Membrane Transport: A Practical Approach

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Membrane Transport

The Practical Approach Series Related Practical Approach Series Titles Animal Cell Culture 3ed* PCR3: PCR in situ hybridization Immunoassays Affinity Separations Differential Display Immunochemistry 1 FMOC Solid Phase Peptide Synthesis Immunochemistry 2 Protein Localization by Fluorescence Signalling by Inositides Microscopy Protein Function 2ed Protein Phosphorylation 2ed Protein Structure 2ed Crystallization of Nucleic Acids and Subcellular Fractionation Proteins Protein Structure Prediction Signal Transduction 2ed Gene Probes 1 DNA Microallay Technology Gene Probes 2 High Resolution Chromatography Ion Channels Post-Translational Modification DNA Cloning 2: expression systems Protein Expression Basic Cell Culture Gel Electrophoresis of Proteins 3ed Protein Blotting In Situ Hybridization 2ed Glycobiology Cell Separation Immunocytochemistry HPLC of Macromolecules 2ed Lipid Modification of Proteins * indicates a forthcoming title Please see the Practical Approach series website at http://www.oup.co.uk/pas for full contents lists of all Practical Approach titles.

Membrane Transport A Practical Approach Edited by

Stephen A. Baldwin School of Biochemistry and Molecular Biology University of Leeds

OXFORD UNIVERSITY PRESS

OXJORD UNIVERSITY PRESS

Great Clarendon Street, Oxford OX2 6DP Oxford University Press is a department of the University of Oxford. It furthers the University's objective of excellence in research, scholarship, and education by publishing worldwide in Oxford New York Athens Auckland Bangkok Bogota Buenos Aires Calcutta Cape Town Chennai Dar es Salaam Delhi Florence Hong Kong Istanbul Karachi Kuala Lumpur Madrid Melbourne Mexico City Mumbai Nairobi Paris Sao Paulo Singapore Taipei Tokyo Toronto Warsaw with associated companies in Berlin Ibadan Oxford is a registered trade mark of Oxford University Press in the UK and in certain other countries Published in the United States by Oxford University Press Inc., New York © Oxford University Press, 2000 The moral rights of the author have been asserted Database right Oxford University Press (maker) First published 2000 All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without the prior permission in writing of Oxford University Press, or as expressly permitted by law, or under terms agreed with the appropriate reprographics rights organization. Enquiries concerning reproduction outside the scope of the above should be sent to the Rights Department, Oxford University Press, at the address above You must not circulate this book in any other binding or cover and you must impose this same condition on any acquirer British Library Cataloguing in Publication Data Data available Library of Congress Cataloguing in Publication Data 1 3 5 7 9 1 08 6 4 2 ISBN 0-19-963705-9 (Hbk.) ISBN 0-19-963704-0 (Pbk.) Typeset in Swift by Footnote Graphics, Warminster, Wilts Printed in Great Britain on acid-free paper by The Bath Press, Bath, Avon

Preface

Biological membranes form the interface between cells and their environment, and the passage of small molecules across this barrier is vital for the supply of metabolites. In the case of most small organic molecules required by the cell, this passage is catalysed by transport proteins embedded in the membrane. Genome sequencing has created a golden age for the membrane biologist, in that for many bacterial species and several eukaryotes we now know the amino acid sequences of all the transport proteins present in the organism—although in many cases we do not have any idea what their substrates may be. Unfortunately, our understanding of how these proteins actually carry out their task of moving molecules across the lipid bilayer has lagged behind our knowledge of their sequences, largely because of the paucity of membrane protein structures so far solved. This situation primarily reflects the difficulty of crystallising membrane proteins for structural studies. However, recent advances in methods for the expression and characterisation of transporters are beginning to redress the balance. It is the objective of this volume to bring together some of these new (and old!) approaches, so that those interested in transport can take advantage of the opportunities presented by genome sequencing information, and gain a better understanding of the molecular mechanisms of their favourite transporters. The volume starts with a topic at the heart of all studies of transporters, the methods needed for the assay of transport itself, both in whole cells and in membrane vesicles (Chapter 1). This methodology is built upon in Chapter 2, which describes methods for the reconstitution of membrane transporters. While concentrating on a single transporter, a Ca2+-ATPase which can be purified in large amounts from sarcoplasmic reticulum, this chapter illustrates approaches that are applicable to membrane transporters in general, such as use of different detergents for membrane solubilisation, and the methods that can be used to incorporate transporters into lipid vesicles suitable for transport measurements. The next four chapters move into the realms of cloned transporter genes and describe a number of different expression systems for membrane transporters, each with its own particular advantages and disadvantages. For example, Chapter 3 details the use of Xenopus oocytes for the expression and characterisation of V

PREFACE

transporters. This system has been enormously successful not only for the characterisation of cloned transporters, in particular from eukaryotes, but also for cloning these transporters by expression. However, though ideal for most transport experiments, oocyte expression does not yield sufficient protein for structural studies, and so the next three chapters describe methods for larger-scale expression of eukaryotic transporters in yeast (Chapter 4) or insect cells (Chapter 5), and of prokaryote transporters in Escherichia coli (Chapter 6). Included in these chapters are descriptions of methods for purification of the expressed proteins, in particular through the addition of affinity tags by recombinant DNA approaches. Once a transporter has been expressed and purified, the way is open for detailed investigation of its structure/function relationships. Methods available include fluorescence spectroscopy, described in detail in Chapter 7, and identification of substrate binding sites by photoaffinity labelling. As Chapter 8 describes, such labelling approaches are also extremely useful for the investigation of the subcellular locations and trafficking of transporters in whole cells, processes which play a key role in the physiological regulation of transport. Additional information on the mechanism of transporter function can also be gained by site-directed mutagenesis of residues that may play a part in substrate binding or other functions, provided that likely residues can be identified as targets for mutagenesis. Here, molecular modelling can provide important clues, and Chapter 9 reviews the methods currently available for computer analysis of membrane protein sequences to this end. The combination of these modelling approaches with site-directed mutagenesis and functional studies offers a powerful means of probing structure/function relationships in membrane transporters, and is likely to remain the mainstay of transporter research for many years. However, an understanding of the mechanism of transport at the molecular level will ultimately depend upon a detailed knowledge of the three-dimensional structures of the transporters concerned. Hence, the volume concludes with two chapters that detail methods for the 2-dimensional (Chapter 10) and 3-dimensional (Chapter 11) crystallisation of membrane proteins for electron and x-ray diffraction analysis. While such approaches are not for the faint-hearted, these chapters show that success is possible, and that crystallisation trials can be conducted using modest equipment in any laboratory. It is my hope that by setting out simple protocols, this book will encourage more researchers to venture into the study of membrane transporters, such that our understanding of these fascinating proteins comes closer to that already available for their water-soluble cousins. Finally, and most importantly, I wish to thank all of the authors for their contributions and patience in bringing this project to fruition. Leeds June 2000

VI

Stephen A. Baldwin

Contents

Protocol list xi Abbreviations xv 1 Assay of membrane transport in cells and membrane vesicles i Simon M. Jarvis 1 2 3 4 5 6

Introduction 1 Principles for measurements of transport rates 1 Transport techniques 3 Plasma membrane vesicles and transport 8 Kinetic analysis of facilitated-diffusion and co-transport systems 13 Determination of the driving forces for symporters—ion gradients and membrane potential 16

2 Reconstitution of membrane proteins: the Ca2+-ATPase of sarcoplasmic reticulum 21 Anthony G. Lee

1 Introduction 21 2 Choice of detergent 22 3 Purification of the Ca2+-ATPase from skeletal muscle sarcoplasmic reticulum 25 4 ATPase assay 29 5 Reconstitution of the Ca2+-ATPase 30 6 Spectrophotometric assay of Ca2+ accumulation 39 7 Calculation of the internal volume of the vesicles 39 8 Simulation of Ca2+accumulation 41 3 The Xenopus oocyte expression system for the cDNA cloning and characterization of plasma membrane transport proteins 47 Sylvia Y. M. Yao, Carol E. Cass, and James D. Young 1 Introduction 47

vii

CONTENTS

2 The Xenopus oocyte system 48 3 Isolation and size-fractionation of poly(A)+ RNA (mRNA) from mammalian tissues 53 4 Preparation of plasmid cDNA libraries suitable for in vitro transcription of RNA and expression in Xenopus oocytes 58 5 Screening cDNA libraries by functional expression selection in Xenopus oocytes 60 6 Functional and molecular characterization of transporter-encoding cDNAs 63 7 Conclusions 76 4 Expression of foreign transport proteins In yeast 79 N. SauerandJ. Stolz 1 2 3 4 5 6

Introduction 79 Expression of foreign genes in yeast: an overview 79 Expression in Saccharomyces cerevisiae 81 Expression in Schizosaccharomyces ponibe 98 Expression of membrane proteins in Pichia pastoris 101 Future perspectives 102

5 Baculovirus-mediated overexpression of transport proteins 707 Gary J. Litherland and Stephen A. Baldwin \ 2 3 4

Introduction 107 An overview of baculovirus-mediated expression systems 108 Practical aspects of the expression procedure 128 Recent developments in and alternative strategies for insect cell expression 138

6 The amplified expression, identification, purification, assay, and properties of hexahistidine-tagged bacterial membrane transport proteins 141 Alison Ward, Neil M. Sanderson, John O'Reilly, Nicholas G. Rutherford, Ben Poolman, and Peter J. F. Henderson 1 Introduction 141 2 Plasmids and E. coli host strains used in the amplified expression of membrane transport proteins 143 3 Growth conditions and detection of amplified membrane transport protein expression 144 4 Detergent choice and solubilization of integral membrane proteins 150 5 Purification of (His)6-tagged proteins 152 6 Reconstitution and activity assays of purified membrane protein 155 7 Physical properties of purified membrane protein 159 8 Conclusions 164 Vlll

CONTENTS 7 Spectroscoplc and kinetic approaches for probing the mechanisms of solute transporters 167 Adrian R. Walmsley 1 Introduction 167 2 Fluorescence spectroscopy for monitoring changes in the conformation of membrane transporters 767 3 Equilibrium studies of ligand binding to membrane transporters 169 4 The kinetics of ligand binding and translocation 172 5 Extrinsic probes to monitor transporter conformational changes 185 6 A steady-state approach to determining rate constants governing the translocation cycle 185 7 Thermodynamics 189 8 Detection and analysis of glucose transporters using photolabelllng techniques 193 Alison K GtZlmgham, Franfoise Koumanov, Makoto Hashimoto, and Geoffrey D. Holman 1 Methods for photolabelling glucose transporters 193 2 Photoactivation methods 194 3 Detection of the covalent incorporation of photolabels into glucose transporter isoforms 197 4 Biotinylated photolabels 204 9 Computer prediction of transporter topology and structure 209 Rong-I Hong and Mark S. P. Sansom 1 2 3 4 5

Introduction 209 Database searching and sequence alignment 210 Prediction of transmembrane helices 214 Example—B. subtilis ABC transporters 217 Conclusions 226

10 Two-dimensional crystallization of membrane proteins 229 Philippe Ringler, Bernard Heymann, and Andreas Engel 1 Introduction 229 2 Two-dimensional crystallization 230 3 Analysis of the result of 2-D crystallization by electron microscopy 257 11 Crystallization of membrane proteins 269 Tina D. Howard, Katherine E. McAuley-Hecht, and Richard J. Cogdell 1 2 3 4

Introduction 269 Crystallization techniques 270 Case studies 273 Preparing crystals for data collection 295 IX

CONTENTS

5 Screening protocols for the crystallization of new membrane proteins 298 6 Crystal packing in membrane proteins 299 7 Useful websites 302 A1 List of suppliers 309 Index 317

X

Protocol list

Transport techniques for cells Transport assays for adherent monolayers of cultured cells 3 Transport assays on suspended cells using the oil-stop or inhibitor oil-stop technique 6 Plasma membrane vesicles and transport Preparation of intestinal brush-border membrane vesicles 9 Transport by membrane vesicles determined by rapid filtration 10 Determination of the driving forces for symporters—ion gradients and membrane potential Determination of the intracellular pH in suspended cells 18 Purification of the Ca2+-ATPase from skeletal muscle sarcoplasmic reticulum Preparation of sarcoplasmic reticulum 25 Purification of the Ca2+-ATPase from sarcoplasmic reticulum 28 ATPase assay Assay of ATPase activity 29 Reconstitution of the Ca2+-ATPase Reconstitution of the Ca2+-ATPase into membrane fragments 31 Reconstitution into pre-formed large unilamellar vesicles (LUVs) 36 Reconstitution into sealed vesicles using cholate and deoxycholate 38 Spectrophotometric assay of Ca2+ accumulation Spectrophotometric determination of Ca2+ accumulation 40 Simulation of Ca2+ accumulation The subroutine necessary for simulation of Ca accumulation using FACSIMILE 43 The Xenopus oocyte system Isolation of Stages V-VI oocytes 49 Microinjection procedure 52 Isolation and size-fractlonatlon of poly(A)+ RNA (mRNA) from mammalian tissues Isolation of total RNA by the guanidinium thiocyanate and CsCl method 54 Isolation of poly(A)+ RNA by oligo(dT)-cellulose affinity chromatography 55 Size-fractionation of poly(A)+ RNA 56 Preparation of plasmid cDNA libraries suitable for in vitro transcription of RNA and expression in Xenopus oocytes 59 Denaturing agarose gel electrophoresis 59 XI

PROTOCOL LIST Functional and molecular characterization of transporter-encoding cDNAs

Subcloning a cDNA into the vector pSP64T 64 Radiotracer flux assay 65 Isolation of total oocyte membranes 72 Western blot analysis 73 Expression in Saccharomyces cerevisiae

Media for Saccharomyces cerevisiae 82 Transformation of Saccharomyces cerevisiae 89 Small-scale isolation of yeast total membranes 92 Uptake experiment with a radiolabelled substrate 93 Large-scale preparation of yeast total and plasma membranes 95 Purification of biotinylated proteins with immobilized avidin 97 Expression in Schlzosaccharomyces pombe

Transformation of Schizosaccharomyces pombe 100 Practical aspects of the expression procedure

Monolayer culture of insect cells 119 Suspension culture of Sf9 cells 320 Storage and resuscitation of insect cells 121 Transposition of recombinant genes into bacmid DNA 125 Isolation of recombinant bacmid DNA 126 Transfection and co-transfection of insect cells with baculovirus DNA using liposomal transfection reagents 328 Co-transfection of insect cells using calcium phosphate 129 Measurement of viral titre, and purification of recombinants, by plaque assay 331 Estimation of viral titre by cell-lysis assay 133 Amplification and storage of recombinant baculovirus 334 Measurement of solute transport into insect cells 136 Preparation of insect cell membranes 137 Growth conditions and detection of amplified membrane transport protein expression in Escherichia col!

Batch culture of recombinant E. coli for the overexpression of membrane proteins 145 Preparation of E. coli mixed membranes using water lysis 348 Separation of the inner and outer bacterial membrane fractions 349 Detergent choice and solubilization of Integral membrane proteins

Solubilization of bacterial membranes containing (His)6-tagged protein 352 Purification of (His)6-tagged proteins

Purification of (His)6-tagged protein using Ni-NTA agarose affinity chromatography 354 Reconstitution and activity assays of purified membrane protein

Reconstitution of detergent-solubilized membrane proteins into E. coli liposomes by detergent dilution 355 Reconstitution of membrane protein using Bio-Beads 156 Counterflow assay for activity of reconstituted GalP(His)6 protein 157 Physical properties of purified membrane protein

Circular dichroism (CD) spectroscopy of reconstituted and detergent-solubilized membrane protein 161 xn

PROTOCOL LIST

Fourier-transform infrared (FTIR) spectroscopy of proteoliposomes 162 Preparation of solubilized membrane protein for mass spectrometry 163 Equilibrium studies of ligand binding to membrane transporters

Titration of GalP with forskolin 171 The kinetics of llgand binding and translocatlon

Stopped-flow mixing experiments 175 A steady-state approach to determining rate constants governing the translocation cycle

Determination of the temperature dependence of the steady-state parameters for cellular transport 187 Photoactlvation methods

Photolabelling glucose transporters with [3H]cytochalasin B 196 Photolabelling glucose transporters using [3H]ATB-BMPA 198 Detection of the covalent incorporation of photolabels Into glucose transporter Isoforms

Immunopretipitation of the photolabelled glucose transporter isoforms 201 Solubilization of gels crosslinked with bis-acrylamide 203 Blotinylated photolabels

Detection of biotinylated GLUT4 using streptavidin precipitation 205 Detection of biotinylated GLUT4 by immunoprecipitation and detection with Amdex™ streptavidin-HRP 206 Example—B. subtllls ABC transporters

Protein sequence analysis 218 Prediction of transmembrane helices 221 Residue periodicity analysis 223 Two-dimensional crystallization of membrane proteins

Preparation of lipid stock solution in detergent-containing buffer 237 Exchange of detergent using a Centricon concentrator device 242 Exchange of detergent using size-exclusion gel filtration on Sephadex G-200 242 Exchange of detergent using sucrose-gradient centrifugation 243 Free (monomeric) detergent concentration measurement using the falling-drop weight method 244 Determination of the free detergent concentration using the sitting-drop method 245 Determination of phospholipid with ammonium ferrothiocyanate 246 Determination of phospholipid by phosphate content 247 Enzymatic determination of choline-containing phospholipids 247 Bicinchoninic acid (BCA) protein assay 248 Tubular crystallization of photosystem-II core complex (PSII) using dilution 251 Pre-treatment of the dialysis membranes 253 Microdialysis set-up using Eppendorf tubes 254 2-D crystallization procedure using Bio-Beads SM2 256 Phospholipase A2 treatment of 2-D crystals 257 Analysis of the result of 2-D crystallization by electron microscopy

Preparation of carbon-parlodion composite films on copper grids 259 Negative staining 259 Correlation averaging of OmpC 2-D crystals 263 Xlll

PROTOCOL LIST Three-dimensional crystallization of membrane proteins Purification of Rps. acidophila 10050 LH2 275 Detergent exchange by ultrafiltration in Rps. acidophila 10050 LH2 276 Crystallization of Rps. acidophila 10050 LH2 in B-OG 277 Purification of RC from purple photosynthetic bacteria 280 Crystallization of trigonal crystals 283 Preparing crystals for data collection Preparation of 'artificial mother liquor' 296 Equilibration of Rps. acidophila 10050 LH2 crystals prior to cryocooled data collection 297

Abbreviations

[A] InA ABC

AcMNPV adhl adhl-Pro AEBSF AML AMS AMP-PNP AmpR ANS AOX1 ARS AQP AS ASA-BMPA ATB-BMPA a.u. AZT BAT Bchla BB-BMPA BCECF BCECF/AM BES BHK Bio-ATB-BMPA

Bio-LC-ATB-BMPA

concentration of activator constant of integration ATP binding cassette Autographa californica multiple nuclear polyhedrosis virus alcohol dehydrogenase gene promoter of the adhl gene 4-(2-aminoethyl)-benzenesulfonylfluoride artificial mother liquor ammonium sulfate 5'-adenylylimidodiphospate ampicillin resistance gene 8-anilino-l-napthalene-sulfonate alcohol oxidase autonomous replication sequence aquaporin ammonium sulfate azidosalicoyl-l,3-bis(D-mannos-4-yloxy)-2-propylamine 2-N-[4-(l-azi-2,2,2-trifluoroethyl)benzoyl]-l,3-bis(D-mannose-4yloxy)-2-propylamine asymmetric unit 3' -azido-3 '-deoxythymidine broad-specificity amino acid transporter bacteriochlorophyll a 2-N-(4-benzoyl)benzoyl-l,3-bis(D-mannos-4-yloxy)-2-propylamine 2', 7' -bis(carboxyethyl)-5,6-carboxylfluorescein 2' ,7'-bis-(carboxyethyl)-5,6-carboxyfluorescein acetoxymethyl ester N,N-bis-[2-hydroxyethyl]-2-aminoethane sulfonic acid baby hamster kidney 4,4'-0-[2-[2-[2-[2-[2-(biotinylamino)ethoxy]ethoxy]ethoxy]-4-(l-azi2,2,2,-trifluoroethyl)benzoyl]amino-l,3-propanediyl]bis-D-mannose 4,4'-0-[2-[2-[2-[2-[2-[6-(biotinylamino)hexanoyl]amino]ethoxy] ethoxy]ethoxy]-4-(l-azi-2,2,2,-trifluoroethyl)benzoyl]amino-l,3propanediyl bis-D-mannose

xv

ABBREVIATIONS BLAST BLOSUM Bluo-gal BMPA BR Brij 35 Brij 58 BSA BV Bz C. CAA CCAC CCCP CD CEN CnEm C8E4 C12E9, etc. CFTR CHAPS CHAPSO C-HEGA-8 C-HEGA-9, etc. Cg-HESO CIP CL CMC COX CTAB CTF CYMAL-6 CYMAL-5, etc. 2-D 3-D DDAO DDBJ DDM DE52 DEPC B-DG DGDG DHPC di(C14:l)PC di(C18:l)PA

xvi

Basic Local Alignment Search Tool Best Local SUMmed alignment percentages 5-bromo-3-indolyl-B-D-galactoside l,3-bis(D-mannos-4-yloxy)-2-propylamine bacteriorhodopsin C-12^23 C

16E20

bovine serum albumin budded virus benzamidine Chloroflexus casamino acids Canadian Council on Animal Care carbonyl cyanide m-chlorophenylhydrazone circular dichroism centromeric sequence alkylpolyoxyethylene octyltetraoxyethylene dodecylnonaoxyethylene, etc. cystic fibrosis transport-regulator protein 3-[ (3-cholamidopropyl)-dimethylammonio]-l-propane sulfonate 3-[(3-cholamidopropyl)-dimethylammonio]-2-hydroxy-l-propane sulfonate cyclohexylethylethanoyl-N-hydroxyethylglucamide cyclohexylpropylethanoyl-N-hydroxyethylglucamide, etc. octyl(hydroxyethane)sulfoxide calf intestinal alkaline phosphatase cardiolipin critical micellar concentration cytochrome c oxidase cetyltrimethylammonium bromide contrast transfer function cyclohexyl-hexyl-p-D-maltoside cyclohexyl-pentyl-p-D-maltoside, etc. two-dimensional three-dimensional decyldimethylamine oxide; dimethyldecylamine-N-oxide DNA Data Bank of Japan n-dodecyl-B-D-maltoside diethylaminoethyl cellulose diethylpyrocarbonate B-decylglucoside; n-decyl-B-D-glucopyranoside digalactosyldiacylglyceride diheptanoyl-sn-phosphatidylcholine dimyristoylphosphatidylcholine dioleoylphosphatidic acid

ABBREVIATIONS

di(C18:l)PC di(C18:l)PE diS-C3-(5) DM DMEM DMPC DMSO DOPC DOPG 8-DOPSO DOTAP DPPC DTE DTT Dx E-64 Ea ECL ECV EDTA ee egg PC EGlc EGTA es ESI-MS F0 AF AFmax 4F2 FaR FBS FCCP FTIR AG° GalP GCG GLUT1 GLUT4 GPI GTH buffer h AH° HBSS hCNTl

dioleoylphosphatidylcholine dioleoylphosphatidylethanolamine dipropylthiadicarbocyanine p-decyl maltoside; n-decyl-p-D-maltopyranoside Dulbecco's Modified Eagle's Medium dimyristoylphosphatidylcholine dimethylsulfoxide dioleoylphosphatidylcholine dioleoylphosphatidylglycerol 2,3-dihydroxypropyloctyl sulfoxide N-[l-(2,3-dioleoyloxy)propyl]-N,N,N-trimethylammonium methylsulfate dipalmitoylphosphatidylcholine dithioerythritol dithiothreitol 1,4-dioxane trans-epoxysuccinyl-L-leucylamido-(4-guanidino)butane activation energy enhanced chemiluminescence extracellular virus ethylenediaminetetraacetic acid equilibrium exchange egg-yolk phosphatidylcholine phospholipids 4,6-ethylidene-D-glucose ethylene glycol bis-(p-aminoethylether)-N,N,N',N'-tetraacetic acid equilibrative nitrobenzylthioinosine-sensitive electrospray ionization mass spectrometry initial fluorescence change in fluorescence at time t maximum fluorescence change 4F2 cell-surface antigen formaldehyde resistance fetal bovine serum carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone Fourier-transform infrared Gibbs free energy (kJ/mol) D-galactose-proton symporter from Escherichia colt Genetics Computer Group, Inc. mammalian glucose transporter isoform 1 insulin-sensitive glucose transporter glycosyl phosphatidylinositol guanidinium thiocyanate homogenization buffer Planck's constant standard enthalpy change (kJ) Hanks'balanced salt solution human concentrative nucleoside transporter 1 xvii

ABBREVIATIONS

HECAMEG 6-O-(N-heptylcarbamoyl)-methyl-a-D-glucoside HEGA-8 octanoyl-N-hydroxyethylglucamide hENTl human equilibrative nucleoside transporter 1 Hepes 4-(2-hydroxyethyl)-l-piperazineethanesulfonic acid B-HG B-hexyl glucoside; n-hexyl-B-D-pyranoside (His)6 hexahistidine HPLC high-performance liquid chromatography HPTO heptane-l,2,3-triol HRP horseradish peroxidase HTG n-heptyl-B-D-thioglucopyranoside I inhibitor LAPS-forskolin 3-iodo-4-azidophenethylamido-7-0-succinyldeacetyl-forskolin IC50 concentration of inhibitor that inhibits by 50% IPTG isopropyl-B-D-thiogalactoside JIPID Japanese International Protein Sequence Database Jmax maximum flux in a transport experiment association, dissociation, and isomerization rate constants Kn kB Boltzmann constant kCB OFF dissociation rate constant for cytochalasin B association rate constant for cytochalasin B KCBON kobs measured or apparent rate constant koff apparent dissociation rate constant kon apparent association rate constant K1 dissociation constant for isomerization of a protein-ligand complex Kd dissociation constant Kj inhibition constant KI equilibrium constant for a protein isomerization step K Ii affinity of the inward-facing binding site of GLUT1 for 4,6ethylidene-o-glucose KIO affinity of the outward-facing binding site of GLUT1 for 4,6ethylidene-D-glucose KM half-saturation or Michaelis constant Kn concentration of activator at which the flux is 50% of the maximum (Jmax), raised to the power n, where n is the activator/substrate stoichiometry Ks apparent dissociation constant for sugar binding Kso dissociation constant for substrate outside the cell L ligand (in kinetic schemes) LacZ B-galactosidase LomB maltoporin LB Luria-Bertani medium LDAO lauryldimethylamine oxide; dimethyldodecylamine-N-oxide LH1 or LHI light-harvesting complex 1 (bacterial) LH2 or LHII light-harvesting complex 2; light-harvesting antenna complex (bacterial)

ABBREVIATIONS

LHCII LHCP LiAc LM LPPC LPR Lubrol PX LUV MALDI-MS MBM MCS MEGA-8 MEGA-9 MEGA-10 Mes MIP MIPS MLV MME MNPV m.o.i. MOMP Mops MPD MWCO NaCac NBD NBMPR

light-harvesting complex II (plant) light-harvesting core protein (apoprotein) lithium acetate lauryl-maltoside; n-dodecyl-p-D-maltopyranoside p-linoleoyl-7-palmitoyl-L-a-phosphatidylcholine lipid:protein ratio C12 and 14E9 5 large unilamellar vesicle matrix-assisted laser desorption mass spectrometry modified Barth's medium multiple cloning site n-octanoyl-N-methylglucamide n-nonanoyl-N-methylglucamide n-decanoyl-N-methylglucamide 2-(N-morpholino)ethanesulfonic acid major intrinsic protein Munich Information Center for Protein Sequences multilamellar lipid vesicle monomethyl ether multiple nuclear polyhedrosis virus multiplicity of infection major outer membrane protein 3-(N-morpholino)propanesulfonic acid 2-methyl-2,4-pentanediol molecular weight cut-off sodium cacodylate nucleotide binding domain nitrobenzylthioinosine (6-[#(4-nitrobenzyl)thio]-9-p-oribofuranosylpurine) NDAO nonyldimethylamine oxide; dimethylnonylamine-N-oxide NG n-nonyl-fJ-D-glucopyranoside Ni-NTA nickel-nitrilotriacetic acid NMG+ N-methyl-D-glucamine/HCl NMR nuclear magnetic resonance (imaging) nmt no message in thiamine NOGA n-octanoyl-p-D-glucosylamine NorA norfloxacin resistance protein NP-40 Nonidet P-40 NPV nuclear polyhedrosis virus ODAO dimethyloctylamine-N-oxide p-OG or OG B-octyl glucoside; n-octyl-B-D-glucopyranoside B-OGal B-octyl galactoside, n-octyl-B-D-galactopyranoside octyl-POE octylpolyoxyethylene, largely C8E5 Omp outer membrane protein OmpF outer membrane protein porin OpMNPV Orgyia pseudotsugata multicapsid nucleopolyhedrosis virus xix

ABBREVIATIONS

ORF OS OTG P PA PAM PBS PC PCA PCR PE PEG PEG 2000 PEG 3500 PEG 2000 MME p.f.u. PGG PhoE Pi p.i. Pipes PIR PLA2 PMA1 PMAl-Pro PMAl-Ter PMSF pmt 8-POE POPC pPolh PS PSI RC PSII RC PTEN PTS R Kb. RC rCNTl rENTl Ree

open-reading frame octanoyl sucrose n-octyl-B-D-thioglucopyranoside protein (in kinetic schemes) phosphatidic acid Point Acceptable Mutations per unit time phosphate-buffered saline phosphatidylcholine pipecolinic acid; 2-piperidine carboxylic acid polymerase chain reaction phosphatidylethanolamine polyethylene glycol polyethylene glycol, Mr = 2000 polyethylene glycol, Mr = 3500, etc. polyethylene glycol monomethyl ether, Mr = 2000 plaque-forming units piperazine/glycylglycine phosphoporin inorganic phosphate post-infection piperazine-N,N'-bis(2-ethanesulfonic acid) Protein Information Resource phospholipase A2 plasma membrane ATPase gene promoter of the PMA1 gene terminator of the PMA1 gene phenylmethylsulfonyl fluoride photomultiplier tube octyl polyoxyethylene palmitoyloleoylphosphatidylcholine polyhedrin promoter phosphatidylserine photosystem I reaction centre photosystem II reaction centre platinum ethylene diamine dichloride peroxisomal targeting sequence the gas constant Rhodobacter reaction centre rat concentrative nucleoside transporter 1 rat equilibrative nucleoside transporter 1 resistance parameter (1/Vmax) for membrane transport performed

r.p.m. Rps. XX

revolutions per minute Rhodopseudomonas

under equilibrium exchangeconditionsR

ABBREVIATIONS

RS1 RT

s AS0 SB-12

SDS-PAGE SEM SGLT1 SNPV SR Sulfo-NHS-biotin SUV T

[T] TBS-T TEST buffer TCA TE buffer TEM Thesit (C12E9) thio-p-OG TLC TM a-Toc UDAO UM v VDAC max

X-gal YNB YPD

regulatory subunit 1 room temperature Resistance parameter (l/Vmax) for membrane transport performed under zero-trans conditions substrate standard entropy change (kJ/mol) lauryl sulfobetaine; Zwittergent-3,12™; dodecyl-DAPS; dodecylN,N-dimethylammonio-3-propane sulfate sodium dodecyl sulfate-polyacrylamide gel electrophoresis standard error of the mean sodium-dependent glucose transporter 1 single nuclear polyhedrosis virus sarcoplasmic reticulum sulfosuccinirnidobiotin small unilamellar vesicle absolute temperature (kelvin) concentration of transporter Tris-buffered saline-Tween Tris base-sodium chloride-Tween-20 buffer trichloroacetic acid Tris/EDTA buffer transmission electron microscope dodecylnonaoxyethylene thio-fJ-octyl glucoside; n-octyl-p-D-thioglucopyranoside thin-layer chromatography transmembrane, transmembrane segment DL-a-tocopherol undecyldimethylamine oxide; dimethylundecylamine-N-oxide n-undecyl-B-D-maltopyranoside rate of transport voltage-dependent anion channel maximal rate of transport 5-bromo-4-chloro-3-indolyl-p-D-galactoside yeast nitrogen base yeast peptone dextrose

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Chapter 1 Assay of membrane transport in cells and membrane vesicles Simon M. Jarvis Department of Biosciences, University of Kent, Canterbury, Kent CT2 7NJ

1 Introduction Membrane transport is a vectorial process involving the translocation of ions or solutes from one compartment to another. With the exception of simple diffusion, specific proteins catalyse transport. The last decade has seen an explosion in studies applying molecular biology techniques to the identification, classification, localization, and regulation of the transport proteins (see other chapters in this book). An ultimate goal of many studies is to relate structure to function. Transport function is characterized by studying properties of substrate specificity, species of co-substrates and counter-substrates, inhibitors, activators, pH dependence, permeant concentration dependence, and temperature dependence. These functional characteristics rely upon measurements of the initial rates of transport. This chapter will mainly cover the basis for transport experiments with radiolabelled isotopes in suspended cells, adherent cultured cells, and membrane vesicles. The advantages of using radioactive solutes are that the uptake of the solute of interest is directly measured, radiolabelled compounds are widely available, and the detection of radioactivity is straightforward. In addition, the use of fluorescent dyes to determine the driving forces of certain membrane transporters will also be highlighted.

2 Principles for measurements of transport rates 2.1 General To measure the rate of movement of a transported substrate across a biological membrane one needs to measure the amount of substrate that has entered or exited the cell or vesicle as a function of time. As mentioned above, the commonest way to estimate the amount of substrate is to use a radiolabelled substrate, although other methods have been used—such as the movement of fluorescent substrate, changes in cell volume as water follows the movement of an osmotically active substance, and a change in pH. The measurement of time

1

SIMON M. JARVIS

implies that the substrate and membrane are brought together and subsequently separated at a defined instant. The time interval needed to obtain a measurement of the initial rate of transport can vary from seconds to minutes. Even for the same permeant in the same cell type, but from different species, there can be major differences, e.g. the uptake of uridine by erythrocytes (1).

2.2 Metabolized permeants The relationship between transport and any subsequent metabolism of the substrate has important methodological implications. Inward fluxes of a permeant are usually measured from time courses of the accumulation of the permeant. In cells that metabolize the permeant, time courses can be complex due to: (a) the permeation step being reversible and in some cases faster than the subsequent metabolic steps, raising the possibility of significant backflux of the permeant during the stopping process; and (b) multiple metabolic pathways for the permeant. For example, a review of the early (pre-1980) nucleoside transport literature has shown that many of the studies with metabolized nucleosides reflected cellular metabolism rather than transport per se, due to the failure to measure initial rates of nucleoside accumulation (2). By definition, an initial, constant rate of permeant uptake (transport plus metabolism) reflects its transport rate regardless of subsequent metabolism. The challenge for the investigator is to ensure that the measured rate of uptake is indeed the initial rate. A key component to achieving this goal is to use an experimental protocol that allows rapid sampling both to initiate transport and to terminate the assay over time periods of 1 sec, if required. In addition, the assay needs to be sufficiently sensitive to detect intracellular accumulation of the permeant at concentrations of 10% and less of the extracellular permeant.

2.3 Non-metabolized permeants The problems of metabolism noted above can be eliminated by choosing a metabolically inert permeant or conditions where metabolism is impaired; transport will then become the determinant of the time course of permeant accumulation for those permeants that are substrates for transporters. For some permeants this condition arises naturally, e.g. uridine and thymidine are not metabolized by human erythrocytes (3). In other cases, a poorly metabolized analogue is used, e.g. formycin B is a C-nucleoside analogue of inosine and 3-0methylglucose an analogue of glucose (4). However, a prerequisite to the use of a non-metabolizable analogue is to demonstrate that the analogue behaves in the same manner as the physiological permeant. Whereas formycin B is a permeant of the equilibrative, nitrobenzylthioinosine (NBMPR; 6-[(4-nitrobenzyl) thio]-9-p-D-ribofuranosylpurine)-sensitive (es) nucleoside transporters of mammals, it is not a permeant for the pyrimidine-preferring N2 Na+-dependent nucleoside transporters (5). Alternatively, metabolism could be reduced or eliminated by using ATP-depleted and enzyme-deficient cells. However, if transport is dependent on ATP for example, conclusions drawn from these conditions may be wrong. 2

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

3 Transport techniques To measure permeant fluxes across biological membranes, the uptake interval is ended by a variety of procedures including: (a) separating cells and vesicles from the permeant by centrifugation, filtration, or rinsing of monolayers; (b) reducing the temperature; (c) addition of a transport inhibitor; or (d) combinations of these,

3.1 Adherent monolayers of cultured cells Transport by monolayers of cells on coverslips or in culture dishes (usually 24or 6-well trays) is initiated by the addition of radiolabelled permeant for a defined interval (see Protocol 1). Uptake is ended by rapid removal of the extracellular permeant followed by washing (dishes) or dipping (coverslips) in medium. Incubation times as short as 5-10 sec can be achieved with these procedures. Multiple wells can be processed at the same time by constructing a

Protocol 1 Transport assays for adherent monolayers of cultured cells Equipment and reagents • Cell-culture plastic ware and medium (with and without 10% foetal calf serum) (Gibco BRL, Becton Dickinson, Sigma) • Radioactive compounds including test permeant (generally tritium-labelled; 20-100 (i/mmol), and [14C]sucrose (-500 m (i/mmol) or inulin-[14C]carboxylic acid (~10 m (i/mmol) (Amersham) • Transport buffers, e.g.: - Na+-HBSS: 140 mM NaCl, 5 mM KC1. 4,2 mM NaHC03, 0.36 mM K2HPO4,1.3 mM CaCl2, 0.44 mM KH2PO4, 0.5 mM MgCl2, 10 mM Hepes pH 7.4;

- K+-HBSS: the same as Na*-HBSS but with the following alterations: NaHCO3 replaced with KHCO3 and NaCl replaced with KC1; - and NMG-HBSS: the same as K+ -HBSS but with 140 mM KC1 replaced by 140 mM N-methyl-D-glucamine/HCl (NMG - } 0.5 M NaOH 0.25% (w/v) trypsin/0.0356 (w/v) EDTA

Method 1. Culture cells according to the conditions suitable for the individual cell lines. Prepare 24-well culture plates for transport assays by seeding between 4 x 104 to 4 x 105 cells into each well. Allow the cells to grow to confluency in 1 ml of culture medium and change the medium every 2-3 days.

3

SIMON M. JARVIS Protocol 1 continued

2. Prior to the transport assay, incubate the monolayers for 30 min at 37 °C with serumfree culture medium to reduce the intracellular pool of substrates, if required. 3. Then rinse the monolayers three times with 1 ml atiquots of the transport buffer. Test for sodium dependency by washing the cells with KT-HBSS or NMG+-HBSS. 4. Initiate uptake by adding 0.2 ml of transport medium containing 10 uCi/ml of the 3 H-labelied permeant and 0.5 uCi of [14C]sucrose or inulin-[14C]carboxylic acid as an extracellular marker, and non-radioactive permeant to the desired concentration in the appropriate salt solution. Carry out determinations in triplicate. In inhibition studies, add the test compound and radiolabelled penneant simultaneously unless there is a need to pre-incubate the cells with the test compound to allow it to interact with the cells. Continue the uptake for a predetermined time with gentle shaking to avoid the creation of unstirred aqueous layers, and terminate by aspirating the transport solution 1 sec before the indicated time. Then rapidly wash the monolayer three times with 1 ml aliquots of ice-cold K+-HBSS containing, if available, a known transport inhibitor of the system being studied, 5. Determine the radioactivity associated with the monolayers at time zero by using ice-cold cells and transport medium. 6. Solubilize the monolayers in 200 ul of 0.5 M NaOH and assay for 3H and 14C.° 7. Count the cells in replicate cultures after treatment with 0.25% (w/v) trypsin/0.03% (w/v) EDTA to detach the cells," Express uptake rates either as moles per mg protein per time or as moles per 106 cells per time. "The 14C counts will represent trapped extracellular space and can be subtracted from the 3H counts after appropriate corrections for the differences in the level of radioactivity, to determine the cell-associated 3H counts. b The protein content of the wells can also be determined using standard procedures.

simple set of devices based on the 24-wcll tissue culture dish, which allows the simultaneous addition of permeant to all 24 wells and their subsequent washing (6). A major problem that can arise with this procedure is the possible loss of permeant during the washing step(s). This can be minimized by the use of cold medium (0-4oC and, if available, the inclusion of a transport inhibitor, e.g. dilazep for nuclcoside transport by certain mammalian nucleoside transporters (5). Some adherent cells are particularly susceptible to being totally or partially washed off the plastic culture vessels, especially if multiple washes are used. This can sometimes be overcome by testing cell-culture plasdcware from different suppliers or coating the plasticware with collagen or gelatin. In some cases the permeant may become trapped in the monolayer and so an estimate of the remaining extracellular space following the washing steps is essential. This is usually estimated using isolopically labelled inulin-[ 14 C]carboxylic acid or [14C|sucrose that can be added directly with a 3H-labelled permeant. The 4

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

cell-associated permeant radioactivity is the difference between the 3H and 14C counts.

3.2 Suspended cells: centrifugation The simplest centrifugation procedure is to pack the cells by centrifugation and wash off the extracellular radioactivity by repeated centrifugation and resuspension in isotope-free medium. Centrifugation should be performed using high-speed (15 000 g) microcentrifuges where cells can be pelleted within 10-20 sec. Low-speed centrifuges that take a significant time to sediment cells (~ 5 min) must not be used. Typically, four washes with 20 volumes of medium are required to reduce the trapped extracellular space from ~ 15% to < 0.001%. As is the case with adherent cells, possible problems of permeant loss and cell lysis during the washing process should be considered. These can be minimized by including a transport inhibitor and using ice-cold medium, but the method is only really applicable to those situations where the flux rate is slow (1). The shortest interval practicable with the centrifugation/washing method is 30 sec.

3.3 Suspended cells: oil-stop centrifugation A variant on the centrifugation method is to separate cells from the extracellular permeant by centrifugation through a layer of oil with a density less than that of the cells but greater than the incubation medium (see Protocol 2). The transport incubation takes place above the oil layer and the assay is terminated by starting the microcentrifuge. The rapid mixing of cell suspension and permeant solution can be achieved using separate pipettes or by means of a dual syringe device (2, 7). For many mammalian cells it takes 2 sec for the cells to be removed from the medium and this time must be added on to the uptake interval (time between mixing cells and radiolabelled permeant and the starting of the microcentrifuge) (8, 9). However, some cells take longer to centrifuge through the oil, e.g. 4 sec for procyclic forms of trypanosomes (10), and the time taken for each cell type must be determined as shown in Figure 1. During the passage of the cells through the oil, extracellular medium containing isotope will be trapped and carried through. The trapped space represents, on average, 10-20% of the total pellet water. It is thus important to estimate this trapped extracellular space as correction of this value is required to calculate the actual uptake of isotope. It is also desirable to measure the intracellular water as this will assist in determining whether equilibration between intracellular and extracellular water has occurred, and whether the transported permeant is capable of being concentrated within the cell. A refinement to the oil-stop centrifugation technique is to stop the transport assay by adding a transport inhibitor or ice-cold unlabelled permeant at a concentration vastly in excess of the isotope concentration. Immediately after addition of the stop solution, the cells are pelleted under the oil. This method has the advantage that transport can be stopped virtually instantaneously and that the time delay that occurs during centrifugation is eliminated (see Figure 1). 5

SIMON M. JARVIS

Protocol 2 Transport assays on suspended cells using the oil-stop or inhibitor oil-stop technique Equipment and reagents • Microcentrifuge (Beckman Instruments) • Oil mixtures composed of rt-dibutyl phthalate, mineral oil, or silicone fluid mixtures (Sigma, Merck)0 • Transport buffers (see Protocol 1) or other buffers of choice, e.g. Krebs-Ringer phosphate buffer, pH 7.4 • Radioactive compounds (see Protocol 1)

1.5 ml microcentrifuge rubes Ice-cold stop solution: assay buffer containing a transport inhibitor or excess unlabelled permeant High-density acid solution: e.g. 0.5 M perchloric acid in 10% (w/v) sucrose or 20% (w/v) perchloric acid Metronome

Method 1. Pipette 200 ul of the oil mixture at the correct density into 1.5 ml microcentrifuge tubes. 2. On top of the oil layer, pipette 100 ul of the transport medium containing radiolabelled permeant (usually 3H label at 10 uCi/ml). Initiate transport by adding 100 ul of suspended cells at a cell density between 106 to 108 per assay, depending on the volume of the cells, their transport activity, and their availability. 3. Terminate transport either by centrifuging the cells for 30 sec at 12000 g or by adding 1 ml of ice-cold stop solution followed by immediate centrifugation. Add (this is essential) the time taken for the cells to pellet through the oil layer in the absence of the addition of stop solution on to the uptake interval (see Figure 1), For short intervals (<10 sec), use a metronome for timing. 4. Remove the oil and aqueous layers by suction, and wipe the inside of the tube with absorbent paper. Dissolve the cell pellet in 200 u1 of 0.5 M NaOH and count for radioactivity,11 5. Estimate the trapped extracellular space by replacing the radiolabelled permeant with [14C]sucrose or inulin-[14C]carboxylic acid and repeating the procedure described above. 6. Obtain blank values (radioactivity that becomes associated with the cells during uptake intervals of 0 sec) by processing cell samples that have been simultaneously exposed to radiolabelled permeant and excess unlabelled permeant or inhibitor. 7. Calculate the intracellular volume of the cells from the distribution ratio of [3H]H2O and |14C|sucrose. Briefly, layer 100 ul of transport buffer containing both isotopes on top of the oil layer and then add 100 ul of cells at the same density as in step 2. After 1 min centrifuge the tube and process as described in step 4.

6

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

Protocol 2 8. To examine the metabolic fate of the transported permeant, underlay the oil layer with 50-100 |ul of the high-density acid solution. Remove the oil and aqueous layer ahove the oil as in step 4 and subject the acid extract to chromatographic analysis. a

For example: n-dibutyl phthalate is used for erythrocytes; a 7:1 (by vol.) n-dibutyl phthalate/ mineral oil mixture (d = 1.018 g/ml) for parasitic protozoa such as trypanosomes; and silicone fluid mixtures (Dow Corning 550 and 200/lcs in a ratio of 86:14 obtained from Merck) for many mammalian cells (d = 1.034 g/ml). Small adjustments in the density of the oil mixture can be made by varying the proportion of the two mixtures should the need arise. For a new cell type, it is recommended that the best oil mixture is tested for empirically by preparing a series of oil mixtures of slightly different densities and then evaluating whether the cells pellet cleanly through the oil, forming a pellet at the bottom of the microcentrifuge tube, and that the oil-water interface is sharp and stable. b Radioactivity associated with the cell pellet will represent both permeant that has entered the cells and that trapped in the extracellular space.

Figure 1 Time courses of adenosine influx by Trypanosoma brucei bmcei procyclic cells using the oil-stop or inhibitor oil-stop method. Transport at 22°C was initiated by adding 100 ul of procyclic cells (107 cells) to 100 ul of transport medium containing 2 uM [3H]adenosine layered on top of an oil mixture (density 1.018 g/ml, composed by mixing 7 volumes of n-dibutyl phthalate (1,043 g/ml) and 1 volume of mineral oil (0.84 g/ml)). Transport was terminated either by pelleting the cells under the oil layer (. '.) or by adding 1 ml of ice-cold assay buffer containing 4 mM adenosine followed by immediate centrifugation (•). The intercept on the ordinate for the two time courses was different. In the presnce of 4 mM adenosine, the time course intercepted with the estimate of the extracellular space. Without the addition of unlabelled adenosine, an intercept of -4 sec was observed, suggesting that it takes 4 sec for the procyclic cells to pellet and that transport continues during this period. (Adapted from ref. 10.)

7

SIMON M. JARVIS

In addition, contamination of the cell pellet with extracellular isotope is reduced by the inclusion of the stop solution diluting out the isotope before centrifugation. Potential problems that may arise are that the stop solution fails to act instantaneously, or the presence of excess unlabelled permeant outside the cells results in a stimulation of isotope efflux. These possibilities must be tested for and eliminated before proceeding to use stop solutions.

4 Plasma membrane vesicles and transport 4.1 Preparation of membrane vesicles Epithelial cells from the renal proximal tubule and the small intestinal villus differ from many other mammalian cells in that they have distinct plasma membrane domains: apical and basolateral. Membrane vesicles can be prepared from each of these domains and used to characterize and localize transporters in the epithelium. The starting material for vesicle preparations is usually minced or chopped tissue or intestinal scrapings, homogenized by a variety of methods including the use of a Dounce homogenizer, a Polytron® homogenizer or equivalent, or a nitrogen cavitation bomb (11, 12). A popular method used to prepare renal and intestinal brush-border membrane vesicles is by a divalentcation, differential centrifugation procedure (see Protocol 3), whereas densitygradient procedures using Percoll are used for preparing basolateral membrane vesicles (11). The membrane vesicles are closed structures that may be heterogeneous in size (diameter 0.1-0.3 um; volume 1-3 ul/rng membrane protein) and orientated in either the right-side out or inside-out mode. Over 90% of renal basolateral membrane vesicles are right-side out (11). Purity of the preparations is judged from the enrichment of enzyme markers, e.g. maltase, alkaline phosphatase, -y-glutamyl transpeptidase for renal and intestinal brush-border vesicles, and Na + , K+-ATPase for basolateral vesicles. The use of membrane vesicles, as opposed to isolated cells or established cell lines, offers a number of advantages. First, in many cases metabolism will be absent, but the lack of metabolism cannot be taken for granted, and experiments to assess the extent of metabolism are always necessary. For example, pyrimidine nucleosides are not metabolized by brush-border membrane vesicles prepared from kidney, but purine nucleosides are metabolized (11, 13). The second major advantage of vesicles in the analysis of transport phenomena is the ability to control the intra- and extravesicular medium at will, and thus the determination of initial rates of transport under well-defined driving forces.

4.2 Transport measurements using vesicles Although a number of different methods have been employed to measure the uptake of solute into vesicles—including dyes to monitor membrane potential or proton concentration changes, flow dialysis, light scattering, and ionsensitive electrodes—the commonest and most generally applicable method is that of rapid mixing and filtration (14). Vesicles are pre-loaded with solutions of 8

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

Protocol 3 Preparation of intestinal brush-border membrane vesicles Equipment and reagents • Polytron®homogenizer (Kinematics AG) or a glass/Teflon Potter homogenizer 0 (Braim B Medical Ltd) • High-speed centrifuge, e,g, a Beckman J2-21 centrifuge with JA-20 and JA-14 rotors (Beckraan Instruments)

• 50 ml syringe • Homogenization buffer: 300 mM mannitol. 1 mM Hepes/Tris pH 7.5 • Glass plate and microscope slides • 100 mMCaCl2

Method 1. Kill the animals by an approved procedure (decapitation/cervical dislocation or anaesthetic overdose), remove the smalt intestine and flush through with ice-cold 300 mM mannitol. 10 mM Hepes/Tris pH 7.5 using a 50 ml syringe. 2. Cut the small intestine lengthways, invert, and place on an ice-cold glass plate resting on ice. Using a glass slide, gently scrape the mucosa surface and suspend in 110 ml of ice-cold homogenization buffer. 3. Homogenize the intestinal scrapings using either the Potter homogenizer (30 strokes at 1500 r.p.m.) or the Polytron® (30 sec at setting 5). 4. Precipitate basolateral membranes by adding 12 ml of 100 mM CaCl2 and stir the mixture for 20 min at 4°C. Remove the basolateral membranes by centrifuging the homogenate for 10 min at 8000gat 4°C. 5. Collect the supernatant and re-centrifuge at 21000 g for 20 min, Resuspend the pellet in -60 ml of homogenization buffer, homogenize as in step 3 and centrifuge at 21 000 g. Perform all centrifugation steps at 4°C. 6. Resuspend the final pellet in ~1 ml of 300 mM mannitol, 10 mM Hepes/Tris pH 7.5 at a protein concentration of -10-20 mg/ml; either use fresh or store at -70°C until needed. 7. Determine the purity of the brush-border membranes by measuring the enrichment in specific enzyme markers, e.g. sucrase and alkaline phosphatase, compared to the initial homogenate (see ref. 11. 25, 26). Enrichment factors of 10-fold are common.

known composition and then rapidly mixed with incubation medium containing radioaclively labelled permeant and other constituents as required. After an appropriate time interval, stop solution is added and the vesicles collected on a filter that is subsequently washed and counted for radioactivity (see Protocol 4). tifflux from vesicles pre-loaded with non-metabolized substrate can be measured using similar procedures. 9

SIMON M. JARV1S

Protocol 4 Transport by membrane vesicles determined by rapid filtration Equipment and reagents • Filtration apparatus (MiHipore) • Nitrocellulose filters, 0.45 ^m pore size (Millipore) • Radioactive compounds (see Protocol 1) including [3H]uridine • Transport buffers: e.g. 10 mM Hepes/Tris pH 7.5 and 150 mM of an appropriate chloride salt, e.g. NaCl

• Stop solution: e.g. containing 100 mM mannitol, 200 mM NaCl, 1 mM phloridzin, and 1 mM Hepes/Tris pH 7,5 • Syringe fitted with a 25-gauge needle • 5 ml disposable test tubes • Metronome

Method 1. Obtain membrane vesicles (see Protocol 3) and pass through a 25-gauge needle to ensure vesiculation. 2. Pipette aliquots (10 ul) of the membrane suspension (100-200 ug of protein) into the bottom of 5 ml disposable test tubes. Pipette transport solutions (20 u1) containing radiolabelled permeant (e.g. 20 uCi [3H]uridme/ml} in 10 mM Hepes/Tris pH 7,5 and the appropriate salt on to the side of the test tube. 3. Initiate transport by vigorous mixing with a vortex mixer. Time the short incubation times using a metronome. 4. Terminate the incubation by adding 1 ml of ice-cold stop solution and immediately filter the diluted vesicles through pre-wetted nitrocellulose filters. Subsequently wash the filters with 5 ml of stop solution and air-dry before adding scintillation fluid. 5. Obtain blank values for the uptake assays, due to trapping of radioactivity on the nitrocellulose filter, by filtering the transport medium without the membrane vesicles. Measure transport at zero time using ice-cold transport medium and membrane vesicles. Subtract the blank value from measurements associated with membrane vesicles to determine uptake rates expressed as moles/ing protein per time.

Transport measurement!; using vesicles have been useful in demonstrating the existence of secondary active transport mechanisms in which the activator gradient is demonstrated ro drive the concentrativc accumulation of the solute. Figure 2 demonstrates the results of an experiment where the uptake of uridinc into intestinal brush-border membrane vesicles was studied in the presence and absence of an initial extravesicular to imravesicular sodium gradient. In the presence of a sodium gradient the imravesicular concentration of uridinc is 10

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

Figure 2 Demonstration of the 'overshoot' phenomenon for the uptake of uridine by rabbit intestinal brush-border membrane vesicles. Vesicles were prepared in 300 mM mannitol, 10 mM Hepes/Tris (pH 7.5) and the uptake of 4.4 uM [3H]uridine measured in the presence of inwardly directed gradients of 100 mM NaCI (•) or 100 mM choline chloride (O). (Taken from ref. 25.)

seen to transiently rise above (overshoot) its equilibrium value and then fall back to equilibrium as the sodium gradient dissipates. In the absence of a sodium gradient, no accumulation of uridine above equilibration is evident. This figure also demonstrates the need to determine initial rates of transport under conditions where the imposed driving forces have not changed significantly. Membrane potentials are typically controlled through the use of ion gradients and ionophores, e.g. valinomycin/K+ voltage clamp, where the potential is given by the Nernst equation. In view of the large surface area to volume ratio of vesicles, time intervals of a few seconds are likely to be needed for initial rate determinations. The rapid filtration technique is only valid if all transport ceases after adding the stop solution and no loss of radioactivity from the vesicles occurs during filtration and washing. Figure 3 illustrates the effect of different stop solutions on the retention of [3H]uridine by rat renal brush-border membrane vesicles. In this experiment renal membrane vesicles were incubated with [3H]uridine for 10 sec, then subsequently diluted 1:33 into various stop solutions and filtered immediately or after a 5-60 sec delay. Ice-cold stop solution containing 1 mM phloridzin was an effective stop solution with no significant change seen in intravesicular [3H]uridine content during the stopping and washing procedure. In contrast, ice-cold stop solution containing 1 mM HgCl2 or buffer alone resulted in the rapid loss of radioactivity from the vesicles, which was further 11

SIMON M. JARVIS

Rgure 3 Testing the effectiveness of various stop solutions on uridine uptake by rat renal brush-border membrane vesicles. Rat renal cortical brush-border membrane vesicles were incubated for 10 sec with [3H]uridine. At time zero, transport by the 30 ul vesicle suspension was stopped by adding I ml of stop solution (Tris/HCI buffer (pH 7.4), 100 mM mannitol, 100 mM NaCI) at room temperature ([ ]), or ice-cold stop solution (•), or of icecold stop solution containing 1 mM HgCI2 (O) or of ice-cold stop solution containing 1 mM phloridzin (•). The solutions were filtered immediately, or after a time delay, and subsequently washed with 5 ml of the appropriate stop solution. The radioactivity retained by the vesicles has been normalized to that obtained when the vesicles were immediately filtered with ice-cold stop solution containing 1 mM phloridzin. (Taken from ref. 26.)

enhanced when buffer at room temperature was employed. The use of an inappropriate stop solution will lead to uncontrolled variability in the assay and the failure to measure initial rates of transport. A further set of control experiments need to be performed to distinguish between uptake that constitutes transport into an intravesicular space and uptake that is due to binding of the substrate to the membrane. Uptake into an intravesicular space will be characterized by the equilibrium uptake being directly proportional to the intravesicular volume, which can be manipulated by increasing the medium osmolarity with impermeable solutes (see Figure 4). Binding of the radiolabelled solute to the nitrocellulose filter can also be a problem. Thus, blank values for radioactivity retained on the filter in the absence of membrane vesicles should be determined and subtracted from measurements of solute associated with the vesicles. If the blank value is high, pre-incubating the filters in unlabelled solute or 0.3% polyethylenimine can sometimes reduce it.

12

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

1/Osmolarity (OsM) Figure 4 Demonstration that the uptake of uridine by rabbit intestinal brush-border membrane vesicles is due to transport into an intravesicular space. Vesicles were prepared in 300 mM mannitol, 10 mM Hepes/Tris (pH 7.5) and suspended in a buffer containing varying concentrations of cellobiose to vary the extracellular osmolarity. Equilibrium uptake of 5 uM [3H]uridine was determined at 20 min and plotted as a function of the reciprocal of the extravesicular osmolarity. The linear relationship and the zero intercept on the vertical axis at infinite extravesicular osmolarity (zero intravesicular space) indicate that uridine associated with the vesicles is due to transport of uridine into an osmotically sensitive intravesicular space and that there is no significant binding of uridine to the membrane. (Taken from ref. 25.)

5 Kinetic analysis of facilitated-diffusion and co-transport systems Many different types of kinetic transport experiments can be performed, but in this part of the chapter I hope to indicate which of them are the most crucial for obtaining the information to characterize a transport system. The starting point for this analysis is the simple carrier model introduced by Lieb and Stein over 20 years ago for non-concentrative transport systems (15). In the simple carrier model, the permeant binding site on the unoccupied transport system is accessible to permeant molecules in the aqueous compartment at only one side of the membrane at any one time. 5.1 Facilitated-diffusion transporters Facilitated-diffusion transporters translocate the permeant down its concentration gradient until equilibration across the two sides of the plasma membrane 13

SIMON M. JARVIS

has been reached. A common experimental procedure is to measure unidirectional flux under zero-trans (zt) conditions, where the permeant concentration at one side of the membrane is varied while that at the opposite side (trans) is effectively fixed at zero. In the equilibrium exchange procedure the unidirectional flux is measured when the concentrations in the aqueous solutions on both sides of the membrane are identical, i.e. at equilibrium. Under these two experimental conditions the measured flux will vary between zero and some maximum value (Vmax). The concentration at which the flux is 50% of Vmax is called the half-saturation concentration, KM. The reciprocals of the maximum fluxes are termed resistance parameters, R, and the constraint of the simple carrier model is that R equilibrium exchange (ee) + ROo = Rzt influx + Rzt efflux; where Roo is the resistance parameter for the unloaded carrier and Rzt is the resistance parameter (l/Vmax) for membrane transport under zero-trans conditions. Thus, knowledge of the Vmax values will allow one to test whether the predictions of the simple carrier model are confirmed by the experimental data. The NBMPR-sensitive (es) nucleoside transporter in human erythrocytes conforms to all predictions of the simple carrier model (16). A typical set of experiments that may be performed is to compare the kinetic parameters for zero-trans influx and efflux to determine whether the transport system exhibits directional symmetry or asymmetry. For example, in freshly isolated human and guinea-pig erythrocytes the KM and Vmax values for zerotrans uridine influx and efflux were shown to be similar, demonstrating that uridine transport via the es nucleoside transporter in these cells is equally efficient in the influx and efflux modes (16, 17). Additional studies measuring the equilibrium-exchange kinetic parameters showed that for many nucleoside permeants of the erythrocyte es transporter the rate-limiting step in the transport cycle was the carrier switching the accessibility of its binding site from one side of the membrane to the other when in the empty state, i.e. ROO had the highest value of all the resistance parameters (16-18). Nevertheless, for some permeants, e.g. 2-chloroadenosine, the Roo/Ree ratio (which quantifies the differential rate of conformational changes of the substrate-loaded and empty carrier) was less than 1, indicating that the conformational change associated with the nucleoside-loaded carrier is dependent on the structure of the permeant (18).

5.2 Co-transporters Active transport is a process that can effect the net transfer of a solute against its electrochemical gradient, and can be divided into primary and secondary active systems. In secondary active transport, the negative free-energy change required to bring about the substrate flux is provided by coupling the transport of substrate to the flux of other solutes, termed activators. The fluxes of substrate and activator can be in the same direction (such transporters are referred to as symporters), or they can be in opposite directions, i.e. antiporters. Kinetic experiments similar to those undertaken with facilitated-diffusion 14

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

carriers can also be performed with co-transport systems, with the additional factor of determining the effect of substrate and activator on the flux of each other. Thus, for Na+ co-transport systems the substrate flux has been examined as a function of external and internal Na+ and substrate concentrations. Vesicles offer an advantage for these studies, as it is relatively easy to manipulate the internal and external concentrations of substrate and activator. For example, in renal brush-border vesicles under zero-trans influx conditions, increasing the concentrations of external Na+ increased the affinity of the Na+/carboxylic acid carrier for succinate without a change in Vmax: consistent with an ordered system in which Na+ binds to the transporter before succinate (19). A comparison of kinetic parameters for zero-trans influx and efflux and equilibrium exchange will also provide information on the directional symmetry of the carrier and the rate of conformational change of the free vs. loaded carrier. The determination of the activator/substrate stoichiometry is also of interest as it provides insight into the possible mechanism of the co-transport system and the concentrating capacity of the transporter. A number of independent methods have been used to determine the activator: substrate stoichiometry. A common approach is the activation method, which generally involves measuring substrate flux at a fixed concentration as a function of the activator concentration. A hyperbolic relationship suggests a 1:1 coupling ratio, whereas a sigmoidal dependence can be fitted to the Hill equation, with the Hill coefficient suggesting the minimum number of activator sites on the carrier. For the succinate carrier in renal brushborder vesicles, a Hill coefficient of 2 was derived from the sigmoid curves (20). A more direct method for determining the coupling ratio is to simultaneously measure activator-dependent substrate flux and substrate-dependent activator flux. Applying this method to the succinate carrier revealed a Na+/succinate coupling coefficient of 2-2.5 (20, 21). One practical limitation of this approach is that the substrate-dependent activator flux may be small relative to the total activator flux—the static head method has been introduced to circumvent this problem (21).

5.3 Data analysis The kinetic parameters (KM and Vmax) for the different modes of transport are routinely determined by computerized curve-fitting methods using a variety of commercially available programs. Two of the most commonly used programs are Enzfitter and Fig P (Elsevier Biosoft), which come with standard equations that define simple Michaelis-Menten kinetics. Additional equations can be entered by the investigator, and for many substrates uptake rates will be more precisely defined by a saturable component and a non-saturable component that is proportional to the substrate concentration and which is likely to represent simple diffusion. Although for many substrates simple diffusion will be a small percentage of the total flux (< 1%), for some substrates, particularly those that are lipophilic, simple diffusion can account for > 10% of the total flux, especially at high permeant concentrations. Particular attention also needs to 15

SIMON M. JARVIS

be paid to the possibility that there may be more than one transporter involved in the translocation of the substrate. The activation method for determining the activator/substrate stoichiometry will follow Hill-type kinetics and as such can be analysed by fitting the data to the Hill equation:

where [A] is the activator concentration, K is the concentration of A at which the flux is 50% of the maximum (Jmax), and n is activator/substrate stoichiometry. Computer programs are also used to fit inhibition data to dose-response curves to determine the concentration of inhibitor that inhibits by 50% (IC50).

6 Determination of the driving forces for symporters—ion gradients and membrane potential The driving force of many symport transporters is either the electrochemical gradient of an ion, e.g. Na+/substrate co-transporters, or the proton electrochemical gradient—which is composed of two components: the plasma membrane potential and the pH gradient over the plasma membrane. Both direct and indirect methods can be applied to determine whether electrochemical gradients drive the uptake of a substrate. 6.1 Ionic requirements These experiments investigate the effect of changing the external concentration of an ion on the uptake of the radiolabelled substrate. For potential Na + dependent transport systems this generally involves replacing sodium with Nmethyl-D-glucamine or choline in the transport buffer. If the uptake of substrate is dependent on a sodium electrochemical gradient, then substrate transport will be reduced in the absence of sodium (see Figure 2). Changes in the proton gradient can possibly be achieved by changing the external pH, but a corresponding change in the internal pH may also occur. Moreover, changes in the external pH may modify either the transporter or the substrate, resulting in a change in the rate of substrate flux that is independent of the change in the proton gradient. Membrane potential is an important driving force in the transport both of charged compounds and of neutral molecules, where transport occurs via an electrogenic process. In membrane vesicle studies the effect of membrane potential on the uptake of substrate can be investigated by modifying the membrane potential with anions of differing permeability. Thus, studies on Na+-dependent uridine uptake by rabbit renal brush-border vesicles demonstrated that as the permeability of the membrane to anions increased (NO-3 > Cl" > SO2-4), the initial rate and the magnitude of the transient overshoot also

16

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

increased (11), consistent with uridine transport proceeding via an electrogenic process. The membrane potential across vesicles can also be manipulated using valinomycin-induced K+ diffusion potentials. For example, it has been shown that membrane potential independently can provide the energy for uridine to accumulate against a concentration gradient in renal brush-border vesicles, provided that Na+ is present (11).

6.2 Use of ionophores and inhibitors An additional method for studying the ionic requirements of a transport process uses ionophores with varying selectivities. For example, monensin is a sodium ionophore, gramicidin is a Na + /K + exchanger, and carbonyl cyanide mchlorophenylhydrazone (CCCP) is a proton-gradient uncoupler. Unfortunately, these ionophores can have multiple effects, and in many cases not only is the ionic gradient disrupted, but in addition the membrane potential. Thus, caution is required when interpreting the results. Inhibitors can also be used to modify the potential driving force. Ouabain is an inhibitor of the Na + , K+-ATPase pump responsible for maintaining Na+ and K+ gradients across mammalian cells. Equilibrating membrane vesicles with Na+ before the start of the transport flux is another approach for eliminating the potential driving force.

6.3 Measurement of membrane potentials and ion gradients The above approaches, although valuable in providing important pointers to the possible driving forces for the uptake of substrate, require verification by methods that can measure the changes in intracellular ions or plasma membrane potential. Not only does this allow a correlation between transport rates and changes in, say, the proton motive force to be investigated, but it also allows the flux of the possible activator to be measured as a function of the substrate. The technique of choice for measuring membrane potential and the concentration of ions, that is applicable for use with both membrane vesicles and cells, involves fluorescent probes that are sensitive to the various parameters. Useful dyes include dipropylthiadicarbocyanine (diS-C3-(5)) and bisoxonol for the measurement of membrane potential, and the pH-sensitive dye 2',7'-bis(carboxyethyl)-5,6-carboxyrfluorescein (BCECF). For a detailed description and advice about the different dyes, the instrumentation required, calibration of signals, artefacts, and technical considerations the reader should consult the following refs 22 and 23 for reviews. In recent studies from this laboratory we have used fluorescent probes to demonstrate that the uptake of purines by trypanosomes is dependent on the proton motive force and is consistent with the presence of multiple H+symporters for the transport of purine nucleosides and nucleobases (10, 24). The basic procedure is to add dye and cells to incubation medium in a cuvette, allow time for the dye to equilibrate across the membrane, and then to record the 17

SIMON M, JARVIS

fluorescence level (see Protocol 5). The effects of" added substances, e.g. ionophores and permcant, can then be observed. In the case of BCIJCF, cells are preincubated with the acctoxymethyl ester of BCECF, and extracellular dye removed by washing before adding the labelled tells to the cuvette, Figure 5 shows an example of the type of experiment that can be performed to investigate whether proton influx is associated with the uptake of" substrate.

Protocol 5 Determination of the Intracellular pH in suspended cells Equipment and reagents • Fluorimeter, e.g. Perkin-Elmer LS 50B • 10 mM stock solution of 2',7'-bis(carboxyethyl)-5.6-carboxyfluarescein acetoxymethyl ester (BCECF/AM) in dimethylsulfoxide (DMSO), Store at -20°C for 4 weeks

Calibration medium: 20 mM KG, 10 mM NaCl, 2 mM MgCl2,10 mM glucose, 100 mM potassium gluconate, 5 mM Mes, and 5 mM Mops, pH 6.0 to 9.0 Assay buffer for trypanosomes: 33 mM Hepes, 98 mM NaCl. 4.6 mM KCl, 0.55 mM CaCl2. 0.07 mM MgCl2, 23 mM NaHCO3, and 14 mM glucose pH 7.3

Method

1, Wash cells twice in the appropriate assay buffer, resuspend to 106 to 10s cells/ml and incubate for 1 h at 25 °C with 5 utM BCECF/AM.a 2. Then wash the cells twice to remove the extracellular dye, resuspend in the same volume of assay buffer, and protect from light. 3. Transfer aliquots (3 ml) of the BCECF-loaded cells to Km2 quartz cuvettes and maintain at the required temperature. 4. Continuously monitor fluorescence at 440 and 490 nra and the 490/440 nm ratio (excitation 440 and 490 nm; emission 530 nm with 10-nm bandwidths). 5. Once a stable fluorescence trace has been established, add the test permeant or tonophore to the cells and monitor the change in fluorescence (see Figure 5). 6. Calibrate the change in 490/440 nm ratio with pHi using the H + /K + ionophore nigericin in a low sodium, high potassium calibration medium. Incubate cells (30 u1) in calibration medium containing 20 uM nigericin, add to cuvettes containing 3 ml of calibration medium of various pH values, then record the fluorescence intensity at 490 and 440 nm for 2 min. Plot the ratio of the fluorescence intensity at the two wavelengths against the pH to produce a standard curve. The standard curve was shown to be linear between pH 6.0 to 8.0 (ref. 24) for T. bntcei brucei bloodstream forms. 1

The dye will permeate the cells and then be cleaved to BCECF, an impermeable compound.

ASSAY OF MEMBRANE TRANSPORT IN CELLS AND MEMBRANE VESICLES

Rgure 5 Adenosine enhances the recovery of intracellular pH in base-loaded procyclic T. brucei brucei. BCECF-loaded cells (see Protocol 5) were base-loaded at the indicated time by adding NH4CI and, once a new stable level of pHi was established, adenosine (final concentration 25 n-M, dotted line) or an equal volume of assay buffer (solid line) was added at the indicated time. Addition of adenosine to the cells accelerated the rate of recovery in pH, compared to buffer alone, consistent with the co-transport of protons with adenosine.

References 1. Jarvis, S. M., Hammond, J. R., Paterson, A. R. P., and Clanachan, A. S. (1982). Biochem. J., 208, 83. 2. Plagemann, P. G. W. and Wohlhueter, R. M. (1980). Curr. Top. Membr. Res., 14, 225. 3. Oliver, J. M. and Paterson, A. R. P. (1971). Can. J. Biochem., 49, 262. 4. Plagemann, P. G. W. and Woffendin, C. (1989). Biochim. Siophys. Acta, 1010, 7. 5. Griffith, D. A. and Jarvis, S. M. (1996). Biochim. Biophys. Acta, 1286, 153. 6. Gazzola, G. C., Dall'Astra, V., Franchi-Gazzola, R., and White, M. F. (1981). Anal. Biochem., 115, 368. 7. Wohlhueter, R. M. and Plagemann, P. G. W. (1980). Int. Rev. Cytol, 64, 171. 8. Wohlhueter, R. M., Marz, R., Graff, J. C., and Plagemann, P. G. W. (1978). In Methods in cell biol. Vol. 20 (ed. D. M. Prescott), p. 211. Academic Press, New York. 9. Harley, E. R., Paterson, A. R. P., and Cass, C. E. (1982). Cancer Res., 42, 1289. 10. De Koning, H. P., Watson, C. J., and Jarvis, S. M. (1998). J. Biol Chem., 273, 9486.

19

SIMON M. JARVIS 11. Williams, T. C., Doherty, A. J., Griffith, D. A., and Jarvis, S. M. (1989). Biochem.J., 264, 223. 12. Mircheff, A. K., Hanna, S. D., Walling, M. W., and Wright, E. M. (1979). Prep. Biochem., 9, 133. 13. Le Hir, M. and Dubach, U. C. (1985). Eur. J. Clin. Invest., 15, 121. 14. Turner, R. J. (1983). J. Membr. Biol., 76, 1. 15. Lieb, W. R. and Stein, W. D. (1974). Biochim. Biophys. Acta, 373, 178. 16. Jarvis, S. M., Hammond, J. R., Paterson, A. R. P., and Clanachan, A. S. (1983). Biochem.J., 210, 457. 17. Jarvis, S. M. and Martin, B. W. (1986). Can. J. Physiol. Pharmacol, 64, 193. 18. Jarvis, S. M. (1986). Biochem. J., 233, 295. 19. Wright, E. M. (1985). Annu. Rev. Physiol, 47, 127. 20. Wright, S. H., Kippen, I., and Wright, E. M. (1982). J. Biol. Chem., 257, 1773. 21. Fukuhara, Y. and Turner, R. J. (1983). Am. J. Physiol., 245, F374. 22. Bashford, C. L. and Smith, J. C. (1979). In Methods in enzymology, Vol. 55 (ed. S. Fleischer and L. Packer), p. 569. Academic Press, London. 23. Negulescu, P. A. and Machen, T. E. (1990). In Methods in enzymology. Vol. 192 (ed. S. Fleischer and B. Fleischer), p. 38. Academic Press, London. 24. De Koning, H. P. and Jarvis, S. M. (1997). Mol. Biochem. Parasitol, 89, 245. 25. Jarvis, S. M. (1989). Biochim. Biophys. Acta, 979, 132. 26. Lee, C.-W., Cheeseman, C. L, and Jarvis, S. M. (1988). Biochim. Biophys. Acta, 942, 139.

20

Chapter 2 Reconstitution of membrane proteins: the Ca2+-ATPase of sarcoplasmic reticulum Anthony G. Lee Department of Biochemistry, School of Biological Sciences, University of Southampton, Medical and Biological Sciences Building, Basssett Crescent East, Southampton S09 3TU

1 Introduction Biological membranes have complex structures, reflecting the wide range of functions they are required to perform: most biological membranes contain many hundreds of different species of protein and lipid. One approach to understanding this complexity is to purify individual components from the membrane and study their properties in isolation. Purification of an intrinsic membrane protein generally requires it to be first dissolved out of the membrane in some suitable detergent, so that it can then be separated from the other membrane proteins. Some membrane proteins are stable for long periods in detergent solution, but many are not; even those that are stable may show altered properties in the unusual environment of a detergent micelle. For these reasons it is better to transfer the protein, once purified, from the detergent back into a lipid bilayer; this is the process of reconstitution. Reconstitution gives the simplest membrane system that is likely to be a fair representation of the native membrane. Reconstitution can be used to define the minimum number of protein components necessary to carry out some defined function of the membrane, and to study the interactions between lipids and proteins in the membrane. Successful reconstitution is dependent on having a suitable detergent—harsh enough to dissolve the protein of interest out of the membrane but not so harsh that the protein is damaged in any way. Even when a suitable detergent can be found, purification will be difficult if the protein is only present in small amounts in the source membrane; successful purification is very much easier if a membrane enriched in the protein of interest is available. Thus, for example, the Ca2+-ATPase can be readily purified from skeletal muscle sarcoplasmic reticulum, the acetylcholine receptor can be purified from the electric organ of electric fish, an anion transporter from red blood cells, bacteriorhodopsin from 21

ANTHONY G. LEE

Halobacterium halobium, photosynthetic reaction centres from a variety of plant and bacterial sources, and so on; all these proteins can be purified in 10-100 mg quantities because an enriched source is available. Purification also requires a suitable assay for the protein: this can be a problem. If the membrane protein is an enzyme, then the enzymatic function can be assayed using conventional assay techniques; for example, the Ca2+-ATPase can be assayed by measuring the rate of hydrolysis of added ATP. However, for membrane proteins that have no enzymatic activity, including many transporters and receptors, routine assay during purification can be difficult. Ligand binding assays may be possible; but if transport is the only possible assay, the protein will have to be reconstituted into a membrane system which allows vectorial transport to be measured, and this is inconvenient. This chapter will illustrate the approach of reconstitution using, as an example, the Ca2+-ATPase purified from skeletal muscle sarcoplasmic reticulum. This system has a number of advantages. The Ca2+-ATPase can be readily purified in 100 mg quantities. It consists of a single polypeptide chain, of molecular weight approximately 115 000, whereas other ATPases such as the (Na + , K+)ATPase contain two subunits: the catalytic a-subunit and a non-catalytic psubunit. Finally, as both an enzyme and a transporter, it is possible to study separately the scalar process of ATP hydrolysis and the vectorial process of Ca2+ transport.

2 Choice of detergent Detergents are amphiphilic molecules capable of disrupting the bilayer component of the membrane. An important property of a detergent is its critical micelle concentration, or CMC. Below its CMC, a detergent is soluble in water in monomeric form, but, at concentrations above the CMC, organized structures are formed, termed micelles. Micelles are typically spherical or disc-shaped, with a hydrophobic core and a polar surface. Proteins are solubilized in detergent micelles, with the hydrophobic, membrane-penetrant part of the protein coated by the hydrophobic part of the detergent. Often membrane proteins are solubilized with some remaining lipid molecules, and this can help to protect the protein from the harsh effects of the detergent environment. For example, the Ca2+-ATPase can be solubilized in active form in the detergent cholate, as long as a minimum number of 30 phospholipid molecules remain associated with each ATPase molecule; it is presumed that 30 phospholipid molecules are required to coat the transmembrane region of the ATPase (1). It can sometimes be helpful to add additional lipid to a solubilization system to help maintain lipid around the protein. There are no obvious rules to follow in the choice of a detergent to purify any particular membrane protein. Structures of some commonly used detergents are given in Figure 1 and some of their most important properties are listed in Table 1. Some of the detergents were originally developed for industrial use; these tend to be complex mixtures such as Triton X-100, which is a heterogeneous 22

RECONSTITUTION OF MEMBRANE PROTEINS

Figure 1 Structures of some commonly used detergents: (A), based on a sterol ring; (B), based on alkyl chains.

23

ANTHONY G. LEE

Table 1 Commonly used detergents and some of their properties Detergent

Monomer M,

Critical micelle concentration (M)

Aggregation number

Octyl-B-D-glucoside (octyl-B-Dglucopyranoside)

292

2.5 x 10~2

84

Dodecyl-p-D-maltoside

511

1.7 x

Digitonin

1229

7.0 X 10-4

Sodium cholate

431

1.0 x 10-2

4

Sodium deoxycholate

415

3.0 x 10-3

22

CHAPS

615

5.0 x 10-3

9

Sodium dodecyl sulfate (SDS)

288

3

8.0 x 10-

62

Dodecyldimethylamine oxide (or LDAO)

229

2-8 x 10-3

ca 80

C12E8

538

8.7 x 10-5

120

C12 & 14E9.5 (Lubrol PX)

620

—

100

C12E23 (Brij 35)

1200

9.0 x 10-5

40

Triton X-100

1625

3.0 X

10~4

98 60

Polyoxyethylene glycol detergents

Tween 20

1240

10-4 5

6.0 x 10-

140 —

mixture of long-chain polyoxyethylene polymers. Other detergents have been synthesized specifically for biological use; these are chemically well denned, such as octyl-p-D-glucoside (octyl-B-D-glucopyranoside) and dodecyl-B-D-maltoside, and a range of chemically defined polyoxyethylene glycols, including the commonly used C12E8. Of the non-ionic detergents, Triton X-100 is often used as a non-denaturing, mild detergent. It is prone to contamination with peroxides and shows a strong absorption at 280 nm, but its major problem is that it is difficult to remove by dialysis because of its low CMC. It can, however, be removed using 'Bio-Beads' (2) or using Extracti-Gel D, an affinity matrix marketed by Pierce for the removal of detergents. Another disadvantage of Triton X-100 is that it is not possible to use salting-out with ammonium sulfate to purify proteins in the presence of Triton X-100, since this causes separation of the solution into a detergent-rich upper phase and a detergent-poor lower phase; the protein preferentially partitions into the detergent-rich phase where it denatures. Octylp-D-glucoside, another non-ionic detergent, has the advantage of a high CMC, making it easy to remove; dodecyl-B-D-maltoside has rather similar properties, but its longer alkyl chain makes it more difficult to remove. The bile salts cholate and deoxycholate have the advantages of cheapness, and their high CMCs mean they can be removed by dialysis. The pKa values of the acidic groups (5.2 for cholate and 6.2 for deoxycholate) are such that they can only be used at pH values of 8 or greater; at neutral or acidic pH values, the uncharged form of the detergent precipitates from solution. They are compatible 24

RECONSTITUTION OF MEMBRANE PROTEINS

with ammonium sulfate fractional ion, but cannot be used in ion-exchange column chromatography. The zwitterionic detergent 3-| (3-cholamidopropyl)dimcthylammonio|-l-propane sulfonate (CHAPS) contains a steroid ring system similar to that in cholate or deoxycholatc, but because it has no net charge it is compatible with ion-exchange purification.

3 Purification of the Ca2+ -ATPase from skeletal muscle sarcoplasmic reticulum Purification of the Ca2-ATPase from skeletal muscle sarcoplasmic reticulum illustrates the use of the detergent cholate and its removal by centrifugatkm into a detergent-free sucrose gradient, followed by dialysis. The first step of the procedure is the isolation of sarcoplasmic reticulum (SR) from muscle, as described in Protocol 1. A number of procedures have been published for the purification of SR; that given in Protocol I is based on the work of Daiho a til. (3). It is important to maintain a low temperature during the preparation, and the homogenizer blades should be sharp. The final sample consists of sealed vesicles in which the Ca2 -ATPase makes up about 70% of the total protein. The activity of the Ca2 -ATPase can be measured using the coupled enzyme assay described in Section 4. In the absence of a Ca2 ionophore, the steady-state rate of ATP hydrolysis is low, because rapid accumulation of Ca2 within the vesicles leads to inhibition of the ATPase. However, the presence of a Ca2 ionophore such as A23187 prevents accumulation of C a 2 , and allows the full ATPase activity to be measured.

Protocol 1 Preparation of sarcoplasmic reticulum3 Equipment and reagents • Low-speed centrifuge capable of handling up to 1 litre of material at up to 8300 g • Ultracentrifuge with afixed-anglerotor (e.g. 6 x 100 ml capacity) • Heavy-duty blender (e.g. a Waring blender) • A piece of dressmaker's muslin • Buffer A: 0.1 MNaCl. 10mMMops/TrispH 7.0 (2 litres)

• Buffer B: 0.6 M KC1. 5 mM Tris brought to pH 6.5 with solid maleic acid (1 litre) • Buffer C: 0.1 M sucrose, 30 mM Mops/Tris pH 7.0 (200 ml) • Hand-operated glass homogenizer, with a tight fitting (0.3 mm clearance) pestle • 50-100 ral tubes • 1%SDS

Method 1. Chop the white muscle (500 g) from the legs and back of a 3-kg New Zealand white rabbit into small pieces using a scalpel blade, taking care to remove the bulk of any fat and connective tissue.

