Molecular Chaperones and Folding Catalysts: Regulation, Cellular Functions and Mechanisms

MOLECULAR CHAPERONES AND FOLDING CATALYSTS MOLECULAR CHAPERONES AND FOLDING CATALYSTS Regulation, Cellular Function a...

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MOLECULAR CHAPERONES AND FOLDING CATALYSTS

MOLECULAR CHAPERONES AND FOLDING CATALYSTS Regulation, Cellular Function and Mechanisms Edited by

Bernd Bukau Institute for Biochemistry and Molecular Biology University of Freiburg Germany

harwood academic publishers Australia • Canada • China • France • Germany • India Japan • Luxembourg • Malaysia • The Netherlands Russia • Singapore • Switzerland

This edition published in the Taylor & Francis e-Library, 2005. “To purchase your own copy copy of this or any of taylor & Francis or Routledge's collection of thousands of ebooks please go to www.eBookstore.tandf.co.uk.” Copyright © 1999 OPA (Overseas Publishers Association) N.V. Published by license under the Harwood Academic Publishers imprint, part of The Gordon and Breach Publishing Group. All rights reserved. No part of this book may be reproduced or utilized in any form or by any means, electronic or mechanical, including photocopying and recording, or by any information storage or retrieval system, without permission in writing from the publisher. Printed in Singapore. Amsteldijk 166 1st Floor 1079 LH Amsterdam The Netherlands British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library. Molecular chaperones and folding catalysts: regulation, cellular functions and mechanisms 1. Molecular chaperones 2. Protein folding I. Bukau, Bernd 572.6′45 ISBN 0-203-30375-X Master e-book ISBN

ISBN 0-203-34398-0 (Adobe eReader Format) ISBN: 90-5702-370-9 (Print Edition) The cover illustration shows schematically a 100 nm window of the cytoplasm of E. coli, depicting the macromolecular components with their estimated sizes. This cellular environment, in which assisted protein folding occurs, is characterized by extraordinary macromolecular crowding. This illustration is a modified version of Figure 1a reprinted from TIBS, Vol. 16, David Goodsell: “Inside a living cell”, pages 203–206, 1991, with permission from Elsevier Science.

To Anette

CONTENTS

Preface Contributors

xi xiii

I. INTRODUCTION 1. Assisted protein folding B.Bukau , F.X.Schmid and J.Buchner

3

II. REGULATION 2. Autoregulation of the heat shock response in prokaryotes L.Connolly , T.Yura and C.A.Gross 3. Inducible transcriptional regulation of heat shock genes: The stress signal and the unfolded protein response R.I.Morimoto 4. Protein kinase cascades involved in heat shock protein expression and function O.Bensaude 5. Thermotolerance and stress response: Possible involvement of Ku autoantigen G.C.Li , L.Li , D.Kim , A.Nussenzweig , S.-H.Yang , P.Burgman , H.Ouyang and C.C.Ling

13 39

59 85

III. CELLULAR FUNCTIONS

A. Overview and physiological aspects 6. Genetic evidence for the roles of molecular chaperones in Escherichia coli metabolism W.F.Burkholder and M.E.Gottesman 7. Genetic dissection of the Hsp70 chaperone system of yeast E.Craig , W.Yan and P.James 8. Functions in development M.Morange

116

155 180

B. Assisted protein folding processes: From ribosomes to proteases 9. Early events in the synthesis and maturation of polypeptides W.J.Welch , D.K.Eggers , W.J.Hansen and H.Nagata 10. Protein transport into and folding within the endoplasmic reticulum I.G.Haas and R.Zimmermann 11. The role of molecular chaperones in transport and folding of mitochondrial proteins P.J.T.Dekker and N.Pfanner 12. Protein import into and folding within chloroplasts E.Muckel and J.Soll 13. Protein folding in the periplasm of Escherichia coli D.Missiakas , C.Dartigalongue and S.Raina 14. Role of chaperones in replication of bacteriophage lambda DNA M.Zylicz , A.Wawrzynow , J.Marszalek , K.Liberek , B.Banecki , I.Konieczny , A.Blaszczak , P.Barski , J.Jakóbkiewicz , M.Gonciarz-Swiatek , M.Duchniewicz , J.Puzewicz and J.Krzewska 15. Control of hormone receptor function by molecular chaperones and folding catalysts D.O.Toft 16. Role of chaperones in uncoating of clathrin coated vesicles E.Eisenberg and L.Greene 17. The role of Hsp104 in stress tolerance and prion maintenance S.Lindquist and E.C.Schirmer 18. Chaperones and charonins: Protein unfolding enzymes and proteolysis M.R.Maurizi , S.Wickner and S.Gottesman

196 226 260

291 310 325

346

365 384 421

IV. MECHANISMS

A. Overview 19. Spontaneous versus assisted protein folding R.Jaenicke and R.Seckler

448

B. Folding catalysts 20. Protein disulphide-isomerase: A catalyst of thiol:disulphide interchange and associated protein folding R.B.Freedman and P.Klappa 21. Peptidyl-prolyl cis/trans isomerases G.Fischer and F.X.Schmid

479

504

C. Chaperonins 22. The ATPase cycle of the GroE molecular chaperones N.A.Ranson and A.R.Clarke 23. The relationship between chaperonin structure and function S.G.Burston and H.R.Saibil 24. Composition and function of the eukaryotic cytosolic chaperonin-containing TCP-1 K.R.Willison

537 570 605

D. Chaperones 25. Structure and mechanism of Hsp70 proteins J.-H.Ha , E.R.Johnson , D.B.McKay , M.C.Sousa , S.Takeda and S.M.Wilbanks 26. The DnaK chaperone system: Mechanism and comparison with other Hsp70 systems A.Buchberger , J.Reinstein and B.Bukau 27. Mechanisms of ATP-independent vs. ATP-dependent chaperones S.Bose , M.Ehrnsperger and J.Buchner 28. Structure and function of periplasmic PapD-like chaperones involved in assembly of bacterial P pili S.J.Hultgren , D.L.Hung , C.H.Jones and S.Knight Index

625 663

693 722

747

PREFACE One of the most intriguing discoveries in molecular biology in the last decade is the existence of an evolutionary conserved and essential system, consisting of molecular chaperones and folding catalysts, which promotes the folding of proteins in the cell. This volume summarizes our current knowledge of the cellular roles, the regulation and the mechanism of action of this system. It has a broad scope, covering cell biological, genetic and biochemical aspects of protein folding in cells from bacteria to man. The first section provides an overview of the diverse families of molecular chaperones and catalysts and the general principles of their action. The second section discusses the regulation of chaperone gene expression in response to stress. The third section summarizes the roles of chaperones and catalysts in cell physiology, followed by a detailed description of their roles in the life span of proteins, from the de novo folding at translating ribosomes to the aggregation and proteolytic degradation of misfolded proteins. The fourth section presents a detailed discussion of our current knowledge on the mechanisms of action of chaperones and folding catalysts. This volume is aimed at researchers working in basic and applied aspects of molecular biology, biochemistry and molecular medicine, and should be useful as an up-to-date reference book and a textbook for specialized university courses. The editor would like to thank the authors for their contributions and their efforts to make this book as up to date as possible, and his secretary Patricia Müller for expert help in preparation of the manuscripts.

CONTRIBUTORS Bogdan Banecki Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Piotr Barski Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Olivier Bensaude Unité de Génétique Moléculaire Département de Biologie École Normale Supérieure 46 rue d’Ulm 75230 Paris Cedex 05 France Adam Blaszczak Polish Academy of Science Institute of Biochemistry and Biophysics Laboratory of Molecular Biology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Suchira Bose Department of Biochemistry University of Bristol School of Medical Sciences Bristol BS8 1TD UK Alexander Buchberger Centre for Protein Engineering Medical Research Council Centre Hills Road Cambridge CB2 2QH UK

Johannes Buchner Institut für Biophysik und Physikalische Biochemie Universität Regensburg Universitätsstr. 31 D-93040 Regensburg Germany Bernd Bukau Institut für Biochemie und Molekularbiologie Universität Freiburg Hermann-Herder-Str. 7 D-79104 Freiburg Germany P.Burgman Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA William F.Burkholder Department of Biochemistry and Molecular Biophysics Institute of Cancer Research College of Physicians and Surgeons Columbia University 701 W168 Street New York, NY 10032 USA Steven G.Burston Department of Genetics Boyer Center for Molecular Medicine Yale University School of Medicine 295 Congress Avenue New Haven, CT 06510 USA Anthony R.Clarke Department of Biochemistry School of Medical Sciences University of Bristol Bristol BS8 1TD UK Lynn Connolly Department of Biochemistry and Biophysics

University of California San Francisco, CA 94143 USA Elizabeth Craig Department of Biomolecular Chemistry University of Wisconsin 1300 University Avenue Madison, WI 53706 USA Peter J.T.Dekker Institut für Biochemie und Molekularbiologie Universität Freiburg Hermann-Herder-Str. 7 D-79104 Freiburg Germany Marlena Duchniewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Daryl K.Eggers Departments of Medicine and Physiology Lung Biology Research Center University of California Box 0854 San Francisco, CA 94143 USA Monika Ehrnsperger Institut für Biophysik und Physikalische Biochemie Universität Regensburg Universitätsstr. 31 D-93040 Regensburg Germany Evan Eisenberg Laboratory of Cell Biology National Heart, Lung, and Blood Institute Bethesda, MD 20892 USA

Gunter Fischer Max-Planck-Gesellschaft Arbeitsgruppe “Enzymologie der Peptidbindung” Weinbergweg 16a D-06120 Halle/Saale Germany Robert B.Freedman Research School of Biosciences University of Kent Canterbury Kent CT2 7NJ UK Malgorzata Gonciarz-Swiatek Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Max E.Gottesman Department of Biochemistry and Molecular Biophysics Institute of Cancer Research College of Physicians and Surgeons Columbia University 701 W168 Street New York, NY 10032 USA Susan Gottesman Laboratory of Molecular Biology National Cancer Institute Bethesda, MD 20892 USA Lois Greene Laboratory of Cell Biology National Heart, Lung, and Blood Institute Bethesda, MD 20892 USA Carol A.Gross Departments of Stomatology, and Microbiology and Immunology

University of California Box 0512, S534 San Francisco, CA 94143 USA Jeung-Hoi Ha Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA Ingrid G.Haas Institut für Biochemie I Universität Heidelberg Im Neuenheimer Feld 328 D-69120 Heidelberg Germany William J.Hansen Departments of Medicine and Physiology Lung Biology Research Center University of California Box 0854 San Francisco, CA 94143 USA Scott J.Hultgren Department of Molecular Microbiology Washington University School of Medicine 660 S. Euclid Avenue, Box 8230 St. Louis, MO 63110 USA Danielle L.Hung Department of Molecular Microbiology Washington University School of Medicine 660 S. Euclid Avenue, Box 8230 St. Louis, MO 63110 USA Rainer Jaenicke Institut für Biophysik und Physikalische Biochemie Universität Regensburg D-93040 Regensburg

Germany Joanna Jakóbkiewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Philip James Department of Biomolecular Chemistry University of Wisconsin 1300 University Avenue Madison, WI 53706 USA Eric R.Johnson Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA C.Hal Jones Department of Molecular Microbiology Washington University School of Medicine 660 S. Euclid Avenue, Box 8230 St. Louis, MO 63110 USA D.Kim Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA Peter Klappa Research School of Biosciences University of Kent Canterbury Kent CT2 7NJ UK Stefan Knight

Swedish University of Agricultural Sciences Uppsala Biomedical Center Department of Molecular Biology P.O. Box 590 S-751 24 Uppsala Sweden Igor Konieczny Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Joanna Krzewska Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland G.C.Li Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA L.Li Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA Krzysztof Liberek Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Susan Lindquist Howard Hughes Medical Institute Department of Molecular Genetics and Cell Biology

University of Chicago 5841 S.Maryland Avenue, MC 1028 Chicago, IL 60637 USA C.C.Ling Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA Jaroslaw Marszalek Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Michael R.Maurizi Laboratory of Cell Biology National Cancer Institute Bethesda, MD 20892 USA David B.McKay Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA Dominique Missiakas Centre National de Recherche Scientifique LIDSM-CBBM 31 Chemin Joseph-Aiguier 13402 Marseille Cedex 20 France Michel Morange Unité de Génétique Moléculaire Département de Biologie École Normale Supérieure 46 rue d’Ulm 75230 Paris Cedex 05

France Richard I.Morimoto Department of Biochemistry, Molecular Biology and Cell Biology Rice Institute for Biomedical Research Northwestern University 2153 Sheridan Road Evanston, IL 60208 USA Eva Muckel Botanisches Institut Christian-Albrechts-Universität Am Botanischen Garten 1–9 D-24118 Kiel Germany Hirsohi Nagata Departments of Medicine and Physiology Lung Biology Research Center University of California Box 0854 San Francisco, CA 94143 USA A.Nussenzweig Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA H.Ouyang Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA Nikolaus Pfanner Institut für Biochemie und Molekularbiologie Universität Freiburg Hermann-Herder-Str. 7 D-79104 Freiburg Germany

Joanna Puzewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland Satish Raina Centre Medical Universitaire Département de Biochimie Médicale 1 rue Michel-Servet CH-1211 Geneva 4 Switzerland Neil A.Ranson Department of Crystallography Birkbeck College Malet Street London WC1E 7HX UK Jochen Reinstein Abteilung Physikalische Biochemie Max-Planck-Institut für Molekulare Physiologie Rheinlanddamm 201 D-44139 Dortmund Germany Helen R.Saibil Department of Crystallography Birkbeck College Malet Street London WC1E 7HX UK Eric S.Schirmer Howard Hughes Medical Institute Department of Molecular Genetics and Cell Biology University of Chicago 5841 S. Maryland Avenue, MC 1028 Chicago, IL 60637 USA Franz X.Schmid

Laboratorium für Biochemie Universität Bayreuth D-95440 Bayreuth Germany Robert Seckler Institut für Biophysik und Physikalische Biochemie Universität Regensburg D-93040 Regensburg Germany Jürgen Soll Botanisches Institut Christian-Albrechts-Universität Am Botanischen Garten 1–9 D-24118 Kiel Germany Marcelo C.Sousa Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA Shigeki Takeda Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA David O.Toft Department of Biochemistry and Molecular Biology Mayo Clinic 200 1st Street SW/1601 Rochester, MN 55905 USA Alicja Wawrzynow Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24

Poland William J.Welch Departments of Medicine and Physiology Lung Biology Research Center University of California Box 0854 San Francisco, CA 94143 USA Sue Wickner Laboratory of Molecular Biology National Cancer Institute Bethesda, MD 20892 USA Sigurd M.Wilbanks Department of Structural Biology Stanford University School of Medicine Stanford, CA 94305–5400 USA Keith R.Willison Chester Beatty Laboratories Institute of Cancer Research 237 Fulham Road London SW3 6JB UK Wei Yan Department of Biomolecular Chemistry University of Wisconsin 1300 University Avenue Madison, WI 53706 USA S.-H.Yang Departments of Radiation Oncology and Medical Physics Memorial Sloan-Kettering Cancer Center 1275 York Avenue New York, NY 10021 USA Takashi Yura

HSP Research Institute Kyoto Research Park Kyoto 600 Japan Richard Zimmermann Medizinische Biochemie Universität des Saarlandes D-66421 Homburg Germany Maciej Zylicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80–822 Gdansk, Kladki 24 Poland

I. INTRODUCTION

1. ASSISTED PROTEIN FOLDING B.BUKAU1, * , F.X.SCHMID2 and J.BUCHNER3 1 Institut

für Biochemie und Molekularbiologie, Universität Freiburg, HermannHerder-Str. 7, D-79104 Freiburg, Germany 2 Laboratorium für Biochemie, Universität Bayreuth, D-95440 Bayreuth, Germany 3 Institut für Biophysik and Physikalische Biochemie, Universität Regensburg, 93040 Regensburg, Germany

1. Protein folding in vitro and in vivo 2. Classification of folding catalysts and molecular chaperones 2.1. Folding catalysts 2.2. Molecular chaperones 3. References

1. PROTEIN FOLDING IN VITRO AND IN VIVO The classical experiments by Anfinsen and others established that the entire information required for the folding of polypeptide chains to the native three-dimensional conformation is encoded in their primary amino acid sequences (Epstein et al., 1963). This information directs the formation of multiple non-covalent and covalent interactions within polypeptide chains and between subunits of protein oligomers which drive the folding process and stabilize the folded structures (chapter Seckler and Jaenicke). It also establishes a balance between structural stability and flexibility, achieved by low conformational stability of the folded protein (Privalov, 1979), which is required for the folding process itself and the activity of the folded protein. This balance implies that correct and incorrect folding, as well as native and nonnative structures, are separated by only relatively small energy barriers. Subtle changes in the amino acid sequence or the folding milieu may therefore have dramatic consequences for the folding process and the structural integrity of proteins. Misfolding of proteins is indeed the major damaging consequence of stress situations such as heat shock. Misfolded proteins frequently expose hydrophobic surfaces that are prone to intermolecular aggregation, a largely irreversible reaction. *Corresponding author

Molecular chaperones and folding catalysts

4

Cells require a system for the proofreading of protein conformations and the control and assistance of a multitude of folding processes for several reasons (see Figure 1 for overview on major folding reactions occuring in the life span of proteins). Proteins are synthesized vectorially, which may require mechanisms to coordinate synthesis with folding and to protect nascent chains from aggregation (chapter Welch et al.). Organellar and secretory proteins have to be translocated to their subcellular destinations prior to folding which necessitates mechanisms to coordinate folding with translocation and to assist the translocation process

Figure 1 Folding processes assisted by molecular chaperones and folding catalysts in vivo. Depicted are the major categories of folding processes that occur in vivo, starting with the folding of newly synthesized proteins (co- and post-translational) and ending with the degradation by cellular proteases. Molecular chaperones and/or folding catalysts have been implicated in all reactions shown.

itself (chapters Dekker and Pfanner; Muckel and Soll; Welch et al.; Haas and Zimmermann). For survival under stress, cells require an efficient conformational proofreading and repair system for misfolded proteins (chapters Lindquist and Schirmer; Li et al.; Maurizi et al.). The importance of this latter function is indicated by disease states such as amyloidoses and prions which result from the accumulation of aggregated protein (chapter Lindquist et al.; Horwich and Weissman, 1997; Lindquist, 1997; Prusiner, 1997; Thomas et al, 1995; Wetzel, 1996), and by the death of cells occuring upon inactivation of the repair system (chapters Connolly et al; Morimoto; Li et al; Craig et al; Burkholder and Gottesman). Intense research efforts in the past decade have led to the discovery of the evolutionary conserved families of molecular chaperones and folding catalysts which constitute the

Assisted protein folding

5

cellular system for folding and repair of proteins (see Table 1 for chaperones) (Buchner, 1996; Gething and Sambrook, 1992; Hartl, 1996). They assist the folding and targeting of newly synthesized proteins, prevent the aggregation of misfolded proteins, allow the refolding of kinetically trapped folding intermediates, mediate the translocation of proteins across membranes, assist the assembly and disassembly of protein complexes, play roles in proteolysis of unstable proteins, and even control the functional states of regulatory proteins. Members of different chaperone families and folding catalysts cooperate in folding reactions which led to the suggestion that assisted protein folding in vivo is promoted by a flexible network of folding helpers (Ehrnsperger et al, 1997; Bukau et al, 1996; Johnson and Craig, 1997).

2. CLASSIFICATION OF FOLDING CATALYSTS AND MOLECULAR CHAPERONES 2.1. Folding Catalysts The formation and isomerization of disulfide bonds and the cis-trans isomerizations of prolyl peptide bonds are slow and frequendy rate-limiting events in the folding of proteins. In vivo, these folding steps can be catalyzed by two classes of enzymes, known as protein disulfide isomerases or thiol/disulfide oxidoreductases (PDI) (chapter Freedman and Klappa) and peptidyl prolyl cis-trans isomerases (PPI) (Chapter Fischer and Schmid). PDIs are active in both the oxidized and the reduced form. In the oxidized form they introduce disulfide bonds into folding protein chains by direct thiol/disulfide exchange. In the reduced form they can attack existing disulfide bonds and thus isomerize incorrectly formed crosslinks. PDIs are localized in the endoplasmic reticulum of eukaryotic cells and the periplasm of bacteria where they are essential for disulfide bond formation in secreted proteins. All PDI proteins investigated share the catalytically active motif CysX-X-Cys in structurally related catalytic domains for which thioredoxin is the prototype. Despite this structural similarity there are striking differences within the PDI family with respect to the redox properties. Some PDI homologs, such as DsbA from E. coli, act as mere catalysts of disulfide bond formation, while others, such as eukaryotic PDI and E. coli DsbC catalyze both formation and isomerization of disulfide bonds very efficiently. These enzymes are typically composed of several thioredoxin-like domains which carry the catalytic thiol/disulfide exchange site as well as additional domains that mediate good binding to the substrate proteins. Peptidyl prolyl cis-trans isomerases catalyze the intrinsically slow rotation about XaaPro peptide bonds and thus accelerate folding reactions that are rate-limited by such isomerizations. Prolyl isomerases are abundant proteins and occur in virtually all organisms and cellular compartments. It is still unknown whether the catalysis of slow steps in protein folding is their major function. Considering the diversity and wide distribution of these enzymes it is almost certain that they are involved in many different cellular functions. The bacterial trigger factors were recently discovered to belong to the prolyl isomerases. They might, in fact, be prime candidates for ribosome-associated


6

folding enzymes that act very early in the life spans of proteins. 2.2. Molecular Chaperones The term “molecular chaperone” had been coined for a group of proteins which assist polypeptide folding in the cell. Chaperones seem to play multiple, housekeeping as well as stress related, roles in cell metabolism, including the folding and

Table 1 Conserved families of molecular chaperones and their co-chaperones1

Folding Prokaryotic Eukaryotic system Members Members

Functions

Hsp100

ClpA, ClpB, ClpX, ClpY

Hsp104, Hsp78

assistance of proteolysis of Maurizi et al.; unstable proteins (bacterial Lindquist and cytosol); prevention of aggregation Schirmer of misfolded proteins; disaggregation of misfolded proteins (eukaryotic cytosol)

Hsp90

HtpG

Hsp90, Grp94, ERp99, endoplasmin, Hsp108, gp96

prevention of aggregation and Bose et al; assistance of refolding of misfolded Toft proteins; regulation of activity of kinases and steroid hormone receptors

Hsp70

DnaK, HscA (Hsc66)

Hsp70, Hsc70, Ssa1–4, Ssb1, 2, Ssc, Ssh1, Lhs1, Kar2, BiP, Grp78

prevention of aggregation and Ha et al; assistance of refolding of misfolded Buchberger et proteins; folding of newly al; Craig et al. synthesized proteins (eukaryotic cytosol); activity control of regulatory proteins; translocation of precursors across membranes

DnaJ3

DnaJ, DjlA, CbpA, HscB

Hsp40, Ydj1, Sec63, Auxilin, CSPs, Mdj1, Hdj1, Hdj2

co-chaperone of Hsp70

Buchberger et al.

GrpE

GrpE

Mge1p

co-chaperone of Hsp70 (bacteria, mitochondria and chloroplasts)

Buchberger et al.

Functions

Book Chapters 2

Folding system

Prokaryotic Members

Eukaryotic Members

Book Chapters

sHSP

IbpA, IbpB

Hsp18.1, prevention of aggregation and Bose et al. Hsp25, Hsp27, assistance of refolding of -crystallin misfolded proteins

2


7

PapD

PapD

—

assembly of bacterial pili

SecB

SecB

—

prevention of folding and Welch et al. targeting of precursor proteins to translocase (bacteria)

Hsp47

—

Hsp47

folding and assembly of collagen

Bose et al.

Calnexin

—

Calnexin

folding of proteins in the ER

Haas and Zimmermann

Calreticulin

folding of proteins in the ER

Haas and Zimmermann

Calreticulin —

Hultgren et al.

Subfamily of Chaperonins HspGO

GroEL

Hsp60; Cpn60

prevention of aggregation and Burston and folding of newly synthesized Saibil; Ranson and misfolded proteins and Clarke (bacteria, mitochondria and chloroplasts)

Hsp10

GroES, gp31

Hsp10, Cpn10

co-chaperone of GroEL

Burston and Saibil; Ranson and Clarke

CCT

TF55

TRiC

folding of newly synthesized and misfolded proteins (eukaryotic cytosol)

Willison

1

Only selected members of each chaperone family are shown.

2 Only the chapters with the strongest focus on the particular chaperone are listed. 3 The DnaJ family consists of a large group of heterogeneous proteins with diverse metabolic

functions. DnaJ proteins share the J domain, a conserved fragment of approx. 78 residues, which is essential for interaction of DnaJ with Hsp70 proteins.

translocation of newly synthesized proteins, the refolding of conformationally damaged proteins, and the control of biological activity of specific regulatory proteins. Originally, the functional classification of chaperones was restricted to two classes of proteins, the Hsp70 and GroEL heat shock proteins, but is now used for an ever increasing number of proteins unrelated in primary sequence. Molecular chaperones are grouped into families on the basis of their evolutionary conservation. Many chaperones are designated according to their approximate molecular weight, e.g. the 70 kDa heat shock protein is a chaperone termed Hsp70. A constitutively expressed cognate is termed Hsc70, and other members of the Hsp70 chaperone family have kept the name provided to them in the context of their historical discovery (DnaK, BiP, SSA1 etc.). We cannot eliminate this confusing nomenclature but suggest to continue using the now established historic names (see Table 1). In view of the growing number of proteins designated as molecular chaperones it is rewarding to define the basic properties that a protein has to fulfill to qualify as a


8

chaperone. The most common definition for a molecular chaperone is that it assists the structure formation of proteins and prevents unproductive side reactions without becoming part of the final structure (Ellis and Hemmingsen, 1989; Ellis, 1987). Chaperones do not catalyze or accelerate folding reactions, but rather increase the number of molecules that are on a productive folding pathway. This activity relies on their ability to inhibit intermolecular aggregation reactions by reversible association with aggregation-prone folding intermediates. In addition, the subclass of ring-like chaperonins such as GroEL, is capable of unfolding protein substrates whereby they may allow kinetically trapped misfolded polypeptides to reenter the productive folding pathway. Chaperones share the ability to transiently associate with non-native conformers of proteins by recognizing exposed hydrophobic patches. There are, however, differences with respect to the molecular mechanism of substrate recognition, as illustrated for four major chaperones (Figure 2). Hsp70, in functional cooperation with DnaJ co-chaperones, is active as a monomer containing a single substrate binding site (chapters Ha et al.; Buchberger et al.). The segment of the substrate polypeptide that contacts Hsp70 is a short stretch of five consecutive residues in extended conformation that becomes enclosed by the chaperone. Tight binding appears to require that the interacting peptide segment is physically separated from the remainder of the substrate and therefore substantial, at least local unfolding. To qualify as substrate for Hsp70, a minimal requirement for a protein is to expose a single chaperone binding site. This mode of interaction explains the wide spectrum of protein conformers, which can associate with Hsp70 ranging from extended (e.g. nascent polypeptide chains) to native. Chaperonins such as the prokaryotic GroEL and the eukaryotic CCT form double rings, composed of 7 (GroEL) to 8 (CCT) subunits/ring, each ring containing a substrate binding site made up of segments from each subunit (chapters Burston and Saibil; Ranson and Clarke; Willison). The ring structure allows the simultaneous association of various segments of a polypeptide chain within one ring, and this feature is most likely a key property allowing chaperonins to unfold protein substrates before release. A broad range of conformers can associate with GroEL, but in contrast to Hsp70 there are no reports for native proteins that are natural substrates.

Figure 2 Topology of substrate binding by molecular chaperones. Shown are the major molecular chaperones and their modes of interaction with substrate polypeptides. The structural nature of the substrate binding sites of Hsp90 and sHSPs remains unclear. Black bars in substrate polypeptides represent hydrophobic segments that serve as binding


9

motifs for chaperones.

The conformation of the polypeptide segments that directly contact GroEL remains unclear. The small heat shock proteins (sHSPs) form oligomers with an average size of 12 to 42 subunits (chapter Bose et al.). Each oligomer can bind several protein substrates, up to one molecule per subunit, and thus serves as a very efficient binding scaffold for misfolded/unfolded substrates. Hsp90 acts as a dimer capable of binding non-native polypeptides (chapters Bose et al.; Toft). While for sHSPs and Hsp90 only little information exists with respect to the molecular basis of substrate recognition, recent data indicate that sHSPs and Hsp90 chaperones share with Hsp70 and GroEL the ability to recognize a broad range of conformations. The different chaperone families are thus not specialized for defined folding states of substrates, e.g. early unfolded or late molten globule-like states. Further differences between chaperone families exist with respect to the regulation of their functional activity. Some chaperones, including the sHSPs, Hsp47 and PapD, act independently of ATP (chapters Bose et al.; Hultgren et al.). It is somewhat mysterious how substrate binding is controlled in these cases. Yet unknown co-proteins or components of ATP-dependent chaperone systems may provide the cooperating partners for this class of chaperones. In contrast, the activity of major chaperones including Hsp70, chaperonins, Hsp90 and Hsp104/ClpB, is controlled by ATP and co-proteins (chapters Lindquist and Schirmer; Maurizi et al.; Ha et al.; Buchberger et al.; Burston and Saibil; Ranson and Clarke; Bose et al.). The role for ATP has been investigated in detail only for Hsp70 and GroEL. Hsp70 uses the energy of ATP to drive conformational changes that alter its affinity for substrates. The ATPase cycle of Hsp70 can be viewed, in its simplest form, as an alternation between two states: the ATP state with low affinity and fast exchange rates for substrates (substrate binding pocket open), and the ADP state with high affinity and low exchange rates for substrates (substrate binding pocket closed). GroEL uses ATP to drive coordinated conformational changes of all subunits of one ring, and subsequently in the other ring, which allow dissociation of substrates and ligands. ATP thus provides a mechanism to tightly control the activity of both chaperone systems, by affecting the kinetics of substrate binding and release. The ATPase activities of these chaperones are prime targets for regulatory proteins which either stimulate or inhibit checkpoints of the ATPase cycle and thereby control the affinity of the corresponding chaperone partner for substrates. Examples are the Hsp70 co-proteins DnaJ (Hsp40), GrpE, Hip and Bag1, and the GroEL co-proteins GroES and gp31. ATP-dependent chaperone systems are thus sophisticated and tightly regulated machines. The possibility to regulate their binding to substrate allows them at least in the case of Hsp70 to play diverse roles in cell metabolism, ranging from general functions in protein folding to highly specific functions e.g. in control of biological activities of regulatory proteins. Members of different chaperone families have been found in association with the same substrate conformer and capable of competing for binding. This principle of kinetic partitioning of substrates between different chaperones, and possibly folding catalysts and proteases, is likely to constitute the basis for a cellular network of folding helpers that assists protein folding (Ehrnsperger et al., 1997; Bukau et al., 1996; Johnson and Craig,


10

1997). Elucidation of the molecular principles and the biological implications of this network is a central goal for future research and will require the combined input of biochemistry, genetics and cell biology.

3. REFERENCES Buchner, J. (1996). Supervising the fold: functional principles of molecular chaperones. FASEB J. , 10, 10–19. Bukau, B., Hesterkamp, H. and Luirink, J. (1996). Growing up in a dangerous environment: a network of multiple targeting and folding pathways for nascent polypetides in the cytosol. Trends Cell Biol. , 6, 480–486. Ehrnsperger, M., Gräber, S., Gaestel, M. and Buchner, J. (1997). Binding of non-native protein to Hsp25 during heat shock creates a reservoir of folding intermediates for reactivation. EMBO J. , 16, 221–229. Ellis, J. (1987). Proteins as molecular chaperones. Nature (London), 328, 378–379. Ellis, R.J., and Hemmingsen, S.M. (1989). Molecular chaperones: proteins essential for the biogenesis of some macromolecular structures. Trends Biochem. Sci. , 14, 339–42. Epstein, C.J., Goldberger, R.F. and Anfinsen, C.B. (1963). The genetic control of tertiary protein structure: studies with model systems. Cold Spring Harb. Symp. Quant. Biol. , 28, 439–449. Gething, M.-J. and Sambrook, J.F. (1992). Protein folding in the cell. Nature , 355, 33– 45. Hartl, F.U. (1996). Molecular chaperones in cellular protein folding. Nature , 381, 571– 580. Horwich, A.L. and Weissman, J.S. (1997). Deadly conformations-protein misfolding in prion disease. Cell 89, 499–510. Johnson, J.L., and Craig, E.A. (1997). Protein folding in vivo: Unraveling complex pathways. Cell , 90, 201–204. Lindquist, S. (1997). Mad cows meet Psi-chotic yeast: the expansion of the prion disease. Cell , 89, 495–498. Privalov, P.L. (1979). Stability of proteins. Adv. Protein Chem. , 33, 167–241. Prusiner, S.B. (1997). Prion diseases and the BSE crisis. Science , 278, 245–251. Thomas, P.J., Qu, B.-H., and Pedersen, P.L. (1995). Defective protein folding as a basis of human disease. Trends Biochem. Sci. , 20, 456–459. Wetzel, R. (1996). For protein misassembly, it’s the “I” decade. Cell , 86, 699–702.

II. REGULATION

2. AUTOREGULATION OF THE HEAT SHOCK RESPONSE IN PROCARYOTES LYNN CONNOLLY1, TAKASHI YURA2 and CAROL A.GROSS3, * 1 Department

of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA 94143 2 HSP Research Institute, Kyoto Research Park, Kyoto 600, Japan 3 Departments of Stomatology and Microbiology and Immunology, University of California, San Francisco, CA 94143

1. Introduction 2. Regulation of the

heat shock response

2.1. Discovery of 2.2. How does

regulate the response to temperature shift?

2.3. Translational regulation of 2.4. Regulation of

stability

2.5. Regulation of

activity

2.6. What are the signals governing expression of the 3. Regulation of the

heat shock regulon?

(24) heat shock response

3.1. Discovery of 3.2. What is the nature of the signal inducing

activity?

3.3. Regulation of 3.4. How is the extracytoplasmic signal transduced to 3.5. The cellular role of 4. Heat shock regulation in other prokaryotic organisms 5. Summary and prospects 6. Acknowledgments 7. References *Corresponding author

?


14

1. INTRODUCTION When cells of any type are shifted to high temperature, the heat shock response (hsr) ensues and the synthesis of a small number of proteins, called the heat shock proteins (hsps), is rapidly induced. In E. coli, the hsr was discovered independently by the Neidhardt and Yura groups, who monitored the rate of synthesis of individual proteins after a temperature upshift using either 1D or 2D gels (Lemaux et al., 1978; Yamamori et al., 1978). A group of about 20 proteins exhibited a large (10 to 20-fold) but transient increase in synthetic rate upon temperature upshift and a corresponding decrease in synthetic rate upon temperature downshift (Lemaux et al., 1978; Yamamori et al., 1978; Neidhardt et al., 1987; Straus et al., 1989; Taura et al., 1989). This group of proteins comprises the E. coli hsps. Their expression is regulated at the transcriptional level (Yamamori et al., 1980; Taylor et al., 1984; Cowing et al., 1985) by the amount and/or activity of the alternative sigma factor, , which directs RNA polymerase to transcribe this set of genes (Lesley et al., 1987; Skelly et al., 1987; Straus et al., 1987). These hsps, including the chaperones DnaK-DnaJ and GroELGroES, are required for normal growth at physiological temperatures. Whereas E. coli in its natural habitat grows at temperatures between 25°C and 40°C, deletion of the gene encoding restricts growth to temperatures below 20°C (Zhou et al., 1988). Overexpression of the GroEL-GroES and DnaK-DnaJ chaperone machines restores high temperature growth, suggesting that these chaperones play a crucial role in adaptation to high temperature. E. coli also has a second heat-controlled regulon, controlled by ( ), another alternative sigma factor (Erickson et al., 1989; Wang et al., 1989) (see Missiakas and Raina, this volume). Many members of this regulon have yet to be identified. The two responses are intertwined because holoenzyme containing ( ) transcribes at extreme temperature. However, each response also has a distinct role in the cell: controlled genes respond to conditions in the cytoplasm of the cell whereas controlled genes respond to the extracytoplasmic state. The regulon plays an auxiliary role in temperature adaptation as cells lacking cannot grow at temperatures above 40° (Raina et al., 1995; Hiratsu et al., 1995; Rouvière et al., 1995). Such strains also exhibit defects in the cell envelope, emphasizing the dual role played by members of this regulon. The heat induction of several additional genes may occur by other mechanisms. controls genes involved in adaptation to stationary phase and is also somewhat induced upon shift to high temperature, suggesting that genes in the regulon exhibit temperature regulation (Hengge-Aronis, 1996). Finally, the psp operon is controlled by a dedicated activator protein that promotes psp transcription by EJ54 following shift to very high temperatures (Brissette et al., 1990). Two global approaches, one monitoring protein synthesis and the other monitoring RNA synthesis, have been used to identify most of the hsps. In the protein based approach, spots on 2D gels have been correlated with known genes (Georgopoulos et al., 1982; Neidhardt et al., 1981; Tilly et al., 1983). In the RNA based transcriptional mapping approach, radioactively labeled cDNA, made to total E. coli RNA, is hybridized

Autoregulation of the heat

15

to membrane filters containing an ordered E. coli genomic library carried in clones (the Kohara library) and clones whose transcription increases are identified (Chuang et al., 1993; Chuang et al., 1993). A compendium of the proteins whose rates of synthesis increase upon temperature upshift is presented in Table 1.

2. REGULATION OF THE

HEAT SHOCK RESPONSE

2.1. Discovery of The gene encoding was discovered in 1975 as a nonsense mutation that affected the synthesis of the GroEL hsp. The mutation was initially thought to be located in the structural gene for GroEL (Cooper et al., 1975). Subsequently, it was found

Table 1 Heat inducible proteins in Escherichia coli

Min Protein Alphanumeric designation

Molec. Function Weight

Kohara Physical Reference(s) Clones Map

21

?

?

Regulon .3

HtpY

.3

DnaK

B 066.0

69

chaperone

101, 102 11.7–15.5 [Bardwell et al., 1984]

.3

DnaJ

H 036.5

39

chaperone

101, 102 11.7–15.5 [Bardwell et al., 1986]

H 094.0

89

protease

148

464.1– 468.4

[Gayda et al., 1985]

24(22)

protease

148

464.1– 468.4

[Maurizi et al., 1990]

10.0 ClpX

46

chaperone

148

464.1– 468.4

[Gottesman et al., 1993]

10.0 HslA

65

?

70

chaperone

152

501.5– 504.2

[Bardwell et al., 1987]

80

?

212

921.6– 936.7

[Chuang et al., 1993]

35.5

dehydrogenase 330, 331 1872–

[Charpentier et

10.0 Lon

[Missiakas et al., 1993]

H 094.1 10.0 ClpP

10.8 HtpG

F 021.5

C 062.5

19.2 HslC 39.3 GapA

I 033.5



16

1873

al., 1987]

H 034.3 39.8 HslK

49

?

334

1901.2– 1904.2


40.3 HtpX

32

?

?

?

[Kornitzer et al., 1991]

437

2741.2– 2743.7

[Kitagawa et al., 1991; Squires et al., 1992]

56.0 ClpB

F 084.1 E 072.0

84

chaperone

56.8 GrpE

B 025.3

26

nucleotide 438, 439 2757.7– exchange factor 2763.6

[Lipinska et al., 1988]

67.0

B 082.0

70

sigma factor

509

3233.0– 3236.2

[Burton et al., 1981]

520

3331.7– 3350.3

[Herman et al., 1995; Tomoyasu et al., 1993]

3331.7– 3350.3

[Herman et al., 1995; Tomoyasu et al., 1993]

69.2 FtsJ

26

69.2 HflB

70

protease

520

75.0 HslO

33

?

620, 621 3549.6– 3552.0

75.0 HslP

30

?

81.2 HtrM (RfaD)

34

epimerase

575, 576 3815.3– 3816.4

[Raina et al., 1991]

[Chuang et al., 1993] [Chuang et al., 1993]

83.0 IbpB (HtpE, HslS)

C 014.7

16.3

chaperone

566, 567 3889.9– 3892.7

[Allen et al., 1992; Chuang et al., 1993]

83.0 IbpA (HtpN, HslT)

G 013.5

15.8

chaperone

566, 567 3889.9– 3892.7

[Allen et al., 1992; Chuang et al., 1993]

89.0 ClpY (HtpI, HslU))

D 048.5

49

chaperone

538, 539 4149.5– 4151.7

[Chuang et al., 1993, Missiakas et al., 1996]

89.0 HslV (HtpO)

G 021.0

21

protease

538, 539 4149.5– 4151.7

[Chuang et al., 1993, Missiakas et al., 1996]

21

?

60

chaperone

90.0 HtrC 94.2 GroEL

B 056.5

[Raina et al., 1990] 648, 649 4400.5–

[Hemmingsen et


17 4405.7

94.2 GroES

16

chaperone

648, 649 4400.5– 4405.7

[Hemmingsen et al., 1988]

94.2 HslW

22

?

648, 649 4400.5– 4405.7


94.8 HslX

51

?

652

4430.8– 4433.4


94.8 HslY

45

652

4430.8– 4433.4


94.8 HslZ

37

652

4430.8– 4433.4


HtpK

C 015.4

al., 1988]

F 010.1

Min Protein Alphanumeric designation

[Aa]

10

Molec. Function Weight

Kohara Physical Reference(s) Clones Map

Regulon HtpT

A 039.5

40

[Aa]

Regulon: 3.9

DegP (HtrA)

50

55.5

77.5

F 033.4

74.9 tkpA

protease

117, 118 181–182

[Lipinska et al., 1988; Strauch et al., 1989]

sigma factor

435

2718

[Lonetto et al., 1994; Nashimoto 1993; Raina et al., 1995]

sigma factor

613

3614– 3625

[Landick et al., 1984; Yura et al., 1984]

PPlase

625, 626

[Danese et al., 1997]

257, 258 1374– 1378

[Lipinska et al., 1988; Yamamori et al., 1982]

Others: 29.2 PspA

E 026.0

28

29.7 HslE

60

?

260

1388.8– 1409.9


29.7 HslF

51

?

260

1388.8– 1409.9


Molecular chaperones and folding catalysts 29.7 HslG

18

41

?

260

1388.8– 1409.9


36

?

265

1448.9– 1454.5


30.6 HslJ

14

?

265

1448.9– 1454.5


69.2 HslM

31

?

520

3331.7– 3350.3


75.0 HslQ

24

?

620, 621 3549.6– 3552.9


75.0 HslR

18

?

620, 621 3549.6– 3552.9


60

LysyltRNA synthetase

646, 647 4381.8– 4383.2

[Léveque et al., 1990]

30.6 HslI (HtpH)

93.5 LysU

D 033.4

D 060.5

that mutant cells had a global defect in the hsr, suggesting instead that the gene encoded a regulator of the hsr (Neidhardt et al., 1981; Yamamori et al., 1982). The sequence of the gene revealed strong homology to (Landick et al., 1984; Yura et al., 1984) and the regulator was shown to be , the first alternative sigma factor identified in E. coli (Grossman et al., 1984). directs core RNA polymerase to promoters that differ considerably from those recognized by RNA polymerase containing , the housekeeping sigma (Cowing et al., 1985). The fact that expression of the hsps is uniquely responsive to the amount or activity of provides a means to regulate their expression separately from other cellular proteins. 2.2. How Does

Regulate the Response to Temperature Shift?

When cells experience a temperature upshift, for example after shift from 30°C to 42°C, the rate of synthesis of the hsps increases 10 to 20-fold by 5 minutes after upshift and thereafter declines to a new steady state rate of synthesis. Interestingly, at steady state, the amount of hsps at 42° is only 2-fold greater than that at 30°. The large increase in rate of hsp synthesis immediately after temperature upshift allows cells to rapidly accumulate the new steady state level of hsps (Lemaux et al., 1978; Yamamori et al., 1978; Straus et al., 1987). The response of hsps to heat induction is controlled at the transcriptional level, primarily by the amount of in the cell. At low temperature, cells contain very little , on the order of 10 to 50 molecules per cell. By 5 minutes after temperature upshift, the amount of increases about 15-fold and thereafter declines to a new steady state level (Lesley et al., 1987; Straus et al., 1987). Changes in the amount of following temperature upshift result from changes in both the stability and synthesis of (Lesley et al., 1987; Straus et al., 1987). During steady state growth, is translated at a very


19

low rate. In addition, is very unstable, with a T½ for degradation of about 1 minute. As a result, little accumulates in the cell. However, for the first 5 minutes following temperature upshift the rate of translation of increases about 5-fold and is stabilized against degradation. Following this time, the rate of translation decreases and rapid degradation resumes. Together, these two regulatory changes permit the transient accumulation of . To a first approximation, changes in the rate of hsp synthesis after temperature upshift primarily mirror changes in the amount of (Lesley et al., 1987; Skelly et al., 1987; Straus et al., 1987). However, careful examination of the kinetics suggest that shutoff of hsp synthesis in the adaptation phase of the hsp response may slightly precede the decrease in the amount of . Regulation of activity (see below) may be involved in this phenomenon. When cells experience a temperature downshift, for example after shift from 42°C to 30°C, the rate of synthesis of hsps declines 10 to 20-fold within 5 minutes after downshift. This rate of hsp synthesis is considerably lower than that normally exhibited by cells growing at 30°C (Straus et al., 1989; Taura et al., 1989). By one to two doublings after downshift, the cell gradually resumes the 30°C rate of synthesis. Presumably, existing hsps are diluted out during the long shut-off period. Hsp synthesis resumes when their amounts approximate that characteristic of the cells growing continuously at low temperature. The rapid drop in transcription of heat shock genes upon temperature downshift results from a decrease in activity, rather than from a decrease in the amount of . Temperature downshift is not the only condition that promotes inactivation of . Overexpression of hsps at constant temperature also reduces activity, suggesting that cells can sense the amount of hsps and adjust the activity of accordingly (Straus et al., 1989; Craig et al., 1991). These studies indicate that the translation, stability and activity of are all regulated by the cell in response to temperature. The extent to which temperature regulation of each of these processes is understood at a mechanistic level is discussed below, and a speculative model of the regulation of activity is presented in Figure 1. 2.3. Translational Regulation of Translational regulation includes both translational induction, which occurs immediately following temperature upshift, and translational repression, which occurs


20

Figure 1 The promoters and translational regulatory regions of E. coli rpoH. (a) Regions A and B of the mRNA are involved in translational induction by modulating the secondary structure shown in (b), whereas region C of is involved in chaperone mediated translational repression and protein stability (see text). (b) A possible secondary structure of the mRNA formed under nonstress conditions. (Reproduced with permission from Yura, 1996).

subsequently during the adaptation phase of the hsr. The cis-elements and the transacting factors required for induction and repression differ, suggesting that these two processes


21

are mechanistically distinct. The mechanism of translational induction has been probed by both deletion and point mutational analysis of a - -galactosidase fusion protein (Kamath-Loeb et al., 1991; Nagai et al., 1991; Yuzawa et al., 1993). These studies indicate that two regions within , termed A and B, are required for translational induction (Figure 1). Region A, located near the start of translation initiation (nucleotide 6–20), has homology to the “downstream box”, which is required for high rates of translation in several prokaryotic systems. Deletion of the downstream box leads to very low, uninducible synthesis of . Region B is a grossly defined, internal region extending from nucleotide 110–210, part of which has the capacity to base pair with a portion of Region A. Deletion of Region B, as well as some point mutations in the region, leads to high constitutive synthesis of . Initial speculation that thermal induction might simply be explained by disruption of base-pairing potential between the two regions, led to an analysis of compensating mutational changes between putative base-pairing partners. These studies indicated that recovery of base pairing is not always sufficient for regulation, leading to the suggestion that sequence, as well as structure, is important for regulation (Yuzawa et al., 1993; Yura, 1996). The current view is that an unknown transacting factor is involved in this regulatory event. The mechanism of translational repression is distinct from that of translational induction. Translational repression requires Region C of (nucleotide 364–433; amino acid 122–144) and the DnaK, DnaJ, GrpE chaperone machine (Straus et al., 1990; Nagai et al., 1994). Deletion analysis indicates that lack of Region C prevents repression, and analysis of a frameshift of Region C indicated that polypeptide rather than nucleotide sequence was involved in the response. Interestingly, a peptide scan of using a library of overlapping 13 amino acid-long peptides identified Region C as the site of two high affinity DnaK binding sites within , leading to speculation that the function of Region C may be to bind DnaK (McCarty et al., 1996). Further support for this notion comes from comparative analysis of the sigma family of polypeptides. Whereas this region of sigma is highly conserved among homologues from diverse bacteria, it is poorly conserved among sigma factors in general (Nakahigashi et al., 1995). It is certainly plausible that a nonconserved region within the sigma family of proteins has become specialized for a regulatory function specific to homologues. Cotranslational binding of DnaK to Region C may then mediate translational repression by an unknown mechanism. 2.4. Regulation of

Stability

The instability of is a key feature of the response to temperature upshift. Because is so unstable (T½=1 minutes) during steady state growth, increases in its rate of synthesis are immediately reflected in commensurate increases in the level of available to promote transcription of the heat shock genes. Great advances in understanding this process have recently been reported. Both in vivo and in vitro studies indicate that is proteolysed by HflB, an ATP dependent protease located in the inner membrane (Tomoyasu et al., 1993; Herman et al., 1995; Tomoyasu et al., 1995).


22

Depleting cells of HflB (FtsH), or inactivating mutant HflB by shift to high temperature stabilizes about 10-fold indicating that HflB is a major protease responsible for degradation. Moreover, HflB can degrade in vitro. Interestingly, HflB is a member of the regulon and the only essential protease thus far reported in E. coli. There are still important, unresolved questions concerning the physiology of degradation. Currently, the rate of degradation of in vitro (T½=18 minutes) is much slower than the in vivo T½ of 1 min. In vivo, the DnaK-DnaJ-GrpE chaperone machine is required for degradation of , and mutations in dnaK, dnaJ or grpE decrease the rate of degradation as much as 10-fold (Tilly et al., 1989; Straus et al., 1990). Region C of , described above as a possible DnaK binding site, may couple these chaperones to the process of degradation. In support of this idea, the Region C frameshift mutant inhibits degradation of in vivo (Nagai et al., 1994). However, the in vitro degradation system currently in use exhibits no requirement for these hsps (Tomoyasu et al., 1995). Moreover, the presence of core RNA polymerase inhibits the in vitro degradation of by HflB, and this inhibition is not reversed by the DnaK-DnaJ-GrpE chaperone machine. Thus, the in vitro system is not yet a faithful mimic of in vivo degradation, either because of missing components or altered conditions. 2.5. Regulation of

Activity

Inactivation of appears to be a primary mode of regulation whenever is present in excess in the cell (Straus et al., 1989; Taura et al., 1989; Straus et al., 1990). This regulatory mode features most prominently on temperature downshift, but also most likely sharpens the shut-off phase of the heat shock response. The DnaK-DnaJ-GrpE chaperone machine is involved in inactivation, as cells carrying mutations in these genes are defective in this process (Straus et al., 1989 and unpublished experiments). Inactivation is reversible as regains activity after extraction from the cell (Straus et al., 1989). These characteristics led to the proposal that the DnaK-DnaJ-GrpE chaperone machine reversibly binds to to inhibit its function (Straus et al., 1989) (Figure 2). Elegant in vitro studies from the Bukau and Georgopoulos laboratories are beginning to establish the molecular basis for inactivation of . Both DnaK and DnaJ can bind independently to (Gamer et al., 1992; Liberek et al., 1992; Liberek et al., 1993; Gamer et al., 1996). In addition, all three also form an ATP-dependent ternary complex with distinct properties from each of the binary complexes (Liberek et al., 1993; Gamer et al., 1996). It is only this ternary complex that shows decreased activity with core RNA polymerase (Liberek et al., 1993; Gamer et al., 1996). Thus, together DnaK and DnaJ function as an anti-sigma factor. When bound to , they inhibit the formation of the -core RNA polymerase complex (Gamer et al., 1996). Understanding the mechanistic details of the interactions of DnaK and DnaJ with is in its infancy. Indeed, further study of this interaction is likely to yield important insights concerning the regulatory loop governing activity, and also


23

Figure 2 Speculative model for the mechanism by which DnaK, DnaJ and GrpE regulate expression of hps by controlling activity and levels. Upon temperature upshift, the increase in misfolded protein substrates leads to a decrease in the free levels of DnaK, DnaJ and GrpE resulting in increased stability. Upon temperature downshift, the increase in the free pool of these chaperones leads to inactivation of . In addition to these effects, a role for DnaK, DnaJ and GrpE in negatively regulating the increase in translation of observed upon temperature upshift has been proposed (see text). (Figure adapted from Gross, 1996).

into the nature of chaperone interaction with native substrates. The DnaKbinary complex is relatively weak (Kd=5 M), and this binding is considerably decreased by ATP (Gamer et al., 1992; Liberek et al., 1992; Liberek et al., 1993; Gamer et al., 1996). Interestingly, the low binding constant reflects a very slow on rate, as the DnaKcomplex is quite stable once formed (T½>30 minutes) (Gamer et al., 1996). In contrast, the stronger DnaJbinary complex (Kd=20nM; measured in the Biacore), actually dissociates more rapidly than the DnaKcomplex (Gamer et al., 1996). The ternary complex, which requires ATP for its formation, somehow stabilizes the -DnaK interaction and effectively competes with for binding to core RNA polymerase. It is


24

currently unknown how DnaJ promotes formation of this ternary complex. However, DnaJ binding to substrate may not be necessary for its effect. Some DnaJ mutants that do not bind still promote an ATP-resistant -DnaK interaction, and may do so catalytically (Liberek et al., 1995). It is not known, however, whether these -DnaK binary complexes inhibit mediated transcription. 2.6. What are the Signals Governing Expression of the Regulon?

Heat Shock

The challenge of the cell is to integrate diverse environmental information to program the level of hsp expression that is appropriate for the perceived cumulative stress level. Exactly how this is accomplished is still a matter of speculation. We have a great deal of information about initial inputs—expression of the regulon is triggered by heat, ethanol and other diverse insults. Likewise, we are fairly knowledgeable about the final outputs— regulation of both the activity and amount of lead to a defined rate of transcription of the heat shock genes. However, the nature of the signal-transduction pathway(s) that couple(s) the two ends of this regulatory loop remains an area of active investigation. There are at least two distinct signal-transduction pathways governing expression of the hsps. The first pathway controls translation of mRNA in a positive way: increased environmental stress leads to increased translation. This pathway is induced by exposure to heat and ethanol, but not by accumulation of unfolded proteins. To date, the only identified player in this pathway is cis-acting mRNA sequences. Neither the trans-acting factors, nor the signaling molecule (s) have been identified. Our understanding of the remainder of the regulatory events governing the amount of active is somewhat more advanced. Regulating stability, activity and translational repression have in common the involvement of the DnaK, DnaJ and GrpE chaperone machine in the signal transduction pathway. Regulation of these diverse processes may be controlled either by a single pathway, or by multiple, interconnected pathways. A homeostatic mechanism coupling the occupancy of the DnaK, DnaJ, GrpE chaperone machine to the amount and activity of has been proposed (Straus et al., 1990; Craig et al., 1991; Bukau, 1993). Cellular stress is monitored by how well can compete with all other unfolded or misfolded proteins for binding to the DnaK, DnaJ, GrpE chaperone machine. Inducing signals increase unfolded or misfolded proteins, thus titrating DnaK, DnaJ and GrpE away from and relieving their negative regulatory effects on stability and translation. As a consequence, the amount of will rise. Conversely, repressing signals will decrease unfolded or misfolded proteins, thus freeing DnaK, DnaJ and GrpE to inactivate . This response is self limiting because under or over production of DnaK, DnaJ and GrpE will restore the free pool of these chaperones to an appropriate level. Thus, the amount of free DnaK, DnaJ, and GrpE is a “cellular thermometer” that measures the “folding state” of the cell. There is some evidence in favor of this model, however, critical experiments to test the proposition that the DnaK, DnaJ and GrpE chaperones play a regulatory role have yet to be carried out.


3. REGULATION OF THE

(

25

) HEAT SHOCK RESPONSE

3.1. Discovery of was originally discovered as the sigma factor responsible for maintaining transcription of rpoH at extreme temperatures. rpoH has four promoters, three of which are transcribed by E (Figure 1a). The fourth promoter, rpoHp3, is recognized by E . rpoHp3 accounts for only 2% of total rpoH transcription at 30°C, but drives over 90% at the lethal temperature of 50°C (Erickson et al., 1987). The continued production of at 50°C is critical to cellular survival, as the dependent hsps represent the majority of proteins expressed under these extreme conditions (Neidhardt et al., 1984; Pack et al., 1986). was purified based on its ability to direct transcription from rpoHp3 (Erickson et al., 1989; Wang et al., 1989), and the structural gene encoding was recently identified (Raina et al., 1995; Rouvière et al., 1995). 3.2. What is the Nature of the Signal Inducing

Activity?

In addition to being induced by the general stresses of heat and solvents, the pathway is uniquely induced in response to alterations in the expression or maturation of outer membrane proteins (OMPs) (Mecsas et al., 1993). Overexpression of OMPs induces activity, and underexpression of OMPs decreases activity. The inducing signal arises either during or after translocation because cytoplasmic accumulation of OMP precursors does not induce activity. Although activity is induced by overexpression of some periplasmic proteins with known folding defects (Missiakas et al., 1996), overexpression of most periplasmic proteins does not induce , indicating that the signal is probably not arising due to titration of the translocation machinery. Expression of a mutant OMP that is properly translocated but fails to be inserted into the outer membrane also induces activity. Taken together, these results suggest that the signal arises in the periplasmic space, after translocation but prior to insertion into the outer membrane. Outer membrane proteins undergo a complex series of folding events during their maturation into trimeric porins. Blocking this pathway at a step after the signal intermediate is generated should cause an increase in activity. Using this and related strategies, several putative periplasmic folding agents have been identified, including the peptidyl prolyl isomerases SurA and FkpA, and the Skp protein (Rouvière et al. 1996; Missiakas et al., 1996). Loss of function mutations in each of these genes induce activity. The role of SurA in maturation of the trimeric porin LamB has been investigated (Rouvière et al., 1996; Lazar et al., 1996). SurA appears to catalyze the formation of a folded monomeric species from unfolded monomer. Cells lacking SurA and cells overexpressing LamB both accumulate the unfolded monomer form at the expense of folded monomer. The observation that two different inducing conditions result in accumulation of unfolded monomer suggests that the signal for induction occurs somewhere prior to the formation of the folded monomer species (Rouvière et al., 1996).


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3.3. Regulation of The activity of is regulated, in part, at the level of transcription. is transcribed from a -dependent promoter and transcription from this promoter reflects the level of activity in the cell under steady state conditions (Raina et al., 1995; Rouvière et al., 1995). However, both the observation that transcription of is low under steady state conditions and that activity increases rapidly in response to induction suggest additional regulatory controls. Homology arguments suggested that is under the control of negative regulators likely to be encoded in the same operon as rpoE, and this turns out to be the case. belongs to the ECF subclass of the family of proteins, most of which regulate extracytoplasmic functions (Rouvière et al., 1995; Lonetto et al., 1994). Operons encoding other ECF sigmas have previously been shown to also encode regulators of the sigma factor activity. In particular, the operon encoding the closely related algU/T sigma factor required for alginate biosynthesis in P. aeruginosa, includes two negative regulators of AlgU/T activity, MucA and MucB (Martin et al., 1993). MucA inhibits AlgU/T activity in vivo and in vitro (Schurr et al., 1996; Xie et al., 1996), and previous work had identified a partial open reading frame encoded immediately downstream of rpoE, termed mclA, that showed significant homology to mucA (Raina et al., 1995; Rouvière et al., 1995; Yu et al., 1995). Three genes, rseABC (for regulator of sigmaE), are encoded immediately downstream of rpoE, and genetic experiments reveal that rseA (formerly mclA) and rseB negatively regulate activity (De Las Peñas, et al., 1997a; Missiakas et al., 1997). Deletion of rseA leads to a 25-fold induction of activity, whereas deletion of rseB gives only 2.3-fold induction, indicating that RseA is the major negative regulator of . RseA is an inner membrane protein, whose cytoplasmic domain binds directly to and inhibits -directed transcription in vivo and in vitro. Thus, the cytoplasmic domain of RseA acts as an anti-sigma factor. The periplasmic domain of RseA interacts with RseB, which is located in the periplasm, and RseC has a slight positive effect activity. 3.4. How is the Extracytoplasmic Signal Transduced to

?

RseA is the central regulatory molecule in the signal transduction cascade to . Cells lacking RseA are unresponsive to induction because they are already maximally induced. Moreover, cells containing only RseA modulate activity in response to inducer, indicating that RseA alone or in conjunction with unknown molecules responds to the inducing signal. Several mechanisms of RseA inactivation by the inducer can be envisioned including modification, degradation, or oligomerization of the anti-sigma factor. RseB may act to fine-tune this RseA-based signal transduction pathway. Binding of RseB to the periplasmic domain of RseA might shift RseA to a conformation where it is most effective as an anti-sigma (Figure 3a). If RseB binding to RseA were competitive with binding to a signal molecule, RseB would be titrated away from RseA as the concentration of the signal increases (Figure 3b). This would leave RseA in a


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conformational state where it is a less effective anti-sigma, and lead to a small increase in activity. At still higher concentrations, the signal molecule would interact either with an intermediate factor or with RseA itself to further increase activity (Figure 3c). The direct induction signal and how it affects RseA is currently unknown. is induced by the build up of early intermediates in the maturation pathway of outer

Figure 3 Speculative model of the signal transduction cascade leading to activation of . (a) In the presence of low levels of signal, is


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sequestered to the membrane by a protein complex consisting of RseA and RseB, leaving activity low. (b) Under conditions of low level signal, RseB is titrated off of RseA, leaving RseA in a conformation that is less active as an anti-sigma factor, resulting in a small increase in activity, (c) When the signal is high, RseA is further inactivated either by interaction with the signal molecule itself or some intermediate factor, resulting in a large induction of activity.

membrane porins, the accumulation of a few periplasmic proteins, and a deficit of any of several periplasmic folding agents (DsbA, FkpA, Skp and SurA) (Mecsas et al., 1993; Rouvière et al., 1996; Missiakas et al., 1996). The Rse proteins may detect the levels of misfolded protein directly. Alternatively, RseA and/or RseB may monitor the levels of free periplasmic folding agents, including SurA, FkpA, and the Dsb proteins. Decreases in the free levels of each of these proteins in response to the accumulation of unfolded or misfolded species in the periplasmic space may additively induce the pathway. Upon generation of a signal, is released from the complex with RseA, leading to a positive feedback loop. The newly active transcribes its own promoter to generate more and RseA. As long as the signal is present, RseA will be unable to interact with , but when the signal is removed or reduced, RseA, possibly in concert with RseB, will again repress , achieving a new steady state level. Although this model bears a superficial resemblance to the regulation of , it is unlikely that RseA targets for degradation, or that RseA interacts with the signal in the same manner as it interacts with . 3.5. The Cellular Role of is an essential sigma factor, at least at temperatures above 18°C, and cells lacking rapidly accumulate a suppressor of this lethality (De Las Peñas et al., 1997b). Cells lacking and containing this suppressor form colonies at 42°C to 43°C with greatly reduced efficiency (10-3 to 10-5), and die more rapidly than wild type cells after exposure to lethal temperatures (Hiratsu et al., 1995; Raina et al., 1995; Rouvière et al., 1995), while cells containing the suppressor alone are temperature resistant (Connolly and Gross, unpublished observations). These phenotypes confirm the importance of the regulon for resistance to thermal stress. Overexpression of sE leads to the induction of at least 10 proteins (Raina et al., 1995; Rouvière et al., 1995). However, only four members of the regulon have been identified. In addition to rpoH, EsE transcribes the periplasmic protease degP, the periplasmic peptidyl-prolyl isomerase fkpA (Danese and Silhavy, 1997), and one of the two promoters upstream of rpoE itself. Why does E. coli need two heat-inducible regulons? Part of the answer might be that the two regulons respond to stress in different cellular compartments. Some inducers, such as heat and solvents, affect all cellular compartments and thus induce both regulons. Other inducers specifically alter protein folding in either the cytoplasmic or extracytoplasmic environments, and uniquely induce or activity, respectively.


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Just as the response has a close parallel in the eukaryotic heat shock response, the pathway also has a eukaryotic counterpart. Accumulation of unfolded proteins in the endoplasmic reticulum (ER) leads to the transcriptional induction of several ER resident folding agents (Cox et al., 1993; Mori et al., 1993). Like the pathway, the ER response, known as the unfolded protein response (UPR), is controlled separately from the cytoplasmic heat shock response. Although the central regulator of the UPR shares no common features with RseA, it remains to be seen whether the two systems share common mechanisms of sensing the initial signal. E. coli has a second signal transduction pathway, the Cpx two-component system, capable of relieving extracytoplasmic stress. Although the Cpx system is not required for growth at high temperature (Connolly et al., unpublished observations), activation of the pathway suppresses the envelope-associated toxicity conferred by certain LamB mutant proteins by inducing the expression of DegP (Cosma et al., 1995; Danese et al., 1995; Snyder et al., 1995). Interestingly, activation of the Cpx pathway also restores the ability to grow at high temperature to cells lacking , in a degP-dependent manner (Connolly, et al., 1997). Overexpression of degP alone does not suppress the rpoE-temperature sensitive phenotype, indicating that other Cpx-controlled genes are required. Future work aimed at elucidating the relationship between the Cpx pathway and the -mediated response should help to clarify the roles of each system in responding to protein misfolding outside of the cytoplasm. Work on the pathway is just beginning. The next few years should provide us with exciting insights into the members of the regulon, the nature of the signal, and the regulatory network that links the cellular compartments. In addition, has already proven to be an invaluable tool in the search for periplasmic folding agents and rapid progress in the understanding of folding processes in this cellular compartment is likely to follow.

4. HEAT SHOCK REGULATION IN OTHER PROKARYOTIC ORGANISMS Study of the heat shock response in a number of different bacteria indicates that the basic E. coli regulatory paradigm is not universal. Although homologues are widespread among gram negative bacteria, additional regulatory mechanisms also affect the primary heat shock response in some of these organisms. Moreover, the gram positive organisms examined to date do not have homologues. homologues have been isolated from a number of Gram negative bacteria (Garvin et al., 1989; Benvenisti et al., 1995; Fleischmann et al., 1995; Naczynski et al., 1995; Nakahigashi et al., 1995; Yura, 1996). All of these homologues can restore growth to E. coli cells lacking functional , indicating that the transcriptional function of the protein is conserved. However, sequence analysis suggests that only some of the regulatory inputs are conserved. All homologues identified to date contain Region C, which binds DnaK with high affinity and is required for control of stability. In contrast, the regions of mRNA implicated in translational control are conserved in but not proteobacteria. If translational control of exists in a proteobacteria, it must be


30

mechanistically distinct from the E. coli model. These observations suggest that diverse mechanisms may control the amount and/ or activity of in different gram negative species. Our knowledge about the heat shock response in gram positive organisms comes from studies of Bacillus subtilis and Clostridium acetobutylicum. (Narberhaus et al., 1992; Narberhaus et al., 1992; Schmidt et al., 1992; Wetzstein et al., 1992; Zuber et al., 1994; Yura, 1996). In these organisms, the major chaperone genes are transcribed by the housekeeping sigma and are preceded by a conserved inverted repeat sequence. This inverted repeat, named CIRCE for controlling inverted repeat for chaperone expression, is the binding site for a putative represser (Yuan et al., 1995). The mechanism of thermal induction of genes regulated by the CIRCE element has not yet been elucidated. CIRCE has also been detected in some gram negative bacteria suggesting that it is rather widely involved in the heat shock response. In Bradyrhizobium japonicum, and CIRCE together control expression of heat shock genes (Babst et al., 1996), suggesting that parallel regulatory strategies may exist in some organisms. In contrast to , the degree of conservation of has not been determined. Although several sigma factors belonging to the ECF family have been described in both gram negative and positive bacteria (Lonetto et al., 1994; Rouvière et al., 1995), their possible role in the heat shock response of these organisms has not been widely studied. Only one of the ECF sigmas in addition to has been implicated in the resistance to thermal stress. Pseudomonas aeruginosa cells lacking the homologue AlgU, show increased killing at 50°C compared to AlgU+ strains (Martin et al., 1994), and the activity of AlgU is induced in response to heat shock (Schurr et al., 1995). However, AlgU carries out additional cellular functions not mediated by . For example, AlgU-cellsshowincreased sensitivity to superoxide-generating compounds (Martin et al., 1994), and AlgU plays a key role in the production of the exopolysaccharide alginate (Deretic et al., 1994). One possibility is that the -mediated response has been co-opted by other signaling systems in P. aeruginosa, and it will be interesting to determine how AlgU and utilize similar signaling molecules to respond to diverse extracellular signals.

5. SUMMARY AND PROSPECTS Although recent studies have given us insight into the mechanisms responsible for the regulation of both and , several basic questions concerning the response to thermal stress in E. coli remain unresolved. For example, the exact nature of the initial signal and sensing mechanism have not been elucidated. Further dissection of the response loops of each sigma factor should provide us with a greater understanding of not only the heat shock response but also of the process of protein folding in each cellular compartment. We have only begun to understand the in vivo role of the chaperones, and to identify periplasmic protein folding agents. The next few years should prove to be an exciting time in the dual fields of thermal stress response and protein folding.


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6. ACKNOWLEDGMENTS We thank Jonathan Tupy for help in preparing figures, and Charlotte Hedlund for excellent assistance in editing and performing the innumerable tasks required to complete this manuscript.

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sigma 32 polypeptide is involved in DnaK-mediated negative control of the heat shock response in Escherichia coli. Proc. Natl. Acad. Sci. USA , 91 , 10280–10284. Nagai, H., Yuzawa, H. and Yura, T. (1991). Interplay of two cis-acting mRNA regions in translational control of sigma 32 synthesis during the heat shock response of Escherichia coli. Proc. Natl. Acad. Sci. USA , 88 , 10515–10519. Nakahigashi, K., Yanagi, H. and Yura, T. (1995). Isolation and sequence analysis of rpoH genes encoding sigma 32 homologs from gram negative bacteria: conserved mRNA and protein segments for heat shock regulation. Nucleic Acids Res , 23 , 4383– 4390. Narberhaus, F. and Bahl, H. (1992). Cloning, sequencing and molecular analysis of the groESL operon of Clostridium acetobutylicum. J. Bacterial. , 174 , 3282–3289. Narberhaus, F., Giebeler, K. and Bahl, H. (1992). Molecular characterization of the dnaK gene region of Clostridium acetobutylicum, including grpE, dnaJ and a new heat shock gene. J. Bacteriol. , 174 , 3290–3299. Nashimoto, H. (1993). The Translational Apparatus , New York, Plenum Press. Neidhardt, F.C. and VanBogelen, R.A. (1981). Positive regulatory gene for temperaturecontrolled proteins in Escherichia coli. Biochem. Biophys. Res. Commun. , 100 , 894– 900. Neidhardt, F.C. and VanBogelen, R.A. (1987). Heat Shock Response. In F.C.Neidhardt, (ed.), Escherichia coli and Salmonella typhimuriam: Cellular and Molecular Biology , American Society for Microbiology,Washington, D.C., pp. 1334–1345. Neidhardt, F.C., VanBogelen, R.A. and Vaughn, V.A. (1984). The genetics and regulation of heat-shock proteins. Annu. Rev. Genet , 18 , 295–329. Neidhardt, F.C., Phillips, T.A., VanBogelen, R.A., Smith, M.W., Georgalis, V. and Subramanian, A.R. (1981). Identity of the B56.5 protein, the A-protein and the groE gene product of Escherichia coli. J. Bacteriol. , 145 , 513–520. Pack, K.H. and Walker, G.C. (1986). Defect in expression of heat-shock proteins at high temperature in xthA mutants. J. Bacteriol. , 165 , 763–770. Raina, S. and Georgopolous, C. (1991). The htrM gene, whose product is essential for Escherichia coli viability at elevated temperatures, is identical to the rfaD gene. Nucleic Acids Res. , 19 , 3811–3819. Raina, S., Missiakas, D. and Georgopolous, C. (1995). The rpoE gene encoding the sigma E (sigma 24). heat shock sigma factor of Escherichia coli. Embo. J. , 14 , 1043– 1055. Rouvière, P.E., De Las Peñas, A., Mecsas, J., Lu, C.Z., Rudd, K.E. and Gross, C.A. (1995). rpoE, the gene encoding the second heat-shock sigma factor, sigma E, in Escherichia coli. Embo. J. , 14 , 1032–1042. Rouvière, P.E. and Gross, C.A. (1996). SurA, a Periplasmic Protein with Peptidyl Propyl Isomerase Activity, Participates in the Assembly of Outer Membrane Porins. Genes Dev. , 10 , 3170–3182. Schmidt, A., Schiesswohl, M., Volker, U., Hecker, M. and Schumann, W. (1992). Cloning, sequencing, mapping and transcriptional analysis of the groESL operon from Bacillus subtilis . J. Bacteriol. , 174 , 3993–3999. Schurr, M.J., Yu, H., Boucher, J.C., Hibler, N.S. and Deretic, V. (1995). Multiple promoters and induction by heat shock of the gene encoding the alternative sigma factor AlgU (sigma E). which controls mucoidy in cystic fibrosis isolates of Pseudomonas aeruginosa. J. Bacteriol. , 177 , 5670–5679. Schurr, M.J., Yu, H., Martinez-Salazar, J.M., Boucher, J.C. and Deretic, V. (1996). Control of AlgU, a member of the sigma E-like family of stress sigma factors, by the


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negative regulators MucA and MucB and Pseudomonas aeruginosa conversion to mucoidy in cystic fibrosis. J. Bacteriol. , 178 , 4997–5004. Skelly, S., Coleman, T., Fu, C.F., Brot, N. and Weissbach, H. (1987). Correlation between the 32-kDa sigma factor levels and in vitro expression of Escherichia coli heat shock genes. Proc. Natl Acad. Sci. USA , 84 , 8365–8369. Snyder, W.B., Davis, L.J., Danese, P.N., Cosma, C.L. and Silhavy, T.J. (1995). Overproduction of NlpE, a new outer membrane lipoprotein, suppresses the toxicity of periplasmic LacZ by activation of the Cpx signal transduction pathway. J. Bacteriol. , 177 , 4216–4223. Squires, C.L., Pedersen, S. and Ross, B.M. (1991). ClpB is the Escherichia coli heat shock protein F84.1. J. Bacteriol. , 173 , 4254–4262. Strauch, K.L., Johnson, K. and Beckwith, J. (1989). Characterization of degP, a gene required for proteolysis in the cell envelope and essential for growth of Escherichia coli at high temperature. J. Bacteriol. , 171 , 2689–2696. Straus, D.B., Walter, W. and Gross, C.A. (1990). DnaK, DnaJ and GrpE heat shock proteins negatively regulate heat shock gene expression by controlling the synthesis of sigma-32. Genes Dev . 4 , 2202–2209. Straus, D.B., Walter, W.A. and Gross, C.A. (1987). The heat shock response of E. coli is regulated by changes in the concentration of sigma 32. Nature , 329 , 348–351. Straus, D.B., Walter, W.A. and Gross, C.A. (1989). The activity of sigma 32 is reduced under conditions of excess heat shock protein production in Escherichia coli. Genes Dev. , 3 , 2003–2010. Taura, T., Kusukawa, N., Yura, T. and Ito, K. (1989). Transient shut off of Escherichia coli heat shock protein synthesis upon temperature shift down. Biochem. Biophys. Res. Commun. , 163 , 438–443. Taylor, W.E., Straus, D.B., Grossman, A.D., Burton, Z.F., Gross, C.A. and Burgess, R.R. (1984). Transcription from a heat-inducible promoter causes heat shock regulation of the sigma subunit of E. coli RNA polymerase. Cell , 38 , 371–381. Tilly, K., Spence, J. and Georgopolous, C. (1989). Modulation of stability of the Escherichia coli heat shock regulatory factor sigma. J. Bacteriol. , 171 , 1585–1589. Tomoyasu, T., Gamer, J., Bukau, B., Kanemori, M., Mori, H., Rutman, A.J., Oppenheim, A.B., Yura, T., Yamanaka, K., Niki, H., Hiraga, S. and Ogura, T. (1995). Escherichia coli: FtsH is a membrane-bound, ATP-dependent protease which degrades the heatshock transcription factor sigma 32. Embo J. , 14 , 2551–2560. Tomoyasu, T., Yamanaka, K., Murata, K., Suzaki, T., Bouloc, P., Kato, A., Niki, H., Hiraga, S. and Ogura, T. (1993). Topology and subcellular localization of FtsH protein in Escherichia coli. J. Bacteriol. , 175 , 1352–1357. Wang, Q.P. and Kaguni J.M. (1989). A novel sigma factor is involved in expression of the rpoH gene of Escherichia coli . J. Bacterial , 171 , 4248–4253. Wetzstein, M., Volker, U., Dedio, J., Lobau, S., Zuber, U., Schiesswohl, M., Herget, C., Hecker, M. and Schumann, W. (1992). Cloning, sequencing and molecular analysis of the dnaK locus from Bacillus subtilis. J. Bacteriol. , 174 , 3300–3310. Xie, Z.D., Hershberger, C.D., Shankar, S., Ye, R.W and Chakrabarty, A.M. (1996). Sigma factor-anti-sigma factor interaction in alginate synthesis: inhibition of AlgT by MucA. J. Bacteriol , 178 , 4990–4996. Yamamori, T., Ito, K., Nakamura, Y. and Yura, T. (1978). Transient regulation of protein synthesis in Escherichia coli upon shift-up of growth temperature. J. Bacteriol. , 134 , 1133–1140. Yamamori, T. and Yura, T. (1980). Temperature-induced synthesis of specific proteins in


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Escherichia coli: evidence for transcriptional control. J. Bacteriol. , 142 , 843–851. Yamamori, T. and Yura, T. (1982). Genetic control of heat-shock protein synthesis and its bearing on growth and thermal resistance in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA , 79 , 860–864. Yu, H., Schurr, M.J. and Deretic, V. (1995). Functional equivalence of Escherichia coli sigma E and Pseudomonas aeruginosa AlgU: E. coli rpoE restores mucoidy and reduces sensitivity to reactive oxygen intermediates in algU mutants of P. aeruginosa . J. Bacteriol. , 177 , 3259–3268. Yuan, G. and Wong, S.L. (1995). Isolation and characterization of Bacillus subtilis groE regulatory mutants: evidence for or/39 in the dnaK operon as a represser gene in regulating the expression of both groE and dnaK J. Bacteriol. , 177 , 6462–6468. Yura, T. (1996). Regulation and conservation of the heat-shock transcription factor sigma-32. Genes to Cells , 1 , 277–284. Yura, T., Tobe, T., Ito, K. and Osawa, T. (1984). Heat shock regulatory gene (htpR). of Escherichia coli is required for growth at high temperature but is dispensable at low temperature. Proc. Natl. Acad. Sci. USA , 81 , 6803–6807. Yuzawa, H., Nagai, H., Mori, H. and Yura, T. (1993). Heat induction of sigma 32 synthesis mediated by mRNA secondary structure: a primary step of the heat shock response in Escherichia coli. Nucleic Acids Res. , 21 , 5449–5455. Zhou, Y.N., Kusukawa, N., Erickson, J.W., Gross, C.A. and Yura, T. (1988). Isolation and characterization of Escherichia coli mutants that lack the heat shock sigma factor sigma 32. J. Bacteriol. , 170 , 3640–3649. Zuber, U. and Schumann, W. (1994). CIRCE, a novel heat shock element involved in regulation of heat shock operon dnaK of Bacillus subtilis. J. Bacteriol. , 176 , 1359– 1363.

3. INDUCIBLE TRANSCRIPTIONAL REGULATION OF HEAT SHOCK GENES: THE STRESS SIGNAL AND THE UNFOLDED PROTEIN RESPONSE RICHARD I.MORIMOTO Department of Biochemistry, Molecular Biology and Cell Biology, Rice Institute for Biomedical Research, Northwestern University, 2153 Sheridan Road, Evanston, Il. 60208, USA

1. Introduction 1.1 Inducible Transcription by a Family of Heat Shock Factors 2. The HSFI Cycle: Proposed Role for Heat Shock Proteins in Autoregulation of the Heat Shock Response 3. Activation of HSF2 is Linked to Proteolysis and the Activity of the UbiquitinDependent Proteasome 4. HSF 3 and HSF4 5. Transcriptional Regulation of Endoplasmic Reticulum Stress Genes (GRP78): The Unfolded Protein Response 6. Conclusions 7. Acknowledgement 8. References

1. INTRODUCTION The activation of heat shock gene expression is a highly regulated response to diverse environmental and physiological conditions including heat shock, oxidative stress, heavy metals, various chemicals, bacterial and viral infection, and exposure to a number of acute and chronic disease states (Ritossa, 1962; Ashburner, 1970; Lindquist and Craig, 1988; Morimoto et al., 1990; 1994). These conditions (Figure 1) can be partitioned into three broad catergories: (1) environmental stress including heat shock, amino acid analogues, drugs, toxic chemicals, and heavy metals, (2) pathophysiologic and disease states including oxidative stress, fever, inflammation, infection, myocardial stress and ischemia, and neural degenerative diseases, and (3) non-stress conditions including the cell cycle, growth factors, serum stimulation, development, differentiation, and activation


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by certain oncogenes. The diverse nature of these conditions has led to questions on the identity and properties of the cellular machinery that detects “stress”. This chapter will address the stress-sensing mechanisms that operate in the cytosol and lumen of eukaryotic cells and that lead to the transcriptional activation of specific genes encoding compartment specific heat shock proteins and molecular chaperones.

Figure 1 Conditions that result in the induction of heat shock gene expression in eukaryotes. Representation of three general classes of conditions known to result in the elevated expression of stress proteins including: (1) environmental and physiological stress, (2) pathophysiological states including conditions of disease and (3) nonstressful conditions such as cell growth and development. Each condition acts on the cell as diagrammed in this figure and in the case of environmental stress and certain pathophysiological states leads to the activation of heat shock gene expression and the synthesis of heat shock proteins.

1.1. Inducible Transcription by a Family of Heat Shock Factors Exposure of eukaryotic cells to elevated temperatures and other stressful conditions leads to the rapid activation of heat shock transcription factor (HSF) and the inducible transcription of genes encoding heat shock proteins and molecular chaperones (Lindquist and Craig, 1988; Morimoto, 1993; Lis and Wu, 1993; Wu, 1995). Analysis of the

Inducible transcriptional regulation of the heat shock genes

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chromosomal structure in the vicinity of the Drosophila Hsp70 and Hsp90 genes led to the identification of binding sites for constitutive and inducible transcription factors (Wu, 1980, 1984). Heat Shock Factor (HSF) was initially identified as the inducible transcription factor which bound specifically to the heat shock element (HSE), a pentameric nucleotide sequence (5'-AGAAn-3') positioned as inverted adjacent arrays in the promoter region of eukaryotic heat shock-responsive genes (Amin et al., 1988; Xiao and Lis, 1988; Perisic et al., 1989; Williams and Morimoto, 1990; Greene and Kingston, 1990; Xiao et al., 1991). A distinction between the heat shock response of yeast and metazoans is the constitutive DNA binding properties of yeast HSF, whereas in Drosophila, plants, and vertebrates, the HSFs are constitutively expressed, negatively regulated for DNA binding, and require heat shock for acquisition for DNA binding (Sorger and Pelham, 1988; Kingston et al., 1987; Zimarino and Wu, 1987; Larson et al., 1988; Mosser et al., 1988; Nover, 1994; Zimarino et al., 1990; Westwood et al., 1991; 1993). The mechanism by which HSFs are negatively regulated involves both cis-acting negative regulatory elements and constitutive phosphorylation which function to repress DNA binding and the transactivation domain (Shi et al., 1995; Knauf et al., 1996; Kline and Morimoto, 1997). The inducibility of the latent Drosophila and mammalian HSFs can also be observed in vitro following exposure to in vitro heat shock, non-ionic detergents, low pH, or exposure to chaotropes (Larson et al., 1988; Mosser et al., 1990). The initial biochemical characterization of HSFs and the subsequent cloning of the respective genes was initially accomplished in S. cerevisiae and D. melanogaster (Sorger et al., 1987; Wiederrecht et al., 1988; Wu et al., 1987). Subsequently, the cloning of HSF genes from larger eukaryotes revealed a multi-gene family. At least three HSFs have been isolated from the human (HSFs 1, 2, 4), chicken (HSFs 1, 2, 3), and tomato genomes (Scharf et al., 1990; Rabindran et al., 1991; Sarge et al., 1991; Schuetz et al., 1991; Nakai and Morimoto, 1993; Nakai et al., 1997). The cloned HSFs vary in size from 301 amino acids (aa) for tomato HSF24, 512aa for tomato HSF8, 503aa and 529aa for mouse and human HSF1, 517aa and 536aa for mouse and human HSF2, 463aa for chicken HSF3, 467aa for human HSF4, 691aa for Drosophila HSF and 833aa for S. cerevisiae HSF (Scharf et al., 1990; Nakai and Morimoto, 1993; Rabindran et al., 1991; Sarge et al., 1991; Schuetz et al., 1991; Nakai et al., 1997). Within a species, members of the HSF family (ie., mouse HSF1-HSF2 or chicken HSF1-HSF2-HSF3) are approximately 40% related in amino acid sequences; this is primarily due to sequence identity within the DNA binding and oligomerization domains (Morimoto, 1993; Wu, 1995). Interspecies comparisons, ie., between human, mouse, and chicken HSF1 reveals a higher level of sequence identity (85–95%) revealing a common ancestral progenitor from which the contemporary HSFs evolved (Nakai and Morimoto, 1993). Functional domains of HSFs (Figure 2) were identified initially by alignment of the amino acid sequences derived from the respective cloned genes and subsequently by functional assays using collections of deletion and point mutants. Comparison of the HSFs from yeasts, Drosophila, tomato, chicken, mouse, and humans identified a conserved ≈100 amino acid DNA-binding domain of the winged helix-turn-helix motif located towards the amino-terminus (Damberger et al., 1994; Harrison et al., 1994; Vuister et al., 1994a; 1994b). The high degree of conservation in the DNA binding domain reflects the conservation of the nucleotide sequences comprising the HSE DNA


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binding motif. Adjacent to the DNA-binding domain is a region of approximately 100 residues corresponding to an extended hydrophobic heptad repeat (HR-A/B) essential for trimerization (Sorger and Nelson, 1989; Peteranderl and Nelson, 1992; Nieto-Sotelo et al., 1990; Perisic et al., 1989;

Figure 2 Functional domains of heat shock factor 1. The organizational structure of mouse HSF1 is indicated with the DNA binding domain designated at the amino-terminus. Adjacent to the DNA binding domain are the conserved hydrophobic heptad repeats involved in oligomerization of HSF1 and the negative regulatory domain. At the carboxyl-terminus is an additional hydrophobic heptad repeat adjacent to the transcription activation domain. The positions of sites of consitutive and inducible serine phosphorylation are indicated. Depicted below is a representation of the conversion of the inert monomeric form of HSF1 to the transcriptionally active trimer.

Clos et al., 1990; Rabindran et al., 1993). The HSFs of S. pombe, Drosophila, and larger


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eukaryotes contain an additional array of hydrophobic heptad repeats (HR-C) positioned near the extreme carboxyl terminus which may function in the negative regulation of DNA binding (Rabindran et al., 1993). A detailed analysis of the transcriptional activation domains of the HSF from S. cerevisiae and K. lactis identified separate regions required for full transcriptional activity (Nieto-Sotelo et al., 1990; Jakobsen and Pelham, 1991; Chen et al., 1993). In contrast, the transcription activation domain of Drosophila and mammalian HSF1 were localized to the extreme carboxyl terminus of the respective proteins (Shi et al., 1995; Newton et al., 1996; Wisniewski et al., 1996; Zuo et al., 1995). The studies on mammalian HSF1 have also identified negative regulatory elements in the vicinity of the DNA binding domain which influences the activity of the transactivation domain (Shi et al., 1995; Newton et al., 1996; Zuo et al., 1995). What is the role for a family of HSFs in the transcriptional regulation of heat shock genes? One possibility is that larger organisms may require multiple HSFs to provide specialized responses to the diverse developmental and environmental cues and insults they may be exposed to during life. Consistent with this speculation, vertebrate HSF1 and HSF2, though structurally related, are distinct by a variety of regulatory and functional criteria. For example, HSF1 corresponds to the general stress-responsive transcription factor whereas HSF2 is activated in response to developmental and differentiation cues (Theodorakis et al., 1989; Sistonen et al., 1992, 1994; Sarge et al., 1993, 1994). Activation of HSF1 is a complex multi-step process which involves oligomerization from an inert monomer to active trimer, acquisition of DNA-binding ability, stress-induced phosphorylation, and nuclear localization (Sarge et al., 1993; Baler et al., 1993; Jurivich et al., 1992; Sistonen et al., 1992; Sistonen et al., 1994; Cotto et al., 1996). Acquisition of trimer formation precedes inducible phosphorylation; furthermore the inducibly phosphorylated state of HSF1 is stress dependent with heat shock, heavy metals and arachidonate treatment resulting in the fully phosphorylated state and exposure to amino acid analogues or salicylate resulting in the activation of the DNA binding competent non-inducibly phosphorylated state (Sarge et al., 1993; Jurivich et al., 1994; Cotto et al., 1996). In contrast, HSF2 is not post-translationally modified by heat shock or other stresses (Sarge et al., 1993; Sistonen et al., 1992, 1994). HSF2 DNA binding is activated, for example, following exposure of human K562 erythroleukemia cells to hemin (Singh and Yu, 1984; Theodorakis et al., 1989; Sistonen et al., 1992). The acquisition of HSF2 DNA-binding activity is accompanied by conversion from an inert dimer to a DNA binding competent trimer; the DNA binding properties of HSF1 and HSF2 is similar but not identical (Kroeger et al., 1993; Sistonen et al., 1994). Constitutive HSF2 DNA binding activity has been observed in embryonal carcinoma cells (Morange et al., 1984; Mezger et al., 1989; Mezger et al., 1994; Murphy et al., 1994). Another observation which supports a role for HSF2 during development is the elevated expression of HSF2 expression and HSF2 DNA binding activity during spermatogenesis (Sarge et al., 1991; 1994) (see Morange, this volume).


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2. THE HSF1 CYCLE: PROPOSED ROLE FOR HEAT SHOCK PROTEINS IN AUTOREGULATION OF THE HEAT SHOCK RESPONSE One of the distinguishing features of the heat shock response is the rapid kinetics of activation, the magnitude of inducibility, and reversibility of the response. Attenuation of the transcriptional response occurs in cells continuously exposed to intermediate heat shock temperatures (42°C) or immediately upon return to control temperatures, whereas following exposure to extreme heat shock temperatures (>43°C), the heat shock response persists and is maintained at high levels (Abravaya et al., 1991a). Comparison of HSF1 DNA binding properties using in vivo genomic footprinting on the human hsp70 promoter reveals that the HSE is unoccupied in cells at controlled temperatures and fully occupied during heat shock due to inducible binding to HSF1 (Abravaya et al., 1991a, b; Sistonen et al., 1994). During the attenuation phase, HSF1 releases rapidly from the HSE, and thereafter HSF1 DNA binding activity is no longer detected. The in vivo equilibrium dissociation rate for HSF1 has a half-life of approximately 10 minutes, whereas the dissociation rate of the HSF: HSE complex formed in vitro is greater than 100 minutes. The disparity of these results have led to the suggestion that other trans-regulatory components may be involved in the release of the activated form of HSF from DNA and dissociation of the trimer to the inert monomer. These speculations are further supported by the properties of recombinant Drosophila, chicken, mouse and human HSF’s in E. coli which are purified as stable trimers exhibiting constitutive DNA binding activity (Clos et al., 1990; Nakai and Morimoto, 1993; Sarge et al., 1993; Kroeger et al., 1993). It has long been speculated from studies in Drosophila and yeast that heat shock proteins may function in an autoregulatory loop to modulate the intensity and duration of the heat shock response (Craig and Gross, 1991). Indeed, this feature appears to be evolutionarily conserved as indicated by the role of heat shock proteins in the regulation of activity (Gamer et al., 1992; see Connolly et al., this volume). Although the sensor of cell stress in eukaryotes has not been established, most models for regulation of the heat shock response have proposed that the appearance of malfolded proteins, induced during heat shock and other forms of stress, sequesters Hsp70, thus allowing HSF1 to be activated (Morimoto et al., 1990; Abravaya et al., 1992; Baler et al., 1992; Morimoto, 1993). Additional support for the autoregulatory hypothesis comes from experimental evidence that links the activation of the heat shock response to increased levels of denatured and misfolded proteins (Anathan et al., 1986; Baler et al., 1992). Exposure to inhibitors of protein synthesis blocks the activation of the heat shock response by interfering with induction of HSF1; these results suggest that the proper synthesis and folding of nascent polypeptides represents a critical target for detection of malfolded proteins (Mosser et al., 1988; Amici et al., 1992; Baler et al., 1992). Taken together, these results support the hypothesis that the appearance and accumulation of malfolded polypeptides are directly involved in the pathway of stress detection and response (DiDomenico et al., 1982). Molecular chaperones such as members of the Hsp70 family are attractive candidates in the autoregulation of the heat shock response as they have a primary role in the association with non-native proteins to prevent their aggregation and to facilitate protein folding (Craig and Gross, 1991; Gething and Sambrook, 1992).


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Although there is a substantial evidence to support an involvement of molecular chaperones in the regulation of HSF1 activation, it is uncertain whether this occurs through direct or indirect effects. HSF1 trimers become associated with Hsp70 during attenuation; additionally HSF1 attenuation is more rapid in cell lines expressing high levels of Hsp70 (Abravaya et al., 1992; Mosser et al., 1993; Rabindran et al., 1994). If indeed it is the balance or ratio of free Hsp70 which is either directly or indirectly involved in maintaining the non-DNA binding form of HSF1 or in the attenuation of HSF1, it might be expected that overexpression of hsp70 would negatively affect the heat shock response. In support of this is evidence that the HSF1 transactivation domain is negatively regulated by Hsp70 (Shi and Morimoto, unpublished observations). Likewise, overexpression of HSF1 obtained by transient transfection results in a constitutively active or more readily activated factor independent of exogenous stress (Sarge et al., 1993). These results reveal that heat shock is not an obligatory step in HSF1 activation and are consistent with the hypothesis that there is a critical balance between HSF1 and its negative regulatory molecules to maintain HSF1 in either the non-DNA binding or DNA binding state. Many of the current observations are consistent with a model for the regulation of HSF1 DNA binding activity which is schematically represented in Figure 3. Under nonstressful conditions, HSF1 is constitutively expressed and maintained as a non-DNA binding monomer through intramolecular interactions perhaps influenced by constitutive serine phosphorylation (Kline and Morimoto, unpublished observations). Heat shock and other stresses result in the appearance of malfolded and aggregated proteins which creates a large pool of new protein substrates which compete with HSF1 for association with Hsp70. Thus, heat shock and other stresses initiate the events that remove the negative regulatory influence on HSF1 DNA binding activity. The conversion of HSF1 monomers to trimers requires a significant change in the conformation of HSF1 which may involve other activities to facilitate or stabilize the DNA binding trimer. HSF1 also undergoes a stress-dependent inducible serine phosphorylation and acquires transcriptional activity. The activated trimer state of HSF1 is associated with HSBP1 (heat shock factor binding protein), a small hydrophobic heptad repeat containing protein, during the period when HSF1 is active as a transcription factor (Satyal et al., in press). The subsequent events are not clearly ordered and include the interaction of HSBP1 with Hsp70 and the association of HSF1 with Hsp70 which has repressive effects on HSF1 transcriptional activity. Ultimately, these events during the regulation of HSF1 transcriptional activity are linked to the conversion of the active trimeric protein to the inert control state.

