltoporin (LamB), 61, 118, 122-123 maltose, 135 mass spectrometry, 64 Maxwell-Boltzmann distributions, 198 MBD (membrane-binding domain), of prostaglandi n H2 syn thase, 219 MCF (mitochondrial carrier family), 138, 249. See a/50 ADP/ATP carrier motif,252

MD simulation. See molecular dynamics (MD) MDOs (membrane-derived oligosaccharides), 129 MDR (multidrug resistance) system, 296, 300 MDRI (multidrug resistance protein 1), 135, 186 mechanosensitive (MS) ion channels, 226, 235 channel opening models, 238, 239 classification of, 235 gating mechanisms, 238-239 large (MscL), 235-236, 237, 238, 239 mini (MscM), 235 small (MscS), 235, 237-238 melillin, 87, 88 membrane binding, See peripheral protein binding membrane damp, 76 membrane components, 1, 3, 8. See a/50 lipid(s); protein(s) lateral mobility of, 7, 8 proportions of, 7, 8,17 tools for studying, 42 membrane fluidity, 24 measurement of, 8-9 membrane fusion proteins, 302-303 Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs (Mistic), J70 membrane permeability, 1-3 membrane rafts, 3 membrane receptors, 139 membrane scaffold proteins (MSPs), 66 membrane solubilization, by detergents, 48-49, 50,129 membrane structure early models of. 5, 6 paradigms of, 3 shift in, 8-10 membrane thickness, 22, 26-28, 98 membrane(s) basic functions of, 1 dynamic nature of, 3 electrochemical gradient across, 53 membrane-binding domain (MBD), of prostaglandin H 2 synthase, 219 mem brane-derived oligosaccharides (MDOs),129 membrane-spanning proteins, in supported biJayers, 57 MEMSAT, 145,147 menaquinol (MOH2), 224 menaquinone (MOl. 224, 226 Metall, 234

MelhanobaCleriu 111 lhennoaulotrophicus, 259 Melhanococcus janllaschii translocon, 175,286-287,288 MexA, 303, 304 MFS. See major facilitator superfamily (MFS) micelles crystallization and, 207, 208 formation of, 45, 46, 47 mixed, 63 size of, 48 surface dilution kinetics 01',128 Michaelis-Menten kinetics, 129, 132 Michel, Hartmut, 107,207 microdomains, in blebs, 64 misfolding diseases, 187-189, 190 Mistic (Membrane Integrating Sequence for Translation of Integral Membrane Protein Constructs), 170 mitochondrial carrier Familv (MCF), 138,249. See a/50 ADP/ATP carrier mitochondrial carrier family (MCF) motif,252 mitochondrial inner membrane, composition of, 7, 8 mitochondrial membranes, 249 mitochondrial proteins, import of, 170-175,177 mixed micelles, 63 MLV (mulLilamellar vesicles), 60 protein binding to, 83 model membranes, 42, 50 for folding studies, 162 of lipid bilayers, 10, 196 peptide insertion into, 87 molecular dynamics (MD), 197-202 aquaporins, 258 ATP synthase, 278, 286 BtuCD complex, 293-294 calculations, 198 examples 01',198,199,200,212, 220 phases of, 199 steps in, J97-198 molecular models, of lipid bilayers, 24, 25, 196 monoamine oxidase, 90 monotopic proteins, 68, 69, 90 Montal, M., 53-55, 56 Monte Carlo (MC) simulations, 202-203 configurational-bias, 203 example of, 203 motifs, 311-312. See a/50 specific motif MPTopo, 144,157