ANTHONY G. LEE

Protocol 1

2. Blend 250 g of the muscle with 1 litre of Buffer A in a Waring blenden To ensure efficient homogenization, operate the blender for 20 sec, and then allow the preparation to settle for 5 sec before repeating the process a total of four times. Repeat for the second 250 g of muscle, and combine the homogenates, 3. Centrifuge the homogenate in a fixed-angle rotor at 8300 g for 35 min. Be careful not to disrupt the pellet, which is very soft. 4. Filter the supernatant through a piece of muslin into a 1 litre beaker. The muslin will clog very quickly, so keep moving the supernatant around within the piece of muslin. 5. Remove the mitochondrial fraction by centrifuging the filtered supernatant at 12 000 g for 30 min in a fixed-angle rotor (two centrifuge runs will be required if an 8 x 50 ml rotor is used). 6. Pool and centrifuge the resulting supernatants at 53000 g for 40 min, using 6-8 tubes of appropriate size (50-100 ml). 7. Resuspertd the pellets in a total volume of 40 ml of Buffer B/g of pellet as follows, Discard the supernatant from each tube. Estimate the weight of pellet by weighing one tube with its pellet, then add a few ml of buffer, scrape the pellet out of the tube into an empty tube, and re-weigh the tube. Multiply the weight of the pellet by the number of tubes, and then calculate the total volume of Buffer B needed. The total weight of pellet will probably be about 10 g, giving a required volume of Buffer B of about 400 ml. Using the minimum amount of Buffer B, scrape all the pellets into one tube, then add the contents of the tube to a 1 litre beaker and add the rest of the Buffer B. Gently stir the resuspension for about 40 mm. 8. Centrifuge the sample at 125 000 g in a fixed-angle rotor for 45 rain. 9. To wash the sample, discard the supernatant and resuspend the pellets in the minimum volume (about 2 ml) of Buffer C, and then tip the suspension into a 3 ml glass homogenizer. Homogenize with a few passes of the homogenizer, and then centrifuge at 125 000 g for 45 min in a fixed-angle rotor. 10. Discard the supernatant, resuspend in the minimum volume of Buffer C and either proceed to the preparation of the Ca2+-ATPase (see Protocol 2) or aliquot in suitable amounts into Eppendorf tubes, snap-freeze in liquid nitrogen, and store at -20°C. Freeze the sample at about 40 mg protein/ml. The preparation should give about 400 mg protein. 11. Measurethe protein concentration of the SR sample by adding 10 ul of the SR suspension to 800 ul of 1% SDS and 190 ul of water, Measure the difference between the optical densities at 280 and 330 nm (the optical density at 330 rm gives a measure of light scatter in the turbid suspensions). The protein concentration is given by the relationship: protein concentration (mg/ml) = OD280 - OD330. a Modified from ref. 3. b Carry out all operations in a cold-room, with ice-cold buffers, and perform all centrifugarions at 4°C. 26

RECONSTiTUTION OF MEMBRANE PROTEINS

The ATPase is purified from SR by solubilization in cholate, followed by separation of proteins on a detergent-free sucrose gradient, as described in Protocol 2, There are a number of important features in the protocol. The CMC of cholate is very dependent on pH and ionic strength and varying these will alter the ability of the detergent to solubilize the ATPase (see ref. 4). The ratio of cholate/protein is important; a ratio of 0.4 mg cholate/mg SR protein ensures good solubilization of the ATPase and gives a final preparation with a molar ratio of lipid:ATPase of about 30:1, the minimum required to maintain ATPase activity (1), Centrifugation into the sucrose gradient achieves two things: it allows purification of the ATPase and it removes most of the cholate, as illustrated in Figure 2. The cholate moves part way into the gradient, forming a detergent front; some of the phospholipid remains at the top of the gradient with the detergent, the rest moves down the gradient with the protein. The purified ATPase is located at the 30-60% interface. It is largely cholate-free and consists of membrane fragments; since it is present as membrane fragments, it is unable to accumulate Ca2+, but can, of course, hydrolyse ATP. The measured ATPase activity should be about 3-4 IU/mg under the conditions of Protocol 3 and should not change on addition of the Ca2+ ionophore A23187, It gives a single band on SDS-PAGE at an Mr of about 100 000 (5). The maximal level of phosphorylation of the ATPase by [r32P]ATP is about 3-5 nmoles [EP]/mg protein,

Figure 2 Purification of Ca2+-ATPase on a detergent-free sucrose gradient. On centrifugal ion, cholate moves part-way into the gradient, forming a detergent front. Some lipid remains at the top of the gradient, with the detergent, and some moves through the detergent gradient with the protein, which forms membrane fragments in the more dense regions of the gradient. 27

ANTHONY G. LEE

Protocol 2 Purification of the Ca2+-ATPase from sarcoplasmic reticulum Equipment and reagents • Potassium cholate • KOH • Choiic acid • Diethyl ether • Vacuum dryer • Sucrose • Ultracentrifiige with a swing-out rotor (14-17 ml tube volume)

• Amberlite XAD-2 ion-exchange resin (12 g), washed four times with deionized water • Buffer C: 0.1 M sucrose, 30 mM Mops/Tris pH 7.0 (200 ml) • Buffer D: 50 mM K2HPO4 pH 8.0,1 M KC1 (500 ml) • Buffer E: 0.25 M sucrose, 50 mM K2HPO4 pH 8,0, 1 M KC1 (2 litres)

Method" 1. Purify potassium cholate by dissolving equimolar amounts of KOH and cholic acid in a small volume of methanol. Precipitate the potassium cholate by adding excess diethyl ether, collect the precipitate on a filter paper and dry under vacuum overnight until no smell of ether remains, 2. Prepare 2-3 ml of a solution of potassium cholate in Buffer D, at 100 mg/ml. 3. Resuspend the SR at about 30 mg protein/ml in Buffer D—up to 150 mg of protein can be purified per sucrose gradient, 4. Prepare six discontinuous sucrose gradients of 3 ml of 60%, 19 ml of 30%, and 13,5 ml of 20% sucrose (expressed as g sucrose per 100 ml final volume of solution), 5. Solubilize the SR by slowly adding the 100 mg/ml cholate solution, swirling constantly to give a final ratio of 0.4 mg cholate/mg SR protein. 6. Layer the solubilized SR on to the sucrose gradients at about 100 mg protein per gradient (the upper limit is about 150 mg protein per gradient). Centrifuge at 95 000 g overnight. 7. Collect the thick band of purified ATPase from the 30-60% interface, dilute with Buffer D, and centrifuge at 37 000 g for 1 h. Resuspend the pellet in a small volume (1-2 ml) of buffer E and dialyse overnight against 1 litre of buffer E containing 10 g of Amberlite XAD-2 ion-exchange resin to remove any remaining cholate. Aliquot the ATPase into Eppendorf tubes and snap-freeze in liquid nitrogen. Store at -20°C a

Carry out all operations except step 1 in a cold-room, with ice-cold buffers, and perform all centrifugations at 4°C.

RECONSTITUTION OF MEMBRANE PROTEINS

implying that only between 30 and 50% of the ATPase is active; similar levels of phosphorylated protein are obtained with SR vesicles. The reason for the high levels of inactive protein in SR vesicles, and in the Ca2+-ATPasepurified from it, is unclear (see refs 6 and 7), but the inactive ATPase cannot be totally denatured since labelling ratios obtained with a number of fluorescence probes such as fluorescein isothio cyanate, 4-(bromomcthyl)-6,7-dimethoxycoumarin, and 0phthalaldehyde are 1:1 with respect to totcil protein (8).

4 ATPase assay The ATPase activity of the Ca2+ -ATPase can be measured using the coupled enzyme assay described in Protocol 3 in which the formation of ADP, generated by hydrolysis of ATP by the Ca2- -ATPase, is linked to the formation of pyruvate from phosphocnolpyruvate, catalysed by pymvate kinase. The pyruvate is then reduced to lactate by lactate dehydrogenase, with conversion of NADH to NAD' which is monitored by the fall in absorbance at 340 nm. To measure the ATPase activity of the ATPase in SR vesicles it is necessary to add the Ca2+ ionophon? A23187 to make The SR membrane leaky to Ca2+ and to prevent a build-up of Ca2 within the lumen of the vesicles; the addition of ionophore is unnecessary when assaying the purified ATPase, since this is present as unsealed, membrane fragments.

Protocol 3 Assay of ATPase activity Equipment and reagents • EGTA

• • • • »

l M and 0.2 M KOH 10 mmol suspension of calcium carbonate 1MHC1 ATP (disodium salt trihydrate) Phosphoenolpyruvate (tricyclohexylaramonium salt)

• NADH • Hepes buffer: 40 mM Hepes/KOH pH 7.2

100 mM MgSO4 in Hepes buffer A spectrophotometer capable of operating at 340 ran, with a temperature control system for measurements at 25 oC Pig heart lactate dehydrogenase Rabbit muscle pyruvate kinase 1 mg/ml of the calcium ionophore A23186 in methanol

Method 1. Make a 100 mM solution of EGTA by suspending 10 ramol of EGTA in about 50 ml of water and adding 1 M KOH to get it to dissolve. Adjust the pH to 7.4 and bring the volume to 100 ml, 2. Make a 100 mM solution of Ca2+ by suspending 10 mmol of calcium carbonate in 50 ml of water and adding 1 M HC1 to get it to dissolve. Adjust the pH to about 7 with 0.2 M KOH and bring the volume to 100 ml 29

ANTHONY G. LEE Protocol 3 con

3. Prepare the assay solution by dissolving 120 mg ATP (disodium salt trihydrate), 23 mg phosphoenolpyruvate (tricyclohexylammonium salt), and 11 mg NADH in 19 ml of Hepes buffer; it will probably be necessary to readjust the pH back to 7.2 after adding the ATP. Then add 5 ml of the 100 mM MgSO4 solution, 1 ml of the 100 mM EGTA solution, 200 U of pyruvate kinase, and 700 U of lactate dehydrogenase. 4. Add SR or purified ATPase (10 ug protein) to 1.73 ml of Hepes buffer plus 0.63 ml of the assay solution, at 25°C in a cuvette. Monitor the absorbance at 340 nm as a function of time. This should hardly change following addition of the protein, indicating the absence of any Ca2+-independent ATPase activity. Start the reaction by adding 24 ul of the 100 mM Ca2+ solution to give a free calcium concentration of about 10 uM, and again monitor the absorbance at 340 nm. Measure the rate of change of absorbance at 340 nm. For SR vesicles this should be low: add 5 ul of the A23187 solution to make the SR vesicles permeable to Ca.2+, and the rate of change of absorbance at 340 nm should increase markedly. Add more Ca2+ and measure the rate again to check that the system contains a maximally stimulating concentration of Ca2T 5. Calculate the ATPase activity from the fall in NADH absorbance at 340 nm, using an extinction coefficient of 6200 1 mol-1 cm-1 for NADH at 340 nm. Typical activities are 3-4 lU/mg protein for the purified ATPase or for SR vesicles in the presence of ionophore. and 0.5-1 RJ/mg protein for SR vesicles in the absence of ionophore.

5 Reconstitution of the Ca2+-ATPase 5.1 Reconstitution into membrane fragments of defined phospholipid composition Protocol 2 gives a preparation of purified ATPase, but one in which the ATPase is still surrounded by the complex mixture of phospholipids characteristic of the native SR membrane. To prepare a fully defined membrane system it is therefore still necessary to replace this complex mixture of SR phospholipids by a single, chosen, chemically defined species of lipid: this is the process of lipid substitution. This can be achieved by mixing the purified ATPase with an excess of the chosen phospholipid in cholate, leaving the mixture for about an hour for the phospholipids to equilibrate (see Kgure 3). The lip id-substituted ATPase can then be separated from excess lipid by sucrose-gradient centrifugation, in a procedure analogous to that used in Protocol 2 to purify the ATPase (9). However, a much simpler procedure is simply to dilute the lipid-A'ITase mixture 500-fold into buffer so that the concentration of cholate drops below its CMC. Membrane fragments then re-form in which the ATPase is surrounded predominantly by the chosen phospholipid; effects of phospholipid structure on the activity of the ATPase determined in this way are identical to those determined using the centrifugaiion procedure (1). Protocol 4 describes the method. 30

RECONSTITUTION OF MEMBRANE PROTEINS

Protocol 4

Reconstltution of the Ca2+-ATPase into membrane fragments Equipment and reagents • Pure phospholipids (e.g. Sigma, Avanti Polar Lipids, etc.), made up as a stock of 20 mg/ml in chloroform or chloroform/ methanol and stored at -20"C • A small, high-power sonicating bath (e.g. Ultrawave, model U100) • Potassium cholate purified as described in Protocol 2 • Reconstitution buffer: 15% sucrose, 5 mM MgSO«, 5 mM ATP, 10 mM Hepes/Tris pH 8.0, containing 26 mg potassium cholate (2.0 ml)

• Buffer E: 10 mMHepes/TrispH 8.0 • Dilution buffer: 250 mM sucrose, 10 mM Hepes/KOH pH 8.0 • Thin-walled, flat-bottomed glass vials (see text below for how to select good vials) • Vacuum desiccator • Nitrogen source » ParafiJm • Whirli-mixer (Fisher) • Assay buffer (see Protocol 3)

Method 1. AJiquot the appropriate amount of lipid into a glass vial. For each 1.25 mg of ATPase to he reconstituted, take 10 umol of lipid. Blow off the solvent under a stream of nitrogen so that the lipid forms a thin film on the side of the vial. Place in a vacuum desiccator and dry for at least 1 h. 2. To the vial add 400 ul of the reconstitution buffer, flush the vial with nitrogen, and seal with Parafilm. Gently whirli-mix the vial to dislodge the lipid from the side of the vial (it may be necessary to warm the vial slightly). Place the vial in the sonication bath and locate it in the bath so that the buffer 'fountains' vigorously within the vial. Continue sonicating until the sample is almost optically clear; this will probably take 10-15 min. 3. To the vial add 1.25 mg purified ATPase in a volume of 3-10 u1. Gently swirl the vial to ensure good mixing, and leave for 15 min at room temperature and then for 45, min at 4 oC.4 4. Add 2 ml of the dilution buffer to the vial, and keep on ice for up to about 2 h until use. 5. For the ATPase assay, take 30 ul of the samples and dilute into 3 ml of assay buffer and assay as described in Protocol 3. Alternatively, the ATPase sample can be diluted into a larger volume of buffer E and the sample centrifuged at 200 000 g for 1 h at 4°C, Resuspend the pellet in the chosen buffer at about 40 mg protein/ml and snapfreeze in liquid nitrogen for later use. aThe lipid must be above its gel-to-liquid crystalline phase-transition temperature for reconstitution. Thus, for example, if reconstituting with dipalmitoylphosphatidylcholine the sample should be incubated at 42°C for 15 min, followed by 45 min at 5°C (10),

ANTHONY G. LEE

Figure 3 The process of lipid substitution. Following equilibration of the ATPase with excess exogenous lipid in detergent, the sample can be diluted into buffer to form membrane fragments in which the ATPase is surrounded predominantly by the exogenous lipid.

Successful reconstitution relies on careful dispersal of the lipid into the cholate micelles. This is best achieved using a sonication bath; the alternative of using a sonication probe is more likely to lead to oxidative damage of the lipids and can produce unwanted metal fragments (from the probe tip) in the sample. The sonication bath should be sufficiently powerful so that 0.5-1 ml of buffer contained in a small glass vial (such as a 10 ml scintillation vial) placed in the bath sprays up on to the walls of the vial. A good 'fountain' effect requires that the glass vial should be thin-walled and, preferably, flat-bottomed, to ensure good propagation of the ultrasonic waves into the sample. It will probably be necessary to test a large number of vials to find a few that are good for the purpose. The required cholate concentration is worked out on the basis of 1 mg of cholate/mg of protein plus 0.5 mg of cholate/mg of exogenous lipid. A good test of the reconstitution procedure is to compare ATPase activities for the ATPase reconstituted into dioleoylphosphatidylcholine (di(C18:l)PC) and into dimyristoleoylphosphatidylcholine (di(C14:l)PC). ATPase activities measured under the conditions of Protocol 3 should be about 3-4 lU/mg protein for the ATPase reconstituted in di(C18:l)PC and about 0.5-1 IU/mg protein for the ATPase reconstituted in di(C14:l)PC (11). Because this procedure gives membrane fragments, the full ATPase activity is measured, uncomplicated by any processes of Ca2+ accumulation, and it is unnecessary to add any Ca2+ ionophore.

32

RECONSTITUTION OF MEMBRANE PROTEINS

5.2 Reconstitution into sealed vesicles of defined phospholipid composition The ATPase can also be reconstituted into sealed vesicles allowing the accumulation of Ca2+ to be measured. Accumulation of Ca2+ is a more complex process than ATP hydrolysis since the level of Ca2+ accumulated depends not only on the rate of hydrolysis of ATP, but also on the size of the vesicles, on the rate of leak of Ca2+, and on the movement of counterions such as H+ or K+, necessary to prevent the build-up of a pH gradient or membrane potential. When dry phospholipids are shaken with water they form multilamellar lipid vesicles (MLVs), or liposomes. These are obviously unsuitable for reconstitution studies since their multilayered structure restricts the access of substrates added to the external medium to only the outer bilayer in the liposome. Small unilamellar vesicles (SUVs) can be made from MLVs by sonication, but SUVs are rather unstable, with a tendency to fuse because of strain in the highly curved bilayer surface. However, it is possible to prepare more stable large unilamellar vesicles (LUV) by a variety of procedures, of which the most convenient is probably that of forcing MLVs under pressure through polycarbonate niters of defined pore size (12). Membrane proteins can be directly incorporated into SUVs in the absence of detergent, but this generally requires the presence of anionic phospholipids and is of variable efficiency; the small internal volume of the SUV is also a disadvantage (13, 14). A better procedure is detergent-mediated incorporation of membrane proteins into pre-formed LUVs. This process has been studied in detail by Rigaud et al. (13), and is described below (see Section 5.2.1). It gives vesicles with very low ionic permeabilities, which show a high level of Ca2+ accumulation. Unfortunately, however, it is not applicable to all lipids or lipid mixtures. Some, such as the phosphatidylethanolamines, do not readily form LUVs. In other cases, such as LUVs containing cholesterol, we have found that detergent-mediated incorporation of the ATPase is inefficient. In these cases, alternative reconstitution procedures are necessary. The most straightforward is a direct extension of the lipid substitution procedure described in Protocol 4 but with the slow removal of detergent, allowing the system time to form sealed vesicles. Detergent can be removed slowly either by dialysis (9) or, more conveniently, by passage through a column of Sephadex G-50 (15); this method is described below in Section 5.2.2. ATPase activities of the sealed vesicles can be assayed using Protocol 3. These measurements also give the percentage sidedness of the ATPase molecules in the vesicles; in sealed vesicles, only those ATPase molecules with their ATP binding sites exposed to the outside of the vesicles will be able to hydrolyse ATP (see Figure 4). The percentage sidedness is calculated from the ratio of the ATPase activity for the sealed vesicles to the activity of the vesicles after the addition of a small amount (0.8 mg/ml) of the detergent C12Eg to make the vesicles leaky to ATP. When using this assay of sidedness it is important to check that C12E8 has no direct effect on the activity of the ATPase; this can be 33

ANTHONY G. LEE

Uptake

Figure 4 The Ca2+-ATPase reconstituted into sealed lipid vesicles. Some ATPase molecules will have an inverted orientation in the vesicles, and so will be unable to react with ATP in the external medium unless the vesicles are made leaky, for example by adding a detergent such as C12E8. Accumulation of Ca2+ by the vesicles is a balance between the rate of uptake and the rate of Ca2+ leakage out of the vesicles. The external radius of the vesicle, r0, can be measured by light scatter. This is related to the internal radius, ri, and the thickness of the bilayer, 8, by r0 = r-( + 8; the internal volume of the vesicle can be calculated from r,.

done using Protocol 4 to reconstitute the ATPase into membrane fragments, and testing for any (direct) effects of C12E8. It is important to determine the size of the reconstituted vesicles since the level of Ca2+ accumulation depends markedly on the internal volume of the vesicles. The most convenient procedure is to use a particle sizer such as the Coulter N4 Plus that measures particle size by light scattering; this technique has the advantages of being rapid and also of giving a measure of the size distribution of the vesicle population. Alternatively, vesicles can be sized by gelpermeation chromatography, on a column of Sepharose 2B calibrated using polystyrene beads of known diameter (Polysciences) (15). The average size can also be determined by measuring the trapping capacity of a water-soluble but impermeable reagent such as [3H]inulin. A variety of procedures are available for measuring Ca2+ accumulation, including the use of 45Ca2+ and spectrophotometric or fluorescence assays using suitable Ca2+-sensitive dyes, as described in Protocol 7. 5.2.1 Reconstitution into pre-formed large unilamellar vesicles (LUV) The procedure described in Protocol 5 is a minor modification of that of Levy et al. (16). The level of Ca2+ accumulated by the ATPase depends markedly on the lipid composition of the vesicles. Relatively low levels of accumulation are 34

RECONSTITUTION OF MEMBRANE PROTEINS

found in vesicles containing di(C18:l)PC as the only lipid, but the addition of 10 mol% dioleoylphosphatidic acid (di(C18:l)PA) or dioleoylphosphatidylserine leads to a marked increase in the accumulation of Ca2+ (see Figure 5); inclusion of more than 10 mol% of an anionic lipid leads to no further increase in the level of Ca2+ accumulation (17). Transport of Ca2+ into sealed vesicles is accompanied by the transport of H+ out of the vesicles, so that increased levels of Ca2+ accumulation are seen in the presence of a proton ionophore such as carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP) to equilibrate H+ across the membrane, as illustrated in Figure 5 (18). Figure 5 also illustrates another important point. The action of the ATPase both depletes the ATP in the sample

Figure 5 Accumulation of Ca2+ by the Ca2+-ATPase reconstituted into LUV. (A) Accumulation of Ca2+ by sealed vesicles composed of the ATPase reconstituted into 100% di(C18:l)PC (c, d) or 90% di(C18:l)PC/10% di(C18:l)PA (a, b) at a lipid:protein ratio of 40:1 (w/w). Ca2+ uptake was initiated by the addition of 0.8 mM ATP. All samples contained 0.25 uM FCCP. In (a) and (c) an ATP-regenerating system was present, but in (b) and (d) this was absent. (B) Accumulation of Ca2+ by sealed vesicles composed of the ATPase reconstituted into 90% di(C18:l)PC/10% di(C18:l)PA, in the absence of an ATP-regenerating system, in (a) the presence or (b) the absence of 0.25 n-M FCCP. In trace (b), 0.25 uM FCCP was added at the time marked by the arrow. In all the experiments shown, the protein concentration was 0.04 mg/ml and the buffer was 10 mM Pipes pH 7.1, 100 mM K2S04, 5 mM MgS04.

35

ANTHONY G. LEE

and generates ADP. Obviously, the accumulation of Ca2- will cease when all the added ATP has been used up, and this will occur after about 15 mm under the conditions used in the experiment illustrated in Figure 5. However, the build-up of ADP also has important consequences, since this leads to increased slippage on the ATPase, a process in which ATP is hydrolysed by the ATPase without transport across the membrane (17), Both these problems can be overcome by adding an ATP-regenerating system to the sample; this can take the form of a mixrure of pyiiivate kinase (200 U) and phosphoenolpyrtivate (0.4 mM), as used in the ATPase assay described in Protocol .3. Large unilamellar vesicles of the required lipid composition are made by forcing MLVs through Nudeopore polycarbonate membranes of 100 nm pore size, SR vesicles are dissolved in the detergent C12E8 and then added to the LUVs in the presence of a low concentration of octyl-B-D-glucoside. Detergent is then removed by the successive addition of SM-2 Bio-Beads to give a preparation of scaled vesicles containing the ATPase. Ca2 accumulation by the vesicles can be

Protocol 5 Reconstitution into pre-formed large unilamellar vesicles (LUVs) Equipment and reagents • SR (see Protocol 1) • LiposoFast extrusion system (Avestin Inc.) • A small, high-power sonicating bath (e.g. Ultrawave, model U100) • 100-nm pore size Nucleopore polycarbonate membranes (e.g. Costar) • Stock solutions of dioieoylphosphatidylchoIine(di(C18;l)PC) and dioleoylphospnatidic acid (di(C18:l)PA) in chloroform/methanol at 20 mg/ml • 20-50 mesh SM-2 Bio-Beads (Bio-Rad, cat. no. 152-3920)

• Buffer A: 10 mM Pipes pH 7,1, 100 mM K2SO4 • Buffer B: BufferAcontainingO.l mM CaCl2 • A stock solution of 1 M octyl-p-o-glucoside in water • Small column • 10 ml glass vials (e.g. scintillation vials) • Nitrogen source • Vacuum desiccator • Parafitm • C12E8

Methoda 1. Thoroughly wash 5 g of SM-2 Bio-Beads. Add the Bio-Beads to 50 ml of methanol and stir for 15 min. Tip the beads into a Buchner funnel and. wash with 200 ml methanol, and then with 200 ml Buffer A. Tip into a small (ca 15 ml) glass column and slowly wash through with 400 ml Buffer A, over a period of 2-3 h. Pour from the column into a beaker and add a small volume of Buffer A so that the beads are just covered. The washed beads can be kept at 4°C for several weeks. 36

RECONSTITUTION OF MEMBRANE PROTEINS Protocol 5 continued

2. Before use, place the sample of Bio-Beads in a vacuum desiccator for about 30 min to thoroughly degas the beads and buffer. Drain off the buffer, and add back just a little Buffer A so that the beads remain wet for weighing. Weigh out four lots of 320 mg, and then add a little buffer to each sample to prevent them drying out, 3. Aliquot 20.2 mg di(C18:l)PC and 2.2 mg di(C18:l)PA into a 10 ml glass vial. Remove the solvent under a stream of nitrogen so that the lipid forms a thin film on the side of the vial. Place in a vacuum desiccator and dry down under vacuum for about 3h. 4. Add 1.4 ml Buffer A to the vial, flush with nitrogen, and seal with Parafilm. Warm in warm water, and then vortex mix for 10-20 sec to remove the lipid from the walls of the vial. Place the vial in a sortication bath, and sonicate for about 5 min. At this stage the sample of MLVs should have a creamy white appearance. 5. Transfer the lipid sample into an Eppendorf tube. A freeze-thaw procedure is used to reduce the size of the MLVs: freeze the tube in liquid nitrogen and then warm the tube in warm water. Repeat the process 10 times. 6. Load the sample into the Avestin extruder and pass 11 times through two 100-nm pore polycarbonate filters, following the manufacturer's instructions. This produces LUVs. Typically 1 ml of sample will be recovered. Dilute the sample by adding 3 volumes of Buffer A. 7. Add, dropwise, the stock solution of octyl-B-D-glucoside to the lipid sample, to give a final concentration of 40 mM octyl-B-D-glucoside; the lipid suspension will become clear. 8. Solubilize the SR with C12E8.Add 1 mg of SR protein to 0.5 ml Buffer B containing 3 mg C12Eg. 9. Reconstitute at a weight ratio of 1:40 protein:lipid. Thus add 0.2 mgof the solubilized SR to 2 ml of the liposome preparation in a glass vial. Gently swirl the sample for about 1 min to ensure mixing and then add the first aliquot of Bio-Beads. Flush the vial with nitrogen, seal with Parafilm, and leave for 1 h, swirling occasionally. Add more Bio-Beads, and repeat the above procedure twice more (four times in all). After the fourth addition of Bio-Beads allow the beads to settle for 5 min and then remove the cloudy proteoliposome suspension, leaving behind the beads. This is done most conveniently using a narrow Gilson pipette tip. The preparation can be kept on ice for several hours until use. ° Based on the method of Levy et al (16(.

measured spectrophotometrically as described in Protocol 7. The ATPase activity of the vesicles, the percentage orientation of the ATPase in the membrane, and the size of the vesicles can be determined as described in Section 5.2 above. Typically, the procedure gives vesicles with a totally random distribution of ATPase molecules, with vesicle diameters of about 80-100 nm. 37

ANTHONY G. LEE

5.2.2 Reconstitution into sealed vesicles using cholate and deoxycholate The procedure described in Section 5.2.1 gives a preparation of large, tightly sealed vesicles showing a very low ionic permeability (13). However, it cannot be applied to the reconstitution of all lipid mixtures. The alternative procedure described in Protocol 6 can be used with a wider variety of lipids but gives somewhat smaller vesicles (typical diameters of about 70 nm). Unfortunately, the vesicles are leaky to Ca2 so that it is necessary to include a high concentration of phosphate in the lumen of the vesicles, to complex the calcium and reduce the rnte of leak (15). The highest levels of accumulation of Cn2 are achieved with mixtures of phosphatidylethanolamines and phosphatidylcholines (15). The procedure does not work well with mixtures containing anionic phospholipids. Lipid is dissolved in cholate and the purified ATPase is dissolved in deoxycholate. The two are mixed and then the detergents are removed by centrifugation through a Sephadex G-50 column, based on the procedure of Penefsky (19) for removing small, water-soluble molecules. The exact time required to pass the sample through the Sephadex column will vary with the type of centrifuge used but can be easily determined by trial and error, applying a mixture of potassium dichromatc and blue Dextran to the column, and monitoring their passage through the column at varying times and speeds. Calcium accumulation by the vesicles can be assayed using Protocol 7.

Protocol 6

Reconstitution into sealed vesicles using cholate and deoxycholate Equipment and reagents • • • • • •

10 ml glass vials (e.g. scintillation vials) Nitrogen source Vacuum desiccator Purified ATPase (see Protocol 2} A slow-speed benchtop centrifuge A small, high-power sonicating bath (e.g. Ultrawave, model U100) • Two 5-ml disposable plastic syringes that will fit into plastic centrifuge tubes in the low-speed benchtop centrifuge • 20 mg/ml stock solutions of dioleoylphosphatidykholine and dioleoylphosphatidylethanolaimne (di(ClS:l)PE) in chloroform/methanol

38

• Glass wool • Parafilm • Phosphate buffer: 0.4 M phosphate pH 7.4 (200 ml) • 10% (w/v) potassium cholate in 40 mM Hepes/KOH pH 7.2.100 mM KC1. Purify the cholate as in Protocol 2, • 10%w/v)potassiumdeoxycholate in 40 mM Hepes/KOH, pH 8.0. Purify the deoxycholate following the method used to purify cholate in Protocol 2. • A boiled de-aerated suspension of coursegrade Sephadex G-50. prepared in deionized water

RECONSTITUTION OF MEMBRANE PROTEINS Protocol 6 continued

Method 1. Aliquot 5 mg of di(C18:l)PC and 20 mg di(Cl8:l)PE into a 10 ml glass vial. Remove the solvent under a stream of nitrogen so that the lipid forms a thin film on the side of the vial. Place in a vacuum desiccator and dry down under vacuum for about 3 h. 2. Add 600 ul of the phosphate buffer and 25 mg of potassium cholate, added as a 10% (w/v) solution, giving a cholate:lipid ratio (w/w) of 1:1. Vortex for 15-30 sec, flush the vial with nitrogen and seal with Parafilm. Then sonicate to optical clarity using the bath sonicaton this will probably take about 10-15 min. 3. To 2 mg of purified ATPase, present as a 20-30 mg/ml suspension (prepared as in Protocol 2)t add 6 ul of the 10% deoxycholate solution to give a final deoxycholate: ATPase ratio (w/w) of 0.6:1. Remove any large unsolubilized aggregates by brief centrifugation at full speed in an Eppendorf inicrocentriftige. 4. Add 1.3 mg of the solubilized ATPase to the solubilized lipid (25 mg), giving a molar ratio of lipid: ATPase of about 1500:1. 5. Equilibrate the Sephadex G-50 with the phosphate buffer. 6. Plug the ends of two 5 ml disposable plastic syringes with glass wool and fill with about 6 ml of the Sephadex G-50 suspension. Centrifuge the syringes in a benchtop centrifuge at about 200 g for 30 sec. 7. Apply the sample to the top of one of the syringes. Repeat the centrifugation and collect the reconstituted vesicles in the eluate. 8. Repeat the procedure using the second syringe to give the final preparation of sealed vesicles.

6 Spectrophotometric assay of Ca2+ accumulation Ca21 accumulation by the vesicles can be measured spectrophotometrically as described in Protocol 7, using the Ca 2- -sensitive dye Antipyralazo III to monitor the drop in external Ca2+ concentration as Ca21 is pumped into the vesicles.

7 Calculation of the internal volume of the

vesicles The external radius of the preparation of reconstituted vesicles can be measured using light scatter. The internal volume of the vesicles can be calculated from the external radius (ra) as illustrated in Hgure 4, The thickness of the bilayer, 6, is taken to be 4 nm. The internal radius r. is given by;

39

ANTHONY G. LEE

The internal volume of a vesicle is given by (4/3)rri3 and the volume of the lipid bilayer in a vesicle is given by (4/3)Tr[(rj + S)J - ri3)\. Thus the internal volume of a vesicle (Vi) can be related to the mass of lipid in a vesicle (Mi) by:

Protocol 7 Spectrophotometrlc determination of Ca2+ accumulation Equipment and reagents

• A dual-wavelength spectrophotometer (e.g. Aminco-Bowman DW2000: SLMAminco) with a stirred, temperaturecontrolled sample compajtment • 2.3 mM stock solution made by dissolving 7.4 mg Antipyralazo III in 4 ml water; filter the solution twice to remove any undissolved solids • 100 mM stock solution of Ca2+ (see Protocol 3)

• 1 mM stock solution of carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP) in methanol • Assaybuffer:10mMPipespH7.1,100mM K2SO4, 5 mM MgS04 • 11.8 mM stock solution of ATP (6 rug/ml) in assay buffer

Method 1. Make up 3 ml of an 80 |uM solution of Antipyralazo III in the assay buffer in a stirred cuvette in a dual-wavelength spectrophotometer, at 25°C

2. Add an aliquot of the reconstituted vesicles to give a concentration of 0,04 mg protein/ml. 3.

Add 0.75 u1 of the stock solution of FCCP.

4.

Measure the absorbance difference with a wavelength pair of 720 and 790 run.

5.

Calibrate the absorbance signal by adding known amounts of Ca 2+ . Do this by diluting the stock Ca2+ solution 1 in 10 to give a 10 mM stock solution. Add three 12 ul aliquots of the 10 mM stock Ca2+ solution to the cuvette; each addition is equivalent to increasing the Ca2+ concentration in the cuvette by 40 uM

6. Add an aliquot of the stock solution of ATP to give a final concentration of 0,8 mM ATP and monitor the change in AA. Using the calibration established in step 5, calculate the level of accumulation of Ca2+, expressed as nmoles Caa+ accumulated/ mg protein, from the change in AA. 7. Repeat the above uptake assay in the absence of FCCP.

40

RECONSTITUTION OF MEMBRANE PROTEINS

where p is the density of the lipid, taken to be 1 g/ml, typical of organic compounds, and all dimensions are in nm. Thus the internal volume (nm3) per mg of lipid in the sample will be:

The internal volume can be converted from nm3 to litres using the relationship that 1 litre = 1024 nm3. To show what these equations mean, a vesicle with an external radius of 50 nm will have an internal volume of 4.1 x 10-19 litre. If the ATPase is reconstituted at a lipid/protein ratio (w/w) of 40:1, the total internal volume will be 140.8 ul/mg protein. The number of lipids per vesicle will be 93 000 and the number of ATPase molecules per vesicle will be 15. If only about a third of the ATPase molecules are active, and if the distribution of ATPase molecules across the membrane is random, then each vesicle will only contain about three molecules of ATPase able to react with added ATP and pump Ca2+ ions into the vesicles. If the vesicle accumulates Ca2+ to an internal Ca2+ concentration of 1 mM, the vesicle will contain just 240 Ca2+ ions.

8 Simulation of Ca2

accumulation

As shown in Figure 4 and Scheme 1, the accumulation of Ca2+ by the Ca2+-ATPase reconstituted into sealed vesicles will be a balance between the uptake of Ca2+ driven by the Ca2+-ATPase and the leak of Ca2+ out of the vesicles. Two forms of leak can be distinguished: a simple passive leak, unconnected to the process of Ca2+ uptake, and slippage, in which the phosphorylated intermediate of the ATPase, instead of releasing Ca2+ into the lumen of the vesicle, releases it to the outside. Accumulation of Ca2+ by reconstituted vesicles will depend on the rate of hydrolysis of ATP, on the rates of passive leak and slippage, on the internal volume of the vesicles, and on the affinity of the phosphorylated intermediate for Ca2+. The relative importance of all these factors is best understood by simulating the accumulation process on a computer.

Scheme 1 A simplified reaction scheme for the Ca2+-ATPase, showing passive leak and slippage pathways.

41

ANTHONY G. LEE

The kinetic pathway for ATP hydrolysis by the ATPase is relatively well understood, and is shown in simplified form in Scheme 1. The scheme proposes that two Ca2+ ions bind in a cooperative fashion to the El conformation of the ATPase, from the cytoplasmic side of the membrane (20,21). Following binding of MgATP to the ATPase, the ATPase is phosphorylated and undergoes a change in conformation to a state in which the two Ca2+ binding sites are of low affinity and inward facing (E2PCa2). Ca2+ is lost from this phosphorylated intermediate to the lumen of the SR. Dephosphorylation of E2P then allows recycling to El (22). Equilibrium and rapid, pre steady-state kinetic measurements have defined the rates and equilibrium constants for all the steps in the uptake pathway, so that the only unknowns in the accumulation process are the rates of simple leak and slippage; these can then be obtained by comparisons of simulations of the time course of Ca2+ accumulation with the experimental data. Simulations of complex kinetic pathways can be mathematically difficult. However, a package such as FACSIMILE (from AEA), running on a PC system, takes care of all the mathematics and just requires a definition of the steps of the reaction sequence to be simulated and the values for the rate constants of these steps. This information is provided in the form of a subroutine. The subroutine necessary to solve a highly simplified model of the Ca2+-ATPase is given in Protocol 8. Despite its highly simplified nature, it illustrates a number of important, non-obvious features of Ca2+ accumulation. The subroutine can readily be extended to include a more realistic model for such steps as the E2-E1 conformational change and Ca2+ binding (21). Steps 1-8 under 'COMPILE EQUATIONS' define the uptake pathway; forward and backward rate constants for these steps are listed under 'COMPILE INITIAL'. Step 9 is the slippage step, and step 10 is simple passive leak. The rates of steps 9 and 10 are varied to observe their effects on the accumulation process. In the experiments shown in Figure 5, the total concentration of ATPase was 0.175 uM and, assuming that 50% of the ATPase is functional and with a random distribution of ATPase molecules across the membrane, the concentration of correctly oriented, active ATPase molecules will be 0.045 uM. At the start of the experiment, before the addition of ATP, the ATPase will be present in its Ca2+-bound form; the initial concentration of ElCa2 is therefore 0.045 |xM. The initial concentrations of ATP and Ca2+ (in the external medium) are 0.8 and 0.12 mM, respectively, in the experiment shown in Figure 5. The only other factor that needs to be accounted for is the relationship between the external volume of the sample and the internal volume of the vesicles. In the experiments shown in Figure 5, the Ca2+-sensitive dye monitors the change in external Ca2+ concentration as Ca2+ is pumped by the ATPase into the lumen of the vesicles. The increase in Ca2+ concentration in the lumen of the vesicles resulting from the transport of a given amount of Ca2+ into the lumen will be much greater than the corresponding drop in Ca2+ concentration in the external medium, because of the small lumenal volume. For the experiments shown in Figure 5, the external vesicle radius (r0) was 47 nm, which, with a lipid concentration of 0.8 mg/ml and a lipid:protein ratio of 40:1 (w/w) gives an internal to external volume ratio of 382.6; this 42

RECONSTITUTION OF MEMBRANE PROTEINS

Protocol 8 The subroutine necessary for simulation of Ca accumulation using FACSIMILE VARIABLE El E2 E1Ca ElCa2 Ca atp ElCa2atp E2PCa2 Cai E2PCa E2P; PARAMETER klf klb k2f k2b k3£ k3b k4f k4b k5f k5b k6f k6b k7f k7b k8f k8b k9f k9b kl0f klOb intconc eaup v; EXEC OPEN 8 'dump.res'; COMPILE

INITIAL;

ElCa2=0.045e-6; Ca=120.0e-6; Cai=1.0e-12; atp=0.8e-3; klf=13.0; klb=47.0; k2f=1.0e8; k2b=1.0e2; k3f=1.0e8; k3b=1.0e2; k4f=1.0e8; k4b=1.0e3; k5f=1.0e2; k5b=1.0e-6; k6f=60.0; k6b=1.8e4; k7f=30.0f k7b=0.9e4f k8f=100.0f k8b=1.0e-6f * k9f and k10f define the rates of slippage and simple leak respectively; * to introduce slippage vary k9f and put k10f=1.0e-10; * to introduce sijmple leak vary k10f and put k9f=1.0e-10; k9f=1.0e-6; k9b=1.0e-6f k!0f=1.0e-10; k10b=1.0e-10; v=382.6;

**; COMPILE EQUATIONS; %klf*klb: E2=E1;

43

ANTHONY G. LEE

Protocol 8 continued %k2f%k2b: El+Ca=ElCa; % k f % k 3 b : ElCa+Ca=ElCa2; %k4f%k4b: ElCa2+atp=ElCa2atp; %k5f%k5b: ElCa2atp=E2PCa2; %k6f%k6b: E2PCa2=E2PCa+Cai+v; %k7f%k7bt E2PCa=E2P+Cai+v; %k8f%X8b! E2P=E2; Ik9f%k9b: E2PCa2=E2+Ca+Ca; % k 1 0 f % k 1 0 b : Cai+v=Ca; **. COMPILE PRINT; caup =((120. Oe-6)-c:a)*l,Oe9/20.0; PSTREAM 2; **; PSTREAM 2 6 5 ; time caup;

**; WHEN TIME = 0 + 10.0 *100%CALL PRINT;

**; BEGIS; STOP;

therefore represents the factor by which the concentration of Ca2 changes on moving from the external medium into the vesicle lumen and is included as the factor v in steps 6, 7, and 10 of the reaction sequence (under 'COMPlLIi EQUATIONS'), The results of these simulations are shown in Figure 6, assuming either just passive leak (see figure 6A) or just slippage (see Figure 6ii). It is clear that passive leak and slippage have very different effects on the time course of accumulation of C a 2 , and that the experimental data (see figure 5) are matched by the slippage model rather than by passive leak; the presence of anionic phospholipid leads to increased levels of accumulation of Ca2- because the rate of slippage decreases (17).

Acknowledgements I thank Kate Dalton, Richard Duggleby, Malcolm East, Sanjay Mall, Jeffrey Pilot, Anthony Starling, Richard Webb, and Ian Williamson who developed most of the protocols described here. 44

RECONSTITUTION OF MEMBRANE PROTEINS

Figure 6 Simulation of the process of Ca2+ accumulation. In (A) it is assumed that leak occurs only by the simple passive pathway, with rates of passive leak (k10f, s-1) of: (a), 1.0 x 10-10; (b), 5.0 x 10-6; (c), 1.0 x 10~5; (d), 3.0 x 10-5. In (B) it is assumed that leak is only by the slippage pathway, with rates of slippage (kgf, s-1) of: (a), 1.0 x 10-6; (b), 50; (c), 150; (d), 500.

References 1. Warren, G. B., Toon, P. A., Birdsall, N. J., Lee, A. G., and Metcalfe, J. C. (1974). Biochemistry, 13, 5501. 2. Holloway, P. N. (1973). Anal. Btochem., 53, 304. 3. Daiho, T., Kubota, T., and Kanazawa, T. (1993). Biochemistry, 32, 10021. 4. Grimes, E. A., Burgess, A. J., East, J. M., and Lee, A. G. (1991). Biochim. Eiophys. Acta, 1064, 335. 5. Gould, G. W., Colyer, J., East, J. M., and Lee, A. G. (1987). J. Biol. Chem., 262, 7676. 6. Gafni, A. and Boyer, P. D. (1984). Biochemistry, 23, 4362. 7. Michelangeli, F., Orlowski, S., Champeil, P., Grimes, E. A., East, J. M., and Lee, A. G. (1990). Biochemistry, 29, 8307. 8. Khan, Y. M., Wictome, M., East, J. M., and Lee, A. G. (1996). Biochem.}., 317, 433. 9. Warren, G. B., Toon, P. A., Birdsall, N. J., Lee, A. G., and Metcalfe, J. C. (1974). Proc. Natl acad. Sri., 71, 622.

45

ANTHONY G. LEE 10. Starling, A. P., East, J. M., and Lee, A. G. (1995). Biochemistry, 34, 3084. 11. Starling, A. P., East, J. M., and Lee, A. G. (1993). Biochemistry, 32, 1593. 12. MacDonald, R. C., MacDonald, R. I., Menco, B. P. M., Takeshita, K., Subbarao, N. K., and Hu, L. R. (1991). Biochim. Biophys. Acta, 1061, 297. 13 Rigaud, J. L., Pitard, B., and Levy, D. (1995). Biochim. Biophys. Acta, 1231, 223. 14. Racker, E. (1985). Reconstitution of transporters, receptors and pathological states. Academic Press, New York. 15. Gould, G. W., McWhirter, J. M., East, J. M., and Lee, A. G. (1987). Biochim. Biophys. Acta, 904, 36. 16. Levy, D., Gulik, A., Bluzat, A., and Rigaud, J. L. (1992). Biochim. Biophys. Acta, 1107, 283. 17. Lee, A. G. and East, J. M. (1998). Biochem. Soc. Trans., 26, 359. 18. Yu, X., Carroll, S., Rigaud, J. L, and Inesi, G. (1993). Biophys. J., 64, 1232. 19. Penefsky, H. S. (1979). In Methods in enzymology. Vol. 56 (ed. S. Fleischer and L. Packer), p. 527. Academic Press, London. 20. Henderson, I. M. J., Khan, Y. M., East, J. M., and Lee, A. G. (1994). Biochem. J., 297, 615. 21. Lee, A. G., Baker, K., Khan, Y. M., and East, J. M. (1995). Biochem. J., 305, 225. 22. Champeil, P. (1996). In Biomembranes, Vol. 5. ATPases (ed. A. G. Lee), p. 43, JAI Press, Greenwich,.

46

Chapter 3 The Xenopus oocyte expression system for the cDNA cloning and characterization of plasma membrane transport proteins Sylvia Y. M. Yao,* Carol E. Cass,1 and James D. Young* * Department of Physiology, Faculty of Medicine, 7-55 Medical Sciences Building, University of Alberta, Canada T6G 2H7 + Department of Oncology, Cross Cancer Institute, Edmonton, Alberta, Canada T6G 1Z2

1 Introduction Oocytes of Xenopus laevis, the South African clawed frog, represent a simple, but powerful system for the transient expression of heterologous proteins. These large and resilient cells have a high translational capacity and are readily microinjected with exogenous RNA or DNA. As a system for studying plasma membrane transport proteins, Xenopus oocytes are readily amenable to both radioisotope flux measurements and electrophysiological recordings. They also subject newly synthesized proteins to various forms of processing, including glycosylation and correct plasma membrane insertion. Unlike other whole-cell heterologous expression systems, Xenopus oocytes have a generally low background of endogenous transport activity, making them uniquely versatile with respect to the range of permeants and types of transporters that can be studied. The ability of Xenopus oocytes to translate foreign mRNA was first described in 1971 (1). Since then, the system has been applied with considerable success to the study of receptors and channels and, following the landmark expressioncloning of the intestinal glucose transporter SGLT1 (sodium-dependent glucose transporter 1) in 1987 (2), to transport proteins. In this chapter, we provide a laboratory guide to the Xenopus oocyte expression system for the cDNA cloning and functional and molecular characterization of plasma membrane transport proteins, illustrating some of these applications with our own studies of nucleoside transport processes in mammalian and other species. Other sources should be consulted for routine molecular biology techniques (3).

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SYLVIA Y. M. YAO ET AL.

2 The Xenopus oocyte system 2.1 Isolation and maintenance of Xenopus laevis oocytes

2.1.1 Sources of Xenopus laevis Mature, laboratory-bred, oocyte-positive, female Xenopus laevis are available from Nasco, Xenopus 1 and, in the UK, from Xenopus Ltd. We keep our Xenopus in covered tanks in chlorine-free tap water (depth 15-25 cm, ~ 3 litres of water per frog) under controlled lighting conditions (light-dark cycle, 12 h each) and a constant temperature of 16-18°C. A pellet diet is available from Tetra (0.2 g/ frog, daily). Multiple small batches of oocytes can be obtained from a single frog by surgical removal of single ovarian lobes (4), allowing a series of experiments to be performed on oocytes from the same animal. Alternatively, as in our laboratory, the animal can be sacrificed, making available more cells for large-scale experiments. The procedure described for handling Xenopus in Protocol 1 conforms to Canadian Council on Animal Care (CCAC) guidelines and may differ from requirements in other countries. Individual Xenopus can vary considerably in their numbers of usable oocytes, and occasional frogs will be encountered in otherwise good condition that yield poor quality oocytes unsuitable for further preparation and processing.

2.1.2 Preparation and maintenance of oocytes The ovaries of adult female Xenopus contain mixtures of oocytes at six different stages (I-VI) of development (5). Only the last two (stages V and VI) are generally used in expression studies. As shown in Figure 1, Stage V and VI oocytes are large cells (1.0-1.3 mm in diameter) with a light vegetal pole and a darker animal pole (containing the nucleus) and are readily distinguished from smaller, more immature forms. We use mixed populations of stage V and stage VI cells. A good Xenopus will have ovaries that contain > 70% stage V/VI oocytes. Since the number of usable oocytes per ovary may decrease with time, we purchase frogs in relatively small batches (e.g. 10 at a time) and use them within 3 months of arrival. Seasonal factors can influence oocyte numbers and quality and may lead to variable levels of expression, particularly for 'difficult' proteins. In our experience, the period September-June gives the most consistent results. As described in Protocol 1, ovary tissue is first treated with collagenase to release the individual oocytes and detach a surrounding layer of follicle cells. This is followed by hypertonic phosphate treatment to complete the removal of the follicle layer. The resulting shrunken cells, which are still surrounded by an inner glycoprotein matrix layer known as the vitelline membrane, are restored to normal osmolarity and stabilized for 24 h before microinjection. The vitelline layer provides stability to the isolated oocytes and does not interfere with radioisotope flux assays or whole-cell electrophysiological recordings. It can be removed (or ruptured) manually (6) if high-resistance seals are required for patch-clamp analysis. 48

THE XENOPUS OOCYTE EXPRESSION SYSTEM

Protocol 1 Isolation of Stages V-VI oocytes Equipment and reagents • Sterile scissors and forceps • Sterile Petri dishes (100 mm) • Stereoscopic microscope (Nikon SMZ-tB, or equivalent) • Shaker (New Brunswick Gyrotory G2, or equivalent)

• Incubator (16-18°C) • Type I collagenase (CLS-1 326 U/mg) (Worthington Biochemical Corp., or equivalent)

Modified Barth's Medium (MBM): 88 mM NaCl, 1 mM KC1, 0.33 mM Caf(NO3)2. 0.41 mM CaCl2. 0.82 mM MgS04. 2.4 mM NaHCO3.2.5 mM sodium pyruvate, 0,1 mg/ml penicillin, 0.05 mg/ml gentamicin sulfate. 10 mM Hepes pH 7.5 (filtersterilized) BSA wash: 1 mg/ml BSA in MBM Phosphate medium: 1 mg/ml BSA, 100 mM K2HPO4 pH 6.5

Method 1. To anaesthetize and then kilt a Xenopus laevis, place the frog directly in ice-water for 15 min, then stun and pith. 2. Place the killed frog. ventral side up, on a dissection tray and swab the abdomen with 70% (v/v) ethanol. 3. Make an incision in the skin of the abdomen with sterile scissors and forceps, 4. Cut open the skin horizontally, 5. Cut open the underlying muscular layer to expose the ovarian lobes. 6. Take out the ovarian lobes with a pair of sterile forceps and place into a sterile Petri dish (100 mm) containing MBM. 7. Wash the lobes twice in MBM. 8. Tear open the lobe membranes and wash again with MBM. 9. Dissect the lobes into smaller clumps (about 5-10 oocytes each) using a pair of fine sterile forceps. 10. Incubate the clumps of oocytes in MBM containing 2 mg/ml type I collagenase for 2 h at room temperature with moderate agitation. 11. Wash the oocytes five times with the BSA wash solution and then five times with MBM. 12. Select Stage V-VI oocytes (see Section 2.1.2) under a dissection microscope (10-20 x magnification). 13. Incubate the oocytes in MBM in a 16-18°C incubator overnight 14. Incubate the oocytes in phosphate medium for 1 h at room temperature with gentle agitation. 15. Wash the oocytes five times with BSA wash solution and five times with MBM. 16. Select the healthy oocytes under the microscope. 17. Incubate the oocytes in MBM at 16-18°C for 24 h before microinjection.

SYLVIA Y. M. YAO ET AL.

Figure 1 Xenopus laevis oocytes. (A) Shows a typical mixed population of defolliculated oocytes isolated by treatment with collagenase and hypertonic phosphate medium from the ovary of a mature female Xenopus laevis as described in Section 2.1.2 and Protocol 1. The small cells with differentiated hemispheres are Stage IV, while those between 1000 and 1300 (um are Stages V and VI. Stage V and VI oocytes are used for expression experiments. (B) Shows the boxed group of cells in (A) at higher magnification: a. a Stage V cell with clearly delineated hemispheres and a light animal pole; b, a slightly larger Stage VI oocyte, distinguished from the Stage V cell by an unpigmented equatorial band between the two hemispheres; c, an unhealthy oocyte with uneven pigmentation; d, a Stage VI oocyte with a small region of discoloration at the vegetal pole. Cells c and d will not survive and should be discarded prior to microinjection.

Type I collagenase from Worthington Biochemical Corp, or type A collagenase from Boehringer Mannheim (now Roche Diagnostics) arc generally suitable for oocyte work, but individual batches should be tested for both enzyme activity (the ability to separate oocytes from each other) and oocyte viability and translation-al capacity. We screen multiple batches of enzyme from different suppliers and order (or reserve) large quantities of the most suitable batch. The collagenase-phosphate treatments should be performed at the lowest enzyme concentration for the minimum duration necessary to achieve the required results. Calcium-free medium can be used to minimize the proteolytic activity of clostripain present in collagenase preparations. Oocytes should be carefully sorted at each stage to remove damaged cells and those showing visible signs of deterioration, such as leakage of egg yolk, changes in coloration (including uneven patches of pigment), or altered sharpness of the vegetal/animal pole boundary. Oocytes survive best in relatively large volumes of medium and should not be crowded, especially after microinjection. A good frog may contain 10000 usable oocytes. This will decrease to perhaps 60007000 after collagenase digestion and 4000 after phosphate treatment, yielding 2000-3000 cells suitable for microinjection. A further 10-20% loss may be expected during microinjection and subsequent handling and incubation, and 50

THE XENOPUS OOCYTE EXPRESSION SYSTEM

some investigators add 5-10% horse serum to the culture medium after microinjection to promote oocyte viability (7). Since a typical radioisotope flux experiment may require upwards of 500 oocytes, a single frog will generally provide sufficient cells for several experiments and/or researchers.

2.2 Oocyte microinjection 2.2.1 Sources of RNA and cDNA (general considerations) Foreign genetic material is introduced into Xenopus oocytes by cytoplasmic (RNA) or nuclear (cDNA) microinjection. RNA can be mRNA isolated from cells or tissues, or synthetic RNA transcript prepared in vitro from a cDNA template, and is microinjected into the vegetal pole of the cell to avoid the nucleus. Alternatively, cDNA can be microinjected directly into the nucleus (8). This eliminates the need to produce synthetic RNA transcript, but it may damage the nucleus. Only a portion of attempted nuclear injections will be successful (because of difficulty in locating the nucleus) and procedures have been devised to identify transfected oocytes, such as coexpression of a released marker enzyme (9). A method employing the vaccinia virus that does not require the nuclear injection of cDNA has also been described (10). We will only consider microinjection of RNA.

2.2.2 Microinjection An optimal system for Xenopus oocyte microinjection requires a good dissecting stereomicroscope, a micromanipulator that can be moved in three dimensions, a cold light source (to avoid overheating) and, if large numbers of microinjections are to be undertaken, an electronically controlled microinjection apparatus to deliver multiple predetermined volumes of RNA. The microinjection system (from Inject + Matic) mentioned in Protocol 2 is driven by air pressure, and has both a vacuum and pressure generator to facilitate cleaning and loading the pipette. Sample delivery is controlled by a foot switch, leaving both hands free to control the microinjector and other equipment. Alternative microinjection systems include the PLI-100 Picoinjector (Harvard Apparatus) and the Nanoject oocyte injector (Drummond Scientific). With experience, 300-500 oocytes can be microinjected in a 1-h session. If only small numbers of microinjections are contemplated, a manual system, such as a syringe driven by a micrometer screw, may suffice (4, 8). Access to a micropipette-puller is also required. It is not necessary to polish pipette tips in a microforge. Microinjections can be performed on the open bench using a standard semi-sterile technique. Oocytes are typically microinjected with 10-50 nl (maximum 100 nl) of RNA dissolved in RNase-free water at a concentration of 1 ug/ul Control oocytes are injected with water alone, and injection volumes can be calibrated using an eyepiece micrometer to measure the diameter of an expelled spherical water droplet while it remains attached to the pipette tip (4). The culture period 51

SYLVIA Y. M. YAO ET AL

Protocol 2 Microinjectlon procedure Equipment and reagents • Stereoscopic microscope (Nikon SMZ-lB. or equivalent) • Fibre-optic light source (Nikon MKH, or equivalent) • Micropipette-puller (Inject + Matic, or equivalent) • Microinjector (Inject + Matic, or equivalent) • Micromanipulator (Singer Instruments, MK1 or equivalent)

• Glass capillary tubes o.d. 0.55 mm, 5u1, length 75 mm (Singer Instruments, or equivalent) • Incubator (16-18°C) • MBM (see Protocol 3) • 1 ug/ul RNA in RNase-free water • Sterile, 35 mm Petri dishes • Sterile. 5 ml glass vials

Method 1. Prepare the micropipette from a glass capillary tube using the micropipette-puller, After pulling, the tip of the micropipette should be broken off at a diameter of —10-20 um." Autoclave micropipettes before use. 2. Insert the micropipette into the micromanipulator. 3. Place 1-4 ul the RNA solution or water on to a Petri dish (35 mm) positioned on top of an ice block. 4. Fill the micropipette with the RNA solution or water. 5. Adjust the injection volume by controlling the duration and magnitude of the force applied by the injector.b 6. Place 5-10 healthy oocytes against the side of a Petri dish lid (35 mm) positioned on top of the ice block. Remove most of the surrounding MBM. 7. Adjust the microscope such that the oocytes are in focus. Gradually lower the micropipette until the tip is level with the oocyte surface, 8. Penetrate the surface of the oocyte with the tip of the micropipette to a depth of 0.1-0.2 mm and inject 10-50 nl of the RNA solution or water. Gently remove the micropipette from the oocyte and repeat for the other cells. 9. Place the microinjected oocytes in a 5 ml sterile glass vial in MBM and store at 16-18°C. Change the medium daily and discard unhealthy oocytes. a

The diameter of the tip can be measured by a micrometer scale under the dissection microscope. b The volume of solution injected can be measured as described in Section 2.2.2. 'A noticeable slight swelling of the oocyte indicates a successful microinjection.

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THE XENOPUS OOCYTE EXPRESSION SYSTEM

required for optimal expression of transport activity may vary for different transporters and should be determined empirically for each RNA. In our experience, 3 days are usually required for synthetic RNA transcript, and 5-7 days for mRNA. Only good-quality, healthy oocytes should be microinjected. Care should also be taken to ensure that RNA solutions (particularly samples of mRNA) are free of particulate matter, or the pipette tips may become blocked. We recommend filtering mRNA preparations through 0.2 um microfilters (Costar) before use. The success rate of transfection following cytoplasmic injection of RNA should be > 95%. Since transport assays are performed on individual oocytes (see Section 6), the small number of cells without transport activity are readily identified and eliminated from data analysis.

3 Isolation and size-fractionation of poly (A)+ RNA (mRNA) from mammalian tissues The expression-cloning of a transport protein in Xenopus oocytes requires a source of mRNA that shows the required functional activity. mRNA is therefore isolated from cells or tissues that are known or suspected to contain the transporter of interest and injected into oocytes. The transfected cells are then assayed for the appropriate transport function as described in Section 6. Multiple activities may be induced, and it may be necessary to modify flux conditions (e.g. by measuring transport with and without a transport inhibitor or Na + ) to make the assay more specific. Antisense hybrid-depletion with synthetic oligonucleotides can be used to eliminate contributions from known proteins with overlapping functional characteristics (11). The next step in the process is to size-fractionate the mRNA to enrich the desired transport activity (membrane transporter mRNAs are usually present in low abundance) and eliminate the majority of unwanted transcripts. As described in Section 4, the mRNA size fraction with greatest functional activity is then used to construct an expression cDNA library. It is essential to prepare high-quality, intact mRNA and to achieve good size-fractionation of that mRNA. Rigorous precautions should be taken against RNase contamination, including the use of baked glassware, RNase-free pipette tips and other disposable plasticware, and water and reagent solutions treated with diethylpyrocarbonate (DEPC). The integrity and purity of mRNA should be suspected if injected oocytes fail to survive or if unusually low levels of expression are obtained. RNA preparations can be monitored by denaturing agarose gel electrophoresis and UV absorption.

3.1 Isolation of total RNA 3.1.1 The guanidinium thiocyanate and CsCI purification method The initial step in the preparation of mRNA is the isolation of total RNA. The guanidinium thiocyanate/CsCl method described in Protocol 3 is relatively time consuming, but is recommended for most applications. It is particularly 53

SYLVIA Y. M, YAO ET AL.

suitable for samples with a low RNA:protein ratio or a high lipid content. The centrifugation time for small samples (< 3 ml) can be reduced to 2 h using a Beckman TL-100 tabletop ultra centrifuge and TLA-100.3 rotor, 3.1.2 Single-step purification method (Gibco BRL Trizol™ reagent) This method is quick, and is an alternative to guanidinium thiocyanate-CsCl for small samples. The manufacturer's protocol should be followed and, if necessary, repeated twice on difficult samples.

Protocol 3

Isolation of total RNA by the guanidinium thlocyanate and CsCI method Equipment and reagents NB: All reagents should be RNase-free and all glassware should be baked at 180° C for 8 h. » Liquid nitrogen • Homogenizer (Caframo RZR1 stirrer type, or equivalent) • 14 x 89 mm polyallomer ultracentrifuge tubes • Ultracentriruge (Beckman L-80 and SW41 rotor, or equivalent) • Refrigerated centrifuge (Sorvall RC-5B and SA-600 rotor. or equivalent)

10% (w/v) sodium lauryl sarcosine GTH buffer: 4 M guanidinium thiocyanate, 0.1 M Tris-HCl pH 7.5 (filtered through a Whatman No. 1 filter). 1% (v/v) Bmercaptoethanol (add immediately before use) CsCI solution: 5.7 M CsCI, 0.1 M EDTA pH 7.5 (filter-sterilized)

Method 1. Freeze the tissue sample in liquid nitrogen immediately after dissection and grind into fine powder. Homogenize in GTH buffer (10 ml solution per g of tissue). Add 10% (w/v) sodium lauryl sarcosine to give a final concentration of 0.5%, 2. Centrifuge the homogenate (5000 g for 10 min at 4°C) in a Sorvall SA-600 rotor. then layer the resulting supernatant on to a CsCI cushion (3 ml CsCI solution per 7 ml of supernatant in a 14 x 89 mm polyallomer ultracentrifuge tube). Centrifuge at 111 000 g for 24 h at 20°C in a Beckman SW41 rotor. 3. Aspirate the supernatant until near the bottom of the tube. then carefully discard the remaining solution. Cut off the bottom of the tube which contains the RNA pellet. 4. Wash the RNA pellets twice in 70% (v/v) ethanol (-20°C) and resuspend in RNasefree water at a concentration of 1 5. Store the RNA solution at -70°C,

54

THE XENOPUS OOCYTE EXPRESSION SYSTEM

3.2 Purification and size-fractionation of poly(A)^ RNA 3.2.1 The PolyAtract™ (Promega) mRNA isolation system Several manufacturers supply kits to isolate poly(A)' RNA (mRNA) from total RNA. In the PolyAtract™ mRNA isolation system {Promega), poly(A)' RNA is first annealed to biotinylated oligo(dT) and then captured on streptavidin-coated paramagnetic beads. The kit produces good quality mRNA, but is expensive for large-scale preparations. A typical yield of mRNA is 3-4% of the starting total RNA. 3.2.2 Oligo(dT)-cellulose affinity chromatography The use of o]igo(dT)-cellulose chromatography (Protocol 4) is more time consuming than PolyAtract™ isolation, but produces similar quality mRNA and can be scaled up for the large amounts of material needed for mRNA sizefractionation and cDNA library construction. Oligo(dT)-cellulose can be reused, but this is not recommended for critical samples.

Protocol 4

Isolation of poly(A)+ RNA by ollgo(dT)-cellulose affinity chromatography Equipment and reagents NB: All reagents should be RNase-free • Oligo(dT)-celIulose (Boehringer Mannheim, now Roche Diagnostics) • Econo-Pac dispocolumn(1.5 x 12cm)(BioRad}

• Binding buffer: 10 mM Tris-HCl pH 7.5, 0.5 M LiCl, 1 mM EDTA, 0.1 % (w/v) SDS

• Washing buffer: 10 mM Tris-HCl pH 7.5. 0.1 M LiCI. 1 mM EDTA, 0.1 % (w/v) SDS • Elution buffer 10 mMTris-HCIpH 7.5. 2 mM EDTA, 0.1 % (w/v) SDS • 10M LiC1 • 3 M sodium acetate pH 6

Method 1. Equilibrate 0.15 g dry oligo(dT)-celtulose in 5 ml of elution buffer for 5 nan, then pour the slurry into an autoclaved dispocolumn. 2. Wash the column with 10 column volumes of binding buffer, 3. Heat the total RNA sample (~ 2 mg) to 65 °C for 10 min. then chill on ice for 5 min, Adjust the RNA sample to 0.5 M LiCl with 10 M LiCl, 4. Load the RNA sample on to the oligo(dT)-cellulose column and save the eluate. Reheat the collected eluate to 65 °C for 10 min and chill on ice for 5 min. Pass through the column for a second time. 5. Wash the column with 5 column volumes of binding buffer, then with 5 column volumes of washing buffer.

55

SYLVIA Y. M. YAO ET AL. Protocol 4

6. Elute the poly(A)+ RNA with 2 column volumes of the elution buffer. 7. Re-purify the eluted poly(A)+ RNA on a second otigo(dT)-ceUulose column (0.05 g dry oligo(dT)-cellulose) using the same procedures described in steps 1-6 above. 8. Precipitate the poly(A)+ RNA by adding 1/10 volume of 3 M sodium acetate (pH 6) and 3 volumes of cold ethanol (-20 °C), then keep at -20°C for 1 h. Collect the poly(A)+ RNA by centrifugation at 10000 g for 30 min at 4°C Wash the RNA pellet with 70% (v/v) ethanol (-20 °C) and store at -70°C as a pellet.

3.3 Size-fractionation of poly(A) + RNA by non-denaturing, sucrose -gradient centrifugation Non-denaturing, sucrose-gradient centrifugation {Protocol 5) is the most common method of mRNA size-fractionation, although non-denaturing, agarose gel elcctrophoresis is also used (12, 13). It is important to start with sufficient mRNA (typically 400 ug), since each fraction needs to be tested for functional activity and should have sufficient material left over, if necessary, for library

Protocol 5 Size-fractionation of poty(A)+ RNA Equipment and reagents NB: All reagents should be RNase-free • Ultracentrifuge (Beckman L-80 and SW41 rotor, or equivalent) • 14 X 89 mm polyallomer ultracentrifuge tubes • 10-21% (w/v) sucrose gradient in TE buffer

• TE buffer: 10 mM Tris-HCl pH 7.4.1 mM EDTA • Fraction recovery system (Beckman, or equivalent)

Method 1. Heat the poly(A)+ RNA (~ 400 ug in 0.5 ml TE buffer) to 65°C for 10 min, then chill on ice for 5 min. 2. Load the heat-denatured poly(A)+ RNA on to a gradient composed of 11 x l ml graded concentrations of sucrose (10-21 % (w/v) in TE buffer) in a 14 X 89 mm polyallomer Ultracentrifuge tube and centrifuge (150000 g for 20 h at 4°C) in the Beckman SW41 rotor. 3. Collect fractions (0.6 ml) using the fraction recovery system (Beckman). Precipitate the poly(A)+ RNA (see Protocol 4, step 8). 4. Dissolve the precipitated poly (A)+ RNA in RNase-free water (~ 1 ug/ul) and store at -70°C. Examine the size range of poly(A)+ RNA in each fraction by denaturing agarose gel electrophoresis (Protocol 6).

THE XENOPUS OOCYTE EXPRESSION SYSTEM

Figure 2 Influx of uridine in Xenopus oocytes microinjected with size-fractionated mRNA. (A) Shows a denaturing agarose gel of rat jejunal mRNA which was size -fractionated by sucrose-gradient centrifugation as described in Section 3.3 and Protocol 5. Samples {-1 ug RNA) containing ethidium bromide were run with BRL size markers on an agarose gel under conditions similar to those described in Protocol 6. The peak sizes of mRNAs in fractions 3-16, estimated by laser densitometry and calculated by reference to positions of the molecular weight standards, were 4.4, 3.7, 3.4, 2.9, 2.3, 2.1, 1.8, 1.6, 1,4. 1.0, 0.8, 0.6. 0.4. and 0.3 kb, respectively. Influx of [ 3 H]uridine (10 uM) was measured after 5 days in NaCI (B) and choline chloride transport medium (C) as described in Protocol 8. Values are means - SEM of 10-12 oocytes. Each oocyte was microinjected with 50 ng mRNA. T, Total mRNA; H2O, oocytes injected with water. (Adapted from ref. 14.)

construction. Figure 2 shows a size-fractional ion of rat jejunal mRNA by sucrosegradient centrifugarion and the testing of individual fractions for uridine transport activity (14), Total rat jejunal mRNA induced the expression of uridine transport (uptake in mRNA-injected oocytes minus uptake in oocytes injected with water alone), and this flux was Na" -dependent (uptake in NaCI transport medium minus uptake in choline chloride medium). Peak activity in fraction 7 was enriched 5,8-fold compared with total mRNA and corresponded to a size range of 1.6 to 3.0 kb (median 2.3 kb). As described in Sections 4 and 5, this fraction was used in the expression-cloning of the pyrimidine-selective, concentrative (Na" -dependent) rat nucleoside transporter rCNTl (rat concentrative

SYLVIA Y. M. YAO ET AL.

nucleoside transporter 1) (15), the first identified member of the CNT family of membrane proteins. 3.4 Denaturing agarose gel electrophoresis of total RNA and mRNA RNA preparations should be monitored by denaturing agarose gel electrophoresis as described in Protocol 6. Good quality total RNA should contain prominent bands of 28S and 18S ribosomal RNA, and the intensity of the 28S band should be approximately double that of the 18S band. Small amounts of ribosomal RNA in the final mRNA preparation, such as that shown for the total mRNA in Figure 2, are acceptable and there should be minimal smearing below 0.5 kb. The purity of mRNA can also be checked by A260/A280 absorption (the ratio should be ~ 2).

4 Preparation of plasmid cDNA libraries suitable for in vitro transcription of RNA and expression in Xenopus oocytes Library preparation is a two-step process involving the synthesis of orientationspecific cDNA from size-selected poly(A)+ RNA, followed by ligation of the cDNA into a suitable plasmid expression vector and transformation and amplification in Escherichia coli. 4.1 cDNA synthesis from size-selected poly(A)+ RNA To be suitable for screening by in vitro transcription and expression in Xenopus oocytes, all the cDNAs in a library should have the same orientation (i.e. the library should be directional). If not, clones in the wrong orientation will not be detected and the resulting antisense RNAs may interfere with the expression of complementary-sense RNAs by antisense hybrid-depletion. It is therefore necessary to produce orientation-specific cDNA from the size-selected poly(A)+ RNA. This involves the synthesis of cDNA with different restriction sites at the 5' and 3' ends and is best carried out using one of the commercial kits designed for this purpose. One such kit is the RiboClone™ (Promega) reverse transcription cDNA synthesis system which uses an oligo(dT)-Xbal primer/EcoRl adapter combination to produce double-stranded cDNA with a 5' EcoRl terminus and a 3' Xbal terminus. 4.2 Construction of a Xenopus expression cDNA library General purpose cDNA libraries prepared from unfractionated mRNA may contain upwards of 107 primary recombinants with an average insert size of perhaps 1.5 kb. Many of these will be truncated at the 5'-terminus (due to incomplete reverse transcription) and be non-functional. The screening of such a library by expression-selection in Xenopus oocytes would be a daunting task. We can minimize the size of the library and increase the likelihood of success 58

THE XENOPUS OOCYTE EXPRESSION SYSTEM

by using size-selected mRNA as described in Section 3, and by subjecting the double-stranded cDNA product from Section 4.1 to a second round of sizefractionation using, for example, a Sephacryl S-500 HR chromatography column (Gibco BRL) with a cut-off of - 0,5 kb.

Protocol 6

Denaturing agarose gel electrophoresls Equipment and reagents • Submarine electrophoresis system (BioRad Mini Sub Cell*, or equivalent) • Enhanced laser densitometer (Pharmacia Ultroscan XL, or equivalent) • UV light box (Fisher Scientific transilluminator FBTV-816, or equivalent) • Camera (Polaroid MP-4. or equivalent) • 10 x Mops buffer: 200 mM Mops pH 7. 50 mM Na acetate, 10 mM EDTA (filtersterilized and stored in a dark bottle) • Loading buffer: 50% (v/v) glycerol, 1 mM EDTA, 0.5% (w/v) Bromophenot Blue • RBS 35 detergent (Pierce)

• RNAmolecularweight standards (0.36-9.49 kb, Novagen) • 0.l M NaOH • Formaldehyde • Deionized formamide: prepared by mixing 50 ml of formamide with 5 g of AG 501-X8 resin (Bio-Rad). Stir for 30 min at room temperature, then filter through Whatman No. 1 filter paper and store at -200C in l ml aliquots. • RNase-free water

• 1 mg[ml ethidium bromide (carcinogenic, handle with care)

Method 1. Clean the gel electrophoresis apparatus (gel tank, gel tray, comb) with RBS 35 detergent, then rinse thoroughly with sterile distilled water. Place the comb and gel tray in the gel tank and submerge with 0.1 M NaOH for 30 nun. Rinse thoroughly with sterile distilled water. 2. Dissolve 1% (w/v) agarose in 1 x Mops buffer by heating in a microwave. Cool the agarose solution to - 60°C, then add formaldehyde to give a final concentration of 2% (v/v). Pour the gel solution into the pre-cleaned gel apparatus and allow the gel to solidify in a fume hood. 3. Mix 5 ul of the RNA sample or RNA molecular weight standards with 3 ul of .10 x Mops buffer, 15 ul of deionized formamide, 5 (ul formaldehyde, and 2 ul RNase-free water. Add 1 ul of ethidium bromide (1 mg/ml) to each sample just before heatdenaturation. 4. Heat-denature the sample at 65 °C for 10 min, then chill on ice for 5 min. 5. Add 3 ul loading buffer to each sample, then load the samples into the gel wells and run the gel at 40 V for 5 h. 6. View and photograph the gel on the UV light box. Keep the negative for scanning by laser densitometry. Calculate the size range of each RNA sample by reference to the RNA molecular weight standards.

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The final ligation, transformation, and amplification steps needed to prepare the primary library are performed using standard procedures. To avoid growth competition between bacteria harbouring plasmids with different inserts, transformed bacteria should only be amplified as discrete colonies on solid medium (i.e. agar plates). A duplicate of the library can be made at this stage using nitrocellulose filter overlays to test the total primary library for functional activity (Section 5.2). The library should be stored at -70°C. In our cDNA cloning of rCNTl (15), the rat jejunal mRNA fraction that induced peak uridine transport activity in Figure 2 was reverse transcribed using the RiboClone™ (Promega) cDNA synthesis system as described in Section 4.1. Orientation-specific cDNAs > 0.5 kb (obtained by Sephacryl S-500 HR column chromatography) were ligated into the EcoRl and Xbal restriction enzyme sites of the plasmid expression vector pGEM-3Z (Promega) and transformed into Escherichia coli JM 109 to give a cDNA library containing 6800 primary recombinants. The properties of pGEM-3Z and other vectors suitable for Xenopus oocyte expression are discussed in Section 6.

5 Screening cDNA libraries by functional expression-selection in Xenopus oocytes cDNA cloning by functional expression-selection in Xenopus oocytes is a relatively labour-intensive method of library screening, but has important advantages over other methods such as PCR amplification, hybridization, or antibody screening. First, it eliminates the problem of false-positives and guarantees the isolation of functional, usually full-length clones with the desired transport characteristics. Second, it can be used for transporters for which no molecular information (N-terminal/internal amino acid sequence, known homologues) or antibodies are available. In addition to SGLT1 (2) and rCNTl (15), at least 14 mammalian transport proteins have been cloned by expression-selection in Xenopus oocytes (13). Functional expression-selection in Xenopus oocytes can also be used to clone cDNAs encoding transport accessory proteins. The mammalian amino acid and glucose transport regulator proteins' broad-specificity amino acid transporter (BAT), 4F2hc, and regulatory subunit 1 (RS1)(16, 17), for example, were identified and functionally characterized through their ability to activate cryptic Xenopus oocyte transporters. The selection of cDNAs encoding regulator proteins can be tailored to a particular heterologous transporter by coexpression of that protein along with the cDNA library that is being screened. Expression-cloning in Xenopus oocytes involves the progressive subdivision of the cDNA library until a single positive clone has been identified. The procedure described in Section 5.2 was used to isolate the rCNTl cDNA (15) and incorporates strategies to minimize the number of clones and pools of clones that need to be tested. Since cDNAs are identified through the functional activity of their RNA transcripts, we first describe the production of in vitro mRNA. 60

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5.1 In vitro synthesis of capped RNA transcript In vitro production of RNA transcript requires linearization of purified cDNA template with a restriction enzyme that cuts downstream (at the 3' end) of the insert. The starting plasmid DNA should be free from bacterial RNA or chromosomal DNA contamination and can be prepared using a plasmid DNA miniprep kit such as the Qiagen plasmid purification system or by standard CsCl centrifugation. After linearization, the cut DNA is recovered by phenol/chloroform extraction and ethanol precipitation. The insert is then transcribed with RNA polymerase in the presence of the m7GpppG cap. For optimal expression, the cap analogue and GTP should be added in a ratio of 4:1. Although inclusion of the cap analogue in the transcription reaction will reduce the amount of RNA obtained, it is required for the stability of synthetic RNAs in Xenopus oocytes (18). Following transcription, remaining template is removed by digestion with DNase I. The RNA is then recovered by phenol/chloroform extraction and ethanol precipitation and stored at a concentration of 1 ug/u1 in RNase-free water at -70°C. Commercial kits that give good yields of RNA (typically 20 ug RNA per 1 (ug DNA) and reliable expression in Xenopus oocytes include the MEGAscript™ and mMessage™ machine (Ambion) transcription systems. Published recipes suitable for oocyte work are also available (11). Agarose gel electrophoresis (Protocol 6) should be used to check the RNA preparation. The product should consist of a single species of RNA having an electrophoretic mobility that is consistent with the size predicted from the nucleotide sequence of the cDNA insert.

5.2 A functional expression protocol for the isolation of transporter-encoding clones The first step in library screening is to verify that the total primary library has functional activity. To do this, colonies from the duplicate of the primary library are pooled and used to make a plasmid miniprep, from which RNA transcript is then prepared and tested in oocytes. If the library has functional activity, we recommend using the following two-stage strategy to identify and isolate individual transporter-encoding cDNAs. First, aliquots of the primary library each corresponding to pools of approximately 500-1000 clones are plated and grown overnight on 150-mm, LB agar plates with ampicillin (100 (ug/ml). Cells from each master plate are transferred on to 132-mm, nitrocellulose filter overlays which in turn are placed cell-side up on fresh LB agar plates with ampicillin (100 ug/ml), and again incubated overnight to produce duplicate sets of colonies. The cells on each nitrocellulose filter overlay are then separately pooled and processed to prepare RNA transcript as described for the total library, while the master plates are stored at 4°C. In the screening of our rat jejunal cDNA library (6800 primary recombinants) for Na+-dependent nucleoside transport activity (15), we tested 20 pools each of approximately 700 clones, corresponding to a total of 14000 cDNAs, or approximately twice the number of primary recombinants in the starting library. Two pools were identified that increased the uptake of 61

SYLVIA Y. M. YAO ET AL.

uridine 8-fold above that of oocytes injected with RNA transcribed from the total library, and 140-fold above that of control oocytes injected with water alone (see figure 3). Colonies from the master plate of a positive pool are individually seeded into the wells of 96-well, flat-bottomed, microtitre plates to produce a grid system. Testing the rows and columns of the grid will then uniquely identify the clone(s) responsible for functional activity of the pool. To do this, cells from either a row or a column are grown together as separate colonies on an agar plate. Pooled DNA is then used to prepare an RNA transcript. If the master plate contains a single positive clone, then only one row and one column will show transport activity. The clone from the well at the intersection of that row and column can then be tested individually to confirm its identity. The single positive clone will typically show greater functional activity than the pool from which it was derived (see Figure 3). An alternative screening protocol is described in ref. 13.

Figure 3 cDNA cloning of Na+-dependent rCNTl from rat jejunum by expression-selection in Xenopus oocytes. Fraction 7 mRNA in Figure 2A was used to construct a size-selected directional cDNA library containing 6800 primary recombinants. Plasmid DNA from the total library and from 20 pools of ~ 700 clones was transcribed in vitro, microinjected into Xenopus oocytes (10 ng/oocyte, see Protocol 2) and tested after 3 days for Na+-dependent [3H]uridine transport activity (see Protocol 8). One of two positive pools (pool 15) was functionally screened as described in Section 5.2 to isolate clone pQQHl encoding full-length rCNTl. Values of 10 uM uridine uptake are means ± SEM of 10-12 oocytes. Open columns, uptake in NaCI transport medium; solid columns, uptake in choline chloride transport medium. Library, RNA transcript from the total library; H20, oocytes injected with water. Pools 16-19 were negative for uridine transport activity.

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6 Functional and molecular characterization of transporter-encoding cDNAs This section discusses plasmid vectors, radioisotope and electrophysiological transport techniques, protein detection, quantification of cell-surface expression, transporter topology, and structure-function studies.

6.1 Plasmid vectors for the expression of transporter-encoding cDNAs in Xenopus oocytes 6.1.1 pBluescript, pGEM-3Z Plasmid vectors for Xenopus oocyte studies should be multiple copy plasmids, have multiple cloning sites (MCS) with many unique enzyme restriction sites, and contain two RNA polymerase promoter sites so that both sense and antisense RNA can be produced. Examples of such vectors include pBluescript (Stratagene) and pGEM-3Z (Promega). We have used these vectors to successfully express nucleoside, amino acid and glucose transporters from species as diverse as mammals, Eptatretus stauti (a primitive marine vertebrate), and the nematode Caenorhabditis elegans. Transport proteins from higher plants have also been expressed in oocytes using these and similar vectors (11). 6.1.2 Oocyte expression vectors: pSP64T and pGEM-HE pBluescript and pGEM-3Z will be suitable for most purposes. Occasionally, however, 'difficult' proteins will be encountered which may benefit from the use of enhanced Xenopus oocyte expression vectors. In pSP64T (18) and pGEM-HE (19), for example, the protein coding region of cloned cDNAs is inserted between flanking 5'- and 3'-untranslated regions of the Xenopus p-globin gene. We have used pSP64T and pGEM-HE to express both lower eukaryote (Saccharomyces cerevisiae) and prokaryote (Escherichia colt) nucleoside transporters in Xenopus oocytes. As shown in Figure 4 for NupC (20), which is an Escherichia coli homologue of rCNTl, pSP64T produced substantially greater uridine influx than pBluescript (Loewen et al, unpublished). Since NupC is proton-dependent, transport activity was further enhanced by acidification of the external medium. Protocol 7 gives a procedure for subcloning a cDNA into pSP64T. Some other enhanced oocyte expression vectors are listed in ref. 21.

6.2 Transport assays in Xenopus oocytes using radioiabelied substrates Transport assays in Xenopus oocytes can be used to: • • • •

screen cDNA libraries for functional activity (see Section 5); verify the transport activity of cDNAs isolated or identified by other means; undertake functional characterization of cloned cDNAs; study structure/activity relationships. 63

SYLVIA Y. M. YAO ET AL

Protocol 7 Subcloning a cDNA Into the vector pSP64T Equipment and reagents • Vector pSP64T • Bgill restriction enzyme (New England Biolabs) • Bgin linker (New England Biolabs} • Calf intestinal alkaline phosphatase (CIP) (New England Biolabs)

DNA polymerase I (Klenow fragment) (New England Biolabs) T4 DNA ligase (Gibco BRL) Phenol-chloroform-isoamyl alcohol (25:24:1) dNTPs (10 mM}

Method 1. Digest pSP64T (see Figure 4) with BgHI for 2 h at 37°C in the buffer provided by the supplier, then add CIP (5 units/ug DNA) and incubate for 1 h at 37°C to remove the phosphate groups from each end of the cut. Extract the DNA with phenolchloroform and precipitate with ethanol (see Protocol 4, step 8). 2. Cut out the full-length cDNA to be subcloned from its original plasmid vector with the appropriate restriction enzyme(s), then treat with DNA polymerase I (Klenow fragment) in the presence of 2 mM dNTPs for 30 min at 37°C to blunt-end the cDNA. Extract and precipitate the cDNA as in step 1. 3. Add Bgtll linker to the blunt-ended cDNA using T4 DNA ligase according to the manufacturer's instructions. Extract and precipitate the cDNA as in step 1. 4. Digest the Bglll-linked cDNA with BgUI for 2 h at 37°C. Extract and precipitate the cDNAas instep 1. 5. Ligate the BgHi-cut cDNA into the BgHI site of the CIP-treated plasmid vector pSP64T using T4 DNA ligase according to the manufacturer's instructions. 6. Transform E. coli with the ligated product. 7. Select clones with the cDNA subcloned into pSP64T in the correct orientation by restriction mapping. Transport activity can be measured either using radioactively labelled permeants (see below), or by electro physiology (see Section 6,3). 6.2.1 Influx The basic flux assay in Protocol 8 is performed at 20°C on groups of 10-12 oocytes and is initiated by the addition of medium containing the appropriate radiolabelled substrate. After incubation. the extracellular label is removed by rapid ice-cold washes with isotope-free transport buffer. Individual cells are then dissolved in detergent for quantification of oocyte-as so dated radioactivity by liquid scintillation counting, RNA-injected oocytes are compared with oocytes injected with water alone to determine the transporter-mediated component of 64

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Figure 4 Expression of proton-dependent Escherichia colt NupC in Xenopus oocytes. NupC cDNA was PCR-amplified from Escherichia coli HB101 chromosomal DNA and ligated into the plasmicl vectors pGEM-3Z and pSP64T. Xenopus oocytes were microinjected with in vitro transcribed RNA (10 ng) or with water and tested for 10 uM [3H]uridine uptake after 5 days, either in standard NaCI transport medium at pH 7.5, or in medium acidified to pH 5.5 (see Protocols 2 and 8). Values are means J. SEM of 10-12 oocytes. The vector map of pSP64T is adapted from ref. 18. The construct NupC/pSP64T was prepared as described in Protocol 7.