3. ACTIVATION OF HSF2 IS LINKED TO PROTEOLYSIS AND THE ACTIVITY OF THE UBIQUITIN-DEPENDENT PROTEASOME In contrast to the observations that HSF1 functions as the predominant stress responsive inducible transcription factor, the function of HSF2 has been an enigma. HSF2 is expressed in a wide range of cell types in all larger eukaryotes, yet HSF2 activity has only been detected as a constitutive DNA binding activity in embryonal carcinoma cells and tissues from murine embryos, during murine spermatogenesis, and in human K562


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erythroleukemia cells upon treatment with hemin (see Morange, this volume). In the latter cell system, HSF2 is inert and activated upon hemin

Figure 3 HSF Cycle—A model of HSF1 regulation. In the unstressed cell, HSF1 is maintained primarily in the cytoplasm in a monomeric, nonDNA binding form. Upon heat shock or other forms of stress, HSF1 translocates and relocalizes within the nucleus and assembles into a trimer; HSBP1 associates with HSF1 trimers; HSF1 trimers bind to the heat shock element located in the promoter regions of heat shock genes and undergoes inducible serine phosphorylation. Transcriptional activation of the heat shock genes leads to increased levels of Hsp70 and to formation of an HSF1—Hsp70 complex. During attenuation of the heat shock transcriptional response, HSF1 dissociates from the DNA and is eventually converted to non-DNAbinding monomers.

treatment to a DNA binding and transcriptionally active trimer that binds in vivo to the promoter of the hsp90 and hsp70 genes (Sistonen et al., 1992). Unlike the robust, yet transient activation of HSF1 and resulting transcription of heat shock genes observed following exposure to heat shock, HSF2 activity and the expression of heat shock genes following hemin treatment can be detected over an extended period of time (Theodorakis et al., 1989; Sistonen et al., 1992). A more complete understanding of the role of HSF2 has come from observations that the activation of HSF2 DNA binding and transcriptional activity appears to be due to the inhibitory effects of hemin on ubiquitin-dependent


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proteasome activity and the demonstration that specific proteasome inhibitors such as the peptide aldehyde, MG132, and the Streptomyces metabolite, lactacystin, also induce HSF2 activity (Mathew, Mathur, and Morimoto, unpublished observations). Exposure of yeast and mammalian cells to MG132, lactacystin or hemin (the latter in a cell specific manner) leads to the elevated expression of heat shock genes hsp90, hsp70 and hdj-1 via transcriptional induction dependent upon the HSE (Lee and Goldberg, personal communication; Mathew, Mathur and Morimoto, unpublished observations). How is HSF2 activity regulated during downregulation of proteaseome activity? The change in HSF2 DNA binding activity is neither accompanied by relocalization of HSF2 nor by changes in the levels of HSF2. Furthermore HSF2 is not polyubiquitinated, yet maintenance of the trimeric state of HSF2 requires ongoing protein synthesis. These data are best explained by the suggestion that HSF2 activation is dependent upon another labile regulatory protein, distinct from HSF2, which itself is degraded by the proteasome. In contrast, HSF1 is a stable protein whose activity is independent of protein synthesis and the activity of the ubiquitindependent proteasome. Thus, HSF2 rather than HSF1 senses the dynamics of protein degradation. The inhibition of protein degradation would be predicted to result in the accumulation of polyubiquitinated non-native proteins that maybe prone to aggregation. Therefore, the elevated expression of heat shock proteins under such circumstances affords a mechanism to prevent the formation of insoluble protein aggregates. HSF2 activation, as a response to down-regulation of proteasome activity, may enable the cell to act in a preventative manner by increasing expression of molecular chaperones whose function is to associate with these non-native proteins. The synthesis of Hsp70 rapidly attenuates upon removal of the proteasome inhibitor, consistent with a role for the accumulated Hsp70 (and perhaps other chaperones) to associate with the increasing levels of polyubiquitinated proteins which build up during the arrest of proteasome activity. Based on these results, a model (Figure 4) is proposed for HSF2 activation in which inhibition of proteasome activity leads to the elevated synthesis and accumulation of heat shock proteins and molecular chaperones which serves to associate with the accumulated proteins, presumably to maintain these non-native proteins in a soluble intermediate folded state. This feature of Hsp70 activity, the maintenance of intermediate non-native proteins in a soluble state, is supported by recent observations of in vitro protein folding experiments with molecular chaperones (Freeman et al., 1995; Freeman and Morimoto, 1996; Freeman, Toft and Morimoto, 1996). There is additional evidence to link the heat shock response and proteolysis. In E. coli, the activities of the ATP-dependent proteases Lon and Clp, which are themselves heat shock proteins (Goff and Goldberg, 1985), are influenced by the heat shock proteins DnaK, DnaJ, GrpE, GroES and GroEL (Sherman and Goldberg, 1992; Sherman and Goldberg, 1996). Likewise, in eukaryotes, ubiquitin and other components of the ubiquitin modification system are induced by heat shock and essential for survival following exposure to stress (Finley, Ozkaynak and Varshavsky, 1987; Seufert and Jentsch, 1990; Jentsch, 1992). The molecular chaperone Ydj-1 (DnaJ homologue) has been shown to associate with an artificially ubiquitinated short-lived abnormal protein and is essential for ubiquitin-dependent degradation (Lee, Sherman and Goldberg, 1996). The link between HSF2 activity and proteolysis offers an explanation for the constitutive


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activation of heat shock genes in the ts85

Figure 4 Model for regulation of chaperone expression by proteasome activity. (1) Malfolded, mutant and short-lived proteins are polyubiquitinated (indicated by addition of Ub). (2) Ubiquitinated proteins are targeted to the proteasome for degradation which can be inhibited by the addition of MG132, lactacystin, or hemin. (3) Inhibition of proteasome activity results in an accumulation of polyubiquitinated proteins. (4) HSF2 is activated from the inert dimer to DNA-binding trimer in response to the accumulation of non-native polyubiquitinated proteins. (5) HSF2 induces expression of molecular chaperones such as Hsp70 and Hdj-1. (6) The increased transcription of molecular chaperones results in increased levels of Hsp70 and Hdj-1 proteins. (7) The molecular chaperones associate with the polyubiquitinated proteins and are suggested to maintain the polyubiquitinated proteins in an intermediate folded state primed for degradation upon resumption of proteasome activity.

mouse cell line which has a thermolabile ubiquitin-activating enzyme (Finley, Ciechanover and Varshavsky, 1984) and a proposed model, based on these studies, for activation of the heat shock response in which inhibition of the ubiquination system would promote accumulation of a non-ubiquinated active transcription factor leading to


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transcription of the heat shock genes (Munro and Pelham, 1985; Finley, Ciechanover and Varshavsky, 1984).

4. HSF3 AND HSF4 Chicken HSF3 was cloned during a molecular screen of a chicken cDNA library for genes homologous to murine HSF1 (Nakai and Morimoto, 1993). The three members of the chicken HSF family (HSF1–3) are closely related (40% sequence identity) and exhibit similar structural motifs for the DNA binding domain and the adjacent hydrophobic heptad repeats. HSF3 is ubiquitously expressed and the DNA binding properties are negatively regulated presumably through intramolecular interactions involving the carboxyl terminal hydrophobic heptad C, as has also been suggested for HSF1. The inert state of HSF3 is a dimer which undergoes oligomerization to an active trimer. However, in contrast with HSF1, acquisition of HSF3 DNA binding activity in response to heat shock, appears to be a cell specific event and has been detected predominantly in HD6 cells and the chicken B lymphoblast cell line, DT40, where it has the properties of a positive activator (Nakai et al., 1995). The co-activation of HSF1 and HSF3 in HD6 cells exposed to heat shock suggests the possibility of HSF regulatory redundancy, perhaps to ensure that the transcriptional activation of heat shock genes is both rapidly induced and sustained at a high level in response to extreme temperatures. In this context, it is worth noting that the optimal heat shock temperature for avian cells is 45°C as compared to 42°C for mammals. The possibility of a complementary role for these apparently redundant HSFs is supported by the rapid kinetics of HSF1 induction followed by HSF3. This observation, together with the demonstration that the in vitro activation properties of HSF3 and HSF1 differ (Nakai and Morimoto, 1993) indicates that these these two factors are similar, yet have distinct regulatory features as measured by acquisition of DNA binding activity. To establish the biological role for HSF3, however, it will be necessary to either inactivate or substantially reduce HSF1 expression which would allow us to demonstrate whether HSF3 is itself sufficient to activate the expression of the endogenous heat shock genes. HSF4 has been recently described as a new member of the human HSF gene family with the unusual feature of lacking the properties of a transcriptional activator despite having in common many of the structural and biochemical features of other previously cloned HSF genes (Nakai et al, 1997). Overexpression of HSF4 leads to the repression of target genes that are regulated through heat shock element promoter sequences. This unusual feature may be related to the fact that HSF4 lacks the carboxyl terminal hydrophobic repeat which is shared among all previously characterized members of the HSF gene family which are negatively regulated and exhibits a highly restricted expression pattern in human tissues. Therefore, it remains to be established whether HSF4 is a novel negative regulator of heat shock gene expression which exerts its activity in a highly tissue restricted manner as a represser whose activity is to balance HSE targeted transcriptional activity. Among the various HSF’s identified in diverse species, only HSF4 lacks the activity of a positive transactivator. Overexpression of HSF4 results in the repression of basal transcription of cellular heat shock genes regulated by the HSE


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(ie., Hsp90, Hsp70, and Hsp27) presumably through direct binding of HSF4 to the HSEs and subsequent inhibitory effects on either the initiation complex or during transcriptional elongation. It is tempting to speculate that the function of HSF4 as a negative regulator of basal transcription of heat shock genes provides a novel means to modulate heat shock gene transcription.

5. TRANSCRIPTIONAL REGULATION OF ENDOPLASMIC RETICULUM STRESS GENES (GRP78): THE UNFOLDED PROTEIN RESPONSE The endoplasmic reticulum (ER) represents yet another subcellular compartment in which nascent polypeptides are imported, folded, and assembled. The accumulation of unfolded proteins (unfolded protein response) in the ER, following exposure to conditions that prevent glycosylation, leads to the elevated transcription of lumen localized chaperones (Grp94, Grp78/Kar2, protein disulfide isomerase) (Lee, 1987; Kozutsumi et al., 1988; Watowich and Morimoto, 1988). Expression of these target genes is regulated via a common unfolding protein response element (UPRE) which is necessary and sufficient to confer the response to the accumulation of malfolded ER proteins (Mori et al., 1992; Kohno et al., 1993). A second component of the UPR is a transmembrane kinase (Ire1p) which is located in the ER (Cox, Shamu and Walter, 1993; Mori et al., 1993). Ire1p is a 1115 amino acid protein of which the amino terminus lies in the lumen of the ER where it presumably recognizes unfolded proteins and the kinase domain lies either in the cytoplasm or nucleus and is likely responsible for transmitting the signal to the downstream components of the UPR. In response to unfolded proteins, Ire1p oligomerizes and is transphosphorylated by other Ire1p molecules (Shamu and Walter, 1996; Welihinda and Kaufman, 1996). The carboxyl terminal tail of Ire1p has been suggested to interact with unfolded proteins. Using the UPRE as the target for a genetic screen, the transcription factor, Hac1p, a member of the bZIP family of transcription factors, was identified which binds directly to the UPRE (Cox and Walter, 1996; Nojima et al., 1994). Disruption of the gene encoding Hac1p results in the loss of the UPR, thus revealing that Hac1p is in the cascade of events in the UPR (Cox and Walter, 1996). HAC1 RNA, however, is present under noninducing conditions and is found on polyribosomes whereas Hac1p is only detected under UPR conditions. Although Ire1p is not an essential gene, splicing of HAC1 mRNA is dependent upon Ire1p to generate a unique exon-exon junction resulting in a change in HAC1 reading frame leading to the synthesis of a stable Hac1p. The differences in the two forms of Hac1p correspond to an extended PEST region which confers degradation by the ubiquitin-dependent proteasome. Thus, Ire1p activated splicing would induce a longer-lasting form of Hac1p. Consistent with these observations, mutations in the ubiquitin conjugating enzymes (UBC) activates the UPR pathway in the absence of ER stress (Seufert and Jentsch, 1990; Seufert et al., 1990). Is Hac1p the only transcription factor that responds to ER stress? Exposure of adipogenic cell lines to low glucose, tunicamycin, and other conditions that induce grp78 expression results in the elevated expression of CHOP (C/EBP homologous protein) (Chen et al., 1992; Price and Calderwood, 1992; Carlson et al., 1993). CHOP was


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initially identified as a DNA damage inducible gene based on its elevated expression following exposure to UV or the alkylating agent methyl methanesulfonate and separately shown to heterodimerize with members of the C/EBP family and function as a dominant negative regulator of transcription (Fornace et al., 1989; Ron and Habner, 1992; Ubeda et al., 1996). CHOP expression is rapidly induced following ER stress relative to the induction of ER chaperones and may be a component of the ER-stress sensing apparatus in mammalian cells (Wang et al., 1996). How CHOP detects the ERstress, however has not been established.

6. CONCLUSIONS Among the fundamental, unanswered, questions that underlie the basis of the heat shock response is the nature of the intracellular signal(s) that indicates the cell is stressed. The expression of heat shock and stress responsive genes has been closely linked with the appearance, within the cell, of malfolded or mutant proteins and the differential activation of heat shock transcription factors. Heat shock factor 1 (HSF1) functions as the ubiquitous stress responsive activator which is maintained in a latent state and responds principally to the appearance of nascent stress-induced malfolded proteins, whereas HSF2 activity is linked to proteolysis and the activity of the ubiquitin-dependent proteasome. Activation of either HSF1 or HSF2 leads to transcription of heat shock element (HSE) regulated genes encoding cytosolic heat shock proteins and molecular chaperones. In contrast, the appearance of unfolded proteins in the lumen of the endoplasmic reticulum (ER) is detected by Ire1p a kinase that spans the ER membrane and senses the increase in unfolded proteins and Haclp a transcription factor which in its activated state binds to the unfolded protein response element (UPRE) and activates the transcription of genes encoding ER-localized chaperones.

7. ACKNOWLEDGEMENTS The studies from our laboratory were supported by grants from the National Institutes of General Medicine. The author extends an appreciation to members of the laboratory who contributed to these studies and provided continuous stimulating discussions.

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heat shock response element. Science , 239 , 1139–1142. Xiao, H., Perisic, O. and Lis, T.J. (1991). Cooperative binding of Drosophila heat shock factor to arrays of a conserved 5 bp unit. Cell , 64 , 585–593. Zimarino, V. and Wu, C. (1987). Induction of sequence-specific binding of Drosophila heat shock activator proteins without protein synthesis. Nature , 327 , 727–730. Zimarino, V., Tsai, C. and Wu., C (1990). Complex modes of heat shock factor activation. Mol. Cell. Biol. , 10 , 752–759. Zuo, J., Rungger, D. and Voellmy, R. (1995). Multiple layers of regulation of human heat shock transcription factor1. Mol. Cell Biol. , 15 , 4319–4330.

4. PROTEIN KINASE CASCADES INVOLVED IN HEAT SHOCK PROTEIN EXPRESSION AND FUNCTION OLIVIER BENSAUDE Génétique Moléculaire, Ecole Normale Supérieure, 46 rue d’Ulm, 75230 Paris Cedex 05, France

1. Introduction 2. Stress-activated MAP kinase cascades 2.1. Stress activation of the ERK type MAP kinases 2.2. Stress-activated MAP kinases related to the yeast osmosensing kinase, hog1 2.3. Multiple activating pathways 2.4. Negative regulation of MAP kinases 2.5. HSP interactions modulate protein kinase cascade activities 2.6. The MAP kinase cascades interfere with the cell survival after stress 3. HSP phosphorylation triggered by environmental stimuli 3.1. Protein kinases and phosphatases in the phosphorylation of small HSPs 3.2. The p38 MAP Kinase, an upstream activator of the small HSP Kinase 3.3. Phosphorylation of the small HSPs and cytoskeleton dynamics 4. Stress-induced phosphorylation of transcription factors 4.1. Transcription Factors Phosphorylated by MAP Kinases 4.2. Phosphorylation of the RNA polymerase II largest subunit by stressactivated CTD kinases 5. Involvement of MAP kinases in stress gene transcription 5.1. Heat shock element (HSE) dependent transcription 5.2. Stress response element (STRE) dependent transcription 5.3. GRP gene transcription 6. Control of protein synthesis 6.1. Dephosphorylation of ribosomal subunit S6 and eukaryotic initiation factor 4E (eIF-4E) 6.2. HSP depletion triggers the phosphorylation of the eukaryotic initiation factor 2 (eIF2 ) 7. Conclusion 8. Acknowledgements


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9. References

1. INTRODUCTION The cell physiology is altered when cells are subjected to heat stress (reviewed in Laszlo, 1992; Nover, 1991; Welch, 1992). The cytoskeleton collapses, RNA splicing and protein synthesis are impaired, and the pattern of gene expression is extensively modified. The extent of the impairements depends on the severity of the stress and may lead to cell death. These alterations constitute the heat shock response and result in a reprogramming of central metabolic processes. The enhanced synthesis of heat shock proteins (HSPs), which contribute to repair and degradation of proteins damaged during stress, is a central manifestation of this reprogramming and contributes to the adaptation of the cells to the stress condition. Numerous posttranslational modifications of proteins occur in response to heat stress that may also be involved in preventing or repairing cellular damages. Changes in protein phosphorylation, glycosylation, methylation, acetylation, farnesylation and ubiquitination have been found to occur during stress (reviewed in Bensaude et al, 1996; Nover, 1991). Among these, protein phosphorylation is by far the best studied modification. Several HSPs are modified by phosphorylation in response to stress treatment. In E. coli for example, heat shock increases the proportion of phosphorylated GroEL and DnaK (Sherman and Goldberg, 1994; Sherman and Goldberg, 1993). But, apart from isolated reports on prokaryote protein phosphorylation, most studies concern the phosphorylation of the eukaryotic HSPs, and among these, phosphorylation of the “small” HSPs (23 to 28 kd) in response to a wide range of stimuli including growth factors, exposure to steroid hormones, cytokines and several stresses (oxidative, heat, osmotic, UV irradiation) (reviewed in Arrigo and Landry, 1994). Indeed, stress modulates the activity of various protein kinases signaling cascades. This chapter will focus on such cascades which relate to the HSPs in three aspects: (i) HSP levels control the activation of the cascades; (ii) HSPs are targets of the activated cascades; (iii) stress gene expression depends on the induction of protein kinases. To provide an accurate picture of the physiological importance of these regulatory circuits the discussion will be extended to stresses other than heat shock which trigger the same signaling pathways.

2. STRESS-ACTIVATED MAP KINASE CASCADES The mitogen activated protein kinases (MAP kinases or MAPK) belong to a network of signal transduction pathways (reviewed in Kyriakis and Avruch, 1996; Robinson and Cobb, 1997; Mahdani and Fink, 1998). Phosphorylation of these kinases is activated by MAP kinase kinases (MEK or MKK), which in turn are activated by MAP kinase kinase kinases (MAPKKK or MEKK). Many of these signal transduction pathways are induced by stress treatment that also induces synthesis of HSPs, and they are part of the cellular system that allows adaptation of metabolism to stress. On the one hand the activation of MAP kinases leads to the phosphorylation of the “small HSPs”, on the other hand it is

Protein kinase cascades involved in heat shock protein

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involved in the regulation of stress gene expression. 2.1. Stress Activation of the ERK Type MAP Kinases The ERK type MAP kinases were first described as growth factor-activated kinases. Subsequent work established that in cultured mammalian cells, they were also activated during heat (Bendinelli et al., 1995; Venetianer et al., 1995; Chen et al., 1995) oxidative (Guyton et al., 1996) and osmotic (Matsuda et al., 1995) stress. In yeast, the activity of the ERK/MAP kinase homologue, encoded by the slt2/mpk1 gene, is modulated by the growth temperature and increases 170-fold when temperature is raised from 23 to 37°C (Kamada et al., 1995; Torres et al., 1991; Zarzov et al., 1996). 2.2. Stress-activated MAP Kinases Related to the Yeast hog1 Osmosensing Kinase A considerable interest has recently arised for two new types of MAP kinase strongly activated by UV irradiation, protein synthesis inhibitors, tumor necrosis factor, oxidative and heat stress (reviewed in Kyriakis and Avruch, 1996): the c-Jun kinases, JNKs or SAPK1 (stress-activated protein kinase) and the p38 MAP kinases. The latter are also designated by a wide variety of acronyms such as RK (Rouse et al., 1994), p40 MAP kinase (Freshney et al., 1994), p38 MAP kinase (Han et al., 1994), CSBP1 and CSBP2 (cytokine-suppressive anti-inflammatory drug—CSAID binding proteins) (Lee et al., 1994) or SAPK2 (Meier et al., 1996) (see Table 1).

Table 1 Mammalian mitogen-activated-protein kinases (MAPK) cascades: Acronyms MAP kinase kinase kinase kinases: Ste20, PAK1, p65PAK, GCK, SOK-1 MAP kinase kinase kinases: MEKKs, MAPKKKs, MKKKs, MUK, SAPKKKs, SEKKs, JNKKKs, RKKKs Raf-1, c-mos MAP kinase kinases: MEKs, MAPKKs, MKKs, SAPKKs, SEKs, JNKKs, RKKs MAP kinase repressers: Cdk inhibitor: p21 WAF1/CIP1/Sdi1 , p21 CIP1 , p21 WAF1 Dual specificity MAP kinase phosphatases: CL100, 3CH134, erp, MKP-1 B23, hVH-3 MAP kinases: Genuine mitogen-activated-protein kinases (MAPK) subclass:


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ERK1, p44MAPK, p44mpk ERK2, p42MAPK, p42mpk Stress-activated protein kinase (SAPK) subclass of MAPK: JNK, SAP kinase, SAPK1; JNK1, SAPK-p46 I; JNK2, SAPK-p54 SAPKp54 I; SAPK-p46 I p38MAPK, p38HOG, CSBP1 and CSBP2, MPK2, SAPK2, RK

II; p54MAP-2 kinase,

MAP kinase activated kinases: MAPKAPK1, p90 ribosomal S6 kinase, p90rsk, RSK2 MAPKAPK2, HSP27 kinase, CREB kinase MAPKAPK3, HSP27 kinase, 3pK

Figure 1 Schematic representation of nuclear and cytoplasmic targets of the signaling network of the ERK, JNK, p38 MAP kinase pathways. PD098059 and SB203580 are highly selective inhibitors of respectively the MEK1/MEK2 kinases, which activate ERK1/2, and the p38 MAP kinase. The nucleus is schematized by a grey box.

The mammalian p38 MAP kinases are highly homologous to the Saccharomyces cerevisiae hog1 gene product that is involved in resistance to hyperosmotic stress. The


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human MAP kinase, CSBP1, complements yeast mutants carrying a deletion in hog1 (Han et al., 1994). Surprisingly, the CSBP2 kinase does not complement the hog1 deletion mutants despite its strong (96%) homology to CSBP1 (Kumar et al., 1995). Recently, a third member of the p38 MAP kinase family, p38 , has been characterized (Jiang et al., 1996). The p38 MAP kinases are furthermore closely related to the JNK. Similar to the CSBP1 gene, the mammalian JNK1 gene rescues a hog1 deficient yeast mutant (Galcheva-Gargova et al., 1994). However, although the JNK2 isoform shares 80% homology with JNK1, it does not rescue the yeast hog1 deficient yeast mutants (Sluss et al., 1994). The JNKs and the p38 MAP kinases are often presented as stress specific kinases and designated as SAPK1 and SAPK2 in contrast to the ERK MAP kinases which would rather be growth factor specific. This differentiation, however, seems not to be consistent with the recent findings described above and with the observation that stress specificity depends on the cell system investigated (Hu et al., 1997). 2.3. Multiple Activating Pathways The MAP kinases are activated by phosphorylation of threonine and tyrosine residues. Multiple stress-specific activators of p38 and JNK MAP kinases have been identified (Fanger et al., 1997). Depending on the inducer, different activating pathways are involved (Adler et al., 1995). Exposure of cells to N-acetylcysteine, an oxygen radical scavenger, or to low concentrations of Triton X-100 inhibits the UV-mediated, but not the heat shock-mediated JNK activation. Conversely, low concentrations of hydrogen peroxide inhibited heat shock-mediated but not UV-mediated JNK activation. Moreover, JNK activation by UV irradiation was impaired in cell lines derived from patients with Cockayne Syndrome of complementation group B (CS-B), whereas induction by oxidative stress and heat stress were normal (Dhar et al., 1996). Separate pathways have been demonstrated in the activation of the JNKs. One mechanism involves the c-Abl tyrosine kinase and is operative after treatment with ionizing radiation and genotoxic agents. Another pathway involving the tyrosine kinase, Pyk2, is required for activation by tumour-necrosis factor (TNF- ), UV and osmotic shock (Kharbanda et al., 1995; Tokiwa et al., 1996). Disruption of the MKK4 gene blocked JNK activation by anisomycin or heat shock but not so much by other stress such UV radiation or osmotic shock (Yang et al., 1997). Tyrosine kinase membrane receptors or the Ras small GTP protein in cooperation with the Raf-1 kinase are upstream activators of the ERK cascade. Upstream activators of the mammalian JNK and p38 cascades can be members of the family of p21-activated kinases, PAK This family includes the Ste20 kinase which controls mating in yeast (Sells and Chernoff, 1997) in cooperation with small GTP-binding proteins of the Rho subfamily, Rac1, Cdc42 (for reviews see (Nagata and Hall, 1996; Symons, 1996) and the human vav oncogene (Crespo et al., 1996). Independent signals, which emanate from distinct cell surface sensors, have been shown to activate the yeast Pbs2p MAP kinase (Posas et al., 1996). A phosphate residue is transferred from a histidine of the sensor kinase, Sln1p, to an aspartate of the response regulator, Ssk1p. Sln1p and Ssk1p are physically linked together by the Ypd1p. This


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phosphorelay is initiated by the stress-induced autophosphorylation of Sln1p. Independently, Pbs2p can be activated by binding to the Src homology 3 (SH3) domain of the osmosensing protein, Sho1p. As a whole, most of the primary sensors which trigger the various MAP kinases cascades appear to be anchored in the plasma membrane where several members of the cascade can associate to each other and form molecular complexes (reviewed in Elion, 1995; Faux and Scott, 1996). It is tempting to speculate that upon heat shock treatment of the cells a temperature-induced increase in membrane fluidity might be the primary event that leads to their activation. Interestingly in yeast, the membrane lipid composition has been shown to be a determinant in the temperature set point of the heat shock response (Carratù et al., 1996). 2.4. Negative Regulation of MAP Kinases The MAP kinase cascades can be regulated at two steps, either phosphorylation or dephosphorylation of kinases (Burgering and Bos, 1995). Two types of protein phosphatases which dephosphorylate the critical threonine and tyrosine residues negatively regulate the MAP kinases (Groom et al., 1996; Keyse, 1995). The protein tyrosine phosphatase SHPTP1, which also downregulates the JNK, is activated by phosphorylation through the stress-activated c-Abl (Kharbanda et al., 1996). Another phosphatase, Pyst 1, which downregulates specifically the ERK type MAP kinases has also been identified (Groom et al., 1996). A serine phosphatase, protein phosphatase 2A, is responsible for the inactivation of the ERK type MAP kinases in some cell lines (Alessi et al., 1995). Although inducers of the heat shock response such as arsenite stimulate the JNK and p38 MAP kinases by inhibiting a dual specificity phosphatase (Cavigelli et al., 1996), the protein phosphatases are generally thought to provide a negative feed back control of the MAP kinase cascades as their expression is inducible by stress. Thus, expression of the S. pombe pyp2 gene encoding a phosphotyrosine phosphatase is stress-inducible and under the control of spc1 MAP kinase (Degols et al., 1996). A similar regulatory loop is also found in mammalian cells as the transcription of the dual specifity phosphatase genes, CL100 (MKP-1, 3CH134, erp) and B23/hVH-3 is rapidly enhanced after heat and oxidative stress and contributes to a downregulation of the MAP kinases of the ERK, p38 and JNK types (Chu et al., 1996b; Keyse, 1995; Sun et al., 1993) with some cell to cell variations (Alessi et al., 1995). Furthermore an anisomycin pretreatment selectively desensitizes components in the stress-kinase activation pathway (Hazzalin et al., 1998). Another negative regulation loop of MAP kinase cascades that relies on a mechanism distinct from phosphatases is mediated by protein p21 WAF1/CIP1/Sdil , a cyclin-dependentkinase inhibitor (reviewed in Harper and Elledge, 1996). This inhibitor also interacts with JNK1 and p38 MAP kinase, but not with ERK1 (Shim et al., 1996), thereby inhibiting the corresponding kinases. The p21 WAF1/CIPI/Sdil protein is induced two-fold by stress including heat shock (Ohnishi et al., 1996). This recently reported negative regulation loop illustrates a new connection between the MAP kinases and the cyclin-dependent kinases (cdk), which constitutes a distinct network of kinases controlling the cell cycle.


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2.5. HSP Interactions Modulate Protein Kinase Cascade Activities Interactions with HSP modulate the activation of protein kinases. For example Hsp70 overexpression prevents the stress-activation of the JNK and the p38 MAP kinases (Gabai et al., 1997). A novel member of the small heat shock protein family has recently been identified in muscle cells (Suzuki et al., 1998). This protein binds and activates a protein kinase the myotonic dystrophy protein kinase, MDPK Various protein kinases associate with Hsp90 and the cdc37 protein in their inactive form (reviewed in Hunter and Poon, 1997; Pratt, 1997). This is exemplified for tyrosine kinases such as src and for the Raf-1 protein kinase, an upstream activator of the ERK type MAP kinases (Morrison and Cutler, 1997). Raf-1 forms a heterocomplex with Hsp90 and p50Cdc37. Preventing the association of Raf-1 with Hsp90 by the antibiotic geldanamycin interferes with trafficking of the newly synthesized Raf-1 from the cytoplasm to the plasma membrane and with the association of Raf-1 with Ras. As a result, the Raf-1-MEK-MAP kinase signalling pathway is disrupted (Schulte et al., 1996; van der Straten et al., 1997). The src family tyrosine kinases are bound to Hsp90 when inactive. Phosphorylation of Hsp90 disrupts its complex with pp60v-src (Mimnaugh et al., 1995). In mammalian cells, Hsp90 chaperones are phosphoproteins for which the turnover of phosphate is increased during heat stress (Legagneux et al., 1991). It is interesting to note that upon heat shock tyrosine phosphorylation increases in cultured mammalian cells (Maher and Pasquale, 1989). In the fission yeast S. pombe, the wis1 and the spc1/sty1 MAP kinases are counteracted by overexpression of the phosphatase 2C (PP2C) encoded by the ptc1 gene (Shiozaki and Russell, 1995) and the phosphotyrosine phosphatases encoded by the pyp1 and the pyp2 genes (Degols et al., 1996). As the ptc1 gene is a multicopy suppressor of swo1–26, a temperature-sensitive mutation of the yeast hsp90 gene (Shiozaki et al., 1994), this observation also illustrates the importance of Hsp90 in the regulation of the MAP kinase cascade. 2.6. The MAP Kinase Cascades Interfere with the Cell Survival after Stress Stresses such as heat shock result in a variety of cellular responses that lead to increased stress resistance (see chapter by Li et al., this volume). Contribution of different stressactivated signal cascades to stress resistance will be presented. The S. cerevisiae slt2/mpk1 gene is essential for survival at high temperature (37°C) in normal medium (Kamada et al., 1995; Torres et al., 1991). Cells disrupted for the slt2/mpk1 gene lyse rapidly at 37°C with a phenotype similar to that of actin and polarized growth mutants (Mazzoni et al., 1993). The temperature sensitivity of the yeast cells disrupted for the slt2/mpk1 gene can be rescued by growing the cells at high osmolarity. The HOG cascade which mediates the response of cells to exposure to high osmolarity is not however, responsible for this rescue but the deletion of antioxidant genes controlled by the HOG cascade contributes to the lethality after heat shock in S. cerevisiae (Davidson et al., 1996). In mammalian cells, ERK and JNK-p38 MAP kinases show opposing effects on apoptotic death. Treatments that induce apoptosis, such as nerve growth factor


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withdrawal from PC 12 pheochromocytoma cells in culture, activate the JNK-p38 MAP kinases and apoptosis was prevented by the expression of recombinant dominant negative p38 or JNK mutants (Xia et al., 1995). In contrast, constitutive activation of the ERK signalling pathway prevented apoptosis. Expression of a constitutive JNK activator (but not of an ERK constitutive activator) in Swiss 3T3 and REF52 fibroblasts enhanced the apoptotic response to UV irradiation (Johnson et al., 1996). Blockade of p38 MAP kinase and JNK activity prevented cell death in rat fibroblasts following UV irradiation, heat shock (Zanke et al., 1996) and exposure to elevated concentrations of sodium salicylate (Schwenger et al., 1997). However, inhibition of the p38 MAP kinase did not affect the toxic effects of tumour necrosis factor in murine cells (Beyaert et al., 1996).

3. HSP PHOSPHORYLATION TRIGGERED BY ENVIRONMENTAL STIMULI In the following paragraphs, we will review first the protein kinases pathways involved in small Hsp phosphorylation and second the biological implications proposed for this modification. 3.1. Protein Kinases and Phosphatases in the Phosphorylation of Small HSPs The phosphorylation status of a protein is determined by the balance between kinases and phosphatases. In mammalian cells, protein kinases which are activated to phosphorylate small HSPs in response to environmental stimuli have been identified (Guesdon et al., 1993; Stokoe et al., 1992b; Zhou et al., 1993). Two proteins of 45-kDa/54-kDa showing stress induced Hsp27 kinase activity have been characterized in hamster cells (Huot et al., 1995). In murine cells, two related 45-kDa/ 55-kDa kinases are activated by the protein synthesis inhibitor, anisomycin, and the epidermal growth factor (Cano et al., 1996). These kinases are easily detected by in-gel kinase assays. The small HSP kinases are activated in vitro by MAP kinases (see previous section). An Hsp27 kinase was identified as MAP kinase activated protein kinase 2 (MAPKAPK2) (Stokoe et al., 1992a). MAPKAPK2 phosphorylates human Hsp27 at serines 15, 78 and 82 (Landry et al., 1992) and murine Hsp25 at serines 15 and 86 (there is no serine at the position corresponding to 78 in the human protein) (Gaestel et al., 1991). Interestingly, these phosphorylation sites share with the MAP kinase activated protein kinase 1 (MAPKAPK1) the Arg-X-X-Ser motif. However, MAPKAPK2 differs from MAPKAPK1 in its requirement for a bulky hydrophobic residue in the -5 position Nterminal to the serine (Stokoe et al., 1993). Several MAPKAPK2 cDNAs have been cloned (Stokoe et al., 1992a; Zu et al., 1994) and recently a related but distinct cDNA has been isolated. This homologue is identical to 3pK, a kinase gene located in the gene region of the small cell lung cancer tumor suppressor (Sithanandam et al., 1996). The corresponding recombinant protein product shows Hsp27 kinase activity and is also designated as MAPKAPK3 (McLaughlin et al., 1996). The in vivo role for this kinase in phosphorylation of small HSPs remains unclear.


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Less is known with respect to the phosphatases controlling small HSP phosphorylation. In cell-free systems, the human small HSPs are dephosphorylated by the calcium/calmodulin-dependent protein phosphatase, calcineurin (PP2B) (Gaestel et al., 1992), and type A protein phosphatase (PP2A) (Cairns et al., 1994). In vivo, however, the dephosphorylation of small HSPs is insensitive to cyclosporin A, a potent inhibitor of PP2B, suggesting that this phosphatase is not involved. The involvement of PP2A in small HSP dephosphorylation in vivo remains to be determined. Taken together, it seems possible that both, kinase activation and phosphatase inactivation contribute to the increased phosphorylation of small HSPs in response to stress. 3.2. The p38 MAP Kinases, Upstream Activators of the Small HSP Kinase (MAPKAPK2) Although the MAP kinases of the ERK type are activated by inducers of the small HSP phosphorylation, the activation of MAPKAPK2 by ERK1/2 became questionable when it became apparent that MAPKAPK2 activation did not strictly correlate with ERK1/2 activation (Guesdon et al., 1993). The MAPKAPK2 is activated in vitro both by the ERK type and by the p38 type MAP kinases (Guay et al., 1997). In mammalian cells, the p38 MAP kinase are activated by a wide variety of stresses including heat shock, oxidative stress (Rouse et al., 1994), hyperosmolarity (Han et al., 1994), endotoxic lipopolysaccharide (LPS) (Han et al., 1994), TNF(Beyaert et al., 1996) and interleukin-1 (IL-1) (Freshney et al., 1994). All these treatments also lead to the phosphorylation of small HSPs (reviewed in Arrigo and Landry, 1994). Most fascinating is the discovery of drugs which inhibit specific MAP kinase cascades (Cohen, 1997). PD098059 is a specific inhibitor in vitro of MEK1 and MEK2, the ERK1/2 activating kinases. In vivo, this compound prevents ERK2 activation but had no influence on p38 MAP kinase and MAPKAPK2 activation. The anti-inflammatory drug, CSAID/SB203580, binds with high affinity to the p38 MAP kinase and inhibits its activity with an extremely high specificity. Stress activation of MAPKAPK2 and phosphorylation of Hsp27 is suppressed in mammalian cells exposed to micromolar concentrations of SB203580 (Ben-Levy et al., 1995; Cuenda et al., 1995; Guay et al., 1997). This result provides strong support for the involvement of a p38 MAP kinase and MAPKAPK2 in phosphorylation of the small HSP during stress. The B-crystallin is another small HSP phosphorylated in response to stress and depending on the stress, either the ERK or the p38 MAP kinase is involved (Ito et al., 1997). The phosphorylation of small HSPs induced by lymphokines is abolished in cells treated with two tyrosine kinase inhibitors, genistein and herbimycin A (Ahlers et al., 1994), suggesting an involvement of a tyrosine kinase upstream in the activation pathway. 3.3. Phosphorylation of the Small HSPs and Cytoskeleton Dynamics Phosphorylation of the small HSPs often parallels the formation of large protein aggregates. It has been suggested that phosphorylation contributes to the dissociation of these aggregates (Kato et al., 1994; Lavoie et al., 1995; Mehlen et al., 1995). But most significantly, phosphorylation of the small HSPs might modulate fluid phase pinocytosis and actin microfilament dynamics (reviewed in Landry and Huot, 1995). In vitro, the


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unphosphorylated chicken small HSP inhibits actin polymerization (Miron et al., 1991), and phosphorylation of the murine Hsp25 abolishes its actin-depolymerizing activity (Benndorf et al., 1994). The human Hsp27 is accumu-lated in motile cell protrusions such as lamellipodia, filopodia, and membrane ruffles. These structures also stain for a number of known and suspected actininteracting proteins thought to regulate the dynamics and structure of F-actin. Collapse of the actin microfilaments occurs in response to growth factors, heat and oxidative stresses (Huot et al., 1995; Lavoie et al., 1995). Hsp27 overexpression in hamster cells enhances cell survival in the presence of the actin polymerization inhibitor, cytochalasin D (Lavoie et al., 1995). This protective effect does not occur with a triple phosphorylation site mutant of human Hsp27. Inhibiting the phosphorylation of Hsp27 with compound SB203580 (see below) suppresses the protective effects of Hsp27 overexpression (Guay et al., 1997). Oxidative stress induces both Hsp27 phosphorylation and severe fragmentation of the F-actin network, however the SB203580 compound blocked these responses in cells which express high levels of Hsp27 (Huot et al., 1997). Expression of small HSPs in murine cells was reported to confer resistance to TNF- and oxidative stress-induced necrotic (Huot et al., 1996; Mehlen et al., 1995) and apoptotic (Mehlen et al., 1996) cell death. Hamster cells stably transfected with DNA encoding human Hsp27 show an increased resistance to heat, oxidative stress and/or to actin depolymerizing agents but this effect is abolished upon replacement of the phosphorylated serines by glycines (Huot et al., 1996; Lavoie et al., 1995). Yet it should be mentioned that in transformed murine fibroblasts transiently transfected with a murine Hsp25 expression vector, replacement of the phosphorylated serines by alanines did not diminish the increased resistance to heat (Knauf et al., 1993) and negative results were obtained in other laboratories which used cell lines constitutively expressing small HSPs (Arata et al., 1995; Beyaert et al., 1996). The SB203580 compound had little effect on fibroblasts which do not spontaneously express Hsp27, however this influence was acquired if these cells were forced to permanently express high levels of Hsp27 after transfection (Huot et al., 1997). Differences in constitutive levels of Hsp27 in the various systems investigated may explain the conflicting results. Interestingly, the Rho family of GTPases which indirectly controls the actin cytoskeleton dynamics (reviewed in Symons, 1996; Zigmond, 1996) also contribute to the control of Hsp27 phosphorylation as upstream activators of the p38 MAPK kinase cascade. Stress-induced phosphorylation of non-heat shock proteins may also modulate the cytoskeleton dynamics. For example stathmin is phosphorylated on MAP kinase sites in response to a wide variety of treatments, including growth factors and stress such as heat shock (Beretta et al., 1995). Stathmin destabilizes microtubules in vitro and it has been suggested that phosphorylation controls this activity in vivo (Belmont and Mitchison, 1996). Thus, the environmentally-induced phosphorylation of the small HSPs and of stathmin may influence the cyto-skeleton dynamic.


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4. STRESS-INDUCED PHOSPHORYLATION OF TRANSCRIPTION FACTORS Numerous putative targets of the ERK/MAP kinases, p38 and JNK have been described. Some of them are involved in transcriptional regulation (reviewed in Cahill et al., 1996; Treisman, 1996). The following paragraphs will review the involvement of MAP kinases in the regulation of gene expression. 4.1. Transcription Factors Phosphorylated by MAP Kinases In mammals, the MAP kinases phosphorylate and modulate the activity of numerous transcription factors including c-Jun, ATF-2, Elk-1 (Raingeaud et al., 1996; Treisman, 1996; and references therein) and CHOP (also known as growth arrest and DNA damageinducible gene 153, GADD153) (Wang and Ron, 1996). Some of these factors are targets for specific kinases, e.g. c-Jun for the JNKs and CHOP for the p38 MAP kinase. Phosphorylation of c-Jun by JNK both targets c-Jun for ubiquitination and prolongs its half-life (Fuchs et al., 1996). The cyclic-AMP response element binding protein, CREB or CRE-BP1, and ATF-1 are phosphorylated downstream the p38 MAP kinase in response to stress or fibroblast growth factor (FGF). Both MAPKAPK1/RSK2 and MAPKAPK2 are CREB kinases in vitro (Tan et al., 1996; Xing et al., 1996). Other factors such as ATF-2, a member of the CREB subfamily of transcription factors but unresponsive to cyclic AMP, or Elk-1, a member of the Ets family, integrate signals from different kinase cascades (Cahill et al., 1996). In addition to transcription factors, chromatin components are MAP kinase targets as well. In particular, hyperphosphorylation of the Sir3p yeast protein through a MAP kinase cascade strengthens transcriptional silencing during mating, starvation and heat shock (Stone and Pillus, 1996). 4.2. Phosphorylation of the RNA Polymerase II Largest Subunit by Stressactivated CTD Kinases In many cells including Drosophila cells, mild heat shock causes dephosphorylation of the C-terminal domain (CTD) of the largest subunit of RNA polymerase II whereas upon severe heat shock, however, the RNA polymerase largest subunit accumulates in a form that remains multiphosphorylated on the CTD (Dubois et al., 1994; Venetianer et al., 1995) (reviewed in Dubois and Bensaude, 1998). Numerous studies have established that the CTD undergoes a cycle of phosphorylation/dephosphorylation in connection with gene transcription (Dahmus, 1996). RNA polymerase II with an unphosphorylated CTD assembles on the promoters and phosphorylation of the CTD appears to be required to release the interactions between the polymerase and the transcription factors sitting on the promoters. Likewise, RNA polymerase assemblies that are hypophosphorylated on their CTD bind to the 5' end of heat shock genes and pause after initiating transcription (O’Brien et al., 1994; Weeks et al., 1993). Phosphorylation occurs upon entry into


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elongation of transcription and is next required to dock the various factors involved in mRNA maturation (splicing, polyadenylation) (Steinmetz, 1997). The CTD is phosphorylated within the preinitiation complex of transcription by a CTD kinase associated with the TFIIH general transcription factor (Seroz et al., 1995; Valay et al., 1995). Furthermore, TFIIH subunits accumulate within the coiled bodies which contain splicing factors (Grande et al., 1997; Jordan et al., 1997). The CTD dephosphorylation observed during mild heat stress relates to the impairement of TFIIH-associated kinase (Dubois et al., 1997). As the heat shock genes remain very efficiently transcribed in these conditions, they might differ from other cellular genes in their requirements for an active TFIIH kinase. Heat stress activates a CTD kinase which copurifies with ERK type MAP kinases. The stressactivated MAP kinases might rescue the impairement of the TFIIH-associated CTD kinase. As initiation of transcription requires an unphosphorylated CTD, the hyperphosphorylation of the CTD observed during severe stress may contribute to a general shut-off of transcription. The CTD phosphorylated in mammalian cells during severe stress or in vitro with purified MAP kinase lacks a particular epitope associated with splicing factors and generated in vitro by phosphorylation with TFIIH (Dubois et al., 1997). Data obtained with insect cells are also suggestive of abnormal phosphorylation of the CTD from polymerases engaged in heat shock gene transcription (Weeks et al., 1993). Thus the abnormal CTD phosphorylation occuring during heat shock may also relate to the impairement of splicing occuring during such stress. Noteworthy, the predominant heatinducible hsp70 genes have no introns in insects and vertebrates.

5. INVOLVEMENT OF MAP KINASES IN STRESS GENE TRANSCRIPTION 5.1. Heat Shock Element (HSE) Dependent Transcription A connection exists at least at some conditions between the MAP kinase cascades and the heat shock transcription factor (HSF)-dependent heat shock response. The stress activation of heat shock gene transcription is inhibited in yeast and rat cells in which the Ras pathway is constitutively active (Engelberg et al., 1994). Activation of ERK kinases has been suggest to inhibit heat shock response in murine fibroblasts (Mivechi and Giaccia, 1995). Indeed, the heat shock transcription factor, HSF-1, is an in vitro substrate for ERK MAP kinases and constitutive activation of these kinases represses heat shock gene transcription at controlled temperatures (Chu et al., 1996a; Knauf et al., 1996) (see chapter Morimoto). In constrast, p38 MAP kinase activation is required for osmotic stress induction of Hsp70 mRNA (but not for heat-shock induction) (Sheikh-Hamad et al., 1998). 5.2. Stress Response Element (STRE) Dependent Transcription The HOG kinase cascade activates the general stress response in S. cerevisiae (reviewed in Ruis and Schüller, 1995). This response is distinct from the HSF-dependent heat shock


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response and requires a regulatory DNA sequence, the stress response element (STRE) with the consensus sequence CCCCT. It mediates the stress induction of transcription of a set of genes including the cytosolic catalase T (ctt1) gene, the DNA damage responsive gene (ddr2) (Kobayashi and McEntee, 1993), the trehalose phosphate phosphatase (tps2) gene (Gounalaki and Thireos, 1994) and the glycerol-3-phosphate dehydrogenase (gpd1) gene. Furthermore, STRE elements are present in the promoters of heat shock genes containing heat shock elements such as ubi4, hsp12, hsp104 and a protein tyrosine phosphatase gene (ptp2) (Ruis and Schüller, 1995). It is interesting to note that many genes controlled by the HOG pathway are involved in either protection from or repair of damage. In S. pombe, the Hog1 kinase homologue, Spc1, mediates the stress induction of the gpd1 and tps2 genes as well as the pyp2 tyrosine phosphatase which inactivates spc1 (see above) (Degols et al., 1996). The Atf1 transcription factor, a homologue of the mammalian ATF-2 factor, mediates this general stress response (Shiozaki and Russell, 1996). In rat pheochromocytoma cells, arsenite treatment leads to enhanced expression of the C/EBP- and CHOP transcription factors (Fawcett et al., 1996). The CHOP factor has been postulated to act as a dominant-negative regulator of CCAAT/ enhancer-binding proteins (C/EBP) and may be involved in an autoregulatory loop. Heat and arsenite stress enhance the transcription of the c-fos proto-oncogene (Bukh et al., 1990; van Delft et al., 1993). This enhanced transcription is accompanied by a marked increase in c-fos mRNA stability (Andrews et al., 1987). The induction of c-fos is likely a consequence of JNK and p38 MAP kinase activation. Indeed, SB203580 suppresses the induction of c-Fos and c-Jun following anisomycin or UV treatments (Hazzalin et al., 1996). The c-fos mRNA accumulation might be responsible for heat-induced FGF gene expression (Erdos et al., 1995) and could lead, in some cases, to a stimulated cell cycle progression after mild stress (van Wijk et al., 1993). 5.3. GRP Gene Transcription Stress may occur in distinct organelles which poses the problem of signal communication between cellular compartments. Signal pathways sensing stress in the ER and transducing it to the nucleus have recently been discovered (see chapter by Morimoto). In yeast, one begins to understand the activation pathway to kar2 expression (the grp78/hsp70 homologue). The ire1/ern1 gene product is one of the sensors for unfolded proteins in the ER (Shamu, 1998; Welihinda et al., 1998). It is a protein kinase homologous to cdc28 and distinct from the MAP kinases. It is oligomerized and phosphorylated during signaling from the ER to the nucleus. The primary function of the Ire1 protein appears to stabilize the expression of the Hac1 transcription factor, that is homologous to the mammalian ATF andCREB transcription factors, by promoting appropriate HAC1 mRNA splicing. In mammalian cells, induction of Grp78 is inhibited by genistein, an inhibitor of tyrosine kinases such as c-Abl, suggesting an involvement of kinases in the signal transduction pathway. However the yeast Ire1p protein is inefficient in inducing the mammalian grp78 gene in an heterologous transfection assay (Cao et al., 1995). Signals from stressed ER leading to the induced synthesis of BiP/Grp78 are also efficient in inducing the transcription factor, CHOP (GADD153) (Wang et al., 1996) which is phosphorylated and activated in turn by p38 MAP kinase (Wang and Ron,


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1996). This transcription factor binds to specific regulatory DNA elements, the CCAAT sequences. Interestingly, transcription of the gene encoding the mammalian BiP/Grp78 following calcium stress is itself transduced by a CCAAT element proximal to the site of transcription initiation (Roy and Lee, 1995). It is unclear at present whether a synergy exists between the different regulatory pathways.

6. CONTROL OF PROTEIN SYNTHESIS Protein synthesis is rapidly arrested in eukaryotic cells submitted to heat shock (reviewed in Duncan, 1996; Brostrom et al., 1998). During recovery from stress, a preferential translation of the HSP mRNAs is observed. The translation of some stress regulated mRNAs might be initiated by a ribosome jumping mechanism operating when eukaryotic initiation factors are deficient (Yueh and Schneider, 1996). It is an attractive possibility that changes in the phosphorylation state of various components of the translational apparatus contribute to these alterations. 6.1. Dephosphorylation of Ribosomal Subunit S6 and Eukaryotic Initiation Factor 4E (eIF-4E) Phosphorylation of the ribosomal S6 protein increases the translation efficiency of mRNAs coding for proteins of the translational apparatus such as the eukaryotic elongation factor, eEF-1 (Stewart and Thomas, 1994). Heat shock induces the rapid dephosphorylation of S6 in growing cells (reviewed in Nover, 1991) and, in sharp contrast, the phosphorylation of S6 in quiescent fibroblasts (Jurivich et al., 1991). Two kinases which phosphorylate S6 in vitro, p70s6k and p90rsk S6 kinases, are activated in vivo by heat shock as well as by mitogenic stimulation (Kozma and Thomas, 1994). The p90rsk which is activated by the ERK/MAP kinases (see above), is referred as MAPK activated protein kinase 1 (MAPKAPK1) or rRSK2. The p70s6k or RSK1 is turned on by a transducing pathway which is specifically inhibited by the immuno suppressant rapamycin (Proud, 1996). p70s6k is the major physiological S6 kinase in mammalian cells. Both S6 kinases have additional targets distinct from S6. One of the p70s6k targets is the protein synthesis represser 4E-BP1 (BP-1; PHAS-I) (von Manteuffel et al., 1996). Phosphorylation of 4E-BP1 would disrupt its interaction with protein synthesis initiation factors, e1F-4E, allowing it to interact with the mRNA cap binding complex and restore cap-dependent protein synthesis (Lawrence and Abraham, 1997). Phosphorylation of 4EBP1 decreases during heat shock, and this decrease is aggravated in the presence of rapamycin which blocks p70s6k activity (Feigenblum and Schneider, 1996). Heat shock also triggers the dephosphorylation of the capbinding factor, eIF-4E. Consequently, an increased association of 4E-BP1 with eIF-4E is observed (Vries et al., 1997). The resulting inactivation of eIF-4E along with the dephosphorylation of 4E-BP1 and phosphorylation of eIF-2 might contribute to the selective translation of heat shock mRNAs in stressed cells (Yueh and Schneider, 1996). Multiple signaling pathways mediate the phosphorylation of eIF-4E in response to growth factor activation (Morley et al., 1997). Rapamycin (inhibitor of p70s6k activation), P98059 (MEK inhibitor),


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SB203380 (p38 MAP kinase inhibitor) all block eIF-4E phosphorylation. Interestingly, they delay recovery of protein synthesis after heat-shock (our unpublished results). Hence, the MAP kinase cascades appear to play an important role in recovery of protein synthesis after stress. 6.2. HSP Depletion Triggers the Phosphorylation of the Eukaryotic Initiation Factor 2 (eIF2 ) Lysates prepared from heat-shocked human cells are inactive for translation in vitro. This inactivation may in part be due to phosphorylation of the a subunit of initiation factor 2 (eIF-2 ) resulting from the stress activation of the heme-regulated inhibitor (HRI) or hemin-controlled eIF-2a kinase (HCR) (De Benedetti and Baglioni, 1986; Farrell et al., 1977). HRI or HCR is a protein kinase, that is activated by hemin deficiency (reviewed in Chen and London, 1995). Phosphorylation of this factor on serine 51 inhibits initiation of translation (Samuel, 1993). Although phosphorylation of eIF-2 is not the sole mechanism responsible for the shut-off of protein synthesis under heat stress conditions, there is evidence for an involvement of such mechanism. Expression of a phosphorylation-resistant eIF-2 mutant (serine 51 is replaced by an alanine) partially protects cells from the inhibition of protein synthesis in response to heat treatment (Murtha-Riel et al., 1993). Noteworthy, eIF2 is itself a heat shock protein (Colbert et al., 1987) which may contribute to a quicker recovery of the cells from translational arrest during the shut off phase of the heat shock response. In contrast to the HCR, the double-stranded RNA-activated protein kinase, PKR, is an eIF-2 kinase which is impaired during heat stress (Dubois et al., 1991). This kinase contributes to the antiviral effects of interferon through the inhibition of protein synthesis. Some viruses such as the influenza virus avoid the translational inhibitory effects of PKR by activating a cellular inhibitory protein, P58IPK. Interestingly, this protein possesses a significant homology to the J domain of DnaJ (Hsp40), and is inhibited through an interaction with Hsp40 (Melville et al., 1997). Interaction with HSPs is also involved in the control of HCR activity. In reticulocyte lysates, the inactive HCR is bound to Hsp90 (Rose et al., 1987), Hsp70 and Hsp56 (Matts et al., 1992). Interaction with Hsp90 is obligatory for HCR to acquire and maintain an activable conformation (Uma et al., 1997). Hsp70 binding to HCR stabilizes the inactive HCR. Limiting the amount of Hsp70 in the lysates reduces the concentration of hemin required to prevent activation of HCR (Gross et al., 1994). Addition of denatured proteins to the lysate activates HCR (Matts et al., 1993). Denatured proteins may bind to Hsp70 present in the lysate and deplete the lysate from “free” Hsp70. The dissociation of Hsp70 from HCR is strongly correlated with the activation of HCR suggesting that sequestration of Hsp70 might be the signal that leads to activation of HCR in response to heat shock. Since Hsp70 chaperones assist the folding of nascent polypeptide chains (Beckmann et al., 1990), the cellular economy might justify the arrest of protein synthesis when the pool of “free” Hsp70 is depleted.


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7. CONCLUSION The heat shock responses involve not only chaperones but also protein kinases, protein phosphatases and detoxifying enzymes, which all contribute to the cellular resistance to stress. The responses to stress activate various signaling cascades of protein kinases, yet the same cascades are activated by different stimuli generated by stress treatment as well as developmental processes. Different stimuli activate the same cascades through different pathways and each cascade targets a wide spectrum of functions including those affecting cell shape, protein synthesis and gene expression. A further level of complexity is brought about by the fact that a given protein may be targeted by different cascades and integrate their specific signals. As a whole, we begin to understand that the protein kinase cascades form an intricate network of interactions which regulates the cellular homeostasis. The signaling cascades represent an early step in the reaction of cells to an economy imbalance generated by stress.

8. ACKNOWLEDGMENTS I wish to thank Drs. A.P.Arrigo, M.-F.Dubois, M.Gaestel, J.Landry and A.Michels for critical reading of the manuscript.

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5. THERMOTOLERANCE AND STRESS RESPONSE: POSSIBLE INVOLVEMENT OF KU AUTOANTIGEN G.C.LI*, L.LI, D.KIM, A.NUSSENZWEIG, S.-H.YANG, P.BURGMAN H.OUYANG and C.C.LING Department of Medical Physics and Department of Radiation Oncology, Memorial Sloan-Kettering Cancer Center, New York, NY 10021

1. Introduction 2. Thermotolerance: a transient phenomenon 2.1. Induction and Development of Thermotolerance 2.2. Characterization of Thermotolerant Cells 3. Thermotolerance and heat shock proteins 3.1. Thermotolerance and heat shock protein Hsp70 3.2. Heat Shock Proteins and RNA Splicing 3.3. Heat Shock Proteins and Apoptosis 3.4. Thermotolerance and heat shock protein Hsp27 4. Regulation of heat shock response: possible involvement of Ku Autoantigen 5. Requirement for Ku80 in growth and V(D)J recombination 6. Conclusion 7. References

1. INTRODUCTION Mammalian cells, when exposed to a non-lethal heat shock, can acquire a transient resistance to one or more subsequent exposures at elevated temperatures. This phenomenon has been termed thermotolerance (Gerner 1983; Gerner, et al., 1975; Henle, et al., 1978; Henle, et al., 1976). On the molecular level, heat shock activates a specific set of genes, the so-called heat shock genes, resulting in the preferential synthesis of heat shock proteins (Lindquist 1986; Lindquist, et al., 1988; Morimoto, et al., 1990). *Corresponding author


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There is much data supporting the hypothesis that heat shock proteins play a key role in modulating the cellular response to heat shock and other environmental stress, and are involved in the development of thermotolerance (Landry, et al., 1989; Laszlo, 1988; Laszlo, et al., 1985; Li, 1985, Li, et al., 1992a; Morimoto, et al., 1994). We begin this review by summarizing our current knowledge about the induction of thermotolerance by heat shock and other stresses. We will examine the biochemical and molecular mechanisms underlying the induction of thermotolerance and consider the role that heat shock proteins play in its development and decay. Finally, we will discuss the possible involvement of Ku protein in the regulation of heat shock and other stress response of mammalian cells.

2. THERMOTOLERANCE: A TRANSIENT PHENOMENON 2.1. Induction and Development of Thermotolerance Exposure of mammalian cells in culture to temperatures above 40°C often leads to reproductive death. Although mammalian cells vary appreciably in their intrinsic thermal sensitivity (Raaphorst et al., 1979), clonogenic survival curves, when plotted as a function of duration of heating, are characterized by an initial shoulder region and then an exponential decrease (Dewey et al., 1980), both of which are dependent on the applied temperature and the duration of exposure. The kinetics of inactivating a wide variety of mammalian cells by heat shock has been analyzed in terms of Arrhenius plots, where 1/Do is plotted against the inverse of the absolute temperature (Do is the time required to reduce the survival on the exponential region to 37% of its initial value). For most cell lines, the Arrhenius plots exhibit two straight segments, with a break at or near 43°C (Bauer et al., 1979; Dewey et al., 1977; Westra, et al., 1971). Above that temperature, the activation energy is between 110 and 150 kcal/mol, consistent with the view that protein damage is responsible for cell death. At lower temperatures, the activation energy is around 300 to 400 kcal/mol. This change in slope of Arrhenius plots below 43°C was interpreted as a manifestation of the ability of the cells to develop thermotolerance (Li et al., 1980; Sapareto et al., 1978). Henle and Leeper (Henle and Leeper, 1976) and Gerner and Schneider (Gerner and Schneider, 1975) first showed that cultured mammalian cells exposed to a nonlethal heat treatment have the ability to develop resistance to a subsequent heat challenge. Thermotolerance can be induced by a short initial heat treatment at temperatures above 43°C followed by a 37°C incubation before the second heat challenge. Thermotolerance can also be induced during continuous heating at temperatures below 43°C (Gerweck, 1977, Harisiadis et al., 1977; Palzer et al., 1973; Sapareto et al., 1978). The effect of thermotolerance can be dramatic, increases in survival levels can be several orders of magnitude (Field et al., 1982; Gerner, 1983; Henle and Dethlefsen, 1978). The mechanism underlying thermotolerance may also be responsible for the variation in thermosensitivity; permanent heat resistance may simply be a genetic alteration of constitutive levels of macromolecules transiently induced in thermotolerant cells. Thermal sensitivity is affected by a number of factors: the cells’ proliferative and

Thermotolerance and stress response

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nutritional status (Gerweck, 1977, Goldin et al., 1981; Hahn 1982), the thermal history, the heat fractionation and sequencing, and the conditions during recovery (Gerner 1983, Henle and Dethlefsen, 1978). The effects of the temperature and the duration of the first heat treatment on the subsequent expression of thermotolerance were studied in detail by many investigators (Li et al., 1982). In general, if the temperature of the priming dose was below 43°C, thermotolerance was nearly fully expressed during the initial treatment. On the other hand, temperatures of 43°C or higher did not permit the development of thermotolerance during this first heat exposure; a subsequent incubation at 37°C was required. Li and Hahn (1980) proposed an operational model of thermotolerance based on these observations. The authors suggest that thermotolerance can be divided into three complementary and sometimes competing processes: an initial event (“trigger”); the expression of resistance (“development”), and the gradual disappearance of resistance (“decay”). In this model, thermotolerance develops in at least two steps. First, the initial event converts normal cells to the triggered state with a rate constant k1. This process very likely involves the activation of the heat shock transcription factor, HSF1 (Lis et al., 1993, Morimoto 1993). Second, these triggered cells are converted to thermotolerant cells with a rate constant kg. Above 42.5°C, k2=0, the triggered cells remain sensitive. When transferred to 37°C, kg > 0, and cells develop thermotolerance. This thermotolerant state is characterized by the elevated expression of heat shock proteins, enhanced protection and faster recovery from thermal damage. Finally, thermotolerant cells can revert to the sensitive state with a rate constant k3, which is generally less than k2. An Arrhenius plot of the induction of thermotolerance as measured by the clonogenic survival assay is shown in Figure 1A; shown for comparison are similar plots for the induced synthesis of one of the heat shock proteins (Hsp70) (Figure 1B), and for the activation of the heat shock transcription factor HSF1 (Figure 1C). The slopes of these plots yield

Figure 1 Phenomenological model of thermotolerance development and decay (upper panel), and Arrhenius plots for induction of thermotolerance,


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induction of heat shock protein Hsp70, and heat shock factor (HSF1). activation (lower panels). (A) Rate at which thermotolerance is induced, as a function of the duration of the triggering, is plotted against the inverse of the absolute temperature of the appropriate treatment; (B) Similar to (A), except that the time required to induce a maximum amount of Hsp70 was used to determine the ordinate. (C) The rate of maximum HSF1 binding activity to the heat shock element (HSE). of the rat heat shock promoter, determined by quantifying bands of HSE-HSF1 complexes that were obtained using the gel-mobility shift assay (Kim et al., 1995).

Table 1 Characterization of the thermotolerant mammalian cell: molecules, cellular structures or functions that are protected after a mild heat shock

Macromolecules, cellular structure and function

References

• DNA Polymerase activity (a)

(Chu, et al., 1987, Dikomey, et al., 1987)

Synthesis

(Van Dongen, et al., 1984)

rRNA synthesis

(Burdon 1986, Nover, et al., 1986)

Splicing

(Yost and Lindquist 1986)

Synthesis

(Hahn, et al., 1985, Mizzen and Welch 1988, Sciandra, et al., 1984)

Denaturation

(Lepock, et al., 1990, Nguyen, et al., 1989, Pinto, et al, 1991)

Removal of aggregates

(Kampinga, et al., 1989, Wallen, et al., 1990)

Con A capping

(Stevenson, et al, 1981)

Na+/K+ ATPase (b)

(Anderson and Hahn 1985, Burdon, et al., 1982)

Permeability

(Maytin, et al., 1990)

Insulin receptors

(Calderwood, et al., 1983)

Cytoskeletal reorganization

(Wiegant, et al., 1987)

cAMP levels

(Calderwood, et al., 1985)

• RNA

• Protein

• Membrane

• Cytoplasm

Thermotolerance and stress response ATP levels (c)

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(Lunee and Cresswell 1983)

• Cell cycle progression

(Van Dongen, et al., 1984)

(a) Not observed by (Jorritsma, et al., 1986, Kampinga, et al., 1985) (b) Anderson and Hahn, 1985, observed thermotolerance development for the ouabain binding capacity only, the heat sensitivity of the ATP hydrolyzing activity was unchanged in thermotolerant cells. (c) Only in L5178Y-S cells, no resistance was observed in Ehrlich ascites cells.

a value of 120 kcal/mole for the activation energy of the associated rate-limiting reaction, suggestive of protein unfolding or protein denaturation. The similarity between these graphs and the Arrhenius plot for cell killing at temperatures above 43°C (Dewey et al., 1977, Westra and Dewey, 1971) strongly suggests a common genesis for the inactivation of cells by heat, the induction of thermotolerance, the induction of Hsp70 synthesis and HSF1 activation. 2.2. Characterization of Thermotolerant Cells When cells are heat-shocked, energy is absorbed throughout the cell, damaging virtually all cellular structures/functions, resulting in cell killing (Laszlo, 1992a).

Table 2 Comparison of the damage and resistance induced in the membrane fraction (PF) and the nuclear fraction of HeLa S3 cells by different resistance inducing agents

TTR10 damage to membranes (by agent) C

enhanced resistance of membranes

damage to nuclei (by agent)

enhanced resistance of nuclei

(1.0)

HTT 2.3

yes

yes

yes

yes

ATT 1.8

yes (a)

yes

no

no

DTT 2.5

n.d.

no

yes

yes

ETT

no

no

yes

yes

2.3

As a measure for the induced thermotolerance, the thermotolerance ratio is given (TTR10: ratio of heating times required to reduce survival to 10%). Thermotolerance is induced as follows: HeLa S3 cells were pretreated with heat (15 min, 44°C+5 hr, 37°C: HTT) , sodium arsenite (1 hr, 100 mM+5 hr, 37°C: ATT), diamide (1hr, 500 M+5 hr, 37°C: DTT) or ethanol (1 hr, 6%+4 hr, 37° C: ETT) (Burgman, et al., 1993, Kampinga, et al., 1995). C: control, non-tolerant cells, n.d.=not determined, (a): taken from (Yih, et al., 1991).


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In thermotolerant cells, many of these structures/functions are more resistant to heatinduced modifications. The data in Table 1 show that thermotolerance develops for nuclear, cytoplasmic and membrane components. Thus, there seems to be no spatial preference in the cell for the expression of heat-induced thermotolerance. However, an increasing amount of data suggest that the development of thermotolerance can be more specific than previously assumed, depending on the agents used for induction. For example, when thermotolerance is induced by sodium arsenite, diamide or ethanol, thermotolerance only develops in cellular fractions that are damaged during the thermotolerance-inducing treatment (Table 2). The subcellular localization of the induced resistance appears to correspond with the localization of the damage induced during the tolerance-inducing treatment. For instance, Lunec and Cresswell (Lunec et al., 1983) observed an enhanced resistance of the ATPsynthesis in thermotolerant cells, but only in a cell line (L5178Y-S) in which the ATPsynthesis was impaired by the tolerance-inducing heat treatment. In another cell line (Ehrlich ascites) this treatment had no effect on the ATP-synthesis, and in the thermotolerant cells no enhanced resistance of this process was observed. In firefly luciferase-transfected cells, Nguyen et al., (Nguyen et al., 1989) found that heat treatments (42°C) that decreased the luciferase activity but not protein synthesis in these cells, only induced resistance for the luciferase activity but not for protein synthesis. On the other hand, after a priming treatment at a higher temperature (45°C), both activities were impaired; and, in the thermotolerant cells resistance against a subsequent heat treatment was observed for both the luciferase activity and protein synthesis. Anderson and Hahn (Anderson et al., 1985) showed that the correlation between damage and induced resistance was even true for multidomain proteins in which the domains differ in heat sensitivity. These authors, working on Na+/K+ATPase, reported that thermotolerance could be induced in a heat sensitive domain of the protein (the ouabain binding domain) without changing the heat sensitivity of a more heat resistant domain (the ATPhydrolyzing domain). Such a direct correlation between damage and tolerance opens the possibility to study the role of inactivation of different proteins/structures in hyperthermic cell killing.

3. THERMOTOLERANCE AND HEAT SHOCK PROTEINS 3.1. Thermotolerance and Heat Shock Protein Hsp70 The mechanism underlying thermotolerance is not well understood, although many studies suggest that the heat shock proteins (HSP) are involved in its development (Landry et al., 1982; Li et al., 1982; Subjeck et al., 1982). Qualitative evidence exists for a causal relationship between HSP synthesis and thermotolerance (Landry et al., 1982; Laszlo and Li, 1985; Li, 1985; Li and Werb, 1982; Subjeck et al., 1982): (i) heat shock induces transiently enhanced synthesis of HSP that correlates temporally with the development of thermotolerance; (ii) the persistence of thermotolerance correlates well with the stability of HSP; (iii) agents known to induce HSP induce thermotolerance; (iv) conversely, agents known to induce thermotolerance induce HSP (Amici et al., 1993;


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Boon-Niermeijer et al., 1988; Boon-Niermeijer et al., 1986; Burgman et al., 1993; Crete et al., 1990; Hahn et al., 1985; Haveman et al., 1986; Henle et al., 1986; Kampinga et al., 1992; Laszlo, 1988; Lee et al., 1987; Li, 1983; Li et al., 1978; Li et al., 1986; Ritossa, 1962; Ritossa, 1963); and (v) stable heat-resistant variant cells express high levels of HSP constitutively (Laszlo and Li, 1985). Quantitatively, of the many HSP preferentially synthesized after heat shock in mammalian cells, the concentration of the 70-kDa heat shock protein (Hsp70) appears to correlate best with heat resistance, either permanent or transient (Laszlo and Li 1985, Li 1985, Li and Werb 1982). However, a good correlation between a 27-kDa heat shock protein (Hsp27) and thermal resistance also has been reported (Landry et al., 1989). In mammalian cells, three types of experiments (e.g., microinjection of affinitypurified anti-Hsp70 antibodies or Hsp70 protein, amplification of hsp70 promoter sequence) were performed before 1990 to vary the intracellular concentration of Hsp70 and to correlate this change with thermal-stress response (Johnston et al., 1988; Li, 1989; Riabowol et al., 1988). More recently, the expression of hsp70 under heterologous promoters has yielded additional insight into its structure and function. Transient expression of Drosophila Hsp70 in monkey COS cells demonstrated that Hsp70 accelerates the recovery of cell nucleoli after heat shock (Munro et al., 1984). Using a retroviral-mediated gene transfer technique, Li et al., have generated rat cell lines stably and constitutively expressing a cloned human hsp70 gene (Li et al., 1992a). These cell lines provide a direct means of studying the effects of Hsp70 expression on cell survival after heat shock. It is clearly demonstrated (Figure 2A) that the clonogenic survival of five independent pools of MVH-infected Rat-1 cells (constitutively expressing human Hsp70)

Figure 2 (A) Expression of human Hsp70 gene confers thermal resistance to


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Rat-1 cells. Survival at 45°C of pooled MVH-infected (expressing human Hsp70), MV6-infected (vector control), and uninfected Rat-1 cells. Survivals from five pooled populations of MVH-infected cells independently derived from separate infection experiments are shown (open symbols). Each pool is derived by pooling 200–600 colonies. Survival values after 30-min heating at 45°C are clustered around 10% for all cells and are, therefore, omitted for clarity (redrawn from Li et al. PNAS, 1992). (B) Cells expressing antisense Hsc70 (designated A12 and A13). show increased thermal sensitivity. Cellular survivals after heat shock at 45 °C were determined by the colony formation assay.

were approximately 100-fold higher (for a 60 or 75 min, 45°C heat treatment) than that of the parental Rat-1 or the MV6-infected Rat-1 cells (vector control). Similar results were obtained for individually isolated clones of Rat-1 cells expressing human Hsp70 (Li et al., 1992a). Furthermore, cells expressing higher levels of human Hsp70 generally survive thermal stress better than cells expressing lower levels, and there appears to be a good correlation between levels of exogenous human Hsp70 expressed and the degree of thermal resistance (Li et al., 1992a). These data provide direct evidence for a causal relation between expression of a functional form of mammalian Hsp70 and survival of cells at elevated temperatures. However, one should note that production of Hsp70 is only part of the program of protein biosynthesis initiated after heat shock, and other components of this response might also enhance cell survival (see chapter Lindquist et al., for role of Hsp104 in yeast). It is generally believed that Hsc70, a constitutively expressed member of Hsp70 family, binds to nascent polypeptide chains to prevent misfolding and aggregation (Hightower et al., 1994; see chapter Welch et al.). It functions cooperatively with Hsp40 and CCT in the folding of nascent polypeptide chains in a large assembly in the cytoplasm of eukaryotic cells (Frydman et al., 1994). Hsc70 also functions to chaperone proteins to organelles such as nuclei, mitochondria and endoplasmic reticulum (Brodsky et al., 1994; Langer et al., 1994). However, experimental evidence for these roles of Hsc70 has been derived mostly from in vitro studies; in vivo experiments using mammalian cells are lacking. In order to understand the functional roles of Hsc70 in mammalian cells, Li and Hightower (Li et al., 1995) used a dexamethasone (Dex)inducible antisense RNA expression vector to generate pseudomutants of NIH3T3 cells containing reduced levels of Hsc70, and to examine and identify phenotypic alteration in these cells. These investigators observed that in Dex-treated cultures, hsc70 antisense RNA blocked the induction of Hsp70 by heat shock. On the other hand, Hsc70 protein synthesis did not decrease in either Dex-treated or untreated cells. Taken together, these data suggest that antisense RNA was either less effective in reducing hsc70 RNA in Dexinduced cultures, or that cells compensated by producing more hsc70 RNA. Alternatively, it is plausible that newly synthesized RNA was targeted effectively before it became translationally active. To verify this hypothesis, Dex-treated cultures were made quiescent via serum starvation, and then restimulated with serum to induce RNA and protein synthesis. Consistent with the above hypothesis, Hsc70 synthesis was blocked in cultures expressing antisense RNA (Li and Hightower 1995).


93

The role of Hsp70/Hsc70 in thermotolerance development was also examined using a cell line that constitutively expresses antisense Hsc70 RNA (Nussenzweig et al., 1993). The antisense RNA was designed to also target Hsp70, which is > 75% homologous to Hsc70. Similar to the finding of Li and Hightower (Li and Hightower, 1995), heatinduced Hsp70 expression is significantly reduced and delayed in the transfected cells relative to the parental Rat-1 cells. For example, after a 45°C, 15 min heat shock, increased Hsp70 protein level is apparent within 4 hours in Rat-1 cells, 2 to 4 hours earlier than in the antisense RNA-transfected cells. In contrast, there were no significantly detectable changes in Hsc70 level in these cell lines. This is also in agreement with previous attempts to regulate gene expression by antisense RNA in which a large excess of antisense over sense transcripts is often necessary to produce significant results. Consistent with the above, the antisense transfected cells are more heat-sensitive than Rat-1 cells (Figure 2B), and show a profoundly reduced thermotolerance level as well as a delay in thermotolerance development. It is well established that heat shock inhibits RNA and protein synthesis. This inhibition is reversible, and the transcriptional and translational activity recovers gradually when heated cells are returned to 37°C. Thermotolerant cells subjected to heat shock exhibit less translational inhibition (Liu et al., 1992; Mizzen et al., 1988). The time required for the recovery of RNA and protein synthesis upon returning the heated cells to 37°C is also found to be considerably shorter for thermotolerant cells than for control nontolerant cells (Black et al., 1989; Laszlo, 1992b; Liu et al., 1992). The role of Hsp70 in these processes was examined using

Table 3 Comparison of heat resistance and recovery from heat-induced inhibition in RNA and protein synthesis relative to control cells

Schematic diagrams of human hsp70 a

Hsp70b Deletion

Heat c Resistance

Recovery after d Heat Shock RNA

protein 0.94 ±0.073

intact hsp70

none

++++

0.86±0.11

hsp70

Bgl

120–428

+++

0.16 ±0.03 0.22 ±0.065

hsp70

Sma-p 438–618

–

0.13 ±0.046

0.15 ±0.03

hsp70

S-C

438–478

+

0.12 ± 0.033

0.22±0.12

hsp70

C-S

478–618

+

0.15 ± 0.025

0.20±0.12

Schematic diagrams of human hsp70 gene and its deletion derivatives are shown on the left with relevant restriction sites. B, BamHI; H, Hind III; S, SmaI; C, ClaII; Bg, Bgl II; nucleotides


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encoding a hexapeptide substance P epitope are represented by the dark box sub-P. The ATPbinding and nucleolar/nuclear localization domains are indicated as a checkerboard or a thinhatched bar, respectively. bDeletion is given in codons (all deletions are in-frame). c++++: 104-fold increase in survival,+++: 102-to 103-fold increase, +: 10- to 102-fold increase, and -: no increase in survival compared to control Rat-1 and MV6 cells. Heat treatment used: 45° C, 60 min. dRecovery of RNA and protein synthesis at 8 hr after 45°C, 25 min heat shock was determined. The relative rate of RNA and protein synthesis was normalized to the unheated control group. Average values ± SD are presented here. At 8 hr after heat shock, the relative RNA synthesis of Rat-1 and MV6 cells is 0.24±0.09 and 0.16±0.03, respectively; the relative protein synthesis of Rat-1 and MV6 cells is 0.35±0.13 and 0.12 ± 0.09, respectively. The differences observed between M21 cells and other cells are statistically significant (pS). and its mixed disulfide with glutathione. Biochemistry , 31 , 9288– 9293. Chen, R. and Henning, U. (1996). A periplasmic protein (Skp). of Escherichia coli selectively binds a class of outer membrane proteins. Mol. Microbiol , 19 , 1287–1294. Creighton, T.E. (1986). Detection of folding intermediates using urea-gradient electrophoresis. Methods Enzymol. , 131 , 156–172. Crooke, H. and Cole, J. (1995). The biogenesis of c-type cytochromes in Escherichia coli requires a membrane-bound protein, DipZ, with a protein disulphide isomerase-like domain. Mol. Microbiol. , 15 , 1139–1150. Danese, P. and Silhavy T.J. (1997). The sigma(E) and the Cpx signal transduction systems control the synthesis of periplasmic protein-folding enzymes in Escherichia coli. Genes Dev. , 11 , 1183–1193. Danese, P., Snyder, W.B., Cosma, C., Davis, L. and Silhavy, T. (1995). The Cpx twocomponent signal transduction pathway of Escherichia coli regulates transcription of the gene specifying the stressinducible periplasmic protease. Genes & Dev. , 9 , 387– 398. Dartigalongue, C. and Raina, S. (1998). PpiD is a new heat shock protein encoding a peptidyl prolyl isomerase. EMBO J. , in press. De Las Penas, A., Connolly, L. and Gross, C.A. (1997). The sigmaE-mediated response to extracytoplasmic stress in Escherichia coli is transduced by RseA and RseB, two negative regulators of sigmaE. Mol. Microbiol. , 24 , 373–385. Derman, A.I., Prinz, W.A., Belin, D. and Beckwith, J. (1993). Mutations that allow disulfide bond formation in the cytoplasm of Escherichia coli. Science , 262 , 1744– 1747. Fischer, G., Bang, H., Ludwig, B., Mann, K. and Hacker, J. (1992). Mip protein of Legionella pneumophila exhibits peptidyl-prolyl-cis/trans isomerase (PPIase). activity. Mol. Microbiol , 6 , 1375–1383. Fischer, G. and Schmid, F.X. (1990). The mechanism of protein folding. Implications of in vitro refolding models for de novo protein folding and translocation in the cell. Biochemistry , 29 , 2205–2212. Geyer, R., Galanos, C., Westphal, O. and Golecki, J.R. (1979). A Lipopolysaccharidebinding cell-surface protein from Salmonella minnesota. Isolation, partial characterization and occurrence in different Enterobacteriaceae. Eur. J. Biochem. , 98 , 27–38. Grauschopf, U., Winther, J.R., Korber, P., Zander, T., Dallinger, P. and Bardwell, J.C. (1995). Why is DsbA such an oxidizing disulfide catalyst? Cell , 83 , 947–955. Guilhot, C., Jander, G., Martin, N.L. and Beckwith, J. (1995). Evidence that the pathway of disulfide bond formation in Escherichia coli involves interactions between the cysteines of DsbB and DsbA. Proc. Natl Acad. Set. USA , 92 , 9895–9899. Grove, J., Tanapongpipat, S., Thomas, G., Griffiths, L., Crooke, H. and Cole, J. (1996). Escherichia coli K-12 genes essential for the synthesis of c-type cytochromes and a


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third nitrate reductase located in the periplasm. Mol. Microbiol. , 19 , 467–481. Hartl, F.U. (1996). Molecular chaperones in cellular protein folding. Nature , 381 , 571– 580. Hendrick, J.P. and Hartl, F.-U. (1993). Molecular chaperone functions of heat-shock proteins. Ann. Rev. Biochem. , 62 , 349–384. Holmgren, A. (1995). Thioredoxin structure and mechanism: conformational changes on oxidation of the active-site sulfhydryls to a disulfide. Structure , 3 , 239–243. Horne, S.M. and Young, K.D. (1995). Escherichia coli and other species of Enterobacteriaceae encode a protein similar to the family of Mip-like FK506-binding proteins. Arch. Microbiol. , 163 , 357–365. Iobbi-Nivol, C, Crooke, H., Griffiths, L., Grove, J., Hussain, H., Pommier, J., Méjean, V. and Cole, J.A. (1994). A reassessment of the range of c-type cytochromes synthesized by Escherichia coli K-12. FEMS Microbiol. Lett. , 119 , 89–94. Jander, G., Martin, N.L. and Beckwith, J. (1994). Two cysteines in each periplasmic domain of the membrane protein DsbB are required for its function in protein disulfide bond formation. EMBO J. , 13 , 5121–5127. Kamitani, S., Akiyama, Y. and Ito, K. (1992). Identification and characterization of an Escherichia coli gene required for the formation of correctly folded alkaline phosphatase, a periplasmic enzyme. EMBO J. , 11 , 57–62. Kawamukai, M., Matsuda, H., Fujii, W., Utsumi, R. and Komano, T. (1989). Nucleotide sequences of fic and fic-1 genes involved in cell filamentation induced by cyclic AMP in Escherichia coli. J. Bacterial , 171 , 4525–4529. Kishigami, S. and Ito, K. (1996). Roles of cysteine residues of DsbB in its activity to reoxidize DsbA, the protein disulphide bond catalyst of Escherichia coli. Genes to cells , 1 , 201–208. Kishigami, S., Kanaya, E., Kikuchi, M. and Ito, K. (1995). DsbA-DsbB interaction through their active site cysteines. J. Biol. Chem. , 270 , 17072–17074. Kleerebezem, M., Heutink, M. and Tommassen, J. (1995). Characterization of an Escherichia coli rotA mutant, affected in periplasmic peptidyl-prolyl cis/trans isomerase. Mol. Microbiol , 18 , 313–320. Kloser, A., Laird, M., Deng, M. and Misra, R. (1998). Modulations in lipid A and phospholipid biosynthesis pathways influence outer membrane protein assembly in Escherichia coli K-12. Mol. Microbiol. , 27 , 1003–1008. Kobayashi, T., Kishigami, S., Sone, M., Inokuchi, H., Mogi, T. and Ito, K. (1997). Respiratory chain is required to maintain oxidized states of the DsbA-DsbB disulfide bond formation system in aerobically growing Escherichia coli cells. Proc. Natl. Acad. Sci. USA , 94 , 11857–11862 Lazar, S. and Kolter, R. (1996). SurA assists the folding of Escherichia coli outer membrane proteins. J. Bacterial. , 178 , 1770–1773 Lipinska, B., Fayet, O., Baird, L. and Georgopoulos, C. (1989). Identification, characterization, and mapping of the Escherichia coli htrA gene, whose product is essential for bacterial growth only at elevated temperatures. J. Bacterial. , 171 , 1574– 1584. Liu, J. and Walsh, C.T. (1990). Peptidyl-prolyl cis-trans-isomerase from Escherichia coli: a periplasmic homolog of cyclophilin that is not inhibited by cyclosporin A. Proc. Natl. Acad. Sci. USA , 87 , 4028–32 Loferer, H., Bott, M. and Hennecke, H. (1993). Bradyrhizobium japonicum TlpA, a novel membrane-anchored thioredoxin-like protein involved in the biogenesis of cytochrome aa3 and development of symbiosis. EMBO J. , 12 , 3373–3383.

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14. ROLE OF CHAPERONES IN REPLICATION OF BACTERIOPHAGE LAMBDA DNA M.ZYLICZ*, A.WAWRZYNOW, J.MARSZALEK, K.LIBERER, B.BANECKI, I.KONIECZNY, A.BLASZCZAK1, P.BARSKI, J.JAKOBKIEWICZ, M.GONCIARZ-SWIATEK, M.DUCHNIEWICZ, J.PUZEWICZ and J.KRZEWSKA Department of Molecular and Cellular Biology, Faculty of Biotechnology, University of Gdansk, 80–822 Gdansk, Kladki 24, Poland 1 Polish Academy of Sciences, Institute of Biochemistry and Biophysics, Laboratory of Molecular Biology affiliated to the University of Gdansk, 80–822 Gdansk, Poland

1. Introduction 2. Early DNA Replication 2.1. Assembly of the O-some Structure 2.2. Assembly of the Preprimosomal Structure 2.3. Activation of the Preprimosomal Structure 2.4. Uni- and Bidirectional 3. 4. 5. 6.

Switch from Early to Late Concluding Remarks Acknowledgments References

DNA Replication DNA Replication

1. INTRODUCTION Genetic and biochemical studies on the involvement of Escherichia coli molecular chaperones in bacteriophage DNA replication gave the first evidence for the synergistic and cooperative action of the DnaK/DnaJ/GrpE molecular chaperone machine (for review see Georgopoulos et al., 1990, 1994). *Corresponding author


326

This particular system is now a paradigm for our understanding of fundamental biochemical processes, such as protein folding, intracellular protein translocation, selective proteolysis and autoregulation of heat shock response (for reviews see Hendrick and Haiti, 1993; Hartl, 1996). The in vitro replication system was also the first functional assay for purification of active DnaK, DnaJ and GrpE proteins (Zylicz et al., 1983; Zylicz and Georgopoulos, 1984; Zylicz et al., 1985, 1987). The replication of bacteriophage DNA depends on interactions between phageencoded replication proteins and the bacterial host’s replication machinery. The single cycle of reproductive growth which generates approximately 100 copies of viral genome may be divided into an early and a late phase (for review, see Furth and Wickner, 1983; Taylor and Wegrzyn, 1995). In the early phase the linear double

Figure 1 Model of the DNA replication cycle of bacteriophage . The phage particle attaches to the host E. coli cell and injects its linear, duplex DNA molecule. (1) the DNA molecule circularizes by the base-pairing of complementary single-stranded ends. The resulting nicked circle is closed by DNA ligase. (2) DNA gyrase introduces supercoils. (3) Transcriptional activation is required for the proper assembly of the ori O P-DnaB preprimosomal structure. (4) The DnaK/DnaJ/GrpE chaperone machine, in an ATP-dependent reaction, releases a fraction of the P protein, thus triggering the unidirectional unwinding of double stranded DNA, catalyzed by the DnaB helicase. Transcription, when transversing the ori sequence dissociates the O-some from the DNA complex, thus allowing DnaB helicase to unwind double-stranded DNA in both directions. (5) In

Role of chaperones in replication of bacteriophage lambda DNA

327

the presence of single-stranded-DNA binding protein (Ssb) and DNA gyrase, the DnaB-dependent unwinding of double stranded DNA proceeds, thus allowing the DnaG primase to synthesize RNA primers, which are subsequently extended into DNA by the DNA polymerase III. At early time after infection replication of most DNA molecules proceeds bidirectionally to generate structure—early replication (theta mode). The rolling-circle DNA replication starts with (4') the blocking of bidirectional unwinding of the double-stranded DNA at the ari sequence by transient inhibition of transcription (and/ or ClpX chaperone activity). The transcriptional activation could be modulated in the presence of Cro repressor and E. coli DnaA (see text for details). (5') In the presence of primase, Ssb, DNA gyrase and DNA polymerase III unidirectional DNA replication proceeds. (61) The collision of a DnaB helicase/DNA polymerase III replication fork complex with an undissociated O-some structure, following the completion of a round of unidirectional DNA replication may be the key event that triggers the initiation of a rolling-circle sigma mode of DNA replication (see text for more details). The concatameric DNA produced by rollingcircle replication can be cut at the cos sides and packaged into phage heads. The addition of tails completes the maturation of phage particles which after cell lysis are ready for a new cycle of infection. The exact stoichiometry of the proteins and the protein subunits are not shown.

stranded bacteriophage DNA is injected into the bacterial cell, where it is rapidly circularized and supercoiled (Figure 1). Then DNA replication is initiated at a single site, ori , and proceeds bidirectionally according to the theta ( ) mode. In the late phase, replication of DNA proceeds by a rolling-circle mechanism (a mode). During this process multiple-length double stranded DNA molecules are generated and then subsequently encapsidated (Figure 1). Only two phage proteins, encoded by genes O and P, appear to participate directly in the initiation and/or propagation of the replication forks. A small fragment of the λ genome, carrying only cro, O, P, the replication origin ori (located inside the O gene) and the promoter pR can replicate autonomously as a plasmid called

dv. The pR promoter is required not only for

expression of O and P, but also for the transcriptional activation of the ori sequence, an event which regulates the frequency of DNA replication (for review see Learn et al., 1993; Taylor and Wegrzyn, 1995). It is believed that the early theta mode of bacteriophage DNA replication is mimicked by plasmid dv DNA (called also plasmid) replication. Most of the host genes required for DNA replication have been identified by challenging the phage to replicate at nonpermissive temperatures in E. coli mutants which are temperature-sensitive for replication of the bacterial chromosome (e.g. dnaB, dnaE, dnaG, dnaZ). The involvement of three other bacterial genes, namely dnaK, dnaJ and grpE were identified using a different approach (Georgopoulos and


328

Herskowitz, 1971; Saito and Uchida, 1977). These genes were originally discovered because a mutation in any of them blocked DNA replication at both permissive and nonpermissive temperatures. Moreover, mutations in the P gene ( / called also ) enable bacteriophage to replicate on hosts carrying these mutations. Subsequent studies established that any mutation (s) in these heat shock genes exerted global, pleiotropic effect on the host’s metabolism, including defects in DNA and RNA synthesis, proteolysis, cell division and overproduction of heat shock proteins (for review see Georgopoulos et al., 1990; Burkholder and Gottesman, this volume). 2. EARLY

DNA REPLICATION

The in vitro reconstitution of the dv plasmid DNA replication system using purified proteins has allowed the identification of intermediate reactions leading to the initiation of DNA replication (Mensa-Wilmot et al., 1989a; Alfano and McMacken, 1989a; Zylicz et al., 1989; Zylicz, 1993). The order of the events in the pathway of replication complex assembly reconstituted in vitro has recently been confirmed in vivo (Wegrzyn et al., 1995a). 2.1. Assembly of the O-some Structure Four dimers of the O initiation protein bind to the ori sequence at four repeating sequences (iterons) forming a large nucleosome-like structure called the O-some (Tsurimoto and Matsubara, 1981; Roberts and McMacken, 1983; Dodson et al., 1985; Liberek et al., 1988; Echols, 1990). Formation of the O-some complex changes the conformation of the ori sequence (Schnos et al., 1988; McMacken, unpublished results). On linear or relaxed DNA templates these topological changes are limited to iteron sequences. However, when in the presence of Mg2+ the O-some structure is formed on negatively supercoiled DNA, the alteration of DNA topology is transferred to the neighbouring 40 bp A/T-rich segment of ori . In this case the A/T-rich region does not become completely unwound, however it is significantly destabilized (McMacken, unpublished results). Such effects help probably to potentiate loading of DnaB helicase onto DNA within the onA sequence (see discussion below). The O protein, like several other DNA-binding replication proteins, has a tendency to aggregate (Wawrzynow et al., 1995a). The aggregated form of O protein is inert in DNA replication. It was shown in vitro that several bacterial molecular chaperones (ClpX, ClpA or DnaK/DnaJ/GrpE) can protect the O protein from aggregation and also dissociate the previously aggregated O (Wawrzynow et al., 1995a; Wawrzynow and Zylicz, unpublished results). Among them, the function of ClpX was studied in more detail. ClpX, the ATP-dependent substrate specificity component of ClpXP protease (Wojtkowiak et al., 1993; Gottesman et al., 1993), in the absence of the ClpP proteolytic subunit, possesses all the properties expected for a molecular chaperone (Wawrzynow et al., 1995a). The ClpX-dependent dissociation of O aggregates requires ATP hydrolysis catalyzed by ClpX ATPase. The chaperone effect of ClpX enhances the specific binding of O to ori , stimulating DNA replication in vitro. The biological role of the ClpX-


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dependent enhancement of the ori O complex formation is still not clear (Wawrzynow et al., 1996). We cannot exclude the possibility that the chaperone function could be required very early, before O protein synthesis is finished. Assuming that the ClpX ATPase can perform a chaperone function in promoting the de novo folding of proteins or repairing active protein structure and in the same time is involved in protein degradation, it is important to determine the regulatory mechanism responsible for the decision to either repair or destroy a protein substrate. A hypothetical model, based on the stability of ClpX-protein substrate complex has been recently proposed to address this problem (Wawrzynow et al., 1996). In vivo, O protein is extremely unstable (half life at 40°C is 1.5 min; Lipinska et al., 1980; Gottesman et al., 1981; Bejarano et al., 1993). The ATP-dependent serine protease ClpXP was identified as being involved both in vivo and in vitro in efficient O degradation (Wojtkowiak et al., 1993; Gottesman et al., 1993). In any purified replication system (s) lacking the RNAP, the binding of O to ori sequence protects the O protein from ClpXP-dependent degradation (Wawrzynow and Zylicz, unpublished results). By contrast, in a crude in vitro replication system based on bacterial protein extract (supplemented with purified O and P) such effects were not observed (Wojtkowiak et al., 1993). Recently, we were able to show that RNA transcription proceeding through the O-some structure, makes O accessible to the ClpXP protease (Wawrzynow and Zylicz, unpublished results). Thus in vitro the stability of the O-some structure depends both on transcription and the activity of ClpXP protease. Genetic experiments have shown that mutations in the clpX gene abolish O degradation, suggesting that other Clp ATPases do not substitute for ClpX in this reaction (Gottesman et al., 1993; Szalewska et al., 1994). However, these findings may not extrapolate to the ClpX chaperone function, and any interpretation of in vivo results obtained with ClpX mutants ought to take into account the possibility that ClpX chaperone activity may be substituted by other chaperones. In particular, it is possible that a close ClpX homologue, ClpY, that has recently been identified in E. coli (Missiakas et al., 1996), can substitute for ClpX. 2.2. Assembly of the Preprimosomal Structure A second phage protein involved in DNA replication is P which is responsible for sequestering a bacterial helicase from the host replication system making it available to the ori sequence. In bacterial cells the DnaB helicase (LeBowitz and McMacken, 1986) is responsible for the propagation of the replication fork during synthesis of the bacterial chromosome (Kaguni and Kornberg, 1984). The DnaB is complexed with the DnaC initiation factor which results in attracting the helicase to the chromosome origin sequence (Marszalek and Kaguni, 1994). After phage infection, when the intracellular level of P initiation protein increases, P protein wins the competition with DnaC for DnaB binding (Mallory et al., 1990) and the P-DnaB complex interacts with the Osome structure to form an ori O P-DnaB preprimosomal complex (Figure 2) (Dodson et al., 1985; Liberek et al., 1988; Alfano and McMacken, 1989a; Zylicz et al., 1989). The stoichiometry of the ori - O P-DnaB preprimosomal complex has not yet been resolved. Probably two P-DnaB complexes (each composed of a multimer of P


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and a hexamer of DnaB; Mallory et al., 1990) are positioned in the A/T-rich region, on the right side of the O-some. The asymmetry of the preprimosomal complex was suggested on the basis of the electron-microscopy studies (Dodson et al., 1985). The observed stability of ori O- P-DnaB complex assembled on the supercoiled DNA could be due to both specific P O protein-protein interactions (Tomizawa, 1971; Zylicz et al., 1984; Wickner and Zahn, 1986; Liberek et al., 1990) and the affinity of their protein components to single stranded DNA. In support of this it is not possible to isolate the stable O- P-DnaB complex in the absence of the DNA template (Zylicz, unpublished results). Futhermore, using UV cross-linking experiments it was possible to demonstrate that both O and P, but not DnaB, are in close contact with the ssDNA (Learn et al., 1997). Proper assembly of an ori O P-DnaB preprimosome has an absolute requirement for a negatively supercoiled DNA template (McMacken, unpublished results). Following the formation of the O-some structure, the DNA replication potential is rapidly lost upon complete removal of free negative DNA superhelical turns. By contrast, the ori O P-DnaB complex once preformed on the negatively supercoiled DNA retains some replication potential even several minutes after removal of free DNA supercoils (t1/2 30– 40 sec on linearized DNA, t1/2 3–4 min on HU coated DNA; McMacken, unpublished results). These results suggest that during DNA replication supercoiling is absolutely required only for proper formation of the ori O P-DnaB. McMacken and coworkers suggest that properly assembled ori - O P-DnaB nucleoprotein structure traps an “open” DNA conformation that is


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Figure 2 Chaperone-dependent activation of the DnaB helicase in the preprimosomal complex. The DnaK/DnaJ/GrpE-dependent activation of preprimosomal complex (reaction 4; Figure 1) can be described by the following steps: (4a) DnaJ chaperone binds to P and DnaB thus attracting (4b) the DnaK*-ADP chaperone to the preprimosomal structure. In the absence of DnaJ, DnaK in the DnaK*-ADP conformation does not form a stable complex with P. The presence of DnaJ changes the conformation of DnaK*-ADP in such mode that more stable preprimosomal-DnaK-ADP complex is formed. (4c) Following the GrpE-dependent ADP/ATP exchange and ATP hydrolysis, a portion of P is released from the preprimosomal complex and DnaK is converted back to the DnaK*-ADP conformation which possesses limited affinity to P but recognizes efficiently the DnaJ protein. Those molecules of DnaB helicase


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which are no longer associated with P proceed to unwind double-stranded DNA rightwards from the ori sequence. (4d) Transcriptional activation displaces or rearranges the O-some structure allowing the DnaB helicase to (4e) unwind DNA leftwards. The ori

iterons which are recognition sites for O are shown as black boxes. The (*) symbol represent the DnaK*-ADP conformation reached only after ATP hydrolysis, and not as a result of the simple preincubation of DnaK with ADP. The stoichiometry of the proteins and the protein subunits are not represented.

induced in the presence of Mg2+ at ori A/T-rich region upon the O-some formation on supercoiled DNA (McMacken, unpublished results). According to this interpretation DnaB helicase is not positioned between the partially unwound DNA strands. DnaK and DnaJ action is needed before DnaB can stably interact with DNA (Learn et al., 1997). Other researchers also speculate that the DnaK/DnaJ/GrpE chaperone machine could be involved in loading DnaB helicase between two partially unwound strands (Wegrzyn et al., 1996b). We suggest that transcriptional activation is required for the “rearrangement” of the protein-DNA complex at ori , localizing the DnaB helicase at the junction of double stranded and single stranded DNA. This hypothesis is supported by the fact that during the RNA polymerase action at least O, P as well as DnaB helicase must be present to initiate the ori -specific DNA replication (Yamamoto et al., 1987). Assembly of the active ori - O- P-DnaB preprimosome on relaxed or coated (by the histone-like protein, HU) DNA needs transcriptional activation (Mensa-Wilmot et al., 1989b). Only when the purified DNA replication system is devoid of HU protein and the DNA template is highly negatively supercoiled (- >0.045) is transcriptional activation not required for efficient DNA synthesis (Alfano and McMacken, 1989a; Zylicz et al., 1989). McMacken and co-workers suggested that transcription activates the DNA replication by counter-acting an inhibitory effect of HU protein, which constrains negatively supercoiled DNA upon binding. For example, an increase of the negative supercoiling behind the RNA polymerase during transcription influences the topology of the A/T-rich ori region in such a way that it helps O and P to trap partially unwound DNA strands. The idea that transcriptional activation of λ replication does not simply rely on dissociation of the HU protein from ori sequence thus allowing formation of the ori - O- P-DnaB complex is supported by the fact that it can emanate from promoters located either upstream or downstream from ori (Furth et al., 1982; McMacken, unpublished results). In the case of the properly assembled ori - O- P-DnaB preprimosome, transcribing RNA polymerase molecules which readily “bypass” the O-some structure, pause indefinitely at ori - O- P-DnaB complex (McMacken, unpublished results). Thus, the O protein remains in the preprimosome structure and is inaccessible to the ClpXP protease (Wawrzynow and Zylicz, unpublished results). Recent genetic studies support these in vitro findings (Wegrzyn et al., 1995a).