Index MQ (menaquinone), 224, 226 MS channels. See mechanosensitive (MS) ion channels MsbA,210 MscL channels, 235-236, 237, 238, 239 MscM channels, 235 MscS channels, 235, 237-238 MSPs (membrane scaFfold proteins), 66 MthK, 259, 311 structure of, 260. 261, 262 Mueller, P., 53-55, 56 multidrug resistance (MDR) system, 296,300 multidrug resistance protein I (MDRI), 135,297 multilamellar vesicles (MLV), 60 protein binding to, 83 multiple sclerosis, 70 multiple sequence alignments, 148 multiprotein assemblies. See protein assemblies multi-spanning membrane proteins, topology of, 186 muscarinic acetylcholine receptor, 139 lVlycobaclerium luberculosis, 235, 236, 237 myelin basic protein, 69, 78 myelin proteolipid, 95, 98 myristate, 73, 74 myristoyl switch, 83, 84 Na+ channels, voltage-gated, 264 Na+K+ -ATPase, 134, 138 transport mechanism, 134 NADPH oxidase, 134 nanodiscs, 66, 67 nanomachines, 307, 312 native gel electrophoresis, 156 NBDs. See nucleotide-binding domains (NBDs) Neher, Erwin, 55, 57 neonatal respiratory distress syndrome, 52 neural networks, J 48 neurotransmitter receptor superfamily, 139-140 neutron diffraction, 191 combined with x-ray crystallography, 194-196 of protein-detergent complexes, 49, 50 techniques, 193 Newton's laws of motions, 197, 198 Nicolson, G. L., 6, 7, 33 nicotinic acetylcholine receptor, 139-140,239 nigericin, 133 nitrate reductase, 222-224

327

nitrate respiratory pathway, 222, 224 NMR. See nuclear magnetic resonance (NMR) Nobel Symposium on Membrane Proteins (2003), 309 nonannular lipids, 94, 204, 205 numbers of, 205 nonpolar domain, 10 non-ribosomally synthesized channels, 132-133 non-ribosomally synthesized ion carriers, 133 nonsteroidal anti-inflammatory drugs (NSAIDs), 216, 222 NPT ensemble, 197, 200 NSAIDs (nonsteroidal anti-inflammatory drugs), 216, 222 NTP ensemble, 202 nuclear magnetic resonance (NMR), 116 fJ-barrels, 116. 168 comparison of crystal structures with,210,211 Fo domain. 274-275 hydrophobic mismatch, 98 lipid bilayer, 24 lipid phase transitions, 39, 40 Mistic, 170 OmpA, 118, 168 OmpX, 118 pore formation, 88, 89 nucleoside channel porin (Tsx), 118 nucleotide(s), 83 nucleotide-binding domains (NBDs) AAC, 251 ABC transporters, 135, 136,292,293, 297,298-299 NVE production phase, 200 NVT ensemble, 197,202 OG (octyl fJ-D-glucoside), 43, 50 oleic acid, 15 oligomerization, 309-310. See also dimer formation protein-protein interactions and, 310 purification process and, 310 x-ray crystallography of, 310 oligomycin, 273 oligomycin-sensitivity conferring protein (OSCP subunit), 273 OmpA porin, 118 protein folding in, 168, 169 structure of, 210 OmpC pol'in, J 18, 119-J20 OmpF porin, 49, 50, 53, 118, 119-120 black films of, 54 blisters and, 65, 66

channel properties of, 121 crystallization of, 208 structure of, 120 OMPLA. See outer membrane phospholipase A (OMPLA) OmpR porin, 119 OmpX porin, 114, 115, 118 structure of. 210 OprM,307 optical spectroscopy, 108 organophosphate phosphate antiporter family, 247, 309 orphan proteins, 156 OSCP subunit (oligomycin-sensitivity conferring protein), 273 osmoporin (OmpC porin), 118, 119-120 osmotic pressure, mechanosensi tive responses to, 235 octyl fJ-D-glucoside (OG), 43,50 outer membrane phospholipase A (OMPLA), 115,214-216 calcium binding sites, 214, 216, 217 catalytic triad, 214, 215 dimer formation. 215, 216, 310 physiological role of, 214-215 protein folding in, 170 structure of, 215 ovispirin, 88 oxidoreductases, 134 OxIT, 248, 31 I oxygen reduction, by cytochrome-c oxidase, 285 p bulge, 93 P450 cytochromes, 129, 130-131 PagP, 210, 21 I pairwise sequence alignments, 142 palmitoyl, 73, 74 palmitoyJ-sphingomyelin (PSpM), 34, 35 Paracoccus denilrificans, 285 paradigms of membrane structure, 3. See aL50 Fluid Mosaic Model; hydrophobic effect shift in, 8-10 passive transporters, 131, 253 patch clamps, 55-56, 57, 58 blisters, 64 mechanosensitive channels, 235, 238 patterns, 311-312. See also specific pattern Pc. See phosphatidylcholine (PC) PCC (protein-conducting channel). See translocon PE (phosphatidylethanolamine), J 7, 38,39,109

