Membrane Protein Purification and Crystallization: A Practical Guide, 2nd Edition

MEMBRANE PROTEIN PURIFICATION AND CRYSTALLIZATION Second Edition A PRACTICAL GUIDE

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MEMBRANE PROTEIN PURIFICATION AND CRYSTALLIZATION Second Edition A PRACTICAL GUIDE Edited by

CAROLA HUNTE Max-Planck-Institut fur Biophysik Frankfurt am Main, Germany and

GEBHARD VON JAGOW AND HERMANN SCHAGGER Zentrum der Biologischen Chemie Universitatsklinikum Frankfurt Frankfurt am Main, Germany

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Contents

Contributors xv Preface xvii

I

STRATEGIES AND TECHNIQUES

1

Purification Strategies for Membrane Proteins GEBHARD VON JAGOW, THOMAS A. LINK, AND HERMANN SCHAGGER I. Introduction 1 11. General Guide for Retaining Catalytic Activity 3

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Contents

III. Choice and Sequence of Purification Techniques 4 IV. Choice of Protein Source, Disruption of Cells, and Preparation of Organelles and Membranes 6 V. Protein Assay 7 A. Catalytic Activity and Other Specific Properties 7 B. Determination of Total Protein 8 VI. Solubilization and Stabilization of Membrane Proteins A. Choice of Buffer 9 B. Choice of Detergent 10 C. Protein Stability and Protease Inhibition 16 References 18

9

Techniques and Basic Operations in Membrane Protein Purification HERMANN SCHAGGER I. Introduction 19 II. Solubilization-Precipitation 20 A. Differential Extraction 20 B. Phase Separation and Differential Precipitation 24 C. Centrifugation 26 III. Concentration of Samples, Exchange of Buffer, and Exchange of Detergent 26 A. Sample Concentration 26 B. Exchange of Buffer 28 C Removal and Exchange of Detergent 28 IV. Chromatographic Techniques 29 A. Hydroxylapatite Chromatography 30 B. Ion-Exchange Chromatography 35 C. Chromatofocusing 38 D. Metal Chelate Chromatography 43 E. Affinity and Dye-Ligand Chromatography 45 F. Covalent Chromatography 48 G. Hydrophobic Interaction Chromatography 49 H. Gel Filtration 49 References 52

3

Production and Purification of Recombinant Membrane Proteins ETANA PAD AN, CAROLA HUNTE, AND HELMUT REILANDER I. Introduction

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Contents

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11. Prokaryotic Expression Systems for Overproduction of Membrane Proteins for Structural Studies 57 A. E. coli Inducible Promoters Used for Overexpression 58 B. Optimizing Expression Levels in E. coli 60 C Inclusion Bodies and Refolding 62 III. Eukaryotic Expression Systems for Overproduction of Membrane Proteins 63 A. Yeast-Based Expression Systems 64 B. Insect Cells as Expression System 70 IV. Use of Fusion Proteins and Affinity Tags for Purification of Membrane Proteins 73 Acknowledgments 77 References 78

A

SDS Electrophoresis Techniques HERMANN SCHAGGER I. Introduction 85 II. Choice of Optimal SDS-Page System and Gel Type 86 A. Wide versus Narrow Molecular Mass Range 86 B. Separation of Proteins with Similar Molecular Masses 86 C. Avoiding Broad Bands 88 D. Analytical versus Preparative Purpose 88 III. Tricine-SDS-PAGE 88 A. Comparison with Laemmli System 89 B. Characteristics of Gel Types 89 C. Gel Preparation 91 D. Sample Preparation and Protein Loading 93 E. Electrophoresis Conditions 95 F. Staining and Drying of High Percentage Polyacrylamide Gels 96 IV. Recovery of Proteins from SDS Gels 97 A. Recovery of Proteins from Fixed and Coomassie-Stained SDS Gels 97 B. Blue-SDS-PAGE 97 C Recovery of Membrane Proteins after Blue-SDS-PAGE 99 V. Processing of Proteins Recovered from Gels for Immunization and Protein Sequencing 100 VI. Electroblotting 101 References 102

C

Blue Native Electrophoresis HERMANN SCHAGGER I. Introduction

105

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Contents II. Techniques 106 A. Native First Dimension: BN-PAGE 106 B. Native Second Dimension: BN-PAGE + Detergent 116 C. Second or Third Dimension: SDS-PAGE 116 III. AppHcations 119 A. Final Purification of Crude Membrane Proteins for Immunization and Protein Sequencing 120 B. Separation of Native Membrane Protein Complexes from Solubilized Biological Membranes 120 References 129

^

Preparative Isoelectric Focusing IRENE TSIROGIANNI AND GEORGIOS TSIOTIS I. Introduction 131 II. Techniques 133 A. Instruments and Accessories 133 B. Analytical Electrofocusing 134 C. Preparative Flat-Bed Electrofocusing 136 III. Application: Purification of Photosystem I 138 IV. Conclusions 139 References 141

7

Membrane Protein Crystallization CAROLA HUNTE AND HARTMUT MICHEL I. Introduction 143 II. Bottlenecks in Membrane Protein Crystallization A. Starting Material 146 B. Protein Purification and Characterization 147 C. Detergents 151 D. Crystallization 154 E. Conclusion 156 References 156

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II Q

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DISCUSSION OF SELECTED ISOLATION PROTOCOLS Lipid-Dependent Inactivation and Reactivation of Bovine Complex III HERMANN SCHAGGER I. Introduction 161 II. Lipid-Dependent Inactivation and Reactivation 162 A. Delipidation 162 B. Reconstitution of Lipid-Dependent Catalytic Activity 163 References 165

Q

Purification of an Affinity-Epitope Tagged G-Protein Coupled Receptor CHRISTOPH REINHART, H MARKUS WEISS, AND HELMUT REILANDER I. Introduction 167 II. Production of Recombinant J82-Adrenergic Receptor in Pichia pastoris A. Media for Pichia pastoris 168 B. Growth and Expression of Pichia pastoris Clones 169 III. Preparation of Pichia pastoris Membranes 169 IV. Solubilization of Membranes 170 V. Purification of the j82-Adrenergic Receptor 170 A. Ligand Affinity Chromatography of j52-Adrenergic Receptor 170 B. Anti-FLAG Ml Antibody Affinity Chromatography 171 C. Affinity Chromatography of Biotinylated j52"Adrenergic Receptor by Monomeric Avidin Resin (Optional) 172 D. Analytical Gel Filtration (Optional) 174 VI. Ligand Binding Assays 174 A. Binding Assay on Membranes 174 B. Binding Assay for Solubilized Receptor 175 References 177

10

168

Purification of NhaA Na^/H^ Antiporter of Escherichia Coli for 3D or 2D Crystallization MIRO VENTURI AND ETANA PAD AN I. Introduction 179 11. Growth of Escherichia coli for Production of His-Tagged NhaA in a 10 1 Fermenter 180

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III. Isolation of Membranes from Escherichia coli TA16/pAXH Strain 182 IV. Two-Step Purification of His-Tagged NHaA 183 A. Affinity Purification of His-Tagged NhaA on the Ni^^-NTA Matrix 183 B. Purification of His-Tagged NhaA by Gel Filtration Chromatography 184 V. NhaA Reconstitution into Liposomes and Activity Assay 185 A. Reconstitution by Detergent Dilution 185 B. Reconstitution by BioBeads 187 C. NhaA Activity Assay: ApH Driven ^^Na^ Active Transport 187 VI. Dynamic Light Scattering Experiments 189 Acknowledgments 189 References 190

1 1

Purification of the Cytochrome bci Complex from Yeast HILDUR PALSDOTTIR AND CAROLA HUNTE I. Introduction 191 II. Preparation of Membranes from Saccharomyces cerevisiae 192 III. Cytochrome bc^ Complex Preparation 193 A. Detergent Solubilization of Membranes 193 B. Purification of the Cytochrome bc^ Complex by Anion-Exchange Chromatography 194 C. Size Exclusion Chromatography 196 D. Protein Determination 196 E. Spectroscopic Quantification of the Cytochrome bc^ Complex 197 F. Determination of Cytochrome bc^ Complex Activity 197 G. Analytical Size Exclusion Chromatography 198 H. SDS-PAGE Analysis 198 IV. Conclusions 200 Acknowledgments 201 References 202

III ^ O

CRYSTALLIZATION OF MEMBRANE PROTEINS Antibody Fragment Mediated Crystallization of Membrane Proteins CAROLA HUNTE AND AIMO KANNT I. Introduction 205 II. Production of Antibody Fv Fragments in the Periplasm of E. coli 210

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III. Purification of Fvs by Streptavidin Affinity Chromatography 210 IV. Fv Mediated Crystallization of Cytochrome c Oxidase from P. denitrificans 211 A. Growth of P. denitrificans 211 B. Isolation of Membranes 211 C. Solubilization of Membrane Proteins 212 D. Indirect Immunoaffinity Chromatography 212 E. Detergent Exchange and Size Exclusion Chromatography 212 F. Crystallization of Cytochrome c Oxidase-Fz? Complex 214 V. Fi;-Mediated Crystallization of the Cytochrome hc^ Complex from the Yeast S. cerevisiae 214 A. Co-Complex Formation and Purification 214 B. Crystallization of Cytochrome hc^ Complex: fi;18Ell Co-Complex 215 VI. Conclusions 217 Acknowledgments 217 References 217

13

Crystallization of Wolinella succinogenes Quinol: Fumarate Reductase C. ROY D. LANCASTER I. Introduction 219 II. Preparatory Steps 221 A. Growth of Wolinella succinogenes 111 B. Isolation of Quinol: Fumarate Reductase 222 III. Crystallization of Quinol: Fumarate Reductase 225 IV. Conclusions 226 References 227

14

Ba3-Type Cytochrome c Oxidase from Thermus thermophilus: Purification, Crystallization, and Crystal Transformation TEWFIK SOULIMANE, REINER KIEFERSAUER, AND MANUEL E. THAN I. Introduction 229 II. Fermentation, Purification, and Molecular Characterization 231 A. Fermentation 231 B. Preparation Protocol 231 C. Molecular Characterization 234 III. Crystallization and Initial Crystallographic Characterization 236

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A. Detergent Exchange 236 B. Crystallization in Sitting-Drops 237 C. Crystallization in Batch 238 D. Initial Crystallographic Characterization 239 IV. Crystal Transformation 240 A. Oil Method 240 B. Transformation and Freezing with the Humidity Machine 243 Acknowledgments 249 References 249

15

Two-Dimensional Crystallization of Membrane Proteins: A Practical Guide WERNER KUHLBRANDT I. Introduction 253 II. Things to Consider Before You Start 254 A. 2D or 3D Crystals? 254 B. Which Membrane Protein Structures Have Been Determined by Electron Microscopy? 254 III. Crystallization Parameters 255 A. The Protein 255 B. The Lipid 260 C. The Detergent 263 D. pH 264 E. Salt Concentration and Ionic Additives 265 F. Nonionic Additives 265 G. Precipitants 265 H. Temperature 266 IV. Methods for Grow^ing 2D Crystals of Membrane Proteins 267 A. 2D Crystallization in Suspension 267 B. 2D Crystallization on a Surface 273 C. 2D Crystallization in situ 276 V. How to Find 2D Crystals 277 VI. Outlook 279 References 280

16

In Cubo Crystallization of Membrane Proteins EHUD M. LANDAU I. Introduction 285 II. Properties of Lipidic Cubic Phases

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III. Practical Aspects of Lipidic Cubic Phases 290 A. Setting Up In Cubo Crystallization 290 B. Establishing In Cubo Crystallization Conditions 295 C. Harvesting Single Crystals from the Lipidic Cubic Phase Matrix 296 IV. Mechanism of Crystallization In Cubo 298 V. Summary 300 Acknowledgments 301 References 301

Index 303

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Contributors

Numbers in parentheses indicate the pages on which the authors' contributions begin.

Carola Hunte (55, 143, 191, 205), Dept. of Molekulare Membranbiologie, Max-Planck-Institut fiir Biophysik, 60528 Frankfurt am Main, Germany Gebhard Von Jagow (1), Zentrum der Biologischen Chemie, Universitatsklinikum Frankfurt, 60590 Frankfurt am Main, Germany Aimo Kannt (205), Aventis Pharma Deutschland GmbH, Assay Development/High Throughput Screening, Industriepark Hoechst, H811, 65926 Frankfurt am Main, Germany Reiner Kiefersauer (229), Max-Planck-Institut fiir Biochemie, Am Klopferspitz 18a, 82152 Planegg-Martinsried, Germany, and Proteros biostructures GmbH, Am Klopferspitz 18a, 82152 Planegg-Martinsried, Germany

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Contributors

Werner Ktihlbrandt (253), Max-Planck-Institut fiir Biophysik, Heinrich-Hoffmann-Strasse 7, 60528 Frankfurt am Main, Germany C. Roy D. Lancaster (219), Max-Planck-Institut fiir Biophysik, Dept. Abteilung Molekulare Membranbiologie, 60528 Frankfurt am Main, Germany Ehud M Landau (285), The Membrane Protein Laboratory, Department of Physiology and Biophysics, Sealy Center for Structural Biology, University of Texas Medical Branch, Galveston, Texas Thomas A. Link (1), Institut fiir Biophysik, J.W. Goethe-Universitat, 60590 Frankfurt am Main, Germany Hartmut Michel (143), Dept. of Molekulare Membranbiologie, Max-PlanckInstitut fiir Biophysik, 60528 Frankfurt am Main, Germany Etana Padan (55, 179), Department of Microbial and Molecular Ecology, Institute of Life Sciences, Hebrew University of Jerusalem, 91904 Jerusalem, Israel Hildur Palsdottir (191), Dept. Molekulare Membranbiologie, Max-PlanckInstitut fiir Biophysik, Heinrich-Hoffmann-Strasse 7, 60528 Frankfurt am Main, Germany Helmut Reilander (55,167), Aventis Pharma, Industriepark Hoechst, D-65926 Frankfurt am Main, Germany Christoph Reinhart (167), Max-Planck-Institut fiir Biophysik, Heinrich-Hoffmann-Strasse 7, 60528 Frankfurt am Main, Germany Hermann Schagger (1, 19, 85, 105; 161), Zentrum der Biologischen Chemie, Universitatsklinikum Frankfurt, 60590 Frankfurt am Main, Germany Tewfik Soulimane^ (229), Rheinisch-Westflische Technische Hochschule Aschen, Institut fiir Biochemie, Pauwelsstrasse 30, D-52057 Aachen, Germany Manuel E. Than (229), Max-Planck-Institut fiir Biochemie, Am Klopferspitz 18A, 82152 Planegg-Martinsried, Germany Georgios Tsiotis (131), Division of Biochemistry, Department of Chemistry, University of Crete, GR-71409 Heraklion, Greece Irene Tsirogianni (131), Division of Biochemistry, Department of Chemistry, University of Crete, GR-71409 Heraklion, Greece Miro Venturi (179), Dept. Molekulare Membranbiologie, Max-Planck-Institut fiir Biophysik, Heinrich-Hoffmann-Strasse 7, 60528 Frankfurt am Main, Germany H. Markus Weiss (167), Novartis Pharm AG, WkL 135.4.83, CH-4002 Basel, Switzerland ^Present address: Paul Scherrer Institut, Life Sciences, OSRA/007, CH-5232 Villigen PSI, Switzerland

Preface to the Second Edition

This volume is written as a practical guide for bench scientists. Protocols of general use for membrane protein purification and crystallization are provided, and special precautions for critical steps in handling membrane proteins are suggested. Part I focuses on the principles of available purification techniques, and the unique features encountered when membrane proteins are involved. In Part II, selected isolation protocols for membrane proteins describe in detail some of the most essential isolation techniques, for instance purification of histidine-tagged and affinity-epitope-tagged membrane proteins, ion exchange chromatography, and size exclusion chromatography. Part III is the first compilation in a monograph of successfully used 2Dand 3D-crystallization protocols for membrane proteins. The introduction of crystallization techniques, and the presentation of specific crystallization

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protocols is the most obvious, but not the only change compared to the first edition. In fact, this second edition comprises mainly new contributions. Only four chapters of the first edition, namely "Purification Strategies for Membrane Proteins" (Chapter 1), "Techniques and Basic Operations in Membrane Protein Purification," (Chapter 2), "SDS-Electrophoresis Techniques" (Chapter 4), and Preparative Isoelectric Focusing" (Chapter 6) have been revised and implemented in the second edition. Chapter 3, "Production and Purification of Recombinant Membrane Proteins," was introduced to attract attention to the considerable advantages of recombinant technologies. Chapter 5 summarizes recent developments in "Blue-Native Electrophoresis," one of the most convenient and effective techniques for small-scale isolation of native membrane protein complexes. Chapter 7, "Membrane Protein Crystallization," outlines the special requirements for crystallization of membrane proteins. Chapter 8, Part II, gives an "easy-to-use" protocol for reactivation of membrane proteins by addition of detergent-solubilized lipid stock solutions, which often can substitute for more time-consuming reconstitution in lipid vesicles. Chapters 9, 10, and 11 contain isolation protcols for membrane proteins that were selected to examplify essential isolation and reconstitution techniques, and methods for characterization of proteins, which have been successfully crystallized. Part III includes Chapter 12 on "Antibody Fragment Mediated Crystallization of Membrane Proteins," a technique that improves the chances to obtain wellordered membrane protein crystals. A remarkable technical aspect of "Crystallization of Fumarate Reductase," Chapter 13, is the use of preparative native isoelectric focusing, as described in Chapter 6, to obtain this membrane protein in a crystallizable form. Crystallization and crystal transformation are exemplified in Chapter 14. Chapter 15 provides an excellent overview on 2D-crystallization of membrane proteins. Last but not least, a promising approach to crystallize membrane proteins from bicontinuous lipidic cubic phase is described in Chapter 16, "In Cubo Crystallization of Membrane Proteins." We thank the authors for contributing their individual chapters in a form that emphasizes the practical aspects of their techniques, which should be of immediate value for bench scientists.

1

Purification Strategies for Membrane Proteins GEBHARD VON JAGOW*, THOMAS A. LINK,** AND HERMANN SCHAGGER* *Zentrum der Biologischen Chemie Universitatsklinikum Frankfurt 60590 Frankfurt am Main, Germany **Institut fur Biophysik J.W. Goethe-Universitat 60590 Frankfurt am Main, Germany

I. INTRODUCTION Why is a separate discussion of membrane proteins advisable? What is different when it comes to isolating membrane proteins and water-soluble proteins? Aggregation problems. Membrane proteins tend to form aggregates even in the presence of detergents, which reduces the efficacy of all separation techniques. It is therefore essential to select an effective solubilizing, but non-denaturing, detergent to ascertain (1) the minimum ionic strength the protein requires to be in a homogeneously dissolved state, (2) the maximum ionic strength that will not cause dissociation of labile proteins from complex structures, and (3) the optimum pH for solubilization and protein stability.

Membrane Protein Purification and Crystallization 2/e: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

Gebhard von Jagow et al.

Interference of detergents with separation techniques. Detergent may restrict the appUcability of certain techniques to membrane protein purification. 1. Methods that differentiate according to molecular mass or size (e.g., gel filtration and centrifugation) have lower resolution because small membrane proteins usually bind more detergent per gram of protein than do larger ones, and differences in size and mass of the protein moiety are masked. 2. Salting out techniques show little discrimination in the presence of neutral detergents. 3. Chromatofocusing and isoelectric focusing in the presence of anionic detergents do not work. Hydroxylapatite and anion-exchange chromatography should not be used in the presence of anionic detergents unless most of the anionic detergent has been replaced with a neutral detergent. 4. Hydrophobic interaction chromatography is problematic with all detergents including neutral ones. Role of detergents in protease degradation. Membrane proteins are best stabilized within membrane structures in which protease attack is slow compared to the rapid degradation often observed after solubilization by detergents. Thus, when working with membrane proteins, it is necessary to add effective protease inhibitors before solubilization. Reversible and irreversible inactivation of proteins by detergents. 1. For some membrane proteins, neither catalytic activity nor structural stability is greatly influenced by delipidating conditions. In these cases, preparation schemes will not differ greatly from those used for watersoluble proteins. 2. Other membrane proteins retain their native structure in the presence of detergents but because of delipidation may lose catalytic activity during the purification process. These delipidated preparations can be ideal for hydrodynamic measurements, protein chemical work, and crystallization attempts. Thus, failure to detect the protein by standard assay is not grounds for despair. Sufficient activity for detection may be restored by replenishing lipid during or before the enzymatic measurements or by using hydrophilic substrate analogs. 3. In some cases, the membrane protein will not only lose catalytic activity during delipidation but structural integrity as well. This is a major problem with membrane proteins but one that can often be overcome by judicious use of detergents. Chances for successful purification are good if a detergent-solubilized membrane protein is obtainable in catalytically active form in the supernatant after centrifugation. Guidelines and rules for preserving catalytic activity are discussed and

CHAPTER 1 Purification Strategies for Membrane Proteins

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summarized here. Detailed examples involving hydroxylapatite and ion-exchange chromatography are given in Chapter 2. The appUcation of detergents in membrane protein purification is a common theme throughout this book. In planning a strategy for membrane protein isolation, it is best to begin by asking fundamental questions about the protein of interest: (1) Is the protein required for functional or protein-chemical analyses? (2) What is known about its abundance and properties? Is the protein required for functional or protein-chemical analyses? This question relates to the quantity of protein to be isolated and hence, on the techniques that may be appropriate. It usually will be necessary to use methods that preserve the native state of the protein in the first steps of the purification. If enzymatic activity is not required, application of denaturing techniques may be advantageous in the final stages when proteins can be identified by apparent molecular masses on SDS-PAGE. Native gel electrophoresis techniques are presently restricted to small-scale preparations and thus will preferentially be used as the last purification step (in fortunate situations as the only purification step). All other methods may be used for small- to large-scale purifications with some differences in cost. Enriching the protein of interest before adding detergents will facilitate purification. If there is a choice, attempts should be made to isolate the protein from sources that contain high levels of it. In all cases, it is recommended that the level of protein be enhanced before adding detergent (such as by using cell culture conditions that favor the expression of the protein of interest or by preparing organelles a n d / o r enriched membranes). Often it is advisable to accept relatively large losses to obtain a suitable starting material. This may ultimately save a purification step in the presence of detergent and may be essential for preserving catalytic activity. What is known about the properties of the protein of interest? The most essential questions are whether a simple enzyme test system is available to follow enrichment during the preparation protocol and whether catalytic activity is retained after solubilization by detergents. It is usually advantageous to perform a series of small-scale preextraction and solubilization experiments to establish conditions under which the protein is in a homogeneously dissolved state, in which catalytic activity is present, and for which optimal enrichment is obtained. This involves testing salt concentrations, pH, protease inhibitors, and protein stabilizing additives.

11. GENERAL GUIDE FOR RETAINING CATALYTIC ACTIVITY A. Dissolve the membrane protein under optimal conditions for stability at a detergent/protein ratio that is not much higher than the minimum

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detergent:protein ratio required for protein solubilization. {Note: The minimum detergent/protein ratio is also the minimum detergent/ lipid ratio. The latter is the essential ratio for solubilization, but the detergent/protein ratio is usually measured for convenience). B. Keep the time for isolation as short as possible. Perform separations (preferentially) in the cold, and replenish protein stabilizing additives or protease inhibitors whenever these are removed or inactivated (e.g., EDTA protecting against Ca-dependent proteases is removed by hydroxylapatite chromatography, and the protease inhibitor PMSF becomes inactive after some hours in water). C. If possible avoid any purification step that would add new detergent and alter the original detergent/lipid ratio. If this condition cannot be strictly met, keep changes in the detergent/lipid ratio to a minimum. {Note: Ideal conditions in this respect are those in which the protein of interest passes through a column together with detergent and lipid at the original detergent/lipid ratio, while contaminants bind to the support. This "pass-through" mode is especially useful in preliminary column chromatography experiments to optimize binding conditions without losing catalytic activity.) In most cases, the pass-through mode will not yield effective enrichment and the "binding-mode" must be used. The detergent/lipid ratio then cannot be kept constant. Complete delipidation can be avoided, however, by using small volumes of washing buffer and maintaining a low level of detergent concentration in the washing buffer. The detergent concentration in the elution buffer is not critical if the protein elutes as a sharp band with the first few milliliters of elution buffer. Gradient elution will be deleterious for proteins sensitive to delipidation, except with steep gradients. The use of a batch procedure is recommended for proteins highly sensitive to delipidation (see Chapter 2, Section IV,A). Extensive delipidation can also be prevented by adding lipid to the buffers. Optimal stabilization can often be achieved at a ratio of 2 to 3 molecules of detergent per molecule of lipid (see Chapter 8). However, because of solubility problems, higher detergent/lipid ratios are generally required.

III. CHOICE AND SEQUENCE OF PURIFICATION TECHNIQUES A typical membrane protein isolation protocol involves several separate steps that are usually applied in the following order. A. Enrichment without use of detergent. Enriched protein content can be obtained, for example, by choosing an optimal source of the protein or by isolating organelles and membranes during the first stages of the purification. B. Solubilization and precipitation techniques. These approaches pro-

CHAPTER 1 Purification Strategies for Membrane Proteins

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vide some degree of purification. Differential extraction and centrifugation are commonly used methods that are occasionally followed by differential precipitation techniques. These are applicable on a large scale and afford low to medium resolution at low cost. C. Hydroxylapatite or ion-exchange chromatography (large or small scale, medium resolution, low to medium cost). D. Chromatofocusing (small to medium scale, high resolution, medium to high cost); or dye affinity chromatography (small to medium scale, high resolution, medium to high cost); or ligand affinity chromatography (small to medium scale, very high resolution; high cost); or immobilized metal affinity chromatography (small to medium scale, very high resolution when His-tagged proteins are applied, medium cost); or covalent chromatography (small to medium scale, high resolution but restricted applicability, medium cost); or hydrophobic interaction chromatography (small to medium scale, low to medium resolution and restricted applicability, medium cost). E. Blue native electrophoresis (small scale, high resolution, yields highly pure proteins, such as those used for generating antibodies and for protein sequencing, less suitable for enzymatic studies); or native isoelectric focusing (small scale, high resolution, yields highly pure proteins, such as for crystallization); or gel filtration (small to medium scale, low resolution, suitable for enzymatic studies); or density gradient centrifugation (small to medium scale, low resolution; suitable for enzymatic studies). Notes: An exchange of buffer by gel filtration may be necessary for transition from step to step—for example, before anion-exchange chromatography or isoelectric focusing. Hydroxylapatite and anion-exchange chromatography have similar separation properties, so consecutive application of these techniques theoretically may not markedly improve protein enrichment. Nevertheless, combining these large-scale, low-cost methods often proves useful. Direct application of ligand affinity chromatography or another highresolution technique may allow a one-step purification after solubilization. A less costly approach, however, is usually used first to remove contaminants such as lipids and DNA, which helps to avoid damaging the expensive columns. Blue native PAGE, an economical and easy-to-use high-resolution technique, can be employed both for final purification and for single-step protein purification directly after solubilization (see Chapter 5). In the end, high-cost

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methods may be less expensive than low- or medium-cost procedures if the number of purification steps is reduced and losses in yield and catalytic activity are avoided.

IV. CHOICE OF PROTEIN SOURCE, DISRUPTION OF CELLS, AND PREPARATION OF ORGANELLES AND MEMBRANES Membrane proteins can be prepared from a variety of sources, such as animal cells, plant cells, fungi (yeast), and bacteria. (Methods for optimization of starting materials for preparation of organelles from eukaryotes and for preparation of membrane extracts are described extensively in Guide to Protein Purification, a volume of Methods in Enzymology; see References.) For production and purification of recombinant membrane proteins, see Chapter 3. For affinity-epitope tagging of G-protein coupled receptors, see Chapter 9. Bacterial membrane protein complexes (e.g., the respiratory chain complexes) are simpler in polypeptide composition than the eukaryotic counterparts while performing essentially the same function. Many proteins have been overexpressed in bacteria by the introduction of suitable plasmids containing genes encoding the target proteins. Animal cells are easily obtainable in large quantities from slaughterhouses. Animal cells can be broken easily with a mixer or blender. Unicellular organisms like yeast or bacteria have much more robust cell walls, so cell disruption is more difficult. The following techniques can be used to break down cell walls. Glass bead milling involves agitating the cells together with fine glass beads (e.g.. Bead Beater from Biospec, Bartlesville, OK, or Dyno-Mill from Bachofen Basel, Switzerland), which destroys the cell walls via abrasion. Homogenization requires a cell suspension to be forced through a narrow valve under high pressure. The rapid pressure drop effectively breaks the cells. On a small scale, one can use a French press cell (SLM Aminco makes one), but large-scale preparation requires the use of a homogenizer (e.g., an APV Gaulin mill). Grinding mills, as described by Weiss et al. (1979), very effectively break filamentous fungi (such as Neurospora crassa). Ultrasonication will break the cells of some microorganisms effectively, although care must be taken not to damage the proteins. Osmotic shock can be used to break cells with less robust walls (e.g., animal and plant cells and some gram-negative bacteria).

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Repeated freezing and thawing will break the membrane in some cells. However, some proteins are cold-sensitive, and membrane protein complexes may dissociate. Enzymatic lysis. Lysozyme, which catalyzes hydrolysis of j5-l,4-glycosidic bonds, is often used to break open gram-positive bacteria. Zymolyase another polysaccharide hydrolase (produced by Kirin Brewery, Japan, and distributed in the United States by ICN) catalyzes partial breakdown of yeast cell walls. This mild procedure allows isolation of intact organelles. Note: Homogenizers and ultrasonication devices can cause significant heating of the sample, as can heavy agitation (glass bead mills). Sufficient cooling must be provided in these cases. Once the cells are broken, cellular fractions and organelles are generally isolated by differential centrifugation. In many cases, additional purification by sucrose density gradients or silica-sol gradients (e.g., FicoU and PercoU) is necessary to remove contaminating material.

V. PROTEIN ASSAY To monitor the purification at different steps, it is essential to measure a specific protein property (often the catalytic activity of an enzyme is involved) and the total protein concentration at each step. Ideally, if the protein is stable, this will immediately reveal whether a specific procedure has improved the purity of the protein.

A. Catalytic Activity and Other Specific Properties Protein properties suitable for following the purification are (1) catalytic activity, (2) spectroscopically detectable prosthetic groups, (3) specific binding of labeled ligands, and (4) binding of antibodies, directed against the entire protein complex or against individual subunits, after SDS-PAGE (Western blot). Any protein-specific assay may suffer from the problem of protein instability even if one follows precautions for handling membrane proteins. Enzymatic activity may decrease during isolation, prosthetic groups and labeled ligands may dissociate, antibodies may detect a dissociated subunit rather than the holoprotein complex, and characteristic polypeptide patterns observed in gel electrophoresis may disappear. Enzyme assay. Enzymatic assays are highly sensitive and rapid. Typically, subpicomole amounts of enzyme provide sufficient signals in optical assays. Sometimes enzymatic activity is lost during isolation, implying that the enzyme has been denatured. However, the inactivation may be

Gebhard von Jagow et al.

due to a loss of essential coenzymes or associated lipids, so it is reversible if the components are replenished during the assay. An easy way to supply lipid is to add lipid-saturated detergent micelles, as described in Chapter 8. In some cases (e.g., if the assay requires measurement of substrate transport across membranes by carrier proteins), more time-consuming reconstitution into proteoliposomes will be required (Kagawa and Racker, 1971). If catalytic activity using hydrophobic physiological substrates is not restored by replacement of components, it may be advisable to use more hydrophilic artificial substrates. Spectrophotometry. Spectroscopic assays are convenient for proteins that contain prosthetic groups. Although less sensitive than enzymatic assays, the spectroscopic approach offers the advantages of ease and speed. Colored proteins, such as hemoproteins, can be readily identified and monitored during chromatography. Unfavorable conditions often are detected in early stages by changes in extinction coefficients and shifts in absorption maxima. Differentiation of hemoproteins by optical difference spectroscopy requires only nanomole amounts of protein. Other groups, such as iron-sulfur clusters or tyrosine radicals, can be detected by electron paramagnetic resonance (EPR) spectroscopy. Flavins become fluorescent after denaturation at acidic pH. Ligand binding. Binding of specific fluorescent or radioactive ligands or inhibitors is especially useful for observing transport proteins that do not contain prosthetic groups. Antibody analysis. Binding of antibodies to the desired protein permits sensitive and highly specific detection. Often the antibodies have to be raised against one known protein component of a less well-characterized complex or against a short protein sequence stretch that was synthesized according to the DNA sequence (peptide antibody).

B. Determination of Total Protein Ultraviolet spectrophotometry at 280 nm. This method is fast and nondestructive, but it is not very specific. The absorbance of a protein solution varies with the tryptophan content of the protein, but for a 1 m g / m l solution it is usually about 1.0. UV spectroscopy is also sensitive to interference from other absorbing components (buffer, nucleotides, etc.). Biuret methods. The Biuret assay, developed by Itzhaki and Gill (1964), involves formation, under alkaline conditions, of a complex with copper(II) bound to the peptide nitrogens. The main disadavantage is the rather low sensitivity ( ^ 1 mg/ml).

CHAPTER 1 Purification Strategies for Membrane Proteins

9

Lowry method. The modification of the original method (Lowry et ah, 1951) by Helenius and Simons (1972) involves the addition of SDS to prevent precipitation of Triton X-100, a detergent often used in membrane protein isolation. In this approach, the copper complex formed under alkaline conditions is used catalytically to reduce phosphomolybdotungstate to heteropolymolybdenum blue (Folin-Ciocalteau reagent). This reaction makes the technique approximately 10-fold more sensitive than the Biuret method. However, it is more sensitive to the amino acid composition of the protein and the lipid content of the sample compared to the Biuret method, and 50% deviations from the real protein concentration are often observed. BCA method. Smith ei ah (1985) improved the Lowry method by substituting bicinchoninic acid (BCA) for the Folin-Ciocalteau reagent. The assay is less sensitive to buffer components and detergents. A microassay has been developed with 10- to 100-fold better resolution than the Lowry test. However, the reaction is very temperature sensitive and must be performed in a water bath under thermostatted conditions. Bradford method. Bradford (1976) used the binding of dyes, such as Coomassie Brilliant Blue G-250, to proteins to estimate protein concentration. This approach is about as sensitive as the Lowry method and as easy to perform. However, it is sensitive to the amino acid composition of the protein. More importantly for membrane proteins, the assay may be biased by the presence of detergent. This can be a major problem when the detergent concentration cannot be determined and corrected (e.g., during gel filtration, where detergent peaks elute according to micellar size).

VI. SOLUBILIZATION AND STABILIZATION OF MEMBRANE PROTEINS A. Choice of Buffer The buffer pH has an important role during protein isolation. The appropriate pH is essential for protein stability and solubility, and it also affects binding and elution properties of many protein separation media. Most useful are the zwitterionic biological buffers developed by Good and Izawa (1972). However, other buffers like phosphate and Tris are also frequently used. Special requirements with individual separation techniques are discussed in Chapters 8 to 11. Detailed information on the properties of buffers for pH values in the 1.5-13 range, along with their effects on protein assays, can be found in Deutscher, 1990. These are the important criteria for selecting a buffer.

10

Gebhard von Jagow et al.

1. The pK of the buffer should be within ±0.5 unit of the desired pH. 2. Potential interactions (beneficial or harmful) with a column matrix must be considered. 3. Whether the buffer is involved with activation or inactivation of enzymes is crucial (metal-dependent enzymes may be deactivated by metal complexing buffers). 4. UV-absorbing buffers should be avoided if a UV detector is to be used. 5. Cost should be considered if scaling up the preparation is desired. The ionic strength is another parameter affecting the solubility of membrane proteins in the presence of detergents. It is a rather freely adjustable parameter, in contrast to pH, which must be chosen according to the stability properties of the protein. The effect of ionic strength on protein solubility is discussed in more detail in Chapter 2.

B. Choice of Detergent Membrane proteins require detergents for extraction from membranes and for solubilization. Detergents are amphiphiUc molecules that form micelles in water. They solubUize proteins by binding to the hydrophobic parts of the proteins on one side and interacting with the aqueous phase on the other side. Properties and uses of detergents in biology and biochemistry have been reviewed (Deutscher, 1990; Neugebauer, 1988). For a detailed discussion of detergents, especially of those used in membrane protein crystallization, see Chapter 7. Detergents may be classified with respect to their hydrophobic and hydrophilic portions. 1. Classification According to Hydrophilic Portion The hydrophilic structures in detergents can be nonionic, anionic, cationic, or zwitterionic (see Table I). Nonionic detergents include the detergents used most often in membrane protein purification, the polyoxyethylene and sugar detergents. Polyoxyethylene detergents contain a polymeric (O—CH2—CH2)„-OH chain of variable length that is neutral at usual pH values and is compatible with all separation techniques. The species with HLB (hydrophilic-lipophilic balance) values ranging from 12 to 14.5 (Table I) are useful for extraction of integral membrane proteins. Those with higher HLB values are "milder" but usually will extract only peripheral membrane proteins. In later stages of purification they may be able to keep integral proteins in the solubilized state. These are fortuitous cases because these detergents barely affect protein-bound lipid, and catalytic activity is well retained.

11

CHAPTER 1 Purification 1Strategies for Membrane Proteins

TABLE 1

Common Detergents''

Detergent

Type

M,

CMC (mM)

N N N N N N N N

540 553 583 1200 625 537 1228 1310

N N N N N N N N N N N

n

HLB

H + >Li+

The affinity of proteins for the column phase can also be regulated by

CHAPTER 2 Techniques and Basic Operations in Membrane Protein Purification

37

adjusting the pH. The minimum buffering strength should be 10 mM at a pH differing not more than 0.3 pH units from the pK value. Low buffering strength cannot prevent p H changes that result from ionic strength changes in buffers and may lead to protein denaturation. The pH always should be adjusted at the temperature chosen for the chromatography (pH values measured at 4 and 25°C differ by 0.1-1 pH units for some buffers). Buffers used for anion-exchange chromatography are primarily cationic buffers—for example, Bistris (pK 6.5), imidazole (pK 7.0), triethanolamine {pK 7.8), Tris (pK 8.1), diethanolamine (pK 8.8), or ethanolamine {pK 9.5). Chloride is usually used as counterion. Buffers carrying carboxyl groups are also useful, since they have only small interactions with the solid support material. Buffers used for cation-exchange chromatography commonly are anionic buffers, such as acetate {pK 4.8), citrate (pK^ 3.1; pi Ni^"^ > Zn^"*^ > Co^"^ > Fe^"^ > Ca^"^. Ni^^ is commonly used for purification of Histidine-tagged proteins, while Fe^ "^ is preferentially used for purification of phosphoproteins because of its high affinity for phosphate groups. Ca^"^ will work only with proteins with high affinity to Ca^"^ itself. Unfortunately, it is difficult to predict which metal ion will prove most appropriate. Some proteins will bind only to Cu^"^. Others bind so strongly that difficulties with elution may occur, and thus use of the weaker binding Ni^"^ or Zn^"^ is advisable. Dilute protein solutions can be applied without loss of resolution. The affinity of individual proteins for the solid support depends on the number and accessibility of specific amino acid residues on the protein surface. The affinity of amino acids for the metal ions decreases in the order His > Trp > Cys > Tyr. 2. Differential Binding and Elution a. Column Preparation and Protein Binding 1. Cast a column, usually 10-30 cm in length, wash with 2 column volumes of distilled water, and then saturate the column with divalent metal ions (e.g., 0.1 M NiS04 dissoved in water; ^ 3 column volumes). 2. Wash the gel with 3-5 column volumes of water. 3. Equilibrate the gel with at least 2 column volumes of a start buffer, which ideally allows binding of the protein of interest, with weaker binding proteins being found in the flow-through. This is especially helpful for membrane proteins, since only small volumes of a washing buffer containing detergent have to be applied next, thus avoiding delipidation and denaturation. Commonly used start buffers are adjusted to the pH range 5.5-8.5—for example, 10-100 mM Sodium phosphate, 50-100 mM Sodium acetate.

CHAPTER 2 Techniques and Basic Operations in Membrane Protein Purification

45

or 25 mM Sodium borate. It is common to include salt 100-500 mM to eliminate any ion exchange effect and to improve solubility of the membrane protein. Add a neutral detergent when membrane proteins are involved—for example, 0.1% dedecylmaltoside or 0.05-0.1% Triton X-100 — and add the eluent ion (see below) at the highest concentration that still allows binding of the desired protein. This leads to flowthrough of all weaker binding proteins, and the protein load per milliliter of column material can be raised considerably. Protein load. By way of example, Kashani-Poor et al., 2001, could load 35 mg of a total mitochondrial extract per 1 ml of Chelating Sepharose Fast Flow medium after addition of 60 mM of the eluent imidazole to the start/equilibration buffer and to the protein sample. Almost all contaminating protein was in the flow-through. Essentially the total protein binding capacity of the column (about 4 mg/ml) was free for the desired protein (complex I), which was then eluted by raising the imidazole concentration in the elution buffer to 140 mM. Chelating agents such as EDTA or citrate should not be included in any buffer. 4. Washing buffer usually is identical to start/equilibration buffer. Elution is achieved by using competitive elution (see below), by reducing pH (most proteins elute between pH 6 and 4; sodium acetate, citrate, or phosphate can be used), or by adding chelating agents such as EDTA (50 mM), which will strip the metal ions from the gel together with the protein. This method does not resolve different proteins. 5. Elute the desired protein by using a modified washing buffer with increased concentration of the competitive eluent—for example, 140 mM imidazole if the imidazole concentration in the washing buffer was 60 mM. Commonly applied concentrations of competitive eluents are up to 500 mM imidazole, up to 50 mM histidine, or up to 2 M ammonium chloride.

E. Affinity and Dye-Ligand Chromatography Affinity chromatography, including chromatographic procedures that use highly specific ligands covalently linked to a solid matrix, is one of the most powerful techniques for isolation of proteins (Hermanson et ah, 1992; Jacoby, 1984; Deutscher, 1990). Despite this fact, single-step purifications using affinity chromatography are not common. Usually a low-cost technique is used first to avoid potential damage to the expensive affinity column. 1. Chromatography Media A wide spectrum of affinity columns having different immobilized groupspecific ligands may be purchased from various suppliers. Alternatively,

46

Hermann Schagger

specific column materials can be constructed, starting with commerially available nonactivated matrices containing spacer arms or with preactivated matrices ready for binding individual Ugands. Commercially available ligands covalently bound to matrices include (1) mono- and group-specific low molecular mass ligands—for example, steroids for purification of steroid receptors, biotin and avidin for biotin-binding and biotin-containing proteins, enzyme cofactors and analogs, vitamins, certain amino acids, "biomimetic dyes," and boronic acid derivatives; and (2) mono- and group-specific macromolecular ligands—for example, protein A and G for purification of immunoglobulins, monoclonal antibodies for purification of antigens, antigens for isolation of specific antibodies, lectins for purification of glycoproteins, and many more. Immobilized biomimetic dyes for dye-ligand chromatography are widely used not only for isolation of nucleotide-containing proteins but also for many other proteins (see Table III). However, the binding properties of a particular protein cannot be predicted. Kits for identification of suitable adsorbents are available from Sigma and Amicon. The more specific the ligand is to a particular procedure, the more likely it will not be commercially available and thus will need to be coupled to a preactivated support by the investigator (see Table IV). Small ligands usually require matrices with extended spacer arms to allow access of the protein. The protein-ligand interaction must be reversible under nondenaturing conditions; dissociation constants normally range from 10"''^ to 10"^ M. Detailed discussion of the properties of gel matrices, choice of spacer between gel matrix, and activation and coupling chemistry is beyond the scope of this book. For descriptions of these techniques, see, for example, Hermanson et al. (1992), as well as product information sheets of the suppliers.

