Membrane Transport: A Practical Approach

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 ( 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,

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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

THE XENOPUS OOCYTE EXPRESSION SYSTEM

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

THE XENOPUS OOCYTE EXPRESSION SYSTEM

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 ( 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

BACULOVIRUS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

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

BACULOVIRLJS-MEDIATED OVEREXPRESSION OF TRANSPORT PROTEINS

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.

136

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.

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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.

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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)

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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.

163

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