Index

328

PEP PTS (phosphoenolpyruvatedependent phosphotransferase system), 135 peptides, transmembrane, 87-88. See also transmembrane (TM) domains; specific peptide peptidoglycan, 113, 1 19 peripheral pl'Otein binding, 76 domains in, 80-83 effects on membrane lipids, 76-78 ligands in, 79, 83 modulation of, 83-84 sites of, 76, 80 peripheral protein(s), 10,68,69-72 amphitropic, 72-73 embedded in membrane, 90 interactions between lipids and, 77, 78-80 reversible interactions with lipid bilayer, 76 permeability barrier, 1-3 blebs and, 64 changes in, pressure-induced, 226 lipid-protein interactions and, 94 porins and, 118 translocon and, 175 permeases, 127 peroxidase (POX), 218, 220, 221 pertussis toxin, 85 PFam-A database, 153 PG (phosphatidylglycerol), 17,38 PGH 2 (prostaglandin H 2), 216 PGHS. See prostaglandin H 2 synthase (PGHS) P-glycoprotein (multidrug resistance protein 1), 135, 297 PH (pleckstrin homology) domains, 80, 81,82 phase diagrams, 31,32 detergent-composition, 47 ternary, 32, 33, 35 phase transitions bacterial membranes, 16, 40 blebs, 64 fallyacids, 15, 17,29 lipid, 28, 29,191,192 lipid monolayers, 51 nanodiscs, 66, 67 PHD, 145, 147, 148 PhoE (phosphoporin), 118, 121, 122 PhoE (phosphoporin) channel properties of, 121 protein folding in, 170 phosphatidic acid, 17, 129 phosphatidyl choline (PC), 15, 17 structure of, 205

phosphatidylethanolamine (PE), 17, 38,39, 109 phosphatidylglycerol (PG), 17,38 phosphatidylinositol (PI), 17 phosphatidylinositol3-phosphate (PIP3),82 phosphoenolpynJvate-dependent phosphotransferase system (PEP PTS), 135 phosphoglycerate kinase, 161 phosphoinositides, 80 phospholipase A2 (PLA2), 71, 128 binding sites, 76 phospholipase C, 71, 72 phosphoJipases, 70 Hap movement in, 71 x-ray crystallography of, 71 phospholipids (PLs), 17-18,20 anionic, 18 interactions with cholesterol, 7, 22, 33 MD simulation of, 199 mixed micelles of, 63 phase changes, 51, 67 properties of, 18 simulations of, 198, 202 species of, 37 structures of, 15, 19, 192, 195 zwitterionic, 18 phosphoporin (PhoE), 118, 121, 122 channel properties of, 121 protein folding in, 170 phosphotidylserine (PS), 17 photoreaction cycle, in bacteriorhodopsin, 105, 107 photoreceptors, retinal, 228, 229 photosynthesis, pmcess of, 107 photosynthetic reaction center antennae in, 111-112,310 cardiolipin in, 205 cofactors in, 109-110 helical bundles in, 102, 107-108 helix-helix interactions in, J 54 hydrophobic mismatching in, 100 light-hal\/esting complexes in, III lipids in, 109 oligomerization in, 310 proteins in, 108-109, 144 reaction cycle in, 112-113 structure of, 108, 109 surface of, 109,206 transport classification, 133-134 photosystems, types of, 108, 112 phototransduction cycle, in rod ceJJs, 229-230 PI (phosphatidylinositol), 17 PIP3 (phosphatidylinositol 3-phosphate), 82 PKC. See protein kinase C (PKC)