TABLE III

Selected Immobilized Biomimetic Dyes and Boronic Acid Derivatives'"

Protein isolated

Product

Manufacturer/ supplier

Dehydrogenases (especially NAD^), kinases and phosphatases, other proteins, removal of albumin IgG from serum

Blue Sepharose C1-6B Affi-Gel Blue Gel Fractogel TSK AF-Blue DEAE Affi-Gel Blue Gel CM Affi-Gel Blue Gel Red Sepharose C1-6B Fractogel TSK AF-Red Affi-Gel 601 (phenyl boronate) Fractogel TSK AF-Phenylboronate

Pharmacia Bio-Rad Merck Bio-Rad Bio-Rad Pharmacia Merck Bio-Rad Merck

Dehydrogenases (NAD"^ and NADP"^), many other proteins Sugars, glycoproteins, nucleotides

"Blue and red gels usually contain Cibacron Blue F3G-A and Procion Red HE-3B dyes. These and many more immobilized dyes are available from many other suppliers. Reactive dye-ligand tests are available (e.g., from Sigma).

CHAPTER 2

TABLE IV

Techniques and Basic Operations in Membrane Protein Purification

47

Selection of Commercially Available Activated Matrices

Ligand(s) to be coupled -SH, —NH2, —OH Also from proteins, peptides, carbohydrates Primary amines and thiols, also from proteins -SH Low M^, thiols and proteins Thiol proteins -NH2 Proteins Proteins / peptides Primary amines (especially proteins) Primary amines (especially low M^)

Primary amines, amino acids, peptides

—CHO Aldehydes (also high M^) —COOH Amino acids, ketoacids, carboxylic acids

Product

Manufacturer/ supplier

Epoxy-activated Sepharose 6B Epoxy-activated Eupergit C Epoxy-activated Separon HEMA

Pharmacia Rohm Pharma Anachem/Tessek

Tresyl-activated Sepharose 4B Fractogel TSK AF-Epoxy 650 FMP-activated Avid gel

Pharmacia Merck BioProbe International

Thiopropyl Sepharose 6B Activated Thiol-Sepharose 45" Affi-Gel 501 (organomercurial support)

Pharmacia Pharmacia Bio-Rad

CNBr-activated Sepharose 4B Activated CH-Sepharose 4B Act-Ultrogel AcA 22/glutaraldehyde Affi-Gel and 15/N-hydroxysuccinimide Reacti-Gel (GX) / carbonyldiimidazole Reacti-Gel (HW-65F) / carbondiimidazole Reacti-Gel (25DF) / carbonyldiimidazole Reacti-Gel (GF-2000)/carbonyldiimidazole Fractogel TSK AF-CDI 650 CM BioGel A Fractogel TSK CM-650 CH-Sepharose 4B EHC-Sepharose 4B

Pharmacia Pharmacia IBF Bio-Rad Pierce Pierce Pierce Pierce Merck Bio-Rad Merck Pharmacia Pharmacia

Agarose-adipic acid hydrazide

Pharmacia

AH-Sepharose 4B EAH-Sepharose 4B Affi-Gel 102 Aminoethyl BioGel P-100 Fractogel TSK AF-Amino 650

Pharmacia Pharmacia Bio-Rad Bio-Rad Merck

'^ See also Section IV,F on covalent chromatography.

The choice of a preactivated matrix (Table IV) that allows coupling of the desired ligand at mild conditions is essential for pH-sensitive small ligands and for protein ligands that are more prone to denaturation. 2. Protein Adsorption and Elution The choice of adsorption and eluting conditions depends of the nature and strength of the ligand-protein interaction. Adsorption and desorption are

48

Hermann Schagger

influenced by ionic strength, pH, temperature, reducing or oxidizing conditions, and the presence or absence of competing ligands. Any of these factors may be altered to promote protein elution. For example, selective elution by competing ligands (nucleotides) has been used for purification of dehydrogenases bound to biomimetic dye-containing columns, and glycoproteins have been removed from lectin columns by sugars. For a recent purification of a membrane protein complex (chloroplast ATP synthase) by dye-ligand chromatography, see Seelert et al, 2000.

F. Covalent Chromatography In principle, any of several functional groups can be used for covalent attachment of proteins to solid supports. However, the harsh conditions necessary to then remove the protein from the column would likely destroy the protein. The only functional group on the protein that is useful for covalent attachment is the sulfhydryl group of cysteine, which reacts with disulfide groups of the support. The disulfide bond thus formed between protein and support is easily split by reducing agents. Before using this approach in a purification protocol, the stability should be tested in the presence of the desired reducing agent. Note: Covalent chromatography will be possible only if at least one—SH group is present on the protein surface and accessible to the support. 1. Thiol Gel Media Thiol Gel Media are available from several companies either in the activated disulfide form [e.g.. Activated Thiopropyl Sepharose 6B and Activated Thiol Sepharose 4B (Pharmacia)], or in the thiol form [e.g., cysteamine agarose, cysteine agarose, and glutathione agarose (Sigma) and N-acetylhomocysteine agarose (Bio-Rad)]. Supports are activated, or reactivated after use, by disulfides, such as 2,2'-dipyridyl disulfide. 2. Description of the Chromatographic Procedure a. Equilibrate the gel in start buffer without detergent (use neutral or slightly acidic pH; choose an ionic strength based on the conditions necessary for protein solubilization and include 1 mM EDTA to trap transition metal ions). Most proteins carrying thiol groups on the surface will bind at pH 8. Selective binding of proteins that have thiol groups with low pX is possible at lower pH (pH range 8-4). b. Pack the column and wash with start buffer containing detergent. c. Prepare homogeneously solubiUzed membrane proteins by methods described above (supernatant obtained after ultracentrifugation). Remove thiols such as glutathione by gel filtration, and apply the sample to the column.

CHAPTER 2 Techniques and Basic Operations in Membrane Protein Purification

49

d. Wash with start buffer in the presence of 0.05-0.5% detergent. The danger of membrane protein denaturation will increase with the volume of buffer used and with the concentration of detergent. In some cases, a batch procedure is advisable (see Section A,3)e. Elute with start buffer containing detergent and 5-25 mM L-cysteine, or 50 mM glutathione, or 20-50 mM dithiothreitol.

G. Hydrophobic Interaction Chromatography The difference between hydrophobic interaction chromatography (HIC) and reversed phase chromatography (RFC) is related to the density of hydrophobic ligands on the surface of the adsorbents (Builder, 1993). The surface of RFC adsorbents can be regarded as a continuous hydrophobic phase; in HIC the hydrophobic Ugand density is much lower. The interaction of hydrophobic domains of proteins with the adsorbent therefore is much higher in RFC, and more drastic conditions for protein elution are required. This will often result in protein denaturation. RFC is especially useful for separation of membrane protein fragments. The various media used in HIC differ in ligand type and degree of substitution of the surface. HIC media are available from many suppliers; the particular material that will allow binding and elution of a given protein under mild conditions must be identified empirically. Froteins are adsorbed to hydrophobic matrices at salt concentrations that are high but below those required for protein precipitation. HIC therefore would be ideal following a salt precipitation step. Desorption usually is achieved by reducing the salt concentration. The strength of hydrophobic interaction also decreases with decreasing temperature and in general with increasing pH. Examples for sucessful application of HIC to membrane proteins are rare because detergents bind very strongly to the matrix. One example is the isolation of cytochrome F-450, described by Kastner and Neubert (1990). The essential steps in the procedure are as follows: (1) removal of most of the detergent by hydroxylapatite chromatography before the HIC step; (2) binding of proteins to Fhenyl-Sepharose or Octyl-Sepharose (Fharmacia) at high phosphate concentration; and (3) elution of proteins at reduced salt concentration and increased detergent concentration.

H. Gel Filtration Gel filtration media are porous beads into which sufficiently small molecules can penetrate. Larger molecules are excluded from the beads, and during gel filtration they pass through the column in the liquid phase (void volume) outside the beads and are therefore eluted earliest. From a practical point of view, the void volume is that volume of buffer that passes through

50

Hermann Schagger

the column first and cannot contain any protein regardless of protein size. Void volumes are typically about 20% of the total column volume. By way of example, consider a mixture of proteins of widely different sizes applied to a 100 ml column. The first 20 ml (void volume) will contain no protein. The next fractions will contain the largest proteins and also protein aggregates, except huge aggregates that cannot even pass between the gel particles in the liquid phase and that remain at the top of the column. Thereafter, proteins that can penetrate the gel beads (i.e., are in the separation range of the particular gel filtration medium being used) emerge from the column. The extent of penetration and retardation on the column depends on the size and shape of the protein. Except perhaps for proteins that have quite asymmetrical shapes, molecules are eluted in the order of decreasing size. 1. Separation Range in the Presence of Detergents The separation range of gel filtration media usually is specified for water-soluble proteins. The behavior of membrane proteins solubilized by detergents on gel filtration columns usually is completely different, owing to the effect of detergent binding to the proteins. Separation ranges of selected gel filtration media are compared in Table V for situations involving no detergent, Triton X-100, and sodium dodecyl sulfate (SDS). 2. Gel Filtration for Separation of SDS-Denatured Proteins a. Concentrate the protein—for example, in the SpeedVac centrifuge or by lyophilization (after SDS-precipitating ions like potassium ions and salts in general have been removed by dialysis against water) — or precipitate the protein directly by adding an equal volume of acetone (first test a small aliquot to see if precipitation occurs), centrifuge, wash the pellet with a small volume of water to remove acetone, and centrifuge again. Note: If acetone precipitation works (the higher the protein concentration, the better) this method is preferred, since not only salt is removed with the supernatant but also detergent. b. Dissolve sedimented proteins to a maximal concentration of 10 m g / m l by incubating for 15-30 min at ^40°C in 5% SDS, 1% mercaptoethanol. Note: Do not heat the sample because many membrane proteins irreversibly aggregate at elevated temperature. c. Perform the gel filtration in the presence of 0.5% SDS at room temperature. The column should be at least 1 m long. The optimal sample volume to be applied is 1-2% of the column volume. Typical gel filtration columns for 5-20 kDa and 10-50 kDa proteins are Sephadex G-50 sf and Sephadex G-lOO, respectively. Sephadex G-50 sf: Proteins with molecular masses at or above 30 kDa will elute at the void volume together with aggregated proteins.

CHAPTER 2

TABLE V

51

Techniques and Basic Operations in Membrane Protein Purification

Separation Ranges of Selected Gel Filtration Media GeV

No detergent total range^ (kDa)

Sephadex G-50 sf Sephadex G-lOO Sephacryl S-200 Sephadex G-150 Sephadex G-200 Ultrogel AcA 34 Sephacryl S-300 Ultrogel AcA 22 Sepharose C1-6B

1.5-30 4-150 5-250 5-300 5-600 20-350 10-1500 100-1200 10-4000

Triton X-100 special range' (kDa)

SDS special range (kDa) 5-20 10-50

10-50^*

50-200"

200-1000

''Sephadex, Sepharose, and Sephacryl media are available from Pharmacia, Ultrogel AcA from IBF and Serva. The useful range of media from other manufacturers and suppliers (e.g., Merck and Bio-Rad) for separation of membrane proteins can be easily estimated by comparing the fractionation range for water-soluble proteins with those given above. '' The total range for separation of water-soluble proteins also seems valid for protein separation in the presence of SDS. ^ The special range refers to the molecular masses of the protein moiety, since the precise degree of Triton binding is rarely known. ''Considering bound Triton, the lower total mass in fact is 100 instead of 10 kDa, and the upper total mass is 150 instead of 50 kDa. " The narrow pore size distribution of Ultrogel AcA results in high resolution over the indicated special range.

Smaller proteins will be separated from one another if the differences in molecular mass are about 10 kDa. Sephadex G-lOO: Proteins larger than 70 kDa and aggregates will be in the void volume. Proteins smaller than 10 kDa will appear as extremely broad peaks. 3. Gel Filtration of Native Membrane Proteins The separation of native membrane proteins is impeded by protein-bound detergent that masks differences in molecular mass. The resolving power is low, even with the optimal gel filtration medium. It is mainly used as a final purification step to remove contaminating small proteins and excess detergent. Note: Gel filtration usually is performed at 4°C, using columns with a length of at least 1 m. The detergent concentration should be kept well above the critical micelle concentration [e.g., use 0.05% Triton (CMC 0.015% at salt concentrations typically used) or 0.1% dodecylmaltoside (CMC 0.01%)] to avoid oligomerization of membrane protein. The ionic strength of the buffer must be above the solubility limit of the protein of interest (usually 20-200

52

Hermann Schagger

mM NaCl). The volume of sample should not be larger than 1-3% of the column volume. Keep the detergent concentration of the sample below 5% and the protein concentration below 10 mg/ml. Regarding resolution limits, small membrane proteins (e.g., 10 kDa) usually elute with the detergent micelles of dodecylmaltoside or Triton X-100. The separation of a 40 kDa and a 10 kDa membrane protein in the presence of Triton is in fact a separation of 130 kDa and 100 kDa protein-detergent complexes (40 kDa plus 90 kDa of bound detegent versus 10 kDa plus 90 kDa). The best gel filtration medium for such separation problems is, theoretically, Sephadex G-200. However, this material is often difficult to work with. With some loss of resolution, other gel filtration columns can be used, such as Ultrogel AcA 34, Sephadex G-150, or Sephacryl S-300, each of which has its own separation characteristics (see Table V).

REFERENCES Bernardi, G. (1971). Chromatography of proteins on hydroxylapatite. In "Methods in Enzymolo g / ' (W. B. Jacoby, ed.). Vol. 22, pp. 325-339. Academic Press, New York. Bordier, C. (1981). Phase separation of integral membrane proteins in Triton X-114. /. Biol. Chem. 256,1604-1607. Brandt, U., Schagger, H., and von Jagow, G. (1989). Purification of cytochrome-c oxidase retaining its pulsed form. Eur. J. Biochem. 182, 705-711. Builder, S. E. (1993). "Hydrophobic Interaction Chromatography—Principles and Methods," Code No. 18-1030-90. Pharmacia. Deutscher, M. P. (ed.) (1990). "Methods in Enzymology," Vol. 182 (Guide to Protein Purification). Academic Press, New York. Fricke, B. (1993). Phase separation of nonionic detergents by salt addition and its application to membrane proteins. Anal. Biochem. 212,154-159. Geier, B. M., Schagger, H., Brandt, U., Colson, A.-M., and von Jagow, G. (1992). Point mutation in cytochrome b of yeast ubihydroquinone:cytochrome-c oxidoreductase causing m)ocothiazol resistance and facilitated dissociation of the iron-sulfur subunit. Eur. J. Biochem. 208,375-380. Giri, L. (1990). Chromatofocusing. In "Methods in Enzymology," Vol. 182 (M. P. Deutscher, ed.. Guide to Protein Purification), pp. 380-392. Academic Press, New York. Hatefi, Y. (1978). Introduction—Preparation and properties of the enzymes and enzyme complexes of the mitochondrial oxidative phosphorylation system. In "Methods in Enzymology," (S. Fleischer and L. Packer, eds.). Vol. 53, pp. 3-4. Academic Press, New York. Hermanson, G. T., Mallia, A. K., and Smith, P. K. (1992). "Immobilized Affinity Ligand Techniques." Academic Press, New York. Jacoby, W. B. (ed.) (1984). "Methods in Enzymology," Vol. 104 (Enzyme Purification and Related Techniques, Part C). Academic Press, New York. Kashani-Poor, N., Kerscher, S., Zickermann, V., and Brandt, U. (2001). Efficient large scale purification of His-tagged proton-translocating NADH:ubiquinol oxidoreductase (complex I) from the strictly aerobic yeast Yarrowia lipolytica. Biochem. Biophys. Acta. 1504, 363-370. Kastner, M., and Neubert, D. (1990). Isolierung von Cytochrom P-450 aus Marmorset-Leber. In "Membranproteine," pp. 109-122. Pharmacia. Okun, J. G., Zickermann, V., Zwicker, K., Schagger, H., and Brandt, U. (2000). Binding of detergents and inhibitors to bovine complex I—A novel purification procedure of bovine complex I retaining full inhibitor sensitivity. Biochem. Biophys. Acta., in press.

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Parish, C. R., Classon, B. J., Tsagaratos, J., Walker, I. D., Kirszbaum, L., and McKenzie, I. F. C. (1986). Fractionation of detergent lysates of cells by ammonium sulfate induced phase separation. Anal Biochem. 156, 495-502. Payne, W. E., and Trumpower, B. L. (1986). A simple, one-step purification for cytochrome h from the hc^ complexes of bacteria. In "Methods in Enzymology'' (S. Fleischer and B. Fleischer eds.). Vol. 126, pp. 325-331. Academic Press, New York. Porath, J., Carlsson, J., Olsson, I., and Belfrage, G. (1975). Metal chelate affinity chromatography, a new approach to protein purification. Nature (London) 258, 598-599. Pryde, J. B. (1986). Triton X-114: A detergent that has come in from the cold. Trends Biochem. Sci. 11,160-163. Schagger, H., Brandt, U., Gencic, S., and von Jagow, G. (1995). Ubiquinol-Cytochrome c reductase from human and bovine mitochondria. In "Methods in Enzymology" (G. M. Attardi and A. Chomyn, eds.). Vol. 260, pp. 82-96. Academic Press, New York. Schagger, H., Link, T. A., Engel, W. D., and von Jagow, G. (1986). Isolation of the eleven protein subunits of the hc^ complex from beef heart. In "Methods in Enzymology" (S. Fleischer and B. Fleischer, eds.). Vol. 126, pp. 224-237. Academic Press, New York. Schagger, H., and Pfeiffer, K. (2000). Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J. 19,1777-1783. Seelert, H., Poetsch, A., Rohlfs, M., and Dencher, N. A. (2000). Dye-ligand chromatographic purification of intact multisubunit membrane protein complexes: application to the chloroplast R-'-FoFi-ATP synthase. Biochem. J. 346, 41-44. Tiselius, A., Hjerten, S., and Levin, O. (1956). Protein chromatography on calcium phosphate column. Arch. Biochem. Biophys. 65,132-155.

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Production and Purification of Recombinant Membrane Proteins ETANA PAD AN Microbial and Molecular Ecology Institute of Life Sciences Hebrew University of Jerusalem 91904 Jerusalem, Israel

CAROLA HUNTE Molekulare Membranbiologie Max-Planck-Institut fur Biophysik 60528 Frankfurt am Main, Germany

HELMUT REILANDER Aventis Pharma, Industriepark Hoechst 65926 Frankfurt am Main, Germany

I. INTRODUCTION The investigation of membrane proteins has become an active area in biochemical and biomedical research. For many experimental approaches, large quantities as well as high concentration of protein are needed. Structural analysis by NMR or x-ray crystallography requires, for example, several milligrams of functional protein in concentrated solutions ('^0.25 mM). However, membrane proteins often exist in their native membranes at low abundance. NhaA, the sodium/proton antiporter of Escherichia coli, comprises for instance less than 0.2% of the total membrane proteins (Taglicht et ah, 1991). Therefore, strong overexpression is needed to attain a high cellular concentration of the protein of interest. While for many soluble proteins

Membrane Protein Purification and Crystallization 2/e: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

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suitable overexpression systems are used routinely, high-level production of membrane proteins is still challenging. Overexpression of both homologous as well as heterologous proteins is often deleterious to the membrane and thus to the growth of the host organism (Miroux and Walker, 1996). A variety of expression systems for membrane proteins in bacteria, as well as in eukaryotic cells, has been developed to overcome these difficulties. Crucial steps in the production of membrane-bound proteins are not restricted to transcription and translation but include the insertion into and the folding of the protein within the membrane. The latter processes are often the rate-limiting steps in production. If correct insertion and folding fail, large quantities of inactive and partly aggregated protein are produced and accumulate within the cell. Although membrane protein insertion and folding follows general principles (Popot, 1993; Popot et ah, 1994), differences in the mechanisms between the prokaryotic and the eukaryotic systems are evident (Dobberstein, 1994). The insertion of a prokaryotic membrane protein is a co-translational process (deGier and Lurirink, 2001). The co-translational insertion of a protein into the membrane involves the signal recognition particle and its receptor (MacFarlane and Mueller, 1995; Seluanov and Bibi, 1997). A membrane potential is needed for correct membrane protein insertion in E. coli, a requisite not known for eukaryotic systems (Bassilana and Gwidzek, 1996; Wickner and Lodish, 1985). To date it is assumed that eukaryotic membrane proteins are cotranslationally inserted into the ER membrane, where they subsequently fold into their final three-dimensional conformation (Borel and Simon, 1996; Do et ah, 1996; Wessel and Spiess, 1988). In many cases, this process is supported by specialized folding proteins, so-called chaperones, like the BiP ("heavy chain binding protein" a member of the Hsp 70-heat shock protein family), the disulfide-isomerase and/or calnexin (Bergeron et al,, 1994; Hartl et ah, 1994; Helenius et al., 1992). In general, molecular chaperones prevent false intraand/or intermolecular interactions during the complicated folding process of proteins (Bergeron et al, 1994; Hartl et al, 1994; Gething and Sambrook, 1992). In only a few cases, there is exact knowledge of when and/or which chaperones are needed for correct folding of membrane proteins. NinaA, which acts as a peptidyl prolyl cis/trans isomerase, has been found to be absolutely essential for production of functional rhodopsin 1 and 2 in the eye of Drosophila melanogaster (Baker et al, 1994; Ondek et al 1992). In the absence of the ninaA protein, the rhodopsins aggregate and accumulate in the ER (Baker et al, 1994). If such specialized chaperones are needed for the correct production and, in particular, for the correct folding of recombinant membrane proteins, heterologous production appears to be very difficult to achieve. Here, the coproduction of specific chaperones might be a solution, which could finally lead to the design of a specialized host cell system (Lenhard and Reilander, 1997).

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Posttranslational modifications should be considered when producing a recombinant protein. Almost all eukaryotic membrane proteins that have been detected in the plasma membrane are glycosylated. Glycosylation of a protein might be important for its function and/or required for its correct folding, for the assembly of subunits, and also as a signal to guide the protein to the proper subcellular location (Geisow, 1992). Additional posttranslational modifications—for example, phosphorylation or palmitylation of receptors—are of functional importance. If posttranslational modifications are an absolute prerequisite for the proper function of a membrane protein, heterologous production can only take place in an expression system that is capable of performing these kind of modifications. Then, prokaryotic expression systems have to be excluded, and production of the respective protein has to be performed in an eukaryotic system (Geisse et ah, 1996). High-level production of active eukaryotic membrane proteins in E. coli, the most widely used bacteria, is still a challenge. This heterologous expression system often results in low yield, or the proteins tend to form insoluble inclusion bodies with inactive protein. Inclusion bodies also form during both homologous as well as heterologous overexpression of prokaryotic proteins in prokaryotic systems. Refolding of functional proteins from inclusion bodies has been described for very few membrane proteins (see Section II,C). In comparison to natural protein sources, expression systems for recombinant proteins offer an enormous potential for selective alteration of the protein structure to improve its properties for various aims. Removal of internal proteolysis or glycosylation sites will reduce heterogeneity of the product. Furthermore, protein or peptide fusions can be added to the protein of interest to facilitate its detection and purification. Expression constructs can be tailored to increase solubility, stability, and yield of the protein.

11. PROKARYOTIC EXPRESSION SYSTEMS FOR OVERPRODUCTION OF MEMBRANE PROTEINS FOR STRUCTURAL STUDIES Various strategies have been exploited in bacteria to achieve overexpression of membrane proteins: (1) use of strong promoters along with the proper transcription initiation and translation signals for overexpression; (2) exploitation of tightly regulated promoters to attain high biomass of bacteria before induction, followed by a limited induction time to obtain high amounts of the protein before the death of the bacteria; (3) use of leaky promoters without induction combined with low temperature to allow sufficient time for insertion and folding processes (Grisshammer and Tate, 1995); and (4) refolding proteins from inclusion bodies.

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The advantages in bacterial expression systems are as follows. 1. Bacterial protein expression is cheaper and faster than eukaryotic systems. 2. Overexpression is obtained without any posttranslational modification that can introduce heterogeneity, potentially harmful for crystallization. However, for proteins that require posttranslational modifications for proper folding and activity, eukaryotic expression systems are needed (see Section III). 3. Escherichia coli is the most frequently used bacteria for protein production due to its highly developed molecular genetics. Furthermore, advancing the understanding of the fundamental aspects of E. coli physiology—that is, protein folding, protein excretion, transcription, and translation—will continue to fuel progress in optimizing this organism for protein expression. This section is limited to expression systems developed in £. coli.

A, E. coli Inducible Promoters Used for Overexpression Plasmid-based inducible promoter systems have been constructed in E. coli using bacterial, phage, and chimeric promoters (Table I). One of the most widely used expression systems in E. coli is based on the T7 RNA polymerase (Studier et ah, 1990). The coding sequence of the enzyme is cloned into the E. coli chromosome downstream of an inducible promoter—for example, the IPTG inducible lac promoter from the excision-deficient A-lysogen DE3 (often used is the UV5 lac promoter that is insensitive to catabolite repression, [see Grossman et ah, 1998)]. The target gene is introduced into this bacteria on a plasmid positioned downstream to the T7 promoter. Therefore, target gene expression via the T7 RNA polymerase is inducible by IPTG. Other variations on this theme exist (for examples see Table I and Durbin, 1999). A thorough study of overexpression of several membrane proteins using this system and the E. coli strain BL21 (DE3) showed cell-toxicity caused by overexpression (Miroux and Walker, 1996). Mutants, C41 (DE3), C43 (DE3), surviving the toxicity were selected and shown to overproduce the proteins without toxic effect. Although neither the mechanism of toxicity nor the relevant mutations are known, the mutations are claimed to reside in the T7 RNA polymerase and to balance its activity with respect to translation. The procedure used for selection of these mutants could be employed to tailor expression hosts in order to overcome toxic effect caused by the T7 RNA polymerase based overexpression systems. The RNA polymerase of phage T7 is inhibited by the phage lysozyme (review by Durbin, 1999). Therefore, cotransformation of cells with one plasmid encoding lysozyme (pLysS and pLysE) and a second plasmid encoding the target protein has been advocated to suppress basal expression toxicity.

CHAPTER 3

TABLE I

Controlled Expression Syst ems Used in E. coli

Promoter used f or^ overexpression of the target gene cloned on plasmid

Regulatory manifold^

Induction

Level

Ref.

JpjQd

Very high XlOO

[1]

Temperature shift to 42°C

Moderately high

PL(^)

T7 RNA polymerase expressed from the lac promoter or derivatives in the presence of lacl^ (overexpressed repressor) The PL promoter in the presence of a)cV'S57 temperature sensitive repression The proU promoter Mutated 77 RNA polymerase in strain BL21 (DE3)'^; balancing transcription translation Lad, lacl^ repression (/l)cP^857 repression

araBAD

AraC repressor

Temperature shift to 42°C L-arabinose

tet

Tet repressor

Anhydrotetracycline

17 RNA polymerase

lac, tac, trc

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Production and Purification of Recombinant Membrane Proteins

NaCl IPTG

[2] [3]

IPTG

Moderately high Moderately high Fine-tuning (but see (5) and below) Very high

[4]

[5] [6]

''In addition to the promoter, other regulatory signals can be manipulated; DNA sequences important for transcription a n d / o r translation. ''Can be on the chromosome or plasmid. ^The site of the mutation has not yet been proven. The strains were successfully used to overexpress membrane proteins (Besette et ah, 1999; Miroux and Walker, 1996). '^IPTG, isopropyl jS-D-thiogalactoside. [1] For review, see Stevens, 2000; [2] Bhandri and Gowri-shankar, 1997; [3] Mulroony and Washell, 2000; Fiermont et al, 2001; [4] Brosius, 1999; [5] Guzman et a/., 1995; [6] Lutz and Bujard, 1997.

Various derivatives of the lac promoters are used for overexpression in addition to the native lacV (Table I). These are hybrids constructed from the — 35 consensus sequence of the trp promoter and the —10 of the lacV spaced by 16 bp in the case of tacV or by 17 bp in the case of trcV, Expression plasmid systems based on the lac promoter are notoriously leaky. Repression is often incomplete due to a combination of plasmid copy number effects and the absence of secondary operators required for the full range of gene control in

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the natural lac operon (review by Durbin, 1999). This problem is often overcome by the use of overexpressed repressor LacI^ or Lacl^l and/or the addition of an operator. Background expression levels should be lower in systems based on positive rather than negative control of expression. Recently, expression plasmids based on the araBAD promoter (FaraSAD) have been constructed (Guzman et al, 1995). Because the ara system can be induced by arabinose and is repressed by either catabolite repression in the presence of glucose or competitive binding of the anti-inducer fucose, these plasmids have very low background levels of expression. In addition, gene expression can be turned on and off rapidly by changing the sugars in the medium. Ideally, one would like to be able to modulate the level of gene expression. Guzman et al. (1995) presented evidence that the pBAD vectors are suitable to express at intermediate levels. However, intermediate levels of gene expression in cultures do not always mean that gene expression is uniform with respect to individual cells. In extreme cases, either every cell in the culture makes the same percentage of the fully induced level, which is observed in the culture, or the culture consists of a mixture of fully induced and completely noninduced cells. In the latter case, the intermediate expression levels observed in the culture reflect the proportion of cells that are fully induced, rather than intermediate expression in any individual cell. The latter alternative was found correct using Aequorea victoria green fluorescent protein (GFP) as a reporter (Siegele and Hu, 1997). Expression in cultures grown at low concentrations of arabinose is bimodal, consistent with an autocatalytic mechanism for induction of transcription of ParaSAD due to active uptake of the inducer. This phenomenon is analogous to the autocatalytic induction of the lac operon (Novick and Weiner, 1957). Expression from the lac promoter can be modulated using different concentrations of the membrane-permeable inducer IPTG in a lacY~ strain, where in the absence of an inducible transport system the level of expression in individual cells in the population is relatively homogeneous. Plasmids with promoters controlled by lac repressor have been used to modulate expression of genes over a 20- to 250-fold range (Hashemzadeh-Bonehi et al., 1998; Malony and Rotman, 1973; Prinz et al., 1997). The importance of performing such experiments with other systems should be emphasized. A recent review (Baneyx, 1999) describes the potential use for protein overexpression of other E. coli promoters, upstream and downstream sequences involved in transcription or translation, and the recent advancements in manipulating the copy number of the target gene (Baneyx, 1999).

B. Optimizing Expression Levels in E. coli Having a good nontoxic expression plasmid does not automatically guarantee that the protein is produced in large quantities. To achieve this goal

CHAPTER 3 Production and Purification of Recombinant Membrane Proteins TABLE II

61

E, coli Genotypes Relevant for Overexpression Relevant genotype

Relevant phenotype

endA Ion ompT argJJ ileY leuU proL extra copies of genes encoding tRNAs frequently limiting translation trxB, gor

Endonucleasel" Lon cytoplasmic protease" OmpT outer membrane protease" ArgU^, IleU"^, leuU"^, for AT-rich genomes ArgU^ and ProL"^ for GC-rich genomes"

dsbC (engineered in a trxB cytoplasm) met lacY lac operator

Thioredoxin reductase ", glutathion reductase " allow disulfide bond formation in the cytoplasm [1] Disulfide bond isomerase" [2] Methionine auxotrophe allows high specific activity labelling with ^^S methionine or selenomethionine for crystallography Lac permease" allows fine tuning with lac derived promoters [3] Addition for tight control with lac repressor

"For codon usage problems, it is also possible to systematically mutate the offending codons to synonymous codons of E. coli. [1] Prinz et al, 1997; [2] Bessette et al, 1999; [3] Hashemzadeh-Bonehi et al, 1998.

it may be necessary to prevent mRNA degradation, to remove undesirable features in the coding sequence that impede translation, to prevent proteolytic degradation, or to assist in protein folding. Hence, in every case of overexpression, empirical trials need to be performed for optimization, altering expression conditions, and observing the yield, solubility, and stability of the recombinant protein. The conditions that should be optimized are temperature, inducer concentrations, and media. In some cases, supplements of salts or trace elements (Zn^"^, Cu^"^, Ca^"^, Fe^"^, Fe^"^) and potential cofactors or prosthetic groups (e.g., heme, FAD, FMN, tetra hydro-biopterine) are needed. Host strain genotype is also important for obtaining optimal expression levels (Table II). Many E. coli strains have been developed for this purpose. The properties manipulated are high competence (usually patented); End A" for cloning expression constructs (endonuclease I is an enzyme that rapidly degrades plasmid DNA); Lon protease"; or OmpT protease" (these proteases can degrade recombinant proteins). Efficient production of heterologous proteins in E. coli is frequently Umited by the rarity of certain tRNAs that are abundant in the organisms from vvrhich the heterologous proteins are derived. Forced high-level expression of heterologous proteins can deplete the pool of rare tRNAs and stall translation. Strains are engineered to contain extra copies of genes that encode the tRNAs that most frequently limit translation of heterologous protein in E. coli.

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Availability of tRNAs allows high-level expression of many heterologous recombinant genes. These can be of high significance for genes derived from AT-rich or GC-rich genomes. Factors involved in protein folding are recruited now to improve overexpression and combat the problem of inclusion body formation. These include chaperones and peptidyl propyl cis/trans isomerases (for further reading, see Baneyx, 1999). Disulfide bonds do not form in the cytoplasm of wild-type E. coli strains, since this environment is reducing, trx^ encodes thioredoxin reductase, a protein responsible for the reduction of oxidized thioredoxins; gov encodes glutathion reductase. Deletion of these genes appears to facilitate accumulation of proteins containing structural disulfide bonds in the cytoplasm (Prinz et al., 1997; Venturi ei al., 2002). Another option is to direct the disulfide bond containing domain to the periplasm. There the environment is oxidizing and contains enzymes catalyzing the formation and rearrangement of disulfide bonds (Cornelis, 2000).

C. Inclusion Bodies and Refolding Overexpression of native proteins, but especially of heterologous recombinant proteins, leads to the formation of insoluble protein aggregates in the cytoplasm known as inclusion bodies. Current estimates are that 15-20% of human gene constructs expressed in E. coli are soluble, 20-40% form inclusion bodies, and the remainder do not express significantly or are degraded (for a recent review, see Rudolph and Lilie, 1996). Inclusion bodies are formed in both eukaryotes and prokaryotes, and there is no direct correlation between the propensity of their formation and intrinsic properties of the protein. For proteins containing disulfide bonds, inclusion body formation can be anticipated because the reducing cytosol inhibits disulfide bond formation (Hauke et al, 1998). Two strategies exist for dealing with inclusion bodies: (1) Minimize or avoid them by increasing the fraction of the protein that remains soluble. This involves modifications in the expression system (both host and vector) (review in Rudolph and Lilie, 1996; Hauke et al, 1998; Stevens, 2000); induction at lower temperatures; lowering the concentration of inducing agent; altering the media composition (e.g., adding sucrose or polyols); coexpressing chaperones and other in vivo folding enhancers; expressing the protein as a fusion with a "solubilizing partner" such as GST, MBP, thioredoxin, or NusA (see below). (2) Since inclusion bodies contain almost exclusively aggregates of the overexpressed protein, driving protein expression to inclusion body formation can be advantageous. Proteins in inclusion bodies are generally protected from proteolytic degradation. The use of inclusion body routes is especially attractive for the production of proteins that are toxic to host cells. Properly folded proteins can be produced from these inclusion bodies using a variety of

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solubilization and in vitro refolding schemes (for review, see Hauke et ah, 1998). Partial success at in vitro refolding has been obtained with hydrophobic interaction chromatography. Unfortunately, refolding screens must still be developed, especially as different proteins have highly variable refolding efficiencies. During generic refolding processes, severe heterogeneity problems can arise owing to the formation of multiple refolded species and aggregates. Hence, efforts aimed at recombinant protein refolding should probe as many factors as possible to maximize the probability of obtaining properly folded protein in its native conformation. Very few membrane proteins were produced by overexpression in E. coli as inclusion bodies and successfully refolded. These include two transport proteins from the mitochondrial inner membrane that appear ready for crystallization attempts (Fiermonte et ah, 1993; 2000) and a G-protein coupled receptor (Echtay et ah, 2000; Kiefer et ah, 1996,1999, 2000). Immobilization on nickel chelating chromatography material was succesfuUy used for refolding Toc75, the translocase of the chloroplast outer envelope, and LHC2, a pigment binding oligomeric assembly (Rogl et ah, 1998).

III. EUKARYOTIC EXPRESSION SYSTEMS FOR OVERPRODUCTION OF MEMBRANE PROTEINS Many eukaryotic expression systems have been tested for heterologous production of membrane proteins, including several single cell yeasts (Saccharomyces cerevisiae, Schizosaccharomyces pombe, and Pichia pastoris), insect cells, and mammalian cells (for reviews, see Grisshammer and Tate, 1995; Reilander et ah, 1999; Schertler, 1992). For the investigation of structure/function relationship of a protein by site-directed mutagenesis, transient expression of the modified protein in mammalian cell lines is the method of choice. It allows fast evaluation of the different altered protein isoforms. The most commonly used cell lines are those from Chinese hamster ovary (CHO), baby hamster kidney (BHK), human embryonic kidney (HEK) 293, and NIH3T3. Transient and stably transfected mammalian cell lines are also well suited for functional investigation of different receptor subtypes as well as for establishment of search systems for new receptor ligands by "high-throughput-screening" (HTS) a n d / o r ultra-HTS (UHTS) approaches (Stratowa et ah, 1995). Due to the presence of intact signalling pathways in most of the mammalian cell lines, analysis of signal transduction of receptors can easily be performed (Wilson et ah, 1999). However, for large-scale production and purification of membrane proteins, mammalian cell systems are less suited. These aims can be achieved by the Pichia pastoris system and the insect cell/baculovirus system, which will be introduced in more detail in this section.

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A. Yeast-Based Expression Systems Yeast-based expression systems are very attractive for heterologous production of membrane proteins. They are easy to manipulate and grow to high cell yield at low cost. Yeasts have been used in fermentation and food industry for decades and have been proven safe. Contrary to E. coli, yeasts have the potential to perform posttranslational modifications, which in some cases are needed for production of functional protein (Buckholz, 1993; Cereghino and Cregg, 1999, 2000; GeUisen and HoUenberg, 1997). Besides the classical expression system of S. cerevisiae, several other yeast systems have recently become popular. These include Schizosaccharomyces pombe, P. pastoris, P. methanolica, Hansenula polymorpha, Kluyveromyces lactis, Yarrowia lipolytica, or Candida utilis. However, only S. cerevisiae, Sz. pombe, and P. pastoris, have been tested for production of membrane proteins [for example: G-protein coupled receptors (GPCRs); see Reilander et ah, 1999 and a multidrug transporter (MRPl) (Cai et al., 1991)]. S. cerevisiae is the best characterized eukaryotic organism in terms of molecular genetics, cell biology, and physiology. Its complete genome sequence has been recently published (Goffeau et ah, 1996), and there are many ongoing studies to elucidate the function of newly discovered genes. A multitude of different strains including protease-deficient strains as well as a variety of expression vectors comprising yeast episomal plasmids (Yeps) and yeast integrating plasmids (Yips) are commercially available and allow easy genetic manipulations. The internal GPCR signal pathway, for example, has been modified and adapted to create a host for high-throughput screening of GPCRs (Pausch, 1997). However, S. cerevisiae has some limitations as a host: (1). It does not produce high cell densities, (2) during fermentation, plasmids, especially Yeps, may be unstable, and (3) glycoproteins may be incorrectly glycosylated—for example, hyperglycosylated. The fisson yeast Sz. pombe is phylogenetically as related to S. cerevisiae as it is to mammalian cells. It has many properties in common with higher eukaryotic organisms (Giga-Hama and Kumagai, 1997). A range of different expression vectors for Sz. pombe have recently become commercially available, some of which are based on the thiamine-repressible nmtl promoter {nmtl, no message in thiamine) (Maundrell, 1993). However, the biotechnological application of this yeast lags behind that of S. cerevisiae and P. pastoris. 1. Pichia pastoris as an Expression System The use of yeast as an expression system commenced when Phillips Petroleum converted the methylotrophic yeast P. pastoris into a producer of recombinant proteins. P. pastoris is one of about a dozen yeast species that can use methanol as its sole energy and carbon source. The metabolic pathway

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needed for methanol consumption involves a set of different enzymes where alcohol oxidase I (AOXl) catalyzes the first step. The expression of the AOXl gene is tightly regulated and specifically induced by methanol. More than 30% of the cell protein is AOXl when the cells grow on methanol in a fermenter. The first steps of the methanol utilizing pathway take place in specialized organelles, the peroxisomes. This compartmentalisation prevents possible oxidative damage to the cell from the production of hydrogen peroxide. In most cases, expression of foreign genes in P. pastoris was accomplished by using the strongly regulated promoter derived from the AOXl gene (Higgins and Cregg, 1998). Other vectors that use the promoter of the constitutive glyceraldehyde-3-phosphate dehydrogenase (GAP) gene (Waterham et ah, 1997) were constructed. Strains and vectors are commercially available (Invitrogen). P. pastoris is always used in combination with vectors for genomic integration. After transformation and integration of the linearized recombinant vectors, the resulting recombinant yeast clones are extremely stable even when induced. The techniques for molecular genetic manipulation of P. pastoris are very similar to those of S. cerevisiae. Additionally, P. pastoris has a strong preference for respiratory growth, a property that facilitates growth to extremely high cell densities of up to 500 g per liter culture medium. To date, more than 100 different proteins have been successfully produced in P. pastoris, including membrane-bound receptors (GPCRs) and transporters (Cereghino and Cregg, 1999, 2000). Four different methods are available for the transformation of P. pastoris. Although more laborious than the others but best characterized is the spheroblast method that produces transformants at high frequency. The other three methods rely on intact untreated cells. The most common whole-cell method of transformation is based on electroporation, which results in almost as many recombinants as the spheroblast method. The other two transformation procedures have the advantage that they do not need special equipment. Both of these methods, the PEG-transformation as well as the LiCl-method, result in the generation of an adequate number of transformants (Cregg and Russell, 1998). To date, all P. pastoris strains available are derivatives of the NRRL-Y11430 strain (Table III). To allow for selection of transformants that carry a vector containing the histidinol dehydrogenase gene (HIS4), most strains bear a mutation in HIS4. These strains are able to grow on complex media, but for their growth in minimal medium, histidine has to be supplemented. The most commonly used host for transformation is strain GS115, which posesses a wild type AOXl and grows rapidly on methanol (Muf^ phenotype). In contrast, the strain KM71 has a reduced growth rate on methanol (Mut^, methanol utilization slow), since the chromosomal AOXl gene was largely deleted and replaced by the S. cerevisiae ARG4 gene. The slow growth of this strain on

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TABLE III

P. pastoris Strains Strain GS115 X33

KM71

KM71H

SMD1168 SMD1168H

SMD1163

Genotype (application) his4 (selection of vectors containing HIS4) wild type (selection of vectors containing the zeocin resistance gene) aoxl::ARG4 his4 arg4 (selection of vectors containing HIS4 resulting in clones with Mut^ phenotype) aoxl::ARG4 arg4 (selection of vectors containing the zeocin resistance gene resulting in clones with Mut^ phenotype) Apep4his4 (selection of vectors containing HIS4) Apep4 (selection of vectors containing the zeocin resistance gene) Apep4 his4 Aprbl (selection of vectors containing HIS4)

Phenotype

Ref.