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2.3. Activation of the Preprimosomal Structure The preprimosomal ori - O- P-DnaB complex assembled on supercoiled DNA is stable, since it can be isolated by gel filtration and subsequently demonstrated to be still active in DNA replication without addition of further O, P or DnaB (Alfano and McMacken, 1988;1989b; Zylicz et al., 1989; Zylicz, 1993). The strong P-DnaB interaction required for assembly of DnaB helicase at ori also inhibits the ATPase and helicase activity of the DnaB (Wickner, 1978; Mallory et al., 1990). Subsequent activation reactions are needed to liberate the DnaB helicase from the inhibitory effects of P. The concerted action of DnaK, DnaJ and GrpE is respon-sible for the “liberation” (Figure 2) (Liberek et al., 1988; Alfano and McMacken, 1989b; Dodson et al., 1989; Zylicz et al., 1989; Hoffmann et al., 1992; Zylicz, 1993). The DnaJ chaperone binds to the preprimosome and further stabilizes its structure, by interacting primarily with P and DnaB (Alfano and McMacken, 1989b; Hoffman et al., 1992; Osipiuk et al., 1993; Zylicz, 1993; Wawrzynow and Zylicz, 1995). The presence of DnaJ bound to P facilitates the binding of the DnaK chaperone to the preprimosomal complex (Wawrzynow et al., 1995b). The binding of DnaK and DnaJ to the preprimosome was shown by using electron microscopy of gold-tagged antibodies against these proteins (Dodson et al., 1989). DnaK has a weak intrinsic ATPase activity (Zylicz et al., 1983; Buchberger et al., this volume). Depending on whether it is in ATP or ADP bound form DnaK possesses a different conformation and protein substrate affinity (Palleros et al., 1992; Banecki et al., 1992; Wawrzynow and Zylicz, 1995; Banecki and Zylicz, 1996). It was suggested that DnaK in its DnaK-ATP form binds to the DnaJ-modified substrate and after ATP hydrolysis, accelerated by DnaJ, a DnaJ-substrate-DnaK-ADP complex is formed (Szabo et al., 1994; McCarty et al., 1995; Gamer et al., 1996; Buchberger et al., this volume). Recent real time kinetics data have suggested a different scenario (Banecki and Zylicz, 1996; Banecki et al., 1996). Using stopped-flow analysis of the fluorescence of a single tryptophan located near the ATP binding site of DnaK, we showed that, upon ATP hydrolysis, DnaK is converted into a so-called DnaK*-ADP conformation, which can interact only transiently with protein substrate. Such a conformation cannot be obtained by simple incubation of DnaK in the presence of ADP. The presence of DnaJ induces other conformational changes of DnaK*-ADP, leading to the formation of a stable substrate-DnaK-ADP complex (Figure 2) (Wawrzynow et al., 1995b; Banecki and Zylicz, 1996). The formation of such a complex is a rate limiting step and depends on the affinity of DnaJ to the particular protein substrate. If the affinity of DnaJ to protein substrate is high a stable DnaJ-(substrate-DnaK-ADP) complex is formed (Wawrzynow et al., 1995b). In summary, DnaJ protein first “targets” DnaK to the preprimosomal structure and subsequently changes DnaK’s conformation to stabilize the P-DnaK-ADP complex. It is important to stress that, according to our experimental results, DnaJ-dependent conformational changes in DnaK occur after ATP hydrolysis when DnaK is already in the DnaK*-ADP form (Banecki and Zylicz, 1996; Banecki et al., 1996; Buchberger et al., this volume, for alternative views). By way of kinetic analysis, it has been shown that


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conformational changes of monomeric DnaK, the form required for DnaJ binding, correlates well with ATP hydrolysis (Banecki and Zylicz, 1996; Banecki, unpublished results) but not with ATP binding as was suggested by Theyssen et al. (1996) and Pierpaoli et al. (1997). Discrepancies in the interpretation of the results could have derived from procedures used by other laboratories, which inadvertently stripped bound nucleotites from DnaK protein while the same time inducing its oligomerization (Banecki, unpublished results). The presence of GrpE and ATP is required for recycling DnaK from the substrateDnaK-ADP complex (Zylicz et al., 1989, Alfano and McMacken, 1989b; Banecki and Zylicz, 1996; Wawrzynow and Zylicz, unpublished results). In the presence of ATP, GrpE accelerates the exchange of ADP for ATP complexed with DnaK, thus triggering the next round of ATP hydrolysis catalysed by DnaK (Figure 2) (Liberek et al., 1991; Banecki and Zylicz, 1996). The ATP hydrolysis changes the conformation of DnaK back to the DnaK*-ADP form which possesses a low affinity for protein substrate (Banecki and Zylicz, 1996). Therefore DnaK dissociates from the preprimosomal complex. It had been postulated previously that binding of ATP, but not ATP hydrolysis, was sufficient for the release of DnaK from protein substrate (Palleros et al., 1993; Schmid et al., 1994; Szabo et al., 1994; McCarty et al., 1995). We suggest that GrpE-dependent exchange of ADP to ATP complexed with DnaK could trigger the release of protein substrate from substrate-DnaK complex but could not recycle the DnaK. Only ATP hydrolysis converts DnaK to DnaK*-ADP conformation which can rebind the DnaJ modified substrate (Figure 2) (Banecki and Zylicz, 1996; Buchberger et al., this volume). Induction of conformational changes of DnaK triggered by ATP hydrolysis may also induce conformational changes in P, resulting in dissociation of P from the preprimosomal complex. The net result of this reaction cycle is the selective release of DnaK and P from the preprimosomal complex (Figure 2) (Liberek et al., 1988; Alfano and McMacken, 1989b; Dodson et al., 1989; Zylicz et al., 1989; Hoffman et al., 1992; Zylicz, 1993). In contrast, most of the O present in the O-some structure does not dissociate from the DNA template during the DnaK/DnaJ/GrpE chaperone dependent activation of the preprimosome (Dodson et al., 1986, 1989; Liberek et al., 1988; Wojtkowiak et al., 1993). The release of P from the DnaB complex is the crucial event leading to the initiation of DNA replication. A mutant of P, called P or , binds so weakly to the DnaB helicase that the requirement of molecular chaperones is significantly reduced (Konieczny and Marszalek, 1995). This observation explains the previous classical genetic experiments showing that DNA replication could proceed in the dnaK, dnaJ and grpE mutant backgrounds, provided that the P gene was mutated to P (Georgopoulos and Herskowitz, 1971; Saito and Uchida, 1977). Konieczny and Marszalek proposed that the phenotype of the P mutation could shed a new light on the evolution of bacteriophage DNA replication and the role of heat shock proteins in this process. According to this hypothesis the ancestor of modern bacteriophage possessed a P protein which resembles P such that heat shock proteins were not required for activation of the preprimosomal complex. The selective pressure for fast growth and development of A, phage produced modern P protein, which was more efficient in sequestering DnaB helicase. The side effect of this adaptation is the stability


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of preprimosome complex and the need of heat shock proteins for activation of the preprimosomal complex (Konieczny and Marszalek, 1995). The release of P from the complex with DnaB helicase leads to the unwinding of DNA near the ori sequence (Figure 2) (Dodson et al., 1986, 1989; Learn et al., 1993). Efficient unwinding of the supercoiled DNA template requires the additional presence of the Ssb protein. Extensive unwinding by DnaB creates positively supercoiled parental DNA, and DNA gyrase action is needed to counteract this effect (Learn et al., 1993). The DnaG primase recognizes the proper DnaB-ssDNA complex and proceeds to synthesize RNA primers at apparently random sites. The DNA polymerase III holoenzyme extends the RNA primers into the DNA. 2.4. Uni- and Bidirectional λ DNA Replication As discussed above, the preprimosomal complex is probably asymmetric, with the Osome structure located within the iteron sequences and with one or more P-DnaB complexes bound to the A/T-rich region located on the right side of theori sequence (Figure 2). Consequently the activated DnaB hexamer can freely unwind DNA duplex rightwards from theori sequence. However, the assembled O-some structure could cause a physical barrier for the passage of DnaB helicase through the O-some complex (Dodson et al., 1986). Supporting this hypothesis, in the in vitro DNA replication system reconstituted with purified proteins, replication is mostly unidirectional with the replication fork moving rightwards from the ori sequence (Figure 2) (Learn et al., 1993). However, addition of purified RNA polymerase together with rNTPs, switches a percentage (up to 30%) of the unidirectionally replicating molecules to a bidirectional DNA replication mode (Learn et al., 1993). McMacken and colleagues postulated that the role of transcriptional activation could be to displace or rearrange the O-some structure, thus allowing the DnaB helicase to unwind DNA to the leftward (Learn et al., 1993). We propose that the presence of the ClpXP protease (or the ClpX chaperone alone) may support RNA polymerase action leading to a more efficient transition from uni- to bidirectional DNA replication (Wojtkowiak et al., 1993). For example, in the presence of ClpXP the rearrangement of O protein into another O-some structure will almost certainly be inhibited, since O will be efficiently degraded upon transcription-mediated disruption of the O-some (Figure 2). Another factor which can influence the directionality of DNA replication is the efficiency of activation of the preprimosomal complex. For example, the release of P from the preprimosomal complex may not be complete (Dodson et al., 1989; Zylicz, 1993). About 50 to 60% of P indeed remain bound to DNA after action of the DnaK/DnaJ/GrpE chaperones. It is unclear whether this is a factor that limits the efficiency of DNA replication. Taylor and Wegrzyn suggest that DnaK/DnaJ/ GrpE chaperone machine only rearranges the preprimosomal complex but does not partially disassemble it (Taylor and Wegrzyn, 1995). According to this hypothesis the DnaB helicase which unwinds DNA may still be complexed with O, P and probably other replication proteins, forming a mobile replication complex which could be inherited, and used by one of two daughter DNA molecules during the second round of DNA replication (Wegrzyn and Taylor, 1992; Wegrzyn and Taylor, unpublished results). This


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hypothesis is based on the fact that in vivo a small fraction of O is not subjected to rapid degradation. The authors of these experiments assume that the stable fraction represents the O present in the replication complex (Wegrzyn et al., 1992). Interestingly, such a stable O fraction disappears after the heat shock induction of the groE operon coding for the GroEL/ES chaperone machine (Wegrzyn et al., 1996a). It is unclear how this chaperone machine affects the halflife of O. We postulate a different interpretation, according to which only the hexamers of DnaB helicase which have dissociated from P are involved in the unwinding of DNA. Release of P from only one of the helicase complexes (specifically distant from the Osome) might result in unidirectional DNA replication (Figure 2). Consequently the DnaB-λP complex proximal to the O-some structure might remain intact on the DNA template and may be inherited by one of two daughter strands. Another possibility is that the inherited replication complex represents a fraction of preprimosomal complexes which has become “frozen” at the ori O- P-DnaB stage. Supporting this interpretation, in the presence of transcription, the O protein bound in the ori O P-DnaB complex is resistant to ClpXP protease (Wawrzynow and Zylicz, unpublished results). 3. SWITCH FROM EARLY TO LATE

DNA REPLICATION

McMacken and colleagues have proposed that unidirectional DNA replication, after one round, could lead to strand separation, resulting in a switch from the early ( ) to the late ( ) mode of DNA replication (Figure 1) (Dodson et al., 1986; Learn et al., 1993). According to this hypothesis, the switch from the to the mode of A DNA replication could be correlated with the inhibition of early transcription (Learn et al., 1993). We propose that the ClpX molecular chaperone could also modulate this reaction. For example, the transient inhibition of ClpX activity could protect the intact O-some structure, thus favouring unidirectional DNA replication, and as a consequence initiates the switch to the rolling circle replication mode. The hypothesis that modulation of O degradation could regulate DNA replication is supported by the fact that the product of the rexB gene, the expression of which is increasing after initiation of DNA replication, prevents the O degradation (Schoulaker-Schwarz et al., 1991). The possibility that rex is involved in the switching from early to late phage DNA replication was suggested by Toothman and Herskowitz (1980). Under stress conditions, like heat shock, involvement of ClpX in other metabolic processes (e.g. proteolysis of host proteins) could also favour unidirectional DNA replication and, consequently, the switch to rolling circle replication. Similarly, mutation in the clpX gene might suppress the early bidirectional mode of DNA replication, therefore after the first round of unidirectional theta replication the switch to rolling circle mode might occur. This hypothesis could explain why mutation in clpX gene only slightly influences plaque formation (Szalewska et al., 1994). Recent findings suggesting that the elimination of dnaA function decreases transcriptional activation of DNA replication (Wegrzyn et al., 1995b) have led to the hypothesis that DnaA replication protein, a key initiation factor required for host DNA


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replication is also involved in DNA replication. In wild-type cells the utilization of DnaA by rapidly replicating phage DNA may inhibit transcriptional activation and thus favour the switch from bidirectional theta to the unidirectional theta replication mode and later to the sigma mode (Wegrzyn et al., 1995c). 4. CONCLUDING REMARKS Molecular chaperones are involved in many biochemical pathways, thus it is not surprising that mutations in these genes have pleiotropic metabolic effects that often compromise DNA replication of the bacterial chromosome and other replicons which are propagated in this bacterium. Detailed biochemical analysis, using in vitro DNA replication systems with purified proteins, indicates that molecular chaperones could be involved in DNA replication at least at three different stages. First, molecular chaperones might function during folding of newly synthesized replication proteins, protecting them from aggregation. Several replication proteins like O, DnaA, protein subunits of DNA polymerase III holoenzyme or RNA polymerase, possess strong tendencies to aggregate. The presence of molecular chaperones like DnaK, DnaJ, ClpX, ClpA, or GroEL could prevent formation of the inactive aggregates and stabilizes the active form of replication proteins (Skowyra et al., 1990; Hwang et al., 1990; Ziemienowicz et al., 1993, 1995; Blaszczak et al., 1995). Another example of this general chaperone mechanism is the ability of molecular chaperones to activate mutant replication proteins (Fayet et al., 1986; Hupp and Kaguni, 1993). It is important to note that several molecular chaperones could substitute for each other in protection reactions. This is probably one of the reasons why in many cases mutation in one of the genes that encode a molecular chaperone does not result in a complete block of the DNA replication process. The second stage at which molecular chaperones could work is the activation of already synthesized replication proteins. Both in vivo and in vitro initiation of plasmid P1 DNA replication is dependent on the functional presence of the DnaK/ DnaJ/GrpE molecular chaperones (Bukau and Walker, 1989; Tilly andYarmolinsky, 1989; Wickner et al., 1992). It was proposed that these chaperones dissociate RepA dimers in an ATPdependent reaction, thus producing RepA monomers that are active to bind to oriP1 sequence (Wickner et al., 1991). Results by DasGupta et al. suggest that monomerization of RepA may not be the only mechanism responsible for activation of RepA protein since dilute solutions of RepA, where the protein is essentially in monomeric form, still require chaperone function for activation (DasGupta et al., 1993). In the case of P1 DNA replication, bacterial molecular chaperones act before nucleoprotein complex formation (RepA-P1 DNA), and the presence of molecular chaperones is not obligatory at any of the subsequent steps leading to the initiation of P1 DNA replication (Wickner et al., 1992). The chaperone-dependent monomerization step required for the activation of initiation proteins for binding to the origin sequences was postulated also for F and RK2 plasmid DNA replication (Kawasaki et al., 1990; Konieczny and Helinski, 1997). In the latter case the ClpX chaperone can dissociate dimers of the TrfA initiation protein in an ATP-dependent reaction thereby allowing efficient TrfA binding to the oriRK2 sequence (Konieczny and Helinski, 1997).


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The third stage at which molecular chaperones could act is the specific initiation process. Initiation of DNA replication is not the only example of such a reaction. Recently it has been shown that ClpX is required for the lytic growth of bacteriophage Mu. The kinetics of the block in bacteriophage growth after induction of a clpX-deleted lysogen suggest that ClpX functions during the initiation of Mu DNA replication (Mhammedi-Alaoui et al., 1994). Bacteriophage Mu employs a transposition mechanism to replicate its DNA that involves the activity of the MuA transposase (Nakai and Kruklitis, 1995). After MuA-dependent catalysis of the strand transfer reaction (ends of the Mu DNA are cleaved and joined to the target DNA) a stable MuA-DNA complex is formed. Initiation of Mu DNA replication is triggered by release of MuA from such complex (Nakai and Kruklitis, 1995). It was shown in vitro that ClpX is required for the ATP-dependent release of MuA from such stable DNA replication intermediate complex (Levchenko et al., 1995; Kruklitis et al., 1996). The molecular mechanism of bacteriophage Mu DNA replication requires the high specificity of chaperone action. ClpX has to interact selectively with the protein substrate (MuA) bound within the replication complex and differentiate it from the other proteins present in this nucleoprotein structure. The analogy with the activation reaction of bacteriophage preprimosomal complex is obvious. The established functions of chaperones in and Mu DNA replication may be unique adaptations of parasites, allowing them to use the host’s DNA replication machinery. The involvement of molecular chaperones in the folding and activation of replication proteins before initiation of DNA replication, however, may represent a more general mechanism of the action of molecular chaperones in DNA replication. 5. ACKNOWLEDGMENTS We are grateful to R.McMacken, K.Taylor, G.Wegrzyn, D.Helinski, D.Missiakas and S.Raina for sharing with us unpublished results. We thank R.McMacken for discussions, critical comments and reading of this manuscript. We thank B.Ashraf for correcting this manuscript. This work was supported by grant 6PO4AO1712 from the Polish State Committee for Scientific Research. 6. REFERENCES Alfano, C. and McMacken, R. (1989a). Ordered assembly of nucleoprotein structures at the bacteriophage replication origin during the initiation of DNA replication. J. Biol. Chem. , 264 , 10699–10708. Alfano, C. and McMacken, R. (1989b). Heat shock protein-mediated disassembly of nucleoprotein structures is required for initiation of bacteriophage DNA replication. J. Biol. Chem. , 264 , 10709–10718. Alfano, C. and McMacken, R. (1988). The role of template superhelicity in the initiation of bacteriophage DNA replication. Nucl. Acids Res. , 16 , 9611–9630. Banecki, B. and Zylicz, M. (1996). Real time kinetics of the DnaK/DnaJ/GrpE molecular chaperone machine action. J. Biol. Chem. , 271 , 6137–6143.


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15. CONTROL OF HORMONE RECEPTOR FUNCTION BY MOLECULAR CHAPERONES AND FOLDING CATALYSTS DAVID O.TOFT Department of Biochemistry and Molecular Biology, Mayo Clinic, Rochester, MN 55905, USA

1. Introduction 2. The Identification of Receptor-associated Proteins 3. The Assembly of Steroid Receptor Complexes 4. Hsp90 and its Associated Proteins Act on a Wide Variety of Cellular Regulatory Proteins 5. References 1. INTRODUCTION The steroid hormone receptors belong to a large family of transcriptional regulators which also includes receptors for vitamin D, retinoic acid, thyroid hormone and a growing number of orphan receptors for which regulatory ligands have not been identified (Mangelsdorf et al., 1995; Mangelsdorf and Evans 1995; Beato et al., 1995; Kastner et al., 1995). These are modular proteins that contain separable domains for binding a regulatory ligand, for binding DNA response elements, and for interaction with other transcription factors. While some members of this protein family can dimerize, bind DNA and perform some functions in the absence of ligand, the unoccupied steroid receptors remain sequestered in an inactive complex. The binding of steroid induces a conformational change in the receptor which results in its liberation from the complex. It can then dimerize, bind to DNA and fulfill the functions of an active receptor. While the concept of inactive and active structural states of steroid receptors was developed several years ago (Gorski et al., 1968; Jensen et al., 1968), the realization that these states relate directly to the activities of molecular chaperones is more recent (Sanchez et al., 1985; Schuh et al., 1985; Catelli et al., 1985). In the simplest terms, the inactive receptor interacts with Hsp70, Hsp90 and some additional proteins that might be classified as cochaperones. These interactions persist until hormone binding occurs and they are lost when the receptor assumes its active state. The study of this system has several

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advantages for the characterization of chaperone functions. It illustrates the cooperative activities of several chaperones, some which were first identified through studies on steroid receptors. Also, one can turn on or off the chaperoning event by the absence or presence of hormone. In addition, the hormone binding activity of the receptor is dependent on chaperones and provides an easily measured end-point for certain chaperone assisted events. The following pages provide a description of the roles of chaperones in this system and of the approaches that are being used to understand functional elements of the system. The likelihood that these studies relate to much more general roles of these chaperones in cell biology is also discussed. 2. THE IDENTIFICATION OF RECEPTOR-ASSOCIATED PROTEINS Studies performed 10 to 15 years ago indicated that the receptors for progesterone (PR) and glucocorticoids (GR) existed in large complexes containing several proteins. The additional proteins were initially thought by some investigators to be contaminants and this made the identification of the receptor proteins confusing. It is interesting that one initial report on the isolation of progesterone receptor had confused it with Hsp90 (Dougherty et al., 1982) and another report had it confused with Grp94 (Kulomaa et al., 1986). These mistakes were soon resolved and several years of study have now revealed nine proteins that interact with steroid receptors in their inactive states (see for review, Pratt and Toft, 1997; Smith, 1995). These are listed in Table 1. These have somewhat arbitrarily been classified as chaperones or co-chaperones. This functional relationship is now quite clear for Hsp70 and DnaJ homologs, but most of the other co-chaperones in Table 1 have been recognized because they bind to Hsp70 or Hsp90 and their functions are still unclear. Thus, the assembly of steroid receptor complexes provides a focus on the chaperoning functions of Hsp70 and Hsp90 with the participation of several proteins that may modulate or extend the functions of these two chaperones. Hip (Hsp70 interacting protein) is a protein that was isolated because it interacts with the ATPase domain of Hsp70 (Höhfeld et al., 1995). This interaction appears to be dependent on the presence of Hsp40 (DnaJ homolog) and ATP/ADP, and it has been proposed that Hip promotes the state for tight substrate binding (ADP state) of Hsp70 (see Buchberger et al., and Ha et al., this volume). Hip was Table 1 Receptor-associated proteins

Chaperone

Co-chaperones

Hsp70

Hsp40/DnaJ homolog Hip/p48 Hop/Sti 1 homolog

Hsp90

Hop/Sti 1 homolog Immunophilins: FKBP52 FKBP51


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CyP-40 p23

independently identified as a component of progesterone receptor complexes and was called p48 according to its migration on gel electrophoresis (Prapapanich et al., 1996). It is found in complexes with Hsp70 plus Hsp90 and Hop, and it appears mainly in intermediate receptor complexes (see below). Hop is a 60 kDa homolog of the yeast heat shock protein Sti 1 (Honoré et al., 1992; Nicolet and Craig, 1989) and it was recognized as an early component in receptor complex formation (Smith et al., 1993b; Smith 1993). Sti 1 is not an essential protein in yeast and, although it has been shown to bind to Hsp90 (Chang and Lindquist, 1994), its function is unknown. In higher eukaryotes, Hop can bind to both Hsp70 and Hsp90 to form a ternary complex (Smith et al., 1993b; Chen et al., 1996) and it has been suggested that Hop (hsp organizing protein) acts to organize the functions of Hsp70 and Hsp90. Gross and Hesseforth (1996) have isolated a protein identical to Hop which they call RFHsp70 (recycling factor for Hsp70). They show that this protein promotes the dissociation of ADP from Hsp70 and its replacement by ATP, but this activity has been questioned by Johnson et al., 1998). Another puzzling group of Hsp90-binding proteins are the high molecular weight immunophilins FKBP51, FKBP52 and CyP-40. These are proteins that have peptidylprolyl-cis/trans isomerase activity and the capacity to bind the immunosuppressant drugs FK506 (FKBP52, FKBP51) or cyclosporin A (CyP-40) (see Fischer and Schmid, this volume). FKBP51 has also been called FKBP54 (Smith et al., 1993a) and FKBP52 has had several names including p59, FKBP59, Hsp56 and HBI (Hsp90-binding immunophilin). These three proteins are found in receptor complexes and they are also bound to Hsp90 in the absence of receptor (Radanyi et al., 1994; Czar et al., 1994; Hoffmann and Handschumacher, 1995). They do not bind together but appear to compete for a common site on Hsp90 (Owens-Grillo et al., 1995). All three proteins contain TPR domains or 34 residue repeating units that are required for Hsp90 binding (Radanyi et al., 1994; Owens-Grillo et al., 1996; Ratajczak and Carello, 1996). TPR domains have been observed in a wide variety of proteins including Hip and Hop and it has been proposed that these domains participate in protein-protein interactions (Lamb et al., 1995). Immunophilins have been implicated as chaperones in protein folding, in the control of cell signalling pathways and in targeting proteins to cellular locations, but their actual functions in receptor complexes are completely unknown. Like the immunophilins, p23 was identified from its binding to receptor complexes and it also binds to Hsp90 in the absence of receptor (Johnson et al., 1994; Johnson and Toft, 1994). p23 is not structurally related to immunophilins or to known heat shock proteins. Its binding to Hsp90 is dependent on the presence of ATP and it thus may be a modulator of Hsp90 activities. However, its precise function is unknown. Most of the p23 in cell extracts is in complex with Hsp90 plus FKBP51, FKBP52 and CyP-40, and this may represent a chaperoning complex needed at the later stages of receptor complex assembly (Johnson and Toft, 1995). The proteins in Table 1 were revealed by their co-purification during isolation of


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inactive steroid receptor complexes. The co-chaperones were then shown to copurify with Hsp70 or Hsp90 as relatively prominent complexes from cell extracts. For example, the abundance of p23 is only 10–20% of the Hsp90 level in cytosol extracts and the majority of p23 is bound to Hsp90 (Johnson et al., 1994). It should be noted that the steroid receptors exist at less than 1% of the Hsp90 level and most known Hsp90 substrates or targets are proteins of low abundance that would only occupy a small fraction of cellular Hsp90. DnaJ homologs, which will be termed collectively as Hsp40, are the only proteins in Table 1 that haven’t been clearly demonstrated in steroid receptor complexes. However, they have been implicated by genetic manipulation of yeast model systems. Mutation of the yeast Ydj-1 protein has been shown to dramatically alter the functional capacity of androgen (Caplan et al., 1995) and glucocorticoid receptors (Kimura et al., 1995). Yeast have also been used to demonstrate the requirement for Hsp90 in the functioning of steroid receptors (Picard et al., 1990; Bohen and Yamamoto, 1993; Kimura et al., 1994; Bohen, 1995a; Nathan and Lindquist, 1995). It will be important to test the significance of the other receptor-associated proteins by genetic manipulation of yeast or other cellular systems. 3. THE ASSEMBLY OF STEROID RECEPTOR COMPLEXES Early attempts to study the interaction of steroid receptors with their associated proteins were frustrated by the instability of these associations in vitro and the observations that receptor complexes don’t form easily by a simple combination of components. This led Pratt and co-workers to suggest that the complexes were formed during receptor synthesis, requiring interactions with nascent polypeptide chains. They and others were able to show that glucocorticoid receptor synthesized in rabbit reticulocyte lysate was in a native multiprotein complex, able to bind hormone (Dalmon et al., 1989; Denis and Gustafsson, 1989). However, subsequent studies showed that translation was not essential, but factors in the reticulocyte lysate provided optimal conditions for complex formation. Thus, native receptor isolated from target tissue could be stripped of its associated proteins by high salt treatment and then reconstituted into a complex by incubation in rabbit reticulocyte lysate (Smith et al., 1990). It has since been shown that other cellular extracts are also effective in this regard, but they must be quite concentrated and able to maintain an effective level of ATP (Stancato et al., 1996a). The formation of receptor complexes required incubation at elevated temperatures, magnesium, potassium, and the hydrolysis of ATP (Smith et al., 1992; Hutchison et al., 1992a). The microsomal fraction, normally present in reticulocyte lysate, was not required. Much new information has been acquired through use of the in vitro system which has led to models of the receptor assembly process. One such model, illustrated in Figure 1, is quite similar to a recent model by Prapapanich et al., (1996). Many liberties have been taken in this model since we are still very early in our understanding of this process. The assembly of the receptor complex is shown to progress through three stages. The first stage, complex 1, represents an initial interaction of Hsp70 with hydrophobic regions in the ligand binding domain of the receptor. This stage has not been described


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experimentally and is simply based on the demonstrated ability of Hsp70 to bind hydrophobic domains in other systems and the fact that Hsp70 is an essential component in the assembly of receptor complexes (Hutchison et al., 1994). Hsp40 is introduced here because it is known as an Hsp70 co-chaperone and it has been implicated by genetic and biochemical studies (Caplan et al., 1995; Kimura et al., 1995; Dittmar et al., 1998). An involvement of Hip at this step is also suggested because of its known interaction with Hsp70. In addition, studies on the time course of complex formation with the PR have shown extensive binding of Hsp70, Hip and Hop at early times and a loss of these components at later times (Smith, 1993). Hop and Hsp90 are incorporated at the second stage of assembly to make an intermediate state. This complex has been observed under three conditions: at early times in the assembly of PR complexes (Smith, 1993), in the absence of an ATP regeneration system when the level of ATP is suboptimal (Smith et al., 1992) and when assembly is inhibited by geldanamycin, a reagent that blocks some functions of Hsp90 (Whitesell et al., 1994; Johnson and Toft, 1995; Smith et al., 1995). Thus, there is good evidence for the existence of this intermediate form. Toward the right in Figure 1, a ternary complex with Hsp90, Hop, and Hsp70 is indicated in brackets. This has been observed in cell extracts and it has been proposed that this is a precursor for receptor assembly (Smith et al., 1993b; Dittmar et al., 1997). Thus, it is possible that the receptor binds pre-formed complexes of these proteins instead of the individual proteins as shown in the model. While not indicated in the model, at stage 2, Hsp90 is underrepresented in comparison to stage 3. In other words, there is more Hsp70 than Hsp90 at stage 2, but little Hsp70 and about twice as much Hsp90 at stage 3 (Smith, 1993; Johnson and Toft, 1995; Smith et al., 1995). The conversion from stage 2 to 3 is the geldanamycin-sensitive step where more Hsp90 plus p23 and the immunophilins are incorporated into the complex. This stage utilizes additional ATP which might be required for a small sustained action of Hsp70 and/or an ATP-dependent activity of Hsp90. Whether or not Hsp90 interacts with ATP in a functional sense has been a controversial issue (Csermely et al., 1993; Jacob et al., 1996) however, recent structural and biochemical studies clearly reveal an ATP binding site near the N-terminus of hsp90 (Stebbin et al., 1997; Prodromou et al., 1997; Grenert et al., 1997). This site also binds the hsp90 inhibitor, geldanamycin. We have found that the binding of ATP to hsp90 promotes a conformational change that allows hsp90 to interact with p23 (Grenert et al., 1997). It is at this last step in Figure 1 when the receptor becomes competent to bind hormone since hormone binding activity, at least for some receptors, is blocked by the inhibitor geldanamycin (Smith et al., 1995). Thus, there is clearly a conformational change in the receptor at this stage. As mentioned earlier, Hsp90 can exist in a pre-formed complex with p23 and any one of the immunophilins FKBP51, FKBP52 and CyP-40 and it may be this complex rather than the individual proteins that interacts with the receptor. The formation of this complex of Hsp90 with p23 and immunophilin also requires ATP (for p23 binding, but not for immunophilin binding) and it is this process that is inhibited by geldanamycin (Johnson and Toft, 1995). Several experiments have indicated the dynamic nature of this system. When individual proteins are labelled with radioisotopes or antibody epitopes, these are rapidly exchangeable with unlabelled components within a few minutes at 30° (Smith, 1993;


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Smith et al., 1995). Thus, it is likely that the complexes are continu-

Figure 1 A model for the assembly of steroid receptor complexes. The receptor (R) is divided into three domains: an amino terminal trans-activation domain, an internal DNA-binding domain, and a carboxy-terminal ligand binding domain interacting initially with Hsp70. In some cases, the proteins are abbreviated by use of a number, eg. 60=Hop/p60. At the right of the figure, two multiprotein complexes are shown which may participate in receptor complex formation. The shaded triangle in the receptor at step 3 indicates the formation of the ligand binding site.

ously dissociating and re-assembling and that the process would be better represented as a cycle rather than a linear pathway. This model brings to mind several important questions that require further study. The largest gap in knowledge concerns function at all levels. Why are so many proteins needed in this process? What are the functions of the individual proteins and how are these functions coordinated among proteins? What are the roles of ATP/ADP and are there other factors or proteins of this process that have not yet been recognized? Efforts to answer these questions have only begun and there are, as yet, few answers. Yeast genetics has been employed to test the significance of some proteins. This is possible because several steroid receptors can be expressed in yeast where they respond


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to hormone to induce expression of a reporter gene (Picard, 1997; Caplan, 1997). Several mutants of Hsp90 have been tested in this system. When Hsp90 is mutated or reduced to low levels, hormone responsiveness is lowered dramatically and this can be explained, in part, by a loss of hormone binding activity of the receptor (Picard et al., 1990; Bohen andYamamoto, 1993; Kimura et al., 1994; Bohen, 1995a; Nathans and Lindquist, 1995). Similar results have been observed with mutations of the DnaJ homologYdj-1 (Caplan et al., 1995; Kimura et al., 1995) indicating that both of these proteins are needed to process the receptor to a native, fully functional state. The yeast Hop homolog Sti 1, can be deleted since it is not essential for viability (Nicolett and Craig, 1989). However, studies on the activity of the glucocorticoid receptor in yeast show that the deletion of Sti1 reduces the activity of this receptor and the functions of Sti1 appear to be linked to those of hsp90 (Chang et al., 1997). Preliminary studies indicate that the yeast homolog of p23 can also be deleted without loss of yeast viability resulting in only subtle effects on glucocorticoid receptor function (Bohen, 1995b). These studies must be pursued further to study compensatory mechanisms and additive effects of various mutations. A second approach has been to study the in vitro reconstitution of receptor complexes with regard to intermediate states and the removal of individual proteins. The ability to reconstitute GR complexes in rabbit reticulocyte lysate is lost after depletion of Hsp70 from the lysate and is re-gained by adding back Hsp70 (Hutchison et al., 1994). Similar results have been found with the removal and replacement of p23 (Johnson and Toft, 1994). In an interesting study by Hutchison et al. (1995), GR was found to form complexes containing Hsp90 in wheat germ lysate, but the receptor lacked hormone binding activity unless exogenous p23 was added. This indicates the importance of p23 in the formation or stability of a native complex and suggests that p23 is lacking in wheat germ lysate or is not compatible with animal steroid receptor systems. The importance of Hop in in vitro receptor complex formation has been indicated by use of a Hop antibody which apparently neutralizes Hop function (Chen et al., 1996). In the presence of antibody, PR complexes accumulate in an intermediate complex similar to complex 2 in Figure 1 which lacks hormone binding activity. An antibody to Hsp70 has also been used to block its activity and the formation of PR complexes (Smith et al., 1992). There is very little information on the functions of FKBP52, FKBP51 or CyP-40. Treatment of reticulocyte lysate with FK506 or cyclosporin A does not alter the formation of steroid receptor complexes (Hutchison et al., 1993). These drugs block the peptidyl prolyl isomerase activity of the immunophilins, indicating that this activity is not essential to receptor complex formation. On the other hand, these immunosuppressant drugs have been shown to alter receptor responsiveness in cell culture systems (Ning and Sanchez, 1993; Tai et al., 1994; Milad et al., 1995; Renoir et al., 1995) and one report indicates an increased hormone binding affinity of PR after treatment with FK506 in vitro (Renoir et al., 1994). Greater efforts must be made to develop chemically defined systems for the formation and study of receptor complexes. This would tell us first of all whether all of the essential components have been identified. One could then add or subtract components, replace them with mutant forms, prepare partial or intermediate complexes, thereby allowing a


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much clearer description of this process and the study of functional contributions of various components. Along this line, Pratt and co-workers have recently succeeded in forming GR complexes using a combination of purified proteins including Hsp90, Hsp70, Hop, p23, and Hsp90 (Dittmar et al., 1996; Dittmar et al., 1998). Four of these proteins are essential for receptor complex formation and the fifth, p23, appears to stabilize the complex in a state that provides the receptor with hormone binding activity. Additional studies are needed on this minimal system to test the importance of Hip or immunophilins and the possible contribution of other factors. 4. HSP90 AND ITS ASSOCIATED PROTEINS ACT ON A WIDE VARIETY OF CELLULAR REGULATORY PROTEINS Hsp90 has been studied in relation to cell stress and it has recently been shown to prevent aggregation of denatured proteins in a manner that does not require ATP or additional protein components (Jacob et al., 1995; Freeman and Morimoto, 1996). However, most studies on Hsp90 have concerned its association with other cellular proteins in either persistent complexes, as with steroid receptors, or transient associations such as that with the tyrosine kinase pp60v-src. These studies have led to the idea that Hsp90 plays a specialized role in chaperoning the folding, processing or trafficking of a unique group of cellular proteins. There are now a wide variety of proteins that have been indicated to be substrates for Hsp90. Most of these are either regulators of transcription or protein kinases (see chapter by Bensaude, this volume). Table 2 lists a number of proteins in these two groups. Some of these have only been linked to Hsp90 through genetics studies, such as Wee 1 and sevenless tyrosine kinases, but most have been recognized in complexes containing Hsp90. Other components in these complexes have not been described in most cases. The complexes for dioxane receptor and for heme-regulated eIF-2a kinase have been described and they show close similarity to steroid receptor complexes (Chen and Perdew, 1994; Nair et al., 1996; Matts et al., 1992). They also contain Hsp70, FKBP52, and p23. These proteins have also been shown in complexes with fes tyrosine kinase and hsf-1 (Nair et al., 1996). In addition, complexes with pp60v-src, fes and Raf-1 contain a phosphoprotein, p50, that is not present in steroid receptor complexes (Brugge et al., 1981; Oppermann et al., 1981; Nair et al., 1996; Stancato et al., 1993; Wartmann and Davis, 1984). p50 also binds to Hsp90 in the absence of the kinase (Whitelaw et al., 1991). A recent study by Stepanova et al., (1996) has revealed the identity of p50 to be Cdc 37. This protein is essential in yeast. It is genetically linked to multiple protein kinases and it is required for the production of Cdc 28/cyclin complexes. Cdc 37 appears to target Hsp90 to the Cdk 4 kinase and the authors suggest that it has a general role in the assisted folding of intrinsically unstable protein kinases. The studies on Hsp90-kinase complexes suggest that some variations exist in the composition of Hsp90 complexes and it will be important to study more examples for comparison. The list of proteins in Table 2 suggests that the functions of Hsp90 might be of special importance to signalling pathways and regulation. However, it is also possible that Table 2 reflects the research bias of investigators. There may be a number of


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Table 2 Hsp90-associated proteins

Associated Proteins

References

Transcription Factors Glucocorticoid receptor

Sanchez et al., 1985

Progesterone receptor

Schuh et al., 1985; Catelli et al., 1985

Estrogen receptor

Joab et al., 1984; Redeuilh et al., 1987

Androgen receptor

Joab et al., 1984; Veldscholte et al., 1992

Mineralocorticoid receptor

Rafestin-Oblin et al., 1989

v-erb A

Privalsky, 1991

Ah receptor

Perdew, 1988; Denis et al., 1988

Myo D1

Shue and Kohtz, 1994

Heat shock factor

Nadeau et al., 1993; Nair et al., 1996

CN-tumor promoter-specific binding protein

Hashimoto and Shudo, 1991

Sim

McGuire et al., 1996

E12

Shue and Kohtz, 1992

p53 tumor suppressor mutant

Sepehrnia et al., 1996; Blagosklonay et al., 1996

Hepatitis B virus reverse transcriptase Hu and Seeger, 1996 Protein Kinases v-src, c-src

Brugge et al., 1981; Oppermann et al., 1981; Hutchison et al., 1992b

v-fps

Lipsich et al., 1982

v-yes

Lipsich et al., 1982

v-fes

Ziemiecki et al., 1986; Nair et al., 1996

v-frg, c-frg

Ziemiecki et al., 1986; Hartson and Matts, 1994

lck

Hartson and Matts, 1994

Weel

Alique et al., 1994

Sevenless

Cutforth and Rubin, 1994

Heme-regulated eIF-2a kinase

Rose et al., 1987; Matts and Hurst, 1989;

eEF-2 kinase

Palmquist et al., 1994

Casein kinase II

Miyata and Yahara, 1992; Shi et al., 1994

v-Raf, c-Raf, Gag-Mil

Stancato et al., 1993; Wartmann and Davis, 1994; Lovric et al., 1994

Control of hormone receptor function by molecular chaperones MAP kinase kinase (Mek)

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Stancato et al., 1997

proteins outside transcription factors and protein kinases that interact with Hsp90. A few known examples are actin (Koyasu et al., 1986), tubulin (Sanchez et al., 1988), proteasomes (Tsubuki et al., 1994; Wagner and Margolis, 1995), G protein -complexes (Inanobe et al., 1994) and calmodulin (Minami et al., 1993). These may also be regulatory interactions or they may simply be transient chaperone-assisted folding processes of a more general purpose. The tools now available should expand the scope of studies in this area. The drug geldanamycin has already provided much new information on processes involving Hsp90 since it can be used to inhibit Hsp90 in intact cells. Thus, geldanamycin has been shown to block the processing and activity of pp60v-src (Whitesell et al., 1994), Raf-1 (Schulte et al., 1995), mutant p53 (Blagosklonny et al., 1995), glucocorticoid receptor (Whitesell and Cook, 1996), progesterone receptor (Smith et al., 1995) and the reverse transcriptase of hepatitis B virus (Hu and Seeger, 1996). Further characterization of these diverse processes should provide a much better understanding of the components and the functional responsibilities of the chaperone pathway involving Hsp70 and Hsp90. 5. REFERENCES Aligue, R., Akhavan-Niak, H. and Russell, P. (1994). A role for Hsp90 in cell cycle control: Weel tyrosine kinase activity requires interaction with Hsp90. EMBO J. , 13 , 6099–6106. Beato, M., Herrlich, P. and Schütz, G. (1995). Steroid hormone receptors: Many actors in search of a plot. Cell , 83 , 851–857. Blagosklonny, M.V., Toretsky, J. and Neckers, L. (1995). Geldanamycin selectively destabilizes and conformationally alters mutated p53. Oncogene , 11 , 933–939. Blagosklonny, M.V., Toretsky, J., Bohen, S. and Neckers, L. (1996). Mutant conformation of p53 translated in vitro or in vivo requires functional Hsp90. Proc. Natl. Acad. Sci., USA , 93 , 8374–8383. Bohen, S.P. (1995a). Hsp90 mutants disrupt glucocorticoid receptor ligand binding and destabilize aporeceptor complexes. J. Biol. Chem. , 270 , 29433–29438. Bohen, S.P. (1995b). Pharmacological and genetic analysis of the roles of Hsp90 and p23 in signaling protein function. Mol. Biol. Cell , 6 , 6a. Bohen, S.P. and Yamamoto, K.R. (1993). Isolation of Hsp90 mutants by screening for decreased steroid receptor function. Proc. Natl. Acad. Sci. USA , 90 , 11424–11428. Brugge, J.S., Erikson, E. and Erikson, R.L. (1981). The specific interaction of the Rous sarcoma virus transforming protein, pp60src, with two cellular proteins. Cell , 25 , 363– 372. Caplan, A.J., Langley, E., Wilson, E.M. and Vidal, J. (1995). Hormone-dependent transactivation by the human androgen receptor is regulated by a dnaJ protein. J. Biol. Chem. , 270 , 5251–5257. Caplan, A.J. (1997). Yeast molecular chaperones and the mechanism of steroid hormone action. Trends Endocrinol. Metab. , 8 , 271–276.


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16. ROLE OF CHAPERONES IN UNCOATING OF CLATHRIN COATED VESICLES EVAN EISENBERG and LOIS GREENE* Laboratory of Cell Biology, National Heart, Lung, and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892

1. Introduction 2. Overall Cycle of Uncoating by Hsc70 3. Role of Auxilin, a DnaJ Homolog, in Uncoating 3.1. Role of Clathrin Domains in Uncoating 3.2. Direct Interaction of Auxilin with Hsc70 3.3. Mechanism of Action of Auxilin 3.4. Contrast Between Substrates and Products of Hsc70 4. Model of Uncoating 5. Time Course of Uncoating 5.1. Initial Burst of Uncoating 5.2. Slow Steady-state Uncoating 6. Concluding Remarks 7. References 1. INTRODUCTION The Hsp70 class of molecular chaperones and the related constitutive proteins, the Hsc70s, have a widely diverse set of functions in the cell in addition to their role in the heat shock response (see reviews by Hendrick and Hartl, 1993; Rassow and Pfanner, 1995; Hartl, 1996). The Hsc70s are involved in both the folding of proteins after synthesis on ribosomes and in maintaining proteins in an unfolded state after synthesis (chapter Welch et al.). They are also involved in unfolding proteins prior to translocation through mitochondrial or endoplasmic reticulum membranes, in driving the translocation process itself, and then in refolding these proteins following their translocation through the membranes (chapters Haas and Zimmerman; Dekker and Pfanner). *Corresponding author: Building 3, Room B1–22, 3 Center Dr., MSC 0301.


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Finally they are involved in the dissociation of protein complexes such as RepA dimers in E. coli (chapter Burkholder and Gottesman) and clathrin coats in mammalian cells and in the formation of protein complexes such as the complex between Hsp90 and steroid receptors in mammalian cells (chapter Toft). Presumably there is a common thread in the mechanism of action of Hsc70 that allows it to carry out such diverse functions. As yet, however, the basic mechanism of action that underlies these varied functions of Hsc70 remains unclear. It has been suggested that the key property of Hsc70 is its ability to bind unfolded regions of proteins in an ATP dependent fashion. However, since attachment and detachment of substrate can occur in an equilibrium system without ATP hydrolysis as well as in a steady-state system driven by ATP hydrolysis, the role of ATP in attachmentdetachment cycles of the various substrates and how it affects the various functions of Hsc70 remains unclear. One of the first constitutive functions discovered for Hsc70 was its ability to uncoat clathrin-coated vesicles in vitro (Schlossman et al., 1984). Monomeric clathrin triskelions with a molecular mass of about 600 kDa, are composed of three identical heavy chains of 190 kDa and three tightly associated light chains (Pearse, 1987; Brodsky, 1988). The triskelion can be structurally and functionally divided into distal and proximal leg sections separated by a flexible hinge region and a globular terminal domain. During receptor-mediated endocytosis clathrin triskelions polymerize and form coats on pits in the plasma membrane that later invaginate into the cell (Keen, 1990; Pearse and Robinson, 1990). Similar clathrin-coated pits also form on the trans-Golgi membrane. In addition to clathrin and receptors, these pits contain assembly proteins or adaptins that serve two functions (Robinson, 1992). They catalyze the polymerization of clathrin triskelions into the planar hexagonal arrays that make up the coat and they bind the receptors that are localized in the clathrin-coated pit. Three major clathrin-assembly proteins have been described (Keen, 1990; Robinson, 1992). AP2 and AP1, which are multimeric subunit complexes of about 270 kDa, occur on the plasma membrane and trans-Golgi membranes, respectively. AP180 is a neuronal specific assembly protein that consists of only one subunit of 120 kDa (Ahle and Ungewickell, 1986; Prasad and Lippoldt, 1988). Following their formation, clathrin-coated pits invaginate, a process that involves the transformation of planar hexagonal clathrin arrays into spherical clathrin coats (Keen et al., 1979; Heuser, 1980; Heuser and Kirchhausen, 1985). This in turn requires the transformation of a number of hexagonal clathrin arrays into pentagonal arrays so that the resulting clathrin-coat has a soccer ball-like structure. As yet it is not clear if clathrin is required for the invagination of the membrane or whether the clathrin coat simply changes its structure as part of the invagination process. In any event, since clathrin coats form spontaneously in vitro (Keen et al., 1979), Rothman and his associates reasoned that removing clathrin from coated vesicles would be an ATP-dependent process and in searching for the enzyme involved, discovered the uncoating ATPase, an enzyme that uncoats clathrin-coated vesicles and artificial clathrin baskets in an ATP-dependent fashion in vitro (Schlossman et al., 1984; Braell et al., 1984). The uncoating ATPase was later found to be a constitutive member of the Hsp70 family (Ungewickell, 1985; Chappell et al., 1986).

Role of chaperones in uncoating of clathrin coated vesicles

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Although the Hsc70 class of proteins is involved in numerous functions in vivo, these functions are often difficult to study in purified systems in vitro. The uncoating reaction, on the other hand, is highly amenable to study in vitro and, since it is ATP dependent, deciphering its mechanism may lead to important information about the basic processes that underlie all of the ATP-dependent functions of the Hsc70 class of proteins. In addition, numerous questions remain about the role of Hsc70 in endocytosis. Although there is evidence that Hsc70 is involved in uncoating clathrin-coated vesicles in vivo (Honing et al., 1994), it is not clear why Hsc70 does not uncoat clathrin-coated pits (Heuser and Steer, 1989), or whether Hsc70 is involved in the formation and invagination of clathrin-coated pits. Therefore, investigation of the uncoating reaction may yield important information about the mechanism of endocytosis. 2. OVERALL CYCLE OF UNCOATING BY HSC70 The basic properties of the uncoating reaction have been studied using both clathrincoated vesicles and artificial clathrin baskets as a substrate for Hsc70; artificial clathrin baskets are prepared using purified clathrin and various types of assembly proteins. Two observations suggest that uncoating is a specific function of Hsc70. First, this process appears to be ATP-dependent (Schlossman et al., 1984; Schmid et al., 1985; Greene and Eisenberg, 1990), although there have been reports that ATP analogs such at ATPγS and AMP-PNP can also support uncoating (Heuser and Steer, 1989; Buxbaum and Woodman, 1995). Second, the uncoating reaction only occurs with certain Hsc70s, as would be expected for a specific reaction. DnaK shows a much reduced level of uncoating activity (Ungewickell et al. 1997), while there is negligible uncoating activity with BiP, the Hsc70 homolog present in the endoplasmic reticulum (unpublished data). The yeast cytoplasmic Hsc70s, SSA1 and SSA2, do carry out uncoating with the same time course as bovine brain Hsc70 but, particularly with SSA1, considerably more enzyme is required for the same amount of uncoating as occurs with bovine brain Hsc70 (Gao et al., 1991). Interestingly, a similar effect occurs with a truncated bovine brain Hsc70 from which the 10 kDa C-terminal domain has been deleted (Ungewickell et al., 1997). Although earlier studies suggested that this truncated Hsc70 cannot carry out uncoating (Chappell et al., 1987; Tsai and Wang, 1994), Ungewickell et al. (1997) recently demonstrated that it can carry out normal uncoating but only at relatively high concentration. The ATP dependence of the uncoating reaction suggests that Hsc70 is acting catalytically, using the energy of ATP hydrolysis to dissociate polymerized clathrin. This, in turn, suggests that a small amount of Hsc70 should rapidly uncoat a large amount of polymerized clathrin and, indeed, in their preliminary studies Rothman and his associates (Schlossman et al., 1984) reported such behavior. However, more detailed studies on the time course of uncoating of clathrin-coated vesicles showed that this time course is strikingly non-linear, consisting of a rapid initial burst of uncoating followed by very slow steady-state uncoating (Greene and Eisenberg, 1990). Furthermore, at pH 7 this initial burst of uncoating is nearly stoichiometric with three Hsc70s uncoating one clathrin triskelion or three clathrin heavy chains (Greene and Eisenberg, 1990). Later studies showed that the same time course and stoichiometry of uncoating occurs with clathrin baskets prepared with mixed assembly proteins, purified AP2, AP180 (open


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triangles, Figure 1), and even myelin basic protein (which acts as an assembly protein) showing that the biphasic time course of uncoating is a general feature of the uncoating reaction (Greene and Eisenberg, 1990; Barouch et al., 1994; Prasad et al., 1994; Prasad et al., 1995). Based on the time course and stoichiometry of uncoating, we proposed a simple model of the uncoating process (Greene and Eisenberg, 1990). In this model Hsc70-

Figure 1 Effect of varying amounts of auxilin on uncoating of clathrin baskets by Hsc70. The time course of clathrin heavy chain (HC) release from baskets consisting of 0.5 M clathrin and 0.5 M AP180, by 0.6 M Hsc70 was measured in the presence of varying concentrations of auxilin: 0 M (●), 0.02 M (□), 0.05 M (■), or 0.10 M (∆) auxilin. Baskets composed of recombinant truncated auxilin consisting of 0.5 M clathrin and 1.5 M recombinant auxilin (▲) were also used as a substrate in the uncoating reaction.

ATP first binds stoichiometrically to the clathrin baskets or vesicles. Then, in association with rapid ATP hydrolysis, a clathrin triskelion complexed with Hsc70-ADP-Pi dissociates from the baskets or vesicles. This accounts for the initial burst of uncoating. The Hsc70-ADP-Pi remains complexed with the clathrin triskelion until the ADP at the active site slowly dissociates, which accounts for the slow steady-state uncoating. Finally ATP rebinds to the Hsc70-clathrin complex, and the monomeric clathrin triskelion rapidly dissociates from the Hsc70 before the bound ATP is hydrolyzed. Since this model was proposed, further studies have confirmed several of its major points, and have shown that these points also apply to the interaction of Hsc70 with other substrates. First, it is now clear that Hsc70 has only one binding site for ATP (Gao et al., 1993). Second, the observation that an initial burst of ATP hydrolysis accompanies the


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initial burst of uncoating (Barouch et al., 1994) confirms that clathrin baskets and vesicles bind to the ATP form of the enzyme rather than the ADP form, and it is also clear that, in the absence of substrate or cofactors, most of the Hsc70 is in the ATP form (Gao et al., 1993). Similar observations have shown that other substrates of Hsc70 also bind to the ATP form of the enzyme (Prasad et al., 1994, Schmid et al., 1994; Greene et al., 1995; McCarty et al., 1995; Wei et al., 1995; Takeda and McKay, 1996). Third, following the initial burst of uncoating, a complex of Hsc70 and clathrin forms that is stable in ADP and Pi and can be isolated by column chromatography confirming that hydrolysis of ATP to ADP traps the dissociated clathrin triskelions on the enzyme in a complex that is slow to dissociate (Prasad et al., 1994). It has also been shown that other substrates dissociate very slowly from Hsc70-ADP (Palleros et al., 1993, Greene et al., 1995; McCarty et al., 1995; Takeda and McKay, 1996). Finally, it has been shown that both clathrin triskelions and other substrates dissociate very rapidly from Hsc70-ATP (Palleros et al., 1993; Prasad et al., 1994; Greene et al., 1995; McCarty et al., 1995; Takeda and McKay, 1996). Indeed, they dissociate from Hsc70-ATP much more rapidly than ATP hydrolysis occurs, confirming that it is ATP binding rather than ATP hydrolysis that causes dissociation of both clathrin triskelions and other Hsc70 substrates from Hsc70 (Palleros et al., 1993; Prasad et al., 1994; Schmid et al., 1994; Greene et al., 1995; McCarty et al., 1995). Although the simple model presented above explains many aspects of the uncoating reaction, it soon became clear that another factor must be involved in the uncoating reaction because clathrin baskets prepared from highly purified clathrin and assembly proteins cannot be uncoated by Hsc70 alone (Prasad et al., 1993; Barouch et al., 1994; Ungewickell et al., 1995). 3. ROLE OF AUXILIN, A DNAJ HOMOLOG, IN UNCOATING The first evidence that a cofactor is required for Hsc70 to uncoat clathrin-coated vesicles came from studies on the ATPase activity associated with uncoating; to avoid contaminant ATPases, these studies must be carried out with artificial clathrin baskets prepared with purified clathrin and purified assembly proteins (Barouch et al., 1994). When the assembly protein, AP2, was used to prepare AP2-clathrin baskets, it was found that these baskets could no longer be uncoated by Hsc70. Further studies revealed that fractions obtained during purification of AP2 contained a 100 kDa protein cofactor that was required for Hsc70 to carry out the uncoating reaction in ATP (Prasad et al., 1993). The 100 kDa cofactor was later identified as auxilin (Ungewickell et al., 1995), a minor neuronal specific assembly protein first isolated and then more recently cloned by Ungewickell and his associates (Ahle and Ungewickell, 1990; Schroder et al., 1995). The requirement for auxilin appears to be a general one, applying to all types of clathrin baskets, whether prepared with AP2, AP180 (Figure 1), or even myelin basic protein (Prasad et al., 1993; Barouch et al., 1994; Ungewickell et al., 1995; Prasad et al., 1995). In its ability to support uncoating by Hsc70, auxilin resembles members of the DnaJ family of proteins that have been shown to act as partner proteins with Hsc70 in a number of the activities carried out by Hsc70 including protein folding, protein translocation, and formation and dissociation of protein complexes (Cyr et al., 1994; Rassow et al., 1995).


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The DnaJ family of proteins is characterized by having a specific J domain usually at the N-terminal region of the protein (Caplan et al., 1993). When the sequence of auxilin was examined, it was indeed found to have a J-domain but at its C-terminal rather than at its N-terminal end. Overall, the J-domain of auxilin is 50–60% similar to the J-domains of E. coli and yeast DnaJ proteins (Ungewickell et al., 1995). Further evidence that this Jdomain is crucial for the activity of auxilin came from studies using recombinant auxilin that lacked the J-domain. While intact recombinant auxilin acts exactly like auxilin isolated from bovine brain, the recombinant auxilin that lacks the J-domain no longer supports uncoating but does bind to clathrin baskets (Ungewickell et al., 1995; Holstein et al., 1996). Interestingly, the N-terminal domain of auxilin shares similarity with tensin (Schroder et al., 1995) and this domain can be deleted resulting in much higher expression of the truncated recombinant auxilin (Holstein et al., 1996). This truncated auxilin still carries out normal uncoating and if its J-domain is deleted, like auxilin without a J-domain, the truncated molecule still binds to clathrin baskets but does not carry out uncoating (Holstein et al., 1996). Therefore, auxilin appears to be a DnaJ homolog with three domains: a C-terminal J-domain, a clathrin-binding domain, and an N-terminal tensin domain (Schroder et al., 1995; Holstein et al., 1996). 3.1. Role of Clathrin Domains in Uncoating The observation that auxilin is required for Hsc70 to uncoat clathrin baskets shed new light on the mechanism of the uncoating reaction itself, in particular on the role of the clathrin light chains and the N-terminal domains of clathrin in the uncoating reaction. It had been postulated that both the N-terminal domains of clathrin and the clathrin light chains were required for Hsc70 to uncoat clathrin baskets (Schmid et al., 1984; Schmid and Rothman, 1985). However, earlier studies on auxilin had suggested that auxilin can bind to clathrin baskets in which the light chains had been removed (Lindner and Ungewickell, 1991). This led to a reinvestigation of the involvement of the clathrin light chains and terminal domains in the uncoating process. Surprisingly, it was found that, when auxilin was present, clathrin baskets prepared from clathrin that lacked light chains and/or terminal domains, were uncoated normally by Hsc70 (Ungewickell et al., 1995). These results strongly suggest that neither the light chains nor terminal domains of clathrin are required for uncoating by Hsc70. The fact that the terminal domains are not required confirms the earlier observation of Prasad et al., (1994) that, during uncoating, Hsc70 acts by binding to the vertex of clathrin. The fact that light chains are not required for uncoating is consistent with the experiments of Acton et al. (1993) in which they found that reduced levels of clathrin light chains in the cell did not affect endocytosis and secretion. There is, however, direct interaction between one of the classes of clathrin light chains (LCa) and Hsc70 as measured by the ability of high concentrations of LCa to activate the Hsc70 ATPase and to inhibit clathrin uncoating by Hsc70 (DeLuca-Flahtery et al., 1990). This interaction could perhaps subtly alter clathrin uncoating by Hsc70 (DeLuca-Flahtery et al., 1990; Anton et al., 1993).


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3.2. Direct Interaction of Auxilin with Hsc70 Since auxilin appears to be a DnaJ homolog, it was of interest to determine whether it had the same effects on Hsc70 as several other DnaJ homologs that have been studied. These other DnaJ homologs activate the Hsc70 ATPase rate by increasing the rate of the ATP hydrolysis step but do not affect the rate of ADP release (McCarty et al., 1995; King et al., 1995b). They also catalytically induce polymerization of Hsc70 in an ATP dependent reaction (King et al., 1995a). Qualitatively, auxilin behaves similarly in its interaction with Hsc70. It causes a 5-fold increase in the Hsc70 ATPase activity and a corresponding increase in the level of bound ADP during steady-state ATP hydrolysis showing that it increases the rate of the ATP hydrolysis step, but does not increase the rate of ADP release from Hsc70 (Jiang et al., 1997). It also induces polymerization of Hsc70 in an ATP-dependent reaction that is reversed by hydrolysis of ATP to ADP (Jiang et al., 1997); the polymerized Hsc70 can be detected by column chromatography although it appears that it does not sediment (Holstein et al., 1996). Interestingly, the J-domain of auxilin also induces Hsc70 to polymerize in an ATP-dependent reaction (Jiang et al., 1997), supporting the view that auxilin is a DnaJ homolog that interacts with Hsc70 through its C-terminal J-domain. However, in contrast to other DnaJ homologs that bind to Hsc70 quite weakly and act catalytically (King et al., 1995a, King et al., 1997b), auxilin binds to the Hsc70 polymer in a 1 to 1 molar ratio, with a dissociation constant of 0.6 M. The observation that this binding is reversed with a half-life of about 4 min when ATP is hydrolyzed to ADP (Jiang et al., 1997) provides direct evidence for the view that DnaJ homologs only interact with Hsc70 in ATP. 3.3. Mechanism of Action of Auxilin Given the observation that auxilin behaves like other DnaJ homologs in its direct effects on Hsc70, it was of interest to determine whether auxilin acts by inducing the binding of clathrin-coated vesicles to Hsc70 just as other DnaJ homologs present substrates to Hsc70. It is difficult to investigate this question at pH 7 because Hsc70 becomes essentially inactive after a single round of uncoating. However, at pH 6, although uncoating is almost completely inhibited, an auxilin-dependent steady-state interaction occurs between Hsc70 and clathrin baskets. This was demonstrated in a study showing that clathrin baskets and auxilin together cause more than 100-fold activation of the steady-state Hsc70 ATPase activity at pH 6 while neither clathrin baskets nor auxilin alone significantly activate the Hsc70 ATPase activity (Barouch et al., 1997). At saturating auxilin, half-maximal Hsc70 ATPase activity occurs at 0.2 M clathrin baskets suggesting that, under these conditions, auxilin induces a strong complex to form between Hsc70 and clathrin baskets. This was confirmed by direct binding studies using sedimentation to measure complex formation between Hsc70 and clathrin baskets at pH 6. Formation of this complex is both auxilin and ATP dependent (Ungewickell et al., 1995; Barouch et al., 1997); it does not occur in ADP, and when the ATP is hydrolyzed to ADP the Hsc70 dissociates from the baskets with a half-life of about 6 min (Barouch et al., 1997). This latter observation shows that, following ATP hydrolysis, the Hsc70-clathrin basket complex is not


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energetically stable; it is metastable since it spontaneously dissociates. Taken together the studies at pH 6 strongly suggest that auxilin acts by inducing clathrin-coated vesicles to bind to Hsc70 in an ATP-depend-ent reaction, and therefore auxilin functions like other DnaJ homologs that present substrates to Hsc70. It is not yet clear whether DnaJ homologs act catalytically or stoichiometrically when they present substrates to Hsc70. Although auxilin has been reported to bind to clathrin triskelions at a 3 to 1 molar ratio (Ahle and Ungewickell, 1990), studies of uncoating at pH 7 showed that both the rate and the magnitude of uncoating saturate at less than a 1 to 5 molar ratio of auxilin to both clathrin triskelions in the baskets and Hsc70 (Figure 1). Even when the auxilin is increased to a 3 to 1 molar ratio to clathrin by preparing clathrin baskets with auxilin, alone, as an assembly protein, neither the rate nor magnitude of uncoating increases (closed triangles, Figure 1). Therefore, auxilin acts catalytically rather than stoichiometrically at pH 7, which raises the question of whether auxilin also acts catalytically when it induces binding of Hsc70 to clathrin baskets at pH 6. Studies at pH 6 showed that auxilin preferentially binds to clathrin baskets rather than Hsc70. These studies also showed that auxilin acts substoichiometrically when it activates the steadystate Hsc70 ATPase activity with maximum ATPase activation occurring at a molar ratio of 1 auxilin per clathrin triskelion in the baskets (Barouch et al., 1997). Even less auxilin is required to induce maximal binding of Hsc70 to clathrin baskets. Maximal binding occurs at about a 1 to 3 molar ratio of auxilin to clathrin triskelions in the clathrin baskets at pH 6 (Barouch et al., 1997) and pH 6.5 (Ungewickell et al., 1995) confirming that auxilin acts catalytically when it induces binding of Hsc70 to clathrin baskets at pH 6. This, in turn, shows that auxilin must induce binding of Hsc70 to clathrin baskets without itself becoming part of the resulting complex. The question then arises as to whether other DnaJ homologs can also catalytically induce the binding of clathrin baskets to Hsc70. Investigation of this question showed that neither the J-domain of auxilin by itself (Holstein et al., 1996) or other DnaJ homologs such HSJ1, YDJ1, and HDJ1 (Cheetham et al., 1996; King et al., 1997b) support uncoating. In fact, these other DnaJ homologs inhibit uncoating, and they do so under conditions where they either do not induce Hsc70 to polymerize (Cheetham et al., 1996) or cause only partial polymerization of Hsc70 (King et al., 1997b) suggesting that they are not inhibiting uncoating simply by polymerizing Hsc70 before it can interact with clathrin baskets. Rather, these homologs may inhibit uncoating by inducing hydrolysis of ATP to ADP before Hsc70-ATP can bind to the auxilin-clathrin basket complex. Therefore, auxilin may specifically support uncoating because it juxtaposes the Hsc70 and clathrin baskets allowing the clathrin baskets to form a transient complex with Hsc70-ATP just before auxilin activates hydrolysis of ATP to ADP. This view of auxilin action is similar to a model of DnaJ action proposed by Bukau and his collaborators (McCarty et al., 1995). It is also supported by the observation that the activity of auxilin can be partially restored by cross-linking the J-domain with the clathrin-binding domain of auxilin, neither of which is active by itself (Holstein et al., 1996). These results suggest that if other DnaJ homologs act like auxilin they would also present only specific substrates to Hsc70, rather than non-specifically presenting all substrates to Hsc70.


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3.4. Contrast Between Substrates and Products of Hsc70 Studies on the auxilin-induced binding of clathrin baskets to Hsc70 reveal major differences in the way Hsc70 interacts with the substrate of the uncoating reaction, clathrin baskets, and the product of the uncoating reaction, free clathrin triskelions. First, although auxilin is required for the binding of clathrin baskets to Hsc70 in ATP it is not required for the binding of free clathrin triskelions. Second, the auxilin-induced binding of clathrin baskets to Hsc70 in ATP is an order of magnitude stronger than the binding of clathrin triskelions (Barouch et al., 1997; Prasad et al. 1994). Third, auxilin only induces clathrin baskets to bind to Hsc70 in the presence of ATP (Ungewickell et al., 1995) while clathrin triske ions bind stably to both Hsc70-ADP (Prasad et al., 1994) and nucleotidefree Hsc70 (Gao et al., 1995a). Finally mutations of Hsc70 that have a marked effect on the auxilin-induced binding of clathrin baskets to Hsc70 have little effect on the binding of clathrin triskelions (Rajapandi et al., 1998). Therefore, Hsc70 interacts very differently with clathrin baskets, the substrate of the uncoating reaction, and free clathrin triskelions, the product of the uncoating reaction. The distinction between the way Hsc70 interacts with clathrin baskets and free clathrin triskelions may have major implications for studies on the interaction of other ligands with Hsc70. Generally, ligands such as peptides or reduced carboxymethylated lactalbumin whose interaction with Hsc70 is affected by ATP have been considered substrates of Hsc70 (Palleros et al., 1991; Palleros et al., 1993). This may be misleading, however, because in their ability to bind to Hsc70-ATP in the absence of DnaJ homologs and in their stable binding to Hsc70-ADP these ligands behave more like clathrin triskelions than clathrin baskets, that is more like products than substrates of Hsc70. It may be that only ligands that can be presented to Hsc70 by DnaJ homologs in ATP are true substrates of Hsc70. 4. MODEL OF UNCOATING Based on our current understanding of the mechanism of action of auxilin, we have developed a model for the uncoating reaction (Figure 2). In this model, the initial burst of uncoating starts with auxilin binding to clathrin baskets. The auxilin then induces Hsc70ATP to bind to clathrin baskets and this is immediately followed byauxilin inducing rapid hydrolysis of the ATP bound to Hsc70. Note that in this model, auxilin activates ATP hydrolysis by Hsc70 only after formation of the Hsc70-ATP-clathrin basket complex. If auxilin activated ATP hydrolysis before this complex formed, the resulting Hsc70-ADP complex would not bind to the clathrin baskets since auxilin does not induce binding of clathrin baskets to Hsc70-ADP. Our model suggests that, following ATP hydrolysis and dissociation of auxilin, three Hsc70-ADPs are bound to a clathrin triskelion in the clathrin basket. Studies at pH 6 suggest that this is a metastable complex that, with time, would completely dissociate (Barouch et al., 1997). On the other hand, Hsc70-ADP does form an energetically stable complex with dissociated clathrin triskelions (Prasad et al.,


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Figure 2 Model for the uncoating of clathrin-coated vesicles by Hsc70. The ATP-bound and ADP-bound forms of Hsc70 are shown by the white and black rectangles, respectively.

1994). Therefore, the power stroke of the uncoating reaction may be a transformation of the metastable Hsc70-ADP-clathrin basket complex to the energetically stable Hsc70ADP-dissociated clathrin complex. More generally, a similar transformation from a metastable Hsc70-ADP-substrate complex to a stable Hsc70-ADP-product complex could act as the power stroke for other reactions involving DnaJ homologs such as the translocation of proteins through membranes and the unfolding of misfolded proteins. Since the clathrin lattice is much more stable at pH 6 than at pH 7 (Van Jaarsveld et al., 1981), the transformation from the metastable Hsc70-ADP-clathrin basket complex to the energetically stable Hsc70-ADP-dissociated clathrin complex may be unable to occur at pH 6, which may explain why uncoating is strongly inhibited at pH 6. However, at pH 7, once all three legs of a clathrin triskelion form stable complexes with Hsc70-ADP, the clathrin will dissociate from the clathrin lattice. Since three Hsc70s are required to dissociate a single clathrin triskelion in this model, the model implies that Hsc70 should act cooperatively in the uncoating process but evidence for such cooperativity has not yet been observed. Since Hsc70-ADP dissociates very slowly from dissociated clathrin triskelions, our model suggests that, in the last step of the uncoating cycle, ADP is released and ATP rebinds to the Hsc70s complexed with the dissociated clathrin triskelion, thereby freeing up the Hsc70 to carry out further cycles of uncoating. In our original model of uncoating we proposed that this slow step was the rate-limiting step in the uncoating reaction and could therefore account for the very slow steady-state uncoating (Greene and Eisenberg, 1990). However, in ATP the rate of dissociation of clathrin from the Hsc70-ADP-Pi-


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clathrin complex is too fast to account for the very slow steady-state uncoating rate (Prasad et al., 1994). Hence, there must be some other explanation for the slow steadystate uncoating. 5. TIME COURSE OF UNCOATING Studies on the time course of uncoating were directed to answering two key questions that are not explained by our model of uncoating. First, we investigated why auxilin acts catalytically, that is why the rate of the initial burst of uncoating saturates at very low levels of auxilin. Second, we investigated the cause of the slow steady-state uncoating. 5.1. Initial Burst of Uncoating Using purified AP180-clathrin baskets plus auxilin, it has been demonstrated that an initial burst of ATP hydrolysis occurs with the same time course as the initial burst of uncoating with three ATPs hydrolyzed per clathrin triskelion uncoated (Barouch et al., 1994). It therefore appears that the initial burst of uncoating represents a single pre-steady-state turnover of ATP by Hsc70. If the rate-limiting step in this single turnover occurred after Hsc70 bound to the clathrin baskets, e.g. if it were the ATP hydrolysis step itself, it would be easy to explain the ability of auxilin to act catalytically since low levels of auxilin could induce Hsc70 to bind to the clathrin baskets much faster than Hsc70 could uncoat them. However, the observation that most of the Hsc70 remains free in solution during the initial burst of uncoating (unpublished data) suggests that it is binding of Hsc70 to clathrin baskets that is rate-limiting, not a step that occurs after binding of Hsc70. This view was confirmed by showing that the rate of the initial burst of uncoating linearly increases with an increase in the Hsc70 concentration even when the Hsc70 concentration is in excess over the clathrin basket concentration (unpublished data); this could not occur if the rate-limiting step in the uncoating reaction occurred after Hsc70 bound to the clathrin baskets. If the binding of Hsc70 to clathrin baskets is the rate-limiting step in the uncoating reaction, simple kinetics predicts that the rate of the initial burst of uncoating should not only linearly increase with an increase in Hsc70 concentration but should also linearly increase with an increase in clathrin basket concentration or the amount of auxilin bound to the clathrin baskets. However, as we pointed out above, the rate of uncoating levels off at very low auxilin concentration (Figure 1). Furthermore, recent experiments showed that the rate of uncoating also does not increase with increasing concentrations of clathrin baskets (unpublished data). One explanation for this unusual kinetic phenomenon is that the rate of the initial burst


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Figure 3 Recent findings on the uncoating cycle. This model alters the model in Figure 2 to include, first, a rate-limiting conformational change in the Hsc70-ATP form, depicted by the transition from the white to the grey rectangle, that occurs prior to the binding of Hsc70 to the baskets; and, second, formation of a steady-state complex between Hsc70, clathrin and assembly proteins after the initial burst of uncoating, which inhibits further uncoating by Hsc70.

of uncoating is controlled by a conformational change that must occur in the Hsc70 before it can bind to the clathrin baskets as illustrated in Figure 3. In this case, increasing the auxilin concentration or the clathrin basket concentration would not increase the rate of uncoating because most of the Hsc70 would be unable to bind to the clathrin baskets until the Hsc70 underwent the rate-limiting conformational change. If, indeed, Hsc70 must undergo a conformational change before it interacts with auxilin and clathrin baskets, it could have major implications for the general mechansim of action of the Hsp70 class of proteins. 5.2. Slow Steady-state Uncoating Detailed studies have ruled out several explanations for the slow steady-state uncoating (Gao et al., 1995b). First, when fresh Hsc70 is added to partially uncoated baskets, a second round of uncoating takes place showing that the partially uncoated baskets are still capable of being uncoated. Second, radioactively labeled clathrin does not rebind to clathrin baskets during slow steady-estate uncoating, ruling out that clathrin is still being released during slow steady-state uncoating but because the clathrin exchanges with clathrin previously released during the initial burst of uncoating, no net uncoating occurs. Therefore, the slow steady-state uncoating is due to an inability of Hsc70 in the


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supernatant to carry out further uncoating. However, this Hsc70 is not irreversibly altered; its uncoating activity can be completely restored by chromatographing the supernatant of the uncoating reaction on a Superose 6 or 12 FPLC column that separates Hsc70 from clathrin and assembly proteins (Gao et al., 1995b). Evidence that assembly proteins are involved in causing the slow steady-state uncoating comes from studies on highly purified clathrin baskets prepared in calcium in the absence of assembly proteins (Gao et al., 1995b). Auxilin is required for the uncoating of these baskets, although less auxilin than is required for the uncoating of clathrin baskets prepared with assembly proteins. Strikingly, however, the time course of uncoating of these pure clathrin baskets is not biphasic. Rather, a nearly constant rate of uncoating occurs until the clathrin baskets are completely uncoated. When assembly proteins are added to the pure clathrin baskets, the time course of uncoating again becomes biphasic confirming that assembly proteins are indeed involved in causing the slow steady-state uncoating. However, a simple mixture of depolymerized clathrin triskelions and assembly proteins does not inhibit uncoating (Jiang et al., 1998). One explanation for the slow steady-state uncoating is that clathrin and assembly proteins are released as a special complex from clathrin baskets during the uncoating reaction, and Hsc70 preferentially binds to this complex, thus preventing Hsc70 from carrying out further uncoating as illustrated in Figure 3. If indeed, Hsc70 interacts with uncoated complexes of free clathrin triskelia and assembly proteins, it may reflect a physiological role for Hsc70 either in preventing these complexes from inappropriately polymerizing in the cytosol or in rapidly catalyzing the formation of clathrin-coated pits from these complexes in nerve cells. In fact, we have recently obtained evidence that these complexes do indeed interact with uncoated vesicle membranes in vitro (Jiang et al., 1998). In addition, in vivo, a stable non-membranous Hsc70-clathrin complex forms in neurons and is transported along the axon (De Waegh and Brady, 1989; Black et al., 1991), and over-expression of auxilin in HeLa cells causes it to form large complexes with clathrin and Hsc70 (Zhao et al., 1998). Interestingly, this complex formation actually inhibits endocytosis in these cells, apparently because the clathrin is inappropriately sequestered away from normal clathrin-coated pits on the plasma membrane (Zhao et al., 1998). 6. CONCLUDING REMARKS The uncoating of clathrin-coated vesicles provides an excellent model system for studying the role of DnaJ homologs and ATP in the mechanism of action of mammalian Hsc70. In addition, the role of Hsc70 and auxilin in receptor-mediated endocytosis is of interest in its own right. It is now clear that uncoating requires the DnaJ homolog, auxilin, as well as Hsc70 and ATP. Auxilin contains both a J-domain and a clathrin-binding domain so it not only interacts directly with Hsc70 but also with clathrin-coated vesicles. Perhaps because other DnaJ homologs do not contain a clathrin binding domain, they cannot substitute for auxilin in inducing uncoating, suggesting that auxilin and perhaps other DnaJ homologs, as well, only present specific substrates to Hsc70 rather than nonspecifically presenting all substrates to Hsc70. There is good evidence that auxilin functions by first inducing formation of a complex


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between Hsc70-ATP and clathrin-coated vesicles and then inducing rapid ATP hydrolysis. Since clathrin baskets do not spontaneously bind to Hsc70-ADP in the presence of auxilin, this leads to formation of a metastable Hsc70-ADP-clathrin basket complex. However, Hsc70-ADP does stably bind to dissociated clathrin triskelions. It is therefore possible that a transition from a metastable Hsc70-ADP-clathrin basket complex to a stable Hsc70-ADP-dissociated clathrin complex provides the power stroke for the uncoating reaction; a similar transition might provide the power stroke in other reactions where Hsc70s in combination with DnaJ homologs utilize the free energy of ATP hydrolysis to do work. This mechanism of action of Hsc70 is based on the observation that Hsc70 interacts very differently with the substrate of the uncoating reaction, clathrin baskets, and the product of the uncoating reaction, clathrin triskelions. Interestingly, other ligands such as peptides and reduced carboxymethylated lactalbumin resemble clathrin triskelions rather than clathrin baskets in their interaction with Hsc70 suggesting that they may act more like products than substrates of Hsc70. One of the key characteristics of the uncoating reaction is its biphasic time course, which consists of a rapid initial burst of uncoating followed by very slow steady-state uncoating. The initial burst of uncoating shows an unusual kinetic phenomenon in which the binding of Hsc70 to clathrin baskets is the rate-limiting step in the initial burst of uncoating yet this rate is independent of both the clathrin basket concentration and the auxilin concentration. A posible explanation for this kinetic phenomenon is that Hsc70 must undergo a conformational change before it binds to clathrin baskets. If so, it will be important to determine if Hsc70 undergoes a similar conformational change in other reactions that it carries out. As for the slow steady-state uncoating, current evidence suggests that it is due to an interaction of Hsc70 with special clathrin-assembly protein complexes produced only during uncoating; this interaction apparently prevents Hsc70 from carrying out further uncoating. It is possible that interaction of Hsc70 with uncoated clathrin-assembly protein complexes in vitro reflects a role for Hsc70 in regulating clathrin dynamics in vivo. Furthermore, since auxilin is a nerve-specific protein, such a phenomenon might play a role in the rapid endocytosis that De Camilli and Takei (1996) have suggested is associated with synaptic transmission. We are currently investigating if auxilin-like proteins support uncoating in nonnervous system tissues and are also attempting to determine why uncoating occurs with varying efficiencies with clathrin-coated vesicles from different tissues in vitro (Buxbaum and Woodman, 1995; Buxbaum and Woodman, 1996). Interestingly, a recently cloned protein, GAK, contains both an N-terminal kinase domain (Kanaoka et al., 1997) and a C-terminal domain highly homologous to auxilin. Recently we have determined that this latter domain acts exactly like auxilin in vitro in regard to its interaction with Hsc70 and clathrin baskets (Greener et al., 1998). Furthermore, unlike auxilin, this protein shows a wide tissue distribution raising the possibility that it could function like auxilin in non-neuronal tissues. However, this protein is also associated with both cyclin G and CDK5, and, therefore its full function in vivo is not yet understood (Kanaoka et al., 1997; Kimura et al., 1997). Future studies will be required to determine if this protein indeed acts like auxilin in non-neuronal tissues and also to determine the role of its kinase domain in vivo.


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7. REFERENCES Able, S. and Ungewickell, E. (1990). Auxilin, a newly identified clathrin-associated protein in coated vesicles from bovine brain. J. Cell Biol. , 111 , 19–29. Able, S. and Ungewickell, E. (1986). Purification and properties of a new clathrin assembly protein. EMBO J. , 5 , 3143–3149. Acton, S., Wong, D. Parhma, P, Brodsky, F. and Jackson, A. (1993). Alteration of clathrin light chain expression by transfection and gene disruption. Mol. Biol. Cell , 4 , 647–660. Barouch, W., Prasad, K., Greene, L.E. and Eisenberg, E. (1994). ATPase activity associated with the uncoating of clathrin baskets by Hsp70. J. Biol. Chem. , 269 , 28563–28568. Barouch, W., Prasad, K., Greene, L. and Eisenberg, E. (1997). Auxilin induced interaction of the molecular chaperone Hsc70 with clathrin baskets. Biochemistry , 36 , 4304–4308. Black, M., Chestnut, M., Pleasure I. and Keen, J. (1991). Stable clathrin: uncoating protein complexes in intact neurons and their axonal transport. J. Neurosci. and , 11 , 1163–1172. Braell, W.A., Schlossman, D.M., Schmid, S.L. and Rothman, J.E. (1984). Dissociation of clathrin coats coupled to the hydrolysis of ATP: role of an uncoating ATPase. J. Cell Biol. , 99 , 734–741. Brodsky, F.M. (1988). Living with clathrin: its role in intracellular membrane traffic. Science , 242 , 1396–1402. Buxbaum, E. and Woodman, P.G. (1995). Selective action of uncoating ATPase towards clathrin-coated vesicles from brain. J. Cell Sci. , 108 , 1295–1306. Buxbaum, E. and Woodman, P. (1996). The speed of partial reactions of the uncoating ATPase Hsc70 depends on the source of coated vesicles. J. Cell Sci. , 109 , 705–711. Caplan A.J., Cyr, D.M. and Douglas, M.G. (1993). Eukaryotic homologues of Escherichia coli dnaJ: A diverse protein family that functions with Hsp70 stress proteins. Mol. Biol. Cell , 4 , 555–563. Chappell, T.G., Konforti B.B., Schmid S.L. and Rothman, J.E. (1987). The ATPase core of a clathrin uncoating protein. J. Biol. Chem. , 262 , 746–751. Chappell, T.G., Welch, W.J., Schlossman, D.M., Palter, K.B., Schlesinger, M.J. and Rothman, J.E. (1986). Uncoating ATPase is a member of the 70 kilodalton family of stress proteins. Cell , 45 , 3–13. Cheetham, M.E., Anderton, B.H. and Jackson, A.P. (1996). Inhibition of hsc70-catalysed clathrin uncoating by HSJ1 proteins. Biochem J. , 319 , 103–108. Cyr, D.M., Langer, T. and Douglas, M.G. (1994). DnaJ-like proteins: molecular chaperones and specific regulators of Hsp70. Trends Biochem. Sci. , 19 , 176–181. De Camilli, P. and Takei, M. (1996). Molecular mechanisms in synaptic vesicle endocytosis and recycling. Neuron , 16 , 481–486. DeLuca-Flaherty, C., McKay, D., Parham, P. and Hill, B. (1990). Uncoating protein (hsc70) binds a conformationally labile domain of clathrin light chain LCa to stimulate ATP hydrolysis. Cell , 62 , 857–887.


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De Waegh, S. and Brady, S. (1989). Axonal transport of a clathrin uncoating ATPase: a role for Hsc70 in the modulation of coated vesicle assembly in vivo. J.Neurosci. Res. , 4 , 433–440. Gao, B., Biosca, J., Craig, E.A., Greene, L.E. and Eisenberg, E. (1991). Uncoating of coated vesicles by yeast Hsp70 proteins. J. Biol. Chem. , 266 , 19565–19571. Gao, B., Emoto, Y., Greene, L. and Eisenberg, E. (1993). Nucleotide binding properties of bovine brain uncoating ATPase. J. Biol. Chem. , 268 , 8507–8513. Gao, B., Eisenberg, E. and Greene, L. (1995a). Interaction of nucleotide-free Hsc70 with clathrin and peptide and effect of ATP analogues. Biochemistry , 34 , 11882–11888. Gao, B., Prasad, K., Greene, L. and Eisenberg, E., (1995b). Uncoating of clathrin baskets by Hsp70: cause of inhibition following one round of uncoating. Mol. Cell Biol. , 6 , 411a. Greene, L.E. and Eisenberg, E. (1990). Dissociation of clathrin from coated vesicles by the uncoating ATPase. J. Biol. Chem. , 265 , 6682–6687. Greene, L.E., Zinner, R., Naficy, S. and Eisenberg, E. (1995). Effect of nucleotide on the binding of peptides to 70-kDa heat shock protein. J. Biol. Chem. , 270 , 2967–2973. Greener, T., Nojima, H., Eisenberg, E. and Greene, L. (1998). Cyclin G associated kinase (GAK) has kinase and auxilin domains that are active in vitro. FASEB J. , 12 , in press. Hartl, F. (1996). Molecular chaperones in cellular protein folding. Nature , 381 , 571– 580. Hendrick, J.P. and Hartl, F.U. (1993). Molecular chaperone functions of heat-shock proteins. Annu. Rev. Biochem. , 62 , 349–384. Heuser, J. (1980). Three-dimensional visulazation of coated vesicle formation in fibroblasts. J. Cell Biol. , 84 , 560–583. Heuser, J. and Kirchhausen, T. (1985). Deep-etch views of clathrin assemblies. J. Ultrastruct. Res. , 92 , 1–27. Heuser, J. and Steer, C.J. (1989). Trimeric binding of the 70-kDa uncoating ATPase to the vertices of clathrin triskelia: a candidate intermediate in the vesicle uncoating reaction. J. Cell Biol. , 109 , 1457–1466. Holstein, S.E.H., Ungewickell, H. and Ungewickell, E. (1996). Mechanism of clathrin basket dissociation: separate functions of protein domains of the dnaJ homologue auxilin. J. Cell Biol. , 135 , 925–937. Honing, S., Kreimer, G., Robenek, H. and Jockusch, B. (1994). Receptor-mediated endocytosis is sensitive to antibodies. J. Cell Set. , 107 , 1185–1186. Jiang, R., Gao, B., Eisenberg, E. and Greene, L. (1998). An Hsc70-clathrin complex formed during uncoating may be involved in rebinding of clathrin to uncoated clathrin vesicles. FASEB J. , 12 , in press. Jiang, R.-F., Greener, T., Barouch, W., Greene, L. and Eisenberg, E. (1997). Interaction of auxilin with the molecular chaperone, Hsc70. J. Biol. Chem. , 272 , 6141–6145. Kanaoka, Y., Kimura, S., Okazaki, I.Ikeda, M. and Nojima, H. (1997). GAK: a cyclin G associated kinase contains a tensin/auxilin-like domain. FEBS Lett , 402 , 73–80. Keen, J.H., Willingham, M.C. and Pastan, I. (1979). Clathrin coated vesicles: Isolation dissociation and factor-dependent reassociation of clathrin baskets. Cell , 16 , 303–312. Keen, J.H. (1990). Clathrin and associated assembly and disassembly proteins. Annu. Rev. Biochem. , 59 , 415–438.