Protocol 8 Radlotracer flux assay Equipment and reagents • 12 X 75 mm glass tubes • 100 ul pipettes • Shaker (New Brunswick GyrotaryG2,or equivalent) • Scintillation counter and vials (Beckman LS 6000 IC, or equivalent) • NaCl transport buffer: 100 mM NaCl. 2 mM KC1, 1 mM CaCl2,1 mM MgCl2,10 mM Hepes pH 7.5 • 5% (v/v) SDS

Choline chloride transport buffer: 100 mM choline chloride, 2 mM KC1.1 mM CaCl2. 1 mM MgCl 2. 10 mM Hepes pH 7.5 Incubation medium: NaCl or choline chloride transport buffer containing the appropriate concentration of unlabelled permeant, traced with 14Cl3H-labelled permeant at a specific activity of 1 uCi/ml ( 14 C)or2 uCl/ml(3H).

Scintillation fluid (Beckman Ready Safe liquid scintillation cocktail, or equivalent)

Method NaCl 1. Place 10-12 healthy Stage V/VI oocytes into a glass tube containing 200 transport buffer at room temperature. For assays to be performed in the absence of sodium, incubate the oocytes in choline chloride transport buffer at room temperature for 15 min before the flux assay. 2. Use a 100 ul pipette to remove most of the solution surrounding the oocytes.

SYLVIA Y. M. YAO ET AL. Protocol 8 conti

3. Add 200 ul of the incubation medium and incubate at room temperature with gentle shaking for the required time (typically 1 min-1 b). 4. After incubation, remove the majority of radiolabelled medium with a pipette and rapidly wash the oocytes five times with 1-2 ml aliquots of the ice-cold NaC1 transport buffer or choline chloride transport buffer. Remove the buffer between washes. 5. Transfer individual undamaged oocytes into scintillation vials, then dissolve in 0.5 ml 5%(w/v) SDS for 2 h with vigorous shaking. 6. Add 3 ml of the scintillation fluid and determine the intracellular radioactivity in the Beckman scintillation counter. Include standards containing 30 ul incubation medium, 0.5 ml 5% (w/v) SDS, and 3 ml scintillation fluid. and blanks containing SDS and scintillation fluid only. uptake (this will be greatest at substrate concentrations equal to or less than the anticipated apparent KM). Incubation periods for kinetic and other quantitative studies should be within the initial linear phase of the uptake curve to approximate zero-trans conditions and measure the initial rates of transport. Oocytes are large, and the initial rates of uptake (influx) are sustained for longer periods (typically 1 min-1 h) than can be achieved with membrane vesicles, bacteria, yeast, or cultured cells. Incubation times should be determined empirically for each transporter. The ice-cold washes can be completed on a group of 10-12 oocytes within 15 sec and are not usually associated with significant substrate loss from the cells. If desired, a transport inhibitor can be added to the wash solution, Radioisotopes used in transport experiments should be pure, since even trace amounts of isotopic contaminants (<2%) can give anomalous results under some conditions. Tritiated compounds are less stable than 14C-labelled derivatives under most circumstances and should be re-purified (e.g. by HPLC) on a regular basis. A SpeedVac system (Savant) can be used to remove [ 3 H]H 2 O from 3Hlabelled isotopes just before use. If possible, select a permeant that is not subject to extensive metabolism in oocytes. It is particularly important to avoid intracellular conversion to a more permeable derivative which might exit the cell. Protocol 8 will allow a wide variety of functional studies to be undertaken, including determination of the apparent Ky, and Vmax (by varying substrate concentration), inhibitor sensitivity (including a pre-incubation step, if necessaiy), substrate specificity (by inclusion of competing non-radioactive substrates in the incubation medium), and cation dependence (by altering the cation composition of the medium). If necessaiy, iso-osmolarity at different substrate or competitor concentrations can be maintained by reducing the concentration of NaCl in the medium. Care should be taken to ensure that high concentrations of substrate or competitor do not alter the pH of the medium. In substrate 66

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specificity experiments, it should be remembered that a demonstration of crossinhibition does not necessarily mean that the competitor is a substrate for the transporter. Adenosine, for example, is transported by rCNTl with a similar apparent KM for influx as uridine (and is therefore a good rCNTl inhibitor), but has a very much reduced Vmax (22). As an alternative to radioisotope uptake assays, the transportability of potential substrates identified in competition experiments can be investigated in trans-acceleration efflux studies (see Section 6.2.2). If the transporter is electrogenic, then current measurements can also be used to demonstrate substrate translocation (see Section 6.3.1). In studies of Na+-dependence, choline (or N-methyl-D-glucamine) can be used as a Na+substitute (see Figure 2). The external medium can be acidified if proton dependence is anticipated (see Figure 4). The transport regulators BAT and 4F2hc each induce multiple transport activities when expressed in Xenopus oocytes (23). Multiple transport activities should also be anticipated in cases where mRNA is expressed in oocytes. Variations in the transport assay, including the use of different substrates, crosscompetition, inhibitors, or Na+-dependence may be helpful in resolving parallel independent routes of permeant uptake. The quality of transport data that can be obtained with Xenopus oocytes is illustrated in Figure 5 for the human equilibrative (Na+-independent) nucleoside transporter hENTl (human equilibrative nucleoside transporter 1) (24). hENTl is structurally unrelated to rCNTl/NupC and was cloned from human placenta by a PCR strategy based on the N-terminus of the human erythrocytic transporter (24). It was the first representative of a second, newly recognized family of membrane transport proteins, the ENT family, and, unlike rCNTl, has a broad substrate selectivity for pyrimidine and purine nucleosides. hENTl does not transport nucleobases and is potently inhibited by nitrobenzylthioinosine (6-[ (4nitrobenzyl)thio]-9-p-D-ribofuranosylpurine; NBMPR) and the vasoactive drugs dipyridamole and dilazep. 6.2.2 Efflux Although it is more usual to study influx, efflux studies can also give important functional information on the kinetics, substrate selectivity, and exchange properties of membrane transport proteins. For example, Figure 6 shows that extracellular uridine causes the trans-acceleration of rCNTl -mediated efflux of uridine, while adenosine results in trans-inhibition (22). This is consistent with the low KM/low V,^ behaviour of adenosine as an rCNTl permeant (see Section 6.2.1), and illustrates the reduced mobility of the adenosine/transporter complex. To measure efflux, it is first necessary to pre-load oocytes with radioactive permeant. This can be achieved by incubating oocytes with an appropriate concentration of a non-metabolized radiolabelled permeant as described in Protocol 8. Alternatively, oocytes can be microinjected with radiolabelled substrate (25). Cells are washed free of extracellular label with ice-cold transport buffer and added in groups of 20 to 1 ml of fresh solution at 20 °C to initiate efflux (22). Small aliquots (5 (xl) are removed in duplicate at intervals and 67

SYLVIA Y. M. YAO ET AL.

Figure 5 Functional characterization of recombinant hENTl expressed in Xenopus oocytes. (A) Time-course of [3H]uridine (10 n-M) uptake by oocytes microinjected with hENTl RNA transcript (10 ng) or water (see Protocol 2) and tested after 3 days. (B) Shows hENTlmediated uridine influx (10 p-M) inhibited by a series of physiological nucleosides and anti-cancer nucleoside analogues. The hatched column represents the uptake of uridine in choline chloride transport medium. (C) Concentration-dependence of uridine influx; inset, the transporter mediated-component of influx (Influx in RNA-injected cells minus that in water-injected cells). (D) Inhibition of hENTl-mediated 10 uM uridine influx by 1 uM NBMPR, dilazep, and dipyridamole; inset, concentration-dependence of NBMPR inhibition. Transport was performed as described in Protocol 8. Values are means ± SEM of 10-12 oocytes. (Adapted from ref. 24.)

quantified for radioactivity. Use HPLC or TLC to ensure that the isotope leaving the cell is in the same molecular form as the original substrate. 6.3 Electrophysiological recordings in Xenopus oocytes Transporters which are electrogenic can also be studied using electrophysiological techniques. Since the movement of charge can be an ionized substrate (such as a cationic amino acid), a co-transported cation (e.g. Na + , H+, or K+) or anion (e.g. Cl-), or some combination of these, a wide range of transporters from different protein families are potentially amenable to electrophysiological recordings. Although transporters have unitary transport capacities (i.e. turnover numbers) that are slow (101--104/sec) compared to channels (^ 106/sec), expression 68

THE XENOPUS OOCYTE EXPRESSION SYSTEM

Figure 6 Trans-stimulation and trans-inhibition of rCNT1-mediated uridine efflux by extracellular uridine and adenosine. The efflux of uridine from oocytes pre-loaded with 10 uM [3H]uridine for 30 min was measured on groups of 20 oocytes suspended in NaCI transport buffer alone or in NaCI transport buffer containing 1 mM uridine or adenosine. Cells were microinjected with 10 ng rCNTl RNA transcript (see Protocol 2) and tested after 3 days. The composition of the transport buffer is described in Protocol 8. (Adapted from ref. 22.)

levels in Xenopus oocytes are sufficient to allow whole-cell recording using the two-microelectrode voltage clamp (21, 26, 27), or patch-clamp recording from detached oocyte macropatches (21, 26). 6.3.1 The two-microelectrode voltage clamp as an alternative to radioisotope transport assays The two-microelectrode voltage clamp in Xenopus oocytes is performed using methods standard for smaller cells. One intracellular microelectrode clamps the oocyte membrane to a predetermined potential (which can be varied), while a second microelectrode delivers current to maintain that potential. The current (in nA) needed to hold the desired membrane potential is the measured parameter. Oocytes are used with the vitelline membrane intact and are prepared as described for radioisotope flux studies. In its simplest form, the two-microelectrode voltage clamp can be used in place of radioisotope flux measurements (see Section 6.2) to investigate the steady-state kinetics, cation/anion coupling, or substrate specificity (including permeants not available in radioactive form). Repetitive measurements can be made on the same cell under different experimental conditions and at different membrane potentials so that, for example, complete voltage- or concentrationdependence curves can be generated from a single oocyte. We measure whole-cell oocyte membrane currents using a CA-1B oocyte clamp (Dagan). Microelectrodes are filled with 3 M KC1 with resistances from 1-2.5 MO. The CA-1B is interfaced to a computer via a Digidata 1200B A/D converter and controlled by Axoscope software (Axon Instruments). Current 69

SYLVIA Y. M. YAO ET AL.

signals are filtered at 20 Hz (four-pole bessel filter) and sampled at a sample interval of 50 msec. For data presentation the current signals are further filtered at 0.5 Hz using pCLAMP software (Axon Instruments). Oocytes are penetrated and the membrane potential is monitored for 15 min. If the membrane potential is unstable or less than -30 mV, these oocytes are not used. Rgure 7 shows that hCNTl (human concentrative nucleoside transporter 1) (28), the

Figure 7 Electrophysiology of the hCNTl-mediated transport of uridine and AZT. Uridine induced a reversible inward current in an oocyte microinjected with hCNTl RNA transcript, but not in the control oocyte injected with water alone. No current was seen when sodium in the transport medium was replaced by choline. An hCNTl-mediated sodium current was also seen with the pyrimidine nucleoside analogue AZT. Electrophysiological recordings were performed as described in Section 6.3.1. Oocytes were prepared as described in Protocol 2 and tested 3 days after microinjection with 10 ng hCNTl RNA transcript or water. The composition of the transport buffers is described in Protocol 8. The uridine and AZT experiments were performed on different oocytes. 70

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human homologue of rCNTl, functions as a Na+/nucleoside co-transport protein (i.e. mediates a uridine-dependent Na+-current) and transports the antiviral nucleoside drug 3'-azido-3'-deoxythymidine (AZT) (Smith et at, unpublished). 6.3.2 Other applications of the two-microelectrode voltage clamp The two-microelectrode voltage clamp also permits the analysis of pre steadystate currents and can be used in the standard whole-cell configuration and in the cut-open oocyte preparation (where both sides of the plasma membrane are accessible for perfusion) to derive rate constants for individual steps of the transport reaction cycle (29, 30). Analysis of transient currents also allows determination of transporter plasma membrane density (7, 31), and can be combined with transport assays in the same cell to calculate unitary transport capacity (see also Section 6.4.2). Changes in transporter membrane abundance and catalytic activity in response, for example, to regulation by protein kinases can be studied in conjunction with capacitance measurements to determine changes in cell-surface area due to endocytosis and exocytosis (31). In this way, it is possible to distinguish between regulation resulting from the direct modulation of transporter activity and regulation resulting from vesicle fusion with, or retrieval from, the plasma membrane. The equipment described in Section 6.3.1 will measure pre steady-state currents and can be used in the cut-open oocyte mode.

6.3.3 Patch-clamp recording from macropatches Currents in Xenopus oocytes can also be measured by patch-clamp recording from excised macropatches. Macropatches achieve a high degree of temporal resolution, but are in the inside-out configuration (i.e. the cytoplasmic and extracellular faces of the oocyte membrane are in contact with the bath and pipette solutions, respectively) and are best suited for studies which require primary access to the cytoplasmic membrane surface (32, 33).

6,4 Protein detection and quantification of cell-surface expression 6.4.1 Protein detection Recombinant protein expression in Xenopus oocytes can be detected immunologically or, if antibodies are not available, by radiolabelling with [35S]methionine or [3H]leucine (11). A simple procedure for the isolation of total oocyte membranes (plasma membrane + intracellular membranes) is given in Protocol 9. Detection of proteins in Western blots of total membranes is described in Protocol W and illustrated for hENTl in Figure 8 (Sundaram et al, unpublished).

6.4.2 Cell-surface expression Quantification of cell-surface content can be combined with Vmax values to determine unitary transport capacities (turnover numbers), thereby providing estimates of absolute catalytic activity. Analysis of cell-surface expression is also 71

SYLVIA Y. M, YAO ET AL

Protocol 9 Isolation of total oocyte membranes Equipment and reagents • Refrigerated centrifuge (Beckman GPR centrifuge and GH-3.7 rotor, or equivalent) • Ultracentrifuge {Beckman TL-100 and TLA100.3 rotor, or equivalent) • Lysis buffer: 7.5 mM Na2HP04.1 mM EDTA pH7.4

• PMSF

• 15 ml centrifuge tubes • 13 x 51 mm polycarbonate ultracentrifuge tube

Method 1. Mix 100 oocytes with 1 ml of ice-cold lysis buffer containing 0.1 mM PMSF in a 15 ml centrifuge tube. 2. Use a 1 mI pipette to break open the oocytes by pipetting up and down 20-30 times. Keep the tube on ice. 3. Centrifuge at about 500 g for 5 min at 4°C in the GPR centrifuge. 4. Transfer the supernatant into a polycarbonate uhracentrifuge tube (13 x 51 mm) and centrifuge at 16 000 g for 30 min at 4°C using the TL-100 ultracentrifuge. Wash the membrane pellet twice with 2 ml aliquots of ice-cotd lysis buffer containing 0.1 mM PMSF. 5. Resuspend the oocyte membrane pellet in lysis buffer without PMSF and store at -70°C.

Figure 8 Western blot of Xenopus oocyte total membranes showing expression of recombinant hENTl. Transport protein was detected in oocyte total membranes using a polyclonal antipeptide antibody raised against an epitope within a large intraceilular loop linking hENTl TMs 6 and 7. Lane 1. oocytes injected with hENTl RNA transcript. Lane 2, oocytes injected with water alone. Digestion with endoglycosidase F leads to a sharpening of the band and a shift to a lower apparent M, (not shown). Oocytes were prepared as described in Protocol 2 and tested 3 days after microinjection with 10 ng of hENTl RNA transcript or water. Preparation of total membranes and Western blot analysis were performed as described in Protocols 9 and 10.

72

THE XENOPUS OOCYTE EXPRESSION SYSTEM

useful in studies of transport regulation (to distinguish between changes in intrinsic catalytic activity and transporter trafficking), and in site-directed muragenesis (to differentiate between constructs with reduced activity and those with impaired targeting to the plasma membrane).

Protocol 10 Western blot analysis Equipment and reagents • Electrophoresis gel system (Bio-Rad MiniProtean II Cell, or equivalent) • Electroblotting system (Bio-Rad MiniTrans-Blot Transfer Cell, or equivalent) • Pre-stained SDS-PAGE standards (19900-102000 Da) (Bio-Rad, or equivalent) • Sample buffer: 125 mM Tris-HCl pH 6,8, 4.1% (w/v) SDS, 20% (v/v) glycerol, 0,001% (w/v) Bromophenol Blue, 2% (v/v) 2mercaptoethanol (add just before use)

Transfer buffer: 25 mM Tris-base, 192 mM glycine, 20% (v/v) methanol TBST buffer: 20 niM Tris-HCl pH 7.5, 137 mM NaCl, 0.1% (v/v) Tween-20 TBST-M: TBST containing 5% (w/v) non-fat dry milk powder (made up fresh) Hybond™-PVDF membrane (Amersham) ECL™ Western blotting system (Amersham) Primary and secondary antibodies

Method 1. Dilute the protein sample with the sample buffer (1:1), 2. Use a Bio-Rad minigel system to run a 12% SDS-PAGE (2-5 ug protein or 10 ul prestained markers per lane) according to the manufacturer's instructions. 3. Pre-treat the Hybond-PVDF membrane in methanol for 5 sec, sterile distilled water for 5 min, and transfer buffer for 10 min. 4. Electroblot the samples from the gel on to the treated PVDF membrane at 30 V overnight at 4°C using the Mini-Trans-Blot Transfer Cell according to the manufacturer's instructions. 5. Block the PVDF membrane with TBST-M buffer for 30 min at room temperature with gentle shaking. Repeat the blocking step with TBST-M buffer. 6. Seal the PVDF membrane in a plastic bag with 4 ml of TBST buffer containing the primary antibody at a dilution of between 1:500 and 1:2000. Incubate overnight at room temperature with gentle mixing, 7. Wash the membrane three times for 5 min each time in TBST with gentle mixing. 8. Incubate the membrane with 10 ml of TBST-M containing 3 \i.l of the secondary antibody at room temperature for 1 h with gentle mixing. 9. Wash the membrane three times for 5 min each time in TBST with gentle mixing. 10. Use the ECL Western blotting system for secondary antibody detection, following the procedures recommended by the manufacturer.

SYLVIA Y. M. YAO ET AL.

Calculation of unitary transport capacity requires the determination of the actual number of transporter molecules at the cell surface. This has been achieved for SGLT1 using freeze-fracture electron microscopy to directly visualize the expressed transporter particles in the plasma membrane lipid bilayer (for SGLT1 there are ~ 5000/n,m2 or ~ 1.5 x 10u/oocyte) (34). Quantitative estimates of membrane abundance can also be provided by electrophysiology (see Section 6.3.1). Most other applications require only the determination of relative cellsurface abundance. Methods that rely on oocyte membrane fractionation are potentially unreliable because of possible contamination of the plasma membrane preparation with intracellular membranes (which may contain the majority of expressed protein). A better strategy (35) is to label plasma membrane proteins with membrane-impermeant biotin derivatives such as sulfosuccinimidobiotin (sulfo-NHS-biotin; Pierce). Total membranes (prepared as described in Protocol 9) are then solubilized, immunoprecipitated with a transporter-specific antibody (e.g. an antipeptide antibody or an antibody against an engineered epitope tag such as myc), and probed by Western analysis (see Protocol 10) with streptavidin coupled to horseradish peroxidase (Amersham).

6.5 Analysis of transporter topology and structure-function relationships Many approaches have been developed to investigate the molecular architecture of transport proteins and to identify structural domains and individual amino acid residues of functional importance. In this section, we describe some representative strategies which have been used in, or can be applied to, the Xenopus oocyte expression system. Transport proteins have multiple transmembrane segments (TMs) and typically contain several consensus sites for N-linked glycosylation. By combining Protocols 9 and 10 with endoglycosidase-F digestion and site-directed mutagenesis, it is possible to determine which of these sites are glycosylated. This, in turn, identifies extracellular regions of the transporter and can be helpful in formulating models for the overall arrangement of the protein in the membrane. In contrast to predictions from hydropathy analyses (15), for example, rCNTl is glycosylated at the C-terminus, which must therefore be extracellular (36). Potential roles of glycosylation in transporter function can be assessed by transport comparisons between the wild-type and aglyco forms of the protein. Topology models can be further refined using glycosylation-scanning mutagenesis to systematically introduce new potential sites of glycosylation into putative exofacial and endofacial loops of the aglyco form of the protein. As an alternative, for example, the factor Xa cleavage site can be introduced into predicted linker regions (37). It is also possible to engineer a transporter mutant devoid of Cys residues, and to use such a construct in conjunction with cysteinescanning mutagenesis and cysteine-directed protein chemistry to provide more 74

THE XENOPUS OOCYTE EXPRESSION SYSTEM

specific information on TM helix topology and packing—to identify pore-lining residues and to study substrate-induced changes in transporter conformation (see, for example, refs 38-43). Structural features involved in particular transport or regulatory functions can be identified in Xenopus oocytes by chimaeric exchanges between appropriate transporter isoforms, followed by site-directed mutagenesis to pinpoint the amino acid residues involved. For example, Figure 9 shows a series of exchanges between hENTl and its rat homologue rENTl (rat equilibrative nucleoside transporter 1) to identify TMs 3-6 as the region of hENTl involved in the binding of dipyridamole, rENTl being dipyridamole-insensitive (44). Xenopus oocyte expression of truncated versions of transporters can provide information on the minimum functional unit (45). Complementation studies with coexpressed fragments of transporters are also feasible in oocytes (46, 47).

Figure 9 Inhibition of hENT1/rENT1 chimaeras by NBMPR and dipyridamole. (A) Shows 10 uM [3H]uridine influx (± 1 H.M inhibitor) expressed as a percentage of the uninhibited flux for each transporter. Oocytes were prepared as described in Protocol 2 and tested 3 days after microinjection with 10 ng of hENTl RNA transcript or water as described in Protocol 8. Values are means ± SEM of 10-12 oocytes. (B) A schematic representation of the chimaeras. hENTl and rENTl both have 11 putative TMs. The number of amino acid residues in each construct is indicated. (Adapted from ref. 44.)

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SYLVIA Y. M. YAO ET AL.

7 Conclusions The Xenopus oocyte system is uniquely suited to the expression-cloning and functional characterization of plasma membrane transport proteins. Primarily used to study mammalian transporters, Xenopus oocytes are also capable of expressing transport proteins from a wide range of other eukaryotic species, including plants, demonstrating the system's general utility as a heterologous expression system. As we have also shown in this chapter, prokaryotic transporters can also be expressed successfully in Xenopus oocytes, allowing functional comparisons between eukaryotic and prokaryotic transporters expressed into the same membrane environment. When combined with recombinant DNA techniques, Xenopus oocyte expression provides a versatile tool for investigating transporter topology and structure-function relationships.

Acknowledgements We are grateful to Dr C. M. Harvey who introduced us to the Xenopus oocyte expression system, to Dr Q. Q,. Huang who performed the expression-cloning of rCNTl, and to our research collaborators Drs S. A. Baldwin and E. Karpinski. We thank Dr E. J. Sanders for taking the photographs shown in Figure 1. Work described in this article was supported by grants from the Canadian MRC, NCIC, NSERC, and CANFAR. JDY is a medical scientist of the Alberta Heritage Foundation for Medical Research.

References 1. Gurdon, J. B., Lane, C. D., Woodland, H. R., and Marbaix, G. (1971). Nature, 233, 177. 2. Hediger, M. A., Coady, M. J., Ikeda, T. S., and Wright, E. M. (1987). Nature, 230, 379. 3. Ausubel, I. and Frederick, M. (ed.) (1998). Current protocols in molecular biology. CD-ROM version. Wiley, New York. 4. Ceriotti, A. and Colmam, A. (1995). In In vitro transcription and translation protocols (ed. M. J. Tymms), p. 151. Humana Press, Totowa, NJ. 5. Dumont, J. N. (1972). J. Morphol, 136, 153. 6. Methfessel, C., Witzemann, V., Takahashi, T., Mishina, M., Numa, S., and Sakmann, B. (1986). Pflugers Archiv, 407, 577. 7. Quick, M. W., Corey, J. L, Davidson, N., and Lester, H. A. (1997). J. Neurosci., 17, 2967. 8. Goldin, A. L. (1992). In Methods in enzymology, Vol. 207 (ed. B. Rudy and L. E. Iverson), p. 266. Academic Press, London. 9. Coady, M. J., Jalal, F., Chen, X., Lemay, G., Berteloot, A, and Lapointe, J.-Y. (1994). FEBS Lett., 356, 174. 10. Yang, X. C., Karschin, A, Labarca, C., Elroy-Stein, O., Moss, B., Davidson, N., and Lester H. A. (1991). FASEB J., 5, 2209. 11. Theodoulou, F. L. and Miller, A. J. (1995). AM. Biotech., 3, 101. 12. Tarnuzzer, R. W., Campa, M. J., Qian, N. X., Engelsberg, E., and Kilberg, M. S. (1990). J. Bid. Chem., 265, 12914. 13. Romero, M. F., Kanai, Y., Gunshin, H, and Hediger, M. A. (1998). In Methods in enzymology, Vol. 296 (ed. S. G. Amara), p. 17. Academic Press, London.

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THE XENOPUS OOCYTE EXPRESSION SYSTEM 14. Huang, Q. Q., Harvey, C. M., Paterson, A. R., Cass, C. E., and Young, J. D. (1993). J. Biol. Chem., 268, 20613. 15. Huang, Q. Q.. Yao, S. Y. M., Ritzel, M. W. L, Paterson, A. R., Cass, C. E., and Young, J. D. (1994).J. Biol. Chem., 269, 17757. 16. Yao, S. Y. M., Muzyka, W. R., Elliott, J. R, Cheeseman, C. I., and Young, J. D. (1998). ;. Biochem., 330, 745. 17. Veyhl, M., Spangenberg, J., Puschel, B., Poppe, R., Dekel, C., Fritzsch, G., Haase, W, and Koepsell, H. (1993). J. Biol. Chem., 268, 25041. 18. Kreig, P. A. and Melton, D. A. (1984). Nucleic Adds Res., 12, 7057. 19. Liman, E. R., Tytgat, J., and Hess, P. (1992). Neuron, 9, 861. 20. Craig, J. E., Zhang, Y., and Gallagher, M. P. (1994). MoL Microbiol, 11, 1159. 21. Mager, S., Cao, Y., and Lester, H. A. (1998). In Methods in enzymology, Vol. 296 (ed. S. G. Amara), p. 551. Academic Press, London. 22. Yao, S. Y. M., Ng, A. M. L., Ritzel, M. W. L., Gati, W. P., Cass, C. E., and Young, J. D. (1996). MoJ. Pharmacol., 50, 1529. 23. Yao, S. Y. M., Muzyka, W. R., Cass, C. E., Cheeseman, C. I., and Young, J. D. (1998). Biochem. Cell Biol, 76, 859. 24. Griffiths, M., Beaumont, N., Yao, S. Y. M., Sundaram, M., Boumah, C. E., Davies, A., Kwong, F. Y., Coe, L, Cass, C. E., Young, J. D., and Baldwin, S. A. (1997). Nature Med., 3, 89. 25. Closs, E. I., Albritton, L. M., Kim, J. W., and Cunningham, J. M.(1993).J. Biol. Chem., 268, 7538. 26. Stuhmer, W. (1992). In Methods in enzymology. Vol. 207 (ed. B. Rudy and L. E. Iverson), p. 319. Academic Press, London. 27. Busch, A. E., Waldegger, S., Murer, H., and Lang, F. (1996). Nephron, 72, 1. 28. Ritzel, M. W. L., Yao, S. Y. M., Huang, M. Y., Elliott, J. F., Cass, C. E., and Young, J. D. (1997). Am.]. Physiol, 272, C707. 29. Parent, L, Supplisson, S., Loo, D. D. F., and Wright, E. M. (1992). J. Membr. Biol, 125, 49. 30. Chen, X. Z., Coady, M. J., and Lapointe, J. V. (1996). Biophys. J., 71, 2544. 31. Hirsch, J. R., Loo, D. D. F., and Wright, E. M. (1996). J. Biol. Chem., 271, 14740. 32. Matsuoka, S., Nicoll, D. A., Hryshko, L. V., Levitsky, D. O., Weis, J. N., and Philipson, K. K. (1996).J. Gen. Physiol., 105, 403. 33. Lu, C.-C, Kabakov, A., Markin, V. S., Mager, S., Frazier, A., and Hilgemann, D. W. (1996). Proc. Natl Acad. Sci. USA, 92, 11220. 34. Zampighi, G. A., Kreman, M., Boorer, K. J., Loo, D. D. F., Bezanilla, F., Chandy, G., Hall, J. E., and Wright, E. M. (1995). J. Membr. Biol, 148, 65. 35. Chillaron, J., Estevez, R., Samarzija, L, Waldegger, S., Testar, X., Lang, F., Zorzano, A., Busch, A., and Palacin, M. (1997).;. Biol. Chem., 272, 9543. 36. Hamilton, S. R., Yao, S. Y. M., Ingram, J., Henderson, P. J. F., Gallagher, M. P., Cass, C. E., Young, J. D., and Baldwin, S. A. (1997).J. Physiol., 499, 50P. 37. Preston, G. M., Jung, J. S., Guggino, W. B., and Agre, P. (1994)J. Biol. Chem., 269, 1668. 38. Yan, R. T. and Maloney, P. C. (1995). Proc. NatlAcad. Sci. USA, 92, 5973. 39. Akabas, M. H., Cheung, M., and Guinamard, R. (1997). J. Bioenerg. Biomembr., 29, 453. 40. Wu, J., Voss, J., Hubbell, W. L., and Kaback, H. R. (1996). Proc. Natl Acad. Sci. USA, 93, 10123. 41. Tang, X. B., Fujinaga, J., Kopito, R., and Casey, J. R. (1998). J. Biol. Chem., 273, 22545. 42. Lo, B. and Silverman, M. (1998)J. Biol. Chem., 273, 29341. 43. Loo, D. D. R, Hirayama, B. A., Gallardo, E. M., Larn, J. T., Turks, E., and Wright, E. M. (1998). Proc. NatlAcad. Sci. USA, 95, 7789. 44. Sundaram, M., Yao, S. Y. M., Ng, A. M. L, Griffiths, M., Cass, C. E., Baldwin, S. A., and Young, J. D.(1998).J. Biol. Chem., 273, 21519.

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Chapter 4 Expression of foreign transport proteins in yeast N. Sauer and J. Stolz Lehrstuhl Botanik II, Universita't Erlangen, Staudstrasse 5, D-91058, Germany

1 Introduction Over the last few years we have learned from sequencing projects for various organisms that up to 5% of all nuclear genes encode putative transport proteins, emphasizing the importance of these proteins for compartmentation and transmembrane fluxes. The number of cloned transporter genes is increasing exponentially, and the functional analysis of these transporters is an inalienable requirement for understanding their specific functions and their influence on the underlying physiological processes. For that reason, expression of transporter genes or cDNAs in heterologous expression systems has become increasingly important, and expression in yeast has, in particular, turned out to be extremely useful. Besides functional analysis, which includes determination of substrate specificities, ion and pH-dependences, KM values, electrochemical properties, and energy-dependences, expression in yeast has also been the basis for the biochemical characterization of recombinant transport proteins. Numerous well-characterized yeast promoters and yeast mutants represent excellent tools for the large-scale preparation of transport proteins and also for the identification of transporter genes by complementation analyses. The aim of this chapter is to point out the opportunities offered by yeast expression systems, to present an overview of the available literature on transporter gene expression in yeast, and to summarize techniques of basic interest for transport physiologists, who want to use yeast cells as an expression system. Successfully used expression vectors and strains will be listed and benchtop protocols will be provided.

2 Expression of foreign genes in yeast: an overview 2.1 Yeast vs. other expression systems Yeast cells can be grown rapidly on simple media to very high densities. Easy-touse transformation protocols with vectors that can be propagated both in yeast 79

N. SAUER AND J. STOLZ

and Escherichia coli allow the rapid production of large numbers of transformants. Yeast cells possess the machinery for processing eukaryotic proteins and use basically the same mechanisms for membrane insertion and targeting as other eukaryotes. If necessary, foreign genes can easily be incorporated into the nuclear genome due to an extremely efficient in vivo homologous recombination process. Together, these technical advantages and the extremely advanced genetics make yeast cells the number one experimental organisms for the expression of transport proteins from plants and other eukaryotes (1-3). Of course there are other expression systems that have or could have advantages for very specific applications. For example, Xenopus laevis oocytes (see Chapter 3) are very useful for electrophysiological studies (4-7), although transporter gene expression in Xenopus is transient and technically demanding, and recent publications have shown that yeast cells can be used for electrophysiological studies as well (8). Bacterial expression systems can be used for large-scale protein production (9) (see Chapter 6). However, expression of foreign transporter genes in E. coli is often toxic to the cells (10). Moreover, when expressed under the control of strong promoters even the products of endogenous bacterial transporter genes tend to aggregate (11). 2.2 Expression of transporters in Saccharomyces and non-Saccharo/nyces yeasts Besides Saccharomyces cerevisiae several other yeast strains, such as Schizosaccha.romyces pombe, Pichia pastoris, Hansenula polymorpha, Klyveromyces lactis, or Yarrowia lipolytica have been used for foreign gene expression in the past (reviewed in ref. 1), and, in some cases, genes of membrane proteins were also successfully expressed (see below). So far, however, only S. cerevisiae (12-26) and S. pombe (27-33) have been used for the expression of transport proteins. P. pastoris, although having clear advantages with respect to the yield in terms of cell mass or expression levels (34), has so far not been used for expression of transporters. The methods described in this chapter will therefore focus on expression in S. cerevisiae and S. pombe. 2.3 Brief history of heterologous transporter gene expression in yeast The first membrane protein that was successfully expressed in S. cerevisiae was the a-subunit of the acetylcholine receptor from Torpedo californica (35). The functional expression in S. pombe of the gene for the archaebacterial protein bacterio-opsin (36), built up hope that this might also work for eukaryotic transport proteins. The first eukaryotic transporter to be characterized by heterologous expression in yeast was the CkHUP1 gene from the unicellular green alga Chlorella kessleri (37), encoding a plasma membrane-localized monosaccharide-H+ symporter. In the following years plant transporters for various substrates were expressed both in S. pombe and S. cerevisiae. The general applicability of yeast for 80

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

the expression-cloning of plant genes was published in 1992 (38), when an Arabidopsis thaliana cDNA library in an S. cerevisiae expression vector was used for complementation. In the same year the first plant sucrose transporter was isolated by complementation of a genetically engineered S. cerevisiae mutant (17). Since then expression of single cDNAs or cDNA libraries in yeast has become a widely used tool for the isolation, identification, and characterization of transport proteins.

2.4 Expression of genes encoding transport proteins from different membranes As mentioned above, the first foreign transporter gene expressed in yeast coded for a plasma membrane sugar transporter. Meanwhile transporters, channels, or pumps for many different substrates, such as monosaccharides (14), amino acids (15, 16), disaccharides (17-19), NH4+ (20), Ca2+ (23), H+ (25), K+ (12, 13), oligopeptides (22), sulfate (26), and H2O (24), have been expressed successfully in yeast. In addition, a human multidrug resistance-associated protein has been characterized by functional expression in S. cerevisiae (21). Many of these proteins are putatively localized in the plasma membrane of their 'home organism' and several have been shown to be indeed plasma membrane proteins. Most of these transport proteins are also targeted to the plasma membrane in yeast, but there are exceptions. A plant plasma membrane H+-ATPase, for example, accumulates in the endoplasmic reticulum of transgenic yeast cells (25). In some cases, however, the accumulation of plasma membrane proteins in internal yeast membranes occurs on purpose as a desired consequence of the selective choice of a yeast strain. Mutants in the secretory pathway have been used to accumulate the human red-cell water channel CHIP28 in post-Golgi vesicles of S. cerevisiae (24, 39). The purified vesicles were then used for physiological characterization and purification of functionally active CHIP28 protein (24, 40). Heterologous expression in yeast has also been used for the expression of non-plasma membrane transporters. Several genes encoding transporters from the inner chloroplast membrane were expressed in S. pombe (31-33). These and other chloroplast transporters are most likely targeted to the yeast mitochondria. The recombinant proteins have been purified after solubilization and have been used for functional analyses after reconstitution into lipid vesicles.

3 Expression in Saccharomyces cerevisiae Various parameters have to be taken into consideration for the optimal expression of a transporter gene in yeast. The choice of the right DNA fragment will be as important for successful expression as the choice of the right yeast, the proper strain, or the ideal vector. Several of these parameters depend, at least in part, on the question that has to be answered. 81

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3.1 Choice of DNA to be expressed In S. ivirvisiae introns are present in approximately 4% of all genes. To avoid problems potentially associated with splicing, heterologous genes should be tree of intron sequences, Intro n-less gen omit: DNA and cDNAs are perfectly suited tor expression in yeast. If cDNAs with different lengths of 5'-untranslated sequences are available, the cDNA with the shortest 5'-scquences should be used for the first trials. As most cDNAs cany J:V»RI or Notl ends introduced as linker sequences, yeast expression vectors with unique EcoRI or Noll cloning sites are ideal for constructing expression plasmids.

Protocol 1 Media for Saccharomyces

cerevisiae

A. YPD-medlum (complete medium) • 1% Yeast extract

• 2% Glucose

• 2% Peptone

B. YNB-medium (selective medium) • 0.67% Yeast Nitrogen Base without amino acids (Difco)

• 2% Glucose

1. Autoclave.

2. Add all supplements your strain needs for growth on selective medium except the one used for selection (compare genotypes of strains in Table I).

C. CAA-medlum (selective medium for URA3 and TRP1 plasmids) • 0.67% Yeast Nitrogen Base without amino acids (Difco)

• 2% Glucose • 1% casamino acids

B. Agar plates 1. For agar plates add 2% agar to YPD-, YNB-, or CAA-medmm, 2. Add supplements to YNB-medium at 50°C before making plates.

E. Supplements (50 x concentrated) 1. Prepare stock solutions according to your strain's requirements. 2. Prepare L-leucine, L-Jysine, L-histidine and uracil as 1 mg/ml stocks and autoclave." 3. Dissolve L-tryptophan (1 mg/ml) by heating to 60-70°C, filter-sterilize, and store in the refrigerator. 4. Prepare adenine as a 1.5 mg/ml stock and autoclaved." " Will go into solution during autoclaving!

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Protein N-glycosylation in yeast, as in plant or mammalian cells, occurs at Asn-X-Ser and Asn-X-Thr sequons (where X may be any of the standard amino acids except proline). Putative sites for N-glycosylation may therefore be recognized by the yeast glycosylation machinery; however, a clear prediction is not possible (1). If a recombinant protein is glycosylated, both the size and composition of the glyco-portion may differ drastically from that in the original protein. In particular, hyperglycosylation, which can occur due to a lack of Golgi mannosidases in yeast, may influence both the function and immunoreactivity of recombinant proteins. To avoid undesired N-glycosylation, site-directed mutagenesis may be employed to change the amino acid sequence accordingly (usually to Gln-X-Ser or Gln-X-Thr). Alternatively, expression in yeast mutants defective in mannan biosynthesis can be tried (1). As in other eukaryotic cells, 0-linked glycosylation can occur at the hydroxyl groups of serine and threonine residues, but a consensus sequence for modification is not readily apparent.

3.2 Host strains 3.2.1 Strains for functional analyses If the function of an already cloned cDNA has to be characterized by heterologous expression in yeast, it is important to choose a strain with little or ideally zero inherent transport activity for the investigated substrate. This facilitates transport studies with radioactive tracers, especially if the expression of the foreign transporter gene is low. For many substrates yeast mutants defective in inherent transporter genes are available (see below). Frequently, however, it is sufficient to grow yeast cells under defined conditions, e.g. the proper carbon source (such as glucose), where other endogenous transporter genes are not expressed (e.g. genes encoding transporters for galactose or maltose). A strain of S. cerevisiae has recently been generated where seven known genes for yeast monosaccharide transporters have been deleted (41). This strain is unable to grow on glucose concentrations up to 5% and is likely to be defective in all metabolically relevant glucose transporters. 3.2.2 Strains for cloning transport proteins by complementation Zero or very little background transport activity due to an inherent yeast mutation is also necessary for cloning new transporters by complementation. For example, S. cerevisiae mutants defective in genes encoding K+ channels can easily be transformed with expression libraries, and strains harbouring vectors with foreign K+ channel cDNAs can be identified by selection for growth on low K+ concentration media (12, 13). Examples of the complementation cloning of several plant transporters are described below.

(i) Sucrose transporters A highly efficient strategy has been used for identifying plant sucrose carriers by expression-cloning in yeast. The yeast strain SUSY7 (17) carries mutations 83

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preventing maltose or sucrose uptake from the medium and extracellular sucrose hydrolysis by the secreted cell-wall invertase. In addition, the gene of the potato sucrose synthase has been integrated into the genome. Sucrose synthase can split intracellular sucrose into UDP-glucose and fructose and can provide monosaccharides for metabolism. SUSY7 grows poorly on media containing sucrose as the sole carbon source, but normal growth is restored upon expression of sucrose transporter genes. (ii) Amino acid transporters Many plant amino acid transport proteins have recently been identified by complementation-cloning in S. cerevisiae (42). Strains that have been employed in these experiments are listed in Table 1. Most yeast strains can use proline as their sole nitrogen source. Strain 22574d is defective in the uptake of this amino acid due to multiple mutations. Selection for transformants expressing proline transporters has to be carried out in a medium containing proline as the sole nitrogen source. Another approach is to use a yeast strain that is auxotrophic for a certain amino acid (e.g. JT16 for histidine) due to a mutation in a biosynthetic gene Table 1 S. cerevisiae strains used for expression of foreign transporter genes Strain (Reference) Genotype

Used for (Reference)

SEY2102 (48)

Mate, ura3-52, /eu2-3,112, Ws4-519,suc2-D9, gal2

functional analysis of disaccharide transporters (19)

RS453

Mata, acte2-l, trpl-1, canl-100, leu2-3,112, his3-l, ura3-52

functional analysis of mono- and disaccharide transporters (14, 20, 49)

RE700A (41)

Mata, hxtlD::HIS3::Dhxt4, hxt5::LEU2, hxt2D::HIS3, hxt3D::LEU2::DhxtS, hxt7::HIS3, ura3-52, MAL2, SUC2, GAL, MEL

functional analysis of a monosaccharide transporter (57)

DBY2617 (50)

Mata, his4-539, lys2-801, ura3-52,suc2-438

functional analysis of disaccharide transporters (19)

SUSY7 (17)

suc2::URA3, ma/0, trp1, LEU2::sucrose synthase

functional characterization (51) and expression cloning of sucrose transporters (17)

22574d (52)

MATa, ura3-1 gap1-1, put4-l, uga4-l

complementation cloning of a plant amino acid transporter (15)

JT16 (53)

MATa, hip1-614, his4-401, ura3-52, ino1, can1

complementation cloning of a plant amino acid transporter (54)

PLAS23-4B (43) MATa, shr3-23, ura3-52, Ws4A29 defective in targeting of yeast amino acid transporters:complementation cloning of plant amino acid transporters (44) SY1 (46)

MATa, ura3-52, leu2-3,112, his4-619, sec6-4, GAL

C13-ABYS-86 (55) MATa, pral-1, prb1-1, prcl-1, cps1-3, ura3A5, Ieu2-3,112, his

84

ts secretory mutant, accumulates postGolgi secretory vesicles (39) protease deficient, useful for protein purification (56)

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

(his4) and, in addition, unable to take up histidine due to mutations in two transporter genes (hip1 and can1). Selection is carried out in a medium with ammonia and limiting concentrations of histidine. Another yeast strain used for cloning amino acid transporter genes is PLAS23-4B (see Table 1). This strain carries a mutation in shr3 causing a defect in the targeting of endogenous amino acid transporters to the plasma membrane (43). PLAS23-4B grows poorly on media containing proline as the nitrogen source plus limiting concentrations of histidine. Plant transporters for proline and a transporter with broad substrate specificity for different amino acids have been identified by their ability to suppress the mutant phenotype of the shr3 strain (44). A drawback of this approach could be that some heterologous transporters may also depend on SHR3 for correct targeting and may thus not be identified in an shr3 mutant yeast. 3.2.3 Protease-deficient strains S. cerevisiae contains a large number of proteases and the vacuolar proteases, especially, may cause problems during protein purification and can lead to artefacts in transporter activity or protein structure. The use of strains with defective alleles of these vacuolar proteases reduces the probability of protein degradation. For protein isolation from other yeast strains, but also in combination with protease deficient strains, a large selection of protease inhibitors can be used (see ref. 45 and Protocol 3). In growing yeast, the activities of most proteases increase when the cells approach stationary phase. Therefore working with log-phase cells is more favourable for biochemical analyses. For an extensive discussion of the protease problem in S. cerevisiae see ref. 45. 3.2.4 Strains defective in secretion (sec strains) A promising system for the expression of plasma membrane proteins in S. cerevisiae has been developed by Slayman and colleagues (46). It exploits a temperature-sensitive, secretory mutant, which accumulates post-Golgi vesicles that can no longer fuse with the plasma membrane after a shift to the nonpermissive temperature. For foreign gene expression a host strain with a sec6-4 (e.g. SY1) or a seel mutation and a plasmid with a galactose inducible promoter are used. After a switch from 30°C to 37°C and from glucose medium to galactose medium, gene expression is turned on and the gene product will accumulate. Secretory vesicles containing the heterologous transporter may be purified from a crude cell lysate (47). This strategy prevents the heterologous protein from being present at the plasma membrane and is therefore especially useful for potentially toxic proteins. Using the sec4-6 mutation, the human water channel CHIP28 (24) and the mouse P-glycoprotein (39) have been functionally expressed in S. cerevisiae.

85

N. SAUER AND J. STOLZ

3.3 Expression vectors 3.3.1 Type of vector Episomal vectors, that carry the origin of replication from the yeast 2u plasmid and marker genes for selection in S. cerevisiae and E. coli, are the first choice for high expression levels of a foreign gene. These vectors are maintained at 10-40 copies per cell. Vectors using centromeric sequences (CEN) plus yeast autonomous replication sequences (ARS) for replication, replicate only once per cell cycle and are stably maintained at 1 copy per cell. Integrative vectors carry no sequences for replication and have to integrate into the genome to be propagated. Typically only one copy is present per cell, and expression levels of foreign genes are therefore usually low, even if transcribed from strong promoters. Because overproduction of a foreign protein could be toxic for the host cells, the use of tightly regulated and inducible promoters will separate the growth phase of the cells from the production phase of the protein. Thus these vectors may have advantages over constitutive promoters for the production of potentially toxic proteins. Some vectors with inducible promoters are presented in Table 2. 3.3.2 5'- and 3'-regulatory sequences Several prokaryotic and eukaryotic promoters exhibit substantial transcriptional activity in yeast. High expression levels, however, are usually obtained with strong yeast promoter sequences, such as the promoter of the plasma membrane ATPase gene (PMA1) or promoters of genes for glycolytic enzymes in S. cerevisiae. In S. pombe strong expression is obtained with the promoter of the alcohol dehydrogenase gene (adhl). Frequently, the role of terminator sequences for the optimal expression of foreign genes or cDNAs is underestimated. As in other organisms the terminator can strongly influence the length and stability of an mRNA molecule. It has been shown that inefficient transcription termination results in low transformation efficiencies and correlates with reduced expression of a reporter gene (58). The use of an expression vector with a yeast promoter/yeast terminator box, interrupted by one or more unique restriction sites for cDNA insertion is strongly recommended. 3.3.3 Auxotrophic markers Mostly haploid strains of S. cerevisiae (designated MATa or MATa, according to the mating type) are employed for heterologous gene expression. Many strains have one or several nutritional requirements that can be used as auxotrophic selection markers (denoted by lowercase letters in S. cerevisiae genetic nomenclature, e.g. leu2-3, ura3-52, his3-l, trpl-1, for a mutation in a leucine, uracil, histidine, or tryptophan biosynthesis gene). Plasmid vectors that carry one of the corresponding wild-type genes (denoted by using uppercase letters for 86

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

dominant alleles, e.g. LEU2, URA3, HISS, TRP1) may be used for selection in such strains. Selection for the presence of the plasmid has to be performed in synthetic minimal media. Variants of the LEU2 and URA3 markers carrying promoters with diminished activity (LEU2-d or URA3-d) are available. These vectors can complement auxotrophic mutations only at very high copy numbers (obtained during selection) and thus will result in increased expression levels of the trans gene. Among the episomal vectors containing the wild-type markers, URA3 and TRP1 vectors have the advantage that transformants can be grown in minimal media enriched with hydrolysed casein (casamino acids; CAA-medium in Protocol 1), which is a source of most amino acids but not of tryptophan and not of uracil. The addition of casamino acids increases the yield of cells per volume at least by a factor of 3.

3.3.4 Resistance markers Expression plasmids harbouring dominant selectable marker genes (conferring resistance to otherwise toxic compounds) can be selected for in any yeast strain regardless of its genotype. This expands the range of heterologous gene expression to wild-type and polyploid industrial yeast strains. Selection is possible in complete media, such as YPD, where cells grow faster and to higher densities compared to minimal media. The vector FREP20RN (see Table 2) confers resistance to formaldehyde and has been used for high level expression of disaccharide transporters (Stolz and Sauer, unpublished data). There is a large selection of expression vectors for S. cerevisiae to choose from. The vectors given in Table 2 thus represent a personal choice of suitable vectors from different categories. They all contain 2u plasmid sequences as the origin of replication in S. cerevisiae.

3.4 Transformation Transformation of S. cerevisiae is as easy and nearly as efficient as transformation of E. coli. Most current transformation protocols are based on the lithium acetate-mediated method for the transformation of intact cells developed by Ito et al. (67). An excellent collection of high-efficiency transformation protocols is available from the Gietz laboratory at: http://www.umanitoba.ca/faculties/ medicine/humanXgenetics/gietz/Trafo.html. Three protocols, frequently used in our lab, are presented below.

3.5 Screening individual transformants for the presence of recombinant transport proteins Individual transformants may differ in the amount of recombinant protein they produce. If antibodies against a foreign transporter are available, individual transformants may be grown in small-scale liquid cultures and analysed by Western blotting for the presence and amount of protein. If expression of the heterologous gene in yeast is strong enough, the gene product may even be 87

N. SAUER AND J. STOLZ

Table 2 Yeast expression vectors with different properties Name

E. coll S. cerevlslae Promoter Marker Marker

Unique cloning sites

Reference

several

59

LEU2 URA3

CYC± TEFL DH1 GPD

Standard expression vectors p4XX series

Amp

H/S3 TRP1

pVT-U series

Amp

URA3

ADH1

BamHI, Xba\, Sacl, Xho\, Pstl, Pvull, Hindlll

60

NEV-N

Amp

URA3

PMAi

Notl

18

NEV-E

Amp

URA3

PMA1

EcoRI

18

NEV-X

Amp

URA3

PMA1

Xhol

18

YEpll2AlNE

Amp

TRPI

ADHi

EcoRI, Notl, BamHI

17

PYADE4

Amp

TRP1

ADH1

BamHI, Smal, EcoRI, C/al, 61 Sa/l, Xhol, Kpnl

pYPGE2

Amp

TRPI

PGK

BamHI, Smal, EcoRI, C/al, 61 Sa/l, Xhol, Apa\, Kpnl

PYSVE9

Amp

TRPi

SV40

BamHI, Smal, EcoRI, Clal, Xhol, Apal, Kpnl

61

Expression vectors with Induclble promoters pYES2

Amp

URA3

GAL1

Hindlll, Kpnl, Sacl, BamHI, EcoRI, Notl, Xhol, Sphl, Xbal

Invitrogen

pYeDPl/10

Amp

URA3

GAL10/CYC1

BamHI, EcoRI,

62

PYGAE2

Amp

TRP1

GAL10

BamHI, Smal, EcoRI, Xhol, Apal, Kpnl

61

pYE4

Amp

TRP1

PH05

EcoRI

63

pYEULCBa

Amp

LEU2-dURA3 CUP1

Bgl||

BamHI, Sa/l, Pstl, EcoRI

64

Expression vector with a resistance gene FREP20RN

Amp

formaldehyde PMA1 resistance FAR

see Figure 4.1

this work

several"

65

Vectors for expression in S. cerevisiae or E. coll pMTL8013

Amp

LEU2-d

PGK::REP2

pMTL8023

Amp

URA3

PGK::REP2

PMTL8033

Amp

URA3-4

PGK::REP2

severalb

65

several"

65

Sacl, BamHI, H/ndlll

66

EcoRI

30

Vectors for expression In S. pombe or S. cerevisiae

88

pEVP11

Amp

LEU2

SAP-E

Amp

LEU2

adh1S. pombe adh1

S. pombe

a

ATG upstream of polylinker

b

Cloning of insert DNA is possible at the authentic start codon of PGK

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

Protocol 2 Transformation of Saccharomyces cerevislae Equipment and reagents • TE buffer: 10 nM Tris-HCl pH 8.0.1 mM EDTA • Carrier DNA: Add high molecular weight DNA from salmon testes (Sigma D-1626) to TE buffer to give 2 mg/ml. Disperse by repeated pipetting and by vigorous stirring on a magnetic stirrer. If available, sonication with a tip-type ultrasonic processor is an easier way to get the DNA into solution. Stop when the viscosity of the solution has greatly decreased. Store aliquots at -20°C and heat-denature in boiling water for 5 ruin prior to use. Chill quickly on ice.

1 M and 100 mM Li-acetate solutions (LiAc) The final pH should be between 8.4 and 8.9. Filter-sterilize, 50% (w/v) PEG solution (MW 3350 or 4000). Filter-sterilize or autoclave, Plasmid DNA: DNA prepared by standard minipreps from E. coli without further purification will give excellent results. YPD liquid medium and agar plates (see Protocol 1)

Selective YPD plates (see Protocol 1)

A. The highest efficiency method, e.g. for transformation with a cDNA library; sufficient for up to 10 transformations 1. Grow cells overnight in 25 ml of YPD medium on a rotary shaker at 29°C. 2. Read the OD of the cells at 600 nm. Dilute into 50 ml pre-warmed (29°C) YPD (see Protocol I) to a final 0D600 of 0,5, 3. Grow the cells to an OD600 of 2.0 (depending on the strain. this will take 3-5 h). 4. Harvest the cells by centrifugation at 3000 g for 3 min. 5. Discard the medium, resuspend the cells in 25 ml of sterile water, and centrifuge again. 6. Discard the water, resuspend the cells in 1 ml of 100 mM LiAc, and then transfer to a 1.5 ml microcentrifiige rube. 7. Pellet at top speed for 15 sec and then remove the supernatant with a pipette. 8. Resuspend the cells to a final volume of 0.5 ml in 100 mM LiAc, then divide into 10 portions of 50 ul each (yielding 10 samples in microcentrifiige tubes for 10 transformations}. 9. Pellet the cells for 15 sec and remove the supernatant with a pipette. 10. To each tube add, in the given order:

• • • •

240 ul 36 ul 25 ul 50 ul

PEG 1 M LiAc carrier DNA water with 1 ug of plasmid DNA

N. SAUER AND J. STOLZ

Protocol 2 continued

11. Vortex each tube vigorously to resuspend the cells. 12. Incubate at 30°C for 30 min 13. Incubate at 42°C for 20 min. 14. Centrifuge for 60 sec at 5000g to pellet the cells. Use a pipette to remove the supernatant. 15. Resuspend the cells in 1 ml of sterile water by pipetting up and down. Do not vortex! 16. Plate aliquots of 2-200 u1 on to selective YNB plates (see Protocol 1). Incubate the plates at 29°C. Transformants will appear within 3 or 4 days.

B. A very quick method, but with reduced efficiency 1. Streak the strain to be transformed on to a YPD plate (see Protocol 1) to obtain single colonies, and grow for 1-2 days at 29°C. 2. For each transformation to be made, take a mid-sized colony with a loop and transfer the cells into 1 ml of sterile water. 3. Pellet the cells, resuspend in 1 ml of 100 mM LiAc and incubate at 30°C for 5 min. 4. Divide the cell suspension into aliquots as needed and pellet by centrifugation as described in Part A above. 5. Perform steps 10, 11, 13, and 14 of Part A given above. 6. Resuspend the cells in 200 ul of water and plate on to YNB medium (see Protocol 1) that selects for the presence of the plasmid.

C. For transformation with dominant marker genes NB: The following changes have to be made when transforming plasmids with selectable dominant marker genes 1. Prepare plates containing YPD plus the selective compound on the day of transformation (e.g. 3 mM formaldehyde, if using FREP20RN; see figure 1). 2. After transformation of the cells (by following either Part A or B), transfer the cell suspension to 5 ml of YPD medium and shake at 30°C for 3-12 h to allow for the expression of the resistance gene. 3. Pellet the cells and plate aliquots on to selective media.

D. The following control transformations should be performed: (a) No plasmid DNA added. This checks for the stability of the marker gene and for potential contaminations. (b) Parental vector (empty vector without insert). The resultant transformants can be used as wild-type controls in further experiments and can be grown under conditions identical to those used for transformants bearing recombinant constructs.

90

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

Figure 1 Circular maps of three expression vectors (A-C) that have been used for the successful expression of foreign transport proteins in S. cerevisiae. SAP-E can be used for expression in S. pombe as well. For detailed descriptions see Table 2. Variants of the vector NEV-E with a unique Xhol (NEV-X) or Notl (NEV-N) cloning site are also described in Table 2, Restriction sites marked with an asterisk are methylated in dam+ E. coli strains. Abbreviations: FaR, formaldehyde resistance; PMAl-Pro, promoter of the PM41 gene; PMA1Ter, terminator of the PMA1 gene; adhl-Pro, promoter of the adh1 gene; AmpR, ampicillin resistance gene.

91

N, SAUER AND J. STOLZ

visible as a protein band on Coomassie-staincd SDS gels of Lotnl membranes (see Protocol .3), when compared to the detergent extract of a control strain. Note: Do not boil samples for solubilization with SDS. Many transport proteins will aggregate under these conditions. Solubilizalion for 10 min on ice

Protocol 3 Small-scale isolation of yeast total membranes Equipment and reagents • Selective medium • 100 x proteinase inhibitor mix: 250 mM PMSF. 100 mM p-aminobenzamidine, Solubilize in DMSO and store frozen, • Storage buffer: 50 mM potassium phosphate (KPi) pH 6.3.1 mM MgSO,, 20% glycerol • Glass beads, 0,5 ram diameter

• Breaking buffer: 25 mM Tris-HCl pH 7,5, 5 mM EDTA • Middle-sized test tubes (1.2 x 10 cm) • Pasteur pipettes. Draw out the tip of the Pasteur pipette over a Bunsen burner flame to reduce its inner diameter to below 0.5 mm, i.e. the size of the glass beads.

Method 1. Grow cells on selective medium to mid-logarithmic phase (OD600 of 1-2). 2. Harvest 40 0D600 units of cells (i.e. 40 ml of cells have to be harvested if the culture, grown on minimal medium, has an 0D600 of 1.0). 3. Wash the cells once with 20 ml of water and once with breaking buffer (0°C). Remove the supernatant completely! 4. Resuspend the cell pellet in 200 ul of breaking buffer (0°C) and transfer to a test tube containing 0.5 g of glass beads (pre-cooled on ice). 5. Add 2 ul of proteinase inhibitor mix and break the cells by four cycles of 30 sec vortexing and 30 sec resting on ice (to reach maximal agitation, the vortex mixer should be set to permanent operation). 6. Add 1 ml of breaking buffer (0°C) and 10 ul of proteinase inhibitor mix. Vortex to mix, 7. Use a Pasteur pipette to transfer the broken cells to a microcentrifuge tube and centrifuge at 3000 g for 2 min at 4°C. 8. Transfer the supernatant (membranes and soluble proteins) to another centrifuge tube. Spin at 75000 g for 30 min at 4°C. 9. Discard the supernatant and resuspend the pellet in 100 ul of storage buffer (0°C) and 1 ul of proteinase inhibitor mix.a 10. Determine the protein content of the isolate with the Bradford assay (68) using BSA as a standard. 11. Analyse the isolated membranes by SDS-PAGE, and Western blot for the presence of your protein. a

92

A pestle that fits tightly into the microcentrifuge tube may be used to aid resuspension.

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

or at room temperature is usually sufficient for efficient dcnaturation and minimizes the risk of irreversible aggregation. Sometimes expression levels are too low and many membrane proteins do not migrate as a sharp band on SDS gels. In these cases it may be difficult to detect the additional protein in extracts from total membranes. If the transporter is targeted to the plasma membrane, preparation of plasma membranes may help to visualize the protein in stained gels. See Protocol 5 for the preparation of plasma membranes from yeast.

3.6 Radiotracer uptake experiments As stated above, eukaryotic plasma membrane transporters are usually targeted to the plasma membrane in yeast. For this reason radiotracer uptake experiments with intact yeast cells allow the simple and direct determination of the activity of a foreign transporter. A standard uptake experiment is presented (with 14C-labelled sucrose as substrate) in 1'ratocol 4. Depending on the type of transporter and depending on what is known about its properties it may be necessary to modify some of the parameters described in Protocol 4, such as the pH or substrate concent ration.

Protocol 4 Uptake experiment with a radlolabelled substrate Equipment and reagents • Selective medium • Transport assay buffer: 50 mM KPi pH 5.5 • 100 mM stock solution of unkbelled sucrose in H2O • [U-14C]sucrose (specific activity 200 • Mix of unlabelled and labelled sucrose: 100 ul 100 mM sucrose plus 10 ul [U-14C]sucrose • 1 M D-glucose solution in H20

• Rotatoryshaker.setto30°C • Filtration device, suitable for multiple nitrations • Vacuum pump, connected to a vacuum trap and to the filtration device • Membrane filters (e.g. Schleicher and Sehuell, ME 27,25 mm diameter, pore size 0.8 um) • Scintillation counter. vials, and fluid

Method 1. Grow transformed yeast cells under selective conditions to logarithmic phase (0D 600 1-2). 2. Harvest the cells by centrirugation for 3 min at 4000 g (for each transport test 10 0D600 units of yeast cells will be needed). 3. Wash the cells once in 20 ml ice-cold water and once in 20 ml ice-cold transport assay buffer by resuspending the pellet and then centrifugrng as above. 4. Resuspend the cells in transport assay buffer to give 10 0D600/ml and keep on ice until needed.

93

N. SAUER AND J. STOLZ Protocol 4 continued

5. Put filters on the filtration device, fill the chimneys of the device with 5-10 ml of water. 6. Pre-warm 1 ml of the cell suspension in a 25 ml Erlenmeyer flask in a rotatory shaker set to 200 r.p.m. and 30°C. 7. After 2 min add 11 ul of the mix of labelled and unlabelled sucrose (i.e. 10 ul of unlabelled and 1 ul of labelled sucrose). Start the timer now! 8. Withdraw aliquots of 100 ul at timed intervals (e.g. 15 sec, 1 min, 2 min, 3 min, 5 min, 7 min, 10 min). Pipette the aliquots immediately into the filtration device and apply a vacuum to filter the cells on to the membrane. 9. Wash the cells by adding about 5-10 ml of water from a squirt bottle and reapply the vacuum. 10. Put the membrane filters from each time point into scintillation vials, add scintillation fluid, and count in a scintillation counter. 11. Pipette a 100 ul aliquot from step 7 directly into a scintillation vial without filtration. The measured radioactivity in this sample will allow you to correlate the radioactivity of the samples counted in step 10 with the amount of sucrose that is present in each, Note: For maximal transport rates with energy-dependent transporters (e.g. with a H+-sucrose symporter) supply the cells with a rapidly metabolizable energy source, such as 10 ul of 1 M D-glucose (added during the pre-warming at step 6; the final concentration of D-glucose is 10 mM). For uptake tests with radiolabelled monosaccharides 10 ul of ethanol can be added instead (the final concentration of ethanol will be 100 mM). Both additions cause optimal energization of the yeast plasma membrane.

3.7 Large-scale preparation of total or plasma membranes Recombimmt transporters in yeast can be purified after solubilization from a crude preparation of yeast total membranes. If the protein is targeted to the plasma membrane, purification of these membranes will greatly enrich the protein prior to solubilization. The add precipitation method for enriching plasma membranes (see ref. 69 and Protocol 5) is a fast and convenient method. If the recombinant transport protein gives a major band on SDS gels after solubilization from plasma membranes, preparative SDS-PAGE may be used as a simple and direct way of protein isolation, e.g. for the production of antisera,

3.8 Addition of protein tags and purification of biotin-tagged transport proteins 3.8.1 Tagging proteins For the rapid and simple purification of recornbinant transport proteins, a variety of tags can be fused to the coding sequence by standard genetic manipulations. Addition of a protein tag to the C-terminus of a transporter should minimize

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

Protocol 5 Large-scale preparation of yeast total and plasma membranes Equipment and reagents • See Protocol 3 for the required materials. In addition you will need: • TM buffer: 10 mM Tris-HCl pH 7.5, 1 mM MgS04 • 2 M acetic acid « 2MNaOH • Dounce homogenizer with a tight-fitting pestle

A mechanical device for breaking the cells with glass beads is necessary for preparative protein isolation from yeast. In our laboratory a Braun-Melsungen glass-bead homogenizer equipped with a C02 cooling system is used. Glass-sintered filter (porosity 1) Magnetic stirrer

Method 1. Grow cells in selective media until late logarithmic phase. 2. Harvest the cells by centrifugation for 5 min at 4000 g, wash once with water and once with breaking buffer at room temperature. 3. Resuspend the cells in breaking buffer to give 1 g fresh weight/ml. Note: Steps 4-6 are described for a Braun-Melsungen glass-bead homogenizer and have to be adapted if another device is used for homogenization. 4. Fill a 70 ml breaking vessel with 90 g of ice-cold glass beads and add 20 ml of the cell suspension. 5. After adding 200 ul of proteirtase inhibitors, break the cells at maximum speed for 1 min in the cell mill. Cool intermittently with solid C02. 6. Remove the glass beads by filtration through a glass-sintered filter (porosity 1), then wash the beads with 10 ml of cold breaking buffer. 7. Centrifuge the combined filtrates twice at 0°C for 5 min at 3000 g to remove unbroken cells. 8. Add proteinase inhibitors to the supernatant and centrifuge at 100000 g for 60 min to pellet the membranes, 9. Repeat step 8, if required, to achieve a more complete removal of soluble proteins. 10. Use the total membrane fraction directly for protein purification (following solubilization of the membranes with detergent) or store at -80°C for later use, (For the latter. resuspend the membrane pellet in storage buffer containing proteinase inhibitors and freeze it in liquid nitrogen). The fraction may also be used for further purification of plasma membranes as described in step 11, below. 11. Resuspend the membrane pellet at 0°C in TM buffer (1 ml/g fresh weight of cells) using a Dounce homogenizer with a tight-fitting pestle.

IM. SAUER AND J. STOLZ

Protocol 5 co

12. Transfer the membrane suspension to a centrifuge tube, add a small magnetic stirring bar. and lower the pH of the suspension to pH 5.2 by the stepwise addition of 2 M acetic acid with continuous stirring at 0 °C. 13. Remove the stirring bar and immediately centrifuge at 10000 g for 1 min at 0°C (the resultant large pellet will contain most of the mitochondrial membranes), 14. Transfer the supernatant to a fresh tube and bring to pH 7.5 with the 2 M NaOH solution. 15. Sediment the highly enriched plasma membranes from the supernatant (if required) by centrifugation at 55000 g for 30 min at 0°C. 16. Resuspend the white plasma membrane pellet in storage buffer, freeze in liquid nitrogen, and store at -8Q°C.

Table 3 Tags used for purification or detection of proteins in yeast Tag

Size Purification (amlno acids)

his tag

6

biotinylated 75 or domains

96

more

Crosspurification of yeast proteins

depends, possible mostly simple

Immuno- Detection genlcity

Crossreactivity In yeast

Remarks {Reference}

very low difficult

?

(31, 49)

easy,

not detected strong

very

with

additional

high affinity

after

sensitive,

soluble

biotin in

to avidin

complete removal of soluble proteins

with avidinbased reagents

biotinylated the growth proteins medium required (56, 70} 7 (73, 74)

GST

219

simple

no

strong

anti-GST antibodies

Strep-Tag

9

simple

?

low

very sensitive. with avidinbased reagents

with (75) soluble biotinylated proteins

HAepitope

9

with mAK

no

low

with mAK 12CA5

no

(40)

c-tnyc epitope

10

?

?

low

with mAK 9E10

no

(76, 77)

GFP

238

?

?

strong

easy and no sensitive, anti-GFP antibodies also available

(78)

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

the potential interference with N-terminal targeting signals. Also, the C-tcrmini of plant transport proteins seem to be very tolerant to the addition or removal of amino acid residues {31, 49, 56, 70|. If possible, thecodon usage tor additional amino acid residues should be in agreement with the corions most frequently used in S. cm'visittc (71) or 5". pombt? (72). TAA should be used, if a stop codon has to be included. An ideal tag should allow purification of the transporter in one step as well as specific detection of the tagged protein in viva and on Western blots. It should be as little immunogenic as possible, if the tagged protein is to be used for antigen production, and it should not interfere with protein structure, function, or localization. As this perfect tag does not exist, the relative benefits of available tags are summarized in Table 3.

3,8.2 Purification of biotin-tagged protein Follow Protocol 5 for the large-scale purification of yeast total membranes. i'roUicol h has been optimized tor the purification of the biotin-iagged sucrose transporter PmSUC2biohis6 from total membranes (56), As the interaction of biotin and avidin is stable over a wide range of pH values, salt concentrations and buffers, adaptation of Protocol 6 to the special requirements of other biotinylated transporters will be easy. The optimal detergent used for the purification of a given transporter has to be found empirically. Mild non-ionic detergents, such as Triton X-100, n-dodecyl-p-i) maltoside (DDM) or n-octyl-p-n-glucoside work well with many different transport proteins. Furthermore, they are well suited for reconstituiion of transport proteins into liposomes and have no negative side-effects during the generation of antisera.

Protocol 6 Purification of biotinylated proteins with immobilized avidin Equipment and reagents • Imnuraopure immobilized monomeric avidin (Pierce):

• Wash buffer 1: 50 mM KPi pH 6.3. 50 mM sucrose, 500 mM KC1, 0.1% (w/v) DDM

• KPi/sucrose buffer: SO mM KPi pH 6.3, 50 mM sucrose

» Wash buffer 2: 50 mM KPi pH 6.3, 50 mM sucrose, 0.1% (w/v) DDM

• Solubilization buffer: 50 mM KPi pH 6.3, 50 mM sucrose, 500 mM KC1, 2% (w/v) DDM

• Elution buffer: 50 mM KPi pH 6.3, 50 mM sucrose, 0.1% (w/v) DDM, 1 mg/ml n-biotin

A. Column preparation 1, Prepare a column (bed volume 1 ml) according to the manufacturer's protocol. 2, Equilibrate the column in 10 column volumes of solubilization buffer by gravity flow. 97

N. SAUER AND J. STOLZ Protocol 6 continuec

B. Purification NB: All steps should be performed in the cold room and on ice! 1. Thaw an aliquot of yeast total membranes (equivalent to 100 mg of membrane protein). 2. Resuspend in 20 ml of 50 mM KPi/sucrose buffer and centrifuge at 50 000 g for 30 min to pellet the membranes. 3. Solubilize the membrane pellet with 10 ml of solubilization buffer. Keep on ice for at least 30 min and shake intermittently, 4. Centrifuge at 100 000 g for 60 min to remove insoluble material. 5. Transfer the supernatant (containing solubilized proteins) to the equilibrated column and let it pass through. 6. Pass the flow-through fractions through the column a second time (to ensure complete binding of biotinylated protein). 7. Wash the column with 10 ml of wash buffer 1. 8. Wash the column with 10 ml of wash buffer 2. 9. Elute the protein with 10 ml of elution buffer. 10. Use the eluate for a second elution (to ensure complete release of biotinylated protein), 11. Analyse aliquots of the eluted fractions by SDS-PAGE (using Coomassie staining, silver staining, or Western blotting with avidin peroxidase to detect biotinylated proteins on the blot).

4 Expression in Schizosaccharomyces pombe Most of the general remarks made for .S. cereviside also hold true for the fission yeast S. pombe. S. potribe is frequently used as an alternative expression host, e.g. because its cod on usage differs from that in S. cemisiae or simply because most plasmids constructed for use in S. cmvisiae are also compatible for use in S. pombe. However, S. pombe is still less developed as an expression system, resulting in a smaller selection of plasmids and host strains. S. pombc should be grown on YE complete medium (0.5% yeast extract, 2% glucose), with YNB used as the selective medium (see Protocol 1).

4,1 Host strains Primarily, two nutritional markers are used for molecular studies in .S. pomhc, ura4~ and leu I'. Strains cany ing the Icul' allele can be transformed with plasmids harbouring the U
EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST Table 4 S. pombe strains used for the expression of foreign transporter genes Strain (Reference)

Genotype

Used for (Reference)

leu 1.32 (79)

leu1-

functional expression and protein purification (27-29, 31, 36) functional expression and protein purification (29, 30, 81)

YGS-5

h-,

leu1-,

ght1-

(80)

copies of a vector carrying the S. cerevisiae URA3 gene. However, it has also been described that the S. pombe ura4~ mutation can only be complemented by the S. pombe URA4 gene. Protease deficient strains of S. pombe are not available. Strain S. pombe leu1.32 (79) has frequently been used for expression studies (see Table 4). Unlike in S. cerevisiae cells, which harbour multiple genes encoding glucose transporters, there is only one glucose carrier present in S. pombe. Strain YGS-5 carries a mutation in the gene encoding the glucose transporter (GHT1) and is unable to grow on medium containing glucose as the sole carbon source (80). Strain YGS-5 will grow on complete medium containing 2% gluconate (potassium salt) instead of D-glucose. When transformed with plant glucose transporter genes, YGS-5 regains the ability to grow on a glucose-containing medium. YGS-5 is an excellent host for the characterization of glucose transport proteins using transport assays, as well as for the expression of transporters for purification purposes.

4.2 Expression vectors Vectors for constitutive expression frequently carry the adhl promoter of the alcohol dehydrogenase gene or, for a moderate level of expression, the SV40 early promoter. Inducible expression is most frequently driven by the thiaminerepressible nmtl promoter (nmt for no message in thiamine). A shift to thiamine-free medium causes strong expression of DNA cloned downstream of this promoter. Vectors with S. cerevisiae 2u sequences are maintained at 5-10 copies per cell. Alternatively, vectors using S. pombe ARS sequences as the origin of replication, will be present at 25-80 copies per cell.

4.3 Transformation As for S. cerevisiae, several protocols are available for the transformation of S. pombe. The procedure given below will yield approximately 1000 transformants per 1 ug of DNA. Much higher transformation efficiencies can be obtained by electroporation (86). S. pombe transformants can be analysed with the same methods that have been described in detail for S. cerevisiae (see Sections 3.5-3.8). 99

N. SAUER AND J. STOLZ

Protocol 7 Transformation of Schizosaccharomyces pombe Equipment and reagents • YE complete medium: 0.5% yeast extract, 2% glucose • YNB selective medium, agar plates, and supplements (see Protocol 1) • 15 ml Falcon tubes • Li-acetatefTris/EDTA (LiAcfTE): 100 mM lithium acetate, 10 mM Tris-HCI pH 7.5. 1 mM EDTA (autoclave!)

la-acetate/Tris/EDTA with polyethylene glycol (LiAc/TE/PEG): 100 mM lithium acetate, 10 mM Tris-HCl pH 7.5.1 mM EDTA. 40% PEG-4000 (filter-sterilize!) Carrier DNA: Prepare as described in Protocol 2, but at a concentration of 10 mg/ml in TE.

Method 1. Grow cells in 100 ml YE-medium to an 0D600 of approx. 1.0 in a 250 ml Erlenmeyer flask. 2. For each transformation, centrifuge 10 ml of this culture in a 15 ml Falcon tube for 5 min at 4000 g. 3. Resuspend the pellet in 10 ml of sterile water and centrifuge the cells again. 4. Resuspend the pellet in 10 ml of LiAc/TE and centrifuge the cells again. 5. Resuspend the cells in 100 ul of LiAc/TE. transfer to a microcentrifuge tube and keep at 30°C for l-3h. 6. Add 5 ul of carrier DNA plus up to 5 ug of plasmid DNA and keep at 30°C for 30 min, 7. Add 700 ul of LiAc/TE/PEG. Shake well to mix and keep at 30°C for at least 60 min. 8. Centrifuge the cells in a microcentrifuge (at 12 000 g) and remove the supernatant with a pipette. 9. Resuspend the pelleted cells in 200 ul of sterile water and plate on to YNB plates. 10. Transformants appear as colonies within 4-6 days at 30°C

4.4 Influence of 5'-untranslated sequences Depending on the source of DNA that has been used tor insertion into an S. pumbe expression vector the 5'-untranslated region will vary in length and base composition. It has been described that both S. cerevisiae and S. pombe have specific requirements for the base composition in their 5'-untranslated sequences tor optimal expression (1). To increase the expression level of a foreign gene, the sequence upstream of the start ATG may be changed by PCR. This worked well for the CkHUP2 galacrosc-H' symporter and the CkHUP3 glucose-H' symportiM- genes from the alga Chlorella kessleri, which could not be expressed with 100

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

Table 5 Schizosaccharomyces pombe expression vectors Name

E. coll marker

S. pombe marker

Origin of replication

Promoter

Unique cloning sites

Reference

Vectors with auxotrophlc markers pEVP11

Amp

LEU2

2u.

adh1

SamHI, Sacl, Hindlll

66

pART1

Amp

LEU2

ars1

adh1

Pstl, Sa/l, BamHI, Smal, Sad

82

SAP-E

Amp

LEU2

2u.

adh1

EcoRI

30

Vectors with resistance markers pTL2Ma

Amp

Neomycin resistance

hCMV

Afllll, EcoRI, Hindlll, Sacl, Smal, Xbal

83

pcD4

Amp

Neomycin resistance

hCMV

SamHI

83

Vectors for Inductble expression pREP1a

Amp

LEU2

arsl

nmt1

Ndel, Sa/l, BamHI, Smal

84

pREP2a

Amp

URA4

ars1

nmt1

Ndel, Sa/l, BamHI, Smal

84

pREP3a

Amp

LEU2

ars1

nmt1

Ba/l, Sa/l,

84

BamHI, Smal pREP4a

Amp

URA4

ars1

nmt1

Ba/l, Safl, BamHI, Smal

84

REP3X

Amp

LEU2

ars1

nmt1

Xhol

85

"Vector contains ATG in the polylinker

the 5'-untranslated sequences of the CkHUP2 or CkHUP3 cDNAs (29). Replacement of these 5'-untranslated sequences by PCR with the 5'-untranslated sequences of the AtSTP1 monosaccharide transporter cDNA from A thdliana (last 15 bases upstream of the start AIG of AtSTPI (including a HindIII cloning site): 5'-AAG CTT GTA AAA GAA ATG-3'), that had been successfully expressed before (28), resulted in the functional expression of CkHUP2 and OKHUP3 (29). Alternatively, changes of 5'-untranslated sequences can also be made according to available consensus sequences (1).

5 Expression of membrane proteins in Pichia pastoris Although many soluble proteins are highly abundant when overexpressed in P. pastoris (34), there are hardly any reports on the functional expression of membrane proteins and no reports on the expression of transporters in this yeast. To our knowledge, the expression of a mouse serotonin receptor (87), human B2101

N. SAUER AND J. STOLZ

adrenergic receptor (88), a human opioid receptor (89), and bovine opsin (90) are the only examples of the functional expression of membrane proteins in P. pastoris published up to now. This yeast is capable of using methanol as the sole source of carbon and energy, and will grow to very high cell densities of up to 100 g fresh weight/litre of medium. The promoter of the key enzyme of methanol assimilation, alcohol oxidase (AOX1), is tightly regulated and is induced upon the addition of methanol to produce alcohol oxidase at levels of up to 30% of total cellular protein. The AOX1 promoter is routinely used for foreign gene expression in P. pastoris, either from episomal plasmids or, after chromosomal integration, from multiple copies of the integrated expression cassette. Marker genes used in P. pastoris are the HIS4 or ARG4 biosynthetic markers, SUC2, which encodes invertase and renders P. pastoris able to grow on medium containing sucrose as the sole source of carbon and energy, as well as some antibiotic resistance genes (34). Strains of P. pastoris with protease deficiencies are available and should be used when purification of the recombinant transporter is desired.

6 Future perspectives Over the last 10 years heterologous expression of transport proteins in yeast has become a valuable tool for the characterization, cloning, and purification of foreign transport proteins, and the influence of this technique on the transporter field will certainly increase in the future for different reasons. First, transporters for a variety of so far undetected substrates will be identified by expression in yeast mutants lacking the corresponding activity. Since the complete sequence of the S. cerevisiae genome is available and the number of proteins with known function is gradually increasing, more and more well-defined transport mutants will be available in the future. Complementation studies with expression libraries will also identify transporter genes for the respective substrates in animals and plants. Second, with the currently available molecular tools, the specific advantages of other yeast types, such as the methylotrophic yeasts Hansenula polymorpha or Pichia pastoris, will be used more frequently in the future. The enormous yields of recombinant proteins that have been described for several genes of soluble proteins expressed in these yeast cells surpass all reports on protein yields in S. cerevisiae or S. pombe by far. Another advantage of these yeasts is their potential of targeting proteins to peroxisomes. Production of peroxisomes is massively induced during growth on methanol (91) and foreign transport proteins may be packaged into the membrane compartment of these organelles upon the addition of peroxisomal targeting sequences (e.g. PTS1: Ser-Lys-Leu-COOH at the C-terminus). This will reduce possible toxic effects of foreign transporter gene expression to the yeast and may also protect the recombinant transporters from undesired aggregation. In addition, in H. polymorpha peroxisomal membranes show strong proliferation in the presence of oleate (92), yielding a 102

EXPRESSION OF FOREIGN TRANSPORT PROTEINS IN YEAST

membrane compartment with an extremely low protein content. These properties are especially useful for the expression of membrane proteins and will extend the attractiveness of yeasts as hosts for foreign transporter gene expression.

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Faulkner, J. D., Anson, J. G., Tuite, M. F., and Minton, N. P. (1994). Gene, 143, 13. Russel, P. and Nurse, P. (1986). Cell, 45, 145. Ito, H., Fukuda, Y., Murata, K., and Kumura, A. (1983). J. Bacteriol, 153, 163. Bradford, M. M. (1976). Anal. Biochem., 72, 248. Dufour, J.-P., Amory, A., and Goffeau, A. (1988). In Methods in enzymology. Vol. 157 (ed. S. Fleischer and B. Fleischer), p. 513. Academic Press, London. Caspari, T., Robl, I., Stolz, J., and Tanner, W. (1996). Plant J., 10, 1045. Sharp, P. M. and Cowe, E. (1991). Yeast, 7, 657. Forsburg, S. L (1994). Yeast, 10, 1045. Mitchell, D. A., Marshall, T. K., and Deschenes, R. J. (1993). Yeast, 9, 715. Ward, A. C., Castelli, L. A., Macreadie, I. G., and Azad, A. A. (1994). Yeast, 10, 441. Murphy, J. T. and Lagarias, J. C. (1997). Photochem. Photobiol, 65, 750. Matijekova, A. and Sychrova, H. (1997). FEBS Lett., 408, 89. Kuchler, K., Dohlman, H. G., and Thorner, J. (1993). J. Cell. Biol, 120, 1203. Niedenthal, R. K., Riles, L., Johnston, M., and Hegemann, J. H. (1996). Yeast, 12, 773. Beach, D. and Nurse, P. (1981). Nature, 290, 140. Milbradt, B. and Hofer, M. (1994). Microbiology, 140, 2617. Caspari, T., Stadler, R., Sauer, N., and Tanner, W. (1994). J. Biol Chem., 269, 3498. McLeod, M., Stein, M., and Beach D. (1987). EMBOJ., 6, 729. Tohda, H., Okada, H., Giga-Hama, Y., Okayama, H., and Kumagai, H. (1994). Gene, 150, 275. Maundrell, K. (1993). Gene, 123, 127. Forsburg, S. L. (1993). Nud Adds Res., 21, 2955. Prentice, H. L. (1991). Nud. Acids Res., 20, 621. WeiB, H. M., Haase, W., Michel, H., and Reilander, H. (1995). FEBS Lett, 377, 451. Weift, H. M., Haase, W., Michel, H., and Reilander, H. (1998). Biochem.]., 330, 1137. Talmont, F., Sidobre, S., Demange, P., Milon, A., and Emorine, L. J. (1996). FEBS Lett., 394, 268. Abdulaev, N. G., Popp, M. P., Smith, W. C., and Ridge, K. D. (1997). Protein Expr. Purif., 10, 61. Veenhuis, M. (1992). Cell Biochem. Funct., 10, 175. Veenhuis, M., Kram, A. M., Kunau, W.-H., and Harder, W. (1990). Yeast, 6, 511.