Mut+His"

[1]

wild type

[2]

Mut^His"

[31

Mut'His + protease-deficient

[2]

Mut+His"; protease-deficient Muf^His^; protease-deficient

[4]

Mut+His"; protease-deficient

[5]

[2]

Adapted and expanded according to Higgins and Cregg (1998). [11 Cregg et al, 1985; [21 Invitrogen; [3] Cregg and Madden, 1987; [4] White et al, 1995; [5] Gleeson et al, 1998.

methanol relies on the activity of a second alcohol oxidase gene, AOX2, which is present in the P. pastoris genome. A0X2 has an explicitly lower activity compared to AOXl. With many expression vectors it is possible to replace the AOXl gene in the GS115-related strains with an expression casette by homologous recombination resulting in a Mut^ phenotype. Proteolysis is a problem during production of recombinant proteins, especially membrane proteins (Weiss et al, 1995). The vacuolar proteases play a major role in this proteolysis. Therefore, the use of the protease-deficient strains SMD1168 or SMD1163 (Table III) is highly recommended. The PEP4 gene encodes for the vacuolar proteinase A, which is required for the activation of several other vacuolar proteases. The PRBl gene codes for proteinase B. SMD1163, a strain bearing the double mutation pep4 prbl, shows a significant reduction in protease activities and, therefore, provides an optimal environment for production of sensitive proteins. The strains X33, KM71H, and SMD1168H (Table III), which all bear no mutation in the HIS4 gene, are designed for transformation with expression vectors conferring resistance either to Zeocin or Blasticidin.

CHAPTER 3 Production and Purification of Recombinant Membrane Proteins

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The most commonly used vectors for heterologous production of proteins in P. pastoris are shuttle vectors (Table IV). Some vectors bear the wild-type HIS4 gene and the bacterial ampicillin resistance gene as selectable markers. The HIS4-based vectors are in general large (up to 10 kb), and the multiple cloning site is not very convenient. The AOXl promoter region present on these vectors is followed by a multiple cloning site suited for insertion of the foreign gene followed by the AOXl transcriptional termination sequence. The 3' flanking region of the AOXl gene contains the HIS4 selection marker, which is needed for screening the integration of the expression cassette at the AOXl locus by gene replacement. Furthermore, it contains sequences required for replication and propagation of the vectors in E. coli (for example, the pPIC9 vector). Generation of recombinant strains bearing multicopy integrations of the expression plasmid in the genome has been shown to result in an increased production of heterologous protein via a gene dosage effect. Special vectors that carry a second selection marker in addition to HIS4 were designed for this application. They contain the E. coli kanamycin gene of transposon 903 inserted between HIS4 and the 3' flanking region of AOXl (for example, the pPIC9K vector). This marker allows identification of recombinant clones bearing multicopy integration of the expression cassettes by screening for increased resistance to geneticin (G418). In the first step Pichia transformants are selected on histidine-deficient medium and then screened for their level of resistance to G418. However, the addition of the kanamycin gene increases the size of these already large plasmids and may yield unstable vectors. Smaller vectors were constructed using a single selectable marker active both in E. coli as well as in P. pastoris. To date, two selection markers are used for this purpose: the ble gene from Streptoalloteichus hindustanus, conferring resistance to Zeocin (pPICZ series) (Higgins et ah, 1998), and the hsd gene from Aspergillus terreus, conferring resistance to Blasticidin S (pPIC6 series). In these vectors, the respective resistance genes are preceded by a S. cerevisiae TEFl gene promoter needed for transcription in P. pastoris and the synthetic EM7 promoter needed for transcription in E. coli. The genes are followed by the 3' transcription processing sequences of the S. cerevisiae CYCl gene. Direct selection of hyperresistant transformants with multicopy integration events has been claimed but not clearly proven. These vectors have several advantages; they are small and contain an extended multiple cloning site, which in most cases allows C-terminal fusion of a c-myc-tag and a His6-tag to the foreign gene. Some of these vectors have special features. The so-called E-series of vectors are designed for directional cloning by rapid Cre-based recombination (pPICZ-E and pPICZa-E). In several vectors the S. cerevisiae a-factor prepro-signal sequence has been included to allow N-terminal fusion to the foreign gene. Although these vectors were constructed for the production of secreted proteins, GPCRs fused to this sequence showed a higher production level (Schiller et ah, 2000, 2001; Weiss et al, 1995).

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TABLE IV

P. pastoris Expression Vectors for Intracellular Production or Secretion Selection marker in yeast

Properties

Ref.

Intracellular pHIL-D2 pA0815

HIS4 HIS4

pPIC3.5

HIS4

pPIC3.5K

HIS4 and kan"^

pPICZ A, B, C

ble'

pPICZ-E

ble'

pPIC6 A, B, C

bsd'

pHWOlO pGAPZ A, B, C

HIS4 ble'

AOXl-promoter AOXl-promoter; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette A0X2-promoter; multiple cloning site for insertion of the foreign gene AOXl-promoter; multiple cloning site for insertion of the foreign gene; G418selection for multicopy clones AOXl-promoter; multiple cloning site for insertion of the foreign gene; Zeocin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional) AOXl-promoter; vector designed for in vitro recombination (echo-cloning system); zeocin-selection AOXl-promoter; multiple cloning site for insertion of the foreign gene; blasticidin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional) GAP-promoter GAP-promoter; multiple cloning site for insertion of the foreign gene; zeocin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional)

[1] [2]

[3] [3]

[41

[1]

[1]

[5] [1]

Secretion pHIL-Sl

HIS4

pPIC9

HIS4

AOXl-promoter; PHOl-signal sequence; multiple cloning site for insertion of the foreign gene A0X2-promoter; S. cerevisiae a-factor signal sequence; multiple cloning site for insertion of the foreign gene

[1]

[3]

69

CHAPTER 3 Production and Purification of Recombinant Membrane Proteins

TABLE IV

Continued Selection marker in yeast pPIC9K

HIS4 and kan'

pPICZaA, B, C

ble'

pPICZa-E

hie'

pPIC6a A, B, C

bsd'

pGAPZaA, B, C

ble'

Properties

Ref.

AOXl-promoter; S. cerevisiae a-factor signal sequence; multiple cloning site for insertion of the foreign gene; G418-selection for multicopy clones AOXl-promoter; S. cerevisiae a-factor signal sequence; multiple cloning site for insertion of the foreign gene; zeocin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional) AOXl-promoter; S. cerevisiae a-factor signal sequence; vector designed for in vitro recombination (echo-cloning system); zeocin-selection AOXl-promoter; S. cerevisiae a-factor signal sequence; multiple cloning site for insertion of the foreign gene; blasticidin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional) GAP-promoter; S. cerevisiae a-factor signal sequence; multiple cloning site for insertion of the foreign gene; zeocin-selection; BamHl and Bglll restriction sites for in vitro multimerization of the expression cassette; His6 and c-myc epitope tags (optional)

[3]

[6]

[1]

[1]

[1]

Adapted and expanded according to Higgins and Gregg (1998). [1] Invitrogen; [2] Thill et ah, 1990; [3] Scorer et ah, 1993; [4] Higgins et al, 1998; Scorer et al, 1993; [5] Waterham et al, 1997; [6] Higgins et al, 1997.

For constitutive production of foreign proteins the GAP promoter (Waterham et al., 1997) has recently been inserted in pPICZ vector backbone. The GAP promoter is a convenient alternative to the AOXl promoter as long as the expressed products are not toxic to the cell. As yet, membrane proteins have not been produced by these vectors.

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As in S. cerevisiae, linearized vectors are used for the generation of stable transformants via homologous recombination fostered by sequences shared between vector and genome. Even in the absence of an antibiotic selective pressure, such transformants are stable. The vectors mentioned above carry at least one genomic DNA fragment (the promoter) with a unique restriction site that can be cleaved and used to direct the vector to integrate into the P. pastoris genome via a single crossover. Vectors that contain the HIS4, therefore, can be forced to insert into the P. pastoris genomic his4 locus. Vectors that bear in addition the 3'AOXl sequences can be integrated in the genome by single crossover at either the AOXl or the HIS4 locus, but they can also replace the AOXl completely by a double crossover event. Recombinant clones resulting from such an event (replacement) show a His"^ Mut^ phenotype. Multiple integrations of the expression cassette occur sponataneously at low frequency (1-10% of His"^ transformants). Detection for these events is possible by screening for clones showing hyperresistance on G418 or Zeocin. As mentioned above, when the AOXl promoter is used for transcriptional control of the foreign gene, a major advantage of P. pastoris is that it can be grown to extremely high cell densities in fermenters. The AOXl promoter is induced by methanol and repressed in the presence of excess glycerol. A standard fermentation protocol is to grow the recombinant cells with an amount of glycerol, just enough to get very high cell density. When glycerol is exhausted, protein production is induced by supplementation of methanol to the medium. The advantage of Muf^ transformants is their faster growth on methanol. However, the concentration of methanol must be tightly controlled to avoid enrichment of toxic formaldehyde, the product of the first enzymatic step in the metabolism of methanol. Mut^ strains grow slowly on methanol because they lack AOXl and their growth relies on AOX2. In principal, cultivation of a Mut^ strain is similar to that of Mut^ strains, but lower methanol feed rates are needed to maintain a methanol concentration of 0.2-0.8% in the fermenter. Alternatively, Mut^ strains can be grown in a mixed glycerol/methanol medium (Stratton et al., 1998).

B. Insect Cells as Expression System Different insects cell lines for stable or transient transfection with appropriate vectors are available. Stably transformed insect cells have a number of potential benefits over mammalian cell-based expression systems (McCarroU and King, 1997; Pfeifer, 1998). These include lower growth temperature, less complex growth conditions, the possibility for a long-term continuous culture, growth in large fermenters, and the possibility of protein production in a null background. Vectors, which allow quick selection of stable cell clones, were constructed for transfection of Spodoptera frugiperda (Sf9 and Sf21 cell lines), Trichoplusia ni ("high five" cell lines), and Mamestra brassicae cell lines. Most commonly, baculovirus immediate early (ie) promoters from Bombyx mori

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multicapsid nuclear polyhedrosis virus (BmMNPV), Autographa californica (AcMNPV), and Orgyia pseudotsugata (OpMNPV) are used to drive transcription of the foreign gene (Jarvis et ah, 1990; Jarvis and Finn, 1996). Other widely used insect promoters include that of actin, of Drosophila melanogaster metallothionein, of Drosophila hsp70, and of copia, which function in the cell lines, albeit at a much lower level. The latter promoters have been mainly used for production of proteins in Drosophila, Schneider S2 and S3 cell lines, and mosquito cells. A variety of selection systems can be applied for the stable transformation of insect cells. They are based on genes conferring resistance to geneticin (G418), hygromycin B, methotrexate, and, recently, Zeocin and Blasticidin. The procedure for the generation of stably transformed insect cells is easy. Cells are seeded into small dishes and transfected with the appropriate recombinant vector using standard transfection methods such as calcium phosphate precipitation, electroporation, or liposome-mediated transfection. After a growth phase of 1-2 days to ensure expression of the resistance gene, selective pressure is applied by administration of the respective antibiotic. Nonresistant (i.e. nontransfected) cells die, and individual recombinant cell clones can be picked for further amplification. One of the most commonly used applications of stably transfected insect cells is the production of receptors or receptor subunits. Since in most cases insect cells do not appear to produce homologous endogenous receptors, the respective proteins are produced in an extremely low background. The insect/baculovirus-expression system has also proven to be successful for overproduction and characterization of several membrane proteins. The recombinant membrane proteins produced in the insect cells are subjected to posttranslational modifications such as glycosylation, phosphorylation, and palmitoylation. Most importantly, simultaneous infection of insect cells with several recombinant baculoviruses is possible. This opens the way for the production of complex multisubunit proteins or for enhancement of production levels of certain proteins by coproduction with a set of appropriate chaperons. 1. The Baculovirus Expression System The baculoviridae comprise a large family of occluded viruses, which are pathogenic for arthropods belonging to the insect orders Lepidoptera, Diptera, and Hymenoptera (Miller, 1996). In the last decade, baculoviruses, especially the Autographa californica nuclear polyhedrosis virus (AcMNPV), have been developed to a popular and versatile expression system (Kost and Condreay, 1999). The virus can be propagated easily in a multitude of cell lines, from which Spodoptera frugiperda (Sf9 and Sf21 cell lines), Trichoplusia ni ("high five" cell lines), and Mamestra brassicae are most frequently used (Hink et ah, 1991). As most baculoviruses, the AcMNPV has a complex, biphasic life cycle with

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two distinct morphological virus types (Miller, 1996). The so-called budded virus form comprises a single enveloped virus, whereas the so-called occluded virus form comprises multiple virus particles that become embedded in a protein matrix composed almost exclusively of one protein called polyhedrin. The development of the expression system was based on the dispensibility for in vitro propagation of at least two very "late" viral gene products: the plO and the polyhedrin gene. Both genes are under control of strong promoters, which were used for the design of expression vectors. However, most baculovirus expression vectors use the polyhedrin gene promoter and locus for insertion of foreign DNA. "Early" promoters have also been used for the heterologous production of proteins but were found to be less effective (Jarvis et al, 1996). The manipulation for direct cloning of the baculovirus genome, a closed, circular, double-stranded DNA molecule of about 130 kb, is a difficult task. Therefore, recombinant transfer vectors containing the flanking region of the polyhedrin gene were constructed for cloning of the foreign gene. The recombinant virus is generated by homologous recombination with these vectors. The viral DNA and the recombinant transfer vector are cotransfected, and the expression cassette recombines into the virus genome, replacing its wild-type polyhedrin gene. The efficiency of this technique is less than 1%, and the screening for recombinant virus, which is performed by plaque assays a n d / o r hybridizations, is tedious and time-consuming (Luckow and Summers, 1988). By using linearized virus DNA for cotransfection, the efficiency was raised to about 30% recombinants per virus progeny. For this approach a unique Bsw36I restriction site was introduced at the polyhedrin locus of wild-type baculovirus DNA (Kitts et ah, 1990). A further development was the use of the baculovirus designated BacPAK6 for cotransfection. This virus DNA contains the E. coli lacZ gene at the polyhedrin locus and two additional Bsu36l sites in the polyhedrin flanking regions (Kitts and Possee, 1993). After Bsu36l digestion the virus DNA is not only linearized but two genomic fragments are removed. One disrupts a gene essential for replication of the virus. Restoration of this gene is only possible after recombination of the linearized and substracted virus genome with the respective polyhedrin-flanking region in the recombinant cotransfected transfer vector. The selection of recombinant baculovirus is based on LacZ (blue/white assay). Recombinant baculoviruses form white plaques on a background of blue plaques. Due to these modifications the efficiency of the system was raised to 80-90% recombinants per virus progeny. The system is commercially available (BaculoGold, Pharmingen). An alternative system is based on linearized BacPAK6 DNA (Orbigen). To increase expression, the gene encoding for protein disulfide isomerase has been inserted at the plO locus of the virus (Sapphire Baculovirus DNA). This might be advantageous for the production of membrane proteins. Another popular system is based on a site-specific transposition of an expression cassette into a baculovirus, which propagates in E. coli (Luckow et

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al, 1993, Invitrogen). Here, the complete baculovirus genome replicates as a large and stable, low-copy number bacmid (pMON14272) in E. coli. This was achieved by recombination of a mini-F replicon into the polyhedrin locus of the virus. Additionally, the target site for the Tn7 transposon, a kanamycin resistance gene, and the lacZoc peptide coding region originating from a pUC-based vector have been introduced into the bacmid. The attachment site for the Tn7 transposon (mini-affTn7) was inserted corresponding to the N-terminus of the lacZoc region without disrupting the reading frame. Transposition of an expression cassette is performed in the E. coli DHlOBac strain harbouring the bacmid as well as the tetracyclin-resistant helper plasmid pMON7124, which in-trans supplies proteins necessary for transposition. The cells are transformed with an ampicillin-resistant vector that contains the foreign gene cloned under the transcriptional control of the polyhedrin promoter and the gentamycin acetyltransferase gene flanked by the left and right arms of Tn7. The mini-Tn7 in the so-called pFastBac vectors, which also contain the inserted foreign gene, can transposit to the vmni-attTn? attachment site on the bacmid. This transposition disrupts the lacZa peptides so that colonies, which contain a recombinant bacmid, are white in a background of blue colonies. Recombinant transformants are selected for in the presence of kanamycin, tetracyclin, gentamycin, X-Gal, and IPTG. The bacmids are isolated from the transformants and directly used for transfection of insect cells, eliminating the need for a tedious screening of positive plaques. In addition, the time required to generate, identify, and characterize a recombinant baculovirus with the transposition-based baculovirus system is reduced to about a week as opposed to six weeks with the former approach. For the procedures introduced here, different vectors are available, some of which also allow fusions to signal sequences, different tags (His-tag, c-myc-tag), as well as other proteins (GST). Some vectors allow simultaneous production of two or three proteins from one expression cassette via multiple promoters and multiple cloning sites.

IV. USE OF FUSION PROTEINS AND AFFINITY TAGS FOR PURIFICATION OF MEMBRANE PROTEINS Peptide affinity tags and fusion proteins are valuable tools that allow efficient and standardized purification of recombinant proteins. These affinity markers are attached to the target protein by gene fusion, thereby adding specific properties to the recombinant protein that can be exploited for protein purification via affinity chromatography, as well as for easy detection and immobilization. Affinity chromatography systems are designed to be highly specific and fast. They allow protein purification with high-yield, highenrichment, and often mild-purification conditions. In general, affinity tags

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and fusion proteins can be added to the amino(N)-terminus or the carboxyl(C)-terminus of the protein. Alternatively, the DNA encoding the respective affinity marker may be introduced within the coding sequence of the target protein, a so-called internal fusion. The latter approach requires information about the secondary structural elements and the topology of the target. In some cases, linker regions have to be included to ensure accessibility of the affinity marker. The introduction of a tag or a fusion protein should affect neither the activity or structure of the protein of interest nor interfere with targeting, folding, or membrane insertion of the protein. In some cases, destabilization of the product and low yield are provoked, since the marker may disturb folding processes of the protein or increase its susceptibility to proteolytic degradation. The prediction of disadvantageous side effects is difficult, and, in practice, each affinity system has to be established empirically. Therefore, it is recommended to prepare different gene arrangements with respect to type and location of the tag and the length of the linker region. Short affinity tags are less likely to interfere with structure and function of the target protein and might be preferred for this reason. However, a stabilizing effect of larger fusion proteins has been reported and might be desirable. The insertion of a highly specific protease cleavage site between protein and affinity tag or fusion domain allows the removal of the attached marker after protein purification. The most common examples of affinity tags and fusion proteins used for membrane proteins will be described in this section. A general requirement for this application is tolerance to detergents. The Hzs-tag is probably the most widely applied affinity tag. Six or 10 consecutive histidine residues are fused to either the N- or the C- terminus of the protein. The tagged protein can be purified by immobilized metal-affinity chromatography (IMAC, Hochuli et ah, 1988). For this purpose metal-chelating matrices, which are loaded with transition metal ions (Co^"^, Ni^"^, Cu^^, Zn^), are used. The hybrid protein binds by interaction of the histidine imidazole groups with the free ligand binding sites in the coordination sphere of the metal ions. After applying the protein mixture to the column, unbound material is washed off with an appropriate buffer. The specifically bound protein is eluted by adding free imidazole to the elution buffer or by lowering the pH. This rapid method allows 100-fold enrichments in a single purification step and results in u p to 95% purity of the affinity-tagged protein (for review, see Bornhorst and Falke, 2000). It has been applied for purification of recombinant proteins from a number of different expression systems. The most commonly used H/s-tag consists of six histidine residues. Longer histidine stretches allow more stringent washing conditions and might help to reduce impurities. However, the longer the tag, the higher the risk that it interferes with the protein function. Affinity matrices are commercially available—for example, Nickel-nitrilotriacetic acid (Ni^"^-NTA, Qiagen) or cobaltcarboxymethylaspartate (Co^'^-CMA or Talon, Clontech). The binding affinity

CHAPTER 3 Production and Purification of Recombinant Membrane Proteins

75

is very high compared to other affinity systems. IMAC materials are reusable, since the metal ions can be stripped off and reloaded for regeneration. A binding capacity of standard affinity matrices of 5-10 mg of protein per ml of matrix is described. Furthermore, the matrices tolerate the presence of detergents and can be used easily for the purification of membrane proteins as, for example, the sodium/proton antiporter from E. coli (Olami et ah, 1997), the plant plasma membrane H"^-ATPase (Jahn et ah, 2001), and the trehalose/ maltose ABC transporter (Greller et ah, 2001). Remarkably, the siderophore receptor FhuA located in the outer membrane of E. coli was successfully purified with an internal H/s-tag. A hexahistidine peptide was inserted in a loop that was known to be surface exposed. This approach allowed purification of the receptor to homogeneity in large quantities and consequently its crystallization suitable for x-ray analysis (Ferguson et ah, 1998). Rogl et ah (1998) succeeded in refolding the chloroplast light-harvesting complex II from inclusion bodies, while the protein was immobilized on an IMAC matrix. In addition, the His-tag is often used in multiaffinity fusion systems, in which several tags or fusion proteins are attached to the same protein (Bornhorst and Falke, 2000). The Strep-tag is another useful but, until today, less well-known affinity marker. The short peptide consists of nine amino-acids and binds to streptavidin, allowing one-step purification of the fusion protein via streptavidin affinity chromatography (for review, see Skerra and Schmidt, 2000). The Sfrep-tag binds to the biotin binding pocket of streptavidin and permits very gentle elution of the recombinant protein from the affinity matrix by competitive displacement. Already small amounts of biotin or analogues thereof are sufficient for efficient elution without changing the overall buffer conditions with respect to pH or ionic strength. This second specificity-conferring step enables purity grades of over 99% in one step. While the original Sfrep-tag with the sequence Ala-Trp-Arg-His-Pro-GlnPhe-Gly-Gly (Schmidt and Skerra, 1993) can only be used as C-terminal fusion, the slightly modified version Strep-tag II (Asn-Trp-Ser-His-Pro-GlnPhe-Glu-Lys) allows in addition N-terminal and internal addition of the affinity tag. Affinity matrices can either be custom-made by coupling commercially available streptavidin to a variety of matrices or they can be obtained ready-made. Strep-Tactin columns (IBA) consist of an engineered streptavidin mutant with optimized binding properties for the Strep-tag II (Voss and Skerra, 1997) immobilized to various matrices, enabling affinity purification via gravity flow, FPLC, or HPLC. Binding of Strep-tagged proteins to streptavidin matrices is highly specific. Thus, it is an attractive alternative to Hfs-tagged proteins when problems with unspecific contamination cannot be solved. Specific elution of the protein from streptavidin matrices is most often performed with desthiobiotin. Biotin itself binds irreversibly to tetrameric streptavidin and thus does not allow the reversible use of the affinity matrix. To avoid contamination with endogenous biotin or biotin-carrying proteins.

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extracts can be pretreated with avidin that will irreversibly bind and thereby mask biotin, while the Strep-ieig is not recognized. The Strep-tag system has been used for the purification of a variety of recombinant proteins, although applications described up to now comprise mainly proteins from bacterial sources. It seems to be a promising approach for membrane protein purification, since the streptavidin affinity matrix tolerates various detergents, the binding properties are highly specific, and the elution conditions are very mild. The betaine carrier BetP from Corynebacterium glutamicum was successfully purified by streptavidin affinity chromatography (Riibenhagen et ah, 2000). Furthermore, the Strep-tag has already been proven to be suitable for the purification of multisubunit membrane protein complexes, as, for example, the cocomplex of cytochrome c oxidase from Paracoccus denitrificans with the specifically bound antibody fragment carrying the Strep-tag (Ostermeier et ah, 1995). 3D crystals of this protein preparation (Ostermeier et ah, 1995) led to the structure determination of the P. denitrificans cytochrome c oxidase (Iwata et ah, 1995). The Flag-tag is an epitope tag that can be used for membrane protein purification. This peptide consists of eight amino acid residues (Asp-Tyr-LysAsp-Asp-Asp-Asp-Lys) and forms the epitope for a set of different monoclonal antibodies. The monoclonal antibody Ml reacts specifically with the Flag-tag at the free N-terminus of the protein, and the binding is calcium dependent. Therefore, Flag-tagged proteins can be bound to a mAB Ml affinity matrix in the presence of calcium and eluted by adding EDTA to the buffer. Furthermore, the N-terminal specificity offers an elegant method to select for recombinant proteins with correctly processed leader sequences (see Chapter 9) or for selection of homogenous products after cleavage of Nterminal fusions. The mAB M2 binds to the Flag-epitope at the N- or C-terminus of the protein, but the binding is not calcium dependent. Elution of protein bound to mAB M2 affinity columns can be achieved by pH shift or by competition with the free Flag-peptide. Antibodies as well as specific affinity matrices are commercially available (Eastman Kodak). In vivo biotinylation can be exploited for protein purification. Biotinylation occurs in all eukaryotic and prokaryotic organisms, and the number of different endogenous biotinylated proteins is low. The protein of interest is fused to a Bio-tag that provides the biotin (vitamin H) accepting domain either in peptide or protein domain format. The highly specific interaction of biotin with streptavidin or avidin allows selective purification of the marked protein. The system can be used in principal for all expression systems and different biotin acceptor domains have been successfully introduced. Biotinylated proteins bind to monomeric avidin with a dissociation constant of about 10"^ M compared to 10"^^ for the tetrameric form. Therefore, immobilized monomeric avidin is often used as affinity matrix, and elution can be easily accomplished by competitive elution with low concentrations of biotin (for review, see Cull and Schatz, 2000). This system was used, for example, for the

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purification of the jS2-adrenergic receptor, which was fused to a 75 amino acid biotinylation domain of Propionibacteriutn shertnanii and expressed in Pichia pastoris (see Chapter 9). The rat neurotensin receptor expressed in E. coli was purified most efficiently with a 13-amino-acid-long Bzo-tag at its C-terminus (Tucker and Grisshammer, 1996). Various fusion proteins — for example, the maltose-binding protein (MBP), the jS-galactosidase (GAL), thioredoxin (TRX), glutathione S-transferase (GST), or protein A—have been attached to proteins to facilitate purification. The fusion of larger domains offers the advantage of not only providing an affinity marker but it often stabilizes the recombinant protein by preventing degradation or promoting solubility. The N-terminal fusion of thioredoxin was essential for stabilizing the membrane-bound protein cytochrome b (5), whereas the C-terminal Hzs-tag was used for purification (Begum et ah, 2000). Again, a stabilizing effect of thioredoxin was reported for the production of the rat neurotensin receptor expressed in E. coli (Tucker and Grisshammer, 1996). In addition, thioredoxin can be directly applied for purification by either using its inherent thermal stability or by introducing secondary affinity properties to thioredoxin—for example, with in vivo biotinylation (for review, see LaVaUie et ah, 2000). Hybrid proteins containing the MBP fusion can be easily purified by affinity chromatography using immobilized amylose matrix (for review, see Sachdev and Chirgwin, 2000). This method has been successful for the purification of membrane proteins (Hampe et ah, 2000; Henderson et ah, 2000). GST fusion proteins are commonly used for protein purification from E. coli sources. The corresponding affinity matrix is glutathione-agarose. Bound protein can be eluted by adding glutathione to the buffer. No standard protocols can be applied, but conditions for purification have to be carefully optimized (for review, see Smith, 2000). A truncated form of the human Na"^/glucose cotransporter was expressed and purified from E. coli using a GST fusion vector and glutathione affinity chromatography (PanayotovaHeiermann et ah, 1999). When constructing the expression cassette for the hybrid protein, it is advisable to include a short sequence for a specific protease site between the target protein and the affinity fusion protein. Specific proteases such as thrombin, factor Xa, or Tobaco Etch Virus (TEV) protease (Dougherty et al., 1989) are used for the selective removal of the fusion partner, a step that might be exploited for purification.

ACKNOWLEDGMENTS Thanks are due to Danieli Tsafi (Protein Expression Facility), Institute of Life Sciences, Hebrew University of Jerusalem, Givat Ram for help and advice, and to the support by the BMBF and International Bureau of the BMBF at the

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DLR (German-Israeli Project Cooperation of Future-oriented Topics) and the Israel Science Foundation. Financial support by the Deutsche Forschungsgemeinschaft (SFB 472) to C.H. is acknowledged.

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4

SDS Electrophoresis Techniques HERMANN SCHAGGER Zentrum der Biologischen Chemie Universitatsklinikum Frankfurt 60590 Frankfurt am Main, Germany

I. INTRODUCTION Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) has become a valuable means for separation of proteins, and several electrophoresis systems that provide excellent resolution are available (see Hames and Rickwood, 1998). The Laemmli system (Laemmli, 1970) has advantages for separation of large proteins, and the Tricine-SDS-PAGE system (Schagger and von Jagow, 1987) is particularly useful for small proteins and peptides. These two electrophoresis systems cover the entire molecular mass range. Adjusting electrophoresis conditions for the special requirements of membrane proteins largely circumvents problems such as membrane protein aggregation during incubation in SDS and low recovery during electroelution and electroblotting.

Membrane Protein Purification and Crystallization 2/e: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

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11. CHOICE OF OPTIMAL SDS-PAGE SYSTEM AND GEL TYPE A. Wide versus Narrow Molecular Mass Range Wide molecular mass ranges usually are best covered by using acrylamide gradient gels and the Tris-glycine system according to Laemmli (1970). If there is no protein of interest larger than 100 kDa, the Tricine system described below is preferred. If there is a narrow molecular mass range of proteins to be separated, gradient gels should not be used. Although protein bands are usually sharper in acrylamide gradient gels compared with uniform gels, the resolution is lower because the separation distances between bands are decreased. The best gel systems and gel types for different separation problems are presented in Table I. Note: Use the Laemmli system if two proteins to be separated have similar molecular masses above 30 kDa; use the Tricine system for smaller proteins. If both systems show equal resolution, use the Tricine system for the advantages in electroelution and electroblotting.

B. Separation of Proteins with Similar Molecular Masses If the optimal gel/system is still not sufficient for resolving two proteins, one can often take advantage of the variable migration behavior displayed by individual proteins under special conditions. TABLE I

Gels and Systems for Optimal Separation Protein range

M, limits (kDa)

Optimal gel/system

Wide

6.0-250 2.5-200 2.5-100 1.0-70

8-16% gradient/Laemmli" 4-20% gradient/Laemmli 10% uniform/Schagger'' 16% uniform/Schagger

Mj. optimum (kDa) Narrow

50-100 40-80 30-60 5-50 2-30 1-20

8% 10% 12% 10% 16% 16%

uniform/Laemmli uniform/Laemmli uniform/Laemmli uniform/Schagger uniform/Schagger uniform/Schagger^

''LaemmU(1970). ''Schagger and von Jagow (1987). Concentration of cross-linker is doubled; gels without urea have nearly the same resolution as those with 6 M urea.

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1. Effect of Acrylamide Concentration Some proteins, especially membrane proteins, show anomalous migration behavior in SDS gels. Such proteins may bind an unusual ratio of SDS molecules per amino acid residue or may have hydrophobic stretches that do not completely unfold. These characteristics are reflected in a change in the apparent molecular mass in gels differing in acrylamide concentration. If circumstances require separation of proteins having similar electrophoretic mobilities, run samples at acrylamide concentrations above and below the "optimal" acrylamide concentration for the given molecular mass (as deduced from Table I). If one protein has anomalous migration behavior, it typically will shift to higher apparent molecular masses at higher acrylamide concentration, and the apparent molecular mass will approach the true molecular mass. For determination of molecular mass by a Ferguson plot, see Ferguson (1964) and Hames and Rickwood (1998). 2. Effect of Cleavage of Internal Disulfide Bridges To ascertain whether either of the two proteins to be separated may contain internal disulfide bridges, incubate samples in SDS in the presence or absence of reducing agents (DTT, DTE, mercaptoethanol). Differences in migration distances, corresponding to a 1-3 kDa shift in the apparent molecular mass, are often observed. 3. Use of Urea to Alter Electrophoretic Mobilities Unusual migration of individual proteins, especially of small watersoluble and membrane proteins, can be induced by including 6-8 M urea in the gels. This presumably reflects changes in SDS binding or conformation of the SDS-denatured protein in the presence of urea, to an extent dependent on the specific protein involved. Some proteins display a reduced relative electrophoretic mobility and higher apparent molecular mass, whereas others seem to be only slightly affected. A side effect of electrophoresis in urea, relevant to all proteins, is the generally reduced migration distances that are observed. This effect may be helpful for increasing the resolution of small proteins. Note: Several complications arise when urea is used. First, urea gels are not suited for determination of molecular mass. Addition of 8 M urea may cause extreme retardation, especially of small proteins (13 5->13 6^18

"The lowest polymerizing acrylamide concentration (which varies with the lot of acrylamide used) is required for complexes as large as 10,000 kDa. Use "analytical gels'' with dimensions 14 X 14 X 0.16 cm (sample wells 0.5 or 1 cm) or "preparative gels" with dimensions 14 X 14 X 0.3 cm (one 14 cm sample well). Gradient gel preparation is exemplified in Table III.

1. Materials Dodecyl-j8-D-maltoside, Triton X-100, and 6-aminocaproic acid can be purchased from Huka; acrylamide and bisacrylamide (the commercial twice crystallized products) and Coomassie blue dyes can be purchased from Serva; and digitonin can be purchased from Roth. All other chemicals can be purchased from Sigma. 2. Stock Solutions and Buffers The detergents dodecyl-j8-D-maltoside, Triton X-100, and digitonin are stored as 10% stock solutions. Digitonin, which precipitates at 4°C and also partly at room temperature, is redissolved by heating shortly before use. Stock solutions of Tricine (1 M), 6-aminocaproic acid (2 M), imidazole (1 M), and imidazole/HCl (1 M; pH 7.0) are stored at 7°C. Buffers for BN-PAGE and further stock solutions are listed in Table I. 3. Gel Types BN-PAGE separates proteins in the molecular mass range from 10 kDa to 10,000 kDa. A selection of gel types for optimal resolution of specific molecular mass ranges is given in Table II. Use only gradient gels overlaid with a large-pore sample gel.

4. Choice of Detergent and Detergent/Protein Ratio for BN-PAGE Any neutral detergent, and also mild anionic detergents like taurodeoxycholic acid, can be used for solubilization of biological membranes. We preferentially use dodecyl-j8-D-maltoside, Triton X-100, and digitonin. Digitonin usually retains physiological protein-protein interactions best, and

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CHAPTER 5 Blue Native Electrophoresis TABLE III

GRADIENT GEL PREPARATION" Gradient separation gel

AB-mix Gel buffer (3 x ) Glycerol Water APS10%^ TEMED Total volume

Sample gel 4% T

5%T

13% T

0.5 ml 2 ml

1.88 ml 6 ml

—

—

3.9 ml 5 ml 3g 3 ml 75/il 7.5 fA 15 ml

3.5 ml 50 fA 5fil 6 ml

10 ml 100/^1 10 fi\ 18 ml

''Volumes for one analytical gel (14 x 14 x 0.16 cm). Linear gradient separation gels are cast at 4°C and maintained at room temperature for polymerization. The volume of the 5% T solution is larger than that of the 13% T solution containing glycerol. This assures that the two solutions initially are not mixed when the connecting tube is opened. The sample gel is cast at room temperature. After removal of the combs, gels are overlaid with gel buffer (1 x) and stored at 4°C. ''APS10% is 10% aqueous ammonium persulfate solution, freshly prepared.

it is especially useful for isolation of physiological supercomplexes. DodecyljS-D-maltoside and Triton X-100, in contrast, usually split the supercomplexes into separate complexes. As a first rule of thumb, bacterial membranes are solubilized by using a detergent/protein ratio of 1 g Triton or 0.75 g dodecylmaltoside or 2 g digitonin per gram of total protein. Mitochondrial membranes require a doubled detergent/protein ratio. However, as a second general rule, don't use a single detergent/protein ratio for first trials, but solubilize several aliquots with increasing amounts of detergent—for example, to obtain Triton/protein ratios of 0.4, 0.75, 1.5, 3.0 (g/g) for bacterial membranes — so at least one detergent/protein ratio is below the solubilization limit. At least one of the higher detergent/protein ratios should then guarantee complete solubilization with low risk of dissociation of subunits. Native electroblotting after BN-PAGE and antibody detection reveals which minimal detergent/protein ratio is required to solubilize and detect the desired protein, and it might also indicate detergent or detergent/ protein-dependent variations of the running distance of the interesting protein in BN-PAGE. Since BN-PAGE separates according to molecular mass, such variations of running distance can provide first hints for a transition from dimeric to monomeric state, for association or dissociation of unknown protein components, and for dissociation of subunits from subcomplexes.

110

Hermann Schagger

A

B Triton X-100/prottln

DDM/protein OJ

1.6

{gig)

0.4

0.6

0.8

10

1.2

14

16

2.4

(gig)

Lmj%

DDM 1J

Comptex

DIgltonIn 10

4.

15

2.0

i

3.0

4.0

8.0

^

(9

^

hlilatVi I1IH2

- ^ tMor»

HI—

^: i i i i ^ i l

iiiia

ig)

— llMon

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5. General Scheme of Sample Preparation for BN-PAGE and Preparation of a Molecular Mass Standard from Bovine Heart Mitochondria Many special protocols for the sample preparation for BN-PAGE have been described: from isolated bovine and yeast mitochondria (Arnold et ah, 1998; Schagger, 1995a; Schagger and Pfeiffer, 2000; Schagger and von Jagow, 1991), from chromaffin granule membranes (Ludwig et al., 1998), from celllines (Schagger et ah, 1996), and from homogenized mammalian tissues (Schagger, 1996). Instead of using 500 mM aminocaproic acid, as described in these previous protocols, we now prefer the addition of 50 mM NaCl for facilitated protein solubilization by detergents as described in the following general scheme. Storage of biological membranes. Biological membranes are best suspended in carbohydrate solution (e.g., >250 mM sucrose or 400 mM sorbitol) or 10-15% glycerol before they are shock-frozen in liquid nitrogen and stored at — 80°C. This minimizes dissociation of multiprotein complexes. Ideally the suspension is buffered with imidazole/HCl, pH 7.0, and the salt concentration is below 50 mM sodium chloride. Avoid potassium or divalent cations, which can aggregate Coomassie dye required for BN-PAGE and thus may coprecipitate membrane proteins.

FIGURE 1

BN-PAGE after differential solubilization of bovine heart mitochondria using different detergents and varying detergent/protein ratios. Solubilization followed the general sample preparation protocol. A. A DDM/protein ratio of 0.6 g/g is sufficient to solubilize all OXPHOS complexes quantitatively. Using such a low detergent/protein ratio retains some complex V in a dimeric form, which is split into the monomeric form by adding more detergent. The apparent mass of complex IV is higher at a low DDM/protein ratio. This shift of apparent mass is too small for a shift from monomeric to dimeric state. We assume that this shift is due to binding of different amounts of lipid. B. Triton X-100 solubilizes complexes II and V almost completely, using a Triton/ protein ratio of 1.0 g/g, but in contrast to DDM solubilizes almost no complexes I, III, and IV. An explanation for this different behavior is the existence of a network of respiratory chain supercomplexes made up of complexes I, III, and IV with the indicated stoichiometries of the components. This network is easily dissociated by DDM but more stable in the presence of Triton (Schagger and Pfeiffer, 2000). C. Digitonin retains physiological protein-protein interactions even better than Triton X-100. Complete solubilization of all complexes is achieved using a digitonin/protein ratio of 2.0 g/g, and complex I is almost quantitatively associated with complex III. Complex IV is partially retained in the supercomplex IJilj^i, comprising one monomer of complex I, a dimer of complex III, and a complex IV-monomer.

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^General sample preparation and ^Mitochondrial marker protein preparation a^. Use aliquots of the biological membranes (50-200 jag of total protein for a 0.16 x 1 cm gel well). Concentrated protein suspensions (>10 mg/ml) can be used directly for solubilization. Diluted suspensions have to be pelleted by centrifugation. a^. Use sedimented bovine heart mitochondria (200 fig protein; isolated according to Smith, 1967) to prepare a standard for molecular mass calibration in the range from 100 to 1000 kDa. Alternatively use a selection of commercially available water-soluble proteins that have been shown to fit the calibration curve of Triton- or dodecylmaltosidesolubilized membrane proteins (Section III,B,3). Ferritin and catalase do not bind Coomassie dye, and fixation/staining of one copy of a duplicate gel is required for molecular mass assignment. b^. Add 40 fil of solubilization buffer (50 mM NaCl, 5 mM aminocaproic acid, 1 mM EDTA, 50 mM imidazole/HCl, pH 7.0) to 50-200 fig sedimented protein, add 1 fd PMSF (0.5 mM stock in DMSO), homogenize, and after some minutes solubilize by adding detergent from 10% stock solutions (DDM or Triton/protein ratio in the range from 0.4 to 3.0 g/g; digitonin/protein 0.8-6 g/g; see above). Solubilization is complete within several minutes. ^M Solubilize the pelleted bovine heart mitochondria as above by adding 5 fil of 10% dodecylmaltoside (DDM/protein ratio of 2.5 g/g). Note: This solubilization in the presence of 50 mM NaCl usually yields better results than previous protocols using 500 mM aminocaproic acid as a dielectric. However, the NaCl concentration should not exceed 50 mM. Higher salt concentrations can lead to extreme stacking of proteins (i.e., proteins concentrate to very sharp protein bands), but unlike the situation in SDSPAGE, the highly concentrated membrane proteins can aggregate. Protease inhibitors used are 5 mM aminocaproic acid, 1 mM EDTA, and PMSF. c^'^. Centrifuge for 30 min at 100,000 g if the molecular mass of the desired protein is not larger than 1000 kDa, or for 20 min at 20,000 g for giant multiprotein complexes with masses u p to 10,000 kDa to avoid sedimentation. d^. Collect the supernatant, add 5-15% glycerol or 200-500 mM aminocaproic acid to increase the density and to facilitate sample application. Shortly before sample application to BN-PAGE, add Coomassie dye from a 5% suspension in 500 mM aminocaproic acid, underlay the 20-40 fil sample to the cathode buffer, and start BN-PAGE (see step 8). d^. As above. Add 2.5 fil of the 5% Coomassie dye suspension to the supernatant of solubilized mitochondria.

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Note: The quantity of added Coomasie dye depends on the quantity of detergent used. Usually 1 /il of the 5% dye suspension is added when proteins are solubilized by addition of 2 /zl of a 10% detergent solution to adjust the optimal dye/detergent ratio of 1:4. Addition of dye is especially helpful for concentrated samples containing high protein, detergent, and lipid concentrations. Excess lipid-detergent micelles, which would impede protein resolution, receive a lot of negative charges when incorporating the anionic dye and are rapidly pulled to the running front. The mixture of a neutral detergent and an anionic dye mimics a mild anionic detergent. Starting from biological membranes this usually is not critical, because the lipids that were extracted from the membrane compensate for a potential denaturing effect. However, some complexes—for example, complexes of the mitochondrial import machinery— contain easily dissociating subunits and should be treated like prepurified membrane proteins described following.

6. Processing of Prepurified Membrane Proteins Isolation of membrane proteins and complexes in a highly pure state is often rather difficult, and only partial purification may be achieved by conventional isolation protocols. BN-PAGE is a valuable technique for final purification of these problematic proteins—for example, for immunization and production of antibodies or for protein sequencing (see Section III,A). Using prepurified protein seems easier than starting from biological membranes. However, there are more potential pitfalls that can be minimized by following three rules. As a general first rule, keep the ionic strength of the sample for BN-PAGE as low as possible, ideally 25-50 mM NaCl, to avoid extensive protein stacking that may lead to membrane protein aggregation. Only a few proteins need higher salt concentrations to stay in solution. If a protein, for example, is eluted from an anion exchanger by 300 mM NaCl, salt reduction by gel filtration or dialysis is absolutely required. Aminocaproic acid, a zwitterionic compound that was used instead of NaCl to assist solubilization of membranes by neutral detergents, does not migrate in the electrical field at neutral pH and therefore does not cause protein stacking. Unfortunately, it cannot replace NaCl during gel filtration or other chromatographic steps. Second rule: Avoid excess detergent for elution of the desired protein from a chromatographic column. The minimal detergent concentrations are 0.05% Triton X-100 and 0.1% dodecylmaltoside. Despite this precautionary measure, chromatographically prerepurified proteins and protein complexes usually are isolated in a partially delipidated state. The stabilizing effect of lipids is largely missing, and addition of the anionic Coomassie dye may facilitate dissociation of residual lipid and lead to dissociation of detergentlabile subunits.