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Kimura, S., Tsuruga, H. Yabuta, N. Endo, Y. and Nojima, H. (1997). Structure, expression and chromosomal localization of human GAK. Genomics , 44 , 179–187. King, C., Eisenberg, E. and Greene, L. (1995a). Polymerization of 70-kDa heat shock protein by yeast DnaJ in ATP. J. Biol. Chem. , 270 , 22535–22540. King, C., Eisenberg, E. and Greene, L. (1995b). Polymerization of Hsp70 by DnaJ in the presence of ATP. Mol. Cell Biol. , 67a , 391. King, C., Eisenberg, E. and Greene., L. (1997a). Interaction between Hsc70 and DnaJ homologs: evidence for weak binding of DnaJ homologs to Hsc70. Mol Bio. Cell , 8 , 96a. King, C., Eisenberg, E. and Greene, L. (1997b). Effect of yeast and human DnaJ homologs on clathrin uncoating by 70 kDa-Heat Shock Protein. Biochemistry , 36 , 4067–4073. Lindner, G. and Ungewickell, E. (1991). Light-chain-independent binding of adaptors, AP180 and auxilin to clathrin. Biochemistry , 30 , 9097–10101. McCarty, J.S., Buchberger, A., Reinstein, J. and Bukau, B. (1995). The role of ATP in the functional cycle of the DnaK chaperone system. J. Mol. Biol. , 249 , 126–137. Palleros, D.R., Welch, W.J. and Fink, A.L. (1991). Interaction of Hsp70 with unfolded proteins: effects of temperature and nucleotides on the kinetics of binding. Proc. Natl. Acad. Sci. USA , 88 , 5719–5723. Palleros, D.R., Reid, K.L., Shi, L., Welch, W.J. and Fink, A.L. (1993). ATP-induced protein-Hsp 70 complex dissociation requires K+ but not ATP hydrolysis. Nature , 365 , 664–666. Pearse, B.M.F. (1987). Clathrin and coated vesicles. EMBO J. , 6 , 2507–2512. Pearse, B.M. and Robinson, M.S. (1990). Clathrin, adaptors and sorting. Annu. Rev. Cell Biol , 6 , 151–171. Prasad, K., Barouch, W., Greene, L. and Eisenberg, E. (1993). A protein cofactor is required for uncoating of clathrin baskets by uncoating ATPase. J. Biol. Chem. , 268 , 23758–23761. Prasad, K., Barouch, W., Martin, B.M., Greene, L.E. and Eisenberg, E. (1995). Purification of a new clathrin assembly protein from bovine brain coated vesicles and its identification as myelin basic protein. J. Biol. Chem. , 270 , 30551–30556. Prasad, K., Heuser, J., Eisenberg, E. and Greene, L. (1994). Complex formation between clathrin and uncoating ATPase. J. Biol. Chem. , 269 , 6931–6939. Prasad, K. and Lippoldt, R. (1988). Molecular characterization of the AP180 assembly protein. Biochemistry , 27 , 6098–6104. Rajapandi, T., Wu, C., Eisenberg, E. and Greene, L. (1998). Characterization of DIOS and K71E mutants of human cytosolic Hsp70. Biochemistry , 37 , in press. Rassow, J. and Pfanner, N. (1995), Molecular chaperones and intracellular protein translocation. Rev. Physiol. Biochem. Pharmacol. , 126 , 199–264. Rassow, J., Voos, W. and Pfanner, N. (1995). Partner proteins determine multiple functions of Hsp70. Trends Cell Biol , 5 , 207–212. Robinson, M.S. (1992). Adaptins. Trends Cell Biol , 2 , 293–297. Schmid, D., Baici, A., Gehring, H. and Christen, P. (1994). Kinetics of molecular chaperone action. Set. , 263 , 971–973. Schmid, S.L., Braell, W.A. and Rothman, J.E. (1985). ATP catalyzes these questration of


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clathrin during enzymatic uncoating. J. Biol. Chem. , 260 , 10057–10062. Schmid, S.L., Braell, W.A., Schlossman, D.M. and Rothman, J.E. (1984). A role for clathrin light chains in the recognition of clathrin cages by uncoating ATPase. Nature , 311 , 228–231. Schmid, S.L. and Rothman, J. (1985). Two classes of binding sites for uncoating protein in clathrin triskelions. J. Biol. Chem. , 260 , 10050–10056. Schlossman, D.M., Schmid, S.L., Braell, W.A. and Rothman, R.E. (1985). An enzyme that removes clathrin coats: purification of an uncoating ATPase. J. Cell Biol . 99 , 723–733. Schroder, S., Morris, S., Knorr, R., Plessmann, U., Weber, K., Vinh, N. and Ungewickell, E. (1995). Primary structure of the neuronal clathrin-associated protein auxilin and its expression in bacteria. Eur. J. Biochem. , 228 , 297–304. Takeda, S. and McKay, D.B. (1996). Kinetics of peptide binding to the bovine 70 kDa heat shock cognate protein, a molecular chaperone. Biochemistry , 35 , 4636–4644. Tsai, M. and Wang, C. (1994). Uncoupling of peptide-stimulated ATPase and clathrinuncoating activity in deletion mutation of hsc70. J. Biol. Chem. , 269 , 5958–5962. Ungewickell, E. (1985). The 70-kd mammalian heat shock proteins are structurally and functionally related to the uncoating protein that releases clathrin triskelia from coated vesicles. EMBO J. , 4 , 3385–3391. Ungewickell, E., Ungewickell, H., Holstein, S., Lindner, R., Prasad, K., Barouch, W., Martin, B., Greene, L. and Eisenberg, E. (1995). Role of auxilin in uncoating clathrincoated vesicles. Nature , 378 , 632–635. Ungewickell, E., Ungewickell, H. and Holstein, S., (1997). Functional interaction of the auxilin J domain with the nucleotide-and substrate-binding modules of hsc70 . J. Biol. Chem. , 272 , 19594–19600. Van Jaarsveld, P., Nandi, P., Lippoldt, R., Saroff, H. and Edelhoch, H. (1981). Polymerization of clathrin protomers into basket structures. Biochemistry , 20 , 4129– 4135. Wei, J., Gaut, J.R. and Hendershot, L.M. (1995). In vitro dissociation of BiP-peptide complexes requires a conformational change in BiP after ATP binding but does not require ATP hydrolysis. J. Biol. Chem. , 44 , 26677–26682. Zhao, X., Greener, T., Greene, L. and Eisenberg, E. (1998). Expression of the clathrin assembly protein, auxilin, in tissue culture cells inhibits endocytosis. FASEB J. , 12 , in press.

17. THE ROLE OF HSP104 IN STRESS TOLERANCE AND PRION MAINTENANCE SUSAN LINDQUIST* and ERIC C.SCHIRMER Howard Hughes Medical Institute and the Department of Molecular Genetics and Cell Biology, The University of Chicago, 5841 S. Maryland Avenue, MC 1028, Chicago, IL 60637, USA

1. Introduction 2. The HSP100/CLP Family of Proteins 2.1. A Large Family of Proteins with Diverse Biological Activities 2.2. Hsp104 Protein Structure 3. Biochemical Properties of Hsp104 3.1. Oligomerization 3.2. ATPase Activity 4. The Role of Hsp104 in Stress Tolerance 4.1. Thermotolerance and the Importance of Hsp104 4.2. The Thermotolerance Function of Hsp104 4.2.1. No Evidence for a Proteolysis Function 4.2.2. A Disassembling/resolubilizing Activity 4.3. Other Stresses 4.4. Combinatorial Stresses 4.5. Physiological Inductions of Tolerance 4.6. Conservation of Tolerance Functions in B-Type Proteins 4.7. The Relationship Between Hsp104 and Other Tolerance Factors 4.7.1. Other Tolerance Factors 4.7.2. Genetic Interactions Between Hsp70 and Hsp104 4.7.3. Genetic Interactions Between Trehalose Synthase and Hsp104 5. Hsp104’s Interaction with a Yeast Prion 5.1. A Role for Hsp104 During Normal Growth 5.2. The Prion Hypothesis 5.3. The Relationship Between [PSI+] And Sup35 5.4. The Role of Hsp104 in the Maintenance of [PSI+] 5.4.1. Overexpression of HSP104 Cures [PSI+] 5.4.2. Deletion of HSP104 Cures [PSI+] 5.4.3. Hsp104 and Sup35 Aggregation in Vivo

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5.4.4. Seeded Polymerization of Sup35 5.4.5. Hsp104 and Sup35 Interaction in Vitro 5.5. A Model For Hsp104-[PSI+] Interactions 5.6. An Explanation For Previous Puzzles 5.6.1. Hsp104 Explains Puzzling [PSI+] phenomena 5.6.2. [PSI + ] Explains a Puzzling Thermotolerance Phenomenon 5.7. Broader Implications 5.8. Hsp104 and Mammalian Prions 6. Concluding Remarks 7. References

1. INTRODUCTION The gene encoding Hsp104 from Saccharomyces cerevisiae was first cloned in an effort to decipher the mechanisms that cells employ to survive a variety of toxic conditions (Sanchez and Lindquist, 1990). Hsp104 is expressed at a low level under most growth conditions, but it is strongly induced by a variety of environmental stresses (Figure 1 and Sanchez et al., 1992). The protein proved to play a key role in helping cells survive many of these stresses, particularly under the most extreme conditions (Sanchez and Lindquist, 1990; Sanchez et al., 1992; Lindquist et al., 1995). Recent work indicates that, unlike many other Hsp chaperones, Hsp104 has little or no capacity to prevent the aggregation of denatured proteins. Nor does it promote the forward folding of denatured proteins. Rather, Hsp 104 helps to disentangle partially aggregated, stress damaged proteins so that they can be refolded by other chaperone systems in the cell (Parsell et al., 1994b; Glover and Lindquist, 1998). Hsp104 was independently cloned in a series of experiments aimed at understanding the mysterious mechanism of inheritance of the genetic element known as [PSI + ] (Chernoff et al., 1995). Recent genetic and biochemical evidence indicates that [PSI + ] is a prion-like element (Wickner, 1994; Chernoff et al., 1995; Patino et al., 1996). The inheritance of the [PSI + ] phenotype is due to the inheritance of an altered physical state of the normal cellular protein, Sup35. Once the altered state has been established, it is self-perpetuating in wild-type cells. Hsp 104 was cloned in a screen for proteins whose overexpression suppresses the [PSI + ] phenotype. The role of Hsp104 in this remarkable and novel mechanism of inheritance is related to its role in thermotolerance—that is, its ability to alter the conformational state of other proteins. The relationship between Hsp 104 and thermotolerance, however, is simpler than the relationship between Hsp104 and [PSI + ]. As the concentration of Hsp104 increases, the ability of cells to survive high temperatures increases, in a direct relationship (Lindquist and Kim, 1996). In contrast, the ability of cells to inherit the [PSI + ] factor depends upon the ratio of Hsp 104 to Sup35. Cells are cured of [PSI + ] if they express either too much


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or too little Hsp104 (Chernoff et al., 1995). Determining the precise molecular mechanisms that dictate the role of Hsp 104 in stress tolerance and prion maintenance represents a major challenge for future investigations.

Figure 1 Hsp104 expression patterns. (A) Patterns of protein synthesis in yeast cells. Cells grown to mid-log phase in glucose media at 25°C were labeled for 30 min with 3H-isoleucine at 25°C (C, control cells) or 15 min after a shift to 39°C (H, heatshocked cells). Total cellular proteins were separated by electrophoresis in SDS gels and label incorporation was visualized by fluorography. (B) Accumulation of Hsp104 in response to stress. Cells grown to mid-log phase in glucose media at 25°C were maintained at 25°C (C), heat shocked at 39°C for 30 min (H), or incubated at 25°C in the presence of 6% ethanol (EtOH) for 1 hr, 0.75M sodium arsenite for 1 hr (As), or 1.5 mM cadmium chloride for 30 min (Cd). Proteins were separated by electrophoresis in SDS gels, transferred to membranes, and reacted with an antibody directed against Hsp104. Immune complexes were visualized by incubation with 125I-labeled protein A followed by fluorography. (C) Accumulation of Hsp104 under specific, normal physiological conditions. Left: cells were grown to mid-log phase at 25°C in synthetic glucose medium (S-Glu) or synthetic acetate medium (S-Ac) and maintained at 25°C (C) or heat shocked at 39°C for 30 min. Middle: cells grown


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at 25°C in glucose medium were collected at various stages in the growth curve, from mid-log phase to stationary phase. Right: diploid cells were sporulated in low nitrogen media treated with zymolyase and mature spores were collected on Percoll gradients. Hsp104 protein was visualized as in B. Data taken from (Sanchez et al., 1992).

2. THE HSP100/CLP FAMILY OF PROTEINS 2.1. A Large Family of Proteins with Diverse Biological Activities Hsp104 belongs to a newly discovered family of chaperones with diverse patterns of expression, subcellular localizations, and biological functions, known as the

Figure 2 Structural features of the HSP100 family. Thicker boxes indicate greater conservation and particularly conserved sequence motifs unique to the family are indicated I–V. Abrupt transitions in conservation demarcate the two ATP-binding domains, the aminoterminal, middle, and carboxyl-terminal regions. The X and Y proteins are more distantly related and their single ATP-binding domain is depicted as lighter in color. The locations of the Walkertype nucleotide-binding motifs, A and B, are indicated. (It is not clear if both B motifs in NBD1 are used.) Class 1 proteins contain two NBDs. Class 2 proteins contain only one NBD and the carboxylterminal region. The M and N subfamilies of Class 2 are distinguished from the X and Y subfamilies by much greater homology to the Class 1 proteins. To the right are listed the percent similarity between the ATP-binding domains of representatives of different subfamilies or the overall percent similarities. The numbers listed were determined using the DNAStar


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program with the Clustal method and PAM250 residue weight table. For class 2 HSP100 proteins, just the NBD2 and C-terminal domains were used for the overall analysis. The proteins representative of each subfamily used in this analysis are (A) Escherichia coli ClpA, (B) E. coli ClpE, (C) Tomato ClpC, (D) Ambidopsis ERD1, (M) mouse SKD2, (N) Pseudomonas aeruginosa AmiB, (X) E. coli ClpX, (Y) E. coli ClpY. A list of sequence motifs characteristic of the different domains, together with database accession numbers for members of the family cloned as of July 1996, can be found in Schirmer et al., 1996; updates appear on Web site: “http://http.bsd.uchicago.edu/~hsplab/index.html”.

HSP100/Clp family. (For a more detailed review of the family, see Schirmer et al., 1996). The distinct subtypes of the HSP100 family characterized to date are depicted in Figure 2. Several members are induced by stress (the B, D, and X subtypes, and some members of the C subtype); others are constitutively expressed. In eukaryotic cells, different members are found in the nuclear/cytosolic compartment(s), in mitochondria, and in chloroplasts. At least four different subtypes (the A, B, X, and Y subtypes, see Figure 2) are found in Escherichia coli (Hwang et al., 1987; Katayama-Fujimura et al., 1987; Squires et al., 1991; Chuang et al., 1993; Gottesman et al., 1993; Wojtkowiak et al., 1993). The ClpA protein of E. coli was the first member of this broad family of proteins to have its biochemical function defined: it is an ATP-regulated subunit of the ClpP protease (Hwang et al., 1987; Katayama-Fujimura et al., 1987). (The name derives from caseinolytic protease). ClpX also functions in proteolysis in conjunction with ClpP (Gottesman et al., 1993; Wojtkowiak et al., 1993). The substrate specificity of ClpP is determined by its association with ClpA or ClpX (Gottesman et al., 1993; Wojtkowiak et al., 1993; Wawrzynow et al., 1995). In the absence of the ClpP protease, ClpX and ClpA alter the conformational states of their substrates. In particular, they promote the disassembly of these substrates from higher ordered structures (Wickner et al., 1994; Wawrzynow et al., 1995). These observations argue that the function of ClpA and ClpX in proteolysis is not simply to determine the substrate specificity of the protease, but also to change the conformational state of the substrate and make it more accessible to digestion by ClpP. The complete sequence of the Saccharomyces cerevisiae genome is known and three members of the HSP100 family are present. Two of these are of the B subtype: Hsp104, found in the nucleus and cytoplasm, and Hsp78, in mitochondria. The third is of the X subtype. This protein may be mitochondrial, but it has not been established. The absence of an obvious ClpP homolog in S. cerevisiae (determined by a homology search of the genome) suggests the yeast ClpX protein may function solely to alter the conformational state of its substrates, rather than to connect them to proteolysis, but its specific functions have not yet been assigned. Members of the HSP100 family have diverse biological functions, but recent work suggests that these functions are unified by a common biochemical mechanism (Schirmer et al., 1996). All of the proteins investigated at the biochemical level to date (including members of the A, B, and X subtypes) employ ATP to promote changes in the


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conformations of other proteins and/or their oligomeric state. In some proteins, this function is coupled to proteolysis, in others to gene regulation, DNA transposition, prion maintenance or stress tolerance. Stress tolerance seems to be a particular specialization of proteins of the B subtype. Bproteins from a very broad range of organisms are induced by stress and function in stress tolerance. These include both B-pro teins of the fungi S. cerevisiae, Hsp104 and Hsp78 (Sanchez and Lindquist, 1990; Schmitt et al., 1996), the B proteins of the bacterium E. coli, ClpB (Squires et al., 1991), of the trypanosomatid Leishmania major, Hsp100 (Hubel et al., 1995), of the land plants Arabidopsis thaliana (Schirmer et al., 1994) and Glycine max (Lee et al., 1994), Hsp101, of the cyanobacterium Synechococcus sp PCC7942, ClpB (Eriksson and Clarke, 1996), and of the closely related C-subtype protein of Bacillus subtilis, ClpC (MecB) (Kruger et al., 1994). It is striking that both Band C-type HSP100s are encoded in the genome of the hyperthermophilic bacterium Aquifex aeolicus, which grows at temperatures up to 95°C (Deckert et al., 1998). Their roles in thermal adaptation, however, have not yet been tested. This review will concentrate on the functions of Hsp104, the only HSP100 B protein whose biological and biochemical functions have been explored in detail. The reader is directed to other recent reviews for a discussion of proteins of other HSP100 subtypes (Gottesman et al., 1995; Schirmer et al., 1996; Maurizi et al., this volume). 2.2. Hsp104 Protein Structure As shown in Figure 2, based on sequence analysis, the Hsp104 protein can be divided into 5 domains. Whether these will correspond to structural domains in the folded protein is currently under investigation. Two of the domains contain highly conserved nucleotide-binding motifs known as the Walker A (P-loop) and Walker B consensus elements. These elements are embedded within larger blocks of highly conserved sequence shared with other members of the family. Although each of these larger domains is itself highly conserved, the two domains have little sequence in common, except for the few residues that constitute the Walker motifs. Thus, the first nucleotide binding domain of Hsp104, NBD1, shares 60% amino-acid identity with the NBD1 of E. coli ClpB and the second nucleotide-binding domain of Hsp104, NBD2, shares 62% identity with the NBD2 of E. coli ClpB, but NBD1 and NBD2 of Hsp104 share only 22% identity. The NBDs of Hsp104 are separated by a middle domain of ~180 amino acids, previously termed the “spacer” region, because of high sequence and size variability in different subfamilies. However, recent analysis indicates that in the ClpB subfamily this region contains conserved sequence motifs (Schirmer et al., 1996). Moreover, dominant Hsp104 mutations cluster in this region (Schirmer and Lindquist, unpublished). These observations suggest that this middle domain has an important biological function. Like the middle region, the N-terminal and C-terminal domains of Hsp104 are less conserved than the NBDs. However, small stretches of common amino acids can be discerned and these are more highly conserved within members of a particular subfamily (Schirmer et al., 1996). Additionally, the C-terminal regions of HSP100 proteins show some similarity to PDZ domains (Levchenko et al., 1997), which function in other


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proteins to mediate protein:: protein interactions. Sequence motifs IV and V (Figure 2) are encompassed within this PDZ-like region. A fragment of the E. coli ClpX protein containing these motifs bound a ClpX proteolytic substrate in an immobilized ELISA assay, indicating this region provides an important function in substrate recognition. It is unclear whether these motifs function in this manner in Hsp104 and other HSP100 proteins, but proteins carrying deletions in the C-terminal domain of Hsp104 do not function in yeast thermotolerance (Schirmer and Lindquist, unpublished). Thus, the Cterminal regions of Hsp104 and of HSP100 proteins in general have important and until recently unappreciated biological functions. 3. BIOCHEMICAL PROPERTIES OF HSP104 3.1. Oligomerization Hsp104 assembles into hexameric ring-shaped particles in the presence of adenine nucleotides, a property it shares with the E. coli ClpA protein (Parsell et al., 1994a; Kessel et al., 1995). In the absence of nucleotides, Hsp104 assembles into hexamers at high protein concentrations (Schirmer, Queitsch, Kowal and Lindquist, unpub-lished). Identical single amino-acid substitutions in consensus nucleotide-binding residues of the two ATP-binding domains have different effects on the ability of Hsp104 to form these particles. In all P-loop containing nucleotide-binding proteins with solved structures, including both ATP- and GTP-binding proteins, a conserved lysine contacts the and phosphates of bound nucleotide (Saraste et al., 1990). A threonine to lysine substitution in this residue of NBD1 (the K218T mutation) has little effect on oligomerization, but the same substitution in NBD2 (K620T) has a strong detrimental effect on oligomerization (Parsell et al., 1994a). 3.2. ATPase Activity Another property that Hsp104 shares with other HSP100 proteins (all tested members, including representatives of the A, B, N, and X subfamilies) is the ability to bind and hydrolyze ATP (Woo et al., 1992; Maurizi et al., 1994; Wawrzynow et al., 1995; Wilson et al., 1995). Hsp104 cleaves ATP with a Vmax of ~2 nmole Pi min-1 g-1 (~20 ATP molecular per hexamer s-1) and binds ATP with a Km of ~5 mM (Schirmer et al., 1998). As with oligomerization, the K218T and K620T mutations have different effects on this activity. The K218T substitution eliminates or severely reduces ATPase activity, while the K620T substitution reduces ATPase activity several fold. However, the ATPase activity of K620T returns to near wild-type levels when the oligomerization defect of the protein is overcome by high protein concentrations (Schirmer, Queitsch, Kowal and Lindquist, unpublished). Thus, it appears that NBD1 is responsible for most of the ATPase activity of the protein, while nucleotide interactions in NBD2 influence that activity by affecting the structural state of the protein (Schirmer et al., 1998). As is the case with several other HSP100 proteins, the ATPase activity of Hsp104 is stimulated by peptides. A few peptides such as -insulin stimulate ClpA, ClpB and Hsp104 (Hwang et al., 1988; Woo et al., 1992; Maurizi et al., 1994; Schirmer and


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Lindquist, unpublished); however most of the peptides which stimulate Hsp104 are different from those that stimulate ClpA (Maurizi et al., 1994; Schirmer and Lindquist, unpublished). The strongest stimulation observed with Hsp104 to date is with polylysine, which increases ATP hydrolysis seven-fold. Interestingly, biologically relevant peptides with very different amino-acid compositions can either stimulate or inhibit ATP hydrolysis by Hsp104 (Schirmer and Lindquist, 1997). One possibility is that different sites for peptide interaction exist in Hsp104, at which peptide binding has different biochemical consequences. 4. THE ROLE OF HSP104 IN STRESS TOLERANCE 4.1. Thermotolerance and the Importance of Hsp104 In all organisms, exposure to mildly elevated temperatures induces tolerance to more severe temperatures (see Li et al., this volume). The magnitude of this effect can be astonishing. In Saccharomyces cerevisiae, for example, a 30 min pretreatment

Figure 3 The role of Hsp104 in stress tolerance.


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(A) Wild-type cells and hsp104 mutants die at similar rapid rates when shifted directly from 25°C to 50°C. (B) Pre-treatments at 37°C for 30 min induce thermotolerance in both mutant and wild-type cells (compare with A), but after a few minutes at 50° C, hsp104 mutants die at ~100 times the rate of wild-type cells. After about 20 minutes at 50°C, the hsp104 mutants die at ~10,000 times the rate of wild-type cells. (C) Mild heat pre-treatments induce tolerance to ethanol but hsp104 mutants die at ~1,000 times the rate of wild-type cells. (D) Mild heat pre-treatments induce tolerance to arsenite, but hsp104 mutants die at similar rates to wild-type cells. (E) Mild heat pre-treatments induce tolerance to cadmium, but hsp104 mutants die at the same rate as wild-type cells. (F) Acetate-grown cells are more tolerant to a direct shift from 25°C to 50°C than glucose grown cells (compare with A), and much of this tolerance is lost in hsp104 mutants. (G) Spores are much more tolerant of heat than log phase cells (compare to A and F), but hsp104 spores are much less tolerant than wild-type spores. Method: Wild-type cells and hsp104 deletion mutants exposed to various stresses for the indicated times were plated on rich glucose media to determine the number of viable cells (colony forming units) remaining. Prior to the stress, cells were grown at 25°C to mid-log phase in glucose medium (A-E), acetate medium (F), or sporulated in low nitrogen medium also at 25°C (G). Data taken from (Sanchez et al., 1992).


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Figure 4 Hsp104 expression is sufficient for tolerance. Cells exposed to 50°C for 4 min were serially diluted, 5-fold at each step, and spotted onto rich glucose agar to determine the number of colony forming units remaining. Left: wild-type cells were maintained at 25°C prior to the heat shock (C) or preheated at 37°C for 30 min (H). Right: hsp104 deletion mutants transformed with plasmids that allow different levels of Hsp104 induction in response to exogenously added hormone (Louvion et al., 1993) were incubated in -estradiol at the indicated concentrations for two hours prior to heat shock. Data taken from (Lindquist and Kim, 1996).

at 37°C increases the ability of cells to survive a severe 50°C heat shock by nearly 10,000-fold (Sanchez and Lindquist, 1990; Sanchez et al., 1992). When log-phase cells carrying hsp104 deletions are grown at 25°C and shifted to 50°C, they die at the same rapid rate as wild-type cells (Figure 3A). When pretreated at 37°C, however, they behave very differently from wild-type. Although the mutants display some tolerance at 50°C, it quickly dissipates. Within 10 min, mutant cells start to die at 100 to 1,000 times the rate of wild-type cells (Figure 3B). Thus, Hsp104 is not the only factor involved in thermotolerance, but it plays a vital role. When total yeast proteins were analyzed by high resolution 2-D gels, no differences could be detected in the accumulation of any proteins, other than Hsp104 itself, either at high or low temperatures (Parsell et al., 1993). This strongly suggests that Hsp104 affects thermotolerance directly, rather than by altering the expression of other tolerance factors. Hsp104’s role in thermotolerance is specific to extreme conditions. Hsp104 deletions grow as well as wild-type cells, not only at normal temperatures, but even at the highest temperatures in the yeast growth range (Sanchez and Lindquist, 1990; Sanchez et al., 1992). They also survive as well as wild-type cells when held for two days at


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temperatures just beyond the growth range (Sanchez and Lindquist, 1990). At 44°C, where cells take many hours to die, mutant cells die at 10 times the rate of wild-type cells (Sanchez et al., 1992). At 55°C, where cells survive for only a few minutes, hsp104 mutants die at 1,000 to 10,000 times the rate of wild-type cells (Blake and Linquist, unpublished). Such experiments demonstrate that Hsp104 is important in stress tolerance, but give little indication of how central its role might be. Disrupting just one element in a web of 50 equally vital factors could produce a severe reduction in survival. The true importance of Hsp104 became apparent in experiments where its expression was divorced from that of other tolerance factors, by placing HSP104 coding sequences under the control of heterologous promoters (Lindquist and Kim, 1996). When Hsp104 is selectively induced to a modest level it increases survival several fold. More remarkably, when it is expressed at a high level, Hsp104 provides as much tolerance as a mild heat pretreatment, at least for a short severe heat shock (Figure 4 and Lindquist and Kim, 1996). 4.2. The Thermotolerance Function of Hsp 104 4.2.1. No Evidence for a Proteolysis Function The homology between Hsp 104 and ClpA, with its known proteolysis function, suggested that Hsp 104 might provide thermotolerance by promoting the proteolysis of irretrievably heat-denatured proteins. That is, it might increase survival by preventing protein aggregates from ‘clogging up’ the cell. Experiments using previously established assays for detecting proteolytic deficits in yeast suggest this is not the case. Hsp104 deletion mutants 1) release previously incorporated amino acids into the media at the same rate as wild-type cells, 2) show similar rates of -galactosidase digestion, using the unstable N-end rule variants engineered by the Varshavsky laboratory, 3) grow on plates containing amino-acid analogs at least as well as wild-type cells, and 4) show no detectable accumulation of novel proteins by high resolution 2-D gels (Parsell et al., 1993; Parsell and Lindquist, unpublished; Taulien and Lindquist, unpublished). Although negative evidence is never conclusive, it seems very unlikely that a 100- to 1,000-fold difference in survival after heat shock could be based on a level of proteolysis that is undetectable by these methods. 4.2.2. A Disassembling/Resolubilizing Activity Rather than promoting proteolysis, Hsp 104 seems to function in stress tolerance by promoting the resolubilization and reactivation of proteins that have been denatured by heat and have begun to aggregate. Such a function was unexpected; it was generally believed that aggregated proteins are irretrievably lost. As surprising as such a function for Hsp104 may be, it is now supported by several different lines of evidence. One line of evidence involves in vivo analysis of a specific heat-damaged substrate. The critical heat-sensitive lethal targets in yeast are unknown. However, a heterologous temperature-sensitive luciferase protein provides a model heat-sensitive substrate that denatures at temperatures which do not affect viability, even in the hsp104 mutants. Luciferase activity is lost at the same rate in mutant and wild-type cells, and in both cell


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types the enzyme is found in an aggregated state. Remarkably, when cells are returned to normal temperatures, luciferase is resolubilized and reactivated in wild-type cells, but not in cells lacking Hsp104 (Parsell et al., 1994b). Another line of evidence involves direct visualization of heat-induced damage by electron microscopy. It takes several hours to kill yeast cells at 44°C. However, after just one hour, when both wild-type and hsp104 mutant cells retain greater than 95% viability, extensive damage is already apparent. Wild-type and hsp104 mutant cells are indistinguishable: both display large masses of electron-dense material (presumably protein aggregates) in their nuclei and cytoplasms (Parsell et al., 1994b). During two hours of recovery at normal temperatures, this material completely disappears from wildtype cells, but not from the mutants. Since there appear to be no differences in proteolysis in wild-type and mutant cells, the disappearance of these aggregates is most likely due to resolubilization. A third line of evidence involves RNA splicing, one of the most heat-sensitive cellular processes known (Yost and Lindquist, 1986; Yost and Lindquist, 1988). When whole cells are shifted directly to moderately high temperatures, splicing is disrupted while transcription and translation continue. Splicing is equally affected in wild-type and mutant cells. Moreover, mild pretreatments protect splicing from disruption as effectively in mutant cells as in the wild-type cells (Yost and Lindquist, 1991). However, when splicing is disrupted by a short, sub-lethal heat shock and cells are returned to normal temperatures for recovery, it is restored much more rapidly in wild-type than in the mutant cells. Thus, Hsp104 does not protect splicing from disruption, but helps to reactivate splicing once it has been disrupted. In vitro experiments also indicate Hsp104’s function is to repair damage rather than prevent it. When splicing-competent cell-free extracts are heated, they are rapidly inactivated. If Hsp104 is present during the heat treatment, it does not prevent inactivation. However, if purified Hsp104 is added after inactivation, it at least partially restores splicing (Vogel et al., 1995). Hsp104 also has no capacity to prevent the aggregation of chemically denatured enzymes upon dilution into aqueous buffers. Nor does it promote the refolding of these substrates directly. Rather, Hsp104 helps Ydj1 and Hsp70 promote the activation of these enzymes by an ATP-dependent mechanism (Glover and Lindquist, 1998). Taken together, these data indicate that the primary function of Hsp104 is to mediate the repair of substrates damaged by heat, rather than to protect those substrates from damage. Heat is a protein denaturant. Thus, Hsp104’s molecular function provides an explanation of its biological role in protecting cells from exposure to extreme heat— when the rate of protein denaturation outpaces the ability of other protective systems to prevent aggregation. The biochemical properties of Hsp104 described earlier are clearly related to these functions. The single-amino acid substitutions in NBD1 and NBD2 that inhibit ATP hydrolysis (Schirmer et al., 1998) or oligomerization (Parsell et al., 1994a) do not promote the resolubilization of heat-damaged luciferase in vitro (Glover and Lindquist, 1998) or in vivo (Parsell et al., 1994b), and are ineffective in promoting survival after a severe heat shock (Parsell et al., 1991).


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4.3. Other Stresses A striking and universal feature of stress tolerance is that one type of mild stress induces tolerance to many others. Thus, mild heat treatments induce tolerance not only to higher temperatures, but also to ethanol, anoxia, heavy metal ions, free radicals, UV light, and a host of other damage-causing agents. (See Watson, 1990; Nover, 1991; Weber, 1992; Parsell and Lindquist, 1993 for review.) For many years it was not clear whether the same or different mechanisms were responsible for tolerance to all of these stresses. Genetic analysis of Hsp104 provided one of the first tools for dissecting this problem. As shown in Figure 1, Hsp104 is induced by a wide variety of stressful conditions. It is one of the most prominent proteins induced by heat (Figure 1A) and is induced to a similar level by ethanol, sodium arsenite, and cadmium (Figure 1B). Hsp104 is just as important in protecting cells from ethanol as from heat (Figure 3C and Sanchez et al., 1992). Indeed these stresses exhibit complete cross tolerance: mild heat pretreatments protect against damage by ethanol, mild ethanol pretreatments protect against heat shock (Sanchez et al., 1992). Moreover, as with heat, the importance of Hsp104 in ethanol tolerance is greater under more severe conditions. It has little effect on survival when cells are incubated with 12% ethanol, but promotes survival by 10,000-fold when they are incubated with 20% ethanol. The expression of Hsp104 gives cells a modest survival advantage upon exposure to sodium arsenite, but this is apparent only with arsenite pretreatments and not with heat pretreatments (Figure 3D and Sanchez et al., 1992). Moreover, Hsp104 provides no advantage upon exposure to cadmium (Figure 3E) or copper, over a broad range of toxic concentrations and, in some cases, actually reduces survival (Sanchez et al., 1992, and Taulien and Lindquist, unpublished), whether Hsp104 is pre-induced by heat (Figure 3E), or by a low concentration of the metal ion itself (Sanchez et al., 1992). In the light of what we now know about Hsp104’s molecular functions and the nature of cadmium and copper toxicity, this makes perfect sense. Heat and ethanol are protein denaturants, and Hsp104 helps to restore denatured proteins to the native state (Parsell et al., 1994b). Cadmium and copper, however, cause covalent oxidative damage that requires degradation of the damaged protein (Stohs and Bagchi, 1995). Therefore, Hsp104 is not involved. Curiously, although Hsp104 is induced by DNA damaging agents, such as UV and MMS, it consistently reduces the rate of survival during exposure to radiation (Boreham and Mitchel, 1994)—that is, hsp104 mutant cells survive better than wild-type cells. The molecular basis of this phenomenon is unknown. The complex and overlapping stressresponse networks of the cell induce a broad range of proteins in response to many stresses (Ruis and Schuller, 1995) but, clearly, some proteins are not always beneficial to the particular stress encountered. 4.4. Combinatorial Stresses The fact that the functions of Hsp104 in stress tolerance are most important under extreme conditions calls into question its general relevance. It seems unlikely that yeast cells will commonly find themselves exposed to 50°C in their natural environment.


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However, it seems equally unlikely that cells will encounter a single type of stress while all other conditions remain optimal. Single stresses reduce the number of variables in laboratory experiments, but they are hardly a realistic mimic of natural conditions. The synergistic effects of combined stresses is apparent in cells exposed to two stresses which themselves cause no loss in viability, 6% ethanol and 40°C for 2 hours. In combination, these two stresses kill 99.99% of the cells, unless the cells have been given a mild pretreatment and contain a wild-type copy of the HSP104 gene (Lindquist et al., 1995). In that case, most of the cells survive. Thus, Hsp104 is a vital tolerance factor for lethal combined stresses, a situation more likely to be encountered in the natural environment. 4.5. Physiological Inductions of Tolerance In addition to stress, Hsp104 is induced in response to certain normal conditions of growth and development. For example, it is expressed at a high basal level when cells are grown on carbon sources that require high rates of respiration, such as acetate (Figure 1C). Such cells survive a direct shift from 25°C to 50°C much better than glucose-grown cells (Sanchez et al., 1992) and this basal tolerance is greatly reduced in hsp104 mutants (Figure 3F). Because yeast cells normally switch into respiratory metabolism when their preferred carbon source, glucose, has been fermented to ethanol, this induction may be part of a developmentally regulated program to ensure that Hsp104 is present to protect against ethanol toxicity. The protein is also induced in glucose-grown cells a few generations before stationary phase, at the time of the diauxic shift. Stationary phase cells exhibit a much higher level of stress tolerance than log-phase cells and, again, this is largely dependent upon Hsp104. Finally, Hsp104 is strongly induced during sporulation and is present in mature spores at a high concentration (Figure 1C). Yeast spores employ many mechanisms for surviving hostile environments. (Note the difference in time scale on the killing curves of Figures 3B and 3G.) Nevertheless, they remain dependent upon Hsp104. Once spores begin to die, hsp104 mutants die at ~200 times the rate of wild-type cells (Figure 3G). Long-term viability of spores stored at low temperatures also depends on Hsp104 (Sanchez et al., 1992). Apparently, sufficient structural fluctuations take place in the proteins of these dormant cells to constitute a threat to long-term survival. It is not clear whether the maintenance of viability in spores involves the continuous action of Hsp104 or a delayed repair function initiated on germination. 4.6. Conservation of Tolerance Functions in B-Type Proteins The tolerance functions of the B- and closely related C-type HSP100 proteins have been conserved throughout evolution. Mutations in the E. coli B protein compromise survival at high temperatures (Squires et al., 1991; see Burkholder and Gottesman, this volume); mutations in the B. subtilis C protein reduce survival at high temperatures, high concentrations of ethanol, and high salt (Kruger et al., 1994); and mutations in the cyanobacterial B protein reduce viability roughly 5-fold and severely reduce the ability of cells to remain photosynthetically active after heat shock (Eriksson and Clarke, 1996). The related cyanobacterial C protein is essential for growth and is induced by high light intensities or CO2 concentrations (Clarke and Eriksson, 1996). Although it is not


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normally induced by heat, when the B gene is defective, the C protein becomes inducible, presumably fulfilling stress tolerance functions of the absent B protein (Eriksson and Clarke, 1996). Except for Hsp104 and, to a lesser degree, ClpB from E. coli, the biochemical functions of the B-type proteins have not been investigated. Given their sequence conservation and common roles in stress tolerance, it seems very likely that all B-type proteins function through a conserved molecular mechanism. Indeed, the Hsp104 homologs of vascular land plants, organisms separated from yeast by nearly a billion years of evolution, are able to partially restore thermotolerance to hsp104 mutants. The Arabidopsis protein increases survival ~100-fold (Schirmer et al., 1994) and the soybean protein ~25-fold (Lee et al., 1994). The ability of plant Hsp104 homologs to function in yeast thermotolerance strongly suggests they have a thermotolerance function in their native organisms, and preliminary data from Arabidopsis strains with altered HSP101 levels support this suggestion (Queitsch, Vierling and Lindquist, unpublished). In contrast, the E. coli B protein provides no thermotolerance in hsp104 mutants (Parsell et al., 1993). Assuming that the E. coli protein’s function in stress tolerance is similar, this observation indicates that the functions of HSP100 proteins are not autonomous, but require cooperation with other factors. In vitro studies (Section 4.2.2) suggest that these cooperating factors include the chaperones Ydj1 and Hsp70. Presumably, the Arabidopsis and soybean proteins are still capable of interacting with helper factors, but do so less effectively than Hsp104, while the E. coli protein does not interact with them productively. In eukaryotes, many Hsps have homologs in different cellular compartments, where they serve similar functions (see chapter by Bukau et al., this volume). The complete genomic sequence of S. cerevisiae indicates there is no Hsp104 homolog in the endoplasmic reticulum. There is, however, a B-type HSP100 protein in yeast mitochondria (Hsp78), and recent work suggests it functions in stress tolerance in that organelle. hsp78 mutations themselves have little phenotype (Leonhardt et al., 1993), but lead to defects in mitochondrial protein import and DNA maintenance when combined with certain Hsp70 mutations (Moczko et al., 1995; Schmitt et al., 1995). However, the recovery of mitochondrial protein synthesis after heat stress depends on Hsp78. This appears to involve a repair function rather than a protective function because Hsp78 does not prevent inactivation of mitochondrial protein synthesis by heat (Schmitt et al., 1996). When Hsp78 is expressed in the cytosol it can partially restore thermotolerance in hsp 104 deletion mutants (Schmitt et al., 1996). Thus far, three human relatives of Hsp 104 have been cloned, one type X, one type M, and one fragment that is too small to assign. The smaller human fragment is more homologous to class 1 HSP100 proteins than to class 2 proteins; however it is too early to predict whether a B-type Hsp 104 homolog exists in humans. Oddly, a 100 kd heatinducible hamster protein that cross reacts with an antibody directed against a C-terminal peptide of yeast Hsp104 (Parsell et al., 1991) is actually an unusual variant of the Hsp70 protein family, with no sequence homology to Hsp104 except for a few amino acids at the extreme C-terminus. In any case, the HSP100/Clp B-type proteins are clearly relevant to human biology and medicine. Many human parasites mount a strong heat-shock response when they transfer


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from other environments to their human hosts and the induction of HSP100 proteins is often particularly intense. A deletion mutant of the Hsp104 homolog of Leischmania is less virulent than wild-type Leischmania cells (Hubel et al., 1997). This extends B-type protein functions in thermotolerance to yet another evolutionarily distinct group of organisms. If Homo sapiens should prove not to have a B-type HSP100 homolog, then this protein may prove a useful target for pharmaceutical intervention against human pathogens. 4.7. The Relationship Between Hsp104 and Other Tolerance Factors 4.7.1. Other Tolerance Factors Although Hsp104 plays a central role in stress tolerance, it is by no means the only factor involved. Other known tolerance mechanisms include the induction of osmolytes to alter the solvent properties of the fluid phase, the alteration of lipid composition to change the fluidity of membranes, the synthesis of metal-chelating agents and free-radical scavengers to sequester or eliminate toxins, and the induction of other Hsps (Parsell and Lindquist, 1993; Ruis and Schuller, 1995). The induction of other Hsps itself represents several different survival mechanisms, as each protein has a distinct biochemical function. Hsps do, however, have a common functional theme: in one way or another they help the cell cope with protein damage. The best-studied Hsps bind unfolded proteins and prevent them from aggregating (e.g., Hsp70, Hsp60, Hsp40) or help clear the cell of damaged proteins by ushering them along the degradation pathway (e.g., ubiquitin and certain ubiquitinconjugating enzymes; see Maurizi et al., this volume). The relationship between these different tolerance mechanisms and that of Hsp104 is just beginning to be explored, but in two specific cases informative stories are emerging. 4.7.2. Genetic Interactions Between Hsp70 and Hsp104 In many organisms, heat-inducible members of the HSP70 family play an important role in thermotolerance (Parsell and Lindquist, 1993; Schröder et al., 1993). Surprisingly, in S. cerevisiae, hsp70 mutations have little effect on thermotolerance (Craig and Jacobsen, 1984). The interpretation of this observation, however, must be tempered by the complexity of the problem. S. cerevisiae express four closely related HSP70 proteins in the cytosolic/nuclear compartment, the Ssa proteins, which form a single complementation unit (see Craig et al., this volume). Deletion of all four proteins is lethal. Deletion of a subset produces a strong temperaturerelated phenotype—failure to grow at the upper end of the normal temperature range—but does not impair the ability of cells to survive a short severe heat shock. In fact, these deletions increase survival compared to wild-type when cells are shifted directly from 25°C to 50°C (Craig and Jacobsen, 1985). This increase in basal thermotolerance is due to Hsp70’s role in regulating other stress-response proteins. Several Hsps, including Hsp104 and trehalose synthase (whose significance is explained below) are overexpressed in ssa mutants at normal temperatures (Hottiger et al., 1992; Sanchez et al., 1993). The essential character of the ssa proteins, and the effects of hsp70 mutants on the expression of other proteins, makes it difficult to unambiguously determine the role of Hsp70 in yeast


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thermotolerance. Two other experiments, however, suggest Hsp70 functions complement those of Hsp104. First, as described earlier, when Hsp104 mutants are given a conditioning pretreatment and then shifted to 50°C, they initially exhibit some tolerance. This early tolerance is reduced when cells also carry mutations in heat-inducible ssa proteins (Sanchez et al., 1993). More compellingly, overexpression of Hsp70 partially restores thermotolerance in cells carrying hsp104 mutations (Sanchez et al., 1993). Thus, in the absence of Hsp104, Hsp70 clearly plays a role in thermotolerance. In a complementary fashion, when Hsp70 function is compromised by mutation, Hsp104 becomes important for normal growth (Sanchez et al., 1993). Cells carrying mutations in certain ssa proteins do not grow at high temperatures, but grow quite well at 25°C (Craig and Jacobsen, 1984). However, if an hsp104 deletion, which itself has no effect on growth, is introduced into these cells, their growth rate is severely reduced (Sanchez et al., 1993). The genetic complementarity between Hsp70 and Hsp104 reflects an underlying biochemical complementarity. Although Hsp70 can resolubilize certain special types of protein aggregates (Skowyra et al., 1990), it generally does not (Schröder et al., 1993; Wickner et al., 1994). Rather, it helps proteins that are already in an unfolded, or partially unfolded state, to remain soluble. As described elsewhere in this volume (Buchberger et al.; Ha et al.), Hsp70 accomplishes this by binding to small, hydrophobic, aggregationprone stretches of amino acids that would normally be buried in the hydrophobic core of a protein. Hsp104 does not prevent protein aggregation, but helps resolubilize aggregates once they occur. Presumably, when Hsp70 function is compromised, some aggregation occurs even at 25 °C and Hsp104 then becomes essential for normal growth. (Indeed electron microscopy of Ssa mutants reveals a low level of electron dense material, indicative of protein damage, at normal temperatures and this damage is exacerbated by hsp104 deletions; Lindquist et al., 1995). Conversely, when Hsp104 is not present to repair damaged proteins after heat shock, overexpression of Hsp70 compensates by reducing the rate at which protein aggregation occurs. While increased expression of Hsps is beneficial at high temperatures, it can be detrimental to growth at normal temperatures (Feder et al., 1992). The need to balance the beneficial and detrimental effects of these proteins in the context of an individual cell’s particular physiology is believed to account for differences in the patterns of Hsp synthesis observed in different organisms. A case in point is the absence of a stressinduced Hsp104 homolog in Drosophila cells. In this organism, the Hsp70 genes have been amplified, with five identical copies per haploid genome. Thus, yeast cells in which Hsp70 overexpression partially compensates for the loss of Hsp 104 in thermotolerance, mimic what appears to be a naturally-occuring adaptation of Drosophila. Only when we gain a more complete understanding of the beneficial and detrimental effects of Hsps will we understand the manner in which the expression of these proteins has changed in the course of evolution. 4.7.3. Genetic Interactions Between Trehalose Synthase and Hsp104 The induction of osmolytes is another common tolerance mechanism. Different


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organisms produce different osmolytes in response to stress. For example, E. coli, S. cerevisae, and Artemia selina (brine shrimp) produce trehalose, whereas certain vascular plants produce sucrose, and mammalian kidney cells produce sorbitol and other solutes (Yancey et al., 1982; Wiemken, 1990; Somero et al., 1992). These compounds accumulate to extraordinary levels, both in response to stress and as part of the natural developmental adaptations for stress tolerance. For example, in yeast trehalose accumulates up to 0.5 M in response to heat shock (Hottiger et al., 1994), and in some fungi may accumulate to 15% of the dry weight of the cell during sporulation (Van Laere, 1989). Trehalose is the only osmolyte whose function in stress tolerance has been investigated genetically. In yeast, trehalose synthase mutants die more rapidly than wild-type cells during exposure to extreme heat (De Virgilio et al., 1994). One study suggests that, in tobacco, drought tolerance can be promoted by trehalose via expression of trehalose synthase (Holmstrom et al., 1996). By changing the properties of the fluid phase, trehalose preserves protein and membrane structure at high temperatures (Crowe et al., 1984; Colaco et al., 1992; Hottiger et al., 1994). Other osmolytes function in a similar manner in vitro and are likely to function in a similar manner in vivo (Yancey et al., 1982) To determine how protection afforded by the production of trehalose relates to that afforded by Hsp104, double mutants were examined. Each mutation alone reduced survival in conditioned cells on the order of 100-fold. In the double mutant, tolerance was reduced by 10,000-fold (Elliott et al., 1996; and Singer and Lindquist, 1998). Thus the genetic data confirm biochemical expectations. These two mechanisms operate on different critical points, trehalose reducing the likelihood that proteins will be denatured by heat, and Hsp104 acting to repair denaturation damage once it occurs. Together, they appear to provide the primary mechanisms by which yeast cells survive sudden exposure to extreme conditions. 5. HSP104’S INTERACTION WITH A YEAST PRION 5.1. A Role for Hsp 104 During Normal Growth An unexpected role for Hsp 104 in yeast cells at normal temperatures was discovered in a screen for factors that reduce nonsense suppression in strains carrying the omnipotent suppressor known as [PSI + ] (Chernoff et al., 1995). [PSI + ] proved to be a remarkable new type of genetic element composed only of protein, in which a heritable change in phenotype derives from a self-perpetuating change in protein structure. Thus, [PSI + ] is conceptually related to the remarkable infectious agents in mammals known as prions. The role of Hsp 104 in the propagation of this novel genetic element becomes intelligible in the context of its function in altering protein conformations. 5.2. The Prion Hypothesis Prions are unconventional infectious agents responsible for several devastating, invariably fatal neurodegenerative diseases, commonly termed the transmissible


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spongiform encephalopathies (TSEs) (Prusiner, 1994; Prusiner and DeArmond, 1994; Weissman, 1994; Gajdusek, 1996; Prusiner, 1996). In humans these include CreutzfeldtJacob disease, Gerstmann-Straussler-Schienker syndrome, fatal familial insomnia and kuru (most famous for its transmission by the cannibalistic eating of infected human brains). In sheep, which were early subjects for experimental investigation, it is commonly known as scrapie; in cattle it is commonly known as “mad cow” disease. Although there are some differences in presentation, the TSEs are all characterized by dementia, motor neuron dysfunction, and an unusual “spongy” histology of brain tissue brought about by extensive vesiculation without lymphocytic infiltration. The devastating economic impact of “mad cow” disease in Great Britain, and the frightening prospect that a new, more virulent form of the disease is now appearing in humans (perhaps transmitted through contaminated beef products; Collinge et al., 1995) have heightened the urgency of the quest for effective medical intervention. Unlike all other known pathogens, prions are believed to consist entirely of protein— specifically, a normal cellular plasma-membrane protein that has undergone a change in conformation from its usual protease sensitive state, PrPC, to a pathogenic, proteaseresistant “scrapie” form, PrPSc. Pathogenicity is believed to derive from the ability of proteins in the PrPSc conformation to interact with normal PrP molecules and induce them to adopt the PrPSc structure. That is, a protein-conformational chain reaction both propagates the disease and generates new infectious material (Prusiner, 1994; Lansbury and Caughey, 1995). As the disease progresses, new infectious material is generated and PrPSc conformers are deposited in the brain and in certain peripheral tissues as dense, detergent-insoluble complexes. The TSEs are also unusual in that they can arise either from sporadic incidents, from the inheritance of mutated PrP proteins, or from infection. Rare sporadic cases may derive from rare, spontaneous misfolding events that produce PrPSc conformers. Inheritance is associated with mutations in PrP that are thought to destabilize the normal folded state, increasing the likelihood of obtaining the PrPSc conformation. Both the sporadic and the heritable diseases give rise to infectious prion particles that contain the same protein, PrPSc. Not surprisingly, the notion that an infectious agent can consist entirely of protein, with no associated nucleic acid, was greeted initially with great skepticism. Recently, however, the hypothesis has gained critical support from many different types of investigation, including biochemical analyses of the infectious material, in vitro conversions of PrPC into PrPSc-like conformers, and genetic analysis of transgenic mice. The prion hypothesis has now gained broad, though by no means universal, acceptance. Recent work in yeast suggests prions may be more common than initially envisioned. For nearly 30 years yeast geneticists have been tracing the inheritance of two mysterious cytoplasmic elements known as [PSI + ] and [URE3]. In biological terms [PSI + ] and [UEE3] are unrelated. [PSI + ] is a factor that reduces the fidelity of ribosome termination during protein synthesis, leading to the suppression of nonsense codons by ribosome read-through (Cox et al., 1988). [URE3] allows cells that are growing in the presence of ammonia to use additional nitrogen sources (Wickner et al., 1995). However, [PSI + ] and [URE3], share several unusual patterns of genetic behavior. Moreover, repeated attempts to link their dominant, cytoplasmic inheritance to any cytoplasmically


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inherited nucleic acid have failed. Four years ago, Reed Wickner made the remarkable suggestion that the puzzling behavior of these elements could be explained through a prion-like model of protein conformation-based heredity (Wickner, 1994). In the short time since that suggestion, several lines of evidence have provided support—one of the most convincing being the central role played by Hsp104 in the inheritance of [PSI + ]. Because a role for Hsp104 has not been established for [URE3], it will not be considered further here. The reader is referred to Wickner’s excellent recent review for further information on this element (Wickner et al., 1995). 5.3. The Relationship Between [PSI + ] and Sup35 [PSI + ] is a dominant, cytoplasmically inherited trait. It is postulated to represent the inheritance of conformationally altered Sup35 protein. Sup35 in its normal conformation is a subunit (together with Sup45) of the translation-release factor that allows ribosomes to recognize termination signals (Stansfield et al., 1995). Just as mutations in PrP can produce heritable spongiform encephalopathies (Prusiner, 1994; Prusiner and DeArmond, 1994), mutations in Sup35 can suppress nonsense mutations by ribosome read-through (Tuite and Stansfield, 1994; Stansfield et al., 1995). Of course, [PSI + ] can not be due to a mutation in Sup35, because such mutations exhibit normal Mendelian inheritance (Tuite and Stansfield, 1994). That is, when a sup35 mutant is mated to a wild-type cell and the diploid is sporulated, the defect in translation termination is found in half of the haploid progeny. When a [PSI + ] cell is mated to a [psi - ] cell, the termination defect is observed in all of the progeny (Cox et al., 1988; Cox, 1994). Remarkably, overexpression of wild-type Sup35 can induce not only the [PSI + ] phenotype, but also new, heritable [PSI + ] elements. Even more remarkably, only transient overexpression of Sup35 is required. Once cells that overexpress Sup35 have converted to [PSI + ], this state is maintained in all of its progeny, even when the plasmid that confers overexpression is lost (Chernoff et al., 1993; Wickner, 1994). The gene encoding Sup35 has an unusual structure (Figure 5). The C-terminal domain encodes the translation-release factor and is essential. The N-terminal domain has an unusual amino acid composition and is not essential (Ter-Avanesyan et al., 1993). Overexpression of just the N-terminal domain is sufficient to convert a cell to [PSI + ]. Deletion of the N-terminal domain prevents cells from ever acquiring the [PSI + ] phenotype. Moreover, it causes the loss of [PSI + ] in cells that have already acquired it (Chernoff et al., 1993; Ter-Avanesyan et al., 1993). Physical analysis of the Sup35 protein demonstrates that it has a different conformational state in [PSI + ] and [psi - ] cells: 1) it pellets at 12,000×g in [PSI + ] cells, while it is mostly soluble in [psi - ] cells, 2) it is protease resistant in [PSI + ] cells while it is protease sensitive in [psi - ] cells, 3) newly synthesized proteins containing the


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Figure 5 Sup35 domain structure and its relationship to [PSI + ]. A. Domain N has an unusual amino acid composition and contains several repeats ‘PQGGYQQYN’ that are similar to repeats found in the mammalian prion protein ‘PHGGGWGQ’. Domain M is highly charged and domain C, which is sufficient for Sup35’s translation termination function, contains four potential GTP-binding sites. A correlation was observed between the fiber forming propensity of domains and their ability to induce [PSI + ]. In cases where domains did not precisely match, the amino acid numbers are given in parentheses. B. The effects of deleting or overexpressing the NM and the C domains are shown. (Data from Chernoff et al., 1993, Ter-Avanesyan et al., 1993, Derkatch et al., 1996, and Glover et al., 1997.)

N-terminal region of Sup35 are rapidly sequestered into large particles in [PSI + ] cells, but remain freely distributed in [psi - ] cells (Patino et al., 1996; Paushkin et al., 1996). These data are consistent with the hypothesis depicted in Figure 6. The N-terminal domain of Sup35 converts cells from [psi - ] to [PSI + ] by promoting a conformational change that interferes with the ability of the C-terminal domain to function in translation termination, perhaps simply sequestering it in an aggregate that makes it inaccessible to ribosomes. The reduced concentration of functional termination factor leads to the readthrough of nonsense codons. The conformational alteration of Sup35, once achieved, is self-sustaining (with the help of Hsp104, see below). Therefore, it behaves as a dominant, cytosolically-inherited genetic element. 5.4. The Role of Hsp104 in the Maintenance of [PSI+] 5.4.1. Overexpression of HSP104 Cures [PSI + ] The relationship between Hsp104 and [PSI + ] was discovered by Yury Chernoff, who screened a yeast plasmid library for factors that would inhibit the [PSI + ] phenotype. The only plasmid uncovered in this initial screen proved to encode Hsp104 (Chernoff et al., 1995). Several criteria were used to demonstrate that Hsp104 does not affect translation directly, but simply eliminates the [PSI + ] element (Chernoff et al., 1995). Remarkably,


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just as the transient overexpression of the N-terminal domain of Sup35 can cause the appearance of new, heritable [PSI + ] elements, the transient overexpression of Hsp104 causes the heritable loss of [PSI + ] (Chernoff et al., 1995;

Figure 6 Model of Sup35 sequestration effects in [PSI + ] cells. Sup35 is one subunit of the eukaryotic translation termination factor. The other is Sup45. In [PSI + ] cells, most Sup35 is found in a self-perpetuating, conformationally altered state (here depicted as an ordered aggregate) that prevents it from functioning in termination. The low level of functional Sup35 reduces the fidelity of protein synthesis, causing some ribosomes to read through nonsense codons. The presence of [PSI + ] is detected by the ability of cells carrying nonsense mutations in auxotrophic markers to grow without otherwise essential nutrients in the media.

Patino et al., 1996; Paushkin et al., 1996). When Hsp104 is placed under the control of a galactose-inducible promoter, a few hours of galactose induction is sufficient to eliminate [PSI + ]: 95% of the cells become [psi-] and pass this trait on to their progeny. The ability of a chaperone protein to cure cells of [PSI + ] strongly supports the hypothesis that [PSI + ] represents a heritable, altered conformation of Sup35. Once this conformation is eliminated, the strain no longer propagates [PSI + ]. 5.4.2. Deletion of HSP104 Cures [PSI + ] In most cases, when overexpession of a protein produces one phenotype, reduced expression produces the opposite. That is, deletion of Hsp104 might be expected to increase the stability of [PSI + ]. This is not the case. Deletion of Hsp104 cures cells of [PSI + ]. In fact, deletion of Hsp104 produces a psi-no-more (PNM) phenotype. When cells carrying an hsp104 deletion are mated to [PSI + ] cells, the diploid is [PSI + ]. However, when that diploid is sporulated, only half of the progeny, those that carry the wild-type HSP104 gene, remain [PSI + ] (Chernoff et al., 1995). 5.4.3. Hsp104 and Sup35 Aggregation in Vivo The unexpectedly similar effects of overexpressing and underexpressing Hsp104 on the inheritance of [PSI + ] provided an opportunity to test the relationship between [PSI + ]


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and the altered, aggregated state of Sup35. If this altered state is responsible for the [PSI + ] phenotype, the aggregates should disappear when [PSI + ] is cured either by overexpression of Hsp104, or by deletion of Hsp104. Note that this is counter to expectations derived from the effects of Hsp104 on heat-induced aggregates: deletion of HSP104 causes proteins damaged by heat to remain in an aggregated state. The conformational state of Sup35 was examined in two ways, by differential sedimentation analysis of yeast lysates and by cytological visualization of Sup35 aggregates with Green Fluorescent Protein (Patino et al., 1996). Results of both analyses agreed. When cells are cured of [PSI + ], Sup35 returns to the soluble state, whether that cure is affected by overexpression of HSP104 or by deletion of HSP104. The effects of point mutations in consensus residues of the Hsp104 ATP-binding domains are also telling. When cells carrying a wild-type copy of HSP104 are transformed with plasmids encoding the lysine to threonine substitutions in NBD1 and NBD2 described in Section 3 (K218T, K620T), the [PSI+] phenotype disappears. Because cells are cured of [PSI + ] by either overexpression of Hsp104 or deletion of Hsp104, it is not possible to determine from these experiments whether these plasmids eliminate the [PSI+] phenotype by increasing Hsp104 function or by interfering with Hsp104 function. The effects of these mutations do, however, provide another means of testing the relationship between [PSI + ] and Sup35 aggregation. Remarkably, the K218T or K620T plasmids eliminate the [PSI + ] phenotype only as long as they are present in the cell. When the plasmids are lost by growth in non-selective media, [PSI + ] reappears. [PSI + ] must still be present in these cells, but somehow translation termination remains normal. Analysis of the Sup35 protein in these cells explains this cryptic behaviour. In cells carrying the individual substitutions, aggregates (intense fluorescent foci) of a Sup35-GFP fusion protein are still present, but a substantial fraction of Sup35 is diffusely distributed and soluble. Apparently, the soluble protein functions in translation termination, eliminating the [PSI + ] phenotype, while the aggregates promote the condensation of most newly synthesized Sup35 protein, restoring [PSI + ] when the plasmids are lost. These observations confirm that the aggregated state of Sup35 is intimately related to the inheritance of [PSI + ]. They also demonstrate that the relationship between Hsp104 and Sup35 aggregates is different from that which exists between Hsp104 and heatdamaged protein aggregates. Hsp104 is actually required for the formation of Sup35 aggregates. 5.4.4. Seeded Polymerization of Sup35 Recent biochemical investigations of Sup35 confirm that it can form aggregates with very special properties. When purified Sup35, produced in E. coli, is denatured in urea and diluted into aqueous buffer, it forms highly ordered, amyloid-like fibrils that bind Congo Red dye (Glover et al., 1997). These have a central, rod-like core of 10–11 nm, with more amorphous material splayed out on the sides. The C-terminal region (see Figure 5) and the M region show no capacity for fiber formation, but the N-terminal domain does, forming fibers that are short and 8 to 9 nm in diameter. Fibers formed by the N-terminal domain appear to be rich in -sheet structure, as are the fibrils formed by PrP and other


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amyloids (Glover et al., 1997; King et al., 1997). The N and M regions together form long smooth fibers that look very much like the central core of the fibers formed by whole Sup35. Thus it would appear that fiber formation is directed by the glutamine and tyrosine-rich N domain,

Figure 7 Model of Hsp104’s function in [PSI+ ] conversion. Here, [PSI+ ] (right), is depicted as a special, ordered aggregate of Sup35 (left). (1) and (2) Hsp104 produces a conformational change in Sup35, either by unfolding Sup35 monomers, or by dissociating Sup35::Sup45 dimers. (3) The conformationally altered state of Sup35 produced by Hsp104 is unstable. In the absence of pre-existing [PSI + ] elements, the protein reverts to its normal, functional state. (4) When [PSI + ] elements are present, they capture the unstable conformers of Sup35. Presumably these elements fragment from time to time, giving rise to new elements that can be transmitted to the progeny of cells that contain them. Because [PSI + ] is transmitted to progeny with high fidelity (loss is less than 10−4 per generation), these elements must either be subject to frequent random fragmentation, or be partitioned to progeny in an orderly way. (5) In cells that do not already contain [PSI + ], overexpression of Sup35 will generate new elements. (6) Without Hsp104 (black bar) Sup35 can not acquire an aggregationprone state. Thus, Hsp104 deletion mutants have a psi-no-more phenotype. This is likely due to high local concentrations of the unstable, aggregation prone conformers of Sup35 in the vicinity of the individual Hsp104 molecules that produce them. Because Hsp104 is hexameric, it is possible that several Sup35 molecules are bound to the same Hsp104 particle at the same time, facilitating their coalescence into [PSI + ] elements. Once the [PSI + ] element has formed continuous over expression of Sup35 is not required, as the


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cycle, once started, is self perpetuating. (7) The mechanism by which over expression of Hsp104 cures cells of [PSI + ] remains a mystery. There are several possibilities. For example, when it is present at high concentrations, Hsp104 might bind Sup35 aggregates and disaggregate them, in a manner analogous to the manner in which it promotes the disaggregation of heat-damaged proteins. Another possibility, is that high concentrations of Hsp104 reduce the local concentration of a limiting number of Sup35 proteins, so that they do not form postulated pre-aggregate subassemblies (depicted as trimers at the bottom of the figure) that might be precursors for association with [PSI + ]. Yet another possibility, among many, is that a limiting cofactor is required to activate Hsp104 in this process and it must associate with Hsp104 in a specific stoichiometry.

that the highly charged M domain packs into the fiber on the outside and the C-terminus remains amorphous. Most remarkably, the N and M domains, as well as the whole Sup35 protein, form fibers very slowly, over many hours of incubation. However, the addition of a very small fraction of preformed fibers greatly increases the rate of fiber formation in fresh protein (Glover et al., 1997). This seeded polymerization process provides a coherent molecular explanation for the inheritance of [PSI + ] (Glover et al., 1997), similar to that proposed previously for mammalian prions (Lansbury and Caughey, 1995). That is, the ordered aggregates of Sup35 reduce the population of free Sup35, reducing the efficiency of translation termination and causing the [PSI + ] phenotype. These are unlikely to form on their own in vivo, and thus [psi - ] cells remain [psi - ] for generation after generation. But once an aggregate has formed, it seeds the polymerization of new protein, which is passed on from mother cells to their daughters as heritable [PSI + ] elements. An important independent study by Paushkin et al., 1997 also supports such a model. Aggregated Sup35 protein purified from [PSI + ] cells causes the soluble Sup35 protein of [psi - ] cells to aggregate, and this aggregated material can be used to seed the aggregation of more soluble Sup35 from fresh [psi - ] lysates. 5.4.5. Hsp104 and Sup35 Interaction in Vitro The genetic interactions between Hsp104 and [PSI + ] and their influence on the aggregation state of Sup35 support the idea that a chaperone controls the conformational state of this yeast prion. But evidence for a direct physical interaction between Hsp104 and Sup35, which could mediate this transformation was wanting. Recent in vitro studies with purified Sup35 and Hsp104 proteins have provided this evidence (Schirmer and Lindquist, 1997). While stable Sup35-Hsp104 complexes have not been detected, three separate approaches indicate they interact directly. 1) Sup35 protein selectively inhibits the ATPase activity of Hsp104. This was surprising in that casein, a test-substrate of the ClpAP pro tease, stimulated the ATPase activity of ClpA (Hwang et al., 1988) and casein even weakly stimulated the ATPase activity of Hsp104 (Schirmer and Lindquist, unpublished). Perhaps the nature of Hsp104’s interaction with Sup35 requires more


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concerted “work” from the chaperone, which reduces its basal rate of ATP hydrolysis. 2) Light scattering is increased in mixtures of purified Sup35 and Hsp104 proteins over the sum of the scattering by the proteins individually. 3) The circular dichroism (CD) spectrum observed for mixtures of the two proteins deviated significantly from that predicted by adding their individual spectra, contrary to the expected result if two proteins do not interact. This effect was dependent on the initial state of Sup35 which was controlled using different buffer conditions. In contrast, mixtures of Sup35 and control -lactalbumin, bovine proteins (aldolase, IgM, IgG, 2-mac-roglobulin, apoferritin, serum albumin, Hsp70) or of Hsp104 and (aldolase, IgM, IgG, 2-macroglobulin, apoferritin, -lactalbumin, bovine serum albumin) produced in each case a spectrum that matched the sum of the proteins individual spectra regardless of which buffers were employed. These data clearly indicate that a direct and specific physical interaction occurs between Hsp104 and Sup35 which changes the state of the protein(s). However, it is as yet unclear what is the exact nature of the change that occurs in the protein (s). In mixed solutions it is impossible to assign CD changes to individual proteins. Furthermore, the increase in light scattering suggests that some part of the spectral shift observed by CD is due to a loss of material from the solvent phase making analysis of the structural changes underlying the shift more complex. It is also unclear whether the interaction of Sup35 and Hsp104 alone in vitro will fully represent the role of Hsp104 in prion maintenance. In vivo, Sup35 is normally bound to a partner protein, Sup45, and other as yet unidentified cofactors might also be involved. Finally, in keeping with genetic observations, variations in the ratios of Hsp104 to these components need to be investigated in vitro. 5.5. A Model for Hsp104-[PSI + ] Interactions Clearly, Hsp104 plays a central role in the formation and maintenance of [PSI + ], The relationship between Hsp104 and heat-induced aggregates is different from that between Hsp104 and Sup35 aggregates. Nevertheless, the role Hsp104 plays in providing thermotolerance and the role it plays in the inheritance of [PSI + ] must be biochemically related by Hsp104’s capacity to alter the conformational state of other proteins. Although it must be taken as highly speculative, a model to explain the role of Hsp104 in the inheritance of [PSI + ] is presented in Figure 7 and its legend. 5.6. An Explanation for Previous Puzzles The remarkable interactions between Hsp104 and [PSI + ] provide an explanation for three long-standing puzzles concerning [PSI + ] and one more recent puzzle concerning thermotolerance. 5.6.1. Hsp104 Explains Puzzling [PSI + ] Phenomena The longest standing puzzle concerns the underlying genetic nature of the [PSI + ] element. Many types of evidence, including the effects of Hsp104, now support the hypothesis that the inheritance of [PSI + ] is based upon the inheritance of a conformationally altered Sup35 protein. This mechanism of inheritance is, of course,


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novel and unexpected. For many years it had been assumed that the inheritance of [PSI + ] must be based upon the inheritance of a specific nucleic acid. The ability of mutagens such as UV light and MMS (methyl methanesulfonate) to cure cells of [PSI + ] seemed to provide strong support for this hypothesis (Tuite and Cox, 1980). How can this be reconciled with current data? DNA sequence analysis reveals the presence of two DNA damage-response elements in the HSP104 promoter. Indeed, Hsp104 is strongly and selectively induced by UV and MMS (Schmitt and McEntee, personal communication). The ability of Hsp104 to cure cells of [PSI + ] thus allows the effects of these mutagens to be accomodated within the proteinconformation based hypothesis of [PSI + ] inheritance. Another puzzle concerns the effects of guanidinium hydrochloride (GuHCl), a protein denaturant, on [PSI + ]. Growth in low concentrations of GuHCl cures virtually all cells of [PSI + ] (Tuite and Cox, 1980). This concentration seems too low to change the conformation of Sup35 directly. More likely, curing results from the induction of Hsp104 by low concentrations of GuHCl (Lindquist et al., 1995). Finally, heat shock also cures cells of [PSI + ]. This, of course, is likely due to the induction of Hsp104. Curiously, however, heat shock cures cells of [PSI + ] much less efficiently than growth in the presence of GuHCl (Tuite and Cox, 1980), and growth at high temperatures does not cure cells of [PSI + ] at any appreciable frequency (Tuite et al., 1981; Chang and Lindquist, unpublished). The reason may be that the induction of Hsp104 by GuHCl is far more selective than its induction by heat (Lindquist et al., 1995). For example, at the low concentrations of GuHCl required to cure [PSI + ] Hsp70 and Hsp26 are not induced (Lindquist et al., 1995). Apparently, in response to heat, yeast cells produce other factors that protect [PSI + ] from efficient Hsp104 curing, thereby protecting prion-based inheritance from minor vagaries of the environment. This protection of [PSI + ] suggests that [PSI + ] is not simply a disease of yeast cells, but represents a biologically advantageous mechanism for changing the fidelity of protein synthesis, and presumably the pattern of gene expression, in a heritable way. Indeed, the mere existence of the N-terminal domain of Sup35 suggests as much. Yeast cells rapidly rid themselves of disadvantageous markers, yet they retain the N-terminal domain of Sup35. Since this domain is dispensible for normal growth and apparently serves only to reduce the fidelity of translation when cells carry [PSI + ], it follows that reducing the fidelity of translation is likely to be of selective advantage under some circumstances. Perhaps [PSI + ] allows ribosomes to read through stop codons on certain natural messages, into new coding information, which might allow cells to exploit a specific new environmental niche. If so, the heritability of the element would ensure that this trait is stably maintained, but the low-level instability of the element would ensure that once a population has attained an appreciable size, [PSI + ] will be lost from some individuals, returning them to normal translation and fitness. 5.6.2. [PSI + ] Explains a Puzzling Thermotolerance Phenomenon The vital role played by Hsp104 in stress tolerance is clear. For several years, however, the observations of Smith and Jaffe (1991) on the effects of a temperature-sensitive heat shock transcription factor (HSF) provided a puzzling contradiction. This mutant can not


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induce Hsps in response to stress, yet it is perfectly capable of acquiring thermotolerance. The existence of non Hsp-based mechanisms of tolerance, such as the synthesis of trehalose, provides a partial explanation. However, these HSF mutants acquire just as much thermotolerance as wild-type cells, which should not be possible in the absence of Hsp104 induction. Analysis of Hsp accumulation in these mutants with specific antibodies seemed to solve the problem. The mutants have high basal levels of Hsp 104 expression and induction of Hsp 104 by heat is not completely blocked. In addition to classic HSEs, heat shock elements, the Hsp104 promoter contains STREs, stress response elements that use a different transcription factor. It seemed a simple matter to prove that thermotolerance in the HSF mutant was due to Hsp104 expression. But, it proved impossible to delete the Hsp104 gene in this strain in order to test this directly. In fact, it was impossible to even add an extrachromosomal wild-type copy of HSP104 to these cells. Either overexpression or deletion of Hsp104 was lethal. This conundrum was solved when it was determined that the HSF mutation was not a missence mutation, but a nonsense mutation that should have been lethal, because HSF is essential in yeast. However, the mutation had been produced in a [PSI + ] strain. Readthrough of the nonsense codon apparently kept the cells alive. However the cells were fixed with a requirement for moderate expression of Hsp104. Either overexpression or deletion of Hsp104 would cure the cells of [PSI + ], uncovering the effect of the lethal nonsense mutation in HSF. Thus, these cells represent a new type of mutant in [PSI + ] phenomenology, PFM, or psi-forever-more (Lindquist and Kim, 1996). 5.7. Broader Implications The study of mammalian neurodegenerative diseases is labor-intensive, expensive, and requires the use of many animals. The study of yeast prion-like elements can now provide a valuable supplement to mammalian investigations that will speed our understanding of self-propagating conformational changes, and may lead to the development of novel approaches for therapeutic intervention in neurodegenerative diseases. The hypothesis that [PSI + ] represents a protein-conformation based mechanism of inheritance, however, has much broader implications. If prions exist in both mammals and yeast, it is likely that they are very broadly distributed and may be responsible for many different epigenetic phenomena in other organisms, which have thus far defied molecular explanation. Finally, because chaperone proteins are induced by stress, their ability to influence the phenotypic state of the cell in this manner provides an extraordinary mechanism for a cell to respond to its environment in a heritable way. 5.8. Hsp 104 and Mammalian Prions The role of Hsp104 in the maintenance of the yeast prion [PSI + ] suggests that chaperone proteins might also regulate the structural state of other prions. Indeed there have been previous indirect suggestions of such interactions (Telling et al., 1995; Kenward et al., 1996). Providing a “proof of principle”, two recent studies have demonstrated that Hsp 104 can affect the conformational state of the mammalian prion, PrP, believed to be the causative agent in the transmissible spongiform encephalopathies (TSEs; See section 5.2).


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In the first, a bacterially expressed fragment of PrP corresponding to the proteaseresistant core that is characteristic of the TSE-associated form (rPrP) was mixed with Hsp104 in vitro. Changes in the ATPase activity of Hsp 104 and changes in the CD spectra of mixed proteins was assessed as for mixtures of Hsp104 and Sup35 (see section 5.4.4). Like Sup35, rPrP inhibited the ATPase activity of Hsp104. Also, mixtures of Hsp104 and rPrP exhibited a specific and highly reproducible shift in their circular dichroism spectra from that predicted by addition of each protein’s individual spectrum (Schirmer and Lindquist, 1997). This version of PrP has been extensively characterized (Mehlhorn et al., 1996; Zhang et al., 1997) and can be folded initially into either helical or -sheet forms. Just as the interaction between Hsp104 and Sup35 depended upon the initial state of Sup35, a spectral shift was observed when Hsp104 was mixed with the -sheet form of rPrP but not when Hsp104 was mixed with the -helical form of rPrP (Schirmer and Lindquist, 1997). Some of the control proteins used were also rich in -sheet, yet spectral shifts were not observed: thus this feature alone is insufficient to explain the shift. Another difference between the two rPrP forms is that the -sheet form sediments by ultracentrifugation at an average Mr which suggests it is in small aggregates containing ~7 proteins each, while the -helical form sediments as a monomer (Zhang et al., 1997). This is particularly interesting in light of the observation that in Hsp104’s in vitro interaction with luciferase, a thermotolerance test-substrate (section 4.2.2), the size of the most reactivation-competent aggregates were composed of 7 or more proteins (Glover and Lindquist, 1998). The second study employed an in vitro assay for the conformational conversion of PrP developed by Byron Caughey and colleagues. This assay uses preparations of the infectious, protease-resistant form of PrP, PrPSc, to seed the conversion of a small amount of radio-labeled protease-sensitive form, PrPC, to a PrPSc-like protease-resistant form. Although the infectious nature of the converted material has not been demonstrated, the conversion reaction mimics many essential features of PrPSc conversion, including the requirement for PrPSc, the production of strain-specific proteolytic fragments, and the species barrier (Kocisko et al., 1995; Raymond et al., 1997). Hsp104 and several other chaperones were added to this conversion reaction to determine if they could affect the rate of conversion. None of the chaperones could convert PrPC to the protease resistant form in the absence of added PrPSc, confirming that even in the presence of proteins that can alter the conformations of other proteins, a “templating” activity of PrPSc is required. In the presence of PrPSc, both GroEL and Hsp104 increased the rate of conversion (DebBurman et al., 1997). The initial state of the PrP was critical to the conversion process: Hsp104 only facilitated conversion if a partially denatured PrPSc seed was used. The post-translational modifications of PrP also influenced the process. A ~30–40% increase in conversion to protease-resistant material was observed when a modified PrP lacking its GPI anchor or a non-glycosylated form was used in the assay (DebBurman et al., 1997). While Hsp104 promoted conversion, osmolytes (chemical chaperones) inhibited it (DebBurman et al., 1997). Both Hsp104 and osmolytes protect cells from severe heat stress (see section 4.7.3). Osmolytes do so by preventing protein aggregation and Hsp104 does so by solubilizing proteins that have already aggregated. With PrP, the role of the osmolytes is consistent—preventing aggregation. In contrast, the role of Hsp104 appears to be to promote aggregation, contrary to its role in thermotolerance, but


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consistent with one of its two effects on [PSI + ] and also consistent with the idea that Hsp104’s function with [PSI + ] differs from its function in thermotolerance (section 1). Hsp104 and PrP are from different organisms and different subcellular compartments. These observations, therefore, do not imply that Hsp104 plays a role in prion-mediated disease. However, the demonstration that a chaperone can affect the conformational transitions of PrP, emphasizes the importance of determining what natural chaperones might interact with PrP in vivo, as a possible means for therapeutic intervention. Perhaps the most striking outcome of these studies is the underlying structural similarities they suggest between the PrP protein and Sup35. At an amino-acid sequence level, these proteins have little in common and their biological functions are completely unrelated. Yet Hsp104, which shows no evidence of interaction with most other cellular proteins, interacts with both PrP and Sup35 in a highly specific manner to promote an unusual change in structure that has profound consequences. 6. CONCLUDING REMARKS All proteins that are produced in large quantities in response to heat, the Hsps, are now known to perform functions that address problems in protein folding. Some of the proteins, such as polyubiquitin and certain of the ubiquitin congugating enzymes, function to rid the cell of damaged proteins through proteolysis. Others, such as Hsp70, DnaJ, and GroEL (or Hsp60), bind unfolded proteins and prevent them from aggregating. Hsp104 complements these functions in a unique and unexpected way, helping to resolve aggregates once they have formed and to return the aggregated proteins to the soluble, functional state. This biochemical function fits the biological role of Hsp104 in stress tolerance. Hsp104 is most important under conditions of extreme stress, when the rate of protein aggregation is likely to outpace the capacity of other protective mechanisms to prevent aggregation. The role of Hsp104 in the inheritance of the novel, protein conformation-based genetic element known as [PSI + ] is more perplexing. In this case, Hsp104 is actually required, at a specific concentration, to maintain Sup35 aggregates. Hsp104’s ability to promote the dissolution of heat-damaged aggregates and to promote the formation of Sup35 aggregates are biochemically related. Mutations in conserved nucleotide-binding residues of Hsp104 have strong effects on thermotolerance and the maintenance of [PSI + ]. Homologous mutations in NBD-1 and NBD-2 have different effects on the two most distinctive biochemical properties of Hsp104, its ability to hydrolyze ATP and its capacity to oligomerize in response to nucleotide. In the future, a detailed understanding of these biochemical properties will no doubt illuminate Hsp104’s remarkable biological functions. A complete picture of Hsp104’s functions in thermotolerance and [PSI + ] maintenance will also require better understanding of the structural nature of Hsp104’s substrates. This presents a major challenge, as methods for studying protein aggregates are still poorly developed. Happily, the problem of protein aggregation is engaging an ever growing number of biochemists due, in part, to a new appreciation of the important role played by protein aggregation in several devastating human diseases (Thomas et al, 1995). Moreover, it is increasingly clear that chaperone proteins play an important role in the


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metabolism of these aggregated proteins (Perutz et al., 1994; Thomas et al., 1995; Gajdusek, 1996; Prusiner, 1996; Wetzel, 1996). Lessons learned in studying the aggregated proteins that cause disease will no doubt provide important insights on Hsp104 functions. Hopefully, the ability to analyze the problem of protein aggregation in an organism as genetically tractable as yeast, will also provide insights of importance to human biology and medicine. The problem is also of great economic importance, not only in the context of human and animal diseases (e.g. mad cow disease), but also because heat and water stress are the greatest barriers to crop productivity. The role demonstrated for Hsp104 in thermotolerance and the functional conservation of its homologs in plants suggest that manipulation of HSP100 proteins will be of great advantage for the survival of crops during heat and drought stresses. 7. REFERENCES Boreham, D.R. and Mitchel, R.E. (1994). Regulation of heat and radiation stress responses in yeast by hsp-104. Radiation Rsch. , 137 , 190–195. Chernoff, Y.O., Derkach, I.L. and Inge-Vechtomov, S.G. (1993). Multicopy SUP35 gene induces de-novo appearance of psi-like factors in the yeast Saccharomyces cerevisiae. Curr. Genet. , 24 , 268–270. Chernoff, Y.O., Lindquist, S.L., Ono, B., Inge-Vechtomov, S.G. and Liebman, S.W. (1995). Role of the chaperone protein Hsp104 in propagation of the yeast prion-like factor [psi + ]. Science , 268 , 880–884. Chuang, S.E., Burland, V., Plunkett, G.D., Daniels, D.L. and Blattner, F.R. (1993). Sequence analysis of four new heat-shock genes constituting the hslTS/ibpAB and hslVU operons in Escherichia coli. Gene , 134 , 1–6. Clarke, A.K. and Eriksson, M.-J. (1996). The cyanobacterium Synechococcus sp. PCC7942 possesses a close homologue to the chloroplast ClpC protein of higher plants. Plant Molec. Biol , 31 , 721–730. Colaco, C., Sen, S., Thangavelu, M., Pinder, S., and Roser, B. (1992). Extraordinary stability of enzymes dried in trehalose: simplified molecular biology. Bio/Technology , 10 , 1007–1011. Collinge, J., Palmer, M.S., Sidle, K.C., Hill, A.E, Gowland, I., Meads, J., Asante, E., Bradley, R., Doey, L.J. and Lantos, P.L. (1995). Unaltered susceptibility to BSE in transgenic mice expressing human prion protein. Nature , 378 , 779–783. Cox, B. (1994). Cytoplasmic inheritance. Prion-like factors in yeast. Curr. Biology , 4 , 744–748. Cox, B.S., Tuite, M.F. and McLaughlin, C.S. (1988). The psi factor of yeast: a problem in inheritance. Yeast , 4 , 159–178. Craig, E.A. and Jacobsen, K. (1984). Mutations of the heat inducible 70 kilodalton genes of yeast confer temperature sensitive growth. Cell , 38 , 841–849. Craig, E.A. and Jacobsen, K. (1985). Mutations in cognate genes of Saccharomyces cerevisiae hsp70 result in reduced growth rates at low temperatures. Molec. Cell. Biol , 5 , 3517–3524. Crowe, J.H., Crowe, L.M., and Chapman, D. (1984). Preservation of membranes in anhydrobiotic organisms: the role of trehalose. Science , 223 , 701–703.


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18. CHAPERONES AND CHARONINS: PROTEIN UNFOLDING ENZYMES AND PROTEOLYSIS MICHAEL R. MAURIZI1,*, SUE WICKNER2 and SUSAN GOTTESMAN2 1 Laboratory

2 Laboratory

of Cell Biology and of Molecular Biology, National Cancer Institute, Bethesda, MD 20892, USA

1. Introduction 2. Chaperone-mediated Proteolysis by ATP-dependent Pro teases 2.1. Protein Degradation in vivo by ATP-dependent Proteases 2.1.1. Lon Proteases 2.1.2. Clp Proteases 2.1.3. FtsH 2.2. Structures and in vitro Activities of ATP-dependent Proteases 2.2.1. Lon Protease 2.2.2. Clp Proteases 2.2.3. FtsH 3. Chaperone-like Activities of ATP-dependent Proteases 3.1. General Considerations 3.2. ClpA Activation of RepA 3.3. ClpX Functions in Phage Replication and Transposition 3.2.1. Disassembly of the Mu Transpososome 3.2.2. Lambda O Protein 3.4. ClpB/Hsp100 Proteins: Disaggregation but no Degradation 3.5. Chaperone Activity of the Lon Protease ATPase Domain 3.6. Chaperone Activity of FtsH Homologs 4. Protein Unfolding as a Step in ATP-dependent Proteolysis 4.1. Requirement for ATP Hydrolysis 4.2. ATP Hydrolysis Drives Unfolding by Lon Protease 5. Involvement of General Chaperones in Protein Degradation 5.1. Decreased Protein Degradation in Chaperone Mutants 5.1.1. General Chaperone Mutants in E. coli 5.1.2. E. coli Chaperones Allow Protein Degradation by Preventing


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Aggregation 5.1.3. Defects in Protein Degradation in Eukaryotic Chaperone Mutants 5.2. Pleiotropic Effects of Mutations in Chaperones 6. Substrate Selection by ATP-dependent Proteases and Chaperones 6.1. Degradation Motifs 6.2. Exposed Bonding Domains as Potential Recognition Motifs 6.3. Do Chaperones and Proteases Recognize Similar Features of Proteins? 7. Concluding Remarks

1. INTRODUCTION When protein structures become disrupted, for example in response to chemical damage or physical stress, they unfold and tend to aggregate. Similar unfolding and aggregating behavior is displayed by proteins that are not necessarily damaged but have unusual surface features or abnormal conformations, such as a few intrinsically unstable proteins, mutant proteins, unbound subunits of multimeric complexes, proteins that have lost stabilizing ligands, and proteins that are in a foreign intracellular milieu. Because unfolded proteins can interfere with the functioning of normal proteins, quality control over protein conformation and structure is important for cell growth. Cells respond to unfolded proteins by attempting to refold the proteins or by degrading them. In either case, the ability to prevent rapid aggregation and to disaggregate existing aggregates is necessary to allow refolding or degradation. What has become clear in recent years is that the two major systems that respond to misfolded proteins are closely integrated: enzymes whose primary role is protein folding are needed for rapid protein degradation, and proteins identified as integral components of pro teases can promote refolding as well as degradation. The best studied refolding enzymes are the molecular chaperones and chaperonins, which we call general chaperones. In E. coli, general chaperones include the DnaK system (see chapters Burkholder & Gottesman; Zylicz et al.; Buchberger et al., this volume) consisting of DnaK (a member of the Hsp70 family), DnaJ, and GrpE, as well as the GroE system (see chapters Burkholder & Gottesman; Burston & Saibil; Ranson & Clarke, this volume) consisting of GroEL (a member of the Hsc60 chaperonin family) and GroES. These general chaperones display a broad range of ATP-dependent protein folding activities and also promote assembly or disassembly of macromolecular complexes. Another general chaperone is ClpB/Hsp104 (Gottesman et al., 1990b; Parsell et al., 1991; Squires & Squires, 1992), which has protein disaggregating activity (Parsell et al., 1994b). ClpB is highly homologous to ClpA, a component of the ATP-dependent ClpAP pro tease (Gottesman et al., 1990b), but has not been shown to participate directly in protein degradation (Park et al., 1993; Woo et al., 1992). In addition to influencing protein folding and aggregation, general chaperones and other protein remodeling enzymes have large effects on protein degradation in vivo. While cells contain a multitude of proteases, most intracellular protein degradation is

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carried out by a limited set of ATP-dependent proteases. Prototypes of all of the known ATP-dependent proteases are found in E. coli and include Lon protease, ClpAP, ClpXP, FtsH (or HflB), and ClpYQ (or HslVU). In eukaryotic cells, homologs of Lon and FtsH are found in mitochondria. The major cytosolic protease, the 26S proteasome, is structurally analogous to ClpAP (Kessel et al., 1995) and evolutionarily related to ClpYQ (Chuang et al., 1993). Sequence analysis as well as enzymatic and physical studies of the ATP-dependent proteases indicate that they are composed of two domains: a proteolytic domain and an ATPase domain that has been shown to have a protein unfolding or remodeling activity (Maurizi, 1992; Wickner et al., 1994). The ATP-dependent activities of this domain, which include unfolding the protein and facilitating its translocation to the proteolytic active sites, are essential for protein degradation. General chaperones and ATP-dependent proteases work on a common pool of substrates and may share a common mode of action in the initial steps of interaction with protein substrates. The distinction between general chaperones and the chaperone components of ATP-dependent proteases may thus be arbitrary, an accident of discovery or assay, although differences in specificities, binding affinities, and kinetics of interaction and unfolding may determine if particular chaperones participate predominantly in refolding or in degradation. Mutations in general chaperones can drastically affect degradation of proteins in vivo, but it has been difficult to demonstrate direct physical association between general chaperones and proteases or even to define a pathway in which chaperones and proteases act in an ordered fashion. The discovery that integral components of ATP-dependent proteases have molecular chaperone activity, however, has provided evidence that chaperone-assisted unfolding can be directly coupled to protein degradation. We have proposed that the chaperone components of ATP-dependent proteases be called “charonins’” after the mythological underworld creature who ferried souls across the river Styx. Chaperone components or domains of ATP-dependent proteases may in fact unfold proteins exclusively for degradation, or they could first attempt to release the protein so it can refold. Conversely, general chaperones may also have a “charonin” function, taking an active role in promoting degradation when a protein fails to refold. We do not know the extent to which chaperone and charonin functions overlap. The relative contributions of different chaperones and proteases to either refolding a particular protein or to elimination of an intransigent unfolded protein are likely to vary widely. In this chapter we will summarize current data about the properties and activities of general chaperones and of the chaperone components of ATP-dependent proteases with the following questions in mind: • Do general chaperones and protein remodeling enzymes act directly or indirectly in affecting proteolysis—that is, do proteases recognize and degrade the “chaperonebound” or the “released” form of unfolded proteins? • What chemical or structural motifs in unfolded proteins are recognized by chaperones and proteases and are they the same? • And perhaps the most critical question—what is the nature of the decision checkpoint between refolding and degradation? As will be seen, we have only begun to obtain answers to some of these questions, and


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we are almost completely in the dark regarding mechanistic issues. However, our base of knowledge is now sufficiently broad and solid to provide context for obtaining more profound answers in the near future. 2. PROTEOLYTIC FUNCTIONS AND STRUCTURES OF ATP-DEPENDENT PROTEASES 2.1. Protein Degradation in vivo 2.1.1. Lon Proteases Lon protease (protease La) from E. coli was identified on the basis of its ability to degrade proteins in vitro in the presence of ATP and was the first ATP-dependent protease to be purified (Goldberg, 1992; Goldberg et al., 1994). E. coli Lon is the product of a single gene, lon, which is required in vivo for the degradation of proteins such as SulA, RcsA, N protein, CcdA, and Tn903 transposase (Gottesman & Maurizi, 1992). lon mutants display phenotypes, such as UV sensitivity, mucoidy, and susceptibility to post-segregational killing after loss of certain plasmids, as a consequence of defects in degradation of specific Lon substrates. Lon can also degrade certain mutant proteins in vivo (Sherman & Goldberg, 1992) and is responsible for = 50% of the degradation of abnormal proteins, such as puromycyl fragments and canavanine-containing proteins (Bukhari & Zipser, 1973; Maurizi et al., 1985). Lon degrades the N protein (Maurizi, 1987) and CcdA (Van Melderen et al., 1996) in a purified in vitro system, which demonstrates that no other proteins are essential for expression of Lon activity. Lon proteases have important functions during growth and development in other bacteria. In Myxococcus, two Lon proteases are found; one is essential during vegetative growth and the other plays an unknown role during stalk formation and myxospore maturation (Gill et al., 1993; Tojo et al., 1993a; Tojo et al., 1993b). In Caulobacter, lon mutations lead to developmental defects attributable in part to the failure to degrade CcrM, an adenine DNA methyltransferase involved in cell-cycle dependent variation in DNA methylation (Wright et al., 1996). Lon protease has been identified in eukaryotic cells where it is encoded in chromosomal DNA but is found, apparently exclusively, in the mitochondrial matrix (Suzuki et al., 1994; Van Dyck et al., 1994; Wang et al., 1993). In vitro properties of eukaryotic Lon suggest it is similar to E. coli Lon in promoting ATP-dependent protein degradation (Desautels & Goldberg, 1982; Wang et al., 1993; Watabe & Kimura, 1985). Yeast cells with mutations in LON (also called PIM1) accumulate deletions in mitochondrial DNA, are respiration deficient, cannot grow on non-fermentable carbon sources, and accumulate dense inclusion bodies within mitochondria (Suzuki et al., 1994; Van Dyck et al., 1994). Yeast LON mutations lead to stabilization of several mitochondrial proteins, including the subunit of the matrix processing peptidase and the , , and subunits of F1 ATPase (Suzuki et al., 1994).