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Chapter 5 Baculovirus-mediated overexpression of transport proteins Gary J. Litherland* and Stephen A. Baldwinf *Department of Anatomy and Physiology, MSI/WTB Complex, University of Dundee, Dow Street, Dundee DD1 5EH f School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT

1 Introduction A major limitation to our understanding of the molecular mechanisms of membrane transport processes is the lack of detailed structural information on transport proteins, such as might be obtained using X-ray crystallography and other biophysical techniques. One reason for the paucity of structural studies on transporters using such techniques has been the difficulty in producing sufficient quantities (10-100 mg) of pure transport protein. The natural abundance of most transporters is low, and purification difficult. While many prokaryotic transporters have been successfully expressed at high levels in Escherichia coli (see Chapter 6), the use of bacteria to overexpress complex, often glycosylated or otherwise modified membrane proteins from eukaryotic sources has rarely been successful. Even in those few cases where functional expression of a eukaryote transporter in Escherichia coli has been reported, such as for the hexose-proton symporter HUP1 from the green alga Chlorella kessleri, the activity of the expressed protein is usually lower than that obtained by expression in eukaryotic (yeast) cells, even following reconstitution under optimal conditions (1). Similarly, while Xenopus oocytes and mammalian cell lines have been used to great effect in the expression and investigation of many eukaryote transporters (see Chapter 3), the levels of expression obtained using these systems are generally insufficient for biophysical studies. Currently, the optimal hosts for the overexpression of eukaryote transport proteins are therefore yeasts and insect cells. The relative merits of yeast and insect cells might be summarized as convenience and cheapness in the case of the former, and a more sophisticated modification machinery and more native (in the case of mammalian proteins) environment in the latter. While the use of yeast systems is described elsewhere in this volume (see Chapter 4), the present chapter will consider the use of 107

GARY J. LITHERLAND AND STEPHEN A. BALDWIN

baculovirus expression systems to facilitate the heterologous overproduction of eukaryotic transport proteins in insect cells. The background theory is briefly covered, and generalized schemes for protein expression are presented. Examples from the literature pertaining to baculovirus-dependent expression of membrane proteins in general and of transporters in particular are discussed. A discussion of the practical aspects of the expression technology follows, and potential problems are highlighted.

2 An overview of baculovirus-mediated expression systems 2.1 The baculoviruses The family Baculoviridae is composed of large (80-220 kbp genome), membranebound, double-stranded DNA viruses that specifically infect arthropods, in particular lepidoptera (2). It is this species selectivity that gave rise to their original usefulness to man, i.e. as insecticides in agriculture. Baculoviruses can be grouped into three classes: nuclear polyhedrosis viruses (NPVs), granulosis viruses, and non-occluded viruses. It is the NPVs that will be discussed here, since this class forms the basis of baculovirus expression technology. The NPVs are so called because they produce paracrystalline, proteinaceous, nuclear occlusion bodies (polyhedra) in which progeny virions are embedded toward the end of the infection cycle (see Figure 1). When multiple virion are occluded within each polyhedron, the virus is classed as a multiple nuclear polyhedrosis virus (MNPV), in contrast with single nuclear polyhedrosis viruses (SNPVs) which embed only a single virus particle within each occlusion body. The MNPV strain most commonly used in the development of expression vectors is that which infects the alfalfa looper Autographa californica, designated AcMNPV. This virus, which has a genome of 134 kbp, is the most intensively studied baculovirus and its derivatives are at the heart of the vast majority of baculovirus expression systems (3-6). 2.2 The baculovirus life cycle A schematic illustration of the AcMNPV life cycle is shown in Figure 1. When occlusion bodies, lying dormant on food, are ingested by host insect larvae, they dissolve in the alkaline environment of the insect mid-gut. This releases the embedded virions, which are highly infectious and enter the mid-gut cells by membrane fusion. Viral genes are expressed during infection in three phases: early, late, and very late (7). Once in the nucleus, the viral DNA is uncoated and early viral genes are expressed, prior to and necessary for efficient viral replication. Replication occurs from approximately 6 hours post-infection (p.i.) until 18 hours p.i., during the late phase of viral gene expression (8). The cell nucleus appears enlarged at this stage of the infection cycle. By 12 hours p.i., progeny extracellular virus (ECV) is released from the cells by budding (budded virus, BV) 108

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

Figure 1 A schematic representation of the baculovirus life cycle. Viral occlusion bodies are ingested by susceptible organisms and are dissolved in the high pH environment of the insect mid-gut lumen. Released virus particles enter intestinal cells by membrane fusion, and upon entering the nucleus are uncoated and replicate. Some replicated virion migrate to the cytoplasm and leave the cell, whereupon they cause infection of adjacent cells. Other particles are incorporated into nuclear occlusion bodies, which accumulate within the cell until it eventually undergoes lysis. When the host dies the occlusion bodies are released into the environment where they may cause further infection.

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GARY J. LITHERLAND AND STEPHEN A. BALDWIN

to infect adjacent cells, thus spreading the infection throughout the insect. During the last stage of infection the very late (occlusion-specific) viral genes are expressed. This results in membrane proliferation within the nucleus and the envelopment of progeny viral nucleocapsids. The production of extracellular virus continues, together with the accumulation of virion within the nucleus, and at a late stage of infection (approximately 70 hours p.i.) many viruses become encapsulated within occlusion bodies (polyhedra), consisting largely of a paracrystalline matrix of the 29-kDa protein polyhedrin (9). The cell finally undergoes lysis at approximately 4-5 days post-infection, releasing the occlusion bodies, which may remain dormant but viable for several years in the soil or on plant tissue. 2.3 Expression technologies based on baculoviruses At the time of cell lysis, polyhedrin can account for as much as 50% of the cellular protein (10). Since viral occlusion is unnecessary for viral replication, the polyhedrin gene can be removed without affecting the infectivity of the virus. Transcription of the polyhedrin gene occurs from the extremely efficient polyhedrin promoter (pPolh), and it is this promoter that is at the heart of most commercially available baculovirus expression systems, and which allows abundant expression of recombinant proteins. However, a second, very late viral gene, responsible for the expression of the p10 protein, is also sometimes used in baculovirus expression vectors (3, 4, 11). The p10 gene product is thought to be involved in cell lysis and so, like the polyhedrin gene, is dispensable for viral infection and replication. It is expressed earlier in the infection time course than polyhedrin and this, coupled with delayed cell lysis by baculoviruses lacking p10, has been reported to result in the increased accumulation of recombinant protein in expression systems using the p10 promoter (e.g. the BacTen™ baculovirus system marketed by Quantum Biotechnologies Inc.) (11). Some expression systems utilize both the polyhedrin and p10 promoters to drive the expression of two heterologous proteins simultaneously, facilitating the study of proteinprotein interactions. Although use of the polyhedrin and p10 promoters can lead to very high levels of expression, a potential disadvantage of these promoters is that expression occurs at such a high level and late stage in infection that the capacity of the cell post-translationally to modify the expressed protein is overwhelmed. To circumvent these problems vectors that utilize other promoters may be employed, such as those of the immediate-early, ie1, and basic protein genes, which are expressed in the early and late phases of baculovirus infection, respectively. While these promoters are likely to yield lower expression levels, post-translational modifications (such as glycosylation) are likely to be more efficiently performed because these genes are expressed earlier after infection, and therefore in 'healthier' cells. The yield of biologically active expressed protein may therefore be higher than for expression using the polyhedrin and p10 promoters (12, 13). 110

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

There are many commercially available baculovirus expression systems, but all are designed according to one of two themes. Since the baculovirus DNA is large (134 kbp in the case of AcMNPV), it is usually impractical to clone the gene of interest into this molecule directly. Instead, a multiple-stage procedure is used to introduce the gene into the viral DNA. The first stage is to clone the gene into a transfer vector that can replicate in £. coli. The next stage involves the transfer of the gene from the vector to the baculovirus DNA, and it is at this point that the various available methods diverge.

2.3.1 In vitro co-transfection methods The most widely used type of baculovirus expression system utilizes homologous recombination within the host insect cell to produce recombinant baculovirus (see Figure 2). The transfer vector contains the polyhedrin promoter (or an alternative baculovirus promoter), usually preceding a versatile multiple cloning site, and further flanking regions that are derived from the baculovirus genome. After constructing a recombinant transfer vector by placing the gene of interest downstream of the polyhedrin promoter, the next step is to cotransfect insect cells with a mixture of the transfer vector and purified baculovirus DNA. Homologous recombination then occurs between identical sequences in the baculovirus and vector DNA molecules; a double crossover event results in the generation of a recombinant baculovirus species containing the gene of interest under the control of the polyhedrin promoter. The major problem with this method is that the frequency of homologous recombination between the co-transfected species is generally less than 1%. The recombinant progeny virus must therefore be identified in, and isolated from, a high-background population of wild-type virus. This entails the examination of viral plaques and the purification of those identified as recombinant, by morphology (14), DNA hybridization (5), immunological detection (15), or by using the polymerase chain reaction (16, 17). For example, where recombination results in the removal of the polyhedrin gene from the wild-type baculovirus DNA, recombinants may be identified by light microscopy by virtue of their non-occluded phenotype—occlusion bodies are highly retractile and give the cells a 'shiny' appearance. However, this procedure is time-consuming and can be technically difficult. Moreover, confirmation of plaques as representing recombinant virus requires additional screens. Fortunately, there have been several refinements made to the above method, aimed at either facilitating the identification of recombinant plaques, or at increasing the proportion of progeny virus that is recombinant: (a) Just as blue/white selection using the chromogenic substrate 5-bromo-4chloro-3-indolyl-p-D-galactoside (X-Gal) for assaying p-galactosidase (LacZ) activity has become a standard procedure for identifying recombinant bacterial colonies, the same process can be used to detect plaques made by recombinant baculovirus progeny, providing that either the parent baculovirus or the transfer vector carries lacZ. The presence of lacZ in the parent virus DNA aids the visualization 111

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112

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS Figure 2 Production of recombinant baculovirus by co-transfection of insect cells with baculovirus DNA and a transfer vector. The foreign gene of interest is cloned downstream of the polyhedrin promoter (pPolh) in a transfer vector. The vector contains additional flanking baculovirus sequences to act as sites for homologous recombination following co-transfection into insect cells with linearized baculovirus DNA. The latter ideally lacks a gene essential for viral replication, such that recombination with the transfer vector is required for the production of infectious viral particles. As a result, almost all the resultant baculoviruses represent recombinants and bear the foreign gene. Virus is harvested from the cell medium after 2-3 days. At this point it is usually necessary to purify viruses from individual plaques after secondary infection, to isolate clonal recombinants. The virus stock is amplified by further infection of insect cells, its titre determined and then used for the large-scale infection of cells. Recombinant protein is isolated from the resultant cell lysates, often using affinity chromatography facilitated by the presence of transfer vector-encoded affinity tags.

of plaques per se. Some baculovirus systems, by direct analogy to the bacterial systems, use a loss of blue colour to indicate recombination (18), but in the context of plaque screening this is not ideal since white plaques are often difficult to detect. One variation of this method utilizes a baculovirus that produces blue plaques whether recombined or not, which facilitates the visualization of the non-occluded phenotype resulting from recombination (19). The third, and most powerful, variation on this theme places the lacZ gene on the transfer vector within the recombined region (20, 21). As a result, only recombinant plaques are blue, which greatly facilitates their identification against potentially high-background populations. (b) Other modifications to the original method have been directed at increasing the proportion of recombinant progeny virus. One of the first approaches employed to minimize the background of non-recombinant viruses resulting from co-transfection experiments was to use baculovirus DNA linearized at a suitable restriction site, because linear baculovirus DNA is much less infectious, while retaining the ability to undergo recombination within the cell. Such linearization was achieved by introduction of an artificial Bsu36I site adjacent to the polyhedrin promoter: the wild-type viral genome contains no Bsu36I sites (22). Following co-transfection, a double crossover event between the transfer vector and linearized viral DNA at homologous DNA sequences flanking the polyhedrin promoter and the linearization site should produce circular, and thus infectious, baculovirus DNA, while few progeny should arise from nonrecombinant DNA. While this approach did not completely eradicate the background of non-recombinant virus resulting from co-transfection experiments, it did significantly reduce this background (22). (c) A further advance in this technology has been made by introducing a second Bsu36I site downstream of the polyhedrin locus in the baculovirus genome (23). This second site lies in a gene, ORF1629, that is essential for viral replication, and so digestion with Bsu36I yields a linear fragment lacking the essential gene. In theory, co-transfection of this truncated, linearized baculovirus DNA with a 113

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transfer vector that contains sequences complementing the deficiency should produce infectious baculoviral DNA only if the latter has undergone a double crossover event, and has thereby regained the essential gene. Of course, such recombination events will also transfer a foreign gene present in the expression locus of the transfer vector to the viral genome. In contrast, re-circularized baculovirus DNA lacking the essential sequences would not be viable, and so would produce no progeny virus. Several commercially available forms of linearized baculovirus DNA are, in fact, produced from an AcMNPV derivative bearing a third Bsu36I site within a copy of the lacZ gene at the polyhedrin locus (23), to increase the probability of complete linearization upon restriction (e.g. BaculoGold™ linearized baculovirus DNA from Pharmingen, BacPAK™ viral DNA from Clontech, BacVector-1000 Triple Cut Virus DNA from Novagen). By this means, the proportion of recombinant virus obtained following cotranfection of insect cells routinely approaches 100%. 2.3.2 Generation of recombinant baculovirus DNA in microorganisms A second approach for recombinant baculovirus production utilizes the selection of recombinants in microorganisms to circumvent the most technically difficult and time-consuming part of previously described methods, i.e. the detection, isolation, and purification of recombinants from a high-background population of wild-type virus. This approach was originally made possible by the construction of a modified baculovirus genome that could be propagated in the yeast Saccharamyces cerevisiae (24). In this method, yeast cells harbouring the modified baculovirus DNA are transformed with a recombinant transfer vector encoding the cDNA to be expressed. Homologous recombination is particularly efficient in yeast, and recombinants can be readily selected for by virtue of their resistance to the toxic arginine analogue canavanine: recombination results in the loss of ochre suppression of a mutation in the arginine permease gene. The process is rapid due to the fast generation time of yeast compared to insect cells, it is convenient and inexpensive in consumables, and, best of all, it totally obviates the need for plaque purification since the vast majority of canavanineresistant yeast colonies should harbour recombinants. Selected recombinant yeast colonies are cultured, the baculovirus DNA isolated in quantity, and then used directly to transfect insect cells. One aspect of this type of method that makes it particularly powerful is that the use of yeast facilitates the generation of multiple recombinant baculoviruses simultaneously; hence it is possible to study several gene products at the same time, or mutants thereof, for the elucidation of structure-activity relationships. Such undertakings would be very difficult using co-transfection technologies, due to considerations of time and expense. The only minor problems attendant with this system are the low transformation efficiency of yeast and the time-consuming steps required for purifying the recombinant baculovirus DNA from yeast cells before insect cells can be transfected. Recently, this advance in baculovirus expression technology has been taken a 114

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step further by the construction of modified baculovirus derivatives (bacmids) capable of propagation in £. coli (25) rather than in yeast (seeFigure3). The genetic manipulation of £. coli is even more straightforward than that of yeast, and the bacterium grows more quickly (this is particularly pertinent for growth on solid media). A range of antibiotic selections are used to maintain the baculovirus DNA within E. coli and to indicate recombination with the transfer vector. The natural frequency of homologous recombination in this organism is lower than that in S. cerevisioe, but this potential problem is obviated by using transposon sequences to direct the recombination event. The system therefore has all the advantages of the yeast system and is even more rapid and convenient. Since protocols for the use of the yeast baculovirus system have been described elsewhere in this series (26), procedures pertaining to the £. coli baculovirus system, marketed as the BAC-TO-BAC™ Baculovirus Expression System by Life Technologies, will be detailed here instead.

2.4 Heterologous expression of transporters and other membrane proteins in insect cells The baculovirus expression system has been widely used for the overproduction of membrane proteins, and is attractive for a number of reasons. Since insect cells are eukaryotic, they can perform a range of post-translational modifications that may be important for the function of eukaryote membrane proteins, such as glycosylation, palmitoylation, myristoylation, glycosyl phosphatidylinositol (GPI) anchoring, and site-specific proteolysis. The N-glycosylation pathway of Spodoptera frugiperda cells (the most widely used insect host for baculovirus expression systems—see Section 3.1) appears to differ from that of mammalian cells, such that glycoproteins produced in the baculovirus system typically lack complex, biantennary, N-linked oligosaccharide side chains containing penultimate galactose and terminal sialic acid residues. However, cells derived from the salt-marsh caterpillar Estigmene acrea, such as the Ea4 cell line (available from Novagen), are capable of producing complex oligosaccharides and are susceptible to baculovirus infection (27). Moreover, recent studies have demonstrated that metabolic engineering can be used to extend the glycoprotein processing capabilities of other lepidopteran cells, and so in future it may be possible to produce more 'authentic' mammalian glycoproteins in the baculovirus system (28). The limited host range of baculoviruses means that their use poses no hazard to humans. Commercial vectors and kits to facilitate the generation of recombinant baculoviruses are widely available. Moreover, the culture and handling of insect cells is relatively simple and may be scaled-up to fermentor level. It should be noted, however, that whereas polyhedrin can constitute up to 50% of total cellular protein at a late stage of infection of insect cells with wild-type virus, the expectation of similar yields for expressed membrane proteins is usually unrealistic. Just as with other expression systems, the yield of membrane proteins is typically an order of magnitude or more lower than may be 115

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Figure 3 Production of recombinant baculovirus by transposon-mediated recombination in Escherichia coli. To produce recombinant baculovirus in E. coli, the cDNA of interest must first be subcloned into a pFASTBAC™ donor vector downstream of the baculovirus polyhedrin promoter (pPolh). The recombinant plasmid is transformed into DH10BAc™, an E. coli strain containing the baculovirus E co/i-insect cell shuttle vector (bacmid) bMON14272 plus a helper plasmid. Upon transformation, helper plasmid-encoded proteins facilitate the transposition between mini-Tn7 sequences on the vector and the baculovirus DNA, resulting in the formation of recombinant bacmids harbouring the cDNA of interest. Clonal recombinant bacmid DNA prepared from single bacterial colonies can then be used to transfect insect cells, yielding recombinant baculovirus particles suitable for the infection of large-scale insect cell cultures and the consequent production of recombinant protein.

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expected for a soluble protein. Moreover, high levels of protein production are sometimes undermined by a disappointing level of activity, presumably due to an overburdening of the machinery involved in membrane protein insertion into membranes and post-translational modification, or to factors inherent to impending cell death. Future developments of the technology may help to alleviate these problems, but there are no guarantees of success. Though there are exceptions, some general trends have been noted (29), including a reciprocal relationship between expression levels and number of transmembrane domains, which may deter workers attempting to overexpress transport proteins with ten or more transmembrane domains. In addition, proteins that require little post-translational modification tend to be expressed to higher levels. These trends are likely to reflect the difficulty that dying insect cells have in performing the proper folding and processing of unusually large amounts of nascent membrane proteins. The unpredictability of success in the expression of membrane proteins is highlighted by the dramatic expression changes seen between highly related proteins, and even mutants of the same protein (30). Importantly, however, the addition of affinity tags has been reported not to adversely affect expression levels (31-34). The high-level expression of membrane proteins from various systems, including insect cells, has been admirably reviewed elsewhere (29), thus we shall simply touch upon a few relevant highlights amongst the literature. Of particular interest here is the high-level expression of mammalian transporters, especially the passive glucose transporter GLUT1 (35, 36), but also including the serotonin transporter (34) and the Na+-glucose co-transporter SGLT1 (37). While fairly low levels of expression—usually in the five to a few tens of pmol range/mg total membrane protein—have been observed for the expression of a number of membrane receptors and other proteins, in our laboratory the GLUT1 glucose transporter has been expressed at levels of 200 pmol functional protein/mg membrane protein (35). It should be noted, however, that in our experience equal or even greater amounts of non-functional GLUT1 are simultaneously expressed (35). Although the presence of this inactive protein does not interfere with most studies of the functional expressed protein, it might compromise some downstream applications, particularly structural studies. Interestingly, in the case of the cardiac Ca2+-ATPase, not only was the protein reported to be expressed at a high level, corresponding to 20% of the microsomal protein, but at least 70% of the protein was found to be functionally active (38). Similarly, several seven-transmembrane domain type receptors have been functionally expressed using the baculovirus system, and these have subsequently been isolated in quantities potentially sufficient for spectroscopic and/or crystallographic studies. These include the (3-adrenergic receptor (39), muscarinic acetylcholine receptor (40), endothelin B receptor (41), 5-HT2c receptor, and the substance P receptor (42). Various other membrane proteins have also been expressed to very high levels in baculovirus-infected cells, including the gap junction protein (43), prostaglandin synthase (44), aquaporin (45), caveolin (46), and myelin proteolipid protein (47). 117

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3 Practical aspects of the expression procedure 3.1 Introduction Methods used for the expression of proteins in insect cells are described in this section, which includes protocols for the use both of the insect cell co-transfection and the £. coli baculovirus strategies for the production of recombinant baculoviruses. Certain methods, such as cell culture and virus handling and transfection, are common to both, while those unique to one or other strategy will be indicated as appropriate. The insect cells most commonly used as hosts for AcMNPV in baculovirus expression systems originated from the IPLBSF-21 cell line, derived from the pupal ovarian tissue of the fall army worm S.frugiperda. These cells, IPLBSF21-AE (Sf21) and its clonal derivative Sf9, are available from many manufacturers (Sf9 cells are also available from the American Type Culture Collection as ATTC Number CRL-1711). These cells grow well in suspension or monolayer cultures, with a convenient doubling time of approximately 20 hours at 27°C. Sf21 cells are somewhat larger than Sf9 cells and may give higher levels of expression for some proteins (48), although a rigorous comparison of the two cell lines' abilities to express a variety of different membrane transport proteins has not yet been performed. Another alternative for AcMNPV propagation is the cell line BTI-TN-5B1-4, derived from ovarian cells of the cabbage looper Trichoplusia ni and marketed as 'High 5'™ by Invitrogen. These cells are reported to express significantly greater amounts of some secreted recombinant proteins than the S. frugiperda cells (49), although they are somewhat more difficult to adapt to suspension culture, which is the method of choice for large-scale protein production.

3.2 Methods for insect cell culture As mentioned previously, insect cells are quite robust and convenient to maintain in cell culture. They require constant temperature but do not require C02, since their growth media are buffered at a pH of approximately 6.2. Sf9 cells exhibit a doubling time of 18-24 hours in a complete insect cell medium such as Grace's Insect Medium or TC-100, the latter being used routinely in our laboratory. These media must be supplemented with 10% fetal bovine serum (FBS). Serum-free media are available, but are expensive and the cells must be 'adapted' to such media. These media are recommended for simplifying the isolation of recombinant secreted proteins and so are unnecessary in the production of integral membrane proteins. As Sf9 cells adhere relatively loosely to tissue culture vessels, they can be conveniently subcultured without the use of trypsin or other enzymes. The methods described below pertain to the Sf9 line, but also apply to other cell lines, and are modified from Summers and Smith (10). 3.2.1 Monolayer culture Sf9 cells are normally subcultured two or three times per week, by diluting resuspended cells 4- to 8-fold in fresh medium (see Protocol 1). The insect cells do 118

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not become contact-inhibited, and so should be subcultured at 90% conflucncy or below. If cultures arc allowed to overgrow, the proportion of released cells will increase, and such cultures are not optimal for virus replication or protein production. Dying cells can be recognized morphologically by their nonuniform shape and granular appearance, in contrast to healthy cells that are well rounded and appear bright when examined by phase-contrast microscopy.

Protocol 1

Monolayer culture of insect cells Equipment and reagents • Laminar air-flow tissue culture cabinet • 27°C incubator" • Complete TC-tOO medium: TC-100 medium11 supplemented with 10% v/v fetal bovine serum (FBS), 50 units/ml penicillin, and 50 M-g/ml streptomycin; pre-warmed to 27°C

• Haemocytometer • 0.2% (w/vj Trypan Blue (Sigma) in phosphate-buffered saline (PBS) • Tissue culture flasks or plates • Inverted microscope with phase-contrast optics

Method 1. Grow Sf9 cells at 27°C in loosely capped, tissue culture flasks in complete TC-100 medium until approximately 80-90% confluent. Use the microscope to check that the cells are healthy (rounded and shiny; not granular). Do not allow the cells to overgrow the surface of the plate, 2. Discard the medium by aspiration. Add an equal volume of fresh complete medium, then repeatedly and gently tap the sides of the flask or briskly pipette the medium over the surface of the flask to dislodge the cells.c 3. Continue this process until almost all cells are dislodged and visible clumps are dispersed, while avoiding excessive foaming of the medium, 4. Remove a small aliquot and determine the cell number using a haemocytometer. (Optional: To a second aliquot of the cells add an equal volume of 0.2% (w/v> Trypan Blue to determine cell viability,d) 5. Dilute the cells in the appropriate volume of complete TC-100 medium. 6. Seed the diluted cells into tissue culture flasks at the following densities (corresponding to ~ 1:4 dilution): for a 25 cm2 flask—I .5 x 106 cells in 5 ml medium; for a 75 cm2 flask—5 x 106 cells in 15 ml medium; for a 150 cm2 flask—1 x 107 cells in 30 ml medium, 7. Allow the cells to settle and adhere to the bottom of the flask on a flat surface (in the tissue culture cabinet) for approximately 30 min at room temperature.

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8. Replace the flasks in the 27°C incubator until further subculture is required. "Cells will grow satisfactorily between 26 and 28°C, However, for optimum production of recombinant protein by infected cells, cultures should be performed at 27 ± 0.5°C. b Available in powdered or liquid form from many manufacturers, such as Life Technologies. Other commercially available, insect cell culture media may be substituted, e.g. Grace's supplemented medium (Tricfiopliwia ni Medium-Formulation Hink: TNM-FH}, Trypsinization is unnecessary to dislodge Sf9 cells. J Non-viable cells will take up the stain and appear blue. Healthy log-phase cultures should contain > 97% viable, unstained cells. e Dilutions of between 4- and 8-fold can be used, although we prefer the former because excessive dilution can inhibit growth. Since the doubling time for Sf9 cells at 27°C is 18-24 hours, cells diluted 1:4 will require subculturing after approximately 2 days; cells diluted 1:8 will need to be subcultured after 3-4 days.

3.2.2 Suspension culture Sf9 cells grow well in suspension culture and can be transferred from monolayer to suspension culture and back again without adaptation if grown in serum-containing media. Indeed, many workers subculture insect cells in suspension culture routinely (sec Protocol 2), as large quantities of cells am be maintained without consuming high numbers of tissue culture flasks. One important factor in this type of culture is aeration, which is provided by magnetic stirring. However, since too vigorous a stir will produce harmful shearing forces, this limits the culture volume:total volume ratio to be used in spinner flasks (though this is less of a problem in FBS-conLaining media). Proper aeration is particularly important during infection, as this is critical for the efficient production of recombinant protein.

Protocol 2 Suspension culture of Sf9 cells Equipment and reagents • Spinnerflasks"(Techne) • Magnetic stirring systemb (Techne) • 27°C incubator

• Complete TC-100 medium (see Protocol 1) • Sf9 cells from an existing spinner culture or from monolayer culture (see Protocol 1)

Method 1, Seed Sf9 cells at a density of approximately 2.S x 105 cells/ml" into a total volume of 250 ml of complete TC-100 medium in a 1000 ml spinner flask. 2. Incubate the flask at 27°C with constant stirring at 40-80 r.p.m.d until the cell density reaches approximately 2 x 106 cells/ml.

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BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS Protocol 2 continue

3, Subculture the cells by removing 200 ml of the suspension and replacing it with 200 ml of fresh complete medium. Ensure that the cell viability is > 97% by Trypan Blue exclusion (see Protocol 1). 11

Siliconizing spinner flasks with a non-toxic siliconizing agent may minimize the attachment of cells and debris at the medium meniscus. b Choose a magnetic stirring system that produces little or no heat, and which can therefore be located within an incubator. c To avoid a lag phase during the initial establishment of a spinner culture, seed with monolayer-cultured cells at a density of 5 x 105 cells/ml. Once the culture is established. subsequent seeding densities can be reduced to 2.5 x 105 cells/ml. To ensure adequate aeration, the total culture volume should never exceed half the capacity of the spinner flask. d The required stirring speed will depend upon the configuration of the spinner flask. If cell clumping occurs, increase the speed of stirring. The addition of the surfactant Pluronic-F68 (Life Technologies) to a concentration of 0.1% will decrease membrane shearing during stirring and may increase cell viability at higher stirring rates. * Subculturing will be required approximately twice weekly.

3.2.3 Storage and resuscitation of insect cells Sf9 cells may be stored indefinitely in liquid nitrogen if frozen slowly in a medium containing DMSO (see Protocol 3).

Protocol 3 Storage and resuscitation of insect ceils Equipment and reagents • Laminar air-flow tissue culture cabinet • -20°C and-80°C freezers, and liquid nitrogen cell-storage facility • Insulated freezing box (e.g. expanded polystyrene container) • Sterile cryogenic storage vials (e.g. Nunc Cryovials)

• Freezing mix; complete TC-100 medium (see Protocol 1) containing 20% (v/v) DMSO (add DMSO to the medium, rather than vice versa, and mix quickly), chilled to 4°C • Log-phase culture of Sf9 cells (See Protocols \ and 2)

Method 1. Concentrate the Sf9 cells by gentle centriftigation, e.g. at 1000 g4 for 5-10 min in a benchtop centrifuge at room temperature. Discard the supernatant fluid and resuspend the cells in complete medium at a density of approximately 4 x 106 cells/ml.b 2. Add an equal volume of Freezing mix to the cells.

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Protocol 3 continued

3. Rapidly transfer 1 ml aliquots of the cells to cryovials and place the vials in an insulated freezing box. 4. Allow the cells to freeze slowly at -20°C in a freezer for 2 hours, then transfer to a -80°C freezer overnight. 5. Transfer the vials to liquid nitrogen for long-term storage. 6. Resuscitate the cells, either after a few weeks to check viability4 or to establish a new culture, by thawing an aliquot quickly in a water bath at 37°C. 7. Immediately spray the outside of the vial with 70% ethanol to decontaminate, open the vial carefully in the tissue culture cabinet and transfer the cells to a small (25 cm2) flask with 5 ml of pre-warmed complete medium. Allow the cells to attach to the flask by incubating at 27°C for 2 hours. 8. Aspirate the medium to remove the DMSO and dead cells, and replace with 5 ml fresh pre-warmed medium. Incubate the flask at 27°C until the cells have grown to a confluency of 80-90% before subculturing as described in Protocol 1. "Use the minimum time required for complete pelleting, to avoid damage to the fragile cells. b Determine cell density and viability as described in Protocol 1. Only cultures that are at least 95% viable are suitable for freezing. c Work rapidly, because DMSO is cytotoxic! d It is recommended that a vial of cells is tested for viability after a week or two of storage, to check that the freezing procedure has been successful.

3.3 Generation of recombinant bacutovirus DNA The first step in the expression of a heterologous transport protein in insect cells is to clone the corresponding cDNA into an appropriate transfer vector, so as to place it under the control of a suitable baculovirus promoter. The construction of recombinant transfer vectors is performed by standard procedures in £. coli, as described by Sambrook et al (50). Vectors that employ the polyhedrin promoter are described in the protocols that follow. Many such vectors are commercially available (e.g. from Clontcch laboratories, Invitrogcn, Life Technologies, Novagen, Pharmingcn, and Stratagene), some of which allow the expression of modified protein, for example via the addition of terminal oligohistidine peptides, glutathione S-transferase, cellulose binding domains, or other tags for purification purposes. In our own laboratory, we have primarily used the vectors pAcYMl (51), pFASTBAC1, and pFASTBAC HT (Life Technologies) for the expression of mammalian glucose and nucleoside transporters (see figure 4). However, vectors that utilize other promoters are also available, such as those of the very late gene p10, of the late gene encoding the basic protein, and of the immediate-early ie1 gene. As described in Section 23 use of these vectors, available for example from Quantum Biotechnologies (e.g. pTen 12, pTen21), Pharmingen (e.g. pAcMP2, pAcMP3), and Novagen (pAcPIE l vectors) 122

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Tn7R

Figure 4 Examples of vectors employed for the production of recombinant baculoviruses by the insect cell co-transfection and E. coli baculovirus technologies. (Top) The transfer vector pAcYMl was one of the first widely used vectors for co-transfection with baculovirus DNA. Sequences derived from the plasmid pUC8 (hatched) enable replication and ampicillin selection (AmpR) within £. coli. The remainder of the vector (white) is derived from AcMNPV, enabling homologous recombination with linearized baculovirus DNA and the expression of foreign genes under the control of the polyhedrin promoter (pPolh). The vector contains a unique SamHI site for the insertion of foreign DNA. (Bottom) The donor vector pFastBac HTa (Life Technologies) is designed for the E. coli baculovirus system. The vector contains sequences for replication (ori) and ampicillin selection (Amp?) in E, coli. and a polyhedrin promoter to allow protein expression in insect cells. Downstream of this promoter there is an extensive multiple cloning site (MCS) that also encodes an amino-terminal, hexa-histidine tag lollowed by a protease cleavage site to enable lag removal. The polyhedrin promoter region, together with a gentamicin resistance gene (GmRl and a SV40 polyA signal, form an expression cassette bounded by the left and right arms {Tn7L and TnTRl of a mini Tn7 transposon element. The latter allows transposition of the recomPinant gene cassette into a bacmid propagated in £. coli strain DHlOBAc1" (Life Technologies). The GmR gene enables The selection of recombinant bacmid-containing colonies after transposition.

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may have advantages over polyhedrin based systems, in particular with respect to the yield of biologically active expressed protein. As described in Section 2.3, following construction of a recombinant transfer vector, there are two major routes that can be followed for generating recombinant baculoviruses. The most widely used method involves the co-transfection of insect cells with transfer vector and linearized baculovirus DNA. Recombinants are generated by homologous recombination between these two species within the insect cell (see Protocol 6). More recently, methods have been introduced which involve recombination between baculoviral DNA and transfer vector within microorganisms, either E. coli (see Protocol 4) or yeast (24, 26). Each of the two routes has advantages. For example, a much wider range of vectors, bearing different promoters and encoding a variety of tags, can be employed in the insect cell co-transfection approach, while the preparation of recombinants in E. coli obviates the need for plaque purification of recombinants and so is a much faster process. Both methods are therefore described in detail below. 3.3.1 Transposition of recombinant genes into baculovirus DNA propagated within E. coli The system for the production of recombinant baculovirus in £. coli was developed by Luckow et al. (25) and is available commercially as the BAC-TO-BAC™ Baculovirus Expression System from Life Technologies. We have used it successfully for the rapid production of recombinant baculoviruses encoding mammalian equilibrative nucleoside transporters (hENTl and rENTl) and the human glucose transporter GLUT1. The rapidity with which recombinants can be produced renders this system particularly useful for screening large numbers of transporter mutants generated by site-directed mutagenesis (52). The first step is to construct a recombinant donor vector bearing the cDNA of interest using a pFASiBAC™ (pMON14327 derivative) donor vector (see Figure 4). This donor vector bears an ampicillin resistance gene for selection in E. coli, plus an expression cassette consisting of a gentamicin resistance gene (GmR), the polyhedrin promoter, a multiple cloning site, and an SV40 polyA signal inserted between the left and right arms of the bacterial transposon Tn7. The recombinant vector is then transformed into E. coli DHIOBAC™, which contains the baculovirus E. coli-insect cell shuttle vector (bacmid) bMON14272. The bacmid is maintained in E. coli by virtue of its possession of a kanamycin resistance gene and contains a segment of DNA encoding the LacZa peptide, allowing the bacmid to complement a lacZ deletion on the chromosome to form blue colonies in the presence of a chromogenic substrate such as X-gal when induced with isopropylp-D-thiogalactoside (IPTG). Within the 5' end of the lacZa gene is a short segment containing the attachment site for Tn7 (mini-attTn7) that does not disrupt the reading frame of the LacZa peptide. Tn7 transposition functions are provided in trans by a helper plasmid (pMON7124) also harboured by E. coli DHIOBAC™. Following transformation with a recombinant donor plasmid (selected for using 124

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gentamicin), transposition of the mini-Tn7 element bearing the foreign gene then occurs. Insertion of the element into the mini-uU'fn? attachment site on the bacmid disrupts expression of the LacZex peptide, such that colonies harbouring recombinant bacmids arc white rather than blue following induction with IPTG and treatment with X-gal. Selection of recombinants is therefore a relatively simple procedure.

Protocol 4 Transposition of recombtnant genes into bacmid DNA Reagents • DH10BAC™ competent cells" (Life Technologies) • Recombinant pFASTBAC™ vector bearing the cDNA of interest • SOC medium: Dissolve 2 g bactotryptone and 0.55 g yeast extract in 97 ml H2O, add 1 ml of 1 M NaCl and 1 ml of 1 M KC1, then autoclave. After cooling, add 0.5 ml 1 M MgCl2, 0.5 ml 1 M MgSO4, and 1 ml 2 M glucose. Filter-sterilize. • Antibiotic stock solutions: kanamycin (10 mg/ml in H2O, filter-sterilized); gentamicin (7 mg/ml in H20, filter sterilized); tetracycline (10 mg/ml in ethanol). Store all at -20°C

• Luria-Bertani (LB} medium: 10 g bactotryptone, 5 g yeast extract, 10 g NaCl per litre, adjusted to pH 7.0 with a few drops of 5 M NaOH and then autoclaved • X-gal stock solution: 20 mg/ml in dimethylformamide. Store at -20°C, protected from light, • IPTG stock solution: 200 mg/ml in H20. filter-sterilized. Store at -20°C. • LBagar plates supplemented with antibiotics: 1,5% bacto-agar in LB medium containing 50 ug/ml kanamycin, 7 gentamicin, 10 ug/ml tetracycline, 100 ug/ml X-gal,b and 40

Method 1. Thaw the bacmid-containing competent cells (DH10BAC™) on ice; use 20 ul per transposition' in a 1.5 ml microcentrifuge tube, 2. Add 10-50 ng recombinant donor plasmid to the cells and gently mix by tapping the tube. 3. Incubate the mixture on ice for 30 min. 4. Heat-shock the cells by incubation in a water bath at 42 °C for 45 sec. 5. Chill the cells on ice for 2 min. 6. Add 980 ul pre-warmed (37°C) SOC medium to the cells. 7. Allow the cells to recover in a shaking incubator (220 r.p.m.) at 37°C for 4-8 hours.* 8. Serially dilute an aliquot of the cells in SOC medium to give 10-1 and 10-2 dilutions. 9. Spread 100 ul diluted and undiluted cells on to the LB agar plates supplemented with antibiotics as indicated above.

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GARY J. LITHERLAND AND STEPHEN A. BALDWIN Protocol 4 continued

10. Incubate the plates for 36 hours at 37°C. Select white colonies harbouring recombinant bacmids/ "These cells contain the bacmid bMON14272 and the helper plasmid pMON7124, b An alternative, which is reported to yield more intense blue staining. is S-bromo-3-indolyl-BD-galactoside (Bluo-gal). c The suppliers recommend 100 ul cells, but 20 ul should be sufficient if transposing with ;> 10 ng vector, d We have observed that extended (> 4 hours) recovery times may be necessary for good transposition efficiency. f Further dilutions may be necessary if the number of colonies on the plates is too high to allow good growth and thus proper screening. ^Perhaps because the cells must replicate the large bacmid molecule, they grow relatively slowly. We have found that incubating the cells for at least 36 hours at 37°C is necessary to enable easy identification of blue colonies, the colour of which appears most intense in the centre of larger colonies. The suppliers of the system advocate re-streaking white colonies on to fresh plates to confirm their phenotype, but the delay caused by this precaution is probably unnecessary provided that the colonies are allowed to grow to a large size on the master plate. Usually, 10-25% of the colonies will harbour recombinants. Once rccombinant clones arc identified, the next step is to isolate the recombinant baculovirus DNA. This is done using a version of the alkali lysis plasmid mini-prep method, modified for the isolation of very large plasmids (see Protocol 5).

Protocol 5 Isolation of recomblnant bacmid DNA Reagents • LB medium (see Protocol 4} supplemented with 50 ug/ml kanamycin, 7 ug/ml gentamicin, and 10 ug/ml tetracycline • Solution 1:15 mM Tris-HCl pH 8.0.10 mM EDTA.lOO^g/mfRNaseA

• Solution II: 0,2 M NaOH, 1% (w/v) SDS • 3 M potassium acetate pH 5.5 • TE buffer; 10 mM Tris-HCl pH 8.0.1 mM EDTA

Method 1. Following Protocol 4, select large white colonies (approximately 3 mm in diameter) from a plate with around 100 colonies, Ensure that white and blue colonies can be clearly distinguished. 2. From single white colonies set up 2 ml cultures in LB medium, supplemented with antibiotics, from which to isolate bacmid DNA. Incubate with shaking at 220 r.p.m. at 37°C until stationary phase is reached (this may take up to 24 hours).

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BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS Protocol 5 continued

3. Transfer the cultures to 1.5 ml microcentrifuge tubes and sediment the cells in a microcentrifuge at 13 000 g for l min. 4. Aspirate the supernatant fluid and resuspend the cells (by pipette) in 0.3 ml of Solution I. Add 0.3 ml of Solution II and gently mix. Incubate the samples at room temperature for 5 min, until reduced turbidity of the sample indicates that most of the cells are lysed. 5. Add 0.3 ml of 3 M potassium acetate dropwise, mixing gently. Incubate the samples on ice for 10 min. 6. Sediment the precipitate for 10 min at 13000 g in a microcentrifuge. Carefully transfer the supernatant fluid to a fresh tube containing 0.8 ml of isopropanol. Mix the sample by inversion and incubate on ice for 10 min. 7. Centrifuge the sample for 15 min in a microcentrifuge at 13 000 gat room temperature. 8. Discard the supernatant fluid and wash the pellet with 0,5 ml of 70% (v/v) ethanol, inverting the tube. Centrifuge the sample for 5 min at 13000 g at room temperature, 9. Carefully aspirate the supernatant, removing as much as possible. Allow the pellet to dry in air for up to 10 min. 10. Add 40 ul TE buffer to the pellet. Do not mix by tip or vortex, but allow the pellet to slowly dissolve, tapping the tube occasionally and gently. The DNA should be dissolved within 10 min. 11. Dispense the DNA into 5 ul aliquots and store at -20 °C,

Following the preparation of the recombinant E. coli-insect cell shuttle vector, the clonal bacmid DNA may be used directly to transfect Sf9 cells, in order to generate infectious virus particles. The protocol for this procedure is very similar to that used for the preparation of recombinant baculovirus by cotransfection of insect cells with conventional transfer vectors plus linearized baculovirus DNA, although the titre of virus resulting from the initial transfection of cells with bacmids (2-4 x 107 p.f,u./ml) is likely to be much higher than that resulting from co-transfection experiments. A common protocol for both procedures is therefore described below (see Protocol 6).

3.3,2 Co-transfection of insect cells with baculovirus DNA and transfer vector As described in detail in Section 2.3.1, recombinant baculoviruses can be produced by homologous recombination following the co-transfection of insect cells with a recombinant transfer vector and (usually) linearized baculovirus DNA. Transfection of insect cells is performed in the same manner as for mammalian and other cultured cells, i.e. by forming a complex between the 127

GARY j. LITHERLAND AND STEPHEN A. BALDWIN

Protocol 6 Transfection and co-transfection of insect cells with baculovirus DNA using liposomal transfectlon reagents Equipment and reagents • 1 mg/ml suspension of D0TAP liposomes in Mes-buffered saline pH 6.0 (BoehringerMannheim)n • Solution A: (for each transfection) 5 ul of recombinant bacmid DNA (Protocol 5) in 100 ul TC-100 medium without FBS or antibiotics, or 1 ug of recombinant transfer vector plus 200 ng linearized baculovirus DNA in 100 ul TC-100 medium without FBS or antibiotics

• Solution B: (for each transfection) 6 ul of DOTAP in 100 ul TC-100 medium without FBS or antibiotics • TC-100 medium without FBS or antibiotics • Recombinant bacmid DNA (see Protocol 5) or linearized baculovirus DNA (e.g. Baculogold™ DNA, Pharmingen} and recombinant transfer vector • For other materials, see Pratocok 1 and 2

Methods 1. Seed approximately t x 106 Sf9 cells, from a log-phase culture, per 35-mm tissue culture dish, or into each well of a 6-well plate, in 2 ml of complete TC-100 medium. Allow the cells to attach to the surface for 1 hour at 27°C. 2. Gently mix Solutions A and B in a sterile tube, then incubate for approximately 30 min at room temperature. 3. For each transfection, add 0.8 ml of TC-100 medium without FBS or antibiotics to each 0.2 ml sample of DOTAP-DNA mixture following the incubation in step 2, and gently mix. 4. Wash the cells in each dish or well with 2 ml TC-100 medium without FBS or antibiotics. After aspirating the wash medium add, to each dish, 1 ml of the diluted lipid-DNA complex prepared in step 3. 5. Incubate the cells for 5-8 hours at 27°C without shaking. 6. Aspirate the transfection mixtures and replace with 2 ml of complete TC-100 medium. Incubate the cells for 72 hours at 27°C without shaking. 7. Decant the medium and centrifuge at 1000 g for 5 min to clarify the viruscontaining supernatant fluid. Store the resultant virus stock at 4°C prior to plaque or cell lysis assay (see Protocols 8 and 9), amplification (see Protocol 10), or infection of insect cells to investigate transporter expression (see Section 3.4),b °0r alternative liposomal transfection reagent. ''If recombinant bacmid DNA was used for transfection, the viral titre may be sufficient for use without amplification, although in order to perform infection experiments at a known multiplicity of infection (m.o.i.) it will be necessary to determine the viral titre by plaque assay or other means. If viruses have been produced by co-transfection of transfer vector and linearized baculovirus DNA, amplification will usually be necessary, and plaque purification is recommended to ensure that a single recombinant clone is present.

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DNA of interest and a liposonud transfection reagent, and then incubating the cells with this complex (see Protocol 6). There are many sources of transfection reagents available, most of which will efficiently facilitate transfection of insect cells. Some suppliers recommend products such as the CEU.FECTIN™ reagent (Life Technologies), and procedures optimized for the trans feet ion of insect cells. However, we routinely use cationic liposomes prepared from N-|l-(2,3dioleoyloxy|propyl]-N,,V,N-tnmethylammonium methylsulfate (DOTAP) for transfectioii into a range of eukaryotic cell types, including insect cells. In addition, many laboratories use the calcium phosphate transfection technique with success (see Protocol 7).

Protocol 7 Co-transfection of insect cells using calcium phosphate Equipment and reagents • Transfection buffer 25 mMHepespH 7.1, 140 mM NaCl, 125 mM CaCl2, filtersterilized

Linearized baculovirus DNA {e.g. Baculogold™ DNA, Pharmingen) and recombinant transfer vector For other materials, see Protocols 1 and 2

Method 1. Seed approximately 1 x 106 Sf9 cells (from a log-phase culture) per 35-mm tissue culture dish, or into each well of a 6-well plate, in 2 ml of complete TC-100 medium. Allow the cells to attach to the surface for 1 hour at 27 °C. 2. Aspirate the medium from each well and replace with 1 ml of fresh complete TC100 medium. Leave the cells at room temperature, 3. Mix 1 ml of the Transfection buffer with 200 ng of linearized baculovirus DNA and 2 ug recombinant transfer vector in a sterile microcentrifuge tube. Add the mixture dropwise while swirling the medium in the wells," 4. Incubate the plates for 5 hours at 27 °C without shaking. 5. Aspirate the medium from the cells, wash with 2 ml of complete TC-100 medium and finally replace with 2 ml of complete medium, 6. Incubate the cells at 27°C for 4-5 days without shaking,b 7. Transfer the medium to a sterile centrifuge tube and centrifuge at 1000 g for 5 min to clarify the virus-containing supernatant fluid. Store the resultant virus stock at 4°C prior to plaque assay (see Protocol 8) or amplification (see Protocol 10). " A precipitate of calcium phosphate will be observed upon mixing the calcium-containing Transfection buffer with the phosphate-containing medium. ''Calcium-phosphate transfection is less efficient than the use of lipid reagents, thus a longer incubation is necessary for an adequate virus yield.

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The alternative to the use of liposomal transfection reagents is transfection of Sf9 cells by calcium phosphate treatment (see Protocol 7). This method is quite satisfactory but is less efficient than the use of liposomes. For this reason, it is necessary to allow secondary infection of the cell culture in order to obtain a sufficient viral yield. Consequently, the protocol takes approximately 48 hours longer to complete than that described above (see Protocol 6). 3.3.3 Baculovirus handling, amplification, and purification Protocols 6 and 7 should result in the production of recombinant baculovirus particles in the insect cells, and their release into the medium by budding. The resultant viral titre will depend largely upon the efficiency of transfection, but is unlikely to be sufficient for the purposes of transport protein production. Thus the viral stocks must be titred (see Protocols 8 and 9) and amplified (see Protocol 10). Knowledge of the titre of amplified stocks is also essential to optimize the production of recombinant protein, because the m.o.i. of insect cells influences the extent and time course of protein expression (see Section 3.4). In the case of recombinants that have been produced by transposition in E. coli (see Protocol 4) a simple cell-lysis assay, such as that described in Protocol 9, may be sufficient to determine the titre. Because all virus generated from transfection with recombinant bacmid DNA will be recombinant, there will be no contamination with wild-type virus and hence no need for purification. However, if the baculoviruses have been produced by co-transfection methods, even the use of linearized baculovirus DNA bearing a lethal deletion cannot be guaranteed to produce 100% recombinants. Thus a more laborious and technically demanding procedure termed the plaque assay (see Protocol 8) is required to identify and purify clonal stocks of recombinant viruses, in addition to being of use in determining viral titre. In the plaque assay, cell monolayers are infected with a low ratio of virus particles, such that only a few cells become infected. The cells are then overlaid with agarose so that the subsequent spread of virus is limited. Thus, when an infected cell lyses, only the immediately neighbouring cells become infected. After several cycles of infection, the original site of infection is surrounded by a group of lysed cells termed a plaque, which is visually distinguishable from the surrounding, healthy cells. Since each plaque originates from a single baculovirus, the number of plaques present can be used to determine the viral titre of the stock solution (plaque-forming units (p.f.u.)/ml) and clonal populations of virus can be prepared by isolating individual plaques. If plaque purification of recombinant viruses is unnecessary, simpler and more rapid methods can be used to estimate the titre of baculovirus stocks. An expensive, though elegant and rapid, approach is provided by the BacPAK™ Baculovirus Rapid Titer Kit marketed by Clontech, which exploits the early expression of the AcMNPV envelope glycoprotein gp64 to allow visualization of infected cells. The cell-lysis assay (a modified end-point dilution method (3)) described below (see Protocol 9) is less rapid, but provides an approximate measure of viral titre (satisfactory for the optimization of protein expression) 130

BACULOVIRUS-MED1ATED OVEREXPRESSION OF TRANSPORT PROTEINS

Protocol 8 Measurement of viral titre, and purification of recomblnants, by plaque assay Equipment and reagents • Exponentially growing culture of Sf9 cells in complete TC-100 medium at 5 x 10s cells/ml (30 ml per titration) • 35-mm tissue culture dishes or 6-weIl plates • Water bathsetat40°C

• Low melting-point agarose" • Neutral Red (Sigma) solution: 0.33%, sterile stock solution in PBS (pH 7.3) • 0.5 ml of each clarified baculovirus supernatant to be titred • For other materials see Protocols 1 and 2

A. Assay of viral titre 1. To each tissue culture dish or well add 2 ml of the Sf9 cell suspension, i.e. 106 cells." Gently rock to ensure an even distribution of the cells. Note that 12 dishes will be required per virion stock that is being titred. 2. Incubate for 1 h at 27°C to allow the cells to settle and attach. 3. Prepare 10-fold serial dilutions of the virus stock to be titred, e.g. by sequentially diluting 0.5 ml samples into 4.5 ml complete TC-100 medium, resulting in dilutions from 10-1 to 10-8. 4. Sequentially remove the medium from each well and immediately replace with 1 ml of the appropriate dilution of virus. Assay duplicate samples of each dilution. from 10-3 to 10~8, i.e. 12 dishes or wells per stock to be titred. 5. Incubate for 1 hour at room temperature to allow the virus to adsorb to the cells. 6. During step 5, resuspend 1.5 g of low melting-point agarose in 50 ml distilled water in a small bottle. Autoclave it for 15 min then cool to 40 °C. Warm 50 ml complete TC-100 medium to 40 °C, then mix with the agarose to give a 1.5% agarose solution. Store in the water bath at 40 °C until required. 7. After the 1-hour virus incubation, carefully aspirate the medium from the cells, starting with the highest dilution of the virus stock, working quickly to avoid desiccation of the cells. Then gently overlay with 2 ml of the agarose solution per well, running it down the edge of the well and taking care not to disturb the cell monolayer. 8. Leave the dishes undisturbed for 1 h at room temperature to allow the overlay to solidify before adding 1 ml of complete TC-100 medium to each dish. 9. Incubate the plates at 27°C in a humidified incubator" for 4-10 days. Milky plaques of slight contrast are formed by recombinant virus. 10. Monitor the plates each day until the number of plaques does not increase for two consecutive days. (Optional: dilute the Neutral Red stock solution to 0.03% by adding 1 ml to 10 ml of sterile PBS (pH 7.3) just before use." Add 1 ml of the diluted stock to each dish, then incubate at 27°C for 3 h. Aspirate the stain, invert the dishes then leave in the dark overnight at room temperature to make the plaques more visible,")

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Protocol 8 continued

11. Choose appropriate duplicate wells from which to count the plaques, A suitable number would be from 5 to 20 plaques per well, 12. Calculate the titre (p.f.u./ml) in the original, undiluted viral stock as follows: p.f.u./ml = average number of plaques per well x (dilution factor)"1.

B. Preparation of clonal virus stocks 1. In wells containing just a few plaques, mark well-isolated ones by circling with a pen on the underside of the plate. 2. Pick about H/of the marked plaques by pushing a sterile Pasteur pipette through the agarose into the plaque, and gently sucking an agarose plug into the pipette tip. 3. Transfer each plug into 1 ml of complete TC-100 medium in a separate microcentrifuge tube. Vortex gently, and leave overnight at 4°C to elute the virus particles from the agarose, 4. Seed 35-mm dishes or wells with 5 x 10s Sf9 cells in 2 ml of Complete TC-100. 5. Incubate for 1 h at 27 °C to allow the cells to settle and attach. 6. Aspirate the medium, then gently add 100 ^1 of the eluted virus suspension from step B3 to the middle of the dish, 7. Incubate at room temperature for 1 h, then add 2 ml of complete TC-100 medium. 8. Incubate at 27°C for 3-4 days, then transfer the medium to a sterile centrifuge tube and centrifuge at 1000 g for 5 min to clarify the virus-containing supernatant fluid. Store the resultant virus stock at 4°C prior to titre determination or amplification, 9. Perform a Western blot assay on the cells remaining from step B8 to confirm expression of the desired recombinant protein, and thus that the chosen plaque contained recombinant baculovirus. "Commercial 'DNA grade' agaroses often contain contaminants that are toxic to insect cells, and so agarose certified for use in cell overlays should be employed, e.g. Sea-Plaque (FMC Bioproducts} or BacPlaque (Novagen) agarose. b The cells should be about 50% confluent. c If a humidified incubator is not available, the dishes can be sealed in a plastic storage box containing a moist paper towel to minimize medium evaporation. d Do not store the diluted stain—it is light-sensitive and will come out of solution. 'Neutral red stain is taken up and thus stains living cells, but not dead cells. Plaques should therefore appear as clear circles against a pink or red background. ^If linearized baculovirus DNA bearing a lethal deletion is used in transfection experiments, the vast majority of plaques should contain recombinant baculovirus. The identity of individual clones as recombinants can be assessed as described in step B9. Alternatively, in those instances where a lacZ gene in the parental virus is replaced by the foreign gene upon recombination, recombinants can be identified by blue/white selection, as described elsewhere in this series {26}.

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Protocol 9 Estimation of viral tltre by cell-lysis assay Equipment and reagents * HLA assay plates (Nunc)

* For other materials, see Protocols 1 and 2

• Cryovials (Nunc)

Method 1. Harvest Sf9 cells from an exponentially growing culture, sediment at 1000 g for 5 min at room temperature, and resuspend gently at a density of 0.25 x 10s cells/ml in complete TC-100 medium, 2. Serially dilute a sample of viral stock, making dilutions (200 ^1 volume) in com* plete TC-100 medium from 10~l through to 10-10 in sterile Nunc cryovials. 3. Add 1 volume of cell suspension to each virus dilution and mix gently. 4. Add triplicate 10 ul aliquots of each mixture to wells in each row of a Nunc HLA plate. This should result in approximately 50% confluency after settling, adjust the cell resuspension density if this is not the case, 5. Add medium to the empty wells on the plate, or a drop (100 ul} in each corner, to maintain humidity within the plate. 6. Replace the lid of the plate and place in a sealed humid box, inside a 27°C incubator. 7. Incubate the plate at 27°C for 6 days and observe the extent of cell lysis, 8. Observe the cells each day until no further increase in cell lysis is observed." "At the end of the experiment there should be a clear difference in the appearance of cells between two adjacent rows, indicating the dilution below which there is no vhion present in the sample added to the cells. Healthy cells are of a relatively uniform size and appear rounded and shiny. Following infection, cells become swollen and misshapen, and exhibit a more granular appearance before finally lysing. This can be used to approximate the viral titre using the following simple approach: e.g. consider the situation where there is lysis in all wells of dilution 10-(x-1), but not in all of 10-x. For this to occur, on average there must be less than 1 virus particle in 10 ul of the 10-x dilution, but more than 0.1 particles. Correspondingly, there must be less than t x 10x virions in 10 M-I of the mixture of original virus stock plus cell suspension, but more than 1 x 10(x-1), It follows therefore that there are less than 2 X 1 0 " particles in 10 ul of the original virus stock, and more than 2 x 10 (x-1) . Thus the titre lies between 2 x 10(x-1) and 2 x 101*+ *' virus particles/ml. in a format less technically demanding and labour-intensive than the plaque assay. Amplification of a baculovirus stock is a matter of simply infecting a culture of Sf9 cells at a known titre and harvesting the virus-containing media between 3 and 5 days later, In this procedure, however, it is important to use a low m.o.i. to avoid the accumulation of viruses with defective genomes. When it is

GARY J. LITHERIAND AND STEPHEN A. BALDWIN

established by titre assay that a sufficient viral titre has been attained, the viral stock maybe stored for future infections at 4°Cin the short term, and at -80 "C in the long term. Frozen or refrigerated stocks show little lass in infectivity, so long as they are protected from light (53),

Protocol 10 Amplification and storage of recombinant baculovirus Equipment and reagents • For materials, see Protocols 1 and 2

Method 1. Transfer virus-containing supernatant fluid to a sterile tube. Clarify the fluid by centrifuging at 1000 g for 5 min at room temperature. Transfer the clarified medium, which may be sterile-filtered, to fresh tubes and determine the titre (see Protocols 8 or 9). 2. To amplify the virus, infect a monolayer exponential culture of Sf9 cells (see Protocol 1} at an m.o.i. of 0.01-0.1." Harvest the virus after 72-120 hours. Note that this should result in at least a 100-fold amplification that may yield virion at a high enough concentration for assaying protein expression. Infect larger Sf9 cultures if a larger stock is required. 3. Store the virus at 4°C in the dark (in the short to intermediate term).11 0

m.o.i. = the ratio of infective virion to Sf5 cells. To estimate the viral inoculum required use the following formula: inoculum required (ml) = (m.o.i. required (p.f.u./cell)) x (total cell number)/ (viral titre (p,f.u./ml)). 11 An aliquot can be stored in the long term at -80°C if FBS is present in the medium (at least 2% (vfv)). It is advisable to store an aliquot of virus at -80'C in case of stock contamination.

3.4 Baculovirus-mediated heterologous expression of transport proteins 3.4.1 Optimization of infection protocols to maximize protein expression When viral stocks of sufficiently high titre have been obtained, they can be used to infect cells for expression of transport proteins. As with any oilier expression system, the amount of transport protein produced by bactilovirusinfected insect cells is dependent on several parameters, which need to be optimized for each particular protein. The most important of these parameters are the growth phase of the cells prior to infection, the multiplicity of infection (m.o.i.) used, and the time (post-infection) of harvest (54). Monolayer cultures should be infected before they are confluent, i.e. flasks or plates should infected

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

shortly (1-2 h) after seeding with between 1 and 1.2 X 106 cells (ideally from an exponentially growing suspension culture, see Protocol 2) per cm2. If cells are to be infected in suspension cultures, these should be infected in the early exponential growth phase (i.e. at approximately 106 cells/ml). The optimum m.o.i. must be empirically tested, but values between 3 and 10 are usually required to achieve the simultaneous infection of all cells in the culture. Similarly, the optimum time post-infection to harvest cells will depend on the properties of the protein being expressed, including its susceptibility to proteolytic degradation, and so again it must be empirically tested by taking samples at various times following infection. Such testing requires a means of detecting expressed transport protein, as described in the following sections. 3.4.2 Irmmunological detection of expressed transport proteins While soluble proteins may be produced using the baculovirus expression system in amounts sufficient for their detection and quantification on Coomassie blue-stained gels of insect cell extracts, this is usually not the case for transport proteins. However, expressed protein can readily be detected by standard Western blotting procedures (55) using transporter-specific antibodies raised, for example, against synthetic peptides. Alternatively, commercially available antibodies against oligopeptides, such as hexahistidine tags, introduced into the protein by recombinant DNA technology, may be usefully employed. Simple dot blots of infected cell lysates may be sufficient to follow the time course of protein expression following infection, but will not enable investigation of possible transporter degradation at later stages after infection. For this reason it is preferable to perform Western blotting of samples following SDS-PAGE. In preparing samples for such analysis, care must be taken to avoid artefacts resulting from proteolytic degradation in the gel sample buffer, which has been reported to activate the viral cysteine protease V-CATH (56). This is likely to be an especial problem for the analysis of membrane proteins, where boiling gel samples (which will inactivate the protease) is often avoided to prevent protein aggregation. Addition of a cysteine protease inhibitor such as trans-epoxysuccinylL-leucylamido-(4-guanidino)butane (E-64) to gel samples has been reported to prevent such degradation (56). 3.4.3 Functional assay of expressed transport proteins While Western blotting can provide an index of protein expression in the baculovirus system, many studies have revealed that only a proportion of the expressed protein may be functional. The remaining protein is presumably inactive as a result of misfolding or aggregation. Monitoring transporter protein function during the time course of protein expression is therefore of great importance. If a tight-binding ligand is available in radiolabelled form, equilibrium dialysis or other techniques can be employed as a means of quantifying functional protein in membrane preparations (see Protocol 12) from infected insect cells. Such assays have the advantage over transport assays in that they do not rely upon the integrity of the insect cell membrane, which is known to 135

GARY J. LITHERLAND AND STEPHEN A. BALDWIN

become 'leaky' during the later stages of the infection time course. For example, we have successfully used this approach to quantity expression of the glucose transporter GLUT! using the ligand cytochalasin B (35). If a tight-binding transporter ligand is not available, a transport assay measuring inward flux of the radiolabelled substrate is an alternative method of assessing the functionality of expressed protein. Protocol J1 describes a typical transport assay for use with insect cells (.'57), However, there are several caveats associated with the use of such transport assays. First, as in many other cell

Protocol 11 Measurement of solute transport into insect cells Equipment and reagents • Microcentriftige (Beckman Instruments) • Silicone-paraffin oil mixture (d = 1,032 g/ml) (see Chapter 1, Protocol 2) • Transport buffer: e.g. TC-100 medium or other isotonic buffer"

Radiolabelled substrate: typically3H- or 14 C-labelled at 10 |ACi/ml in Transport buffer" For other materials, see Protocols 1 and 2

Method 1. Harvest infected SfS cultures at the required time by sedimentation at 1000 g for 5 min at room temperature. 2. Wash the cells three times (remove an aliquot for viable cell counting (see Protocol 1)) and resuspend in Transport buffer at 5 x io* cells/ml. 3. Layer 100 jJ of a radiolabelled transport substrate" in Transport Buffer over 200 ul of the silicone-paraffin oil mixture in 1.5 ml raicrocentrifuge tubes. 4. Start the assay by adding 100 ul of the suspension of washed insect cells to the substrate in the tube. 5. Terminate the assay after the appropriate time" by centrifuging the cells through the oil layer for 30 sec at 12 000 g, to separate them from the substrate. 6. Carefully remove the top aqueous layer from the tube, wash the sides of the tube gently with Transport buffer, and remove the wash—without disturbing the oil layer or the cells beneath. Then remove most of the oil layer, without disturbing the cell pellet, 7. Remove the cells from the tube by solubilizing in 100 ul of 1 M NaOH and determine the uptake of radioactivity by liquid scintillation counting. "The buffer chosen, the concentration of the substrate used, its specific radioactivity, and the assay time will obviously depend upon the transport system under investigation, and must be established empirically. For additional discussion of transport assays of this type, including the use of inhibitor-containing 'stop sortitions' and the estimation of trapped extracellular radioactivity, see Chapter 1, Section 3,3.

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BACULOVIRLJS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

types, the endogenous transport activity of the host insect cells may mask that of the heterologously expressed protein, particularly during the early stages of the infection time course, when little transport protein has yet been expressed. Second, the promoters usually employed in baculo virus expression systems, such as the polyhedrin promoter, act at a late stage during the infection process, which results in eventual cell lysis. The insect cell membranes become 'leaky' prior to such lysis, and this phenomenon, coinciding with the likely time of maximum heterologous protein expression, may interfere with transport measurements. For these reasons, the rirnmg (post-infection) of the assay may be crucial, and the optimum time may be different for each expressed transporter. It is important to keep other parameters, particularly m.o.i,, constant between experiments.

3.5 Large-scale production of recombinant protein from insect cells Once the optimal conditions for the expression of functional transporters have been determined, the system may be scaled up to produce sufficient protein for purification and/or structural analyses. This may feasibly be clone using either monolayer cultures or suspension cultures in spinner flasks (sec Protocol 2). While the latter may be more cost-effective, scale-up from small-scale cultures may require further optimization of parameters (e.g. aeration) to produce the maximum yield of functional protein. Whichever culture method is used, the next step will be the large-scale isolation of membranes as a starting material for protein purification or other experiments, A suitable method for such preparation is described in Protocol 12.

Protocol 12 Preparation of Insect cell membranes Equipment and reagents

• Lysis buffer: 50 mM sodium phosphate pH 7.4.100 mM NaCl, 1 mM EDTA, 0.2 mM 4(2-aminoethyl)-benzenesulfbnylfluoride (AEBSF), 10 >iM leupeptin, 1 (uM pepstatin. 1 mM benzamjdine, 1 [ig/ml aprotinin

Parr celt for nitrogen cavitation Benchtop ultracentrifuge (e.g. Eeckman) and Beckman TLA100.3 rotor or equivalent

Method NB: Cany out steps 3-7 at 4°C. 1. Harvest Sf9 cells (107 cells from a suspension culture) by centrifugatton at 1000 g for 5 min and discard the supernatant fluid. 2. Resuspend the cells in 3 ml of ice-cold Lysis buffer.

GARY J. L1THERLAND AND STEPHEN A. BALDWIN Protocol 12 continued

3. Disrupt the cells using a Parr cell. Expose the cells to 800 p.s.i. under nitrogen for 10 min, Lyse the cells by explosive decompression. Repeat this step once {optional} to provide a greater proportion of lysed cells. 4. Sediment the unbroken cells and debris by centrifugation at 1000 g for 10 min, 5. Transfer the supernatant fluid to ultracentrifuge tubes and sediment the membranes by centrirugation at 100000 g for 1 hour at 4°Cusing a Beckman TLA100.3 rotor or equivalent. 6. Gently resuspend the membranes in 300 |u,l of Lysis buffer using a pipette tip, avoid frothing. 7. Estimate the protein concentration before resolution of the proteins by SDS-PAGE.

4 Recent developments in and alternative strategies for insect cell expression One problem that commonly arises when using the baculovims expression system (as indeed with many other systems) to overexpress membrane proteins is that the protein produced is only partially active (34, 35). This phenomenon may reflect a rate of" protein synthesis. resulting from the use of powerful viral promoters, that exceeds the rate-limiting mechanisms of protein folding, such that misfblded or aggregated protein accumulates. A possible solution to this problem may be to reduce the growth temperature of the insect cells. Sf9 cells will grow at reduced temperatures (e.g. 20"C), albeit slowly, and the consequent reduction in the rale of protein production may help the folding and trafficking apparatus of the insect cell to cope with the unnatural load. Alternatively, increasing the amounts of specific molecular chaperones or other enzymes that help membrane proteins to fold correctly may increase the levels of functional protein. Several recent papers have reported success in this regard. For example, Lenhard and Reilander (58) introduced the Dmsuphilu rndarwgaster. membrane-bound, peptidyl-prolyl ds/lruns isomernse NinaA into Sf9 cells and showed that coexpression of this protein with the human dopamine transporter substantially increased the amount of properly folded, active transport protein. Similarly, by coexpression with baculovirus encoding the molecular chaperone calnexin, Tare and colleagues have enhanced expression of the functional serotonin transporter threefold (59). As discussed in Section 2.3, the use of less powerful promoters than the polybedrin promoter, that are expressed earlier post-infection, may also lead to the production of more functional expressed protein, albeit in lesser quantities. For example, vectors have recently been introduced that allow the stable constitutive expression of proteins in a variety of insect cell lines under the control of the Orgyia pseudotsugata multicapsid nucleopnlyhedrosis vims (OpMNPV) immediate-early 2 (ie2) promoter (60). For roxic proteins, inducible systems for insect cell expression are also available. Encouragingly, expression of the 138

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

functional human glucagon receptor under the control of a metallothionein promoter in the Drosophfta Schneider 2 (S2) cell system at a level of 250 pmol/mg membrane protein has already been reported (61). Developments like these hold much promise for the continued exploitation of insect cell expression systems for structure-function studies on membrane transporters in the future.

References 1. Opekarova, M., Robl, I., Grassl, R., and Tanner, W. (1999). FEMS Microbiol. Lett.. 174, 65. 2. Arif, B. M. (1986). Curr. Top. Microbiol. Immunol, 131, 21. 3. O'Reilly, D. R., Miller, L. K., and Luckow, V. A. (1992). Baculovirus expression vectors: a laboratory manual. W. H. Saunders, NY. 4. King, L. A. and Possee, R. D. (1992). The baculovirus expression system: a laboratory guide. Chapman and Hall, London. 5. Luckow, V. A. and Summers, M. D. (1988). Virology, 167, 56. 6. Blissard, G. W. and Rohrmann, G. F. (1990). Annu. Rev. Entamol, 35,127. 7. Friesen, P. D. and Miller, L. K. (1986). Curr. Top. Microbiol Immunol, 131, 31. 8. Granados, R. R., Lawler, K. A., and Burand, J. P. (1981). Intervirology, 16, 71. 9. Rohrmann, G. F. (1986). J. Gen. Virol, 67,1499. 10. Summers, M. D. and Smith, G. E. (1987). A manual of methods for baculovirus vectors and insect cell culture procedures. Texas Agricultural Experiment Station Bulletin No. 1555, College Station, Texas. 11. Chaabihi, H., Cetre, C., and Berne, A. (1997). J. Virol. Meth., 63, 1. 12. Jarvis, D. L., Weinkauf, C., and Guarino, L A. (1996). Prot. Express. Purif. 8,191. 13. Bonning, B. C., Roelvink, P. W., Vlak, J. M., Possee, R. D., and Hammock, B. D. (1994). ]. Gen. Virol, 75, 1551. 14. Oka, A., Sugisaki, H., and Takanami, M. (1981). J. Mol. Biol, 147, 217. 15. Capone, J. (1989). Gene Anal. Tech., 6, 62. 16. Malitschek, B. and Schartl, M. (1991). BioTechniques, 11, 177. 17. Webb, A., Bradley, M., Phelan, S., Wu, J., and Gehrke, L. (1991). BioTechniques, 11, 512. 18. Vlak, J. M., Schouten, A., Usmany, M., Belsham, G. J., Klinge, R. E., Maule, A. J., Van, L. J., and Zuidema, D. (1990). ViroZogy, 179, 312. 19. O'Reilly, D. R., Passarelli, A. L., Goldman, I. F., and Miller, L. K. (1990)./. Gen. Virol, 71, 1029. 20. Richardson, C., Lalumiere, M., Banville, M., and Vialard, J. (1992). In Baculovirus expression protocols (ed. C. Richardson and J. Walker), Humana, Clifton, NJ. 21. Zuidema, D., Schouten, A., Usmany, M., Maule, A. J., Belsham, G. J., Roosien, J., Klinge, R. E., Van, L. J., and Vlak, J. M. (1990)J. Gen. Virol, 71, 2201. 22. Kitts, P. A., Ayres, M. D., and Possee, R. D. (1990). Nuckic Acids Res., 18, 5667. 23. Kitts, P. A. and Possee, R. D. (1993). Biotechniques, 14, 810. 24. Patel, G., Nasmyth, K., and Jones, N. (1992). Nucleic Acids Res., 20, 97. 25. Luckow, V. A., Lee, S. C., Barry, G. F., and Olins, P. O. (1993). J. Virol., 67, 4566. 26. Patel, G. and Jones, N. C. (1995). The baculovirus expression system. In DNA cloning 2: a practical approach (ed. D. M. Glover and B. D. Hames), p. 205. Oxford University Press, Oxford. 27. Wagner, R., Geyer, H., Geyer, R., and Klenk, H. D. (1996). J. ViroZ., 70, 4103. 28. Jarvis, D. L., Kawar, Z. S., and Hollister, J. R. (1998). Curr. Opin. Biotechnol, 9, 528. 29. Grisshammer, R. and Tate, C. G. (1995). Quart. Rev. Biophysics, 28, 315. 30. Parker, E. M. and Ross, E. M. (1991). J. BioZ. Chem., 266, 9987. 31. Mills, A., Allet, B., Bernard, A, Chabert, C., Brandt, E., Cavegn, C., Chollet, A., and Kawashima, E. (1993). FEES Lett., 320, 130.

139

GARY J. LITHERLAND AND STEPHEN A. BALDWIN 32. Mouillac, B., Caron, M, Bonin, H., Dennis, M., and Bouvier, M. (1992). J. Biol Chem., 267, 21733. 33. Ng, G. Y. K., George, S. R., Zastawny, R. L, Caron, M., Dennis, M., and O'Dowd, B. F. (1993). Biochemistry, 32, 11727. 34. Tate, C. G. and Blakely, R. D. (1994). ]. Biol. Chem., 269, 26303. 35. Yi, C. K., Charalambous, B. M., Emery, V. C., and Baldwin, S. A. (1992). Biochem.]., 283, 643. 36. Cope, D. L., Holman, G. D., Baldwin, S. A., and Wolstenholme, A. J. (1994). Biochem. J., 300, 291. 37. Smith, C. D., Hirayama, B. A., and Wright, E. M. (1992). Biochim. Biophys. Acta, 1104, 151. 38. Autry, J. M. and Jones, L. R. (1997). J. Biol. Chem., 272, 15872. 39. George, S. T., Arbabian, M. A., Ruoho, A. E., Kiely, J., and Malbon, C. C. (1989). Biochem. Biophys. Res. Commun., 163, 1265. 40. Parker, E. M., Kameyama, K., Higashijima, T., and Ross, E. M. (1991). J. Biol. Chem., 266, 519. 41. Doi, T., Hiroaki, Y., Arimito, L, Fujiyoshi, Y., Okamoto, T., Satoh, M., and Furuichi, Y. (1997). Em.]. Biochem., 248, 139. 42. Nishimura, K., Frederick, J., and Kwatra, M. M. (1998). J. Receptor Signal Transduct. Res., 18, 51. 43. Stauffer, K. A., Kumar, N. M., Gilula, N. B., and Unwin, N. (1991). J. Cell Biol., 115, 141. 44. Barnett, J., Chow, J., Ives, D., Chiou, M., Mackenzie, R., Osen, E., Nyugen, B., Tsing, S., Bach, C., Freire, J., Chan. H., Sigal, E., and Ramesha, C. (1994). Biochim. Biophys. Acta, 1209, 130. 45. Yang, B., van Hoek, A. N., and Verkman, A. S. (1997). Biochemistry, 36, 7625. 46. Li, S., Song, K. S., Koh, S. S., Kikuchi, A., and Lisanti, M. P. (1996). J. Biol. Chem., 271, 28647. 47. Fukuzono, S., Takeshita, T., Sakamoto, T., Hisada, A., Shimizu, N., and Mikoshiba, K. (1998). Biochem. Biophys. Res. Commun., 249, 66. 48. Hink, W. F., Thomsen, D. R., Davidson, D. J., Meyer, A. L., and Castellano, F. J. (1991). Biotechnol. Progr., 7, 9. 49. Davis, T. R., Trotter, K. M., Granados, R. R., and Wood, H. A. (1992). Biotechnology, 10, 1148. 50. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). Molecular cloning: a laboratory manual (2nd ednj.Vol. 1. Cold Spring Harbor Laboratory Press, New York. 51. Matsuura, Y., Possee, R. D., Overton, H. A., and Bishop, D. H. L. (1987). J. Gen. Virol, 68, 1233. 52. Fung, W. K. Y. (1998). PhD Thesis, University of Leeds. 53. Jarvis, D. L. and Garcia, A. (1994). Biotechniques, 16, 508. 54. Licari, P. and Bailey, J. E. (1991). Biotechnol. Bioeng., 37, 238. 55. Towbin, H., Staehelin, T., and Gordon, J. (1979). Proc. NatlAcad. Sti. USA, 76, 7350. 56. Hom, L. G. and Volkman, L. E. (1998). Biotechniques, 25, 18. 57. Hogue, D. L, Hodgson, K. C., and Cass, C. E. (1990). Insect Biochem. Molec. Biol, 24, 517. 58. Lenhard, T. and Reilander, H. (1997). Biochem. Biophys. Res. Commun., 238, 823. 59. Tate, C. G., Whiteley, E., and Betenbaugh, M. J. (1999). J. Biol. Chem., 274, 17551. 60. Hegedus, D. D., Pfeifer, T. A., Hendry, J., Theilmann, D. A., and Grigliatti, T. A. (1998). Gene, 207, 241. 61. Tota, M. R., Xu, L., Sirotina, A., Strader, C. D., and Graziano, M. P. (1995).;. Bioi. Chem., 270, 26466.

140

Chapter 6 The amplified expression, identification, purification, assay, and properties of hexahistidine-tagged bacterial membrane transport proteins Alison Ward,* Neil M. Sanderson,* John O'Reilly,* Nicholas G. Rutherford,* Bert Rodman,1" and Peter J. F. Henderson* *School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT Department of Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands

1 Introduction In bacteria, 3-15% of genes are predicted to encode proteins involved in membrane transport (1), a process which is vital for both the capture of nutrients and the excretion of waste products, toxins, and antibiotics. The energy required for these processes is derived from the electrochemical gradients of ions across the cell membrane (in bacteria the majority operate by proton-driven symport or antiport mechanisms but some utilize a sodium gradient) or from the hydrolysis of ATP (2). Whilst the 3-D structures of thousands of soluble proteins have been determined, to date only 21 membrane protein structures have been solved to atomic resolution (3, 4). The elucidation of the structures of membrane proteins using physical techniques is, therefore, a key area of research. Structure determination is difficult for the majority of membrane proteins due to their extreme hydrophobicity, which means they are refractory to direct manipulation, and can only be removed from the membrane and studied in the presence of detergent. In addition, many membrane proteins are only expressed at low levels, frequently corresponding to less than 0.1% of total cell protein. Working with 16 prokaryotic membrane transport proteins in our laboratory, practical approaches have been devised to clone the gene of interest and to express in Escherichia coli sufficient undenatured protein for structural studies. 141

ALISON WARD ET AL.

The strategy is summarized in Table 1 and its implementation described below, indicating where variations may be needed to overcome the inevitable differences between individual proteins. From the hydropathic profile of the predicted amino acid sequence the majority of the proteins studied (see Table 2) are predicted to comprise 12 membrane-spanning a-helices, but some of these, and of their relatives not described here, are predicted to comprise 10-14 a-helices (5). Table 1 Strategy for obtaining working quantities of membrane transport protein in E. co//once the gene is identified (a) Transfer the gene of interest to the multicloning site in plasmid pTTQIS, so that expression is amplified downstream of the tac promoter. Other vectors may be used (see Section 2). (b) Introduce an appropriate restriction site into the terminating codons of the gene of interest. Design and insert an oligonucleotide that will introduce an -H6 amino acid sequence at the C-terminus of the protein (see Section 2). (c) Optimize growth conditions for the uninduced/induced vector in an appropriate E. coli host strain (see Table 6.3), and implement activity assays. (d) In parallel with (c), check for the appearance of an overexpressed protein in the cell membrane (see Section 3). (e) Identifiy the overexpressed protein as the one required (see Section 3). (f) Carry out solubilization trials in different detergents (see Section 4). (g) Purify the identified protein using an Ni-NTA affinity column initially, and additional steps if required (see Section 5). (h) Reconstitute the purified protein into liposomes to test transport activity, and any other assays of integrity, such as ligand binding (see Section 6). Table 2 Membrane transport proteins cloned and overexpressed in E. coli Substrate(s)

Protein

+

Organism

Galactose-H

GalP

Escherichia coli

Xylose-H+

XylE

Escherichia coli

AraE

Escherichia coli

Glucuronide-H

GusB

Escherichia coli

Proline/betaine-H+

ProP

Escherichia coli

L-Fucose-H+

FucP

Escherichia coli

L-Rhamnose-H+

RhaT

Escherichia coli

Nucleosides-H+

NupC

Escherichia coli

Nucleosides

NupG

Escherichia coli

Aromatic amino acids Bicyclomycin

PheP

Escherichia coli

Bcr

Escherichia coli

Quinolones

NorA

Staphylococcus aureus

Multidrugs

Bmr

Bacillus subtilis

Multidrugs

Bit

Bacillus subtilis

Multidrugs? Lactose-H+

MJ1560

Methanococcus jannaschii

LacS

Streptococcus thermophilus

Arabinose-H+ +

142

THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURFICATION AND ASSAY

One advantage of studying bacterial transport proteins is that amplification of their expression to levels up to 20% of inner membrane protein can often be achieved, yielding 20-50 mg/25 litre culture. In addition, bacteria are much cheaper and easier to grow in large quantities compared to mammalian cells. Eukaryotic transport proteins often have bacterial homologues, so physicochemical studies in bacteria may provide a route to understanding the molecular mechanism of transport in higher organisms.

2 Plasmids and E. coli host strains used in the amplified expression of membrane transport proteins All the vectors and host strains that have been used for the amplified expression of transport proteins in prokaryotes have been reviewed (6). Furthermore, Miroux and Walker (7) have described mutants of £. coli host strains selected for enhanced heterologous expression of transport proteins cloned into plasmid pET vectors. They also discussed the minimization of inclusion body formation. Our own experience is that the plasmid pTTQIS (8) (see Figure 1) has proved successful for all the prokaryote genes that we have tested. Expression via its tac promoter is controlled by repression with LacI, sufficient copies of which are

Figure 1 The plasmid vector pTTQIS (8). Restriction enzyme sites in the multiple cloning site are illustrated by BamHI, Psfl, and H/ndlll. The actual multiple cloning site consists of (in the order 5'-3') EcoRI, Ec/l36, Sad, Asp718, Kpnl, Aval, Smal, Xmal, BamHl, Xba\, Acc\, Sa/l, Sse8387, Pstl, Sphl, and Hindlll.