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Third rule: Don't add Coomassie dye to the sample. Apply a noncolored sample or a sample slightly colored by addition of the dye Ponceau S to BN-PAGE (5% glycerol or 200-500 xnM aminocaproic acid might be added to increase the density and facilitate sample application). The source of the Coomassie dye, which, of course, is required for BN-PAGE, is cathode buffer B or B/10. We speculate that the beneficial effect of applying a noncolored sample results from a preseparation of the desired protein from the bulk of detergent before Coomassie dye from the cathode buffer reaches the protein. Proteins with an isoelectric point below p7 7 migrate to the anode, while the bulk of the detergent stays in the sample well. Coomassie dye from the cathode buffer initially is retarded by these detergent micelles, but it catches u p with the protein later when the protein has already entered the gel. 7. Maximal and Minimal Protein Loading The maximal protein load to a 1 x 0.16 cm sample well is around 200 /ig protein (e.g., 200 /xg of a chromatographically prepurified multiprotein complex or 200 fig protein extracted from biological membranes; total protein before solubilization is around 400 /ig). Using bacterial membrane extracts directly, the protein load should be reduced to 50-100 /ig, since the maximal load seems to depend on the amount of DNA present in the solubilized sample. DNA seems to occlude the pores of the sample gel, and proteins cannot enter the gel. Use of DNases does not help much. Protein entry is improved, but deep blue ragged artefact bands appear in the gel. The best way to remove DNA seems to be mild disruption of cells and differential centrifugation. The minimal protein load to BN-PAGE depends on the sensitivity of the protein detection method, which is around 0.1-1 jag for one single polypeptide using Coomassie staining, and on the ng scale, using silver-staining or antibody detection. Note: It is essential to use the same detergent/protein ratios for solubilization of minimal protein amounts as for normal or high load, and the final detergent concentration must be clearly above the critical micelle concentration (CMC) of the specific detergent. For the latter reason, it might be necessary to reduce the final volume—for example, to add only 5 jA instead of 40 fA solubilization buffer (see 5.b^). 8. Electrophoresis Conditions BN-PAGE usually is performed at 4-7°C, since broadening of bands was observed at room temperature. Perform BN-PAGE in a vertical apparatus, and use deep blue cathode buffer B. Begin and maintain electrophoresis at 100 V until the sample is within the sample gel. Then continue with voltage limited

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to 500 V and current limited to 15 mA. After about one-third of the total running distance, cathode buffer B can be removed and replaced by cathode buffer B/10. This optional buffer exchange improves detection of faint bands. It is especially recommended when electroelution or electroblotting of native proteins is desired. Running times are about 3-4 hr using analytical gels and about 4 - 6 hr using preparative gels. 9. Electroelution of Native Proteins Ideally a band of the desired protein should be detectable during BNPAGE. This is usually achieved by starting BN-PAGE using deeply blue cathode buffer B and switching to slightly blue cathode buffer B/10 after about one-third of the total running distance. With respect to the recovery of protein from the gel, it is essential to stop BN-PAGE as soon as the desired protein band is detected. At that point the protein still has some mobility and can therefore be efficiently extracted. The desired protein must not reach the normal running distance. Mobility and extraction efficiency drop considerably when the protein approaches the pore size limit of the acrylamide gradient gel. Extract native proteins at 4°C with the H-shaped electroelutor/concentrator used for SDS-denatured proteins (Chapter 4, Section IV,C,1). The same membranes (cutoff value of 2 kDa) are used here to seal the anodic and cathodic arms of the elutor chamber because of their high mechanical stability compared to normal dialysis membranes. a. Excise the blue bands of native proteins from the gel, and squeeze them through a syringe into the cathodic arm of the elutor chamber. b. Fill both arms of the chamber and the horizontal connecting tube with electrode buffer (25 mM Tricine, 3.75 mM imidazole, pH 7, and 5 mM aminocaproic acid is added for protease inhibition). c. Extract at 500 V for several hours. Limit the current to about 2 mA per elutor vessel to prevent damage in case a wrong buffer is used in error. Partially aggregated proteins collect as a thin blue layer on the anodic dialysis membrane. To minimize protein aggregation, such as for functional studies, reduce the voltage to 50-100 V. 10. Removal of Bound Coomassie Dye Dye removal by gel filtration. Removal of dye from chloroplast ATPsynthase CFoFi, from the hydrophilic CF^ part and from the hydrophobic CFQ part, was described by Poetsch et al. (2000). The electroeluates were subjected to FPLC gel filtration on Sephacryl S-100 HR (Pharmacia) using a low ionic strength buffer containing 25 mM Tricine, 7.5 mM Bistris (or 3.75 mM imidazole to allow for a Lowry protein determination), 20% glycerol, 0.02%

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Hermann Schagger

NaN3, 5 mM MgCl2, pH 7.0, and 8 mM of the zwitterionic detergent Chaps. It is essential to use a detergent that forms small micelles (about 6 kDa in the case of Chaps) and small mixed detergent/Coomassie dye micelles. A prerequisite for a separation of protein and detergent/Coomassie micelles is that the latter are sufficiently small to enter the pores of the gel filtration matrix in contrast to the larger proteins. Dye removal by binding to Bio-Beads. Dye removal from electroeluted CF^Fo by Bio-Beads was performed in the presence of dedecylmaltoside and soybean lecithin (Poetsch et ah, 2000). This technique is similar to the detergent removal protocol by Lacapere et al. (1997). 11. Electroblotting of Native Proteins Like native electroelution native electroblotting requires short runs of BN-PAGE for efficient transfer of proteins and final use of slightly blue cathode buffer B/10 to avoid a reduction of the protein-binding capacity of PVDF membranes by competing free Coomassie dye. Dont use nitrocellulose membranes, which bind Coomassie dye rather strongly and cannot be destained under the conditions described here. a. Wet PVDF membranes (e.g., Immobilon P^^) with methanol, and soak in electrode buffer (50 mM Tricine, 7.5 mM imidazole; the resulting pH is around 7). b. Place the gel and then the PVDF membrane on a 3 mm stack of papers soaked with electrode buffer. The lower electrode is the cathode in this arrangement. Finally, add another 3 mm stack of papers soaked with electrode buffer on top, mount the anode, and place a 5 kg load on top before transfer is started. c. Allow transfer to occur at 20 V for 1 hr. The proteins are then visible as blue-stained bands on PVDF. d. Reduce the background using 25% methanol, 10% acetic acid.

B. Native Second Dimension: BN-PAGE + Detergent After BN-PAGE in the first dimension, usually a denaturing resolution of the protein subunits by a second dimension SDS-PAGE follows (see C and Section III,B,1, Fig. 4). However, as described here, also a second native resolution can follow. A modified BN-PAGE in a second dimension was introduced for the dissociation of supercomplexes of the respiratory chain into the individual complexes I, III, and IV (Schagger and Pfeiffer, 2000). These supercomplexes contained up to 67 individual subunits, and it seemed advisable to dissociate these supercomplexes first before the protein subunits of the individual complexes finally were resolved by SDS-PAGE in a third dimension.

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All stock solutions for BN-PAGE can also be used for BN-PAGE in the second dimension. However, 0.02% dodecylmaltoside or 0.03% Triton X-100 is added to cathode buffer B. 1. Excise one lane from first BN-PAGE (0.16 mm gel), and place it on a glass plate with "ears" at the position where the sample gel is normally found. 2. Mount the side and bottom spacers, which should be slightly thinner (e.g., 0.14 mm), to avoid only sliding down of the gel strip, place the second glass plate on top, fix the assembly with clamps, and put it upright. 3. Cast a 5-20% acrylamide gradient gel (analogue to Table III) below the gel strip at 4°C, overlay with water, and let polymerize at room temperature. 4. Add a 3.5% acrylamide native gel mixture to embed the native gel strip, but don't cover it. 5. Perform BN-PAGE at 4°C with the voltage set to 500 V and the current limited to 15 mM, and use cathode buffer B with addition of 0.02% dodecylmaltoside or 0.03% Triton X-100, which dissociate supercomplexes in different ways (Fig. 2).

C. Second or Third Dimension: SDS-PAGE The preferred SDS-PAGE system for resolution of protein subunits following first or second BN-PAGE is Tricine-SDS-PAGE (Schagger and von Jagow, 1987; see also Chapter 4), because 2-3% lower acrylamide compared to the Laemmli tris-glycin system (Laemmli, 1970) can be used, protein bands are sharper, and electrotransfer of proteins to PVDF membranes is facilitated. 1. Protocol for the Analytical Scale a. Cut out 0.5 cm lanes from first or second BN-PAGE, place them on a glass plate with "ears" at the position where the sample gel is normally found, and soak with 1% SDS (±1% mercaptoethanol) for 1-2 hrs. If cleavage of disulfide is not essential, and if there is no extremely strong band in BN-PAGE, this incubation is not required. SDS from the cathode buffer is sufficient to dissociate moderate amounts of multiprotein complexes within the native gel strip. b. Position 0.7 mm spacers when BN-PAGE was performed with 1.6 mm gels, place the second glass plate on top, fix the assembly with clamps, remove mercaptoethanol carefully, since it inhibits efficiently the polymerization of acrylamide, and put the whole assembly upright. The 0.5 cm lanes are squeezed to a width of about 1 cm.

118

FIGURE 2 Two-dimensional native resolution of OXPHOS complexes from bovine heart mitochondria. First BN-PAGE used bovine heart mitochondria that were solubilized by using a digitonin/protein ratio of 4 g/g (see Fig. 1C). Besides the bands of complexes I-V, additional bands of dimeric complex V and two major supercomplexes containing a complex I-monomer and a complex 111-dimer (111112),or an additional complex IV (IlII121Vl),were resolved as schematically drawn on top. Identical lanes from BN-PAGE were used in (A) and (B) for second BN-PAGE using cathode buffer B with 0.02% dodecylmaltoside (A) or 0.03% Triton X-100 (B). Second, BN-PAGE with added DDM dissociated bovine supercomplex IlII121Vlcompletely into complexes I, 111, and IV (Fig. 2A), while addition of Triton retained complex I-IV interactions (Fig. 28). All single complexes that retain their masses after second BN-PAGE are found on a diagonal. 3D SDSPAGE of excised vertical lanes originating from I,II121V,-supercomplexesrevealed the characteristic polypeptide patterns of the single complexes (A) or of complex I associated with complex IV (IIIVl) in (B). It also revealed that the second BN-PAGE can remove some labile subunits partially or completely as indicated by boxes.

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c. Cast a separating gel below the native gel strip, usually a 10% or a 16.5% acrylamide Tricine-SDS-gel (Chapter 4, Table III), overlay with water, which may contact the native gel strip, and let polymerize. The gap between 10% gel and native gel strip should be about 3 mm, while the gap to a 16.5% gel usually is about 1.5 cm. After removal of water, the 16.5% gel is overlayed by a 1 cm 10% acrylamide Tricine SDS gel, which improves transition of proteins from the native to the SDS-gel, especially when a lot of large proteins are transferred. d. Overlay the native gel strip with water, and push the strip down to the 10% acrylamide gel by using a piece of plastic that is slightly thinner than the spacers used. Air bubbles below the native gel strip are efficiently removed this way. e. Remove water and fill the gaps to the left and right of the native gel strip by a 10% acrylamide native gel mixture (see Table III). This guarantees a homogenous field strength over the total width of the gel. Don't cover the native gel strip with this native gel mixture. f. Run the second- or third-dimension SDS-PAGE (gel dimensions 14 X 14 X 0.7 mm) at room temperature with the voltage set to 200 V and the current limited to 50 mA. When the current drops below 50 mA, the voltage can be set to 300 V with the current still limited to 50 mA. The running time is about 5-6 hrs. The voltage has to be considerably reduced when using thicker gels. 2. Preparative Approach for Protein Sequencing Quantities of protein subunits obtained by BN-PAGE and SDS-PAGE in the second dimension often are too low for protein sequencing, even when 3 mm BN-gels and 1.6 mm SDS-gels are used. Sufficient protein for sequencing can be obtained either by using preparative BN-gels, electroelution, and final SDS-PAGE or by following the more convenient way (Arnold et ah, 1998). a. Separate the desired protein complexes by using one or several preparative gels (3 mm) for BN-PAGE. b. Excise the visible band of the complex of interest, cut it into 4 - 6 pieces, and incubate with 1% SDS ( ± 1 % mercaptoethanol) for 1-2 hrs. c. Load the pieces as a stack of 4 - 6 pieces (maximal height 1.5 cm) onto a 1.6 mm Tricine-SDS gel, as described above. d. Following SDS electrophoresis, blot protein subunits onto PVDF membranes (Chapter 4). Intensely Coomassie blue-stained protein bands usually are directly accessible to aminoterminal protein sequencing.

III. APPLICATIONS BN-PAGE and two-dimensional electrophoresis (BN-PAGE/SDS-PAGE) are microscale techniques that are mostly used for analytical purposes. However, they can also be scaled u p to obtain highly pure protein for immunization and aminoterminal protein sequencing.

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A. Final Purification of Crude Membrane Proteins for Immunization and Protein Sequencing Protocols for isolation of membrane proteins often end up with partially purified proteins. BN-PAGE can be used in these cases as a final purification technique to obtain highly pure proteins for immunization and protein sequencing (Fig. 3). See Section 11,6 for special precautionary measures.

B. Separation of Native Membrane Protein Complexes from Solubilized Biological Membranes 1. Isolation of Multiprotein Complexes from Mitochondria and Chloroplasts: PDC, OGDC, OXPHOS, TIM, and TOM BN-PAGE was originally developed for isolation of the five membrane multiprotein complexes of the system of oxidative phosphorylation (OXPHOS complexes I-V) from bovine mitochondria (Schagger and von Jagow, 1991; Schagger et ah, 1994; Schagger, 1995a) and to study the development of cytochrome c oxidase isoforms during the perinatal period of the rat (Schagger et ah, 1995). It has been used also for isolation of OXPHOS complexes from yeast and plant mitochondria (Arnold et aL, 1998; Jansch et ah, 1996) and from chloroplasts (Burrows et aL, 1998; Kiigler et ah, 1997). An example for a wide range separation of proteins from bovine heart mitochondria by BN-PAGE and SDS-PAGE in the second dimension is given in Fig. 4. BN-PAGE has also been used for the analysis of the TOM and TIM protein complexes of the mitochondrial protein import machinery of yeast (Dekker et ah, 1998; Dietmeier et aL, 1997; Hill et aL, 1998; Kurz et aL, 1999; Ryan et aL, 1999; Van Wilpe et aL, 1999) and plants (Caliebe et aL, 1997; Jansch et aL, 1998). 2. Isolation of Complexes from Other Membranes: Neurotransmitter Receptors, Bacterial OXPHOS Complexes, Vacuolar ATPase The assembly of nicotinic acetylcholine receptors and the combinations of distinct subunits have been analyzed by BN-PAGE after prepurification of digitonin extracts from oozytes by Ni^'^-NTA agarose chromatography (Nicke et aL, 1999). The oligomeric states and the assembly of P2Xi, P2X3, and glycin receptors have been analyzed using a similar protocol (Griffon et aL, 1999; Nicke et aL, 1998). Resolution of OXPHOS complexes from bacterial membranes essentially followed the BN-PAGE protocol using mitochondrial membranes, except that the required detergent/protein ratio usually was lower by a factor of 2 (Schagger, unpublished). The vacuolar ATPase holo-enzyme and its membrane sector were isolated from chromaffin granula membranes (about 10 mg protein) by preparative BN-PAGE (Ludwig et aL, 1998). Using this micropreparative approach, the

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f\

B • Core Proteins l/ll Cytochrome b Cytochrome c. "Rieske'* ISP

13 4 kDa 9 5kl)a 9 2 kDa 1 0 ^ 7 2 kDa 6,4 kDa

FIGURE 3

BN-PAGE used as a final purification method. A. One ml (2 mg) of a partially purified complex III from bovine heart (see Chapter 8) was applied to a preparative gel for BN-PAGE (dimensions 14 x 14 x 0.28 cm; linear 5-13% acrylamide gradient). The sample contained 0.05% Triton, 100 mM NaCl, 10 mM Mops, and pH 7.2, and vvras applied w^ithout addition of Coomassie dye to avoid dissociation of labile subunits (see Section 11,6). The staining of native complex III occurred during the run by Coomassie dye originating from cathode buffer B. B. The purity of complex III before BN-PAGE (lane 2) and after electroelution (lane 1) is compared by Tricine-SDS-PAGE using a uniform 16.5% acrylamide gel (see Chapter 4). Reprinted from Schagger (1995a).

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Hermann Schagger A

BN-PAGE BN-PAGE o

3.0 J

Complex kDa

3.5 4Acryiamide

%

13

FIGURE 4

PDC 10000 h OGDC 2500 yoim 1400 1 1000 VMon . 700 yi 490 h IV II

200 130

30 kDa lOH

ly

i

CO Q

Separation of multiprotein complexes from bovine heart mitochondria. A. Multiprotein complexes from bovine heart mitochondria (100 /ig) were solubilized by dodecylmaltoside using a detergent/protein ratio of 2.4 g/g and separated by BN-PAGE using a linear acrylamide gradient gel from 3.0% to 16%, overlaid by a 2.5% sample gel. PDC: Pyruvate decarboxylase complex. OGDC: Oxoglutarate decarboxylase complex. B. The subunits of the native complexes resolved by BN-PAGE were separated by Tricine-SDS-PAGE using a 13% T, 3% C gel type. The identity of PDC and OGDC was confirmed by aminoterminal sequencing of the 68 kDa subunit E2 of PDC, and the 96 kDa and 48 kDa subunits El and E2, respectively, of OGDC.

protein subunits of the membrane sector of the mammalian enzyme could be characterized, and two membrane sector associated proteins were identified and sequenced. 3. Identification of Labile Protein-Protein Interactions: Dimeric ATPSynthase, Respiratory Chain Supercomplexes, TIM, and TOM Linked by Imported Preprotein Some recent publications demonstrate that BN-PAGE is also a valuable technique for the identification of labile physiological protein-protein interactions. A prerequisite to the detection of such interactions is the use of a mild detergent a n d / o r the use of a detergent/protein ratio that is just sufficient for complete solubilization (see Section II). F^Fo-ATP synthase from yeast mitochondria was recently shown to exist in a dimeric state. This dimeric state is retained when a low Triton X-100/protein ratio of 0.6 g / g is used, while increasing the Triton/protein ratio rapidly produces the monomeric form from the dimer (Arnold et al., 1998). The dimer is less sensitive to digitonin, but a gradual transition from dimeric to a partly monomeric state is observed when the digitonin/protein ratio is increased

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from 1.5 to 6 g / g (Schagger and Pfeiffer, 2000). The dimeric state is not produced by the Coomassie dye a n d / o r by the conditions of BN-PAGE, since (1) the monomer, but not the dimer, is detected by BN-PAGE when the genes for subunits ^ or g are deleted, and (2) the dimer contains three additional protein subunits compared to the monomer, two of which—namely e and g — show clear honiology to subunits of monon\eric bovine F I F Q - A T P synthase. Respiratory chain complexes in yeast and mammalian mitochondria form supercomplexes that again are detected preferentially by using digitonin for solubilization or by using low Triton/protein ratios (Schagger and Pfeiffer, 2000). Two supercomplexes (III2IV1 and III2IV2), each comprising a dimeric Complex III and one or two copies of Complex IV, exist in yeast. A different type of protein-protein interaction has been recently shown by using BN-PAGE. TOM, the translocase of the outer mitochondrial membrane, and TIM, the translocase of the inner mitochondrial membrane, have been trapped as a larger unity linked together by a preprotein just being imported (Dekker et al, 1997). The feature of BN-PAGE to retain pysiological protein-protein interactions seems to be an essential one, since there is only a limited number of other techniques that can help with the identification of labile protein-protein interactions: (1) coimmunoprecipitation, which in contrast to BN-PAGE requires a suitable antibody; (2) gel-filtration or ultracentrifugation, which has a much lower resolution compared to BN-PAGE; (3) the two-hybrid-methods, which helped to identify thousands of new protein-protein interactions but can produce false negative and false positive results; and (4) chemical crosslinking of proteins and isolation of the cross-linked product by affinity chromatography using an antibody to the known protein. Outlook: Many labile protein-protein interactions will not be detectable by BN-PAGE even when using close to physiological conditions, mild detergents, and low detergent/protein ratios. However, it seems possible that BN-PAGE might be useful as an efficient isolation technique following cross-linking. 4. Analysis of Isolated Complexes a. Determination of Molecular Mass and Oligomeric State of Native Membrane Proteins The separation of multiprotein complexes from solubilized bovine heart mitochondria suggested that BN-PAGE separates proteins according to their molecular masses (Fig. 4). This was confirmed by plotting the distance of migration in BN-PAGE versus Ig Mr of the mitochondrial marker proteins (Fig. 5 and Table IV). Surprisingly, a number of commercially available water-soluble proteins can alternatively be used for molecular mass caUbration, although they do not bind lipid and detergent, and some of them don't even bind Coomassie dye.

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Hermann Schagger IgMr

140 FIGURE 5

mm

Molecular mass calibration curve for BN-PAGE. (•) Multiprotein complexes from solubilized mitochondria (Table IV) and (D) commercially available proteins and protein complexes (Table V) were used.

The proteins listed in Table V have been tested to fit the calibration curve of the bovine heart marker proteins solubilized by Triton X-100 and dodecylmaltoside (Schagger et ah, 1994). Water-soluble proteins with a pi in the acidic range, some of vv^hich do not bind Coomassie dye (e.g.. Ferritin or Catalase), have a sufficient electrophoretic mobility to fit the calibration curve. However, those water-soluble proteins that do not bind Coomassie dye and additionally have a p7 more basic than 5.4 are not useful as molecular mass markers — for example. Carbonic anhydrase II from bovine erythroctes, p7 5.9. Some water-

TABLE IV

Molecular Mass Markers from Solubilized Bovine Heart Mitochondria (BHM) No.

Protein

Mass (kDa)

1 2 3 4 5 6 7

PDC, Pyruvate dehydrogenase complex OGDC, 2-Oxoglutaratedehydrogenase complex Complex I monomer Complex V monomer Complex III dimer Complex IV monomer Complex II monomer

10,000 2500 1000 ZOO'' 490 200 130

"The sum of masses of the subunits of complex V is around 600 kDa, assuming a 1:1 stoichiometry of all subunits. However, an apparent mass of 700 kDa for complex V fits the calibration curve best.

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TABLE V

Commercially Available Molecular Mass Markers No.

Protein

Source

pi

Mass (kDa)

Blue stain

1 2

Pyruvate dehydrogenase complex'' 2-Oxoglutarate dehydrogenase complex Ferritin (dimer) Ferritin (monomer) Catalase (dimer) Catalase (monomer)'' Lactate dehydrogenase (dimer) Lactate dehydrogenase (monomer) Hexokinase Conalbumin Albumin (dimer) Albumin j8-Lactoglobulin A and B

Porcine heart Porcine heart

nd nd

10,000 2500

Yes Yes

Horse spleen

4.2--4.5

Bovine liver

5.4

Rat muscle Yeast Chicken egg

8.6 4.5--4.8 5.9

Bovine serum Bovine milk

4.9 5.1--5.3

880 440 460 230 280 140 99 78 132 66 35

No No No No Yes Yes Weak Yes Yes Yes Weak

8d 8 9d 9 lOd 10 11 12 13d 13 14

" PDH can enter the gradient gel only when the lowest polymerizing acrylamide concentration is used for the sample gel (see Table II). ''Catalase is solubilized by dilution with 15% glycerol, 50 mM imidazole/HCl, pH 7, and freezing/thawing.

soluble proteins having a pi in the strongly basic range Trypsinogen from bovine pancreas (pJ 9.3) and Cytochrome c from horse heart (pJ 10.7) bind Coomassie dye and migrate to the anode, but their electrophoretic mobility is too low to fit the calibration curve. Others, like lactate dehydrogenase, p i 8.6, are useful as molecular mass markers. Mitochondrial OXPHOS complexes solubilized by digitonin have shorter running distances in BN-PAGE compared to Triton- or dodecylmaltosidesolubilized complexes (Fig. IC). We assume that this shift of apparent molecular masses is due to less delipidation by digitonin. The molecular mass determination of a digitonin-solubilized protein requires use of a standard curve from digitonin-solubilized mitochondrial marker proteins or appropriate corrections. BN-PAGE is a high-resolution technique compared to gel filtration and analytical ultracentrifugation, which can easily discern monomeric or dimeric states of a complex (Arnold et al, 1998; Schagger et al, 1994), or a 200 kDa mass difference between supercomplexes I1III2 (1.500 kDa) and IJII2IV1 (1.700 kDa; see Fig. 1). Also, dissociation of a single subunit representing, for example, 5% of the total mass of a complex can be detected. The electrophoretic technique is faster than gel filtration and analytical ultracentrifugation, and it requires only /ig amounts of a partially purified protein. BN-PAGE was also applied for control of purity and homogeneity of column

126

Hermann Schagger

eluates for the detection of subcomplexes generated by dissociation of subunits and to find out whether two proteins found in the same fraction are assembled to form a complex or exist as isolated proteins (Schagger et ah, 1994). h. Identification of OXPHOS-Deficiencies in Human Tissues and Cell-Lines: Diagnostics of Mitochondrial Encephalomyopathies The mitochondrial protein fraction of total cellular protein is as high as 5-10% in skeletal muscle or even higher in heart muscle. So it seemed possible to relinquish careful but low-yield isolation of patient mitochondria and to use directly a crude mitochondrial sediment obtained after breaking tissue specimens or cells (10 mg wet weight) by using a tightly fitting homogenizer and centrifugation at 10.000 g. This direct way was required because only 10-20 mg of skeletal muscle (wet weight) is obtained by needle biopsies. The detailed protocols for processing different tissues (Schagger, 1996) and cell lines (Schagger et ah, 1996) will not be repeated here. Two-dimensional electrophoresis (BN-PAGE/SDS-PAGE) has been successfully applied for the localization of OXPHOS deficiencies in mitochondrial encephalomyopathies (Bentlage et ah, 1995; Dionisi-Vici et al, 1997; Schagger et al, 1996) and other mitochondrial disorders (Kretzler et al., 1999; Schagger, 1995b; Schagger and Ohm, 1995). Usually such deficiencies are apparent from the loss of assembly of individual or multiple OXPHOS complexes. However, there are also rare cases where a specific complex is almost normally assembled, but the corresponding catalytic activity is significantly reduced (Pfeiffer et ah, manuscript in preparation). One example comparing the OXPHOS system from a MELAS patient (mitochondrial encephalomyopathy with /actic acidosis and stroke-like episodes) and a control is shown in Fig. 6. The MELAS patient carried a heteroplasmic A3243G mutation (60%) in a mitochondrial tRNA^^"" that affected the mitochondrial protein biosynthesis in general and especially Complexes I and IV. c. Miscellaneous Analyses: Catalytic Activities, Analysis after ID-Crystallization

Electron

Microscopic

Proteins and multiprotein complexes electroeluted after BN-PAGE usually retain their tertiary and quarternary structures. However, catalytic activity often is impeded or disturbed by binding of Coomassie dye to the protein or to the substrate, and special protocols (some examples are described below) are required to measure catalytic activities. Enzymatic measurements using non-Coomassie-stained proteins are clearly preferred when large-scale chromatographic purifications are possible.

CHAPTER 5

Blue Native Electrophoresis

^~ - I - - 4 - ^ Complex I

V

HI

BN-PAGE

127

• fV

75 (kDa)|

20

13

s

B

75

20

13

I I FIGURE 6

Two-dimensional resolution (BN-PAGE/SDS-PAGE) of the OXPHOS proteins from human skeletal muscle. OXPHOS complexes were extracted from 20 mg quadriceps muscle of (A) a control person and (B) of a MELAS patient, and separated by BN-PAGE in the first dimension. Resolution of the protein subunits was achieved by Tricine-SDS-PAGE using a 16.5% acrylamide gel. Silver-staining revealed an almost complete absence of complex I and a decreased amount of complex IV.

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Hermann Schagger

Catalytic activities of complexes J, IT, IV, and V quantified directly in BN gels: Zerbetto et al. (1997) and Jung et ah (2000) used histochemical staining of blue native gels to quantify the OXPHOS complexes from muscle and central nervous system, respectively, of patients suffering from mitochondrial encephalomyopathies. A histochemical assay for complex III is not yet available. Catalytic activities of proteins electroeluted from BN gels: Extracted complex III (bci complex) was tested for ubihydroquinoneicytochrome c reductase activity directly and also after addition of phospholipid (Schagger and von Jagow, 1991). Without added phospholipid the activity was low (30 sec~^ at 25°C); however, activity increased more than 10-fold within 15 min when egg yolk phospholipid dissolved in Triton X-100 (see Chapter 8) was added to a molar ratio of 5000 phospholipids per complex dimer. Catalytic activity further increased during overnight incubation and was then comparable to that of the best chromatographically prepared complex III (1000 sec"^). The rate of activation of electroeluted complex III was extremely slow compared to chromatographically delipidated complex III (Schagger and von Jagow, 1991), and 25 times more phospholipid was needed, presumably because protein-bound Coomassie dye must be removed first. Addition of detergent without phospholipid does not activate complex III, because bound phospholipid, which is essential for catalytic activity, is also removed. Extracted complex IV (cytochrome c oxidase) was tested for cyanidesensitive ferrocytochrome c oxidase activity, which increased linearly with addition of reduced cytochrome c to reach a maximal rate of 60 sec~^ at the highest feasible concentration of 60 /nM cytochrome c. (K^ and V^^^x of chromatographically purified complex IV are 20 fiM and 350 sec"^, respectively). This indicated a shift of the apparent K^ for cytochrome c, which is probably due to binding of the negatively charged Coomassie dye to the basic protein cytochrome c. ATP synthesis activity of chloroplast ATP-synthase (CFoFi) after different preparative steps was compared. CFoF^ electroeluted from BN gels showed about 50% of DCCD-sensitive ATP-synthesis activity compared to a noncolored CFQFI reference sample. This residual activity could not be restored after removal of bound Coomassie dye (Poetsch et ah, 2000). 2D crystallization and electron microscopy: Electroeluted CFQEI and its membrane sector CFQ have also been used successfully for 2D crystallization and electron microscopy, and CFQ electroeluates were used for analysis of the secondary structure by CD spectroscopy (Poetsch et ah, 2000). Binding of Coomassie dye did not disturb CD analyses and had no effect on the high a-helical content of CFQ.

CHAPTER 5

Blue Native Electrophoresis

129

REFERENCES Arnold, L, Pfeiffer, K., Neupert, W., Stuart, R. A., and Schagger, H. (1998). Yeast FiFo-ATP synthase exists as a dimer: Identification of three dimer specific subunits. EMBO ]. 17, 7170-7178. Bentlage, H., De Coo, R., Ter Laak, H., Sengers, R., Trijbels, F., Ruitenbeek, Schlote, W., Pfeiffer, K., Gencic, S., von Jagow, G., and Schagger, H. (1995). Human diseases with defects in oxidative phosphorylation: I. Decreased amounts of assembled oxidative phosphorylation complexes in mitochondrial encephalomyopathies. Eur. J. Biochem. 227, 909-915. Burrows, P. A., Sazanov, L. A., Svab, Z., Maliga, P., and Nixon, P. J. (1998). Identification of a functional respiratory complex in chloroplasts through analysis of tobacco mutants containing disrupted plastid N d h genes. EMBO J. 17, 868-876. Cahebe, A., Grimm, R., Kaiser, G., Lubeck, J., Soil, J., and Heins, L. (1997). The chloroplastic protein import machinery contains a Rieske-type iron cluster and a mononuclear iron-binding protein. EMBO J. 16, 7342-7350. Dekker, P. J. T., Martin, F., Maarse, A. C , Bomer, U., Miiller, H., Guiard, B., Meijer, M., Rassow, J., and Pfanner, N. (1997). The Tim core complex defines the number of mitochondrial translocation contact sites and can hold arrested preproteins in the absence of matrix Hsp70-Tim44. EMBO J. 16, 5408-5419. Dekker, P. J. T., Ryan, M. T., Brix, J., Miiller, H., Honlinger, A., and Pfanner, N. (1998). Preprotein translocase of the outer mitochondrial membrane: Molecular dissection and assembly of the general import pore complex. Mol. Cell. Biol. 18, 6515-6524. Dietmeier, K., Honlinger, A., Bomer, U., Dekker P. J. T., Eckerskorn, C , Lottspeich, F., Kiibrich, M., and Pfanner, N. (1997). Tom5 functionally links mitochondrial preprotein receptors to the general import pore. Nature 388,195-200. Dionisi-Vici, C , Ruitenbeek, W., Fariello, G., Bentlage, H., Wanders, R. J. A., Schagger, H., Piantadosi, C , Sabetta, G., and Bertini, E. (1997). New familial mitochondrial encephalopathy with macrocephaly, cardiomyopathy and complex I deficiency. Ann. Neurol. 42, 661-665. Griffon, N., Biittner, C , Nicke, A., Kuhse, J., Schmalzing, G., and Betz, G. (1999). Molecular determinants of glycine receptor subunit assembly. EMBO J. 18, 4711-4721. Hill, K., Model, K., Ryan, M. T., Dietmeier, K., Martin, F., Wagner, R., and Pfanner, N. (1998). Tom40 forms the hydrophilic channel of the mitochondrial import pore for preproteins. Nature 395, 516-521. Jansch, L., Kruft, V., Schmitz, U., and Braun, H. P. (1996). New insights into the composition, molecular mass and stoichiometry of the protein complexes of plant mitochondria. Plant ]. 9, 357-368. Jansch, L., Kruft, V., Schmitz, U. K., Braun, H. P. (1998). Unique composition of the preprotein translocase of the outer mitochondrial membrane from plants. /. Biol. Chem. 273,17251-17257. Jung, C , Higgins, C. M. J., and Xu, Z. (2000). Measuring the quantity and activity of mitochondrial electron transport chain complexes in tissues of the central nervous system using blue native polyacrylamide gel electrophoresis. Anal. Biochem. 286, 214-223. Kretzler, M., Haltia, A., Solin M.-L., Schagger, H., Taanman J.-W., Kritz, W., Kerjaschki, D., Schlodorff, D., and Holthofer, H. (1999). Altered gene expression and functions of mitochondria in human nephrotic syndrome. FASEB J. 13, 523-532. Kiigler, M., Jansch, L., Kruft, V., Schmitz, U. K., and Braun, H. P. (1997). Analysis of the chloroplast protein complexes by blue-native polyacrylamide gel electrophoresis. Photosynth. Res. 53,35-44. Kurz, M., Martin, H., Rassow, J., Pfanner, N., and Ryan, M. T. (1999). Biogenesis of Tim proteins of the mitochondrial carrier import pathway: Differential targeting mechanisms and crossing over with the main import pathway. Mol. Cell Biol. 10, 2461-2474. Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature Til, 680-685. Lacapere, J.-J., Stokes, D. L., Mosser, G., Ranck, J.-L., Leblanc, G., Rigoud, J.-L. (1997). Twodimensional crystal formation from solubilized membrane proteins using Bio-Beads to remove detergent. Ann. N.Y. Acad. Sci. 834, 9-18.

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Hermann Schagger Ludwig, J., Kerscher, S., Brandt, U., Pfeiffer, K., Getlawi, F., Apps, D. K., and Schagger, H. (1998). Identification and characterization of a novel 9.2 kDa membrane sector associated protein of vacuolar proton-ATPase from chromaffin granules. /. Biol. Chem. 273, 10939-10947. Nicke, A., Rettinger, J., Mutschler, E., and Schmalzing, G. (1999). Blue native PAGE as a useful method for the analysis of the assembly of distinct combinations of nicotinic acatylcholine receptor subunits. /. of Receptor and Signal Transduction Research 19, 493-507. Nicke, A., Baumert, H. G., Rettinger, J., Eichele, A., Lambrecht, G., Mutschler, E., and Schmalzing, G. (1998). P2Xi and P2X3 receptors form stable trimers: a novel structural motif of ligandgated ion channels. EMBO ]. 17, 3016-3028. Poetsch, A., Neff, D., Seelert, H., Schagger, H., and Dencher, N. A. (2000) Dye removal, catalytic activity and 2D crystallization of chloroplast H'^-ATP synthase purified by blue native electrophoresis. Biochim. Biophys. Acta 1466, 339-349. Ryan, M. T., Miiller, H., and Pfanner, N. (1999). Functional staging of ADP/ATP carrier translocation across the outer mitochondrial membrane. /. Biol. Chem. 274, 20619-20627. Schagger, H., and Von Jagow, G. (1987). Tricine-sodium dodecyl sulfate polyacrylamide gel electrophoresis for the separation of proteins in the range from 1-100 kDalton. Anal. Biochem. 166, 368-379. Schagger, H., and Von Jagow, G. (1991). Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal. Biochem. 199, 223-231. Schagger, H., Cramer, W. A., and Von Jagow, G. (1994). Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis. Anal. Biochem. 217, 220-230. Schagger, H., Noack, H., Halangk, W., Brandt, U., and von Jagow, G. (1995). Cytochrome c oxidase in developing rat heart: Enzymatic properties and aminoterminal sequences suggest identity of the fetal heart and the adult liver isoform. Eur. J. Biochem. 230, 235-241. Schagger, H. (1995a). Native electrophoresis for isolation of mitochondrial oxidative phosphorylation protein complexes. Methods Enzymol. 260,190-202. Schagger, H. (1995b). Quantification of oxidative phosphorylation enzymes after blue native electrophoresis and two-dimensional resolution: Normal complex I protein amounts in Parkinson's disease conflict with reduced catalytic activities. Electrophoresis 16, 763-770. Schagger, H., and Ohm, T. (1995). Hunian diseases with defects in oxidative phosphorylation: II. FIFQ ATP-synthase defects in Alzheimer's disease revealed by blue native polyacrylamide gel electrophoresis. Eur. }. Biochem. 227, 916-921. Schagger, H. (1996). Electrophoretic techniques for isolation and quantitation of oxidative phosphorylation complexes from human tissues. Methods Enzymol. 264, 555-566. Schagger, H., Bentlage, H., Ruitenbeek, W., Pfeiffer, K., Rotter, S., Rother, C , Bottcher-Purkl, A., and Lodemann, E. (1996). Electrophoretic separation of multiprotein complexes from blood platelets and cell lines: Technique for the analysis of diseases with defects in oxidative phosphorylation. Electrophoresis 17, 709-714. Schagger, H. (2000) Blue native gels to isolate protein complexes from mitochondria. In "Methods in Cell Biology." In press. Schagger, H., and Pfeiffer, K. (2000). Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO ]. 19, 1777-1783. Smith, A. L. (1967). Preparation, properties, and conditions for assay of mitochondria: Slaughterhouse material, small scale. Methods Enzymol. 10, 81-86. Van Wilpe, S., Ryan, M. T., Hill, K., Maarse, A. C , Meisinger, C , Brix, J., Dekker, P. J. T., Mocko, M., Wagner, R., Meijer, M., Guiard, B., Honlinger, A., and Pfanner, N. (1999). Tom22 is a multifunctional organizer of the mitochondrial preprotein translocase. Nature 401, 485-489. Zerbetto, E., Vergani, L., and Dabbeni-Sala, F. (1997). Quantification of muscle mitochondrial oxidative phosphorylation enzymes via histochemical staining of blue native polyacrylamide gels. Electrophoresis 18, 2059-2064.

6

Preparative Isoelectric Focusing IRENE TSIROGIANNI AND GEORGIOS TSIOTIS Division of Biochemistry Department of Chemistry University of Crete GR-71409 Heraklion, Greece

I. INTRODUCTION The net charge of an amphoteric molecule is determined by the pH of its local environment. In the case of protein molecules this is the sum of all positive and negative charges of the ionizable amino acid side chains and prosthetic groups. For every protein there is a specific p H at which the net charge is zero. This isoelectric pH value, termed pJ, is a characteristic physicochemical property of every protein. The amphoteric nature of proteins can be used as the basis for isolectric focusing (lEF). In this method proteins are separated according to their pJ in a pH gradient gel. Stable, linear pH gradients are the keys to successful lEF.

Membrane Protein Purification and Crystallization Ije: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

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Irene Tsirogianni and Georgios Tsiotis

The pH gradient is formed by mixtures of compounds called ampholytes. Ampholytes are mixtures of polyamino-polycarboxylated compounds. They are small (about 300-1000 Da in size), multicharged, organic buffer molecules with closely spaced pi values and high conductivity. When a voltage is applied across a solution of ampholytes, they partition into a smooth pH gradient, increasing monotonically from the anode to the cathode (Laas, 1989). lEF is a high-resolution technique that can resolve proteins differing in pi by less that 0.05 pH units. The technique is rapid and nondenaturating and can be run preparatively as well as analytically. Preparative fractionations on the scale from hundreds of milligrams to grams of proteins, with recoveries of greater than 90%, are possible. Purification by factors of 10 to 100 in preparative applications place lEF intermediate between ion-exchange and ligand-binding chromatography. lEF, is well suited for use at any stage of a preparative scheme, and it is particularly effective in the early stages of purification. Commercially available analytical and preparative scale equipment can also be used for purification of membrane proteins in the presence of nonionic detergents. Ionic detergents, such as SDS, LDAO, or Sulfobetains, are not compatible with lEF. Detergent concentrations slightly above the critical micelle concentration (CMC) usually are sufficient to keep proteins in a solubilized state. Electrofocusing differs from conventional electrophoresis by using pH gradient gels. Either the pH gradient within the gel preexists if ampholytes covalently bound to acrylamide are used for casting gels (ready-to-use gels are available, e.g., from Pharmacia or Serva), or the pH gradient is formed during electrofocusing if mobile carrier ampholytes are used. This chapter describes isoelectric focusing using mobile ampholytes. During the experiment the carrier ampholytes arrange according to their isoelectric points, and a pH gradient develops. Proteins behave similarly; migration of a protein will cease when the protein approaches a pH zone corresponding to its p7 (Fig. 1). Any protein with a definite pi finally will be focused to a sharp band, especially when high electric fields are applied to minimize diffusion. In addition to the positive effect on resolution, high electric fields also result in shortened run times, but there is a considerable heat production. Because bands usually are sharper than in conventional electrophoresis, resolution is often higher, and smaller amounts of protein can be detected. Analytical electrofocusing, using a wide pH range gel, is an essential first step before working on a preparative scale. Knowing approximate p7 values of the proteins of interest then allows the investigator to choose a narrow pH range for optimal resolution in preparative gels. Additional information will be revealed about the protein stability near their pi because some proteins precipitate at their pi.

CHAPTER 6 Preparative Isoelectric Focusing

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Illustration of the focusing effect. A protein with pi 6 will be negatively charged (— 1 net charge units) when it is applied to a pH gradient gel at the pH 7 position; therefore, it will migrate to the anode in an electric field. The same protein will be positively charged (+ 2 net charge units) when applied to the gel at the pH 4 position, and thus will migrate to the cathode. On application of an electric field the two parts of the same protein sample will focus at the pH 6 position of the gel, where the intrinsic protein charge becomes zero.