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2.1.2. Clp Proteases ClpAP (also referred to as protease Ti (Woo et al., 1989) was isolated from E. coli lon mutant cell extracts based on its ability to degrade casein in an ATP-dependent manner (Katayama-Fujimura et al., 1987; Maurizi, 1992). ClpAP is composed of two separate gene products, ClpA, an ATPase, and ClpP, a peptidase, which can be purified separately and reconstituted to obtain the active protease (Maurizi, 1992). In vivo, ClpAP appears to play a minor role in the degradation of abnormal proteins, such as canavanyl proteins (Gottesman et al., 1990a) but targets certain -galactosidase fusion proteins quite specifically (Gottesman et al., 1990a; Tobias et al., 1991). ClpAP is responsible for degradation of abnormal -galactosidase proteins with bulky hydrophobic amino terminal residues (Leu, Phe, and Trp) and thus appears to recognize the degradation motif displayed by those proteins whose in vivo stability follows the N-end rule (Tobias et al., 1991). The only natural substrate for ClpAP identified to date is MazE (Aizenman et al., 1996), a low molecular weight protein homologous to antidote proteins, such as PemI, which are part of plasmid post-segregational killing systems (Jensen & Gerdes, 1995). The function of MazE in E. coli and the consequence of its degradation are not known, although it appears to be regulated as part of the stringent response (Aizenman et al., 1996). ClpP is also activated in vivo and in vitro by ClpX (Gottesman et al., 1993; Wojtkowiak et al., 1993), a protein with sequence similarity to ClpA. The clpP and clpX genes are co-transcribed in vivo (Gottesman et al., 1993), but the two proteins are easily separated in cell extracts and must be added together in the presence of ATP to reconstitute the active protease (Wojtkowiak et al., 1993; Grimaud, et al., 1998). ClpXP does not degrade the same set of proteins degraded by ClpAP indicating that the ATPase component confers specificity to the protease. In vivo, ClpXP degrades the E. coli starvation sigma factor, RpoS (Schweder et al., 1996) several unidentified proteins made during carbon starvation (Damerau & St. John, 1993), and several phage proteins: the O protein (Wojtkowiak et al., 1993), PhD, an antidote protein made by phage P1 (Lehnherr & Yarmolinsky, 1995), and mutant forms of the Mu repressor (MhammediAlaoui et al., 1994). Another member of the Clp protease family, ClpYQ, was originally identified by sequence analysis (Chuang et al., 1993). ClpY is homologous to ClpX (Gottesman et al., 1993) and is encoded in an operon along with ClpQ, which is homologous to the subunit of the 20 S proteasome (Chuang et al., 1993). A mixture of purified ClpY and ClpQ has been shown to have peptidase activity (Rohrwild et al., 1996; Yoo et al., 1996) as well as ATP-dependent protein degrading activity (Kessel et al., 1996; Huang and Goldberg, 1997). In vivo, ClpYQ may affect the levels of the acute heat shock sigma factor, , but no other targets have been identified (Missiakas et al., 1996). 2.1.3. FtsH (HflB) The ftsH gene is essential in E. coli and encodes an ATP-dependent protease found in the cytoplasmic membrane (Tomoyasu et al., 1995). Mutants of ftsH are temperature sensitive for growth, possibly as a result of defective translocation of membrane spanning


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proteins, and these mutations can be partially suppressed by overproduction of GroEL/GroES or Hsp90 (Shirai et al., 1996). ftsH was originally identified as hflB (high frequency lysogenization), because of its effect on the frequency of lambda lysogenization (Banuett et al., 1986). Mutations in ftsH lead to stabilization of two regulators of lambda lysogenization, CII (Banuett et al., 1986) and CIII (Herman et al., 1997). FtsH is responsible for the rate limiting step in degradation of (Herman et al., 1995; Tomoyasu et al., 1995) (see chapter Connolly et al., this volume) and for degradation of unassembled subunits of the membrane-associated secretory protein, SecY (Kihara et al., 1995). Homologs of FtsH have been identified in several organisms. In yeast, the FtsH homologs YTA-10 and YTA-12 can degrade newly synthesized proteins in isolated mitochondria, and this activity depends on an intact Zn-metalloprotease site (Arlt et al., 1996). 2.2. Structures and in vitro Activities of ATP-dependent Proteases 2.2.1. Lon Protease Lon (subunit Mr 89,000) is a soluble oligomeric protein whose exact structure remains to be definitively determined (Goldberg et al., 1994; Watabe & Kimura, 1985). From sequence analysis, Lon appears to be divided into two domains, an amino terminal ATPase and a carboxy terminal pro tease (Chin et al., 1988; Goldberg et al., 1994; Gottesman et al., 1995). The ATPase domain contains a Walker consensus ATP-binding motif, and mutations in that site eliminate the ATP-dependent protein degrading activity of Lon (Fischer & Glockshuber, 1994). The carboxy terminal region has no obvious sequence similarity to other pro teases, although Lon is inhibited by reagents that inactivate classical serine pro teases (Goldberg et al., 1994). A mutation in Ser679 eliminates protease and peptidase activity (Fischer & Glockshuber, 1993; Yu et al., 1991), but the mutant Lon-S679A retains ATPase as well as a chaperone-like activity (see below). Purified Lon protease can cleave low molecular weight peptides in the absence of nucleotide, and this function is activated by non-hydrolyzable analogs of ATP (Waxman & Goldberg, 1982; Waxman & Goldberg, 1986). Degradation of various partially denatured proteins in vitro shows a more complex response to nucleotide. For some denatured proteins, there is a stoichiometric coupling of two ATP molecules hydrolyzed per peptide bond cleaved (Goldberg & Waxman, 1985; Menon et al., 1987). The requirement for ATP hydrolysis, however, is dependent on a combination of the size and the structure of the protein substrate, such that ATP hydrolysis is absolutely required for degradation of some (generally higher molecular weight) proteins (Goldberg & Waxman, 1985; Menon et al., 1987), but non-hydrolyzable analogs can activate degradation of smaller (Mr=30,000) proteins (Maurizi, 1987; Van Melderen et al., 1996). Protein substrates bind to Lon and activate the ATPase activity (Waxman & Goldberg, 1986) but stimulation of ATP hydrolysis is not dependent on peptide bond cleavage, because ATPase activity of a mutant Lon lacking the catalytic serine residue was still activated by protein binding (Fischer & Glockshuber, 1993).


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2.2.2. Clp Proteases ClpP shares no motifs common to the trypsin and subtilisin families but does contain an active site serine, histidine, and aspartate triad characteristic of serine proteases (Maurizi et al., 1990a; Maurizi et al., 1990b). ClpQ (HslV), which by sequence comparison is unrelated to ClpP, belongs to a newly discovered family of threonine proteases that includes the Archae and eukaryotic proteasomes (Seemuller et al., 1995). ClpP (subunit Mr 21,400) is composed of two axially aligned rings of seven subunits each. The ClpP structure is reminiscent of the β subunits of the proteasome, which has 14 active sites located on the interior surface of an aqueous cavity formed by the joining of two concave seven-membered rings (Lowe et al., 1995). ClpQ (subunit Mr 19,000) also is composed of two superimposed rings, but, despite its homology with the proteasome, ClpQ has only six subunits in each ring and the proteolytic cavity should then have 12 rather than 14 active sites (Kessel et al., 1996; Rohrwild et al., 1997). How this difference affects efficiency of cleavage or the size of the average peptide products generated by ClpQ and proteasomes should provide valuable information about the mechanism of these proteases. The ATPases that define the Clp family can be divided into two subfamilies, one represented by ClpA and ClpB, which possess two ATPase domains, and the other represented by ClpX and ClpY, which possess a single ATPase domain (Gottesman et al., 1995). ClpA and ClpB are highly homologous (64% overall similarity and 88% similarity in approximately 200 amino acids in each of the core ATPase regions) (Gottesman et al., 1990b). ClpX and ClpY are homologous to each other (55% overall similarity) but somewhat more distantly related to ClpA (~50% similarity in the ATPase region and Cterminal domain) (Gottesman et al., 1993; Gottesman et al., 1995). Despite its similarity to ClpA, ClpB has not been shown to activate proteolysis (Park et al., 1993; Woo et al., 1992) by ClpP or any other proteases. ClpA subunits (Mr 84,000) associate to form six-membered rings in response to binding of ATP or non-hydrolyzable analogs of ATP. The two ATPase domains of ClpA appear to form structurally distinct rings (Kessel et al., 1995). In the ClpAP complex, a hexamer of ClpA binds on each face of ClpP forming a barrel-like molecule. The axial channels through ClpA and ClpP into the proteolytic active sites appear constricted and may have to expand in response to protein binding or ATP hydrolysis in order to allow passage of unfolded proteins. ClpX (subunit Mr 46,000) is purified as a heterogeneous mixture of oligomers and forms stable hexameric rings in the presence of ATP or ATP S (Grimaud et al., 1998). Complexes of ClpX and ClPP resemble ClpAP, with a hexameric ring of ClpX bound to each face of ClpP. ClpX and ClpA have similar affinities for ClpP, and complexes with CIPA and ClpX bound to opposite faces of ClpP have been observed (Grimaud et al., 1998). ClpY (subunit Mr 49,000) associates in the presence of ATP to form two types of oligomeric rings (Kessel et al., 1996). Both six-membered and seven-membered rings of ClpY have been observed (Kessel et al., 1996; Rohrwild et al., 1997). The significance of the heterogeneity in ring structures is not known, but it might affect substrate selection. Since the central channels through the six- and seven-membered rings should have different diameters, the size of proteins that interact with or are translocated to ClpP may


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be different. ClpP alone has activity against low molecular weight peptides, but its active site appears to be inaccessible to large or folded polypeptides and proteins (Thompson & Maurizi, 1994; Woo et al., 1989). In the presence of ClpA, ClpP can degrade longer peptides as well as proteins; the requirement for ATP hydrolysis depends on some combination of the size and structure of the substrate. Non-hydrolyzable analogs of ATP stimulate cleavage of some polypeptides (10–30 amino acids) at remarkably rapid rates (>11,000 peptide bonds per minute per tetradecamer of ClpP) (Thompson & Maurizi, 1994). Protein degradation by ClpP in the presence of ClpA requires ATP hydrolysis (Thompson et al., 1994; Woo et al., 1989). ClpA in the absence of ClpP has a substantial ATPase activity (~600 ATP per min per ClpA6) which is activated only 20–40% in the presence of protein substrates with or without ClpP (Hwang et al., 1988; Singh & Maurizi, 1994). Thus, ATP hydrolysis is not coupled in a concerted mechanistic manner to peptide bond cleavage, and the stoichiometric ratio of ATP hydrolysis to peptide bond cleavage (4–8 ATP per peptide bond) is variable (Maurizi et al., 1994). The peptide bond cleavage rate in protein substrates is 120–180 per minute per ClpAP complex, about 50 times slower than the fastest rates seen with peptides, indicating that the rate limiting step in protein degradation is not peptide bond cleavage but some reaction associated with the ATP hydrolysis. In the presence of ClpX, ClpP can degrade specific substrates (such a O protein) in vitro, and this activity also requires ATP hydrolysis (Wojtkowiak et al., 1993). ClpQ by itself can degrade small fluorogenic peptides (Wu, Gottesman, Maurizi, unpublished) but peptide cleavage can be activated by ClpY in the presence of ATP (Rohrwild et al., 1996; Yoo et al., 1996). ClpY and ClpQ can also degrade proteins, such as casein, when ATP hydrolysis is allowed but not in the presence of non-hydrolyzable analogs of ATP (Kessel et al., 1996; Huang and Goldberg, 1997). 2.2.3. FtsH FtsH is composed of a single polypeptide chain (Mr 71,000) that forms an oligomeric integral membrane protein (Tomoyasu et al., 1995). Sequence analysis predicts that FtsH has two membrane spanning regions and that the C-terminal 530 amino acids constitute a large cytoplasmic domain containing a 200-amino acid region conserved among the AAA family of ATPases (Tomoyasu et al., 1995) and a putative carboxy terminal Znmetallopeptidase domain. Purified detergent solubilized FtsH has ATPase activity and could degrade a specific protein substrate, , in the presence of ATP (Tomoyasu et al., 1995), suggesting that FtsH alone is capable of carrying out ATP-dependent protein degradation. 3. CHAPERONE-LIKE ACTIVITIES OF ATP-DEPENDENT PROTEASE COMPONENTS 3.1. General Considerations The evidence that components or domains of ATP-dependent proteases have activity


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similar to that of molecular chaperones is now incontrovertible. Direct evidence for chaperone-like activities has been obtained in purified systems, and biological activity has been shown for ATPase domains in the absence of enzymatically active proteolytic domains. Both ClpA and ClpX are able to bind to thermally-unfolded proteins and protect them from irreversible denaturation, though neither protein has been shown to catalyze refolding of thermally unfolded proteins, as has been shown for general chaperones (Wawrzynow et al., 1995; Wickner et al., 1994). More convincing data has been obtained in studies showing that ClpA, ClpX, and proteolytically inactive mutants of Lon protease can help assemble or disassemble specific protein complexes. These studies show clearly that the ATPase components of proteases can remodel protein structure. 3.2. ClpA Activation of RepA ClpA interacts with an inactive dimeric form of the RepA protein of phage P1 and converts it to an active monomeric form that binds specific oriP1 DNA sequences (Wickner et al., 1994). The inactive dimer of RepA binds tightly to ClpA in the presence of non-hydrolyzable analogs of ATP, and activation and release of the active monomer requires ATP hydrolysis. RepA activation was originally shown to be catalyzed by the DnaJ/K chaperone system and to be accomplished non-enzymatically by dissociation of RepA dimers with mild denaturants such as urea (Wickner et al., 1991). ClpA thus has the activities of a molecular chaperone in binding and catalyzing partial unfolding of proteins. ClpA will also promote the ATP-dependent degradation of RepA if ClpP is present (Wickner et al., 1994). Do the same unfolding reactions that lead to activation of RepA also lead to degradation? The answer is not entirely clear but several bits of data suggest a close link. Dimers of RepA bind with the same stoichiometry to ClpA and to ClpAP complexes in the presence of non-hydrolyzable ATP analogs, and the dimer of RepA has a lower Km for degradation than the monomer, suggesting that ClpA acts on the same form of RepA for both reactions (Pak and Wickner, unpublished). Also, when oriP1 DNA, which protects active RepA from degradation, is present from the start of a reaction with ClpAP and RepA, some of the RepA is activated and binds DNA, but some of the RepA is rapidly degraded. Thus, some RepA unfolded by ClpA can be captured and degraded before it is released to refold into an active form. The timing of unfolding and release versus unfolding and capture is the key, still missing, to a more complete understanding of chaperone-mediated proteolysis. 3.3. ClpX Functions in Phage Replication and Transposition 3.3.1. Mu Transposase ClpX displays chaperone-like activity with several proteins in vitro and experiments suggest that, in vivo, ClpX may have activity, presumably as a chaperone, independently of ClpP (Mhammedi-Alaoui et al., 1994). Purified ClpX promotes disassembly of Mu transposase (MuA) from the highly stable strand transfer complex, which is an obligatory intermediate in Mu transposition (Kruklitis et al., 1996; Levchenko et al., 1995). Displacement of the MuA tetramer from the strand transfer complex requires ATP


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hydrolysis and results in no covalent modification of the protein. The released MuA is active in a subsequent round of Mu recombination (Levchenko et al., 1995). In vivo, ClpX is essential for Mu growth, however ClpP is not, suggesting that it is the chaperone activity of ClpX that is required for Mu replication (Mhammedi-Alaoui et al., 1994). 3.3.2.

O Protein

Zylicz and co-workers demonstrated in a purified system that ClpX can protect O protein, which is a specific substrate for degradation by ClpXP in vivo and in vitro (Wawrzynow et al., 1995), from heat-induced aggregation and can disaggregate preformed aggregates of O protein. Binding of O protein stimulates ATP hydrolysis by ClpX, which is required for the disaggregation reaction. 3.4. ClpB/Hsp104: Disaggregation but no Stimulation of Degradation Purified ClpB, which has over 80% sequence similarity with ClpA, does not appear to interact with ClpP (Maurizi, unpublished), and biochemical studies have failed to show protein degradation mediated by E. coli ClpB (Woo et al., 1992) or the homologous yeast Hsp104 (Parsell et al., 1994a). E. coli ClpB mutants (Squires et al., 1991) and yeast Hsp104 (Parsell et al., 1994b) mutants tend to accumulate protein inclusion bodies, and Hsp104 mutants are sensitive to acute heat stress or to combinations of two or more moderate stresses (Parsell et al., 1994b; Sanchez et al., 1992). These and other data suggest that members of the ClpB subfamily catalyze the disaggregation of proteins (Schirmer et al., 1996). For example, activity of thermally unfolded firefly luciferase is recovered at low temperature in wild type yeast cells but not in HSP104 mutants (Parsell et al., 1994b), and purified Hsp104 can increase the recovery of RNA splicing activity when added back to heated yeast extracts lacking Hsp104 (Vogel et al., 1995). Hsp104 also appears to be necessary for the propagation of a prion-like particle in yeast cells (Paushkin et al., 1996) (for details see chapter Lindquist et al., this volume). Aggregates of PSI, a translation termination protein, are stable in yeast cells and are passed on to daughter cells, where they aggregate with newly synthesized PSI, thereby maintaining the steady state concentration of free PSI at a sub-optimal level which leads to read-through of stop codons. Moderate levels of Hsp104 are required to maintain this self-propagating aggregate of PSI, whereas aggregation is not initiated in the absence of Hsp104, and preexisting aggregates disappear when Hsp104 is in excess (Paushkin et al., 1996). A homolog of ClpB/Hsp104, HSP78, is found in mitochondria, where it does not appear to have an essential function (Schmitt et al., 1995). However, deletion of HSP78 in yeast cells carrying a temperature sensitive mt-Hsp70 results in a petite phenotype and impaired growth on non-fermentable carbon sources even at the permissive temperature. Overproduction of HSP78 suppresses a defect in import of mitochondrial proteins in the mt-Hsp70 strain but failed to repair a defect in degradation of abnormal proteins (Schmitt et al., 1995). HSP78 may have an auxiliary role in mitochondrial protein assembly and may be more restricted in its ability to recognize unfolded or unassembled proteins.


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3.5. Chaperone Activity of the Lon Protease ATPase Domain An intriguing, although indirect, demonstration of chaperone-like activity for an ATPdependent protease is the ability of multicopy yeast Lon to complement mutations in yeast genes, AFG3 (YTA-10) and RCA1 (YTA-12), which encode membrane-associated protease/chaperones involved in assembly of heterologous inner membrane complexes in yeast mitochondria (Rep et al., 1996). Complementation did not depend on the proteolytic activity of the overexpressed Lon, because a mutant Lon in which the active site serine was mutated to alanine showed enhanced assembly of cytochrome c oxidase. Mutating the ATPase site of Lon did abolish its ability to promote assembly (Rep et al., 1996). These results suggest that Lon can interact with intermediates in an assembly pathway and possibly prevent them from aggregating before they can find their appropriate partners in the complex. Whether Lon plays a passive role by interacting with exposed bonding domains of the unassembled subunits or can promote remodeling of the subunit structure is not clear from the available data. These data also confirm the ability of ATP-dependent proteases to specifically bind substrates that are recognized by other molecular chaperones. 3.6. Chaperone Activity of FtsH Homologs Yeast YTA-10 (AFG3) and YTA-12 (RCA1), homologs of E. coli FtsH, form an 850 kDa membrane-bound complex with both proteolytic activity and chaperone-like activity in assembly of mitochondrial inner-membrane protein complexes (Arlt et al., 1996). Mutations in YTA-10 or YTA-12 cause defects in respiration and impair assembly of ATP synthase in mitochondria (Arlt et al., 1996). Activity could be reintroduced with plasmids encoding either gene, and mutants of YTA-10 or YTA-12, in which a critical histidine residue in the Zn-metalloprotease site was altered, were able to promote ATP synthase assembly and restore respiration. Unassembled subunits of membrane complexes were degraded in wild type cells, but were stable when the cells were mutated in YTA-10 or YTA-12. Thus, it appears that the YTA10–12 complex mediates not only assembly of the inner membrane complexes but also degradation of any unassembled subunits (Arlt et al., 1996). 4. PROTEIN UNFOLDING AS A STEP IN ATP-DEPENDENT PROTEOLYSIS 4.1. Requirement for ATP Hydrolysis Much enzymatic and structural data has suggested that the energy-dependent step in protein degradation involves an ATPase component of the protease acting on the protein prior to cleavage of the polypeptide chain. First, ATP hydrolysis and peptide bond cleavage are not coupled stoichiometrically or mechanistically, since the ATPase activity is activated by protein binding independently of peptide bond cleavage, and increasing the rate of peptide bond cleavage occurs upon nucleotide binding but does not require ATP hydrolysis (Fischer & Glockshuber, 1993; Thompson & Maurizi, 1994). Second, the


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ATPase components confer specificity, which suggests that productive interaction with the substrate should first occur at a recognition site in the ATPase domain. Third, the structures of the proteases suggest that the openings to the proteolytic sites are blocked by the ATPases and only unwound polypeptide chains can use the limited access channels. With Lon, ClpAP, and the proteasome only relatively short peptides, which presumably can pass easily through the access channels, can be degraded without ATP hydrolysis. 4.2. ATP Hydrolysis Drives Unfolding by Lon Protease Recent studies have shown that the presence of secondary structure in a substrate determines whether ATP hydrolysis is needed for its degradation. CcdA, an F plasmidencoded protein that is a physiological substrate for Lon (Van Melderen et al., 1994), is a small (72 amino acid), dimeric protein with an extended structure composed of ~55% helix (Van Melderen et al., 1996). CcdA is degraded by purified Lon protease, and degradation at 25–37°C requires ATP hydrolysis (Van Melderen et al., 1996). A variant of CcdA, CcdA41, which is composed of the carboxy-terminal 41 amino acids, has some of the in vivo and in vitro activities of CcdA, but lacks stable secondary structure. CcdA41 is degraded by Lon at a similar rate and cleaved at the same peptide bonds as CcdA; however, CcdA41 degradation is activated by non-hydrolyzable analogs of ATP and ATP hydrolysis does not alter its rate of degradation. Furthermore, with CcdA at 50° C, which is the melting temperature for the secondary structure, degradation by Lon also proceeds with non-hydrolyzable analogs of ATP at =25% of the rate with ATP. These data suggest that ATP hydrolysis is used to drive disruption of the secondary structure of protein substrates. The ability of ATP-dependent proteases to disrupt protein structure is limited, and they can degrade only proteins that have less stable tertiary structures or have been “denatured” to some extent, for example, ccasein, which exists in a molten globule state. Proteins that have stable quaternary or tertiary structures are generally resistant to proteolysis even by ATP-dependent proteases. Susceptible proteins should have a structure that we can describe as being “deformable,” meaning that the energy barriers between various conformational states of the protein bound to the protease are relatively low. ATP hydrolysis is required to drive the transitions between states. Presumably the protease then has a mechanism for recognizing and trapping the more extended, unfolded states needed to allow the polypeptide chain to be translocated into the proteolytic sites. 5. INVOLVEMENT OF GENERAL CHAPERONES IN PROTEIN DEGRADATION 5.1. Decreased Protein Degradation in General Chaperone Mutants 5.1.1. General Chaperone Mutants in E. Coli In E. coli, mutations in the major heat shock regulator, , lead to lower expression of Hsps, including both chaperones and proteases, and result in defects in the degradation of “abnormal proteins” such as truncated polypeptides produced by disruption of translation


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with puromycin and misfolded proteins formed by incorporation of the arginine analog, L-canavanine (Baker et al., 1984; Goff et al., 1984). These abnormal proteins, as well as several mutant proteins, are also degraded more slowly in E. coli mutants with defects in major chaperones such as DnaK, DnaJ, GrpE, or GroEL (Jubete et al., 1996; Kandror et al., 1994; Keller & Simon, 1988; Sherman & Goldberg, 1992; Straus et al., 1988). A significant portion of the proteins whose degradation is affected by chaperones are targeted by ATP-dependent proteases (Jubete et al., 1996; Straus et al., 1988). When degradation of individual canavanine-containing proteins was analyzed by pulse-chase labelling and one-dimensional SDS-PAGE electrophoresis, the same proteins were affected by both lon and dnaJ mutations (Jubete et al., 1996). Mutations in lon or chaperone genes affect degradation of specific mutant proteins as well. Lon-dependent degradation of the temperature sensitive PhoA61 protein was almost completely blocked by a mutation in dnaK (Sherman & Goldberg, 1992), and a mutation in groEL blocked degradation of an abnormal fusion protein, CRAG, which appears to be a substrate for ClpP (Kandror et al., 1994). Curiously, CRAG degradation in vivo is not affected by mutations in either ClpA or ClpX, suggesting either that this protein is degraded by ClpP itself or that degradation by ClpP can be activated by a more general chaperone (Kandror et al., 1994; Kandror et al., 1995). No in vitro data supports the latter suggestion however. CRAG degradation was also correlated with the intracellular levels of trigger factor (Kandror et al., 1995). These results demonstrate that various enzymes that catalyze protein remodeling can act in concert with ATP-dependent proteases to affect the rate of protein degradation in E. coli. 5.1.2. E. Coli Chaperones Allow Protein Degradation by Preventing Aggregation General chaperones could affect protein degradation by 1) promoting an active conformation of the pro tease or 2) affecting the accessibility or the conformation of the protein substrate. Purified Lon and Clp proteases are active in vitro (see above) in the absence of chaperones such as DnaK or GroEL (Goldberg et al., 1994; Maurizi et al., 1994). In vivo, natural substrates of Lon, such as SulA and N protein, were degraded at the same rates in wild type and in dnaJ mutant cells (Jubete et al., 1996), and a LacZ fusion protein targeted by ClpA was also degraded in a dnaJ mutant (Maurizi, unpublished). Thus, chaperones are not essential to promote active conformations of either of these proteases, which recognize at least some substrates that are not bound to and have not been modified by chaperones. Chaperones may facilitate protein degradation by preventing or reversing aggregation of potential protein targets. In E. coli cells, degradation of RcsA, a specific substrate of Lon protease, is dependent on a functional DnaJ (Jubete et al., 1996). In cell extracts from dna/mutants, RcsA and SulA are found in an insoluble fraction, whereas in dnaJ + cells, the major fraction of these proteins is soluble even when they accumulate to nonphysiological levels in lon mutant cells (Jubete et al., 1996). Thus, DnaJ contributes to RcsA and SulA degradation by maintaining the protein in a non-aggregated state. DnaJ’s effect on abnormal protein turnover may reflect a similar mechanism. A large fraction of abnormal canavanine-containing proteins is found in an insoluble form, presumably


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inclusion bodies. In wild-type cells, degradation is accompanied by a redistribution of the canavanyl proteins from the insoluble to the soluble fraction. In dnaJ mutants there is reduced degradation and an increase in insoluble protein with time. ClpB/Hsp104, which has a protein disaggregating activity, can also affect degradation of proteins in vivo. T4 lysozyme expressed in E. coli cells is unstable but degradation is blocked in clpB mutant cells, and the accumulated lysozyme is found in inclusion bodies (Inoue & Rechsteiner, 1994). Since ClpB does not activate proteolysis directly, these data further indicate that chaperones can promote degradation of a protein by preventing its aggregation. 5.13. Defects in Protein Degradation in Eukaryotic Chaperone Mutants In eukaryotic cells, degradation in mitochondria, in the endoplasmic reticulum (ER), and in the cytosol is also dependent on molecular chaperones. In vitro synthesized proteins taken up by isolated mitochondria but mistargeted to the matrix are degraded by PIM1 protease, the yeast Lon homolog (Suzuki et al., 1994). Degradation of these abnormal proteins is reduced in the absence of Mdj1 or mtHsp70 (Wagner et al., 1994), yeast mitochondrial homologs of E. coli DnaJ and DnaK, respectively. Thus, there is a functional interaction between yeast mitochondrial chaperones and ATP-dependent proteases. Protein that accumulates in PIM1 mutants wild-type for the chaperones is found in the soluble fraction. However, in the chaperone mutants, the protein is found either bound to mt-Hsp70 or in insoluble aggregates (Wagner et al., 1994). Protein degradation is also defective in yeast mutants lacking the specific cytosolic chaperone, Ydj1 (Lee et al., 1996; Yaglom et al., 1996). Degradation rates of abnormal proteins containing azetidine carboxylate as well as ubiquitin-proline- -galactosidase and Gcn4- -galactosidase, which are subject to ubiquitin-dependent degradation, were reduced 50–80% at the non-permissive temperature in a temperature sensitive Ydj1 mutant (Lee et al., 1996). Degradation of other ubiquitin-dependent substrates (namely, the N-end rule substrates, arginine- or leucyl -galactosidase and the B-type cyclin Clb5 -galactosidase) were not affected, indicating that the chaperones act on specific substrates or on specific components of the degradative pathway. Ydj 1 mutants are also defective in degradation of native forms of the Gl cyclin, Cln3 as well as -galactosidase fusions to this protein (Yaglom et al., 1996). For Cln3, degradation is preceded by phosphorylation, which was also blocked in the YDJ1 mutant (Lee et al., 1996; Yaglom et al., 1996). Unassembled immunoglobulin (Ig L) light chains are degraded in an intracellular compartment, probably the ER (Beggah et al., 1996). Degradation of these chains is dependent on interaction with BiP, a DnaK/Hsp70 homolog found within the ER, and degradation of variants of the Ig L chains was inversely proportional to their rates of release from BiP (Beggah et al., 1996). 5.2. Pleiotropic Effects of Chaperone Mutations Several factors contribute to the extreme difficulty of interpreting the effects of general chaperones on protein degradation in vivo; these cautions should be kept in mind when considering the results of in vivo studies. 1) Chaperone mutants are generally sick and do not grow well, especially at temperatures above 32°C; consequently, extragenic


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suppression of phenotypes is common, though poorly documented. 2) Unfolded protein accumulating in chaperone mutants leads to induction of other chaperones and also ATPdependent pro teases. 3) The accumulated abnormal protein in chaperone mutants can be expected to interfere with the activity of other chaperones and proteases. 4) Proteins targeted by chaperones and proteases are structurally heterogeneous and prone to aggregation, thus the stability of two different proteins may be affected oppositely by mutations in chaperones. Although chaperones appear to facilitate degradation of proteins, particularly abnormal ones, examples of chaperone mutations enhancing protein degradation have also been reported. In E. coli, a temperature sensitive mutant of LacI is degraded more rapidly in dnaK mutants than in dnaK + strains (Keller & Simon, 1988), and the catabolite regulatory protein (CRP) is degraded more rapidly in a dnaJ mutant than in dnaJ + cells (Ohki et al., 1992). It is possible that the chaperone mutations lead to induction of a protease that targets these proteins, but it is more likely that the chaperones are required for proper folding or oligomerization of the proteins and that in the absence of the chaperones, the misfolded or unassembled proteins are susceptible to proteolysis. 6. SUBSTRATE SELECTION BY ATP-DEPENDENT PROTEASES AND CHAPERONES 6.1. Degradation Motifs ATP-dependent proteases probably do not have unique specificity, although different proteases show a high degree of selectivity towards different substrates. The basis for selectivity may lie in sequence or structural motifs recognized by the proteases, or it may reflect extrinsic factors such as localization, the presence of competing substrates, or the participation of auxiliary proteins. Protein degradation in eukaryotic cells (other than that occurring in mitochondria) depends on ubiquitin conjugated proteins being uniquely recognized and degraded by the 26 S proteasome (Hershko & Ciechanover, 1992). However, the existence of the ubiquitin-dependent degradation simply redirects the question of selectivity to the ubiquitination system- how are proteins targeted by the ubiquitination enzymes? That question remains to be answered. In prokaryotic cells, proteases interact directly with substrate proteins. Two degradation motifs have been proposed: 1) bulky hydrophobic amino terminal residues (the N-end rule) (Varshavsky, 1992), and 2) alanine-rich hydrophobic carboxy terminal sequences (Keiler et al., 1996; Parsell et al., 1990). Other degradation signals, such as PEST sequences (Rogers et al., 1986) and “degradation boxes” (Glotzer et al., 1991) have not been shown to occur in rapidly degraded proteins or to influence degradation in prokaryotic cells. Proteins with the N-end degradation signal (degron) are degraded by ClpAP, but this signal is not obligatory in ClpAP substrates (Gottesman et al., 1990a; Thompson & Maurizi, 1994; Tobias et al., 1991). Conditions that give rise to proteins bearing the N-end degron are not known. Proteins with the alanine-rich tail degradation signal are targeted by several proteases, including ClpXP and ClpAP (Gottesman et al., 1998), but again the substrate range of these proteases is not limited to proteins carrying that motif. A specific alanine-rich tail (AANDENYALAA) is enzymatically linked to


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nascent polypeptide chains whose synthesis cannot be completed because their messenger RNAs have been truncated (Keiler et al., 1996), which represents an elegant example of signal transduction in a degradative pathway. Further confounding this issue is the observation that the carboxy-terminal amino acids of MuA, a substrate for the ClpX chaperone activity, can be transferred to a heterologous protein causing it to become a substrate for ClpXP (Levchenko et al., 1997). Although some ClpX or ClpXP substrates have carboxy terminal regions similar to that of MuA, there are at least three dissimilar classes of ClpX substrates based on carboxy terminal sequences, suggesting that ClpX might have binding sites for several different motifs. Carboxy-terminal sequences could constitute part of or all of specific recognition motifs that target a limited number of proteins for degradation by ClpXP. Alternatively, high specificity of interaction is reserved for specialized functions (such as disassembly of the Mu strand transfer complex). Degenerate versions of these sequences might be found in various proteins, sometimes at the carboxy terminus or sometimes, as in RpoS, not. When exposed, these sequences may help to target proteins for degradation by ClpXP. 6.2. Exposed Bonding Domains as Potential Recognition Motifs Proteins targeted for rapid degradation generally have some structural perturbation. The truth of that statement for many unstable proteins (proteins containing amino acid analogs, proteins mistargeted to cells or organelles, various fusion proteins, thermally unfolded proteins) is self-evident. This generalization may apply equally to rapidly degraded normal proteins, which include many important cellular proteins whose degradation is an integral part of regulating their activity. All of the proteins known to be rapidly degraded in vivo function as parts of high molecular weight protein or proteinnucleic acid complexes. In a number of cases, the proteins are stabilized by interaction with their functional partners and degraded when they are free, or, when the protein is degraded from a complex, it is usually capable of forming several different complexes and only one such complex is targeted. Unsatisfied bonding domains in abnormal proteins and in the free subunits or in exposed regions of certain hetero-oligomeric complexes of normal proteins are the target sites for ATP-dependent proteases. Such bonding sites generally have an amphipathic character, but otherwise do not have a defined structural or chemical composition for all proteins. 6.3. Do Chaperones and Proteases Recognize Similar Features of Proteins? There is much in common between the unstructured proteins recognized by chaperones and proteases. Lon protease, GroEL, and DnaK can bind simultaneously to the same immobilized abnormal protein (Sherman & Goldberg, 1992), and immunoprecipitation has been used to show a soluble complex containing an unstable abnormal protein, DnaK, and Lon protease (Sherman & Goldberg, 1991). Since the chaperones could be released from the complex with ATP but not non-hydrolyzable analogs, the binding appears to reflect normal interactions with unfolded proteins (Sherman & Goldberg, 1991). Whether chaperones and proteases bind to exactly the same sites or whether they recognize different exposed regions in abnormally folded proteins is not known. Studies of chaperones binding to peptides of 6–13 amino acids has revealed some


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positional preferences but no real consensus for high affinity binding. For E. coli DnaK and other Hsp70s, peptides with a core of 4–5 hydrophobic residues, with at least one (often central) leucine and basic residues at the ends, tend to bind best (Gierasch, 1994; Rüdiger et al., 1997b) (see chapter Buchberger et al., this volume). Crystal structure data on DnaK with a bound peptide indicate that the peptide backbone as well as side chains of the bound substrate make contact with the binding pocket (Zhu et al., 1996). Sequence comparisons indicate that the overall character of the binding pocket of DnaK is conserved among other Hsp70 proteins, particularly in the central hydrophobic binding position. However, as pointed out in a recent review (Rüdiger et al., 1997a), there is some variation in the disposition of specific residues that make up the subsites in the binding cavities and there are differences in the surface charge distribution at the sides of the binding pockets, which may account for differences in specificity of peptide binding between these proteins. Further thermodynamic and structural studies should bring the issue of substrate recognition by Hsp70 proteins into clearer focus. 7. CONCLUDING REMARKS The critical elements in intracellular proteolysis are 1) the ability to recognize abnormal structural features of proteins and to bind tightly to a region of the protein, probably exposed amphipathic stretches, and 2) the ability to unfold the bound protein to an extended conformation that can penetrate the access channel to the proteolytic site. The first element is common to both chaperones and ATP-dependent proteases, although there is a range of specificities, and consequently affinities, between different chaperones and proteases for particular structural or chemical features exposed in individual proteins. The unfolding activity is also common to both chaperones and proteases, but there is an additional property of the proteases that with a high frequency drastically alters the outcome of the unfolding reaction. Proteases have a means of holding on to proteins and directing them into the proteolytic active sites. Once the degradation pathway is selected, no significant amounts of partially cleaved protein are released and the protein is degraded into peptides of 5–15 amino acids. We can propose several models to explain how proteases might trap proteins for degradation. (1) A “flypaper” model. In this model, the protease has multiple interaction sites for proteins and peptides. A protein that interacts with the protease will bind at one or more of those sites depending on the extent to which it is unfolded and, with time, will interact with additional sites unless it can rapidly assume a stable folded structure. If the interaction sites line a channel leading to the proteolytic sites, the bound protein will move by a simple facilitated diffusion mechanism, similar to that proposed for transport of proteins through membrane channels. ATP hydrolysis by the protease could cause the “sticky” side to alternate with a “non-stick” side allowing the protein to migrate from one site to another. Such a change in binding character in response to ATP hydrolysis has been shown to occur in the protein binding cavity of GroEL (Roseman et al., 1996). (2) A “trapdoor” model. In a trapdoor model, a channel in the protease is lined with


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flexible structures that can bend inward to allow passage of a portion of the protein. In response to nucleotide hydrolysis, these structures could close around narrow regions of the protein and, because they cannot be bent backwards, not allow the protein to withdraw. One major difference between this and the previous model is that it does not require interaction sites to line the access channel into the proteolytic sites. (3) A “clamping” model. The crystal structure of DnaK suggests that it can fold around the polypeptide chain of a substrate, essentially clamping the protein to the chaperone (Zhu et al., 1996) (see chapter Buchberger et al., this volume). The clamp might be formed in one of a few stable conformations of a chaperone/protease, and transitions from one state to another could be facilitated during nucleotide binding, hydrolysis, and release. A clamping mechanism might apply to proteases such as Lon, which undergoes substantial conformational change upon protein binding and for which protein binding results in a large activation of ATP hydrolysis. (4) A cork-screw mechanism. A substrate anchored to a point on the ATPase could be moved through the access channel in a kind of cork-screw motion by rotation or ratcheting of the ATPase subunits with respect to the protease subunits. The symmetry mismatch between six-membered rings of ClpA and seven-membered rings of ClpP allows only one pair of subunits to be in direct register at any given time, and conformational changes upon ATP hydrolysis could lead to different pairs of subunits coming into alignment with each catalytic cycle (Kessel et al., 1996). The protein binding properties of the channel would also have to change during the cycles to allow the protein to move through. A possible method of anchoring protein substrates to the protease would be by covalent attachment to a site on the ATPase. No evidence for such an attachment has been found, but it has not been ruled out experimentally. Whichever model applies, the attached protein must be unwound and translocated into the proteolytic active sites. The multiple active sites could serve as additional points of attachment for substrates, and interactions there could serve to reel the extended polypeptide chain into the proteolytic cavity. If ATP-dependent proteases have their own built-in chaperones, how is it that general chaperones also affect degradation? There is no evidence for direct interaction between general chaperones and ATP-dependent proteases, and we think it is likely that chaperones facilitate proteolysis primarily by binding misfolded proteins and preventing their aggregation. General chaperones, such as DnaK and GroEL, are present in much higher concentrations than the proteases and most likely encounter misfolded proteins more frequently than do the proteases. However, the available data do not differentiate between direct (the pro tease interacts with the substrate bound to the chaperone) or indirect (the pro tease interacts with the substrate after release from the chaperone) mechanisms for the subsequent steps. Chaperones and proteases probably recognize similar sites, and a direct mechanism would require at least two interaction sites on the substrate. For substrates that are released slowly from chaperones, proteases may interact with other exposed sites on the chaperone-bound protein and initiate degradation. For proteins that aggregate rapidly, maintaining the misfolded protein at a minimal concentration so it remains free and accessible to proteases may be the kinetically most significant factor. It may be that both mechanisms are at work with different substrates and the difference depends on the kinetics of release from the chaperone and on the


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(1993). A human mitochondrial ATP-dependent protease that is highly homologous to bacterial Lon protease. Proc. Natl. Acad. Sci. USA , 90 , 11247–11251. Watabe, S. and Kimura, T. (1985). Adrenal cortex mitochondrial enzyme with ATPdependent protease and protein-dependent ATPase activities. J. Biol. Chem. , 260 , 14498–14504. Wawrzynow, A., Wojtkowiak, D., Marszalek, J., Banecki, B., Jonsen, M., Graves, B., Georgopoulos, C. and Zylicz, M. (1995). The ClpX heat-shock protein of Escherichia coli, the ATP-dependent substrate specificity component of the ClpP-ClpX protease, is a novel molecular chaperone. EMBO J. 14 , 1867–1877. Waxman, L. and Goldberg, A.L. (1982). Protease La from Escherichia coli hydrolyzes ATP and proteins in a linked fashion. Proc. Natl. Acad. Sci. USA , 79 , 4883–4887. Waxman, L. and Goldberg, A.L. (1986). Selectivity of intracellular proteolysis: protein substrates activate the ATP-dependent protease (La). Science , 232 , 500–503. Wickner, S., Gottesman, S., Skowyra, D., Hoskins, J., McKenney, K. and Maurizi, M.R. (1994). A molecular chaperone, ClpA, functions like DnaK and DnaJ. Proc. Natl. Acad. Sci. USA , 91 , 12218–12222. Wickner, S., Hoskins, J. and McKenney, K. (1991). Monomerization of RepA dimers by heat shock proteins activates binding to DNA replication origin. Proc. Natl. Acad. Sci. USA , 88 , 7903–7907. Wojtkowiak, D., Georgopoulos, C. and Zylicz, M. (1993). Isolation and characterization of ClpX, a new ATP-dependent specificity component of the Clp protease of Escherichia coli. J. Biol. Chem. , 268 , 22609–22617. Woo, K.M., Chung, W.J., Ha, D.B., Goldberg, A.L. and Chung, C.H. (1989). Protease Ti from Escherichia coli requires ATP hydrolysis for protein breakdown but not for hydrolysis of small peptides. J. Biol. Chem. , 264 , 2088–2091. Woo, K.M., Kim, K.I., Goldberg, A.L., Hai, D.B. and Chung, C.H. (1992). The heatshock protein ClpB in Escherichia coli is a protein-activated ATPase. J. Biol. Chem. , 267 , 20429–20434. Wright, R., Stephens, C., Zweiger, G., Shapiro, L. and Alley, M.R. (1996). Caulobacter Lon protease has a critical role in cell-cycle control of DNA methylation. Genes and Dev. , 10 , 1532–1542. Yaglom, J.A., Goldberg, A.L., Finley, D. and Sherman, M.Y. (1996). The molecular chaperone Ydj1 is required for the p34/CDC28-dependent phosphorylation of the cyclin Cln3 that signals its degradation. Mol. Cell Biol. , 16 , 3679–3684. Yoo, S.J., Seol, J.H., Shin, D.H., Rohrwild, M., Kang, M.-S., Tanaka, K., Goldberg, A.L. and Chung, C.H. (1996). Purification and characterization of the heat shock proteins HslV and HslU that form a new ATP-dependent protease in Escherichia coli. J. Biol. Chem. , 271 , 14035–14040. Yu, A., Antonov, V.K., Gorbalenya, A.E., Kotova, S.A., Rotanova, T.V. and Shimbarevich, E.V. (1991). Site-directed mutagenesis of La protease. A catalytically active serine residue. FEBS Lett. , 287 , 211–214. Zhu, X., Zhao, X., Burkholder, W.F., Gragerov, A., Ogata, C.M., Gottesman, M.E. and Hendrickson, W.A. (1996). Structural analysis of substrate binding by the molecular chaperone DnaK. Science , 272 , 1606–1614.

IV. MECHANISMS

19. SPONTANEOUS VERSUS ASSISTED PROTEIN FOLDING RAINER JAENICKE * and ROBERT SECKLER Institut für Biophysik und Physikalische Biochemie, Universität Regensburg, D93040 Regensburg, Germany

1. Protein Structure and Protein Self-organization 1.1. The Protein Folding Problem 1.2. Native, Intermediate and Denatured States 1.3. Denaturation-Renaturation 2. Folding of Small Single-Domain Proteins 2.1. Rate-Determining Late Folding Events 2.2. Early Intermediates 2.3. Multiple Pathways 2.4. Protein Engineering Analysis of Protein Folding 2.5. Chymotrypsin Inhibitor 2 (CI-2) 3. Folding of Domain Proteins 4. Association 4.1. Sequential Folding and Association 4.2. Specificity of Association 4.3. Superstructures 5. Off-pathway Reactions 5.1. Thermodynamics Versus Kinetics 5.2. Productive Versus Non-productive Pathways 6. The in vitro vs. in vivo Issue 6.1. Cytosolic Solvent Parameters 6.2. Folding Catalysts 6.3. Chaperones 7. Practical Aspects 8. Recent Developments and Conclusions 9. References *Corresponding author

Spontaneous versus assisted protein folding

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1. PROTEIN STRUCTURE AND PROTEIN SELF-ORGANIZATION It has almost become a truism for more than a generation of biochemists that the acquisition of the native functional state of proteins is a spontaneous and autonomous process depending solely on the amino acid sequence and its (quasi-) physiological environment. As has been demonstrated by Anson and his colleagues, who pioneered the field long before the three-dimensional structure of a single protein was known, the natural cellular environment can be mimicked by wellchosen buffers. Based on this discovery, the time course of protein folding and association has been investigated in great detail leading to plausible models describing sequential and multiple pathways of protein self-organization. 1.1. The Protein Folding Problem Because of the large size of the molecules and the unresolved problem how to summarize the weak attractive and repulsive interatomic forces in a unique potential function, theorists have been unsuccessful in solving the protein folding code which would allow to translate the one-dimensional amino acid sequence of a given protein into the corresponding unique three-dimensional structure of the polypeptide chain. Thus, the only way to “predict” protein structures still is homology modeling based on sequence alignments and known spatial structures of related proteins. Obviously, this computeraided, knowledge-based approach is postdictive rather than predictive, and the result commonly does not provide us with the high resolution required in order to work out mechanistic details, e.g., of enzyme function or stability. To reach this level, X-ray diffraction of protein crystals and/or NMR analysis in solution are indispensable. The question whether or not the physical state of a protein obtained from crystal data is compatible with its functional state in vivo has been extensively discussed since Kendrew, Perutz and Phillips came up with their first final analyses of the structures of myoglobin, hemoglobin and lysozyme. Hardly any scientific result has ever been challenged to such an extent. The final (fundamentally positive) answer came from comparative studies using model proteins crystallized in different space groups, on one hand, and NMR spectra, on the other. The accumulated evidence allows us to conclude that, under optimum conditions, both methods yield threedimensional structures at atomic resolution that complement each other to the extent that local interactions of functional groups, including single molecules of the surrounding solvent, can be defined. Regarding the formation of the native functional state of protein molecules, the situation is more complicated, because of fundamental differences between the situation in the cell, on the one hand, and optimum conditions of in vitro reconstitution experiments, on the other. One obvious discrepancy comparing in vivo and in vitro folding is that the nascent polypeptide chain may fold cotranslationally, i.e., in a vectorial fashion from its N- to its C-terminal end, whereas folding in vitro commonly starts from the integral protein in its denatured state. This differs from the native state in two ways: first, it represents an astronomically large ensemble of different configurations, and


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second, it is solvated to a higher extent than the native state. Considering the amount of hydrophobic residues present in most soluble proteins, water is a poor solvent. Structure formation in the cytosol or in aqueous buffer systems is driven by this very fact. Making use of heteropolymers with approximately equal amounts of polar and unpolar residues, nature allows solvation to balance by exposing hydrophilic groups to the aqueous solvent, at the same time minimizing the hydrophobic surface (Richards, 1992). Solubilization of the inner core by mixed solvents or altered temperature leads to unfolding (Privalov, 1992). However, it is obvious from the balance of hydrophilic and hydrophobic amino acids in common proteins that complete solvation cannot be accomplished. For this reason it is doubtful whether a polypeptide chain will ever be “fully randomized” (Jaenicke, 1987). Even in the process of translation, the nascent protein is limited in its conformational space because the space-filling properties of the growing polypeptide and its side chains do not allow all / angles in the Ramachandran plot. 1.2. Native, Intermediate and Denatured States Because of the high internal flexibility of the molecules, the detailed structural characterization of unfolded or partially unfolded proteins is difficult. However, X-ray small angle scattering and multidimensional heteronuclear NMR revealed that even in extensively unfolded states, e.g., at high concentrations of chaotropic agents, considerable residual structure remains. This result confirms previous assumptions that local hydrophobic clusters with features of the native state are preserved, at least in equilibrium with less-structured states. There is no evidence for significant long-range structure in the denatured polypeptide chain. In contrast to the multitude of unfolded states, the native state of a given protein is commonly assumed to be well defined within the limits of the B-factors of X-ray analysis or the conformational dynamics calculated from multidimensional NMR. It is clear that there must be a certain range of flexibility for the obvious reason that in many cases proteins serve as multifunctional devices which have evolved, e.g., to bind, transform and release educts and products of metabolic reactions (Huber, 1988). The driving forces guiding the folding polypeptide chain to its final structure are the same next-neighbor and through-space short-range interactions that are responsible for protein stability: H-bonds, hydrophobic and Coulomb interactions, and van der Waals forces (Dill, 1990; Jaenicke, 1991). From the point of view of thermo-dynamics, the minimum of free energy of the native conformation corresponds to the state with maximum packing density connected with minimum hydrophobic surface area and cavity space (Richards, 1992; Matthews, 1995); the entropic driving force in both hydrophobic collapse and subunit assembly comes from water release involving nonpolar residues and ion pairs (Lauffer, 1975; Jaenicke, 1987). Considering the kinetics of the folding process, it is evident that the acquisition of the native state cannot be a stochastic process; rather, folding must proceed along a pathway or a limited number of alternative pathways (Levinthal, 1968; Baldwin & Eisenberg, 1987). Successive kinetic intermediates along the U N transition have been clearly established, indicating that folding is a hierarchical condensation reaction (Figure 1): next-neighbor interactions first form fluctuating native and non-native secondary structural elements; during this step (which occurs in the sub-milliseconds time range)


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kinetic nuclei gain increasing stability. As soon as the intermolecular interactions surpass the thermal energy, the polypeptide chain collapses into a persistent native-like secondary structure which still lacks the close

Figure 1 Hierarchy of protein structure illustrating the levels of selforganization and stability of globular proteins.

packing characteristics and the low hydrophobic surface area of the native tertiary structure. The term “molten globule” has been used to describe these early, partially folded equilibrium intermediates on the folding pathway, which also form at low pH or medium denaturant concentration (Ptitsyn, 1995). Its properties may be summarized as follows: high content of secondary structure, overall compactness with highly mobile aromatic side chains, exposure of hydrophobic surface, tendency to aggregate, lack of highly cooperative (thermal) unfolding, and rapid equilibration with the unfolded state. Evidently, the given characteristics are not well defined. Attempts to distinguish between native-like structures or irregular collapsed unfolded forms have led to much confusion regarding various molten globule “states”; for a clarification of the terminology, see Baldwin (1991). At a later stage of folding, stable tertiary structure is formed by intramolecular shuffling. If the resulting tertiary structure still exposes excessive hydrophobic surface area or specific charge patterns, intermolecular quaternary contacts will finally cause assembly. 1.3. Denaturation-Renaturation The underlying principle that allows us to analyze single steps along the folding pathway of proteins is that most proteins undergo at least partial refolding after preceding denaturation, with the native structure as the final state of the cycle (with N, U, N* as native, unfolded and renatured states, respectively).

(1)


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Experimental evidence proving that the native and renatured states are indistinguishable has been accumulated using all available physical, biochemical and immunochemical methods. Regarding the difference between directionality of translation from the N- to the C-termini, on the one hand, and integral refolding of the complete polypeptide after preceding denaturation, on the other, three sets of experiments may be taken as proof that vectorial and integral folding are compatible: (i) Merrifield synthesis of ribonuclease and other polypeptides from the C- to the N-terminal end has been shown to yield authentic, active proteins (Merrifield, 1993). (ii) Truncation experiments, as well as limited proteolysis of domain proteins and circular permutations of domains clearly indicate for a variety of systems that removing N- or C-terminal extensions, and altering the succession of domains may leave the overall structure (and function) unchanged (Jaenicke, 1987, 1996). In the case of the domain constructs, this conclusion is in accord with the concept of independent “folding-by-parts” (Wetlaufer, 1981) which has long been considered as one solution to Levinthal’s paradox (1968) that the self-organization of proteins would require an astronomical period of time if folding were the stochastic exploration of the conformational space for the global minimum of potential energy, (iii) Immobilization of proteins at their N- or C-terminal end has been used as another means to invert the direction of folding. Tagging polypeptide chains (with oligo-glutamic acid or oligoarginine extensions fixed at their natural N- or C-termini) to an ion-exchange resin has been shown to allow successful denaturation/renaturation of proteins both in the N→C and C→N direction; in both cases, the products of reconstitution were equally active, showing identical physical properties after release from the column (Stempfer et al., 1996). In summary, translation and denaturation/renaturation yield identical final products so that, as a working hypothesis, in vitro renaturation experiments may be assumed to mimic the cellular process of protein self-organization. There have been occasional reports of proteins which—for unknown reasons—are inaccessible to refolding. Examples are glutamate dehydrogenase (Müller & Jaenicke, 1980; West & Price, 1988), blood coagulation factor XIIIa (Rinas et al., 1990), and creatinase (Schumann & Jaenicke, 1993). For most proteins, optimization of in vitro renaturation has been shown to reach high yields (Jaenicke, 1987). In comparing these results, the protein in its initial state is used as 100% reference. In this context, it is commonly assumed that stucture formation in vivo yields exclusively native protein. However, a number of observations clearly indicate that there is misfolding and misassembly in the cell (Hurtley & Helenius, 1989). Obviously, protein secretion or translocation through the endoplasmic reticulum (ER), and subsequent specific degradation provide an inherent quality control, leading to an apparent yield of 100%. Under unbalanced physiological conditions, in vivo folding, like in vitro reconstitution, gives rise to wrong conformers which may reduce the yield


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considerably, in limiting cases down to 0%. This holds true especially for overexpressed recombinant proteins which, as a consequence of kinetic partitioning between folding and aggregation, may be trapped at early stages of folding, ending up in inclusion bodies (Mitraki & King, 1989; Rudolph, 1990). The fact that proteins, due to their low free energy of stabilization, are basically close to their denaturation transitions, implies that the native state in vivo may occupy a whole set of substates where “misfits” are continuously removed by proteolysis. Thus, again, the yield of (re-)folding does not represent a significant difference between structure formation in vivo and in vitro. The following discussion focuses, first, on spontaneous folding, reviewing the various levels of the hierarchy of protein structure and self-organization. Second, rate-limiting steps and their enhancement by folding catalysts will be considered. Finally, we shall return to the question how far in vitro experiments do provide insight into the cellular processes of structure formation, and at which point the classical reductionist’s approach of physical biochemistry needs to be revised in order to cope with the complexity of the multicomponent multiparameter system of the living cell. 2. FOLDING OF SMALL SINGLE-DOMAIN PROTEINS The elucidation of the folding path of any well-chosen protein would require the complete description of the nascent (unfolded) and final (native) states, together with all intermediates along the Un→N transition: (2) where Un is the ensemble of unfolded conformations, Ii a series of intermediates in sequential order, and N the functional native state. Due to (i) the structural degeneracy of Un, (ii) the limited time resolution and (iii) the question whether there is one sequential pathway or multiple pathway funnelling in a complex energy landscape, this formidable task has not been solved for any protein so far. The most detailed mechanisms which have been worked out in the past, refer to small single-chain one domain proteins with and without cystine crossbridges. Considering the best-known standard systems: basic trypsin inhibitor (BPTI), ribonuclease (RNase A and RNase T1), cytochrome c, hen eggwhite lysozyme, chymotrypsin inhibitor CI-2, and barnase (Figure 2), each has been a role model for a particular facet of the folding problem. This indicates that there is no consensus pathway of protein folding, except for the above mentioned general idea of hierarchical condensation. Evolution seems to have selected individual solutions to the problem how a nascent polypeptide chain can reach the native state, facing the kinetic competition between folding, association, chemical modification and degradation. A couple of examples may illustrate the present state of the art. 2.1. Rate-determining Late Folding Events There are two slow reactions along the folding pathway which serve two quite different


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functions: the oxidation of cysteine residues to form cystine crosslinks contributes to stability; the cis-trans isomerization of proline residues serves as a switch providing the necessary local flexibility of the polypetide chain in late shuffling processes during tertiary structure formation. Due to the enormous number

Figure 2 Architecture of model proteins used in protein folding studies. The regular secondary structure elements of globular proteins may consist entirely of -helices, like in myoglobin (a), or -strands, like in.the fast folding cold-shock protein from Bacillus subtilis (b). Globular proteins may be built from alternating helices and strands, like the / -barrel family of proteins (c, triose phosphate isomerase), but most of them contain helices and -sheets in less symmetrical


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arrangements (d-k). Most of the small proteins depicted here (d, pancreatic trypsin inhibitor BPTI; e, ribonuclease A; f, chymotrypsin inhibitor 2; g, lysozyme; h, ribonuclease T1; i, barnase) have been model subjects of studies on protein folding and stability. A view on a subunit of a larger protein, lactate dehydrogenase (LDH, j), discloses its two-domain nature, with the nucleotide binding domain in the upper right, and the substrate binding domain in the lower left. The same subunit is shown in bold lines in k as part of the complete LDH tetramer. Figures were prepared with MolScript (Kraulis, 1991). using coordinates taken of protein data bank entries 1MBO, 1NMG, 1YPI, 4PTI, 1ROB, 2CI2, 1LZ1, 1RNT, 1RNB, and 1LDN.

of combinatorial alternatives in crossbridge formation, on one hand (Anfinsen & Scheraga, 1975), and the high activation energy of proline isomerization, on the other, both reactions are catalyzed in the cell (see below). Early experiments to probe the conformational transitions during stucture formation following disulfide cross-linking of bovine pancreatic trypsin inhibitor (BPTI), were the first successful approach to unravel the folding pathway of a small protein (Creighton, 1992). Optimizing the technique by acid quenching (to block SH/SS exchange reactions) and HPLC (to separate intermediates), it turned out that the intermediates accumulating in the folding pathway contain a subset of the correct cystine crosslinks (Weissman & Kim, 1991). X-ray crystallography, two-dimensional 1H-NMR, and circular dichroism showed that intermediates with one or two disulfides exhibit a compact conformation that is very similar to that of the native protein (Eigenbrot et al., 1990; van Mierlo et al., 1994). The redox system necessary to introduce the last crosslink via disulfide exchange can be provided by a cysteine residue in the pro-sequence of the BPTI precursor which takes care of the high local SH-concentration required for the reaction with the sterically inaccessible cysteine residues in the major two-disulfide intermediates (Weissman & Kim, 1992). In the cell, oxidative folding is strongly accelerated and the accumulation of BPTI folding intermediates is prevented by protein-disulfide isomerase (Creighton et al., 1993). Because of the required oxidation potential in the cell the reaction takes place only in the ER or in the periplasm. As a consequence, hardly any proteins with disulfide bonds are found in the cytosol. Using ribonuclease (RNase A) as a representative example for a single-chain protein with the size of an average domain, denaturation/reduction and subsequent controlled reoxidation experiments gave the first proof for the one-to-one relationship of the primary and tertiary structure of proteins. However, with 4 disulfide bonds, i.e., 105 possible combinations of SH groups and 7193 mixed disulfide intermediates, no folding mechanism could be established. Instead, the oxidized (native) enzyme was chosen as a paradigm in order to identify rate-determining slow steps on the folding pathway (Anfinsen & Scheraga, 1975). The primary aim was to explain the apparent discrepancy of thermodynamic two-state and kinetic multi-state behavior which clearly pointed to a kinetic scheme with at least three states:


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(3a)

(3b) (with U=unfolded state, Us, UF=slow and fast folding species, and N=native state). The explanation on the basis of Brandts hypothesis that the isoenergetic cis-trans isomerization of proline residues might be involved in the two-state transition (Brandts et al., 1975), has been confirmed by a wide range of experiments (Schmid et al., 1993). With respect to spontaneous versus assisted folding, the essential point is that proline cistrans isomerization, next to disulfide crosslinking, is the second rate-limiting process in the overall folding reaction of single-chain proteins. Its significance is high as indicated by the findings that (i) the nascent trans-conformation of the peptide bond is isomerized to cis in approximately 7% of all prolyl

Figure 3 Protein folding kinetics studied by NMR spectroscopy. NMR spectra of -lactalbumin, a structural homolog of lysozyme, recorded at varied time increments after initiation of refolding. The positions of the peaks reflect the chemical environment of methyl and methylene protons. Reproduced from Balbach et al. (1995) with permission.

residues in native proteins (in RNase, two out of four prolyl residues are in cisconformation) and (ii) there is now clear evidence that proline cis-trans isomerization plays a role in protein folding in vivo (Fischer, 1994; Rassow & Pfanner, 1996). In connection with the question of how the acquisition of the native-like conformation and disulfide formation are interconnected, RNase T1 mutants were used to demonstrate


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that low stability and high flexibility are essential for correct oxidative protein folding (Frech & Schmid, 1995). 2.2. Early Intermediates RNase has been the first paradigm in the elucidation of protein folding, applying pulsed hydrogen exchange in conjunction with NMR spectroscopy (Udgaonkar & Baldwin, 1990; Kim & Baldwin, 1990; cf. Figure 3). Present evidence shows that the -sheet is formed rapidly and cooperatively, shortly after the start of folding. In contrast to a simple sequential model, its stability increases over the first 0.4 s so that one must assume that “the early intermediate” actually represents a broad distribution of species which gradualy changes until side-chain interactions lock the fluctuating polypeptide chain in the stable -sheet conformation. Similar “funnelling” behavior has been observed for other proteins (see below). Based on the observation that the folding mechanism corresponds to the reverse unfolding reaction (Kiefhaber & Baldwin, 1995), the questions whether partially folded intermediates are significantly populated on the folding pathway and how they look like, has also been analyzed starting from the well-defined N-state of RNaseA rather than the degenerate Un states. Monitoring circular dichroism and chemical shift dispersion (pulsed hydrogen-deuterium exchange), the start of unfolding is accompanied by a wide-spread unlocking reaction of side chains which become free to rotate even though the peptide hydrogen bond network still remains intact. The faster exchanging protons correspond to “weak points” in the three-dimensional structure; they are found mainly at the ends of the -strands. Early intermediates on the folding path are difficult to characterize because of their short lifetime. Spectroscopic approaches (UV absorbance, fluorescence, CD), as well as protein-chemical methods (limited proteolyis, antibody binding, H-D or H-T exchange) are commonly restricted to the analysis of global structural changes with a relatively low time resolution, having in mind that the rates of helix-coil transition and -structure formation are in the s time range. However, initiating the folding reaction by a light pulse may shift the time resolution to the s or even ns range, measuring the change in absorption of an intrinsic chromophore to monitor the kinetics. Cytochrome c (Cyt c) has been the model case for this approach (Jones et al., 1993), using the ns photo-dissociation of the heme-CO complex to trigger the folding of the unliganded enzyme. By monitoring time-resolved absorption spectra, transient binding of native and non-native axial amino-acid ligands is observed in the s time range, i.e., before folding begins. Obviously, folding of Cyt c is preceded by the formation of transient loops of the polypeptide chain with non-native contacts between residues or clusters of residues. This is what has been hypothesized earlier as “collapsed form” of the folding protein which would subsequently relaxe or shuffle to reach the native tertiary structure. Another approach to unravel early folding events, with broader application but lower time resolution (ms), has been 2D NMR spectroscopy with rapid pulse labeling, using stopped-flow multimixing techniques combined with the trapping and subsequent identification of exchangeable amide protons (Udgaonkar & Baldwin, 1990; Roder &


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Elöve, 1994). Since structure formation is accompanied by reduced exchange rates, the method provides a direct measure of secondary structure or core formation upon folding. Early results seemed to corroborate the above mentioned heterogeneity of protein fractions and folding kinetics, this time in the seconds time range. However, avoiding kinetic barriers along the folding path by using proper solvent conditions, the time constant for Cyt c folding, was found to be reduced to essentially the same rate as the molecular collapse (< 15 ms) in a single kinetically unresolved step, i.e., without populating observable intermediates (Sosnick et al., 1994). This observation is in contrast to the view that particular steps in protein folding (including the supposedly rate-limiting molten globule to native transition) are intrinsically slow. Instead, it appears that folding intermediates may be kinetically trapped by barriers that are optional rather than integral to the folding process. Major barriers may result from misfolding of the polypeptide chain in the initial collapse step. Fast folding is not unique to cytochrome c: chymotrypsin inhibitor-2, ubiquitin, RNaseA, the Ig binding domain of streptococcal protein G and the cold-shock protein from Bacillus subtilis have been shown to fold at similar rates (Baldwin, 1995; Jaenicke, 1996). Whether the fast rate in all these systems depends on the fact that the (re-)folding polypeptide chain, in collapsing to its early intermediate state, avoids non-native ligations needs further experimental verification. In the case of the refolding of cytochrome c, protection against NH proton exchange shows multiphasic kinetics with at least three phases. Taking the NH exchange rates as a measure, the N- and C-terminal helices are formed within ca. 20 ms, whereas most of the other NH protons follow in the 200 ms time range; the slowest exchange rates (10 s) are attributable to groups involved in tertiary contacts. It is important to note that not only the formation of the helices at the N- and C-terminal end is an early event, but also their specific docking. Being the most stable structural elements in the native molecule, this clearly contradicts the idea that cotranslational (“vectorial”) folding from the N- to the Cterminal end of the nascent polypeptide chain might be decisive for the folding pathway and/or the final structure of the functional protein. In contrast to helix formation in cytochrome c, in the case of ubiquitin and the cold-shock protein from B. subtilis, structure formation has been shown to be involved in very fast early folding events (Briggs & Roder, 1992). On the other hand, the all- interleukin 1 b refolds slowly, with a half time of about 20 min (at 4°C) (Varley et al., 1993). Evidently, there is no clear correlation between either the size or the stability or the structural type of a protein and its folding rate. 2.3. Multiple Pathways Egg lysozyme has been an ideal model for protein folding because it is similar to RNase regarding its size and crosslinking pattern, with two important differences: (i) lysozyme does not contain cis proline so that folding of the oxidized protein is not perturbed by the rate-determining late folding events; (ii) the enzyme shows certain properties of a twodomain protein, in spite of the fact that the two lobes that constitute the “active cleft” do not represent contiguous stretches along the amino acid sequence. Strong evidence that two independent “domains” can be differentiated came from X-ray data at elevated


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pressure (Kundrot & Richards, 1987). In accordance with the topological “ - and -domains” both pulse labeling H-D exchange experiments and 2D NMR allow two regions of the enzyme that differ in their NH protection to be distinguished (Mirankar et al., 1991). Thus, the kinetic mechanism contains an intermediate in which one lobe has reached its compact native-like state, whereas the other is still unfolded or in the process of folding. A more detailed analysis (monitoring H-D exchange by mass spectrometry) indicated that the enzyme does not fold in a single cooperative event. Instead, different parts of the structure become stabilized with different kinetics, the a-helical domain folding faster than the -sheet domain. Furthermore, different populations of molecules fold by kinetically distinct pathways, so that the results force us to believe that folding of oxidized lysozyme is not a simple sequential reaction, but involves parallel alternative pathways (Evans & Radford, 1994). Comparing the kinetics of the reduced polypeptide chain, it has been shown that decreasing the structural constraints leads to a drastic decrease in the folding rate (Goldberg & Guillou, 1994). Thus, the question remains whether the cystine crosslinks may be important determinants in the above kinetic mechanism. In summarizing the previous results, there is clear evidence that folding intermediates with native-like structure accumulate. The fact that they are not well synchronized seems to contradict the classical sequential framework model (Kim & Baldwin, 1990). Theories of the folding process suggest that neither the folding pathway nor the set of folding intermediates is unique, and that folding intermediates only accumulate because they are trapped kinetically by partial misfolding. This view resembles the jigsaw puzzle model (Harrison & Durbin, 1985) which, in the past, has been neglected because experimental data give clear evidence that there are folding pathways with successive intermediates and hierarchical order. Further experiments are needed in order to distinguish between the classical and the new view. As has been pointed out, one important issue in this context is the consideration of secondary structural elements as a major factor determining the folding pathway. One may assume that robust and fast folding pathways have been selected through evolution. Thus, random misfolding in computer simulations may not necessarily apply to the folding behavior of real proteins (Baldwin, 1995). 2.4. Protein Engineering Analysis of Protein Folding With barnase, a 100 amino acid, single-domain extracellular RNase from Bacillus amyloliquefaciens, protein engineering has entered the analysis of protein folding (Fersht, 1995a). As the enzyme contains -helices and an antiparallel -sheet, but no cysteine and cis-proline, it offers itself as a paradigm for folding studies on a stable protein without covalent crosslinks. In using kinetic and equilibrium unfolding/refolding measurements on more than 60 mutants, structure formation in transition states and intermediates was mapped. The fundamental assumptions that the mutations neither perturb the structures of the folded and unfolded states nor the folding pathway, and that the target groups make no additional interactions with partners in their spatial environment, were studied by measuring the structure and stability increments of the standard states and the first significant transition states of unfolding. The structure of the transition state (resulting from urea denaturation) is that of the native-like enzyme, with


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its hydrophobic core weakened and several of the tertiary interactions and loops lost, but with the majority of the secondary structure elements, including tight turns, maintained. This implies that the last events in folding must be the consolidation of the hydrophobic core, the closing of loops and slight rearrangements of tertiary contacts, without any indication for parallel pathways. The same approach as in the case of the transition state for unfolding was applied in order to characterize the structure of an intermediate on the refolding pathway (for details, cf. Jaenicke, 1996). This time, the order of events is: early formation of the N-terminal -helix, the -sheet, part of the core, and docking of the Cterminus to this “nucleus”; subsequent steps stabilize the rest of the core and the loops, with the tertiary contacts as the coda. The theory has been confirmed and supplemented by independent evidence taken from H-D exchange NMR studies. The general conclusions may be summarized as follows: There is a compulsory pathway of folding which is, at least in part, sequential. Secondary structure formation is driven by the local minimization of hydrophobic surface area; it precedes tertiary structure formation. Tertiary interactions become increasingly defined as water release consolidates the hydrophobic core. 2.5. Chymotrypsin Inhibitor 2 (CI-2) Chymotrypsin inhibitor 2 from barley is a 64-residue, single-domain protein of mixed secondary structure. Protein engineering of CI-2 has been employed to map the interactions characterizing the transition state of its folding reaction (Fersht, 1995a). This reaction proceeds rapidly and without a populated intermediate for those CI-2 molecules having all proline residues in the native trans conformation at the onset of refolding. For a large number of mutant proteins with amino acid substitutions scattered throughout the sequence, folding and unfolding rates were extrapolated to zero denaturant concentration and compared to the extrapolated free energy of folding determined in equilibrium unfolding experiments. In addition, amino-terminal CI-2 fragments comprising 5–63 of its 64 residues as well as a nicked form of the protein and its point mutants have been examined by equilibrium and kinetic methods. Major conclusions are the following: (i) the CI-2 folding reaction conforms to a nucleation-collapse model, where residues in an -helix form a local nucleation site, around which all of the secondary structure collapses in a single cooperative process; (ii) the transition state of CI-2 folding is like an expanded form of the native structure without fully formed secondary structure elements; (iii) the nascent CI-2 polypeptide chain has very little tendency to fold before completion of its synthesis (Fersht, 1995b). 3. FOLDING OF DOMAIN PROTEINS Beyond a certain limiting length of the polypeptide chain, proteins consist of domains which may be considered as independent folding units giving rise to separate phases in the process of folding/unfolding. In the simplest case of two-domain proteins, this may be described by a three-state model


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(4) where I represents an intermediate with one domain still intact and the other unfolded (Figure 4A). If the whole molecule represents one single cooperative unit, the population of I remains undetectable, and Eqn. (3a) is sufficient to analyze the equilibrium transition.

Figure 4 Folding of multidomain and multisubunit proteins. A, equilibrium unfolding of bovine B-crystallin, as observed by fluorescence spectroscopy (◯) and analytical ultracentrifugation (●) at pH 2. The amino-terminal domain is stably folded whereas the carboxy-terminal domain is denatured in the intermediate populated around 3 M urea. B, reactivation of LDH after acid denaturation is speeding up with increased protein concentration, until a first-order folding reaction becomes rate-limiting in the uni-bimolecular reaction scheme. The addition of the coenzyme has no effect on the reactivation kinetics.

As taken from in vitro translation experiments, independent domain folding takes place also in the cell: using immunoglobulin and serumalbumin, it has been shown that intradomain cystine-bridges are formed sequentially during translation (Bergman & Kuehl, 1979; Peters & Davidson, 1982). Essentially, folding-by-parts must be considered a most significant acquisition of evolution for a number of reasons: (i) it enhances the folding rate by synchronous folding at multiple sites along the nascent polypeptide chain, (ii) it is a most efficient way to exclude wrong intramolecular interactions in the case of large protein molecules, (iii) it protects the nascent polypeptide chain from proteolysis, and (iv) it may be considered a simple mechanism to proceed from monomeric to multimeric proteins by domain swapping (Bennett et al., 1995; Schlunegger et al., 1997; see below). Regarding the folding mechanism, domain proteins may be considered as the sum of their constituent parts; this means that what has been discussed in connection with the sequence of consecutive steps in single-domain proteins, holds unchanged. In the overall kinetic scheme, the rate-determining reactions are of first order. Depending on whether the reaction is two-state or three-state, one or two pairs of rate constants will be detectable. There are examples where biological function requires the cooperation of domains,


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e.g., in dehydrogenases which contain NAD- and substrate-binding sites residing in two separate domains (D1, D2). In such cases, domain folding may precede domain pairing as the rate-determining step, according to

(5) In cases where proline isomerization or other slow reactions participate in the overall mechanism, the kinetic mechanism will lead to schemes of even higher complexity (Garel, 1992). Using intrinsic markers (fluorophores, epitopes for monoclonal antibodies, ligands), sequential folding steps may be resolved on the time scale from a few milliseconds to seconds, in accordance with the sequential model. 4. ASSOCIATION In advancing from domain proteins to protein assemblies, we approach cellular substructures and finally the macroscopic world: surface layers, the microtrabecular lattice, the tubulin-dynein system, flagella, ribosomes, the extracellular matrix, muscle, are all selfassembly systems involving proteins or protein conjugates. From the point of view of the structural hierarchy of proteins, oligomerization corresponds to domain pairing, except that the docking process is dominated by non-covalent interactions. 4.1. Sequential Folding and Association Model reactions simulating quaternary structure formation made use of proteolytic fragments. In certain cases they were found to exhibit high specificity of subdomain or domain interactions which allow them to recognize and complement each other. In general, the association process is entropy-driven as a consequence of water release from the subunit interfaces. There may be a significant contribution to protein stability from quaternary stucture formation (Lauffer, 1975; Jaenicke, 1991). Complementation requires the correct recognition sites to be preformed; this means that fragments or domains that are expected to trigger the assembly process, must fold autonomously. The overall reconstitution can then be visualized as a sequential folding-association reaction, where folding provides the correct docking surfaces allowing the consecutive association reaction to take place. At low concentrations, association becomes rate-determining (Jaenicke, 1987). Thus, quaternary structure formation may affect both the stability of proteins and their rate of folding. In describing the complete association pathway, the steps preceding subunit docking are the same as in domain proteins. The overall mechanism consists of three stages: first, formation of elements of secondary and super-secondary structure, second, collapse to subdomains and domains, ending up with structured monomers, and third, association to form the correct stoichiometry and geometry of the native quaternary structure. Evidently, the “collision complex” of the structured monomers may still undergo intramolecular rearrangements in order to reach the state of maximum packing density


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and minimum hydrophobic surface area. Thus, the uni-bimolecular folding/association mechanism may involve further first-order steps belonging to slow shuffling processes at the level of the native-like assembly. Focusing on the rate-limiting steps, in the simplest case for a dimer, the overall reaction would then obey a uni-bi-uni-molecular reaction according to

(6) with M, M as unfolded and structured monomers, N as the native dimer and k1, k2, as first- and second-order rate constants (Jaenicke, 1987; Garel,1992). How the single steps along the folding/association pathway can be monitored depends on the specific structure-function relationship for a given system (cf. Figure 4B). In most cases, biological function relates to the native quaternary structure such that the final ratedetermining step can be measured by the regain of activity. Preceding steps may be accessible to spectral analysis, cross-linking and a wealth of other methods (Rudolph et al., 1996). 4.2. Specificity of Association Considering the crowding of a great variety of different components in the cell, one important aspect of protein folding and association is the specificity of subunit recognition, i.e. the question of whether or not other proteins may interfere with the formation of the correct native quaternary structure. A qualitative criterion for the fidelity of subunit recognition was gained from renaturation experiments in crude mixtures where reactivation in the presence of excess foreign protein can be considered a direct measure of correct quaternary structure formation. For example, in refolding recombinant antibodies, no significant differences in the yield as well as the kinetics are observed in the homogenous system compared to the crude mixture obtained upon braking up the E. coli cell (Buchner & Rudolph, 1991). A quantitative investigation made use of pairs of topologically related enzymes. Applying strictly synchronized reactivation conditions, neither hybrid intermediates nor chimeras as endproduct were detected (Gerl et al., 1985). As has been mentioned, the same high degree of specificity holds at the level of domains (Wetlaufer, 1981; Opitz et al., 1987). Clear evidence that hybrid formation may occur comes from isoenzymes, (e.g. the five isoforms of lactate dehydrogenase, LDH), or from multifunctional enzymes where certain gene products have been found as subunits in different complexes (e.g. protein disulfide isomerase (PDI) in prolyl hydroxylase). LDH isoenzymes show exceedingly high structural homology leading to complementary subunit surfaces. In cases in which isoenzymes in different compartments are involved, both target sequences and different folding mechanisms may contribute to specificity. For example, in the case of dimeric mitochondrial and cytosolic malate dehydrogenases, the second-order subunit assembly of m-MDH is preceded by slow folding (which may even be retarded by the signal sequence), whereas for c-MDH first-order reconstitution indicates diffusioncontrolled association (Jaenicke, 1987). It might be attributed to these mechanistic


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differences that all attempts to trap hybrid dimers of MDH, either in the process of reconstitution or in equilibrium experiments failed (Jaenicke, unpublished results). 4.3. Superstructures So far, the relationship of folding and association was discussed mainly considering oligomers. However, successful attempts to reconstitute highly complex biological systems such as phage or the ribosome or giant multienzyme complexes have clearly demonstrated that, in going from oligomeric to multimeric structures, no fundamental differences occur (Jaenicke, 1987; 1996). Regarding their structural analysis, unpredicted progress in these fields has been accomplished thanks to technical developments in the fundamental methods, i.e., electron microscopy, X-ray analysis and NMR. The greatest impact in this connection came from cryo-electron microscopy, synchrotron radiation, the perfection of the image plate and the analysis of tractable structural elements. There are good prospects to fit these elements so that insight into the structure-function relationship up to the level of complex macroassemblages may be gained. Evidently, increasing numbers of subunits lead to a wealth of topological variants which may require specific assembly programs for their regulation. Available data show that cells contain more or less all platonean bodies as compact core structures; in addition, there are hollow cylindrical or helical rods and shells and, at the other extreme, one- and two-dimensional assemblies. A wide variety of methods have been used to elucidate assembly pathways: X-ray crystallography; electrone microscopy; chemical cross-linking, hybridization, recombinant and mutant techniques etc. An example will be discussed in connection with the in vivo and in vitro assembly of the tailspike protein of Salmonella bacteriophage P22. 5. OFF-PATHWAY REACTIONS 5.1. Thermodynamics vs Kinetics The problem of off-pathway reactions in the process of protein folding relates to the questions whether the folded structure is determined by its thermodynamic stability, and how it is possible for a polypeptide chain to fold up along a pathway that allows rapid folding and, nevertheless, arrive at the thermodynamic most stable structure. The compromise may be a consequence of natural selection, insofar as evolution may have selected sequences that have the ability both to fold rapidly and to arrive at thermodynamically stable structures. However, there are two challenges to this hypothesis: First, proteins uncapable of reversible unfolding/refolding and, second, proregion dependent folding of (pre-)proproteins or proenzymes. To illustrate the first point, the kinetic partitioning between the assembly and aggregation of the tailspike protein (TSP) of Salmonella typhimurium bacteriophage P22 may serve as an example (cf. Figure 7). TSP is a homotrimer which is noncovalently bound to the neck of the virus capsid and essential for phage adsorption to the bacterial host. The protein has served as a model system for the folding and assembly of large


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multi-subunit proteins. Its folding pathway comprises subunit folding, followed by the formation of a protrimer, in which the chains are stably associated but not fully folded, and a final shuffling reaction from the protrimer to native TSP as the rate-limiting step (Goldenberg & King, 1982; Fuchs et al., 1991). Along the whole reaction sequence, offpathway aggregation competes with proper folding and association. Numerous point mutants enhancing or suppressing aggregation at high temperature have been isolated, proving that single amino-acid substitutions may profoundly affect the partition ratio between the two competitive processes (Mitraki & King, 1992). The in vitro refolding and the thermal stability of a number of these mutant proteins have been characterized in detail, with the general result that the physicochemical observations precisely complement the genetic analysis (Danner & Seckler, 1993). On the basis of the in vitro results and the recent high-resolution structure of the N-terminally shortened protein (Steinbacher et al., 1994), the folding pathway and the high thermal stability of the tailspike protein, as well as the effects of the folding mutations can be rationalized (Beissinger et al., 1995). In connection with the effect of the pro-region of cymogens on the final structure of the processed molecules, the kinetic competition between on-pathway reactions leading to the native state, and non-productive pathways leading to aggregation gains importance. In this context, pro-regions could function either by increasing the rate of the forward folding reaction or by decreasing the rate of aggregation. Chaperones are known to suppress protein aggregation; in contrast, pro-regions function by directly increasing the rate of the forward folding reaction. They are required for folding under conditions in which off-pathway reactions are suppressed; in addition they interact strongly with the product of the folding reaction, i.e., the native state of the processed protein. For example, denatured a-lytic protease, subtilisin, carboxypeptidase A, among others, do not fold to the native state in the absence of the pro-region. Instead, upon removal of the denaturant, the protein folds to a stable intermediate with substantial secondary structure but little organized tertiary structure; upon addition of the pro-region, the intermediate is rapidly converted to the native state (Winther et al., 1994; Baker & Agard, 1994). 5.2. Productive vs Non-productive Pathways As has been mentioned, there are three stages where side reactions on the folding path may compete with proper folding and association of proteins: the hydrophobic collapse, the merging and swapping of domains and the docking of subunits (Figure 5). An example for the first was discussed in connection with the initial phase of cytochrome c folding where transient interactions of non-native and native amino-acid ligands to the heme iron were found to precede correct folding; cases illustrating the second and third are the domain swapping in crystallins, on one hand, and inclusion body formation, on the other. At all three levels, correct folding requires specific substructures to be preformed in order to proceed on the correct folding path. Collapse and domain merging involve intramolecular rearrangements. Due to the high local concentrations of the reacting groups, they are not significantly affected by neighboring molecules, i.e., they obey first-order kinetics with the slowest isomerization reaction determining the overall rate. In the case of domain proteins, the relative stabilities of the


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domains and the contributions of the domain interactions to the overall stability are crucial. The significance of the linker peptide connecting two well-defined domains has been studied by grafting experiments, e.g., by mutually exchanging the linkers of - and -crystallins (Mayr et al., 1994; Trinkl et al., 1994; Jaenicke, 1994). In both cases, domain contacts dominate over

Figure 5 Side reactions of protein folding. Polypeptide chains may become trapped during folding in conformations representing local minima on the multidimensional conformational landscape (A, reproduced from Mirny et al. (1996) with permission). At high protein concentration, interactions between structural elements maybe formed intermolecularly rather than intramolecularly, leading to off-pathway aggregation (B).

subunit contacts. At concentrations up to 0.5 mM, the recombinant separate domains do not interact with each other, which stresses the above local concentration argument. In going from single-chain domain proteins to protein assemblies, kinetic competititon of first-order folding and second-order association becomes important as soon as the protein concentration reaches the level where folding becomes rate-limiting (Zettlmeissl et al., 1979) (Figure 6). As has been discussed, the reason for this is that subunit assembly requires the monomers to be close to their proper conformation before they coalesce to form the native quaternary structure. If folding intermediates expose wrong contact sites, they will give rise to aggregation instead of association, because the folding polypeptide chain does not distinguish between intra- and intermolecular interactions. Accordingly, not only a shift in the kinetic mechanism but also a decrease in yield of active protein will be observed. The occurrence of inclusion bodies rather than soluble protein in overexpressing recombinant genes illustrates the consequence. The underlying kinetic mechanism

(7)


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resembles the previous sequential uni-bimolecular mechanism which represented the limiting case at high dilution. With increasing protein concentration, when k1

Figure 6 Kinetic competition between folding and aggregation. A, The reactivation yield of LDH (●) decreases with increasing protein concentration, concomitant with an increase in the amount of aggregated material ( ). There is little difference in the specific circular dichroism (□) between renatured LDH and small aggregates proving that the aggregates contain much secondary structure (compare Figure 5 B). B, The protein concentration dependence of LDH renaturation yields (●, data from A) can be modeled as the result of a kinetic competition between unimolecular folding and bimolecular aggregation, as indicated by the solid line (Kiefhaber et al., 1991).

determines the overall rate, the limiting value for k2 will correspond to diffusioncontrolled association. If this explanation is correct, and if the native protein is in a lower energy minimum than the aggregates, it should be possible to manipulate the kinetic competition by adding weakly destabilizing agents. In fact, it has been shown, that the yield of reconstitution may be improved, e.g., in the presence of moderate concentrations of arginine or urea (Rudolph, 1990). Considering Eqn. (7), three questions need to be answered: (i) what is the committed step in aggregate formation, i.e., at which stage along the sequential reaction are aggregates formed, (ii) when is the structured monomer committed to end up as the native protein, and (iii) what is known about the structure of aggregates and their constituent polypeptide chains? With respect to the first two problems, commitment to aggregation was shown to be a fast reaction, whereas the kinetics of the commitment to renaturation follows precisely the slow kinetics of the overall reaction (Goldberg et al.,


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1991). This means that early, collapsed intermediates are much more sensitive to aggregation than later species, but only the native state is fully protected from misassembly. Regarding the structure of aggregates, electron microscopy and circular dichroism indicate that wrong subunit interactions give rise to irregular networks with a broad distribution of highly structured particles at least 10 times the size of the native proteins. They resemble the native protein in its spectral properties, as far as perturbations by turbidity allow a quantitative analysis (Figure 6). Inclusion bodies may represent surprisingly homogeneous precipitates, often with less than a dozen components. Whether disulfide bonds contribute to their compactness in the cell, or whether they are preparative artifacts is still controversial (Mitraki & King, 1989; Valax & Georgiou, 1993). 6. THE IN VITRO VS. IN VIVO ISSUE Aggregation in the cell causes inclusion body formation, because of the high local concentration of folding intermediates, i.e., for the same reasons that are responsible for aggregation in vitro. From this parallelism one may conclude that there is no difference between folding in vitro and folding in the cell, except that overexpression might be considered unphysiological and, therefore, atypical for the standard cell. In this context, it is important to note that there is misfolding and misassembly also in the cell (Mitraki & King, 1989; Helenius et al., 1992). Obviously, secretion or trafficking through the ER and Golgi, and subsequent specific degradation provide an inherent quality control, clearing the cell from “misfits”. Thus, the recovery of refolding does not represent a significant difference between structure formation in vitro and in vivo. As has been discussed before, the same holds for the directionality of protein biosynthesis in the cell which is in contrast to the integral folding of the complete polypeptide chain in vitro. As indicated by the successful “renativation” of proteins under a wide variety of conditions, cytosolic solvent conditions, cotranslational and posttranslational modification, transcriptional or translational control etc. do not play significant roles in the folding process. In addition, tertiary structure formation and subunit association have been shown to tolerate a wide range of variations with respect to sequence and chain connectivities: circular permutations of parts of the sequence, chain extensions, covalent joining of subunits, fragmentation, derivatization, hybridization by peptide interchange etc. (Jaenicke, 1993). Obviously, certain core regions of a protein determine the overall topology, whereas peripheral parts of the protein may be altered or even lacking (Privalov, 1994). Regarding the time requirements, nascent proteins acquire their native structure with half-times in the minutes range (at most), whereas in vitro refolding rates may vary in a wide range from seconds to days. One reason is that optimum conditions regarding the yield of refolding do not necessarily represent optimum conditions for the folding rate; in addition, in the cell, accessory proteins are involved in regulating and/or catalyzing the rates of folding and association. That they assist rather than direct folding is generally accepted. To date, protein folding in the cell has been inaccessible to a detailed analysis of its kinetic mechanism and structural intermediates on its folding pathway. Thus, it is still


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unresolved whether refolding polypeptides in vitro proceed via the same pathway as nascent chains released from the ribosome within the cell. In the case of the tailspike endorhamnosidase from Salmonella bacteriophage P22, detailed in vitro refolding studies indicate that the acquisition of the native structure closely resembles the self-organization of the protein in vivo (Figure 7A). The properties of refolding intermediates resemble those of intermediates observed after pulse-labeling in vivo and the rate-determining folding reaction occurs with identical rates

Figure 7 Bacteriophage P22 tailspike endorhamnosidase. A, the folding and assembly reactions of phage P22 tailspike protein upon biosynthesis in vivo and upon denaturation by acid and urea in vitro comprise similar, if not identical, intermediates and occur with identical rates


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under comparable conditions. B, folding yields of the tailpike protein decrease with increasing temperature both in vivo and in vitro and are affected by single amino-acid substitutions, which alter the thermal stability of thermolabile folding intermediates. C, sites of mutations that decrease (black dots) or increase (open circles) folding yields at high temperature are spread through most of the parallel β-helix (Steinbacher et al., 1994), suggesting that this main structural component of a tailspike subunit is formed early in the folding pathway.

both in the test tube and in the bacteriophage-infected cell (Fuchs et al., 1991; Danner et al., 1993). In vitro translation of firefly luciferase has shown that the ribosome-bound polypeptide chain is essentially inactive; activity appears within a few seconds after release of the enzyme from the ribosome. In contrast, the reactivation of chemically denatured luciferase in the cell-free translation system occurs with a half-time of 14 minutes (Kolb et al., 1994), and the reactivation of pure, completely unfolded luciferase in buffer takes many hours (Herbst et al., 1996). Although the reason for such variable refolding rates is still unknown, the results support the idea that the nascent polypeptide chain folds cotranslationally, corrobo-rating earlier results for immunoglobulin chains (Bergman & Kuehl, 1979; Peters & Davidson, 1982). In summary, the resulting three-dimensional structure is the same, in vitro as well as in vivo. Differences refer to the kinetics of folding and association and to the partitioning between folding/association on the one hand, and aggregation on the other. Shifts of the corresponding partition coefficient are caused by the presence of helper proteins or accessory proteins in the cell. They do not interfere with the “central dogma” that the three-dimensional structure of proteins is determined by their amino-acid sequence. Thus, there remain two questions: (i) are there differences in the mechanism of protein folding within and without the cell, and (ii) what are the effects of cytosolic solvent parameters, on one hand, and folding catalysts and chaperones, on the other? 6.1. Cytosolic Solvent Parameters The solvent conditions in common cells vary only in a narrow range, in spite of the fact that in the biosphere life faces a wide range of extremes of physical conditions. In extremophiles, they may require adaptation of the entire cell inventory to temperatures close to the boiling point of water and presssures ~1200 atmospheres, as well as water activities as low as 0.6. On the other hand, the intracellular pH is kept constant, close to neutrality, even in the case of extreme acido- or alkalophiles (Jaenicke, 1991). In connection with protein self-organization, expressing recombinant proteins from extremophiles in mesophilic hosts, or vice versa, provides information with respect to the impact of solvent parameters on protein folding and association. It turns out that alterations in the folding conditions often have surprisingly little effect. An example is the expression of active enzymes from hyperthermophilic bacteria in E. coli where temperature differences between host and guest may amount to = 60ºC (Jaenicke et al., 1996) After complete denaturation, e.g., of glyceraldehyde 3-phosphate dehydrogenase, renaturation at 5–100°C yields the fully active protein indistinguishable from its initial


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native state. From this we may conclude that not only the native tetramer but also the intermediates on its folding pathway must be stable over the whole temperature range. Little is known with respect to other solvent parameters. In the case of halophiles, the few proteins that have been investigated require salt for folding and assembly (Jaenicke, 1991). For non-halophilic proteins, the ionic strength is not as critical, apart from stabilizing Hofmeister effects that may be used to optimize the folding conditions. What is crucial, is the effect of specific structurally or functionally essential ions, either in metallo-enzymes or in proteins stabilized by other ligands. Such individual requirements can easily be quantified in vitro, thus allowing the cellular situation to be mimicked (Jaenicke, 1987). Regarding effects of hydrostatic pressure, no systematic experiments have been performed so far. The same holds for viscosity effects in the crowded cytosol. 6.2. Folding Catalysts As has been mentioned, there are three possible rate-determining steps in the selforganization of proteins: disulfide shuffling, proline cis-trans isomerization and assembly. On the other hand, there are two enzymes, localized in the appropriate cellular compartments to catalyze the first two, and there is a whole “machinery” of chaperones to assist the third. The biological significance of all three reactions has been established, however, their molecular mechanism is still far from being understood. Protein disulfide isomerase (PDIs, in E. coli DsbA) and peptidyl prolyl cis-trans isomerases (PPIs) are abundant and ubiquitous in all organisms, suggesting that disulfide shuffling and proline isomerization are catalyzed both in eu- and prokaryotes. These catalysts are described in detail in other chapters of this volume (Fischer and Schmid; Missiakas and Raina; Freedman and Klappa). Here, we only briefly summarize their activities and discuss their roles in the assisted folding of proteins in a more general sense. PDIs such as DsbA catalyze the oxidative refolding of a number of small substrate proteins by the rapid unidirectional transfer of an active site disulfide to the substrate protein and subsequent disulfide isomerization:

(8)

The catalytic efficiency of the enzyme is evident: adding catalytic quantities of DsbA leads to rapid refolding under conditions where no spontaneous refolding can be achieved in its absence. With respect to substrate specificity, inhibition studies with peptides of various lengths and sequences suggest that PDIs show rather broad peptide-binding capacity. There is clear evidence that DsbA catalyzes protein folding in vitro. It must also act catalytically in the cell since the levels of oxidized substrates can exceed the level of DsbA by almost three orders of magnitude. Being oxidoreductases and isomerases at the same time, the catalytic cycle of PDIs requires reoxidation of the active-site disulfide, making use of low-molecular-weight compounds like oxidized glutathione or of a protein electron acceptor (Bardwell, 1994; Bader et al., 1998).