PLA2 (phospholipase A2), 71, 128 binding sites, 76 planar bilayers, 53-55 formation of, 53-54, 55, 56 plants P450 cytochromes in, 130 photosystems in, 108 pldA gene, 215 pleckstrin homology (PH) domains, 80, 81,82 PLs. See phospholipids (PLs) pmf (proton motive force), 272, 279, 296 polar clamp, 155, 156 polyethylene glycol, 57 polymerized lipids, 57 polyphosphorylated inositol, 80 polystyrene beads, detergent adsorption to, 50 poly topic proteins, 68, 69, 90 classification of, 152 genomic analysis of, 152, 153 pmtein insertion in, 184 topogenesis in, 184, 186 polyunsaturated fatly acids, 15 MD simulations of, 200-201,202 POPC (l-palmitoyl-2-0Ieoyl phosphatidylcholine) in lipid rafts, 34, 35 MD simulation of, 200, 220 pore architecture, in potassium channels, 259, 261,262 pore formation cholesteml, 55 colicins, 86, 87 peptides, 88, 89 pore(s) conduction, 253 gated, 131, 132, 288 general import, 178 porins, 8, 118-119,241. See also specific porin Ii-barrels in, 113, 119, 120, 121, 132-133,157 channel properties of, 119, 121 classification of, 118 detergent-solubilized, 119 mutations, 119 number of, 119 purification of, I 19 specific, 120-122 structure of, 114, 119, 122 trimeric, 157 porters, 133 positive-inside rule, 144-145, 146, 184, 185 combined with hydrophobicity plots, 145 factors overr'iding, 185, 186

Index posttranslational translocation, J70 potassium channels, 258-259 conformational changes in, 3 J I diversity of, 258 gating mechanisms, 259 inward-rectifying, 258 selectivity of, 258 stnJcture of, 259 potential energy, 197, 198,202 POX (peroxidase), 218, 220, 221 precipitation, detergent removal by, 50 pressure in phase diagrams, 31 lateral. in protein folding, 167, 169 permeability changes caused by, 226 primary active transporters, 133-134 prion protein (PrP), 186, 187 GPI anchor of, 75 probability distributions, 192, 193, 194, 197 probes, in phase diagrams, 31-32 production phase, 199, 200 prokaryotes. See also bacteria; specific species anaerobic respiration in, 222, 224 tJ-barrels in, 114 cell membranes in, 1,2 lipid diversity in, 21, 22, 37 mechanosensitive channels in, 235 protein export in, 170 translocation systems in, 160, 175 transJocons in, 175 transporters in, 248 prostaglandin H 2 (PGH2), 216 prostaglandin H 2 synthase (PGHS), 90, 214,216-222 catalytic domain, 2 J7, 220, 221 EGF-domain, 219, 220 MD simulation of, 220 mechanism of action, 218 structure 01',218,219,220 protease protection assay, 171 protease sensitivity, 143 protective antigen (PA), 85 protein assemblies, 10,271-272. See also specific complex as nanomachines, 307, 312 genomic analysis of, 156 respiratory chain, 279 Protein Data Bank, 141, 142 protein folding, 160, 161 a-helical,162-164 bacteriorhodopsin, 164, 166-167 tJ-barrels, 167-168, 169 bilayer stresses and, 167, 169 environmental factors in, 91 helix packing in, 164,165 hydrophobic mismatch in, 100