11. TECHNIQUES A. Instruments and Accessories All electrofocusing equipment is from Pharmacia. Multiphor unit II Power supply (ECPS 3000V/150A) Thermostatic circulator GelBond PAG film Electrofocusing kit for analytical electrofocusing Electrofocusing kit for preparative electrofocusing Granulated gel (Ultrodex) Electrofocusing strips

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B. Analytical Electrofocusing Analytical electrofocusing in thin acrylamide gels is used to analyze purity and homogeneity, and for determination of isoelectric points. A critical step for selection of optimal separation conditions in preparative experiments is determination of the pH stability and the approximate p i of the protein of interest.

1. Preparation of Gels a. Prepare stock solutions as in Table I. Caution: Acrylamide is very toxic! Wear gloves whenever handling solid acrylamide or its solutions, b. Clean glass plates with detergent and ethanol. c. Prepare the gel mold by application of a "slot former" strip to the glass plate equipped with a silicone rubber U-frame. An alternative method for applying samples to thin polyacrylamide gels is to place filter paper strips soaked with sample directly on the gel surface. d. Coat another glass plate with Repel-silane (Pharmacia, Sweden), and dry with a paper tissue. e. Cut a GelBond PAG film to form the gel support foil. f. Add a few drops of water to the glass plate, and carefully lay the sheet of GelBond PAG film on top (hydrophobic side down). Roll the support foil on the glass plate to remove air bubbles and to ensure good contact with the glass plate. g. Clamp together the glass plate and the GelBond PAG film with the slot former inside. h. Mix solutions for the gel (25 ml) as in Table II. Degas for 5 min before addition of TEMED and ammonium persulfate. i. Cast the gel using a pipette. Polymerization is complete within 60 min. j. Remove clamps and remove the slot former, k. Switch on the thermostatic circulator 30 min before starting the experiment and set the temperature to 10°C. 1. Add some drops of kerosene to the template and position the gel.

TABLE I

Stock Solutions Acrylamide: N,N"-methylenebisacrylamide Ammonium persulfate Ampholine carrier ampholytes Cathode solution Anode solution

29.1%: 0.9% (w/v) 10% (w/v) (Pharmacia or Serva) 20 mM arginine, 5 mM lysine 25 mM aspartic acid, 25 mM glutamic acid, 2 M ethylenediamine

135

CHAPTER 6 Preparative Isoelectric Focusing

TABLE II

Gel Preparation Acrylamide, bisacrylamide stock solution Ampholine 3.5-10 Water Nonionic detergent at a concentration above the CMC Ammonium persulfate N,N,N',N'-Tetramethylethylenediamine (TEMED) Total volume

3 ml 1.8 ml 20.1 ml variable 90/il lO/il

25 ml

m. Prepare and apply strips for electrode solutions. Remove excess solution with paper tissue, n. Prerun the gel as indicated below to establish the pH gradient. Prerun conditions Power setting 4W Voltage setting max Current setting max Time 15 min 2. Electrofocusing a. Centrifuge the sample for 10 min at 12,000 g before applying to the gel, to avoid streaking caused by aggregates. b. If necessary, remove salt by diafiltration or by gel filtration (high salt concentration reduces the quality of the protein zones). If diafiltration is used, dilute the sample 10-fold with water or low salt buffer and concentrate the sample by ultrafiltration. For gel filtration use PD-10 columns (Pharmacia) if the sample volume is less than 2.5 ml. For larger sample volumes, use self-packed Sephadex G-25 columns. c. Apply 1-5 jUg of the protein of interest in a volume of 5-10 ^l. In the analysis of photosystem I, the sample contains a low concetration of detergent (0.05% C12-M) and 50 mM potassium phosphate (see Section III). Simultaneously apply pJ marker proteins, such as Serva IFF Marker 3-10, usually at a protein load of about 0.3 jug per protein band. d. Carefully layer the sample on the gel using a pipettor. e. Run the gel under the conditions indicated below. Running conditions Power setting 6W Current setting max Voltage setting max

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TABLE III

Solutions for Fixing and Staining Fixing solution Staining solution Destaining solution

20% trichloroacetic acid Serva Blue W (100 mg in 250 ml water) 0.2% trichloroacetic acid

f. The focusing time depends on the mobiUty of the protein and on the pH range chosen. For most proteins, 60-90 min is sufficient when a wide pH range (e.g., pH 3-10) is used. Narrow pH ranges with proteins close to their p7 values require longer focusing times. 3. Staining of Proteins a. Prepare solutions as in Table III. Note: Although staining with Serva Blue W is generally used in our laboratory, other methods are also available, including Coomassie blue R 250 and G 250 staining (Righetti and Chillemi, 1978) and silver staining for increased detection sensitivity (Merril et ah, 1981). b. Immediately after electrofocusing immerse the gel in fixing solution for 30 min. c. Wash the gel in destaining solution for a few minutes. d. Stain for 10 min. e. Destain for 5 min, and repeat the destaining step at least twice. f. Wash the gel with water. 4. Determination of Isoelectric Points Determine the apparent p7 values by comparison with a standard protein mixture with known p i values run on the same gel. One may use, for example, Serva lEF Marker 3-10, containing proteins in the range p7 3.5-10.5. In the case of colored proteins, determine the pJ by measuring the pH directly within the focused and colored protein zone, using a surface glass pH electrode (e.g., from Ingold).

C. Preparative Flat-Bed Electrofocusing The use of a stabilizing matrix is essential for electrofocusing. Polyacrylamide seems to be optimal for analytical electrofocusing. Ultrodex (Pharmacia), a dextran matrix, is an excellent stabilizing medium for preparative electrofocusing.

CHAPTER 6 Preparative Isoelectric Focusing

137

1. Gel Preparation a. Mix 110 ml of a 2% ampholyte solution and add the appropriate detergent. Note: Concentration of ampholytes below 1% (w/v) often result in unstable pH gradients. Ampholytes at concentrations above 3% (w/v) are difficult to remove from gels and can interfere with protein staining. b. Set 10 ml aside to soak six electrofocusing strips. Mount a stack of three strips at both ends of the tray (these stacks serve as a support for the electrode strips) and note the weight of the tray plus electrofocusing strips. c. Add 4 g of Ultrodex to the remaining 100 ml of ampholyte solution. Homogenize by gently stirring and pour the suspension into the tray. Weigh the tray again and note the weight. d. Dry at 50°C and check the degree of evaporation occasionally. e. Stop evaporation when the evaporation limit is reached (e.g., 35%; see product literature from the supplier). Note: The water content of the gel is crucial. If the water content is too high, it is not possible to use the paper print technique (Winter et ah, 1980) for visualization of the focused proteins; if too low, the gel bed may crack. 2. Sample Application The sample can be applied before or after establishing the pH gradient and at any position within the gel. Our approach is to apply the sample as a narrow zone near the cathode. The maximum volume applied is 5 ml per 100 ml of the initial gel slurry. According to Radola (1974) the loading capacity for a single protein is 2-4 mg/ml of gel bed when narrow pH ranges are used. a. Prepare 5 ml of sample solution, using the remainder of the ampholyte solution. [Note: High salt content of a sample may impair the development of straight protein bands (see Section II,B).] b. Form a trough by scraping away gel at the point of application; mix the removed gel with the sample. c. Pour the sample back into the trough and allow to settle for a few minutes. d. Run the gel for 14-16 hr at 8 W constant power, 8°C; these conditions work well for 100 ml of initial slurry and a narrow pH gradient (2 pH units). The optimum time for electrofocusing depends on the sample pi and on the pH range used. 3. Recovery of Separated Proteins Proteins containing chromophores are focused to easily detectable colored bands:

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a. Measure the pH directly in the gel using a small pH electrode. b. Scrape out the focused bands and mix with an appropriate elution buffer (see Section III). Note: After proteins are eluted from the matrix by diffusion, carrier ampholytes can be removed by dialysis, gel filtration, and anion-exchange chromatography (see Section III). For noncolored proteins, use the following paper print technique (Radola, 1974) as described by Winter et al (1980). a. b. c. d.

Cut a piece of filter paper so that it will completely cover the gel bed. Roll the dry filter paper onto the bed surface and leave for 2 min. Remove the print and dry at 110-120°C for 10-15 min. Wash the print for 15 min in 10% trichloroacetic acid; repeat twice more with fresh trichloroacetic acid. e. Stain with 0.2% Coomassie blue R 250 in 50% methanol, 10% acetic acid. f. Destain with frequent changes at early times in 50% methanol, 10% acetic acid, until the background color disappears.

III. APPLICATION: PURIFICATION OF PHOTOSYSTEM I The technique of isoelectric focusing is illustrated by the purification of photosystem I (Tsiotis et ah, 1993; Fig. 2). Photosynthetic membranes from Synechococcus were solubilized by using the nonionic detergent dodecyl-jSmaltoside (C12-M) at 1.4%. The chlorophyll concentration was 1.2 mg/ml (13.2 mg total protein/ml), and the C12-M/chlorophyll ratio was 12:1. PSI and PSII from the membrane extract were purified by a sucrose density gradient centrifugation in the presence of detergent (C12-M), which resulted in separation of two green bands. The upper band was a mixture of photosystems I and II (PSI and PSII), whereas the lower band contained predominantly photosystem I in trimeric form. The lower band was used for further purification, and an approximate pJ around 5 was determined by analytical electrofocusing using a wide pH range (Fig. 3). For optimal resolution in preparative electrofocusing, a narrow pH 4-6 range of ampholytes (Servalyte 4-6) is chosen, and 0.05% C12-M is added to the ampholine solution. PSI is now found predominantly at the pH 4.6-4.8 position; occasionally a second green band is located at pH 5.0-5.2, and a third band, containing carotenoids, is observed close to the anode. The pH 4.6-4.8 band is then eluted by diffusion using a 10-fold volume of 20 nriM MES, pH 6.5,10 mM MgCl2, and 0.05% C12-M. The ampholytes are removed by a combination of anion-exchange chromatography and gel filtration. The eluate is loaded onto a Q-Sepharose fast

139

CHAPTER 6 Preparative Isoelectric Focusing Extraction of membrane proteins by Cj2-M

I

Sucrose density gradient centrifugation (+ Cj2-M)

Upper green band (PSI/PSII)

Trimeric PSI \ Isoelectric focusing

I I

Anion exchange column

Purified PSI FIGURE 2

Flow diagram for the purification of photosystem I.

flow column (Pharmacia) equilibrated with the above buffer. After washing with 3 volumes of the same buffer, PSI is eluted by the wash buffer plus an additional 200 mM MgS04. The eluate is then concentrated 10-fold by ultrafiltration using Centricon microconcentrators (Amicon; cutoff 30 kDa) and low-speed centrifugation.

IV. CONCLUSIONS Previous isolation of PSI by anion-exchange chromatography (Mono Q, Pharmacia) and gel filtration led to apparently homogeneous preparations; however, crystallization attempts were not successful. Preparative isoelectric focusing allows separation of two forms of PSI with different isoelectric points. A homogeneous trimeric PSI complex was isolated in large quantity (5-10 mg) and in a state of purity that permitted crystallization (Tsiotis et ah, 1993). Perhaps the removal of a minor form of PSI with slightly differing pi may be the key factor that allowed for crystallization.

140

Irene Tsirogianni and Georgios Tsiotis pH - 3

- 6

IliHiilllii

IIIBIBlll

FIGURE 3

Analytical isoelectric focusing of photosystem I (lower green band after sucrose gradient centrifugation). A wide p H range of ampholytes (pH 3-10) was used with the following conditions: prerun, 15 min, at 4 W; electrofocusing, 60 min, at 6 W; temperature, 6°C (see text).

Another advantage of preparative electrofocusing is the free choice of buffer and detergent during elution from Ultrodex. Buffer salt concentrations and pH are critical factors in protein crystal growth (McPherson, 1990). The most critical parameter for crystallization of membrane proteins, however, is the choice of appropriate detergent (Michel, 1990). Crystals of PSI from Synechococcus FCC 7002 were obtained in three different buffers (potassium phosphate, Tris, MES) with the following detergents: (i) dodecyl-jS-maltoside; (ii) decyl-jS-maltoside; (iii) mixture of dodecyljS-maltoside and ocytl-j8-glucoside; and (iv) mixture of dodecyl-jS-maltoside and decyl-j8-maltoside. The detergent concentrations used were below the CMC (Tsiotis, 1991). Other examples of successful crystallization after preparative electrofocusing include the following: PSI complexes from Plactonema boryanum FCC

CHAPTER 6 Preparative Isoelectric Focusing

141

pH

FIGURE 4

Preparative electrofocusing of photosystem I. A 100 ml bed of Ultrodex was used. Electrofocusing was done for 16 hr at 8 W, 6°C.

73110, Anabena variabilis PCC 7937, and Anacystis nidulans (G. Tsiotis and H. Michel, unpublished); fumarate reductase from Wolinella succinogenes (Lancaster et ah, 1999; see also Chapter 13), and bacteriorhodopsin from Halobacterium halobium (Schertel, 1989).

REFERENCES Laas, T. (1989). Isoelectric focusing in gels. In 'Trotein Purification. Principles, High Resolution methods and Applications'' (J. C. Janson and L. Ryden, eds.), pp. 376-403. VCH, Weinheim. Lancaster, C. R. D., Kroeger, A., Auer, M., and Michel, H. (1999). Structure of fumarate reductase from Wolinella succionogenes at 2.2 A resolution. Nature 402, 377-385. McPherson, A. (1990). Current approaches to macromolecular crystallization. Eur. J. Biochem. 189, 1-23. Merril, C , Goldmann, D., Sedman, S. A., and Michael, E. (1981). Ultrasensitive stain for proteins in polyacrylamide gels shows regional variation in cerebrospinal fluid proteins. Science 111, 1347-1348. Michel, H. (1990). Crystallizing membrane proteins: Experiments on different systems. In ''Crystallization of Membrane Proteins" (H. Michel, ed.), pp. 73-78. CRC Press, Boca Raton, Florida. Radola, B. J. (1974). Isoelectric focusing in layers of granulated gels. Biochem. Biophys. Acta 386,181 -195. Righetti, P. G., and Chillemi, F. (1978). Isoelectric focusing of peptides. /. Chromatogr. 156,243-251. Schertel, G. (1989). "Kristallisation von Bacteriorhodopsin. Charakterisierung des M-Intermediates im Kristall." Ph.D. Thesis. Universitat Miinchen.

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Irene Tsirogianni and Georgios Tsiotis Tsiotis, G. (1991). "Isolierung und Charakterisierung der Chlorophyll a-haltigen Proteinkomplexe aus einer Phycobilisomen-freien Mutante des Cyanobakteriums Synechococcus PCC 7002. Kristallisation des Photosystems I." Ph.D. Thesis, Universitat Frankfurt. Tsiotis, G., Nitschke, W., Haase, W., and Michel, H. (1993). Purification and crystallization of photosystem I complex from a phycobilisome-less mutant of the cyanobacterium Synechococcus PCC 7002. Photosynth. Res. 35, 285-297. Winter, A., Perlmutter, H., and Davies, H. (1980). Preparative flat-bed electrofocusing in a granulated gel with the LKB 2117 Multiphor. LKB. Application Note 198.

7

Membrane Protein Crystallization CAROLA HUNTE AND HARTMUT MICHEL Dept. Molekulare Membranbiologie Max-Planck-Institut fur Biophysik 60528 Frankfurt am Main, Germany

I. INTRODUCTION Proteins fulfill many functions in the cell, based on a large variety of specific architectures. Therefore, for the understanding of the molecular mechanism of a protein, the respective high-resolution structure provides crucial information. In addition, membrane proteins are the target for a majority of drugs, and structures can be used for rational design and development of the compounds. The most widely used method for determination of protein structures at high, near atomic resolution is x-ray crystallography that requires well-ordered three-dimensional crystals grown from protein solution. Whereas several thousand structures of soluble proteins have already been determined by x-ray crystallography, fewer than 50 integral

Membrane Protein Purification and Crystallization 2/e: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

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Carola Hunte and Hartmut Michel

membrane protein structures became known since 1985, when the first membrane protein structure of the bacterial photosynthetic reaction center was presented (Deisenhofer et ah, 1985). This is in contrast to the high proportion of integral membrane proteins predicted from a genome analysis of eubacterial, archaean, and eukaryotic organisms, in which 20 to 30% of the open reading frames encode integral membrane proteins (Wallin and von Heijne, 1998). 3D crystallization of membrane proteins for high-resolution structural studies is challenging due to the amphipathic surface of the molecules. Embedded in the membrane bilayer, the contact sites of the protein with the acyl chains of the phospholipids are hydrophobic, whereas the polar surfaces are exposed to the polar head groups of the lipids and to the aqueous phases. Membrane proteins can be solubilized from the lipid bilayer in their native conformation by the use of detergents. The latter are amphiphilic molecules consisting of a polar or charged head group and a hydrophobic tail. At low concentration, detergent molecules are present in aqueous solution as monomers, and they form a monolayer at the water-air interface. However, above a certain concentration of the detergent, which is called critical micelle concentration (CMC), self-association occurs and micelles are formed. In these spherical aggregates the hydrophobic tails of the detergent molecules are oriented toward the micelle interior, and the hydrophilic head groups are pointing outward. Membrane proteins are soluble in micellar detergent solutions and form protein-detergent complexes. A detergent micelle is covering the hydrophobic surface of the membrane protein in a belt-like manner, while the polar detergent head-groups are facing the aqueous phase (for review, see Garavito and Ferguson-Miller, 2001; Zulauf, 1991). Membrane proteins can be crystallized within a phospholipid bilayer, resulting in two-dimensional crystals, which are used for structure determination by electron microscopy (for review, see Kiihlbrandt, 1992; Walz and Grigorieff, 1998; and see Chapter 15). Several structures of membrane proteins at medium resolution ('^5-10 A) have already been obtained. Structure determination at near atomic resolution by electron microscopy is possible (Henderson et al, 1990; Kiihlbrandt et al., 1994; Murata et «/., 2000), but it is still cumbersome. The amphipathic surface of membrane proteins allows for two ways to form 3D crystals, a necessity for x-ray crystallographic analysis (Michel, 1983; see Fig. 1). The so-called type I crystals are in principal ordered stacks of 2D crystals, which contain the ordered protein molecules within the membrane plane. Furthermore, type II 3D crystals can be formed by membrane proteins integrated in detergent micelles similar to soluble proteins. In these crystals contacts are made by the polar surfaces of the protein, which protrude from the detergent micelle. Here, crystallization conditions are very similar to those for soluble proteins. Mixed type I and II crystals are possible, although most membrane protein crystals belong to type II. Obviously, the detergent micelle

145

CHAPTER 7 Membrane Protein Crystallization

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Basic types of three-dimensional membrane protein crystals. Type I crystals consist of ordered stacks of two-dimensional crystals, which are present in a bilayer. In the detergent solubilized state the hydrophobic surface of membrane proteins is covered by a micelle. This membrane protein-detergent complexes form so-called type II three-dimensional crystals. Stable crystal contacts are made between the hydrophilic surfaces of the protein. In a modification of type II (type II b), tightly and specifically bound antibody fragments add a hydrophilic domain to the complex. This surface expansion provides additional sites for crystal contacts and space for the detergent micelle.

requires space in the crystal lattice, and it cannot contribute to rigid crystal contacts. Thus, proteins w^ith small hydrophilic domains are especially difficult to crystallize, as for example, many transporters or channels that consist mainly of transmembrane helices w^ith very short solvent exposed loops. A very promising approach to obtain well-ordered type II crystals is to enlarge the polar surface of the membrane protein by the attachment of polar domains of specifically binding antibody fragments (Ostermeier et ah, 1995). Up to now.

146

Carola Hunte and Hartmut Michel

nearly 30 membrane protein structures have been determined by x-ray crystallography, among them 12 structures of unrelated polytopic membrane proteins from the inner membranes of bacteria and mitochrondria, as well as from eukaryotic membranes. A list of structures and crystallization conditions of the different protein types is given in Tables I and XL Since the success of crystallization cannot be foreseen, it is advisable to pursue 2D and 3D crystallization trials in parallel. 2D crystallization trials need less protein, and it might be faster to obtain crystals. However, once good 3D crystals are available, fast structure determination at high-resolution by x-ray crystallography is more likely. There are several bottlenecks in the crystallization of membrane proteins, which will be discussed in the following sections.

11. BOTTLENECKS IN MEMBRANE PROTEIN CRYSTALLIZATION A. Starting Material The first problem encountered is the availability of sufficient amount of starting material, since 3D crystallization consumes at least 10 mg of pure protein per trial. Nearly all membrane protein structures that have been determined are of proteins that occur in high abundance in their natural environment, such as proteins of the respiratory chain or of photosynthetic membranes (Table I). Often these proteins are pigmented. Another group of high abundance is that of eubacterial outer membrane proteins. The latter contain mainly jS-strands in their membrane-spanning section, and they are of extraordinary stability. However, most membrane proteins are present in the cell only in low amounts. Therefore suitable overexpression systems for membrane proteins are needed. Insertion of the protein into a membrane limits the production yield, unless one chooses production via inclusion body formation and refolding. The latter has been very successfully applied for the unusually stable jS-barrel membrane proteins from the outer bacterial membranes (Pautsch et ah, 1999; see Schulz, 2000). Progress has been made especially in the development of overexpression systems for membrane proteins from bacterial sources. This resulted, for example, in the structure determination of the mechanosensitive ion channel from Mycobacterium tuberculosis that was produced in Escherichia coli (Chang et al, 1998). Further development of high-level overexpression system for production of recombinant membrane proteins from eukaryotic source is crucial for the successful exploration of their structures. Eukaryotic membrane proteins pose a larger problem for overexpression, since their signal sequences and targeting are highly differentiated, and posttranslational modifications occur frequently (see Chapter 3). Recombinant techniques for the production of proteins have clear advantages. Proteins can be engineered to allow easy purification by addition of affinity tags. In addition, proteins can be modified to provide a

CHAPTER 7 Membrane Protein Crystallization

147

homogenous protein source—for example, by removal of glycosylation sites. Furthermore, with the growing number of available genome sequences, the recombinant system allows an additional strategy, namely to clone, express, and purify the homologous proteins from several organisms and to test which protein is suitable for crystallization. This strategy was successfully used for structure determination of the mechanosensitive channel (Chang et ah, 1998), for which MscL homologs from nine prokaryotic species were subjected to crystallization trials. Homologous proteins from different species might not only feature differences in essential criteria like production yield or protein stability, but small variations in surface properties might cause the crystallization success, since crystal contacts are made by the protein surface.

B. Protein Purification and Characterization Starting from a good natural or recombinant protein source, the purification procedure has to ensure pure and homogenous protein preparations in sufficient yield. In general, the methods for protein purification are the same as those applied for soluble proteins except that the proteins are purified in a detergent solubilized state. Membrane proteins refolded from inclusion bodies are also finally obtained as protein-detergent complexes. Up to now, membrane protein crystallization for structure determination was always carried out from a detergent solubilized protein. 2D-crystallization of membrane proteins starts as well from detergent solubilized protein, but it is then reinserted into a phospholipid bilayer by removal of the detergent. Membrane proteins are kept soluble in micellar detergent solutions. The usual working range concentration is 1.5-2-fold CMC. The choice of detergent for solubilization and purification is important and should be considered with care. Most purification materials tolerate a wide range of detergents. Affinity purification systems have the advantage of high speed and high specificity compared to standard chromatographic procedures like anion exchange or size exclusion chromatography. For specific applications, recombinant antibody fragments can be used in indirect immunoaffinity chromatography for one-step purification of membrane protein-antibody complexes, as shown for bacterial cytochrome c oxidase (Kleymann et al., 1995). The aim is to ensure reproducible, pure, and homogenous protein preparations devoid of heterogeneity caused by proteolysis, different oligomerization states or posttranslational modifications like, for example, glycosylation. Crystallization of the water channel AQPl was performed with fully deglycosylated protein samples (Sui et al., 2000). Analytical size exclusion chromatography using, for example, the SMART system (Pharmacia) is a fast tool to analyze oligomerization states or aggregation tendency of proteindetergent complexes. Crystallization might be hampered by flexible termini or loops, and proteolytic cleavage or recombinant products omitting these termini might be useful. For instance, crystallization of the potassium channel

148

TABLE I

Crystallization and Structures of Membrane Proteins: Polytopic Membrane Proteins from Inner Membranes of Bacteria and Mitochondria and from Eukaryotic Membranes. A Complete and Regularly Updated List is Available at http://www.mpibp-frak~~.mpg.de/micheYpublic/memprotstruct.html. Detergent/additive/precipitatingagent (crystallization conditions)

Photosynthetic reaction centers and photosystems LDAO/heptane-l,2,3-triol/ammonium sulfate from Rhodopseudomonas viridis from Thermochromatium tepidum Photosystem I from Synechococcus elongatus Bacterial rhodopsins Bacteriorhodopsin from Halobacterium salinarum Bacteriorhodopsin from Halobacterium salinarum Bacteriorhodopsin from Halobacterium salinarum Sensory Rhodopsin I1 from Nu tronobacterium pharaonis Light-harvesting complex I1 from pea chloroplasts

Bacterial light-harvesting complexes Light-harvesting complex 2 from Rhodopseudomonas acidophila from Rhodospirillum molischianum

OG/-/PEG 4000, NaCl LM/-/water (reverse of salting in)

Key references Deisenhofer et al., 1985; Deisenhofer et al., 1995; Lancaster and Michel, 1999 Nogi et al., 2000 Jordan et al., 2001

(electron crystallography using naturally occurring two-dimensional crystals) (type I crystal grown in cubic lipidic phase of monooleyl-rac-glycerol in the presence of sodium/potassium phosphate) OG/benzamidine/sodium phosphate (epitaxial growth on benzamidine crystals) (type I crystal grown in cubic lipidic phase of monoolein glyceride in the presence of KC1)

Henderson et al., 1990; Kimura et al., 1997

(electron crystallography of two-dimensional crystals prepared from Triton X-100 solubilized material)

Kuehlbrandt et al., 1994

OG/benzamidine/phosphate

McDermott et al., 1995

UDAO/ heptane-l,2,3-triol/ammonium sulfate

Koepke et al., 1996

Cytochrome c oxidasehespiratory complex IVhbiquinol oxidase from Paracoccus denitrificans: LM/-/PEG-MME 2000 four-subunit and two-subunit enzyme in complex with antibody fragment (Fv) DM with some residual chelate/-/PEG 4000 from bovine heart mitochondria NG/-/PEG 2000 from Thermus thermophilus OG/ethanol/PEG 1500, MgC1, ubiquinol oxidase (cytochrome bo) from Escherichia coli

Pebay-Peyroula et al., 1997

Essen et al.. 1998 Luecke et al., 2001

Iwata et al., 1995; Harrenga and Michel, 1999; Ostermeier et al., 1997 Tsukihara et al., 1996; Yoshikawa et al., 1998 Soulimane et al., 2000 Abramson et al., 2000

Carola Hunte and Hartmut Michel

Membrane protein

Cytochrome bc, cornpledrespiratory complex I11 from bovine heart mitochondria MEGA-10 or diheptanoyl phosphatidyl choline/-/PEG 4000 from bovine heart mitochondria %/-/PEG 4000 from bovine heart mitochondria pure DDM or mixture with HECAMEG/PEG 4000 from the yeast Saccharomyces cerevisiae in UM/-/PEG 4000 complex with antibody fragment (Fv)

Zhang et al., 1998 Iwata et al.. 1998

Fumarate reductasekuccinate dehydrogenasehespiratory complex I1 Fumarate reductase from E . coli c,,E~/-/PEG 1000 Fumarate reductase from Wolinella succinogenes mixture of DDM and DM/benzamidine/PEG 4000

Iverson et al., 1999 Lancaster et al., 1999

Xia et al., 1997; Kim et al., 1998

~~

Potassium channel KcSA from Streptomyces lividans KcSA, in complex with antibody fragment (Fab)

LDAO/-/PEG 400 DM/-/PEG 400

Doyle et al., 1998 Zhou et al.. 2001

Mechanosensitive ion channel from Mycobacterium tuberculosis

LM/-/triethylene glycol

Chang et al., 1998

(type I crystal obtained by removing the detergent C,,E, by dialysis)

Toyoshima et al., 2000

NG/heptane-1,2,3-triol/ammoniumsulfate

Palczewski et al., 2000

(electron crystallography of two-dimensional crystals) NG/-/PEG-MME 550 OG/dithiothreitol/PEG 2000, MgCl,

Murata et al., 2000

P-type ATPases Ca-ATPase from rabbit sarcoplasmic reticulum G-protein coupled receptors Rhodopsin from bovine rod outer segments Aquaporins and relatives Aquaporin AQPl from human red blood cells Aquaporin AQPl from bovine red blood cells Glycerol facilitator GlpF from E . coli

CHAPTER 7 Membrane Protein Crystallization

Hunte et al., 2000

Sui et al., 2001 Fu et al., 2000

Detergents -C,E,: octylpentaoxyethylene, C,E,: octyltetraoxyethylene, C,,E,: dodecyl octaoxyethylene, C, ,R,: dodecyl nonaoxyethylene (Thesit), DDAO N,N-dimethyldecylamine-N-oxide,DM: decyl-P-D-maltoside, HECAMEG: methyl-6-O-(N-heptylcarbamoyl)-alpha-D-glucopyranoside,HEW. dihyH,G: n-hexyl-P-D-glucopyranoside, LG: n-dodecyl-P-Ddroxethyl sulfoxide, H,G: n-heptyl-P-D-glucopyranoside,H,DAO: hexyldimethylamine-N-oxide, glucopyranoside, L M dodecyl-8-D-maltoside, LDAO N,N-dimethyldodecylamine-N-oxide,MEGA-10: decanoyl-N-methyl-glucamide,N G nonyl-P-Dglucopyranoside, OG: octyl-8-D-glucopyranoside, OM: octyl-8-D-maltoside, UDAO N,N-dimethylundecylamine-N-oxide,UM: undecyl-P-D-maltoside, 8-HESO n-octyl-2-hydroxyethylsulfoxide; Precipitants-MPD: 2-methyl-2,4-pentanediol, PEG polyethylene glycol, PEG-MME: polyethylene glycolmonomethylether.

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Carola Hunte and Hartmut Michel

T A B L E II

Crystallization and Structures of M e m b r a n e Proteins: M e m b r a n e Proteins from the Outer M e m b r a n e of Gram N e g a t i v e Bacteria and Related M e m b r a n e Proteins

Membrane protein 16-stranded porins from Rhodobacter capsulatus OmpF and PhoE from Escherichia coli OmpK36 (osmoporin) from Klebsiella pneumoniae anion selective Omp32 from Comamomonas acidovorans 18-stranded porins Maltoporin from £. coli Maltoporin from Salmonella typhimurium Sucrose specific ScrY porin from S. typhimurium alpha-hemolysin from Staphylococcus aureus 8-stranded p-barrel membrane anchor OmpA-fragment from E. coli 22-stranded receptors FhuA from E. coli Ferric enterobacterin receptor (fepA) from E. coli

Detergent/additive/precipitating agent (crystallization conditions) C8E4/-/PEG 600 mixture of 8-HESO and octylpolyoxethylene; or DDAO/-/PEG 2000 8-HESO/-/PEG 2000

Weiss et al, 1991 Cowan et al, 1992

OG/VLi^SO^, (NH4)2S04

Zeth et al, 2000

mixture of DM and C12E9/-/PEG 2000 mixture of CgE^ and HxDAO/-/PEG 1500 mixture of OG and HxDAO/-/PEG 2000

Schirmer et al, 1995

O G / - / a m m o n i u m sulfate, PEG-MME 5000

Song et al, 1996

C8E4/2-propanol, HxDAO/ammonium sulfate

Pautsch and Schulz, 2000

DDAO/inositol/PEG-MME 2000 LDAO/heptane-l,2,3-triol/PEG1000

Ferguson et al, 1998 Buchanan et al, 1999

12-stranded p-barrel membrane anchor outer membrane phospholipase OG/-/MPD A (OMPLA) from E. coli TolC channel (3 x 4p-strands) from £. coli

Key references

DG, HxG, HpG, OG/heptane-l,2,3-triol/PEG-MME2000

10-stranded p-barrel outer membrane protease OmpT from £. coli OG/-/MPD

Dutzler et al, 1999

Meyer et al, 1997 Forst et al, 1998

Snijder et al, 1999

Koronakis et al, 2000

Vandeputte-Rutten et al, 2001

C. Proteins inserted into but not crossing the membrane ("monotopic membrane proteins") Prostaglandin H2 synthase-1 (cyclooxygenase 1) from sheep O G / - / P E G 4000 Picot et al, 1994 Cyclooxygenase-2 from Mus musculus from Homo sapiens Squalene cyclase from Alicyclobacillus acidocaldarius Abbreviations see Table I.

OG/-/PEG-MME 550 CgEs/VPEG 4000

Kurumbail et al, 1996 Luong et al, 1996

C8E4/-/sodium citrate

Wendt et al, 1997

CHAPTER 7 Membrane Protein Crystallization

151

KcSA was achieved only after proteolytic cleavage of 35 carboxy-terminal amino acids (Doyle ei ah, 1998; Zhou et ah, 2001). Restrained mutagenesis at the polar surface of the outer membrane proteins OmpA and OmpX was used for their crystallization (Pautsch et ah, 1999). It cannot be generalized whether the amino- or carboxy-terminal introduction of affinity tags—that is, short flexible peptides for affinity purification—are disadvantageous for crystallization. Quantitative removal of such tags by introduction of specific protease cleavage sites should be considered for crystallization purposes. Many membrane proteins possess autonomously folded extramembranous domains and some of the proteins are anchored in the membrane by a single transmembrane helix. These defined domains can often be isolated from the intact protein by proteolysis or can be produced as recombinant protein while retaining relevant biological activity. If the fragments are stable in aqueous solution, they can be treated and crystallized as soluble proteins without the use of detergent. This approach is often used for structural analysis of receptors and receptor ligand complexes (for review, see Hendrickson, 1996). Similar to soluble proteins, many membrane proteins change their conformation considerably during biological activity. It is essential for successful crystallization to be aware of different conformations or mobile domains. Preferentially one should be able to fix certain conformations, for example, by the use of specific inhibitors. The mobile extrinsic domain of subunit pRIPl of the cytochrome hc^ complex was not resolved in the first available structure of the complex from bovine mitochondria (Xia et ah, 1997). The domain's structure was only visible after its immobilization either by crystal contacts or by addition of Q^ site-specific inhibitors (Zhang et ah, 1998; Hunte et ah, 2000). There are various means to improve stability of a protein. If no conditions and detergents can be found to keep the protein stable or suitable for crystallization, a selection for stable mutants could help (Bowie, 2001). Thorough biochemical characterization of the protein preparation in combination with comprehensive screening for the most suited detergent may be the most efficient strategy to cope with the difficulties of membrane protein crystallization.

C. Detergents Since the early years of membrane protein crystallization, choosing the right detergent has been the key to success. Well-ordered crystals of the photosynthetic reaction center from the purple bacterium Rhodopseudomonas viridis could only be grown using N,N-dimethyl dodecylamine-n-oxide (LDAO) as detergent. Even use of the decyl homolog did not lead to crystals. In the case of the yeast cytochrome hc^ complex, which was crystallized with the help of antibody fragments, crystals did not grow in the presence of dodecyl-maltopyranoside (LM) but did grow in undecyl-maltopyranoside

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(UM) solutions. The alkyl chain of the latter is only one carbon atom shorter, and it seems obvious that the favored crystal packing cannot be obtained with the larger detergent micelle. Interestingly, the highest number of structures from polytopic membrane proteins from the inner membranes of bacteria and mitochondria and from eukaryotic cell membranes was solved in the presence of octyl-glucoside, one of the detergents with the shortest alkyl chains (see Table III). Obviously, the detergent micelle has to fit optimally into the crystal lattice of the protein. On one hand, this could mean that the detergent micelle should be as small as possible to allow a spacious crystal packing without interference of the detergent micelles. On the other hand, attractive polar interactions between

TABLE III

Detergents Used for Crystallization of Membrane Proteins. Numbers are given for all membrane protein structures available up to now (all), polytopic membrane proteins from inner membranes of bacteria and mitochondria as well as from eukaryotic membranes (A), membrane proteins from the outer membrane of gram negative bacteria and related membrane proteins (B) and monotopic membrane proteins (C); the latter are proteins that are only inserted into the membrane but do not cross it. Detergent

All

A

B

6 1 3 3

5 1

1

3 17 1

3 9 1

6

MEGA-10

1

1

—

LM UM DM

5 2 4

5 2 3

1

8-HESO

3

—

3

HECAMEG

1

1

—

C12E9

2 6 1 1

1

1 5

LDAO UDAQ DDAQ HxDAQ (as additive) NG OG glucopyranoside mixture

octylpolyoxyethylene

3 3 2

1 1

1

Abbreviations see Table I; for continuous update see http: / / www.mpibp-frankfurt.mpg.de / michel/ public/ memprotstruct.html

CHAPTER 7 Membrane Protein Crystallization

153

neighboring detergent micelles could be helpful by contributing to the stability of the crystal lattice. Such contacts can occur when the detergents have rather short alkyl chains. That attractive interactions between detergent micelles play a role is also indicated by the fact that crystallization often occurs close to conditions in which phase separation takes place, which is caused by attractive interactions between detergent micelles (Zulauf, 1991). At the so-called cloud point, a clear detergent solution turns turbid upon changes in temperature and separates into a detergent-rich and a detergent-depleted phase. In a phase diagram plotting detergent concentration versus temperature, the curved border between the detergent phase and the two immiscible aqueous phases is called the consolution boundary. Some detergents have an upper (e.g., octyltetraethylene oxide) and others a lower (e.g., octylglucoside) consolution boundary, meaning that some detergents separate into two phases upon heating and others upon cooling. Phase separation depends on the type of detergent and is affected by salt, polar additives, temperature, or precipitant (see Garavito and Ferguson-Miller, 2001). It is often experienced in crystallization screens of membrane proteins, and it might interfere with the screen. However, often crystallization occurs close to the consolute boundary. Crystal formation of the bacterial cytochrome c oxidase, for example, normally starts at the physical boundary between the two phases. The same is true for the first crystals of the yeast cytochrome bc^ complex (see Chapter 12). Detergent micelles can be made smaller by adding small amphiphilic molecules such as heptane-l,2,3-triol. This approach has been successfully used for the crystallization of bacterial photosynthetic reaction centers and light-harvesting complexes, for which rather harsh detergents with small polar head groups can be used. Heptane-l,2,3-triol is one of the most commonly used additives in crystallization trials (Tables I and II), but it is without positive effect in combination with mild detergents such as alkyl maltosides. Not only the length of the lipophilic tail of a detergent but also its head group properties are important for crystallization success. In some cases, crystals of the same protein can be obtained in different detergents, but diffraction quality varies in relation to the detergent used. The two subunit cytochrome c oxidase, for example, could be crystallized in LM, resulting in a diffraction limit of 8 A resolution; crystals grown in nonyl-maltoside (NM) were even less ordered. However, crystals grown in UM or cyclo hexyl-hexylmaltoside (CYMAL-6) showed a diffraction limit of 2.5 A and 2.6 A, respectively (Ostermeier et ah, 1997). In conclusion, even small chemical differences in the detergent can cause essential differences in the crystallization behavior of detergent-membrane protein complexes. Therefore, emphasis should be put on the screening of various detergents for crystallization. A good strategy is to prescreen the stability of the protein of interest in a broad range of available detergents and to perform crystallization trials in all of the selected detergents

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that keep the protein stable. Obviously, the type of detergent that is good for solubilization and purification of a protein need not be identical with the one suited for crystallization. Therefore, the most efficient approach for a detergent screening is to keep the established purification procedure as it is and perform detergent exchange with the pure protein or in the final purification step. For this purpose, it is advisable to bind the protein to a column chromatography material: a specific affinity matrix in case of recombinant proteins or to anion exchange material. The protein is then washed with excess amounts of buffer containing the new detergent and finally eluted in buffer containing the new detergent. This method was successfully used for detergent exchange of the yeast cytochrome bc^ complex (Hunte et ah, 2000) and resulted in complete removal of LM, the initial detergent, as shown by mass spectroscopic analysis (Lange, C. and Hunte, C, unpublished results). Size exclusion chromatography or exchange by ultrafiltration is not recommended, since they do not yield complete detergent exchange. Membrane proteins have been successfully crystallized in various nonionic and zwitterionic detergents mainly belonging to the groups of alkyl dimethylamine-oxides, alkyl glucosides, alkyl maltosides, and alkyl polyoxyethylenes (see Table III). Additionally, detergent mixtures in the protein solution or by addition of detergents in the crystallization trials can be useful. In the future so-called amphipoles may be an alternative to the classic detergents. These short-chain amphipatic polymers possess a strongly hydrophilic backbone decorated with hydrophobic side chains, and they were designed to keep membrane proteins soluble in aqueous solution (Tribe et ah, 1996). Amphipols stabilized detergent solubilized ATPase of the sarcoplasmatic reticulum against inactivation and aggregation (Champeil et ah 2000).

D. Crystallization Reassuringly, membrane protein crystals are obtained with standard crystallization techniques and conditions that are routinely used for soluble proteins (see Tables I and II) and finding the optimal crystallization conditions appears not to be the limiting step (for reviews, see Garavito et ah, 1996; Klihlbrandt, 1988; McPherson, 1990; Ostermeier and Michel, 197; Michel, 1983, 2001). The most commonly used precipitants to achieve supersaturation of the membrane protein solution and thereby nucleation for crystal growth are polyethylene glycols, polyethylene glycol monomethylether, or ammonium sulphate, and phosphate salts. The vapor diffusion method with sitting drops is most frequently applied. Data collection and structure determination is in general the same as with soluble proteins. Flash cooling is often used to enhance crystal stability in the x-ray beam or for trapping reaction intermediates (e.g., Yoshikawa et ah, 1998; Luecke et ah, 2001). In some cases establishing cryo-conditions for membrane proteins is difficult. This might be due to the presence of detergent micelles and large solvent content of the crystals.

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It must be kept in mind that most frequently well-ordered crystals are of type II—that is, crystals contacts are formed via the surfaces exposed to the aqueous environment. Therefore, expansion of the hydrophilic surface by tightly and highly specifically binding antibody fragments to the hydrophilic portion of the membrane protein is very attractive. Specifically binding antibody fragments form stably attached domains, and this approach has been successfully used for the crystallization and structure determination of the four- and two-subunit bacterial cytochrome c oxidase (Iwata et ah, 1995; Ostermeier ei ah, 1995; Ostermeier et al., 1997), the yeast cytochrome hc^ complex (Hunte et ah, 2000), and the potassium channel KcSA (Zhou et ah, 2001). Whereas the first two proteins were crystallized with recombinant antibody fragments, the latter structure was obtained with a proteolytically prepared Fab fragment. In all cases, the antibody fragments mediate important, or the only, crystal contacts. In case of the potassium channel, the complex structure could even be solved by molecular replacement using a known Fab fragment structure. To allow the formation of rigidly attached domains, fragments should bind with high affinity and should preferentially recognize discontinuous epitopes (see Chapter 12). Excellent high-affinity monoclonal antibodies can be obtained by traditional hybridoma technology. In the future, recombinant libraries may give faster access to suitable material. Provided a careful selection of suitable antibodies, the antibody fragment can fix a defined protein conformation or stabilize a flexible domain to aid crystallization attempts. Once crystals are obtained, it is required to improve their quality to obtain x-ray diffraction at high resolution. This includes control of protein purity and homogeneity as well as optimization of crystallization conditions. Small changes in pH, salt, precipitant concentration, additives, protein concentration, type, or purity of detergent have to be tested empirically. Furthermore, crystals have to be either grown to a large size for data collection at 4°C or adapted to cryo-protecting agents for flash-cooling and data collection at low temperature to obtain high-resolution data. When mixed with aqueous solvents, some lipids form a bicontinuous three-dimensional bilayer designated lipidic cubic phases. The idea was advanced that proteins can integrate into the lipid bilayer and be stabilized by the phospholipid environment, which might provide a stronger stabilizing lateral pressure as compared to a detergent micelle. The proteins should diffuse freely within the bilayer and should be able to form crystal contacts via their hydrophilic domains. Monomeric bacteriorhodopsin purified from purple membranes was incorporated in such a bilayer and successfully crystallized (Landau and Rosenbusch, 1996; see Chapter 16). These crystals are type I crystals, well-ordered stacks of 2D crystals. Although the mechanism of crystallization in lipidic mesophases is still not exactly understood (Caffrey, 2000), well-diffracting crystals of membrane proteins could be obtained and used for structure determination (Liicke et ah, 1998; Kolbe et ah, 2000). The

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technique is a valuable addition to the tools for membrane protein crystallization, and further development and understanding of the technique might help to crystallize membrane proteins that are unstable in detergent micelles or without added lipids.