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As for the isomerase activity, catalysis may be operative at two levels, folding of the nascent chain and rescue of misfolded (and aggregated) polypeptides. The latter has been shown to be involved in the dramatic acceleration of the isomerization reactions necessary for the completion of disulfide formation on the folding pathway of BPTI. Apparently, in certain proteins PDI is capable of gaining access to buried thiol groups after the protein substrate has acquired a substantial percentage of its tertiary structure. In the case of the oxidative renaturation of antibodies, this holds only for the first phase of the reaction (Lilie et al., 1993). If PDI can act late on the refolding pathway, it could help explain why the correct disulfide bonds are made: The proteins have already folded so as to place those cysteines that are to participate in disulfide-bond formation in close proximity, supporting the idea that PDIs are true folding catalysts in the sense that they enhance the rate of folding without determining the final structure. In this context they are involved in both formation and isomerization of disulfide bonds in their specific compartments in the cell. PPIs catalyze the rotation around the X-pro peptide bond. Translation yields the alltrans configuration of the polypeptide chain. In cases where the final structure of a protein contains cis-peptide bonds, the high activation energy EA of their isomerization may give rise to slow folding, depending on the neighboring groups. By far the most frequently occurring cis-rotamer in known protein structures involves proline residues:

(9)

In the case of essential prolines EA is of the order of 85 kJ/Mol, so that at room temperature and in the absence of a catalyst the reaction becomes rate-limiting in the overall folding process (Schmid et al., 1993). A variety of structurally unrelated families of enzymes have been discovered which are capable of catalyzing proline isomerization. Three of them, the cyclophilins (CyPs), FK 506 binding proteins (FKBPs) and trigger factor have been investigated in detail (Fischer, 1994). All three are abundant and ubiquitous proteins involved in a variety of cell biological phenomena. Whether their action is in all cases related to their specific catalytic function as PPIs is still unresolved. The catalytic efficiency of the PPIs depends very much on the accessibility of the Xprolyl bonds involved in the isomerization reaction. For small prolyl peptides, the catalytic efficiency approaches the diffusion-limited second-order rate constant for the association of a protein with a small molecule (Schmid et al., 1993). Little is known regarding the sequence specificity; as in the case of PDIs, the substrate specificity seems to be low. The significance of PPI in catalyzing the in vitro refolding of small model proteins containing essential proline residues is well-established. Concerning the biological significance of PPI catalysis, three observations support the idea that the in vitro results reflect cotranslational or posttranslational folding events in the cell: (i) the striking evolutionary conservation of PPI function, together with the proper loca-tion and ubiquity


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of the enzyme in every organism and subcellular compartment; (ii) PPI has been shown to be involved in mitochondrial protein folding in cooperation with molecular chaperones (Rassow et al., 1995; Matouschek et al., 1995); (iii) in the case of the maturation of procollagen to form the mature collagen triple helix, PPI accelerates the rate by a factor of approximately three, whereas PPI inhibition leads to retardation (Bruckner et al., 1981; Bächinger, 1987). In transferring this kind of experiment into intact cells, it was shown that the time for half-completion of the triple helix in chicken embryo fibroblasts increased significantly upon PPI inhibition; the obvious explanation is that the ratelimiting prolyland hydroxyprolyl isomerizations during the in vivo folding of collagen are catalyzed by PPI. A PPIase associated with E. coli ribosomes has recently been identified as the trigger factor, a protein originally suggested to trigger secretory proteins to adopt a translocation-competent conformation, but recently found to interact with nascent chains of non-secretory as well as secretory proteins (see chapters by Welch et al. and Fischer and Schmid, this volume). Whether its main cellular function is to catalyze proline isomerization or to prevent premature folding and aggregation of newly synthesized polypeptides, remains to be determined. The given results suggest the following folding mechanism: The nascent all-trans polypeptide chain collapses rapidly to form a compact state with elements of secondary structure and sufficient stability to expel water from its hydrophobic interior; subsequent slow steps lead to the native-like state which finally undergoes trans→cis isomerization of proline residues to reach the biologically active, native conformation. Assuming this mechanism to hold in the cell, extrapolation from model peptides would predict protein folding to be much slower than estimated from in vivo studies. There are three explanations why proline isomerization does not necessarily limit in vivo protein folding: (i), trans prolines may be trapped in their initial configuration; (ii), the occurrence of proline residues in solvent-exposed turns causes their state of isomerization to be of minor functional significance, and (iii), constraints of the chain conformation may greatly decrease the energy barrier of proline isomerization, thus speeding up the reaction. These arguments do not consider the idea that PPIs may operate at the site of protein translation, thus enhancing the trans→cis isomerization at defined sites depending on the sequence specificity of the enzyme. Finally, it is worth mentioning that PDI and PPI catalysis occur simultaneously, in a specific and synergistic way (Schönbrunner & Schmid, 1992). Whether a similar synergism operates in the course of the de novo synthesis and folding in the cell remains to be shown. 6.3. Chaperones As has been shown in connection with side reactions competing with folding and association in vitro, the limit at which kinetic partitioning between folding and association becomes significant is commonly far below the average protein concentrations in vivo (Mitraki & King, 1989). Thus, in the cell, mechanisms must be effective that inhibit the unproductive aggregation and, at the same time, promote correct protein folding and association. Molecular chaperones, the components which serve this function, are a large family of unrelated proteins; they block the above side reaction, without being components of the final structures (Ellis & van der Vies, 1991). The


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requirement for “folding helpers” differs, depending on the system; in general, multidomain oligomeric proteins seem to need chaperone assistance for unperturbed assembly. Apart from their involvement in folding, molecular chaperones also limit damage caused by stress conditions such as heat. Accordingly, their cellular level was found to be strongly enhanced under shock conditions; however, not all chaperones are heat shock proteins. It has become clear that molecular chaperones play significant roles in a variety of cellular processes such as protein targeting, translocation through membranes and compartmentation. Since the activities and mechanisms of action of molecular chaperones are subject of many chapters of this volume we will not further discuss their roles in protein folding here. 7. PRACTICAL ASPECTS As has been mentioned, aggregation in vitro and inclusion body formation in the cell correspond to each other. There are various strategies to cope with the problem. Making use of weaker promoters, the local concentration may be reduced. Discontinuous pulse dilution has been devised in order to optimize the yield of soluble protein: a certain amount of the denatured protein is subjected to dilution and reactivated at low concentration; after refolding, new portions of the denatured protein are added in a stepwise fashion until the whole batch is transferred. This way, the concentration of folding intermediates is always kept below the critical concentration of aggregation. Additives such as arginine may strongly increase the yield by shuffling aggregates back on the productive folding path. The repertoire of reconstitution methods has been extended by systematic experiments, applying folding catalysts and chaperone proteins (Rudolph et al., 1997). One might assume that overexpressing a specific protein together with chaperones and folding catalysts would finally yield 100% of the desired protein in its native state. Attempts toward this goal have so far been unsuccessful. Since sequestration of the folding polypeptide chain seems to be the underlying principle of chaperone action, renaturation in reverse micelles and immobilization on solid matrices might be promising alternatives. Successful developments in this direction have recently been reported (Jaenicke, 1995; Stempfer et al., 1996; Tuena de Gómez-Puyou and Gómez-Puyou, 1998). 8. RECENT DEVELOPMENTS AND CONCLUSIONS Fifty years after the protein folding problem was first formulated, there is still no way to predict the spatial structure of a given polypeptide chain from its sequence, nor has any protein been fully described regarding its detailed folding pathway. There are various reasons for that, apart from the insufficient size of the data set, and extrinsic factors not encoded in the primary structure. Practical applications of denaturation/renaturation placed high priority on the cellular aspects of folding. Anfinsen anticipated them at both levels, folding catalysis and folding on a template, i.e., chaperone-assisted folding (Epstein et al., 1963). However, before approaching the problem at the complex cellular


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level, the problem had to be solved under ideal conditions, varying the relevant parameters in dilute solution. The outcome has been the mechanistic concept of multiplepathway sequential folding with very fast early events (hydrophobic collapse), middle events (local shuffling toward the native tertiary structure), and late events determining the rate of the overall reaction; the latter are proline cis-trans isomerization, disulfide cross-bridge formation and subunit assembly. As has been shown, all three are facilitated by accessory proteins, either enzymes or chaperones. From their ubiquity and abundance in all organisms, cells and cell-compartments, the two isomerases may be assumed to be essential for the formation of the native structure of proteins in the cell. In the case of the chaperones, the fundamental importance (beyond the stress response) has been clearly shown, (i) for the kinetic partioning between folding and association on the one hand, and aggregation on the other, (ii) for protein targeting, e.g., translocation of mitochondrial proteins across membranes, and (iii) for processes involved in morphogenesis, such as growth and self-assembly of phages. However, the mechanistic details of their action are still not fully understood. For a brief review on the present state of the art, see Beissinger and Buchner (1998). 9. REFERENCES Anfinsen, C.B. and Scheraga, H.A. (1975). Adv. Protein Chem. , 29 , 205–300. Bächinger, H.-P. (1987). J. Biol Chem. , 262 , 17144–17148. Bader, M., Muse, W., Zander, T. and Bardwell, J.C.A. (1998). J. Biol. Chem. , in press. Baker, D. and Agard, D.A. (1994). Biochemistry , 33 , 7505–7509. Balbach, J., Forge, V., van Nuland, N.A., Winder, S.L., Hore, P.J., and Dobson, C.M. (1995). Nature Struct. Biol. , 2 , 865–870. Baldwin, R.L. (1991). Chemtracts: Biochem. Mol Biol . 2 , 379–390. Baldwin, R.L. (1995). J. Biomol. NMR 5 , 103–109. Baldwin, R.L. and Eisenberg, D. (1987). In: Protein Engineering (Oxender, D.L. and Fox, C.F., eds.). Alan R.Liss, Inc., New York, pp. 127–148. Bardwell, J.C.A. (1994). Mol. Microbiol . 14 , 199–205. Beissinger, M. and Buchner, J. (1998). Biol Chem. , 379 , 245–259. Beissinger, M. Lee, S.C., Steinbacher, S., Reinemer, P., Huber, R., Yu, M.-H. and Seckler, R. (1995). J. Mol. Biol . 249 , 185–194. Bennett M.J., Schlunegger, M.P. and Eisenberg, D. (1995). Protein Sci. , 4 , 2455–2468. Bergman, L.W. and Kuehl, W.M. (1979b). J. Biol. Chem . 254 , 8869–8876. Brandts, J.F., Halvorson, H.R. and Brennan, M. (1975). Biochemistry , 14 , 4953–4963 Briggs, M. and Roder, H. (1992). Proc. Natl Acad. Sci. USA , 89 , 2017–2021. Bruckner, P., Eikenberry, E.F. and Prockop, D.J. (1981). Eur. J. Biochem . 118 , 607– 613. Buchner, J. and Rudolph, R. (1991). Bio/Technology 9 , 157–162. Creighton, T.E. (1992). In: Protein Folding (Creighton, T.E., Ed.), W.H.Freeman, New York, pp. 301–351. Creighton, T.E., Bagley, C.J., Cooper, L., Darby, N.J., Freedman, R.B., Kemmink, J. and Sheikh, A. (1993). J. Mol. Biol . 232 , 1176–1196. Danner, M. and Seckler, R. (1993). Protein Science , 2 , 1869–1881.


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20. PROTEIN DISULPHIDE-ISOMERASE: A CATALYST OF THIOL: DISULPHIDE INTERCHANGE AND ASSOCIATED PROTEIN FOLDING ROBERT B.FREEDMAN* and PETER KLAPPA Research School of Biosciences, University of Kent, Canterbury, Kent CT2 7NJ, UK

1. Introduction 2. Molecular Properties of PDI 2.1. Analysis of PDI Protein Sequences 2.2. Evidence for the Domain Organization of Mammalian PDI 2.3. PDI as a Member of the Thioredoxin Superfamily of Proteins 2.4. Chemical Properties of the Thioredoxin-like Domains within PDI 3. PDI as a Catalyst of Thiol:disulphide Interchange and Protein Folding 3.1. Activity Towards Simple Peptide Substrates 3.2. Activity Towards Well-Defined Protein Substrates 3.3. Activities of Mutant PDIs and Domains 3.4. Does PDI Catalyse Folding or Just the Disulphide-Isomerizations Associated with Folding? 4. Binding Properties of PDI 4.1. Interaction of PDI with Non-Peptides 4.1.1. Interaction of PDI with Thyroid Hormone 4.1.2. Interaction of PDI with Oestrogens 4.2. Interaction of PDI with Peptides and Proteins 4.2.1. Interaction of PDI with Peptides 4.2.2. Interaction of PDI with Proteins 4.2.3. Interaction of PDI with ER Proteins 5. Is PDI a Molecular Chaperone? 6. References *Corresponding author


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1. INTRODUCTION Protein disulphide-isomerase (PDI), an enzyme found in the endoplasmic reticulum of eukaryotic cells, was the first catalyst of protein folding to be identified. In their initial studies on protein refolding in vitro, Anfinsen and colleagues recognised that the conditions required for successful refolding of reduced ribonuclease (high dilution, high pH etc.) were not physiological, and that the time-scales involved (hours to days) were much longer than those for protein folding in the cell (Epstein et al., 1963). They inferred that the process was catalysed in the cell and sought potential cellular catalysts. In the mid 1960s, they and other groups showed that refolding of reduced ribonuclease and other proteins could be catalysed by microsomal preparations from vertebrate secretory tissues such as liver, pancreas etc. (Goldberger et al., 1963), purified the enzyme responsible (DeLorenzo et al., 1966) and proposed a role for it in protein folding in the cell (Givol et al., 1964). The catalytic properties of the purified enzyme from mammalian liver were described over the next 10 years, and were consistent with its being involved in the catalysis of native disulphide bond formation during the folding of nascent secretory proteins. It was 20 years after this initial work on PDI before it was generally accepted that protein folding in the cell is a process mediated and facilitated by cellular factors. By that time, the evidence for the involvement of PDI in cellular catalysis of protein folding was stronger than before, but still circumstantial (Freedman, 1984). PDI was known to be most abundant in cells actively synthesizing and secreting disulphide-bonded proteins and to be located in the lumen of the endoplasmic reticulum, the compartment in which such proteins fold immediately after synthesis (Lambert and Freedman, 1985). Furthermore, in several physiological situations, the pattern of expression of PDI matched the pattern of expression of the disulphide-bonded proteins which are its potential targets (Freedman, 1984). Within the last 10 years, the role of PDI in cellular protein folding has been established beyond doubt by the classical methods of biochemistry, cell biology and molecular genetics. In whole cells, and in cell-free systems for in vitro translation and translocation, PDI can be cross-linked to nascent and newly translocated secretory proteins (Roth and Pierce, 1987; Klappa et al., 1995). In such cell-free systems, the presence of PDI is required for the rapid formation of native disulphide bonds; removal of lumenal resident proteins leads to defective folding which is repaired by reconstitution with purified PDI (Bulleid and Freedman, 1988). Finally, the gene encoding PDI in S. cerevisiae has been cloned and shown to be required for the proper folding, targeting and secretion of disulphide-bonded proteins (LaMantia and Lennarz, 1993; Dunn et al., 1995). More recently, this cellular role for PDI has been demonstrated by a further stringent test which also has practical application. In both eukaryotic and prokaryotic systems it has been shown that overexpression of recombinant PDI (in the ER or periplasm, respectively) can significantly increase production of functional recombinant proteins containing multiple disulphide bonds (Wittrup, 1995; Humphreys et al., 1995). In other words, there are cases where the yield of a correctly folded recombinant protein is limited by inefficient formation of the native disulphide bonds, and this limitation can be

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overcome by co-overexpression of PDI. This chapter reviews the structural, catalytic and ligand binding properties of PDI. It considers, in particular, recent work aimed at establishing the structural basis of how PDI acts as a catalyst of protein folding. 2. MOLECULAR PROPERTIES OF PDI 2.1. Analysis of PDI Protein Sequences The cloning and sequencing of rat PDI cDNA (Edman et al., 1985) immediately led to the recognition of internal sequence homologies within the protein and the

Figure 1 Domain structure of PDI. The border between the a' and c domain is not exactly known.

suggestion of a structural organization based on duplicated sequence modules (see Figure 1). The homologous a and a' sequence modules show significant sequence identity to thioredoxin, and on the basis of this homology, PDI was recognised as a member of a superfamily of proteins containing thioredoxin-like sequences (see 2.3 below). Since the internal sequence repetition does not include the whole of PDI, a more detailed analysis was undertaken (Freedman et al., 1998) in an attempt to define more precisely the sequence modules within PDI and their boundaries. This analysis drew on i) the pattern of sequence conservation in a large set of multiply-aligned PDI sequences from a range of species, ii) the pattern of hydrophobicity/ hydrophilicity in this sequence set, iii) the intron/exon structure of the human PDI gene, and iv) the claim that part of the PDI protein sequence was homologous to part of a steroid receptor sequence. The results of this analysis suggested the existence of up to 6 structural domains (Freedman et al., 1994), but did not define their boundaries unambiguously; the analysis provided the basis for experiments aimed at determining the domain organization of PDI (see 2.2 below), on the assumption that the sequence modules defined within PDI would correspond to structural domains. 2.2. Evidence for the Domain Organization of Mammalian PDI As yet, no high resolution structure has been determined for PDI from any source. Although crystals have been obtained by some groups over the years, they have not yet yielded high resolution x-ray diffraction data. In view of its molecular size—a homodimer of polypeptides each of over 55 kD—PDI remains beyond the range of structure determination by current NMR methods. In the absence of a high resolution


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structure, even quite low resolution information would be helpful. Is PDI organized into clear structural domains? If so, how many are there and where are their boundaries? Can they be obtained as independent folded structures and subjected to high resolution structure determination, so that the overall structure of PDI can be built up domain-bydomain? In an attempt to answer these questions two complementary methods have been used recently by the laboratory of Creighton and by us. First, native purified PDI has been subjected to limited proteolysis, the fragments identified and the data interpreted on the argument that the preferential sites for proteolysis are likely to lie at the boundaries between structural domains. Secondly, putative domains inferred from sequence analysis (see 2.1 above), have been expressed as recombinant polypeptides and their structural properties characterised; the underlying assumption is that a fragment which adopts a stable conformation with secondary and tertiary structure, and cooperative unfolding and refolding behaviour, corresponds to a domain since it behaves as an independent folding unit. Limited proteolysis of bovine PDI by trypsin and by V8 protease indicated a number of preferential sites of proteolysis (Freedman et al., 1998). N-terminal sequencing of derived fragments demonstrated cleavage following R115, R328, E331 and K333 and mass determinations of fragments by mass spectrometry suggested that proteolysis also occurred at E222 and K349. The region from R328 to K349 is clearly highly susceptible to proteolysis and we propose that it is a disordered or exposed region linking the b′ and a′ domains. The proteolysis at E222 probably marks a boundary region between the b and b′ domains. No cleavage sites were found within the region putatively linking the a′ and c domains, suggesting that the acidic C-terminal region of PDI does not form a distinct structural domain. Similarly no sites of cleavage were found at either end of the putative e domain (despite the presence there of residues which are compatible with the cleavage specificity of the enzymes used). Instead, there was preferential proteolysis at R115, in the middle of the putative e domain, and fragments cleaved at this site were stable and could be recovered. This suggests that there is no e domain, and that the cleavage at R115 marks a boundary region between the a and b domains. The conclusions from proteolysis are supported by work on the recombinant expression of putative domains (Freedman et al., 1998). We have expressed each of the putative e, b and b′ domains as fusions with glutathione-S-transferase (GST), recovered the products and released the putative domain from the fusion partner. In parallel, we have generated the putative ‘linked domains’ e-b (101–226) and e-b-b′ (101–335) in the same way. None of the released fragments corresponding to putative individual domains showed significant secondary structure in solution or co-operative conformational change as a function of denaturant concentration. In contrast, after cleavage from the N-terminal fusion partner, both the ‘linked domain’ polypeptides had the properties of folded proteins. Interestingly, in both cases the N-terminal sequence of the polypeptide was not as expected, based on the junction with the fusion partner, but showed that the Nterminus had been trimmed to remove 2 linking residues and part of the putative e domain, leaving the N-terminal sequence corresponding to residues 115–117 in the mature sequence. This result implies that the residues prior to 115–117 do not form part of the folded domain constituted by the remainder of the fragment, and is consistent with


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the conclusions from proteolysis of authentic full-length PDI. A similar approach has been adopted, independently, by Creighton and colleagues (Darby et al., 1996). They found no evidence of structure in recombinant polypeptides corresponding to the e (100–157) or b (148–257) domains alone. They then expressed a polypeptide corresponding to the hypothetical linked e-b domains (residues 100–257) and deliberately subjected it to limited proteolysis by trypsin, V8 protease and thermolysin. Each of these enzymes trimmed the recombinant fragment substantially at both ends; analysis of the products suggests that a core b domain comprises residues 116–218 and the individual proteolysis products extend beyond this, based on the availability of sites consistent with the specificity of the proteases. On the basis of this analysis, a new putative b domain polypeptide was designed and expressed (residues 119–228) which in contrast to earlier constructs (100–157 and 148–257) showed CD, NMR and urea gradient gel electrophoresis properties consistent with being a folded polypeptide. Similarly, recombinant polypeptides corresponding to putative a and a' domains have been expressed and characterised. The results suggest that residues 1–120 and 348–462 both form folded structures when expressed in isolation (Darby and Creighton, 1995b), as do the shorter fragments 5–96 and 347–436 (Freedman et al., 1998). These recombinant polypeptides show some of the functional properties of PDI (see 3.4 below) and hence have been the target for more detailed structural studies. In summary, the work reviewed here reveals a 4-domain structure (a-b-b'-a'), with the inter-domain boundaries falling around residues 116, 220 and 335–350 respectively, plus an ill-defined region extending from residue 462 to the C-terminus. 2.3. PDI as a Member of the Thioredoxin Superfamily of Proteins As noted, sequence similarity between PDI and the small ubiquitous protein thioredoxin immediately suggested a family relationship between them. Thioredoxin is a cytosolic protein which functions as a dithiol reductant in a range of cellular functions. Over time, a substantial number of proteins have been discovered which have sequence and functional similarities to thioredoxin and are recognized as members of a thioredoxin superfamily (Table 1). A larger number of cDNA sequences or genomic open reading frames also are clearly related to this family, even though the corresponding protein has not been isolated or characterised. Some members of the superfamily (thioredoxins, glutaredoxins) are small and of known tertiary structure; they comprise a single domain with a characteristic fold which is also found in a number of enzymes involved in glutathione or sulphur metabolism (Martin, 1995). DsbA, a bacterial periplasmic protein which acts as a disulphide oxidant (see Missiakas and Raina, this volume), also has this fold, but is larger and contains an additional helical domain inserted within the thioredoxin fold (Martin et al., 1993). The structural data on the N-terminal 120 residues of PDI, corresponding to the a domain, confirm that it is a structural domain and has


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Table 1 The thioredoxin/PDI superfamily

Protein

Mr

Structure

Sequence of active site motif

Thioredoxin

11kDa

thioredoxin domain

-CGPC-

Glutaredoxin 12kDa

thioredoxin domain

-CP(Y/F)C-

DsbA

22kDa

thioredoxin domain+helical insert

-CPHC

PDI

55kDa

2 thioredoxin domains+others

-CGHC-

PDI-like ER proteins

50–75kDa 2 or 3 thioredoxin domains+others

-CGHC-CGTC- (PDIp)

the classic thioredoxin fold; residues 1–3 and 117–120 were not fixed in the native state of this fragment confirming that residues 116–117 mark the limit of the a domain (Kemmink et al., 1995, 1996). More recent studies of the global fold of the b domain of PDI by NMR reveal that, like the a domain, the b domain shows the secondary structure pattern characteristic of the thioredoxin fold, despite showing no significant amino-acid sequence similarities to any members of the thioredoxin family. This observation, together with modelling and preliminary experimental studies on the a' and b' regions strongly implies that PDI comprises four sequential domains each with the thioredoxin fold (Kemmink et al., 1997). Interestingly, many cDNA sequences have been cloned from a wide variety of sources showing the existence of gene products which can be inferred to contain two (or three) thioredoxin-like domains. In some mammalian species, four or more of such sequences co-exist (Freedman et al., 1994). The significance of this multiplicity has not been established, but it is natural to assume that it relates either to differences in specificity, or to differently regulated patterns of expression, or to different cellular or sub-cellular location. For example, one member of the family (PDIp) has recently been claimed to be expressed specifically in the acinar cells of the pancreas (DeSilva et al., 1996, 1997), whereas another, ERp57, has been found to interact exclusively with glycosylated secretory proteins after their translocation into the ER (Oliver et al., 1997). It has been shown that this specificity is due to the interaction between ERp57 and calnexin, a glycoprotein-specific chaperone of the ER (Zapun et al., 1998). 2.4. Chemical Properties of the Thioredoxin-like Domains within PDI Early chemical modification studies with alkylating reagents, and inhibition studies by arsenicals and heavy metals, suggested that PDI contained an essential dithiol group which could be oxidised to a disulphide form in the isolated protein (Hillson and Freedman, 1980). Now it is clear that both the a and a' domains of PDI contain such a group with the local sequence…WCGHCK…and that similar vicinal dithiol/ disulphide groups are characteristic of the members of the thioredoxin superfamily. The structural properties of this active site dithiol/disulphide group are known in thioredoxin and DsbA,


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whose structures have been determined to high resolution by X-ray diffraction (Martin, 1995). The motif is found in an exposed turn linking a strand to an extended helix. The sulphur atom of the more N-terminal Cys residue is at the N-terminal pole of the a helix and is exposed at the surface of the molecule, while the sulphur atom of the more Cterminal Cys residue is buried behind it. The chemical properties of this group have been most thoroughly studied in human PDI and in thioredoxin and DsbA from E. coli (see Missiakas and Raina, this volume). The following features are characteristic both of DsbA and the a domain of PDI (Zapun et al., 1993; Nelson and Creighton, 1994; Darby and Creighton, 1995c): (i) the more N-terminal Cys residue of the dithiol group is unusually low in pK and is present exclusively as the -S- thiolate anion at neutral pH, (ii) this group is unusually reactive as a nucleophile both in alkylation reactions and in the formation of mixed disulphides by reaction with, for example, oxidised glutathione (GSSG),

(iii) mixed disulphides formed by the latter reaction are unusually unstable since they are prone to attack by the more C-terminal Cys residue of the active site, leading to displacement of reduced glutathione and formation of the disulphide form of the protein

(iv) the equilibrium constant for reaction between the dithiol/disulphide group of the protein and reduced and oxidised glutathione (GSH/GSSG), via the two partial reactions above, is in the range 1 mM or less, corresponding to a standard redox potential in the range –90 to –180 mV which is unusually high for a protein, indicating that the active site disulphide is unusually oxidising (i.e. weak, unstable). This can be contrasted with thioredoxin where the active site dithiol/disulphide is highly reducing and the equilibrium constant with GSH/GSSG is 10 M. (v) the active site disulphide in DsbA and PDI is so unstable that, unlike in most disulphide bonds in proteins, it actually destabilizes the protein, so that the reduced form of the protein is the more stable to denaturation by urea etc. These chemical properties are clearly interrelated and the fundamental property appears to be the unusually low pK of the exposed thiol group of the N-terminal Cys residue. The structural basis of this can be analysed by comparing the high resolution structures of DsbA and thioredoxin (Martin, 1995); regrettably, a comparable high resolution structure of PDI is not yet available. The steric properties of the active site disulphides in the two proteins are indistinguishable; there are no obvious indications of strain to account for the weakness of the disulphide in DsbA. The low pK must reflect the existence of electrostatic interactions which stabilize the thiolate form of the exposed Cys residue. In both DsbA and thioredoxin, the S atom of the N-terminal Cys is at the N-terminus of an


486

helix and could therefore interact electrostatically with the directed H-bonds along the axis of the helix, which generate an effective ‘helix dipole’. The positive pole of this dipole is at the N-terminus and hence can stabilize an anionic -S- group there, but this applies, in general terms, to both thioredoxin and DsbA and cannot simply account for the difference between them. A major difference occurs between the proteins in the residues within the active site motif (see Table 1); mutation studies in thioredoxin, DsbA and the recombinant a domain of PDI (Lundstrom et al., 1992; Grauschopf et al., 1995; Kortemme et al., 1996) demonstrate that the presence of a His residue at position…CxHC…(as in wild-type DsbA and PDI) has a major effect on the properties of the site, and other mutations in these residues also affect the chemistry and functional properties of the protein (Grauschopf et al., 1995). However, it is unlikely that this is the only source of difference (Warwicker and Gane, 1996). Our electrostatic analysis identified two other factors (Gane et al., 1995): i) the geometry of the helix following the active site is different between thioredoxin and DsbA, leading to a stronger electrostatic interaction with the exposed thiolate group in the latter case, and ii) the presence of the additional inserted domain in DsbA modifies the dielectric properties of the region close to the active site giving rise to stronger electrostatic interactions. It remains to be seen whether similar additional factors exist in PDI to account for the chemical properties of its active site disulphides, and indeed whether there are differences in such properties between the active sites of the a and a' domains. 3. PDI AS A CATALYST OF THIOL:DISULPHIDE INTERCHANGE AND PROTEIN FOLDING 3.1. Activity Towards Simple Peptide Substrates PDI was discovered on the basis of its ability to catalyse reactivation of reduced inactive ribonuclease and has been widely assayed using this and related reactions with protein substrates. However, the underlying processes here are complex, involving a mixture of protein disulphide formation, disulphide isomerization and conformational change. To clarify some aspects of PDI catalysis, it is useful to work with simpler and more defined catalysts. Darby et al. (1994) introduced a model 28-residue peptide containing 2 cysteine residues at positions 2 and 27, analysed thiol: disulphide interchange reactions between it and glutathione by HPLC, and characterised the catalysis of these reactions by PDI and DsbA (Darby et al., 1994; Darby and Creighton, 1995a). The model peptide has no defined secondary or tertiary structure in either the reduced dithiol form or after oxidation to form an intramolecular disulphide; as a result the reactions are ‘pure’ examples of thiol: disulphide interchange and oxidoreductions in a protein-like substrate without being associated with folding or extensive conformational change. In conditions of pH and glutathione concentrations comparable to those within the ER lumen, PDI catalysed (approximately 30-fold) reactions between the reduced peptide and oxidised GSSG leading to formation of both single mixed disulphide species (with glutathione bonded to residues 2 and 27 respectively) and it catalysed (approximately 100-fold) their subsequent disulphide isomerization to form an intramolecular peptide


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disulphide and release reduced GSH. There was little formation of the double mixed disulphide species in the conditions used, nor did PDI effectively catalyse transfer of the glutathione group between one cysteine residue of the peptide and the other. In addition to these reactions, PDI also catalysed the direct oxidation of the peptide to the disulphide state, in a process not directly involving glutathione, but requiring the GSSG-dependent recycling of the resultant reduced PDI. This latter mechanism obviously requires the involvement of the disulphide form of PDI, whereas the former mechanism can be represented as a series of thiol: disulphide interchange steps in which the exposed Cys residue interconverts between the thiol form, and mixed disulphides with glutathione and/ or with the peptide substrate, and the buried Cys residue of the active site is not directly involved. This model peptide system has been most instructive in analyzing basic features of the mechanism of action of both PDI and DsbA. However it is not convenient for routine assay. Recently Ruddock et al. (1996) introduced a related substrate, a small peptide containing two cysteine residues and a single tryptophan residue whose fluorescence emission is quenched when an intramolecular disulphide bond is formed. PDI and DsbA catalysis of glutathione-dependent oxidoreduction using this substrate are comparable to those with the model substrate of Darby et al. (1994) but the overall reaction can be monitored continuously by fluorimetry. 3.2. Activity Towards Well-Defined Protein Substrates PDI is active towards a continuously-growing list of protein substrates, but only a few have been studied in any detail. It was shown many years ago (Creighton et al., 1980) that PDI catalyses the net process of refolding of reduced bovine pancreatic trypsin inhibitor (BPTI) and specifically catalyses processes involving formation or isomerization of protein disulphide bonds together with associated conformational change. Since that time, the folding pathway of BPTI has been characterized in great detail so that the identities and structures of many intermediate species have been determined, and the kinetics and pathways of their interconversion have been established (Goldenberg, 1992; Weissmann and Kim, 1993; Creighton et al., 1995). Although there have been extensive studies with bovine pancreatic ribonuclease and other proteins, BPTI is by far the best-characterised protein system in which to study action of PDI on a protein substrate. In particular, the folding pathway of BPTI involves isomerizations between a number of species containing two of the three native disulphide bonds. The species containing the disulphides 30–51 and 5–55 (30–51, 5–55) is also referred to as N(SH)2 since it has almost complete native folding and rapidly forms the final native disulphide bond (14– 38). However the species N(SH)2 is not readily formed from species containing a single disulphide bond. The major species which accumulate under native conditions are (30– 51, 14–38) and, in some conditions (5–55, 14–38), both of which contain two native disulphide bonds and are extensively folded. These species (termed N' and N* respectively by Weissmann and Kim (1993)) require only the formation of one further native disulphide bond but they cannot do so directly or rapidly since to do so would require extensive unfolding and refolding. Instead they rearrange to form N(SH)2 via


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species (30–51, 5–14) and (30–51, 5–38) and these latter species which contain nonnative disulphides (5–14 and 5–38, respectively) are obligatory intermediates in the folding pathway. These rearrangements provide a stringent test of the activity of PDI since they involve disulphide isomerization with associated conformational change, rather than net formation or breakage of protein disulphide bonds. They can be studied by isolating the intermediates N' and N* and, as suggested by the earlier work, they are strikingly (3500– 6000-fold) catalysed in the presence of excess PDI (Weissmann and Kim, 1993). The dramatic catalytic effect on these reactions found with PDI is not observed with DsbA (Darby et al., 1995a); this bacterial periplasmic relative of PDI appears to act primarily as a net oxidant, a donor of disulphide bonds, rather than as a catalyst of rearrangements. Another contrast is between the dramatic catalysis of the isomerizations between species containing two disulphides (N'→N(SH)2 and N*→N(SH)2), and the minimal catalysis by PDI (2-fold) of the further conversion of N(SH)2 to the native folded protein (Weissmann and Kim, 1993), a process which involves disulphide formation but limited conformational change. Other well-defined systems in which the catalytic activity of PDI can be studied include the oxidation of reduced pancreatic ribonuclease (Lyles and Gilbert, 1991) and the formation of native ribonuclease T1 from both the reduced state (with 4 free thiol groups) and the mixed disulphide state with four bound glutathione groups (Ruoppolo and Freedman, 1995; Ruoppolo et al., 1996). The standard assay for PDI with a protein substrate is its ability to catalyse the reactivation of ‘scrambled’ ribonuclease, a substrate in which the eight Cys residues of bovine ribonuclease have been oxidised essentially at random in denaturing conditions (Freedman et al., 1995). This reactivation clearly requires both disulphide isomerization and protein conformational change, but its details have not been established due to the heterogeneity of the substrate. 3.3. Activities of Mutant PDIs and Domains The possibility of designing and expressing mutant or truncated forms of PDI allows detailed insight into the structural basis of its activity. This is particularly significant in view of the presence of multiple domains and two apparently similar active sites in the a and a' domains respectively. Darby and Creighton (1995b) expressed fragments corresponding to residues 1–120 and 348–462 (the a and a' domains respectively) of human PDI and showed that the a domain comprised a well-folded protein with some disordered residues at each terminus (Kemmink et al., 1995). These constructs (Table 2) were relatively active in catalysing simple thiol: disulphide reactions with no protein folding component. Thus, in the glutathione-dependent conversion of the dithiol to disulphide form of the model 28residue peptide, each showed a specific activity approximately 50–60% that of native full-length PDI. Similarly in the conversion of reduced BPTI to the major two-disulphide intermediate (N'; 30–51, 14–38) each showed activity up to 70% that observed with the full-length enzyme. These results suggest that in these relatively simple reactions, the action of full-length PDI is simply the sum of roughly equivalent activities mediated by each of its a and a' domains.


489

However, a different picture emerges in other assays. The isolated recombinant domains are completely inactive in catalysing the conversion of the N' form of BPTI to the native product, an intrinsically very slow process requiring disulphide isomerization and protein conformational change. Similarly, the recombinant domains only show approximately 10% of the activity of native PDI in catalysing the reactivation of ‘scrambled’ ribonuclease, another process dependent on both disulphide isomerizations and conformational change in the protein substrate. (A slightly lower Table 2 Relative activities of active site mutants of full length PDI and single domain constructs of PDI

Assay

Mutant

Activity (%)

Reference [Species]

sRNase/DTT (SH)

SGHC…CGHC

46

Vuori et al., (1992)

CGHC…SGHC

47

[Human]

SGHC…SGHC

2

CLHS…CGHC

60

LaMantia and Lennarz (1993)

CGHC…CIHS

60

[S. cerevisiae]

CLHS…CIHS

5

SGHC…SGHC

2

Laboissiere et al., (1995)

CGHS…CGHS

92

[Rat]

SGHC…SGHC

–

CGHS…CGHS

3

CGHS…CGHS

18–30

Lu et al., (1992)

SGHS…SGHS

–

Lyles and Gilbert (1994)

CGHC…SGHS

23–50

Walker et al., (1996)

CGHS…SGHS

3–8

[Rat]

SGHC…SGHS

–

SGHS…CGHC

23–27

SGHS…CGHS

9–18

SGHS…SGHC

–

rBPTI/DTT (SS)

sRNase/GSH/GSSG

rRNase/GSH/GSSG

rRNase/GSH/GSSG

Assay

a Domain a′ Domain Activity (%) Reference [Species]

sRNase/DTT (SH)

CGHC CGHC

14

Darby and Creighton

9

(1995b, c) [human]

rBPTI/GSSG/GSH to N′

CGHC

70 CGHC

70

Molecular chaperones and folding catalysts N′ to BPTI

CGHC

– CGHC

Peptide (SH)2 to Peptide (SS)

490

CGHC

– 62

CGHC

54

sRNase, ‘scrambled’ ribonuclease; rRNase, reduced ribonuclease; rBPTI, reduced BPTI; N′, native-like form of BPTI

figure of 3% was determined for this rearrangement activity by Freedman et al. (1998), using more extensively truncated forms of the a and a′ domains, residues 5–96 and 347– 436 respectively). These results strongly suggest that whereas PDI functions as the sum of independent a and a′ domains in simple reactions, its ability to catalyse complex disulphide isomerizations linked to protein conformational change either involves the other domains of the molecule, or depends on specific interactions or co-operation between the a and a′ domains in the intact enzyme (or both). Some light can be thrown on this question by studying mutant full-length PDI molecules in which one active site has been inactivated by mutation. Several groups have generated such mutations (see Table 2) and assayed the mutants in a variety of activities. The earliest work was that of Kivirikko’s group on mutant human PDIs (Vuori et al., 1992). They used the assay based on ‘scrambled’ ribonuclease and showed that the enzyme could be fully inactivated by mutating to serine the N-terminal Cys residues in the both active site motifs (i.e. in both the a and a′ domains of the full-length enzyme). When each of these mutations is made separately, leaving one wild-type active site and one mutant site, the resulting proteins are essentially 50% active in this assay. Thus a full-length protein with a single functional active site is 50% active in this rearrangement assay, whereas the individual active site domains, when present as isolated fragments are only 10% active (see paragraph above). This strongly implies a role for the other domains of the molecule in isomerization activity. The most extensive set of mutants is that of rat PDI generated by Gilbert and colleagues (Lyles and Gilbert, 1994; Walker et al., 1996) assayed in the reactivation of reduced ribonuclease. Mutants with one entirely wild-type active site (CGHC) and the other converted to SGHS are moderately active (up to 50% of the wild-type enzyme, consistent with the results of Vuori et al. (1992)), although detailed kinetic analysis suggests that the kinetic properties of the a and a′ active sites in the intact protein are distinct. In the other mutants, activity is only detected when the wild-type Cys residue is present in the N-terminal position of the active site motif in at least one of the active sites, confirming the crucial importance of the N-terminal Cys in each site. Interestingly, however, mutation of the more C-terminal Cys residue in each site has more limited effects; the mutant with this mutation in both active sites has 18–30% of the activity of the wild-type enzyme, suggesting that the enzyme can act entirely through thiol: disulphide interchange processes involving the exposed Cys residues only. In qualitative agreement with this result, Laboissiere et al. (1995) find that the double mutant with the sequence SGHC at both active sites is minimally active in rearranging ‘scrambled’ ribonuclease, whereas the double mutant with the sequence CGHS at both


491

sites is almost fully active. In their hands, neither double mutant is active towards reduced ribonuclease, implying that activity towards this substrate requires net oxidation by the disulphide form of the PDI active site. The quantitative discrepancy between this result and the finding of Gilbert’s group may arise from differences in assay conditions and method, but has not been resolved. Similarly the result of LaMantia and Lennarz (1993) on the effect of mutation in the C-terminal Cys in each active site is not immediately consistent with the results of the other groups, but this work used yeast PDI and a different assay. Overall the fragment and mutant data point to two conclusions. 1) The full-length molecule is required for full activity in processes involving disulphide isomerization and conformational change, and full-length molecules with a single functional active site are 50% active in such assays. The individual isolated domains are much less active in these assays, but fully functional in simple thiol: disulphide interchange. This strongly suggests direct involvement of the other domains in the more complex activities. 2) Only the Nterminal Cys residue in each active-site motif is absolutely required for activity, although mutations in the more C-terminal Cys residue have some effect. This implies that the enzyme operates through a mixture of thiol: disulphide exchange and direct oxidation mechanisms, dependent on the substrate and conditions, and that species mutated in the C-terminal Cys are moderately effective in the former mechanism. 3.4. Does PDI Catalyse Folding or just the Disulphide-Isomerizations Associated with Folding? This question cannot be directly answered from the studies reviewed here. In the more complex activities studied to date (e.g. the interconversions of the 2-disulphide isomers of BPTI, and the reactivation of ‘scrambled’ ribonuclease) the disulphide isomerizations and conformational changes are intimately associated. However the fact that these reactions have more stringent requirements for catalysis than do processes simply involving thiol:disulphide chemistry, strongly implies that PDI is directly involved in facilitating conformational changes in its protein substrates. If so, it is difficult to imagine how this could be brought about except by direct contacts between PDI and its substrate proteins during folding, which are more extensive than the transient formation of mixed disulphides at the active site. In view of this argument, it is important to review what is known about the ability of PDI to interact with other proteins, and with peptides and polypeptides which might be models for its interactions with folding substrates. In the following section, we consider first some cases in which a specific ligand-binding activity has been proposed for PDI. Next we review studies on the interactions between PDI and peptides and polypeptides, including protein folding intermediates. Finally we examine the cases in which PDI forms long-term stable interactions with other polypeptides to generate species with other activities; detailed investigation of these “special cases” may illuminate several aspects of the interaction between PDI and protein folding intermediates.


492

4. BINDING PROPERTIES OF PDI 4.1. Interaction of PDI with Non-peptides 4.1.1. Interaction of PDI with Thyroid Hormone It has been shown that a bromoacetylated derivative of 3, 3', 5-triiodo-L-thyronine (T3) can specifically label a protein with a molecular mass of 55kDa in a variety of mammalian cell lines. The nucleotide sequence of this human cellular thyroid hormone binding protein, present in the endoplasmic reticulum, has been determined and the encoded protein was found to be identical to PDI (Cheng et al., 1987; Yamauchi et al., 1987). In another independent approach to identify the type I iodothyronine deiodinase (ID-I), a rat cDNA clone coding for rat PDI has been isolated (Boado et al., 1988). Here, a library was screened with polyclonal antibodies directed against solubilized rat liver microsomes with clones coding for ID-I being selected by an indirect enzyme-binding inhibition assay. However, Schoenmakers et al. showed that ID-I is not identical to PDI, since inhibitors and substrates for ID-I did not prevent the labeling of PDI with bromoacetylated [125I]T3. In contrast, a 27kDa microsomal protein with ID-I activity has been detected (Schoenmakers et al., 1989). They also showed that the binding of bromoacetylated [125I]T3 to PDI was not inhibited by unmodified T3 or 3, 3', 5, 5'tetraiodo-L-thyronine. Since PDI has a high reactivity towards alkylating agents (see above) like haloacetyl compounds the most likely explanation is that PDI reacts only with the bromoacetyl moiety. From the lack of competition between modified and unmodified T3 it is tempting to speculate that PDI has no specific binding affinity for thyroid hormone. 4.1.2. Interaction of PDI with Oestrogens PDI activity has been claimed to be selectively inhibited by certain oestrogens (Tsibris et al., 1989). In this approach the effect of oestrogens on the GSH-dependent degradation of insulin by partially purified PDI of different sources was tested with 17 -oestradiol (17 E2) and oestrone being the most potent inhibitors (Ki 1mM). Chemical crosslinking of [125I] insulin to PDI was partially inhibited in the presence of 17b-E2 suggesting that oestrogens can compete with peptides/proteins for the binding to PDI. Furthermore, it has been proposed that segments of PDI share some sequence homologies with human oestrogen receptor, since residues 101–144 (corresponding to exon 3) and residues 163– 211 of mature PDI are similar to residues 350–392 and 304–349 of human oestrogen receptor, respecively. However, the binding of radiolabeled oestradiol to PDI preparations was not saturable, thus the nature of interaction between oestrogens and PDI remains elusive. Furthermore, the quantitative interpretation of results obtained by chemical crosslinking can be very difficult, while this method provides a powerful tool to study the qualitative interaction between proteins. As for the homology between PDI and the human oestrogen receptor, it has been shown recently by NMR that it is unlikely that any of the PDI segments with similarities to the oestrogen receptor comprise individual


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domains (Kemmink et al., 1995: see above). Taken together, these findings suggest that PDI has no specific binding affinity for oestrogens and related compounds. 4.2. Interaction of PDI with Peptides and Proteins 4.2.1. Interaction of PDI with Peptides Morjana and Gilbert (1991) showed that peptides of various length and amino acid composition can inhibit the GSH-dependent degradation of insulin by PDI. The observed inhibition constants indicate that the affinity of PDI for peptides increases with the length of the peptide. Peptides containing cysteine residues are 4–8 fold better inhibitors than other peptides of comparable length. The Ki ranged from 46 M (20 amino acids containing a cysteine residue) to >10 mM (3 amino acids without a cysteine residue). Apart from the presence or absence of cysteine residues the binding affinity seemed to be determined by the length of the peptide while the amino acid composition was of minor significance. This result was confirmed by Noiva et al. (1991) by using a photo-crosslinking approach. It was found that PDI can be photo-crosslinked to glycosylatable peptides containing the sequence -Asn-Xaa-Ser/Thr-with the binding not restricted to peptides containing this N-glycosylation site. Furthermore, the presence of Bolton-Hunter-reagent (3-[4-hydroxy-5-iodophenyl]-propionate) in the peptides tested, seemed to increase the binding affinities. Subsequently, the binding site on PDI for the peptides was determined by using a photoreactive [125I] tripeptide probe which was photo-crosslinked to purified PDI (Noiva et al., 1993). After fragmentation of the crosslinking product the binding site could not be mapped to the thioredoxin-like active sites of PDI but was found in the Cterminal acidic segment of PDI. However, since the binding affinity of PDI for tripeptides is rather low (see above) the question is whether a tripeptide is a suitable probe. It is conceivable that a tripeptide is too short to target a specific binding site. As mentioned above, it can not be ruled out that the Bolton-Hunter-reagent, attached to a peptide, contributes significantly to the binding. Since this effect was observed to be more pronounced in small peptides, it is important to use longer peptides in order to define the complete peptide binding site of PDI. To characterise the substrate binding site of PDI, we investigated the binding of radiolabelled peptides to PDI by chemical cross-linking. We identified a specific interaction with a number of labelled oligopeptides of 10+ residues in length. The interaction was independent of the method of radiolabelling or cross-linking and appeared to be physiologically significant in that it was saturable, reversible and required native PDI; furthermore binding was competitively inhibited by unlabelled peptides and by misfolded or unfolded proteins, but not by native proteins (Klappa et al., 1997). To locate the site at which this binding occurred, we studied interactions of various recombinant fragments of human PDI, expressed in E.coli, with different radiolabelled model peptides. We observed that the b' domain of human PDI is essential and sufficient for the binding of small peptides. In the case of larger peptides, specifically a 28 amino acid fragment derived from bovine pancreatic trypsin inhibitor, or misfolded proteins, the


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b' domain is essential but not sufficient for efficient binding, indicating that contributions from additional domains are required. Hence we propose that the different domains of PDI all contribute to the binding site with the b' domain forming the essential core (Klappa et al., 1998). 4.2.2. Interaction of PDI with Proteins By crosslinking, it has been demonstrated that PDI can interact in vivo with misfolded human lysozyme but not with correctly folded lysozyme (Otsu et al., 1994). This result clearly shows that i) PDI can interact not only with small peptides but also with proteins, ii) the interaction between PDI and proteins/peptides is not re-stricted to artifical in vitro conditions but can be detected in vivo as well, and iii) PDI can discriminate between correctly and incorrectly folded proteins. In an independent approach, nascent chains of secretory proteins after their translocation into the endoplasmic reticulum were chemically crosslinked to PDI, irrespective of whether the secretory protein contained a Cysteine residue or not (Klappa et al., 1995). The interaction between the probed secretory proteins and PDI was found to be transient and restricted to unfolded or partially folded proteins. These findings confirm the observations of Otsu et al. and indicate that PDI can bind to proteins without Cysteine residues. It is therefore tempting to speculate that PDI is not only involved in the formation and rearrangement of disulphide bonds but is also involved in other processes. This idea is supported by the observations of LaMantia and Lennarz (1993) that disruption of the gene coding for yeast PDI interfered with the viability of the yeast cells while mutations of the active site cysteine residues were not lethal but affected the formation of disulphide bonds. In addition to that, Hayano et al. (1995) could demonstrate that a PDI mutant devoid of its isomerase activity was able to accelerate intracellular folding of human lysozyme in vivo. Taken together, these results suggest that neither the active site dithiol of PDI nor a Cys residue on a protein are absolutely required for PDI/protein interaction. Therefore, PDI might be capable of taking part in the folding, prevention of aggregation, and assembly of proteins without disulphide bonds. This was confirmed by the findings that PDI was able to suppress aggregation of Dglyceraldehyde-3-phosphate dehydrogenase (GAPDH), which does not contain any disulphide bonds, during refolding in vitro (Cai et al., 1994). Similar results were obtained for rhodanese (Song and Wang, 1995). When the active site Cysteine residues were inactivated by alkylating agents, PDI lost its ability to form and rearrange disulphide bonds. However, this PDI was still active with respect to suppressing aggregation of GAPDH (Quan et al., 1995). In contrast, when PDI was incubated together with a peptide containing 28 amino acids without Cysteine residues, the efficiency of PDI to prevent aggregation of and refold GAPDH was remarkably reduced (Quan et al., 1995). The Ki of this peptide inhibitor is in the range of 200 µM, which is in excellent agreement to the numbers obtained by Morjana and Gilbert (1991). PDI therefore might interact with unfolded or partially folded proteins, thus preventing aggregation or incorrect interactions between the folding substrates. The site or sites on PDI involved in such interactions include the b' domain (see 4.2.1) but are not yet fully


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defined. It is also not clear whether this property is exploited in the cell in the folding of proteins lacking disulphide bonds, or whether it is simply an aspect of PDIs ability to catalyse folding in association with the generation of native disulphide bonds. 4.2.3. Interaction of PDI with ER Proteins (a) PDI as a subunit of the oligosaccharyl transferase complex It has been reported that a glycosylation site binding protein, which is a component of the oligosaccharyl transferase, is highly similar to PDI (Geetha-Habib et al., 1988). By using a photoreactive tripeptide containing the sequence -Asn-Lys-Thr- it was demonstrated that PDI can interact with glycosylatable peptides. In contrast, Bulleid and Freedman (1990) observed cotranslational glycosylation of proteins in systems depleted of PDI, which makes it unlikely that PDI is an essential component of the oligosaccharyl transferase. This result was extended by Noiva et al. (1991), who showed that the interaction between PDI and tripeptides is not limited to peptides containing Nglycosylation sites (see above). Purified PDI showed no significant preference for glycosylatable tripeptides. However, when intact microsomes were used, PDI was preferentially labeled by glycosylatable tripeptides. The most likely explanation for this is that by glycosylation these tripeptides were trapped within the microsomes, thus increasing the peptide concentration in the lumen while non-glycosylatable peptides might exit the microsomes by diffusion (Noiva et al., 1991). These findings indicate that PDI is neither a component of the oligosaccharyl transferase complex nor involved directly in the glycosylation of proteins. However, it can not be ruled out that PDI might have some indirect influence on the glycosylation of proteins, e.g. by making the glycosylation sites accessible for the oligosaccharyl transferase. (b) PDI interacting with calreticulin Recently it has been reported that PDI can interact in vitro with calreticulin, a molecular chaperone of the ER involved in the correct folding of glycoproteins (Baksh et al., 1995). Furthermore, it has been shown that interaction between calreticulin and PDI can modulate their activities: calreticulin does not bind Ca2+ with high affinity in the presence of PDI while PDI is inhibited in the presence of calreticulin with respect to folding of ‘scrambled’ ribonuclease. However, the biological significance of this interaction is far from clear, particularly in view of the finding (see 2.3) that another member of the PDI family, Erp57, interacts specifically with calnexin in vivo (Zapun et al., 1998). (c) PDI as a subunit of the microsomal triglyceride transferase PDI is an essential subunit of the microsomal triglyceride transfer protein (MTP) (Wetterau et al., 1991). This -heterodimer is obligatory for the assembly of apoBcontaining lipoproteins; the -subunit is identical with PDI. A 30 amino acid sequence at the C-terminus of the -subunit was found to be essential for the formation of the active heterodimer (Ricci et al., 1995). Upon dissociation of the heterodimer by treatment with chaotropic agents, low concentrations of denaturants or a nonionic detergent, the catalytic activity was lost completely. Furthermore, the free 88kDa -subunit showed high tendency to aggregate. Even under conditions when the -subunit did not aggregate


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(chaotropic agents) no catalytic activity was expressed. Even when PDI was added to the -subunit after dissociation of the complex no catalytic activity was obtained, indicating that PDI must interact with the α-subunit cotranslationally. These results were confirmed by Lamberg et al. who overexpressed the MTP subunits in Sf9 cells. Expression of the asubunit alone led to insoluble aggregates with no catalytic activity while, upon coexpression of the / -subunits, soluble heterodimers were formed with MTP activity (Lamberg et al., 1996). By inactivating the active site cysteine residues of PDI/ -subunit it was demonstrated that the ability of PDI to form disulphide bonds is not essential for the formation of active MTP. However, it is still not clear whether PDI has an additional, more direct role in MTP activity. Taken together these results indicate that PDI might keep the -subunit in an active, non-aggregated conformation without the disulphide isomerase activity being involved. (d) PDI as a subunit of the prolyl 4-hydroxylase Prolyl-4-hydroxylase (P4H) is important in the posttranslational formation of 4hydroxyproline in procollagen in the ER (Kivirikko and Myllylä, 1987). This enzyme is formed by an heterodimer in C. elegans and an 2 2 tetramer in vertebrates with the -subunit identical with PDI (Pihlajaniemi et al., 1987; Vuori et al., 1992a; Veijola et al., 1994). P4H is very similar to MTP with respect to dissociation: once dissociated the subunits could not re-associate (Kirivikko et al., 1992). When the -subunit alone was synthesized in a baculovirus expression system no P4H activity was obtained, instead the -subunit had a remarked tendency to form aggregates. This result was confirmed in vitro using dog pancreas microsomes (John et al., 1993). However, upon co-expression of and -subunit aggregation of the -subunit was prevented and a functional 2 2 tetramer was formed (Vuori et al., 1992a, b). Site-directed mutagenesis of the subunit/PDI demonstrated that the active site cysteine residues of PDI were not essential for the assembly and activity of active 2 2 tetramer (Vuori et al., 1992b). Thus, as in the case of MTP (see above), these results indicate that the -subunit/ PDI is essential for keeping the -subunit in a non-aggregated, active conformation. Furthermore, the -subunit/PDI seems to be involved in the retention of P4H in the ER. Since the -subunit does not contain any retention signal, deletion of the -KDEL retention sequence of the -subunit led to substantial secretion of P4H and PDI. The interaction between PDI and P4H/MTP, respectively, might very well reflect the interaction between PDI and its substrates on which PDI acts to facilitate folding and/or disulphide bond formation. Thus, it might be useful to determine how PDI interacts with P4H/MTP in order to elucidate its function in protein folding and disulphide bond formation. 5. IS PDI A MOLECULAR CHAPERONE? “Molecular chaperones are defined as a functional class of unrelated families of protein that assist the correct non-covalent assembly of other polypeptide-containing structures in vivo, but are not components of these assembled structures when they are performing their normal biological function” (Ellis, 1993). This definition implies that molecular


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chaperones prevent or reverse the formation of incorrect interactions between transiently exposed surfaces of folding polypeptides. PDI clearly has the ability to suppress the aggregation of a variety of denatured proteins upon renaturation. It seems also clear that this is not exclusively related to the enzymatic function of PDI, i.e. formation and rearrangement of disulphide bonds, since PDI also suppresses the aggregation of proteins lacking disulphide bonds. It is now established that PDI has a specific binding site for oligopeptides and unfolded proteins to which the b' domain and other domains contribute, depending on the size of the ligand (Klappa et al., 1998). This interaction is clearly essential for PDI’s ability to catalyse disulphide-bond isomerization and folding in misfolded or partially folded protein substrates and could account for PDI’s ability to suppress aggregation of some non-disulphide-containing proteins during renaturation. However, all these results were obtained from in vitro experiments and there are only indirect data available suggesting that PDI is a molecular chaperone in vivo. As for PDI being identical to the -subunit of the prolyl 4-hydroxylase and of the microsomal triglyceride transferase respectively, PDI can not be regarded as a molecular chaperone in these contexts, since PDI is still a component of the assembled complexes. In summary, i) PDI certainly is an enzyme involved in the formation and rearrangement of disulphide bonds, ii) it seems certain that PDI has specific binding affinities for peptides and some unfolded proteins, which contribute to its ability to act as a catalyst of folding in addition to simple thiol-disulphide exchange, though further work is required to define these interactions, and iii) it is not yet known whether PDI indeed is a molecular chaperone in vivo. 6. REFERENCES Baksh, S., Burns, K., Andrin, C. and Michalak, M. (1995). Interaction of calreticulin with protein disulfide isomerase. J. Biol. Chem. , 270 , 31338–31344. Boado, R.J., Campbell, D.A. and Chopra, I.J. (1988). Nucleotide sequence of rat liver iodothyronine 5'-monodeiodinase (5’MD): its identity with the protein disulfide isomerase. Biochem. Biophys. Res. Com. , 155 , 1297–1304. Bulleid, N.J. and Freedman, R.B. (1988). Defective co-translational formation of disulphide bonds in protein disulphide isomerase-deficient microsomes. Nature , 335 , 649–651. Bulleid, N.J. and Freedman, R.B. (1990). Cotranslational glycosylation of proteins in systems depleted of protein disulphide isomerase. EMBO J. , 9 , 3527–3532. Cai, H., Wang, C. and Tsou, C. (1994). Chaperone-like activity of protein disulfide isomerase in the refolding of a protein with no disulfide bonds. J. Biol. Chem. , 269 , 24550–24552. Cheng., S., Gong, Q., Parkinson, C., Robinson, E.A., Appella, E., Merlino, G.T. and Pastan, I. (1987). The nucleotide sequence of a human cellular thyroid hormone binding protein present in endoplasmic reticulum. J. Biol. Chem. , 262 , 11221–11227. Creighton T.E., Hillson, D.A. and Freedman, R.B. (1980). Catalysis by proteindisulphide isomerase of the unfolding and refolding of proteins with disulphide bonds. J. Mol. Biol. , 142 , 43–62. Creighton T.E., Zapun, A. and Darby, N.J. (1995). Mechanisms and catalysts of


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contact with secretory proteins during late stages of translocation. Eur. J. Biochem. , 232 , 755–764. Klappa, P., Hawkins, C., and Freedman, R.B. (1997). Interaction between protein disulphide isomerase and peptides. Eur. J. Biochem. , 248 , 37–42. Klappa, P., Ruddock, L.W., Darby, N.J. and Freedman, R.B. (1998). The b domain provides the principal peptide-binding site of protein disulfide isomerase but all domains contribute to binding of misfolded proteins. EMBO J. , 17 , 927–35 Kortemme, T., Darby, N.J. and Creighton, T.E. (1996). Electrostatic Interactions in the Active-Site of the N-terminal thioredoxin-like domain of protein disulfide-Isomerase. Biochemistry , 35 , 14503–14511. Laboissiere, M.C.A., Sturley, S.L. and Raines, R.T. (1995). The essential function of protein-disulfide isomerase is to unscramble non-native disulfide-bonds. J. Biol. Chem. , 270 , 28006–28009. LaMantia, M. and Lennarz, W.J. (1993). The essential function of yeast protein disulfide isomerase does not reside in its isomerase activity. Cell , 74 , 899–908. Lamberg, A., Jauhiainen, M., Metso, J., Ehnholm, C., Shoulders, C., Scott, J., Pihlajaniemi, T. and Kivirikko, K.I. (1996). The role of protein disulphide isomerase in the microsomal triacylglycerol transfer protein does not reside in its isomerase activity. Biochem. J. , 315 , 533–536. Lambert, N. and Freedman, R.B. (1985). The latency of rat liver microsomal protein disulphide-isomerase. Biochem. J. , 228 , 635–645. Lu, X., Gilbert, H.F. and Harper, J.W. (1992). Conserved residues flanking the thiol/disulfide centers of protein disulfide isomerase are not essential for catalysis of thiol/disulfide exchange. Biochemistry , 31 , 4205–4210. Lundstrom J., Krause, G. and Holmgren, A. (1992). A Pro to His mutation in active site of thioredoxin increases its disulfide-isomerase activity 10-fold. J. Biol. Chem. , 267 , 9047–9052. Lyles, M.M. and Gilbert, H.F. (1991). Catalysis of the oxidative folding of ribonuclease A by protein disulfide isomerase: dependence of the rate on the composition of the redox buffer. Biochemistry , 30 , 613–619. Lyles, M.M. and Gilbert, H.F. (1994). Mutations in the thioredoxin sites of protein disulfide isomerase reveal functional nonequivalence of the N- and C-terminal domains. J. Biol.Chem. , 269 , 30946–30952. Martin, J.L. (1995). Thioredoxin: a fold for all reasons. Structure , 3 , 245–250. Martin, J.L., Bardwell, J.C.A. and Kuriyan, J. (1993). Crystal structure of the DsbA protein required for disulphide bond formation in vivo. Nature , 365 , 464–468. Morjana, N.A. and Gilbert, H.F. (1991). Effect of protein and peptide inhibitors on the activity of protein disulfide isomerase. Biochemistry , 30 , 4985–4990. Nelson, J.W. and Creighton, T.E. (1994). Reactivity and ionization of the active site cysteine residue of DsbA, a protein required for disulfide bond formation in vivo. Biochemsitry , 33 , 5974–5983. Noiva, R., Kimura, H., Roos, J. and Lennarz, W.J. (1991). Peptide binding by protein disulfide isomerase, a resident protein of the endoplasmic reticulum lumen. J. Biol. Chem. , 266 , 19645–19649. Noiva, R., Freedman, R.B. and Lennarz, W.J. (1993). Peptide binding to protein disulfide


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isomerase occurs at a site distinct from the active site. J. Biol. Chem. , 268 , 19210– 19217. Oliver, J.D., van der Wal, F.J., Bulleid, N.J. and High, S. (1997). Interaction of the thioldependent reductase Erp57 with nascent glycoproteins. Science , 275 , 86–88. Otsu, M., Omura, F., Yoshimori, T. and Kikuchi, M. (1994). Protein disulfide isomerase associates with misfolded human lysozyme in vivo. J. Biol. Chem. , 269 , 6874–6877. Pihlajaniemi, T., Helakoski, T., Tasanen, K., Myllyla, R., Huhtala, M.L., Koivu, J. and Kivirikko, K.I. (1987). Molecular cloning of the b-subunit of human prolyl-4hydroxylase. This product and protein disulphide isomerase are products of the same gene. EMBO J. , 6 , 643–649. Quan, H., Fan, G, and Wang, C. (1995). Independence of the chaperone activity of protein disulfide isomerase from its thioredoxin-like active site. J. Biol. Chem. , 270 , 17078–17080. Ricci, B., Sharp, D., O’Rourke, E., Kienzle, B., Blinderman, L., Gordon, D., SmithMonroy, C., Robinson, G., Gregg, R.E., Rader, D.J. and Wetterau, J.R. (1995). A 30amino acid truncation of the microsomal triglyceride transfer protein large subunit disrupts its interaction with protein disulfide-isomerase and causes abetalipoproteinemia. J. Biol. Chem. , 270 , 14281–14285. Roth, R.A. and Pierce S.B. (1987). In vivo cross-linking of protein disulfide isomerase to immunoglobulins. Biochemistry , 26 , 4179–4182 Ruddock, L.W., Hirst, T.R. and Freedman, R.B. (1996). pH-dependence of the dithioloxidizing activity of DsbA (a periplasmic protein thiol: disulphide oxidoreductase). and protein disulphide-isomerase: studies with a novel simple peptide substrate. Biochem. J. , 315 , 1001–1005. Ruoppolo, M. and Freedman, R.B. (1995). Refolding of disulfide isomerization: the mixed disulfide between ribonuclease T1 and glutathione as a model refolding substrate. Biochemistry , 34 , 9380–9388. Ruoppolo, M., Freedman, R.B., Pucci, P. and Marino, G. (1996). Glutathione-dependent pathways of refolding of Rnase T-1 by oxidation and disulfide isomerization— catalysis by protein disulfide-isomerase. Biochemistry , 35 , 13636–13646. Schoenmakers, C.H.H., Pigmans, I.G.A.J., Hawkins, H.C., Freedman, R.B. and Visser, T.J. (1989). Rat liver type I iodothyronine deiodinase is not identical to protein disulfide isomerase. Biochem. Biophys. Res. Com. , 162 , 857–868. Song, J. and Wang, C. (1995). Chaperone-like activity of protein disulfide-isomerase in the refolding of rhodanese. Eur. J. Biochem. , 231 , 312–316. Tsibris, J.C.M., Hunt, L.T., Ballejo, G., Barker, W.C., Toney, L.J. and Spellacy, W.N. (1989). Selective inhibitors of protein disulfide isomerase by estrogens. J. Biol. Chem. , 264 , 13967–13970. Veijola, J., Koivunen, P., Annunen, P., Pihlajaniemi, T. and Kivirikko, K.I. (1994). Cloning, baculovirus expression, and characterization of the a subunit of prolyl 4hydroxylase from the nematode Caenorhabditis elegans. J. Biol. Chem. , 269 , 26746– 26753. Vuori, K., Myllylä, R., Pihlajaniemi, T. and Kivirikko, K.I. (1992). Expression and sitedirected mutagenesis of human protein disulfide isomerase in Escherichia coli. J. Biol. Chem. , 267 , 7211–7214.


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Vuori, K., Pihlajaniemi, T., Marttila, M. and Kivirikko, K.I. (1992a). Characterization of the human prolyl 4-hydroxylase tetramer and its multifunctional protein disulfideisomerase subunit synthesized in a baculovirus expression system. Proc. Natl. Acad. Sci. USA , 89 , 7467–7470. Vuori, K., Pihlajaniemi, T., Myllylä, R. and Kivirikko, K.I. (1992b). Site-directed mutagenesis of human protein disulphide isomerase: effect on the assembly, activity and endoplasmic reticulum retention of human prolyl 4-hydroxykase in Spondoptera frugiperda insect cells. EMBO J. , 11 , 4213–4217. Walker, K.W., Lyles, M.M. and Gilbert, H.F. (1996). Catalysis of oxidative protein folding by mutants of protein disulfide isomerase with a single active-site cysteine. Biochemistry , 35 , 1972–1980.. Warwicker, J. and Gane, P.J. (1996). Calculation of Cys DpKa’s and oxidising power for DsbA mutants. FEBS Letters , 385 , 105–108. Weissmann, J.S. and Kim P.S. (1993). Efficient catalysis of disulphide bond rearrangements by protein disulphide isomerase. Nature , 365 , 185–188. Wetterau, J.R., Combs, K.A., McLean, L.R., Spinner, S.N. and Aggerbeck, L.P. (1991). Protein disulfide isomerase appears necessary to maintain the catalytically active structure of the microsomal triglyceride transfer protein . Biochemistry , 30 , 9728– 9735. Wittrup, K.D. (1995). Disulfide bond formation and eukaryotic secretory productivity. Curr. Opin. Biotech. , 6 , 203–208. Yamauchi, K., Yamamoto, T., Hayashi, H., Koya, S., Takikawa, H., Toyoshima, K. and Horiuchi, R. (1987). Sequence of membrane-associated thyroid hormone binding protein from bovine liver: its identity with protein disulphide isomerase. Biochem. Biophys. Res. Com. , 146 , 1485–1492. Zapun, A., Bardwell, J.C.A. and Creighton, T.E. (1993). The reactive and destabilizing disulfide bond of DsbA, a protein required for protein disulfide bond formation in vivo. Biochemistry , 32 , 5083–5092. Zapun, A., Darby, N., Tessier, D.C., Michalak, M., Bergeron, J.J.M. and Thomas, D.Y. (1998). Enhanced catalysis of ribonuclease B folding by the interaction of calnexin or calreticulin with ERp57. J Biol Chem. , 273 , 6009–6012.

21. PEPTIDYL-PROLYL CIS/TRANS ISOMERASES GUNTER FISCHER1, * and FRANZ X.SCHMID2 1 Max-Planck-Gesellschaft,

Research Unit “Enzymology of Protein Folding”, D-06120 Halle/Saale, Germany 2 Laboratorium für Biochemie, Universität Bayreuth, D-95440 Bayreuth, Germany

1. Introduction 2. Prolyl Isomerization 2.1. Chemistry of Peptide Bond Isomerization 2.2. Prolyl Isomerizations as Slow Steps in Protein Folding 3. Enzymes Catalyzing Prolyl Isomerization 3.1. Activity Assays 3.2. Classification 3.3. Fundamental Properties 3.4. Cellular Interactions of PPIases 4. Catalysis of in vitro Protein Folding Reactions by PPIases 4.1. Acceleration of Proline-limited Folding 4.2. Ribonuclease T1 as a Model Protein for Investigating Catalyzed Protein Folding 4.3. Catalysis of RNase T1 Folding by Cyclophilins and FKBPs 4.4. Catalysis by Parvulins 4.5. Isomerization of Peptide Bonds not Preceding Proline 4.6. Autocatalytic Folding of a PPIase 5. The Trigger Factor as a Folding Enzyme 5.1. PPIase Activity 5.2. Enzyme Kinetics of Trigger Factor-catalyzed Folding 5.3. Chaperone Properties of the Trigger Factor 6. Catalysis of Prolyl Isomerization During de novo Protein Folding 7. Concluding Remarks 8. References *Corresponding author

Peptidyl-prolyl cis/trans isomerases

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1. INTRODUCTION The complexity of protein folding originates in large part from the huge amount of rotations about single bonds that are possible in a polypeptide chain. Even when only the backbone is considered, rotations about two single bonds can occur at every Cα atom. The enormous number of conformations that are thus possible for the backbone of an unfolded protein have long been considered as a major problem for protein folding (Levinthal, 1968). At the same time the peptide bonds introduce considerable rigidity into the protein main chain. The partial double bond character of the carbon-nitrogen bond leads to the planarity of the peptide units, it creates a high barrier to rotation about this bond, and it allows a peptide bond to occur in only two conformations, cis or trans. It also leads to an enhanced stability toward nucleophilic attack. The individual properties of the amino acid preceding the prolyl bond may play a particular role for the height of the rotational barrier separating individual conformations as well as for general folding transitions. Unrestricted motions, acting cooperatively in structure-forming processes may appear kinetically uncoupled from such highly hindered rotations. Well-resolved slow folding reactions result which have a frequent occurrence even in the folding of small globular proteins. The molecular nature of the individual motion that will cause a slow, uncoupled folding event with a relaxation time > 100 ms is not always known. However, there is good evidence now that the slow isomerizations about proline imidic bonds are frequently the rate-determining events in folding, and thus they should be prime targets for a potential catalysis of protein folding. In the cell, folding should not be too slow and partially-folded intermediates should not be present for an extended time in order to minimize the risk of their aggregation. Indeed, several families of enzymes have been discovered which catalyze prolyl peptide bond isomerizations. These so-called peptidyl-prolyl cis/trans isomerases (PPIases) are the only known enzymes, evolved to stabilize a transition state that is separated from a ground state only by a difference in a torsional angle. They are very widespread and occur in virtually all tissues and organisms. As many as seven PPIases distinct in amino acid sequence, localization and enzymatic properties have been found in E. coli cells. Even more are known in yeast (Dozinski et al., 1997). PPIases are a unique class of enzymes, because they catalyze the interconversion of conformers by accelerating the rotation about a chemical bond. This is one of the simplest processes that can lead to products chemically distinct from reactants. Unlike chaperones, PPIases are classical enzymes. They do not require energy (e.g. in the form of ATP) and in their enzymatic properties they follow the simple MichaelisMenten relation. Also, they can perform unlimited numbers of catalytic cycles while leaving the product composition unaltered. In addition, a well-defined active site is located on the surface of the PPIases for binding of the substrates or inhibitors and for catalysis. Enzymes have two major functions in metabolism. They accelerate reactions which otherwise would occur at too low rate, and, in doing so, they convey specificity, because competing, unwanted reactions are not catalyzed and thus suppressed. These two functions may also be important for PPIase-catalyzed protein folding. High specificity


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constants approaching kcat/Km≥107 M-1 s-1 were determined for substrates with the reactive bond freely accessible, indicating a perfectly evolved catalytic function. A proline ring does not suffice to activate the catalytic machinery of PPIases. Therefore, substrate reactivity must be viewed in the context of secondary binding sites within substrate chains which may be restricted by steric limitations of partially folded substrates. These preconditions create features of regiospecific enzyme actions (Kern et al., 1993).

Figure 1 (A) Representation of the prolyl isomers; (B) The family tree of peptidyl-prolyl cis/trans isomerases according to their amino acid sequence homologies.

2. PROLYL ISOMERIZATION 2.1. Chemistry of Peptide Bond Isomerization In folded proteins the peptide bonds occur only in two conformations, cis or trans, and the dihedral angles for the rotation about the CN bonds are tightly clustered around 0° (as) and 180° (trans) (Figure 1A) (Stewart et al., 1990). Peptide bonds not preceding proline are almost always trans in folded proteins, but 5.7% of all Xaa-Pro peptide bonds show the cis conformation in the proteins with known three-dimensional structure (MacArthur & Thornton, 1991). In the following we refer to Xaa-Pro peptide bonds simply as “prolyl bonds” and to their isomerizations as “prolyl isomerizations”. Prolyl bonds introduce considerable conformational heterogeneity into peptides. In principle, peptides or proteins with n proline residues can form 2n isomers unless structural constraints (such as in folded proteins) stabilize one isomer strongly relative to the other. Accordingly, as many as 16 cis/trans isomers could be identified by 1H-NMR


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spectroscopy for the octapeptide IFPPVPGPG, which is derived from the prolactine receptor (Oneal et al., 1996). In this case, the decrease in the fraction of isomers is not a monotone function of a decreasing number of trans peptide bonds in the molecule. Consequently, the most abundant species is the trans/trans/ cis/trans isomer having a cis conformation for the prolyl bond in penultimate position. For a prolyl bond the probability to occur in cis conformation in a polypeptide is in part determined by its local environment and, in particular, by the nature of the amino acid preceding proline. Extensive investigations on Ac-Ala-Xaa-Pro-Ala-Lys-amides, having all 20 amino acids in Xaa position, revealed a high percentage of cis isomer up to 40% when Xaa is an aromatic residue and low cis contents of about 6% when Xaa is an aliphatic residue (including proline itself). In this series of compounds the relaxation times for prolyl isomerization were found to vary about 3 fold around 1000 sec at 4°C (Reimer et al., 1998). With about 90 kJ/mol the activation barrier to prolyl isomerization is high and entirely enthalpic in nature. As a consequence, the rate of cis/trans isomerization is highly temperature dependent and decreases 3–4 fold when the temperature is decreased by 10°C (Stein, 1993). As many as four distinct transition state structures with a 90° twisted peptide bond have been suggested from theoretical calculations for the isomerization reaction. The calculated energy barriers for the isomerization leading to the syn orientation in the activated complex are always lower than those of the anti conformation (Fischer et al., 1994a). Because the cis/trans interconversion is relatively slow at low temperature and because cis and trans prolyl isomers can differ in their molecular shape, it is possible in favorable cases to separate proline peptides into different isomers by capillary zone electrophoresis (Meyer et al., 1994; Moore & Jorgenson, 1995; Ma et al., 1995) or by reversed phase chromatography in the cold (Kalman et al., 1996; Melander & Horvath, 1982). These results are straightforward for probing isomer specific recognition (Meyer et al., 1994) since the transient stability of the resolved isomers allows their separate application in chemical reactions. Contigent on the chemical and structural differences of the isomers, many examples exist for the initiation of conformational specific recognition by cis/trans isomerizations. It preferentially occurs when macromolecules are allowed to react with prolyl peptides. Well-known consequences for peptide bond hydrolysis have been reported due to isomer specificity of a wide range of proteases (Brandts & Lin, 1986; Fischer et al., 1983), even if the isomeric bond is remote from the scissile bond. Often complete lack of reactivity for the “wrong” prolyl isomer was obtained. Thus, fractions of proline-containing peptide hormones, like bradykinin, can be refractory to proteolytic inactivation even during passage through organs (Merker & Dawson, 1995). Conformational preferences in receptor binding is exemplified by the cis peptide bond preference for the binding of dermorphin analogues to the opioid receptor (Yamazaki et al., 1993). Surprisingly, selectivity in conformational recognition can already be achieved by simple, bidentate terephtaloyl amides which can bind preferentially to cis prolyl bond derivatives (Vicent et al., 1991). The rate constants of prolyl isomerizations depend on the solvent, and a change from water to organic solvents led to an about 20 fold increase in the isomerization rate of acetyl-Gly-Pro-methylester (Eberhardt et al., 1992). A similar acceleration of cis/trans isomerization was found for proline peptides which are incorporated into micelles and


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phospholipid vesicles. In organic chemistry, micellar catalysis often follows the pseudophase model which quantitatively describes the dependence of the reaction rate on the detergent concentration (Fendler & Fendler, 1975). In fact, prolyl isomerization investigated in the presence of detergents differing in their ionic character also fits the pseudophase model. Interestingly, the rate constants kmic determined under saturating conditions do not vary greatly with respect to the charge of the micelle-forming detergent indicating close contact of the prolyl bond to the hydrocarbon chains buried in micelles (Kramer & Fischer, 1992). Under cellular conditions chemical catalysis of prolyl isomerization is difficult to obtain. The isomerization is independent of pH in the physiological range unless dissociable groups are present adjacent to proline. Acid catalysis requires protonation of the imide nitrogen, which show a pKa of about –7 and thus it can be observed only in very strong acids (Sigel & Martin, 1982; Steinberg et al., 1960; Schmid & Baldwin, 1978). Recently, in a five-membered metal chelate containing the nitrogen of the proline ring of acetyl-proline amides transiently coordinated to Cu(II) the rotational barrier to CN rotation was shown to decrease by up to 18 kJ mol-1. A catalytic rate enhancement by this Cu (II) complex of 10 fold was measured for the interconversion of all-trans polyproline to the all-cis form in CDCl3 (Cox et al., 1996). Moderate success was obtained by constructing antibodies that were complementary to the twisted peptide bond thought to mimic the transition state of PPIase catalysis (YliKauhaluoma et al., 1996). The a-dicarbonyl moiety of FK506 served as a model to design haptens for these experiments. However, the kcat values of the monoclonal antibodies produced were rather similar to those already found for the micellar catalysis described above. 2.2. Prolyl Isomerizations as Slow Steps in Protein Folding Ribonuclease A (RNase A) was the first protein for which a proline-limited folding reaction was observed. In 1973, Garel & Baldwin (1973) discovered that in its unfolded state RNase A consists of a kinetically heterogeneous mixture of fast-folding (UF) molecules, which could refold in less than a second, and slow-folding (Us) molecules, which refolded in the range of several minutes. Similar UF and Us species have since then been detected in the folding of many other proteins (Kim & Baldwin, 1982; Schmid, 1992; Schmid, 1993). Brandts and coworkers were the first to point out that this pronounced heterogeneity in the folding rates of UF and Us could be linked with prolyl isomerization. They suggested that the fast- and slow-folding molecules differ in the cis/trans isomeric state of one or more Xaa-Pro peptide bonds (cf. Figure 1) (Brandts et al., 1975). In a native protein (N) usually each prolyl peptide bond is in a defined conformation being either cis or trans in every molecule as defined by the ordered structure in the native state. After unfolding (N UF, Eq. 1), however, the prolyl bonds become free to isomerize slowly as in short oligopeptides (in the UF

Us i reaction, Eq. 1):


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(1) A mixture of unfolded species is thus created. It consists of a single unfolded form with correct prolyl isomers (UF) and one or more unfolded species with incorrect prolyl isomers (Us i). As outlined above, the UF molecules can refold rapidly to the native conformation. The refolding of the Us i molecules, however, is slow, because refolding is coupled with the re-isomerizations of the incorrect prolyl bonds. Of course, non-native isomers do not block the very first steps of refolding as suggested initially. Incorrect prolyl isomers can usually be well accommodated in partially folded, flexible intermediates, but not in native proteins. Thus a protein chain can begin to fold while some prolines are still in the nonnative state. At some point, however, correct prolines are required, and therefore the final steps of folding are necessarily limited in rate by their slow isomerizations (Cook et al., 1979; Schmid & Blaschek, 1981; Schmid, 1992; Schmid et al., 1993). The conformational folding with incorrect prolyl isomers can proceed to different extents, depending on the location of these non-native isomers in the structure and on the conditions used for refolding. Generally, incorrect prolyl bonds interfere least with conformational folding when they are located at the surface of the folding protein or in flexible chain regions. Solvent conditions that strongly stabilize folded proteins will also stabilize partially folded structure in intermediates with incorrect isomers. For RNase T1 it was found that protein chains with correct and with incorrect prolines folded with very similar rates of 1–5 s-1 (Mayr et al., 1996). Of course, the products of these folding reactions were different: native protein was formed in the folding of the chains with correct prolines, but in the presence of incorrect prolines intermediates were formed, which still had to undergo the very slow proline-limited final steps of folding. Conformational folding steps and prolyl isomerization are thus mutually interdependent. The presence of incorrect isomers in the chain can block folding at a certain stage, and at the same time the presence of folded structure can affect the equilibrium and the kinetic properties of the subsequent Xaa-Pro peptide bond isomerization. This close interrelationship between structure formation and prolyl peptide bond isomerization is a key feature of these slow folding steps and is of central importance for understanding the role of PPIases in these processes. 3. ENZYMES CATALYZING PROLYL ISOMERIZATION 3.1. Activity Assays There are only a few methods available to measure the enzymatic activity of PPIases. They are often based on methods monitoring the re-establishment of the fraction of isomers to actual reaction conditions following rapid disturbance of the cis/trans isomeric equilibrium. These preconditions always lead to coupled assays since a rapid jump in the conditions is followed by the relatively slow monitoring of an isomer sensitive signal. Recording of time courses must be performed up to completion of the cis/trans


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isomerization. PPIase activities are calculated by determining the rate constants of the time-course traces. Based on these facts, isomer-specific proteolysis provides a useful method for designing sensitive PPIase assays (Fischer et al., 1984). Since chymotrypsin has a very high conformational specificity for a trans peptide bond at the P2 subsite, it cannot cleave even at high concentrations the approx. 8% fraction of cis conformer present in aqueous solution of the chromogenic oligopeptide Suc-Ala-Ala-ProPhe-4-nitroanilide. In contrast, under these conditions the anilide bond of the trans isomer is cleaved in much less than a second. Thus, after this rapid reaction, which is complete within the time of manual mixing, a slow first-order reaction is observed, which is limited in rate by the irreversible cis to trans interconversion of the Ala-Pro bond in the assay peptide. In the presence of PPIases, this slow process is accelerated and its firstorder rate constant increases. This rate constant can be used to calculate the enzymatic bimolecular rate constant kcat/Km. Evaluation of the full set of steady state enzyme constants, kcat and Km, requires specialized analysis (Kofron et al., 1991). An improved performance of the protease-coupled assay resulted from increasing the fraction of cis isomer up to 70% by starting the assay from a LiCl/trifluoroethanol stock solution of the assay peptide (Kofron et al., 1991). Some disadvantages of the protease-coupled assay such as the risk of proteolytic degradation of the PPIases under study and the limited substrate variability were avoided by using NMR-techniques (London et al., 1990; Justice et al., 1990; Kern et al., 1993; Kern et al., 1995) and proteolytically uncoupled fluorescence (GarciaEcheverria; et al., 1992) or absorbance techniques (Garcia-Echeverria; et al., 1993), but at the expense of a much lowered sensitivity and reduced variability in the conditions. 3.2. Classification In the course of identifying catalysts in pig kidney cortex for slow steps in protein folding, a proline-containing oligopeptide 4-nitroanilide was utilized to record kinetic traces typical of catalysis of prolyl isomerization. Catalysis resulted from a protein with a molecular mass lower than 20 kDa. It displayed properties typical of an enzyme thus dating the discovery of PPIases to 1984 (Fischer et al., 1984). In 1989 this enzyme was found to be identical to the already known CsA receptor cyclophilin of 18 kDa molecular mass (pig Cyp18) (Fischer et al. 1989; Takahashi et al., 1989). Subsequent research yielded a FK506-binding protein (FKBP) (see below) that represents a PPIase of a 11.8 kDa catalytic core not related to Cyp18 in its amino acid sequence (Siekierka et al., 1989; Harding et al., 1989). The currently known families of PPIases are depicted in Figure 1B. All enzymes are classified as isomerases under the EC number 5.2.1.8. The enzymatic properties and the biological functions of cyclophilins and FKBPs were reviewed recently (Fischer, 1994b; Galat & Metcalfe, 1995; Kay, 1996). A proposal for the nomenclature of the isoenzymes has been made there (Fischer, 1994b). The major cyclophilins and FKBPs tend to be high in their intracellular concentrations because kidney tubules and endothelial cells contain about 10 g of Cyp18 per mg of total protein (Ryffel et al., 1991). This is comparable to 20 M of FKBP12 present in the cytosol of lymphocyte cells (Siekierka et al., 1991). Immunosuppressive compounds, the cyclic undecapeptide cyclosporin A and the pipecolic acid-derived peptidomacrolides FK506 and rapamycin, can bind tightly (pM to


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M level of Ki values) to the respective active sites of cyclophilins and FKBPs, thereby inhibiting the enzymatic activity of the proteins in a reversible and competitive manner. Cross-inhibition does not occur. Formation of the PPIase/inhibitor complexes proved complicated in kinetics. Solvent-dependent conformational changes of the inhibitors as well as isomerizations of the initially formed enzyme/ inhibitor complexes can lead to lags in the development of the full inhibitory capacity of up to several hours (Kock et al., 1992; Kofron et al.; 1992; Zarnt et al.; 1995, Janowski et al., 1996). Due to the ability to bind specifically immunosuppressants, proteins of both PPIase families were alternatively designated as immunophilins (Standaert et al., 1990). However, several PPIases do not exhibit high affinity to the immunosuppressants, and other proteins devoid of PPIase activity can bind immunosuppressants (Fischer, 1994b). The inhibition of the PPIase activity of the putative cytosolic receptors of the immunosuppressant in T-lymphocytes, Cyp18 and FKBP12, seems to be insufficient to block signal transduction and to induce both immunosuppression and toxic effects of the inhibitors on cell growth (Tropschug et al., 1989; Schreiber et al., 1993). Further extension of the number of PPIase families has recently been obtained. In E. coli, the trigger factor was identified as a 48 kDa PPIase during screening the cellular protein synthesis apparatus for PPIases (Stoller et al., 1995). It is situated adjacent to nascent secretory and non-secretory polypeptide chains at the 50 S subunit of the ribosome (Valent et al., 1995; Hesterkamp et al., 1996a). Homologs of the trigger factor have been detected in other prokaryotes, but not yet in eukaryotes (Hesterkamp & Bukau, 1996b). In the complete genome of Mycoplasma genitalium, the smallest known genome for any free-living organism, a gene was identified coding for a trigger factor homologue (Fraser et al., 1995). It represents the only PPIase identified so far by homology search in this gene complement. There are no sequence similarities between the E. coli trigger factor and Cyp18. Weak similarities were found, however, with the FKBPs. By using a hydrophobic cluster analysis Callebaut & Mornon (1995) could identify a small, but significant overall identity (value of 28.3%) between FKBP12 and amino acid residues Gln148 to Thr249 of the trigger factor (Stoller et al., 1996; Hesterkamp & Bukau, 1996b). However, in consideration of the inability of high concentrations of FK506 to inactivate full-length E.coli trigger factor as well as its catalytic domain (Stoller et al., 1995; Stoller et al., 1996), positioning of these enzymes in a special subgroup of the FKBP family seems to be a reasonable choice (Figure 1B). In 1994, a cytoplasmic PPIase was identified in E. coli lacking any significant sequence similarity to the other families of PPIases. With only 92 amino acids, it forms the archetype of the parvulin family (from Latin, parvulus, very small) of PPIases (Rahfeld et al., 1994a). High concentrations of both cyclosporin A and FK506 do not affect the PPIase activity of parvulin (Rahfeld et al., 1994b). In a first approach, juglone, a chinoid walnut dye, was found to irreversibly inhibit E.coli parvulin and yeast ESS1/PTF1 leaving other PPIases completely unaffected (Hennig et al., 1998). Database searches with the protein sequence of parvulin showed highly significant similarity with a number of domains in larger bacterial, yeast, invertebrate and human proteins (Rahfeld et al., 1994a; Rudd et al. 1995; Hani et al., 1995). Several of the prokaryotic homologues were thought to be involved in maturation or transport of specific proteins (Lazar & Kolter, 1996; Kontinen & Sarvas, 1993).


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The cellular functions of PPIases have been investigated genetically by gene disruption experiments. They usually revealed either dispensability or strong redundancy of PPIase functions. A notable exception was found in a conditional lethal phenotype for a mitochondrial Cyp20 in yeast that was identified genetically (Davis et al., 1992). Similarly, complementation of yeast cells defective in a parvulin homologue by the 18 kDa human parvulin Pin1 showed an essential function of this enzyme for normal cell growth. Pin1 variants with either a C-terminal truncation or a triple mutation of critical amino acids (both inactivating the PPIase activity) cannot complement the yeast mutant cells with respect to mitotic arrest followed by nuclear fragmentation (Lu et al., 1996). There is evidence that eukaryotic parvulins are interchangeable in terms of enzymatic function (Maleszka et al., 1996). 3.3. Fundamental Properties The catalytic activity of all PPIases seems to be independent of metal ions, cofactors or posttranslational modifications, unless sensitivity is conveyed by an extra domain in the sequence. Cyp 18 treated with Zn2+ was reported to acquire DNA binding properties but at the expense of losing its enzymatic activity (Krummrei et al., 1995). In addition, for this PPIase a Ca2+/Mg2+-dependent nuclease activity has been suggested (Montague et al., 1994). It seems likely that this additional catalytic function does not exist in the native state of Cyp 18 because a protein sample which was active in a nuclease assay did not bind CsA. While being of unclear biological significance this nuclease activity may serve as a probe for integrity of the Cyp18 structure (Schmidt et al., 1996). The specificity constant kcat/Km represents a very useful kinetic parameter for the assessment of the effectiveness of PPIase catalysis since non-productive substrate binding does not affect this parameter. Two microscopic specificity constants have to be considered, (kcat/Km)trans and (kcat/Km)cis, because catalysis occurs in both directions of the reversible isomerization. To calculate both constants it is essential to know whether or not an irreversible reaction, such as a folding step of the protein substrate, is coupled to the prolyl isomerization. The magnitude of (kcat/Km)cis can be more easily determined by an assay that is based on isomer specific proteolysis of the substrate since under the used conditions a trans to cis isomerization would never occur. Similar to many proteinases, PPIases have an extended binding site covering several peptide units of the substrate. To be recognized by Cyp18 a substrate peptide should contain at least three amino acids, including the proline (Fischer et al., 1994c). Accordingly, PPIases do not have affinity for an isolated proline ring. Although systematic investigations are still lacking, Cyp18 is believed to be maximally active toward substrates encompassing a sequence of 5 peptide bonds with the proline in the penultimate position. It is possible, however, that PPIases also bind to protein substrates at positions that are much further apart from the reactive prolyl bond (Schutkowski et al., 1996). Considering the high degree of conservation of the amino acid residues within the active cleft, which extends the binding site of the docked tetrapeptide (Zhao & Ke, 1996) in both directions of the substrate chain, enhanced affinity for large substrates is anticipated (Fischer, 1994b). The internal equilibrium constant of the Michaelis complexes Kint=Cyp 18* [trans]/


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Cyp18* [cis]=1.45 has been determined for Cyp 18 with a tetrapeptide substrate by line shape analysis of the 1H-NMR spectrum (Kern et al., 1995). The magnitude of Kint was shown to match the situation that an enzyme has to maintain in vivo. Enzymes evolved to operate near equilibrium have a Kint near unity (Burbaum et al., 1989). Thus, the experimental Kint for Cyp18 points to a cellular function of the enzyme for the catalysis in a reversible manner. Obviously this is at variance with a function in the de novo protein folding since this process has an equilibrium constant far from unity. The kinetic specificity of Cyp18 and FKBP12 towards peptide substrates is manifested essentially in the turnover numbers kcat, while Km values range invariantly around 0.6±0.5 mM (Decenzo et al., 1996; Kofron et al., 1991; Kern et al., 1995). Km still remains in this range for mutant FKBPs with strongly reduced (kcat/Km) cis values (Decenzo et al., 1996). Provided a favorable contribution of the amino acid in the position preceding proline, Cyp18 (kcat>104 s−1) predominates over FKBP12 in the turnover number by one or two orders of magnitude. According to these data, the proficiency of PPIases as catalysts is quite low. Defined by dividing the second-order rate constant by the rate constant k0 of the spontaneous reaction of a substrate, the proficiency factor (and the rate enhancement factor kcat/k0) of Cyp18 resides on the lower end of an enzyme scale (Radzicka &Wolfenden, 1995). The value of kcat/Km>107 M-1 s-1 that already approaches diffusion control for the bimolecular step of enzymatic catalysis indicates that the proficiency of PPIases will become physically limited. Thus enzymes catalyzing reactions with high spontaneous rates cannot evolve while maximizing proficiency. Except for cyclophilins which have a limited variability in kcat/Km within one order of magnitude, other families of PPIases prefer strongly substrates with hydrophobic residues in the position preceding proline (Stoller et al., 1995; Harrison & Stein, 1990; Bergsma et al., 1991; Albers et al., 1990). On the PPIase side, the amino acid residues that are thought to be important for catalysis seem to be highly conserved for the cyclophilins and parvulins, but less for the FKBPs (Figure 2). It was hypothesized that for cyclophilins the evolution of the intracellular compartimentalization occurred much later than the division of prokaryotes and eukaryotes took place. In contrast, distinct prokaryotic members of the FKBP family were used to diverge into specialized enzymes (Trandinh et al., 1992). Domains active as PPIases occur in many large proteins and in two cases proteolytically resistant, active FKBP and trigger factor cores could be isolated from larger proteins (Chambraud et al., 1993; Stoller et al., 1996; Hesterkamp & Bukau, 1996). Far from completeness, Figure 3 schematically describes the protein modules found in the different families of PPIases. Besides the catalytic domain most of the characteristic sequence patterns are known to direct compartimentalization of the protein. Interestingly, the presence of a WW domain in all eukaryotic parvulins with the consequent affinity for prolin-rich protein segments (Sudol et al., 1995), may confer additional specificity to the PPIase domain of parvulins.


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Figure 2 Fully conserved amino acid residues through the entries of the PPIase database. Portions of the depicted residues can be used as signature sequences of the respective enzyme family. (A) cyclophilins; (B) FKBPs and (C) parvulines.