329 in vilro studies 01',161-162,170

stages in, 161, 162, 163, 164 thermodynamics of, 160, J63,I69 protein insertion, 160, 169, 178-184 biological scale for, J83 cross-linking studies of, 178, 179, 182 protein kinase C (PKC), 72 activation of, 73 binding sites, 80 surface dilution kinetics of, 128 protein misfolding diseases, 187-189, 190 protein sequences databanks of, 141, 142, 154 genomic information on, 141 protein sorting, hydrophobic mismatch in, 100 protein toxins. See toxins protein translocation, 171,175,178 apparatus, 160,175 colicins, 85 cotranslational, 170, 176 GlpT, 242, 245, 247 LacY, 242, 243 posttranslational, 170 process of, 170-175,177 proteins involved in, 171, 175, 178 toxins, 84, 85 translocon-ribosome complexes, 288-289 proteinO, trafficking, in rafts, 36, 37 protein(s), 1,7. See also specific protein protein(s) biogenesis of, 160, 170, 182 bioinformatics tools 1'01',141,142 classification of, 68-69,127, 14J. 213 denaturation of, 161 export from cytoplasm, 170-172, 174,175,177 functions of, 127 genomic analysis 01',145-154 in blebs, 64 in detergent-resistance membranes, 36,37,49 lipid requirements of, 94, 95 orientation of, 143-144, 145 orphan, 156 proportion of, 7, 8 protrusion beyond membrane, 312 raft, 9, 34 used as markers, 36 reconstituted into nanodiscs, 66 topogenesis in, 170, 184-187 transport, 131-132 protein-conducting channel (PCC). See translocon protein-lipid interactions, 94-98, 127 x-ray crystallography and, 203-207

protein-protein interactions, 7, 9 disruption of, during purification process, 310 in AAC, 253 oligomerization and, 310 proteoliposomes, 60, 62 proteomics, 156, 157, 234 protomers, 271 proton motive force (pmf), 272, 279, 296 proton pathways aquaporins, 256-258 bacteriorhodopsin, 106,285 cytochrome-bcl complex, 283, 284-286 F 1 Fo-ATPase, 277 LacY, 243, 244 proton symporters, bacterial. 137 proton/drug antiporters, 299, 300 PrP (prion protein), 186,187 GPI anchor of, 75 PS (phosphotidylserine), 17 Pseudomonas aeruginosa, 303, 306, 307 PSI, 108 PSIBLAST, 142 PSI!, 108 PSpM (palmitoyl-sphingomyelin), 34, 35 PufX,112 pulmonary surfactant, 52 purple bacteria antennae system in, 111-112 photosynthetic reaction centers of, 108, 109, 112 purple membranes, 103 composition of, 104 Q cycle, 281, 282 quenching processes, 26 quinone(s), reaction centers, 109 quinone-binding sites, cytochrome-bcl complex, 281, 282

RCK domain, 261, 262 reaction center (RC), 107. See also photosynthetic reaction center reaction cycle, photosynthetic, 112-113 reconstitution, 42, 50 detergent removal for, 50 goal of, 309 recoverin, 83, 84 redox loop, 214, 218, 224 redox potentials, nitrate respiratory pathway, 223, 224 redox sites, cytochrome-bcl complex, 280, 281 reduction potentials, heme subunit of reaction center, 110 reporter enzymes, 144