E. Conclusion In the last several years, structural membrane protein research has been a rapidly evolving field with an increasing number of structures at near atomic resolution being determined. This progress was fostered by the availability of new detergents and by improvements in the field of overproduction of functional membrane proteins, both in their native environment and refolded from inclusion bodies. However, well-ordered 3D crystals are the prerequisite of structure determination by x-ray crystallography. Membrane protein crystallization is based on empirical screening steps and is still far from routine. Important factors for the successful crystallization of a membrane protein are stability and homogeneity of the protein and the choice of the detergent. Removing bottlenecks, especially in overproduction of proteins from eukaryotic organisms and in stabilization of proteins, is still needed so that structure determination of membrane proteins will be an approachable aim to study the mechanism of membrane proteins.

REFERENCES Abramson, J., Riistama, S., Larsson, G., Jasaitis, A., Svensson-Ek, M., Laakkonen, L., Puustinen, A., Iwata, S., and Wikstrom, M. (2000). The structure of the ubiquinol oxidase from Escherichia coli and its ubiquinone binding site. Nature Structural Biology 7, 910-917. Bowie, J. U. (2001). Stabilizing membrane proteins. Current Opinion in Structural Biology 11, 397-402. Buchanan, S. K., Smith, B. S., Venkatramani, L., Xia, D., Esser, L., Palnitkar, M., Chakraborty, R., van der Helm, D., and Deisenhofer, J. (1999). Crystal structure of the outer membrane active transporter FepA from Escherichia coli. Nature Structural Biology 6, 56-63. Chang, G., Spencer, R. H., Lee, A. T., Barclay, M. T., and Rees, D. C. (1998). Structure of the MscL homolog from Mycobacterium tuberculosis: a gated mechanosensitive ion channel. Science ISl, 211^

Cytochrome c oxidase : Fv7E2C50S co-complex purification by streptavidin affinity chromatography. Samples of solubilized P. denitrificans membranes (lane 1), E. coli periplasmic extract containing f i;7E2C50S (lane 2), mixture of solubilized membranes and periplasmic extract (lane 3), flow-through during loading of mixture (lane 4), proteins removed during washing step (lane 5), and fraction of desthiobiotin elution (lane 6) were subjected to SDS-PAGE (15% gel, 10 /ig of protein per lane, Coomassiestaining). Molecular mass standard proteins (M) were co-separated. As indicated on the right side, the highly purified co-complex is present in the eluate and contains the three major subunits of the cytochrome c oxidase as well as heavy and light chain of the Fv. The two latter polypeptides are not resolved, since they have a very similar molecular mass of about 14 kDa. Subunit IV, with a mass of 5 kDa, is not visible on the gel.

cherichia coli, an integral membrane protein with 12 transmembrane helices and very short solvent exposed loops. These antibodies were used to probe conformational changes of the antiporter upon regulatory pH shift (Venturi et al, 2000). Native antibodies are not suitable for crystallization attempts because they are flexible in their "elbow" region between the variable and constant domains (Lesk and Chothia, 1988). Furthermore, their bivalent binding mode is in most cases undesirable. Monovalent antibody fragments can be generated by proteolytic cleavage of the whole antibody, resulting in two Fab (fragment antibody binding) molecules per antibody molecule. Proteolytic Fahs are relatively easy to obtain, but care must be taken to produce material suitable

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for crystallization. Proteolytic cleavage might result in heterogeneous fragment preparations. Furthermore, residual parts of the flexible "elbow," and in some cases glycosylation, can be an obstacle for crystallization attempts. Yet, a large number of proteolytic Fabs alone and in complex with the respective hapten or antigen have been crystallized and the structure of the complexes determined. Recently, the crystallization of a membrane protein with a proteolytic Fab, namely of the KcsA K^-channel, was the first successful example (Zhou et al., 2001). Here, the antibody fragment was not only essential for crystallization by providing crystal contacts but was as well utilized for structure determination, since the phases were solved by molecular replacement based on the known atomic model of a Fab. Recombinant antibody fragments appear to be more versatile, since they can be engineered to facilitate one-step affinity purification of the antigenantibody complex (Kleymann ei aL, 1995). Well-established systems ensure pure and homogenous preparations of the protein complexes. Furthermore, recombinant antibody fragments can be used either in Fv (fragment i;ariable, ^ 2 8 kDa) or Fab ( ^ 5 6 kDa) format. These alterations in size might be critical for the success of crystallization, since the bound fragment should outreach the detergent micelle, which covers the transmembrane portion of the protein and does not contribute to rigid crystal contacts (Fig. 3). In addition, formation of antigen-antibody complexes might be useful to stabilize specific conformations of a protein to aid crystallization attempts. Three-dimensional crystallization of complex target antigens, such as viral flexible proteins, has been achieved with specifically bound Fabs (Kwong et ah, 1998). Recombinant antibody fragments can be obtained by cloning the encoding genes from hybridoma cell lines (Kleymann et ah, 1995; Ostermeier and Michel, 1995). As a future perspective, direct selection of recombinant antibody fragments from phage display libraries (for review, see Viti et ah, 2000) or by ribosome display (for review, see Hanes et ah, 2000) should ensure fast access to suitable fragments even for native membrane proteins. A variety of expression systems are in use for the production of recombinant fragments. In this chapter, the production of Fvs in E. coli by periplasmic expression is described (Kleymann et ah, 1995). The oxidizing milieu of the periplasm is essential for the proper formation of disulfide bridges and thus for the correct folding of the antibody fragments. However, the relatively small volume of the periplasm results in low yield. A promising approach to overcome these limitations is the cytoplasmic expression of fragments in an E. coli host, which has an impaired reducing system and therefore an oxidizing cytosol (Venturi et ah, 2002). In the following sections, the indirect immunoaffinity purification of cytochrome c oxidase from P. denitrificans, as well as the Fi;-mediated crystallization of this complex and of the yeast cytochrome bc^ complex (Hunte et ah, 2000) are described.

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II. PRODUCTION OF ANTIBODY Fv FRAGMENTS IN THE PERIPLASM OF E. coli The recombinant Fv consists of two polypeptide chains, the variable domains of the heavy and light chains, which are noncovalently associated. The protein is produced by periplasmic expression, using the plasmid pASK68 and the E. coli strain JM83 as host (Kleymann et ah, 1995). The vector carries under control of the lac promoter an artificial dicistronic operon for the simultaneous expression of the genes encoding heavy and light chain of the Fv. The N-terminal fusion of OmpA and PhoA signal sequences to the heavy and light chain, respectively, will target the expression products to the periplasm, where assembly and folding of the Fv take place. For expression 6 x 50 ml I3amp medium (see Sambrook et ah, 1989) in 250 ml flasks are inoculated with 6 X 5 /il glycerol stock of the E. coli strain ]M83 carrying the desired plasmid. The cultures are grown overnight (o/n) at 30°C while shaking at 220 rpm. 6 x 2 1 LB«^p medium in 5 1 flasks are preincubated at 22.5°C. These cultures are started by addition of 50 ml of the o/n culture per flask. After ^^4 hr incubation at 22.5°C and 200 rpm, an OD550 of 0.5 is reached and IPTG is added to a final concentration of 1 mM to induce expression. Incubation is continued under the same conditions for 3 hrs. Then the cells are transferred to precooled centrifugation buckets and sedimented by centrifugation for 12 min at 4000 X g and 4°C. The sedimented cells are gently resuspended in ice-cold extraction buffer (100 mM Tris-HCl, pH 8.0, 500 mM sucrose, 1 mM EDTA) at 1/100 of the culture volume. After 30 min incubation, cells and cell debris are spun down at 5000 x g for 45 min at 4°C. The periplasmic extract is poured off and can be stored in appropriate aliquots at — 20°C for several months. Low temperature and a relatively short expression time are essential requirements for the production of the Fvs. The yield depends on stability and folding properties of the respective fragment; it varies between 0.1 mg/1 to 2.5 mg/1 of culture. The quality of the expression can easily be analyzed by subjecting the periplasmic extract to SDS-PAGE analysis. Expression levels of 1 mg/1 of culture are visible in Coomassie-stained gels. The presence of heavy and light chain (^ 14 kDa each) can be checked by Western immunoblot, since the expression vector pASK68 provides a myc-tag and a Strep-iag fused to the C-terminus of the light and heavy chain, respectively. The myc-tag can be detected using the monoclonal antibody 9E10. For visualization of the Streptag streptavidin-alkaUne phosphatase can be used.

III. PURIFICATION OF Fvs BY STREPTAVIDIN AFFINITY CHROMATOGRAPHY A 25 ml streptavidin affinity column is prepared by covalent attachment of recombinant core-streptavidin/mutantl (Schmidt and Skerra, 1994; Voss

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and Skerra, 1997) to activated CH-Sepharose 4B (Pharmacia). Then 5-10 mg protein per ml matrix is coupled according to the manufacturers instructions. Streptavidin or streptavidin chromatography material is commercially available (e.g., IBA, Gottingen). The streptavidin affinity column (2.5 cm I.D.) is equilibrated with 5 column volumes of Tris buffer (100 mM Tris-HCl, pH 8.0) at a flow rate of 0.7 ml/min. Biotin and biotin-carrying proteins are removed from the periplasmic extract by avidin treatment prior to applying the extract to the column to avoid irreversible contamination of the column material. 400 /ig avidin per liter of culture is added from a stock solution (10 m g / m l ; store aliquots at — 20°C). After 30 min incubation on ice, the aggregated proteins are sedimented for 30 min at 35,000 x g at 4°C. The clear supernatant is sterile filtered to reduce the risk of microbial contamination (0.2 //m) and loaded onto the column. Unbound material is removed with equiUbration buffer until the signal of the UV-recorder reaches approximately 10% of the base level. Elution is performed using 2.5 mM Desthiobiotin in equilibration buffer. The eluted protein is collected and concentrated using spin filtration devices (10 kDa cutoff) to a final protein concentration of --5 m g / m l . Desthiobiotin can be removed by dialysis or by subsequent dilution and concentration steps with equilibration buffer. Samples should be sterile filtrated and kept at 4°C. Depending on the Fv, conditions of the storage buffer have to be adjusted — for example, by addition of salt.

IV. Fv MEDIATED CRYSTALLIZATION OF CYTOCHROME c OXIDASE FROM P. Denitrificans A. Growth of P. denitrificans P. denitrificans (ATCC 13543) cells are grown in succinate medium (Ludwig, 1986) in 5 1 shaker flasks at 32°C. The medium contains 50 mM K2HPO4, 75 mM NH4CI, 11.5 mM Na2S04, 1.25 mM MgCl2, 1 mM citric acid, 85 mM succinic acid, and l m l / 1 trace element solution. The pH is adjusted with KOH to 6.2. The trace element solution consists of 100 mM CaCl2, 90 mM FeCl3, 50 mM MnCl2, 25 mM ZnCl2, 10 mM C0CI2, 5 mM CUCI2, 5 mM H3BO3 and 10 mM Na2Mo04 dissolved in half-concentrated HCl. The cells are harvested at mid-log phase by centrifugation, resuspended in 200 mM Tris-HCl, pH 8, 1 mM EDTA, and stored at -80°C.

B. Isolation of Membranes For the preparation of membranes, cells are thawed, supplemented with 0.05 m g / m l Pefabloc SC (Biomol), and disrupted using a microfluidizer (pressure 16,000 psi, 5 cycles; Microfluidics Corporation, Newton, Mass.). Cell debris is removed by centrifugation at 20,000 x g for 15 min. The supernatant

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is diluted 1:4 in 20 mM Tris-HCl, pH 8, 1 mM EDTA, and centrifuged for 4 hrs at 200,000 x g using a Ti45 rotor (Beckman). The supernatant is removed by suction, and the reddish, soft membrane pellets are resuspended in 20 mM Tris-HCl, pH 8, 1 mM EDTA, with a blender (Ultra-Turrax T25, IKA Labortechnik) and stored in 25 ml aliquots at — 80°C. The protein concentration is determined using the Lowry assay (Lowry et ah, 1951; BioRad). The typical concentration should be around 50 mg of protein per ml membrane suspension.

C. Solubilization of Membrane Proteins P. denitrificans cytoplasmic membranes (30 ml) are solubilized by addition of 20 ml of a 10% (w/v) solution of n-dodecyl-jS-D-maltopyranoside (equivalent to 1.3 mg detergent/mg protein). 50 /iM Pefabloc SC (Biomol) and 4 mg Avidin (Gerbu) are added to respectively prevent proteolysis and to bind biotin traces. After stirring the mixture for 5 min on ice, 50 ml E. coli periplasmic extract containing the recombinant monoclonal antibody fragment Fz;7E2C50S (Kleymann et ah, 1995; Ostermeier et ah, 1995) is added and the mixture is stirred for additional 10 min on ice. The mixture is centrifuged for 45 min at 350,000 g using Ti70 rotor (Beckman). The supernatant is collected and passed through a 1.2 /^M sterile filter. For the preparation and crystallization of the so-called two-subunit cytochrome-c oxidase, which lacks subunits III and IV, the P. denitrificans membranes are solubilized with 7 ml of a 30% (w/v) dodecyl-]V,N-dimethylamineN-oxide solution (Fluka). This treatment effectively removes the two subunits and yields a "core complex" containing only subunits I and II.

D. Indirect Immunoaffinity Chromatography The filtrate from Section IV,C is loaded onto a 25 ml streptavidin affinity column (see Fig. 1) after preequilibration with 20 mM Tris-HCl, pH 8, 50 mM NaCl, 0.05% n-dodecyl-^-D-maltopyranoside (or 0.1% dodecyl-N,N-dimethylamine-iV-oxide for purification of the two-subunit enzyme) at a flow rate of 0.7 ml/min. Unbound material is removed by washing with 1.5-2 column volumes of equilibration buffer until the absorbance at 280 nm declines close to base level. The co-complex consisting of Fv and cytochrome-c oxidase is then eluted with 2.5 mM desthiobiotin (Sigma) in the same buffer. This single-step purification yields a very pure and concentrated protein preparation (see Fig. 2).

E. Detergent Exchange and Size Exclusion Chromatography For preparation of the two-subunit cytochrome c oxidase, the detergent needs to be replaced after the affinity column because the enzyme slowly degrades in the presence of dodecyl-]V,N-dimethylamine-N-oxide. Detergent

CHAPTER 12 Antibody Fragment Mediated Crystallization of Membrane Proteins

FIGURE 3

213

Antibody fragments are essential for the crystallization of the yeast cytochrome bc^ complex, as crystal contacts (white arrows) of the dimeric yeast cytochrome bc^ complex: fi;18Ell co-complex are mediated by the Fv (encircled in red) [from Hunte, 2001]. The fragment binds to the extrinsic domain of the Rieske protein and generously provides space for the detergent micelle, in which the transmembrane portion of the complex (dotted lines) is embedded, as schematically depicted in the insert. The central molecule of the shown crystal packing is colored in green and yellow, the neighboring molecules in gray. {See also color plate.)

exchange is achieved by binding the protein to an anion-exchange column, followed by washing and elution with a buffer containing the new detergent. Here, the exchange to n-undecyl-jS-D-maltopyranoside (Biomol) is described, but the method works with any other nonionic detergent. After addition of n-undecyl-jS-D-maltopyranoside to a final concentration of 0.2% to the eluted co-complex of the previous step, the protein is loaded onto a 10 ml Q-Sepharose FF column. For detergent exchange 120 ml 20 mM Tris-HCl, pH 8, 50 mM NaCl, 0.2% n-undecyl-jS-D-maltopyranoside are passed through the column. Then the protein is eluted with the same buffer containing 600 mM NaCl. For optimal co-complex yield the antibody fragment is present in excess throughout the preparation. In a final step, surplus Fv is removed by size exclusion chromatography (SEC). The eluted co-complex is concentrated to ^ 1 ml by spin-filtration (Centriprep, 50 kDa cutoff, Amicon), filtrated (0.2 /im), and applied to a 60 x 0.7 cm TSK 3000 HPLC SEC column (Tosohaas), which was equilibrated with 10 mM Tris-HCl, pH 7.2, 20 mM NaCl, and detergent at approximately the 2-fold of the critical micellar concentration (i.e..

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0.02% n-dodecyl-jS-D-maltopyranoside or 0.07% n-undecyl-j8-D-maltopyranoside). Separation is performed at room temperature at a flow rate of 1.0 ml/min. The co-complex is fully separated from unbound Fv, the corresponding fractions are pooled, concentrated to ^100 juM by spin-filtration (50 kDa cutoff, Centricon, Amicon), and stored at — 80°C.

F. Crystallization of Cytochrome c Oxidase-Fi? Complex P. denitrificans cytochrome c oxidase was originally crystallized as a complex containing all four subunits and bound Fv (Ostermeier et al., 1995). Crystals were grown at 14°C in the presence of N-dodecyl-j8-D-maltopyranoside by vapor diffusion (sitting drop) against 10-14% polyethylene glycol 2000 monomethyl ether in 400 mM ammonium acetate, pH 8. The procedure yielded tetragonal crystals (space group P4, a = b = 206.7 A, c = 83.5 A, one molecule per asymmetric unit) that appeared after four weeks and diffracted to a resolution better than 3 A. However, the crystals were difficult to grow and showed a high degree of anisotropy in their diffraction pattern. Better crystals can be obtained when a preparation containing only the two catalytically essential subunits I and II is used (see above). Crystal growth is of better reproducibility, and they diffract x-rays to below 2.5 A resolution. They are grown in the presence of N-undecyl-jS-D-maltopyranoside (UM) at 16°C by vapor diffusion (hanging drop) against 7-8.5% polyethylene glycol 2000 monomethyl ether in 100 mM sodium acetate, pH 5.5. The first small crystals are visible in one to two days; crystal growth is complete after two weeks. The reddish brown crystals have a size of up to 0.3 x 0.3 x 1 mm and are orthorhombic {Vl^l^l^, a = 93.5 A, b = 151.0 A, c = 156.7 A, one molecule per asymmetric unit).

V. Fr-MEDIATED CRYSTALLIZATION OF THE CYTOCHROME hc^ COMPLEX FROM THE YEAST S. cerevisiae A. Co-Complex Formation and Purification The yeast cytochrome bc^ complex is crystallized with the fz;18Ell. The parental mablSEll was obtained by standard hybridoma technique by immunizing the mouse with detergent-solubilized and purified cytochrome bc^ complex (Hunte, unpublished results). Both antibody and derived fragment do not recognize the protein in Western Immunoblot after SDS-PAGE separation of the complex. The structure determination of the co-complex revealed that the antibody binds to the extrinsic domain of the Rieske-protein, one of the three catalytic subunits of the enzyme (Hunte et ah, 2000). The mobile extrinsic domain of the Rieske protein is fixed to the complex by a single transmembrane helix. Purification attempts of the co-complex by streptavidin

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affinity chromatography, as it was shown for the bacterial cytochrome c oxidase, were not successful. They resulted in partial disintegration of the complex. Therefore, the yeast cytochrome bc^ complex and the fi;18Ell are purified separately, the enzyme by anion-exchange chromatography as described in Chapter 11 and the antibody fragment by streptavidin affinity chromatography (see Fig. 1). For co-complex formation, cytochrome bc^ complex and frlSEll are mixed in SEC buffer (50 mM Tris, pH 8.0, 250 mM NaCl, 0.05% UM) at a final concentration of 30 fiM and 35.7 /iM, respectively. Surplus antibody fragment is removed by size exclusion chromatography, subjecting this mixture to a TSKsw 4000 HPLC column (60 x 0.7 cm, Tosohaas), which was preequilibrated with SEC buffer. Separation is performed with a flow rate of 0.6 ml/min at 4°C. The co-complex and the unbound fragment are clearly separated. The co-complex containing fractions are pooled and concentrated using a spindevice (20 kDa cutoff, Centrisart, Sartorius).

B. Crystallization of Cytochrome bc^ Complex: Fz;18Ell Co-Complex The cytochrome bc^ complex: fz;18Ell co-complex was initially crystallized by the vapor diffusion technique using a hanging drop setup (Hunte et ah, 2000). Twenty-four well Linbro-plates are prepared with 1 ml reservoir solution per well, and vacuum grease is applied to the upper ridges of each well. On a silane-treated cover slide 1 /xl of co-complex (20 mg/ml. A) is mixed with 1 III of reservoir solution (4-5% PEG4000, 100 mM Tris, pH 8.0, 0.05% UM, 10 /iM stigmatellin). The cover slide is inverted and placed on top of the ring of grease so the well is tightly sealed. Crystals grow at 4°C within 2-3 weeks (Fig. 4A, B) to a final size of approximately 70 x 70 x 70 jum. They diffract x-rays at a synchrotron source to 2.3 A resolution and belong to the space group C2 with cell constants of a = 214 A, b = 162 A, c = 147 A, j^ = 117°. The crystals crack or lose diffraction upon addition of cryo-protecting agents, and they do not grow in the presence of these additives. At 4°C diffraction declines very fast upon exposure with synchrotron x-ray sources. To obtain high-resolution data, large crystals have to be grown, since they can be exposed at several positions when using a small beam size. Larger crystals were obtained using the microseeding technique. A single, well-grown crystal of the initial vapor diffusion setup is taken out of the drop with microtools. It is washed with several aliquots of seeding buffer (8% PEG4000,100 mM Tris-HCl, pH 8.0, 0.05% UM, 10 iM stigmatellin). With the help of a pipette tip or a microspatula, the crystal is fully ground in a small aliquot of the same buffer. Larger particles sediment after a short period of incubation without agitation. The supernatant is subjected to a dilution series in seeding buffer (10^-10^ x dilution), and the optimal seeding stock is tested as follows: 2 ^l co-complex (50 mg/ml) are mixed with 1 /il crystallization solution (reservoir buffer with PEG4000 concentration increased to 11-13%).

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FIGURE 4

Crystals of the yeast cytochrome bc^ complex: Fi;18Ell co-complex. (A) Crystals were initially grown using the vapor diffusion technique and 4 - 5 % buffered PEG 4000 precipant solution (see Section IV). They appeared after 2 - 3 weeks incubation at 4°C accompanied by granular precipitate. Crystals often grew in areas of phase separation, resulting in a globular form. (B) Setups with less phase separation produced pyramidlike crystals. Both types typically grew to a maximal size of 70 x 70 x 70 fiirv. (C) Microseeding (see Section IV) was used to grow large single crystals with an average side length of 500 fivn. In these setups the protein is completely used up for crystal growth, leaving a clear background. (See also color plate.)

A cat whisker is dipped in the seeding stock and then pulled through the mixed droplet. The cover slide is sealed on a crystallization well filled with 1 ml reservoir solution (PEG4000 concentration 5%). Crystals can be seen after 1-8 hrs, indicating a strong batch-crystallization component rather than a vapor diffusion process. The optimal seeding stock produces 1-5 crystals per drop. The sitting drop technique is used for scale-up, increasing the final size

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of the droplets to a maximum of 20 /il per depression well of the crystallization stages (W. Jockel company, Weiterstadt, Gemany). Crystallization chambers (Boll, Munich) are filled with 10 ml reservoir solution. Under optimal conditions the crystals grow to full size within a few days, and the protein in the drop is fully depleted. With this approach crystals up to 1 mm in length can be grown (Fig. 3C). They are sufficient to collect at 4°C complete x-ray diffraction data sets up to 2.5 A resolution at synchrotron sources (e.g., at beamlines ID14EH3, ESRF, Grenoble and Xll, DESY, Hamburg). As the crystals are highly isotropic, data sets at higher resolution are achieved by merging the diffraction data from several crystals. Ten crystals were used for the 2.3 A resolution structure of the yeast cytochrome bc^ complex (Hunte et al 2000; Lange et ah 2001).

VI. CONCLUSIONS The cytochrome c oxidase from P. denitrificans can be rapidly purified by indirect immunoaffinity chromatography in complex with an antibody fragment. Recombinant antibody fragments (Fvs) have been the key for crystallization and structure determination of this membrane protein as well as of the yeast cytochrome bc^ complex. The approach of antibody fragment mediated crystallization appears to be especially attractive for obtaining well-ordered 3D crystals from membrane proteins with small hydrophilic domains, as recently shown for the K^-channel. Furthermore, an appropriate antibody fragment can help to stabilize a flexible protein or to fix a defined conformation of a protein, thereby enhancing the chances for their crystallization.

ACKNOWLEDGMENTS Financial support to C.H. by the Deutsche Forschungsgemeinschaft (SFB 472)I isis acknowledged acknowledged.

REFERENCES Hanes, J., Jermutus, L., and Pluckthun, A. (2000). Selecting and evolving functional proteins in vitro by ribosome display. Meth. Enzytn. 328, 404-430. Hunte, C, Koepke, J., Lange, C, Rossmanith, T., and Michel, H. (2000). Structure at 2.3 A resolution of the cytochrome bc^ complex from the yeast Saccharomyces cerevisiae co-crystallized with an antibody Fv fragment. Structure 8, 669-684. Hunte, C. (2001). Insights from the structure of the yeast cytochrome bc^ complex: Crystallization of membrane proteins with antibody fragments. FEES Lett. 504, 126-132.

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Carola Hunte and Aimo Kannt Kleymann, G., Ostermeier, C , Ludwig, B., Skerra, A., and Michel, H. (1995). Engineered Fv fragments as a tool for the one-step purification of integral multisubunit membrane protein complexes. Bio/Technology 13, 155-160. Kwong, P. D., Wyatt, R., Robinson, J., Sweet, R. W., Sodroski, J., and Hendrickson, W. A. (1998). Structure of an HIV gpl20 envelope glycoprotein in complex with the CD4 receptor and a neutralizing human antibody. Nature 393, 648-659. Lange, C , Nett, J., Trumpower, B. L., and Hunte, C. (2001). Specific roles of protein-phospholipid interactions in the yeast cytochrome bc^ complex structure. EMBO /. 20, 6591-6600. Lange, C , and Hunte, C. (2002). Crystal structure of the yeast cytochrome hc^ complex with its bound substrate cytochrome c. Proc. Natl Acad. Sci. USA 99, 2800-2805. Lesk, A. M., and Chothia, C. (1988). Elbow motion in the immunoglobulins involves a molecular ball-and-socket joint. Nature 335,188-190. Lowry, O., Rosebrough, N., Farr, A., and Randall, R. (1951). Protein measurement with the folin phenol reagent. /. Biol Chem. 193, 265-275. Ludwig, B. (1986). Cytochrome-c oxidase from Paracoccus denitrificans. Meth. Enzymol 126,153-159. Ostermann, T. (2000). PhD thesis. University of Frankfurt. Ostermeier, C , Iwata, S., Ludwig, B., and Michel, H. (1995). Fv fragment-mediated crystallization of the membrane protein bacterial cytochrome-c oxidase. Nat. Struct. Biol 2, 842-846. Ostermeier, H., and Michel, H. (1996). Improved cloning of antibody variable regions from hybridomas by an antisense-directed RNase H digestion of the P3-X63-Ag8.653 derived pseudogene mRNA. Nucl Acids Res. 24, 1979-1980. Ostermeier, C , Harrenga, A., Ermler, U., and Michel, H. (1997). Structure at 2.7 A resolution of the Paracoccus denitrificans cytochrome-c oxidase complexed with an antibody Fv fragment. Proc. Natl Acad. ScL USA 94, 10547-10553. Padan, E., Venturi, M., Michel, H., and Hunte, C. (1998). Production and characterization of monoclonal antibodies directed against native epitopes of NhaA, the Na'^/H'^-antiporter of Escherichia coll FEBS Lett. 441, 53-58. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Laboratory Press. Schmidt, T. G. M., and Skerra, A. (1994). One-step affinity purification of bacterially produced proteins by means of the "Strep tag" and immobilized recombinant core streptavidin. /. Chromatogr. A 676, 337-345. Schmidt, T. G. M., and Skerra, A. (1993). The random peptide library-assisted engineering of a C-terminal affinity peptide, useful for the detection and purification of a functional Ig Fv fragment. Prot. Eng. 6, 109-122. Schmitt, J., Hess, H., and Stunnenberg, H. G. (1993). Affinity purification of histidine-tagged proteins. Mol Biol Reports 18, 223-230. Studier, F. W., Rosenberg, A. H., Dunn, J. J., and Dubendorff, J. W. (1990). Use of T7 RNA polymerase to direct expression of cloned genes. Meth. Enzymol 185, 60-89. Venturi, M., Seifert, C , and Hunte, C. (2002). High-level production of functional antibody Fab fragments in an oxidizing bacterial cytoplasm. /. Mol Biol 315,1-8. Venturi, M., Rimon, A., Gerchman, Y., Hute, C , Padan, E., and Michel, H. (2000). The monoclonal antibody 1F6 identifies a pH-dependent conformational change in the hydrophilic NH2 terminus of NhaA Na"^/H"^-antiporter of Escherichia coU. J. Biol Chem. 275, 4734-4742. Viti, F., Nilsson, F., Demartis, S., Huber, A., and Neri, D. (2000). Design and use of phage display libraries for the selection of antibodies and enzymes. Meth. Enzym. 326, 480-505. Voss, S., and Skerra, A. (1997). Mutagenesis of a flexible loop in streptavidin leads to higher affinity for the Strep-iag II peptide and improved performance in recombinant protein purification. Prot. Eng. 10, 975-982. Zhou, Y., Morais-Cabral, J. H., Kaufman, A., and MacKinnon, R. (2001). Chemistry of ion coordination and hydration revealed by a K"^ channel-Fab complex at 2.0 A resolution. Nature 414, 43-48.

13

Crystallization of Wolinella succinogenes Quinol: Fumarate Reductase C. ROY D. LANCASTER Dept. Molekulare Membranbiologie Max-Planck-Institut fur Biophysik 60528 Frankfurt am Main, Germany

I. INTRODUCTION Quinolifumarate reductases (QFR) and succinate:quinone reductases (SQR) catalyze the reduction of fumarate to succinate with concomitant oxidation of hydroquinone (quinol) to quinone, as well as the reverse reaction. SQR (respiratory complex II) is involved in aerobic metabolism as part of the citric acid cycle and the aerobic respiratory chain. QFR is involved in anaerobic respiration with fumarate as the terminal electron acceptor (Kroger, 1978; Kroger et al., 1992), and it is part of the electron transport chain catalyzing the oxidation of various donor substrates (e.g., NADH, H2 or formate) by fumarate. These reactions are coupled to ADP phosphorylation via an electrochemical proton potential.

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QFR and SQR complexes are collectively referred to as succinateiquinone oxidoreductases (EC 1.3.5.1) and are predicted to share similar structures. The complexes consist of two hydrophilic and one or two hydrophobic membraneintegrated subunits (reviewed by Hagerhall, 1997, and Ohnishi et ah, 2000). The larger hydrophilic subunit A carries covalently bound FAD (8a-(N3)histidyl flavin adenine dinucleotide), and subunit B contains three ironsulphur centers. QFR of Wolinella succinogenes and SQR of Bacillus subtilis contain only one hydrophobic subunit (C) with two haem b groups. In contrast, SQR and QFR of Escherichia coli contain two hydrophobic subunits (C and D) that bind either one (SQR) or no haem b group (QFR). At resolutions of 2.2 A and 3.3 A, respectively, the crystal structures of W. succinogenes QFR (Lancaster et ah, 1999) and E. coli QFR (Iverson et ah, 1999) have been determined. In the case of W. succinogenes QFR, three crystal structures were determined, based on three different crystal forms, all of the monoclinic space group P2i. In all three crystal forms, two 130kd QFR monomers of A, B, and C subunits are arranged in an identical manner with respect to one another, thus forming a dimer. The structure obtained from crystal form "A," which has the unit cell dimensions a = 85.2 A, b = 189.0 A, c = 117.9 A, and jS = 104.5°, was refined at 2.2 A resolution (PDB entry IQLA, Lancaster et ah, 1999). It shows that subunit A, the flavoprotein, is composed of four domains, the first two of which are the bipartite FAD binding domain (residues Al-260 and A366-436, with "A" indicating the A subunit), into which the second domain, the capping domain (A260-366) is inserted. A W. succinogenes QFR crystal grown in the presence of fumarate was found to be of crystal form ''B," with unit cell dimensions of a = 118.4A,b = 85.1 A, c = 188.9 A, and jS = 96.5°. The structure was refined at 2.33 A resolution (PDB entry IQLB, Lancaster et ah, 1999). This allowed the localization of the fumarate binding site between the FAD binding domain and the capping domain next to the plane of the FAD isoalloxazine ring. The structure of the enzyme in the third crystal form, "C," with unit cell dimensions a = 81.1 A, b = 290.2 A, c = 153.6 A, and P = 95.7° (Lancaster et ah, 2000) was refined at 3.1 A resolution (PDB entry 1E7P, Lancaster et ah, 2001). Compared with the previous crystal forms, the capping domain is rotated in this structure relative to the FAD-binding domain. This leads to interdomain closure at the fumarate reducing site, suggesting that the structure encountered in this crystal form represents a closer approximation to the catalytically competent state of the enzyme. In summary, all three crystal forms have contributed toward our understanding of the structure and mechanism of action of W. succinogenes QFR. All three crystal forms are obtained under the same crystallization conditions, as will be presented below. There are two principle types of membrane protein crystals (Michel, 1983; see Michel, 2001, for a recent overview). Type I crystals can be thought of as ordered stacks of two-dimensional crystals formed in the planes of the membrane. Type II crystals contain detergents bound in a micellar manner.

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They are held together via polar interactions between the polar surfaces of the membrane protein. All of the three QFR crystals discussed here are of type II. In this case, the size and shape of the detergent micelle are critical in enabling or disabling the polar contacts between the polar surfaces of QFR. The crystals described here were obtained with a detergent mixture of dodecyl-jS-Dmaltoside and decyl-j5-D-maltoside. Less useful crystals could also be obtained using either dodecylmaltoside alone or dodecylsucrose. No crystals were obtained with N,N-dimethyldodecylamine-N-oxide (LDAO). Detergent micelles can be made smaller when small amphiphilic molecules like heptane1,2,3-triol (Gast et al., 1994; Timmins et ah, 1991) or benzamidine are added. The latter is used as an additive in the case of QFR. QFR crystals can also be obtained in the absence of benzamidine, but these do not diffract X-rays. The crystallization procedure involves the simultaneous equilibration of the protein droplet along a polyethylene glycole gradient, as detailed below. A major bottleneck in membrane protein crystallization is the availability of a biochemically well-characterized, homogenous, and stable membrane protein preparation in quantities of more than 10 mg of membrane protein. Similar to some other respiratory or photosynthetic membrane proteins, this is not a serious obstacle for W. succinogenes QFR. From 60 liters of W. succinogenes wild-type culture, 150 mg of stable material can be obtained after isolation and preparative isoelectric focusing. The latter technique allows the removal of minor impurities from the QFR preparation and thus increases the reproducibility of the subsequent crystallization procedure.

11. P R E P A R A T O R Y S T E P S A . G r o w t h of Wolinella

succinogenes

W. succinogenes is grown with formate and fumarate, as described by Bronder et al (1982) and Kroger et al (1994) . The growth medium (60 liters) has the following composition: fumaric acid (90 mM), sodium formate (0.1 M) K2HPO4 (20 mM), (NH4)2S04 (5 mM), NH4CI (5 mM), sodium acetate (20 mM), glutamate (1 mM), and Tris (50 mM). The solution is adjusted to pH 7.8 by the addition of KOH (about 0.1 M). After the addition of MgCl2 (0.25 mM), CaCl2 (0.05 mM), and trace element solution (2 ml/liter), the medium is sterilized, inoculated (1%), flushed with N2, and kept at 37°C until the stationary growth phase is reached (about 17 hr). Bacteria are harvested and kept at — 80°C. The trace element solution contains (in mg/liter) disodium EDTA (5200), FeCl^ • 4H2O (1500), ZnCl^ (70), MnCl^ • 4H2O (100), H3BO3 (62), CoCl2-6 H2O (190)-CuCl2-2H2O (17), NiCl2-6 H2O (24), Na2Mo04-2H20 (36).

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6. Isolation of Quinol: Fumarate Reductase Quinol: fumarate reductase is purified as described by Unden et al. (1980), with modifications described by Lancaster et al, (1999). Unless stated otherwise, all purification steps are performed at 0°C, and all buffers are flushed with N2 and contain 0.2 mM phenylmethylsulfonyl fluoride (from a 0.5 M stock solution in DMSO) to reduce protease activity. 1. Cell Homogenate The frozen bacteria are thawed and washed with 50 mM Tris-acetate, pH 7.35, and centrifuged at 9000 g for 20 min. The pelleted cells are resuspended (10 g protein/1) in 50 mM Tris-acetate, pH 7.35,1 mM dithiothreitol (DTT), 2 mM malonate, and passed through a French press at 130 MPa at flow rate of 10 ml/min. 2. Triton X-100 Extract The membrane fraction is precipitated from the cell homogenate by centrifugation at 100,000 g and 4°C for 45 min and resuspended in 50 mM Tris-acetate, pH 7.35, 1 mM DTT, and 2 mM malonate with Triton X-100 at a detergent/protein ratio of 1:1 (w/w). After stirring for 1 hr at room temperature under nitrogen and further centrifugation at 100,000 g and 4°C for 45 min, the Triton X-100 extract (supernatant) is separated from unsolubilized material. 3. DEAE Anion Exchange Chromatography A DEAE CL-6B column (300 ml volume) is equilibrated with 50 mM Trisacetate, pH 7.35, 1 mM DTT, 2 mM malonate, and 0.05% Triton X-100. The Triton X-100 extract is applied and the column is washed with the equilibration buffer. A linear NaCl gradient (0-200 mM) in this buffer is applied at a flow velocity of 80 ml/h, and QFR is eluted at a NaCl concentration of approximately 100 mM. Fractions containing QFR are identified by their characteristic red-brown color and their enzymatic activity (see point 5 below). On a second DEAE CL-6B column, Triton X-100 is replaced by a mixture of 0.05% dodecyl-j8-D-maltoside/0.05% decyl-j8-D-maltoside. 4. Preparative Isoelectric Focusing The enzyme is then subjected to isoelectric focusing. The preparative flat-bed electrofocusing procedure for the preparation of photosystem I, described by Tsiotis (1994, see also Chapter 1/6 in this edition), is used with the following modifications (Lancaster et ah, 1999): the pH range of the ampholytes is 5-8 (servalyt 5-8), and dodecyl-j8-D-maltoside (0.05%) and decyl-j8-D-maltoside (0.20%) are added to the ampholine solution. In more recent work (Lancaster

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et ah, 2000), we have lowered the detergent concentrations to 0.01% dodecyl-j8D-maltoside and 0.1% decyl-jS-D-maltoside. The gel is poured from a suspension of 8 g of Ultrodex (Pharmacia) in 200 ml of a 2% ampholine solution and dried at 60°C for approximately 2^ hrs until 30% evaporation is reached. Under these conditions, the maximum sample volume is 15 ml, and not more than 150 mg of QFR should be applied. After 12-16 hours of isoelectric focusing at 16 W and 5°C, the gel band containing QFR (at pH = 6, see Fig. lA) is cut out and suspended in a buffer (pH 7.3) containing 0.01% dodecyl-jSD-maltoside, 0.1% decyl-jS-D-maltoside, 20 mM HEPES, 1 mM EDTA, and 20 mM malonate or other dicarboxylate of interest. After sedimentation of the gel material by gentle centrifugation (1000 g, 10 min), the gel extraction procedure is repeated three times. The pooled supernatants are concentrated in Centrisart (300 kd cutoff) microconcentrators (Sartorius) to a volume of less than 2.5 ml. The ampholytes are then removed by gel filtration on PD-10 columns (Pharmacia). The absorbance at 415 nm is used to calculate the QFR concentration for crystallization (A415 = 21 corresponds to 10 m g / m l QFR). In the absence of fumarate as the dicarboxylate, the ratio of absorbances at 280 nm and 415 nm is used to monitor the comparability of successive preparations and should be between 0.95 and 1.2 for successful crystallization attempts. 5. Activity Measurements of Quinol: Fumarate Reductase a. The following protocol measures succinate oxidation activity using methylene blue as electron acceptor (Unden et ah, 1980). This activity is independent of QFR subunit C, the integral membrane diheme cytochrome h. 1. Prepare an anaerobic reaction buffer (pH 7.9, 37°C) containing 990/il of 10 mM Tris-HCl, 0.2 mM methylene blue, 10 mM succinate. 2. Start the reaction by addition of 10 /^l enzyme sample. 3. Record the reduction of methylene blue by succinate at 578 nm (£ = 17.1mM-^ cm"^). A typical specific activity for the wild-type enzyme is 28.8 U / m g (Lancaster et ah, 2000), with 1 U corresponding to 1 /imol of succinate oxidized per minute and the protein concentration determined using the biuret method with KCN (Bode et ah, 1968). b. The following protocol measures quinol: fumarate reductase activity using dimethyl-naphthoquinol (DMNH2) as electron donor (Unden et ah, 1980). 1. Prepare an anaerobic reaction buffer (pH 7.4, 37°C) containing 960 /il 50 mM potassium phosphate. 2. Add 10 )ul 20 mM dimethylnaphthoquinone (DMN, in ethanol). 3. Add 10 jul 5 m g / m l KBH4 for quinone reduction. 4. Add 10 jul of enzyme sample.

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FIGURE 1

A. Preparative isoelectric focusing of quinol: fumarate reductase. Shown is a 200-ml gel after application of 150 mg of membrane protein and 14 hrs of electrofocusing at 16 W and 5°C. The QFR band is found at pH 6 (lower arrow); impurities are frequently found at p H 7 (see upper arrow) and p H 6.5. In addition Coomassie staining of a corresponding analytical isoelectric focusing gel (not shown) indicates the removal of an impurity at p H 5.5, B. Crystal forms " A " and " B " of QFR are morphologically hard to distinguish. C. In the case of crystal form " C " of QFR, the crystals are more rod-shaped, less plate-like, compared to the two previous crystal forms. D. Crystal packing in crystal form " A " (a = 85.2 A, b = 189.0 A, c = 117.9 A, and p = 104.5°). The Ca traces of the A, B, and C subunits are colored in blue, red, and green, respectively. The ten green helices indicate the 10 transmembrane helices of the QFR dimer. Top: The ac plane. The non-90° angle is enclosed by the two shorter cell axes. Bottom: The be plane. E. Crystal packing in crystal form " B " (a = 118.4 A, (See also color plate.)

CHAPTER 13 Crystallization of Wolinella succinogenes QuinoLFumarate Reductase 5. Start the reaction by the addition of 10 ju\ 100 mM fumarate. 6. Record the oxidation of DMNH2 by fumarate as the absorbance difference at 270 and 290 nm (A£(oxidized-reduced) = ^5.2 mM"^cm"^) using a dual wavelength spectrophotometer. A typical specific activity for the wild-type enzyme is 7.4 U / m g (Lancaster et al, 2000).