The three-dimensional structures of the parent parvulin and trigger factors are not yet known. However, there is a wealth of information concerning the high-resolution molecular structures of both FKBP12 and FKBP12/inhibitor complexes, and for cyclophilins complexed with cyclosporins or linear oligopeptides. Based on antiparallel -strands supplemented with short helical stretches, Cyp18 and FKBP12 show a protein fold resembling -lactoglobulin and retinol-binding protein or the pleckstrin-homology domain, respectively. The structure of Cyp18 was solved for the apoprotein as well as in several complexes with ligands. In general, the protein conformation did not change significantly upon complexation with substrates or inhibitors (Ke, 1992; Kallen & Walkinshaw, 1992; Ke at al., 1994; Zhao & Ke, 1996). A hydrophobic pocket on the surface of the eight-stranded -barrel structure forms the binding cleft. Bound oligopeptides are exclusively found with cis conformation of the respective prolyl bond. Reasoning that the minimal reaction pathway for PPIase catalysis would comprise Michaelis complexes of both the cis and trans isomer of the substrate, microscopic rate constants can be calculated by 1H-NMR based line shape consistent with preferred catalysis of trans to cis isomerization (Kern et al., 1995). Currently it is believed that the cyclophilins differ in the enzyme mechanism from the FKBPs (Fischer et al., 1993; Kramer & Fischer, 1997). For the FKBPs desolvation and substrate-assisted catalysis may be important factors in the mechanism of catalysis. For cyclophilin, an electrophilic assistance for example by Arg55 could in part account for its high activity. The positively charged side chain of this amino acid residue is located in hydrogen bonding distance perpendicularly to the plane of the proline ring (Zhao & Ke, 1996). It could possibly immobilize the lone electron


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Figure 3 The domain structure of several members of cyclophilins, FKBPs and parvulins. Numbers represent percentage of identical amino acids in the catalytic cores according to the BESTFIT sequence comparison program. Symbols characterize: ; transmembraine ; signal sequences ; WW domain ; metal binding ; ER targeting ; histone-like ; proline-rich ; catalytic domain.

pair at the nitrogen atom and thus, as in acid catalysis, abolish the partial double bond character of the amide bond. Indeed, the replacement of Arg55 by Ala decreases the activity of Cyp18 about 1000-fold (Zydowsky et al., 1992). In the refolding of dihydrofolate reductase a prolyl isomerization was suggested to be catalyzed intramolecularly also by an arginine (Texter et al., 1992). 3.4. Cellular Interactions of PPIases As stated above the major cytosolic cyclophilin Cyp18 may play a role in structure formation of cellular proteins that is not coupled to an irreversible folding step. Thus compartimentalized PPIases are likely to have pleiotropic biological effects. A promising approach to dissect the different functions is based on the identification of proteins intracellularly associated with PPIases. The complexes between cyclophilin and CsA and between FKBP and FK 506 or rapamycin have apparently important functions in immunosuppression. They have been reviewed recently (Dumont & Su, 1996; Cardenas et al., 1995) and will not be considered here. Copurification, affinity chromatography and the yeast two-hybrid screen provided valuable tools to characterize protein/PPIase complexes. In view of low stability of known Michaelis complexes and the lack of fairly stable intermediates of the enzyme


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reaction it is difficult to say a priori whether or not a cellular binding protein will represent the original substrate of the PPIase within the cell. It is also uncertain whether the complex of a binding protein with a PPIase detected by the above methods will have any in vivo relevance. A useful probe may lie in complementation studies which make use of variants of the binding partners independently altered in their proteinaceous functions. Evidence of similar phenotypes will point to a common signalling pathway, and thus indicate concerted action of the proteins within the cell. Representative examples for the PPIase/protein complexes are given in the following. By using the yeast two-hybrid system, the zinc finger transcription factor YY1 was found to interact with both Cyp18 and FKBP12 (Yang et al., 1995). These interactions could be disrupted by CsA and FK506, respectively, suggesting that the PPIase active sites were involved in these interactions. The interaction of YY1 with FKBP12, but not with Cyp18, involves a C-terminal stretch of 83 amino acids. An enhanced cellular level of FKBP12, realized by overexpression, is able to competitively prevent binding of YY1 to its DNA target site. Evidence for a direct interaction between FKBP12 and YY1 in vitro has not yet been obtained. Several of the proteins associated with heterooligomeric steroid receptors in their inactive state were described as FKBPs and as a cyclophilin, each of high molecular mass (Ratajczak et al., 1993; Peattie et al., 1992) (see chapter by Toft, this volume). Conflicting reports exist about the influence of CsA and FK506 on the receptors (Milad et al., 1995). Affinity to Hsp90, another component of the receptors, was demonstrated for members of both families of PPIases, FKBP52 (Lebeau et al., 1992) and Cyp18 (Nadeau et al. 1993). For Cyp18, binding is dependent on the Mg2+/ATP concentration but does not require the hydrolysis of ATP and cannot be blocked by CsA. Obviously, in the inactive steroid receptor complexes PPIases interact with chaperones, but the function of this interaction is unknown. Another example for PPIase complexes with cellular proteins is the interaction of FKBP12 with the skeletal muscle ryanodine receptor. The first evidence came from the observation that the two proteins copurify (Collins, 1991). It now appears that most of the intracellular calcium release channels are associated with FKBP12. Involvement of the enzymatic function of FKBP12 in the regulation of the channel may be inferred from the effects of administration of the FKBP12 inhibitor rapamycin which causes immediate channel activation (Kaftan et al., 1996). Since a large distance has been found between the putative ion-conducting site and the FKBP12 binding site in the transmembrane assembly, only a long range interaction may be hypothesized for a possible involvement of FKBP12 in channel activation (Wagenknecht et al., 1996). An exchange procedure of receptor bound FKBP12 with 3 different FKBP12 variants nearly devoid of PPIase activity does not confirm the necessity of enzyme catalysis in the modulation of channel activity (Timerman et al., 1995). However, the activity of the FKBP12 variants was measured with artificial substrates. Considering the lack of knowledge about the efficiency toward natural substrates it remains difficult to predict the residual activity of the FKBP12 variants toward natural substrates. FKBP12 also forms a complex with the transforming growth factor- type 1 receptor. This interaction was identified by monitoring a rat heart cDNA library in the yeast twohybrid screen (Wang et al., 1994). It is suggested that the complex may play a role in


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type I receptor-mediated signalling. Co-immunoprecipitation of FKBP12 and a cytoplasmic domain of the receptor was used to confirm the interaction of both proteins in vitro. In contrast to the situation with the calcium release channel, the low activity D37G FKBP12 variant cannot maintain complex formation with the growth factor- type 1 receptor. Inactivating FKBP12 activity by rapamycin or FK506 abolished complexation as the D37G mutation did. Recently, it was shown that host cell Cyp18 is required for HIV-1 infectivity prior to reverse transcription of the virus RNA but subsequent to receptor binding and membrane fusion (Braaten et al., 1996). A two-hybrid screen identified a Cyp18/ pr55gag interaction (Franke et al., 1994; Thali et al., 1994). Incorporation of Cyp18 into HIV-1 virions was found to occur via contact with a proline-rich segment in the capsid domain of Pr55gag (Franke et al., 1994). This conserved array totaling four prolines occurs in the periodicity (P(Xaa)4P222(Xaa)2P(Xaa)5P). Mutant proteins of the HIV-1HXB2 having site-directed mutations, either P222A or G221A, failed to bind to a GST-Cyp18 fusion protein. Virions equipped with these mutant proteins cannot sequester Cyp18 into the released virions, emphasizing the importance of the Gly221-Pro222 bond for Pr55gag /Cyp18 complex formation. Analogously, small in-frame deletions in the capsid domain of Pr55ggag drastically decreased the efficiency of packaging Cyp18 into viral particles (Thali et al., 1994). It seems that the PPIase activity of Cyp18 is required for packaging, because there was a good correlation between the differential effects that several derivatives of CsA had on the enzymatic activity of Cyp18 and on the packaging process (Bartz et al., 1995). However, it is unclear whether or not catalysis of prolyl isomerization in Pr55gag contributes to the enhancement of HIV-1 infectivity mediated by the Pr55gag/Cyp18. In this respect, the level of affinity for the active site of Cyp18 is 6 fold higher for a 25-mer peptide encompassing all the relevant proline residues than it was found for a 10-mer peptide containig only 3 of the conserved prolines (Schutkowski et al., submitted). There is also convincing evidence for a specific cellular binding protein of a parvulin. The intracellular binding partner of the NIMA (never in mitosis) protein kinase, screened by the two hybrid system using the Aspergillus nidulans protein kinase with a human cDNA bank, resulted in the identification of the parvulin-like PPIase Pin1 (Lu et al., 1996). Colocalization in HeLa cells and co-immunoprecipitation experiments provided further evidence for this interaction. Pin1 specifically contacts the residues 280–396 (NID domain) of NIMA kinase known to mediate protein/ protein interactions, but does not function as substrate or inhibitor for the kinase. Taken together, these results suggested that binding of Pin1 to NID mediates inhibition of the mitotic function of NIMA protein kinase. In terms of PPIase/polypeptide interactions, a puzzling finding may be presented finally. It was shown by CsA sensitive cross-linking that the proline-free peptide CGYDVSTAQKIIAKL can react specifically with cytosolic Cyp 17 of yeast, indicating some unexpected affinity of the compounds to each other (McNew et al., 1993). Sequence specific, non-proline directed binding of oligopeptides to Cyp18 was also demonstrated in kinetic competition experiments (Fischer & Schutkowski, unpublished results). The above interaction may not cause a catalytic event given the inability of Cyp18 to catalyze the trans to cis isomerization of the stable non-proline cis peptide bond


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Tyr38-Ala39 in a RNase T1 variant (see chapter 4.5). However, amino acid sequences lacking proline may adopt partial structural characteristics typical of proline, as it was already found for a seven residue stretch of cytochrom C551 (McArtur & Thornton, 1991). It is not unlikely that such segments could bind to PPIases. 4. CATALYSIS OF IN VITRO PROTEIN FOLDING REACTIONS BY PPIases 4.1. Acceleration of Proline-limited Folding The immunoglobulin light chain, porcine ribonuclease (RNase) and the S-protein fragment of bovine RNase A were the first proteins for which a catalysis of folding by a PPIase, porcine cytoplasmic cyclophilin18, could be demonstrated (Lang et al., 1987). In all these protein folding reactions, however, the activity of this PPIase was much lower than towards prolines in tetrapeptides. It is now clear that the rapid formation of folding intermediates and the concomitant decrease in accessibility are the major reasons for the decreased activity of the cyclophilins in protein folding. A catalysis of proline-limited steps by various PPIases was observed in the folding of many proteins, including the collagen triple helix (Bächinger et al., 1980; Bächinger. 1987; Davis et al., 1989), barnase (Matouschek et al., 1990), carbonic anhydrase (Freskgard et al., 1992; Kern et al., 1994), -lactamase (Lejeune, unpublished results), chymotrypsin inhibitor CI2 (Jackson & Fersht, 1991), yeast iso-2 cytochrome c (Veeraraghavan & Nall, 1994; Veeraraghavan et al., 1995), the immunoglobulin light chain (Lang et al., 1987; Lilie et al., 1995), and trp aporepressor (Mann et al., 1995). Most of these proteins are small, and the molecules with correct prolyl isomers refold in the time range of milliseconds. Both cis to trans and trans to cis isomerizations are catalyzed equally well in the folding of these proteins, which rules out that the catalytic function of the PPIases is restricted to the cis to trans direction. The importance of the accessibility of the prolines was demonstrated for iso-2 cytochrome c. In aqueous buffer the folding of this protein is barely catalyzed by cyclophilin. When, however, guanidinium chloride is added in increasing, but still non-denaturing concentrations to destabilize folding intermediates the prolyl bonds becomes better accessible, and the catalysis of folding is markedly improved (Veeraraghavan et al., 1995), even though the catalytic activities of the PPIases decrease with increasing denaturant concentration. Of course, not all slow steps in protein folding are prolyl isomerizations. The very slow refolding of large proteins is often limited in rate by other slow conformational rearrangements, by domain pairing reactions, or by subunit associations. 4.2. Ribonuclease T1 as a Model Protein for Investigating Catalyzed Protein Folding Much work on the mechanisms of proline-limited protein folding and of its catalysis by PPIases has been carried out with RNase T1 (Pace et al., 1991) as a model protein. Almost the entire folding of RNase T1 involves prolyl isomerizations (Kiefhaber et al., 1990b; Odefey et al., 1995), and it is a good substrate not only for PPIases, but also for other folding enzymes and chaperones (Schmid, 1993; Schmid et al., 1993).


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RNase T1 is a small single-domain protein of 104 amino acids (Heinemann & Saenger. 1982; Pace et al., 1991; Martinez-Oyanedel et al., 1991) with two disulfide bonds (Cys2Cys10 and Cys6-Cys103). It contains four prolyl peptide bonds; two are trans (Trp59Pro60 and Ser72-Pro73) and two are cis (Tyr38- Pro39 and Ser54-Pro55) in the native protein. RNase T1 is most stable near pH 5 and further stabilized when NaCl is added (Oobatake et al., 1979; Pace et al., 1988). Importantly, in the absence of the disulfide bonds, RNase T1 can still fold to a native-like conformation in 1 M NaCl. Variants of RNase T1 have been designed to create simple models for investigating particular aspects of protein folding. These variants differ in the conformational stability and in the number and location of cis prolyl bonds and of disulfide bonds (Kiefhaber et al., 1990; Mayr et al., 1994a; Mayr et al., 1994b). The folding mechanism of the wild-type protein with the two cis prolines and two disulfide bonds has been studied most extensively (Mayr et al., 1996). In unfolding of RNase T1, the rapid conformational unfolding step is followed by the isomerizations of the two cis prolyl peptide bonds at Pro39 and Pro55 until, at equilibrium, both Pro39 and Pro55 are 80–90% trans. Four unfolded species are thus formed, one UF species with the native isomers at Pro39 and Pro55 and three Us species with one or two incorrect isomers. The UF species with both Pro39 and Pro55 in the correct cis conformation refolds to the native state N in less than a second (Mayr et al., 1996). The three Us species with one or two incorrect prolyl isomers also regain most of their secondary structure and presumably part of their tertiary structure early in refolding. The products of these reactions are, however, partially folded intermediates that still contain the same incorrect prolines as the Us species from which these reactions were initiated. The reisomerizations of the incorrect prolyl isomers occur in the subsequent slow steps and are coupled with further folding. The tight coupling between conformational folding and the isomerizations of the prolyl peptide bonds is evident in the folding of RNase T1. As discussed above, very rapid partial folding is possible in the presence of nonnative proline isomers (Kiefhaber et al., 1990a; Kiefhaber et al., 1992), but the correct isomers are required to complete folding. The final events of folding are thus about 1000-fold decelerated. The protein chains therefore remain in a partially folded state for an extended time and could, in principle, undergo nonproductive reactions, such as aggregation. In RNase T1, partial folding has an additional unfavorable effect: it decelerates the trans to cis isomerization at Pro39 in the final step of folding (Kiefhaber et al., 1990a). In the folding of pancreatic RNase A, however, prolyl isomerization is accelerated in a folding intermediate (Cook et al., 1979; Schmid & Blaschek. 1981), and in the folding of dihydrofolate reductase intramolecular catalysis of a prolyl isomerization was proposed to occur (Texter et al., 1992). After breaking of the two disulfide bonds of RNase T1 by reduction and subsequent carboxymethylation, the resulting RCM-RNase T1 is unfolded in aqueous buffer, but folds reversibly to a native-like ordered conformation in 1 M NaCl (Mücke & Schmid, 1992). The RCM form of the S54G/P55N variant ((-P55)-RCM-T1) is a particularly valuable model protein for studying assisted protein folding. Its folding mechanism is very simple, because it involves a single trans to cis isomerization only (of Pro39) and because the access to this proline is not impaired by premature structure formation, as in


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the presence of the disulfide bonds. Since all RCM forms are unfolded in aqueous buffer, spontaneous as well as assisted and catalyzed unfolding and refolding can be studied in the absence of denaturants, simply by varying the NaCl concentration (Mücke & Schmid, 1994a; Mücke & Schmid, 1994b). This is important because several PPIases are very sensitive to residual concentrations of denaturants, such as guanidinium chloride or urea. 4.3. Catalysis of RNase T1 Folding by Cyclophilins and FKBPs The slow reactions in the folding of RNase T1 (with intact disulfide bonds) are catalyzed by PPIases with varying efficiency. Pro55 is solvent-exposed in the native protein and presumably also in folding intermediates, and therefore catalysis at this proline is generally good. Pro39 is buried in the native protein, and its isomerization is not well catalyzed, because rapid conformational folding blocks the access for PPIases. The catalysis of Pro39 isomerization is improved when intermediate formation is inhibited by destabilizing mutations, or by breaking of the two disulfide bonds (as in the reduced and carboxymethylated RCM form of RNase T1). In these variants the PPIases are no longer hindered in their access to the refolding protein molecules (Mücke & Schmid, 1992). Many different PPIases of the cyclophilin and FKBP families from a variety of organisms catalyze the refolding of RNase T1 (Schmid et al., 1993). The best catalysis of Pro39 isomerization in the refolding of RNase T1 with intact disulfide bonds was observed with the cytoplasmic cyclophilin 18 from E. coli. In the presence of 29 M cyclophilin 18 the rate of this folding reaction was 300 fold increased (Schönbrunner et al., 1991). In protein folding the PPIases function as enzymes. They catalyze cis to trans isomerization in either direction and do not determine the isomeric states of the prolyl bonds in the protein substrates. Catalysis of prolyl isomerization in protein unfolding cannot be measured easily, because PPIases are inactivated under conditions, such as the presence of denaturants and elevated temperatures, which are normally used to denature proteins. These problems could be overcome by using the RCM forms of RNase T1 as substrates (Mücke & Schmid, 1992). As outlined above, these proteins are only folded in the presence of = 1 M NaCl and can be unfolded by a dilution to e.g. 0.4 M NaCl at pH 8 and 15°C. Under these conditions the PPIases are stable and retain their full enzymatic activity. Unfolding, or, to be more precise, the prolyl cis to trans isomerizations in the unfolding chains of RCM-RNase T1 were indeed catalyzed and the efficiency of Cyp 18 was the same in unfolding and refolding experiments under identical conditions near the midpoint of the unfolding transition. Analogous results were also obtained for the catalysis of folding by FKBP12 (Scholz et al., 1998). The extents of unfolding in the kinetic experiments with and without PPIases were found to be identical, indicating that these enzymes do not affect the equilibrium between native and unfolded protein. Clearly, as expected for enzymes, PPIases catalyze prolyl isomerization in protein folding in either direction, and they have no directional information for folding. The isomeric states of the prolines are determined only by the structure and the conformational stability of the folding protein under the employed conditions.


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4.4. Catalysis by Parvulins An additional family of PPIases, the parvulins, was discovered by Rahfeld et al. (1994). These small proteins show no sequence homologies with the cyclophilins and the FKBPs, and they are not inhibited by the respective inhibitors cyclosporin A and FK 506. Parvulin from E. coli also catalyzes the folding and unfolding of our model protein (P55)-RCM-T1 with an efficiency that is slightly higher than that of the FKBPs, but lower than that of the cyclophilins (Scholz et al., 1997a). 4.5. Isomerization of Peptide Bonds not Preceding Proline For normal peptide bonds (not preceding proline) the trans state is favored at least 100fold over cis (Ramachandran & Mitra, 1976; Jorgensen & Gao, 1988), which implies that in unfolded proteins on average one incorrect cis peptide bond occurs in every 100 residues. In native proteins such cis peptide bonds are infrequent, but still several of them have been found by X-ray crystallography. Therefore, non-prolyl cis/trans isomerizations should also occur in protein folding and might also be catalyzed by enzymes, possibly by the PPIases. Because of the strong preference for the trans state, it is virtually impossible to design peptide substrates with cis peptide bonds to search for such a function. Also, until recently no protein folding reactions were known which are limited in rate by the isomerization of normal peptide bonds. In our model protein RNase T1 we could generate a cis Tyr-Ala bond by replacing the cis-Pro39 of the wild-type protein by an alanine (Mayr et al., 1994). This mutation caused a major change in the folding mechanism of RNase T1. In the unfolded state virtually all molecules contain a trans Tyr38-Ala39 bond, and in refolding its very slow trans to cis isomerization determines the reaction rate (Odefey et al., 1995). The time constant of this reaction is about 500 s at 25°C. Because nascent proteins are synthesized probably as all-trans chains, the folding of the proteins which contain cis peptide bonds (Herzberg & Moult, 1991) in their native conformations should be very slow and limited in rate by trans to cis peptide bond isomerizations as in the folding of the Pro39Ala variant of RNase T1. None of the known PPIases (cyclophilin18, FKBP12, or parvulin) could catalyze this folding reaction, suggesting that normal peptide bonds might not be substrates for these enzymes (Scholz et al., 1998). It will be intriguing to learn, whether enzymes similar to the PPIases can be found, which can catalyze these isomerizations of normal peptide bonds. 4.6. Autocatalytic Folding of a PPIase As folding enzymes the PPIases can, in principle, catalyze their own folding. Indeed, for human cytosolic FKBP12 it could be shown that its folding is an autocatalytic process for the mature protein and, even more pronounced, for a variant with an aminoterminal extension of 16 residues (Veeraraghavan et al., 1995; Scholz et al., 1996). In the native form FKBP contains seven trans prolyl peptide bonds, and the cis to trans isomerizations of some or all of them determine the rate of its folding. The reaction product catalyzes its


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own formation in an autocatalytic reaction, and therefore its rate increases with reactant concentration. Indeed, a more than 10-fold acceleration of folding was observed when the concentration of the FKBP 12 fusion protein was increased from 0.05 M to 10 M (Scholz et al., 1996). Different residues flank the seven trans prolyl bonds of FKBP12 (Siekierka et al., 1989), they probably remain not equally accessible during folding, and only two of them (Phe15-Pro16 and Ile91-Pro92) should be good substrates for the native FKBP molecules. Therefore, in the autocatalysis of its own folding, FKBP12 probably accelerates the prolyl isomerizations with different efficiencies. In the large proteins which contain FKBP-, cyclophilin- or parvulin-like domains (Fischer, 1994; Galat & Metcalfe, 1995; Rahfeld et al., 1994; Hani et al., 1995) the functions of these PPIase modules are not yet known. It is tempting to speculate that they could serve a role as intermolecular or even intramolecular catalysts of the folding or of conformational rearrangements which are necessary for the biological functions of these proteins. At present there is, however, no experimental evidence in support of such a role. 5. THE TRIGGER FACTOR AS A FOLDING ENZYME 5.1. PPIase Activity In the cells the trigger factor is located near the site of protein biosynthesis at the ribosome and can interact with newly formed protein chains. Therefore there is no doubt that it might be the prime candidate for a folding enzyme that accelerates prolyl isomerizations in the de novo folding of nascent proteins. Intriguingly, the trigger factor catalyzes the folding of RNase T1 much better than all other PPIases, which are known to date. For the cyclophilins and FKBPs the PPIase activities are much lower in protein folding than towards exposed prolyl peptide bonds in short tetrapeptides. Trigger factor, however, catalyzes prolyl isomerizations in folding proteins more efficiently than in peptides. The addition of as low as 2.5 nM trigger factor leads already to a doubling of the folding rate of (-P55)-RCM-T1, and in the presence of 20 nM trigger factor folding is 14-fold accelerated. This remarkable catalytic efficiency of the trigger factor as a folding enzyme is reflected in a specificity constant k cat /K m of 1.1×106 M-1s-1. This value is almost 100-fold higher than the respective value for human FKBP12 (Scholz et al., 1998). 5.2 Enzyme Kinetics of Trigger-factor-catalyzed Folding To determine the basis for the excellent catalysis of folding by the trigger factor, K M and k cat of catalyzed folding were determined separately (Scholz et al., 1997b). In these experiments the trigger factor behaved like a classical enzyme. It obeyed the MichaelisMenten equation in its catalysis of folding, and the initial rates of catalyzed folding showed saturation behavior. The analysis of the kinetic data gave values of 0.7 M for the K M value and 1.3 s-1 for the catalytic rate constant k cat of trigger-factor-catalyzed folding. The data can be compared with the K M of 220 M and the k cat value of 620 s-1, as measured for the catalysis by Cyp 18 of the trans to cis prolyl isomerization in a short


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tetrapeptide (Kern et al., 1995). This comparison indicates that the turnover number of the trigger factor in protein folding is very low, and that its high activity as a folding catalyst originates clearly from its tight binding to protein substrate, as reflected in the low K M value. For other PPIases such kinetic analyses cannot be performed, because their binding to protein or peptide substrates is very weak. 5.3. Chaperone Properties of the Trigger Factor The catalysis of RCM-T1 folding by the trigger factor is strongly inhibited by the reduced and carboxymethylated form of bovine -lactalbumin (RCM-La), a permanently unfolded, but soluble protein. RCM-La, which is also a good substrate for the chaperone GroEL (Okazaki et al., 1994; Hayerhartl et al., 1994), competes efficiently with RCM-T1 for binding to the trigger factor, and half-maximal inhibition is observed in the presence of 0.6 M RCM-La. In the presence of 1.0 M RCM-La the Michaelis-Menten kinetics of catalyzed folding are significantly shifted to higher substrate concentrations, as expected for competitive inhibition. This suggests that indeed the folding substrate RCMT1 and the non-folding inhibitor RCM-La compete for the same binding site on the trigger factor. This high-affinity binding site for unfolded proteins is probably distinct from the catalytic site of trigger factor. The isolated FKBP domain of the trigger factor is fully active as a PPIase towards a short tetrapeptide (Stoller et al., 1996), but in protein folding its activity is about 800-fold reduced, and, moreover, this residual activity of the FKBP domain is no longer inhibited by RCM-La. This suggests that the active site and the highaffinity binding site for unfolded substrate proteins reside on different domains of the intact trigger factor. It is possible that the additional protein binding site (s) on the intact trigger factor decelerates the dissociation of the protein substrate so strongly that a change in the ratelimiting step from bond rotation (in tetrapeptide substrates) to product dissociation (in protein substrates) occurs. Such a change might explain the observed low k cat value of 1.3 s-1 of the trigger factor in RCM-T1 folding. Additionally, some of the binding events should be non-productive, because the reactive prolyl peptide bonds are not positioned correctly within the PPIase site. Indeed, a lowering of both K M and k cat, as observed for the intact trigger factor, points to non-productive binding of a substrate to an enzyme (Fersht, 1985). The trigger factor thus seems to resemble a chaperone. The cooperation of protein binding and prolyl isomerization in catalyzed folding probably forms the basis for the very high catalytic efficiency of the trigger factor in protein folding. It is unknown at present, whether the chaperone properties of the trigger factor are also required for additional functions, such as accepting newly-synthesized protein chains at the ribosome. Prolyl isomerizations are presumed to be late steps in protein folding (Schmid, 1992), and therefore it seems surprising that a PPIase binds to a nascent protein chain very early, possibly cotranslationally, while it is still at the ribosome. The trigger factor might, however, remain associated with the folding chains and could even accompany the folding protein chain to other chaperone systems, such as GroEL. There is indeed evidence that the trigger factor can bind to GroEL in a substrate-dependent manner (Kandror et al., 1995). It is also possible that not all newly formed proteins require the


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DnaK or GroE chaperone systems and consequently might fold rapidly to completion right after their synthesis at or near the ribosomes. 6. CATALYSIS OF PROLYL ISOMERIZATION DURING DE NOVO PROTEIN FOLDING Studies on the folding of collagen (Steinmann et al., 1991) provided the first evidence for an involvement of prolyl isomerization and of PPIases in a cellular protein folding reaction. The formation of the collagen triple helix is limited in rate by successive prolyl isomerizations both in vitro and in vivo. The maturation of collagen in chick embryo fibroblasts is retarded when CsA is added and Steinmann and colleagues (Steinmann et al., 1991) proposed that CsA inhibits a cyclophilin which catalyzes collagen folding in the endoplasmic reticulum. Similarly, in rabbit reticulocyte lysate the folding of luciferase was retarded by CsA (Kruse et al., 1995). Effects of CsA on proteins other than cyclophilins could of course not be ruled out in these experiments. Additional convincing evidence for a role of cyclophilins in cellular folding came from the work of Rassow et al. (1995) and Matouschek et al. (1995). Both groups used the mitochondrial protein import system to find a function of cyclophilins in protein folding. Proteins that are targeted to the mitochondrial matrix must unfold outside the mitochondria, cross the two mitochondrial membranes and then refold in the matrix. To investigate folding after transport through the membranes, both groups used a fusion protein between the presequence of subunit 9 of the Neurospora crassa F1F0-ATPase and mouse cytosolic dihydrofolate reductase (Su9-DHFR). This construct has been used before to study other aspects of protein import into isolated mitochondria. After import the refolding of DHFR in the matrix showed half times of about 5 min in yeast mitochondria at 30°C as well as in N. crassa mitochondria at 25°C. These folding reactions were about 5-fold decelerated when the mitochondria had been pretreated with 2.5–5 M CsA to inhibit the PPIase activity of mitochondrial cyclophilin. In vitro, the rates of folding of Su9-DHFR differed to a similar extent when purified cyclophilin from N. crassa mitochondria was either present or absent. The refolding of DHFR was also retarded in mitochondria from yeast and N. crassa mutants which lack a functional mitochondrial cyclophilin, and the folding of the model protein Su9-DHFR in the mutant mitochondria proceeded with similar kinetics as in wildtype mitochondria which were pretreated with CsA. This provided additional evidence that the mitochondrial cyclophilins catalyze proline-limited protein folding reactions in organello. 7. CONCLUDING REMARKS Due to the electronic nature of the prolyl peptide bond, a high rotational barrier exists in polypeptides for the dihedral angle ω which cannot be greatly lowered through low molecular weight substances within the cell. This effect gives rise to the occurrence of kinetic intermediates in protein folding being decelerated in adopting the native state as well as to native state isomerizations. The detection and characterization of several


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families of PPIases have been vital in advancing our understanding of such prolinelimited slow phases in in vitro protein folding. In their catalytic function these proteins act as Michaelis-Menten type enzymes. They exert catalytic effects in both directions, cis to trans and trans to cis, of the reversible prolyl isomerization. With regard to steady state catalytic constants some PPIases are reminiscent of perfectly evolved enzymes which have high turnover numbers k cat in conjunction with high Michaelis constants Km. Protein folding is catalyzed with variable efficiency in vitro. Among PPIases the E.coli trigger factor represents the most remarkable example of catalytic power when the trans to cis isomerization in the refolding of a RNase T1 variant is considered. Kinetic studies of the folding of FKBP12 gave rise to intermolecular autocatalysis in that the already folded molecules can catalyze isomerizations in molecules that are still unfolded. Several cellular binding proteins for PPIases have been identified by different techniques. Conclusions whether they would represent cellular substrates cannot yet be drawn unequivocally. Nevertheless, protein folding in cells assisted by PPIases was already detected in several cases. 8. REFERENCES Albers, M.W., Walsh, C.T. and Schreiber, S.L. (1990). Substrate specificity for human rotamase FKBP: a view of FK506 and rapamycin as leucine-(twisted amide)- proline mimic. J. Org. Chem. , 55 , 4986–4986. Bächinger, H.-P. (1987). The influence of peptidyl-prolyl cis/trans isomerase on the in vitro folding of type III collagen. J. Biol. Chem. , 262 , 17144–17148. Bächinger, H.P., Bruckner, P., Timpl, R., Prockop, D.J. and Engel, J. (1980). Folding mechanism of the triple helix in type-III collagen and type-II pN collagen. Eur. J. Biochem. , 106 , 619–632. Bartz, S.R., Hohenwalter, E., Hu, M.-K., Rich, D.H. and Malkovsky, M. (1995). Inhibition of human immunodeficiency virus replication by nonimmunosuppressive analogs of cyclosporin A. Proc. Natl. Acad. Sci. USA , 92 , 5381–5385. Braaten, D., Franke, E.A. and Luban, J. (1996). Cyclophilin A is required for an early step in the life cycle of human immunodeficiency virus type 1 before the initiation of reverse transcription. J. Virol. , 70 , 3551–3560. Brandts, J.F., Halvorson, H.R. and Brennan, M. (1975). Consideration of the possibility that the slow step in protein denaturation reactions is due to cis/trans isomerism of proline residues. Biochemistry , 14 , 4953–4963. Brandts, J.F. and Lin, L.-N. (1986). Proline isomerization studied with proteolytic enzymes. Methods Enzymol , 131 , 107–126. Bergsma, D.J., Eder, C., Gross, M., Kersten, H., Sylvester, D., Appelbaum, E., Cusimano, D., Livi, G.P., McLaughlin, M.M., Kasyan, K., Porter, T.G. Silverman, C., Dunnington, D., Hand, A. Pritchett, W.P. Bossard, M.J. Brandt, M. and Levy, M.A. (1991). The cyclophilin multigene family of peptidyl-prolyl isomerases. Characterization of three separate human isoforms. J. Biol. Chem. , 266 , 23204– 23214. Burbaum, J.J., Raines, R.T., Albery, W.J. and Knowles, J.R. (1989). Evolutionary


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22. THE ATPase CYCLE OF THE GroE MOLECULAR CHAPERONES NEIL A.RANSON1 and ANTHONY R.CLARKE* Department of Biochemistry, School of Medical Sciences, University of Bristol, Bristol, BS8 1TD, UK 1Present address: Department of Crystallography, Birkbeck College London, Malet Street, London, WC1E 7HX, UK

1. Introduction 2. GroEL-protein Interactions: Some Basic Considerations 3. ATP-induced Rearrangements 3.1. Equilibrium Properties and Transient Kinetics of the ATP-induced Conformational Switch 3.2. Negative Co-operativity and the Establishment of Asymmetry in the GroEL Toroids 4. The Binding of ADP 4.1. Weak and Tight Interactions—A Further Manifestation of Negative Cooperativity 5. The GroEL ATPase Cycle in the Absence of GroES 6. GroES-GroEL Interactions and Their Effects on the Hydrolysis of ATP 6.1. Binding of GroES to the ADP and ATP States of GroEL 6.2. ATP Hydrolysis in the Presence of GroES 6.3. The Dissociation of GroES During the ATPase Cycle 6.4. Bullets and Footballs 7. The Coupling of Protein and Nucleotide Binding Affinities 7.1. The Effects of ATP, ADP & AMP-PNP on Protein Binding Affinity 7.2. The Effect of Unfolded Protein on Co-operativity Within the GroEL Oligomer 8. The Coupling of the ATPase Cycle to Chaperonin-assisted Protein Folding 8.1. Models of Chaperonin Action 8.1.1. Encapsulation and Folding 8.1.2. Unfolding and Release 8.2. Towards a Global Model for GroE-mediated Folding 9. Concluding Remarks 10. Acknowledgements 11. References


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Abbreviations ATP—

adenosine 5' triphosphate

ADP—

adenosine 5' diphosphate

*Corresponding author

AMP-PNP—

adenylylimidodiphosphate

LDH—

bacillus stearothermophilus lactate dehydrogenase

DHFR—

dihydrofolate reductase

GroEL—

Escherichia coli chaperonin-60

GroES—

Escherichia coli chaperonin-10

mMDH—

porcine mitochondrial malate dehydrogenase

rubisco—

ribulose bisphosphate carboxylase oxgenase

1. INTRODUCTION One of the central tenets of biogenesis is that the primary sequence of a protein contains all of the information necessary to specify the precise, three-dimensional structure of the native state. At its simplest, protein folding can be viewed as a directed conformational search which locates the kinetically accessible state of lowest free energy. Given that the principle function of the GroE proteins is to enhance the efficiency of the folding process, it is somewhat curious that energy derived from ATP hydrolysis is required to facilitate a reaction which is intrinsically thermodynamically favourable. Where does this chemically derived energy go and how is it harnessed? Any attempt to address this intriguing and novel mechanistic question inevitably requires us to elucidate the steps in the cycle of ATP binding, hydrolysis and product release. Both cryo-electron microscopy and recent crystallographic data reveal profound changes in the structure of the GroE complex as this chemistry proceeds, i.e. energy liberated in the cleavage of orthophosphate from ATP is used to drive rearrangements of GroEL which, in turn are coupled to the folding of substrate proteins (see Burston and Saibil, this volume). In this chapter we concentrate on four aspects of this energytransducing system: (1) the kinetics of binding, hydrolysis and dissociation of nucleotides, (2) the behavioural features of GroE conformers which arise during these steps, (3) the elaborate system of conformational communication between GroEL subunits in response to nucleotide binding and (4) the dynamics of GroES and substrate interactions in the ATPase cycle. GroEL, and its co-protein GroES, were initially identified as the products of E. coli genes in which mutations could block the assembly of bacteriophage (Georgopoulos et

The ATPase cycle of the GroE molecular chaperones

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al., 1973), but the proteins are expressed and required for cell viability at all temperatures (Fayet et al., 1989). GroEL is a tetradecamer of identical 57,259 Da. subunits arranged as two stacked heptameric rings, and GroES is a heptamer of identical 10,497 Da. subunits which associate to form a single ring (for a comprehensive review of the structures of GroEL, GroES and their complexes see the chapter by Burston and Saibil). Each GroEL subunit has a weak, Mg2+-dependent (or Mn2+-dependent, Diamant et al., 1995) ATPase activity (Hendrix, 1979), which is inhibited by approximately 50% in the presence of GroES (Chandrasekhar et al., 1986). The binding affinity of ATP is also highly dependent on the concentration of K+ ions (Viitanen et al., 1990; Todd et al., 1993). The detailed mechanistic study of the GroE proteins and the part they play in protein folding was initiated by the work of Goloubinoff et al. (1989), who recon-stituted the chaperonin system in vitro from purified components. The essential finding was that GroEL alone formed an extremely stable complex with unfolded bacterial rubisco, thus entirely arresting the folding reaction. On addition of both Mg-ATP and GroES rubisco was discharged from the complex and went on to fold with a final yield of around 80%. This ATP-dependent enhancement of folding yield has since been demonstrated and kinetically examined in vitro with a range of substrates including DHFR and rhodanese (Martin et al., 1991), LDH (Badcoe et al., 1991), glutamine synthetase (Fisher, 1992), citrate synthase (Buchner et al., 1991), mMDH (Miller et al., 1993; Staniforth et al., 1994b). There are four, general findings from this type of experiment which cast light on the coupling of the ATPase cycle to the folding reaction. Firstly, apo-GroEL is a strong inhibitor of folding owing to the high stability of the GroEL: substrate complex; hence some nucleotide-bound state arising in the ATPase cycle must be responsible for substrate dissociation. Secondly, while some substrates can be discharged by the addition of ATP alone, others require the additional impetus provided by GroES. Thirdly, the timescale of folding is usually much longer than that of the hydrolytic reaction ( ~25 seconds per subunit), implying multiple cycles of ATP turnover during chaperoninassisted folding reactions. Fourthly, under optimal conditions, assisted folding reactions give very large enhancements of yield at folding rates comparable with those measured in the absence of the chaperone. 2. GroEL-PROTEIN INTERACTIONS: SOME BASIC CONSIDERATIONS The last observation is mechanistically more interesting than it first appears. It can be argued, a priori, that the yield of the folding reaction can be enhanced by a passive process in which the unfolded protein chains are tightly bound to the surface of GroEL. This lowers the concentration of unfolded chains which are free in solution and will enhance the yield of folding, given that two criteria are satisfied: (i) the effective stability of the interaction between GroEL and the unfolded substrate is not larger than the free energy of folding of the polypeptide chain (∆G(F-U)) i.e.


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and (ii) the loss of yield in the spontaneous folding reaction results from the aggregation of unfolded chains. In these circumstances the folding yield (Y), at the outset of the folding reaction, can be described by the competition between the velocities of folding (Vf) and aggregation (Va). This can be represented as follows:

where kf and ka are the unimolecular and multi-molecular rate constants describing these processes, [U] is the concentration of free, unfolded substrate and N is the molecularity of the aggregation process. Hence, any reduction in [U] will increase Y. The price to be paid for this energy-independent process is a reduction in folding rate. If folding can occur while the protein substrate is bound to the GroEL surface (Gray & Fersht, 1993; Itzhaki et al., 1995), then the reduction in rate is defined by the difference in binding energy between the chaperone and the unfolded state, and the chaperone and the transition state for folding (∆GB(U-t)) such that the observed rate (kobs) becomes:

If the unfolded substrate must dissociate before folding and the binding equilibrium is relatively rapid then:

Either way, the observed folding rate for a protein which binds to GroEL in the unfolded state will be slower than the intrinsic folding rate. While this passive mechanism for the enhancement of folding yield—with a concomitant reduction in folding rate—is observed when bacterial LDH folds in the presence of GroEL alone (Badcoe et al., 1991), it is violated in all reported systems when ATP and GroES are included. Hence, however nucleotides modify the binding affinity between GroEL and the protein substrate, the simple binding schemes outlined above cannot explain the action of the GroE proteins during folding. 3. ATP-INDUCED REARRANGEMENTS 3.1. Equilibrium Properties and Transient Kinetics of the ATP-induced Conformational Switch The observation that substrate proteins can be released from GroEL by adding hydrolysable Mg-ATP to a highly stable binary complex does not reveal which GroEL: nucleotide state is responsible for altering the binding properties. However, the


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subsequent observation that the non-hydrolysable analogue Mg-AMP-PNP is able to discharge protein substrates (Badcoe et al., 1991), identifies ATP binding, rather than any post-hydrolytic process, as the critical step in the substrate release mechanism. This finding was reinforced by transient kinetic experiments in which the binary complex between unfolded barnase and GroEL was challenged with ATP and GroES (Corrales & Fersht, 1996b). The formation of the nucleotide-containing complex initiated folding of the enzyme which proceeded at a rate which is faster than ATP hydrolysis. In addition, the fact that ADP is scarcely able to promote substrate dissociation (Staniforth et al., 1994) is entirely consistent with this conclusion. This led to the minimal hypothesis that one of the roles of the ATPase activity of GroEL was to switch the protein between two states: an ATP-stabilized conformation which has a low affinity for substrate proteins and an apo- or ADP-stabilized state with a high substrate affinity. Nucleotide binding and hydrolysis would then drive the system cyclically between these forms, leading to a continual tendency to bind and release unfolded polypeptide chains (Jackson et al., 1993; Todd et al., 1994). The early observation that the steady-state rate of ATP hydrolysis showed a sigmoidal saturation curve established not only that the complex can be converted between two conformational states but that the switch was positively co-operative. Steady-state results also reveal that the affinity of GroEL for ATP is high (K1/2~ 10–20 M) and that GroES is able to increase the co-operativity of the system by shifting the Hill constant from around 2.5 to 4.0 (Gray & Fersht, 1991; Yifrach & Horovitz, 1994). The labelling of GroEL with a conformationally sensitive fluorescent dye marked a considerable step forward in detecting and characterizing the binding of nucleotides. If the GroEL oligomer is labelled with a single pyrene maleimide group the function of the protein is not detectably altered but the fluorescence intensity is highly sensitive to rearrangements induced by nucleotide binding (Jackson et al., 1993). Owing to the slow turnover of ATP, the extrinsic fluorescence enhancement on binding of the ligand can be recorded prior to the hydrolytic step. These experiments showed that binding rather than hydrolysis of ATP is co-operative, i.e. the binding curves were remarkably similar to the rate versus [ATP] curves recorded by Gray & Fersht and Yifrach & Horovitz. If, in the classical terminology of Monod, Wyman & Changeux (Monod et al., 1965) GroEL exists in a T and an R conformation, with ATP-binding stabilizing the latter, then the pyrene fluorescence probe ought to be capable of reporting the dynamics of the cooperative rearrangement of the oligomer. The transient kinetics of this interaction have been studied using stopped-flow fluorescence spectroscopy (Jackson et al., 1993) by mixing labelled apo-GroEL (T-conformation) with differing concentrations of ATP and measuring the first-order rate constant for the formation of the R-conformer. The data show that the observed rate constant for this transition reaches a maximum of 180 s−1, and that the half-maximal rate is achieved at an ATP concentration of 4mM. The simplest kinetic mechanism which is consistent with these results can be written:


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and the observed rate constant (kobs) for formation of the hyper-fluorescent Rconformation is given by:

Figure 1 A minimal model of ATP binding and hydrolysis by GroEL. The initial binding of ATP is weak (K1/2=4 µM) and is followed by a conformational rearrangement occurring at a maximal rate of 180 s−1, which converts the chaperonin to a state which binds ATP tightly (Kd=10 M). Hydrolysis of the bound nucleotide then converts the chaperonin back to a weak nucleotide binding state.

These transient data show the initial formation of a weak collision complex between ATP and the predominant T-conformer, with a dissociation constant of 4mM, followed by a rearrangement of the complex to the R-conformation at a rate of 180 s−1. Thus in the terminology of Monod, Wyman & Changeux (Monod et al., 1965), KT=4mM. This rather loose interaction is in stark contrast to the tight equilibrium binding of ATP (K1/2=10 µM) in which the nucleotide is ‘pulled on’ to GroEL by virtue of the T→R transition. It is clear from the transient kinetic data that the rate of the R→T transition (i.e. the value of k2, the intercept on the vertical axis) is very small, showing that the equilibrium between T and R conformations lies heavily toward the latter when the complex is occupied by ATP. The transient kinetics of this process have also been measured using an intrinsic, genetically introduced tryptophan probe at residue 485 (Tyr→Trp) and are essentially identical to those revealed by the pyrene label (Kad and Clarke., unpublished data). In the light of these results from transient experiments which define KT, the equilibrium binding data can be fully interpreted in accordance with the MWC model which envisages a concerted switch of the protein from a state in which all subunits are in


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the T- to one in which all are in the R-conformation (see Figure 2). In an equilibrated system, the proportion of oligomers in the R-confor-

Figure 2 Positive co-operativity in GroEL. GroEL exists in at least two states, a T-conformer with (square complex) with a weak affinity for nucleotide, and an R-conformer (circular complex) with a tight affinity. The binding of ATP is represented as promoting a T→R transition according to the MWC model of cooperatively as discussed in the text.

mation (PR) as a function of the concentration of ATP is defined by:

where c=Kr/KR (the ratio of the dissociation constants of ATP for the T- and R-


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conformers, α=[ATP]/KR, L is the ratio of T- to R-conformers ([T]/[R]) in the unliganded state and n is the number of binding sites for ATP in the co-operative unit. The value of each of these constants is given in the scheme in Figure 2. From a biological standpoint it is important to consider why such a co-operative mechanism should have evolved in GroEL. The most compelling reason is that the binding site for a single, respectably sized protein substrate is effectively constructed from seven binding surfaces on seven individual subunits within a ring. If GroEL must bind and then displace such a substrate during the ATPase cycle, the only way to achieve this efficiently would be to coordinate the behaviour of the subunits in a ring such that all are ‘on’, or all are ‘off’ at a given instant. Hence, a concerted switch of all subunits is central to the mechanism. Such an argument is consistent with the positively co-operative unit comprising seven rather than fourteen subunits (Horovitz et al., 1994; 1995; Bochkareva & Girshovic, 1994; Burston et al., 1995). A further feature of the cooperativity parameters is the relative values of L and of c. Since the equilibrium distribution of T-rings and R-rings (T7/R7) is determined by the value of L.cn (where n is the number of ATP molecules bound), then the binding of a single ligand is sufficient to balance the equilibrium. The binding of seven ATP molecules leads to a predominance of the R7 state of 3×1017 to 1. This massive stabilization of the R-conformation when fully liganded with ATP reflects a second mechanistic property, the very tight interaction of substrate proteins with the Tconformer. When a ring is loaded with a substrate protein, the effective value of L will be much larger, hence the transition to the R-conformation—which is required for displacement of the substrate—becomes more difficult (Yifrach & Horovitz, 1996). Hence, the large change in ATP binding energy between the T and R confomers is required to break the GroEL-substrate interaction and discharge the polypeptide from the binding surface of GroEL. The co-operative, ATP-induced rearrangement of GroEL has been visualised by both negative stain and cryo-electron microscopy (Saibil et al., 1991; Chen et al., 1994; Roseman et al., 1996). Reassuringly these structures offer a clear physical explanation for the effect of ATP binding on the affinity of GroEL for protein substrates. In the ATPconformation (the R7 state) the apical domains are all rotated such that their proteinbinding surfaces are occluded and less available for interaction with the polypeptide. The structural aspects of these conformational rearrangements and their relationship with the functional properties of GroEL are discussed more extensively in the chapter by Burston and Saibil. 3.2. Negative Co-operativity and the Establishment of Asymmetry in the GroEL Toroids The basic description of ATP binding given above views GroEL as behaving like a single positively co-operative unit comprising seven sites. It was recognized at an early stage that there must be some element of negative co-operativity in the 14-mer complex, since the binding of GroES to GroEL is seen to be asymmetric in most conditions (Roseman et al., 1996; Chen et al., 1994; Xu et al., 1997). That is, the binding of the GroES co-protein to one ring reduces its ability to bind to the other. As GroES can only associate with


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GroEL after the latter has bound nucleotide, this poses the question of whether this asymmetry is imposed by the association of the co-protein or is established prior to this step, i.e. by nucleotide binding itself.

Figure 3 The biphasic dependence of the initial velocity of the GroEL (R196→A) ATPase upon the ATP concentration (Adapted from Yifrach & Horovitz, 1994). Data were fitted to the equation V°= (0.5*Vmax1*[S]+Vmax2*L2*[S]*(1+[S]/K)7)/(K+[S]+K*L2(1+[S]/K) 8), where S is the substrate (ATP) concentration, K is the dissociation constant of the R-state for ATP (12.5 M), L2 is an equilibrium constant derived from [RR]/[TR] (L2=0.0096), and the subscripts 1 & 2 denote the two rings of GroEL. This analysis assumes that the ATP binds exclusively to the R-conformer, that in these conditions the TT state can be ignored, and that the affinity of ATP to a ring in the Rstate is similar for both RR and TR complexes.

Initially, two lines of evidence emerged which suggested that nucleotide binding was sufficient to enforce asymmetry upon the chaperonin oligomer. Horovitz and colleagues determined the concentration dependence of the initial rate of ATP hydrolysis by a GroEL mutant Arg196→Ala (shown in Figure 3), an amino acid substitution designed to modify co-operative contacts between subunits within a ring. If all of the fourteen available GroEL sites were able to bind and hydrolyse ATP at the same time, then this experiment should give a sigmoidal plot (reflecting the positively co-operative binding of ATP by GroEL) which reaches a plateau at high ATP concentrations as the ATPase sites of GroEL become saturated with ligand. However, the concentration-dependence of the initial rate of ATP hydrolysis is strongly biphasic, showing a rapid increase in initial rate followed by an inhibition of the rate at higher ATP concentrations. They proposed that these two phases correspond to the increase in ATPase activity as one of the two rings of GroEL is saturated with ATP, and the inhibition of the ATPase rate as the second ring


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becomes occupied at higher substrate concentrations. The authors concluded that negative co-operativity would have to exist between the GroEL rings in order that binding of nucleotide to one ring weakens the affinity of the second ring. The inhibition of the ATPase rate by ADP (Bochkareva & Girshovic, 1994) is also biphasic and points to the same conclusion.

Figure 4 Initial velocity of the GroEL ATPase as a function of GroEL concentration. ATPase velocity was estimated by measuring release of 32Pi. after four seconds, and increases linearly until only half of the available ATPase sites are occupied.

This question has also been examined by determining the stoichiometry of ATP binding to GroEL (Burston et al., 1995). Shown in Figure 4 is a plot of initial velocity as a function of protein concentration; a standard method for determining the number of active sites per protein in classical enzyme kinetics. In this case, the initial velocity of the GroEL ATPase was determined as a function of the GroEL concentration. A saturating concentration of ATP (175 µM; K1/2=10 µM) was included in each reaction, and this was mixed with varying concentrations of GroEL (expressed for simplicity in this instance as the GroEL subunit concentration) to initiate the reactions. The initial velocity of the ATPase reaction increases linearly until a GroEL concentration of 350 µM is reached. In other words, the velocity increases until there are two GroEL subunits for every ATP molecule, after which it flattens. This indicates that at any given time, only half of the GroEL ATPase sites are active, presumably the GroEL subunits constituting one of the two rings of the chaperonin. In support of this conclusion, addition of 7 ATPs per pyrenelabelled GroEL oligomer gives the maximal fluorescence enhancement after which the signal decays with a single exponential rate of 0.12 s-1 as the ATP is hydrolysed (see Figure 5). Addition of any greater quantity of ATP leads to a steady state phase of


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hydrolysis prior to the exponential phase in which the last 7 ATPs are turned over. This confirms the half-sites reactivity of GroEL and, in addition, shows that only 7 ATPs are required to generate the hyper-fluorescent R-conformation. Thus asymmetry is enforced at the level of negative co-operativity in nucleotide binding and the ‘one-sided’ association of GroES results from this. In summary,

Figure 5 Time resolved fluorescence of ATP binding to pyrenyl-GroEL. 100 M pyrenyl-GroEL (subunits) was present in each reaction, and either 50 M (A) or 100 M (B) ATP was added. Addition of ATP to pyrenyl-GroEL causes a 60% enhancement in fluorescence (λex=340 nm, λem=355nm), which decays as the nucleotide is hydrolysed.

seven sites on a single ring of GroEL are filled as a positively co-operative unit and the acquisition of the R-conformation in this ring inhibits ATP binding and hydrolysis on the other, i.e. negative co-operativity exists between rings preventing the formation of a symmetrical complex. The more recent work of Yifrach, Horovitz and colleagues has furthered our understanding of the GroEL ATPase through the development of a mathematical description of the co-operativity (Yifrach & Horovitz, 1995). This “nested cooperativity” model describes GroEL in terms of both the Monod-Wyman-Changeux (MWC) and Koshland-Nemethy-Filmer (KNF) models of co-operativity (see Figure 6). Each heptameric ring is in equilibrium between the T and R state, and binding of ATP to a single ring promotes the T→R transition within that ring by a concerted MWC mechanism. The two rings of GroEL however, undergo sequential transitions between TT (both rings in the T-state; i.e. with a low affinity for ATP), TR (ATP bound to one ring) and RR (ATP bound to both rings) states via a KNF mechanism where mixed oligomers are allowed. For wild type GroEL, the positively co-operative binding of ATP to one ring occurs with half-saturation at 16 µM, forming a TR complex with high ATPase activity. Weaker binding of ATP to the second ring produces the RR state which has a lower ATPase activity.


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4. THE BINDING OF ADP 4.1. Weak and Tight Interactions—A Further Manifestation of Negative Co-operativity The interaction of ADP with GroEL has also been studied in some depth, both in the presence and in the absence of GroES. In its absence, pyrene-labelled GroEL

Figure 6 A nested model of co-operativity in the GroEL ATPase (Adapted from Yifrach & Horovitz, 1995). As discussed in the text.

detects a weak and positively co-operative binding of ADP which occurs with a K1/2 of 2.3 M Jackson et al., 1993). However, the ATPase activity of GroEL is strongly modified by the inclusion of only 5 M ADP in steady-state reactions (Yifrach & Horovitz, 1996) and, as mentioned previously, the biphasic inhibition of the GroEL ATPase by ADP has been used as evidence for asymmetric binding of nucleotides (Bochkareva & Girshovic, 1994). These experiments imply that ADP, like ATP, can exhibit positive co-operativity in binding within a ring, but once the first ring is occupied negative co-operativity inhibits binding to the second. The stoichiometry of the tight phase of ADP binding has been determined by ultrafiltration which gives a value of seven tight sites per GroEL oligomer in accordance with the implications described above (Burston et al., 1995). The K1/2 for the high affinity occupation of the first ring has been crudely estimated to be 5 (±3) M using an extremely small fluorescence quench (0.25%) reported by pyrene-labelled GroEL (Burston et al., 1995); this occurs before the onset of the large (12%) enhancement caused by the weak binding to the second ring at high ADP concentrations. This phenomenon of negative co-operativity provides a mechanism for the biphasic inhibition of the initial ATPase rate observed by Bochkareva and Girshovic (1994) since the first phase may be caused by tight binding of ADP to the ring distal to that occupied by ATP and the second by weak competition with ATP binding. Recent data using a single-tryptophan mutant of GroEL (Tyr485→Trp), which acts as a fluorescence reporter, also shows tight equilibrium binding of ADP with a dissociation constant of 70 M (Kad and Clarke, unpublished data). Moreover, the transient kinetics shows that binding occurs in two steps: an initial weak collision complex is formed (Kd=650 µM) which triggers a rapid rearrangement, at a rate of 720 s−1, to a conformation which is very different from the ATP-stabilized R-conformer with respect to its kinetic behaviour. While ATP drives the conformational equilibrium heavily in favour of the rearranged state, the ADP-stabilized conformer remains in a rapid and


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poised equilibrium with the initial collision state, with a reverse conformational rate of 60 s−1. Hence, the binding-rearrangement reaction for the binding of the first 7 ADPs can be described:

The ADP-stabilized conformer is shown by cryo-electron microscopy to be different in structure from that generated by the association of ATP (Roseman et al., 1996) and retains a high affinity for unfolded protein substrates (Staniforth et al., 1994a). 5. THE GroEL ATPase CYCLE IN THE ABSENCE OF GroES In the absence of GroES the steady-steady rate of ATP turnover by GroEL is 0.04–0.06 s1 per subunit or 0.08–0.12 s-1 per active subunit (Viitanen et al., 1990; Jackson et al., 1993; Todd et al., 1993; Horovitz et al., 1993). The rate-determining step in the steadystate ATPase cycle was found to be the hydrolytic reaction by the following reasoning, (i) The single exponential rate constant for a single turnover reaction (7 ATPs per complex) is identical to the steady-state rate constant per active site and there is no burst of hydrolysis preceding the steady-state phase. These observations demonstrate that the release of products is non-limiting, (ii) Conformational changes induced by ATP binding are rapid in comparison to the steady-state rate, showing that steps preceding bond breakage cannot be limiting (Jackson et al., 1993; Todd et al., 1994, Burston et al., 1995). The above observations of nucleotide binding, conformational switches and hydrolysis suggest the following model for the ATPase cycle of GroEL shown in Figure 7. Asymmetric association of ATP to one of the two rings of GroEL occurs in a positively co-operative manner (Gray & Fersht, 1991; Bochkareva et al., 1992; Jackson et al., 1993), and promotes a conformational shift from a T-state with low affinity for ATP, to an R-state with a high affinity (Jackson et al., 1993; Yifrach & Horovitz, 1994 & 1995). Subsequent hydrolysis occurs at 0.12 s-1 per active subunit (Burston et al., 1995), the slowest step in the reaction cycle. The resulting 7 molecules of ADP are stably bound (Burston et al., 1995) and a second seven molecules of ATP bind to the unoccupied ring. If ATP can hydrolyse on one ring of GroEL while the other is occupied by ADP, then the next round of hydrolysis creates a symmetrical GroEL14: ADP14 complex which is unstable (Jackson et al., 1993) owing to strong inter-ring negative co-operativity, and promotes dissociation of 7 ADPs from the complex. However, recent results show unambiguously that ATP may only turn over upon dissociation of ADP from the other ring. This phenomenon is demonstrated in experiments in which the hydrolysis of ATP on one ring is shown to be completely blocked by the binding of ADP to the other. ADP does not prevent binding of ATP to the opposite ring, but it does prevent turnover, i.e. the mode of inhibition is asymmetric and non-competitive (Kad et al., 1998). Such a mechanism provides the basis of a reciprocating, ring-switching mechanism as depicted in Figure 7, where ATP hydrolysis is carried out alternately by each GroEL ring.


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This series of reactions imposes an obligatory change in the conformation of each of the GroEL rings during a revolution of the ATPase cycle, i.e. each GroEL ring is driven reciprocally between states which have a high and a low affinity for substrate proteins.

Figure 7 The GroEL ATPase cycle. Starting at the bottom right, ATP binds to a GroEL: ADP ring to form an asymmetric ADP7: GroEL14: ATP7


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complex. ADP must dissociate before hydrolysis can proceed. This series of reactions enforces ATP hydrolysis on alternating GroEL rings. Broken rectangles represent unliganded, and solid rectangles represent ADP-liganded GroEL rings respectively. Both species have a high affinity for non-native protein substrate. ATP liganded rings with low affinity for non-native protein substrate are represented by ovals. Note that this cycle represents the system at low ATP concentrations where the possibility of generating an ATP14 complex is avoided.

6. GROES-GROEL INTERACTIONS AND THEIR EFFECTS ON THE HYDROLYSIS OF ATP 6.1. Binding of GroES to the ADP and ATP States of GroEL While apo-GroEL and GroES do not interact, the addition of adenine nucleotides generates two types of GroEL: GroES complex observable by electron microscopy. Depending on the reaction conditions, the presence of ATP can create either an asymmetric structure with GroES bound to only one of the two GroEL toroids (Saibil et al., 1993; Chen et al., 1994; Roseman et al., 1996), or a symmetrical complex with GroES bound at each end of the GroEL tetradecamer (Llorca et al., 1994; Azem et al., 1995; Llorca et al., 1997; Behlke et al., 1997). With ADP only the former, asymmetric complex is formed (Roseman et al., 1996; Xu et al., 1997). In an attempt to understand the mechanistic features of GroEL/GroES interactions and to elucidate their functional relevance, the dynamic and equilibrium properties of such complexes, formed in the presence of each nucleotide, have been studied in considerable detail. The association of GroES with pyrene-labelled GroEL in the presence of ADP leads to a large enhancement of the extrinsic fluorescence which can be used to report the formation of the asymmetric (1:1) protein-protein complex (Jackson et al., 1993). This process occurs slowly, yielding a bimolecular rate constant of 1×105 M-1 s-1. Incubation of the components at micromolar ADP concentrations requires many hours to allow the GroEL-GroES interaction to reach equilibrium. In the equilibrated system the binding affinity of ADP in the GroEL: GroES complex is extremely high with a dissociation constant below 70 nM and a stoichiometry of 7. Thus, although the GroEL: ADP7: GroES complex forms very slowly, it is extremely stable. This conclusion is confirmed by the low rate of dissociation of this complex (4×10-3 s-1) (Burston et al., 1995) and the low rate of exchange of radio-labelled ADP and GroES (3×10-5 s-1) (Todd et al., 1994). However, in the presence of ATP the formation of the GroEL: GroES complex is much faster. Shown in Figure 8 is a stopped-flow fluorescence experiment in which a subsaturating concentration of ATP (20 M) is mixed with pyrene-labelled GroEL in the presence and absence of GroES. With no GroES in the system, the transient obtained is described by a single rate constant of 2.8 s-1 representing the rate of rearrangement of GroEL to the R-conformation at low ATP concentration. Addition of either 0.375 or 3 M GroES introduces a new phase into the fluorescence enhancement which occurs at 16s-1, i.e. it is rapid and


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insensitive to the concentration of the co-chaperonin. This result is compatible with the following reaction scheme:

Figure 8 Stopped flow fluorescence of the binding of ATP and GroES to pyrenyl-GroEL. Sub-saturating concentrations of ATP (20 M) cause a sub-maximal fluorescence enhancement in pyrenyl-GroEL. Subsequent binding of GroES is then reported by a greater enhancement of fluorescence as discussed in the text.

GroES must bind to the GroEL14: ATP7 complex very rapidly and before the transition to the R-conformation to yield GroEL14: ADP7 GroES7. The rate of rearrangement of this complex, to give the high fluorescence GroEL (R)14: ATP7: GroES7 form, is faster than in the absence of GroES and is not limited by the prior association of GroES, since increasing its concentration does not alter the kinetics. The co-protein must associate with the GroEL14: ATP7 complex at a rate faster than 16 s-1 in these conditions. This corresponds to a collision rate constant in excess of 4×107 M-1 s−1 (i.e. >16 s−1 / (0.375×10−6 M)) which approaches the upper limit for protein diffusion in aqueous solution and is at least two orders of magnitude faster than the formation of the GroEL14: ADP7: GroES7 complex. Although these kinetic observations appear somewhat arcane, they may have considerable bearing on the dynamics of the ATPase cycle and its relationship to the folding mechanism. Firstly they show that GroES has a kinetic preference for binding to an ATP ring, and secondly that the co-protein can bind before the rearrangement of subunits which leads to the low protein affinity R-conformation. With respect to the overall chaperonin mechanism this provides a crucial ‘trapping’ reaction in which a


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stably bound protein substrate could be encapsulated by GroES before being displaced into the cavity by the ATP-driven rearrangement of the GroEL apical domains. Although kinetic experiments shed no light on whether the toroid structure associated with the formation of the binary complex (GroEL14: ATP7) is different from that found in the ternary species (GroEL14: ATP7: GroES7), it should be noted that the structure of GroEL in these two complexes appears different under the electron microscope (Chen et al., 1994; Roseman et al., 1996) (see Burston and Saibil, this volume). In keeping with the ability of GroES to bind to GroEL prior to the hydrolysis of ATP and to influence conformational transitions, GroES has significant effects on the measured ATP binding properties of GroEL as shown by enhanced Hill coefficients in both steady-state and equilibrium binding studies (Gray & Fersht, 1991; Jackson et al., 1993; Kovalenko et al., 1994). 6.2. ATP Hydrolysis in the Presence of GroES When GroEL, GroES and ATP are mixed and the rate of formation of total ADP/ Pi. (both free and bound) is measured, there is a first-order ‘burst’ of ATP hydrolysis occuring at a rate of 0.12 s-1 with an amplitude equal to 7 sites per GroEL oligomer, followed by a steady-state phase which proceeds at rate of 0.042 s-1 per active site, i.e. half the sites in the oligomer (Todd et al., 1994; Burston et al., 1995). Thus, despite the presence of GroES in the complex, hydrolysis initially occurs at the same rate as is seen in the absence of co-protein, implying that a new, non-hydrolytic ratelimiting step has been introduced into the reaction cycle. To confirm this, isolated single-turnover reactions were designed to examine the next hydrolytic step. The transient phase of the above experiment describes the rate of ATP hydrolysis by a GroEL ring to which GroES is bound; it thus reports the rate of conversion of the GroEL14: ATP7: GroES7 complex to produce GroEL14: ADP7: GroES7. When this product complex was challenged with a further 7 ATP molecules to form the ATP7: GroEL14: ADP7: GroES7 species, hydrolysis of ATP on the ring distal to GroES occurred in a single-exponential phase with a rate constant of 0.12 s-1 (Burston et al., 1995). Hence, the hydrolytic step in the presence of GroES, whether on the ring proximal or distal to the co-chaperonin, occurs at a rate of 0.12 s-1. This rate is identical to the rate of hydrolysis in GroEL alone, yet the steady-state rate of the hydrolytic cycle in the GroEL: GroES system is two-to-three times slower. The fact that the rate-determining step is not observed when the isolated turnover reactions are carried out, shows that either a product release step or a structural rearrangement is the slowest process in steady-state hydrolysis in the presence of the co-protein. The relationship of this step to the rate of dissociation of GroES is discussed below. 6.3. The Dissociation of GroES During the ATPase Cycle Lorimer and colleagues have carried out a comprehensive study of the GroEL ATPase cycle (Todd et al., 1994) which provides extensive information on the lifetimes of a number of complexes containing GroEL, GroES and nucleotide. They show, by exchange of radio-labelled ADP and GroES, that the asymmetric GroEL14: ADP7: GroES7 complex has a half-life of approximately five hours. However, the addition of ATP reduces its


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half-life to about 15 seconds, commensurate with the turnover of a single ring of ATP. Addition of the non-hydrolysable ATP analogues ATP S or AMP-PNP also promotes complex dissociation, although somewhat more slowly (t1/2 of ~60 seconds). Their results show that the dissociation of GroES

Figure 9 The rate of decay of GroEL: GroES complexes followed by pyrene fluorescence. Decay is promoted by either ATP hydrolysis (k=0.042 s−1) or by addition of alkaline phosphatase (k=0.004 s−1). For discussion see text. Adapted from Burston et al., 1995.

and ADP from a GroEL ring is linked to the binding and/or hydrolysis of ATP on the opposite ring. Further, if ATP is bound on the GroES-associated ring of GroEL it is committed to hydrolysis, i.e. the rate of cleavage is faster than the rate of dissociation, and the GroES remains tightly associated through the hydrolytic step. Similarly, the rate of dissociation of the chaperonin complex has been determined directly by pyrene fluorescence (Burston et al., 1995). The addition of a sub-saturating concentration of ATP to a solution of pyrene-labelled GroEL leads to an increase in fluorescence intensity which is further enhanced by the addition of GroES (see Figure 9). The ATPase reaction then reaches a steady state with a high-fluorescence GroEL: GroES: nucleotide complex predominating. If a large excess of unlabelled GroEL is then added, any GroES dissociating from the labelled protein will rebind to the unlabelled GroEL which then serves as an optically silent GroES trap. As shown in Figure 9, the rate of loss of fluorescence fits to a single-exponential decay at a rate of ~0.04 s−1, and the fluorescence returns to that of the GroEL: ATP complex. This rate is the same as the steady state velocity of ATP hydrolysis in the presence of GroES. If, instead of adding the unlabelled trap, all free and rapidly dissociating (uncommitted) ATP and ADP is destroyed by the addition of a large concentration of alkaline phosphatase, the GroEL: GroES complex decays very slowly. Hence, in the steady state reaction, GroES obligatorily dissociates once per cycle at a rate coincident with the rate-limiting step and


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this requires binding of ATP to the ring distal to GroES. 6.4. Bullets and Footballs One of the more controversial questions concerning the GroE hydrolytic cycle is the nature of the GroES exchange intermediate, i.e. it is indisputable that GroES binds to and dissociates from GroEL on each turnover of the steady-state ATPase reaction, but it is an attractive proposition that this process is mediated by a quasi-symmetrical GroES7: GroEL14: GroES7 complex in the following way:

In such a mechanism GroES is forced to exchange from one ring of GroEL to the other through an intermediate which is symmetrical with respect to the co-chaperonin but not the nucleotide. This process would open and close each cavity on successive rounds of ATP turnover. Evidence from electron microscopy shows that two types of GroEL: GroES are formed. The asymmetrical “bullet shaped” complex with GroES bound to one of the two available GroEL rings was observed in the first electron micrographs of GroE complexes (Hendrix, 1979), and has since been extensively characterized (Chen et al., 1994; Roseman et al., 1996; Xu et al., 1997). More recently, a symmetrical complex shaped like an American football with GroES bound to each end of the GroEL tetradecamer (Llorca et al., 1994; Schmidt et al., 1994; Llorca et al., 1997) has been imaged. Hence, the fact that both complexes exist is not in question, but the significance of the “football” intermediate has been the subject of intense debate, with evidence both for and against the symmetrical GroES7: GroEL14: GroES7 complex being an important intermediate in the biologically functional GroE reaction cycle. Two lines of evidence have been cited in favour of a symmetrical intermediate. Firstly, an analysis of the rates of refolding of several chaperonin dependent substrate proteins as a function of the GroEL: GroES stoichiometry has been completed (Azem et al., 1995). This study concludes that the rate of refolding is maximal in conditions where the symmetrical complex is found, as judged by negative stain electron microscopy and glutaraldehyde cross-linking studies. Secondly, a comprehensive study of the kinetics of the chaperonin-assisted refolding of barnase shows that GroES: GroEL stoichiometries in excess of 1 introduce a faster phase of folding which rises to 50% of the total folding amplitude at a 2:1 stoichiometry (Corrales & Fersht, 1996a). Against this, when the dissociation of GroES in steady-state hydrolysis is measured using a GroES trap (Burston et al., 1995) the rate of 0.04 s-1 is unaltered when the GroES: GroEL stoichiometry is reduced below 1:1. Additionally, in this experiment all free GroES is rapidly absorbed onto the trap, thus preventing rebinding to the labelled GroEL. Similarly, surface plasmon resonance studies by Hartl and colleagues (HayerHartl et al., 1995) indicate that if a stable GroEL14: ADP7: GroES7 complex is formed, addition of ATP causes dissociation of the complex via hydrolysis on the GroEL ring to which GroES is not bound, without binding of a second molecule of GroES. The latter pair of experiments show that the formation of a “football” intermediate is not an obligatory step in discharging a GroES ring during the ATPase cycle, i.e. they


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demonstrate the existence of the following reaction:

However, the results do not show that such a structure is not formed as an exchange intermediate when GroES is in excess. In conclusion, the mechanistic significance of the symmetrical GroES7: GroEL14: GroES7 intermediate remains somewhat unclear, and further experiments are required if its role in assisted protein folding is to be fully elucidated. In particular it is necessary to examine the mechanism in reaction conditions which mimic the ATP: ADP and GroEL: GroES ratios and concentrations within the cell. Recent experiments with a GroEL mutant in which hydrolysis of ATP occurs very slowly have clarified the nature of the step in which GroES is released (Rye et al., 1997). When the ADP stabilized GroEL-GroES complex (GroEL: ADP7: GroES) is challenged with ATP, the co-protein is released at a much faster rate than the ATP can be hydrolysed. Hence GroES dissociation is driven by the binding of ATP to the opposite ring and is mediated by the consequent structural rearrangement of GroEL. Given the experimental results described in the above sections, it is tempting to propose a concensus model for the ATPase reaction mechanism at the macromolecular level. Such a model is forced to account for the following phenomena: (1) Asymmetry of nucleotide interactions. (2) The action of ADP as a non-competitive inhibitor of ATP hydrolysis. (3) The obligatory dissociation and reassociation of GroES in the hydrolytic cycle. (4) The role of GroES in committing bound ATP to undergo hydrolysis. (5) The ability of ATP to drive GroES dissociation from the opposite ring prior to its hydrolysis. (6) The kinetic preference for GroES to bind to an ATP-occupied ring. The reaction scheme shown in Figure 10 represents a minimal model for the reaction steps which constitute the GroE reaction cycle. 7. THE COUPLING OF PROTEIN AND NUCLEOTIDE BINDING AFFINITIES 7.1. The Effects of ATP, ADP & AMP-PNP on Protein Binding Affinity The affinity of GroEL for protein substrates in different stages of folding has long been a matter of intense interest. An early study concluded that apo-GroEL had


557

Figure 10 The GroEL ATPase cycle in the presence of GroES. Binding of ATP (T) to an unliganded GroEL ring (rectangle) promotes a conformational shift to a state with low affinity for non-native protein (oval) and promotes GroES dissociation from the opposite ring. Binding of GroES occurs rapidly to a GroEL: ATP ring creating the likely acceptor state for non-native protein. A round of ‘committed’ hydrolysis occurs to yield a stable GroEL: ADP7: GroES complex. Subsequent binding of ATP promotes dissociation of GroES from the distal ring and rebinding of GroES to the ATP occupied ring would lead to encapsulation of the protein substrate. Hydrolysis of the


558

bound ATP and rebinding of ATP to the opposite ring would lead to substrate release. As in Figure 7 this represents the system at low ATP concentrations. It is possible that at high ATP concentrations GroES: GroEL: ATP7 complex should be added to the group of species (shown in square brackets) which accumulates before the hydrolytic step.

the highest affinity for the most unfolded intermediate on the refolding pathway of an oligomeric enzyme, and a much lower affinity for the late, compact, monomeric folding intermediate which immediately precedes dimerization (Badcoe et al., 1991). A considerable body of evidence has now been accumulated which suggests that GroEL recognizes a reasonably well-structured and rapidly formed folding intermediate often referred to as the “molten globule” state (e.g. Martin et al., 1991; Hayer-Hartl et al., 1994). However, the definition of which protein states are bound tightest may be somewhat arbitrary and case-dependent, since an incisive study of the conformation of barnase bound to GroEL using proton-deuterium amide exchange kinetics measured by NMR (Zahn et al., 1996) concludes that both the fully unfolded and the partially structured intermediate state interact tightly. Most would agree that any conformational state which has extensive exposure of hydrophobic surface will bind with high affinity. Estimates of the dissociation constants for complexes between non-native proteins and apo-GroEL vary from 10–12 M for large polypeptides up to 10−8 M for small ones (Badcoe et al., 1991; Todd et al., 1994; Staniforth et al., 1994; Lilie & Buchner, 1995; Sparrer et al. 1996; Zahn et al., 1996). Regardless of which type of pre-native state is recognised by GroEL, the effect which nucleotide binding has on the protein binding affinity has been studied extensively. One of the continuing themes of this discussion of the GroEL ATPase cycle has been the cooperative binding of ATP triggering a shift from a T-state with a low affinity for ATP, to an R-state with a high affinity. This rearrangement causes an accompanying change in the protein-binding affinity of GroEL, with the T-state having high, and the R-state having low affinity for the unfolded protein (Badcoe et al., 1991; Jackson et al., 1993). This proposal has been examined by measuring the affinity of GroEL for lactate dehydrogenase (LDH) through the retardation of the substrate folding rate (Staniforth et al., 1994a). The affinity of GroEL for unfolded LDH is tightest to the apo and the ADP states, is weaker to GroEL: AMPPNP and weakest to GroEL: ATP (see Table 1). This is entirely in accordance with the proposal that the affinity of GroEL for protein and ATP are reciprocally coupled. Accordingly, the GroEL ATPase cycle will enforce a switch between weak and tight protein binding state. The conclusion that ATP triggers a weakening of substrate binding by two or three orders of magnitude has also been made by Buchner and colleagues using surface plasmon resonance as a probe for complex formation (Lilie & Buchner, 1995; Sparrer et al. 1996). The presence of GroES further weakens substrate binding, thus potentiating the effect of ATP (Staniforth et al., 1994; Schmidt et al., 1994).


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Table 1 Role of nucleotide and GroES on the affinity of GroEL for unfolded LDH

Nucleotide None

−GroES

+GroES 5 nM

5 nM

ADP (4 mM)

18 nM

25 nM

AMP-PNP (4 mM)

53 nM

83 nM

ATP

118nM

439 nM

The practical consequences of this nucleotide-induced modulation of protein binding affinity is to facilitate a cycle of binding and release of protein substrates from GroE complexes. The release of substrate has been characterized by using mutant GroEL protein traps (Fenton et al., 1994) which bind but do not release non-native proteins, even in the presence of ATP and GroES. These experiments show that rhodanese and ornithine transcarbamylase (OTC) are expelled from the chaperonin in a non-native state (Weissman et al., 1994) and that many cycles of binding and dissociation must occur before the folded, chaperonin-independent state is acquired. Similar findings have been made by monitoring the distribution of isotopically labelled rubisco after a single round of ATP hydrolysis (Todd et al., 1994), by using chemically modified GroEL (Taguchi & Yoshida, 1995; Ranson et al., 1997), and by analysis of the kinetics of rhodanese refolding (Smith & Fisher, 1995). 7.2. The Effect of Unfolded Protein on Co-operativity Within the GroEL Oligomer The analysis of co-operativity in the GroEL ATPase by Horovitz and co-workers has been advanced still further by directly examining the effect of unfolded protein on these co-operative transitions (Yifrach and Horovitz, 1996). The stimulation of the GroEL ATPase by addition of non-native protein has been observed previously (Martin et al., 1991; Jackson et al., 1993). However, these measurements were made by addition of non-native protein which then proceeded to refold, and resulted in a highly complicated kinetic system. In experiments using a stably unfolded protein substrate, the effect of protein binding on the steady-state ATPase activity of GroEL has been determined. If the binding of one ligand (e.g. ATP) to a protein, in a co-operative manner, affects the affinity of the protein for a second ligand (unfolded protein), then the reverse should also be true. This is precisely the outcome they observed; stably unfolded -lactalbumin (in a reduced, calcium depleted form) binds preferentially to the T state and pushes the equilibrium back


560

from the RR state towards the TR and TT states. The fact that T state rings have a higher ATPase activity means that an unfolded substrate will enhance the rate of turnover. This is a possible mechanism for adapting the GroEL molecule to a role in responding to cellular stress; as the concentration of misfolded protein increases, binding to GroEL is made more likely. 8. THE COUPLING OF THE ATPase CYCLE TO CHAPERONIN-ASSISTED PROTEIN FOLDING So far, we have concentrated on the kinetic properties of the ATPase cycle and the characteristic complexes which arise therein. In this section we consider potential mechanisms for the coupling of ATP turnover to protein folding in a way which will enhance the folding yield. Rapid progress has recently been made in elucidating how the nucleotide-driven switches in affinity, for both GroES and unfolded substrates, may account for the ability of GroE to force protein substrates to fold productively and two main models of the GroE mechanism now predominate. 8.1. Models of Chaperonin Action 8.1.1. Encapsulation and Folding In an encapsulation model the unfolded substrate protein enters the central cavity of GroEL and remains there until committed to fold. It was originally proposed that DHFR and rhodanese reached the native state while associated with the GroEL-GroES complex owing to the inability of casein to block refolding when the GroES capping protein was included in the reaction (if substrate proteins were leaving GroEL in the reaction cycle, then the presence of casein should block refolding by blocking the rebinding of substrate protein to the binding sites of GroEL) (Martin et al., 1991). Several experiments localised the binding of substrate protein to the central cavity of GroEL (Langer et al., 1992; Braig et al., 1993) and further experiments led to the proposal of a detailed model of chaperonin action which involved rebinding and release within this cavity until the native state or near-native state was attained, after which the polypeptide was released into bulk solution (Martin et al., 1993). An electron microscopy study visualised an expansive GroES-capped GroEL cavity and introduced the term “Anfinsen cage” to describe protein folding within this protected space in which the substrate is prevented from making incorrect intermolecular interactions which lead to irreversible aggregation (Martin et al.,


561

1991; Saibil et al., 1993). 8.1.2. Unfolding and Release The unfolding model arose from the argument that GroEL has its highest affinity for unfolded states of proteins and that protein-protein binding energy may be used to disrupt non-native interactions (Hubbard & Sander, 1991; Jackson et al., 1993; Todd et al. 1994; Ranson et al., 1995; Chan & Dill, 1996, Sfatos et al., 1996). In this hypothesis, an enhancement of folding yield could be achieved by actively unfolding misfolded proteins in a tight binding phase, while in the weak phase they are released to allow them to refold spontaneously. Hence, if a population of unfolded molecules is transferred to folding conditions, a proportion will fold to a highly organised native-like state which does not interact tightly with GroEL and goes on to fold spontaneously. The rest becomes trapped in unproductive, misfolded conformations and interact tightly with GroEL by virtue of greater exposure of hydrophobic residues. Having been bound they are unfolded, and then released in an ‘untrapped’ state and given a second chance to acquire a native-like structure. Repetition of this cycle will deliver all molecules to their native structure. Several lines of evidence support this proposal, including the switch in affinity between tight and weak protein binding states (Jackson et al., 1993), the release of non-native protein by GroEL (Weissman et al., 1994; Todd et al., 1994; Taguchi & Yoshida, 1995; Burston et al., 1996; Ranson et al., 1997) and the degree of enhancement of amide exchange in GroEL: substrate complexes (Zahn et al., 1994; Zahn et al., 1996). However, both proposals are attractive in that they account for the absence of substrate specificity in GroEL and explain the enhancement of yield without the penalty of slowing down the folding reaction unduly (see section 2). Also, both mechanisms require an input from ATP binding and hydrolysis; in the encapsulation mechanism it provides the energy to capture the substrate and open and close a GroES-capped cavity while in the unfolding/release hypothesis this energy is required to bind, unfold and discharge. 8.2. Towards a Global Model for GroE-mediated Folding Recent developments have begun to tie together these seemingly disparate models, and produce a more unified view of the action of GroE (for reviews see Ellis & Hartl, 1996; Clarke, 1996; Hartl, 1996). By a series of order-of-addition and protease digestion/protection experiments, the productive complex in chaperonin-assisted protein folding has been isolated as being a as complex of GroEL, GroES and substrate protein, with the GroES and substrate protein located on the same side of the GroEL tetradecamer (Weissman et al., 1995). They have also subsequently shown that substrate protein can reach the native state in this complex, by using mutant GroEL molecules which cannot release GroES once it has bound (Weissman et al., 1996). These results have been mirrored by those of Hartl and colleagues who have shown that a monomeric substrate protein can reach the native state whilst covalently tethered within the GroEL cavity (Mayhew et al., 1996). This conclusion is underlined by a kinetic analysis of the refolding of mMDH at a range of concentrations of GroEL: GroES. These results show that the GroEL: GroES complex to which unfolded mMDH binds in the assisted, ATP-driven reaction has a high


562

substrate affinity with an effective KM of ~10-8 M and a stoichiometry of 1:1. At an mMDH concentration of 1 M and a GroEL: GroES concentration in the range 0 to 1 µM the observed rate of folding increases owing to the disruption of misfolded enzyme in the form of low molecular weight aggregates (Ranson et al., 1995). The rate of folding then remains constant up to 40 M GroEL: GroES. At these elevated concentrations of chaperone, only 0.025% of mMDH is in free solution yet there is no diminution of the observed folding rate (Ranson et al., 1997). Recent work (Rye et al., 1997) has confirmed that both mMDH and bacterial rubisco reach a native-like state within the chaperonin cage, and oligomerise upon release to form the native, active enzyme complex. These observations provide convincing evidence that (1) the substrate protein can reach the folded, monomeric state whilst associated with GroEL and (2) the nature of the association allows rapid folding i.e. the substrate must be encapsulated rather than bound, so that folding is not inhibited by tight interactions between GroEL and the unfolded state. The previously reported release of non-native protein is also seen in the mMDH system with a dissociation rate of 0.04 s−1. That is, the complex ejects both GroES and the substrate protein once every cycle of ATP hydrolysis (Burson et al., 1996; Ranson et al., 1997). At first sight, it appears somewhat paradoxical that mMDH can fold to a committed, monomeric state in association with the GroE complex, yet is continually being ejected into bulk solution to be rebound by GroE to go through further cycles of encapsulation and obligatory ejection. However, such a mechanism will prevent covalently ‘damaged’ substrates (which are unable to fold) from saturating the GroE apparatus by remaining stably bound, rather than being degraded through proteolysis (Weissman et al. 1996; Mayhew et al., 1996; Ranson et al., 1997). The general dynamics of the binding, encapsulated folding and ejection cycle with respect to the renaturation of mMDH (Ranson et al., 1997) are shown in Figure 11. GroEL, therefore, appears to bind its substrate proteins in trans to GroES within a high-affinity ring; the presence of GroES would occlude the binding site for substrate protein on the cis ring (Chen et al., 1994; Fenton et al., 1994). This binding step will afford an opportunity for incorrect interactions to be reversed (Jackson et al., 1993; Todd et al., 1994; Ranson et al., 1995; Todd et al., 1996; Walter et al., 1996; Corrales & Fersht, 1996; Zahn et al., 1996a&b). Subsequent ATP binding leads to a reduction in substrate binding affinity and the rapid association of GroES. This displaces the substrate from its binding surfaces and encapsulates it in a cis complex (Weissman et al., 1995). The encapsulation phase may proceed through a symmetrical intermediate with GroES bound to each end of the GroEL oligomer (Llorca et al., 1994; Schmidt et al., 1994; Todd et al., 1994; Azem et al., 1995) or via ATP-driven dissociation and rebinding of GroES (Burston et al., 1995; Hayer-Hartl et al., 1995)—see Figure 10. The substrate will then dwell in the cavity and have an opportunity to fold without inhibitory contacts with hydrophobic surfaces on the apical domains (Weissman et al., 1995; 1996; Mayhew et al., 1996; Burston et al., 1996; Ranson et al., 1997). The proportion of molecules able to fold per ‘visit’ will be dependent solely on the ratio of the lifetime of the cis complex and the rate of protein folding. In the case of mMDH ~3% of polypeptide chains fold to a committed, GroE-independent state per cycle. For fast folding subunits this percentage may become quite large. For example, the subunits


563

which constitute the glutamine synthetase dodecamer com-mit to the folded state at a rate of ~0.09 s−1 (Fisher & Yuan, 1994), so that 70% will fold in a single cycle. Further turnover of ATP forces the cavity to open with the expulsion of GroES, ADP and the protein substrate whether committed or not (Burston et al., 1996; Ranson et al., 1997; Rye et al., 1997). The possible fates of ejected, non-native protein are diverse. It can rebind to another (or the same) chaperonin molecule to attempt another round of encapsulated folding. It also has an opportunity to fold in free solution depending on the time between encounters with GroE. At physiological

Figure 11 The dynamics of mMDH binding, encapsulation and folding by GroEL.

concentrations of GroE, however, the transit time between one encounter with GroE and the next is much shorter than the dwell time in the encapsulated state. Thus owing to the fact that sponatneous folding occurs at an intrinsic and fixed rate in either environment, a minority of molecules will reach their native states when free in solution. Thirdly, the release of non-native substrate protein ensures that damaged proteins which can never refold can be released from GroEL into the bulk solvent, rather than rapidly “outtitrating” the cell’s chaperonin capacity upon cellular stress. Once in solution, they can partition to the proteolytic apparatus of the cell, a process in which GroEL is been implicated in vivo (Kandror et al., 1994). 9. CONCLUDING REMARKS This discussion of the GroEL ATPase cycle and its connection with assisted protein folding began with a question: What is the role of ATP binding and hydrolysis in aiding a thermodynamically favourable process? The answer lies in the enforced rearrangements of GroEL which are imposed by its differential interaction with ATP and ADP. These


564

serve two purposes: firstly, the binding and hydrolysis of ATP provides energy to drive formation and disruption of the chaperonin cage in which assisted-folding takes place (as summarized in Figure 12). Secondly, it provides energy to disrupt the tight interaction between substrate protein and GroEL which

Figure 12 Coupling of the ATPase cycle with substrate binding and ejection. The substrate is bound by an open end of an asymmetric GroEL: GroES complex (step (I)) where the acceptor ring is most likely to be in a tight binding, ADP-induced conformation. ATP hydrolysis on the opposite ring (step (ii)) then allows ATP to associate with the substrate-containing site. This has three consequences (step (iii)); it forces dissociation of ADP and GroES from the opposite ring, it weakens interactions with the substrate and allows reassociation of GroES on the substrate side to displace the unfolded protein molecule into the cavity where productive folding can occur. ATP is then hydrolysed (step (iv)) a process which then allows ATP to associate with the trans ring and eject GroES, ADP and the protein substrate from the opposite ring to complete the cycle.


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is used to reverse incorrect interactions. GroES has three major roles. It co-ordinates ATP hydrolysis in GroEL, strengthening co-operativity and therefore potentiating the conformational shift between states with a high and low affinity for substrate protein. Secondly it commits ATP to hydrolysis and serves to maintain a reciprocating cycle of reactions. Thirdly, its binding to GroEL displaces substrate protein into a cavity, which GroES itself is essential in forming. The question of whether the major function of GroE in the cell is to disrupt misfolded states or to encapsulate sticky folding intermediates, thereby preventing aggregation, is difficult to answer unequivocally. Both activities have been demonstrated with different substrates and both activities may be inevitable consequences of the mechanism described here. 10. ACKNOWLEDGEMENTS The authors would like to thank the Wellcome Trust for Programme Grant Support for their work. A.R.Clarke is a Lister Institute Research Fellow. 11. REFERENCES Azem, A., Diamant, S., Kessel, M., Weiss, C. and Goloubinoff, P. (1995). The protein refolding activity of chaperonins correlates with the symmetrical GroEL14: (GroES7)2 heteroligomer. Proc. Natl. Acad. Sci. USA. , 92 , 12021–12025. Badcoe, I.G., Smith, C.J., Wood, S., Halsall, D.J., Holbrook, J.J., Lund, P. and Clarke, A.R. (1991). Binding of a chaperonin to the folding intermediates of lactate dehydrogenase. Biochemistry , 30, 9195–9200. Behlke, J., Ristau, O. and Schonfeld, H.J. (1997). Nucleotide dependent complex formation between the Escherichia coli chaperonins GroEL and GroES studied under equilibrium conditions. Biochemistry , 36, 5149–5156. Bochkareva, E.S. and Girshovich, A.S. (1994). ATP induces non-identity of two rings in chaperonin GroEL. J. Biol. Chem ., 269 , 23869–23971. Bochkareva, E.S. Lissin, N.M., Flynn, G.C., Rothman, J.E. and Girshovic, A.S. (1992). Positive cooperativity in the functioning of chaperonin GroEL. J. Biol. Chem ., 267 , 11637–11644. Braig, K., Otwinowski, Z., Hegde, R., Boisvert, D.C.Joachimiak, A., Horwich, A.L. and Sigler, P.B. (1994). The crystal structure of the bacterial chaperonin GroEL at 2.8Å. Nature , 371, 578–586. Braig, K., Simon, M., Furuya, F., Hainfeld, J.F. and Horwich, A.L. (1993). A polypeptide bound by the chaperonin GroEL is localized within a central cavity. Proc. Natl. Acad. Sci. USA , 90 , 3978–3982. Buchner, J., Schmidt, M. Fuchs, M. Jaenicke, R., Rudolph, R., Schmid, F.X. and Kiefhaber, T. (1991). GroE facilitates refolding of citrate synthase by supressing aggregation. Biochemistry , 30, 1586–1591. Burston, S.G., Weissman, J.S., Farr, G.W., Fenton, W.A. and Horwich, A.L. (1996). Release of both native and non-native proteins from a as-only GroEL ternary complex.