143

ALISON WARD ET AL.

obtained by including the lacP gene on the plasmid (see Figure 1). During exponential growth in the absence of isopropyl-p-D-thiogalactoside (IPTG) as inducer, expression is repressed, but as the culture moves into stationary phase 'leaky', IPTG-independent expression is often seen. The recombinant plasmid is transferred to the host strain E. coli NO2947 or the commercially available NM554 (Stratagene) for expression, though an advantage of using plasmid pTTQIS is that the resulting construct should be independent of the host (provided appropriate precautions are taken to modify its DNA in restriction-compatible intermediates). The strain DH5a, which is modification-plus, restriction-minus, is used during construction of the plasmid. Other plasmid systems that we have used are plasmid pAD2587, though its \PL promoter requires a specific host strain, AR120 (9), and a derivative of plasmid pBR322 containing the galP promoter (10,11) expressed in E. coli JM1100 (10). The latter host undergoes a morphological transition to long L-forms during expression, in which energization of transport activity may be diminished, though the expressed protein is typically fully functional as determined, for example, by ligand binding assays. At this stage the construct can be tested for overexpression of the protein as described in Section 3. Either before or after this test, a hexahistidine (His)6 tag can be introduced. A restriction site is chosen that will be unique to the whole construct, and engineered into the C-terminal codon of the protein by judicious design of an appropriate oligonucleotide and polymerase chain reaction (PCR). An oligonucleotide is designed with matching flanking sites, six in-frame histidine codons, optional epitope codons (e.g. RGSH6), an optional protease-susceptible site before the inserted amino acids (e.g. Factor Xa), and an optional additional unique restriction site to facilitate the recognition of a successful insert. This oligonucleotide is then ligated into the unique C-terminal restriction site in the plasmid. After the construction of the plasmid it is important to confirm by sequencing that no mutations were introduced into the construct.

3 Growth conditions and detection of amplified membrane transport protein expression The overexpression of membrane proteins in E. coli is often associated with toxicity and cell death, e.g. the expression of histidine-tagged norfloxacin resistance protein (NorA) from Staphylococcus aureus. In such cases it is often more productive to produce cells in batch culture in flasks (see Protocol 1), rather than producing them using a fermenter. Even where there is no toxicity associated with membrane protein overexpression, e.g. the glucuronide transporter of E. coli, batch culture in flasks may still produce the best results for reasons that remain unclear. The conditions for optimal overexpression, such as growth media, concentration of inducer, and time of induction should be determined experimentally on a small scale before embarking on larger scale cultures. Examples of growth conditions for transformed £. coli are given in Table 3. 144

THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURFICATION AND ASSAY

Protocol 1 Batch culture of recombinant E. coll for the overexpresslon of membrane proteins Equipment and reagents • Luria-Bertani (LB) medium: 10 g/1 Bactotryptone, 10 g/1 NaCl, 5 g/1 Bacto-yeast extract. Adjust pH to 7.5 with NaOH and then sterilize by autoclaving. • Temperature-controlled orbital shaker

• Terrific broth, modified (Sigma): 12 g tryptone, 24 gyeast extract, 9.4 g KjHPO4, 2.2 g KH3PO4, 8 ml glycerol per litre • Refrigerated superspeed centrifuge

Method 1. Pick a single bacterial colony from a freshly streaked plate and transfer into a 250 nil baffled conical flask containing 50 ml LB and antibiotics appropriate for plasmid selection." 2. Grow the cells in an orbital shaker at 37 °C, 220 r.p.m. for 12-16 h. 3. Transfer the cells to a sterile 50 ml plastic centrifuge tube and collect the cells by centrtfUgation at 12 000 gav for 10 min, at room temperature, 4. Resuspend the cells in 1 ml of LB and use this to inoculate 800 ml of Terrific broth (again with appropriate antibiotic selection) in a 2 litre baffled flask. 5. Grow the cells in an orbital shaker at 37°C, 220 r.p.m., and monitor their growth by measuring the absorbance of the culture at 680 nni (A^6. Induce the cells at mid log phase (approx. A580 = 0.6) with an appropriate amount ofinducer.b 7. Harvest cells by centrifugation at 12000 g^, for 20 min at room temperature, at stationary phase or when increasing cell death and lysis occur. " This method describes the overexpression of histidine-tagged norfloxacin resistance protein of Staphylococcus aurens (NorA) in E. coli from a pTTQ18-based plasmid in the E. coli cell strain Blr (Novagen) with carbenicillin (100 n.g/ml) selection. b For overexpression of NorA, the cells are induced with 0,2 mM IPTG for 4-5 h.

For the small-scale preparation (culture volumes no greater than 100 ml) of mixed membranes, i.e. inner plus outer membranes, water lysis of E. coli cells is conveniently carried out according to Protocol 2. This method is more reproducible than sonication. and quicker than the preparation of vesicles using Kaback's method (12). The protein is assayed by the Schaffner-Weissmann method (13), and then suitable quantities are solubilized in SDS and the proteins separated by SDS-PAGE (14). The membrane protein of interest may not be resolved as a righr band on the gel but instead may be diffuse in appearance. 145

Table 3 Media used in the overexpression of membrane proteins in 25-litre fermenter cultures Protein expressed

GalP(His)6

XylE(His)6

AraE(His)6

GusB(His)6

ProP(His)6

FucP(His)6

NorA(His)6

Bmr(His)6

Strain/plasmid

JM1100 pBR322

N02947 pTTQIS

N02947 pTTQIS

N02947 pTTQIS

WG389 pBR322

BLR pTTQIS

BLR pTTQ18

BLR pTTQIS

%/nocu/uftt (v/v)

3.0

1.0

1.0

1.0

3.0

3.0

6.0

6.0

b

Inoculum medium

2TY" Tet + HisThy"

LB" Carb"

Media constituents

(Final concentration g/litre)

Na2HP04 (anhyd. salt)

9.0288

KH2P04 NH4CI

d

e

LB'Carb"

LB'Carb*

2TY" Garb"

LB Garb

LBd Carbe

LBd Carba

6.0

6.0

6.0

6.0

—

—

6.0

3.8092

3.0

3.0

3.0

3.0

—

2.2

3.0

2.7

1.0

1.0

1.0

1.0

—

—

1.0

K2HP04

—

—

—

—

—

—

9.4

—

Yeast extract

0.1332

—

—

—

0.1332

10.0

24.0

—

Bactottyptone

0.1332

—

—

—

0.1332

10.0

—

—

NaCI

0.0666

0.5

0.5

0.5

0.5666

5.0

—

0.5

Casamino acids

—

2.0

2.0

2.0

2.0

—

12.0

2.0

Thiamine

0.0002

—

—

Histidine

0.12

— —

—

—

0.1 —

— —

— —

— —

Thymine

0.03

Proline

—

Tryptophan

—

CaCI22H20

0.6664

MnCI2-4H20

0.00668

MgS047H20

0.13332

FeS04:7H20

0.00668

Glucose (carbon source)

5.0

Glycerol (carbon source)

—

Tetracycline (|xg/ml)

15

Ampicillin (ug/ml)

—

f

IPTG(mM)

—

0.02944

0.02944

0.02944

0.4930

0.4930

0.4930

1.84

1.84

1.84

100 1.0

100 0.4

100 0.6

All E. coli cultures are grown at 37°C. a2TY: Bacto-tryptone 10 g/litre, yeast extract 10 g/litre, sodium chloride 5 g/litre. Tetracycline at 15 (ug/ml. Histidine 80 (ug/ml, Thymine 20 (ug/ml. "Luria-Bertani medium: Bacto-tryptone 10 g/litre, yeast extract 5 g/litre, sodium chloride 10 g/litre. "Carbenicillin at 100 (ig/ml. the final concentration of IPTG is given for pTTQIS-based plasmids.

0.1 0.1 0.6664 0.00668 0.13332 0.00668 3.6

100

0.02944 0.4930

0.92

10.0

1.84

100 0.2

100 0.2

100 0.6

ALISON WARD ET AL.

Protocol 2 Preparation of E. coll mixed membranes using water lysis Equipment and reagents • Refrigerated superspeed centrifuge • Shaking water bath or incubator • Hand-held homogenizers for 30 ml and 1 ml volumes • 50 ml of E coli cell cultures from which membranes are to be prepared • 0.2MTris-HClpH8.0 Sucrose buffer: 1 M sucrose, 1 mM EDTA, 0.2 M Tris-HCl pH 8.0

• Membrane resuspension buffer: 0.1 M sodium phosphate pH 7.2,1 rnM 2-mercaptoethanol • DNase ' MgCl2 • EDTA • 1.3 mg/ml lysozyme in sucrose buffer, freshly prepared

Method 1. Transfer the cells to a 50 ml plastic centrifuge tube and centrifuge at 12000gav, at 10 0C for 10 min. 2. Resuspend the cell pellet in 10 ml of 0.2 M Tris-HCl pH 8.0 and shake at 25°C for 20 min. 3. At zero time add 9.7 ml of sucrose buffer. 4. At 1.5 min add 1 ml of 1.3 mg/ml lysozyme, 5. At 2 min add 20 ml of deionized water, and leave the solution shaking at 25°C for 20-60 min," 6. Sediment the spheroplasts formed at 20 000gav, at 10°C for 20 min. Note that the supernatant constitutes the periplasmic fraction and can be retained for analysis. 7. Resuspend the spheroplasts in 30 ml of deionized water with a hand-held homogenizer and allow to stand at 25 °C for 30 mm. 8. Sediment the membranes at 30000 gav. at 4°C, for 20 min. Note that the supernatant obtained is the cytoplasmic fraction and can be retained for analysis. 9. Wash the membranes three times in 15 ml of membrane resuspension buffer, using a hand-held homogenizer to resuspend the pellet. 10. Finally, resuspend the washed membranes in l ml of membrane resuspension buffer, adding MgCl2 to a final concentration of 1 mM and DNase to a final concentration of 20 ug/ml. 11. Incubate the membranes at 37°C for 30 min and then stop the DNase reaction by adding EDTA to a final concentration of 1 mM. 12. Snap-freeze at -70°C in ethanol. 13. Store at -70°C "Follow the formation of spheroplasts by phase-contrast microscopy at 800 x magnification, and note the number and motility of any intact cells.

148

THE AMPLIFIED EXPRESSION, IDENTIFICATION. PURFICATION AND ASSAY

Whether expression has been successful is quickly ascertained by comparing Coomassie ISlne R-250-stained. SDS PAGK-separated, membrane preparations from induced vs. um'nduced cultures (or induced host containing the original unmodified vector vs. induced host containing the construct with the gene of interest). Since the transport proteins we study always migrate at an anomalous rate, faster than expected from their predicted M1, values, additional confirmation of the expressed protein's identity is required. Possible methods are listed below: • electroelution and determination of the N-terminal amino acid sequence; • partial proteolysis and determination of N-terminal amino acid sequences of derived peptides {this may be essential if the N-terminus of the protein is blocked, e.g. by N-formylation of methionine); • Western blotting after SDS-PACE with an antibody of proven efficacy against -H6 'tag', or antibody raised against peptides derivd from the predicted amino acid sequence; • if specific labelling reagents are available, e.g. radioactive cytochalasin B, forskolin or 3- |125l|iodo-4-azidophenethylamido-7-0-succinyldeacetyl (IAPS)forskolin in the case of the £ coli galactose H' symponer GalP, then prior photoaffinity-labelling of the overexpressed protein with these, followed by appropriate detection after SDS-PAGE. Note that these membrane preparations contain both inner and outer membrane proteins. Some of the outer membrane proteins can be abundant and so can dominate the overall profile of the proteins, making detection of the desired membrane protein more difficult, especially if it migrates at a similar Mr value. This is overcome by making larger scale membrane preparations using the f'rench press and separating inner and outer membrane fractions by sucrose density-gradient centrihigation (see Protocol 3 and Figure 2).

Protocol 3 Separation of the inner and outer bacterial membrane fractions Equipment and reagents • Cell pellet (50-60 g maximum wet weight," stored at - 70 °C in 20 mM Tris-HCl, 0.5 mM EDTA, 10% (v/v) glycerol, pH 7.5 • 20 mM Tris-HCl buffer pH 7.5 » Tris-EDTA buffer: 20 mM Tris-HCl pH 7.5, 0.5 mM EDTA

Tris-EDTA-sucrose buffers: Tris-EDTA buffer containing sucrose at concentrations of 55, 50, 45, 40. 35, 30, and 25% (w/w) French Press (Aminco-SLM Instruments Inc.) and pressure cell pre-cooled to 4°C

Methoda,b,c,d,e 1. Thaw the pellet and keep on ice until required.

ALISON WARD ET AL. Protocol 3 continued

2. Homogenize the cell suspension with Tris-EDTA buffer to give a volume of 200-300 ml. Add more buffer if the slurry is very thick. 3. Pass the slurry through the French Press at 20000 lb/in2. Collect the outflow and adjust the volume to approximately 400 ml with more Tris-EDTA buffer. 4. Sediment the debris pellet by centrifugation at 10 000 gav for 45 min and retain this for analysis. 5. Sediment the membranes from the supernatant fluid at 131000 gav for 90 min. 6. Prepare sucrose gradients in 65 ml centrifuge tubes in 10 ml layers of: 55, 50. 45,40, 35, and 30% (w/w) sucrose in Tris-EDTA buffer. Store in the cold room until required. 7. Resuspend the membrane pellet from step 5 in a small volume of 25% (w/w) sucrose Tris-EDTA buffer. 8. Layer the membrane fraction on to the sucrose gradient. Note that each centrifuge tube takes about 4 ml of the membrane fraction. 9. Centrifuge at 113000 g^ for 18 h. with minimal acceleration and no braking. 10. Draw off the membrane layers. The golden inner membranes are at the 35-40% interface and the white outer membranes are at the 50-55% interface.11 11. Resuspend the membranes in Tris-HCl buffer and sediment at 131000 gav for 2 h. 12. Wash the membranes three times, to remove traces of EDTA and sucrose, by resuspending in Tris-HCl buffer and sedimenting at 131000 gav for 60 min. 13. Resuspend in Tris-HCl buffer and aliquot in 250 ul volumes into cryotubes. 14. Snap-freeze at -70°C in ethanol. 15. Store at -70°C. " For smaller quantities than this scale down the volumes in the protocol accordingly, ""This method describes the preparation of membrane fractions using a Beckman fixed-angle Ti45 rotor for the high-speed centrifugation step. c Cany out all procedures at 4°C, d Some inner membrane material may be located with the higher sucrose density 'outermembrane' band and this should therefore be regarded as an 'inner membrane-depleted' fraction, ( In addition to checking the fractions for the presence of the expressed protein, tests may be carried out for the presence of the normal membrane markers.

4 Detergent choice and solubilization of integral membrane proteins Integral membrane proteins are removed from lipid bilayers by the action of detergents. Detergents are amphipathic molecules comprising a polar head group and a hydrocarbon tail. At a determined concentration, referred to as the critical micelle concentration (CMC), detergent monomers aggregate to form ordered structures, called micelles, into which membrane proteins can insert. 150

THE AMPLIFIED EXPRESSION. IDENTIFICATION, PURFICATION AND ASSAY

1

2

3

4

Figure 2 Membrane fractions containing overexpressed NorA(His)6 generated during the separation of inner and outer membranes. NorA(His)s was expressed from a pTTQIS-bascd plasmid, in Terrific broth with 100 ug/ml carbeniciHin-selection and 0.2 mM IPTG-induction for 4-5 h. Membrane fractions were prepared from harvested cells as outlined in Protocol 3. separated on a 15% SDS-PAGE gel and stained with Coomassie Brilliant Blue. Mixed membranes (Track 1) were prepared after French pressing and removal of the cell debris (Track 2). Outer membranes {Track 3) were separated from inner membranes (Track 4) by sucrose density gradient centrifugation, followed by washes to remove the sucrose.

Detergents are classified into three broad categories: • ionic, which carry a net charge associated with their head group, e.g. the anionic detergent sodium dodccyl sulfate (SDS); • non-ionic, which have uncharged hydrophilic head groups, e.g. n-dodecyl-p-Dmaltoside (DDM); • zwiUmonic. which have both positive and negative charges but carry no net charge, e.g. 3-(3-cholamidopropyl)-dimethylammonio-1 -propane sulfonate (CHAPS). Detergents used in the solubilization and purification of membrane proteins must main Lain the structural integrity of the protein and its activity on reconstitution. In many cases it may not be possible to select a detergent that is suitable for both solubilization and purification. In such cases detergent exchange may be carried out (see later). The solubilization of membrane proteins is a multistep process. Below the CMC detergent monomers partition into the lipid bilayer. As the concentration of the detergent increases to levels at, or above, the CMC the membrane breaks down to generate mixed micelles of protein/detergent, protein/detergent/lipid, lip id/detergent, and detergent alone. Membrane protein solubilization trials should be carried out in the presence of stabilizing additives such as glycerol and NaCl, using a wide range of deterI5I

ALISON WARD ET AL.

gents at concentrations at and above the CMC, and protein concentrations in the range from 1 to 10 mg/ml. The solubilization mix is incubated (usually on ice) before recovering the solubilizecl material by ultracentrifugation (108000 grav, for 1 h, at 4"Q. Individual membrane proteins will show different solubilization requirements, especially pH and salt concentration. The process of solubilization can be monitored using SDS-PAGIi and Western blotting. As an example, the solubilization conditions for the hisridi lie-tagged £ culi galactose II"1" symport protein (GalP) are given below (see Protocol 4).

Protocol 4 Solubilization of bacterial membranes containing (Hls)6tagged protein Equipment and reagents • Bacterial membranes of known protein concentration (see Protocol 3) • n-Dodecyl-p-D-maltopyranoside (DDM) (Calbiochem or Melford)

Solubilization buffed: 20 mM Tris-HCl*1 pH 8.0. 20 mM imidazole1, 300 mM NaCld, 20% (v/v) glycerol,r 1% (w/v) DDMf

Method 1. Add the membrane preparation to the solubilization buffer to give a final protein concentration of 5 rag/ml. 2. Vortex for 5 sec and incubate on ice for 1 h. 3. Centrifuge at 108 000 gav for 1 h at 4 °C. Carefully decant the supernatant from the pellet and retain these fractions on ice. Cany out protein determinations by the method of Schaffner and Weissmann (13), 4. Perform SDS-PAGE by the method of Henderson and Macpherson (14). 11

For the solubilization buffer, the molarities and concentrations given are final (i.e. after the addition of the membranes). " 20 mM Tris-HCl can be replaced with l0 mM Hepes in the buffers. c 20 mM imidazole is added to prevent non-specific binding to Ni-NTA resin (Qiagen). J NaCl (0-1 M) aids solubilization. ' Glycerot stabilizes the solubilized membrane protein. ^DDM was found to be the detergent best suited to solubilize GalP(His)6 inner membranes from E. coli strain JM1100 (pPER3).

5 Purification of (His)6-tagged proteins 1 lexahisticline tags (see above) facilitate affinity purification using nickel chelateaffmity diromatography (see Protocol 5]. The attachment of a hex;ihistidine tag to the C-terminus of 12a-helical prokaiyotic transport proteins has been successful for the purification of all the proteins studied so far in our laboratoiy (Figure 3). 152

THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURFICATION AND ASSAY

Figure 3 15% SDS-PAGE gel illustrating the purification of Bmr(His)B from mixed membranes using Ni-NTA affinity chromatography. Mixed membranes, 20 mg (Track 2) were solubilized at 4 mg/mt membrane protein in 10 mM Hepes pH 7.9, 20 mM imidazole pH 8.0, 1% (w/v) DDM, and 20% (v/v) glycerol. After the removal of non-solubilized material by ultracentrifugation, the solubilized material (Track 3) was bound to 1 ml of packed Ni-NTA resin at 4°C for 12-16 h. The resin was then centrifuged at 180 g^ for 1 min, and the supernatant collected (Track 4). The resin was then washed with 30 ml of buffer (10 mM Hepes pH 7.9, 20% (v/v) glycerol, 20 mM imidazole, 0.05% (w/v) DDM) and the supernatants collected (Track 5). Finally, the specifically bound Bmr(His) 6 was eluted, after packing the resin into a 2 ml disposable column. using 4 ml 10 mM Hepes pH 7.9, 20% (v/v) glycerol, 200 mM imidazole. Fractions of 1 ml were collected (Tracks 6-9). M, markers are shown in Track 1. The predicted M1 of Bmr{His) e is 43536, but as analysed by SDSPAGE the protein has an anomalous M, of 32000. Analysis by MALDI mass spectrometry (see Figure 8) reveals the experimental M, to be 43739.

If protein preparations require further purification there are several options available: • an ion-ex change chromatography, e.g. using Q-Sepharose or MonoQ. (Pharmacia); • cation-exchange chromatography, e.g. using S-Sepharose (Pharmacia); • gel filtration, e.g. using Superdex 200 (Pharmacia); • hydroxyapatite chromatography. Gel filtration requires only that the protein sample is in a small volume before being applied to the column. VVhilst good resolution of protein can be obtained, it does result in at least a 30-fold dilution of protein so that a good concentrative method is needed following this procedure. For anion- and cation-exchange chromatography samples must be in a nonionic detergent and desalted before applying to a column. With these methods, protein is bound to the column and then eluted using increasing concentrations of salt. Roth are concent!'alive steps.

ALISON WARD ET AL.

Protocol 5 Purification of (Hls)6-tagged protein using Nf-NTA agarose affinity chromatography Equipment and reagents • Nickel-nitrilotriacetic acid (Ni-NTA) agarose (Qiagen) • Wash buffer: 20 mM Tris-HCl. 20 mM imidazole, 20% (v/v) gtycerol, 0.05% (w/v) DDM, pH 8.0fl

• Elution buffer: 20 mM Tris-HCl, 200 mM imidazole, 10% (v/v) glycerol, 0.05% (w/v) DDM. pH 7.5" • Disposable polystyrene columns, 0.75 cm internal diameter, 2 nil capacity, fitted with a polyethylene disc, 45 um pore size (Pierce)

Method 1. Wash 1 ml of packed Ni-NTA agarose three times with 5 ml of deionized water and twice with 5 ml of wash buffer, by centrifugation at 180 gav for 1 min. 2. Equilibrate the Ni-NTA agarose in 5 ml wash buffer for 1 h on ice. 3. Sediment the Ni-NTA agarose by centrifugation at 180 gav for 1 min. 4. Add 4.5 ml of a solution of detergent-solubilized, (His)6-tagged protein (e.g. from Protocol 4 step 3) to the sedimented agarose and gently mix for 2 h at 4°C. 5. Sediment the Ni-NTA agarose at 180gavfor 1 min and retain the supernatant. 6. Wash the sedimented Ni-NTA agarose, using a batchwise procedure, at least 10 times with 5 ml aliquots of ice-cold wash bufferc. 7. Resuspend the washed Ni-NTA agarose in 1-2 ml of wash buffer and transfer to a disposable column, 8. Allow the wash buffer to drain away and elute the (His)fi-tagged protein with 4 ml of the elution buffer, and collect 1 ml fractions. 9. Analyse the fractions by SDS-PAGE. " For affinity chromatography, the concentration of DDM is reduced from that used for membrane solubilizarion to 0.05% (w/v}, which is still above the CMC. b In the elution buffer, the concentration of glycerol may be decreased to 5% (v/v) for some proteins. The bound protein is eluted by displacement with imidazole. Elution can also be achieved by protonation of the (His)6 tag with 100 mM sodium citrate pH 4, or 100 mM glycine pH4. c Detergent exchange and/or buffer exchange can be undertaken while the purified protein is still bound on the Ni-NTA agarose, using at least 30 column volumes of the exchange solution.

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THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURFICATION AND ASSAY

6 Reconstitution and activity assays of purified membrane protein Before the isolated protein can be used for structural studies it is necessary to show its integrity after the potentially denaturing purification procedures. The most important test is to measure its transport activity. The procedure is to reconstitute the protein into lipid bilaycr membranes, diminishing the concentration of detergent as much as possible by dilution, dialysis, or adsorption to polystyrene beads (see Protocols 6A and 6B). While many procedures have been established, we must emphasize the importance of pre-treating the liposomes with carefully controlled concentrations of the same or different detergents (15, 16),

Protocol 6A Reconstitution of detergent-solubillzed membrane proteins into E. coll liposomes by detergent dilution Equipment and reagents • Thermobarrel Extruder (Lipex Biomembranes Inc.) plus polycarbonate membrane filters (Poretics Products), pore size 0.1 (i.m, diameter 25 mm • E. coil total lipid extract (Avanti Polarlipids, Inc.) • Chloroform

Membrane protein solubilized in detergent Phosphate buffer: 50 mM potassium phosphate pH 7,6,1 mM dithiothreitol fl-octyl glucoside (octyl-fJ-r>glucopyranoside) (13.6 % (w/v))

Method 1. In a test tube, dissolve 20 mg of the lipid in chloroform and dry under nitrogen or argon to produce a film. 2. Rehydrate the film by adding 1 ml of the phosphate buffer at 500C. 3. Incubate the sample at 50°C for a few minutes and then vortex the tube. Repeat these heating and vortexing steps until the dried lipid film has been totally removed from the walls of the tube, 4. Pass the sample at least twice through the extruder at 50°C under nitrogen at a pressure of 200-400 lb/in2 to produce unilamellar vesicles, 5. Incubate at 4°C and add 130 u1 ofB-octylglucoside plus an amount" of detergentsolubilized membrane protein. Mix for 15 min. 6. Dilute to a volume of 130 ml with phosphate buffer, 7. Centrifuge at 108000gavfor 1 h at 4°C 8. Discard the supernatant and dry the insides of the centrifuge tubes carefully with tissue. Resuspend the pellets in 1 ml of phosphate buffer.*1 155

ALISON WARD ET AL.

Protocol 6A continued

9. Assay the protein content of the vesicles using the method of Schaffher and Weissmann (13). " For the counterflow assay (see Protocol 7) a lipid to protein (w/w) ratio of 100:1 is normally desirable. For FTIR (see Protocol 9) 1 mg protein is used per 20 mg lipid. h For FTIR (see Protocol 9) proteoliposomes are not resuspended in buffer.

Protocol 6B Reconstitution of membrane protein using Bio-Beads' Equipment and reagents • Thermobarrel Extruder (Lipex Biomembranes Inc.) and polycarbonate membrane filters (Poretics Products), pore size 0.1 um, diameter 25 mm • Bio-Seads SM-2 (Bio-Rad)11 • Purified E col; lipidsL

• Membrane protein solubilized in detergent • 10% (w/v) Triton X-100 stock solution in deionized water ( = 154 mM) • Phosphate buffer: 10 mM potassium phosphate pH 7.6

Method 1. Resuspend 20 mg of purified £. colt lipid in a test tube in 1 ml chloroform, 2. Dry the lipid under a stream of nitrogen to form a thin film. 3. Rehydrate the lipid in 1 ml of phosphate buffer by vortexing. Note that heating at 50°C may be required for complete rehydration. 4. Pass the lipid at least twice through the extruder at 50°C under nitrogen at a pressure of 200-400 Ib/in2 to produce unilamellar vesicles. 5. Dilute the liposomes in phosphate buffer to 4 mg/ml, 6. Destabilize the liposomes by adding Triton X-100 to a final molarity of 1.5 mM. 7. To 1 ml of 4 mg/ml destabilized liposomes add 100 u1 of 0,4 mg/ml solubilized membrane protein to give a final lipid to protein ratio of 100:1 (w/w).J 8. Incubate the solution with shaking at 25°Cfor 15 rain. 9. Add 40 mg Bio-Beads and incubate with shaking at 25 °C for 30 min. 10. Remove the solution and transfer to a clean 1.5 ml microcentriftlge tube containing 80 mg Bio-Beads, 11. Incubate, with shaking, at 4°C for 60 min. 12. Repeat step 10 and incubate, with shaking, at 4°C for 12-16 h. 13. Remove the Bio-Beads and pellet the proteoliposomes by centrifugation at 108 000 gav at 4°C for 60 min.''

THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURF1CATION AND A S S A Y

Protocol 68 continued

14. Resuspend the proteoliposomes in 0.5 ml phosphate buffer and assay for protein/ " Based on the method of Knol et al (16) b Prepare the Bio-Beads by washing twice with methanol and then resuspending in water. c Purify the E. colt lipids by acetone/ether extraction. d This method describes the reconstitution for CD or functional studies. For FTIR studies 20 mg lipid (5 ml liposomes) and 1-2 mg protein can be used, with a corresponding scale-up in the amount of Bio-Beads used. f If the reconstitution has been performed for PTIR, a sample of proteoliposomes should be removed for protein assay prior to ultracentrifugation, as proteoliposomes are not resuspended in buffer for this method (see Protocol 9). ^"Recovery of protein for the E. coli glucuronide transport protein using this method has been 80-100%, compared with only 40-50% using the rapid dilution method (see Protocol 6A).

The transport activity can then be assayed by measuring the counterflow of rail ioisolope-labe lied and tmlabclied substrates, i.e. the method we describe in detail here (sec Protocol 7 and Figure 4). Also, for the many Transport proteins for which activity in vivo is driven by electrical and/or ion gradients, the reconstituted proteoliposomes can be treated with valinomycin ' appropriate gradients of K' and Na 1 to'drive'the proteindependent transport of labelled substrate against its concentration gradient (18). Alternatively, oxidases/ATPascs capable of pumping protons and thus

Protocol 7 Counterflow assay for activity of reconstituted GatP(His)6 proteina This protocol is based on procedures routinely used for assaying the GalP(His)6 protein, and so galactose is cited as the substrate; however, other substrates must be substituted when assaying different proteins. Equipment and reagents • Proteoliposomes {see Protocol 6A) • Phosphate buffer: 50 mM potassium phosphate pH 7.6,1 mM dithiothreitol • Phosphate buffer plus D-galactose: 50 mM potassium phosphate pH 7.6,1 mM dithiothreitol, 20 mM D-galactose

• D[1-3H]galactose (Arnersham Life Science), 50 uM, 4 uCi • Vacuum manifold • Nitrocellulose filter membranes (Millipore) type GS, 0,22 um pore size

Method 1. Make proteoliposomes as in Protocol 6A, using 0.2 mg of GalP(His)G solubilized in 0.05% DDM, but use phosphate buffer plus D-galactose.

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ALISON WARD ET AL Protocol 7 continued

2. At zero time, add 40 JA! of the galactose-loaded proteoliposomes to 920 p-1 of phosphate buffer plus 40 u1 of D-[l-3H|galactose, giving a final concentration of the radiolabelled galactose of 2 uM. 3. At each time point. filter an 80 ul sample on the vacuum manifold using the nitrocellulose filter membranes and wash with 4 ml of ice-cold phosphate buffer. 4. Determine the radioactivity appearing in the proteoliposomes retained on the filter by liquid scintillation counting. " Based on the method of Newman and Wilson (17).

Time (min) Figure 4 Entrance-counterflow assay for reconstituted GalP(His)6. Entrgnce-counterflow assays were based on the method of Newman and Wilson (17) and carried out at 4°C (• and 20°C (•]. At zero time, the galactose-loaded proteoliposomes (40 uI, - 4-20 ug protein) were diluted into a mixture of 920 ul potassium phosphate (pH 7.6), 1 mM DTT with 40 uI (3H]galactose (50 uM, 4 uCi). The final concentration of the radiolabelled galactose was 2 uM. At each time point, an 80 ul sample was filtered on a vacuum manifold using nitrocellulose filter membranes (Millipore type GS, 0.2 (um pore size) and washed with 4 ml of ice-cold 50 mM potassium phosphate (pH 7.6), 1 mM DTT. The radioactivity appearing in the proteoliposomes retained on the filter was determined by liquid scintillation counting.

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generating H+ ion gradients can be reconstituted into the same preparations. The specific activity can then be compared with that determined in the original cells or membrane preparations. For some transport proteins tight-binding ligands are available, which can be used for binding assays to the isolated and/or reconstituted protein, e.g. cytochalasin B or forskolin for GalP. This is often more convenient than transport assays.

7 Physical properties of purified membrane protein Once a purified protein is obtained, its properties can be determined. We find that all the transport proteins we have purified and reconstituted so far tenaciously retain a circular dichroism (CD) spectrum characteristic of substantial a-helix content, even when solubilized in detergent (see Figure 5). Quantitative analyses

.

Wavelength (nm) Figure 5 CD spectra of detergent-solubilized GalP(His)6 (dashed line) and GalP(His)6 proteoliposomes (solid line). The circular dichroism measurements were obtained using a Jasco J-715 spectropolarimeter at 20°C with constant nitrogen flushing. The samples were analysed in Hellma quartz-glass cells of 1 mm path length. Spectra were recorded with 1 nm sampling intervals at a scan rate of 50 nm/min. The sensitivity was set at 20 mdeg with a response time of 1 sec. For detergent-solubilized GalP(His)6, a sample of the purified protein was exchanged into 10 mM potassium phosphate (pH 7.6), 1 mM DTT, 0.05% DDM and an aliquot of this suspension (300 ul, protein concentration ~ 25 ug/ml) scanned from 190-260 nm. Each spectrum was an average of 10 scans. Spectra were solvent-subtracted and smoothed. For reconstituted GalP(His)6, a sample of proteoliposomes (300 (il) was scanned in a similar fashion in detergent-free buffer. Note for the GalP(His)6 in 0.05% DDM the negative indentations at 208 and 222 nm, with a positive signal at 190 nm, are indicative of a high percentage a-helix. The distortion of the 208 nm indentation for the sample of proteoliposomes is due to light scattering.

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ALISON WARD ET AL.

Temperature (°C) Figure 6 Thermal unfolding of GalP(His)6 (25 ug/ml) in 0.05% DDM monitored by CD. Tertiary interactions in the detergent-solubilized protein were monitored by heating the sample between 10°C and 90°C at 0.1°C intervals (scan rate 100°C/h). Structural changes were monitored by recording the effect of the temperature change on the CD maxima observed at 222 nm. The sigmoidal unfolding curve is indicative of the initial loss of tertiary structure followed by the loss of secondary structure.

by several algorithms estimate a-helix contents of 75-95%. Also, the a-helix content can be obtained by FTIR spectroscopy after reconstitution and evaporation of water, but the proportion of a-helix (about 50%) is less than indicated by CD, even though it is easily the dominant type of secondary structure. That the spectra truly represent a folded protein is confirmed by reduction of the CD absorption peaks under denaturing conditions, e.g. heating (see Figure 6).

7.1 Circular dichroism (CD) spectroscopy Provided the correct buffer is used (see Protocol 8), the CD spectrum of the protein in proteoliposomes or detergent can be measured. There is a danger of light scattering, or heterogeneity of the sample, interfering with the quantitative analysis in terms of secondary structure content in the case of proteoliposomes. This is less of a problem with detergent-solubilized protein, provided that the micelles do not scatter light significantly.

7.2 Fourier-transform infrared (FTIR) spectroscopy FTIR spectroscopy can be performed using detergent-solubilized and purified protein after exchange into 2H2O (19), or proteoliposomes (see Figure 7} may be used as outlined in Protocol 9. Analysis of membrane protein secondary structure by FTIR spectroscopy may be more reliable than analysis performed using CD because, unlike the latter, quantitative estimates of secondary structure are not dependent on a knowledge of protein concentration.

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Protocol 8 Circular dichrolsm (CD) spectroscopy of reconstituted and detergent-solubilized membrane protein Equipment and reagents • Proteoliposomes (see Protocol 6) or purified protein • Phosphate buffer: 10 mM potassium phosphate pH 7,6.1 mM dithiothreitol with detergent as appropriate

CD spectropolarimeter Quartz glass cells (1 mm path length; Hellma)

Method 1. Make proteoliposomes as in Protocol 6, using 0.2 mg of purified membrane protein solubilized in 0.05% DDM, but use 10 mM of phosphate buffer instead of 50 mM." 2. Dilute the proteoliposomes in 10 mM of phosphate buffer to give a final protein concentration of 25 ug/ml. 3. Add 300 ul of the diluted proteoliposomes to the glass cell. 4. Set up the CD instrument so that sample spectra are obtained under constant nitrogen flushing at a scan rate of 50 nm/min. 5. Scan the sample between 190 and 260 nm, averaging at least 10 accumulations. 6. Scan a sample of phosphate buffer alone and subtract this from the proteoliposome spectrum. 7. Convert the arbitrary CD units to values of mean residue ellipticity (deg cm-2 dmol-1) to allow the prediction of secondary structure content using one of the various computer programs available e.g. Jasco Secondary Structure Estimation Program, 8. To examine the stability of the protein, increase the temperature of the sample chamber from 10°C to 90 °C in 0.1°C intervals (scan rate 100°C/h). Monitor changes in secondary structure by recording the effect of the temperature change on the CD maxima observed at 222 nm. "Purified membrane protein in detergent may be used, in which case begin the protocol at step 3. The buffer used for CD must always be 10 mM phosphate. Biological buffers and chloride ions all absorb strongly in CD,

7,3 Mass spectrometry of membrane proteins SDS-PAGn docs not provide a reliable measure of the M- of membrane transport proteins, because of their anomalous migration (sec above). The lower M, observed leads ro the danger t h a t post-translalional processing might occur undetected, and be responsible for loss of activity and/or structural integrity during purification. Recently Hufnagel et al. (20) devised a protocol to prepare bcicleriorhodopsin for matrix-assisted laser desorption, mass spectrometry 161

ALISON WARD ET AL.

(MALDI-MS), and we have extended this to the analyses of membrane transport proteins (see Protocol 10). The important observation is that expected Mr values are obtained by MALDI-MS in rather broad, but discrete, peaks (see Figure 8). So far, however, only multiple peaks—albeit in the correct range—are obtained with electrospray ionization mass spectrometry (ESI-MS). ES1-MS was more satisfactory when the sample comprised proteolytic fragments derived from

Protocol 9 Fourier-transform Infrared (FTIR) spectroscopy of proteollposomes Equipment and reagents • Proteoliposomes (see Protocol 6) * Phosphate buffer: 10 mM potassium phosphate pH 7.6,1 mM dithiothreitol

• FTTR spectrometer • NaCl crystal discs

Method 1. Make 1 ml of liposomes as in Protocol 6A, using 2 mg of detergent-solubilized protein, but use 10 mM phosphate buffer instead of 50 mM. 2. Sediment the proteoliposomes using an ultracentrifuge (100 000 g,v for 1 h at 4°C) and remove the supernatant. Dry the inside of the centrifuge tube carefully with tissue. 3. Smear the sedimented sample between the crystal discs using a spatula. 4. To reduce the water content, allow the sample on the crystal to dry by exposing it to the air for 5 min. 5. Set up the FTTR instrument to record spectra with 2 nm sampling intervals over the range 400-4000 nm, with each spectrum averaged from at least four accumulations. 6. Scan the sample. Also perform a scan in the absence of the sample. Subtract this background scan (corresponding to the presence of water vapour) from the sample spectrum. 7. Use second-derivative analysis of the resulting amide I band—usually found centred at around 1656 cm-1 in the spectra of proteins that are predominantly helical—to identify the number and positions of the individual bands corresponding to discrete structural components. 8. Derive the spectral areas of each of these bands using a program such as Peaksolve.f this involves fitting the experimental bands to mixed Lorenzian-Gaussian bandshape functions. Note that the fractional areas of the component bands, assigned to different types of secondary structure, are taken to represent the proportion of the peptide chain in that structure. 1

Galactic Industries Corporation (Kore Technology Ltd)

162

THE AMPLIFIED EXPRESSION, IDENTIFICATION, PURFICATIOIM AND ASSAY

)

tMX)

Wavenumber (cm-2) Figure 7 FIR spectrum of the XylEiHisfe E. colt xylose-H " symport protein. Proteoliposomes of XylE(His) 6 were prepared as outlined in Protocol 6A. and FIR spectroscopy performed as in Protocol 9. (A) This shows the FIR spectrum obtained in the region of interest, 1500-1800 cm-1, indicating the positions of lipid, amide I, and amide II absorption. (B) Shows the second derivative of (A) from which structural information is obtained. Using Peaksolve {Galactic Industries, Kore Technology Ltd), XylE(His) 6 is predicted to comprise approximately 50% u-helix.

Protocol 10 Preparation of solubilized membrane protein for mass spectrometry* Equipment and reagents • Precipitation solvent: 10 ml acetone, 1 ml aqueous ammonia, and 100 mg trichloroacetk acid pre-dissolved in 100 M,! deionized water • Nitrogen or argon cylinder

Mass spectrometry solvent: chloroform:methanol:water:formicacid (100:100:33:2 (v/v/v/v))"1 Sonieation water bath or probe Anala R hexane (BDH)

Method t.

Add 100 \ti of purified detergent-solubilized protein, containing not less than 10 nmol protein, to 1.9 ml of the precipitation solvent.

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ALISON WARD ET AL. Protocol 10 continued

2. Vortex for 2 min. 3. Incubate on ice for 30 min. 4. Centrifuge in a microcentrifuge at 2000 gav for 3 min. 5. Resuspend the precipitate in 1 ml of ice-cold acetone using a sonication bath. 6. Centrifuge in a microcentrifuge at 2000 gav for 3 min. 7. Resuspend the pellet in 0.5 ml of hexane, 8. Remove the hexane by drying under a stream of nitrogen or argon. 9. Resuspend the pellet in a minimal amount of the mass spectrometry solvent. 0

From the method of Hufnagel et al (20). This method has been used successfully to prepare dodecyl-B-D>maltoside solubilized and purified E. coli 12a-helix inner membrane proteins for both MALDI (matrix-assisted laser desorption) and ESI (electrospray ionization} mass spectrometry. b This solvent may not work for all membrane proteins and other solvent systems should be tried (21). partial proteolysis, but it has not yet provided unequivocal identifications of the proteins.

8 Conclusions Out of the 16 membrane transport proteins listed in Table 2, we have only failed to amplify the expression of one, i.e. the L-rhamnosc-H transporter, Rha'l. Significantly, this protein is thought to comprise 10 membrane-spanning <*helices, not 12; and the N-tcrminus is in the periplasm (22) not the cytoplasm, which is thought to be the usual location. Of the remaining proteins only NupC proved resistant to amplified expression following the attachment of a -(His)fi tag to the C-terminus. With careful manipulation in Ji. coli, potentially using a variety of plasmid vectors, it is possible to amplify membrane protein expression up to 50% of the inner membrane protein. Such systems provide milligram quantities of protein for further study and structural analysis.

Acknowledgements We arc grateful to the following for their contributions towards this work: A. Aggeli, A. Ashcroft, M. Bacon. S. A. Baldwin, E. Barksby, N. Hoden, M. T. Cairns. J. Clough, C. L. Dent, S. M. I'erguson, M. Gallagher, M. A. T. Groves. F. Gunn, C. K. Hoyle, J, Keen, J. Knol, W. J, Liang, G. J. Lirhrrlnnd, V. Lucas, G. Martin, B. }. Mckeown, S. 1.. Palmer, K. Petro, P, Roberts, G. Smith, A. Steel, C. Tate, R. Venter, and J. Wood. We would also like to thank the EU, BBSRC, Wellcome Trust, and 164

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SmithKline-Beecham for financial support. This work was carried out under the auspices of the BBSRC-funded North of England Structural Biology Consortium.

References 1. Paulsen, I. T., Brown, M. H., and Skurray, R. A. (1998). J. Bacteriol., 180, 3477. 2. Mitchell, P., Roepe, P., Kaback, H. R., Henderson, P. J. F., Ames, G. F., Dean, D. A., Davidson, A. L, Nikaido, H., Erni, B., Jacobson, G., Sutrina, S. L, Schnetz, K., Ralk, B., Saier, M. H., and Maloney, P. (1990). Res. Microbiology, 141, 384. 3. Sakai, H. and Tsukihara, T. (1998). J. Biochem., 124, 1051. 4. Wendt, K. U., Lenhardt, A., and Shultz, G. E. (1999) J. Mol.Biol., 286, 175. 5. Griffith, J. K. and Sansom, C. E. (1997). The transporter facts book. Academic Press, London. 6. Grisshammer, R. and Tate, C. G. (1995). Q. Rev. Biophys., 28, 315. 7. Miroux, B. and Walker, J. E. (1996). J. MoL Biol., 260, 289. 8. Stark, M. R. J. (1987). Gene, 51, 255. 9. Gunn, F. J., Tate, C. G., and Henderson, P. J. F. (1994). Mol. Micrdbiol., 12, 799. 10. Roberts, P. (1992). PhD thesis. Cambridge University. 11. Voegele, R. T., Jung, H., Marshal, E. V., Culham, D. E., Ferguson, S., Henderson, P. J. F., Liang, W. J., Tripet, B., Hodges, R. S., and Wood, J. M. (1999). Biochemistry, 38, 1676. 12. Kaback, H. R. (1971). In Methods in enzymology. Vol. 22 (ed. W. B. Jakoby), p. 99. Academic Press, London. 13. Schaffher, W. and Weissmann, C. (1973). AnoZ. Biochem., 56, 502. 14. Henderson, P. J. F. and Macpherson, A. J. S. (1986). In Methods in enzymology, Vol. 125 (ed. S. Fleischer and B. Fleischer), p. 387. Academic Press, London. 15. Knol, J., Sjolleme, K., and Poolman, B. (1998). Biochemistry, 37, 16410. 16. Knol, J., Veenhoff, L. M., Liang, W. J., Henderson, P. J. F., Leblanc, G., and Poolman, B. (1996). J. Biol. Chem., 271, 15358. 17. Newman, M. J. and Wilson, T. H. (1980). J. Biol. Chem., 255, 10583. 18. Jung, H., Tebbe, S., Schmid, R., and Jung, K. (1998). Biochemistry, 37, 11083. 19. Patzlaff, J. S., Moeller, J. A., Barry, B. A., and Brooker, R. J., (1998). Biochemistry, 37, 15363. 20. Hufhagel, P., Schweiger, U., Ekerskorn, C., and Oesterheldt, D. (1996). Anal. Biochem., 213, 256. 21. Schindler, P. A., Vandorsselaar, A., and Falick, A. M. (1993). Anal. Biochem., 213, 256. 22. Tate, C. G., Muiry, J. A. R., and Henderson, P. J. F. (1992). J. Biol. Chem., 267, 6923.

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Chapter 7 Spectroscopic and kinetic approaches for probing the mechanisms of solute transporters Adrian R. Walmsley Division of Infection and Immunity, Institute of Biomedical and Life Sciences, University of Glasgow, Glasgow G12 8QQ

1 Introduction This chapter deals with the use of spectroscopic methods to monitor conformational changes in membrane transporters and kinetic approaches to timeresolve these changes. It outlines the use of fluorescence spectroscopy to monitor ligand-membrane protein interactions and the use of stopped-flow fluorescence spectroscopy to determine mechanisms of ligand binding and translocation. It also outlines the use of rapid-quench techniques for measuring the rate constants governing the translocation of the substrate across the membrane. To a large extent, these studies are illustrated by reference to the kinetics of substrate and inhibitor binding and translocation by various transporters studied in the author's laboratory.

2 Fluorescence spectroscopy for monitoring changes in the conformation of membrane transporters Fluorescence spectroscopy can be used to monitor the interaction of ligands with integral membrane proteins when the fluorescence of the protein-ligand complex differs from that of the free protein or ligand. Proteins contain natural fluorophores, notably the aromatic residues tyrosine and tryptophan. Tryptophan residues are excited maximally by light with a wavelength of 280 nm and emit light maximally with a wavelength of 340 nm. The excitation and emission maxima of tyrosine are 275 nm and 310 nm, respectively. However, for most proteins the tryptophan fluorescence predominates. The tryptophans can be specifically excited at 295 nm. When both tryptophan and tyrosine residues are 167

ADRIAN R. WALMSLEY

present within a protein, the maximum fluorescence signal can often be obtained by exciting the protein with light at about 280-285 nm. The binding of a ligand can induce a change in the fluorescence of the protein, due to movements of these aromatic residues. For example, when a tryptophan residue moves from a polar position, on the surface of the protein, to a more apolar position, buried within the protein interior, an increase in the tryptophan fluorescence can occur. This is due to shielding of the tryptophan residue from water molecules that quench the fluorescence by collision with the residue. The movement of a tryptophan residue from a polar to an apolar environment is commonly accompanied by a decrease in the excitation maximum, within the range 305 nm to 350 nm. Studies with collisional quenchers, such as acrylamide, also indicate a correlation between the excitation maximum and the degree of solvent exposure of tryptophan residues. However, interpretations of the molecular events involved in the conformational change of a protein are difficult when there are a number of aromatic residues. In any event, changes in the protein fluorescence can be used to determine the binding affinity and stoichiometry of a protein-ligand interaction. Other fluorophores can be used to monitor protein-ligand interactions. We have found that the hydrophobic fluorophore 8-anilino-l-napthalene-sulfonate (ANS) is a useful probe of conformational changes in membrane proteins (1). The probe binds to non-specific hydrophobic sites on the protein and can be excited either directly (X excitation = 370 nm, \ emission = 480 nm) or via tryptophan residues by fluorescence energy transfer (\ excitation = 295 nm, \ emission = 480 nm) (see Figure 1). In the former case, ligand-induced conformational changes in the protein that also propagate changes in these nonspecific hydrophobic sites can be monitored as changes in the ANS fluorescence. In the latter case, the movement of tryptophan residues can be monitored as a change in ANS fluorescence. Because there is little fluorescence of ANS excited at 295 nm, the background, against which changes in the fluorescence are monitored, is lower than that for direct measurements of the protein fluorescence. Changes in the tryptophan fluorescence are reflected in the ANS fluorescence because of the change in the amount of fluorescence energy available for transfer between the donor (i.e. tryptophan) and the acceptor (i.e. ANS) molecules. Furthermore, the tryptophan residue may also move to a position closer to the bound ANS molecule, resulting in an increase in the ANS fluorescence. This is a consequence of the fact that the efficiency of the fluorescence energy transfer is inversely proportional to the sixth power of the distance (r) between the fluorophores (i.e. 1/r6). We have found ANS useful in 'amplifying' the small sugar-induced changes in the tryptophan fluorescence of the o-galactose-proton symporter GalP from Escherichia coli. However, great care must be exercised when using this technique with membrane-bound proteins because the integration of ANS into the membrane also results in an increase in ANS fluorescence. This is not a problem in titration experiments where the cells or vesicles are equilibrated with the ANS prior to titration with the ligand that binds specifically to the transporter. 168

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300

400

500

Wavelength (nm) Rgure 1 Membranes (200 mg/ml) enriched in GalP were excited at 297 nm and the emission spectra recorded between 300-500 nm. The spectra shown are 100 n-M ANS (A), membranes (B), membranes equilibrated with 100 (xM ANS (C), and membranes equilibrated with both 100 IAM ANS and 200 mM o-galactose (C). The binding of ANS to GalP is characterized by a broad emission peak between 425-575 nm, the ANS (maximal excitation 370 nm, emission 450 nm) being excited via tryptophan residues in the protein. The substrate induces a large increase in the ANS fluorescence.

3 Equilibrium studies of ligand binding to membrane transporters As discussed above, ligand-induced changes in the protein fluorescence can be used to monitor the interaction of a ligand with a membrane transporter. By titrating the protein fluorescence with the ligand, the dissociation constant (Kd) for the ligand-protein complex can be determined. However, from a practical standpoint, the protein must be present in the membrane at a sufficient level to enable changes in its protein fluorescence to be monitored. For a purified, reconstituted protein there will be no background fluorescence from other proteins. However, this will not be the case for studies with native membranes, where the transporter protein must naturally occur at high levels. Alternatively, molecular-genetic approaches are frequently employed to induce the overproduction of the protein of interest. For example, measurement of the kinetics of the binding of cytochalasin B (a high-affinity inhibitor of sugar transport) to mammalian glucose transporters (GLUT1) has been possible using both protein reconstituted into lipid membranes and native membranes, where GLUT1 constitutes about 6% of the membrane protein (2). Another practical consideration is 169

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that, in contrast to soluble proteins, care must be taken in avoiding artefactual signals due to changes in the light-scattering properties of the vesicle systems employed. These changes in light scattering can occur as a consequence of vesicle sedimentation during the experiment, or be due to the shrinkage or swelling of the vesicles if the titration requires the addition of sufficient ligand to perturb the osmolariry of the medium appreciably. In studies using vesicle systems it is advisable to use a stirred cuvette and to monitor for changes in light scattering. In protein fluorescence titrations, light scattering can be conveniently monitored as changes in the absorbance at the emission wavelength (e.g. 340 nm). Alternatively, select a wavelength as low as convenient for maximum sensitivity (scattering being proportional to 1/X4), but at a position clear of any absorption bands. Care must also be taken when titrating a protein with a ligand that has optical (e.g. absorbance or fluorescence) properties of its own, since this may introduce a progressive inner-filter effect that complicates the changes in protein fluorescence. For example, if the ligand has an absorbance at 340 nm, then its addition will progressively decrease the protein fluorescence. A simple means to distinguish a true quench in the protein fluorescence, due to the binding of the ligand to the protein, from an inner-filter effect, is to position a cuvette directly in front of that containing the protein and to add ligand to this first cuvette. The first cuvette acts as a filter of the excitation light and if the ligand causes an inner-filter effect this will be seen as an apparent decrease in the fluorescence of the protein in the second cuvette.

3.1 Titration of the o-galactose-H+ symporter (GalP) with the antibiotic forskolin As an example, the titration of the GalP protein with forskolin is described here (3). Forskolin is a fungal metabolite that acts as a high-affinity inhibitor of both mammalian and bacterial sugar transporters. The GalP protein has been overexpressed in E. coli, so that the proportion of GalP in the membrane is elevated from less than 1% up to 50% of the membrane protein. Inside-out vesicles can be prepared from these GalP-overproducing strains, by disrupting the cells in a French press, and then using them directly for protein fluorescence studies. Titration of the protein fluorescence can be used to provide a measure of the Kd for the GalP-forskolin complex at equilibrium (see Figure 2). Alternatively, the JCd can be determined by equilibrium dialysis. The procedure described above can be adapted to determine the substrate Kd, where the substrate may cause a much smaller perturbation in the protein fluorescence. For example, the binding of sugars to GalP produces a much smaller quench in the protein fluorescence than with the high-affinity inhibitors cytochalasin B and forskolin. An approach to determining the sugar Kd is to repeat the forskolin (or cytochalasin B) titration in the presence of a series of sugar concentrations. The apparent Kd for the forskolin (measured in the presence of the sugar) will increase in a hyperbolic manner with the sugar concentration (for the situation where the sugar and inhibitor compete for the 170

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B

100

[Forskolinl

Figure 2 (A) Fluorescence emission spectra of membranes containing overexpressed levels of GatP. The upper trace (curve A) represents the emission spectrum of membranes (250 (ug/ml) in the absence of ligand. The lower trace (curve B) was recorded in the presence of 10 p.M forsholin. The protein was excited at 280 nm. (B) Fluorescence titration of GalP with forskolin. The quench in protein fluorescence was monitored (excitation 280 nm; emission 330 nm) as forskolin was added in small increments to membranes (250 ug/ml) until no further change was observed. The dissociation constant for forskolin was determined from the titration curve. The smooth line represents the fit to a hyperbolic equation with a K^ of 1.2 uM. The total fluorescence decrease was 8.3%.

Protocol 1 Titration of GalP with forskolin Equipment and reagents • Ruorimeter and quartz cuvette • Phosphate buffer: 50 mM sodium phosphate, 100 mM NaCI, 1 mM EDTA, pH 7.4

Inside-out inner membrane vesicles produced by French press disruption of Escherichia colt strains overexpressing GalP

Method 1. Resuspend the inside-out vesicles in the phosphate buffer to a protein concentration of 250 fig/ml. Place 2 ml of this suspension in a quartz cuvette and allow to equilibrate to 20°C. Stir the cuvette to maintain a homogeneous suspension of the vesicles, but not too vigorously so as to cause protein denaturation.

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Protocol 1 continued

2. Set the bandwidths of the excitation and emission beams to 3 nrn and 10 nm. respectively." Set the excitation wavelength to 280 nm and scan the emission between 310 and 450 nm. 3. Make microlitre additions of a concentrated (e.g. 10 mM) solution of forskolin to the cuvette and scan the protein fluorescence emission; ideally, so that the additions increase the concentration in a hyperbolic manner to span the concentration range from 0.2 Kd to 10 JCd. Ensure that these additions produce less than a 10% volume change, to minimize corrections for dilution, 4. Plot the fluorescence of the protein as a function of the fbrskoiin concentration. 5. Analyse the data; this is best done in terms of a quadratic equation describing the titration curve for a second-order binding process:6 ~ = AFn"ii/[T| • {(Kd + [T] + [/orskolin]) - V
"This allows maximization of the fluorescence signal (i.e. emission slit = 10 nm) but avoids photobleaching of the protein (i,e, excitation slit = 3 nm). * [T] and [fwsJcoftn] represent the initial transporter and forskoh'n concentrations, F0 is the initial fluorescence, AF is the change in fluorescence for a given concentration of forskolin, AF,™ is the total fluorescence change, and JQ is the dissociation constant. The titration experiment has the potential to yield both the K^ and [Tj. In practice, the transporter concentration can only be measured precisely under 'tight-binding' conditions, which occur when [T] is greater than the Kd. Under these conditions, most of the ligand added during the early part of the titration is bound. The titration curve (e.g. fluorescence change) increases almost linearly, rapidly reaching a plateau as the protein becomes saturated with bound ligand. Conversely, the Kd is more precisely determined under 'weak-binding1 conditions, which occur when [T) is less than the Kd, and there is a equilibrium between bound and free ligand. Under these conditions, the titration curve increases in a hyperbolic manner. Indeed, under these conditions Equation I tends towards that for a hyperbola: [2]

Generally, values for these parameters (i.e. Kd and [Tj) are generated by fitting the data directly to Equation 1 (or Equation 2) using a non-linear, regression curve-fitting program, such as SigmaPlot from SPSS Inc. Such an analysis of the data will provide the parameter values and their standard errors. same binding site) and the JQ for the sugar can b
PI

4 The kinetics of ligand binding and translocation 4.1 The stopped-flow instrument A detailed appraisal of the kinetic mechanism for the binding of ligands Lo a transporter protein often requires the use of specialized rapid-reaction 172

SPECTROSCOPIC AND KINETIC A P P R O A C H E S

techniques, such as stopped-flow fluorescence spectroscopy. The stopped-flow instrument allows the rapid mixing of the transporter with the ligand and enables one to time-resolve changes in the fluorescence on a millisecond time scale. A modern stopped-flow instrument, such as that supplied by Applied Photophysics, is shown in Iigure 3. It consists of a 150 W Xenon-lamp, two mo no chroma tors, to select the excitation and emission wavelengths, a photomultiplier tube (pmt) bolted to the second monochromntor, and a thermostatically controlled mixing block. Light is guided from the lamp to the mixing block via an optical light guide, illuminating the mixing chamber, and the emitted light is collected via a second optical light guide, which passes the light through the second monochromator to be detected by The pmt. Alternatively,

B

Figure 3 (A) The Applied Photophysics SX.18MV stopped-flow instrument. (B) A schematic diagram of the SX.1SMV stopped-flow instrument.

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the pmt can be bolted directly to the mixing block and the emission light wavelength selected by an intervening cut-off glass filter. This has the advantage of allowing the collection of more emitted light, increasing the sensitivity of the instrument. On the other hand, there are circumstances where it is preferable to focus on a specific wavelength of light, rather than simply collecting light above a cut-off wavelength. For example, in studies involving fluorescence resonance-energy transfer, where we wish to monitor the emission of the fluorescence donor and acceptor independently (see below). This problem can be overcome by using two photomultiplier tubes. The mixing block consists of a pneumatic drive (powered by nitrogen pressure from a gas cylinder) that when activated drives two glass syringes containing the reactants. The syringes are housed in a thermostatically controlled water bath and are loaded by two plastic syringes via Teflon valves. The glass syringes are mounted in a vertical position, which reduces cavitation (i.e. the production of air bubbles) and is convenient for removing air bubbles from the reactant solutions. The activation of the pneumatic piston drives the solutions through a mixing chamber and optical cell, filling the stop syringe, and the plunger movement activates the computer to record the reaction data. Generally, the time taken to mix the reactants and flush-out the optical cell (i.e. the instrument dead-time) is about 1 millisecond. The stop syringe automatically empties its contents after filling, in readiness for the next run, and the instrument can be set to undertake several consecutive runs for averaging. This feature is particularly useful when collecting data over a long timebase (e.g. 1000 seconds) or in generating temperaturedependency data. In the latter case the temperature of the water bath can be microprocessor-controlled and linked to the data acquisition, or more simply the instrument can be activated to make repeat measurements immediately after switching the bath to a lower/higher temperature (i.e. during the temperature re-equilibration time). More detailed information on the architecture of a generic stopped-flow instrument, and the determination of its dead-time, can be found in the companion volume Spectrophotometry and spectrofluorimetry in this 'A Practical Approach' series of books (4). Generally, ligand-binding experiments are set up to be performed under pseudo first-order conditions, in which the ligand concentration is in excess (e.g. 10-fold) over the protein concentration. Under these conditions, an exponential change in the fluorescence of the protein with time can be fitted, by non-linear regression, directly to an exponential equation defining a first-order binding process: AF = AFmax • Exp(kt)

[4]

where the AF is the change in fluorescence at time, t, AFmax is the maximum fluorescence change, and k is the rate constant. However, the reaction profile may be, and often is, more complex than a simple monophasic reaction. The equation to be fitted can include additional exponential terms to account for a multiphasic reaction. Although adding additional exponential terms to the fitted equation will improve the fit, this may only be apparent due to an 174

SPECTROSCOPIC AND KINETIC APPROACHES

increase in the degree's of freedom of the fit. Generally, the rate constants for two phases/processes must be 5- to 10-fold different to allow their accurate resolution. In practice, it is difficult to resolve more than three exponential phases from a stopped-flow trace, unless some of" these differ in their fluorescence response (e.g. one phase involves a fluorescence quench and another, a fluorescence enhancement). The ATP-induced. conformational changes of the ArsA protein, the catalytic subunit (i.e. ATPase) of the arsenite pump from I-. nili.

Protocol 2 Stopped-flow mixing experiments Equipment and reagents • Stopped-flow instrument • Appropriate transporter protein, buffer, and ligand solutions

• Plastic syringes • Glass syringes

Method 1. Load the two plastic syringes with the reagents (e.g. protein vesicles in one and the buffer solution in the other). Invert the syringes and tap to collect air bubbles at the nozzle. Slowly expel the air bubbles to leave a drop of solution on the end of the nozzle. Insert the syringes into the Teflon valves of the stopped-flow instrument. Remove air, if necessary, from the reactant solutions by pumping the solutions backward and forward between the glass and plastic syringes. Allow the solutions to equilibrate to the required temperature. 2. Set the excitation wavelength and slit width on monochromator 1. Set the emission wavelength using either a cut-off filter positioned directly in front of the pmt (bolted to the sample handling unit) or via monochromator 2. If necessary, set the flow volume of the stop syringe (e.g. usually to 100 pi but smaller volumes can be used, down to 50 p.1, with precious samples). 3. Activate the stopped-flow instrument several times to clean out and refill the observation cell with protein solution. 4. In fluorescence mode, set the pmt voltage (e.g. between 0 and 999 volts) to give a suitable signal voltage (e.g. usually 4 volts}, and back-off the signal to zero volts. Note that changes in fluorescence will be measured relative to this 4 V signal, 5. Replace the buffer solution with a solution of the ligand. Activate the stopped-flow instrument and collect data." 6. Repeat the experiment for a range of ligand concentrations. 7. Use non-linear regression software to fit the traces as discussed below, "The Applied-Photophysics stopped-flow apparatus allows data to be collected over a logarithmic timebase, avoiding the problem of repeating experiments if the 'correct' timebase isn't initially chosen with an unknown reaction or one with several phases.

ADRIAN R. WALMSLEY

provide an example of such a multiphasic process (see Figure 4) (5). Manual mixing procedures revealed a transient increase in the protein fluorescence, over 100-200 seconds (phases 1-3), followed by a slow decay in the fluorescence, over 1000 seconds (phase 4). This transient has been attributed to the formation of a pre steady-state intermediate. Stopped-flow experiments revealed more complex behaviour at shorter times. Over the first 10 seconds the profile was

D

Figure 4 (A) A stopped-flow record for the mixing of ArsA/500 uM ATP with 5 mM MgCI2. The record was generated over a long timebase, of 1000 seconds, and shows the characteristic transient increase (phases 1-3, see B-E) and decrease (phase 4) in fluorescence. (B-E) A stopped-flow record for the mixing of ArsA/500 (xM ATP with 5 mM MgCI2 shown over five timebases that differ from one another by an order of magnitude. The traces illustrate phase 3 (A), phase 2 and 3 (B), phase 1 and 2 (C), and phase 1 (D). The smooth curve through each trace is the best-fit to a triple-exponential equation with rate constants of 48.0 (± 1.02) s"1, 5.2 (± 0.26) s'1, and 0.027 (± 0.0023) s-1. One vertical division in traces A, B, C, and D represents a fluorescence change of 0.25, 0.5, 1.25, and 1.25%, respectively.

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SPECTROSCOPIC AND KINETIC APPROACHES

clearly multiphasic; with a very fast increase in fluorescence, followed by a moderately fast decrease and then a slow increase in fluorescence. Although the two fast phases merged, they were well resolved from the slow phase. Accordingly, the initial part of the trace was analysed as a double-exponential (during the first 4 seconds) and then as a single-exponential over the remainder of the trace. The three phases occurred with rate constants of 52 sec-1 (phase 1), 5.2 sec-1 (phase 2), and 0.026 sec-1 (phase 3), respectively. It was also possible to fit the entire data set to a triple exponential, which indicated similar rate constants of 48 sec-1, 5.2 sec-1, and 0.027 sec-1, for the three phases respectively. Consequently, it proved possible to time resolve the four phases of the reaction.

4.2 Ligand-binding kinetics The aim of most ligand-binding experiments is to exploit the changes in the protein fluorescence induced by the binding of ligands (e.g. substrates or inhibitors) to determine the kinetics of the interaction, and to deduce the kinetic mechanism for the binding process. An isomerization of the ligandprotein complex can be distinguished from the binding event because it occurs at a constant rate. Consequently, the 'mixing experiment' is repeated for a range of ligand concentrations. For a one-step binding process the apparent rate constant (kobs) will increase in a linear manner: Scheme I

where P is the protein (in conformation 1), L the ligand, kobs the measured rate constant, and k1 and k-1 are the association and dissociation rate constants, respectively. However, care should be taken to extend the range of concentrations to as high as is feasible because deviation from a linear dependency of the data may only then become apparent. The dissociation constant, Kd, can be calculated from the association and dissociation rate constants:

However, the high-affinity binding of a ligand is often attributable to its slow dissociation from the protein. Consequently, it is often difficult to determine the dissociation rate constant by extrapolation of the concentration dependency of the binding rate to zero ligand concentration. This difficulty can be overcome by measuring the dissociation rate constant directly in a displacement experiment (see below). The initial interaction of forskolin with GalP is an example of such a mechanism (but the overall binding mechanism is similar to Scheme II) (see Figure 5) (3). In the case of a two-step binding process, with rapid equilibrium binding (e.g. 177

ADRIAN R. WALMSLEY

_

10 Forskolin Figure 5 (A) A representative stopped-flow record of the time course for the binding of forskolin to GalP. The trace was generated by mixing membranes containing overexpressed GalP (100 ug/ml) with forskolin (2.5 (uM). The rate of quench in protein fluorescence (excitation = 297 nm, emission > 335 nm) was biphasic and best fitted by a double-exponential function as shown by the random scatter of the residual variance of the data about the best-fit doubleexponential (Panel (i)), compared to that of a single-exponential (Panel (ii)). (B) Measurement of kon and koff for forskolin binding. The rate of the fast phase determined from a fit to a double-exponential (O) increases linearly with increasing concentrations of forskolin, consistent with it being a single-step process. Linear regression analysis of the data yielded an apparent association rate constant (kon) from the slope of 6.2 per mM s-1, a dissociation rate constant (koff) from the ordinate intercept of 10.4 s-1, and an apparent dissociation constant (Kd) from koff/kon of 1.7 uM. The slower rate (A) was independent of concentration, indicative of an isomerization step, and ranged from 1.5 to 3.6 s-1.

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ki, k_! » manner:

k2, k_2), the apparent rate constant (kobs) will increase in a hyperbolic kill] ^2 P! + L ^ Pr -L v* P2-L

Scheme II

m)

[7]

where K1 is the dissociation constant and k2 and k-2 are the forward and back rate constants for a rate-limiting isomerization of the PL complex. In each case the data should be fitted to the appropriate equation (e.g. Equation 5 or 7) by nonlinear regression to provide values for the rate and equilibrium constants with their associated standard errors. Here, care must be taken to extend the range of concentrations for which measurements are made to as low as is feasible. Otherwise, measurements might only be made under near-saturating ligand concentrations and kobs will apparently have no concentration dependence, and may be mistaken for an isomerization process. The concentration dependence of the binding of MgATP to the ArsA protein (of the arsenical transporter discussed above) is an example of such a mechanism. The apparent rate constant for phase 1 increased in a hyperbolic manner with the MgATP concentration, indicative of the binding step, while the rate constants for phases 2-4 were independent of the MgATP concentration, indicative of isomerizations of the MgATP-ArsA complex (see Figure 6) (5). 60 -

so 40 -

30 -

20 -

10 -

500

1000

[ATP]

1500

2000

2500

3000

(uM)

Figure 6 The rates of MgATP-induced conformational changes of ArsA. A series of stoppedflow records were generated by pre-equilibrating ArsA with the indicated ATP concentration and then mixing with 5 mM MgCI2 in a stopped-flow device. The rates of the fast increase (•, phase 1) and decrease (•, phase 2) in the fluorescence of ArsA, induced by the binding of MgATP, are plotted as a function of the ATP concentration. The rate constant for phase 1 (•) increases in a hyperbolic manner and the curve through the data points is the best-fit to a hyperbolic equation, with minimal and maximal rates of 7.3 (± 3.10) s-1 and 53.7 (± 3.25) s-1, respectively, and a Kd of 178 (± 45.7) uM. The rate constant for phase 2 (•) was independent of the ATP concentration, varying non-systematically between 4.5 s-1 and 6.5 s-1, indicative of an isomerization process.

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The protein may adopt different conformations, existing as an equilibrium mixture of these conformational forms, but only one of which is able to bind the ligand. For example, GLUT1 is thought to exist in two conformations, an inward- and an outward-facing conformation with respect to the cell membrane. The inhibitors, cytochalasin B and forskolin, only bind to the inward-facing conformation, thus shifting the equilibrium towards this form. Under circumstances where the binding of the ligand to a protein is dependent upon a slow conformational change, kobs may decrease in a hyperbolic manner with increasing ligand concentration: P2-I

Scheme III

[8]

An example of such a binding mechanism is that of D-malate to the periplasmic-binding protein, DctP, a component of the C4-dicarboxylate binding protein-dependent transport system from Rhodobacter capsulatus (see Figure 7) (6). The binding protein can adopt conformations in which the substrate-binding site is either open or closed, and in the absence of the substrate these forms exist in equilibrium. Since the substrate can only bind to the open conformation, the equilibrium needs to be displaced to fully complex the protein with substrate. Knowledge of the kinetics of ligand binding, in conjunction with a measure of the overall equilibrium constant, can be usefully employed in defining the minimum number of steps in the binding process. As described above, titration of the protein fluorescence will provide such a measure of the overall equilibrium constant, and the Kd obtained is equivalent to K1/(l + K2) and K2(1 + l/K1:) for Schemes II and III (K1 and K2 are defined as k-1/k1 and k-2/k2, respectively). Since the kinetic experiments provide values for K1 and K2, it is possible to calculate the overall Kd and to compare this with the measured value. If the measured value is less than the calculated value then this suggests that there

Figure 7 The concentration dependence for the binding of D-malate to DctP. The curve through the data points represents the best fit of the data to Equation 7, which yielded values for k1 (L1, and K2 of 11.1 s-1, 167 s-1, and 25.5 uM, respectively.

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may be an additional isomerization of the PL complex. The effect is to tighten the binding by the factor 1/(1 + Kj), where Kj is the equilibrium constant for this additional isomerization step (i.e. Kd = Kcalc x l/(l + K1)). Conversely, if the measured value is more than the calculated value, then a pre-existing equilibrium between ligand binding and non-binding states may need to be displaced. The effect is to reduce the overall affinity, by the factor (1 + 1/K1), because this equilibrium needs to be overcome in forming the PL complex (i.e. Kd = Kcalc x (1 + 1/Ki)). For example, the overall Kd for the cytochalasin B-GalP complex, determined by titration of the protein fluorescence, is greater than that calculated from the kinetics of the binding of cytochalasin B (cf 1.9 H.M vs. 0.41 uM) (2). This indicates that GalP exists as an equilibrium mixture of at least two conformational forms that differ in their affinities for cytochalasin B. The simplest suggestion is that GalP can adopt inward- and outward-facing conformations but only the inward-facing form is able to bind cytochalasin B (by analogy to GLUT1).

4.3 Ligand-displacement kinetics The dissociation rate constant for a protein-ligand complex can often be determined by displacement using an alternative ligand, which differs in its fluorescent properties when complexed with the protein. Usually conditions are arranged so that the competing ligand is in vast excess over the bound ligand, so that its binding is rapid and it efficiently displaces the bound ligand by competition for the binding site. Under these conditions the reaction is rate limited by the dissociation of the bound ligand. An example is the displacement of forskolin bound to the o-galactose-H+ symporter, GalP, by D-galactose (see Figure 8). Forskolin produces a large quench in the protein fluorescence of GalP compared to that with D-galactose. Consequently, when the GalP-foskolin complex (generated by pre-mixing 100 ug/ml protein with 10 uM forskolin) is mixed with an excess of D-galactose (100 mM) there is an increase in the protein fluorescence as the forskolin is displaced (3). Competition experiments can also be set up to determine the Kd for a ligand. For example, the binding of cytochalasin B to sugar transporters can be readily determined by stopped-flow fluorescence spectroscopy. The inclusion of a transported sugar in the assay reduces the apparent rate of binding of cytochalasin B (see Figure 9) (7). If the sugar competes with the cytochalasin B for binding to the same site, then the apparent rate constant will vary as a function of both the sugar and cytochalasin B concentrations:

where kobs is the apparent rate constant for cytochalasin B, kcB.oN and )CCB.OFF are the association and dissociation rate constants for cytochalasin B, respectively, and Ks is the apparent Kd for the sugar. The data is best analysed by non-linear regression analysis of Equation 9, with k0bs, [sugar] and [cytochalasin B] as variables, and KCB.ON. KCB.OFF and Ks as parameters to be fitted. 181

ADRIAN R. WALMSLEY

0.4

0,8

1.2

1.6

2.0

Time (s)

o Figure 8 The displacement of forskolin from GalP by ogalactose. Trace A shows the increase in protein fluorescence (excitation 297 nm, emission > 335 nm) when membranes (100 mg/ml) equilibrated with forskolin (10 ^M) were mixed in the stopped-flow apparatus with o-galactose (200 mM). The resulting change in protein fluorescence was fitted to a double-exponential function, revealing dissociation rate constants of 12.3 s-1 and 2.4 s-1. Trace D is the normalized residual variance of the data about the best-fit curve. Traces B and C show that mixing with buffer + L-galactose (200 mM) or buffer alone, respectively, do not displace forskolin from the protein.

4.4 Measurements of transporter reorientation Many transporters are proposed to operate by cycling between conformations in which the substrate-binding site is inward- and outward-facing and the substrate is transported across the membrane during this conformational change. Reorientation occurs both in the presence and absence of transported substrates. Ti

V-s

T 0-S -S Scheme IV

A kinetic model for this process is shown in Scheme IV, with T0 and Ti representing the outward- and inward-facing conformations, respectively; S is the substrate, and the rate constants governing the cycle are designated k1-k4. Such a model has been proposed for the operation of the GLUT1 and other sugar transporters (8). The binding of sugars to the transporter is a rapid equilibrium process and reorientation of the unleaded-transporter is the ratelimiting step (i.e. k4 and k_4 are the slowest steps). For this reason the equilib182

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[Cytochalasin B] Figure 9 The binding of Cytochalasin B to the i-arabinose transporter (AraE) from E. coli in the presence of D- and L-arabinose. The Cytochalasin B binding rate is plotted as a function of the concentration of Cytochalasin B in the absence of i-arabinose (A) and in the presence of 2.5 mM (•) and 150 mM (A) L-arabinose, and 150 mM o-arabinose (D).

rium exchange (governed by k2 and k_2) of sugars across the membrane is more rapid than either net influx (governed by k4) or efflux (governed byfe_4).This situation has been exploited in determining the rate constants governing reorientation (9, 10). The outward-facing site has a much higher affinity for the non-transported sugar analogue 4,6-ethylidene-D-glucose than the inward-facing site. Consequently, when leaky vesicles, into which GLUTl has been reconstituted, are mixed with 4,6-ethylidene-D-glucose the transporters are largely sequestered into the outward-facing conformation. This redistribution of transporter conformations is concomitant with a decrease in the protein fluorescence, which is attributed to T0 having a lower intrinsic fluorescence than T,. The quench in protein fluorescence, induced by mixing GLUTl vesicles with 4,6ethylidene-D-glucose, can be time-resolved by stopped-flow fluorescence spectroscopy. As the 4,6-ethylidene-D-glucose concentration is increased kobs decreases in a hyperbolic manner (see Figure 10) according to the following equation:

10

where EGlc is 4,6-ethylidene-D-glucose, k4 and k_4 are the outward-inward and inward-outward reorientation rate constants, respectively, and Kto and kli are the affinities of the outward- and inward-facing sites for 4,6-ethylidene-D-glucose. 183

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15

Figure 10 The concentration dependence of kobs when reconstituted GLUT1 was mixed with 4 6-ethylidene-oglucose (EGlc). The data were fitted to Equation 10 to yield values for k4, k4, K10 and Kj, of 9.5 s'1, 5.2 s'1, 2.5 mM, and 116.8 mM, respectively. The smooth curve through the data points was generated using these constants.

The data were fitted to Equation W to provide values for the rate and equilibrium constants. Such an analysis can be extended by including a transported sugar to accelerate the rate of T0-Tj interconversion.

4.5 Single-turnover experiments to determine the translocation rate constants The rate constants governing the translocation of substrate across the membrane can be determined in single-turnover experiments. A transport inhibitor, which competes with the substrate for the binding site, is used to orientate the transporters (e.g. to pull them into the outward-facing conformation). The transporters are then diluted into a solution of radiolabelled substrate. The resulting drop in inhibitor concentration allows a proportion of the transporters to bind the substrate and turnover, to produce an initial burst in substrate uptake. Relaxation of the transport system can only occur when the substrate has equilibrated across the membrane, and the time taken is given by the formula: where k, and k4 are the rate constants for inward and outward reorientation of the unloaded transporter; k_2 is the rate constant for outward reorientation of the substrate-loaded transporter; ki is the inhibition constant for the inhibitor (I) and Kso is the kd for substrate (S) outside the cell (denoted o). Thus, if kobs is determined as a function of the substrate concentration, then rate constants k4, k_4, and k_2 and the true affinity of the outward-facing transporter can be determined. In practice, it may be necessary to perform these experiments using rapid-reaction equipment, such as quenched-flow and rapid-filtration instruments.

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More elaborate single-turnover experiments can be initiated for active transport systems. For example, a single turnover of the Na+,K+-ATPase can be initiated by the release of a sub-stoichiometric amount of ATP by flashphotolysis of caged ATP (11).

5 Extrinsic probes to monitor transporter conformational changes The preceding discussion focused on the use of the intrinsic fluorescence of the protein to monitor ligand-protein interactions. However, there are many examples where membrane transporters have been labelled with fluorescent probes to monitor ligand-induced conformational changes. For example, the Na+,K+-ATPase has been covalently labelled with fluorescein 5'-isothiocyanate to monitor conformational changes by stopped-flow fluorescence spectroscbpy (12). Conformational changes in GalP can be monitored from changes in the fluorescence of non-covalently bound ANS (1).

6 A steady-state approach to determining rate constants governing the translocation cycle If the steady-state parameters (e.g. Vmax, the maximal rate of transport; KM, the half-saturation constant) for substrate flux are measured at a series of different temperatures then it may be possible to extract the rate constants governing the translocation cycle depicted in kinetic Scheme IV. Given that substrate binding and release are fast compared to transporter reorientation, simplified expressions for the steady-state parameters for exchange and net flux can be derived according to kinetic Scheme IV: v exchange =

-

exchange

/

1

=

where k_1/k1 (= KMo) and k-3/k3 (= KMi) are the true dissociation constants of the substrate complexes of the outward- and inward-facing transporter and [T] is the transporter concentration. Since the temperature dependence of the rate constants for transporter reorientation can be expected to be described by the Arrhenius equation, an analytical relationship for the temperature dependence of the various Vmax can be derived. Thus, according to the Arrhenius equation:

where k represents a particular rate constant (with k(273K) its value at 273 kelvin), Ea is the corresponding activation energy, T the temperature (in kelvin), and R the gas constant. Hence:

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ADRIAN R. WALMSLEY

where Vmax (temp) is the Vmax at a particular temperature T and El and E2 represent the activation energies of k2, ^-2- k4. or k-4 depending upon the type of transport (for example, E1 and E2 represent k2 and k-4 during net influx). The method is applicable when the various rate constants differ in their temperature dependencies, leading to curved Arrhenius plots. It has been utilized in analysing the translocation cycle of the mammalian glucose transporter, GLUT1, from human erythrocytes (13). The temperature dependence of Vmax for the net flux and equilibrium exchange of D-glucose in erythrocytes was determined, revealing a curvature in the corresponding Arrhenius plots (see Figure 11). The data from influx, efflux, and equilibrium exchange experiments

Figure 11 The temperature dependencies of Vmax for the influx of D[14C]glucose into human red cells under zero trans (•, O) and equilibrium exchange (•, D) conditions. These data were used in a non-linear regression analysis of the glucose translocation cycle, generating the rate constants for reorientation of the transporter with and without sugar. This analysis yielded the fitted rate constants and their activation energies shown in Table 1. Table 1 Fitted rate constants and their activation energies Parameter

Rate constant (per sec) (0°C)

Activation energy (kJ/mol)

k2

1113 ± 494 90.3 ±3.47 12.1 ± 0.98 0.726 ± 0.498

31.7 ± 5.11 88.0 ± 6.17 127.0 ± 4.78 173.0 ± 3.1

K-2

k4 k-4

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SPECTROSCOPIC AND KINETIC APPROACHES

Protocol 3 Determination of the temperature dependence of the steady-state parameters for cellular transport Reagents • Cells, buffers, radiolabelled substrate Method 1. Deplete the cells of the substrate under investigation. For example, deplete glucose by repeatedly washing the cells with buffer. 2. Measure the initial rates of influx into the cells using an appropriately radiolabelled substrate (so-called zero-trans influx). Initiate the experiment by diluting the cells into a solution of the radiolabelled substrate. Terminate uptake by mixing with a large excess of cold inhibitor (e.g. phloretin for glucose transport into eiythrocytes). Collect the cells by centrifiigation, wash with a solution containing a transport inhibitor, to prevent substrate efflux, collect by centrifugation., and disrupt with trichloroacetic acid (TCA). Separate the supernatant from the solid precipitate by centrifugation and assay for radioactivity by liquid scintillation counting. If desired, Prior to collecting and disrupting the cells, remove a sample for a protein assay. To measure true initial rates, ensure that uptake is linear with time and represents less than 10% of the equilibrium filling of the cells with the substrate. 3. Measure initial rates of (zero-trans) efflux from cells loaded with radiolabelled substrate. To load the cells with substrate, repeatedly wash with a solution of the substrate, and collect the cells by centrifugation. To initiate the experiment, dilute the cells into a large volume of buffer, so that the extracellular substrate concentration is negligible. Terminate efflux by mixing with a large excess of ice-cold inhibitor and collect the cells by centrifugation. Alternatively, rapidly filter the cells. Either measure the radioactivity in the extracellular medium or in the cells (as described above). 4. Measure the initial rates of equilibrium exchange flux. These measurements are most conveniently made by loading the cells (as for efflux measurements) with the unlabelled substrate and then mixing the cells with radiolabelled substrate at an equivalent concentration. Note that under these conditions there is no net flux of the substrate, just an exchange of radiolabelled for non-labelled substrate across the membrane. Collect and treat the cells as for uptake experiments. 5. Repeat the measurements for a range of concentrations approximately spanning 0,2-10 KM and determine the KM by fitting the data, by non-linear regression, to an equation for a hyperbola:

where v is the rate of transport, Vmax is the maximal transport rate, KM is the halfsaturation constant. and S is the substrate.

ADRIAN R. WALMSLEY Protocol 3 continued

6. Repeat the measurements over as wide a temperature range as possible. 7. Plot the Vmilx data in the form of an Arrhenius plot (i.e. Infy^) vs. 103/T (kelvin)). Note that if the plots are curved then it may be possible to apply the analysis described above to determine the rate constants and activation energies for individual steps of a defined mechanism. 8. Determine the rate constants (and associated activation energies) governing kinetic Scheme IV by the following fitting procedure: (a) Estimate the rate constants k-4 and k2 (at 0°C) from the value of Vmax for zerotrans influx and equilibrium exchange, respectively. (b) Estimate the activation energies forfork_4and k2 from the slopes of the two extremes of the Arrhenius plot for Vmax for zero-trans influx. (c) Use these estimates as the starting points for obtaining k-4 and k2 (at 273 K) and their activation energies by non-linear regression fitting to the equation: Vmax (zt-influx) = [T]/k4(273k) • exp[(Ea/R)(l/273 - 1/T)] + k2(273k) exp [(Ea/R)( 1/2 7 3 - l/T)] [21] (d) Repeat this procedure with the temperature dependency data for zero-trans efflux to provide estimates of k4, k_2 and their activation energies. (e) Set the rate constant k2 and its activation energy to the values from the fit to Equation 20, and set the rate constant k_2 and its activation energy according to the data obtained for Vmax equilibrium exchange. Use these estimates as starting points to find best-fits for k2, k-2, and their activation energies by non-linear regression curve fitting of the Vmax equilibrium-exchange temperature dependency data to an equation analogous to Equation 21. (f) Using the values obtained above for k2, k -2 .k4,k-4. and their respective activation energies, perform a global analysis of the data (i.e. fit the data simultaneously to the three equations for the temperature dependence of Vmax under zero-trans influx, efflux, and equilibrium exchange conditions), thereby generating a set of values for the rate constants and activation energies that are simultaneously consistent with all the data: Vmax = [T]/A(k2(273kl • exp[(Ea/R)(l/273 - 1/T)]} + • exp[(Ea/R)(l/273• exp[(Ea/R)(l/273 exp[(Ea/RKl/273 -1/7)]) [22] where A, B, C, and D have values of 0 or 1 depending upon the type of transport. Enter this information as a variable along with the Vmax and corresponding temperature values (e.g. y = Vmax; x - temperature; and z = 1 (zt -- influx), 0 (zt - efflux), or -1 (equilibrium exchange). If z = 1 then: A = D = 1 and B = C = 0; if z = Othen:B = C = 1 and A = D = 0; if z = -\ then: A = B = 1 and C = D = 0). (g) Using the values generated for k2, k_2, k4, k_4 calculate the dissociation constants for the binding of substrate to the outward-facing and inward-facing transporter from the measured KM values according to Equations 15-17.