Index

330

Resistance Nodulation cell Division (RND) superfamily, 297 respiratory chain complexes, 279, 280 classification of, 279 protrusion beyond membrane, 312 respiratory distress syndrome, neonatal, 52 retinal, 104, lOS, 106 analogs, 105 in pmtein folding, 166 in rhodopsin, 231,232,233 isomeriz.ation of, 105. 107 retinal photoreceptors, 228. 229 retinitis pigmentosa. 187, 189. 190, 232 reverse signal-anchors. 184, 185 Rhodobacter capsulatus, 113, 114. 119, 145 RJ1Odobacter sphaeroides cytochrome-c oxidase, 284, 285 integral lipids. 206 reaction centers, 108,109,110,111, 112 Rhodopseudornonas acidophila, I I I Rhodopseudomonas blastica, 119 Rhodopseudomonas sphaeroides. See RJ1Odobacter sphaeroides RJ1Odopseudomonas viridis. See Blastochloris viridis rhodopsin, 95, 226, 227 activation of, 229 cytoplasmic domain, 234 DHA in, 201 dimer formation, 234 ex tracell ula r- transmem brane interface in, 232 function of, 228-230 hydmgen bonding in, 232, 311 hydrophobic mismatching in, 99, 100 mutations, 189, 190 01 igomerization state of, 234 polar clamp in, 156 (-etinal in, 231, 232, 233 structure of, 103, 140,209,211. 227, 228, 230-235 TM helices in, 232. 243 ribosome, translocon bound to, 288-289,290 ripple phase, 28, 29 RND (Resistance Nodulation ceJl Division) superfamily, 297 RnfA, 145, 146 RnfE, 145, J46 Robertson, J. David, 6 rocker-switch type mechanism, in translocation, 242, 245, 247 rod cells, 228, 229 hyperpolarization in, 228. 229 phototransduction cycle in. 229-230

mtational diffusion, 24 RxxRR motif, 248 SAl protein. 182 Sakmann, Bert, 55, 57 salt bridge, in AAC, 251, 252 saponins, 43. 55, 56 sarcoplasmic reticulum calcium ATPase (SERCAI) conformational changes, 265, 266, 267, 268 structure of, 264, 265, 266, 267 Sav 1866, 135, 290, 297-299, 312 coupling helices, 298-299 drug binding to, 297-298 structure of, 297, 298 scattering length distribution, 193 Schiff base, 104, 105, 106 Schindler, H., 55 SCQP (structural classification of proteins), 142 ScrY (sucrose pOlin). 118 SDS. See sodium dodecyJsuJfate (SDS) SDS polyacrylamide electrophoresis (SDS-PAGE), 171, 172 SDSL (site-directed spin labeling), 97 sea anemone toxins, 87 Sec (secretion) proteins, 174, l75, 176, 312 SecA, 88-90, 173, 174, 286, 290 SecB, 173, 174 SeeD, 174, 175,290 SecDFYajC, 286 SecE, 175 SecF, 175, 290 SecG, 175,286 secondary active transporters, 137-138 SecY, 175,311 structure of, 286-287, 288 tl'anslating versus nontranslating states, 289 SecYEG, 156, 174, 175,286 ribosome binding, 289-290 structure of, 288, 289 sensory rhodopsin II, 154 SERCA 1. See sarcoplasmic reticulum calcium ATPase (SERCAI) serine z.ipper motif, 155 serotonin, 139 serpentine receptors, 140 short-chain/long-chain unilameJlar vesicles (SLUVs), 62 signal peptidase, 176, 184. See also leader peptidase (Lep) signal peptides, 52 interaction with monoJayers, 52 signal recognition particle (SRP), 172, 173,174

signal transduction in rafts, 36 integral proteins in, 92 light-induced, efficiency of, 229 signal(s) initiating protein export, 170, 171, 172. 173, 177 stop transfer, 176, 178, 184, 186 that govern topogenesis, 184, 185, 186, 187 signal-anchors, 173, 176, 182, 184, 185 reverse, 184, 185 signaling proteins calcium, 264 in raf-ts, 36 isoz.ymes as, 129 membrane receptors as, 139 simulation,191,210-212.Seealso molecular dynamics (MD); Monte Carlo (MC) combined with x-ray crystallography, 210-212 lipid bilayers, 196-197 Singer. S. J., 6, 7, 33 site-dil-ected spin labeling (SDSL), 97 sitosterol, 2 J, 22 small unilamellar vesicles (SUVs), 60, 61 smectic liquid crystals, diffraction studies of, 193, 195 SMS,61 sn-glycerol-3-phosphate, 17 snorkeling by transmembrane domains, 92 of amino acid residues, 184 sodium channels, voltage-gated, 264 sodium cholate, 43 sodium deoxycholate, 43 sodium dodecylsulfate (SDS), 43 precipitation of, 50 protein unfolding in, 166, 169 solubility of, 43 sonication, raft isolation with, 36 special pair, J 10, J J2 specific porins, 120-122 spectroscopic rulers, 168 sphingolipids, 17, 18,20 sphingomyelins, 20, 21, 98 sphingosine, 20, 2 J spider toxins, 87 spin probes, 97, 238 squalene-hopene cyclase, 90 Src peptide, binding by, 78, 80 SRP (signal recognition particle), 172, 173.174 Staphylococcus aureus. 85. See also Savl866 stator, 274 stearic acid, 14, 15, 200, 202