III. CRYSTALLIZATION OF QUINOL: FUMARATE REDUCTASE The sitting drop method is used with crystallization stages from the W. Jockel company, Weiterstadt, Germany, and crystallization chambers from Boll, Munich. The fully oxidized QFR from W. succinogenes is crystallized in the presence of 1 mM KsFeCCN)^, 0.01-0.05% dodecyl-jS-D-maltoside and, 0.10-0.20% decyl-j8-D-maltoside, 1.2-2.4% benzamidine, 5-6% polyethylene glycole (PEG-3350), 75 mM NaCl, 5-6% dimethylformamide (DMF), 1 mM dimethylnaphthoquinone (DMN), 1-10 mM malonate, fumarate, or other dicarboxylate, and 10 mM HEPES/10 mM citrate, pH 6.4, at a protein concentration of 9.5-15 m g / m l in a 40-50 fA droplet. This is equilibrated by vapor diffusion against an 8 ml reservoir containing 150 mM NaCl, 10% PEG-3350, and 20 mM citrate, pH 5.6. Alternatively, the reservoir buffer can be replaced with 100 mM NaCl, 12% PEG-4000, and 100 mM citrate, p H 5.6, with half of these concentrations present in the protein droplet and the omission of DMF. In practice, this setup is achieved by performing the following steps. 1. Add the reservoir buffer to the crystallization chamber. 2. Prepare a double-concentrated protein solution, where all components not present in the reservoir buffer are present at twice their final concentration. This usually involves further concentration of the QFR sample with CentriSart microconcentrators (300 kd cutoff). 3. Mix equal volumes of the double-concentrated QFR solution and the reservoir buffer in an Eppendorf tube. 4. Spin down any precipitate at maximum speed in an Eppendorf centrifuge (2 min, 4°C). 5. Swiftly pipette the supernatant into the depression wells of the crystallization stages. 6. Close and seal the crystallization chamber.

Figure 1 Continued, b = 85.1 A, c = 188.9 A, and j? = 96.5°.). Top: The ah plane. Bottom: The ac plane. The non-90° angle is enclosed by the two longer cell axes. F. Crystal packing of crystal form "C" (a = 81.1 A, b = 290.2 A, c = 153.6 A, and P = 95.7°). In the he plane. Two QFR dimers (in color) form the asymmetric unit.

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Nucleation is enhanced by applying the microseeding technique (McPherson, 1999). Crystals from previous crystallization attempts are sonicated for 30 s in harvesting buffer containing 1 mM KsFeCCN)^, 0.01% dodecyl-^D-maltoside, and 0.10% decyl-jS-D-maltoside, 1.0% benzamidine, 12% polyethylene glycole (PEG-3350), 150 mM NaCl, 6% dimethylformamide (DMF), 1 mM dimethylnaphthoquinone (DMN), and 20 mM citrate pH 6.0 (modified from supplementary material to Lancaster et aL, 1999), using a microtip and a Branson sonifier 250 (output level 6, duty cycle 50%). The material is then diluted appropriately (1:10 to 1:10^), and 1 pA. of the respective dilution is carefully added to the protein droplet. The best crystals are not visible after a few days and take several weeks to grow (see Fig. IB and IC).

IV. CONCLUSIONS QFR crystals grown by the procedure described above were of sufficient quality to allow for the collection at 4°C of complete x-ray diffraction data sets from individual crystals. This proved to be very important for solving the phase problem, because it allowed the reliable incorporation of the anomolous signals arising from the 11 native Fe centers per QFR monomer into the phase calculation. Although a number of heavy-atom derivative data sets could be collected, their phasing power was insufficient due to the binding of mostly only one heavy atom per 130 kd QFR monomer (supplementary material to Lancaster et ah, 1999). The incorporation of the Fe anomalous signal for the crystal form "A" data resulted in an experimental electron density map at 2.2 A, which was easily interpretable for large parts of the QFR complex (Lancaster et ah, 1999). When comparing the subtle differences in packing for crystal forms "A" and "B" (Fig. ID and IE), it is understandable that these two forms are not easy to discriminate morphologically and sometimes coexist in the same crystal. Apart from the interactions between the two C subunits within one QFR dimer, which is noncrystallographic, there are no packing interactions between C subunits in the crystal forms A and B. An imaginative membrane bilayer can be fitted through all C subunits of one QFR row in the crystal, a one-dimensional feature reminiscent of the corresponding two-dimensional feature in type I membrane protein crystals. Compared to these two crystal forms, the packing in crystal form "C" is very different. First, rather than having one QFR dimer in the asymmetric unit, as is the case for the first two crystal forms, this crystal form contains two QFR dimers—that is, 520 kd—in the asymmetric unit (shown in color in Fig. IF). It is not possible to fit an imaginative membrane bilayer through one QFR row in the crystal, and there are crystal packing contacts between C subunits. This very different crystal packing has its

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consequences for the relative orientation of the FAD-binding domain and the capping domain in subunit A, which in turn has important consequences for the interpretation of the structure with respect to the mechanism of fumarate reduction (Lancaster et ah, 2001).

REFERENCES Bode, C , Goebell, H., and Stahler, E. (1968). Zur Eliminierung von Triibungsfehlern bei der Eiweissbestimmung mit der Biuret-Methode. Z. Klin. Chem. Klin. Biochem. 6, 419-422. Bronder, M., Mell, H., Stupperich, E., and Kroger, A. (1982). Biosynthetic pathways of Vibrio succinogenes growing with fumarate as terminal electron acceptor and sole carbon source. Arch. Microbiol. 131, 213-223. Cast, P., Hemelrijk, P., and Hoff, A. J. (1994). Determination of the number of detergent molecules associated with the reaction center protein isolated from the photosynthetic bacterium Rhodopseudomonas viridis. Effects of the amphiphilic molecule, 1,2,3-heptanetriol. FEBS Lett. 337, 39-42. Hagerhall, C. (1997) Succinate:quinone oxidoreductases. Variations on a conserved theme. Biochim. Biophys. Acta 1320, 107-141. Iverson, T. M., Luna-Chavez, C , Cecchini, G., and Rees, D. C. (1999). Structure of the Escherichia coli fumarate reductase respiratory complex. Science 284, 1961-1966. Kroger, A. (1978). Fumarate as terminal acceptor of phophorylative electron transport. Biochim. Biophys. Acta 505,129-145. Kroger, A., Geisler, V., Lemma, E., Theis, P., and Lenger, R. (1992). Bacterial fumarate respiration. Arch. Microbiol 158, 311-314. Kroger, A., Geisler, V., and Duchene, A. (1994). Isolation of Wolinella succinogenes hydrogenase. Chromatofocusing. In ''A Practical Guide to Membrane Protein Purification'' (G. von Jagow and H. Schagger, eds.), pp. 141-148. Academic Press, San Diego, California. Lancaster, C. R. D., Kroger, A., Auer, M., and Michel, H. (1999). Structure of fumarate reductase from Wolinella succinogenes at 2.2 resolution. Nature 402, 377-385. Lancaster, C. R. D., Gross, R., Haas, A., Ritter, M., Mantele, W., Simon, J., and Kroger, A. (2000). Essential role of Glu C66 for menaquinol oxidation indicates transmembrane electrochemical potential generation by Wolinella succinogenes fumarate reductase. Proc. Natl. Acad. Sci. U.S.A. 97, 13051-13056. Lancaster, C. R. D., Gross, R., and Simon, J. (2001). A third crystal form of Wolinella succinogenes quinol: fumarate reductase reveals domain closure at the site of fumarate reduction. Eur J. Biochem. 268, 1820-1827. McPherson, A. (1999). ''Crystallization of Biological Macromolecules." Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Michel, H. (1983). Crystallization of membrane proteins. Trends Biochem. Sci. 8, 56-59. Michel, H. (2001). Crystallization of membrane proteins. In "International Tables of Crystallography Volume F, Macromolecular Crystallography" (M. G. Rossmann and E. Arnold, eds.). Chapter 4.2. Kluwer Academic Publishers, Dordrecht, pp. 94-99. Ohnishi, T., Moser, C. C , Page, C. C , Dutton, P. L., and Yano, T. (2000). Simple redox-linked proton-transfer design: New insights from structures of quinol-fumarate reductase. Structure 8, R23-R32.

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C. Roy D. Lancaster Timmins, P. A., Hauk, J., Wacker, T., and Welte, W. (1991). The influence of heptane-l,2,3-triol on the size and shape of LDAO micelles. Implications for the crystallization of membrane proteins. FEES Lett. 280, 115-120. Tsiotis, G. (1994). Photosystem I from cyanobacteria isolated in crystallizing form by preperative isoelectric focusing. In "A Practical Guide to Membrane Protein Purification'' (G. von Jagow and H. Schagger, eds.), pp. 149-159. Academic Press, San Diego, California. Unden, G., Hackenberg, H., and Kroger, A. (1980). Isolation and functional aspects of the fumarate reductase involved in the phosphorylative electron transport of Vibrio succinogenes. Biochim. Biophys. Acta 591, 275-288.

14

jBa3-Type Cytochrome c Oxidase from Thermus thermophilus: Purification, Crystallization and Crystal Transformation TEWFIK SOULIMANE Rheinisch-Westfalische Technische Hochschule Aachen, Institut fur Biochemie Pauwelsstrasse 30, D-52057 Aachen, Germany Current address: Paul Scherrer Institut, Life Sciences, OSRA/OO? CH-5232 Villigen PSI, Switzerland REINER KIEFERSAUER Max-Planck-Institut fur Biochemie Am Klopferspitz 18a 82152 Planegg-Martinsried, Germany Proteros Biostructures GmbH Am Klopferspitz 19 82152 Planegg-Martinsried, Germany MANUEL E. THAN Max-Planck-Institut fur Biochemie Am Klopferspitz 18a 82152 Planegg-Martinsried, Germany

I. INTRODUCTION Cytochrome c oxidase (cytochrome-c-02-oxidoreductase; EC 1.9.3.1), the terminal complex of the respiratory chain of mitochondria and many bacteria, catalyzes the electron transfer from cytochrome c to molecular oxygen, thereby reducing the latter to water. The energy liberated in this exergonic reaction is invested into the generation of an electrochemical proton gradient across the inner mitochondrial or bacterial membrane. The stored free energy is then employed by the FoFi-ATP-synthase for the synthesis of ATP from ADP and inorganic phosphate, the main energy source of most biological reactions (for reviews see Michel et al., 1998; Fergusson-Miller and Babcock, 1996).

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Thermus thermophilus HB8 (ATCC 27634) is an extremely thermophilic gram negative bacterium. It grows at temperatures between 47° and 85°C. Depending on the fermentation conditions and the 02-supply, this organism expresses two different oxidases known as the caa^- and the ba ^-type (Fee et al., 1980; Zimmermann et ah, 1988). The f7fl3-oxidase from T. thermophilus is a monomeric enzyme consisting of three subunits (I, II, and Ila) as well as two heme irons and three coppers (Soulimane et ah, 2000a). The biochemical, biophysical, and structural properties of this aberrant member of the hemecopper oxidase superfamily have been summarized in a recent review by Than and Soulimane (2001). We here describe a new procedure for the preparation of monodispersed highly active t'flj-type cytochrome c oxidase from Thermus thermophilus. The enzyme is isolated from the membrane using Triton X-100 and purified by subsequent anion-exchange chromatographies first in Triton X-100 and then in dodecyl-jS-D-maltoside. The preparation is finished within seven days, yielding approximately 100 mg pure enzyme starting from 100 g biomass. Crystallization experiments were set up after an exchange of the dodecyl-j8-Dmaltoside to a series of different detergents, yielding crystals of different space groups and quality. The best crystals were grown in nonyl-jS-D-glucoside using PEG 2000 as precipitant. However, these crystals show drastic changes in their cell constants if measured in a standard capillary and a relatively weak diffraction power in their native (as grown) state. In order to obtain a useful dataset, it was necessary to transform these crystals by dehydration. The first water-mediated transformations of protein crystals were already investigated in the early years of protein crystallography. Kendrew and Perutz produced different crystal forms of hemoglobin by connecting the capillary, in which the crystal was mounted, to different types of salt reservoir (Bragg and Perutz, 1952; Huxley and Kendrew, 1953). After a long gap the first example of a successful crystal transformation from the low- to the highresolution form was described by Schick and Jurnak (1994), Increasing the PEG concentration in the storage buffer resulted in a shrinkage of the crystal, which was explained as the displacement of crystalline water by PEG. With the enhanced use of the cryotechnique, accompanied by the increase of post-growth crystal treatments, other examples of successfully transformed crystals followed (Cramer and Miiller, 1997; Cramer et al., 2000; Esnouf et al., 1998; Fethiere et al, 2000; Izard et al, 1997; Petock et al, 2001; Qian et al, 2000; Soulimane et al, 2000a; Stammers et al, 1994; Tahirov et al, 1998; Weiss and Hilgenfeld, 1999). In our case, dehydration was achieved either by using the steady evaporation of crystal solvent through an oil layer that surrounds the crystal or by generating a crystal environment of tightly regulated humidity using a novel humidity apparatus. The use of such a machine for improving the crystal quality was shown on different crystal probes (Estebanez-Perpina et al, 2000; Kiefersauer, 1998; Kiefersauer et al, 2000). This crystal shrinkage by dehydra-

CHAPTER 14 Bflg-Type Cytochrome c Oxidase from Thermus thermophilus

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tion resulted for the ba ^-oxideise in an increase in diffraction power by about 1A and a diffraction limit of about 2.1 A using synchrotron radiation. Interestingly, the kinetics of dehydration is important for the course of the unit cell shrinkage, resulting either in preferred states of the crystal or in the complete breakdown of the crystalline lattice. Successful flash-ifreezing of the crystals, which is necessary for data collection at the synchrotron, is only possible after transformation to one of the preferred states at lower solvent content. The methodological aspects of different crystal transformation procedures are described in detail.

11. FERMENTATION, PURIFICATION, AND MOLECULAR CHARACTERIZATION A. Fermentation Fermentation of Thermus thermophilus HB8 (ATCC 27634) has been performed at the Gesellschaft fiir Biotechnologische Forschung (GBF) Braunschweig. For the expression of ba^-cytochrome oxidase, the cells were grown at 70°C in 100 1 of culture medium using a stainless steel jar fermenter under restricted oxygen conditions 0.05 W~^ min"^ (volume air per volume medium per minute) according to Castenholz (1969), yielding about 500 g biomass. The cells were harvested in the early to middle exponential growth phase and stored at — 80°C.

B. Preparation Protocol 1. Lysozyme Digest: Preparation of Spheroblasts 1. Suspend 100 g frozen cells in 500 ml of 0.25 M Tris-HCl, pH 7.6, containing 200 mM KCl at room temperature by stirring for 30 min. 2. Homogenize for 1 min at full speed in a Braun (type MX-32) homogenizer. 3. Add 400 mg lysozyme to the suspension and keep it under stirring for 120 min. 4. Centrifuge the suspension containing the broken cells at 17700 xg (Beckman J2-21 centrifuge; JA-10 rotor, 10,000 rpm) for 30 min at 4°C. 5. Discard the supernatant and resuspend the pelleted spheroblasts in 500 ml of 100 mM Tris-HCl, pH 7.6. Note: The supernatant (step 5) can be used for the purification of cytochrome C552, the substrate of the ba^-oxidase, according to Soulimane et ah (1997). The crystal structure of this basic protein has been solved at 1.28 A resolution using MAD-phasing (Than et al, 1997).

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2. Solubilization of Cytochromes and Chromatography on DEAE-Biogel 1. Pack the anion exchanger DEAE-Biogel Agarose (Biorad, Richmond CA 94804) into a 20 x 10 cm column and equilibrate it with 5 1 of 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100. 2. Adjust the suspension from step I,B/l/5 to 5% Triton X-100 for the solubilization of the membrane proteins comprising the spectroscopically visible respiratory chain complexes of T. thermophilus. 3. Keep the suspension under stirring for 3 hrs at 4°C. 4. Centrifuge the suspension at 17700 x g (Beckman J2-21 centrifuge; JA-10 rotor, 10,000 rpm) for 60 min at 4°C. 5. Discard the pellet, dilute the supernatant with 5 1 H2O, and apply the solution onto the DEAE-biogel column. 6. Wash the column containing the firmly bound ba^-oxideise and other cytochromes with 3 1 of 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100 and then elute the protein with 4000 ml of a linear gradient from 0 to 250 mM NaCl in Tris-HCl, pH 7.6 and 0.1% Triton X-100 at a flow rate of 120 m l / h (see Fig. 1). Note: The stirring of the sphearoblast suspension in Triton X-100 for 3 hrs is important for the quantitative solubilization of the te3-oxidase. Also, the washing step of the column with 3 1 of 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100 is important for the complete removal of a yellow pigment. The reduced-minus-oxidized difference spectra of the first peak (fractions 70-150; Fig. 1) indicate the presence of te3-oxidase. Furthermore, fractions containing the cafl3-cytochrome c oxidase (peak 2) as well as other complexes of the respiratory chain from T. thermophilus, such as cytochrome C548/554/ cytochrome ??562/ or complex II (peak 3) are observed. The purification of the te3-oxidase is continued by an anionic exchange chromatographic step on DEAE-Sepharose CL-6B. 3. Chromatography on DEAE Sepharose CL-6B 1. Pack the anion-exchange DEAE-Sepharose CL-6B (Pharmacia) into a 10 X 3 cm column and equilibrate it with 2 1 of 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100. 2. Collect the fractions containing the spectroscopically visible ba^-oxideise from step II,B,2 (peak 1; Fig. 1) and concentrate the solution to a volume of 30 ml using the Minitan concentrator (30 kD membrane; Millipore, Eschborn). 3. Dilute the to3-oxidase solution to a conductivity of 5 mS with 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100 and apply it onto the DEAE-Sepharose CL-6B.

233

CHAPTER 14 Bfl3-Type Cytochrome c Oxidase from Thermus thermophilus 0,45

100

FIGURE 1

150

200

Fraction

250

300

350

Elution profile of the solubilized ^flg-cytochrome c oxidase from T. thermophilus on a DEAE-Biogel column. The enzyme was eluted with 0-200 mM sodium chloride. The volume of each fraction is 10 ml. Peaks 1 and 2 represent the fractions containing the ba^-type and caa^-type cytochrome-c oxidases, respectively. Peak 3 represents a mixture of different cytochromes including cytochrome C554/543/ and cytochrome b^^2' as well as complex II.

4. Wash the column containing f7a3-oxidase with 1 1 of 10 niM Tris-HCl, pH 7.6 containing 0.1% Triton X-100 and then elute the enzyme writh 2000 ml of a linear gradient from 0 to 250 mM NaCl in Tris-HCl, p H 7.6 and 0.1% Triton X-100 at a flow rate of 120 m l / h . Note: The ba^-oxidase is at this stage spectroscopically pure. Analysis by SDS-PAGE, however, demonstrates the presence of protein impurities. The ba ^-oxidase is further purified by two subsequent anion exchange chromatography steps using fast protein liquid chromatography (FPLC). 4. Fast Protein Liquid Chromatography (FPLC) 1. Pack the anion-exchange Fractogel EMD TMAE-650 (S) (MERCK Darmstadt) into a 10 X 0.5 cm column and equilibrate it with 200 ml 10 mM

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2. 3. 4.

5.

Tris-HCl, pH 7.6 containing 0.1% Triton X-100 using FPLC (LKB 2150 Ultrochrom GTi). Collect the fractions containing bfl3"Oxidase from step II,B/3 and concentrate the solution to a volume of 10 ml using the Minitan concentrator (30 kD membrane; Millipore, Eschborn). Dilute the ba^-oxidase solution to a conductivity of 5 mS with 10 mM Tris-HCl, pH 7.6 containing 0.1% Triton X-100 and apply it onto the Fractogel EMD TMEA-650 (S) column. Wash the column containing ba^-oxideise with 300 ml of 10 mM Tris-HCl, pH 7.6 now containing 0.05% dodecyl-jS-D-maltoside and then elute the enzyme with a linear gradient of 0-250 mM NaCl in Tris-HCl, pH 7.6 and 0.05% dodecyl-jS-D-maltoside at a flow rate of 2 ml/min. Repeat steps 1 to 4 using dodecyl-jff-D-maltoside.

Note: The exchange of Triton X-100 can be controlled by following its absorption at 280 nm using a UV-detector. Complete purity, as judged from a series of analytical experiments, is obtained after a gelfiltration step on a Superdex 200-column in a buffer containing 0.05% dodecyl-jS-D-maltoside. 5. Gelchromatography on Superdex 200 1. EquiUbrate a Superdex 200 column (Pharmacia) with 50 mM Tris-HCl, pH 7.6 containing 0.05% dodecyl-j8-D-maltoside. 2. Concentrate the collected ba^-oxidase fractions from step II,B,4 by ultrafiltration (Centricon 30, Amicon) and apply 0.5 ml (20 mg) of the protein onto the Superdex 200 column. 3. Elute the enzyme with 50 mM Tris-HCl, pH 7.6 containing 0.05% Dodecyl-jS-D-maltoside. Note: The ba^-oxidase fractions were collected, concentrated by ultrafiltration (Centricon 30000; Amicon), and used for spectroscopical and protein chemical investigations and crystallization experiments.

C. Molecular Characterization Analytical characterizations from the different purification steps are summarized in Table I. Starting from 100 g wet biomass obtained after growth under restricted aerobic conditions, about 100 mg of the ba^-enzyiae are routinely isolated. In relation to the amounts present after the first DEAEchromatography the yield of the purified enzyme is 72%. The heme A/protein ratio is 11.9 nmol heme A/mg protein, approximately equal to the theoretical value of 11.8 based on the molecular mass of 84.884 Da for the protein monomer.

CHAPTER 14 Bag-Type Cytochrome c Oxidase from Thermus thennophilus TABLE I

235

Purification of bag-Cytochrome c Oxidase from Thermus thennophilus Starting from 100^ Biomass

Solubilization DEAE-Biogel DEAE-Seph. l.FPLC 2. FPLC Gelfiltration

Total Protein

Heme A''

Protein^

H/P

mg

nmol

mg

nmiol/mg

1450 1341 1198 1115 1050

217 139 110 98 88

Yield

9257 6.5 9.6 10.8 11.4 11.9

100 92 83 77 71

" Heme A concentrations were obtained from the dithionite reduced-minus-oxidized spectrum at 612 nm using a molar absorption coefficient of 6300 M~^ cm"^ (Zimmermann ei al, 1988). Absorption spectra of cytochrome c oxidase were recorded with a Perkin Elmer Lambda 5 UV/VIS spectrophotometer. ^ Protein concentration and amino acid content have been determined by amino acid analysis on a Biotronik amino acid analyser LC 5001 (Eppendorf-Nethleler-Hinz, Maintal, Germany) equipped with a fluorescence detection of o-phtalaldehyde-derivatives. The protein sample was hydrolyzed in 5.7 N HCl containing 0.2% thioglycolic acid for 90 min at 150°C. Tryptophan was determined after hydrolysis in 5.7 N HCl containing 5% thioglycolic-acid for 90 min at 150°C. The cysteine content was determined after oxidation of the entire protein with performic acid to cysteinsulfonic acid before hydrolysis.

SDS/PAGE of the purified enzyme reveals after denaturation in SDS and staining with Coomassie blue the presence of the well-known subunits I and 11. The subunit Ila, which has been recently identified during the crystal structure determination (Soulimane et ah, 2000a, b), cannot be observed. Using reversed phase high-performance liquid chromatography (RP-HPLC), all three subunits (I, II, and Ila) of the tej-oxidase have been detected and identified as part of the enzyme (Fig. 2; Soulimane et ah, 2000b). The three subunits, I, II, and Ila, consist of 562 residues (62 591 Da), 168 residues (18 499 Da), and 34 residues (3 794 Da), respectively. The quantitative Edman degradation of the purified enzyme exclusively releases the known N-terminal residues of subunit II. Subunit I and subunit Ila are blocked at their N-terminus. No impurities were detected. Homogeneity of the preparations of the terminal oxidase was tested by gelfiltration chromatography. The enzyme elutes as single symmetric peak upon gelfiltration chromatography on Superdex 200 and is therefore considered to be monodisperse. The te3-type oxidase elutes with an apparent molecular weight of 90 kDa. The molecular weight fits nicely to the theoretical molecular mass derived from the sequence of the three subunits assumed to constitute the monomer.

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m FIGURE 2

.LL.

5

Separation of the ?7fl3-type cytochrome c oxidase subunits by reversed phase-HPLC. The polypeptides were purified on a C4 column (250 x 4,6 mm) using a HewlettPackard 1050 HPLC System with multiple wavelength detector and the following solvents: (A) 5% formic acid, (B) 95% formic acid, (C) n-propanol, and (D) acetonitril. Gradients were formed with a microprocessor-controlled quaternary p u m p (HP) by low-pressure mixing of four solvents in 60 minutes: (A) 14% to 0%, (B) 70% = constant, (C) 6% to 30%, and (D) 10% to 0% (by vol.). Peaks 1-6: 1. injection peak; 2. nonseparated enzyme; 3. subunit II; 4. heme; 5. subunit Ila; 6. subunit I. © Reprinted from Frot. Sci. 9, Soulimane, T., Than, M. E., Dewor, M., Huber, R. and Buse, G. Primary structure of a novel subunit in t>a3-cytochrome oxidase from Thermus thermophilus. 2068-2073, (2000b), with permission of Springer (2000).

III. CRYSTALLIZATION AND INITIAL CRYSTALLOGRAPHIC CHARACTERIZATION A. Detergent Exchange Crystallization of ba ^-oxidase w^as achieved using sitting-drop and the batch method in different detergents. The detergent exchange has been performed as foUov^s (for the detergents tested for crystallization setups, see Table II). 1. Pack the anion exchange Fractogel HMD TMAE-650 (S) (MERCK Darmstadt) into a 10 x 1 cm column and equilibrate it with 100 ml 10

237

CHAPTER 14 Bflg-Type Cytochrome c Oxidase from Thermus thermophilus

TABLE II

Summary of the Crystallization Trials Using Different Detergents CMC (mM)

Group of detergents

Name

M r (g/mol)

Alkyl-p-D-glucoside

Octyl-jS-D-glucoside Nonyl-j5-D-glucoside Decyl-j5-D-glucoside Cyclohexylpropyl-jg-D-glucoside Octyl-j5-D-thioglucoside Nonyl-jS-D-thioglucoside Nonyl-j?-D-maltoside Decyl-j5-D-maltoside Dodecyl-i?-D-maltoside Cyclohexylpropyl-j5-D-maltoside Nonyl-j5-D-thiomaltoside MEGA-9 HEGA-9

292 306 320 304 308 322 468 483 511 466 484 336 366

3.2 25 39

N-dodecyl-N,N'-dimethyl-amineN-oxide (LDAO)

229

1.4

Alkyl-p-D-thioglucoside Alkyl-p-D-maltoside

Alkyl-p-D-thiomaltoside Alkanoyl-N-methylglucamide (MEGA-w) Alkanoyl-N-hydroxyethylglucamide (HEGA-w) N-Alkyl-N,N'-dimethylamin-N-oxide

30.3

6.5 2.6 28 9 2.9 6 1.6 0.15 34.5

Crystal (8.0 A^) (2.7 A^) (8.0 A'') (3.0 A«) (3.8 A'') (2.7 A')

— — — — — (8 A'')

— (10 A«)

The resolution of the crystals obtained is given in parentheses: " using a rotating anode x-ray generator and ^ using synchrotron radiation; CMC, critical micellar concentration (as provided by the vendor).

2. 3. 4. 5.

mM Tris-HCl, pH 7.6 containing 0.05% dodecyl-j?-D-maltoside using FPLC. Apply the purified ?7fl3-oxidase (10 mg) in 50 mM Tris-HCl, pH 7.6 containing 0.05% dodecyl-jS-D-maltoside. Wash the column with 200 ml of 10 mM BisTris-HCl, p H 7.0 containing the new detergent at a concentration of twice the CMC (Table II). Elute the enzyme with 10 mM BisTris-HCl, p H 7.0 and 100 mM KCl containing the new detergent at a concentration of twice the CMC. Concentrate the sample to about 1 ml corresponding to a concentration of 8-10 m g / m l by ultrafiltration (Centricon 30 kD, Amicon) and use the enzyme for crystallization experiments.

Note: If the complete sample is not needed for crystallization experiments, then the rest of the enzyme can be frozen in 100 fil aliquots in liquid nitrogen and stored at - 7 4 ° C .

B. Crystallization in Sitting-Drops 1. Prepare (i) a solution of 14% polyethylene glycol 2000 in 30 mM BisTris-HCl, pH 7.0, without detergent; (ii) the enzyme sample in 10 mM BisTris-HCl, pH 7.0 containing detergent in a concentration of

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2. 3.

4. 5. 6.

twice the CMC (8-10 mg/ml); and (iii) a Cryschem MVD24 multichainber plate for vapor diffusion crystallization on individual sitting drops. Mix in a separate Eppendorf tube 50 jx\ ha^-enzyvcve in 10 mM BisTrisHCl, pH 7.0 and 100 mM KCl with 25 /xl of 14% polyethylene glycol 2000 in 30 mM BisTris-HCl, pH 7.0 (precipitating agent). Fill up the reservoir chamber of each Cryschem well with 1 ml of 14% polyethylene glycol 2000 in 30 mM BisTris-HCl, pH 7.0 and the plastic post holding the protein sitting drop with 6 /zl of the protein/ precipitant mixture. Seal the plate with a special tape that is supplied by the manufacturer. Incubate the plate at 20°C. Crystals appear after an incubation time of 1-4 weeks.

Note: Filter all solutions with a microfilter of 0.22 /im. The Cryschem plate are commercially available (manufactured by Cryschem Inc. or C. Supper Company). Each well contains a plastic post in the center to hold the protein sitting drop. The detergent concentration in the protein preparation is always twice its CMC, and no detergent is added to the precipitating solution.

C. Crystallization in Batch 1. Mix in a 1 ml Eppendorf tube 50 jul ha^^-erxzyvcve in 10 mM BisTris-HCl, pH 7.0 and 100 mM KCl containing detergent in a concentration of twice its CMC with 25 piX of 16% polyethylene glycol 2000 in 30 mM BisTris-HCl, pH 7.0 (precipitating agent). 2. Incubate the mixture at 20°C. 3. Crystals appear after an incubation time of 1-4 weeks. Notes: Both crystallization methods produce crystals of equal quality but different sizes and forms (data not shown). The largest crystals were greater than 1.5 mm in size. Crystals with a size of about 0.5 m m were used for the crystal transformation experiments and structure determination. As shown in Fig. 3, crystals have been obtained using the detergents octyl-j?-D-glucoside and octyl-j8-D-thioglucoside diffracting to 8 A and 3.8 A, respectively (Figs. 3A and 3C; Soulimane et ah, 1995). Crystals have also been obtained with the detergents nonyl-j8-D-glucoside and nonyl-jS-D-thioglucoside (Figs. 3B and 3D) both diffracting to 2.7 A using synchrotron radiation. However, the crystals obtained with nonyl-jS-D-thioglucoside were twinned and the crystals obtained with nonyl-jS-D-glucoside show a variability in their cell constants during data collection in standard glass capillaries. Well-ordered crystals have also been obtained using cyclohexyl-propyl-jSD-glucoside diffracting to 3 A resolution. Crystals were also grown in nNonanoyl-N-methylglucamide (MEGA-9) and dodecyl-dimethylamineoxide (LDAO) diffracting to about 8 A. No crystals grew with the detergents containing a maltoside group and with detergents of the n-alkyl-oligoethylene

CHAPTER 14 Bflg-Type Cytochrome c Oxidase from Thermus thermophilus

FIGURE 3

239

?7fl3-Cytochrome c oxidase crystals from T. thermophilus grown by vapor diffusion using the sitting-drop-method with 30 mM Bis Tris-HCl, pH 7.0 and 16% polyethylene glycol 2000 as precipitant in the presence of (A) octyl-j5-D-glucoside, (B) nonyl-j5-Dglucoside, (C) octyl-jS-D-thioglucoside, and (D) nonyl-jS-D-thioglucoside as detergent. (See also color plate.) glycol-monoethers (C^E^) class. Twelve different detergents from this group were tested. The detergents tested for crystallization experiments as well as the resulting crystals are summarized in Table II.

D. Initial Crystallographic Characterization The native crystals of the ?7fl3-cytochrome c oxidase grown in the presence of nonyl-jS-D-glucoside (Fig. 3B) contain one molecule per asymmetric unit and belong to space group P432i2 with unit cell dimensions of a = b = 112116 A, c = 174-190 A, corresponding to a solvent content between 62% and

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Tewfik Soulimane et al.

68% (Matthews, 1968). They show a high inherent flexibiUty in their crystal packing and differ in their cell constants as well as their diffraction power significantly between different crystals. Even within a given dataset recorded either at room temperature, 16°C, or 4°C on capillary mounted crystals, the cell dimensions change by up to 10 A for the c-axis. No correlation to the experimental or environmental settings can be found for a given experiment. The limiting resolution of the diffraction images correlates to some degree with the cell constants and is between 7 and about 3 A using a rotating anode x-ray generator. The crystal size had to be at least 0.5-0.7 mm in order to observe a reasonable diffraction power. Crystals older than four to six weeks show reduced diffraction quality and are impossible to transform successfully (see below).

IV. CRYSTAL TRANSFORMATION Here, crystal transformation is used in the sense of changing the crystal quality by post-growth crystal treatment. Crystal quality comprises both the crystal form and the crystal order. In a protein crystal the packing has a very complex nature and can easily be changed by water-mediated crystal shrinkage. Compared to other methods, such as high-pressure experiments, it is a very effective method to reduce uniformly the distance between lattice points. Normally, crystal shrinkage results in a continuous degrading of the crystal order (see e.g., Dobrianov et al., 2001). Nevertheless in some cases the crystal order can be improved by crystal dehydration as monitored by x-rays. New crystal contacts can be formed, especially hydrogen bonds. The mechanism for this phenomena is at present not understood in detail. In the following sections, materials and methods are described to achieve a successfully crystal transformation, and a first analysis of different humidity stages of the te3-oxidase crystals is presented.

A. Oil Method The oil method for the crystal transformation uses the continuous evaporation of crystal solvent through an oil layer, which functions as a diffusion barrier. This leads to a steady dehydration of the crystals that is stopped by flash-freezing of the transformed crystal once it has reached its optimal diffraction behavior (Fig. 4). 1. Replace the mother liquor surrounding the crystal with an appropriate oil (e.g., Perfluoropolyether (PPPE) oil or a 9:2 mixture of paraffin oil and n-heptane). This can be done by carefully placing an oil layer on top of the crystal and removing the residual mother liquor using a small capillary and filter paper. 2. Mount the crystal in a standard cryo-loop onto the x-ray camera. 3. Follow the transformation behavior via diffraction images. Using synchrotron radiation as well as a CCD-detector gives a much better time

CHAPTER 14 Bflg-Type Cytochrome c Oxidase from Thermus thermophilus

241

B FIGURE 4

Transformation of ba^-oxideise crystals with the oil-method: The crystals show initially a very poor diffraction pattern that may be even worse than that of capillary-mounted crystals due to the initiation of the transformation (A). A few minutes later, the diffraction pattern improves (B), and the crystal was frozen, which leads again to an improvement in diffraction power of the crystal (C). All exposures were recorded without rotation of the crystal and an exposure time of 120 s, using a rotating anode x-ray generator. The edge of the image plate corresponds to a resolution of 3.0 A. © Reprinted from /. Appl Cryst., 33, Kiefersauer, R., Than, M. E., Dobbek, H., Gremer, L., Melero, M., Strobl, S., Dias, J. M., Soulimane, T., and Huber, R. A novel free-mounting system for protein crystals: Transformation and improvement of diffraction power by accurately controlled humidity changes. 1223-1230, (2000), with permission from International Union of Crystallography (2000).

resolution as compared to rotating anode generators and an image plate detector, w^hich might be essential to find the sometimes very short-lived optimal state. 4. A remote-controlled shutter placed previously onto the outlet of the cryostream system is opened when the crystal show^s optimal diffraction behavior leading to shock-freezing of the crystal and preservation of the crystal in this optimal state. 5. Record standard diffraction images of the frozen and thereby stabilized crystal according to the experimental protocol required. Note: The kinetics of the transformation is highly dependent on the nature of the oil used, as well as many environmental conditions (e.g., temperature and humidity of the surrounding air), and it may be difficult to control. For the &fl3-oxidase crystals, the transformation was performed within 10-15 min using a mixture of paraffin oil and n-heptane. It was necessary to follow the progress of the dehydration at the synchrotron with a reduced beam intensity in order to prevent radiation damage to the crystal during this period. During this steady evaporation of crystal solvent, the crystals go through several alternating states of higher and lower diffraction power (see below). The resulting frozen crystals that were analyzed for their crystal structure with MAD-methods (Soulimane et ah, 2000a) had unit cell constants of

242

Tewfik Soulimane et ah

dryair—•!=(

I ^ ^ I fc (i)

FIGURE 5

(iii)

(A) Schematic representation of the humidity machine. Shown is the humidity generator (I), the transport tube of the humid gas stream (II), and the crystal holder (III). (B) The crystal holder in more detail. The humid gas stream is fed into the crystal holder and flows through the gas channel to the crystal. The temperature of the outer part of the crystal holder (T„) is monitored by a PtlOO element. The inner part

CHAPTER 14 Bfl3-Type Cytochrome c Oxidase from Thermus thermophilus

243

a = b = 112.1 A, c = 161.4 A—in other words, significantly smaller than the capillary mounted "native crystals."

B. Transformation and Freezing with the Humidity Machine The water content in protein crystals is a function of the relative humidity (r.h.) of the surrounding atmosphere. The correlation of r.h. with dew point Tdp and temperature Tg of an arbitrary gas is described by the Magnus formula.

r.h. = exp< —a^b

T,-T,^ X 100% (b^ + T,^)(b^ + TJ_

(1)

with the constants a^ = 17.5043 and b^ = 241.2 K. From Eq. 1 follows that in practice a control of r.h. means the control of temperature. For the proper use of the machine, condensation has to be avoided. According to Eq. 1 this is guaranteed only if Tg > T^p. 1. The Humidity Apparatus The humidity apparatus can be divided into three functional parts (Fig. 5A). a. Humidity generator: Mixing dry air with a stream of air saturated with water produces a humid air-stream with dew point T^p (Kiefersauer et ah, 2000). The ratio of both flows is adjusted by valves. A useful range of the flow of the combined air-stream is 0.6-1.0 1/min.

Figure 5 continued, carries the micropipette or loop. The crystal holder does not interfere with the optical path for x-rays or light. Also, the combination with a cold gas-stream for cryoexperiments is possible. (C) Mounting the crystal onto the micropipette. The droplet containing the crystal is placed near the pin of the micropipette in the humid air stream (i). In the next step the pin is gently contacted with the crystal surface, and some mother liquor is sucked in the micropipette by capillary forces (ii). At this stage, the vacuum is switched on, and the crystal is sucked onto the tip. The loop holder is removed, and the crystal is free from solution (iii). These operations are done by a micromanipulator under a stereo microscope (M3Z, Leica, Switzerland). Panel 5C © Reprinted from /. Appl. Crysi., 33, Kiefersauer, R., Than, M. E., Dobbek, H., Gremer, L., Melero, M., Strobl, S., Dias, J. M., Soulimane, T., and Huber, R. A novel free-mounting system for protein crystals: Transformation and improvement of diffraction power by accurately controlled humidity changes. 1223-1230 (2000), with permission from International Union of Crystallography (2000).

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b. Transport of the humid gas: A flexible Teflon tube connects the humid air-stream to the crystal holder. The flexibility facilitates mounting and measurement of the probe at different locations. The tube has a length of 1 m and is 6 mm in diameter. To avoid condensation the tube is isolated and warmed above T^p. For this purpose a resistant wire is wrapped round the tube and connected to a power supply unit. c. Crystal holder: Important features of the crystal holder are its compactness, the possibility to combine x-ray, optical as well as humidity measurements without geometrically interference, and the use for shock-freezing of the sample (Fig. 5B). The crystal holder consists of an inner part, which is fixed on the goniometer head. It carries a micropipette or a standard cryo-loop for holding the crystal. The outer part forms the gas channel that is connected to the humid air stream. The humidity seen by the crystal also depends on the distance of the crystal relative to the outlet of the gas channel. It can be adjusted via a position screw. Typically the crystal is positioned 2 - 3 mm outside of the gas channel. The outer part is freely rotatable relative to the inner part without axial movement. Therefore, leads or supplies are not damaged when the insert is rotated during the measurement. The temperature of the crystal holder, T^r, is monitored with a PtlOO element. The temperature of the air stream, Tg, is adjusted to T^r by passing through the crystal holder (for more details, see Kiefersauer et ah, 2000). Note: Control of the humidity apparatus via a personal computer facilitates the processing of complex humidity time protocols like humidity gradients. 2. Mounting the Crystal Initially, the humidity has to be adjusted to correspond to the growth conditions of the crystal. Otherwise the crystal can be dissolved or damaged irreversibly. For this purpose the vapor pressure of the mother liquor of the crystals has to be determined. 1. Place a cryo-loop filled with mother liquor in front of the gas nozzle of the humidity apparatus. 2. Inspect the size of the droplet with time under different humidity settings. 3. If the size of this droplet stays constant, the current humidity corresponds to the water pressure in the mother liquor (97.5% r.h. for the tflj-oxidase crystals). The procedure for mounting the crystal depends on the type of crystal holder. 1. Loop-holder: At first the solution in which the crystal is stored has to be replaced by the carrier solution used for the loop (see Section

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IV,A,1). The crystal is then picked up with the loop, the loop is placed on the magnetic base, and the shells of the outer part of the crystal holder are closed (Kiefersauer et al., 2000). To allow faster equilibration of the crystal with the humid gas stream, the thickness of the layer of solution around the crystal should be as thin as possible. 2. Pipette-holder: The mounting of the crystal on the pin of the micropipette is demonstrated in Fig. 5C. The crystal is sucked u p by the capillary using a weak vacuum (0.1 bar). The micropipette is made similar to pipettes used by patch clamp technique. To get larger openings, the tip is broken off and ground to create a flat mounting surface for the protein crystal. The diameter of the tip should match the size of the crystal.

3. Freezing the Crystal For freezing and hence stabilizing crystals that were transformed with the humidity apparatus, the loop crystal holder is used. 1. Mount the crystal in a standard cryo-loop and place it onto the loop crystal holder of the humidity machine, according to Section IV,B,2. 2. Transform the crystal by changing the humidity according to an appropriate protocol and follow the transformation by diffraction images. The oil layer functions as an diffusion barrier such that the changes in the crystal packing may be delayed. 3. Once the crystal has reached the desired state, the humid air stream is turned off and the cryo-gas stream is turned on — for example, by opening a previously installed shutter. 4. The transformed and shock-frozen crystal can now be removed from the crystal holder and be handled by standard cryo-techniques. 4. Transformation of the bag-Oxidase During the transformation experiments with the bfl3-oxidase we found that these crystals show significant differences in their reaction toward humidity changes depending on the speed of the dehydration. In addition to the usually performed "equlibrium transformation experiments" (see Section IV,B,4,a), we devised "humidity gradient experiments" (see Section IV,B,4,b) in order to characterize the influence of the kinetics of the dehydration on the changes in the crystals. The equilibrium method has the advantage that one can evaluate the current state of the crystal prior to any further reduction in humidity and should be the first method employed. However, humidity gradient experiments may be necessary if the speed of dehydration is an important parameter.