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were pooled and fitted simultaneously to the three equations for the temperature dependence of Vmax, providing values for k2, k_2, k4, and k_4 and their activation energies. Although the fitting procedure described above is a powerful technique for obtaining pre steady-state information from steady-state measurements, great care must be exercised in its use because non-linear Arrhenius plots could occur for a variety of different reasons: (a) The transporter undergoes irreversible heat inactivation at higher temperatures, leading to a reduction in Vmax. (b) Change occurs in the environment of the transporter. Many membrane transporters undergo a change in the apparent activation energy at a transition point in the state of the lipid membrane, as it changes from a fluid to a crystalline state at lower temperatures. (c) The transporter can exist in two different conformational forms that are both active but have different activation energies. (d) The transporter is reversibly inactivated, so that the transporter exists as an equilibrium mixture of active and inactive molecules. In the case of the glucose transporter it was possible to measure some of the individual rate constants, by stopped-flow spectrofluorimetry, at a series of temperatures and to show that these yielded linear Arrhenius plots. Hence, the non-linearity of the Arrhenius plots of Vmax could not be attributed to denaturation of the transporter or membrane phase transitions.

7 Thermodynamics The thermodynamics of a reaction can be obtained from the temperature dependency of an equilibrium constant (K), which varies according to the van't Hoff equation: l

where T is the absolute temperature (kelvin), AH° is the standard enthalpy change for the reaction (kj), and R is the gas constant. Thus the equilibrium constant increases exponentially with temperature. The integration of Equation. 23 gives the following equation:

where InA is a constant of integration. The enthalpy change (AH°) can be obtained conveniently from the slope of a linear plot of InK vs. 103/T: slope = -AH°. Knowledge of the enthalpy change allows further thermodynamic information to be obtained, by application of standard relationships. The Gibbs free 189

ADRIAN R. WALMSLEY

energy (AG°) of a reaction (kj/mol), which is a measure of how likely a reaction is to occur, is given by the following equation: AG° = -RTlnK = AH° - TAS°

[25]

0

where AS is the standard entropy change (kj/mol). Essentially, AH° is the heat required for, or generated, in the course of a reaction. When heat is generated (AH° is negative), the process is termed exothermic; and when heat is required to drive a reaction (AH° is positive), it is termed endothermic. The simplest definition of the entropy (AS0) of a reaction is the change in disorder of the system. When AG° is negative, the reaction will have a tendency to occur spontaneously. Thus, a reaction is likely to be spontaneous if it is exothermic and involves an increase in the disorder of the system. For example, the binding of D-glucose, to the GLUT1 protein involves little change in the enthalpy but a large entropy change. The reaction is entropically driven. This behaviour was postulated to be due to the exchange of hydrogen bonds between the sugar and water for those with the transporter, leading to little change in the enthalpy. However, the release of ordered water from around the sugar and from the sugar-binding site of GLUT1 leads to an increase in entropy that is not compensated for by the condensation of the sugar and protein. Transitions of the transporter between inward- and outward-facing conformations can be considered as equivalent to chemical reactions whose rate is limited by the need to attain sufficient energy to reach an intermediate transition state. The enthalpies, entropies, and free energies associated with these transition states can be calculated from the Boltzmann equation: h

where k is a rate constant (e.g. k1 — k4 or k_1 — k_4 in Scheme IV), with corresponding activation energy Ea; kB is the Boltzmann constant, T is the absolute temperature, h is Planck's constant, and R is the gas constant. The Gibbs free energy (AG), entropy (AS), and enthalpy (AH) changes for formation of the transition state are related by the following equations: AH = Ea + RT, AG = AH - TAS

[27] [28]

Such an analysis has been applied to the glucose transporter, GLUT1 , to provide a profile of the thermodynamic changes involved in glucose binding and translocation (see Figure 12). The results have largely been interpreted in terms of the hydration state of the transporter. The slight change in enthalpy (endothermic outside and exothermic inside) and the moderate increase in entropy for glucose binding have been attributed to the dissociation of water from both glucose and the binding site of the transporter, as glucose effectively exchanges hydrogen bonds with water for a similar number of hydrogen bonds with the transporter. The glucose preclusion of water, which transiently stabilizes the 190

SPECTROSCOPIC AND KINETIC APPROACHES

Figure 12 Gibbs free energy (c), enthalpy (b), and entropy (a) diagrams associated with the transport of glucose via the mammalian GLUT1 glucose transporter at 37°C. Standard Gibbs free energies are shown except for glucose binding, for which basic Gibbs free energies, with glucose at 5 mM on both sides of the membrane, are given.

binding site, would then tend to reduce the energy barrier to reorientation of the glucose-loaded transporter relative to that of the unloaded transporter. The increase in enthalpy and entropy in going from the inward- to outward-facing conformation could be attributed to the inward-facing transporter being more highly hydrated.

References 1. Walmsley, A. R., Martin, G. E. M., and Henderson, P. J. F. (1994). J. Biol. Chem., 269, 17009. 2. Walmsley, A. R., Lowe, A. G., and Henderson, P. J. F. (1994). Eur.J. Biochem., 221, 513. 3. Martin, G. E. M., Rutherford, N. G., Henderson, P. J. F., and Walmsley, A. R. (1995). Biochem.J.,308, 261. 4. Harris, D. A. and Bashford, C. L. (ed.) (1987). Spectrophotometry and spectrofluorimetry: a practical approach. IRL Press, Oxford. 5. Walmsley, A. R., Zhou, T., Borges-Walmsley, M. I., and Rosen, B. P. (1999). J. Biol. Chem. 274, 16153. 6. Walmsley, A. R., Shaw, J. G., and Kelly, D. J. (1992). J. Bid. Chem., 267, 8064. 7. Walmsley, A. R., Petro, K. R., and Henderson, P. J. F. (1993). Eur.j. Biochem., 215, 43. 8. Walmsley, A. R. (1988). Trends Biochem. Set., 13, 226. 9. Appleman, J. R. and Lienhard, G. E. (1985). J. Biol. Chem., 260, 4575.

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192

Appleman, J. R. and Lienhard, G. E. (1989). Biochemistry, 28, 8221. Forbush, Bliss III (1984). Proc. NatlAcad. Sci. USA, 81, 5310. Smimova, I. N., Lin, S-H., and Faller, L. D. (1995). Biochemistry, 34, 8657. Lowe, A. G. and Walmsley, A. R. (1986). Biochim. Biophys. Acta, 857, 146.

Chapter 8 Detection and analysis of glucose transporters using photolabelling techniques Alison K. Gillingham, Frangoise Koumanov, Makoto Hashimoto, and Geoffrey D. Holman School of Biology and Biochemistry, South Building, University of Bath, Claverton Down, Bath BA2 7AY

1 Methods for photolabelling glucose transporters The use of photolabels to study biological systems has emerged as a powerful technique for tagging proteins, and the range of approaches and types of ligand that can be employed have been extensively reviewed (1). Although various affinity labelling approaches have been tried (for example using the highly reactive isothiocyanate group substituted into maltose (2)), these have generally been unsuccessful in labelling glucose transporters. This difficulty can be largely attributed to the low affinity of glucose transporters for their substrate and the poor tolerance for groups substituted around the glucopyranose ring. Specificity studies (3) indicate that there is a close approach of the binding site around the sugar as it enters from the external solution. These properties of the glucose-transporter binding site have limited the range of approaches that can be used in photoaffinity labelling studies. Often photoreactive substituents are rather bulky and hydrophobic, and based on benzoic acid substituted with azides or diazirine substituents. However, two successful approaches have emerged and these are described in detail in this chapter. First, it has been discovered that two naturally occurring ligands of glucose transporters, cytochalasin B and forskolin, have such high affinity for the protein that UV irradiation of the protein leads to crosslinking (this has been attributed to the activation of tyrosine and tryptophan side chains that absorb relatively long-wavelength UV light). This approach therefore circumvents the need to introduce bulky photoreactive substituents into the ligand, although iodoazido derivatives of forskolin have been synthesized and used successfully in labelling glucose transporters (4). Second, our group has developed an approach that utilizes the specificity of the transporter and the hydrophilicity of sugar-based ligands to confine the 193

ALISON K. GILLINGHAM ET AL.

interaction between the transporter and the labelling compound to the exofacial membrane surface. We have substituted into the C-4 hydroxy position of the hexose, as specificity studies indicated that bulky groups would be tolerated there. In addition, we have employed bis-hexoses as photolabels since the introduction of a second hexose can balance the hydrophobicity of the photoreactive moiety with the increased number of hydrophilic hydroxy groups in the bishexose structure. Both the cytochalasin B and bis-hexose labelling methods are described in detail in the protocols below. The two approaches are complementary for several reasons: First, cytochalasin B labels the endofacial-binding site of the glucose transporters, while the bis-hexoses label the exofacial surface (5, 6). Second, cytochalasin B can label transporters in isolated membrane fractions and, because the compound is hydrophobic, its label will crosslink to all the glucose transporters within the cell of interest. By contrast, the bis-hexoses will only label those transporters that reside at the cell surface. As the compounds are impermeant, they do not reach those glucose transporters that are sequestered in intracellular membrane compartments of the cell. This property has been used to advantage in studying the insulin-induced translocation of GLUT4 to the cell surface of insulin target cells. Furthermore, the time courses and kinetics of subcellular trafficking of photolabel-tagged GLUT4 can be studied with these ligands (7, 8).

2 Photoactivation methods The ligands used for photochemistry are usually substituted with either nitrene precursors (from aryl-azides) or carbene precursors (from benzophenones and aryl-diazirines). The photoreactive moieties therefore absorb light at wavelengths between 260 and 350 nm (up to 500 nm for some nitro-substituted arylazides). The most effective wavelength for irradiating biological samples is therefore between 300 and 350 nm, where UV damage of the protein is minimized. UV light sources suitable for irradiation in this range are available from several manufacturers and are based on a mercury filament. However, if one uses a mercury lamp then one needs a high-power lamp (0.5-1 kW) as lowpower lamps only emit the short mercury line at 256 nm. The high-power mercury lamps have a broad emission spectrum; however, to select wavelengths suitable for irradiating biological samples, low-wavelength UV cut-off filters have to be employed to reduce the intensity of wavelengths below 300 nM. Irradiation through a thick (2-5 mm) layer of glass reduces most of this harmful irradiation. An alternative to the high-power mercury lamp is provided by the mercury phosphor lamps. These use the short mercury line at 256 nm to excite a phosphor that emits a higher wavelength for excitation at 300-350 nm. These phosphor lamps are low power but emit a more easily controlled output. Our laboratory uses a Rayonet photochemical reactor with 16 low-power (5 W) 300 nm or 350 nm lamps. The samples to be irradiated are placed in the centre 194

DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS

of the photoreactor approximately 20 cm from the lamps. The UV output reaching the samples is approximately 9200 W/cm2. The 300 nm lamps are used for activating the interaction between cytochalasin B and the transporters, and for most cell-surface labelling studies using the bis-mannose compounds. For experiments in which we study the subcellular trafficking of photolabel-tagged glucose transporters, we either irradiate through a thick layer of glass or, more conveniently, irradiate using the 350 nm lamps. The activation of the ligand results in covalent insertion of the ligand into the transporter. The type of amino acids that are labelled is somewhat dependent on whether a nitrene or a carbene precursor compound is used. The arylnitrenes tend to undergo nucleophilic side reactions that result in extensive non-specific labelling. In addition, carbene-labelled proteins are more stable than nitrene-labelled proteins under HPLC conditions in trifluoroacetic acidacetonitrile. These are the conditions necessary to purify labelled fragments and determine the amino acid sites of labelling within a protein (9). The carbene precursor compounds (see Figure 1) are preferred as these insert rapidly into protein but do not react significantly with solvent. The 3-trifluoromethyl3-phenyl diazirines are known to be stable under a wide range of chemical and biological conditions and are very versatile in the range of additional substituents that can be introduced into the phenyl ring (10). Irradiation times for the benzophenone compounds have to be very long and this is limiting when rapid processing of intact cells is required (as in cell-surface labelling experiments). However, the levels of non-specific side reactions are low with these ligands. Irradiation times for diazirines have to be a little longer than for the aryl-azides, but the levels of non-specific interactions are much lower. On balance we find (as do others, see ref. 11) that the aryl-diazirine derivatives are the most useful of the currently available photolabel precursor compounds. 2.1 Photolabelling with cytochalasin B Cytochalasin B has mainly been used for the study of GLUT1 in erythrocyte membranes, and a suitable method for carrying out this type of labelling experiment is described in Protocol 1. Cytochalasin B labelling has also been carried out using intact cells (12); but since long irradiation times are necessary to obtain measurable levels of incorporation of ligand, this approach has not been widely used.

Figure 1 Mechanism of insertion of the photolabel into target proteins upon irradiation.

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ALISON K. GILLINGHAM ET AL.

Protocol 1 Photolabelling glucose transporters with [3H] cytochalasin B Equipment and reagents • UV photoreactor with 300 nm lamps » 35 mm2 plastic Petri dishes (Falcon, Nunc, or Greiner) • [4(n)-3H]Cytochalasin B (Amersham, specific activity 699 GBq/mmol) • 10 mM stock solution of cytochalasin E or cytochalasin D (Sigma) in ethanol (used to reduce non-specific background labelling)

• Human erythrocytes or erythiocyte ghosts prepared according to Baldwin and Lienhard(13) • PBS buffer: 154 mM NaCl, 12,5 mM NaH2PO4, pH 7.04 • Incubation buffer: for labelling human erythrocytes or ghosts use phosphatebuffered saline

Method 1. In a 35 mm2 Petri dish mix together 1 ml of cell suspension (human erythrocytes at 10% cytocrit or isolated rat adipocytes at 40% cytocrit) and 1 fxl of a 10 mM stock of cytochalasin D or E (final concentration 10 u.M),u 2. Add 0.5 uM of [3H]cytochalasin B (final concentration), and incubate for 20 mm at

18C.b 3.

Transfer the Petri dish to the photoreactor and irradiate for 45 sec with 300 nm UV light.

4.

Wash the cells four times at room temperature with an appropriate incubation buffer (see above), to remove unbound cytochalasin molecules. For erythrocytes, wash with 1 ml buffer in an eppendorf tube, centrifuging at 3000 g and discarding the supernatants. For adipocytes, wash with 15 ml buffer in a 20 ml plastic tube, briefly centrifuging at 300 g and discarding the infranatants.

5.

Analyse photolabel incorporation as appropriate (see Protocols 3-4),

"Additives such as 500 mM D-glucose or i-glucose may be included in the incubation buffer before photolabelling. ''The final ethanol concentration should not exceed 0.1%.

2.2 Photolabelling glucose transporters with membrane-impermeant substrate analogues Bis-hexo.ses have been developed to study cell surface-loaned glucose transporters (14, 15). These include 1,3-bis (i)-mannos-4-yloxy(-2-propylamine (BMPA), which has an aminopropane spacer between two mannose moieties. The amino group of this spacer can be coupled to a range of pholoactlivable groups. The resulting compounds arc hydrophilic and cell-impermeant. are relatively stable in the dark, and arc soluble in most physiological buffers (14), They exhibit higher affinity for glucose transporters than their parent compound, D-mannose, but have much lower affinity than the inhibitors cytochalasin B (5. 16) and 3-iodo-4azidophenethylamido-7-O-succinyldeacetyl-forskolin (lAPS-fbrskolin; (17)). Thus 196

DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS

an important consideration for photolabels which are sugar analogues and have relatively low affinity for the glucose transporters, is the choice of photoactivable molecule, since one with high selectivity will reduce non-specific labelling. The azidosalicoyl derivative of BMPA (ASA-BMPA) allows selective labelling of the human erythrocyte glucose transporter, GLUT1 (18). However, this photolabel displays poor affinity towards the insulin-sensitive glucose transporter GLUT4. These findings led to the investigation of photoreactive ketones as possible alternative candidates for reagents able to affinity-label GLUT4 and other glucose transporters. The highly reactive benzophenone derivative 2-N-(4benzoyl) benzoyl-BMPA (BB-BMPA), which under UV irradiation produces a radical that can insert into the target protein, has been described (19). This photolabel displays improved selectivity for GLUT4, but requires a relatively long irradiation time to achieve effective photolabelling. An alternative is to use a diazirine substituent as the carbene precursor, as in the compound ATB-BMPA (2-N-[4-(l-azi-2,2,2-trifluoroethyl)benzoyl]-l,3-bis-(D-mannose-4-yloxy)2-propylamine (ATB-BMPA)) (20). Since its first report, ATB-BMPA has been used extensively for studying the properties of glucose transporter isoforms. The photolabel displays a high affinity for the erythrocyte glucose transporter GLUT1 (with a Ki of approximately 300 uM) as well as for the insulin-sensitive isoform GLUT4 in rat adipocytes (Ki for basal and insulin-stimulated glucose transport is 247 uM) (21). In combination with isoform-specific antibody immunoprecipitation the photolabel has been successfully employed in the quantification of cell-surface glucose transporters, and in characterizing changes in their number under various conditions. Here we describe procedures for photolabelling different cell types using tritiated ATB-BMPA. When studying glucose transporters from rat adipocytes the cells are isolated according to Taylor and Holman (22). Mouse 3T3-L1 adipocytes are cultured in 35 mm2 plastic Petri dishes as reported previously by Calderhead et al. (23). Rat soleus muscles are isolated as described by Lund et al. (24) and Dudek et al. (25), and adult rat cardiomyocytes are prepared according to Fischer et al. (26).

3 Detection of the covalent incorporation of photolabels into glucose transporter isoforms Several approaches can be employed to analyse the extent of UV irradiationinduced crosslinking of the photolabel into the glucose transporter. These include: (a) Direct analysis by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and the construction of a photoincorporation profile by gel slicing, solubilization, and liquid scintillation counting. These studies can provide information 197

ALISON K. GILLINGHAM ET AL.

Protocol 2 Photolabelling glucose transporters using [3H]ATB-BMPA Equipment and reagents • Rayonet photochemical reactor (RPR-100) with 300 and 350 nm lamps • [3H]ATB-BMPA (specific radioactivity 370 MBq/mmol) (as described by Clark and Holman(20)) • Incubation buffers (chosen according to cell type): (a) 1% BSA/Hepes buffer for rat adipocytes: 140 mM NaCl, 4.7 mM KCI, 125 mM MgSO4. 2.5 mM CaClj, 2.5 mM NaH2PO4, 10 mM Hepes pH 7.4.1% (w/v) bovine serum albumin (BSA) (b) Krebs-Ringer-Hepes buffer for 3T3-L1 adipocytes: 136 mM NaCl, 4,7 mM KCl, 1.25 mM CaCl2,1.25 mM MgSO,,, 10 mM Hepes pH 7.4 (c) PBS for human eiythrocytes—see Protocol 1 (d) Haemolysis buffer: 5 mM NaH2PO4 pH 7.8,1 mM EDTA supplemented with 20 ug/ml 4-(2-aminoethyl)benzenesulfonylfluoride (AEBSF)

(e) Krebs-Henseleit bicarbonate buffer for isolated rat soleus muscle: containing 2 mM pyruvate. 38 mM manm'tol, 0.1% (w/v) BSA (f) Krebs-Ringer-Hepes buffer for isolated adult rat cardiomyocytes: 128 mM NaCl, 6 mM KCI, 1 mM CaCl2, 1 mM Na2HP04, 0.2 mM NaH2P04, 1.4 mM MgSO4,10 mM Hepes pH 7.4, with 2% (w/v) fatty acid-free BSA Serum-free Dulbecco's modified Eagle's medium (DMEM) Monocomponent porcine insulin (Novo Nordisk) 35 mm2 plastic Petri dishes (Falcon, Nunc, or Greiner) Water bath at 18°C (this can be a simple sandwich box and thermometer, kept at 18 °C by the addition of ice) Centrifuge 1.5 ml assay tubes Liquid nitrogen

Method NB: As the photolabelling conditions vary between cell types each photolabelling procedure is described separately in this section.

A. Photolabelling isolated rat adipocytes: 1. Prepare rat adipocytes and adjust the cell suspension to a cytocrit of 40%. Incubate the adipocytes in 1% BSA/Hepes buffer for 20 mirt at 37°C in the absence or presence of 20 nM insulin.

2. Transfer 1 nil aliquots of the cells to 35 mm2 Petri dishes and incubate for 5 min at 1S°C in a water bath to slow transporter recycling. 3. Add 9.25 MBq of [3H]ATB-BMPA to the cells (to yield a final concentration of 40 u M ) ,

swirl to mix and irradiate for 1 min in the photoreactor using 300 nm lamps. 4. Then either wash the cells once with fresh incubation medium to remove excess

photolabel as in Protocol 1 and incubate at 37°C for studying glucose transporter recycling, or wash four times with 1% BSA/Hepes buffer at 18°C and use for immunoprecipitation with relevant antibodies, or for subffactionation studies.

198

DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS Protocol 2 continued

B. Photolabelling 3T3-L1 adlpocytes: 1. Incubate the differentiated 3T3-L1 adipocytes for 2 hours in serum-free Dulbecco's modified Eagle's medium (DMEM) in 35 mm2 Petri dishes. 2- Incubate for 30 min in the absence or presence of 100 nM insulin in KrebsRinger-Hepes buffer at 37"C. 3. Wash the cells with Krebs-Ringer-Hepes buffer at 18°C to slow the glucose transporter recycling, by adding 5 ml buffer, swirling gently and then removing. 4. Irradiate the cells for 1 min in the presence of 3.7 MBq of [3H]ATB-BMPA in a final volume of 250 u1 of Krebs-Ringer-Hepes buffer (the final concentration of the photolabei is 40 uM). 5. Wash the cells four times with Krebs-Ringer-Hepes buffer as in step B3 to remove excess label. 6. Analyse (see Protocols 3-6). C. Photolabelling intact human erythrocytesa: 1. Wash intact human erythrocytes obtained from 1- to 3-week-old blood (e.g. outdated blood from a blood bank) in ice-cold PBS five times by centrifugation at 2300 g for 10 min. 2. Bring the erythrocyte suspension to a 20% cytocrit with PBS. 3. Transfer 200 ul of the 20% erythrocyte suspension to a 35 mm2 Petri dish. 4. Add 370 KBq of [3H]ATB-BMPA in 200 ul of PBS (final concentration of the photolabei is 2.5 uM, and the erythrocytes are now at 10%). 5. Irradiate for 1 min in the photoreactor with 300 nm lamps. 6. Transfer the erythrocytes to a 1.5 ml assay tube and wash five times with ice-cold PBS as in Protocol 1 to remove excess photolabel. 7. Lyse the cells in haemolysis buffer for 20 min on ice. 8. Recover the membranes by centrifugation at 20 000 g for 20 min at 4°C. 9. Wash the pellet once in haemolysis buffer, re-centrifuging for at 4°C for 20 min at 20000 g (as in step 8). The membranes are now ready for further analysis (see Protocols 3-6). D. Photolabelling Intact soleus muscle: 1. Incubate the isolated soleus muscle in Krebs-Henseleit bicarbonate buffer at 30 °C in the absence or presence of 20 nM insulin and continuously gas with 95% 02/5%CO2. 2. At the end of the incubation transfer the muscles to a dark room and incubate for 8 min at 18°C in Krebs-He nseleit bicarbonate buffer containing 37 MBq/ml [3H]ATBBMPA (final concentration of the label is 100 uM). 3. Irradiate twice for 3 min each time in the photoreactor, turning the muscle tissue over manually after 3 min. 199

ALISON K. GILLINGHAM ET AL Protocol 2 continued

4. Immediately following irradiation blot the tissues with filter paper, trim, and then freeze them in liquid nitrogen,

£. Photolabelling isolated adult cartilomyocytes: 1. Incubate the isolated cardiomyocytes in Krebs-Ringer-Hepes buffer in the absence or presence of 30 nM insulin for 30 min at 37°C in a gently shaking water bath. Continually gas the cells with 100% 02,c 2. Allow the cardiomyocytes to settle to form a loose pellet and remove the supernatant buffer, leaving approximately 500 (ul of cell suspension (2 x 105-5 x 105 cells). 3. Transfer the cells to a 35 mm2 Petri dish and incubate for 5 min at 18°C to stop transporter recycling. 4. Add 18.5 MBq of [3H]ATB-BMPA (final concentration of the label is 100 nM) and irradiate for 1 min in the photoreactor with 350 nm lamps. 5. Transfer the cells to 1.5 ml assay tubes and wash four times at room temperature with 1 ml volumes of Krebs-Ringer-Hepes buffer or PBS buffer, briefly centrifuging to pellet the cells then discarding the supernatants, 6. Analyse (see Protocols 3-6). "The same protocol can be applied to photolabelling erythrocyte membranes. In this case irradiate 300 ^g of erythrocyte ghosts, obtained as described by Baldwin and Lienhard (13). for 1 min with 370 KBq of [3H]ATB-BMPA in a final volume of 300 ul of 5 mM sodium phosphate buffer pH 7.2. Then wash the membranes and recover them by centrifugation in 5 mM sodium phosphate buffer pH 7.2, at 20000 g for 20 min at 4°C. b D-glucose or other sugars may be used as inhibitors at a final concentration of 500 mM, and cytochalasin B at a final concentration of 10 uM. "The shaking water bath should be very slow to prevent increased basal glucose uptake caused by mechanical agitation.

concerning the specificity of the labelling reaction. For example, using this method it has been demonstrated that, in addition to the glucose transporter GLUT4, ATB-BMPA also covalently binds to a protein of approximately 97 kDa in rat adipose cells. Competition studies show that ATB-BMPA is not displaced from this protein by n-glucose or cytochalasin B, and the protein is therefore assumed to be unrelated to the glucose transporter family (21). (b) Photolabelling in combination with isoform-specific immunopivcipitiition to analyse more accurately the phorolabelling of different transporters in various cell types and to avoid interference by non-specific labelling (see Protocols 3 and 4), These studies can provide information about the levels of labelled transporters at the surface compared with the total amount of transporter present in the cell. 200

DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS

Protocol 3 Immunoprecipitatlon of the photolabelled glucose transporter isoforms Equipment and reagents • PBS (seeProtocol!) • Solubilization buffer: 2% (w/v) Thesit (CUE9) in PBS supplemented with the proteinase inhibitors antipain, aprotinin. pepstatin, and leupeptin, each at a final concentration of 1 mg/ml • Wash buffer A: 1% (w/v) Thesit in PBS • Wash buffer B: 0.1% (w/v) Thesit in PBS • Protein A immobilized on Sepharose CL4B (Sigma)

• Refrigerated centrifuge • Rabbit anti-GLUT4 C-terminal peptide antibody (whole serum) • Rabbit anti-GLUTl C-terminal peptide antibody (whole serum) » 10% SDS-PAGE equipment and reagents • Electrophoresis sample buffer: 62,5 mM Tris-HCl pH 6.7, 2% SDS, 50% glycerol, 0.02% Bromophenol Blue, supplemented with 10% (3-mercaptoethanol

Method 1. Make a 50% slurry of protein A-Sepharose beads: (a) In a 1,5 ml assay tube weigh out the desired amount of protein A-Sepharose beads (e.g. for 10 ul slurry use 2.5 mg of protein A-Sepharose powder). (b) Add 1 ml of PBS and leave on ice for 10 min to allow the protein A-Sepharose beads to swell. (c) Centrifuge the slurry at 6000 g in a microcentrifuge for 1 min. (d) Discard the supernatant and resuspend the protein A-Sepharose beads in 1 ml of PBS. (e) Repeat steps (c) and (d) twice, finally resuspending the protein A-Sepharose beads in an equal volume of PBS to obtain a 50% slurry. 2. Conjugate the anti-GLUT4 or anti-GLUTl antibodies to the protein A-Sepharose slurry obtained in step 1: (a) To 40 ul of the protein A-Sepharose slurry add 50ulA of anti-GLUT4 or antiGLUTl antibodies. (b) Add 1 ml of PBS and incubate for 2 hours at 4°C with end-to-end rotation to couple the antibodies to the protein A-Sepharose beads." (c) At the end of the incubation centrifuge the tubes at 6000 g for 1 min to pellet the protein A-Sepharose beads and carefully discard the supernatant. (d) Wash the pellets three times with 1 ml of ice-cold PBS to remove the remaining unconjugated antibodies, (e) Keep the protein A-Sepharose-antibody conjugate in PBS at 4 °C. 3. Solubilize the photolabelled material (cells or membranes) by adding 1 ml of the solubilization buffer to the samples obtained in Protocols I and 2,

201

ALISON K. GILLINGHAM ET AL Protocol 3 continued

4. Incubate for 20 min at room temperature (for adipose cells) or 4°C (for erythrocytes, 3T3-Lt adipocytes, muscle samples (homogenized first), and cardiomyocytes}. 5. Centrifuge the samples at 20 000 g for 20 min at 4 °C. 6. Centrifuge the protein A-Sepharose-anti-GLUT4 antibody conjugate obtained in step 2 (e) at 6000 g for 1 min and carefully discard the supernatant. 7. Transfer the detergent-solubilized supernatant from step 5 to the beads and incubate for 2 hours at 4 °C with end-to-end rotation." 8. At the end of the incubation centrifuge the samples at 6000 g for 1 min. 9. For sequential imiminoprecipitation, transfer the supernatant obtained in step 8 to protein A-Sepharose beads conjugated with antibodies against another glucose transporter isoform, e.g. GLUT1, and again incubate for 2 hours at 4°C with endto-end rotation. If sequential immunoprecipitation is not required proceed to step 11. 10. At the end of the second incubation centrifuge the samples at 6000 g for 1 min and remove the supernatant.11 11. Wash the protein A-Sepharose pellets: (a) with 1 ml of Wash buffer A and centrifuge at 6000 g for 1 min. Repeat four times. (b) with 1 ml of Wash buffer B and centrifuge at 6000 g for 1 min. Repeat four times. (c) once with PBS and centrifuge at 6000 g for 1 min. 12. Add 100 ul of the sample buffer supplemented with 10% |J-mercaptoethanol to the washed pellet and incubate for 20 min at room temperature. 13. Mix vigorously to remove the glucose transporter-antibody conjugate from the protein A-Sepharose beads. 14. Centrifuge at 6000 g for 1 min to pellet the protein A-Sepharose beads and load the supernatant on to a 10% SDS-PAGE gel. " This incubation can be also performed overnight. "The supernatant obtained after incubating the samples with the anti-GlUTl antibodyprotein A-Sepharose conjugate (step 10} can be kept to further analyse it for the remaining amount of photo labelled material.

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DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS

Protocol 4 Solubilization of gels crosslinked with bis-acrylamide Equipment and reagents • Coomassie Blue stain reagent: 0.01% (w/v) Coomassie Brilliant Blue R-250.10% (v/v) glacial acetic acid, 30% (v/v) methanol, 60% (v/v) H20 • Destain reagent; 10% (v/v) glacial acetic acid, 30% (v/v) methanol, 60% (v/v) H20 • Gelslicer—this consists of a series of razor blades positioned between metal blocks embedded in Perspex. The number of razor blades required depends upon the number and thickness of the gel slices desired.

• Oven set to 80°C

• Solubilization reagent: aqueous 2% (v/v) ammonia solution in 30% (v/v) H2O2. Make this reagent fresh just before use and keep in ice during the addition to each vial. • Molecular weight standards • Polyethylene liquid scintillation vials • Scintillation fluid (OptiPhase 'Safe'; Walkc) • Scintillation p-counter

Method 1. Fix proteins by briefly staining the gel in Coomassie Blue stain, and then destain with the destaining reagent until the protein bands on the gel are clearly seen. 2. Wash briefly with distilled H20. 3. Cut the gel into separate lanes according to the loading wells. Do the same for the lane containing the molecular weight standards." 4. Cut each lane into 6.6-mm wide slices and place them in scintillation vials.a,b 5. Dry the gel slices at 80°C for 2 h in open vials, then allow them to cool to room temperature. 6. Add 500 ul of the Solubilization reagent to each vial and immediately close the vials. 7. Allow the slices to solubilize by incubating them with the Solubilization reagent for 1-2 hat 80°C. 8. When the slices are completely solubilized remove them from the oven and allow to cool to room temperature.' 9. Add 8 ml of scintillation fluid and count the radioactivity in each gel slice using a liquid scintillation counter, "Regularly wet the gel with distilled water, otherwise it wilt become sticky and difficult to cut. '"The lane with the molecular weight standard is also cut and the number of the slice containing each of the proteins of known molecular weight is recorded, so that it can be matched with the radioactive profile detected in the sample lanes. c It is necessary to cool vials before counting to eliminate occasional chemiluminescence.

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ALISON K. GILLINGHAM

4 Biotinylated photolabels One of the disadvantages of the original bis-mannose derivatives was their dependence on radioactivity as the mode of detection. A problem associated with this is that despite the efficiency of the label to crosslink to the transporter (approaching 100%), only a small fraction of the total label added to a solution occupies the binding site (this is because of the relatively low affinity of the hexose analogues for the binding site). Therefore, large quantities of tritiated compound have to be added to solutions and extensive washings of cells and membranes have to be performed to reduce background radioactivity. Thus, new cell-impermeant bis-mannose photolabels have now been developed with biotinyl groups attached to ATB-BMPA (see Figure 2) by either a polyethoxy spacer (Bio-ATB-BMPA) or an additional hexanoic acid spacer (Bio-LC-ATB-BMPA). The JQ values for inhibition of glucose transport activity in insulin-stimulated rat adipocytes using these photolabels are very similar to those previously reported for ATB-BMPA, being 359 ± 10 and 273 ± 28 uM for Bio-ATB-BMPA and Bio-LCATB-BMPA, respectively (27). Photolabelling with the biotinylated derivatives can be performed exactly as described in Protocol 2, such that the final concentration of the label is 500 n-M. Following UV irradiation-induced crosslinking, the biotinylated glucose transporter can be detected utilizing the interaction of the biotin moiety with streptavidin or avidin molecules. For biotinylated transporters, streptavidin precipitation can be performed followed by isoform-specific Western blotting (see Protocol 5). Alternatively, samples may be subjected to isoform-specific immunoadsorption followed by Western blot detection of biotinylated transporters using suitable conjugates such as streptavidin-horseradish peroxidase

Figure 2 Structure of ATB-BMPA and the biotinylated derivative Bio-LC-ATB-BMPA.

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DETECTION AND ANALYSING OF GLUCOSE TRANSPORTERS

(streptavidin-HRP) or Extra-Avidin-HRP. Particularly sensitive detection can be achieved using the Amdex™ high-performance streptavidin-HRP conjugate, where multiple enzyme molecules are linked to streptavidin via a Dextran backbone (see Protocol 6).

Protocol 5 Detection of blotlnylated GLUT4 using streptavidin precipitation Equipment and reagents • Streptavidin-agarose beads (Pierce)

• Solubilization buffer: 2% Thesit in PBS

• PBS (see Protocol 1) • l% BSA/Hepesbuffer(see Protocol 12)

• Wash buffer A: 1% Thesit in PBS

• 0.1% BSA/Hepes buffer: 140 mM NaCl. 4.7 mM KC1.1.25 mM MgSO4, 2.5 mM CaCl2, 2.5 mM NaH2PO4.10 mM Hepes pH 7.4, 0.1%(w/v)BSA

• Wash buffer B: 0.1% Thesit in PBS • Sample buffer: 62.5 mMTris-HClpH 6.7, 2% SDS, 50% glycerol, 0.02% Bromophenol Blue

Method 1. Following photolabelling, wash the cells (250-500 ul) four times in a large volume of 1% BSA/Hepes buffer and once in 0.1% BSA/Hepes buffer. 2. Add 1 ml of the solubilization buffer, vortex briefly, and incubate for 20 min at room temperature (for adipocytes) or at 4°C (for 3T3-L1 adipocytes, cardiomyocytes, erythrocytes. and muscle), 3. Centrifuge at 20 000 g for 20 min at 4 °C and retain the supernatant containing the solubilized membranes (1-3 mg/ml) (see Protocol 3). 4. Wash 50 ul of a 50% slurry of streptavidin-agarose beads twice with 1 ml PBS and combine with the total cell lysate (generated in step 3). 5. Incubate overnight at 4°C with end-to-end rotation. 6. Following incubation, wash the pellet four times with Wash buffer A, four times with Wash buffer B, and once with PBS. 7. Elute bound protein with 30 ul of the sample buffer by heating at 95°C for 20 min." Centrifuge at 6000 g for 1 min at room temperature and collect the eluate. Repeat this step and pool the eluates. 8. Analyse the sample by SDS-PAGE and Western blotting against GLUT4,b ° To avoid transporter aggregation and loss on heating, the choice of the type of tube used appears to be important. We routinely use Sarstedt microcentrifuge tubes for this treatment. ''This method can also be employed for the analysis of other GLUT isoforms, providing the relevant antibodies are available.

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Protocol 6 Detection of biotinylated GLUT4 by Immunoprecipitatlon and detection with Amdex™ streptavldin-HRP Equipment and reagents • PBS (see Protocol 1) • 1% BSA/Hepes buffer (see Protocol 2) • 0.1% BSA/Hepes buffer (see Protocol 5) • Sohibilization buffer: 2% Thesit in PBS • Wash buffer A: 1% Thesit in PBS • Wash buffer B: 0.1% Thesit in PBS • TBS-T (Tris-buffered saline-Tween): 0.09% NaCl, 1 M Tris-HCl pH 7.4, supplemented with0.1%Tween20

• Sample buffer: 62.5 mM Tris-HCl pH 6.7, 2% SDS, 50% glycerol, 0,02% Bromophenol Blue • SDS-PAGE equipment and reagents • Amdex™ streptavidin-HRP (Amersham Pharmacia Biotech) • ECL™ Western blotting equipment and reagents (Amersham Pharmacia Biotech) • Nitrocellulose membrane, 0.45 ^m poresize (Gelrnan Sciences) • 5% (w/v) non-fat, dried milk powder

Method 1. Following photolabelling, wash cells {250-500 (il) four times in a large volume of 1% BSA/Hepes buffer and once in 0.1% BSA/Hepes buffer. 2. Add 1 ml of the solubilization buffer, voitex briefly, and incubate for 20 min at room temperature. 3. Centrifuge at 20000 g for 20 min at 4°C and retain the supernatant containing the solubilized membranes (1-3 mg/ml), 4. Immunoprecipitate GLUT4 as described in Protocol 3. 5. Elute bound protein from the immunoprecipitates with the sample buffer by heating at 95°C for 20 min. Centrifuge at 6000 g for 1 min and collect the eluate. 6. Separate the eluted proteins by SDS-PAGE (10% acrylamide) and transfer to nitrocellulose, 7. Block the nitrocellulose in 5% (w/v) non-fat dried-milk powder in TBS-T buffer for 30-60 min at room temperature. 8. Rinse the nitrocellulose thoroughly in TBS-T buffer and incubate for 90 min at room temperature with a 1:4000 dilution of Amdex™ streptavidin-HRP in 1% BSA/TBS-T buffer (do not use milk powder as this will interfere with the signal thus resulting in high background). 9. Wash the membrane extensively in TBS-T buffer and detect bound streptavidinHRP by enhanced chemiluminescence (ECL™) according to the manufacturer's instructions.

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References 1. Baley, H. and Knowles, J. R. (1977). In Methods in enzymology, Vol. 46 (ed. W. B. Jakoby and M. Wilchek), p. 69. Academic Press, London. 2. Taverna, R. D. and Langdon, R. G. (1973). Biochim. Biophys. Acta, 323, 207. 3. Holman, G. D., Pierce, E. J., and Rees, W. D. (1981). Biochim. Biophys. Acta, 646, 382. 4. Wadzinski, B. E., Shanahan, M. F., Seamon, K. B., and Ruoho, A. E. (1990). Biochem.]., 272, 151. 5. Shanahan, M. F. (1982). J. Biol. Chem., 257, 7290. 6. Holman, G. D., Parkar, B. A., and Midgley, P. J. W. (1986). Biochim. Biophys. Acta, 855, 115. 7. Clark, A. E., Holman, G. D., and Kozka, I. J. (1991. Biochem. L, 278, 235. 8. Yang, J., Clark, A. E., Kozka, I. J., Cushman, S. W., and Holman, G. D. (1992). J. Biol. Chem., 267, 10393. 9. Yoshida, E., Nakayama, H., Hatanaka, Y., and Kanaoka, Y. (1990). Chem. Pharm. Bull, 38, 982. 10. Hashimoto, M., Kanaoka, Y., and Hatanaka, Y. (1997). Heterocycks, 46, 119. 11. Brunner, J. (1996). Trends Cell Biol., 6, 154. 12. Oka, Y. and Czech, M. P. (1984). J. Biol. Chem., 259, 8125. 13. Baldwin, S. A. and Lienhard, G. E. (1989). In Methods in enzymology. Vol. 174 (ed. S. Fleischer and B. Fleischer), p. 39. Academic Press, London. 14. Midgley, P. J. W., Parkar, B. A., and Holman, G. D. (1985). Biochim. Biophys. Acta, 812, 33. 15. Holman, G. D. and Midgley, P. J. W. (1985). Carbohydrate Res., 135, 337. 16. Carter-Su, C, Pessin, J. E., Mora, R., Gitomer, W., and Czech, M. P. (1982) J. Biol. Chem., 257, 5419. 17. Wadzinski, B. E., Shanahan, M. F., and Ruoho, A. E. (1987). J. Biol. Chem., 262, 17683. 18. Holman, G. D., Parkar, B. A., and Midgley, P. J. W. (1986). Biochim. Biophys. Acta, 855, 115. 19. Holman, G. D., Karim, A. R., and Karim, B. (1988). Biochim. Biophys. Acta, 946, 75. 20. Clark, A. E. and Holman, G. D. (1990). Biochem. J., 269, 615. 21. Holman, G. D., Kozka, I. J., Clark, A. E., Flower, C. J., Saltis, J., Habberfield, A. D., Simpson, I. A., and Cushman, S. W. (1990). J. Biol. Chem., 265, 18172. 22. Taylor, L. P. and Holman, G. D. (1981). Biochim. Biophys. Acta, 642, 325. 23. Calderhead, D. M., Kitagawa, K., Tanner, L. L, Holman, G. D., and Lienhard, G. E. (1990). J. Biol. Chem., 265, 13800. 24. Lund, S., Holman, G. D., Schmitz, 0., and Pedersen, O. (1993). FEES Lett., 330, 312. 25. Dudek, R. W., Dhom, G. L., Holman, G. D., Cushman, S. W., and Wilson, C. M. (1994). FEES Lett., 339, 205. 26. Fischer, Y., Rose, H., and Kammermeier, H. (1991). Life Sci., 49, 1679. 27. Koumanov, F., Yang, J., Jones, A. E., Hatanaka, Y., and Holman, G. D. (1998). Biochem.]., 330, 1209.

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Chapter 9 Computer prediction of transporter topology and structure RONG-I HONG and MARK S. P. SANSOM Laboratory of Molecular Biophysics, University of Oxford, Rex Richards Building, South Parks Road, Oxford 0X1 3QU

1 Introduction One of the resources which may be exploited in studying membrane transport proteins is the enormous (and increasing) body of genome sequence data. At the time of writing, complete genomic sequences are known for 21 prokaryotes (1), a yeast (2), and for Caenarhabditis elegans (3). Current estimates suggest that integral membrane proteins comprise about 20-30% of most genomes (4, 5). This provides a strong motivation to exploit this information in terms of trying to understand the relationship between membrane protein sequence and structure (and eventually function). Prediction of protein structure is fraught with difficulties and despite intense efforts, ab initio methods for structure prediction remain elusive (6). However, for membrane proteins (or at least those whose membrane domains are composed largely of a-helices), the prospects of prediction are somewhat brighter as the lipid bilayer restricts the number of degrees of freedom of the system. Prediction of transmembrane (TM) a-helices is widespread and a number of different methods and hydrophobicity scales have been devised to aid such predictions (7). Other methods allow the prediction of preferred orientations of such TM helices, thus placing restrictions on how they may be packed together in the intact protein. More recently, simulation-based methods have been developed that allow the structural dynamics of predicted TM helices to be examined in a lipid bilayer environment. Together, although such methods do not yet allow the unambiguous prediction of the structure of a transport protein, they do provide an important tool in studying membrane proteins, especially if only limited or low-resolution structural data are available. The focus of this chapter is on those membrane proteins (the majority) which are made up of bundles of a-helices. For prediction methods for transbilayer p-barrels see ref. 8. In this chapter we review some of the methods currently available for 209

RONG-I HONG AND MARK S. P. SANSOM

Figure 1 Domain structure and proposed transmembrarie topology of ABC transporters, with two transmembrane domains (each containing six TM helieesj and two nudcotide binding domains (NBD) on the intracellular face of the membrane.

computer analysis of membrane protein sequences. We illustrate this via their application to ATP binding-cassette (ABC), transporter protein sequences from the Btinllus subttlis genome (9). The ARC transporters make up one of the larger superfamilies of membrane transport proteins, and are present in both prokaryotes (10) and eukaryotes. They are responsible for the transport of a wide range of solutes. Of particular biomedical importance are their roles in antibiotic resistance in microorganisms and in the resistance of tumour colls to chemotherapeutic agents ( 1 1 ) . They share a common transmembrane topology (see Figurf J) made up of 12 TM helices and two nudcotide (ATP) binding domains. The crystal structure of an isolated nucleotidc binding domain (NBD) from a bacterial ABC transporter lias been described (12). These domains may all be part of a single polypeptidc chain (as in, for example, the cystic fibrosis transport-regulator protein, CFTR) or the two groups of six TM helices may occur on distinct polypeplide chains (as is the case in many bacterial transporters) (13). In those cases where the 12 helices arc on a single polypeptide chain there is evidence for internal symmetry, with TM7 resembling TM1, etc.

2 Database searching and sequence alignment Before we can proceed with structure prediction, we need to obtain sequences of the transport proteins in which we are interested from the various databases, to compare and align those sequences, and to examine possible evolutionary relationships by cluster analysis. One may already have a database of sequences at hand. Alternatively, one may extract sequences of interest from a number of different databases. If the latter is the case, keyword searching is probably the simplest way to identify such sequences. For example, if one is interested in finding ABC transporter proteins in different organisms, one could submit the string 'ABC transporter' to the search engine of a genome database to extract sequences of known ABC transporters from the given genome. Howrever, there may be some ambiguities or even mistakes in the sequence annotations in different databases. Thus, a more systematic search requires the use of sequence-based searching. This will identify related sequences that may not be picked up by keyword searching. We will therefore start by describing search methods. 210

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2.1 Search algorithms The most reliable way of finding proteins that are related to a 'target' protein is to search for those proteins that exhibit sequence similarity with the target. Several different algorithms for sequence database searching have been developed to achieve this goal. Among them, BLAST (Basic Local Alignment Search Tool) (14) and FASTA (15) are probably the most widely used programs for exploring all the available sequence databases. These two algorithms were developed on the basis of different concepts. However, they were both designed with the assumption that the best overall sequence similarities can be detected by local alignments (16). Both BLAST and FASTA have been constantly improved and refined since their first releases. Both programs are widely used in different sequence databases, and are available at public domains (see Table 1). Some new capabilities have been implemented into the latest version of BLAST (17). The newer Gapped BLAST allows a sequence database search with gaps, i.e. deletions and insertions, which is unavailable in the original BLAST. Furthermore, the new PSI-BLAST iteratively searches sequence databases using a position-specific score matrix. This gives higher sensitivity, enabling one to detect weak, but biologically relevant, sequence similarities that may be missed by a conventional pairwise comparison. For example, Beamer et al. (18) successfully used PSI-BLAST to detect distant relatives of mammalian LPS-binding and lipid transporter proteins. However, it is important to have a statistical measure of the significance of a weak similarity. Statistical measures of sequence similarity, like the £ value of BLAST2, provide good indicators of distantly homologous sequences. The newer version of the FASTA package has been enhanced to provide statistical estimates for length-corrected sequence similarity scores, which is reported to be more effective than raw scores at identifying distantly related homologues (19). Table 1 Software and databases for sequence-similarity searches Software BLAST FASTA

http://www.ncbi.nlm.nih.gov/BLAST/ http://alphalO.bioch.virginia.edu/ http://www2.ebi.ac.uk/fasta3/

Nucleic acid sequence databases DDBJ

http://www.ddbj.nig.ac.jp/

EBI

http://www.ebi.ac.uk/

GenBank

http://www.ncbi.nlm.nih.gov/Genbank/index.html

Protein sequence databases MIPS OWL PIR SWISS-PROT

http://www.mips.biochem.mpg.de/ http://www.biochem.ucl.ac.uk/bsm/dbbrowser/OWL/OWL.html http://www-nbrf.georgetown.edu/pir/ http://expasy.hcuge.ch/sprot/

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2.2 Scoring matrices In addition to comparison algorithms, scoring matrices play an important role in determining the sensitivity of pairwise comparisons of two protein sequences. Two major series of scoring matrix are widely used: PAM (Point Acceptable Mutations per unit time); and BLOSUM (Best Local SUMmed alignment percentages). The PAM matrix series were developed from observed amino acid substitution frequencies by Dayhoff and co-workers (20). Thus, the higher number denoted for each matrix, e.g. PAM250, implies a higher degree of substitution. The BLOSUM matrix was developed by Henikoff and Henikoff (21) based on analyses of conserved regions in aligned, protein sequence families. Therefore, a high denoted BLOSUM number, e.g. BLOSUM62, implies a high degree of conservation. Vogt et al. (22) analysed alignment accuracies as a function of matrix. In their study, PAM, BLOSUM, and other matrices were used to align protein sequences which were also matched by superposition of known three-dimensional structures. The comparisons showed relatively similar results for the PAM and BLOSUM series. This result suggests that no single matrix is optimal for all protein sequence comparisons. The default scoring matrices are BLOSUM62 (in BLAST) and BLOSUM50 (in FASTA).

2.3 Sequence databases Using these similarity searching tools, a number of sequence databases can be explored. Major nucleic acid sequence databases are collected and maintained at GenBank (NCBI) in the USA (23), EMBL Nucleotide Sequence Database (EBI) in Europe (24), and the DNA Data Bank of Japan (DDBJ) in Japan (25) (see Table 1). There are also two major protein sequence databases: SWISS-PROT (26); and PIR (Protein Information Resource) (27). SWISS-PROT, and its supplement TrEMBL which contains all the translations of EMBL nucleotide sequence entries not yet integrated in SWISS-PROT, are designed to provide a high level of annotations for protein functions, domains, and other information. The PIR-International Protein Sequence Database is a new joint venture between the Munich Information Center for Protein Sequences (MIPS) (1) and the Japanese International Protein Sequence Database (JIPID) aimed at creating a comprehensive and wellverified database; it is also organized according to biological principles including structural, functional, and evolutionary relationships. In addition to these two major protein sequence databases, there is also a non-redundant composite of SWISS-PROT, PIR, GenBank (translation), and NRL-3D, called OWL, which is designed in accordance with a philosophy of providing a database which is 'small and hence efficient in similarity searches' (28) (see Table 1). As these databases provide up-to-date sequences and the most reliable sequence-searching programs it is inadvisable to maintain all local copies if only a relatively small number of membrane protein sequences are to be analysed. By using the World Wide Web, sequence searching can be easily achieved 212

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without having to worry about installing programs and maintaining databases. As these databases are always mirrored at a number of institutions, it is more efficient to use the nearest mirror site.

2.4 Multiple alignments Having generated a long list of hit sequences via similarity searching, these results should be carefully examined and interpreted using both statistical measures and one's biological knowledge. As mentioned above, the statistical measures provide a good way of assessing the reliability of alignments. Examination of the annotation of each extracted sequence provides a simple 'biological filter'. Having gathered together all the sequences of interest, patterns of amino acid conservation are detected by refining the initial sequence alignments, i.e. by running multiple sequence alignment (see Table 2). The amino acid conservation patterns in the multiply aligned sequences are useful for further sequence analysis, e.g. phylogenetic analysis, secondary structure prediction, and fold prediction. Aligning a large set of N protein sequences is, in principle, the extension of pairwise alignment to the alignment of N sequences simultaneously. In theory, this task can be achieved by the generalization of pairwise dynamic programming to N-dimensional dynamic programming. However, due to finite CPU time and computer memory, such a simultaneous approach limits the maximum number of sequences in a set and the number of residues in each sequence that can be compared. Lipman and co-workers (29) developed a program, MSA, which used a rather sophisticated approach to reduce the volume of the N-dimensional dynamic programming matrix. Even with this refinement, MSA is only suitable for up to eight protein sequences with average lengths of 200 to 300 residues. This limits the practical use of the truly simultaneous approach. An alternative approach is to align multiple sequences progressively. The first step is to construct a succession of optimal pairwise alignments by dynamic programming. The two most similar sequences are aligned first. Then, the next most similar sequence is added to the existing alignment. This process is iterated until all sequences have been aligned. This approach has been widely implemented in a number of multiple sequence alignment programs: PILEUP Table 2 Servers for multiple alignment of protein sequences AMPS CLUSTALW

http://barton.ebi.ac.uk/manuals/amps/amps.html http://www2.ebi.ac.uk/clustalw/ http://www.clustalw.genome.ad.jp/ http://helix.nih.gov/science/clustalw.html

HMMER

http://hmmer.wustl.edu/

MAXHOM

http://www.embl-heidelberg.de/predictprotein/ http://cubic.bioc.columbia.edu/predictprotein/

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RONG-I HONG AND MARK S. P. SANSOM (in the Wisconsin Package™ of Sequence Analysis software from the Genetics Computer Group, Inc. (GCG)); AMPS (30); MAXHOM (31); and CLUSTALW (32). In addition to dynamic programming algorithms, other mathematical methods have been used to solve this problem of multiple sequence alignment. Notredame and Higgins (33) developed a program, SAGA, based on a genetic algorithm. Hidden Markov Models have been implemented in the HMMER package for multiple sequence alignments (34). Sonnhammer et al. (35) extended the latter work to construct a database, Pfam, which contains multiple alignment and Hidden Markov Model-based profiles to detect remote homologous sequences. However, it seems that programs based on dynamic programming, in general, perform better than other methods.

3 Prediction of transmembrane helices (see Table 3) The prediction of TM helices is a crucial step in membrane protein modelling. As mentioned in the Introduction, predicting TM helices is generally believed to be easier than predicting secondary structures in globular proteins, since the lipid bilayer provides extra restraints upon the sequences of individual TM helices and on the topology of a membrane protein as a whole (36). Many attempts have been made to develop algorithms for locating putative TM helices, either from single sequences or from the multiple alignment of protein sequences. 3.1 Secondary structure and topology prediction A number of algorithms have been developed to predict the location of TM helices within a protein sequence. Early methods (37, 38) simply searched for runs of approximately 21 predominantly hydrophobic acids in a sequence, i.e. a sequence element of sufficient length to form a hydrophobic a-helix capable of spanning the approximately 30-A thick hydrophobic core of a lipid bilayer. This resulted in much discussion of the optimum hydrophobicity scale for use in such predictions (39). More recent work has analysed the sequences of known Table 3 Servers for the prediction of transmembrane helices DAS

http://www.biokemi.su.se/~server/DAS/

MEMSAT8

http://globin.bio.warwick.ac.uk/~jones/memsat.html

PHD

http://www.embl-heidelberg.de/predictprotein/predictprotein.html http://cubic.bioc.columbia.edu/predictprotein/

SOSUI

http://asuza.proteome.bio.tuat.ac.jp/sosui

TMAP

http://www.embl-heidelberg.de/tmap/tmapjnfo.html

TMHMM

http://www.cbs.dtu.dk/services/TMHMM-1.0/

TMPRED

http://www.ch. embnet.org/software/TMPRED_form. html

TOPPRED2

http://www.biokemi.su.se/~server/toppred2/

"The MEMSAT program cannot be run over the World Wide Web, but a PC version of the program is available by anonymous FTP, accessed via this site.

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TM helices in order to develop second-generation, pattern recognition algorithms for TM helix prediction. A number of such methods have been developed, all of which require a single query sequence as input. For example, von Heijne (40) observed the distribution of positively charged amino acid residues (the socalled 'positive inside rule') and analysed residue hydrophobicity along a protein sequence to locate putative helices from a single sequence. On the basis of data derived from well-studied transmembrane proteins, Jones and co-workers (41) used constrained dynamic programming in MEMSAT to predict both the location and orientation (in-out vs. out-in) of TM helices. Cserzo et al. (42) developed the program DAS by using a dense alignment surface method, which employed low-stringency dot-plots of the query sequence against a collection of non-homologous membrane proteins to locate putative helices. An amphiphilicity index and two sets of physicochemical indices were applied by Hirokawa (43) in designing SOSUI to detect putative helices. Sonnhammer and co-workers (44) applied a Hidden Markov Model to predict the location and orientation of putative TM helices. Their method, TMHMM, avoids using a fixed hydrophobicity cut-off. Thus it avoids a procedure which has the disadvantage of underestimating potential TM helices. In general, secondary structure prediction is more accurate if multiply aligned related sequences are used (45). This also seems to apply to the prediction of TM helices. TMAP (46) uses calculations of hydrophobicity propensity and terminal polar propensity for each aligned residue, allowing the prediction of TM helices from multiple sequence alignments. Rost and co-workers developed PHDhtm (47). This takes multiple alignments of protein sequences as input to a neural network, which is followed by dynamic programming refinement to locate putative TM helices. A list of TM helix prediction methods and associated websites is given in Table 3. The overall success rate of most TM helix prediction methods is approximately 80%. However, if the same protein sequence is analysed using a range of different algorithms, subtle differences in the resultant predictions are obtained, especially in terms of identifying the precise start and end residues of each TM helix. This possible source of error may be important in subsequent modelling and simulation studies. A comparison of some TM helix prediction programs has been made by Sonnhammer and co-workers (44). Their studies suggested that, in general, PHDhtm yielded better prediction results than other methods. However, it was noticed that PHDhtm tends to predict the TM helices to be shorter than they are shown to be when structural data are available. None the less, this program is reasonably good at identifying the location and orientation of TM helices, and it is relatively easy to use. But, to introduce a note of caution, examine Figure 2 in which the results of seven different prediction methods applied to bacteriorhodopsin are compared with one another and with the secondary structure assignment obtained by analysing the high-resolution X-ray structure of this protein. It is clear that no single method outperforms the others for all seven TM helices. Whilst not pretending that this is a systematic comparison of available methods, it does reinforce the message that TM helix 215

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Figure 2 Comparison of TM secondary structure/topology prediction methods applied to bactcnorhodopsin. The sequence of bacteriorhodopsin is given, aligned with the results of TM helix prediction by PHD, TMAP, TopPred2, DAS. TMHMM, SOSUI, and MEMSAT. The darkgrey cylinders (labelled DSSP) are the locations of the TM helices as determined from tne nigh-resolution, X-ray diffraction structure of bacteriorhodopsin (PDB ID code 1AP9; (59)).

predictions should be treated in the same critical fashion as the results of any other analysis, be they computational or experimental. It should be remembered that current prediction methods are based on a relatively small database of the experimental three-dimensional structure for membrane proteins. It is likely that improvements in prediction accuracy may occur as this database continues to expand.

3.2 Periodicity analysis Having defined the locations within the sequence of putative TM helices, and their orientations (in-out vs. out-in) relative to the bilayor. one may try to 216

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predict which residues of the helix face the surrounding lipid and which point in towards the centre of the helix bundle. A number of algorithms are available for this. They are derived from early work on the photosynthetic centre of Rhodobacter sphaeroides and related species (7), which revealed that membraneexposed resides are less highly conserved than interior residues (7, 48, 49). This work was extended by Donnelly and colleagues to G-protein coupled receptors (50). Based on these observations, Donnelly et al. (51) constructed tables of hydrophobicity and of the variability/conservation of amino acid residues along a TM helix. These parameters are used to estimate which residues are facing the lipid bilayer (and which the membrane interior) for predicted TM helices. This is done using the program PERSCAN, which detects a-helical periodicities (e.g. of residue conservation) in sequence alignments by a Fourier analysis method.

3.3 Model building To aid the interpretation of these analyses, it is useful to build molecular models of predicted TM a-helices. Such models can be used to identify possible interactions on a given surface (inward facing or outward facing) of a TM helix, and thus may aid in designing experiments to probe the location of TM helices within an intact membrane protein. The generation of model TM helices can either be achieved using, for example, standard helix-backbone geometry and preferred side-chain conformations (52-54) using a variety of molecular modelling packages (e.g. Quanta or Insightll from Biosym/MSI), or by using rather more computationally intensive procedures, based on restrained in vacua molecular dynamics simulations, as described in, for example, ref. 55. The latter methods are preferable if one intends to move on from single-helix to helix-bundle modelling; but if one simply wishes to build models of single TM helices, either approach will suffice. Display of model TM helices may be performed using one of a wide variety of molecular graphics programs. Rasmol has the advantage of being in the public domain (http://www.umass.edu/microbio/rasmol/), and is implemented on a variety of platforms from PCs to unix workstations. Static diagrams (for instance, for publication) may be generated by a variety of programs. Of these, Molscript (56) is quite widespread (although a bit tricky to learn how to use). A useful list of molecular graphics programs is maintained on the website: http://www.ebi. ac.uk/biocat/.

4 Example—B. subtills ABC transporters 4.1 Biological background As was discussed above, the ABC transporters form a large superfamily of membrane transport proteins, present in prokaryotes and eukaryotes. Their ubiquity and their biomedical importance make them particularly attractive for the methods of analysis discussed above. For example, the genome of B. subtilis has approximately 4.2 x 106 base pairs, and encodes 4100 proteins. Interestingly, 217

RONG-I HONG AND MARK S. P. SANSOM 77 members of the AKC-trans porter protein family were identified in this simple organism (9), The large number of ABC transporters suggests that B. subtilis has evolved a sophisticated nmgc of transporters to enable exchanges with a complex and dynamic environment. These transporters were therefore chosen to illustrate the methods available for the computer analysis of membrane protein sequence,

4.2 Sequence searching and alignment The whole genome of Bucillus subtilis was completed in 1997 (9). All the gene sequences nnd their translated open-reading frames (ORFs) are available to the public at the SubtiList web-sewer. http://www.pas[eur.fr/Hio/SubtiList/. At the time of writing, there were 4107 proteins in release R15.1 of this database. To illustrate how information on transporter structure can be obtained by computer predictions, the procedures summarized in Vmtncol I were used to

Protocol 1 Protein sequence analysis Method 1. Identify and then extract sequences potentially corresponding to the integral membrane protein components of a transporter family of interest from a database." 2. Use each extracted sequence to perform a sequence-similarity search against the SwissProt or TrEMBL databases, using e.g. FASTA or BLAST.b 3. Assess the results of the similarity searches by the E-vames obtained for each match, and/or using expert knowledge, to exclude spurious matches and produce sets of related sequences. 4. For each of the putative integral membrane components of the transporter protein family identified in step 1, construct a multiple sequence alignment.c 5. Analyse the multiple sequence alignments (and also individual sequences) for predicted TM helices, using the procedures described in Protocol 2. Exclude from further analysis any proteins predicted by these means to be non-transmembrane. By this means generate a subset of the putative transporter family members, identified in step 1, that are likely to be homologous integral membrane components of the family.11 6. Construct a multiple sequence alignment' of identified family members, to identify subfamilies and facilitate further analysis (see Protocols 2 and 3). " For example, by a text word search for 'ABC transporters' in the B. subtito 'SubtiList' database. b See Table 1 for sites on the World Wide Web where such similarity searchers can be performed. ( See Table 2 for sites on the World Wide Web where such alignments can be performed. d For example, the ABC-transporter family from B. subtilis.

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investigate the family of ABC transporters present in this organism. The transporter sequences were extracted from the SubtiList server by a keyword search for 'ABC transporters'. This yielded 144 protein sequences. It was possible that not all these were transmembrane proteins, and so computer analysis was used to assess the likelihood that they were integral membrane proteins. As we have seen, many TM helix prediction programs require multiple sequence alignments. Therefore, for each of the 144 sequences, homologous sequences were identified using Blast2 to search against the SwissProt 35 sequence database. The results of sequence similarity searching by Blast2 were further refined using MaxHom, with the E value cut-offset at 0.001. By these means multiple sequence alignments were generated for each of the 144 B. subtilis sequences. These alignments, or the individual sequences within each group where appropriate, were submitted to DAS, MEMSAT, TMPRED, and PHD and only those sequences predicted by all four methods to contain multiple TM helices were retained for further analysis (see below). This reduced the 144 sequences to 48. Many of the excluded sequences corresponded to soluble periplasmic binding proteins that are associated with ABC transporters. It is possible that these could have been excluded by a more refined keyword search, but it was decided that a relatively crude keyword search followed by TM helix prediction was less likely to result in 'false negatives'. Those ABC protein sequences which were predicted to contain membranespanning domains (see Section 4.3) were gathered together and further analysed to provide insights into their possible functional relationships. Sequence comparisons were carried out using CLUSTALW (see Table 2). A phylogenetic tree was constructed on the basis of the sequence comparison results—three clusters were identified (see Figure 3), in addition to 17 proteins which did not fit within any obvious cluster. In Cluster 1, most of the proteins are oligopeptide transporters. In Cluster 2, CydC and CydD are the only two proteins whose functions have been identified (they are required for the expression of cytochrome bd). However, the nucleotide-binding domain is present in all the proteins in Cluster 2. The functions of most of the proteins in Cluster 3 remain to be identified. However, four proteins in Cluster 3 of known function are transporters for basic solutes (OpuBB and OpuBD are choline ABC transporters, while OpuCB and OpuCD are glycine betaine/carnitine/choline ABC transporters). Thus, some possible correlation between sequence-generated clusters and transport function appear to exist. An alternative approach to exploring such correlations, which has been employed by, for instance, Saurin and Dassa (57), is to focus on those ABC transporters for which periplasmic binding proteins have been identified.

4.3 TM helix prediction As described in the previous section, analysis of protein sequences for transmembrane helices can be performed to identify the genuine integral membrane protein members of a transporter family. A number of algorithms are available 219

RONG-I HONG AND MARK S. P. SANSOM

Figure 3 Cluster analysis of B. subtilis ABC transporters. CLUSTALW was used to produce a phylogenetic tree for 47 ABC transporter sequences from the B. subtilis genome. Three pronounced clusters are seen. Subsequent TM helix prediction suggests that these clusters differ in their number of predicted TM helices.

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for performing such analyses, as detailed in Protocol 2. Once genuine family members have been identified in this way, comparison of the predicted topology of different family members can provide further insight into their relationships. As an example, the prediction results for Cluster 1 of the B. subtilis

Protocol 2 Prediction of transmembrane helices Method Prediction of transmembrane helices can be performed by analyses either of single protein sequences or of multiple alignments. The latter usually provide a more reliable prediction; however, if only a single sequence is available, it is wise to compare the results of several prediction algorithms.

A. From a single sequence 1. Prepare a text file containing the sequence in a format suitable for the programs listed in step 2." 2. Analyse the file using one of the following programs that are available on the World Wide Web:11 • DAS • MEMSAT • SOSUI

• TmHMM • TMPRED • TOPPRED2

B, From multiple sequences 1. Prepare a multiple sequence alignment file using one of the programs detailed in Table 2. 2. Analyse the alignment file using one of the following programs that are available on the World Wide Web:6 • PHDr • TMAP 0 Details of the file formats required are available on the Web servers used for TM helix predictions. Typically, analysis of single sequences requires the input of plain text or FASTA format files, which, if necessary, can readily be generated using a word processor; whereas multiple sequence alignments are performed on files in formats such as msf, which can be generated by the multiple sequence alignment programs detailed in Table 2. " See Table 3 for a list of servers available for predicting transmembrane helices. All programs can be used interactively on the World Wide Web except for MEMSAT, which must be downloaded and run locally (see Table 3). c PHD can use either a single sequence or a multiple sequence alignment as input.

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ABC-trnnsponer family are displayed in Figure 4. In this cluster, five out of the seven proteins are known to be transporters for oligopeprides. Interestingly, comparison of the locations of predicted helices within different members of the cluster suggests the existence of three subclusters (see figure 4).

Figure 4 TM helix predictions for the seven B. subtilis ABC transporters of Cluster 1 (see Figure 3). Predictions were performed using PHD. The extents of the TM helices are indicated by the grey boxes. These predictions suggest the existence of three subclusters, shown as dark, pale, anct mid-grey boxes.

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4.4 Periodicity analysis Once the trans membra no helix topology of members of a transporter family has been predicted, computer methods fan be applied to the analysis of the possible arrangements and orientations of individual helices within transmembrane helical bundles. For example, the periodic patterns of residue properties along predicted TM helices can be analysed using the PERSCAN package (see Protocol 3). Using multiple protein-sequence alignments as input, the PKRCON module detects the periodicity of residue conservation, whereas PHRHYD detects the periodicity of hydrophobicity (see ttgiira .5). In Cluster 1 of the B subtilis ABC-transporter family it is observed that DppC, OppC, and AppC have similar patterns of periodicity in both residue conservation and hydrophobicity for their TM helices, which indicates that the overall arrangement and orientation of the TM helices within the intact helix bundles of these three proteins are similar. Comparing the different TM segments, it is observed that TM1, TM2, TM3, and TM4 show sequence periodicities strongly supportive of an IYhelical conformation, whereas the periodicity results are less clear-cut for TM4 and TM6 (data not shown).

4.5 Models Putative trans membrane helices can be modelled following periodicity analysis. As an example, models of the six predicted TM helices from OppC were generated by restrained in vucuo molecular dynamics. This protein is a member of

Protocol 3 Residue periodicity analysis Method 1. Prepare a multiple sequence alignment file using one of the programs detailed in Table 2. 2. Analyse the periodicity of residue properties along the predicted transmembrane helices using one or more of the following members of the PERSCAN" suite of programs: • PERCON: quantifies the periodicity of residue conservation; • PERHYD: quantifies the periodicity of residue hydrophobicity; • PERMUT; quantifies the periodicity of lipid accessibility parameter; • PERVAR: quantifies The periodicity of residue variability. 3. From the analyses performed in step 2, identify the faces of putative transmembrane helices likely to be in contact with the lipid, and the faces likely to be in contact either with the transmembrane solute translocation channel or with adjacent helices (see Section 4.5) "This suite of programs can be obtained from Dr D. Donelly at [email protected]

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Figure 5 Periodicity analysis of helix TM3 from the three transporters corresponding to the first subcluster (dark-grey in Figure 4) of Cluster 1. (A) Sequence conservation; (B) hydrophobicity. In each case a power spectrum is shown. A peak at 100° corresponds to a sequence pattern that repeats once every 3.6 residues, i.e. with the periodicity expected for an a-helix.

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Cluster 1 of the B. subtilis ABC-transporter family detailed in Figurc 3. The structure of the TM3 model is shown in Figure 6. It can be seen that this has clear hydrophilic and hydrophobic surfaces, as identified by the periodicity analysis (Fitfurt; 5B). Of course, this model was generated in vacua. To get 3 more realistic

Figure 6 Model helix generated by restrained zin vacua molecular dynamics simulations. (A) Helical wheel for TM3 of OppC showing most conserved (unshaded) and most hydrophobic (shaded) faces in the corresponding sequence alignment (from analysis in Figure 5). (B) Molecular model of TM3 of OppC. generated by restrained in vacuo molecular dynamics simulations. The residues of the 'conserved' face are shown in pale grey and those of the 'hydrophobic' face in dark grey.

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RONG-I HONG AND MARK S. P. SANSOM model of the structure of this helix in a bilayer environment it is possible to refine such models via multi-nanosecond molecular dynamics simulations which include helix, lipid, and water atoms. However, this takes us beyond the range of this chapter, and the interested reader is referred to ref. 58 for a review.

5 Conclusions We have shown how the computational analysis of sequences and modelling can be used to aid experimental studies of membrane transport proteins. Much information can be gathered just by examining protein sequences. Homologous proteins can be detected by searching databases using sequence comparison methods. The sequences of these proteins may be used to generate alignments that can be employed in secondary-structure prediction and periodicity analysis. From the latter, constraints on the possible spatial arrangement of TM helices within a bundle may be deduced. Finally, three-dimensional models of TM helices may be constructed to facilitate, for example, the design of mutagenesis experiments, or as starting structures for simulations of helix/bilayer interactions.

Acknowledgements We acknowledge the financial support of The Wellcome Trust (grants to MSPS) and of The Ministry of Education, Taiwan (to RIH). Our thanks to Ian Kerr for his interest in this work.

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COMPUTER PREDICTION OF TRANSPORTER TOPOLOGY AND STRUCTURE 16. Pearson, W. R. (1996). In Methods in enzymology. Vol. 266 (ed. R. F. Doolittle), p. 227. Academic Press, London. 17. Altschul, S. F., Madden, T. L, Schaffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D. J. (1997). Nud. Adds Res., 25, 3389. 18. Beamer, L. J., Fischer, D., and Eisenberg, D. (1998). Prot. Sci., 7, 1643. 19. Pearson, W. R. (1998). J. Mol. Biol., 276, 71. 20. Dayhoff, M. O., Schwartz, R. M., and Orcutt, B. C. (1978). In Atlas of protein sequence and structure, Vol. 5, Suppl. 3, p. 345. National Biomedical Research Foundation, Washington, DC. 21. Henikoff, S. and Henikoff, J. G. (1992). Proc. NatlAcad. Sci. USA, 89, 10915. 22. Vogt, G., Etzold, T., and Argos, P. (1995). J. Mol. Biol., 249, 816. 23. Benson, D. A., Boguski, M. S., Lipman, D. J., Ostell, J., and Ouellette, B. F. F. (1998). Nud. Acids Res., 26, 1. 24. Stoesser, G., Moseley, M. A., Sleep, J., McGowran, M., Garcia-Pastor, M., and Sterk, P. (1998). Nud. Acids Res., 26, 16. 25. Tateno, Y., Fukami-Kobayashi, K., Miyazaki, S., Sugawara, H., and Gojobori, T. (1998). Nud. Acids Res., 26, 8. 26. Bairoch, A. and Apweiler, R. (1998). Nud. Acids Res., 26, 38. 27. Barker, W. C., Garavelli, J. S., Haft, D. H., Hunt, L. T., Marzec, C. R., Orcutt, B. C., Srinivasarao, G. Y., Yeh, L-S. L., Ledley, R. S., Mewes, H.-W., Pfeiffer, F., and Tsugita, A. (1998). Nud. Acids Res., 26, 27. 28. Bleasby, A. J., Akrigg, D., and Attwood, T. K. (1994). Nucl. Adds Res., 22, 3574. 29. Lipman, D. J., Altschul, S. F., and Kececioglu, J. D. (1989). Proc. Notl Acad. Sci. USA, 86, 4412. 30. Barton, G. J. and Sternberg, M. J. E. (1987). J. Mol. Biol., 198, 327. 31. Sander, C. and Schneider, R. (1991). Proteins: Struct. Func. Genet., 9, 56. 32. Thompson, J. D., Higgins, D. G., and Gibson, T. J. (1994). Nucl. Acids Res., 22, 4673. 33. Notredame, C. and Higgins, D. G. (1996). Nud. Adds Res., 24, 1515. 34. Eddy, S. R. (1995). In Proceedings of the Third International Conference on Intelligent Systems for Molecular Biology, p. 114. AAAI Press, Menlo Park. 35. Sonnhammer, E. L. L., Eddy, S. R., and Durbin, R. (1997). Proteins: Struct. Func. Genet, 28, 405. 36. Taylor, W. R., Jones, D. T., and Green, N. M. (1994). Proteins: Struct. Func. Genet, 18, 281. 37. Kyte, J. and Doolittle, R. F. (1982). J. Mol. Biol., 157, 105. 38. Engelman, D. M., Steitz, T. A., and Goldman, A. (1986). Annu. Rev. fliophys. Biophys. Chem., 15, 321. 39. Cornette, J. L, Cease, K. B., Margalit, H., Spouge, J. L., Berzovsky, J. A., and DeLisi, C. (1987). J. Mol. Biol., 195, 659. 40. von Heijne, G. (1992). J. Mol.Biol., 225, 487. 41. Jones, D. T., Taylor, W. R., and Thornton, J. M. (1994). Biochemistry, 33, 3038. 42. Cserzo, M., Wallin, E., Simon, I., von Heijne, G., and Elofsson, A. (1997). Prot. Eng., 10, 673. 43. Hirokawa, T., Boon-Chieng, S., and Mitaku, S. (1998). Bioinformatics, 14, 378. 44. Sonnhammer, E. L. L., von Heijne, G., and Krogh, A. (1998). In Proceedings of the Sixth International Conference on Intelligent Systems for Molecular Biology, p. 175. AAAI Press, Menlo Park. 45. Russell, R. B. and Barton, G. J. (1993). J. Mol. Biol., 234, 951. 46. Persson, B. and Argos, P. (1994). J. Mol. Biol., 237, 182. 47. Rost, B., Casadio, R., Fariselli, P., and Sander, C. (1995). Prot. Sci., 4, 521. 48. Yeates, T. O., Komiya, H., Rees, D. C., Allen, J. P., and Feher, G. (1987). Proc. NatlAcad. Sci. USA, 84, 6438. 227

RONG-I HONG AND MARK S. P. SANSOM 49. Komiya, H., Yeates, T. 0., Rees, D. C., Allen,]. P., and Feher, G. (1988). Proc. NatlAcad. Sci. USA, 85, 9012. 50. Donnelly, D., Johnson, M. S., Blundell, T. L, and Saunders, J. (1989). FEES Lett., 251, 109. 51. Donnelly, D., Overington, J. P., Ruffle, S. V., Nugent,]. H. A., and Blundell, T. L. (1993). Prot. Sci, 2, 55. 52. Sutcliffe, M. J., Hayes, F. R. F., and Blundell, T. L. (1987). Prot. Eng., 1, 385. 53. Ponder, J. W. and Richards, F. M. (1987). J. Mol. Biol., 193, 775. 54. Dunbrack, R. L. and Karplus, M. (1993). J. Mol. Biol, 230, 543. 55. Kerr, I. D. and Sansom, M. S. P. (1993). Eur. Biophys.J., 22, 269. 56. Kraulis, P. J. (1991). J. Appl Cryst., 24, 946. 57. Saurin, W. and Dassa, E. (1994). Prot. Sci., 3, 325. 58. Biggin, P. C. and Sansom, M. S. P. (1998). Eiophys. Chem., 76, 161. 59. PebayPeyroula, E., Rummel, G., Rosenbusch, J. P., and Landau, E. M. (1997). Science, 277, 167.

228

Chapter 10 Two-dimensional crystallization of membrane proteins PHILIPPE RINGLER, BERNARD HEYMANN, and ANDREAS ENGEL Maurice E. Muller-lnstitut fiir Mikroskopische Strukturbiologie am Biozentrum, Universitat Basel, CH-4056 Basel, Klingelbergstrasse 70, Switzerland

1 Introduction While structure determination of soluble proteins by X-ray crystallography and nuclear magnetic resonance (NMR) imaging has progressed at a remarkable rate, it has been far less successful for membrane proteins, as manifested by the small number solved to atomic resolution. The fact that membrane proteins must be solubilized using detergents leads to large complexes unsuitable for study by NMR. Also, solubilized membrane proteins are often destabilized by the detergent and are thus unsuitable for 3-D (three-dimensional) crystallization. 3-D crystals that have allowed atomic resolution by X-ray crystallography have so far all been obtained from proteins known for their high structural stability. These include four bacterial porins (for a review, see ref. 1), the photosynthetic reaction centre (2), two cytochrome c oxidases (3, 4), two forms of the bovine mitochondrial cytochrome bca complex (5, 6), the purple bacterium light-harvesting complex II (7), bacteriorhodopsin (8), and a microbial K+ channel (9). In addition, a membrane inserting toxin (a-haemolysin) has recently been crystallized in 3-D (10). In most cases, solubilized membrane proteins have been crystallized using conventional methods that foster the interaction of hydrophilic protein surfaces. In the case of the cytochrome c oxidase, the formation of a complex with Fv fragments has been essential in assembling 3-D crystals (11). Thus, the immune system has been exploited to engineer a highly specific antibody fragment providing additional hydrophilic contacts. A novel method for crystallizing membrane proteins in a cubic lipid phase was used to obtain highquality 3-D crystals of bacteriorhodopsin (12). The inherent design of membrane proteins for the 2-D (two-dimensional) environment of the lipid bilayer suggests that the most appropriate geometry for a regularly packed protein would be a 2-D crystal. Electron crystallography is 229

PHILIPPE RINGLER ET AL.

ideally suited for the study of such a specimen at different levels of resolution, up to resolving atomic positions (for a review see ref. 13). As demonstrated recently, 2-D protein crystals and their conformational changes can also be studied at high resolution by atomic force microscopy (14). Suitable 2-D crystals are more easily obtained than 3-D crystals, because membrane proteins have a propensity to pack into a lipid bilayer. Two-dimensional crystals of membrane proteins have been obtained in three different ways: (a) by inducing packed regular arrays of a highly abundant protein in its native membrane (see Section 2.1); (b) by reconstituting the purified membrane protein into a lipid bilayer at high protein density (see Section 2.2); and (c) by adding precipitants to promote protein-protein interactions, analogous to 3-D crystallization, but yielding 2-D crystals (see Section 2.3). This chapter discusses various techniques for obtaining 2-D crystals of membrane proteins and their analysis by electron crystallography.