Index sterols, 17, 18, 20-2 I, 23 STGA, 206, 207 stigmasterol, 2 I, 22 stop transfer signals, 176, 178, 184, 186 stopped-flow absorption spectroscopy, 166 Streptomyces lividans, 259 stll.lctural classification of proteins (SCOP),142 substr·ate concentration, calculation of, 129 substrate-analogs, crystallization in presence of, 3 J 0 sucrose porin (ScrY), I 18 sugars, group translocation of, 135 supported bilayers, 57, 59 surface concentration terms, 23 surface dilution effects, 128 surface pressure vs. area isotheml, in monolayer formation,S I surface tension measurement of, 43 micelle formation and, 47 surfactants amphiphilicity of, micelle formation and,47 definition of, 43 hemiAuorinated,45 pulmonary, 52 SUVs (small unilamellar vesicles), 60, 61 Swiss-Prot, 141, 142, 154 switch helix, in TonB-dependent transporters, 295 symport, J 31, 137, 138 in LacY, 243, 244 symporters, 137-138 targeting complexes, in protein export, 176 TAT (twin arginine translocation) pathway, 176 Tay-Sachs disease, 17, 18 TBDTs (TonB-dependent transporters), 294 structure of, 295, 311 Tb-MscL,235,236,237 TC (transport classification) system, 132-133,134,158 TOG (t!-D-galactopyranosyl I -thio-{l-D-galactopyranoside), 243 temperature in phase diagrams, 3 I micelle formation and, 47 temperature factor (B values), 207, 3 II TEMPO, 32 terpenes, 73, 74

331

tetanus neurotoxin, 85 tetraheme cytochrome subunit, reaction center, I 10 TF (trigger factor), 174 thapsigargin (TG), 264, 266, 310 Thermochromatium tepidum, 108, 110, 112 thermodynamics in lipid phase transitions, 30 of F 1 Fo-ATPase, 276 of protein folding, 160,163, 169 TIM complexes, 178 time average (Aave), 197 time scales, in simulations, 202 time-averaged probability distributions, 192,193,194,197 TM domains. See transmembrane (TM) domains TMHMM, 145, 147, ISO, 156 TNBS (trinitrobenzenesulfonic acid), 27 TolC tunnel, 296, 297, 300, 302, 304-307 drug efflux systems dependent on, 306 opening mechanism, 306-307 partners of, 307 protrusion beyond membrane, 312 structure of, 300, 304-305, 306, 307, 310 TOM complex, 178 TonB box, 124, 295 TonB protein, 133,294 energy coupling, 296 structure of, 296, 297 TonB-dependent transporters (TBDTs), 294 structure of, 295, 3 11 topogenesis, 170, 184- 185, 187 topologv models helical bundles, 147 protein orientation, 144,145,147, 156 TopPred, 145 toroidal modeJ, 88 torsion angle, 15, 16 torsional potential, 199 toxins, 84-85. See also specific toxin channel-forming, 132-133 peptide, 87 TRAM (translocating chain-associated membrane protein), 174, 175, 176 tra11s fatty acids, 15,200 trans torsion angle, IS, 16 transcription translation systems, ;1'1 vitro, 170 transducers, 226-227 transducin, 228, 229, 234