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a, Equilibriutn or Slow Transformation of the hsL^-Oxidase Crystals 1. Mount a crystal on top of the micropipette at a humidity that is equal to or somewhat larger than the water pressure in the mother liquor—for example, 98% r.h. for the ba^-oxiddise crystals (see Section IV,B,2). 2. Transfer the crystal holder onto the x-ray camera. 3. Decrease the humidity in small steps, such as 0.5-1.0% each, and follow the change in diffraction quality and cell constants via diffraction images and auto-indexing with an appropriate software package. For each step, the response of the crystal to the humidity change is observed until a new equilibrium is reached (15-30 min). At the vicinity of a preferred crystals state the humidity value may be fine-tuned at smaller steps. 4. If the crystal shows the required diffraction quality, it may either be frozen or measured under tight regulation of the humidity at a value corresponding to the desired crystal state. Note: Not all preferred crystal states may be reached by a slow or equilibrium transformation (see below). The diffraction quality of the crystals may be lower between preferred crystal states such that upon reduction of the humidity an initial decrease in diffraction power may finally result in a much better crystal quality at even lower humidity values. At humidity values below the preferred crystals states—that is, after an excessive dehydration of the crystals—the diffraction power becomes very poor, correlating with a very high mosaic spread such that no further auto-indexing may be possible. Extensive transformation studies with different crystals showed that cell dimensions above 178 A for the c-axis coincide with very poor diffraction power. Furthermore, very subtle changes in the relative humidity lead here to dramatic changes in the cell dimensions. Therefore, we defined the crystals with a c-axis of 178 A (corresponding to about 97.5% relative humidity) as the "native crystals" for all further transformation experiments. With the equilibrium transformation method, the best diffraction quality was reached after an decrease in relative humidity by about 1.8-2.0% relative to the native crystals and corresponded to a c-axis of 172 A. b. Fast Transformation of the hdL^-Oxidase Crystals Using a Humidity Gradient If the relative humidity is reduced below 95.5% with the equilibrium transformation method, the ^flj-oxidase crystals show an irreversible damage of the crystal lattice. Since a preferred crystal state corresponding to a much smaller unit cell was found with the oil method (see Section IV,A), the kinetic

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effects of the transformation were studied. The corresponding fast transformation experiments were performed as follows. 1. Mount the crystal at the crystal holder of the humidity machine, as under Section IV,B,4,a. 2. Apply a steady humidity gradient (e.g., 1% reduction in r.h. per 4 min), and follow the changes in the crystal packing via continuous recording of short exposure diffraction images. 3. Determine the limiting resolution as well as the cell constants for each image and plot those against the corresponding humidity value as in Fig. 6. Note: The preferred crystal states are determined as local optima of the diffraction power of the crystals. The relative humidity as well as the cell constants corresponding to these preferred crystal states are reproducible between different crystals and different humidity gradients and should be characteristic for a given crystal system (Fig. 6). The limiting resolution obtained at any given preferred state was, however, highly dependent on the velocity of the humidity changes and the crystal being used. Interestingly, there seem to be two different regions in the response of i7fl3-oxidase crystals to changes in their solvent content. The first region ranges from the native crystals to the first optimum (c-axis = 172 A) and represents a continuous transition of the crystals where a reduction in solvent content leads to a steady increase in diffraction power. The second region can only be reached if the transformation is sufficiently fast (oil transformation and gradient experiments). It is characterized by the presence of discrete optima for the diffraction quality, which correspond to characteristic cell constants (Fig. 6). Only crystals that were transformed to one of the preferred states at lower solvent content (i.e., with a c-axis around 160 A) could reproducibly be frozen. Native crystals or such that were only transformed to the first optimum (c-axis = 172A) did show severe damage to the crystal lattice upon shock-freezing, corresponding to a poor diffraction power and an excessive mosaic spread. Dehydration of the &a3-oxidase crystals leads initially to an unhindered approach of the molecules in the unit cell, resulting in decreased cell constants and an increase in crystal packing forces, corresponding to the continuous transition range in Fig. 6. The second phase of the transformation, which is characterized by discrete optima in diffraction power, results in a further approach of the molecules, which then start to collide. This leads to a much lower increase in crystal packing forces as well as the displacement of some amino acid residues, which might explain the requirement for a suitable kinetics in the transformation process. This correlates also with the fact that the 172 A crystal state shows the best diffraction quality for the &a3-oxidase crystals. The analysis of the intermolecular hydrogen bonds between the different molecules in the crystal shows nicely the effect of the two phases of

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Tewfik Soulimane et al 8,0

lOU.U -

176,3

175,0 -

y ^ 172,3

•^ 170,0 o c

1 165,0

^166,3

c o = 160,0 o O

^^"^^

^A/^'''^^^

155,0 if^n n

60.0

160,5

158.5

1

65,0

70,0

75,0

80,0

85,0

1

—

90,0

95,0

100,0

Rel. Humidity [%]

FIGURE 6

Transformation of ^flg-oxidase crystals using a humidity gradient. The diffraction power (upper panel) and the cell constant c (lower panel) are drawn as a function of the humidity. The relative humidity was varied in form of a gradient from 97.5% down to 30% with a steady decrease of l % / 4 min. Diffraction images were recorded and analyzed with MOSFLM (Leslie, 1991), resulting in the indicated values for the resolution limit and cell constant. Several preferred crystal states are clearly visible, which correspond to the resolution optima in the upper panel. These preferred states, as well as the "native" crystal, are marked with closed symbols, and the cell constants are given. After reaching 30% r.h., the humidity was again increased to 97.0%, the crystal equilibrated overnight, and two more diffraction images were taken. The crystal returned to its original state (plus signs). © Reprinted from /. Appl. Cryst. 33, Kiefersauer, R., Than, M. E., Dobbek, H., Gremer, L., Melero, M., Strobl, S., Dias, J. M., Soulimane, T., and Huber, R. A novel free-mounting system for protein crystals: Transformation and improvement of diffraction power by accurately controlled humidity changes. 1223-1230 (2000), with permission from International Union of Crystallography (2000).

CHAPTER 14 Bflg-Type Cytochrome c Oxidase from Thermus thermophilus TABLE III

249

Intermolecular H-Bonds in Different States of baj-Oxidase Crystals Number of H-bonds*" Crystal state

Strong

Weak

MAD-structure: c-axis = 161A first optimum: c-axis = 172 A native crystals: c-axis = 178 A

26 26 6

12 8 4

' Number of intermolecular H-bonds that are responsible for the crystal packing forces determined (including the rating as strong or weak) by CONTACT (Collaborative Computational Project, 1994).

the transformation on the crystal packing forces (Table III). For a more detailed discussion of the molecular effects of the transformation procedure on the ba^ oxidase crystals, see Than (2000).

ACKNOWLEDGMENTS The authors would like to thank Professor R. Huber and Professor G. Buse for their support and interest in this work as well as Marianna Tatarek-Nossol for her excellent technical assistance.

REFERENCES Bragg, W. L., and Perutz, M. F. (1952). The external form of the haemoglobin molecule. II. Acta Cryst. 5, 323-328. Castenholz, R. W. (1969). Thermophilic blue-green algae and the thermal environment. Bacteriol. Rev. 33, 476-504. Collaborative Computational Project (1994). The CCP4 suite: Programs for protein crystallography. Acta Cryst. D50, 760-763. Cramer, P., and Miiller, C. W. (1997). Engineering of diffraction-quality crystals of the NF-K:B P52 homodimer-DNA complex. FEES Lett. 405, 373-377. Cramer, P., Bushnell, D. A., Fu, J., Gnatt, A. L., Maier-Davis, B., Thompson, N. E., Burgess, R. R., Edwards, A. M., David, P. R., and Kornberg, R. D. (2000). Architecture of RNA polymerase II and implications for the transcription mechanism. Science 288, 640-649. Dobrianov, I., Kriminski, S., Caylor, c. L., Lemay, S. G., Kimmer, C, Kisselev, A., Finkelstein, K. D., and Thome, R. E. (2001). Dynamic response of tetragonal lysozyme crystals to changes in relative humidity: Implications for post-growth crystal treatments. Acta Cryst. D57, 61-68. Esnouf, R. M., Ren, J., Garman, E. F., Somers, D. O'N., Ross, c. K., Jones, E. Y., Stammers, D. K. and Stuart, D. I. (1998). Continuous and discontinuous changes in the unit cell of HIV-1 reverse transcriptase crystals on dehydration. Acta Cryst. D54, 938-953.

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Tewfik Soulimane et ah Estebanez-Perpina, E., Fuentes-Prior, P., Belorgey, D., Braun, M., Kiefersauer, R., Maskos, K., Huber, R., Rubin, H., and Bode, W. (2000). Crystal structure of the caspase activator human granzyme B, a proteinase highly specific for an Asp-Pl residue. Biol. Chem. 381, 1203-1214. Fee, J. A., Choc, M. C , Findling, K. L., Lorence, R., and Yoshida, T. (1980). Properties of a copper containing ''Cjaag'' complex: A terminal oxidase of the extreme thermophile Thermus thermophilus HB8. Proc. Natl Acad. Sci. USA 77, 147-151. Ferguson-Miller S., and Babcock G. T. (1996). Heme/copper terminal oxidases. Chem. Rev. 96, 2889-2907. Fethiere, J., Andersson, A., Keinanen, K., and Madden, D. R. (2000). Crystallization of an AMPA receptor binding domain without agonist: Importance of carbohydrate content and flashcooling conditions. Acta Cryst. D56,1625-1629. Huxley, H. E., and Kendrew, J. C. (1953). Discontinuous lattice changes in haemoglobin crystals. Acta Cryst. 6, 76-80. Izard, T., Sarfaty, S., Westphal, A., Kok, A. D., and Hoi, W. G. (1997). Improvement of diffraction quality upon rehydration of dehydrated icosahedral enterococcus faecalis pyruvate dehydrogenase core crystals. Protein Sci. 6, 913-915. Kiefersauer, R. (1998). Kapillaren-freies Montieren von Proteinkristallen unter kontroUierter Feuchte. Vorrichtung und methodische Nutzung. Ph. D. Thesis. TU Miinchen, Germany. Kiefersauer, R., Than, M. E., Dobbek, H., Gremer, L., Melero, M., Strobl, S., Dias, J. M., Soulimane, T., and Huber, R. (2000). A novel free-mounting system for protein crystals: Transformation and improvement of diffraction power by accurately controlled humidity changes. /. Appl. Cryst. 33, 1223-1230. Leslie, A. G. W. (1991). Recent Changes to the MOSFLM Package for Processing Film and Image Plate DATA. SERC Laboratory, Daresbury, Warrington, WA 4AD, UK. Matthews, B. W. (1968). Solvent contents of protein crystals. /. Mol. Biol 33, 491-497. Michel, H., Behr, J., Harrenga, A., and Kannt, A. (1998). Cytochrome c oxidase: Structure and spectroscopy. Ann. Rev. Biophys. Biomol Struct. 17, 329-356. Petock, J. M., Wang, Y-F., DuBois, G. C , Harrison, R. W., and Weber, I. T. (2001). Effects of different post-crystallization soaking conditions in the diffraction of Mtcpl crystals. Acta Cryst. D57, 763-765. Qian, C , Lagace, L., Massariol, M., Chabot, C , Yoakim, C , Deziel, R., and Tong, L. (2000). A rational approach towards successful crystallization and crystal treatment of human cytomegalovirus protease and its inhibitor complex. Acta Cryst. D56, 175-180. Schick, B., and Jurnak, F. (1994). Extension of the diffraction resolution of crystals. Acta Cryst. D50, 563-568. Soulimane, T., Gohlke, U., Huber, R., and Buse, G. (1995). Three-dimensional crystals of cytochrome-c oxidase from Thermus thermophilus diffracting to 3.8 A resolution, FEBS Lett. 368, 132-134. SouHmane, T., von Walter, M., Hof, P., Than, M. E., Huber, R., and Buse, G. (1997). CytochromeC552 from Thermus thermophilus: a functional and crystallographic investigation. Biochem. Biophys. Res. Comm. 237, 572-576. Soulimane, T., Buse, G., Bourenkov, G. P., Bartunik, H. D., Huber R., and Than, M. E. (2000a). Structure and mechanism of fcflg-Cytochrome c Oxidase from Thermus thermophilus. EMBO J. 19,1766-1776. SouHmane, T., Than, M. E., Dewor, M., Huber, R., and Buse, G. (2000b). Primary structure of a novel subunit in bflg-Cytochrome Oxidase from Thermus thermophilus. Prot. Sci. 9, 2068-2073. Stammers, D. K., Somers, D. O., Ross, C. K., Kirby, L, Ray, P. H., Wilson, J. E., Norman, M., Ren, J. S., Esnouf, R. M., Garman, E. F., Jones, E. Y., and Stuart, D. I. (1994). Crystals of HIV-1 reverse transcriptase diffracting to 2.2 A resolution. /. Mol. Biol. 242, 586-588.

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Tahirov, T. H., Oki, H., Tsukihara, T., Ogasahara, K., Yutani, K., Libeu, C. P., Izu, Y., Tsunasawa, S., and BCato, I. (1998). High-resolution crystals of methionine aminopeptidase from pyrococcus furiosus obtained by water-mediated transformation. /. Struct. Biol. Ill, ^%-ll. Than, M. E., Hof, P., Huber, R., Bourenkov, G. P., Bartunik, H. D., Buse, G., and Soulimane, T. (1997). Thermus thermophilus Cytochrome-C552: a new highly thermostable Cytochrome-c structure obtained by MAD phasing. /. Mol. Biol. 271, 629-644. Than, M. E. (2000). Rontgenstrukturanalyse der ba^ Cytochrom-c Oxidase aus Thermus thermophilus und ihres Substrates Cytochrom-C552. Ph.D. Thesis. TU Miinchen, Germany. Than, M. E., and Soulimane, T. (2001). Bflg-Cytochrome c oxidase from Thermus thermophilus. In ''Handbook of Metalloproteins'' (A. Messerschmidt, R. Huber, T. Poulos and K. Wieghardt, eds.) pp. 363-378. John Wiley & Sons Ltd, Chichester, UK. Weiss, M. S., and Hilgenfeld, R. (1999). Dehydration leads to a phase transition in monoclinic factor XIII crystals. Acta Cryst. D55, 1858-1862. Zimmermann, B. H., Nitsche, C I., Fee, J. A., Rusnak, P., and Munck, E. (1988). Properties of a copper-containing cytochrome ba^: A second terminal oxidase from the extreme termophile Termus thermophilus. Proc. Natl. Acad. Sci. U.S.A. 85, 5779-5783.

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Two-Dimensional Crystallization of Membrane Proteins: A Practical Guide WERNER KUHLBRANDT Max-Planck-Institut fur Biophysik Heinrich-Hoffmann-Strase 7 60528 Frankfurt am Main, Germany

I. INTRODUCTION Membrane proteins still present a major challenge in structural biology. Electron microscopy (EM) of two-dimensional (2D) crystals is an increasingly important technique for studying their structure and function. An earlier review (Kiihlbrandt, 1992) published about 10 years ago outlines the basic techniques and concepts. While these are unchanged and valid, the number of membrane proteins crystallized in 2D has risen encouragingly in the past three or four years, no doubt largely due to the recent progress in expressing membrane proteins in homologous or heterologous systems. The evaluation of more than 60 2D crystallization protocols now provides a sound base for some general guidelines. Recent technical developments are also discussed in

Membrane Protein Purification and Crystallization Ije: A Practical Guide. Copyright 2003, Elsevier Science (USA) All rights of reproduction in any form reserved.

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this chapter and may help to facilitate the future production of 2D crystals. Several other reviews published in the past 10 years (Jap et ah, 1992; Mosser, 2001; Rigaud et al., 2000; Walz and Grigorieff, 1998) address various aspects of 2D crystallisation.

11. THINGS TO CONSIDER BEFORE YOU START A. 2D or 3D Crystals? You have worked hard to isolate and purify a membrane protein and want to find out its structure. Given that NMR techniques for routine structure determination of membrane proteins are still some way off, there are only two choices: x-ray crystallography of 3D crystals and electron microscopy of 2D crystals. Should you attempt 2D or 3D crystallization first? The answer depends mostly on how much protein you have and how much can be produced on a regular basis. If 5-10 mg of pure protein can be prepared routinely, 3D crystallization will be the first choice. More typically, 1 or 2 mg have been purified, and it has taken several months. In this case, 2D crystallization is more promising. However, the outcome of any crystallization experiment is unpredictable, especially with membrane proteins, and it is therefore always a good idea to try both, if the amount of material permits it.

B. Which Membrane Protein Structures Have Been Determined by Electron Microscopy? So far, 2D crystals have yielded atomic models of three membrane proteins: bacteriorhodopsin (Grigorieff et al., 1996; Henderson et ah, 1990; Kimura et ah, 1997), plant light harvesting complex (Kiihlbrandt et ah, 1994), and aquaporin (Murata et ah, 2000). Currently, 2D crystals of five or six other membrane proteins have the potential for structure determination at nearatomic resolution. But electron microscopy of 2D or tubular crystals of membrane proteins has also produced a wealth of structures at 5-9 A resolution, including 3D maps of the Ca^"^ ATPase in the E2 conformation (Zhang et ah, 1998), the Neurospora proton ATPase (Auer et ah, 1998), photosystem II (Rhee et ah, 1998), the nicotinic acetylcholine receptor (Miyazawa et ah, 1999), cardiac gap junctions (Unger et ah, 1999), halorhodopsin (Kunji et ah, 2000), the N a ' ' / H + antiporter from E. coli, NhaA (Williams, 2000), microsomal glutathione transferase 1 (Schmidt-Krey et ah, 2000), and the c-subunit ring of an F^-ATPase (Vonck et ah, 2002). Many other 2D membrane protein crystals obtained more recently are at a less advanced stage of analysis and have yielded projection structures at 6-8 A resolution. Recent examples include bacterial NO reductase (Gohlke et ah, 1997), the monomeric porin OmpG (Behlau et ah, 2001), the bacterial transporter EmrE (Tate et ah, 2001), an oxalate transporter (Heymann et ah, 2001), a chloride channel (Mindell et ah, 2001), the enzyme II mannitol transporter (Koning et ah, 1999), and the bacterial protein translocase Sec YEG (CoUinson et ah, 2001).

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Even 17 years after the X-ray structure of the bacterial reaction center was pubUshed (Deisenhofer et ah, 1985), the 3D crystaUization of membrane proteins is far from routine, and this is not Hkely to change in the foreseeable future. If 3D crystals are obtained, they are rarely very good to begin with, and improving their quality to the point at which structure determination by x-ray crystallography becomes feasible can take years, during which little or no publishable structural information is forthcoming. By comparison, 2D crystals of membrane proteins form quite readily. At least 28 bacterial, 3 archaeal, and 20 eukaryotic membrane proteins have so far yielded twodimensional crystals (Kiihlbrandt, 2002), which means that the chances of success are quite high. Some examples are shown in Fig. 1. Another point in favor of 2D crystals is that most of them form under conditions that are close to physiological. All 2D crystals have provided interesting and important information on the structure and function of the protein. 2D crystals are therefore an excellent option for studying molecular architecture and mechanisms of membrane proteins.

III. CRYSTALLIZATION PARAMETERS The basic concept of a typical 2D crystallization experiment is very simple. The purified protein in detergent solution is mixed with lipid, also in detergent solution. The detergent is then removed, whereupon lipid and protein coalesce into proteoliposomes or reconstituted membrane sheets in which many membrane proteins will form 2D lattices under favorable conditions. A typical 2D crystallization experiment thus starts with three main components: the protein, the detergent, and the lipid. In the course of the experiment, the three components are then reduced to two as the detergent is removed. As in all protein crystallization experiments, the parameters of 2D crystal formation need to be carefully controlled. The most important parameters are the protein concentration, the choice of lipid, the lipid-to-protein ratio (LPR), the detergent, and the ionic strength, pH, and temperature. Table I contains an overview of promising conditions, based on a survey of more than 60 published 2D crystallization protocols (Kiihlbrandt, 2002). Even though there is as yet no failsafe general method for 2D crystallization, the range of conditions that will yield 2D crystals is surprisingly small.

A. The Protein The most important component, but the one over which one has the least control, is the protein itself. Every protein is different, and membrane proteins in particular are notorious individualists. A very few occur in crystalline form

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FIGURE 1 2D crystals of membrane proteins. A. 2D crystals of the sodium/proton antiporter NhaA at low magnification, showing the characteristic shape of elongated tubular vesicles flattened against the carbon support film. Scale bar, 1 /im. B. Freezefracture replica of NhaA incorporated into lipid vesicles. The fracture face on the upper right shows a membrane densely packed with the protein, which becomes ordered toward the center of the picture. In the lower half, two small fragments of other crystalline vesicles are visible, showing the typical lattice repeat of 2D crystals of NhaA. Scale bar, 0.1 /xm. C. Crystalline spherical vesicle of Fo ATPsynthase c-subunit, surrounded by small noncrystalline lipid vesicles. Scale bar, 0.25 jam. D. Tubular vesicle of Fo ATPsynthase c-subunit. The edge of the square inset showing the lattice is 300 nm. (A) is courtesy of Dr. Christine Ziegler, (B) of Dr. Winfried Haase, (C) and (D) of Dr. Janet Vonck, all at the Max-Planck Institute of Biophysics, Frankfurt am Main, Germany. From Kiihlbrandt (2002)

CHAPTER 15 Two-Dimensional Crystallization of Membrane Proteins

-^-^

FIGURE 1

Continued.

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Werner Kiihlbrandt

TABLE I

2D Crystallization Conditions for Membrane Proteins Most likely to work

Range to try

1 mg/ml Detergent in which protein is stable and active, DDM DMPC or natural lipid extract 0.2-0.5

0.5-2 m g / m l C12E8, DM, OG, OTG

pH Salt

Neutral 100 mM NaCl 10 mM MgCl2

Nonionic additives

20% glycerol

Temperature

25°C-30°C

Dialysis time

4 - 7 days

3.5-10 0-600 mM NaCl 0-100 mM MgCl2 Other divalent ions, EDTA 0-40% glycerol, low molecular weight PEG 4°C-37°C Dependent on protein stability Up to 2 months

(a) Crystallization mix Protein concentration Detergent

Lipids LPR ( w / w )

DOPC, POPC, lipids extracted from native membrane 0-1.5

(h) Detergent dialysis

(c) Detergent absorption Preincubation pH, salt Temperature Absorption time Postincubation

1-8 hours As above 4°C Hours Days

As above Up to 25°C Weeks

in the native membrane, some crystallize easily, others do so only reluctantly, and some not at all. Persuading a reluctant protein to crystallize can be difficult, but an apparent reluctance may simply mean that the right conditions have not yet been found. Of course, it is not known at the outset whether such "right conditions" exist, but it now seems that most membrane proteins that are available in a suitable form and in sufficient quantity for 2D crystallization trials can be induced to form 2D crystals. 1. Protein Expression For many years, the limited availability of membrane proteins from natural sources has been a major bottleneck. By their nature, most membrane proteins are much less abundant than cytoplasmic or cytoskeletal proteins because the cell membrane only represents a tiny fraction of the total cell

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volume. To make matters worse, the membrane typically contains a large number of different proteins that are present in comparatively small copy numbers. Their isolation and purification in crystallizable quantities therefore requires expression of recombinant proteins in heterologous or homologous systems (see review by Grisshammer and Tate, 1995). Expression has finally become routine for some bacterial membrane proteins, and this is reflected by the recent surge of successful 2D crystallizations. However, the expression of most eukaryotic membrane proteins remains an unsolved problem. Only few have been expressed in workable quantities, and only two of these have been crystallized, both in 2D: truncated cardiac gap junction (Yeager et ah, 1999) and Arabidopsis H'^-ATPase (Jahn et al, 2001; Lebeau et al, 2001). Since 2D crystallization requires less material by one or two orders of magnitude and is more likely to work than 3D crystallization, it will gain in importance for investigating the structure and function of eukaryotic membrane proteins that are difficult to express in large quantities. 2. Protein Concentration An important parameter related to the protein over which one does have control is its concentration. For 2D crystallization by detergent dialysis or detergent absorption, a protein concentration around 1 m g / m l is a good starting point. Occasionally, slightly higher or lower concentrations work better. For the recently explored surface crystallization protocols, protein concentrations down to 0.1 m g / m l or even 0.01 m g / m l are sufficient. 3. Purity Protein purity is an obvious prerequisite for any crystallization. 2D crystallization is more forgiving in this respect than 3D crystallization, even though it is difficult to be specific about the levels of impurities that will be tolerated. Evidently, it is possible for membrane proteins to form 2D crystals, even well-ordered 2D crystals in very heterogenous systems. This is demonstrated by the fact that a fair number of membrane proteins form 2D crystals in situ — that is, without ever being removed from the membrane—and therefore in the presence of all other membrane components. The best example is bacteriorhodopsin, which forms some of the best 2D crystals known in vivo, in a membrane that must contain hundreds of different membrane proteins. Unfortunately, the tendency to form 2D lattices under such conditions depends very much on the protein. Some will form strong lattice contacts and therefore cope with comparatively high levels of impurities, whereas the interactions of others will be much weaker and therefore more prone to disruption by intercalating foreign proteins and lipids. To be on the safe side, the protein should be as pure as possible while maintaining it in a stable and active conformation.

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4. Monodispersity Next to purity, monodispersity of the protein solution is usually an important prerequisite for crystallization. Monodispersity means that only one of several possible protein oligomers is present in solution. Again, this is particularly relevant for 3D crystals. 2D crystals have been known to form in systems where different oligomeric forms exist in equilibrium with one another, as demonstrated most recently by the 2D crystallization of SecYEG (CoUinson et ah, 2001). Analytical ultracentrifugation had shown that the protein solution contained monomers and tetramers in equilibrium, yet 2D crystals were obtained in which the complex exists as a dimer. 5. Stability Protein stability is a major concern in 2D and 3D crystallization. Many membrane proteins are unstable in detergent solution. The stability often depends critically on the choice of detergent and on the presence of lipids. Strict requirements for a particular detergent or lipid may make 3D crystallization difficult, because either may interfere with crystal contacts. These requirements are easily accommodated in 2D crystallization because the detergent will be removed and lipids are needed anyway for 2D crystal formation. Glycerol is frequently added to enhance protein stability during 2D crystallization, in particular to fragile eukaryotic proteins. Many membrane proteins are much more stable in a lipid bilayer than in detergent solution, as is shown by 2D crystals in which the protein remains stable and active for many months. This is another advantage over 3D crystallization where the protein usually remains exposed to high levels of detergents during crystallization and even in the crystal lattice. 2D crystallization is therefore especially appropriate for sensitive, unstable membrane proteins, in particular for those from higher eukaryotes.

B. The Lipid After the protein, the second most important component in a 2D crystallization experiment is the lipid. Lipids are a natural part of the environment of all membrane proteins in the cell. It is therefore important to be familiar with the biochemistry, analysis, and isolation of membrane lipids. Christie's, (1982) book is an excellent, comprehensive reference on this subject. 1. Lipid Classes Lipid classes are defined by head groups. The most commonly encountered class of lipids are the phospholipids. The phosphatidyl cholines (PC), phosphatidyl ethanolamines (PE), phosphatidyl serines (PS), and sphin-

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gomyelin are zwitterionic, which means that the effective charge of their head groups depends on the pH. Others are negatively charged at neutral pH, including the phosphatidyl glycerols (PG), phosphatidyl inositols (PI), and phosphatidic acids (PA). The cardiolipins (which can be thought of as two PG molecules linked via a glycerol) carry two negative charges on their head group. Other major constituents of biological membranes are the ceramides and gangliosides. These are uncharged sphingolipids, which are found predominantly in neuronal tissue and brain. Plants have characteristic glycolipids, monogalactosyl diacylglycerol (MGDG), and di-galactosyl diacylglycerol (DGDG), which are not found in other organisms. Their head groups are hydrophilic but uncharged. 2. The Fatty Acids Most natural lipids have unbranched C18 or C16 fatty acid chains. These are either fully saturated or singly or multiply unsaturated. The length and saturation of the fatty acid tails determine the fluidity of the lipid bilayer. Membrane fluidity depends on the transition temperature at which the lipid changes from a crystalline to a fluid state. Lipids with fully saturated fatty acid chains of more than 14 carbon atoms have transition temperatures above 20°C, whereas most biological lipids with unsaturated fatty acid chains have transition temperatures below 0°C (see Cevc and Marsh, 1987; Kiihlbrandt, 1992). It has often been pointed out that 2D crystallization of membrane proteins in a lipid bilayer requires the latter to be fluid, and this is one of the reasons why slightly elevated temperatures are preferred (see Table I). Multiply unsaturated fatty acid chains have nonconjugated cis double bonds that make them prone to oxidation. Lipid oxidation is a common concern when different batches of natural lipids fail to give reproducible results. With polyunsaturated lipids it is therefore important to take precautions to prevent oxidation, such as storing them under argon or nitrogen at low temperature. It is advisable to work in an inert gas atmosphere or to add an enzymatic oxygen scavenging system such as glucose oxidase to the crystallization mix. Monounsaturated synthetic lipids such as di-oleoyl phosphatidyl glycerol (DOPC), di-oleoyl phosphatidyl glycerol (DOPG), and palmitoyl oleoyl phosphatidyl choline (POPC) are less sensitive to oxidation than the polyunsaturated natural lipids, but they have suitably low transition temperatures. They have therefore become increasingly popular as an alternative to natural lipids in 2D crystallization experiments. 3. Lipid-Protein Interactions Biological membranes contain a variety of different lipids, classified by their head groups and hydrophobic fatty acid tails. Both moieties may interact specifically with the protein, as can be studied in several of the recent

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high-resolution x-ray structures. Some proteins require the presence of Upids during all stages of isolation and purification. Even very fragile proteins that last for only a short time in detergent become surprisingly stable once lipid is added to the solution. If this is the case, it is likely that the success of the crystallization will also depend on a particular lipid. Only the minimal amount of lipid consistent with protein stability should be added, however, since the lipid may be difficult to remove later on. 2D crystallization requires a carefully controlled, usually rather low amount of lipid. Some proteins will crystallize only with lipids from their own membrane. This is the case for example, with sensory rhodopsin (Kunji et ah, 2001), squid rhodopsin (Davies et ah, 1996), and light-harvesting complex of photosystem II (LHC-II) (Nussberger et ah, 1993). These essential lipids should be extracted from isolated membranes. This is easily done by standard chloroform/ methanol extraction of bulk membranes, followed by column chromatography to separate lipid classes (Christie, 1982). Unless the protein requires a particular lipid, there is usually a choice of using either an extract from natural sources or synthetic lipids. If nothing is known about lipid requirements of the protein, it may be best to aim for a lipid composition that it is as close as possible to the membrane where it normally occurs, on the assumption that the protein will then find and select the lipid it needs. Commercially available lipid extracts are frequently used, most commonly E. coli polar lipids for proteins of bacterial origin, or egg yolk and plant lecithin for proteins from animal or plant tissues, respectively. However, the reproducibility of lipid batches can be problematic, since plant and animal lipid extracts are often oxidized and very impure. Fortunately, most membrane proteins are not particular with respect to lipid requirements. A number of bacterial membrane proteins have been crystallized with animal lipids and vice versa. Pure synthetic compounds such as DOPC, POPC, DOPG, and even DMPC (di-myristoyl phosphatidyl choline), or mixtures of these, often work surprisingly well. In these cases, the lipid seems to be little more than a hydrophobic matrix into which the protein inserts without specific lipid-protein interactions. 4. Lipids Copurified with the Protein Frequently, one or several membrane lipids bind tightly to an isolated membrane protein throughout isolation and purification. Occasionally, these copurified lipids are sufficient for 2D crystal formation, as was found with LHC-II (Nussberger et ah, 1993) and a photosystem II subcomplex (Nakazato et ah, 1996; Rhee et ah, 1997; Rhee et ah, 1998). The presence of tightly bound lipids in isolated membrane proteins should be established early on, since they are likely to be essential for protein integrity and stability. They should not be removed by excessive purification—for example, by column chromatography—which exposes the protein

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to a large excess of detergent. The lipid content of a purified membrane protein can easily be monitored by ID or 2D thin-layer chromatography (TLC) of a small sample (2-4 jA) of concentrated detergent-protein solution (for details of the procedure see Christie, 1982). However, the identification and exact quantitation of accompanying lipids in a small volume of a protein solution is not easy. Laser-desorption mass spectroscopy is a useful method for identifying lipids by comparison against known standards, although the fatty acid chain heterogeneity of natural lipids can complicate the evaluation. Semiquantitative analysis of lipid content is possible by ID thin-layer chromatography and comparison against a dilution series of pure compounds. 5. Lipid-to-Protein Ratio (LPR) Usually, lipids need to be added to the purified protein for 2D crystallization. The amount of lipid in relation to the protein is given by the lipid-to-protein ratio (LPR). Next to the protein concentration and the kind of lipid used, the LPR is the most critical variable in 2D crystallization. The LPR range that has produced good 2D crystals is, however, surprisingly small: Most form at an LPR of 0.3-0.5 by weight (Kiihlbrandt, 2002). For a membrane protein of 40,000 Da and an average molecular mass of ^ 700 for the lipid, this translates into 15-30 lipid molecules per protein, or 7-15 in each membrane leaflet. This is the range one would expect of a biological membrane with ^50% protein content. There are few exceptions from this rule: Microsomal glutathione transferase-1 only forms good 2D crystals at a very low ratio of ^ 3 lipid molecules per protein (Schmidt-Krey et ah, 1999). a-TIP, the aquaporin of plant vascular membranes requires a similarly low number (Daniels et ah, 1999). At the other end of the scale, enzyme II C mannitol (Koning et ah, 1999), the membrane part of human Band 3 (Wang et ah, 1993) and the tetracyclin transporter TetA (Yin et ah, 2000) have higher lipid requirements of ^50-60 lipid molecules per protein, corresponding to an LPR of close to 1.0.

C. The Detergent The detergent is less important in 2D crystallization than the protein or the lipid, simply because all or most of it is removed as a prerequisite for 2D crystal formation. Most detergents are chosen at the stage of isolation and purification to keep the protein stable and active. For 3D crystallization a detergent change is often necessary, since the type of detergent and its concentration are two key parameters in 3D lattice formation. For 2D crystallization, a detergent change is usually unnecessary. 2D crystals have been grown from a large variety of detergents. Most commonly these are mild, nonionic detergents that are kind to the protein, such as dodecyl maltoside (DDM), decyl maltoside (DM), and octa-ethyleneglycol mono-dodecyl ether

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(C12E8), but harsher detergents octyl glucoside (OG), lauryl dimethyl amine oxide (LDAO), and even sodium dodecyl sulfate (SDS) have also yielded 2D crystals. Poly disperse detergents such as Triton X-100 and polyethyleneglycol mono-octyl ether (Cg-POE), which are avoided in 3D crystallization are nevertheless useful for 2D trials, because the detergent does not play a major structural role in the 2D crystal lattice. Dodecyl maltoside (DDM) has become the most widely used detergent in membrane protein biochemistry and crystallization. Decyl maltoside (DM) has similar properties but a 10-fold higher critical micellar concentration (CMC), in line with the useful rule of thumb that for each CH2 group added to the fatty acid chain, the CMC of a detergent within one chemical series (e.g., the alkyl maltosides) drops by a factor of about 3. DM is therefore more readily removed by dialysis. DDM and DM, or mixtures of both, are good detergents for attempting the 2D crystallization of a new membrane protein. The initial detergent concentration affects the rate and duration of dialysis and therefore the size of protein-lipid vesicles obtained. The exact detergent concentration in a membrane protein solution at the beginning of a crystallization experiment is often unknown because the material is too precious for this sort of measurement. If necessary, thin-layer chromatography (TLC) of a small protein sample, compared against a dilution series of the detergent, offers a convenient and sufficiently accurate estimate of its concentration. The effective detergent concentration in a solution can also be estimated from the contact angle of 10 /il drops of a dilution series on a hydrophobic surface such as parafilm. Above the CMC, the drop has a shallow contact angle. This changes abruptly to about 90° at the CMC or below. For more accurate methods, see Ringler et al. (2000) and Zulauf (1990). These do, however, require larger volumes of detergent solution. Fortunately, the initial detergent concentration rarely matters in a 2D crystallization experiment, because the detergent will be removed anyway by dialysis or absorption. A high detergent concentration therefore simply means that dialysis will take longer. If this is a problem, it may be possible to reduce the amount of detergent in the protein solution by judicious addition of detergent-absorbing polystyrene beads.

D. pH A strong point in favor of 2D crystallization is that it is nearly always achieved at close to physiological conditions. The majority of 2D crystallizations are carried out with standard buffers, such as Tris or HEPES, at 10-50 mM concentration in the pH range between 6.5 and 8.0, most often close to pH 7.4. Where there are significant derivations from physiological pH, they tend to be decisive for crystal formation. Some membrane proteins require a pH around 9 to form 2D crystals. These include the bacterial cytochrome bo complex (Gohlke et al, 1997), Omp21 (Baldermann and Engelhardt, 2000), and

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the membrane part of mitochondrial complex I (Sazanov and Walker, 2000). By contrast, the sodium/proton antiporter, NhaA (Williams et ah, 1999), the oxalate transporter, OxlT (Heymann et ah, 2001), and sensory rhodopsin II (Kunji et ah, 2001) require acidic pH. In the case of NhaA it is known that low pH induces a stable, inactive conformation of the transporter. This seems to crystallize more readily than the active, presumably more flexible form.

E. Salt Concentration and Ionic Additives Like the pH, the preferred salt concentration in 2D crystallization experiments tends to be close to physiological. Most protocols call for 100-200 mM NaCl or KCl and 2-50 mM MgCl2. In a few cases 1-5 mM EDTA is added to neutralize divalent cations if these have undesirable effects. Substrates, such as oxalate in the case of OxlT, (Heymann et ah, 2001), inhibitors, such as orthovandate for the P-type ATPases (Iwane et ah, 1996; Mohraz, 1999; Rice et ah, 2001; Young et ah, 1997), or cofactors, such as NAD"^ for complex I (Sazanov and Walker, 2000) may be required to induce or maintain a crystallizable conformation of the protein. 3 mM sodium azide is nearly always added to prevent the growth of microorganisms in the crystallization mix. This is an advisable precaution, in particular if crystallizations are to be carried out at room temperature or above. Another option is to use cacodylate buffer (Brisson and Unwin, 1984), which is bactericidal.

F. Nonionic Additives Many fragile membrane proteins, including most of those from eukaryotic organisms, benefit from the addition of 5-30% glycerol to the crystallization mix. Glycerol is often added to stabilize sensitive proteins to protect them against denaturation, presumably by preventing unfavorable hydrophobic interactions. As an agent that mildly disrupts water structure in the hydration shells around solubilized proteins and detergent micelles, it also promotes micelle fusion (Kiihlbrandt, 1992). Mild precipitating agents such as MFD (Williams et ah, 1999) are sometimes added for the same reason.

G. Precipitants Precipitating agents such as ammonium sulfate, high salt, or polyethylene glycol, which force the protein-detergent mixed micelles out of solution, are rarely used in 2D crystallization. Exceptions are those protocols that rely on encouraging polar interactions between large extramembraneous domains, much as in 3D crystallization. Examples include the in situ crystallization of cytochrome oxidase (Prey et ah, 1978) and KdP-ATPase (Iwane et ah, 1996), the freeze-thaw protocol for cytochrome b^f (Mosser et ah, 1997), and the surface crystallization of Neurospora H'^ATPase (Auer et ah, 1999). Strikingly, and not

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surprisingly, all of these tend to yield multilamellar crystals rather than true 2D sheets and vesicles. In some cases, however, it has been possible to arrest crystal growth at the monolayer stage.

H. Temperature As pointed out above, the lipid environment of the hydrophobic part of the protein in a 2D crystal is equally as important as, if not more important than, the solution medium surrounding the hydrophilic solvent-exposed parts. Crystallization parameters influencing the lipid environment are therefore crucial. The most important of these is the crystallization temperature. This affects 2D crystal formation in several ways: First, if the detergent is removed from the crystallization mixture by dialysis, the rate of dialysis is obviously temperature-dependent. Higher temperatures will speed up the process, and lower temperatures will slow it down. Second, the temperature has a direct effect on the fluidity of the lipid and therefore on the rate of two-dimensional diffusion of the protein in the lipid bilayer. Third, the CMC of most detergents is temperature-dependent (da Graca Miguel et ah, 1989; Majhi and Blume, 2001), and ramping the temperature across the CMC once or several times in the course of the experiment has been found to encourage the fusion of mixed micelles into proteoliposomes and 2D crystals. Finally, the strength of hydrophobic interactions increases with increasing temperature (Kiihlbrandt, 1992). Therefore, the interactions between the hydrophobic surface areas of the protein with one another and with the lipid, as well as the strength of lipid-lipid interactions, increase as the temperature rises. These interactions are likely to affect the size and order of the 2D crystals obtained. For example, large vesicles may form at room temperature, whereas only small vesicles may be obtained at 4°C. For these reasons, slightly elevated temperatures or temperature ramps between room temperature and 37°C are used whenever possible. This works well for robust bacterial membrane proteins, including the bacterial terminal oxidases (Hendriks et ah, 1998; Warne et ah, 1995), the sodium/proton antiporter, NhaA (Williams et ah, 1999), the oxalate transporter OxlT, and the tetracycline transporter (Yin et ah, 2000). Of course, not every membrane protein will tolerate elevated temperatures for extended periods. Many of the fragile bacterial and eukaryotic membrane proteins will tend to denature under such conditions, certainly as long as they are in detergent solution. It may then help to first reconstitute the protein into liposomes at lower temperature by detergent absorption (see below) and then induce crystallization by raising the temperature to 37°C for an hour or two. Detergent absorption is less temperature-dependent and comparatively rapid and therefore particularly suitable for the 2D crystallization of fragile proteins. A few membrane proteins have been crystallized in 2D by dialysis at 4°C (Karrasch et ah, 1995; Koning et ah, 1999; Kunji et ah, 2000), but in these cases octyl

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glucoside (OG) was used to reduce dialysis time. Once crystalline sheets or vesicles have formed, they are usually quite stable and may be kept at 4°C for weeks or months.

IV. METHODS FOR GROWING 2D CRYSTALS OF MEMBRANE PROTEINS 2D crystals of membrane proteins can be grown in two different ways: in suspension or on a surface. In both cases, incorporation of the protein into a lipid bilayer is usually induced by detergent removal. So far, most 2D membrane protein crystals have been grown in isotropic suspension.

A. 2D Crystallization in Suspension Normally, a 2D crystallization experiment starts with a solution of protein, lipid, and detergent. When crystals are grown by detergent absorption, the components are mixed in an Eppendorf tube and incubated for several hours to give the lipid time to equilibrate with the detergent/protein micelles. This is usually unnecessary for 2D crystallization by detergent dialysis, which takes longer. The total volume of crystallization mixture prepared for an experiment depends on the availability of the protein and on the crystallization device to be used. Volumes of 50-100 fil are suitable for initial screens, but volumes of 20 jA or less can be used if the amount of protein is limiting. A 100 /il volume of a crystal suspension can be sufficient for an entire structure determination, provided that the crystals are stable. In the next step, the detergent concentration is brought below the CMC. This can be done in three ways: by dilution, detergent absorption, or dialysis. 1. Detergent Dilution Dilution of lipid-protein-detergent mixtures is simple and quick. It is, however, practicable only with high-CMC detergents (0.5-lmM) and high protein concentrations. Detergent dilution has yielded 2D crystals of bacterial porins and photosystem I (Dolder et ah, 1996; Hasler et al., 1998) but has not been used much otherwise. 2. Detergent Absorption Detergent absorption (Fig. 2) is a convenient and universal way of reducing the detergent concentration in a small volume of membrane protein solution. It is almost as quick as dilution but has the advantage of not

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9M #

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FIGURE 2

2D crystallization strategy for detergent absorption.

FIGURE 3

Detergent equilibrium.

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CHAPTER 15 Two-Dimensional Crystallization of Membrane Proteins

FIGURE 4

Detergent dialysis.

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