2 Two-dimensional crystallization 2.1 Two-dimensional crystallization in native membranes Some membrane proteins have a natural propensity to form regular arrays within the native membrane. Since the membrane protein does not dissociate from the lipid bilayer, its native orientation is maintained. Additionally, the proteins are not exposed to harsh detergent and are therefore kept in their native conformation. One disadvantage is that these spontaneously formed 2-D crystals are rarely highly ordered because other components can be trapped and disturb the crystal growth. Another disadvantage is that this approach is limited to cases where the membrane protein occurs at high density. However, the quality of naturally occurring 2-D crystals can be improved by either detergent extraction of lipids, fusion of crystalline patches, or incubation with additives. Examples of naturally occurring 2-D crystals are the bacteriorhodopsin forming the highly crystalline purple membranes from Halobacterium salinarium (15), the gap junction channels formed by members of the connexin family (16-19), and water channels (20, 21). 2-D crystals have been produced by various treatments of native membranes in the case of the photosystem I reaction centre (22), the mitochondrial porin (voltage-dependent anion channel, VDAC) (23), the acetylcholine receptor channels in post-synaptic membranes (24), and the Ca2+-ATPase from the sarcoplasmic reticulum (25). The native purple membranes of Halobacterium salinarium were used for the first structure determination of a membrane protein at 7 A resolution by electron crystallography (15). The quality and size of the crystal patches were improved by adding small amounts of octyl-p-o-glucopyranoside (OG) and dodecyl triammonium chloride, which resulted in the fusion of the patches into large (2 um) 2-D crystals (26). Detergent extraction of a fraction of the native 230

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

lipids increased the packing density of the membrane protein resulting in better-ordered 2-D crystals (27). Detergent extraction improved the crystallinity of cardiac gap junctions, a hexagonal crystal of connexons (19). A combination of overexpressing connexin and detergent extraction led to gap junctions that allowed their projection map to be determined at 7 A resolution (28). Finally, detergent extraction was essential for the induction of photosystem I 2-D crystals (22), while Mannella (23) has produced 2-D crystals of the mitochondrial porin in the native membrane by the use of phospholipase A2 (see Protocol 15). The nicotinic acetylcholine receptor spontaneously crystallizes within membrane vesicles isolated from the electric organs of the ray Torpedo marmorata (29). Tubular crystals appear after several weeks of incubation in buffer at 4°C. The first step in tubular crystallization is the formation of dimers, via disulfide (S-S) bridges between two delta subunits of the receptor. Dimers then align into ribbons, which gradually rearrange into a helical array (A. Brisson, personal communication). 2-D crystallization of some membrane proteins can be induced by additives. For example, Ca2+-ATPase from sarcoplasmic reticulum can be crystallized within the membrane by induction with vanadate (25, 30). Vanadate is an ionic inhibitor that probably locks the flexible enzyme into a single conformation. Tubular crystals of Ca2+-ATPase form when the protein rearranges within the membrane into dimer ribbons, in a manner similar to the tubular crystallization of the nicotinic acetylcholine receptor. Native tetragonal crystals were initially observed in lens fibres and identified as crystals of the major intrinsic protein (MIP) (31), the first water-channel protein sequenced (32). Tetragonal arrays observed in membranes of cells overexpressing aquaporin 4 further demonstrate the propensity of water-channel proteins to form 2-D crystals (33). 2.2 Crystallization of purified membrane protein by reconstitution into lipid bilayers In contrast to 3-D crystallization the purified protein is reconstituted into a native-like environment during two-dimensional crystallization. Nevertheless, to produce high-quality 2-D crystals, the protein of interest must be purified away from other proteins and contaminants as in 3-D crystallization. This involves solubilization of the original membrane with detergent and commonly requires several subsequent separation steps (see Chapter 11). Finally, the pure protein is obtained in a detergent solution, often with residual lipids. In fact, the latter often contribute to the stability of the membrane protein and may be essential for successful 2-D crystallization. The nature of the detergent, in both protein purification and subsequent crystallization trials, is a critical determinant of success. An extensive list of 2-D crystals obtained via reconstitution of detergent-solubilized membrane proteins is given in Table 1. The reconstitution of membrane proteins into bilayers is achieved by mixing lipids and protein, both solubilized in detergents, and then decreasing the 231

Table 1 Examples of 2-D crystals obtained via reconstitution of detergent-solubilized membrane Buffer/pH

Additive

1.0

—

PLA2

E. co// lipids

0.5

Hepes/6.0

MgCI2

OG

DOPC

1-0.3

PBS/7.0

Human erythrocyte

Ci2E8

DMPC, cholesterol

1

Band 3

Human erythrocyte

8-POE or C12E8

DMPC

BR

Halobacterium halobium expressed in £ coli

Triton X-100

Ca2+-ATPase

Rabbit sarcoplasmic reticulum

Cytochrome aa3

Protein

Origin

Detergent

Lipid added

AQP1

Bovine erythrocyte

OG

DMPC

AQP1

Human erythrocyte

OG

AQP1

Human erythrocyte

Band 3

LPR

Resolution

Reference

6 A (3D)

34

25-37#°C

6 A (3D)

35

—

deglycosylated

7 A (3D)

36

PBS/8.0

PEG 200/ MgCI2

deglycosylated

20 A (3D)

37

0.75-1.25

Hepes/7.0

MgCI2

22-37°C

—

38

Halobacterium halobium lipids

—

Na-acetate/5.0

—

dialysis in the dark

3.6 A (2D)

39

C12E8 or Triton X-100

egg PC, PS,

0.5

lmidazole/7.4

Na3V04, KCI, MgCI2

Bio-Beads method

40

Paracoccus denitrificans

DM

egg PC, brain PS

0.85

Bistrispropane/ 7.0

—

tubular crystals "—

41

Cytochrome aa3

Rhodobacter sphaeroides

DM

egg PC, brain PS

1

Bistrispropane/ 7.0

isopropanol

—

—

41

Cytochrome b6f

Chlamydomonas reihardtii

HECAMEG

eggPC, DOPG

0.2

Tricine/8.0

CaCI2, e-aminocaproic acid, benzamidine, PMSF, AS

Bio-Beads method and freezethawing

8A(2D)

42

Cytochrome

Neurospora crassa

Triton X-100

soybean PC, bovine brain PS

0.5-2

20 A (2D)

43

PA, cholesterol

-/5.5

Remarks

Cytochrome

Tricine/8.0

ZnCI2, isoascorbic acid

tubular crystals, Bio-Beads method

16 A (3D)

44

0.29

Tris/9.0

—

tubular crystals

6 A (2D)

45

—

Hepes/7.0

MgCI2

double-layered 2D crystals

—

46

Hepes/7.4

—

membrane fusion

7A(2D)

28

PBS/8.0

glutathione

8 days' dialysis at 20 °C

3 A (2D)

47

Tris/6.8

PEG4000/AS

on air/water interface

10.3 A (2D) 48

—

PEG 4000/AS

on C surface

8 A in plane 49 (3D)

—

Tris-succinate/ 6.5

CaCI2, phospholipases

Bio-Beads

—

50

1

Hepes/7.0

MgCI2

151

25 A (3D)

51

Hepes/7.0

MgCI2

37"C

20A(2D)

52

0.05

NH4HC03/ 7.8

MgCI2

in the dark

8.5 A (2D)

53

DMPC

0.5

Tris-HCI/8.0

—

—

7A(2D)

54

soybean PC or LLPC

—

Tris-HCI/8.0

CaCI2

—

18 A (20)

55

Bovine heart

Triton X-100

soybean PC/PE, CL, a-Toc

Cytochrome bo ubiquinol oxidase

E. coli

Triton X-100

egg PC, brain PS

Gap junction

Ovine lens

OG

—

Gap junction Rat heart (a± connexin, (expressed in Cx43) BHK cells)

Tween-20 and DHPC

—

Glutathione transferase

Rat liver microsomes

Triton X-100

bovine liver lecithin

H*-ATPase

Neurospora crassa

DDM

—

H*-ATPase

Neurospora crassa

DDM

—

H+-ATPase (CFOF1)

Spinach chloroplast

Triton X-100

—

LamB (maltoporin)

E. coli

8-POE

E. coli lipids

LamB (maltoporin)

E. coli

8-POE

E. coli lipids

LHI

Rhodospirillum rubrum

OG

DOPC

LHII

Rhodovulum sulfidophilum

OG

LHII

Ectothiorhodospira spp.

DDM

bCl

complex

0.15

—

0.1-1

Table 1 (Continued) Lipid added

LPR

Buffer/pH

Additive

Remarks

Resolution

Reference

—

glycine/7.0

glycerol

35-40 °C, batch method

3.4 A (3D)

56

thylakoid lipids

1

—

DGDG

batch method

30 A (2D)

57

DM

E. coli phospholipids

0.2-0.5

Mes/6.0

MgCI2

23-37#°C

9 A (2D)

58

Campylobacter jejuni

8-POE

POPC/DMPC

1

Hepes/7.4

MgCI2

—

20 A (2D)

59

NADU: ubiquinone oxidoreductase

Bovine-heart mitochondria

Cholate

—

—

Na-acetate/5.5

AS

—

22 A (2D)

60

NADH: ubiquinone reductase

Neurospora crassa

Triton X-100

soybean PC

0.3-0.4

Tris-acetate/ 7.0-9.0

—

—

40 A (3D)

61

OmpC

E. coli

SDS

lipidA

0.6

Tris-HCI/8.0

MgCI2

—

25 A (2D)

62

OmpF

E. coli

8-POE

DMPC

0.16-0.72

Hepes/7.0

MgCI2

37°C

22A(2D)

63

PhoE

E. coli

SDS

DMPC

0.25

Tris/7.5

—

phospholipase

6 A (3D)

64

PSI RC

Phormidium laminosum

OTG

DMPC

0.31

Hepes/7.0

MgCI2

25-37°C

20 A(2D)

65

PSI RC

Synechococcus Triton X-100 spp. or SB-12

Tris-succinate/ 6.5

Phospholipases

Bio-Beads method

16 A (2D)

22

Protein

Origin

Detergent

LHCII

Pea chloroplasts

Triton X-100 — or Triton X-100 and NG

LHCII

Pisum sativum Triton X-100 (overexpressed or Triton X-100 LHCP and NG in E. co/0

MIP (AQPO)

Ovine lens

MOMP

—

PSIRC

Synechococcus OTG spp.

PSII RC

Spinach thylakoids

DDM

—

PSII RC

Spinach thylakoids

OTG

DMPC

PSII RC

Spinach thylakoids

Triton X-100

—

PSII RC

Spinach thylakoids

HTG

thylakoid lipids

DMPC

Mes/6.0

Ammonium ferric citrate

26-37 °C

19 A (3D)

66

bis Tris/6.5

taurine, MgCI2

—

25A(2D)

67

Hepes/7.5

—

dilution method

20 A (2D)

68

—

Hepes/7.5

MgCI2

tubular crystals

17 A (2D)

69

—

Mes/6.5

CaCI2, zinc acetate, ascorbate, butylated hydroxytoluene

—

8A(2D)

70

0.5-1

— 0.1-1

6 s rH

•z.

RC

Rhodopseudomonas viridis

LDAO

—

—

PBS/5.0

—

23°C

—

71

RC-LHI

Rhodobacter sphaeroides

OG

DMPC, DOPC or plant PC

0.4-0.9

Hepes/7.5

—

20-35 °C

25 A (2D)

72

RC-LHI

Rhodospirillum rubrum

OG

DOPC

0.5

Rhodopsin

Bovine rod cells C8E4

Rhodopsin

Frog rod cells

Ubiquinone: Neurospora cytochrome c crassa reductase

-l O

O 1— O ff\ \J1

—1 NH4HC03/7.9

MgCI2

—

16 A(2D)

73

jz

0.3-1.6

Hepes/7.0

MgCI2

—

9 A (2D)

74

—1

—

Tes/7.5

—

membrane fusion

15 A (3D)

28

Bio-Beads method

—

M

' soybean PC Tween-80 or — Tween-80 and Tween-20 Triton X-100

soybean PC, brain PS

—

Tris-acetate/

7.0

—

0

•z.

0 -n

75

m CO 33

•z. m "O 33 O

M OJ

PHILIPPE RINGLER ET AL.

detergent concentration. During this process, the small micellar structures coalesce into larger structures leading to the formation of vesicles and sheets. Both vesicles and sheets consist of lipid bilayers with varying amounts of protein. At high protein concentrations, the interaction between proteins in the membrane may lead to regular packing and 2-D crystals. Crystals may still be obtained with an excess of protein, but a fraction of the protein ends up in amorphous aggregates.

2.2.1 An important parameter: the lipid/protein ratio At the start of a typical reconstitution experiment, an excess of detergent ensures a homogeneous distribution of protein and lipid in micelles. As the detergent concentration is decreased, lipid and protein interact due to the exposure of their hydrophobic surfaces. With an excess of lipid over protein, the protein is mainly incorporated into lipid bilayers, similar to its native state. In an excess of protein over lipid, some of the protein aggregates, probably in a denatured form. An important parameter is therefore the lipid:protein ratio (LPR), which should be low enough to promote crystal contacts between protein molecules, but not so low that the protein is lost to aggregation. When the membrane protein is reconstituted from a mixture of solubilized components, crystal-ordering of proteins may occur during reconstitution. In some cases it is difficult to distinguish parameters affecting the incorporation of protein into the lipid bilayer from those leading to crystalline order. In other cases these two processes are quite distinguishable. For crystal packing during reconstitution, the LPR of the reconstitution experiment is critical. In any case, as this parameter is quite unpredictable, it has to be determined by carefully designed reconstitution series. While the lipid content of the reconstitution mixture is generally a well-controlled parameter, the content of monodisperse protein is sometimes unknown, because protein assays do not indicate the amount of aggregates. Therefore, it is advisable to determine the fraction of aggregated protein in a given protein batch by, for example, negative stain electron microscopy, ultracentrifugation, or light scattering. Alternatively, it has also been possible to reconstitute with an excess of lipids and then improve crystal packing by mild digestion of the lipids with phospholipase A2 (see Section 2.2.9).

2.2.2 The choice of lipids The lipid mixture used for reconstitution has an influence on the crystallization results. Crystallization is more likely to occur when the lipid bilayer is in the fluid phase and thus allows some lateral mobility of the inserted membrane proteins. While saturated lipids are chemically more stable and preferred, unsaturated lipids (such as those from Escherichia coli) have been successfully used to produce highly ordered crystals. A good compromise is dimyristoylphosphatidylcholine (DMPC)—a lipid frequently used (see Table 1) with success, which has saturated fatty acids but with a phase-transition temperature close to room temperature (23°C). Native lipids are often ideal in terms of stability and 236

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

transition temperatures, and they also provide mixtures of head group charges and molecular geometries similar to the membranes from which the protein originated. Bacteriorhodopsin, which can only be crystallized from a solubilizcd stale in the presence of the native purple membrane lipids, is a classic example (76). Since synthetic lipids, E. coli lipids, soybean lecithin, and egg lecithin have all been successfully used for 2-D crystallization, no general recommendations can be made as to which lipid or lipid mixture is most suitable for any one particular membrane protein. Polyunsaturaled lipids with fatty acid chains containing unconjugated double bonds are easily oxidized, and therefore the commercial source (i.e. Avanti Polnr-Lipids) of these lipids is often a chloroform solution (at 10 or 20 mg/ml) or a lyophilized powder requiring storage at -20°C in the dark. Prior to their use for 2-D crystallization trials, lipids have to be transferred to detergent-containing buffer solutions (see Protocol 1). Lyophilized lipids can easily be weighed and re-dissolved at 1-10 mg/ml in buffer containing a high concentration of

Protocol 1 Preparation of lipid stock solution In detergent-containing buffer Equipment and reagents • 10 or 20 mg/ml lipid stock solutions in chloroform (Avanti Polar-Lipids) • Nitrogen gas bottle with reduced-pressure outlet device

• 5 ml borosilicate glass vials • Glass syringe (Hamilton, 10 or 100 ul) • Organic solvents (chloroform, diethyl ether, and methanol, HPLC grade)

Method 1. Wash the syringe and the vials first with chloroform/methanol (1:1 (v/v)) then with pure chloroform, and pipette the appropriate amount of chloroform-solubilized lipid into the glass vial, 2. Evaporate the organic solvent and dry the Upid on to the glass walls with a nitrogen stream at room temperature. 3. Re-solubilize the lipid in a small volume of diethyl ether and evaporate again with a nitrogen stream.0 4. Re-solubilize the lipid at 10 times the final concentration in buffer solution containing detergent at a concentration 10 x (CMC + 3 X concentration of lipids (mol/1)). 5. Dilute to the desired final concentration with buffer devoid of detergent,*1 " Diethyl-ether resolubilization dilutes the remaining chloroform and enables its efficient removal in the second evaporation step. Vacuum evaporation is an alternative for eliminating all organic solvents. * Ultrasound and mild heating can be used to clarify the solution.

237

PHILIPPE RINGLER ET AL.

detergent—at least above the critical micellar concentration, CMC + 3 x [lipid]. Alternatively, the lipids can be co-solubilized with a chloroform/methanol mixture and dried in a glass tube (see Protocol 1). All organic solvent must be removed before solubilization in detergent buffer as it interferes with the crystallization. This is achieved by blowing dry nitrogen gas over the sample followed by evaporation in a high-vacuum system. 2.2.3 The use of additives for 2-D crystallization and the influence of pH Several proteins require a particular buffer and polyvalent salts to produce ordered arrays. Ca2+-ATPase crystallized in the presence of vanadate (30, 40), while the photosystem-I reaction centre from Phormidium laminosum (65) and bovine lens connexin (46) required Mg2+. Magnesium was also implicated in the successful crystallization of a porin from Campylobacter jejuni (59). Crystallization of the cytochrome be, complex from a Triton X-100 solution into tubular crystals was found to require the presence of zinc, a-tocopherol, and cardiolipin (44). Optimal conditions for the 2-D crystallization of photosystem I from Synechococcus included the presence of ammonium ferric citrate, use of the detergent octyl-p-D-thioglucoside (OTG), and a particular temperature profile during dialysis (65, 66). In contrast, AQP1 formed aggregates with ammonium ferric citrate in the dialysis buffer (unpublished data), while highly ordered crystals were produced at pH 6 in the presence of the divalent magnesium cation (77). Since counter ions modulate electrostatic interactions, the surface charge of the protein plays an essential role. Therefore, the pH of the buffer may be of key importance. In many practical cases this has been optimized experimentally (see Table 1). 2.2.4 Choice of detergent and determination of the stability of solubilized protein Many membrane proteins are destabilized on extraction from their native membranes, especially when short-chain (high CMC) detergents are used. While the proteins can be effectively solubilized with detergents that replace the lipid and keep the hydrophobic surfaces of the protein shielded from water, delipidation may destabilize the protein that may then be denatured by shortchain detergents. The use of smaller but harsher detergents is a prerequisite for dialysis-driven 2-D crystallization (see Section 2.2.7), and is also important for 3D crystallization, as they do not impair protein-protein interactions essential for crystal ordering. The choice of detergent is critical: there is a fine balance between disrupting the membrane to solubilize a membrane protein and preserving its structural integrity. Table 2 summarizes the main characteristics of commonly used detergents in the field of membrane protein solubilization. As discussed above, reconstitution is closely linked to the properties of the detergents used both during purification and the reconstitution itself. Therefore, by following the protocols below, the detergents used for purification can be exchanged for a different detergent used for reconstitution. 238

Table 2 Detergents used in the solubilization of membrane proteins Biological detergent

Formula

Anlonic Cholic acid, sodium salt Deoxycholic acid, sodium salt Lauryl sulfate, sodium salt (sodium dodecyl sulfate, SDS)

Aggregation number

Molecular weight (g/mol)

CMC (mM)

C24H3905Na

430.6

9.5 (pH 9.0), 14 (pH 7.5)

2-4

C24H3904Na

414.6

5

C12H25NaS04

288.4

2.6 (pH 7.5), 8.27 (H20)

4-10 60-100

Taurocholic acid, sodium salt

C26H44NNa07S

537.7

3-11 (0.05 M Nad)

4

Taurodeoxycholic acid, sodium salt

C26H44NNa06S

521.7

1-4 (0.05 M NaCI)

6

C19H42NBr

364.5

1

169

C15H34NBr

308.3

14

—

CHAPS

C32H58N207S

614.9

8(H20)

CHAPSO

C32H58N208S

630.9

8(H20)

10(H20) 11(H20)

DHPC (diheptanoylphosphatidylcholine)

C22H44N08P

481

1

—

LDAO (lauryldimethylamine-Noxide)

C14H31NO

229.4

1-2

76

Zwittergent 3-08 (3-(/V,/Vdimethyloctylammonio)propansulfonate)

C13H29N03S

279.4

330

—

Zwittergent 3-10 (3-decyldimethylammonio)propansulfonate)

C15H33N03S

307.5

25-40

41

Cationic Cetyltrimethylammonium bromide (CTAB, Hexadecyltrimethylammonium bromide Dodecyltrimethylammonium bromide Zwitterionic

Table 2 (Continued) Biological detergent

Formula

Molecular weight (g/mol)

CMC (mM)

Aggregation number

Zwittergent 3-12 (3-(N-dimethyllaurylammonio)propansulfonate) (lauryl sulfobetain, SB-12)

C17H37N03S

335.5

2-4

55

Zwittergent 3-14 (3-(N,N-dimethylmyristylammonio)propansulfonate)

C19H41N03S

363.6

0.1-0.4

83

Zwittergent 3-16 (3-(/V-dimethylpalmitylammonio)propansulfonate)

C21H45N03S

391.7

0.01-0.06

155

BIGCHAP

C42H75N3016

878.1

3.4

10

Deoxy-BIGCHAP

C42H75N3015

862.1

1.1-1.4

8-16

40

Non-ionic

Brij 35 (polyethyleneglycol-dodecylether, C12E23)

—

—

0.05-0.1

Digitonin

C56H92029

1229.3

—

5-6

rc-Decyl-p-D-glucopyranoside

C16H3206

320.4

2.2 (H20), 2.3 (0.01 M PBS)

—

rvDecyl-hexaethyleneglycolether (C10E6)

C22H4607

422.6

0.9 (0.05 M NaCI)

—

n-Decyl-fJ-D-maltopyranoside (DM)

C22H4201:L

482.6

1.8 (0.15 M NaCI)

—

n-Dodecyl-nonaethyleneglycolether (C12E9)

C30H62010

582.8

0.046 (0.01-0.2 M NaCI)

—

n-Dodecyl-p-o-gl ucopyranoside

C18H3606

348.5

0.19 (H20), 0.13 (0.05 M NaCI)

—

n-Dodecyl-hexaethyleneglycolether (C12E6)

C24H5007

450.6

0.087 (0.05 M NaCI)

—

n-Dodecyl-octaethyleneglycolether (C12E8)

C28H5809

538.8

0.05-0.1 (0.1-0.2 M NaCI)

120-127

n-Dodecyl-pJ-D-maltopyranoside (lauryl maltoside, DDM)

C24H460±1

510.6

0.17 (H20)

98

HECAMEG (methyl-6-0-(A/)-heptyl-carbamoyl)-a-Dglucopyranoside

C15H29N07

335.4

19.5

—

n-Heptyl-p-D-glucopyranoside

C13H2606

278.3

79

—

-o m

rn -H

n-Heptyl-p-o-thioglucopyranoside (HTG)

C13H2605S

Lubrol (C12E9-10)

294.4

30

582

0.1 (0-0.05 M NaCI)

Mega-8 (W-octanoyl-W-methylglucamine)

C15H31N06

321.4

79 (H20). 58 (0.05 M NaCI)

Mega-9 (WfionanoyWmethylglucamine)

C16H33N06

Mega-10 (/V-decanoyl-N-methylglucamine)

C17H35N06

335.5 349.5

6-7 (H20)

n-Nonyl-B-D-glucopyranoside (NG)

110 (0-0.1 M NaCI)

25(H20)

306.4

6.2-6.5 (0.15 M NaCI), 3.5 (1 M NaCIO

Nonidet P-40 (NP-40) (Octylphenoxypolyethoxyethanol)

—

603.0

0.05-0.3

—

n>Octyl-B-D-glucopyranoside (OG)

C14H2806

292.4

24.4 (H20), 23.4 (0.1 M NaCI)

—

n-Octyl-(3-D-thioglucopyranoside (OTG)

C14H2806S

308.4

9(H 2 0)

Octyl polydisperse oligooxyethylene (8-POE)

—

—

6.6

— —

C16H3405

306.45

6

—

(C8E3-11, mean n = 5)

Octyl tetraoxyethylene (C8E4) Tween-20 (polyoxyethylene (20) sorbitan monolaurate

—

1227.54

0.059

—

Tween-80 (polyoxyethylene (80) sorbitan monolaurate

—

1309.68

0.012

—

Triton X-100 (polyethylene glyco-p-isooctylphenyl ether)

—

625

0.3 (H20), 0.29 (0.1 M NaCI)

100-155

Triton X-100 hydrogenated

—

631

0.25 (0.05 M NaCI)

—

Triton X-114 (cloud point 22°C)

—

537

0.2

—

PHILIPPE RINGLER ET AL.

Protocol 2 Exchange of detergent using a Centricon concentrator device Equipment and reagents • Centricon (50 000 or 100 000 molecular weight cut-off (MWCO), Amicon)

Centrifuge and fixed-angle rotor (e,g, Sorvall, SS-34 rotor)

Method 1. Dilute the membrane protein solution 50 times in the new detergent buffer (40 ul concentrated sample in 2 ml final volume)." 2. Concentrate the 2 ml to 40 ul by spinning at 5000 g (Centricon-50) or 1000 g (Centricon-100) in the SS-34 rotor for the appropriate time (1-2 h at 4°C). 3. Repeat steps 1 and 2 twice to reach the 50 x 50 x 50 = 125000-fold theoretical dilution factor for the initial detergent. " The total amount of membrane protein treated per Centricon should be sufficient in order to neglect the usual protein losses resulting from unspecific adsorption to the device (5-10 ug). Note that some membrane proteins precipitate in Centricon devices, a possibility which has to be determined for each protein, A stringent prerequisite for this method is a sufficiently targe membrane protein complex (i.e. > 50 kDa or > 100 kDa for Centricon-50 or Centricon100, respectively).

Protocol 3 Exchange of detergent using size-exclusion gel filtration on Sephadex G-200 Equipment and reagents • Packed column with Sephadex G-200 (Pharmacia)"

Detergent-containing buffer

Method 1. Concentrate the sample as in Protocol 2 step 2 to a small volume (e.g. 250 ul, final concentration typically 5-10 mg/ml), 2. Pack the column with pre-swollen Sephadex G200 resin (e.g. 5 ml) and preequilibrate the column bed with the solution of the new chosen detergent. 3. Load the concentrated sample (the volume should be smaller than 5% of the total bed volume) and elute with the new detergent-containing buffer.

242

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

Protocof 3 continued

4. Collect the fractions corresponding to the solubilized membrane protein and check the concentration with detergent-compatible protein assays (see Protocol lO).1" "Instead of a Sephadex G-200 column, affinity chrornatography on Ni-NTA agarose (Qiagen) may be used, provided that the protein has a poly-histidine tag. b Protein concentration drops by 5 to 10 times.

Protocol 4 Exchange of detergent using sucrose-gradient centrifugation" Equipment and reagents • Peristaltic pump and gradient maker

Method 1. Concentrate the sample to a small volume (e.g. 250 ul. final concentration typically 5-10 mg/ml). 2. Prepare a gradient of sucrose (10-50%; total volume 5 ml) in buffer containing the new detergent. 3. Pour the sample on the top of the gradient and spin at 50 000 g for 2 hat 4°C. 4. Collect the fractions and assay the protein. "This protocol is derived from the one used to exchange OG with decyl-B-D-maltopyranoside (DM) on MIP channels (58). The manner in which the detergent concentration is decreased for reconstitution and subsequent 2-D crystallization is an important consideration. The commonly used techniques for detergent removal are dilution (78) (Section 2.2.6), dialysis (79) (Section 2.2.7). and selective adsorption of the detergent on solid supports (80) (Section 2.2.8).

2.2.5 Analysis of detergent, lipid, and protein Optimal 2-D crystallization trials require accurate knowledge of the three key components: detergent, lipid, and protein. In this section we discuss several analytical methods tor measuring the concentration of these compounds. Detergent in aqueous solution decreases the surface tension, and this offers a simple and convenient way to determine the detergent concentration. The surface tension in a solution can be measured in many different ways, of which two are given here. The mass of a drop breaking away from a tube is determined by the surface tension, allowing its assessment by the falling-drop weight method. In the sitting-drop method, the decrease in surface tension due to 243

PHILIPPE RiNGLER ET AL.

Protocol 5 Free (monomerlc) detergent concentration measurement using the falling-drop weight method Equipment and reagents NB: Rinse the glassware with purified water to remove residual organic solvents before use, and use disposable plastkrware. • Pipettor • Glass pipettes of 50-200 p.1 volume and 0.7-2 mm radius • Buffer used for reconstitution trials

• Stock solution of detergent in buffer at a concentration of about 5-10 times the CMC

Method 1. Fix a glass pipette in the pipettor, 2. Fill the pipette with a test solution, 3. Mount the pipettor in a retort stand above the balance pan. 4. Place a tube or weighing boat on the weighing pan underneath the pipette. 5. Zero the balance and make sure the reading does not change over several seconds. 6. Slowly push the liquid from the pipette to form a hanging drop," push more liquid out until the drop detaches and falls in the tube or on the weighing boat.6 7. Wait until the mass reading stabilizes then record the weight. Continue pushing out drops in the same way until too little liquid is left to produce a falling drop. " It should take several seconds for the drop to form to ensure partitioning of the detergent into the surface. b Note that precious sample can be recovered from the balance for further analysis.

detergent is manifested in the spreading of a sample drop on a hydrophobic surface. In both cases, the sample must be diluted below the CMC to observe a measurable surface tension change. The dilution factor then allows the total detergent concentration to be estimated, The calculation of the free detergent concentration from the falling-drop weight measurements is given in the following. The falling-drop weight, Mt, is approximately proportional to the logarithm of the free detergent concentration, [D,-]:

where MD and M1 are the intercept and slope of the line. D1 is practically equal to Lhe total detergent concentration below the CMC. Above the CMC, |D1| is approximately equal to the CMC, giving a horizontal line lor M1 at high detergent concentrations. These two lines intersect approximately at the CMC (seeFigure1)), 244

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

Figure 1 Example of falling-drop weight method standard curve. The falling-drop weight (M, in mg) is plotted against the logarithm of the ratio [D]/CMC, with [D] the concentration of detergent (mM) and CMC the critical micellar concentration of the detergent (mM), The dotted lines intersect at the CMC [arrow). Below the CMC, the free detergent concentration [D.] corresponds to the total detergent concentration [D], whereas above the CMC, [D1] remains approximately constant and corresponds almost to the CMC.

Protocol 6 Determination of the free detergent concentration using the sitting-drop method Equipment and reagents • Teflon film

• Camera, or video recorder

Method 1. Deposit 10 ul volumes of sample on the Teflon film. 2. Take a photograph of the drops," 3. Measure the diameters of the drops and compare with a standard series of the same detergent* "This must be done quickly to avoid excessive evaporation from the drops. b Solutions for a standard curve: make a series of detergent solutions with the buffer as diluent in the range 0.1 to 3 times the CMC and allow the solutions to equilibrate overnight at the temperature at which the measurements will be made. When purifying membrane proteins in the presence' of various detergents it is of great interest to know how much endogenous lipids are co-purified with the protein of interest. Below, three methods for the specific titration of lipids, which should help in designing 2-D crystallization trials, are described. Phospholipids like lecithin, lysoledthin. and sphingomyelin can be hydrolysed by plmspholipase D and the liberated choline measured by the Trindcr reaction. Briefly, the free choline is oxidized to betaine and peroxygen in the

PHILIPPE RINGLER ET AL.

Protocol 7 Determination of phospholipid with ammonium ferrothiocyanate* Equipment and reagents • Ultraviolet-visible spectrophotometer • Ammonium ferrothiocyanate reagent: dissolve 27.0 g ferric chloride hexahydrate (FeCl3-6H20) and 30.4 g ammonium thiocyanate (NR4SCN) in 1 litre of distilled water (stable for months at room temperature).

• Chloroform and methanol • Solutions for a standard curve: prepare a series of solutions of 0-0.2 mg/ml of dipalmitoylphosphatidylcholine (DPPC) in chloroform.

Method 1. Extract the aqueous sample containing phospholipid with chloroform and methanol (1:1 or 1:2. (v/v))2. Collect the chloroform extract (lower phase) in a glass tube and diy under a stream of nitrogen gas. 3. Re-dissolve the extract in 2 ml of chloroform. 4. Add 2 ml of ammonium ferrothiocyanate and mix well. 5. Allow phase separation and collect the chloroform phase in a new tube. 6. Measure the absorbance at 488 nmb (to construct the standard curve, add 2 ml of the reagent to 2 ml of each standard solution and proceed as above), " Derived from the method of Stewart (81). "The absorbance maximum and the absorptivity may vary for different Hpids, therefore it is advisable to use a standard lipid similar to that analysed for. Also, the extent of recovery from the extraction of lipid from the aqueous sample may be assessed by adding known lipid to the sample before extraction.

presence of choline oxidasc. The peroxygen then reacts with phenol and 4aminonmipyrine to produce quinonciminc in the presence of pcroxidase. The absorbance of quinoiieimine at 505 nm is proportional to the initial concentration ot'cholinc-containing lipids (linearity range from 0 to 10 mmol/1). The precise concentration of the solubilized membrane protein is always a critical parameter in designing crystallization trials. The protein assay presented here is compatible with the presence of detergents. However, it simply gives the total protein concentration irrespective of its aggregation state. Hence, samples need to be checked by negative stain electron microscopy, or the protein concentration needs to be determined before and after ultra centrifugal ion (e.g. at 50 000 % for 1 h at 4 °C). 246

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

Protocol 8 Determination of phospholipid by phosphate content3 Equipment and reagents • • • •

Solution A: 10% Mg(NO3)2 in 95 % ethanol Solution B: 0,5 M HC1 Solution C: 10% ascorbic acid Solution D: 0,42% ammonium molybdate in 0.5 M sulfuric acid • Solution E: mix 3.3 ml of solution C with 20 ml of solution D (for 20 samples) and keep on ice

• Fume hood and flame device • Pyrex or Kimax tubes (tubes should not be cleaned with soap to avoid the presence of residual phosphate) • Glass beads (20 mm diameter) • Heating water bath • Ultraviolet-visible spectrophotometer

Method 1. Place 10-100 ul of the sample (containing up to 70 umol of P) in the test tubes. 2. Add 30 ul of Solution A. 3. Shake over a flame under the fume hood until dry and the brown fumes are gone. 4. Allow to cool and add 300 u1 of Solution B, 5. Cover the tubes with glass beads and heat in a boiling water bath for 15 min. 6. Allow to cool, then add 700 n.1 of Solution E, mix and heat for 20 min at 45°C (or 1 hour at 37 DC). 7. Allow to cool and read the absorbance at 820 nm..b "Derived from the method of Ames (82). "The assay is very sensitive and l0 nmol of P will give an A2BO of 0.24, The standard curve can be established with phosphate buffer.

Protocol 9 Enzymatic determination of choline-containlng phospholipids" Equipment and reagents • Reagent 1 (standard): 4 mM choline (equivalent to 3.1 g/1 of phospholipids) • Reagent 2 (buffer): 20 mM Trisbase pH 7.8, 3 mM surfactant, and 10 mM phenol

• Reagent 3 (dried enzymes): 2000 U/l cholinesterase, 600 U/l phospholipase D, 1000 U/l peroxidase, 0.5 mM 4aminoantipyrme • Ultraviolet-visible spectrophotometer

Method 1. Reconstitute Reagent 3 with 25 ml of Reagent 2 (buffer).

247

PHILIPPE RINGLER ET AL. Protocol 9 continue*

2. Mix 10 ul of the lipid solution to be tested (or the standard) with 1 ml of Reagent 3 solution. 3. Incubate for 10 min at 37°C and measure the absorbance at 505 nm (492-546 nm).1" " Phospholipides enzymatiques PAP 150 from BioMerieux. ref. from (83), ' The colour intensity is stable for 1 h at room temperature.

Protocol 10 Bicinchoninic acid (BCA) protein assay" NB: The BCA assay is incompatible with strong chelating agents (EDTA) that can be expected to cause a depletion in copper ions necessary for this assay method. Also strong reducing substances (DTT or p-mercaptoethanol) should be avoided because they cause too much colour in the blanks. Tris and asparagine buffers at concentrations of up to 0.05 M may be used if care is taken to adjust the pH to between 11 and 11.25. Equipment and reagents • Reagent A: 4%Na2CO3H20,0.8%NaOH. 0.8% Na2 tartrate, 2% BCA, adjusted to pH 11.25 with Na2CO3.

• Reagent B: 4% cupric sulfate, pentahydrate • BSA (bovine serum albumin) or other

suitable protein Borosilicate glass test tubes Ultraviolet-visible spectrop hotometer Heating water bath

Method 1. Prepare a set of protein standards of known concentration by diluting a stock of BSA (bovine serum albumin) or other suitable protein in the same diluent as the unknown sample (concentration range from 0.02 to 0.2 mg/ml). 2, Prepare the working reagent by mixing 50 parts of Reagent A with 1 part of Reagent B. 3, Mix 0.1 ml of each standard or sample with 2 ml of the working reagent and incubate for 30 min at 60°C or room temperature for 2 h.b 4. Allow to cool and read the absorbance at 562 nm vs. water reference.' a

The BCA Protein Assay is a patent of Pierce. "BCA is a highly specific reagent for cuprous ions (Cu+) and forms a water-soluble purple complex. The Cu+ ions are produced by the reduction of cupric (Cu11) ions in alkaline medium by the peptidk bonds of the three amino acids cysteine, tryptophan, and tyrosine (Biuret reaction). Care has to be taken to avoid the presence of other reducing agents like thiols (dithiothreitol or mercaptoethanol) or high amounts of sugar (sucrose or glucose), ' Since colour development will continue slowly, it is necessary that all absorbance readings be taken within the shortest possible time.

248

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

2.2.6 Dilution method Diluting a solution of protein, lipid, and detergent decreases the concentrations of all components by equal factors, until the free detergent concentration drops below 'saturation'. Figure 2 shows an example where the concentration of octyl polyoxyethylene (8-POE) was decreased by dilution and the formation of structures of different sizes was monitored using dynamic light scattering (78). The dilution experiment led to the formation of vesicles with egg-yolk phosphatidylcholine (egg PC), or vesicles and 2-D crystals with egg PC and the porin OmpF. The latter assembled only if the dilution rate was slow. The grey regions in Figure 2a indicate the size distributions for the two cases. With only lipid present, the aggregation of mixed lipid-detergent micelles led to a heterogeneous mixture of structures shortly before the critical micellar concentration (CMC) was reached. Upon further dilution these structures became more homogeneous, and finally a single population of vesicles was formed. With both lipid and OmpF present, micellar aggregation started earlier during dilution (Figure 2a). Again a heterogeneous mixture of structures was observed. However, on further dilution the heterogeneity persisted, leading to the formation of crystalline sheets of variable size, as well as vesicles. Invariably, the large structures all exhibited crystallinity, while small vesicles had a similar size as those produced in the absence of proteins. A different representation of the same experiment is shown in Figure 2b to illustrate the relationship between detergent concentration and structure sizes. This representation is commonly used to describe the 'three-stage' model of Lichtenberg et al. (84). Stage I is characterized by a detergent concentration too low to disrupt the lipid bilayer. Stage II is the region of detergent concentration where lipid bilayer and mixed micellar structures coexist. In Stage III the detergent concentration is sufficient to fully solubilize all the components of the membrane. These specific regions are delineated by the 'saturation' and 'solubilization' points that define the onset and completion of the solubilization of large structures on addition of detergent. The micelle-bilayer transition region (Stage II) was found to be the key to reconstitution and, by implication, to 2-D crystallization (78, 85, 86). Neutron scattering (87), dynamic light scattering (88), and cryo-electron microscopy (88, 89) have shown, for several lipid-detergent systems, that this transition involves the formation of worm-like extended lipid micelles, probably capped by detergents, that must convert to vesicles on detergent removal. The abrupt increase in light scattering in the dilution experiment (see Figure 2a) is inferred to arise from the formation of these worm-like structures, which are thought to be important intermediates in the formation of 2-D crystals. Crystallization by the dilution method requires a significant dilution of the protein and, therefore, rather high initial protein concentrations. On the other hand, the dilution method allows the process to be arrested when the saturation point is reached, extending the time in which an ordered assembly of the components can take place. In addition, the dilution method is suitable for 249

PHILIPPE RINGLER

Figure 2 Protein reconstitution and formation of lipidic structures of different monitored using dynamic light scattering upon dilution. (a) On dilution of the mixed micellar suspension containing a detergent (octyl polyoxyeUnylene) and either lipid (phosphatidylcholine from egg yolk (egg PC)) or lipid and a membrane protein (pofin OmpF), at least two populations of structures with different sizes are formed during the transition from micelles to bilayer (the grey areas indicate suspected size ranges). In the case of only egg PC being present (continuous line), further dilution results in a homogeneous population of vesicles. With both egg PC and OmpF present (broken line), the two populations are still apparent on further dilution, with the smaller structures being vesicles with Title incorporated protein and the large structures bilayer with densely packed and crystalline OmpF. (b) The data here are represented as a function of detergent concentration to illustrate the relationship with the 'three-stage' model, with large bilayer structures at low detergent concentration, small micelles at high concentration, and a mixture of structures at intermediate concentrations. {Data from ref. 78 with permission.)

250

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

Protocol 11 Tubular crystallization of photosyster.i-ll core complex (PSII) using dilutiona Equipment and reagents * 1 mg/ml purified PSII solution of chlorophylls in 50 mM Mes pH 6,10 mM NaCl. 0.4 M sucrose, 0,4% octyl-p-othioghicopyranoside (OTG)

• 10 mg/ml dimyristoylphosphatidyl choline (DMPC) in 1.3% OTG • Dilution buffer: 50 mM Hepes pH 7,5

Method 1. Mix the protein solution and the lipid solution to vary the lipid:protein ratio to between 0.1 and 1. 2. Dilute the resulting solutions with the dilution buffer over a 4-fold range and incubate at temperatures of between 6°C and 22°C. 3. Monitor the solutions by withdrawing a 3 u1 sample every 8 hours and examining by electron microscopy using negative staining.11 " Method given in ref. 68. ''Tubes appeared to be reproducible within these temperature/dilution ranges after a period of 1-5 days.

low CMC detergents whose concentration is brought close to The CMC by the use of Bio-Beads prior to the dilution experiment. The dilution method in Prolxol 11 yielded tubular crystals of photosystcm-II core complex suitable for structural analysis by negative stnin electron microscopy at medium resolution (20 A) (68). 2.2.7 Dialysis method Dialysis is the most widely used technique in 2-D crystallization trials, usually in the form of small sample compartments dialysed against large buffer volumes. To improve the reproducibility of crystallisation conditions, a temperaturecontrolled, continuous-flow dialysis apparatus was developed (79) (sec Hffjrc 3), The advantage of this system is a precise control of the temperature profile that was found to be quite critical in some cases (38, 65). Additionally, a maximal gradient of detergent concentration is maintained across the dialysis membrane which improves reproducibility. A drawback of the dialysis method is the long dialysis times needed to remove low CMC detergents, making it only practical for medium-to-high CMC detergents (typically CMC > 1 mM). Our in-house designed dialysis apparatus enables us to perform up to 30 different 2-D crystallization trials with 10 different dialysis buffers. This property is of interest if a large range of crystallization conditions needs to be screened. 251

PHILIPPE RINGLER ET AL.

Figure 3 Temperature-controlled, continuous open-flow dialysis apparatus. (a) The dialysis chamber (stainless steel} consists of two buffer reservoirs, the first being used to equilibrate the temperature of the dialysate. A planar dialysis membrane (arrow) separating the sample chamber (6 mm diameter, typically loaded with 100 u1 sample, and manufactured from Teflon) from the second buffer reservoir allows the passage of soluble detergent molecules and salts. Sample chambers (arrow) are sealed with a transparent plastic membrane fixed with an aluminium plate and thermally isolated by a glass plate (arrow) that permits observation of sample turbidity. Extraction of aliquots for electron microscopy during the dialysis experiment simply requires dismounting the glass plate. The Peltier element regulates the temperature of the dialysis chamber, excess heat being eliminated by water-cooling. A pre-sei sequence of linear tempecature gradients is executed by a microprocessor that also controls the peristaltic pump responsible for continuous-flow dialysis. The system allows many critical parameters such as temperature, flow rate, pH, and ionic strength to be properly controlled, (b) Photograph of the temperature-controlled, continuous open-flow dialysis apparatus showing 30 sample chambers (only 6 chambers in front are closed and ready to be used).

As an alternative to the dialysis apparatus shown in Figure 3, an inexpensive3 microdialysis arrangement with Eppendorf tubes (see figure 4 and Protocol )3) or dialysis buttons may be used. This method enables the dialysis of small volumes {< 50 ul) of protein-lipid-detergent. Another interesting microdialysis device using a bent glass capillary tube has been described (90). A 100 mm glass tube of 2,5 mm inner diameter and about 6.5 mm outer diameter is bent by 90° about 10 mm from the end. The end is flame polished into a smooth surface which forms a tight seal with the dialysis membrane'. The dialysis membrane is fixed with a ring of silicon tubing and 252

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Figure 4 Microdialysis with Eppendorf tubes, (a) The Eppendorf tube is cut near the top with a razor blade, (b) The dialysis sample is pipetted into the reservoir formed by the Eppendorf cap. (c) A 3 cm x 3 cm piece of wet dialysis membrane is placed symmetrically over the top of the cylindrical chamber, and the ring pressed down to seal the dialysis chamber (see arrow), (d) The assembly is then ready to be placed on the buffer surface where it will float with the dialysis membrane in contact with the buffer.

Protocol 12 Pre-treatment of the dialysis membranes Equipment and reagents • Dialysis tubing with molecular weight cutoff (MWCO) of 6000-8000 (Spectra/Por 1, Spectrum) • 50% ethanol

- 10 mM Na2C03,1 mM EDTA • 0.05% NaN3 in distilled water (w/v)

Method 1. Add 50% ethanol and the dialysis tubing to a 2 litre beaker. Place a 1 litre beaker full of water inside the 2 litre beaker to push down the tubing into the alcohol. Boil the alcohol for 1 hour. 2. Rinse and squeeze the tubing with distilled water several times. 3. Boil the tubing again in 10 mM Na2CO3. 1 mM EDTA for 1 h, 4. Rinse with distilled water as before. 5. Boil the tubing in distilled water for 1 h. 6. Store in cold distilled water with 0.05% NaN3 at 4°C.

20-50 u1 dialysatc is fed into the capillary from the open end using a syringe. The dialysatc is shaken down against the dialysis membrane to remove air bubbles and the device is then placed in a glass beaker containing dialysis buffer. One interesting advantage of this device is that a sample for electron microscopy can be taken at any time using a fine glass capillary without disturbing the progress of the experiment.

PHILIPPE RINGLER ET AL.

Protocol 13 Microdlalysis set-up using Eppendorf tubes Equipment and reagents • Pre-treated dialysis tubing (see Protocol 12) • Razorblade

• Eppendorf tubes of volume 200 ul 500 ul. or 1.7 ml

Method 1. Cut off a portion near the top (2 mm) of the Eppendorf tube with the razor blade (see figure 4a). 2. Pipette the sample to be dialysed (15-150 ul) into the cylindrical chamber (see Figure 4b) 3. Cut a 3 cm X 3 cm piece of wet dialysis membrane, shake to remove excess storage solution and place symmetrically over the top of the cylindrical chamber (see Figure 4c) and then snap the ring to seal the dialysis chamber (see Figure 4d)." 4. Shake down the sample solution on to the dialysis membrane and let the dialysis assembly float on a large volume of dialysis buffer (dialysis membrane facing the buffer), 5. After incubation, puncture the dialysis membrane and harvest the sample by micropipetting," "Care should be taken not to allow the membrane to wrinkle when closing the ring as this can generate leaks. "Once punctured the dialysis assembly cannot be used again, therefore a new set-up is required for further dialysis of the sample. 2.2.8 Bio-Beads method Adsorption of the detergents Lo polystyrene beads (Bio-Reads SM2, Bio-Rad) is a powerful alternative to the dilution and dialysis methods for obtaining 2-1) crystals of integral membrane proteins (80). Hydrophobic adsorption of detergents on to polystyrene beads allows detergent to be removed independent of the respective CMC and has been used to generate highly ordered 2-D crystals of membrane proteins (e.g. crystallization of the rvtochrome b6f complex by Mosser el at (42)). Bio-Beads can be added to very small sample volumes with almost no dilution of the protein or lipids. This property is of great interest for membrane proteins for which purifying large quantities is particularly lime consuming. However, it should be noted that handling small quantities of BioBeads is difficult, because the beads must not dry in the procedure. According to Rignud's studies, the non-specific adsorption of lipids is about 100-200 times lower than the specific adsorption of detergents (2-4 mg phospholipids/g BioHeads SM2). Since crystallization experiments Lire generally performed at low lipid to protein ratios ((.PR ^ 0.2:1 to 1:1), weak lipid adsorption may have some 254

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Table 3 Adsorption capacities of Bio-Beads SM2 Adsorption (mg/g wet beads)

Capacity (mmoles/g wet beads)

n-Octyl-fJ-D-glucopyranoside

125 ± 10

0.45 ± 0.03

HECAMEG

110 ± 10

0.33 ± 0.03

C12E8

190 ± 10

0.35 ± 0.03

n-Dodecyl-fi-D-maltopyranoside

100 ± 10

0.20 ± 0.02

200 ± 10

0.31 ± 0.03

1-2 2-5

0.001-0.0025 0.0025-0.006

Detergent

(DDM) Triton X-100 Phosphollpid Liposomes Micelles Protein Cytochrome b6f

0.1

Melibiose permease

0.2

Ca2+-ATPase

0.2

Bacteriorhodopsin

0.0

hTATPase(CFOF1)

0.0

effects. However, lipid adsorption can be reduced by pre-incubating the beads with an excess of sonicated liposomes prior to their use for detergent removal. The adsorption capacities of various detergents to the Bio-Beads SM2 are summarized in Table 3. 2-D crystallization trials can be performed using either a one-step addition of Bio-Beads, resulting in a fast removal of the detergent, or by addition of the same Bio-Beads mass in several steps to slow down the process. The rate of detergent removal is not only directly linked to the weight of BioBeads used but also to the working temperature. The rate of adsorption of detergents doubles every 15°C. To maintain a reproducible adsorption property, the freshly blotted Bio-Beads must be precisely weighed and must not be allowed to dry out.

2.2.9 Factors affecting crystal quality An important parameter affecting the quality of crystals obtained by detergent removal is the temperature. In several cases a higher temperature (up to 40 °C) improved the crystal quality of proteins already reconstituted into lipid bilayers (see Table 1). The conclusion is that mild heat treatment promotes reordering of the molecules into more regular arrays. Temperature is also an important consideration in the storage of 2-D crystals. Some crystals degrade slowly and need to be stored at 4°C, while others appear to be reasonably stable even at room temperature. The presence of residual 255

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Protocol 14 2-D crystallization procedure using Bio-Beads SM2 Equipment and reagents • Bio-Beads SM2 (Bio-Rad)

• Buffer

• Methanol

Method 1. Prior to use, briefly wash the beads first with methanol then with the buffer intended for crystallization (either in a batch procedure or packed in a small plastic column). Store washed beads, if required, at 4°C as a 50% slurry in water supplemented with 0,2% azide. 2. Pipette the approximate quantity of Bio-Beads' slurry needed. Remove excess solution by blotting briefly with filter paper and weigh the precise quantity of beads.9 3. Add the wet Bio-Beads directly to the protein/Hpid/detergent solutions and place the samples on a rotating device (10 r.p.m,) or agitate slowly with a very small magnetic stirrer. 4. At various times, stop stirring (dense Bio-Beads sediment quickly) and take small aliquots from the supernatant to check the remaining detergent concentration using the methods cited above (see Protocols 5 and 6). 5. With same procedure used in step 4, check the formation of vesicles and the growth of 2-D crystals by electron microscopy (see Section 3). " Dry beads, which, appear whiter than the wet beads and do not stick together, must be removed.

detergent has been, observed Lo decrease crystal quality, and complete detergent removal after crystallization using Bio-Beads is advised in such cases. The use of excessive lipid (i.e. high LPR) may result in poor crystals. However. the excess lipids may be necessary to ensure that the native conformation of the protein is maintained. In such cases the excess lipids can be removed after crystallization using phospholipase A2 [Protocol 15) (91-94).

2.3 Crystallization of purified membrane protein by precipitation: a procedure that is similar to 3-D crystallization methods In a few cases, a solubilized membrane protein has been crystallized into 2-1) sheets without removal of detergent, under conditions similar to those used in 3-D crystallization experiments. Bacteriorhodopsin solubilized with Triton X100 was found to crystallize into orthorhombic lattices when incubated in a solution containing positively charged detergent at pH 4-6 and at a temperature of 26-27 "C (95). 256

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Protocol 15 Phospholipase A2 treatment of 2-D crystals Equipment and reagents • Dialysis buffer: 10mM Tris-HCl pH7.5,5 mM CaCl2,5 mM MgCl2 • Dialysis tubing and device (see Protocols 12 and 13}

• 1200 U/ml bee-venom phospholipase A2 (Sigma) in dialysis buffed

Method 1. Add phospholipase A2 to the 2-D crystal solution at a final concentration of 0.1-0.5 U/mg of membrane protein.'1 2. Pour the mixture into a dialysis device and dialyse for 18-24 h at 4°C against a large volume.1 3. Pellet the 2-D crystals by centrifugation at 1000 g for 10 min and resuspend the pellets in a suitable buffer for observation in transmission electron microscopy (TEM) by negative staining (see Protocol 17). " One unit will hydrolyse 1.0 umole of t-a-phosphatidylcholine to L-a-lysophosphatidykholine and fatty acid per min at pH 8,9 at 25°C using soybean L-w-phosphatidylcholine}, 'The slow rate of phospholipase reaction may facilitate ordering of the protein components in the membrane plane by the gradual removal of lipids. 'The released lysophospholipids and fatty acids do not reach their critical micelle concentration under the constant extensive dialysis and therefore do not disrupt the 2-D crystals. Bio-Beads (see Section 2.2.8) may be added to the reaction mixture to ensure elimination of lysolipids.

The pea thylakoid light-harvesting complex II (LHCII) was resolved to atomic resolution after formation of 2-D crystals in a so-called batch method (56, 96), As with bacteriorhodopsin, the temperature profile proved to be critical for the crystallization of LHC1I (96), Recently, Cyrklaff et al. (48) have shown that 2-D crystals can also be grown on the surface of .1 drop from detergenl-sohibilized and purified Neurospont crassa plasma membrane H '-ATPase. These methods are best interpreted as variations of 3-D crystallization, and not as the proper reconstitution of membrane proteins into a native-like environment, although the crystals formed appear to contain a significant amount of native lipids.

3 Analysis of the result of 2-D crystallization by electron microscopy 3.1 Negative staining Even if large membrane protein 2-D crystals can sometimes be observed by light microscopy, the shape of the crystals and their degree of order can best be 257

PHILIPPE RINGLER ET AL.

assessed by electron microscopy. Thus, access to an electron microscope is needed for screening 2-D crystallization experiments. The quickest method for preparing specimens for screening 2-D crystals is that of negative staining. Introduced by Brenner and Home in 1959 (97), this method is rapid, needs a small amount of sample (less than 5 ul), is remarkably simple, and can provide structural information to a resolution of about 20 A or even better (for a review see ref. 98). A negatively stained 2-D crystal is embedded in a dry, microcrystalline, heavy-atom replica. As heavy atoms (U, W, Au, Pt, Pb, and Os) used for negative staining scatter electrons much more than the biological atoms (C, H, O, N, P, and S), the contrast is drastically increased and also inverted (hence 'negative' staining). In addition, the heavy-atom salts partially substitute the water in the native environment of the molecules, thus embedding the specimen and protecting it from collapse upon drying in air. Furthermore, negative stain salts are more tolerant to electron irradiation than the biological material. Most commonly used heavy-atom salts are uranyl acetate or formate, and sodium or potassium phosphotungstate. Uranyl salts are more suitable for more proteinic samples, whereas tungstate is useful for lipid structures. The pH of tungstate solutions can be adjusted, whereas uranyl salts precipitate at pH >4-5.

3.1.1 Preparation of electron microscopy grids Copper grids (e.g. from Electron Microscopy Sciences) are the most commonly used specimen support grids for electron microscopy. Typically 200-400 mesh/ in. copper grids, 3.05 mm in diameter and 0.7 mm thick are coated with a specimen support film (mostly either a carbon/parlodion composite film or a thin carbon film). Freshly prepared grids are known to be hydrophilic, or they can be rendered hydrophilic by glow-discharge (see below). If a hydrophobic carbon surface is required for the uptake of 2-D crystals formed on lipid monolayers, carbon-coated grids should be aged for 2 weeks in a dry environment before use.

3.1.2 Negative staining of 2-D crystals To obtain optimal staining and to enhance the adsorption of the sample to the carbon surface of the specimen grids, glow-discharging of these grids under reduced air pressure prior to adsorption of the specimens is highly recommended (99). This procedure renders the grids hydrophilic by ionizing the carbon surface. The optimal stain for a particular specimen has to be chosen and evaluated by trial and error. The ionic conditions of the sample buffer and the presence of remaining detergents, while strongly interfering with specimen adsorption, may have little effect on the quality of the negative stain replica surrounding the biomolecules, as long as a sufficient number of washing steps (two to six) are employed prior to applying the negative stain solution to the specimen. 258

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Protocol 16 Preparation of carbon-parlodlon composite films on copper grids Equipment and reagents • Electron microscopy copper grids 200-400 mesh/in (Electron Microscopy Science) • 2% parlodion in n-butylacetate (Electron Microscopy Science)

Hydrophilk paper (usually recycled paper) Carbon sheet (50 nm thick) Graphite rods Vacuum evaporator

Method 1. Prepare a large (20 cm diameter) water surface on a round Petri dish, and spread a relatively large drop (100 ul) of the parlodion solution. 2. Place around 50 copper grids on to the surface of the flat floating parlodion surface (with the darker copper grid surface facing the parlodion), 3. Completely cover all the grids with a hydrophilic paper (usually recycled paper) and wait until the entire paper surface is soaked. 4. Remove the piece of paper with attached grids and parlodion and let it dry on the bench for 1 h. 5. Evaporate a uniform sheet of carbon (50 nm thick) using a vacuum evaporator at 7.10-5 mbar and a graphite contact point heated by 30 V, 35 A for 6 sec.

Protocol 17 Negative staining Equipment and reagents • Precision forceps (Electron Microscopy Science) • 0.75% uranyl formate (Eastman Kodak Co.) in water

Filter paper (drying block from Schleicher and Schuell, cat. no. 556)

Method 1. Deposit the carbon/parlodion-coated grids (dark, carbon side facing the top) on a glass block covered with ParafUm into a glow-discharge unit and glow-discharge for 15 sec," 2. Hold the specimen grid horizontally with precision forceps and deposit 1-5 ^1 of the specimen solution to the carbon-coated side of the grid. Let it adsorb for 30-60 sec.

259

PHILIPPE RINGLER ET AL Protocol 17 conti

3. Blot off the excess liquid by touching the grid surface with a filter paper, wash the specimen grid on a drop of distilled water for 2 sec and blot again. Repeat this washing once more, 4. Stain the specimen by lowering the specimen grid on to a drop of negative stain solution (see below) for 20 sec, briefly blot the specimen as described above and repeat this step once again. Finally, blot thoroughly by holding the grid in contact with the filter paper. The preparation is then ready for inspection in the electron microscope. " After glow-discharging, grids should be used within the next 30 min.

Figure 5 TEM analysis of 2-D crystallization trials by negative staining, (a) Low magnification micrograph (2500 X) showing large sheets and vesicles of reconstituted aquaporin from f. coli (AqpZ) [scale bar represents 2 um). (b) Typical micrograph of negatively stained twodimensional crystalline arrays of AqpZ that are suitable for image processing. A rectangular lattice can be seen when observed at glancing angle (scale bar represents 100 nm). (c) This micrograph displays membrane proteins and lipids that were solubilized with the detergent oetylglucoside. Mixed detergent-lipid micelles appear as small dots, while some Iipid-protein-detergent structures have elongated (worm-like) shapes (< 10 nm, scale bar represents 100 nm). (d) Partially dialysed sample often exhibit stacked multilamellar lipid structures (scale bar represents 100 nm). (e) Lipid vesicles (20-50 nm diameter) form when no membrane protein was incorporated (empty vesicles). Such structures appear when the LPR is too high, or when the protein aggregates (scale bar represents 100 nm). 260

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

3.2 Analysis of the results from 2-D crystallization trials After negative staining, examination of the samples is carried out by transmission electron microscopy. The presence of large vesicles or sheets can be checked at low magnification (2500 x) because of the high contrast obtained when operating at large underfocus even for very thin objects (see Figure 5a). Further observations are carried out at 50000 x magnification where the lattices can often be seen directly (Figure 5b), and micrographs can be taken for structural analysis (see Section 3.3). If no crystals are found, the presence of single protein particles solubilized in remaining detergents, protein aggregates in multilamellar lipid structures, or lipid vesicles can all be identified (Figure 5c, d, e). These pieces of information may be valuable for re-designing the crystallization trials. Electron micrographs of 2-D crystals are examined by optical diffraction with a laser diffractometer (see Figure 6) to assess the quality of the crystals and the adjustment of the microscope (100). Optical diffraction allows observation of the concentric rings resulting from the contrast transfer function (CTF) of the microscope (see Figure 7a). The shape and quality of the CTF reveals the coherence of the beam, the defocus used, the astigmatism, and the drift (Figure To, c, d, e). With negative staining, the resolution achieved is usually around 20 A. Therefore, if the selected micrograph shows that the first zero of the CTF falls beyond 15 A, no further CTF correction will be needed for processing the image.

Figure 6 Design of an optical diffractometer (according to Aebi et a/. (100)). The folded optical path allows the diffractometer to be placed on a small bench and provides for convenient manipulation of the micrographs with simultaneous observation of the diffraction pattern. (1) He-Ne laser (wavelength 594 nm); (2) high-power microscope objective; (3) 5 Um diameter aperture (pinhole); (4) mirrors; (5) spherically corrected achromatic lens (diameter 8 cm, focal distance 36 cm); (6) aperture for selecting optimum area of micrograph; (7) film holder with x, y table. Displacement along the optical axis allows the size of the diffraction pattern in plane (8) to be adjusted; (9) observation telescope.

261

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Figure 7 The optical (or likewise the calculated} diffraction pattern reveals both the quality of the crystal and the contrast transfer function (CTF) of the electron microscope. The latter depends on the adjustment of the microscope and the stage. For a stigmatic image, concentric rings are seen (Thon's rings), their density and succession depending on the focus, (a) Ideal underfocus (close to the focus): the essential diffraction spots are all contained within the first circular zone, (b) Too much underfocus; spots are lost as they overlap with the first zero (dark circular zone) of the CTF. Spots in the next bright ring are phase shifted by IT (containing structural information with inverted contrast), (c) Astigmatism invokes a diffraction pattern with non-circular zero ;ones of the CTF. (d) Diffraction pattern showing hyperbolic-shaped CTF when astigmatism occurs close to the focus, (e) A linear drift (resulting from transiation of the specimen during the exposure time) induces a banding pattern in the diffractogram that is perpendicular to the drift direction. High-resolution details are smeared out along the drift direction and cannot be fully recovered.

Optical diffraction patterns of 2-D crystals show the crystal parameters, such as dimensions of the unit cell and tryst allographic symmetry. Furthermore, the presence of stacked 2-D crystals is revealed by two or more sets of diffraction spots from the same area, Areas of high crystallinity are marked and subsequently digitized on a microdensitometer. Typically, at 50000 x magnification and scanning the micrographs at steps of 20 u.m (4 A/pixel), 1024 x 1024 pixel images suffice for digital image processing of micrographs from negatively stained samples. 262

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

3.3 Image analysis of 2-D crystals: correlation averaging (Figure.' 8 shows a flow chart for obtaining an averaged picture using the image processing package SLMPER (101). (For an introduction to standard techniques of image processing sec ref, 102, and for the correlation averaging see ref, 103,] Correlation averages from areas containing 2-D crystals often indicate some key features of rhe membrane protein, such as shape, symmetry, and approximate

Protocol 18 Correlation averaging of OmpC 2-D crystals Equipment • Computer running SEMPER 6, library programs and TIFF subroutines can be provided.

Method 1. Load the initial, raw-digitized (1024 x 1024 pixels, TIFF format, see figure 8a) picture and Fourier transform it via: tiff rea nam 'rawpicture.tiff' to 1; fou to 2 2. Produce and display the power spectrum (see Figure 8b) via: ps to 3;ful;mas ins rad 10 val 0; sur;max=max/12 ;dis pre 3. Search and refine the lattice parameters U and V of this 2-D crystal via: lib lattice (point diffraction spots with cursor and indicate their lattice's indexes); pea 3 thr max/4 sra 30; w=0,0;bas uvo tol .1 4. Produce and display a Fourier filtered image (see Figures 10.8c,d) via: win; ima; fit sub; sur f u l ; dis 5. Extract a reference picture (see Figure 8e) via: xwi; ext 3 to 4 pos x,y ; ori res; mas rad 70 wid 10; dis 6. Process a cross-correlation between the picture (see Figure 8a) and the reference (see Figure 8e) and display a correlation map (see Figure 8f) via: xcf 4 wit 1; hp ove 30; dis 7. Search the cross-correlation peaks and select the ones fitting the best to the found lattice via: pea 4 thr max/4 sra 15; lib rec; xwi; w=x,y; lat mar dis; bas tol .1; str 999 to 998 8. Average the raw picture at positions selected by the cross-correlation and produce a first projection map (see Figure 8g) via: mot 1 to 4 siz 128 fp wit 998; fit sub; sur ful; dis tim 4 neg 9. Extract a reference from the averaged picture (see Figure 8f} via: ext siz 1024; mas rad 50 wid 5; dis 10. Cross-correlate again by iteratively doing steps 6, 7, 8, and 9 (see dashed arrow of Figure 8) until the average does not improve anymore. 11. Export the final averaged picture as a TIFF file via: sca byt; tiff wri nam 'average.tiff

263

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Rgura 8 Processing an electron micrograph. After digitization of the micrograph into a field of 0.4 x 0.4 ums containing 1024 x 1024 pixels (a), its Fourier transform (FT) and the corresponding power spectrum (or diffraction pattern (b) is calculated, (c} The distinct diffraction spots are indexed on to a reciprocal lattice which locates small windows that allow just the information in the diffraction spots to pass, (d) After calculation of the inverse Fourier transform from the windowed FT, the filtered image is obtained, \e) The best (clearest, most contrast) area of the filtered image is selected as reference, (f) The cross-correlation function between the raw micrograph (a) and reference {e) exhibits sharp peaks that indicate where raw data and reference are aliKe, (g) Extracting all patches centred at the correlation peaks and averaging them results in a clean map that can serve as a reference for a refinement cycle (scale bar represents 60 A). If lattices are bent or exhibit other defects, the correlation peaks can be used to eliminate bad patches (see, for an example, ref, 103)),

264

TWO-DIMENSIONAL CRYSTALLIZATION OF MEMBRANE PROTEINS

size. The example in Protocol 18 demonstrates the steps of correlation averaging as applied to processing 2-D crystals of the outer membrane protein OmpC from £. coli. The diffractogram of the raw picture is calculated by the fast Fourier transformation algorithm and by squaring the transform to determine the lattice parameters. These are used, first, to produce a Fourier peak-filter version of the 2-D crystal picture, a step that removes most of the non-periodic noise. The most distinct area on this filtered picture is selected and used as a reference to search similar features in the raw picture using the cross-correlation function. The cross-correlation peak positions which fit approximately to the lattice are used to extract image patches that are subsequently averaged to produce a first projection map. This projection map can then be used as a reference for a refinement cycle. This method provides projection maps of good quality and is particularly suited for beginners in the field of image processing. Further image processing steps can be achieved by averaging the structure factors from several images and taking advantage of symmetry relationships.

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Chapter 11 Crystallization of membrane proteins TINA D. HOWARD, KATHERINE E. MCAULEY-HECHT, and RICHARD J. COGDELL Division of Biochemistry and Molecular Biology, Insitute of Biomedical and Life Sciences, University of Glasgow, Glasgow G12 8QQ

1 Introduction As yet, unfortunately, there are no high-resolution structures for any membrane transporters. This probably represents the major barrier to our understanding the precise molecular details of how this important class of membrane proteins actually work. An excellent example of the urgent need for structural information comes from the extensive studies on the lac-permease from Escherichia coli (1). Almost every amino acid in this transporter has been changed by sitedirected mutagenesis (H. R. Kaback, personal communication) and yet the exact details of how it works are still a mystery. It has been argued that the crystal structure of a transporter may not be such a 'holy grail', since it will only give a static picture and transport is, by definition, a dynamic process. However, the recent success in understanding how ATP is made by examining the structure of the mitochondrial F1-ATP synthetase (2, 3) illustrates how, when a structure can be 'caught' in several functional states, the dynamics of the reaction can be inferred. Since for many transporters there are specific inhibitors which can 'lock' the protein in different functional conformations, there is an excellent chance that when the structure of a transporter is determined it will be possible to deduce the mechanism. There are currently three possible methods with sufficient power to determine the high-resolution 3-D structure of a protein, NMR (4), X-ray crystallography using 3-D crystals (5), and electron crystallography using 2-D crystals (see Chapter 10) (6). In this chapter we will discuss the approach using 3-D crystals. We will describe how to tackle the problem of crystallizing membrane proteins, which is based upon our success in obtaining high-resolution crystal structures of two integral membrane proteins from purple photosynthetic bacteria, i.e. reaction centres and light-harvesting complexes (7-10). X-ray crystallography has now been used to successfully determine the structure of several thousand water-soluble proteins. However, even though 269

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membrane proteins are thought to represent about 40% of all proteins, only a handful have so far had their structure determined to high resolution (7-31). Why is this? First of all it is still widely believed that it is either impossible to crystallize membrane proteins or, at very best, horribly difficult. This immediately stops many people from even trying to tackle the problem. This belief is not justified. By far the biggest hurdle is getting enough (i.e. regular milligram batches) fully native protein to work with. Many membrane proteins, including transporters, are only present in low abundance. A reliable, robust system for overexpressing membrane proteins in a native state is urgently needed (see Chapters 4-6) (32, 33). It has been our experience that once you get enough protein which is sufficiently pure and active, then the process of crystallization is not very much more difficult than with water-soluble proteins. In general, proteins crystallize when they are induced to precipitate slowly. This is usually achieved by the equilibration with precipitants, e.g. ammonium sulfate or polyethylene glycol. The major difference between working with membrane proteins and water-soluble ones is that you have to account for the fact that membrane proteins are solubilized, i.e. they are embedded in a detergent micelle. The physicochemical properties of the detergent as well as the protein must therefore be considered. In many cases with membrane proteins at concentrations of the precipitant well below that required to precipitate the protein, there is a reaction with the detergent. Very often the precipitant causes the detergent to 'phase separate'. Usually when this happens the membrane protein denatures in the 'oily' detergent phase and the potential conditions for crystallization are never achieved. Several years ago, working independently, Michel (34, 35) and Garavito (36, 37) realized the nature of this problem. They were able to demonstrate that some specific small molecules, e.g. heptane-1,2,3triol (HPTO) could be added to the crystallization mixture in such a way that the 'phase separation' point could be shifted to be above the precipitation point. Now they were able to access the potential range of concentrations of the precipitants where crystallization was possible. In both cases they were able to achieve crystallization; Michel with purple bacterial reaction centres (38) and Garavito with porin (39).

2 Crystallization techniques 2.1 General crystallization techniques The methods used to crystallize a membrane protein are the same as those used for the crystallization of soluble proteins, i.e. vapour diffusion, microdialysis or batch techniques (40-43). Vapour diffusion is the most commonly used technique and it has proved to be very successful in the crystallization of both soluble and membrane proteins (44). Dialysis, too, has been used to crystallize several membrane proteins, e.g. the porins (39) and the photosynthetic reaction centre (45). The membrane protein complex, chlorophyll binding protein CP43, has been crystallized using a microbatch method, where the crystals were grown 270

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under oil (46). This method may also be useful in screening trials as it allows very small sample volumes (e.g. 2 ul) to be used (47).

2.1.1 Vapour diffusion A typical sitting-drop, vapour diffusion set-up is shown in Figure 1a. A small volume of the protein solution is added to the central well and the reservoir is filled with a larger volume (typically 1 ml) of a precipitant solution. The protein solution also contains precipitant but at a concentration lower than that of the reservoir and below the concentration required for the growth of crystals. The vessel is sealed and incubated at a constant temperature. Slowly the two solutions begin to equilibrate via the vapour phase. This results in a slow concentration of the protein solution which will, hopefully, induce crystallization. The rate of equilibration can affect the presence and the quality of crystals obtained and several techniques to change the kinetics of equilibrium exist. These include varying the precipitant concentrations in both the droplet and the reservoir solutions, increasing or decreasing the volume of the droplet, altering the concentration of the protein, and performing the experiment at different temperatures. Vapour diffusion can also be carried out using a 'hanging drop' procedure, where the protein droplet is suspended above the reservoir, on the underside of a coverslip. However, this method is limited to small droplet volumes, especially when using solutions containing detergent due to the decrease in surface tension of the solution and the resulting difficulties in preventing the droplet from spreading on the coverslip.

2.1.2 Microdialysis Microdialysis achieves the same supersaturation of the protein solution as vapour diffusion, but by restricting the passage of higher molecular weight solutes through a dialysis membrane. In this case the protein solution is slowly equilibrated against a reservoir solution containing a higher concentration of, for example, polyethylene glycol (PEG) across the dialysis membrane. The diffusion of molecules is dependent on the molecular weight cut-off (MWCO) of the dialysis membrane and the molecular weight of the particular PEG employed; the loss of water through the membrane and/or the diffusion of PEG into the proteinaceous well solution results in an increase in the precipitant concentration in the protein solution. Earlier crystallization experiments used a range of 'home-made' set-ups (see 42, 48 and references therein). However, the procedure is now made considerably easier with the commercial availability of microdialysis buttons from either Cambridge Repetition Engineers or Hampton Research. Buttons; with the volume of the wells ranging from 5 ul up to 350 ul being readily obtainable, this allows the crystallization procedure to be carried out on various scales. Figure 1b shows dialysis buttons (5 ul, 10 ul, and 20 ul volumes) along with the O-ring used to hold the dialysis membrane in place and two types of applicator used in assembling the unit. The set-up of the microdialysis button also provides 271

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Figure 1 Typical apparatus used for crystallizations, (a) A sitting-drop, vapour diffusion plate. Each plate can hold 24 crystallization trials. The protein droplet is held in a depression in the central post and this is surrounded by the reservoir. The plate is sealed using clear tape. (b) Microdialysis buttons. The volumes of tile buttons are. from right to left, 5 ul. 10 ul. and 20 ul. Also shown is the 0-nng that holds the dialysis membrane in place and two 0-ring applicator tools.

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scope for changing the reservoir solutions, permitting further variations of conditions if crystals do not appear. Use of the vapour diffusion technique is generally more convenient than microdialysis since it is easier to survey the crystallization trials. Typical vapour diffusion crystallization trays can be placed directly under a microscope in order to view their progress. On the other hand, viewing microdialysis trials is more time consuming and may involve interfering with components within the system, since the dialysis membrane may have to be removed to monitor the progress of the crystallization. Dialysis lends itself to use of the 'salting in' phenomenon, where the solubility of a protein can decrease with a decrease in ionic strength (42)—frequently observed near the isoelectric point of a protein. The protein of interest is retained by a low MWCO dialysis membrane and the salt concentration of the reservoir solution is progressively altered; as the solubility of the protein decreases crystallization can occur (43).

2.2 Further techniques designed specifically for membrane proteins The general crystallization methods discussed above are only applicable when the membrane protein has been sufficiently solubilized and exists as a homogeneous solution of protein-detergent micelles, providing unfavourable phase separations can be avoided (49). Many membrane proteins are unstable in detergent solutions, although often they can be reconstituted in a phospholipid bilayer, regaining their stability and activity. 2.2.1 Lipidic cubic phases Recently, Landau and Rosenbusch (50, 51) devised a new strategy for the production of 3-D crystals of membrane proteins, by reconstituting them into phospholipids which were manipulated into the form of quasi-solid lipidic cubic phases (52). When certain lipids, water, and protein are mixed in appropriate proportions the resulting membrane system forms transparent, three-dimensional structures, which contain interconnecting water channels. These lipid matrices can provide nucleation sites and support crystal growth by promoting lateral diffusion of the protein in the membrane. This method has been successfully used to crystallize bacteriorhodopsin (23), light-harvesting complexes, and reaction centres (P. Nollert, personal communication).

3 Case studies This section is based on our experiences of crystallizing membrane proteins, namely the photosynthetic reaction centre and antenna complexes from purple photosynthetic bacteria, although other examples from the literature are also considered. These case studies demonstrate the factors we have found to be important in the crystallization process. 273

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3.1 Rhodopseudomonas acidophila 10050 LH2 Papiz et al. obtained X-ray diffraction from two crystal forms of Rps. acidophila 10050 LH2 (53). Rhombohedral crystals (space group R32) produced with B-octyl glucoside (p-OG) present as the detergent diffracted to a maximum resolution of 3.5 A. However, only a small fraction of the crystals produced diffracted to this resolution. Investigations into the effects of various amphiphilic compounds produced several different crystal forms (54), although the rhombohedral form from (3-OG and benzamidine hydrochloride remained the most promising. Crystals diffracting reliably to 3.5 A were produced with several small modifications to the purification procedure, e.g. using higher grade (Analar, Fluka) lauryldimethylamine oxide (LDAO) throughout the purification, rejecting more fractions from each of the columns, and ceasing to adjust the pH of the protein solution immediately prior to crystallization trials (55). The major step taken to achieve a structure with a resolution of 2.5 A (8, 9) was to omit the ion-exchange steps from the procedure; using instead a sucrose gradient step to separate the reaction centres and most importantly, an ultrafiltration step to exchange the detergent. Solubilization was also carried out for longer at a lower temperature (4°C). 3.1.1 Purification The purity of the protein is a vital factor in the crystallization process (42, 56, 57). There is little point in carrying out numerous crystallization screens with an impure protein sample. The failure to produce crystals or the production of poorly diffracting crystals can often be the result of impurities and structural inhomogeneities within the protein sample. The LH2 protein complex from Rps. acidophila 10050 can easily be purified for crystallization trials in high yields, because it is by far the most abundant protein in the membrane of this purple bacterium. The purification involves four basic steps—cell breakage, solubilization, separation on a sucrose density gradient, and finally gel filtration chromatography (Protocol 1, adapted from ref. 9). Within the LH2 protein complex, 27 bacteriochlorophyll a (Bchl a) molecules exist in two distinct environments producing two characteristic absorption bands in the near infrared, at 800 nm (18 Bchl a) and at 858 nm (9 Bchl a). The presence of these two signals can be used to monitor the structural integrity of the protein during the purification procedure. The deep-red colour of the complex (due to carotenoid absorption bands) provides a simple way to visually follow the progress of the protein through the purification process. 3.1.2 Crystallization The detergent used in the solubilization and purification of a protein is often not the best one for crystallization (see Section 3.4.2). For the crystallization of Rps. acidophila 10050 LH2, LDAO used in the solubilization procedure is frequently exchanged for (3-OG (or other detergents), using ultrafiltration by centrifugation. Any additional additives necessary for crystallization, e.g. salts, can also be introduced during this exchange procedure. 274

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Protocol 1 Purification of Rps. acldophlla 10050 LH2 Equipment and reagents • Mechanical French Press {e.g. Aminco)

• DNase

• L7 ultracentrinige (Beckman)

• MgCl2 • 30% (w/v)LDAO solution (Fmka)

• Ti 70 ultracentriruge rotor (Beckman) • Polycarbonate ultrabottles (Nalgene, 25 x 89 mm from Beckmanf

• Tris buffer: 20 mM Tris-HCl pH 8.0

• Homogenizer, ~ 30-50 ml capacity

• 0.8 M sucrose in TL buffer

• Centricons (MWCO 50 kDa; Amicon)

• 0.6 M sucrose in TL buffer

• Superdex-200 column (16 X 650 mm; Pharmacia) connected to an FPLC system

• 0.4 M sucrose in TL buffer

• Rps. acidophila 10050 cells (freshly harvested or defrosted from frozen)

• TL buffer: 0.1% (w/v)U3AO in Tris buffer

• 0.2 M sucrose in TL buffer

Method 1. Pass the cells through a French press (950 p.s.i.), twice, in Tris buffer with a little DNase and MgCl2, Centrifuge at 180 000 g for 1 h at 4°C to pellet the membranes, leaving the soluble proteins in the supernatant. 2. Resuspend the pellet in a minimum volume of Tris buffer and homogenize the suspension. Measure the ODg5g of the sample and adjust to 50 cm-1 with Tris buffer. 3. Add sufficient LDAO solution to produce a 2% (w/v) solution. Stir gently for 4 h at 4°C, Dilute the solubih'zed sample by a factor of — 4h and spin at 180 000 g for 90 min at 4°C. 4. During this spin, pour the discontinuous sucrose gradients in the ultracentriftige tubes; 5.3 ml of 0.8 M sucrose solution, 6 ml of 0.6 M sucrose solution. 6 ml of 0.4 M sucrose solution, and 6 ml of 0.2 M sucrose solution. 5. Carefully decant the supernatant from step 3 and apply this to the top of the sucrose density gradients and centrifuge for 12-18 hat 150000 gat 4°C. 6. Remove the tubes from the rotor and observe two dark-red bands within the gradient. The top band is the light-harvesting complex 2 (LH2), the bottom one is 'cote complex'/ Remove the desired protein from the gradient carefully using a Pasteur pipette. Measure the A8SS of the sample and concentrate to an AS58 of 50-75 cm-1 using a Centricon. 7. Apply 1-2 ml to a Superdex-200 column equilibrated in TL buffer at 4°C on an FPLC system.11 Elute the protein complex at ~0.6 ml/min, collecting 0.75 ml fractions.

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Protocol 1 continued

8. Record the absorption spectrum of the fractions between 250 and 950 nm.c Pool the fractions where the ratio of A85S to A-_ass is greater than 3 and concentrate the protein as required for the next step using a Centricon device. 0

Since it is necessary to completely fill the ultracentrifuge tubes (~ 28 ml)—for small sample volumes use a smaller rotor, e.g. Ti 70.1, and smaller capacity tubes, e.g. ~10 ml, to avoid excessively diluting the samples. b Dilute the sample to reduce the concentration of detergent and prevent any denaturation occurring. c The 'core complex' contains the reaction centre surrounded by light-harvesting complex 1 (LH1). The latter is thought to be a cyclic hexadecameric structure related to that of LH2 (58, 59). ''if no FPLC system is available, a peristaltic pump, column, and fraction collector will suffice. £ The purity of the complex is assessed by the ratio of the absorbance of the bacteriochlorophyll o peak at 858 nm to that of the general protein absorbance maximum around 265 nm.

Protocol 2 Detergent exchange by uttrafiltration in Rps. acidophlla 10050 LH2 Equipment and reagents • Rps. oridophilo 10050 LH2 (A^ = 100 cm-1)a in TL buffer (see Protocol 1} • Tris buffer (see Protocol 1) • TL buffer (see Protocol 1} • Centricon (MWCO 50 kDa; Amicon)

Buffer containing new detergent and/or salts, e.g. 32 mM NaCl, 1% B-OG (Boehringer-Mannheim (now Roche Molecular Biochemicals) in Tris buffer

Method 1. Place 300 ul of Rps. acidophila 10050 LH2 solution in the ultrafiltration device and dilute with 1.7 ml of Tris buffer. 2. Reduce the volume to ~ 100 ul by centrifuging at 5000 g for - 45 min at 4°C. 3. Add a further 2 ml of Tris buffer and again reduce the volume to -100 spinning at 5000 g for - 45 min at 4*C.h

^1 by

4. Add 1 ml of buffer containing the new detergent and reduce the volume to - 100 jU by spinning at 5000 g for — 25 min at 4°C.r 5. Place the retentate vial on to the Centricon, invert the device and spin at 2000 g for 2 min at 4'C.

276

CRYSTALLIZATION OF MEMBRANE PROTEINS

Protocol 2 continued 6. Remove the protein solution and make up the volume to 300 n-1 with buffer containing the new detergent, salts etc. " Equivalent to ~ 3.5 mg/ml— the near infrared absorption of LH2 provides a simple, nondestructive way of measuring protein concentration. h Generally, we have found the use of detergent-free Tris to be satisfactory, but to remove the final traces of LDAO the addition of 0.5 mM sodium dithionite into the final wash reduces the N-oxide bond of LDAO removing its ability to act as a detergent (54). r Add 1.9 ml of detergent-free Tris and repeat steps 2-4 if required, The integrity of the light-harvesting complex in the new detergent can be quickly assessed by monitoring changes in the absorption spectrum, both by eye and spcctrophoio metrically in the near infrared. Crystallization trials are prepared as soon as possible after purification and detergent exchange to minimize variations in, for example, temperature which may result in in homogeneities and dcnaturation within the sample (see Protocol 3). The trays are then incubated at the required temperature to await the appearance of protein crystals; some LH2 crystals are shown in I'igurcs 1 ].2A, H.

Protocol 3 Crystallization of Rps. acidophlla 10050 LH2 in p-OG Equipment and reagents For Part A • Tris buffer (see Protocol!) • Rps. acidophila 10050 LH2 (Agsg - 100 cm" in, for example, 32 mM NaCl, 1% p-OG (Boehrmger-Mannheim (now Roche Molecular Biochemicals) in Tris buffer • 10 mg benzamidine hydrochloride" (Aldrich) • 4 M di-potassium hydrogen phosphate • 2.1 M ammonium sulfate pH 9.35" • Crystal Clear Sealing Tape (Hampton Research) • Incubator at 16°C • Crystallization tray suitable for sittingdrop method (NBS Biologicals)

For Part B • As forftirtAwith the followingtwo exceptions • Rps. acidophila 10050 LH2 (AS58 = 100 cm -1 ) in 0.2% (w/v) LDAO (Fluka) in Tris buffer replaces that with p-OG • 4 M di-potassium hydrogen phosphate also contains 8% (v/v) glycerol

A. Crystallization of Rps. acidophlla 10050 LH2 in \.

Add 300 ul of Rps. addophUa 10050 LH2 solution to a microcentrifuge tube containing benzamidine hydrochloride and vottex briefly to dissolve (several very shoit bursts). Immediately add 100 |il 4 M K^HPC^ and invert the tube several times to mix,

277

TINA 0. HOWARD ET AL.

Protocol 3 continued

2. Centrifuge at 13000 g in a microcentrifuge for 10 min at 4°C. Whilst this is spinning add 1 ml of the (NH4)2SO4 solution to each reservoir of the crystallization tray. 3. Place 1.5 ul of the (NH4)2SO4 solution from each reservoir into the sitting-drop depression above each reservoir. Remove the sample from the microcentrifuge and carefully dispense 15 ul of the protein solution into each depression within the tray. 4. As soon as pipetting is completed, seal the tray with tape—pressing firmly around the edges of the wells.d 5. Incubate at 16°C. Note that the crystals are typically visible after 2-3 weeks, although it takes around 4-6 weeks for the procedure to go to completion. B, Crystallization of RPS. acldophila 10050 LH2 In LDAO 1. Follow Part A using the modified solutions listed above. " Pre-weighed in a microcentrifuge tube, equivalent to a final concentration of 2.5% (w/v). b Adjust the pH of the solution immediately prior to use, using 4 M NaOH. r Adjust the volumes and the ratio of volumes of the reservoir solution to the prepared protein solution as required (see Section 3.4.1)—maximum volume of a drop is ~ 50 ul. * To minimize the evaporation of solutions when using very small volumes, the wells may be sealed individually, or in blocks of 4 or 6.