transfer free energy change (Dc.Gtl')' 142,143 transient confinement zones, 76 translocase. See translocon translocating chain-associated membrane protein (TRAM), 174, 175,176 translocon(s), 170, 175, 176,286 dimer formation, 288 function of, 287 pathways through, 178, 179,288 structure of, 286-287,288, 289-290 TM insertion, 288 translocon-ribosome complex, 288-289, 290 ribosome contacts, 289-290 transmembrane (TM) domains, 69, 84, 90 amino acids in, 92 deletion of, for crystallization, 209 formation of secondary structure in, 92, J02 helical bundles, 102 prediction of, 141-143, 148 snorkeling by, 92 thickness of, 37 translocon, 288 types of, 81 transmembrane (TM) electr-on transfer carriers, 134 transmembrane (TM) helices in rhodopsin, 232, 243 numbering of, 243 transmembrane (TM) peptide segment bitopic proteins, 68 transport classification (TC) system, 132-133,134,158,297 tr·ansport mechanisms, 131, 132,134 AOPfATP carTier, 252-253 lac permease, 132 Na+K+-ATPase, 134 transport proteins, 131-132 classification of, 132-133, 134, 158 genomic analysis of, 157 transporters, 241-242 classification of, 242 role of, 241 transverse diffusion, 24 rate of. 25 transverse relaxation optimized spectroscopy (TROSY), 116 transverse view of membrane, 9 trigger faclor (TF), 174 trimeric pOi-ins, 157 trinitrobenzenesulfonic acid (TNBS), 27 tripartite drug efAux, 300 triplets, in helix-helix interactions, 154, 155

Index

332

tripod amphiphiles, 45 Triton X-I 00, 36, 43, 50 in protein-folding studies, 170 micelles, 63 surface dilution kinetics oE, 128 Trojan peptides, 87 TROSY (transverse relaxation optimized spectroscopy), 116 Tsx (nucleoside channel) porin, 118 twin arginine translocation (TAT) pathway, 176 ubiquinone, I"eduction by cytochmme C2, 112 uncoupling protein, 138 unilamellar vesicles, 60 giant (GUVs), 62 large (LUVs), 60, 61 short-chain/long-chain (SLUVs), 62 small (SUVs), 60, 61 uniport, 131 unit membrane, 5, 6 valinomycin, 133 van der Ploeg, P., J 97 van der Waals forces, 199 in helix packing, 164 x-ray crystallography and, 203 VceC, 307 vestibules, 253 vibrational energy, 199

Vibrio cholerae, 307 viral enveloped formation, domain segregation fOl~ 37 vitamin 8 12, J35, 291 structure of, 291 vitamin B l2 transporter (BtuBC), 135 vitamin B l2 uptake system, 291 energized phases of. 291 transport across inner membrane, 291-293,294 transport across outer membrane, 294-296 voltage clamp, 53 voltage gating, 262-263 voltage paddle, 262, 263 volume of lipid, determination of, 192 von Heijne positive-inside rule. See positive-inside rule

Walker A and B motifs, 135, 136 WalkeJ~ John E., 272 water channels. See aquaporins; glyceroaquaporins water solubility, of detergents, 43 water, hydrogen bonding in, 4 Wimley-White (WW) scale, 139, 143 Wuthrich, Kurt, 116 x-ray crystallography artifacts in, 209

combined with neutron diffraction data, 194-196 combined with simulations, 210-212 compared with NMR studies, 210, 211 conformational changes and, 310-311 crystallization pmcess in, 207-210 history of, 207, 309 ligand binding in, 209-210 lipid-protein interactions, 203-207 of bilayer thickness, 100 of helical bundles, 102 of integral proteins, 90 of lipid bilayer, 23, 24, 28, 191 oE oligomers, 310 of pore formation, 88, 89 resolution, 204, 207, 209 techniques, 193 uses of. 191 YajC, 175,290 yeast cytochrome-bcl complex, 280, 282, 283, 284, 285 inlegrallipids, 206, 208 lipid-anchored proteins, 75 mitochondrial Fl-c complex, 273 translocons, 286 yeast two-hybrid analysis, 156 YidC, 174, 175, 176, 286