The nystal structure of R. acidophiki 10050 LH2 was solved using crystals grown with p-OG (8, 9); .subsequently it has been discovered that crystals with similar diffraction properties could be grown from LDAO on a sensible time scale if a small amount of glycerol was included in the well (10). The order of addition of the components is important to prevent phase separation (see, for example, Hgun* 2D) and precipitation. This is discussed more thoroughly, with particular refere-ncc to the addition of amphiphiles, in Section 3.4.3.

3.2 Reaction centres The photosynthetic reaction centre (RC) from the purple bacterium Rhodopscudomontis viridts was the first integral membrane protein to have its 3-D structure determined by X-ray crystallography (60, 61). This achievement proved. at last, that it was possible To crystallize and determine the structure of membrane proteins. The RC from another species of purple bacteria, Rhodobacter sphacmidcs, has also been crystallized and both naturally occurring and genetically engineered mutants of this complex have been studied by X-ray crystallography (7, 31, 60-70). The first structures of the RC from Rh. sphacroides were of relatively low resolutions (3,0-2.8 A); although this was adequate to determine the overall 278

CRYSTALLIZATION OF MEMBRANE PROTEINS

structure of the complex, a higher resolution was required to look at the fine detail of the protein-pigment interactions. Over the years, the diffraction properties of RC crystals have improved dramatically, mainly by changing the crystallization conditions to find new crystal forms. The highest reported diffraction for RC crystals is now 1.9 A, an improvement of more than 1 A from the first structures!

Figure 2 (A) Rhombohedrai crystals of the membrane protein R. scidophilia 10050 LH2. (B) A tetragonal crystal form of R. acidophilia 1.0050 LH2. i'C} Trigonal crystals of the reaction centre from Rb. sphaemitfes. {D} An example of phase separation in the presence of a salt crystal; the dark droplets consist of coloured LH2 protein which is present in the detergent-rich phase.

279

TINA D. HOWARD ET AL.

3.2.1 Purification The first step in obtaining crystals of the reaction centre is to prepare a sufficient quantity of purified protein, and there arc many published procedures for purifying the RC from the chromatophores of Rb. spheroides (e.g. 71, 721. The method we have found To be most useful for purifying RCs from an RC-only strain of Kb. spheroides (73, 74) is given in Protocol 4. The purity of the RC sample can easily be measured by evaluating the absorbance ratio /U sr) : A soo and this should be close to 1.2 (no more than 1.4) for crystallization trials. This is a ratio of general protein absorption by aromatic amino acids at 279 nm, compared to RC-specific absorption at 800 nm from the bacteriochlorophyll pigment molecules. The purified protein may be used immediately tor crystallization, using LDAO as the detergent, or the detergent may he exchanged prior to crystallization using an ultrafiltration technique (sec, for example. Protocol 2).

Protocol 4 Purification of RC from purple photosynthetic bacteria Equipment and reagents • Tris buffer (see Protocol 1) • DNase • MgCl2

• Mechanical French Press (e.g. Aminco) • TL buffer (see Protocol 1) • 1.5% (w/v)LDAO • 150 mM NaCl • 30% (w/v) ammonium sulfate • DE52 ion-exchange media (5 cm x 15 cm column)

• Q.-Sepharose ion-exchange media (1.6 cm x 10 cm column) • Stepwise 50-300 mM NaCl gradients in TL buffer • Continuous NaCl gradient, increasing from 0-400 mM NaCl • Superdex-200 gel-filtration media (1.6 cm x 70 cm column) • Amicon stirred cell • Ultraflltration membrane (MWCO SOkDa)

Method NB: Cany out all steps (except step 4) at 4°C and in the dark. 1. Harvest the cells from 40 litres of purple bacteria by centrifugation at 10 000 g for 20 minutes. Resuspend the pellet in approximately 200 ml of Tris buffer. 2. Add a pinch of DNase and MgCl2 and rupture the cells using a French Press at 950 p.s.i. 3. Pellet the cell membranes by ultracentrifugation at 185000 g for 90 min and resuspend the pellet in 400 ml of Tris buffer. 4. Solubilize the RCs by adding 1.5% (w/v) LDAO and 150 mM NaCl and incubate in the dark, while stirring, at 28°C for 1 h, 5. Ultracentrifuge the sample for 90 min at 185000 g. The RCs are now in the supernatant.

280

CRYSTALLIZATION OF MEMBRANE PROTEINS Protocol 4 continued

6. Precipitate the RCs by adding 30% (w/v) ammonium sulfate to the supernatant, with stirring. Centrifuge for 10 minutes at l0 500g, resuspend the pellet in TL buffer and dialyse overnight against 5 litres of TL buffer. 7. Load the RC sample on to a DE52 column (5 cm x 15 cm), pre-equilibrated with TL buffer, Elute the RC with a stepwise gradient of increasing NaCl concentrations (50-300 mM) in TL buffer. The RC elutes in 200-300 mM NaCl, Dialyse the sample overnight to remove the NaCl, 8. Load the RC sample on to a OjSepharose FPLC column (1.6 cm x 10 cm) and elute with a continuous NaCl gradient, increasing from 0 to 400 mM NaCl. Concentrate the sample to a smaSl volume (typically 4 to 6 ml) using an Amicon stirred cell with a 50 kDa MWCO membrane. 9. Load 2 ml of RC sample on to a Superdex-200 FPLC column (1.6 cm x 70 cm) and run at 0.6 ml/min. Concentrate the purified protein using either an Amicon stirred cell or a Centricon micro concentrator. The protein is now ready for crystallization trials.

3.2.2 Crystallization The first crystal structures of the RC from Kb. sj'dnsToirfc.s were determined from orthorhombic crystals, space group P2,2|2 1 , which wore grown using LI PEG 4000/NaCl precipitant mixture (64, 66). This orthorhombic form can be obtained using cither LDAO or p-OG detergents, and both sitting-drop vapour diffusion and dialysis techniques have been used (see TuMi; 1). The phase diagrams for the crystallization of the RC using PEG 4000 have been determined both with and without the presence of the amphiphile, HPTO (75, 76). The amphiphile is required for crystallization to occur when using the detergent LDAO. However, the protein complex will crystallize from B-OC in the absence of HPTO concomitantly with phase separation. The trigonal crystal form was first reported by Buchanan ct nl. in 1993 (68). A year later the structure of the RG was published by Frmler et al at a resolution of 2.65 A (69). The main difference in the crystallization of the trigonal form compared to the orthorhombic form is the choice of precipitant: the orthorhombic crystals are grown from PEG 4000, whereas the trigonal crystals are obtained from potassium phosphate (or other salts). The amphiphile HPTO is required for the growth of these trigonal crystals, but the detergent can be any one of several. Trigonal RG crystals can be grown over a fairly wide pH range, from pH 7,0 to pH 93, Generally a pH of 8.0 is chosen, using a Tris-HGl buffer, and this method is detailed in Prnioi'oi 5, The pH affects the crystal growth kinetics: crystallization requires 2-6 weeks at pH 8.0, but crystals appear within 1-2 weeks at pH 8.5. Crystals grown using the method described in IVutDrt)! 5 are shown in figure 2C. 281

Table 1 Crystallization conditions for the RC from Rb. sphaeroides Solution

[RC]

[PEG 4000]

[NaCI]

p [Detergent]

[HPTO]

Method

Reference

Orthorhombic crystal form, space group P212121

Z

Drop Reservoir

5 mg/ml

12% 22%

0.36 M 0.60 M

0.06% LDAO

3.9%

sitting-drop

77

Drop Reservoir

30 MM

10% 22% 12-14.5%

0.15 M 0.25 M 0.22 M

0.80% p-OG

1.0%

sitting-drop

78

0.80% p-OG

—

dialysis

45

10% 25%

0.30 M 0.30 M

0.80% p-OG

—

sitting-drop

79

—

0.85% p-OG

2.5% 0.4% Bz

sitting-drop

70

0.09% LDAO

3.0%

sitting-drop

68

0.10% LDAO

3.5%

sitting-drop

7

2 mg/ml Drop Reservoir

20 MM

10 mg/ml

6% 32%

Trigonal crystal form, space group P3±2± Drop Reservoir

10.6 mg/ml

1.0 M KP, pH 7 1.3-1.5 M KP,

Drop Rservoir

12.5 mg/ml

0.75 M KP, pH 8 1.4-1.6 M KP,

— —

O

i. p-

Tetragonal crystal form, space group P43212 Drop Reservoir

o

CRYSTALLIZATION OF MEMBRANE PROTEINS

Protocol 5 Crystallization of trigonal crystals Equipment and reagents • 2 M di-potassiurn hydrogen phosphate pHS.O • 1.4-1.6 M di-potassium hydrogen phosphate • 10mMTris-HClpH8.0,0.1%(w/v)LDAO • RC sample. 20 mg/ml in above buffer

HPTO Crystallization plate (see Protocol 3} Clear sealing tape (see Protocol 3}

Method 1. Weigh 14 mg of HPTO into a microcentrifuge tube to give a final concentration of 3.5% (w/v). 2. Add 250 nl of the RC sample and vortex to mix. 3. Add 150 ni of 2 M di-potassium hydrogen phosphate and mix immediately. 4. Centrifuge at 10000gin a microcentrifuge for Smin at 4°C. 5. Meanwhile, add 1 ml of 1.4-1.6 M di-potassium hydrogen phosphate to the reservoirs of the crystallization tray. 6. Add 15 ul of RC/precipitant solution to each sitting-drop depression of the crystallization tray. 7. Seal with clear tape and incubate at 18°C. Crystals appear within 2-4 weeks.

The most recent crystal form to be published for the RC from Kb. sphccroirfi's is The tetragonal form, space group P4:S212 (31, 70). These crystals diffract to a high resolution of 1.9 A, The crystallization conditions are similar to those for the orthorhombic form, i.e. PEG 4000 and B-OG are both used, but the- concentrations differ and two amphiphiles are added, HPTO and bcnzamidine. These examples demonstrate that it is possible to crystallize the RC in a number of different crystal forms as described in more detail by Allen et al (77) and briefly discussed here. Since these different crystal forms diffract to different resolution limits, it is recommended that all the crystal forms obtained when searching for the 'ideal' crystallization conditions arc screened for diffraction. It is not only necessary to consider resolution limits but also the ease of data collection. The orthorhombic and trigonal crystal forms both suffer severe radiation damage at room temperature when exposed to X-rays. This means that multiple crystals are required to collect a complete data set, leading to an accumulation of errors and degrading the quality of the resulting data. Under these circumstances, the trigonal crystals are preferred since their higher symmetry means that fewer degrees of data have to be collected for high completeness. This is less of a consideration if the crystals can be cryocooled to 283

TINA D. HOWARD ET AL.

100 K but this has proved difficult for the trigonal crystal form. Even with the use of cryoprotectants, there is still some damage to the crystals which leads to high mosaicity and loss of resolution. However, the tetragonal crystals were cryocooled and still diffracted to high resolution (70).

3.3 Summary of crystallization conditions used for other membrane proteins Although the number of membrane proteins whose structures have been solved is still small, many references chart the progress of their successful crystallizations. Table 2 is intended to briefly illustrate the conditions and techniques used in producing crystals which diffract to a high resolution.

3.4 Important factors in crystallization Factors which particularly influence the crystallization processes in both lightharvesting complexes and reaction centres are discussed in more detail below. Table 2 illustrates that although there are similarities in crystallization conditions for different membrane proteins, the overall combination of components tends to be protein-specific. Whilst we have found that the LH2 complex from Rps. atidophila 7050 crystallizes with only minor adjustments to the crystallization conditions used for Rps. addophila 10050 (85), Michel and co-workers have crystallized the equivalent protein from Rs. molischianum using ammonium sulfate with either undecyldimethylamine oxide (UDAO) or LDAO in the presence of HPTO (25). In addition to exploring a wide-range of crystallization conditions to try to crystallize a given membrane protein, it is also worth trying to crystallize related proteins: the same protein from a different bacterium may be more amenable to crystallization. For example Section 6.1 and Section 6.2 point out how the ease of crystallization of RCs and cytochrome c oxidases (COXs) from different sources can vary dramatically. 3.4.1 Choice of precipitant There are no set rules as to which are the 'best' precipitants. As many precipitants as possible should be tried in the quest for good crystals. We have had success with either salts, such as ammonium sulfate, sodium citrate, and potassium phosphate, and with the polyethylene glycols. The use of ammonium sulfate as a precipitant has been shown to affect the pH of the crystallization drop owing to the diffusion of ammonia vapour (86). Derivatives of PEGs, i.e. monomethyl ethers (MME-PEG), have been reported as being useful in crystallizations (e.g. see ref. 84). RC crystals can be grown from either PEG 4000 or from high salt concentrations. The main factor involved in growing trigonal crystals of the RC would appear to be a high ionic strength in the crystallization medium. We have obtained trigonal crystals from several different salts, i.e. potassium phosphate, 284

CRYSTALLIZATION OF MEMBRANE PROTEINS

ammonium sulfate, and trisodium citrate. In each case the crystals are isomorphous with unit cell dimensions within 1% of a = b = 142.4 A, c = 187.1 A. The diffraction properties of the crystals grown from different salts are similar, but can be ordered with respect to resolution as: sodium citrate ~ potassium phosphate > ammonium sulfate, with values ranging from 2.1 A to 2.8 A (measured at a synchrotron source). The large crystals of Rps. acidophila 10050 LH2 which were used to solve the structure (8, 9) were produced in the presence of potassium phosphate and ammonium sulfate, as detailed in Protocol 3. Crystals are obtained over a range of concentrations of the precipitant, certainly within the range 1.8 M to 2.8 M ammonium sulfate. The crystal growth rate is increased when the precipitant concentration is raised. However, crystals grown from the more concentrated ammonium sulfate solutions tend to be generally smaller and more variable in size and diffraction quality than those produced at lower concentrations; a compromise between these two extremes yields crystals of sufficient size and diffraction quality on an acceptable time scale (usually 6-8 weeks). To produce a continuous supply of fresh Rps. acidophila 10050 LH2 crystals, the precipitant concentrations are varied from 2.3 M to 2.0.M in each set of crystallization trials. Protein crystallization protocols, using the vapour diffusion technique, frequently add similar volumes of protein solution (without precipitant) and reservoir solution (containing the precipitant) into the drop; this Rps. acidophila 10050 LH2 procedure is different. Typically 15 ul of the protein solution containing a high concentration of potassium phosphate is added to 1.5 ul of the reservoir solution, in this case ammonium sulfate solution. Again it is possible to alter the speed at which the crystals appear by varying the ratios of these volumes a little. Without the addition of ammonium sulfate solution to the drop, crystals appear much faster, albeit less consistently. Only limited success has been achieved in the crystallization of the antenna complexes using PEG 2000 as a precipitant, with either a polyoxyethylene detergent or dodecyl maltoside. In both cases very fine needle-like crystals are obtained, variations in additives are being investigated to produce 'chunkier' crystals for data collection.

3.4.2 Choice of detergent Two major factors are important with respect to the choice of detergent. First, the membrane protein must be stable in the detergent over an extended period of several weeks at typical temperatures between 4°C and 25°C. Second, the micelle size of the detergent must be small enough so that it does not sterically prevent the hydrophilic parts of the membrane protein coming together to form protein-protein contacts in the crystal. Often the best detergent for isolating and purifying your proteins will not be the optimal one for crystallization. It is a good idea therefore to try to work with detergents that can be readily exchanged (see Protocol 2). The properties of a particular detergent are governed by its critical micelle concentration (CMC); this is the concentration at which monomers aggregate 285

Table 2 Summary of crystallization conditions published for other membrane proteins Protein source

Detergent

Additives amphiphiles

Precipitant

Buffer PH

Method temperature

Space group

Resolution (A)

Reference

P-OG

NaCI NaN3

PEG 2000

Tris/NaP, pH4-9

SD/^D 21-23 "C

P42, P6322 P21, C2

4.5-2.9

41, 80

NaCI NaN3

PEG 2000

NaP, pH6.5

SD 21-23 °C

P42

3.2

41, 11

CS-HESO

NaCI NaN3

PEG 2000

Tris pH9.8

uD

P321

2.7

81

CS-HESO

Porins £. co//, OmpF

CS-HESO E. co//, OmpF

P-OG

E. co//, OmpF Rb. capulatus

C8E4

LiCI NaN3

PEG 600

Tris pH 7.2

HD 20°C

R3

1.8

82, 14

Rps. blastica

C8E4

LiCI

PEG 600

Tris pH7.2

HD 20°C

R3

2.0

13

P. nitrificans

P-OG

CaCI2 KCI, NaN3

PEG 600

Tris PH7.5

SD

PI

3.1

14, 83

P. nitrificans

P-OG

CaCI2 KCI, NaN3

PEG 4000

Tris PH7.5

SD

C2

3.1

14, 83

E. co//, LamB

DM

MgCI2

PEG 20000

Hepes PH7.0

uD

C2221

2.4

15

S. typhimurium ScrY

P-OG HDAO, p-HG

LiCI MgCI2

PEG 2000

Tris pH 7.7

SD 17°C

2.4

16

P21, C2

LM

NaN3

n/s

n/s

complexed Fv antibody

2.8

17, 18

Cytochrome c oxidase P. nitrificans (4 subunits)

P4

P. nitrificans (2 subunits)

UM or CYMAL-6

NaCI

PEG 2000 MME

NaAc pH5.5

complexed fv antibody

Bovine

DM

n/s

PEG 4000

Nap

Batch, 4°C

NaCI

PEG 4000 glycerol

MES/MOPS pH 6.5/7.5

MPD

Na/K P, pH5.6

LCP

NaP,

SD

i

2.7

19

2.8

20

P212121

3.0

21

P63

2.5

23, 24

P42±2

2.4

25

P63

4.0

26

P212121 P2121

Miscellaneous membrane proteins Cytochrome bc± Chicken heart

B-OG

LM

Bacteriorhodopsin H. halobium

LH2 Rs. molischianum

UDAO

HPTO NaN3

AMS

monoolein

20 °C

Photosystem 1 S. elongatus a-haemolysin S. aureus

(B-OG

PEG 5000 MME

AMS

NaCac pH 6.0

HD

C2

1.9

27,84

PGHS-1 Ovine

B-OG

NaCI flurbiprofen

PEG 4000

NaP, pH6.7

HD

1222

3.5

28

Squalene cyclase All. acidocaldarius

C8E4

LDAO

Na citrate

Na citrate pH4.8

HD

P321

2.9

29

K* channel S. lividans

LDAO

CaCI2 KCI, DTE

PEG 400

Tris/Hepes pH 7.5

SD

C2

3.2

30

Abbreviations: n/s, not stated; Lid, microdialysis; SD, sitting-drop vapour diffusion;HD, hanging-drop vapour diffusion; LCP, lipidic cubic phase.

TINA D. HOWARD ETAL.

and form micelles in solution (87). For a detergent with a low CMC the concentration of detergent present in micelles will be approximately equal to the total amount of detergent added to solution, since very little exists as monomers. When the detergent has a high CMC the micellular component of the detergent will be equal to the total detergent present minus that present as monomers (i.e. the total amount minus the CMC). Table 3 illustrates the range of detergents that have been used in membrane protein crystallization trials for Rps. acidophila 10050 LH2, achieving varying success rates. The RC from Kb. sphaeroides is a further example of another membrane protein that has a fairly low specificity for detergent in the crystallization process. We have obtained trigonal crystals using LDAO, dodecyl maltoside, octyl- and nonyl-glucosides, heptyl-thioglucoside, and SB-12 (Zwittergent 3-12). Changing the detergent from LDAO to the previously listed detergents did not improve the diffraction properties of our trigonal crystals, and, in several cases, an increase in crystal disorder was noted, e.g. for the detergents dodecyl maltoside and SB-12. In some cases it may prove beneficial to exchange the detergent after crystallization has occurred. For example, Allen et al. (88, 89) were able to improve the order of their orthorhombic RC crystals by exchanging the original detergent, LDAO, for (3-OG using a microdialysis technique. The improvement in crystalline order was accompanied by a change in unit cell dimensions. Not all membrane proteins are as easy to crystallize as the RC and, in some cases, the type and structure of the detergent can be of extreme importance. Such an example is the crystallization of bovine heart cytochrome c oxidase (COX) (90). Different crystal forms could be obtained by changing the detergent: a hexagonal form was grown from Brij 35 (C12E23) and a more ordered tetragonal form was obtained from either C12E8 or C12E7 detergents. The diffraction improved from 10 A to 5 A on going from the hexagonal to the tetragonal form. Closely related detergents, C12E6 and C12E5, were also evaluated but with poorer results and the enzyme was unstable in C12E5. All these detergents have a similar structure but different ethylene glycol chain lengths. The results indicate that there is an optimum chain length for this particular protein. Since this may be true for other membrane proteins, a comprehensive screening of detergents is recommended. The standard advice for testing a new detergent is to try a concentration of the detergent just above the CMC. Values of CMCs can be determined by several methods, e.g. changes in dye absorbance or fluorescence upon solubilization and measurement of light scattering, surface tension, or hydrodynamic properties (see refs 91, 92 and references therein). The CMC value obtained can vary with the technique used and is frequently determined for a solution of pure detergent in water. The presence of other components in the system, e.g. amphiphiles, proteins, salts (including buffer salts) can all have an effect on the aggregation process and hence the effective CMC value in a particular environment. Literature values for (3-OG, one of the more popular detergents, vary over twofold from 14 mM to 30 mM, with the majority of the reported values lying between 22 and 25 (92-96). Whilst CMC values do provide a useful guide for 288

Table 3 Summary of crystal forms and resolution of Rps. acidophila 10050 LH2 crystals Appearance

Space group

Resolution (A)

Reference/Comments

0.75% p-OG (26 mM)

rhombohedral

R32

2.5

8, 9; room temperature, synchrotron

0.75% p-OG (26 mM)

rhombohedral

R32

2.0

10; cryocooled, synchrotron

0.15% LDAO (7 mM)

rhombohedral

R32

2.2

10; cryocooled, synchrotron

0.15% UDAO (7 mM)

rhombohedral

R32

3.4a

same structure as in LDAO and B-OG

5% p-HG (190 mM)

large needles

P422

3.4

cryocooled, in-house source

0.75% B-OGal (26 mM)

rhombohedral

R32

3.5a

quick scan

0.07% P-DG (2.2 mM)

rhombohedral

n.d.

very small

0.3% thio-p-OG (9 mM)

rhombohedral

n.d.

—

0.7% HECAMEG (20 mM)

rhombohedral

n.d.

—

2.6% NOGA (8 mM)

rhombohedral

3.5"

large ~1 mm

0.03% MEGA-10 (0.9 mM)

rhombohedral

n.d.

very tiny , problems dissolving the detergent at concentrations anywhere near the CMC

square plates

n.d.

very thin in one dimension, many twinned, crystallization conditions to be optimized

Detergent Structural determinations

Same crystal form

R32

Different crystal forms 5% ODAO (290 mM) 0.0095% LM (0.18 mM)

cubic

10

crystallization conditions to be optimized

0.009% LM (0.17 mM)

very fine needles

n.d.

reproducible, too thin for data collection

0.008% Brij 35 (0.06 mM)

needles

n.d.

reproducible, too thin for data collection

0.02% Brij 58 (0.17 mM)

blunt-ended needles

n.d.

too thin for data collection"

n.d., not determined. "No further optimization attempted. "Also obtained using PEG 2000 as a precipitant, MgCI2 and spermidine.

TINA D. HOWARD ET AL.

initial crystallization trials, it may be productive to test detergent concentrations below or above a published value. We have produced quality crystals under conditions where the concentration of detergent is below the published CMC value. Table 4 gives information on pseudonyms, structures, sources, and CMC values of a number of different types of detergents: the full range of compounds available are too numerous to try to include here. Broadly, detergents can be categorized into non-ionic, ionic, and zwitterionic; further subdivisions are often used, e.g. non-ionic detergents can have sugar (glucoside, maltoside etc) or sulfur-based headgroups (e.g. sulfoxide) or contain polyoxyethylene chains. Progressively a range of different carbon chain-length substitutions of the common detergents are now becoming available as well as substituted derivatives (e.g. thio-sugars) with enhanced chemical and thermal stability. Overall the 'classic detergents' such as (3-OG, dodecyl maltoside, and polyoxyethylenes frequently provide starting points for crystallization trials, but there are several new detergents available, e.g. CYMAL series, HEGA, and CHEGA series. The CYMAL series contains a cyclohexyl ring as part of the hydrocarbon chain coupled to the familiar maltoside group and was developed to incorporate more hydrophobicity into a shorter chain length (121, 122). The usefulness of the MEGA series of detergents has always been limited by their solubility; the presence of an extra hydroxyl group in the HEGA series permits more concentrated solutions to be obtained (121); C-HEGA combines both of the above modifications into the glucosamide molecule. Also compounds which mimic more closely the natural phospholipids present in membranes are being developed. These are designed to resemble the environment in the membrane so as to stabilize membrane proteins in aqueous solutions, e.g. amphiphols (123) and phosphocholine compounds (121, 124). Porin crystals diffracting to 3.5 A have previously been produced using shortchain phospholipids to displace the native lipids and circumvent the need for detergents and amphiphiles (125). When aiming for the highest possible resolution structure, it is well worth noting that the purity of detergents from different sources/companies can vary considerably, both chemically and stereochemically, as can the price (not always proportionally!). Section 5.2 contains details of a number of detergent screening kits available commercially. 3.4.3 Choice of additives and amphiphiles Traditionally, additives to crystallization trials have included co-factors necessary for maintaining the stability of proteins and salts affecting the ionic strength and hence solubility of the protein. The addition of non-hydrolysable substrate analogues to 'lock' a dynamic protein in a static form has already been referred to in the Introduction, in solving the structure of ATP synthetase (2, 3); similarly the structures of other membrane proteins, maltoporin and prostaglandin-H-synthetase, have been solved as complexes with various substrate analogues (15, 28, 126). 290

CRYSTALLIZATION OF MEMBRANE PROTEINS

Also described in Section 1, Michel (34, 35) and Garavito (36, 37) introduced the so-called 'small amphiphile' concept for crystallizing membrane proteins. Since then a very wide range of dissimilar small amphiphilic molecules have been successfully used as additives in membrane protein crystallizations. They have been shown to affect the crystallization process by changing the phase diagram of the detergent (91, 127), the size of the detergent micelle (128), the number of detergent molecules in the detergent-protein micelle (129), and they can affect the crystal form obtained (54). Of the more successful amphiphilic additives, three have been used in numerous published membrane protein crystallizations, include heptane-1,2,3triol (HPTO), benzamidine hydrochloride (Bz), and D,L-pipecolinic acid (PCA). The list of possible compounds is too long to include here (see, for example, ref. 43); a few examples will suffice. Song and Gouaux have use n-decyldimethylphosphine oxide and n-decyldiethylphosphine oxide to help prevent twinning and promote the growth of larger crystals of the heptameric form of a-haemolysin (84). Compounds as diverse as CsBr and propionamide have been used to improve crystal quality in another photosynthetic protein, phycoerythrin 545, which, although water-soluble, requires detergent for crystallization (130). Glycerol is needed to crystallize Rps. acidophila 10050 LH2 in LDAO; dioxane, spermidine, spermine have all achieved degrees of success within our groups' crystallizations of bacterial photosynthetic proteins. The important advice is to try as many compounds as your supply of protein will permit! The mode of action of these additives is not really known. It is interesting to note that no amphiphiles are required in the composition of 'artificial mother liquor' used for handling crystals of LH2 (Section 4.1). Crystals still diffract well after several weeks of storage in such solutions, suggesting that either the amphiphilic molecules are intimately associated with the protein-detergent micelle and are not free to diffuse out of the crystal, or that the presence of the amphiphile is somehow important during the crystallization process but not required to stabilize the crystals afterwards. This is a contrast to the early work by Michel's group with Rs. molischianum LH2 (86) where an initial crystal form redissolved as HPTO crystals began to appear in the drop, reducing the HPTO concentration of the solution. The effect of adding HPTO to RCs in different detergent solutions has been measured by re-solubilization of detergent-free RCs. This demonstrated that HPTO reduces the number of detergent molecules bound to the reaction centre by a factor of approximately two (128). This occurred for the detergents commonly used in the crystallization of RCs: fi-OG and LDAO. Its presence also has an effect on the phase diagram (127). HPTO has already been shown to affect the size and shape of LDAO micelles (128). 3.4.4 Choice of pH Again a wide range of pH values should be tried, pH can affect both the stability of the protein and the process of crystallization. Also worth considering is the effect of pH on the chemical stability of the detergent, e.g. the presence 291

Table 4 A selection of the detergents used in crystallizatioin trials Detergent (Common abbreviations)

Suppliers9

Reference

99

(mM)

CMC % (w/v)

n-Hexyl-p-oglucoside (P-HG)

250

6.6

C,F,Hk,S

n-Octyl-p-o-glucoside (p-OG)

14-30

0.4-0.9

An,Ba,BM,C,D,F,Hk

92-99

n-Nonyl-p-D-glucoside (p-NG)

6,6.5

0.2

An,Ba,C,D,F,Hk

99, 100

n-Decyl-p-o-glucoside (P-DG)

2.2, 2.3

0.7

An,C,F

101, 102

An,C,F,Hk

99, 103

Non-ionic—sugar-derived headgroup

rvHeptyl-p-D-thioglucoside(triio-p-HpG)

29

n-Octyl-p-D-thioglucoside (thio-p-OG)

9

2.8

An,BM,C,D,F

104, 105

6-O-(/V-Heptylcarbamoyl) methyl-a-oglucoside (HECAMEG)

19.5

0.65

An,C,Hk

106

n-0ctyl-B-D>galactoside (B-OGal)

29.5

0.9

AI,An

107

n-0ctanoylsucrose (OS)

24.4

1.1

C,Hk

108

n-Dodecyl-p-D-maltoside (lauryl maltoside, LM)

0.17

0.009

An,Bm,C,D,C,F,Hk

95, 98

n-Octanoyl-B-D-glucosylamine (NOGA)

80

2.4

c

109

n-Octanoyl-W-methylglucamide (MEGA-8)

58, 79

1.9,2.5

AI,An,C,D,Hk

110, 111

n-Nonanoyl-W-methylglucamide (MEGA-9)

25

0.8

An,C,D

110

n-Decanoyl-A/-methylglucamide (MEGA-10)

6-7

0.25

An,Ba,C,D

110

CyclohexIbutanoyl-W-hydroxyethylglucamide (C-HEGA-10)

35

1.3

An.Hk

99

20.6

0.52

Ba

112, 113

Non-ionic—sulfur-derived headgroup

2,3-Dihydroxypropyloctyl sulfoxide (DOPSO)

Non-Ionic—polyoxyethylenes

AI,An

101, 114 97 97

Dodecyltricosoxyethylene (Brij 35; C12E23)

0.09

0.01

AI.An

Hexadecyleicosoxyethylene (Brij 58; C16E20)

~0.08C

0.01

AI.An

Hexadecyleicosoxyethylene (Brij 56; C16E10)

0.19

Non-ionic—bile-salt derivative Sucrose monocholate

4.7

0.35

D

Dojondo, pers. comm.

162-223

2.8-3.9

AI,F,S

ZwKterionlc />Nonyldirnethylamine-N-oxide (NDAO)

50.8

0.95

AI,F

n-Undecyldimethylamine-Woxide (UDAO)

6

0.13

AI.F

n-Dodecyldimethylamine-Woxide (NDAO) lauryldimethylamine-Woxide (LDAO)

0.4-2

0.01-0.07

AI,An,C,F,Hk

98, 111, 115 98,115 116 98, 111, 116, 117

3-(3-cholamidopropyl)-dimethylammonio-l-propane sulfonate (CHAPS)

4-10

0.25-0.6

An,BM,C,F,Hk,S

97, 98, 118

9-15

0.4-0.6

An,BM,C,F

102. 119

nOctyldimethylamine-Woxide (ODAO)

Ionic—bile salts Sodium cholate a

Al,Aldrich; An, Anatrace; B, Bachem; BM, Boehringer-Mannheim; C, Calbiochem; D, Dojindo; F, Fluka; Hk, Hampton detergent kit; S, Sigma. "Extensive lists of detergents and CMCs are contained in various references and reviews e.g. websites (Sect/on 7), Calbiochem detergent booklet, Anatrace catalogue, Bachem information sheet and references 113 and 119. c Estimated from CMCs of C16E17 and C16E32 quoted in reference 88.

TINA D. HOWARD ET AL.

of a basic labile group may result in hydrolysis at high pH over a prolonged time. When initially screening the LH2 antenna complex for crystallization, pH values between 5.0 and 10.0 were tried. The complex was found to be stable for extended periods only above pH 8.0 and well-ordered crystals were obtained above pH 9.0 (9). A pH of 9.35 produces Rps. acidophila 10050 LH2 crystals which diffract to a high resolution, for the related complex from Rps. acidophila 7050 a higher pH (> 9.5) is found to be necessary (85).

3.5 Choice of temperature Frequently 4 °C and 20 °C are selected as starting points to investigate the effect of temperature on the crystallization process. Rps. acidophila 10050 LH2, Kb. sphaeroides RC, and a range of Kb. sphaeroides RC mutants all crystallize when incubated at 16°C. Since the vapour pressure of a liquid decreases with temperature, a lowering of temperature reduces the rate of equilibration via the vapour phase and hence the rate of crystallization. It must be remembered that the overall effect of a temperature change may be the result of a combination of factors, i.e. change of the CMC of detergent, changes in the solubilities of additives, shifts in the phase diagram, conformational changes within the protein, and thermostability of the protein. It is important that the system is left at constant temperature to reach equilibrium and (hopefully) produce crystals. Although the progress of the crystallization process needs to be monitored, it is worth remembering that a drop with a volume of 15 ul can heat up rapidly under a microscope light!

3.6 Concentration of the protein solution Initially the correct protein concentration range for crystallization must be determined; too low and it will never reach saturation and be able to crystallize, too high and the protein will rapidly precipitate out of solution as an amorphous mess. Again, the protein concentration required varies from protein to protein, but an initial concentration in the drop of 2-10 mg/ml is a sensible place to start screening. After the initial screens the concentrations can be adjusted accordingly, e.g. lowering the protein concentration may slow down the rate of crystallization and help to produce fewer, larger crystals and this should be tried if showers of crystals are observed, a fact we have utilized in developing the crystallization procedure for Kb. sphaeroides LH2. The conditions necessary for the optimum nucleation and optimum growth of crystals may differ (42, 131); the aim of producing a few well-formed crystals is to obtain a few nucleation sites before the system passes into the crystallization region of the phase diagram, where the crystals grow until the protein supply is exhausted. Seeding techniques aim to optimize the conditions for the growth of crystals, bypassing the need for nucleation by adding small ready-formed crystals as seeds (131). When concentrating a membrane protein solution using ultrafiltration, it is important that the size of the detergent micelles present in the solution is 294

CRYSTALLIZATION OF MEMBRANE PROTEINS

considered; a membrane of a suitable MWCO must be selected to avoid inadvertently altering the concentration of detergent in solution. The aggregation number for p-OG is 78 (121), which means micelles are in the order of 22 kDa and LM, with an aggregation number of 78-92 (121), has micellular size of around 43 kDa. Very small-scale crystallization trays are available, holding 5 ul drops as sitting drops (e.g. Douglas Instruments), these are useful for screening a large number of conditions if the amount of protein is very limited. The drawback with such small volumes is that the protein may be depleted from the solution before the crystals reach a useful size. Exploiting the same methodology, we have used drops up to 50 ul to obtain very large Rps. acidophila LH2 crystals in deuterated detergent for use in neutron diffraction experiments where relatively large crystals are required. Chayen (47) has overcome the problem of rapid evaporation from small volumes by using a layer of light oil to reduce evaporation, allowing samples as small as 1-2 ul to be used in trials, along with the development of an semiautomated procedure.

4 Preparing crystals for data collection 4.1 'Artificial mother liquor' Obtaining crystals is only one step of many towards attaining the ultimate goal of a three-dimensional structure. Handling the crystals correctly is also very important in order to obtain high-resolution diffraction data. As well as being susceptible to mechanical stresses, e.g. vibrations and shearing, the crystals are also very susceptible to dehydration on the molecular scale. Water molecules form an integral part of the structure and thus any alteration in the hydration state of the protein can introduce heterogeneity into the structure of the crystal. To mount a crystal for data collection with no minute traces of precipitate etc., it can be useful to first transfer the crystal to an 'artificial mother liquor'. Soaking the crystal in a suitable simulated mother liquor can quite significantly increase the resolution limit and quality of diffraction data obtained (9). Before a new structure can be solved, the 'phase problem' must be addressed (5); one way of approaching this is to prepare isomorphous derivatives of the protein containing heavy metals whose positions can be readily identified within the unit cell. A suitable mother liquor is required to soak the crystals with an appropriate heavy metal compound and produce heavy metal derivatives of the protein (132). All too frequently, it is necessary to transport crystals to a suitable synchrotron X-ray source, transferring them into a small amount of 'artificial mother liquor' in a 500 ul microcentrifuge tube provides a much more robust mode of transporting them than in situ in the sitting-drop tray with the potential hazard of tipping up the tray! The precise composition of the 'artificial mother liquor' is crucial to maintain the crystals without redissolving or dehydrating them; the method used for Rps. acidophila 10050 LH2 is detailed in Protocol 6. 295

TINA D. HOWARD ET AL

Protocol 6 Preparation of 'artificial mother liquor' Equipment and reagents • 2.3 M ammonium sulfate pH 9.4" • Artificial mother liquor (AML): 20 mM Tris-HCl pH 8.0,1.5 M K2HPO4, 32 mM NaCl, 0.5% B-OG

• Large-scale, sitting-drop pot1" • Sealing tapec • Incubator at 16°C

Method 1. Place 10 ml of the (NH4)2SO4 solution in the bottom of the pot. 2.

Pipette 3 X 300 ul of 20 mM Tris-HCl pH 8.0. 1.5 M K2HPO4, 32 mM NaCl, 0.5% B-OG into the hollows in the bridge.

3. Replace the lid and seal with tape, 4. Incubate at 16°C for at least 36 h. 5. Decant the 'artificial mother liquor' (AML) and store at 16°C in a tightly closed vial. "Adjust the pH of the solution immediately prior to use. ' A circular pot with close fitting lid (diameter ~ 75 mm, height - 30 mm) with an attached bridge containing 3 x 350 ul hemispherical depressions (133). c Use tape which will peel off easily to facilitate opening the pots without splashing the equilibrated solutionsi

4.2 Cryoprotection At room temperature the lifetime of a crystal in an X-ray beam is often very limited because of radiation damage. Thus a large number of crystals are required to be able to collect enough data to determine a structure. In addition to the fact that a large supply of crystals requires the availability of sufficient protein, large-scale protein purification, etc., merging diffraction data from different crystals can increase the mosaic spread of data, thus reducing the overall quality of the diffraction data. Radiation damage can be considerably reduced by recording data at very low temperatures, e.g. 100 K (134, 135), permitting complete data sets to be collected from a smaller number of crystals, even a single crystal. There are many hurdles to overcome to successfully freeze crystals to this temperature without the formation of ice. Practical problems include finding a suitable cryoprotectant in which the crystal is stable and also how to introduce a high level of cryoprotectant without 'shocking' the crystal, which can cause local disruptions and irregularities within the crystal structure. Stepwisc additions of the ctyoprotective agents to the mother liquor are often carried out to overcome the danger of osmotically shocking the crystal; we have developed

296

CRYSTALLIZATION OF MEMBRANE PROTEINS

a method of adding the cryoprotectant slowly and continuously using a very low molecular weight dialysis membrane and microdialysis buttons. For Rps. aadaphila 10050 LH2, sucrose was an obvious candidate when selecting a cryoprotectant since the protein has already been shown to be stable for prolonged periods in the presence of high levels of sucrose during the purification procedure (see Protocol 1). Slowly dialysing sucrose into the 'artificial mother liquor' containing the crystal through a very low molecular weight cut-off membrane introduces the cryoprotectant into the crystal satisfactorily, as detailed in Protocol 7. (The microdialysis buttons used in this procedure are those designed for use in protein crystallizations equilibrating the two solutions across the dialysis membrane (see figure 1h). Using this procedure we have been able To extend the resolution of the Rps, acidophila 10050 LH2 structure from 2.5 A to 2,0

A (10).

Protocol 7 Equilibration of Rps. acidophila 10050 LH2 crystals prior to cryocooled data collection Equipment and reagents • 'Artificial mother liquor'(AMI, see

Protocol*) • Saturated sucrose solution (room temperature) in Tris buffer" • 3 M K2HPO4, 64 mM NaCI

• Microdialysis buttons. 10 fil capacity (Cambridge Repetition Engineers or Hampton Research) • Applicator tool (Hampton Research)" • Forceps

• Dialysis membrane, MWCO 3500 Da, diameterlS mm (SpectraporS series; NBS Biologicals} or pre-cut dialysis membraoe discs, MWCO 3500 Da, diameter 33 mm {SpectraporS series; Hampton Research or NBS Biologicals) • Plastic screw-top vial (nominal capacity 7ml:.tg.BibbySteriEn) • Scalpel blade

Method 1. Make cryoprotected 'artificial mother liquor' by mixing the saturated sucrose solution and 3 M K2HPO4, 64 mM NaCI in equal proportions. After the addition of the relevant detergent/ e.g. 0.5% p-OG, adjust the pH of solution to lOA"* 2. Cut a length of dialysis tubing* (~ 3 cm) and score down one of the long edges with a scalpel, ease the layers apart and open out the membrane to form a rectangle. Using forceps wet the membrane in distilled water and lie it flat on absorbent towels to remove the excess water/ 3. Pipette 5 jxl of the equilibrated AMI into a dialysis button and transfer the crystal of interest into the well*

297

TINA D. HOWARD ET AL. Protocol 7 ci

4. Add sufficient 'artificial mother liquor' to just fill the well within the micro dialysis button. Place the dialysis membrane over the well and position the chosen applicator tool over the button. Roll the O-ring down on to the button until it rests in the notch where it holds the membrane firmly in place. Trim off any excess membrane protruding from the sides of the set-up. 5. Dispense 1 ml of cryoprotected 'artificial mother liquor' into the screw-top vial. Invert the button and place the dialysis membrane surface in contact with the cryoproterted 'artificial mother liquor'. 6. Replace the lid and incubate overnight at 16 "C." "Vigorous manual shaking followed by repeated additions of sucrose produces an obviously more viscous solution than by magnetic stirring which frequently fails to thoroughly mix such dense solutions. h A golf tee and a 1.5 ml freezing vial (e.g. Sigma) have also been used successfully to ease the securing O-ring into position on the dialysis button. 'Load up' the vial with a number of Orings prior to beginning the transfer of crystals, 'When using LH2 crystals produced in LDAO, a detergent-free cryoprotectant must be used, because 0.1% LDAO undergoes a phase separation in the presence of 1.5 M K2HPO4 * Equivalent to the measured pH of equilibrated artificial mother liquor prepared as in Protocol 6. * Dialysis membranes can be pre-treated to remove contaminants, e.g. glycerol, for this particular system no pre-treatment is found to be necessary. f Take care not to let the dialysis membrane dry out completely since this can result in pinholes in the membrane which remove the molecular weight selectivity producing rapid equilibration of the solutions (136). g It is possible to place several crystals within the same button. ''Around 6 hours is sufficient for cryoprotecting crystals, but the crystals are stable and easy to transport in this set-up. They appear to suffer no detrimental effects on remaining as such for days (even weeks) providing the temperature is carefully controlled.

5 Screening protocols for the crystallization of new membrane proteins In this section we will discuss some of the possible routes ro crystallizing a new membrane protein. Again, most of these techniques are standardly applied to soluble proteins with the additional consideration of screening for a suitable detergent.

5.1 Sparse-matrix screens It is possible to screen crystallization conditions using grid screens where, for example, the precipitant concentration is varied along one direction of the plate and the pH is varied in the other direction. Grid screening can give a very comprehensive search through possible crystallization conditions but at the expense of large amounts of protein, since the screens have to be repeated for each 298

CRYSTALLIZATION OF MEMBRANE PROTEINS

different precipitant, at different temperatures, etc. A more efficient use of the protein is to use the sparse-matrix or incomplete-factorial type screen, in which a large number of different conditions (varying pH, precipitants, and additives) are sampled using a small amount of protein sample (137, 138). Hampton Research offer a ready-made sparse-matrix screen which has been designed especially for membrane proteins, called the Membfac screen. This kit provides 48 crystallization conditions to try, and the screen can be repeated with different detergents. Another sparse-matrix screen was designed for the membrane protein a-haemolysin using, specifically, the detergent C8E4 (84). This screen varies salt concentrations, PEG concentrations, and pH within a trial size of 60 conditions. Further screening kits have recently become available from Molecular Dimensions Limited (sole distributors Stratech Scientific Ltd.)

5.2 Detergent screening kits A number of companies produce 'screening kits' containing small amounts of a range of detergents. These are particularly useful when searching for the ideal detergent to use in crystallization. The smaller kits include six or eight popular detergents (available from, e.g. Dojindo, Anatrace) or there is the Master Detergent Kit from Anatrace with a total of 43 different detergents; even more extensive screening can be achieved using the three different detergent screens supplied by Hampton Research, each containing 24 different detergents.

6 Crystal packing in membrane proteins There are two types of packing arrangements in membrane protein crystals as defined by Michel (35). Type-I crystals have layers of molecules with hydrophilic contacts between the layers and hydrophobic intermolecular contacts within the layers. In Type-II crystals the main packing interactions are hydrophilic and the hydrophobic regions of the molecules are surrounded by detergent micelles, and are not directly involved in packing. Having a small detergent micelle size is therefore very important in Type-II crystals.

6.1 Crystal packing in reaction-centre crystals The RC from Kb. sphaeroides has three protein subunits L, M, and H. The L- and M-subunits are mostly embedded in the membrane, whereas the H-subunit has only one transmembrane helix, the remainder of the subunit is located in the cytoplasm and is hydrophilic in nature. This is an important point since it is the H-subunit that is involved in all the crystal contacts in both the trigonal and orthorhombic crystals (see Figure 3). The contacts are either H-subunit-H-subunit contacts or interactions between the H-subunit and the periplasmic surfaces of the L- or M-subunits. The transmembrane region of the protein is not involved in the crystal contacts and is surrounded by detergent molecules, although these are mostly too disordered to be seen in the electron density maps produced by X-ray crystallography. The detergent surrounding the RC has been 299

TINA D. HOWARD ET AL

Figure 3 Representation of the packing of reaction-centre molecules in two different crystal forms. The L- and M-subunits are shown in grey and the H-subunit is in black. (A) Orthorhombic crystal form, P2,2,21; (B) trigonal crystal form, P3,21.

seen in the orthorhombic crystal form using neutron diffraction, and this is shown in Figure 4 (139). From 1-igure 4 we can clearly see that the size and shape of the detergent micelle is a critical factor in the crystallization of a membrane protein. The detergent micelle must be small enough so that it does not interfere with the crystal contacts. Since the RC has the hydropliilic H-subunit, the detergent specificity in the crystallization of RCs is not as critical as for other membrane proteins that lack large hyclrophilic domains. An example of this can be provided by considering the RC from Gtlontflexus aurantiacus. This RC has only two

CRYSTALLIZATION OF MEMBRANE PROTEINS

Figure 4 Two views of the detergent ((B-OG) micelle surrounding the Rb. sphaeroictes reaction centre in the orthorhombic crystal, as determined from neutron diffraction studies (139).

protein subunits corresponding to the L- and M-subunits, the H-subimit is missing. At least three crystal forms have been grown, but the authors reported problems with crystal twinning, growth of platelet crystals, and highly mosaic crystals (140). Unlike the Rb. sphaewidcs RC crystals, the morphology and quality of diffraction obtained for the C. mmmtiacus RC crystals is greatly nffected by the choice of detergent and amphiphile. The problems encountered in the crystallization of this protein complex may be attributed to the absence of the Hsubunit. Since the presence of an hydrophilic domain is apparently advantageous in crystallizing rhe RC, it has been suggested that increasing the hydrophilic surface area of other more hydrophobic membrane proteins might aid crystallization. There are currently two approaches that have been used to achieve ihis effect, as discussed in the following sections.

6.2 Use of antibodies for crystallization One possible way of increasing the hydrophilic surface area of a membrane protein is to complex the protein with an antibody fragment. It is possible that the resulting membrane protein-anti body fragment may crystallize where the membrane protein alone docs not, since the antibody may be able to provide the crystal contacts required to form a 3-D lattice. This approach allowed the crystallization of bacterial cytochrome c oxidase (COX) which diffracted to 2.8 A (17, 18). The crystal packing involves polar contacts provided by the antibody fragment. In the same year as the bacterial cytochrome r oxidase structure was published, Tsukiharn ft til. published the structure of the metal site of the COX from bovine heart mitochondria (20). This latter enzyme was crystallized without the aid of an antibody fragment; overall it had taken well over a decade to find the successful crystallization conditions. 301

TINA D. HOWARD ET AL.

6.3 Use of fusion proteins This approach has been tried with the membrane transporter lactose permease (1, 141), an integral membrane protein which is extremely hydrophobic. In this case, the carrier protein chosen to aid the crystallization process was cytochrome b562. This carrier protein not only gives the Jac-permease a more polar surface but also has the advantage of being coloured, which greatly simplifies detection of the !ac-permease during handling and crystallization. The fusion protein was expressed and purified and maintained lactose transport function. However, no significant progress with crystallization has been published. The carrier protein may also be chosen to aid purification of the fusion product if it can be purified by affinity chromatography, e.g. a poly-histidine tag permits purification on nickel chelate media (see, for example, ref. 140). Although there are few examples of attempts to crystallize membrane proteins as fusion products (143), it is a conceivable route to obtaining crystals.

7 Useful websites Many websites can be found which give useful advice on crystallization and crystallography. A few we have found useful are listed below; along with several company sites detailing chemicals (especially detergents) and equipment necessary to crystallize a protein. Enrico Stura's offers a great deal of advice on all aspects of crystallization. Hampton Research offers practical tips, lists of useful references, conferences, etc., in addition to product information, and the site is regularly updated. • http://bmbsgil3.leeds.ac.uk/wwwprg/stura/cryst.html (for Enrico Stura's European site, numerous screens, detergent details, links, etc.); • http://www-structure.llnl.gov/Xray/101index.html (for Bernhard Rupp's introductory course to crystallography, both practical and theoretical); • http://www.sdsc.edu/pb/xtal/vl/ (for numerous aspects of crystallography); • http://www-bioc.rice.edu/~berry/papers/crystallization/crystallization.html An Internet search will produce hundreds more! Company websites • • • • • •

302

http://www.anatrace.com http://www.bachem.com http://biochem.roche.com http://www.calbiochem.com http://www.dojindo.co.jp http://www.douglas.co.uk (for consumables, automation/crystallization robots, useful links)

CRYSTALLIZATION OF MEMBRANE PROTEINS

• http://www.hamptonresarch.com (for lots of crystallization tips and useful references) • http://www.dialspace.dial.pipex.com (for NBS Biologicals) • http://www.spectrumlabs.com • http://www.stratech.co.uk

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List of suppliers

Accurate Chemical and Scientific Co., 300 Shames Drive, Westbury, NY 11590, USA. Web site: www.accurate-assi-leeches.com AEA Technology, Harwell, Didcot, Oxon OX11 ORA, UK. Agar Scientific Ltd., 66a Cambridge Road, Stanstead, Essex CM24 8DA, UK. Tel: 01279 813519 Althin Medical Ltd., Unit 25, Science Park, Milton Road, Cambridge CB4 4FW, UK. Ambion, Inc., 2130 Woodward Street, Austin, TX 78744, USA. American Type Culture Collection (ATTC), 10801 University Blvd, Manassas, VA 20110-2209, USA. Amersham Pharmacia Biotech Amersham Pharmacia Biotech, Amersham Place, Little Chalfont, Buckinghamshire HP7 9NA, UK. Tel: 0800 515 313 Web site: www.apbiotech.com Amersham Pharmacia Biotech, 800 Centennial Avenue, PO Box 1327, Piscataway, NJ 08855, USA. Web site: www.apbiotech.com

Amicon Amicon Ltd., Upper Mill, Stonehouse, Gloucestershire GL10 2BJ, UK. Amicon Inc., 72 Cherry Hill Drive, Beverley, MA 01915, USA. Amicon, Millipore Corporation, 80 Ashby Road, Bedford, MA 01730, USA. Web site: www.millipore.com/analytical/amicon/index.html Aminco SLM-Aminco, 810 West Anthony Drive, Urbana, IL 61801, USA. DG Electronics, 16/20 Camp Road, Farnborough, Hampshire GU14 6EW, UK. Tel: 01252 373074 Fax: 01252 517783 Anatrace, 434 West Dussel Drive, Maumee, OH, 43537, USA. Anderman and Co. Ltd., 145 London Road, Kingston-upon-Thames, Surrey KT2 6NH, UK. Tel: 0181 541 0035 Fax: 0181 541 0623 Applied Photophysics Ltd., 203-205 Kingston Road, Leatherhead, Surrey KT22 7PB, UK. Tel: 01372 386537 Fax: 01372 386477 Avanti Polar-Lipids, 700 Industrial Park Drive, Alabaster, AL 35007, USA. Avestin Inc., PO Box 8530, Ottawa, ON, KlG 3H9, Canada.

309

LIST OF SUPPLIERS

Axon Instruments, 1101 Chess Drive, Foster City, CA 94404, USA. Bachem (UK), 69 High Street, Saffron Walden, Essex, CB10 1AA, UK. Baxter Health Care Corp., Thetford, Norfolk, UK. BDH (Merck Ltd.), Hunter Boulevard, Magna Park, Lutterworth, Leicestershire LE17 4XN, UK.

Bio 101 Inc. Bio 101 Inc., c/o Anachem Ltd, Anachem House, 20 Charles Street, Luton, Bedfordshire LU2 OEB, UK. Tel: 01582 456666 Fax: 01582 391768 Web site: www.anachem.co.uk Bio 101 Inc., PO Box 2284, La Jolla, CA 92038-2284, USA. Tel: 001 760 598 7299 Fax: 001 760 598 0116 Web site: www.biol01.com Bio-Rad Laboratories Ltd.

BD Pharmingen, 10975 Torreyana Road, San Diego CA 92121, USA. Beckman Coulter Inc.

Sectarian Coulter Inc., 4300 N. Harbor Boulevard, PO Box 3100, Fullerton, CA 92834-3100, USA. Tel: 001 714 871 4848 Fax: 001 714 773 8283

Web site: www.beckman.com Beckman Coulter (UK) Ltd., Oakley Court, Kingsmead Business Park, London Road, High Wycombe, Buckinghamshire HP11 1JU, UK. Tel: 01494 441181 Fax: 01494 447558 Web site: www.beckman.com Becton Dickinson and Co.

Becton Dickinson and Co., 21 Between Towns Road, Cowley, Oxford 0X4 SLY, UK. Tel: 01865 748844 Fax: 01865 781627 Web site: www.bd.com Becton Dickinson and Co., 1 Becton Drive, Franklin Lakes, NJ 07417-1883, USA. Tel: 001 201 847 6800 Web site: www.bd.com Bibby Sterilin, Tilling Drive, Stone, Staffordshire ST15 OSA, UK. Tel: 01785 812121 Fax: 01785 811064 310

Bio-Rad Laboratories Ltd, Bio-Rad House, Maylands Avenue, Hemel Hempstead, Hertfordshire HP2 7TD, UK. Tel: 0181 328 2000 Fax: 0181 328 2550 Web site: www.bio-rad.com Bio-Rad Laboratories Ltd., Division Headquarters, 1000 Alfred Noble Drive, Hercules, CA 94547, USA. Tel: 001 510 724 7000 Fax: 001 510 741 5817 Web site: www.bio-rad.com Bio-Whittaker UK Ltd., 1 Ashville Way, Wokingham, Berkshire RG41 2PL, UK. Biogenex, 4600 Norris Canyon Road, San Ramon, CA 94583, USA. Biomen Diagnostics, Pentos House, Falcon Business Park, Ivanhoe Road, Finchampstead, Berkshire RG40 4QQ, UK. BioMerieux Instruments et reactifs de laboratoire, 69280 Marcy-l'Etoile, France. Biosym/MSI Web site: www.msi.com Boehringer-Mannheim (now Roche Molecular Biochemicals) Boehringer-Mannheim, Bell Lane, Lewes, East Sussex BN7 1LG, UK. Boehringer, 9115 Hague Road, Indianapolis, IN 46250, USA.

LIST OF SUPPLIERST

BPL Bio Products, Dagger Lane, Elstree, Hertfordshire WD6 3BX, UK. Braun B Medical Ltd., 13-14 Farnborough Close, Aylesbury Vale Industrial Park, Stocklake, Aylesbury, Buckinghamshire HP20 1DP, UK. Tel: 01298 393900 B. Braun-Melsungen, Carl Braun Strasse 1, D-34212 Melsungen, Germany. Tel: +49 5661 710 Fax:+49 5661 711290 British BioCell International Ltd., Golden Gate, Ty Glas Avenue, Cardiff CF4 5DX, UK. Tel: +44 (0) 1222 747232 Caframo, Wiarton, Ontario NOH 2TO, Canada. Calbiochem-Novabiochem (UK), Boulevard Industrial Park, Padge Road, Beeston, Nottingham, NG9 2JD, UK. Cambridge Bioscience, 24-25 Signet Court, Newmarket Road, Cambridge CBS SLA, UK. Cambridge Repetition Engineers, Greens Road, Cambridge, CB4 3EQ, UK.

CP Instrument Co. Ltd., PO Box 22, Bishop Stortford, Hertfordshire CM23 3DX, UK. Tel: 01279 757711 Fax: 01279 755785 Web site: www.cpinstrument.co.uk Dagan Corporation, 2855 Park Avenue, Minneapolis, MN 55407, USA. Dako Dako Ltd, Denmark House, Angel Drove, Ely, Cambridge CB7 4ET, UK. Dako Corp., 6392 Via Road, Carpinteria, CA 93013, USA. Diachem International Ltd., Unit 5, Gardiners Place, West Gillibrands, Skelmersdale, Lancashire WN8 9SP, UK. Diatome Ltd., 2501 Bienne, PO Box 551, Switzerland. Dojindo, Kumamoto Techno Research Park, Tabaru 2025-5, Mashiki-machi, Kamimashiki-gun, Kumamotot, 861-22, Japan. (UK Distributors NBS Biologicals) Douglas instruments, 25 Thames House, 140 Battersea Park Road, London SW11 4MB, UK.

CellPro, St-Pietersplein 11/12, B-1970 Wezembeek-Oppem, Belgium.

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Cobe International, Blood Component Technology, Mercuriusstraat 30, 1930 Zaventum, Belgium. Corning Inc., Science Products Division, 45 Nagog Park, Acton, MA 01720, USA. Web site: www.corningcostar.com Costar Corning Costar Corporation, One Alewife Center, Cambridge, MA 02140, USA.

Dupont Dupont (UK) Ltd, Industrial Products Division, Wedgwood Way, Stevenage, Hertfordshire SGI 4QN, UK. Tel: 01438 734000 Fax: 01438 734382 Web site: www.dupont.com Dupont Co. (Biotechnology Systems Division), PO Box 80024, Wilmington, DE 19880-002, USA. Tel: 001 302 774 1000 Fax: 001 302 774 7321 Web site: www.dupont.com

311

LIST OF SUPPLIERS

Eastman Chemical Co., 100 North

Gelman Sciences, Pall Gelman Laboratory,

Eastman Road, PO Box 511, Kingsport, TN 37662-5075, USA. Tel: 001 423 229 2000

MI, USA.

Web site: www.eastman.com Eastman Kodak Co., Rochester, NY 14650, USA Tel: 1-800-225-5352

Electron Microscopy Sciences, 321 Morris Road, Box 251, Fort Washington, PA 19034, USA.

Genetics Computer Group Inc.

Web site: www.gcg.com Greiner Labortechnik Ltd, Brunei Way, Stroudwater Business Park, Stonehouse, Gloucester GL10 3SX, UK. Tel: 01453 825255 Hamilton Company, PO Box 10030 Reno, NV 89520, USA.

Hampton Research, 25431 Cabot Road, Suite 205, Laguna Hills, CA 92653-5527,

Elga Ltd, Lane End, High Wycombe, Buckinghamshire HP14 3JH, UK.

USA.

Elsevier Biosoft, 68 Hills Road, Cambridge CB2 1LA, UK

Harvard Apparatus Inc., 84 October Hill Road, Holliston, MA 01746, USA.

Fisher Scientific

Hellma (England) Ltd., Cumberland House, 24-28 Baxter Avenue, Southend-onsea, Essex SS2 6H2, UK.

Fisher Scientific UK Ltd, Bishop Meadow Road, Loughborough, Leicestershire LE11 5RG, UK. Tel: 01509 231166 Fax: 01509 231893 Web site: www.fisher.co.uk Fisher Scientific, Fisher Research, 2761 Walnut Avenue, Tustin, CA 92780, USA. Tel: 001 714 669 4600 Fax: 001 714 669 1613 Web site: www.fishersci.com Fluka Ruka, PO Box 2060, Milwaukee, WI 53201, USA. Tel: 001 414 273 5013 Fax: 001 414 2734979 Web site: www.sigma-aldrich.com Ruka Chemical Co. Ltd, PO Box 260, CH-9471, Buchs, Switzerland. Tel: 0041 81 745 2828 Fax: 0041 81 756 5449 Web site: www.sigma-aldrich.com FMC Byproducts, BioWittaker Molecular Applications Aps, Risingevej 1, DK-2665 Vallensbaek, Strand Denmark

312

Hybaid Hybaid Ltd, Action Court, Ashford Road, Ashford, Middlesex TW15 1XB, UK. Tel: 01784 425000 Fax: 01784 248085 Web site: www.hybaid.com Hybaid US, 8 East Forge Parkway, Franklin, MA 02038, USA. Tel: 001 508 541 6918 Fax: 001 508 541 3041 Web site: www.hybaid.com HyClone Laboratories, 1725 South HyClone Road, Logan, UT 84321, USA. Tel: 001 435 753 4584 Fax: 001 435 753 4589 Web site: www.hyclone.com Inject + Matic, 4, Pictet-de-Rochemont 1207, Geneve, Switzerland. Invitrogen Invitrogen BV, PO Box 2312, 9704 CH Groningen, The Netherlands. Tel: 00800 5345 5345 Fax: 00800 7890 7890

Web site: www.invitrogen.com

LIST OF SUPPLIERS

Invitrogen Corp., 1600 Faraday Avenue, Carlsbad, CA 92008, USA. Tel: 001 760 603 7200 Fax: 001 760 603 7201 Web site: www.invitrogen.com Jasco (UK) Ltd, Unit 14 (Zone D), Chelmsford Road Industrial Estate, Great Dunmow, Essex CM6 1XG, UK. Jasco Europe s.r.l, via Confalonieri 25, 22060 Cremalla (Co), Italy. Kinematica AG, Luzernerstrasse 147a, CH-6014 Littan-Luzern, Switzerland. Tel: 41 57 1257 Kirkegaard and Perry Laboratories Inc. (KPL), 2 Cessna Court, Gaithersburg, MD 20897, USA. Web site: www.kpl.com Kore Technology Ltd., 291 Cambridge Science Park, Milton Road, Cambridge CB4 ONF, UK. Lelca, Davy Avenue, Knowlhill, Milton Keynes MK5 8LB, UK. Tel: 01908 666663 Web site: www.leica.com Life Technologies Life Technologies ltd, PO Box 35, Free Fountain Drive, Incsinnan Business Park, Paisley PA4 9RF, UK. Tel: 0800 269210 Fax: 0800 838380 Web site: www.lifetech.com Life Technologies Inc., 9800 Medical Center Drive, Rockville, MD 20850, USA. Tel: 001 301 610 8000 Web site: www.lifetech.com Llpex Biomembranes Inc., 3550 West nth Ave, Vancouver, British Columbia V6R 2K2, Canada. Melford Laboratories Ltd., Bildeston Road, Chelsworth, Ipswich, Suffolk IP7 7LE, UK.

Merck Ltd., Merck House, Poole, Dorset BH15 1TD, UK. Tel: 0800 223344 Web site: www.merck-Ltd.co.uk Merck Sharp & Dohme Merck Sharp & Dohme Research Laboratories, Neuroscience Research Centre, Terlings Park, Harlow, Essex CM20 2QR, UK. Web site: www.msd-nrc.co.uk MSD Sharp and Dohme GmbH, Undenplatz 1, D-85540, Haar, Germany. Web site: www.msd-deutschland.com Millipore Millipore (UK) Ltd, The Boulevard, Blackmoor Lane, Watford, Hertfordshire WD1 8YW, UK. Tel: 01923 816375 Fax: 01923 818297 Web site: www.millipore.com/local/UK.htm Millipore Corp., 80 Ashby Road, Bedford, MA 01730, USA. Tel: 001 800 645 5476 Fax: 001 800 645 5439 Web site: www.millipore.com Nanoprobes Inc., 25 E. Loop Road, Sye. 124, Stony Brook, NY 11790-3350, USA. Nasco Inc., 901 Janesville Avenue, Fort Atkinson, WI 53538, USA. NBS Biologicals, 14 Tower Square, Huntingdon, Cambridgeshire, PE18 7DT, UK. NEN™, Life Science Products, 549-3 Albany St, Boston, MA 02118, USA. Web site: www.nenlifesci.com New Brunswick New Brunswick Scientific (UK) Ltd., Edison House, 163 Dixons Hill Road North Mymms, Hartfield Hertfordshire AL9 7JE, UK. New Brunswick Scientific Co. Inc., 44 Talmadge Road, Edison, NJ 08818, USA. New England Biolabs, 32 Tozer Road, Beverley, MA 01915-5510, USA. Tel: 001 978 927 5054 313

LIST OF SUPPLIERS

Nikon Corp. Nikon Corp., Fuji Building, 2-3, 3-chome, Marunouchi, Chiyoda-ku, Tokyo 100, Japan. Tel: 00813 3214 5311 Fax: 00813 3201 5856 Web site: www.nikon.co.jp/main/index_e.htm

Pharmacia Pharmacia Biotech (Biochrom) Ltd, Unit 22, Cambridge Science Park, Milton Road, Cambridge CB4 OFJ, UK. Tel: 01223 423723 Fax: 01223 420164 Web site: www.biochrom.co.uk

Nikon Inc., 1300 Walt Whitman Road, Melville, NY 11747-3064, USA. Tel: 001 516 547 4200 Fax: 001 516 547 0299 Web site: www.nikonusa.com

Pharmacia and Upjohn Ltd., Davy Avenue, Knowlhill, Milton Keynes, Buckinghamshire MK5 8PH, UK. Tel: 01908 661101 Fax: 01908 690091 Web site: www.eu.pnu.com

Novagen, 601 Science Drive, Madison, WI 53711, USA. Novo Nordisk A/S, Novo Alle, 2880 Bagsvaerd, Denmark. Tel: +45 4444 8888 Fax: +45 4449 0555 Nycomed Nycomed Amersham pic, Amersham Place, Little Chalfont, Buckinghamshire HP7 9NA, UK. Tel: 01494 544000 Fax: 01494 542266 Web site: www.amersham.co.uk Nycomed Amersham, 101 Carnegie Center, Princeton, NJ 08540, USA. Tel: 001 609 514 6000 Web site: www.amersham.co.uk Nycomed AS Pharma, Diagnostic Division, PO Box 4284 Torshov, N-0401 Oslo, Norway.

Pharmacia, 23 Grosvenor Road, St Albans, Hertfordshire All 3AW, UK. Pharmingen (distributed by Cambridge Bioscience) Pierce Pierce & Warriner (UK) Ltd., 44, Upper Northgate Street, Chester, Cheshire, CHI 4EF, UK. Web site: www.piercenet.com Pierce, 3747 N. Meridian Road, PO Box 117, Rockford, IL 61105, USA. Pierce Europe, PO Box 1512, 3260 BA OudBeijerland, The Netherlands. Pierce Chemical Co., 3747 N. Meridian Road, Rockford, IL 61105, USA.

Ortho Diagnostic Systems, PO Box 653, Enterprise House, Station Road, Loudwater, Buckinghamshire HP10 9XH, UK.

Polaroid Polaroid Europe Ltd, 3 Furzeground Way, Stockley Park, Uxbridge, Middlesex UB11 1DW, UK.

Oxoid Ltd, Basingstoke, Hampshire, UK.

Polaroid Corporation, Technology Square, Cambridge, MA 02139, USA.

Perkin Elmer Ltd, Post Office Lane, Beaconsfield, Buckinghamshire HP9 1QA, UK. Tel: 01494 676161 Web site: www.perkin-elmer.com PerSeptive Biosystems Inc., 500 Old Connecticut Path, Framingham, MA 01701, USA. Web site: www.pbio.com

314

Polysciences, 400 Valley Road, Warrington PA 18976, USA. Poretics Corporation, ill Lindbergh Avenue, Livermore, CA 94550-9261, USA. Promega Promega UK Ltd, Delta House, Chilworth Research Centre, Southampton SO16 7NS, UK.

LIST OF SUPPLIERS

Tel: 0800 378994 Fax: 0800 181037 Web site: www.promega.com

Savant InstrumentsInc., 100 Colin Drive, Holbrook, NY 11741, USA.

Promega Corp., 2800 Woods Hollow Road, Madison, WI 53711-5399, USA. Tel: 001 608 274 4330 Fax: 001 608 277 2516 Web site: www.promega.com

Schleicher and Schuell Inc., Keene, NH 03431A, USA. Tel: 001 603 357 2398

Qiagen Qiagen UK Ltd, Boundary Court, Gatwick Road, Crawley, West Sussex RH10 2AX, UK. Tel: 01293 422911 Fax: 01293 422922 Web site: www.qiagen.com Qiagen Inc., 28159 Avenue Stanford, Valencia, CA 91355, USA. Tel: 001 800 426 8157 Fax: 001 800 718 2056 Web site: www.qiagen.com Quantum Biotechnologies Inc., 1801 de Maisonneuve Blvd. West, 8th Floor, Montreal (Quebec), Canada H3H 1J9. Rayonet, The Southern New England Ultraviolet Company, Hamden, Connecticut 06514, USA. Roche Diagnostics Roche Diagnostics Ltd., Bell Lane, Lewes, East Sussex BN7 1LG, UK. Tel: 01273 484644 Fax: 01273 480266 Web site: www.roche.com Roche Diagnostics Corp., 9115 Hague Road, PO Box 50457, Indianapolis, IN 46256, USA. Tel: 001 317 845 2358 Fax: 001 317 576 2126 Web site: www.roche.com Roche Diagnostics GmbH, Sandhoferstrasse 116, 68305 Mannheim, Germany. Tel: 0049 621 759 4747 Fax: 0049 621 759 4002 Web site: www.roche.com Sarstedt, Aktiengeselischaft & Co., D-51588 Numbrecht, Germany.

Serotec Ltd., 22 Bankside, Station Approach, Kidlington, Oxford OX5 1JE, UK. Shandon Scientific Ltd., 93-96 Chadwick Road, Astmoor, Runcorn, Cheshire WA7 1PR, UK. Tel: 01928 566611

Web site: www.shandon.com Sigma-Aldrich Sigma-Aldrich Co. Ltd., Fancy Road, Poole, Dorset BH12 4QH, UK. Tel: 01747 822211 Fax: 01747 823779 Web site: www.sigma-aldrich.com Sigma-Aldrich Co. Ltd, The Old Brickyard, New Road, Gillingham, Dorset XP8 4XT, UK. Tel: 01202 722114 Fax: 01202 715460 Web site: www.sigma-aldrich.com Sigma Chemical Co., PO Box 14508, St Louis, Mo 63178, USA. Tel: 001314 771 5765 Fax: 001 314 771 5757 Web site: www.sigma-aldrich.com Singer Instrument Co. Ltd., Roadwater, Watchet, Somerset, TA23 ORE, UK. Sorvall Centrifuges (distributors): MediTech International, Inc., 2924 NW 109th Avenue, Miami, FL 33172, USA. Web site: www.sorvall.com Spectrum Laboratories Inc., 23022 La Cadena Drive, Laguna Hills, CA 92653, USA. (Available in UK from NBS Biologicals) Spectrum Medical Industry Inc., 60916 Terminal Annex, Los Angeles, CA 90054, USA. 315

LIST OF SUPPLIERS

SPSS Inc., 233 S. Wacker Drive llth Floor, Chicago, Illinois 60606, USA. Tel: 001 312 651 3000 Fax: 001 312 651 3668 Stedim, Z.I. des Paluds, BP1051-13781, Aubagne, France. Stratagene Stratagene Europe, Gebouw California, Hogehilweg 15,1101 CB Amsterdam Zuidoost, The Netherlands. Tel: 00800 9100 9100 Web site: www.stratagene.com Stratagene Inc., 11011 North Torrey Pines Road, La Jolla, CA 92037, USA. Tel: 001 858 535 5400 Web site: www.stratagene.com Stratech Scientific Ltd., 61-63 Dudley Street, Luton, Bedfordshire LU2 ONP, UK.

Troplx, 47 Wiggins Avenue, Bedford, MA 01730, USA. Web site: www.tropix.com Ultrawave Ltd., 1 Oxford Street, Cardiff, CF2 1YY, UK. United States Biochemical, PO Box 22400, Cleveland, OH 44122, USA. Tel: 001 216 464 9277 Vector Vector, 30 Ingold Road, Burlingame, CA 94010, USA. Vector, 16 Wulfric Square, Bretton, Peterborough PE3 8RF, UK. Wallac UK, Milton Keynes, UK. Wallac Inc., 9238 Gaither Road, Gaithersburg, MD 20877, USA. Web site: www.wallac.com

Minerva House, Calleva Park, Aldermaston, Berkshire RG7 SNA, UK. Tel: 0118 981 7775

Western Laboratory Service Ltd., Unit 8, Redan Hill Estate, Redan Road, Aldershot, Hampshire, UK. Tel: 01252 312128

Tetra, TetraWerke Dr., rer., nat, Ulrich Baensch GmbH, D 49304, Melle, Germany.

Worthington Biochemical Corp., Halls Mill Road, Freehold, NJ 07728, USA.

The Binding Site Ltd., PO Box 4073, Birmingham B29 6AT, UK.

Xenopus I, 716 Northside, Ann Arbor, MI 48105, USA.

Therapeutic A. L. Centre, Oxford University, Old Road, Headington, Oxford OX3 7JT, UK.

Xenopus Ltd., Holmesdale Nursery, Mid Street, South Nutfield, Redhill, Surrey, RH1 4JY.UK.

TAAB Laboratories Equipment Ltd., 3

316

Index

Italic numbers denote reference to illustrations. ABC transporters 210, 217-20, 222-3, 224, 225 affinity chromatography of tagged proteins affinity tags used in yeast 94, 96, 97-8 biotinylated proteins, purification with immobilised avidin 97-8 his-tagged proteins, purification by nickel chelate affinity chromatography 152, 153,154; see also his-tag vectors ANS 168, 169, 185 ATPase, assay for activity 29-30; see also purification and reconstitution avidin, see affinity chromatography

bacteria, see Escherichia coli baculovirus expression system baculovirus life cycle 108-10 baculovirus promoters, choice for expression experiments 110 insect cells used in the baculovirus expression system cell lines suitable as hosts 118 membrane preparation

from insect cells 137-8 monolayer cultures of insect cells 118-20 post-translational modification of proteins by insect cells 115 storage and resuscitation of insect cell cultures 121-2 suspension cultures of insect cells 120-1 transport measurements in insect cells 136-7 membrane proteins successfully expressed using the baculovirus system 117 recombinant baculovirus amplification and storage 133-4 cell-lysis assay for estimation of viral litre 133 isolation of bacmid DNA 126-7 plaque assay for measurement of viral titre 130-2 plaque purification of recombinants 130-2 preparation by cotransfection of insect cells with baculovirus DNA and recombinant transfer vector 111-14, 127-30 preparation by transposonmediated recombination in E. colt 114-15, 116, 124-6

preparation in yeast 114 transfection of insect cells using calcium phosphate 129-30 using liposomal transfection reagents 127-9 transfer vectors 122-4 bicinchoninic acid, see protein assay biotin, see affinity chromatography of tagged proteins, photolabelling of glucose transporters

Ca2*, see ATPase, purification, reconstitution, transport chimaeras 75 circular dichroism spectroscopy 159-61 computer analysis, see prediction of membrane protein structure crystallization of membrane proteins three-dimensional (3-D) crystallization additives and amphiphiles, choice of 270, 290-1 artificial mother liquor 295-6 co-crystallization with antibody fragments 229, 301 cryoprotection of crystals 296-8 317

INDEX crystallization of membrane proteins (cant.) crystallization conditions published for membrane proteins 284, 286-7 crystallization using microdialysis 271, 272 crystallization using vapour diffusion 271, 272 detergents, choice of 285, 288-90, 292-3, 299 lipid cubic phases 229, 273 light-harvesting complex 2 (LH2) from

protein into lipid bilayers 231, 236-8 detergents, choice of 238-41 examples of 2-D crystals obtained via reconstitution of detergent-solubilised membrane 232-5 see also detergents, electron microscopy, phospholipids cytochalasin B 169, 181, 183; see also photolabelling of glucose transporters

Rhodopseudomonas acidophila 10050 274,

277-8, 289 packing arrangements in membrane protein crystals 299-301 pH, choice of 291, 294 precipitants, choice of 284-5 preparation of crystals for data collection 295-8 protein concentration, choice of 294-5 reaction centre (RC) from Rhodobacter sphaeroides

278-9, 281-4 screening protocols for membrane protein crystallization 298-9 temperature, choice of 294 websites dealing with crystallization and crystallography 302-3 two-dimensional (2-D) crystallization additives used in crystallization 238 crystallization using BioBeads for detergent removal 254-6 crystallization using dialysis for detergent removal 251-4 crystallization using dilution of solutions of protein, lipid and detergent 249-251 crystallization in native membranes 230-1 crystallization by reconstitution of purified

318

databases see sequence databases detergents exchange using Centricon concentrator device 242, 276-7 exchange using size-exclusion gel filtration 242-3 exchange using sucrosegradient centrifugation 243 monorneric detergent, measurement of concentration using falling-drop weight method 244, 2-45 using sitting-drop method 245 properties 22-5, 150-1, 239-41, 292-3 removal using Bio-Beads 36-7, 254-6 see also crystallization of membrane proteins electron crystallography, see electron microscopy, crystallization of membrane proteins (twodimensional (2-D) crystallization) electron microscopy grids, preparation of 258-9 negative staining of 2-D crystals 257-60 transmission electron microscopy of 2-D crystals 261

image analysis of 2-D crystals

261-5 Escherichia coli culture 144-7 expression vectors 143-4 host strains for expression 144 membrane preparation preparation of mixed membranes using water lysis 145, 148 separation of inner and outer bacterial membrane fractions 149-50 solubilisation of membranes 150-2 strategy for obtaining transporters by expression in E. coli 142 transport assays with reconstituted proteoliposomes 157-9 transporters successfully expressed in E. colt 142 see also reconstitution, affinity chromatography of tagged proteins expression systems, see baculovirus expression system, Escherichia coli, Pichia pastoris, Saccharomyces cerevisiae, Schizosaccharomyces pombe, Xenopus oocyte

expression system expression vectors, see baculovirus expression system, Escherichia coli,

his-tag vectors, Saccharomyces cerevisiae, Schizosaccharomyces pombe, Xenopus oocyte

expression system

fluorescence spectroscopy extrinsic probes 185 conformational changes in membrane transporters, monitoring using intrinsic protein fluorescence 167-9, 182-3 ligand binding to membrane transporters, monitoring

INDEX by intrinsic protein fluorescence equilibrium studies 169-72 kinetic studies 172-84 single-turnover experiments 183-4 stopped-flow fluorescence instrumentation 172-7 forskolin 170-2, 177, 178, 181, 182, 193, 196 Fourier transform infrared spectroscopy 160, 162, 163

sarcoplasmic reticulum from skeletal muscle 25-6 Xenopus oocyte membranes 72 yeast membranes 92, 94, 95-6 metal chelate affinity chromatography, see nickel chelate affinity chromatography mRNA, see RNA multiple sequence alignment 213-14

GalP 142, 146, 157-8, 159, 160, 168, 169, 170-2, 178, 181, 182 GLUT1 117, 169, 182-4, 186, 190-1, 195-202 GLUT4 194, 197-202, 205-6

nickel chelate affinity chromatography 152-4; see also his-tag vectors

Hill equation 16 his-tag vectors 122, 123; see also affinity purification of tagged proteins

kinetics of transport, see transport, kinetics LH2, see crystallization of membrane proteins, purification

mass spectrometry 161-2, 163-4, 165 membrane protein structure, see circular dichroism spectroscopy, crystallization of membrane proteins, Fourier transform infrared spectroscopy, prediction of membrane protein structure, topology of membrane transporters membranes, preparation E. cdli membranes 148-50 insect cell membranes 137-8 intestinal brush-border membrane vesicles 9

pH, measurement of intracellular value 17-18 phospholipids choice for 2-D crystallization 236-7 concentration, determination of ammonium ferrothiocyanate method 246 choline-containing lipids, enzymatic determination 245-6, 247-8 inorganic phosphate content 247 removal using phospholipase A2 256-7 stock solutions, preparation of 237-8 photolabelling of glucose transporters ATB-BMPA, method for labelling adipocytes, erythrocytes and myocytes 198-200 biotinylated photolabelling reagents 204-5 biotinylated GLUT4 detection by immunoprecipitation followed by Amdex™ streptavidin-HRP 206 detection using streptavidin precipitation 205 bis-hexoses 196-7

cytochalasin B 195-6 photoactivation methods 194-5 radiolabelled transporters detection following gel electrophoresis 197, 200, 203 immunoprecipitation 200-2 Pichia pastoris 101-02 prediction of membrane protein structure modelling transmembrane helices 217, 225 residue periodicity analysis 216-17, 223-6 trarismembrane helix prediction 214-17, 219, 221-2 protein assay, bicinchoninic acid (BCA) method 248 purification Ca2+-ATPase of sarcoplasmic reticulum 27-9 light-harvesting complex (LH2) from Rhodopseudomonas addophila 10050 275-6 reaction centre (RC) from Rhodobacter sphaeroides 280-1 see also affinity chromatography of tagged proteins

RC, see crystallization of membrane proteins, purification reconstitution Ca2t-ATPase into membrane fragments 30-2 into pre-formed large unilamellar vesicles 34-7 into vesicles by gel permeation chromatography 38-9 proteins expressed in E. cdli incorporation into liposomes using BioBeads 156-7 incorporation into liposomes by detergent dilution 155-6

319

INDEX RNA electrophoresis on denaturing agarose gels 58, 59 isolation of total RNA from mammalian tissues 53-4 purification of poly(A)* RNA by oligo(dT)-cellulose affinity chromatography 55-6 size-fractionation of poly(A)+ RNA 56-8 synthesis in vitro of capped RNA transcripts 61 see also Xenopus oocyte expression system

Saccharomyces cerevisiae affinity tags for purification or detection of proteins expressed in yeast 94, 96, 97-8 choice of DNA to be expressed 82-3 culture media 82 expression vectors 86-8, 91 host strains for expression 83-5 membranes small-scale preparation of total membranes 92 large-scale preparation of total and plasma membranes 94, 95-6 transformation 87, 89-90 transport assays 93-4 transporters successfully expressed in yeast 80-1 see also affinity chromatography of tagged proteins Schizosaccharomyces pombe expression vectors 99, 101 host strains for expression 98-9 transformation 99-100 transporters successfully expressed in yeast 80-1 sequence alignment, see multiple sequence alignment sequence databases ABC transporters as an

320

example of database analysis 217-19 protein and nucleotide sequence databases 210-13 scoring matrices 212 search algorithms 210-13 thermodynamics, see transport (thermodynamics) topology of membrane transporters 74-5; see also prediction of membrane protein structure transport kinetics activator/substrate stoichiometry 16 analysis of data from transport assays 15-16 co-transport systems 14-15 facilitated-diffusion systems 13-14 ion gradients and membrane potential 16-19 rate constants governing translocation cycle, determination from steady-state measurements of substrate flux at different temperatures 185-9 simulation of Ca2+ accumulation 41-4, 45 see also fluorescence spectroscopy, transport (thermodynamics) assay techniques adherent cell monolayers 3-5 Ca2+ accumulation by reconstituted vesicles, spectrophotometric determination 40 insect cells 136-7 membrane vesicles 8-13, 40, 157-9 suspended cells 5-8 Xenopus oocytes 63-8 yeast 93-4 thermodynamics 189-91

vectors, see baculovirus expression system, Escherichia coli, his-tag vectors, Saccharomyces cerevisiae, Schizosaccharomyces pombe, Xenopus oocyte expression system vesicles calculation of internal volume 39-40 large unilamellar 34-7 preparation of intestinal brush-border membrane vesicles 9 preparation of skeletal muscle sarcoplasmic reticulum 25-6 see also reconstitution

western blotting 73, 135

Xenopus oocyte expression system cDNA libraries for Xenopus expression preparation 58-60 screening 60-2 electrophysiological recordings 68-71 expression vectors 63, 64, 65 membranes, preparation from oocytes 72 microinjection 51-3 oocytes, preparation and maintenance 48-51 subcloning a cDNA into the vector pSP64T 64 transport assays 63-8 see also RNA x-ray crystallography, see crystallization of membrane proteins (three-dimensional (3-D) crystallization)

yeast see Saccharomyces cerevisiae, Schizosaccharomyces pombe, Pichia