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580. Hayer-Hartl, M.K., Martin, J. and Hartl, F.U. (1995). Asymmetrical interaction of GroEL and GroES in the ATPase cycle of assisted protein folding. Science , 269, 836–841. Hayer-Hartl, M.K., Ewbank, J.J., Creighton, T.E. and Hartl, F.U. (1994). Conformational specificity of the chaperonin GroEL for the compact folding intermediates of lactalbumin . EMBO J. , 3, 3192–3202. Hendrix, R.W. (1979). Purification and properties of groE, a host protein involved in bacteriophage assembly. J. Mol. Biol. , 129, 375–392. Horovitz, A., Bochkareva, E.S., Kovalenko, O. and Girshovic, A.S. (1993). Mutation Ala2ÆSer estabilizes intersubunit interactions in the molecular chaperone GroEL. J. Mol. Biol. , 231, 58–4. Hubbard, T.J.P. and Sander, C. (1991). The role of heat-shock and chaperone proteins in protein folding—possible molecular mechanisms. Prot. Eng. , 4, 711–717. Itzhaki, L.S., Otzen, D.E. and Fersht, A.R. (1995). Nature and consequences of GroELprotein interactions. Biochemistry , 34 , 14581–14587. Jackson, G.S., Staniforth, R.A., Halsall, D.J., Atkinson, T., Holbrook, J.J., Clarke, A.R. and Burston, S.G. (1993). Binding and hydrolysis of nucleotides in the chaperonin catalytic cycle: Implications for the mechanism of assisted protein folding. Biochemistry , 32, 2554–2563. Kad, N.M., Ranson, N.A., Cliff, M. and Clarke, A.R. (1998). Asymmetry, commitment and inhibition in GroE ATPase cycle impose alternating functions upon the two GroEL rings. J. Mol. Biol. , 278, 267–278. Kandror, O., Busconi, L., Sherman, M. and Goldberg, A.L. (1994). Rapid degradation of an abnormal protein in Escherichia coliinvovles the chaperones GroEL and GroES. J. Biol. Chem. , 269, 23575–23582. Langer, T., Pfeifer, G., Martin, J., Baumeister, W. and Hartl, F.U. (1992). Chaperoninmediated protein folding—GroES binds to one end of the GroEL cylinder, which accomodates the protein substrate within its central cavity. EMBO J. , 11 , 4757–4765. Llorca, O., Marco, S., Carrascosa, J.L. and Valpuesta, J.M. (1994). The formation of symmetrical GroEL-GroES complexes in the presence of ATP. FEBS Lett. , 345 , 181– 186. Llorca, O., Marco, S., Carrascosa, J.L. and Valpuesta, J.M. (1997). Symmetric GroELGroES complexes can contain substrate in both GroEL rings. FEBS Lett. , 405 , 195– 199. Martin, J., Mayhew, M., Langer, T. and Hartl, F.U. (1993). The reaction cycle of GroEL and GroES in chaperonin-assisted protein folding. Nature , 366 , 228–233. Martin, J., Langer, T., Boteva, R., Schramel, A., Horwich, A.L. and Hartl, F.U. (1991). Chaperonin-mediated protein folding at the surface of GroEL through a molten globule-like intermediate. Nature , 352, 36–42. Miller, A.D., Maghlaoui, K., Albanese, G., Kleinjan, D.A. and Smith, C. (1993). Eschericia coli chaperonins cpn60 (GroEL) and cpn10 (GroES) do not catalyse the refolding of mitochondrial malate dehydrogenase. Biochem. J. , 291, 139–144. Monod, J., Wyman, J., Changeux, J.P. (1965). On the nature of allosteric transitions: a plausible model. J. Mol. Biol , 12, 88–118. Okazaki, A., Ikura, T., Nikaido, K. and Kuwajima, K. (1994). The chaperonin GroEL


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does not recognize apo-α-lactalbumin in the molten globule state. Nature Struct. Biol. , 1, 439–446. Ranson, N.A., Burston, S.G. and Clarke A.R. (1997). Binding, encapsulation and ejection; substrate dynamics in a chaperonin-assisted folding reaction. J. Mol. Biol. , 266, 565–664. Ranson, N.A., Dunster, N.J., Burston, S.G. and Clarke, A.R. (1995). Chaperonins can catalyse the reversal of early aggregation steps when a protein misfolds. J. Mol. Biol. , 250, 581–586. Rye, H.S., Burston, S.G., Fenton, W.A., Beechem, J.M., Xu, Z.H., Sigler, P.B. and Horwich, A.L. (1997). Distinct actions of cis and trans ATP within the double ring of the chaperonin GroEL. Nature , 388, 792–798. Saibil, H.R., Zheng, D., Roseman, A.M., Hunter, A.S., Watson, G.M.F., Chen, S., auf der Mauer, A., O’ Hara, B.P., Wood, S.P., Mann, N.H., Barnett, L.K. and Ellis, R.J. (1993). ATP induces large quaternary rearrangements in a cage-like chaperonin structure. Curr. Biol. , 3, 265–273. Schmidt, M., Rutkat, K., Rachel, R., Pfeifer, GL., Jaenicke, R., Viitanen, P.V., Lorimer, G.H. and Buchner, J. (1994). Symmetric complexes of GroE chaperonins as part of the functional cycle. Science , 265, 656–659. Sfatos, C.D., Gutin, A.M., Abkevich, V.I. and Shakhnovich, E.I. (1996). Simulations of chaperone-assisted protein folding. Biochemistry , 35, 334–339. Smith, K.E. and Fisher, M.T. (1995). Interactions between the GroE chaperonins and rhodanese—multiple intermediates and release and binding. J. Biol. Chem. , 270, 21517–21523. Staniforth, R.A., Burston, S.G. Atkinson, T. and Clarke, A.R. (1994a). Affinity of chaperonin-60 for a protein substrate and its modulation by nucleotides and chaperonin-10. Biochem. J. , 300, 651–658. Staniforth, R.A., Cortés, A., Burston, S.G., Atkinson, T., Holbrook, J.J. and Clarke, A.R. (1994b) The stability and hydrophobicity of cytosolic and mitochondrial malate dehydrogenases and their relation to chaperonin-assisted folding. FEBS Lett. , 344, 129–135 Taguchi, H. and Yoshida, M. (1995). Chaperonin releases the substrate protein in a form with a tendency to aggregate and ability to rebind to chaperonin. FEBS Lett. , 359, 195–198. Todd, M.J. and Lorimer, G.H. (1995). Stability of the asymmetric Escherichia coli chaperonin complex—guanidine chloride causes rapid dissociation. J. Biol. Chem. , 270, 5388–5394. Todd, M.J., Viitanen, P.V. and Lorimer, G.H. (1994) Dynamics of the chaperonin ATPase cycle: implications for facilitated protein folding. Science. , 265, 659–666. Todd, M.J., Viitanen, P.V. and Lorimer, G.H. (1993). Hydrolysis of adenosine-5'triphosphate by Escherichia coli GroEL: effects of GroES and potassium ion. Biochemistry , 32, 8560–8567. Viitanen, P.V., Lubben, T.H., Reed, J., Goloubinoff, P., O’ Keefe, O.P. and Lorimer, G.H. (1990). Chaperonin-facilitated refolding of ribulosebisphosphate carboxylase and ATP hydrolysis by chaperonin-60 (GroEL) are K+ dependent. Biochemistry , 29, 5665–5671.


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Walter, S., Lorimer, G.H. and Schmid, F.X. (1996). A thermodynamic coupling mechanism for GroEL-mediated unfolding. Proc. Natl. Acad. Set. USA , 93, 9425– 9430 Weissman, J.S., Kashi, Y., Fenton, W.A. and Horwich, A.L. (1994) GroEL-mediated protein folding proceeds by multiple rounds of binding and release of non-native forms. Cell. , 78, 693–702. Weissman, J.S., Hohl, C.M., Kovalenko, O., Kashi, Y., Chen, S., Braig, K., Saibil, H.R., Fenton, W.A. and Horwich, A.L. (1995). Mechanism of GroEL action: productive release of polypeptide from a sequestered position under GroES. Cell. , 83, 577–587. Weissman, J.S., Rye, H.S., Fenton, W.A., Beechem, J.M. and Horwich, A.L. (1996). Characterisation of the active intermediate of a GroEL-GroES mediated protein folding reaction. Cell , 84, 481–490. Xu, Z.H., Horwich, A.L. and Sigler, P.B. (1997). The crystal structure of the asymmetric GroEL-GroES-(ADP)7 chaperonin complex. Nature. , 388, 741–750. Yifrach, O. and Horovitz, A. (1994). Two line of allosteric communications in the oligomeric chaperonin GroEL are revealed by the single mutation Arg196<arrow>Ala. J. Mol. Biol. , 243, 397–401. Yifrach, O. and Horovitz, A. (1995). Nested co-operativity in the ATPase activity of the oligomeric chaperonin GroEL. Biochemistry , 34, 5303–5308. Yifrach, O. and Horovitz, A. (1996). Allosteric control by ATP of non-folded protein binding to GroEL. J. Mol. Biol. , 255 , 356–361. Zahn, R., Perrett, S., Stenberg, G. and Fersht, A.R. (1996a). Catalysis of amide protonexchange by the molecular chaperones GroEL and SecB. Science , 271 , 642–645. Zahn, R., Perrett, S. and Fersht, A.R. (1996b). Conformational states bound by the molecular chaperones GroEL and SecB—a hidden unfolding (annealing) activity. J. Mol. Biol. , 261 , 43–61. Zahn, R., Spitzfaden, C, Ottiger, M., Wuthrick, K. and Pluckthun, A. (1994). Destabilization of the complete protein secondary structure on binding to the chaperone GroEL. Nature , 368, 261–265.

23. THE RELATIONSHIP BETWEEN CHAPERONIN STRUCTURE AND FUNCTION STEVEN G.BURSTON1 and HELEN R.SAIBIL2 , * 1 Department

of Genetics Boyer Center for Molecular Medicine, Yale University School of Medicine, 295 Congress Avenue, New Haven, CT 06510, USA 2 Department of Crystallography, Birkbeck College, Malet Street, London WC1E 7HX, UK

1. Introduction 2. Architecture of the Oligomeric GroEL 2.1. Overall Structure of the GroEL Oligomer 2.2. Domain Structure of GroEL 2.3. The GroEL Central Cavity 3. Nucleotide Binding to GroEL 3.1. The Nucleotide Binding Site on GroEL 3.2. Cryo-electron Microscopy of Nucleotide-bound GroEL Structures 4. The Structure of the Co-chaperonin, GroES 4.1. GroES Monomer Structure 4.2. Structural and Chemical Properties of the GroES Heptamer 4.3. Subunit-subunit Interface 5. Protein-protein Interactions 5.1 . The Binding of Polypeptide Substrate to GroEL 5.2. The Interaction Between GroEL and GroES 6. Chaperonins from Archaebacteria and Mammalian Cytosol: TF55 Thermosome and CCT 7. Summary of the Functional Consequences of Chaperonin Structure 8. Acknowledgments 9. References *Corresponding author

The relationship between chaperonin structure and function

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1. INTRODUCTION The intriguing structure of chaperonins, first indicated by electron microscopy (EM) studies in the late 1970’s, suggested that they would have a novel and fascinating mode of action. The interest of structural biologists and biochemists alike was ensured by the subsequent demonstration of their essential biological function of stabilizing unfolded, or partially folded states of a wide range of polypeptides, thus preventing irreversible loss through aggregation, and assisting their folding to a native conformation (Ellis & van der Vies, 1991; Jaenicke, 1993; Clarke, 1996a; Hartl, 1996) (see chapters by Welch et al. and Burkholder and Gottesman). However, the way in which the structural complexity of these macromolecules (approximate molecular mass ~lMDa) is organized to transduce the energy of ATP hydrolysis into the refolding process has presented an enormous challenge to researchers. This chapter will focus on the variety of structural data which has been collected, as well as discussing some of the site-directed mutagenesis which has identified critical regions in the chaperonin architecture. We will then attempt to place this structural data in the context of the mechanistic biochemistry which has elucidated many features of chaperonin function (see chapter by Ranson and Clarke, this volume). 2. ARCHITECTURE OF THE OLIGOMERIC GroEL GroEL (the Escherichia coli chaperonin-60 homologue) has a subunit molecular weight of 58.6kDa and was initially identified as a protein involved in bacteriophage assembly (Georgopoulos et al., 1973). It was later shown to interact transiently with newly synthesized proteins in vivo (Bochkareva et al., 1988; Horwich et al., 1993) as well as improve the refolding efficiencies of a wide variety of unfolded polypeptides in vitro (Goloubinoff et al., 1989; Martin et al., 1991; Buchner et al., 1991; Badcoe et al., 1991; Fisher, 1992). 2.1. Overall Structure of the GroEL Oligomer Negative stain EM revealed the architecture of the GroEL oligomer (Hendrix, 1979; Hohn et al., 1979), characterized by rectangular side views with four stripes of electron density and circular end views with seven-fold symmetry. The correct relationship between these two views was elucidated by Hutchinson et al., (1989) who collected EM images at different tilt angles and showed that GroEL was arranged as two double rings of seven subunits with a two-fold axis of symmetry perpendicular to the seven-fold axis. The presence of two layers of density in each ring suggested that each subunit was divided into two major domains. Three-dimensional reconstruction from negative stain electron micrographs of the GroEL homologue from Rhodobacter spheroides revealed the two major domains to be linked by a small bridge of density on the outside of the cylindrical oligomer, creating a cage-like structure with internal cavities (Saibil et al., 1993).


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However, negative stain EM suffers from the limitation that the sample is dehydrated and thus flattened, and the stain used to contrast the structure does not allow imaging of internal features. These limitations have been overcome by the use of cryo-electron microscopy (cryo EM) in which a thin layer of solution on the grid is rapidly plunged into liquid ethane at liquid nitrogen temperature. The water surrounding the structure becomes vitrified instead of forming ice crystals, thus preserving the native protein without distortion, in a stable, frozen-hydrated state. The grid can then be viewed in the EM provided the grid temperature is maintained below –150°C (Dubochet et al., 1988). The low contrast, and hence signal-to-noise ratio, of these cryo-EM images is improved by averaging large numbers and combining them by tomographic reconstruction to provide three-dimensional (3D) density maps. This technique has been used to provide 3D reconstructions of apo-GroEL, GroEL bound to its nucleotide, unfolded polypeptide and co-chaperonin ligands and also the R197A mutant (Chen et al., 1994; Roseman et al., 1996; White et al., 1997). The initial hurdle in the X-ray determination of any structure is obtaining good crystals. Early trials revealed that the wild-type protein was difficult to crystallize. However, a double mutant of GroEL with Arg-13 changed to Gly and Ala-26 changed to Val produced the crystals suitable for X-ray diffraction (Braig et al., 1994). The crystals were orthorhombic with space group C2221 and had a single heptameric ring in the asymmetric unit. The molecular two-fold axis corresponded to the lattice twofold with the sevenfold perpendicular to this axis. Initial phases were obtained from a single isomorphous derivative to yield a 6 A map. Sevenfold averaging was then used to extend the phasing further to 2.7 Å. Some regions of disordered structure could not be wellresolved using these techniques because of the deviation from seven-fold symmetry in the crystal lattice. The X-ray structure (Braig et al., 1994) revealed the tetradecamer to be a porous cylinder of 146 A in length and 137 A in diameter confirming the earlier EM studies. Refinement of the structure without sevenfold symmetry improved the features, but some regions were still poorly defined because of disorder (Braig et al., 1995). The overall shape of the oligomer is shown in Figure 1(a). 2.2. Domain Structure of GroEL The subunits of GroEL are folded into three distinct domains as shown in Figure 1 (b). The large equatorial domain (residues 6–133 and 409–523) is predominantly -helical and accounts for all of the contacts between the two heptameric rings and most of the inter-subunit contacts within a ring. Also part of the equatorial domain are the mobile Nterminal and C-terminal residues which are crystallographically too disordered to resolve but appear to project into the central channel, probably providing a barrier between the central cavities of the two rings. The small intermediate domain (residues 134–190 and 377–408) consists of antiparallel chains linking the equatorial domain in the midplane of the GroEL cylinder to the apical domain at the end of the cylinder. An exposed region capable of hinge rotation is present at each end of the intermediate domain, at the boundaries with the other two domains (Figure 1b). Rotations about these hinges provide the large conformational change observed by EM during the functional cycle (see section 3 below; Roseman et al., 1996). Another notable feature is the diagonal projection of the intermediate domain


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relative to the equatorial plane which provides a site of contact between the intermediate domain of one subunit with the apical domain of the adjacent subunit on the right when viewed from the side. The contact contains a salt bridge between Arg-197 of the intermediate domains and Glu-386 on the apical domain of the adjacent subunit. This was revealed to be an important site of allosteric communication by characterization of the mutant Arg-197→Ala. This mutant GroEL showed a reduction in both positive and negative cooperativity of ATP binding and hydrolysis (Yifrach & Horovitz, 1994).

Figure 1 Left: Structure of the GroEL 14-mer. This image was created from the atomic structure of Braig et al. (1995) displayed as surface contoured electron density filtered to 25 Å resolution. One subunit is outlined, and the two inter-ring contacts are numbered. Right: Structure of a GroEL subunit in approximately the same orientation as the one outlined in (a). The domain structure, ATP binding site, contacts and hinge regions are indicated. Charged residues in the inter-ring contacts are shown in space-filling form, red for negatively charged and blue for positively charged residues. The space-filling groups shown in yellow are hydrophobic residues whose mutation abolishes substrate and GroES binding, and those in cyan affect GroES binding. The pink residue is Arg 197, which contacts Glu 386 in the intermediate domain of the neighbouring subunit. Mutation of any of the blue residues shown in ball and stick form in the intermediate domain causes global defects in function. Adapted from Figure 1 of Roseman et al., (1996) by permission of Cell Press.

The apical domain (residues 191–376) surrounds the entrance to the central cavity and


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has been shown by electron microscopy (Chen et al., 1994) and site-directed mutagenesis studies (Fenton et al., 1994) to contain the binding site for both unfolded polypeptide chain and the co-chaperonin, GroES. This domain contains both and structures and has a great deal of flexibility, indicated by its high crystallographic temperature factors (B factors), particularly on top and in the regions lining the central cavity (Braig et al., 1995). 2.3. The GroEL Central Cavity Each heptameric ring of GroEL contains a central cavity within the cage-like struc-ture formed by the seven subunits of the ring. The diameter of the entrance to the cavity formed by the seven apical domains is 45 A and widens at the level of the intermediate domains. It appears to be continuous through the 146 A length of the cylinder. However, as mentioned above, the disordered N-terminal and C-terminal residues are likely to project from the equatorial domains into the channel thus creating a block between the rings. The N-terminus has also been shown to be crucial for subunit assembly (Horovitz et al., 1993). It has been estimated that a non-native protein such as a folding intermediate of Mr~20,000 could fit into the central cavity of GroEL (Braig et al., 1994), although it should be noted that binding of GroES and nucleotide results in an expanded but enclosed central cavity as described below (section 5.2). A recent cryo-EM structure of the unliganded GroEL showed asymmetry in which the central cavity of one ring was significantly more open than that of the opposite ring (Figure 3; Roseman et al., 1996). Another interesting feature of GroEL is the presence of portals on the side of the central channel (Figure 1a). These side windows are elliptical and are formed by the gap between the top of the equatorial domain, the crossing of the intermediate domain diagonal to the equatorial plane and the apical domain of the neighboring subunit to the right. They are 36 Å×13 Å at the widest point and 20 Å×10 at the narrowest (Braig et al., 1994). These allow solvent molecules and small ligands, such as nucleotides, access to the interior of the central channel even when GroES is bound to that ring. 3. NUCLEOTIDE BINDING TO GroEL The binding and hydrolysis of ATP by GroEL, and the resultant conformational changes which take place during this hydrolytic cycle are essential in order to drive the assisted folding reaction (Martin et al., 1991 & 1993; Jackson et al., 1993; Todd et al., 1994; Weissman et al., 1994 & 1996). The binding and hydrolysis of ATP by GroEL were observed to be positively co-operative within a heptameric ring (Gray & Fersht, 1991; Bochkareva et al., 1992; Jackson et al., 1993; Todd et al., 1993) and negatively cooperative between the two heptameric rings (Yifrach & Horovitz, 1994; Bochkareva & Girshovich, 1994; Burston et al., 1995). These two forms of cooperativity within the same protein complex have been described in a nested model of co-operativity (Yifrach & Horovitz, 1994 & 1995) in which the binding of ATP within a ring occurs in an “allor-none” fashion described by the Monod-Wyman-Changeux model of co-operativity (Monod et al., 1965), while the negative co-operativity between the rings is best described as a sequential Koshland-Nemethy-Filmer model (Koshland et al., 1966).


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Binding of ATP to a GroEL oligomer results in a conformational change in GroEL from the unliganded conformation which has a high affinity for unfolded protein substrates to a conformation with low substrate affinity (Jackson et al., 1993; Martin et al., 1991; Staniforth et al., 1994a; Sparrer et al., 1996; Yifrach & Horovitz, 1996) pointing to significant allosteric communication within the GroEL oligomer which is essential for its function. ADP appears to bind less cooperatively than ATP within a heptameric ring but with a high degree of negative cooperativity between the rings (Burston et al. 1995). 3.1. The Nucleotide Binding Site on GroEL The X-ray structure of the R13G/A26V GroEL double mutant with ATP S bound to each subunit has been solved to a resolution of 2.4 A (Boisvert et al., 1996). The overall architecture of the GroEL-ATPT S structure is essentially the same as that of unliganded GroEL in that it contains two heptameric rings sharing seven-fold rotational symmetry and a two-fold axis perpendicular to the seven-fold. The ATP-binding site is located at the top of the equatorial domain facing the central cavity. Figure 2 shows the amino acid residues in the equatorial domain which interact with the nucleotide analogue. Residues 87–91 (Asp-Gly-Thr-Thr-Thr) interact with the - and -phosphates of ATP (Boisvert et al., 1996) and lie on a highly conserved loop region (Kim et al., 1994). The specificity for adenosine is imposed via hydrogen bonds from adenine-N6 to the side-chain oxygen of Asn-479 and the backbone amide of Ala 480 to adenine-N1. ATP S also makes two interactions with GroEL mediated via metal ions. A magnesium ion chelates the non-bridging oxygen from each of the three phosphates and completes the octahedral coordination via the carboxylate of Asp-87 and two water molecules. A second metal ion also links the -phosphate to the carbonyl of Thr-30 and Lys-51. This is assumed to be either a K+ ion which decreases the cooperativity of ATP binding (Viitanen et al., 1990; Todd et al., 1993) or Ca2+, a component of the crystallization buffer. The peptide nitrogen of Gly-32 makes a hydrogen bond with both the 5'- -phosphate bridging oxygen and non-bridging oxygen of ATP S. Residues Gly-32, Thr-30 and Lys-51 are near the base of a stem-loop formed by -strands 2 and 3 ( 2/ 3 stem-loop) which interacts with the adjacent subunit by forming an extended b-sheet with the N- and C-terminal regions( -strands 1 and 18) of the neighboring subunit within the ring. Differences between the GroEL-ATP S crystal structure and the unliganded GroEL crystal structure are very small in contrast to the change in substrate polypeptide binding properties and large movements seen using cryo-EM (Chen et al., 1994; Roseman et al., 1996) and detected by fluorescence changes (Jackson et al., 1993; Burston et al., 1995). The reasons for the discrepancy are not known but may be a result of the high protein concentration in the crystal lattice. 3.2. Cryo-electron Microscopy of Nucleotide-bound GroEL Structures Negative stain EM of the ATP-free and ATP-bound states of GroEL (Saibil et al., 1993) revealed a conformational change upon binding ATP in agreement with observations made using an extrinsic fluorescence probe (Jackson et al., 1993). Subsequent cryo-EM studies showed that this structural change was due to a opening out of the apical domains


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about 5–10° relative to the equatorial axis causing a slight elongation of the GroEL cylinder (Chen et al., 1994). More recent cryo-EM image reconstructions (Roseman et al., 1996) are shown in Figure 3. The nucleotide-bound structures show an opening and twisting of the apical domains and elongation compared to the unliganded structure. Interestingly, they also show significant deviation-from two-fold symmetry, providing a structural correlation for the biochemical evidence pointing to asymmetry in nucleotide binding and

Figure 2 The ATP binding site from the X-ray structure of Boisvert et al. (1996). The GroEL subunit is viewed from the opposite side from that shown in Figure 1 (b). The protein backbone is represented as a light blue ribbon with the phosphate binding loop (residues 87–91) in dark blue. The non-hydrolyzable analogue ATP S is shown as a balland-stick model using the following atom colours: blue (nitrogen); red (oxygen); yellow (sulphur); purple (phosphorus). and green (carbon). The magnesium ion which coordinates the phosphate oxygens as well as the carboxylate of Asp-87 is shown in gold. Some important amino acid residues involved in the binding of ATP are shown as yellow ball-and-stick side chains. Asp-87 and Thr-91 are at the extreme ends of the phosphate binding loop. Mutation of Asp-87 blocks ATP hydrolysis (Fenton et al., (1994) while Thr-91 is at the start of a helix which extends downwards to the inter-ring interface and is likely to be involved in allosteric communication between the rings. Asn-479 is involved in determining the specificity of adenosine as the preferred base while Ile-493 provides a hydrophobic locality for the purine. Figure produced using InsightII (BioSym Technology).


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hydrolysis (Yifrach & Horovitz, 1994; Bochkareva & Girshovich, 1994; Burston et al., 1995). The ATP-bound structure, from samples vitrified within 4 seconds of mixing, is indistinguishable from the steady-state ATP structure. (The rate-limiting step is the hydrolytic step (Jackson et al., 1993)). These results refine the previous negative stain (Saibil et al., 1993) and cryo-EM observations on GroEL-ATP (Chen et al., 1994) by providing more accurate and higher resolution three-dimensional information. The apical domains of the top ring are opened up in the presence of ATP, but the subunits of the lower ring are rotated inwards, showing the asymmetry between the rings resulting from negative cooperativity (Figure 3). The ADP-bound structure

Figure 3 3-D cryo-EM reconstructions of GroEL complexes. The top row shows the GroEL, GroEL-ADP and GroEL-ATP complexes. Both of the nucleotide bound structures are more extended than GroEL with the upper apical domains (see Figure 1) twisted outwards. The


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bottom row shows the “bullet”-shaped GroEL-GroES-ADP, GroEl-GroESATP and the “American football”-shaped GroEL-GroES-AMP-PNP which can be observed under certain conditions. Compared with the top row it can be seen that GroEL adopts a completely different structure when bound to GroES, with the twist of the subunits reversing handedness. Additionally a large cavity is opened up underneath GroES in which protein substrates can be encapsulated. The contour levels for each structure was chosen to enclose the correct molecular volume. Each reconstruction was produced from an EM data set of approximately 1000 images. Figure adapted from Roseman et al. (1996) by permission of Cell Press.

shows the largest opening of the rings and is generally a more expanded structure. Superposition of unliganded GroEL, GroEL-ADP and GroEL-ATP shows a successive clockwise twisting of the apical domains which would cause the substrate binding site to be rotated out of the central cavity and become buried in the intersubunit interface (Figure 4a). These observations explain how nucleotides control the binding affinity of GroEL for non-native substrates, by controlling the accessibility of the hydrophobic binding sites (Roseman et al., 1996). A recent study of the GroEL mutant R197A has resolved the TR (ATP bound to one ring) and RR (ATP bound to both rings) states (White et al., 1997). The asymmetric GroEL-ATP complex in Figure 3 is in the TR state, and the RR state (not shown) has both rings in a very open conformation. Comparison of the EM reconstructions of GroEL-ATP with GroEL or GroEL-ADP showed a loss of density in the region of the inter-ring contacts (Figure 5; Roseman et al., 1996). This difference was found to be statistically signficant by comparing the data sets with a student’s t test. By superimposing the EM reconstruction over the X-ray structures, the altered region was identified as the inter-ring contact containing Lys-105 and Glu434. Since Lys-105 is directly connected via an -helix to the nucleotide phosphate binding residues 87–91, Roseman et al. (1996) propose that during ATP binding and hydrolysis small displacements of this helix may withdraw the lysine residue, thus weakening the interaction energy across the contact region. The other inter-ring contact, between Glu-461 and Arg-452, shows significant variation between GroEL-GroES-ADP and GroEL-GroES-ATP complexes. Moreover, mutation of Glu-461 to lysine blocks polypeptide release and decreases the stability of the GroEL-GroES interaction (Fenton et al., 1994). This contact may mediate inter-ring effects of nucleotide binding upon the GroES and polypeptide substrate binding sites. 4. THE STRUCTURE OF THE CO-CHAPERONIN, GroES GroES, a member of the Hsp10, or chaperonin-10 class of chaperones, is a heptamer of identical ~10 kDa subunits which acts as a co-protein in the functional cycle of GroEL (Chandrasekhar et al., 1986). The expression of GroES is essential for cell viability and has been shown in vitro to be critical for the folding of certain substrate polypeptides under conditions where little or no spontaneous folding is observed (Goloubinoff et al., 1989; Martin et al., 1991; Schmidt et al., 1994a). It has been observed in EM studies to


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bind tightly to one end of the GroEL cylinder in the presence of adenine nucleotides (Saibil et al., 1991; Langer et al., 1992; Chen et al., 1994). A second GroES binds weakly to the opposite ring of GroEL (Schmidt et al., 1994b; Llorca et al., 1994) in the presence of ATP ot AMP-PNP (see section 5.2). The X-ray crystallographic structure of GroES has been determined to a resolution of 2.8 Å (Hunt et al., 1996; Figure 6a). The protein crystallized in the orthorhombic P212121 space group with one heptamer in the asymmetric unit. Two heavy atom derivatives, one of which clusters to the acidic roof of the GroES dome (see below), were used to create an isomorphous replacement map which in turn

Figure 4 (a) Outlines of the top view of apical domains in GroEL, GroEL-ADP and GroEL-ATP. The grey bar shows the position of the hydrophobia binding site facing the central cavity, and the dark grey and black bars show how ADP and ATP binding respectively twist the domain so that the site becomes progressively occluded. (b) 3D reconstruction of GroEL-GroES-ATP with outlines of one subunit in GroEL, GroEL-ATP and in the GroES complex. The hinge rotation causing the apical domain twist is around 100° (compare to Figure 8a). Figures adapted from Roseman et al. (1996) by permission of Cell Press.


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allowed the structure to be solved making use of seven-fold non-crystallographic symmetry averaging. It was then refined without seven-fold averaging. The structure of a second chaperonin-10 from Mycobacterium leprae has also been solved to 3.5 Å with seven-fold averaging (Mande et al., 1996) and shows essentially the same topology and physical characteristics as the E. coli version described by Hunt et al. (1996). 4.1. GroES Monomer Structure Each subunit is composed of an irregular anti-parallel -barrel which encompasses a hydrophobic core with a -hairpin which extends outwards at the top towards the

Figure 5 (a) GroEL-ADP and (b) GroEL-ATP, aligned with the atomic structure of the interring contact regions of GroEL. The EM reconstructions are shown as wire frame surfaces (white) to reveal the atomic models inside. One inter-ring contact is at the front centre of each structure and is seen as solid density in the ADP reconstruction and as a hole in the ATP reconstruction. The positively and negatively charged residues in the contacts are shown in blue and red, respectively. The helix (green) connecting the central contact (Lys105) to the ATP binding site (Thr-91) extends diagonally outward from the contact, with an ATP molecule (purple) bound in each site. (Reproduced from Roseman et al., (1996) by permission of Cell Press.)

center of the ring. Electron density corresponding to a region termed the “mobile loop” region (residues 16–33), which had been shown previously by NMR studies to interact with GroEL in the presence of ADP (Landry et al., 1993), was only visible in one of the seven subunits. The mobile loop density of this one ring formed a -hairpin sandwiched between the wall of one GroES heptamer and the roof of another in the crystal lattice. Lys-34 (Lys-36 in M. leprae numbering) which has been shown to modulate allosteric transitions in GroEL (Kovalenko et al., 1994) lies adjacent to this mobile loop region and


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is likely to interact directly with GroEL (Mande et al., 1996). 4.2. Structural and Chemical Properties of the GroES Heptamer The GroES heptamer resembles a dome-shaped cap which is 30Å high and 70–80 Å in diameter. The inside of the dome is approximately 20 Å high and 30 Å in diameter (Hunt et al., 1996). Subunits are arranged such that the -barrels of each monomer are parallel to each other and to the seven-fold axis. The inter-subunit contacts are between the first -strand in each subunit and the -barrel of the adjacent subunit. The -hairpin structures of each monomer which extend towards


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Figure 6 (a) A side view of GroES with the backbone displayed in ribbon form. The mobile loop is shown for the subunit in which electron density for this region could be seen. Electron density could not be


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seen for the mobile loop region (residues 16–32) in the other subunits; therefore the discontinuity at residue 15 and residue 33 is indicated with an (L) on the foremost subunit. The N- and C-terminus are also marked on two subunits to distinguish them from the ends of mobile loop. (b) A top view of GroES in ribbon format. The one mobile loop region is again marked as the N- and C-terminus of one subunit. The -hairpin of each subunit which extends toward the top centre can be clearly seen. Both figures were produced using the program RIBBONS (Carson, 1987).

the top and center of the dome contain two glutamate residues (Glu-50 and Glu-53) at the tip of the hairpin, resulting in a dense region of negative charge (Figure 7a). A significant amount of disorder was observed in this region, presumably due to charge repulsion of these clustered acidic side chains. It is not clear how these interactions are stabilized at physiological pH and ionic strength in the bacterial cell. It has been postulated that this flexibility at the roof of the dome may allow non-native proteins to leave the central cavity of GroEL without dissociation of GroES (Hunt et al., 1996). However, biochemical evidence to support such a hypothesis has not been reported. The opening facing the GroEL cavity on the underside of the dome contains the highly conserved residue Tyr-71, forming a region of hydrophobicity near the point at which GroES interacts with GroEL (Figure 7b). These seven tyrosine residues could be available for interaction with the unfolded polypeptide substrate when sequestered in the cis cavity (i.e. on the same ring of GroEL as GroES). The crystal structure indicates a high degree of conformational flexibility in these residues, consistent with a possible role in solvating a dynamic polypeptide folding within the central cavity of GroEL. 4.3. Subunit-subunit Interface Substantial deviation of the GroES heptamer from seven-fold symmetry was observed in the crystal structure (Hunt et al., 1996). Although the hydrophobic core of the -barrel deviated little from seven-fold symmetry, the packing of adjacent subunits was highly irregular due to structural plasticity at the interface between subunits. The principal interaction across the interface is between the first -strand of one subunit with the final -strand of the adjacent subunit. If seven C-terminal residues are cleaved from GroES by limited proteolysis in urea, the cleaved monomers are unable to oligomerize (Scale & Horowitz, 1995). The long, charged side chains of Lys-74, Glu-76 and Arg-37 also serve to maintain a flexible subunit-subunit interface by weakening the interaction between subunits. It has been observed that GroES heptamers dissociate into monomers at ~10-8 M oligomers (Zondlo et al., 1995) corresponding to a binding energy of about –7 kcal mol-1 of interface. This flexibility may be present to allow conformational changes in GroES induced by


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Figure 7 A surface map of the GroES oligmer produced by GRASP (Nicholls et al., 1991). The positive and negative surface potential is represented by blue and red, respectively. The 8 A diameter orifice at the top of the oligomer can be seen surrounded by a high degree of negative charge provided by the two glutamate residues on the hairpin of each subunit.

movements of the GroEL domains to which it is bound, and which may be critical to the folding of a protein substrate in the cis ring of the GroEL-GroES complex. 5. PROTEIN-PROTEIN INTERACTIONS 5.1. The Binding of Polypeptide Substrate to GroEL Central to a structural description of the chaperonin-assisted folding mechanism is the identification of the site and physical nature of the interaction between GroEL and unfolded polypeptide substrate. Since an X-ray crystallographic approach is hampered by the fact that unfolded polypeptides do not have seven-fold symmetry and may be disordered in a crystal lattice, efforts have focused upon electron microscopy and sitedirected mutagenesis. The first attempt to locate the site of the bound polypeptide was made using negative


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stain electron microscopy to study GroEL bound to either unfolded rhodanese or unfolded alcohol dehydrogenase (Langer et al., 1992). End views of an average of ~650 images revealed some extra, poorly defined electron density in the region of the opening of the central cavity which was assigned to bound, unfolded polypeptide. However, because of the difficulty in identifying a bound polypeptide chain using this method, Braig et al. (1993) took the approach of covalently labelling dihydrofolate reductase (DHFR) with colloidal gold particles in order to localize the protein substrate in EM images. When the unfolded, gold-labelled DHFR was bound to GroEL, gold particles were observed in the centres of end views, corroborating the notion that GroEL binds substrates in its central cavity. Side views showed that some GroEL molecules could bind a gold-labelled DHFR inside the central cavity of both rings simultaneously, indicating that unliganded GroEL contains a potential binding site for polypeptide on each of its two rings. Binding of IPMDH was observed in the trans ring (opposite GroES) by labelling with an antibody to the substrate (Ishii et al., 1994). A more direct view of the bound substrate density was obtained using cryo-EM of mitochondrial malate dehydrogenase (mMDH) bound to GroEL (Chen et al., 1994). This study showed a region of additional electron density which could be assigned to the MDH located in one end of the GroEL cylinder between the apical domains and bulging outwards into the solvent. The location and physical characteristics of the interaction between polypeptide chain and GroEL has been examined using site-directed mutagenesis (Fenton et al., 1994). A large number of site-directed mutants of GroEL were screened for their effects upon the binding and refolding of unfolded ornithine transcarbamylase (OTC), a substrate that requires GroES to fold efficiently (Zheng et al., 1993). The mutants were also screened for GroES binding, ATPase activity and ability to rescue E. coli cells where expression of the chromosomal GroEL had been totally repressed. It was noted that a number of mutants abolished the ability to bind unfolded OTC. These mutants all mapped to hydrophobic residues in the apical domain lining the central cavity (e.g. Y199E, Y203E, F204E, L234E, L237E—see Figure 1b). Subsequently the interaction between GroEL and either a mutant of subtilisin, BPN′, which is unable to fold, or -casein, a disordered protein, has been analyzed by scanning microcalorimetry (Lin et al., 1995). Both proteins were able to bind to GroEL with micromolar affinity and showed a negative heat capacity upon binding which are characteristic of a hydrophobic interaction. The conformation of GroEL-polypeptide binary complexes has been probed using amide hydrogen/deuterium exchange, NMR and electrospray mass spectroscopy. Studies using the scrambled three-disulphide -lactalbumin (Robinson et al., 1994) and DHFR (Gross et al., 1996; Goldberg et al., 1997) suggested that the bound polypeptide was highly labile but showed a small degree of protection from amide hydrogen exchange in regions of the polypeptide with a high degree of protection in the folded structure. Although these studies indicated the possible existence of native-like secondary structure when bound to GroEL it was impossible draw a definitive conclusion. More recently the crystal structure of an apical domain (residues 191–376) with a 17 residue N-terminal peptide tag has been solved (Buckle et al., 1997), in which the N-terminal tag appears to be mimicking an unfolded polypeptide substrate by binding to the peptide binding site of a neighbouring apical domain in the crystal lattice. The binding surface maps to the same


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region as those identified by site-directed mutagenesis (Fenton et al., 1994), specifically Leu-234, Leu-237 and V-264, and would appear to extend back to residues Tyr-199, Tyr203 and Phe-204. The interactions with the apical domain are predominantly hydrophobic in nature although there is some hydrogen bonding between the main chain of the tag and side chains of the apical domain. Whilst the tag is essentially in an extended conformation there appears to be significant flexibility in the polypeptide binding site in order to accommodate any general hydrophobic surface, such as that of molten globuletype non-native proteins. 5.2. The Interaction Between GroEL and GroES GroES is able to form a stable interaction with GroEL only in the presence of adenine nucleotides (Chandrasekhar et al., 1986). When the complex was viewed in the electron microscope GroES appeared to cap one end of the GroEL cylinder (Saibil et al., 1991; Langer et al., 1992; Ishii et al., 1992). It was also noted that the presence of GroES inhibited the steady-state ATP hydrolytic rate of GroEL by approximately 40–50% in the presence of K+ ions (Gray & Fersht, 1991; Jackson et al., 1993; Todd et al., 1993). This was originally thought to result from half-sites reactivity due to inhibition of the ATPase in the ring bound to GroES. However, the finding that ATP binding and hydrolysis are asymmetric in GroEL alone (Yifrach & Horovitz, 1994; Bochkareva & Girshovich, 1994) led to the conclusion that the presence of GroES is in fact altering the rate-limiting step in the ATP hydrolytic cycle of GroEL (Burston et al., 1995). The 1:1 stoichiometry of GroES to GroEL in the presence of ADP has been confirmed by protease protection (Langer et al., 1992) and fluorescence studies (Jackson et al., 1993). However, it was noted by negative stain EM that in the presence of ATP and analogues a significant number of GroEL cylinders were found with a GroES oligomer bound to each end in the shape of an American football (Schmidt et al., 1994). The functional significance of these “football” structures remains a mystery (Lorimer, 1997), although their presence as a transient species in the GroEL functional cycle has been detected (Corrales & Fersht, 1996). The site of interaction between GroES and GroEL has been visualised by cryo-EM. The GroEL ring bound to GroES shows an approximately 60° opening between the apical and equatorial domains such that GroES is able to interact with the ends of the apical domains that were previously oriented towards the central cavity (Chen et al., 1994). The most striking feature of this complex is that the central cavity becomes enlarged approximately three-fold compared to that in GroEL alone, and capped, such that a protein of molecular weight < 60,000 could be encapsulated (Clarke, 1996b). Secondly, the site of interaction between GroEL and GroES was in the same region of GroEL to which unfolded MDH was bound (Chen et al., 1994). Close coupling of sites for substrate and GroES was demonstrated by the construction of site-directed mutants in the apical domain (e.g. Y203E, L237E, V263S) which affected the binding of both GroES and unfolded OTC (Fenton et al., 1994; Figure 1b). The region of GroES which interacts with GroEL has also been identified by NMR. Landry et al. (1993) found that a one-dimensional 1H-NMR spectrum of GroES contained sharp peaks on a background of broad resonances. The broad resonances were


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expected since GroES is an oligomer of 70 kDa and has too slow a rotational correlation time in free solution to give rise to sharp signals. The sharp peaks indicated therefore that GroES contained some regions of polypeptide which are more mobile than the rest of the structure. They assigned the sharp peaks to residues 17–32 (VETKSAGGIVLTGSAA) of GroES, which they termed the “mobile loop”, and showed that it was susceptible to proteolysis. When GroES was associated with GroEL in the presence of ATP the linewidths of the mobile loop were broadened and the loop was no longer protease sensitive. Given that a number of mutants of GroES which were unable to support growth of bacteriophage 1 also mapped to this region (Georgopoulos et al., 1973), this provided strong evidence that the mobile loop region interacts directly with GroEL. A peptide corresponding to this region was synthesized and shown to bind to GroEL. Since the exchange rate of this peptide between free and GroEL-bound forms was sufficiently rapid then transfer Nuclear Overhauser Effects (trNOEs) could be used to monitor binding and conformation of the bound state. The conformation of the peptide was determined to be a hairpin loop and could be competed off with intact GroES. The overall conformational changes of GroEL-GroES complexes during the hydrolytic cycle have recently been examined (Figure 3; Roseman et al., 1996). Thin bridges of electron density were observed connecting GroES and GroEL, suggesting that the seven mobile loops of the GroES heptamer may be responsible for most of the interaction with the GroEL apical domains. The GroES-GroEL complexes show a dramatic prolongation of the apical domain twisting observed in the GroEL-ATP cryo-EM structure. Indeed the change in twist is so large that the subunit is reversed in handedness, suggesting that the exposed surface in the central cavity which interacts with bound polypeptide substrate may be very different (Figure 4b). In the GroEL-GroES complex formed with ADP, the during apical domains have a slightly different tilt compared to that formed in the presence of ATP. This shifts the site of interaction with GroES slightly and distorts the GroES subunit interfaces. The apical domains of the open trans-ring in the presence of ADP show a twisting outwards when compared to those in complexes formed with ATP (Figure 3). Changes in the exposure of amino acids in the interior of the cavity upon GroES binding and nucleotide binding and hydrolysis may well be critical structural changes during the functional cycle of GroE. The crystal structure of the asymmetric GroEL7•GroEL7•ADP7•GroES7 has recently been determined (Xu et al., 1997; Figure 8a). The conformation of the cis ring forming the enclosed cavity reveals details of the domain movements generally predicted from the cryo-EM (Roseman et al., 1996). The interface between GroES and GroEL occurs via residues identified by previous mutational analysis (Fenton et al., 1994) and NMR (Landry et al., 1993). In particular, Leu-234, Leu-237, Leu-259, Val-263 and Val-264 of GroEL, which lie on two helices which have shifted to the top of the subunit in comparison to the unliganded structure, and interact with residues Ile-25, Val-26 and Leu-27 of GroES. The twisting and raising of the GroEL apical domain to form the GroEL•GroES interaction site dramatically changes the interaction surface available to the polypeptide. In addition to residues 234, 237 and 264 being recruited to interact with the co-chaperonin, six other residues (Tyr-


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Figure 8 (a) Side view of GroEL-GroES-ADP (Xu et al., 1997). Domains are coloured red (equatorial), green (intermediate) and yellow (apical). The GroES co-chaperonin (green) is bound to the top ring of GroEL. An equatorial (blue), intermediate (light blue) and apical domain (violet) have been highlighted for comparison with the side view of the thermosome (Figure 9a). Reproduced from Ditzel et al. (1998) by permission of Cell Press. (b) Schematic drawing of the chaperonin ATP-mechanism based upon the thermosome structure (Ditzel et al., 1998). The thermosome numbering is used and the equivalent GroEL residue number is shown in parentheses in the following. The -phosphate of ATP is stabilized via hydrogen bonds to the conserved Thr-96 (Thr-89) and 97 (Thr-90) and coordination to the magnesium ion. The carboxylate side chains of Asp-63 (Asp-52) and Asp-390 (Asp-398) polarize the water molecule which makes a nucleophilic attack on the phosphorus atom. The resultant pentavalent intermediate subsequently collapses to liberate the orthophosphate and ADP products. Figure reproduced from Ditzel et al., (1998) by permission of Cell Press. (c) The modified nucleotide binding pocket in the GroEL-GroES-(ADP)7 crystal structure. The protein backbone is represented as a light blue ribbon with the phosphate-binding loop (residues 87–91) in dark blue. The bound ADP is shown as a ball-and-stick model using the following atom colours: blue (nitrogen); red (oxygen); purple (phosphorus) and green (carbon). The magnesium ion which coordinates the nucleotide phosphate oxygens as well as the carboxylate side chains of Asp-87 and Asp398 is shown in gold. In


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comparison with Figure 2 (from the GroEL-ATP S structure) the insertion of Asp398 from the intermediate domain into the nucleotide binding pocket can be seen. Mutation of this residue to alanine resulted in a drastic reduction of ATP hydrolytic activity (Rye et al., 1997).

199, Ser-201, Tyr-203, Phe-204, Leu-259, and Val-263) become buried either between apical domains or along the interface with GroEL. This removal of the polypeptide binding site upon ATP and GroES binding provides a means of releasing the polypeptide into the GroEL central cavity. The residues which now line the wall of the central cavity are polar in nature, thus providing an environment which strongly favours intra-molecular interactions within the polypeptide rather than a hydrophobic interaction between the non-native polypeptide and the chaperonin. A further important aspect of the conformational change upon binding GroES and nucleotide, not detected in the low resolution cryo-EM, was observed. The intermediate domain rotates downwards to interact with the equatorial domain, thus effectively locking the nucleotide into its binding pocket. As a consequence of this movement Asp-398 moves down into the nucleotide binding site such that the carboxylate of the aspartate side chain coordinates with the Mg2+ ion (Figure 8c). Replacement of this aspartate residue with alanine results in the 60-fold reduction in ATP hydrolytic activity (Rye et al., 1997) demonstrating the importance of this domain movement for the hydrolysis of ATP. Xu et al. (1997) also propose that the allosteric communication across the ring-ring interface is due to an en bloc movement of the trans equatorial domains in a complimentary fashion to tilting movements of the cis equatorial domains, thus preserving the integrity of interactions across the ring-ring interface and avoiding steric clashes. 6. CHAPERONINS FROM ARCHAEBACTERIA AND MAMMALIAN CYTOSOL: TF55 THERMOSOME AND CCT A second subfamily of chaperonins, weakly related to GroEL, has been discovered in archaebacteria (e.g. Sulfolobus shibitae, Sulfolobus solfataricus), namely TF55 (Trent et al., 1991) and its homologue the thermosome (Phipps et al., 1991 & 1993), and in the eukaryotic cytosol, CCT (for reviews see Willison & Kubota, 1994; Willison & Horwich, 1996; chapter Willison in this volume). By negative stain EM it could be seen that TF55 had a similar overall architecture to that of GroEL with two stacked rings (Trent et al., 1991; Marco et al., 1994a; Knapp et al., 1994). However, in contrast to GroEL, TF55 contains two nine-membered rings, while the related thermosome from Pyrodictium, Thermoplasma and Archaeoglobus all have eight subunits per ring (Phipps et al., 1991). Additionally, both TF55 and the thermosome have two distinct, although closely related subunits (Knapp et al., 1994; Kagawa et al., 1995). They bind unfolded polypeptides in vitro and have ATPase activity similar to GroEL. However, no GroES homologue has been found to function with this class of chaperonins. Although TF55 and the thermosome are only very weakly related to GroEL they do show ~40% homology with subunits of CCT and it has been hypothesized that the thermosome is the evolutionary ancestor of CCT (Willison & Horwich, 1996).


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The function of the eukaryotic cytosolic chaperonin CCT, appears to be more specific than that of either GroEL or TF55. It has been found to mediate the folding of - and tubulin, actin, and Ga-transducin in vitro and in vivo (Yaffe et al., 1992; Lewis et al., 1992; Gao et al., 1992; Sternlicht et al., 1993; Farr et al., 1997) and can assist the folding of firefly luciferase in vitro (Frydman et al., 1992). At the level of primary structure CCT is significantly more complex than either GroEL or TF55 having up to 9 different polypeptide subunits. Each complex consists of two stacked hetero-oligomeric rings and top views show 8-fold symmetry by negative stain EM (Lewis et al., 1992; Marco et al., 1994b). Electron microscopy of CCT suggests that it has the same overall subunit architecture as GroEL. Side views show four stripes corresponding to the apical domains and equatorial domains of each ring (Lewis et al., 1992; Gao et al., 1992; Frydman et al., 1992). The central cavity of CCT appears larger than that of GroEL, measuring 60 Å in diameter as opposed to 45 Å. Electron micrographs of CCT complexed with bound actin revealed the substrate localized to the central cavity (Marco et al., 1994b). The primary amino acid sequences of GroEL and CCT have been compared and show a great deal of similarity at the N- and C-termini (Kim et al., 1994), especially among the conserved residues of the ATP binding pocket (Boisvert et al., 1996). These regions make up the equatorial domain of GroEL (Braig et al., 1994) suggesting that the equatorial domains of CCT may be very similar to those of GroEL. The location of the ATP binding site on CCT subunits has been supported by the results of mutagenesis in this region which lead to tubulin deficiency and temperature sensitivity (Miklos et al., 1994; Ursic et al., 1994). However, comparison of the sequences corresponding to the apical domains shows little or no homology indicating that the divergent properties of CCT and GroEL result largely from the differences in these domains. Like TF55, CCT performs its function without requirement of a GroES homologue although a number of co-factors have been identified (Gao et al., 1993; Tian et al., 1996). Like GroELs, CCT requires ATP hydrolysis to refold its substrates and substrates undergo cycles of binding and release as they do with GroEL (Tian et al., 1995; Farr et al., 1997). 6.1 High-resolution Structural Studies of the Thermosome Recently, X-ray structures of an isolated apical domain of the -subunit of the thermosome (Klumpp et al., 1997) and the complete thermosome (Ditzel et al., 1998) from the archaeon Thermoplasma acidophilum have been determined. The overall thermosome structure consists of two stacked octameric rings which are composed of alternating and subunits in a ( )4 arrangement and encloses a central cavity similar to GroEL (Figures 9a and b). However, the overall shape of the oligomer is more spherical than the GroEL cylinder, with a height of 158 A along the pseudo 8-fold axis and a diameter of 164 Å along the equatorial 2-fold axes. The sequence identity between the and subunits is 60% and they share a 46% sequence similarity with the eubacterial chaperonin GroEL. Each subunit adopts the same basic fold as a GroEL subunit with an equatorial, intermediate and apical domain. The main difference between the evolutionary diverse structures lies in the apical domain, as might be expected given that the group II chaperonins function with no GroES homologue. Compared to the GroEL subunit structure the thermosome apical domains

Molecular chaperones and folding catalysts show a 35 residue deletion which comprises helices

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Figure 9 Thermosome structure, (a) Side view of the thermosome. The domains are coloured in red (equatorial), green (intermediate) and yellow (apical). The 16 molecules of bound ADP are also shown bound to the equatorial domains in yellow. The stacking of subunits in register forming and pairs across the inter-ring interface can be seen. (b) Top view of the thermosome. -subunits are coloured red or violet and the -subunits are coloured yellow. The two strands of each subunit which lie in the centre and the N-terminal half of the helix which projects away from the centre back into the main core of the apical domain comprise the lid segment. Movements of the lid segment upon twisting of the apical domains during the thermosome functional cycle may open and close the entrance to the central cavity. (c) Comparison of the configuration of the lid segments from the structures of the thermosome (red; Ditzel et al., 1998) and the isolated apical domain (green; Klumpp et al., 1997). It has been speculated that in the polypeptide acceptor state could be generated by opening of the entrance to the central cavity, and that the lid segment may adopt the same fold as that found in the isolated apical domain. This would result in a clustering of hydrophobic residues (whose side chains are shown) which may form a polypeptide binding site.


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(a) and (b) reproduced from Ditzel et al. (1998) by permission of Cell Press, (c) reproduced from Horwich & Saibil (1998) by permission of McMillan.

K and L in GroEL and an insertion of 28 residues which introduces two -strands and extends the N-terminus of helix H. This insertion in the thermosome subunits protrudes towards the pseudo 8-fold axis at the ends of the complex providing an obstruction to the entrance to the central cavity termed the lid domain (Ditzel et al., 1998). The structure of each subunit within the complex resembles that of a subunit in the cis-ring of the GroELGroES-ADP complex (Xu et al., 1997) in that a segment of the intermediate domain has closed down to form the upper part of the nucleotide binding site. The intra-ring contacts of the equatorial domains are conserved between the group I and group II chaperonins. However, the inter-ring contacts are different. The thermosome has its two rings in register producing pairs and pairs (Figure 9a), in contrast to the staggered arrangement of rings in GroEL (see Figure 8a). Of considerable interest is the structure of the lid domain which arches over to block the entrance to the central cavity of each ring. The X-ray structure of the isolated apical domain (residues 214–365) region (Klumpp et al., 1997) shows the same core structure as that in the intact thermosome (Figure 9c). However, the lid segment in the isolated domain has a helix-turn-helix structure whereas in the intact thermosome the same region is strand-strand-turn-helix (Figure 9c). The lid protrusions in the intact thermosome have their 8 hydrophobic side chains buried in side-to-side packing contacts. However, the helix-turn-helix structure of the isolated domain clusters these hydrophobic residues together. It has been speculated (Ditzel et al., 1998) that in a polypeptide-acceptor conformation of the thermosome a counter-clockwise rotation of the apical domains relative to their position in the crystal structure (believed to be the ‘folding-active’ state) could open the lid and give access to the binding sites, perhaps adopting the same conformation as that seen in the isolated domain structure. This could perhaps generate a polypeptide binding site via the clustered hydrophobic residues. Ditzel et al. (1998) have also performed experiments in which they co-crystallized or soaked in various nucleotides. The nucleotide-binding site was found to be located on top of the equatorial domain in the same position as that found in GroEL (compare Figures 8a and 9a and also Figures 8b and 8c). The Mg-ADP-bound crystal structure showed domain motions about the hinge regions between the three domains compared to the unliganded structure. The presence of the nucleotide phosphates displaces Asp-94 from its position in the triphosphate-binding site which it occupies in the unliganded structure. Residues 89–98 are shifted towards the neighbouring intermediate domain forcing the residues 85–88 to undergo a transition from relaxed loop to a helix. However, movements in the 2/ 3 stem-loop noted in the GroEL-ATP S structure (Boisvert et al., 1996) could not be seen in the thermosome-ADP structure (Ditzel et al., 1998). In contrast to GroEL-GroES-ADP where the nucleotide is not exchangeable with free nucleotide (Todd et al., 1994; Xu et al., 1997), the nucleotide-binding site in the thermosome crystal structure is still accessible to solvent, even when the intermediate domain is partly clamped down onto the equatorial domain. Soaking experiments using the ATP transition state analogue Mg-ADP-AIF3 were also performed providing some structural evidence


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on the mechanism of ATP hydrolysis (Ditzel et al., 1998). One fluorine was coordinated to the Mg2+ ion while the remaining two fluorine atoms form hydrogen bonds to threonines 96 and 97. A water molecule forms hydrogen bonds with Asp-390 (equivalent to Asp-398 in GroEL) and Asp-63 (Asp-52). The carboxylate side chains of the aspartate polarize the water molecule such that a nucleophilic attack is made on the phosphorus atom. The resulting pentavalent intermediate can subsequently collapse yielding a free orthophosphate and ADP. The role of Asp-390 is strongly supported by evidence that mutation of Asp-398 to Ala in GroEL resulted in a 60-fold reduction in the rate of ATP hydrolysis (Rye et al., 1997). 7. SUMMARY OF THE FUNCTIONAL CONSEQUENCES OF CHAPERONIN STRUCTURE The challenge remains today to explain how chaperonins are able to harness the energy of ATP hydrolysis in order to improve the efficiency of protein folding in vivo. However, the structural work described above, together with the extensive biochemical and biophysical studies described in the chapter by Ranson and Clarke, provide insights into the way in which chaperonins achieve their cellular task. The earliest stages of protein folding involve the collapse of a polypeptide chain around a hydrophobic core and are usually coincident with the formation of some elements of secondary structure (for review see Dobson et al., 1995; Dill et al., 1995). For many small proteins this occurs rapidly and efficiently, before the rate-limiting step late in the folding pathway. However, it has been observed for a number of proteins, especially larger oligomeric proteins, that a significant degree of misfolding occurs during these early stages (Sosnick et al., 1994; Jaenicke, 1995; Ranson et al., 1995). This ensemble of misfolded conformers contains a high degree of exposed hydrophobic surface and they are thus prone to irreversible aggregation. While some of these misfolded polypeptides can slowly isomerize, by reversing incorrect interactions, to a conformation that is able to rapidly fold to the native state, this process has to compete directly against collision with other misfolded states leading to aggregation. Thus the rate-limiting step of refolding for many large proteins may be the partial unfolding of kinetically trapped conformations, or even dissociation of small, low-order aggregates, to a fast-folding species. The chaperonin, therefore, has the task of preventing these occurrences and/or actively reversing them in order to improve the efficiency of refolding. GroEL has a number of unique physicochemical features in order to perform this function. The initial step is the “recognition” of polypeptide substrates. Identification of the binding site by electron microscopy and mutagenesis (Fenton et al., 1994), coupled with the high resolution X-ray structure has located the site of interaction to the apical domains (Chen et al., 1994; Braig et al., 1994; Buckle et al., 1997). Site-directed mutagenesis of hydrophobic residues in this region strongly affects both polypeptide and GroES binding (Fenton et al., 1994), suggesting that the binding is due primarily to hydrophobic interaction between the apical domains and substrate polypeptide. This has determined directly using microcalorimetry (Lin et al., 1995) and binding studies (Hutchinson et al., 1997). Since early folding intermediates and misfolded protein chains


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expose hydrophobic surface which would otherwise be buried, this provides a means by which chaperonins can “recognize” a wide range of substrate polypeptides, without requiring any specific structural feature which would vary from substrate to substrate. It has also been proposed that the binding energy may be used to reverse incorrect interactions which have formed early in the folding/misfolding pathway (Jackson et al., 1993; Todd et al., 1994; Weissman et al., 1994; Ranson et al., 1995; Todd et al.., 1996; Zahn et al., 1996). The binding of ATP to the GroEL-polypeptide complex causes rotations of the two hinge regions between GroEL domains, raising and twisting the apical domains, and thus with-drawing part of the hydrophobic peptide binding site (Roseman et al., 1996; Xu et al., 1997). These movements permit the rapid association of GroES (Burston et al., 1995) to the remaining hydrophobic surface. Binding of ATP and GroES results in the rapid displacement of the polypeptide into the central cavity (Rye et al., 1997) as the hydrophobic binding site becomes buried (Roseman et al., 1996; Xu et al., 1997). The central cavity is now lined by a relatively polar surface (Xu et al., 1997) which provides an environment in which the protein may fold to a native or near-native conformation (Weissman et al., 1995 & 1996; Mayhew et al., 1996; Rye et al., 1997). This “folding active” complex has a defined lifetime regulated by the rate of ATP hydrolysis within the cis ring which weakens the interaction between the cis ring and GroES (Rye et al., 1997) due to the loss of interactions between the ?-phospate of the nucleotide (Xu et al., 1997). Subsequent ATP binding to the trans ring (Rye et al., 1997) is communicated to the cis ring via movements of the trans equatorial domains (Xu et al., 1997) and inter-ring contacts (Roseman et al., 1996) evicting GroES and both native and non-native polypeptides from the enclosed cavity (Todd et al., 1994; Weissman et al., 1994; Smith & Fisher, 1995; Burston et al., 1996; Ranson et al., 1997). This discharge of polypeptide from the cis cavity allows kinetic partitioning between the native state, other molecular chaperones, the same chaperonin or proteolytic machinery in vivo (Kandror et al., 1994; Buchberger et al., 1996). Iterative cycles of binding, encapsulation and release can be used to actively reverse misfolding/low-order aggregation and thus increase the yield of the native form (Jackson et al., 1993; Todd et al., 1994; Weissman et al., 1994; Ranson et al., 1995; Corrales & Fersht, 1996; Ranson et al., 1997). In a similar manner, although via a different structural mechanism, substrates for CCT in the eukaryotic cytosol also undergo rounds of binding and release (Tian et al., 1995; Farr et al., 1997). Further characterization of the group II chaperonins should yield more details of the similarities and more interestingly the differences between two groups of chaperonins. 8. ACKNOWLEDGMENTS S.G.Burston is a Wellcome Trust International Travelling Fellow. H.R.Saibil thanks the Wellcome Trust and the Biotechnology and Biological Sciences Research Council (UK) for support. The authors thank Art Horwich and Paul Sigler (Yale) for helpful discussions, and Lars Ditzel (Martinsried) for figures of the thermosome and GroELGroES-ADP structures.


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(1990). Chaperonin-facilitated refolding of ribulose-bisphosphate carboxylase and ATP hydrolysis by chaperonin-60 (GroEL). are K+ dependent. Biochemistry , 29, 5665– 5671. Weissman, J.S., Kashi, Y., Fenton, W.A. and Horwich, A.L. (1994). GroEL-mediated protein folding proceeds by multiple rounds of binding and release of non-native forms. Cell , 78, 693–702. Weissman, J.S., Hohl, C.M., Kovalenko, O., Kashi, Y., Chen, S., Braig, K., Saibil, H.R., Fenton, W.A. and Horwich, A.L. (1995). Mechanism of GroEL action—productive release from a sequestered position under GroES. Cell , 83, 577–587. Weissman, J.S., Rye, H.S., Fenton, W.A., Beechem, J.M. and Horwich, A.L. (1996). Characterization of the active intermediate of a GroEL-GroES-mediated protein folding reaction. Cell , 84, 481–490. White, H.E., Chen, S., Roseman, A.M., Yifrach, O., Horovitz, A. and Saibil, H.R. (1997). Structural basis of allosteric changes in the GroEL mutant Arg197->Ala. Nature Struct. Biol. , 4, 690–694. Willison, K.R. and Kubota, H. (1994). The structure, function and genetics of the chaperonin containing TCP-1 (CCT). in eukaryotic cytosol. In “The Biology of Heat Shock Proteins and Molecular Chaperones” (R.I.Morimoto, A.Tissieres and C.Georgopoulos, eds.). pp. 299–312. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY Willison, K.R. and Horwich, A.L. (1996). Chaperonins in archaebacteria and eukaryotic cytosol. In “The Chaperonins” (R.J.Ellis, ed.). pp 107–136. Academic Press, San Diego, CA. Xu, Z., Horwich, A.L. and Sigler, P.B. (1997). The crystal structure of the asymmetric GroEL•GroES• (ADP)7 chaperonin complex. Nature (London) , 388, 741–750. Yaffe, M.B., Farr, G.W., Miklos, D., Horwich, A.L., Sternlicht, M.L. and Sternlicht, H. 91992). TCP1 complex is a molecular chaperone in tubulin biogenesis. Nature (London) , 358, 245–248. Yifrach, O. and Horovitz, A. (1994). Two lines of allosteric communication in the oligomeric chaperonin GroEL are revealed by the single mutation Arg196->Ala. J. Mol. Biol. , 243, 397–401. Yifrach, O. and Horovitz, A. (1995). Nested cooperativity in the ATPase activity of the oligomeric chaperonin GroEL. Biochemistry , 34, 5303–5308. Yifrach, O. and Horovitz, A. (1996). Allosteric control by ATP of a non-folded proteinbinding to GroEL. J. Mol. Biol. , 255, 356–361. Zahn, R., Perrett, S., Stenberg, G. and Fersht, A.R. (1996). Catalysis of amide protonexchange by the molecular chaperones GroEL and SecB. Science , 271, 642–645. Zheng, X.X., Rosenberg, L.E., Kalousek, F. and Fenton, W.A. (1993). GroEL, GroES and ATP-dependent folding and spontaneous assembly of ornithine transcarbamylase. J. Biol. Chem. , 268, 7489–7493. Zondlo, J., Fisher, K.E., Lin, Z., Ducote, K.R. and Eisenstein, E. (1995). Monomerheptamer equilibrium of Escherichia coli chaperonin GroES. Biochemistry , 34, 10334–10339.

24. COMPOSITION AND FUNCTION OF THE EUKARYOTIC CYTOSOLIC CHAPERONIN-CONTAINING TCP-1 KEITH R.WILLISON Institute of Cancer Research, Cancer Research Campaign Centre for Cell and Molecular Biology, Chester Beatty Laboratories, 237 Fulham Road, London SW3 6JB, UK

1. Introduction 2. The Component Parts of CCT 2.1. CCT Gene Family 2.2. CCT is a Group II Chaperonin 2.3. CCT Gene Linkage in Lower Eukaryotes 2.4. Co-chaperones of CCT 2.5. Polypeptide Binding Sites on CCT 3. CCT Mediated Folding 3.1. Spectrum of Substrates 3.2. Tubulin Folding 4. CCT Function 4.1. CCT Functional Cycle 4.2. Co-translational Folding 4.3. Cell Biology 5. Conclusions 6. Acknowledgments 7. References 1. INTRODUCTION Since the discovery and early characterization of the eukaryotic cytosolic chaperonin containing TCP-1 (CCT) in 1992 (Lewis et al., 1992; Gao et al., 1992; Yaffe et al., 1992; Frydman et al., 1992), we have reviewed developments in this field annually (Horwich and Willison, 1993; Willison and Kubota, 1994; Kubota, Hynes and Willison, 1995a; Willison and Horwich, 1996). In contrast to the huge efforts being expended on the structure and reaction mechanism of the GroEL chaperonin of E. coli, there is still surprisingly little work being conducted on CCT. This may be partially due to a


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widespread perception that CCT is merely the eukaryotic equivalent of GroEL and that they must be very similar protein machines, sharing reaction mechanisms and functions in assisting general protein folding. However, evidence is accumulating that CCT is different to GroEL in many respects, particularly in its substrate specificity and interaction with co-chaperones. A stumbling block to the study of CCT is the absence of a high resolution structure, without which we will be unable to understand the significance of its multi-subunit nature and why 8 separate genes evolved to encode the core complex. The purpose of this chapter is not to re-review the early literature on CCT, but to give an up-to-date opinion on the composition and genetic structure of CCT, its natural substrates and mechanism of action. If, as we suspect from present data, CCT is a sequence specific chaperonin for the actins and tubulins, it may be possible in the future to determine the structures of their folding intermediates trapped on CCT. If this experimental approach proves successful, it should provide insight into the general nature of chaperone action and inform us how some partially folded proteins are truly perceived in the eukaryotic cytosol. 2. THE COMPONENT PARTS OF CCT 2.1. CCT Gene Family The studies of Kubota et al. (1994, 1995a, 1995b, 1997a, and 1997b) established the correspondences between the protein subunits of CCT and the genes encoding them. The obtention of the complete sequences of 9 members of the mouse CCT gene family provided the ground work for studies in all other eukaryotic species. It is now clear, both from the molecular genetics of the CCT gene family and from our own studies on the protein composition of mouse and human CCT, using a combination of protein sequencing, mass spectrometry and antibody characterization (Hynes et al., 1995, 1996a, 1996b), and from the protein sequencing studies of Rommelaere et al., (1993) on rabbit reticulocyte cytosolic chaperonin, that there are 8 core components of CCT, and that they are encoded by 8 separate genes. The “extra” CCT subunits in preparations isolated from mouse or bovine testis are now known to be the consequence of the presence of tissue specific CCT genes; mouse contains aCCT related gene, CCT 2, apparently only expressed in testis (Kubota et al., 1997a). A particularly gratifying result, a consequence of the completion of the Saccharomyces cerevisiae genome sequencing project in 1996, was the confirmation of the presence of 8 CCT genes in this organism (Stoldt et al., 1996), each one being clearly the orthologue of one of the 8 core murine genes isolated in our laboratory (Kubota et al., 1997b). It is likely that all eukaryotes possess these 8 CCT genes, since orthologues of one or more CCT genes have been discovered in amphibians (Sun et al., 1995, Dunn and Mercola, 1996), C. elegans (Leroux and Candido, 1995), Tetrahymena (Soares et al., 1994), Drosophila (Ursic and Ganetzky, 1988) and plants (Mori et al., 1992), for example. 2.2. CCT is a Group II Chaperonin There exist two distinct families of ring-shaped chaperonins composed of similar sized

Composition and function of the eukaryotic cytosolic

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subunits around 550 amino acid residues in length. Group I chaperonins, found in eubacteria, mitochondria and chloroplasts, have 7-fold rotational symme-try and E. coli GroEL is the best studied member of this group (see chapters by Burston & Saibil, and Ranson & Clarke, this volume). The structures of GroEL (Braig et al., 1994) and its ATP bound form (Boisvert et al., 1996) and its co-chaperonin GroES (Hunt et al., 1996) and the asymmetric GroEL-GroES-(ADP)7 complex (Xu et al., 1997) have been determined. Group II chaperonins are found in Archaebacteria, TF55 (Trent et al., 1991, 1994) and the thermosome (Phipps et al., 1991, 1993; Waldmann et al., 1995a,b) and the eukaryotic cytosol, CCT (Kubota et al., 1994) also known as TRiC (Frydman et al., 1992) or cytosolic chaperonin (Gao et al., 1992). The lines of evolution of the two groups of chaperonins are consistent with the view that most components of the eukaryotic cytosol are more closely related to archaebacterial relatives and that the organelles of eukaryotic cytosol, mitochondria and chloroplasts, are derived from eubacterial ancestors by symbiosis. The Archaebacterial chaperonins comprise two symmetry groups, with TF55 from Sulfolobus species having 9 subunits per ring (Knapp et al., 1994, Marco et al., 1994b, Kagawa et al., 1995) and the Thermosome from Pyrodictium, Thermoplasma and Methanococcus species all having 8 subunits per ring (Phipps et al., 1991, 1993; Waldmann et al., 1995b; Andra et al., 1996). CCT has 8 subunits per ring (Marco et al., 1994a; Waldmann et al., 1995a; Saibil 1996; Llorca et al., 1998) and we have proposed a model whereby each position in the ring is occupied by one of the 8 constitutively expressed CCT subunit species (Liou and Willison, 1997). We have suggested that it is likely that CCT evolved from a thermosome having 8-fold symmetry (Willison and Horwich, 1996) and recently Nitsch et al. (1997) found the 8-membered rings of the thermosome of thermoplasma acidophilum to contain two subunit species in alternating positions in each ring. Ditzel et al. (1998) have determined the crystal structure of this )4 arrangement with 42-point thermosome and shown it to have an( )4 ( symmetry. Some such thermosome type was the likely immediate precursor of CCT. However, there is so far no evidence from sequence comparisons to indicate that any particular Archaebacterial subunit is more closely related to any of the CCT subunits than any other. Indeed, all the Group II amino acid sequences are about 30–40% identical to each other in pairwise comparisons (Kubota et al., 1994). A most instructive way to examine the evolutionary relatedness of all the chaperonins is to compare the blocks of sequences which constitute their equatorial, nucleotide binding, domains based on the structure of GroEL (Braig et al., 1994; Boisvert et al., 1996) and the sequence analysis of Kim et al., (1994). Figure 1 shows an example of this type of phylogenetic analysis as first performed by Kubota et al., (1994, 1995a). The separateness of the two groups is apparent, as is the more homogeneous nature of Group I. The remarkable result though is the fact that the eukaryotic CCT subunits diverged from each other approximately 2 billion years ago, around the time of the origin of the eukaryotic cell. Depending upon the number of chaperonin genes present in the precursor of the eukaryotic cell, probably one or two as found in present day Archaebacteria, there may have been two or three rapid gene duplication events to provide the sequences from which the 8 modern CCT genes could evolve. A further fascinating question concerning the evolution of CCT is how it managed to evolve into a machine which could bind actins and tubulins which are


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unrelated in sequence, structure and evolutionary origin. The complete sequencing of Methanococcus Table 1 Correspondence between subunit proteins and genes of CCT Protein Source

CCT

CCT

GCT

CCT

CCT

CCT 1

CCT 2 CC

Mouse testis

S3

S4

S5

S9

S2

S6/S7

S10

S8

Bovine testis

P4

P1

P5

P3

—

—

—

P2

6b

4

6a

3

1

—

2

Ccta

Cctb

Cctg

Cctd

Ccte

Cctz1

Cctz2

Cct

Yeast/Worm

(CCT1)

(CCT2)

CCT3)

(CCT4)

(CCT5)

(CCT6)

—

(CC

Yeast/ORFs

YDR212w YIL142w YJL014w YDL143w YJR064w YDR188w —

YJL

Worm/ORFs

CE 07941 CE 25632

—

Rabbit reticulo 5 -cycte Gene Mouse

—

CE 25697 CE 25698

pT 2

pT 2

pT 5

u38846

D43950* L27706

Mouse cDNAs Tcp-1

pT 2

Human cDNAs x52882

w72593* x74801

—

—

pT 12

pT 2.2 pC D78333 N2

The gene and protein nomenclatures for CCT subunits have been described in Kubota et al. (1994) genes for CCT -CCT proteins, respectively); Kubota et al. (1995b) (Criq for CCT ); and Kubot (Cct l and Cct 2 for CCT l and CCT 2 respectively). Numbers are shown for each mammalian C protein prepared from each source; mouse testis (Kubota et al., 1994, 1997a). Bovine testis (Frydm and rabbit reticulocyte (Rommelaere et al., 1993). Clone names are shown for Cct genes of mouse a accession numbers for human sequences. The gene nomenclatures shown in parentheses (CCT1-CC recommended for budding yeast (S. cerevisiae) genes (Stoldt et al., 1996) and C. elegans genes (Le Candido, 1995). The worm ORFs correspond to the EMBL entries for the complete protein sequenc CE 07941 for CCT1. Some human cDNAs are asterisked indicating partial cDNA sequences and no of human cDNA CCT partial sequences are available at NCBI UniGene (http://www.ncbi.nlm.nih.g search using T-COMPLEX as query. This table is modified from Kubota et al., (1995a).


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Figure 1 Phylogenetic analysis of chaperonins based on the amino acid sequences of their putative ATPase domains. The chaperonin family proteins were aligned over the four conserved putative ATPase domains which have been suggested to be involved in ATP binding and hydrolysis (Kim et al., 1994) based upon the crystal structure of GroEL (Braig et al., 1994, Boisvert et al., 1996). The evolutionary tree is based on amino-acid substitutions in the conserved domains and the actual alignments used for this analysis can be found in Kubota et al., (1995a, b). The tree was constructed by the neighbourjoining method (Saitou and Nei, 1987). Rubisco subunit binding protein (RBP) and TF55 have two subunit species and CCT has eight subunit species. The other chaperonins are homo-oligomeric. MM, Mus musculus (Mouse); SC, Saccharomyces cerevisiae (yeast); SS, Sulflobus shibatae (archaebacterium); EC, Escherichia coli (eubacterium); ML, Mycobacterium leprae (eubacterium); BN, Brassica napus (plant). References of sequences are as follows: CCT (Willison et al., 1986; Kirchhoff and Willison, 1990); CCT , CCT , CCT , CCT , CCT and CCT , (Kubota et al., 1994); CCTq, (Kubota et al., 1995b); TF55 (Trent et al., 1991): E. coli GroEL (Hemmingsen et al., 1988); M leprae homologue of GroEL (Mehra et al., 1986); M.musculus Hsp60 (Venner and Gupta, 1990); S. cerevisiae (Hsp60 (Reading et al., 1989); B.napus RBPa and RBPb (Martel et al., 1990). The lengths of branches represent genetic distances between chaperonin proteins determined by the numerical output of the phylogenetic computer analysis.

jannaschii (Bult et al., 1996) shows no actin/hexokinase/Hsp70 ATP binding fold (Flaherty et al., 1991) in this organism although it does contain two tubulin related genes (ORFs 0370 and 0622) and Archaeoglobus fulgidus (Klenk et al., 1997) is similar; no DnaK genes and two Fts-Z tubulin related genes (ORFs 0535 and 0570). However eubacteria do contain the actin/hexokinase/Hsp70 fold and tubulin related genes, as do modern eukaryotic cells, so it is possible that the actin/hexokinase/Hsp70 fold evolved in


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a separate lineage to thermophilic archaebacteria and that a fusion event (Lake, 1991; Doolittle, 1996) brought the precursors of the two cytoskeletal families together in the precursor of the eukaryotic cell. Perhaps when we understand how CCT interacts with folding intermediates of actins and tubulins we will be able to speculate further how the elements of the modern eukaryotic cytoskeleton suddenly evolved and then became virtually fixed in sequence over the subsequent 2 billion years (Willison and Horwich, 1996).

Figure 2 CCT gene linkage in the yeast and worm. The chromosomal locations of all 8 CCT genes in Saccharomyces cerevisiae and in Caenorhabditis elegans are shown representationally. The exact sequences, orientations and locations of the yeast CCT genes can be found at http://speedy.mips.biochem.mpg.de/mips/yeast/. The more precise locations of the worm CCT genes can be found in the C. elegans ACeDB database (Richard Durbin and Jean Thierry-Meig) as described by Leroux and Candido, (1995) and cosmids containing the CCT genes can be accessed by running TBLASTN with cct sequences at www.sanger.ac.uk/projects/c_elegans/blastserver.shtml.See Table 1 for the yeast and worm CCT ORFs. Cosmids for all 8 worm CCT genes have been isolated. Five yeast CCT genes are found in two linkage groups; CCT 3, 8, 5 within 150,000 bps on chromosome X and CCT 6 and 1, 50,000 bp apart on chromosome IV. Three worm CCT genes are closely linked on chromosome II; CCT 1, 4 and 2.

2.3. Gene Linkage in Lower Eukaryotes The CCT genes of mice and humans are distributed across the autosomes and there is no


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linkage of any pairs of CCT genes (Ashworth et al., 1994; Kubota et al., in preparation). This result is not unexpected for a gene family which expanded 2 billion years ago, since there have occurred around 300 chromosomal rearrangements between mouse and human in the 80 million years since they last shared a common ancestor. However, there are strong suggestions of extant ancestral linkage groups in yeast and the worm (Figure 2). The linkage data hints at the existence of two subgroups of CCT genes and, given further data from other species in the future, it may prove possible to discern the order of the duplications and, possibly, the ancestral linkage groups in the original eukaryotic cell and/or its precursor. 2.4. Co-chaperones of CCT The GroES co-chaperonin plays a central role in the mechanism of action of GroEL particularly in helping to form the expanded central cavity in the cis ring and also causing release of bound substrate from the hydrophobic peptide-binding residues into the cavity (Xu et al., 1997) (Chapters Ranson and Clarke; Burston and Saibil, this volume). However, no ring-shaped GroES-like co-chaperonins have been discovered for any of the Group II family and there is no GroES homologous sequence in the Methanococcus jannaschii genome which has a single gene encoding a Group II TF55-like chaperonin (Open reading frame: MJ 0999) (Bult et al., 1996) nor in the Archaeoglobus fulgidus genome (Klenk et al., 1997) which has two Group II thermosome-like chaperonins (Open reading frames AF2238 and AF1451). As we argued in Section 2.2, the GroEL-like Group I chaperonins are a homogeneous, later evolving group of proteins and all cells and organelles containing them also appear to possess a GroES type ring co-chaperonin. It is likely that GroES is a later addition in the evolution of Group I chaperonin function, since the GroES genes seem to be able to duplicate, amplify and diverge; T4 phage even encodes a Gp31 protein which is unrelated in sequence to GroES but nevertheless is able to substitute functionally for it (Hunt et al., 1997). Recently Klumpp et al. (1997) determined the crystal structure of the apical polypeptide binding domain of the thermoplasma acidophilum thermosome -subunit and discovered the presence of a large helical protrusion which they suggest may control access to the central cavity of this chaperonin. The complete structure of the 16-mer confirms that these helices close off the central cavity by forming a lid (Ditzel et al., 1998). Recent negative stain EM analysis of CCT suggests that ATP induces assymmetric movements of the apicol domains in opposite rings and these events could be related to access of the cavity by substrates (Llorca et al., 1998). 2.5. Polypeptide Binding Sites on CCT Electron microscopic studies of GroEL (Chen, 1994; Saibil, 1996; chapter Burston and Saibil, this volume) and group II chaperonins (Phipps et al., 1991, 1993; Marco et al., 1994b) and CCT (Marco et al., 1994a; Waldman et al., 1995a; Saibil, 1996; Llorca et al., 1998) suggest they have the same overall architecture despite the variation in the number of subunits per ring. However, the side views of all the group II chaperonin cylinders suggest that the putative apical polypeptide binding domains are rather smaller than that of GroEL (Braig et al., 1994). Also, CCT seems to have a larger central channel with a


612

diameter of 60Å compared with 45 A for GroEL. We have previously discussed comparisons between GroEL and CCT based on sequence similarities and molecular modelling (Kim et al., 1994; Willison and Horwich, 1996) and argued that the ATPase, equatorial domain is highly conserved in all chaperonins, whereas the putative apical polypeptide binding domains of all the CCT subunits are very divergent and Waldmann et al. (1995b) came to similar conclusions. It is suggested that each CCT subunit has specific binding sites for specific regions of its substrates. This suggestion is presently unproven, but testing each CCT apical domain for substrate binding specificity is a high priority in the field. The determination of the structure of the isolated thermosome apical domain by Klumpp et al. (1997) has been a major advance and shows that the core of the apical domain does in fact resemble GroEL but that the hydrophobic substrate binding regions found in GroEL are absent (chapter Burston and Saibil, this volume). However, as mentioned previously in section 2.4, a helical projection, displaying a novel helix-turnhelix motif, is found in the new structure which may contain the substrate binding sites. In the light of this structure, the amino acid sequences of the thermosome apical domain may be aligned very well with the equivalent regions in the 8 CCT subunits (see Figure 1, Klumpp et al., 1997). There is indication that some of the short stretches of about 10 residues which are conserved between mouse and yeast CCT subunit apical domains (Kim et al., 1994), but which vary between the 8 different CCT subunits within a species, could be potential substrate binding regions (Counsell and Willison, 1998; submitted for publication). Interestingly these regions contain a high proportion of charged residues which leads Klumpp et al. (1997) to discount them as substrate binding sites based on the GroEL model of hydrophobic substrate interaction sites. However, if CCT is a sequencespecific chaperonin it is possible that the subunits may have rather different substrate interaction sites to GroEL. Nevertheless Dobrzynski et al. (1996) identified residues 150– 350 in -tubulin which interact wich CCT (TriC) and which contains a hydrophobic-rich domain. The identification of the specific sequences in substrates which interact with CCT is an urgent problem to solve. 3. CCT-MEDIATED FOLDING 3.1. Spectrum of Substrates In contrast to the large number of proteins which interact with GroEL in vitro and in vivo (Hartl, 1996), a restricted range of substrates has been identified so far for CCT, notably actins, tubulins and their homologues. CCT can capture actins and tubulins upon dilution from denaturants (Gao et al., 1992; Frydman et al., 1992) or upon in vitro translation in reticulocyte lysate (Yaffe et al., 1992; Frydman et al., 1996; Liou et al., 1998). CCT can be recovered from cells with actin and tubulins remaining tightly bound (Sternlicht et al., 1993; Hynes et al., 1995, 1996a) and incubation with ATP at 37°C can cause release of these substrates (Willison et al., submitted). So far, all temperature sensitive mutations in yeast CCT cause defects in actin and/or tubulin behaviour or localization (Ursic and Culbertson, 1991; Miklos et al., 1994; Chen et al., 1994; Vinh and Drubin, 1994; Ursic et al., 1994). Thus, it seems fairly certain that actins and tubulins are bona fide substrates of CCT. There are other CCT binding proteins including luciferase (Frydman et al., 1992),


613

neurofilament fragments (Roobol and Garden, 1993), hepatitis capsids (Lingappa et al., 1994), Gα transducin (Farr et al., 1997) and various unidentified polypeptides (Hynes et al., 1995, 1996a; Willison et al., submitted). The spectrum of proteins which can bind CCT could be as few as some tens of actin and tubulin family members or, as many as hundreds of different proteins, but one has the impression that there are not as many substrates as suggested for GroEL (Hartl, 1996). The identification of the true substrates will be revealed once we know the motifs on the apical polypeptide chain binding regions of CCT subunits which are responsible for binding unfolded substrate proteins. 3.2. Tubulin Folding Gao et al. (1992) discovered the requirement for co-factor proteins in CCT mediated folding of tubulins. It appears that it is not possible to produce assembly competent tubulin through the action of CCT alone (Lewis et al., 1996). Initial studies from Cowan’s laboratory suggested interaction of co-factor A directly with CCT to increase its ATPase activity during folding of -tubulin (Gao et al., 1993, 1994), but subsequent studies showed requirement for co-factor A in sequestration of -tubulin subunits downstream of their release from CCT (Melki et al., 1996). A yeast homologue of cofactor A, Rb12, rescues cells overexpressing toxic levels of -tubulin (Archer et al., 1995). Campo et al. (1994) independently discovered co-factor A and showed that it caused release of -tubulin monomers from C300 complexes. Cowan’s laboratory have now examined the -tubulin pathway in great detail, and a remarkable view of the tubulin folding pathway is emerging in which -tubulin folding requires CCT and four co-factors (FA, FD, FE, FC). -tubulin folding intermediates pass from co-factor to cofactor before attaining an assembly competent state, that is, the ability to form heterodimers with -tubulin (Tian et al., 1996). The 3-D structures of the various, proposed -tubulin intermediates are unknown and, hence it is not yet clear whether the role of the co-factors is to facilitate completion of folding or to stablize aggregationprone tubulin monomers. Tian et al. (1997) now argue that the folding pathways of and -tubulins merge through their interaction together in a complex with FC, FD and FE from which / -tubulin dimers emerge and that tubulin subunits are maintained in activated conformational states by the set of co-factors. However, in addition to co-factor A, several of the co-factors behave operationally as MAPs as pointed out by Tian et al. (1996) for co-factors C and E. Hirata et al. (1998) have just shown an essential role for the S.pombe homologue of FD, Alp1, in microtubule assembly and that Alp1 is associated with microtubules in vivo. It seems that the major roles for the co-factors may not be in folding per se but rather in tubulin monomer and heterodimer sequestration and flux into and out of microtubules. There is no evidence to suggest that any of the four tubulin co-factors identified by Cowan and coworkers interact directly with CCT. In conclusion, it seems clear that tubulin is not released from CCT in a native state, although a portion of -tubulin molecules can reach a quasi-native state without leaving the chaperonin (Tian et al., 1995b; Farr et al., 1997). If, as we think, tubulins are suspended on CCT via multiple contacts, it may be the case that release is ordered and progressive. Once the CCTmediated aspects of tubulin folding are understood it will be easier to comprehend the


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roles of the co-factors in tubulin biogenesis. 4. CCT FUNCTION 4.1. CCT Functional Cycle The nature of the functional cycle of CCT is poorly understood. In vitro, CCT can bind actins, tubulins and luciferase, diluted from denaturant or upon in vitro translation of these substrate proteins in rabbit reticulocyte lysate. Presently there are conflicting models as to the mechanism by which CCT binds actin and tubulin substrates and subsequently uses ATP hydrolysis to release them. There is also conflict concerning the conformation of the released substrates; are they fully native or are they still folding intermediates? Hartl’s laboratory claim that human -actin interacts with TRiC whilst still bound to the ribosome and that folding of actin is facilitated to its native state in a single round of interaction (Frydman et al., 1994, 1996). Cowan’s laboratory suggest that tubulins are released from CCT in a nonnative state and in a series of hand-over reactions proceed to competence for / -tubulin heterodimer formation (Tian et al., 1995a, 1995b, 1996, 1997; Lewis et al., 1996). Horwich’s laboratory have used GroEL and CCT traps to show that, in single rounds of folding, CCT can release a proportion of nonnative tubulins which can be rebound by another chaperonin; simply put, CCT works like GroEL (Farr et al., 1997). Liou et al. (1998) have shown a single-ring mediated assembly and disassembly cycle of CCT in reticulocyte lysate which occurs constitutively at 30°C. This activity could be linked to CCT replication or foliling but its significance for the reaction mechanism of CCT is presently unknown. Much further work is required to resolve the issues. One clear fact is that substrate release is ATP dependent. CCT (Chromobindin A from bovine adrenal medulla and TRiC from bovine testis) has an ATP hydrolysis rate of 14–18 molecules of ATP hydrolyzed/ molecule CCT/min at 37°C (Martin and Creutz, 1987; Frydman et al., 1992). This ATPase activity is relatively slow and, indeed, various aspects of CCT behaviour seem slow. Tian et al. (1995a) showed that actin takes some minutes to attain a conformation which is capturable by CCT, and actins and tubulins interact with CCT for several minutes at 30°C in rabbit reticulocyte lysate (Liou et al., 1998). Sternlicht et al. (1993) measured interactions of actins and tubulins in vivo in pulse-labelled CHO cells and found that these were occurring for several minutes, up to 10 minutes in the case of tubulin. Formation of stable binary complexes between CCT and substrate is favoured in the presence of ADP, suggesting that ADP-bound CCT is the in vivo acceptor state (Melki and Cowan, 1994). Lin and Sherman (1997) have examined genetic interactions between mutations in the ATP binding sites of four CCT subunits in Saccharomyces cerevisiae, Cct 1, 2, 3 and 6, and proposed a simple sequential KNF model of co-operative ATP binding progressing in one direction around the CCT ring (Figure 3). They also suggest a functional hierarchy among CCT subunits in their interaction with nucleotide and that nucleotide binding may initiate at a fixed position in a CCT ring. These are important observations because they raise the possibility that CCT could facilitate ordered capture and release of a substrate bound by two or more subunits as suggested by Liou and Willison (1997) and hence add informational content to a folding reaction which might not be realized by simple


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diffusion and collision reactions onto and off the chaperonin.

Figure 3 Models for subunit order within a single CCT ring. The top diagram shows a model proposing an unique structure for the CCT ring derived from the examination of CCT microcomplexes, present at low abundance in mouse testis and human cells, which are composed of pairs of CCT subunits. Liou and Willison (1997) used these association patterns to suggest a single solution to the topological arrangement of the 8 CCT subunits in the ring. Independently Lin and Sherman (1997) investigated genetic interactions between various mutations in the conserved ATP binding motifs of the Cct1p, Cct2p,


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Cct3p and Cct6p subunits of Saccharomyces cervisiae CCT and discovered a functional hierarchy amongst the subunits. In wild-type CCT the following sequence of co-operative ATP binding/hydrolysis is proposed; ATP→→Cctlp→→Cct3p→→Cct2p→Cct6p. Based upon the nature and strength of the interactions Cct2p and Cct6p are suggested to be neighbouring subunits whereas the number of intervening subunits between Cct1p and Cct3p, Cct3p and Cct2p and Cct6p and Cct1p is undetermined. The two models are mutually consistent.

4.2. Co-translational Folding The role of molecular chaperones in folding of newly synthesized proteins in the eukaryotic cytosol is discussed in detail in the chapter by Welch et al., this volume. We therefore restrict our discussion to the specific role of CCT in this process. As already mentioned above, Frydman and Hartl have presented a model for early events in the CCT (TRiC) folding pathway (Frydman et al., 1994; Frydman and Hartl, 1996; Hartl, 1996) suggesting that TRiC captures and folds actin co-translationally and that there may be functional coupling between the Hsp70/ Hsp40 chaperone machine and TRiC. Our laboratory also found Hsp70 co-purifying with CCT (Lewis et al., 1992; Kubota et al., 1994) and it is possible that Hsp70 bound substrates could occupy CCT. Frydman and Hartl (1996) show that -actin cannot bind TRiC until approximately 150–200 amino acid residues have been translated; therefore, the co-translational interaction can only last as long as it takes to synthesize the remaining C-terminal portion of -actin, say 10–20 seconds, depending upon translation elongation rates. As we discussed in Section 4.1, CCT and substrates remain in contact for several minutes in vitro and in vivo, so it seems that a significant part of CCT function must occur post-translationally. Eggars et al. (1997) have challenged the model of co-translational folding on TRiC through the use of anti-puromycin antibodies to precipitate nascent polypeptide chains, finding Hsp70 associated with them but not any CCT subunits. It is important that TRiC be shown to associate with polysomes in vivo by other techniques hopefully including electron microscopy. 4.3. Cell Biology In addition to its role in the folding of newly synthesized proteins, CCT might have other important functions in cell metabolism. Knowledge of these functions, however, is only emerging. Brown et al. (1996) have shown that antibodies to CCT subunits bind to centrosomes and can inhibit centrosome function after microinjection into living cells. This result implicates CCT in microtubule assembly processes, possibly via interactions with -tubulin, a known CCT binding protein (Melki et al., 1994; Moudjou et al., 1996). Studies in neuronal cells suggest roles for subsets of CCT proteins in neurites (Roobol et al., 1995), possibly in transport of actin or tubulin monomers (Campenot et al., 1996). The human CCTd subunit has appeared as a factor involved in the stimulation of HIV1 TAR RNA (Wu-Baer et al., 1996) suggesting roles for individual CCT subunits in the life cycle of HIV.


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5. CONCLUSIONS CCT appears to be an important component of the protein folding machinery in the eukaryotic cytosol. CCT is made from eight, essential, gene products and present data suggest it is rather specific for folding of actins and tubulins, although there are strong hints of other activities and functions. Hopefully, progress on the struc-tural front will be made over the next few years and this will illuminate the mechanism of CCT activity. Perhaps GroEL and CCT are as similar and different as the Type I and Type II restriction enzymes of bacteria, rather unspecific versus site-specific proteins. 6. ACKNOWLEDGMENTS This work is funded by the Cancer Research Campaign. I thank Hiroshi Kubota for help with Figure 1 and Michel Leroux and Jonathan Hodgkin for help with Figure 2 and Sylvia Holt for help with the manuscript. 7. REFERENCES Andra, S., Frey, G., Nitsch, M., Baumeister, W. and Stetter, K.O. (1996). Purification and structural characterization of the thermosome from the hyperthermophilic archaeum Methanopyrus kandleri. FEBS Lett. , 379 , 127–131. Archer, J.E., Vega, L.R. and Solomon, F. (1995). Rbl2, a yeast protein that binds to tubulin and participates in micro tubule function in vivo. Cell , 82 , 425–434. Ashworth, A., Malik, A.N., Walkley, N.A., Kubota, H., and Willison, K.R. (1994). The Tcp-1 related gene, Cctg, maps to mouse Chromosome 3. Mammalian Genome , 5 , 509–510. Boisvert, B.C., Wang, J., Otwinowski, Z., Horwich, A.L. and Sigler, P.B. (1996). The 2.4 A crystal structure of the bacterial chaperonin GroEL complexed with ATP S. Nature Struct. Biol. , 3(2), 170–177. Braig K., Otwinowski, Z., Hegde, R., Boisvert, D.C.Joachimiak A., Horwich, A.L. and Sigler, P.B. (1994). The crystal structure of the bacterial chaperonin GroEL at 2.8 Å. Nature , 371 , 578–586. Brown, C.R., Doxsey, S.J., Hongbrown, L.Q., Martin, R.L. and Welch, W.J. (1996). Molecular Chaperones and the Centrosome—a Role For TCP-1 In Microtubule Nucleatin. J. Biol. Chem. , 271 , 824–832. Bult, C.J. plus 40 authors (1996). Complete Genome Sequence of the Methanogenic Archaeon, Methanococcus jannaschii. Science , 273 , 1058–1073. Campenot, R.B., Lund, K. and Senger, D.L. (1996). Delivery of Newly Synthesized Tubulin to Rapidly Growing Distal Axons of Rat Sympathetic Neurons in Compartmented Cultures. J. Cell Biol. , 135(3), 701–709. Campo, R., Fontalba, A., Sanchez, L.M. and Zabala, J C. (1994). A 14 kDa release factor is involved in GTP-dependent -tubulin folding. FEBS Lett. , 353 , 162–166 Chen, X., Sullivan, D.S., and Huffaker, T.C. (1994). Two yeast genes with similarity to


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composition of plastid chaperonin-60 and b polypeptide-encoding genes are highly divergent. Gene , 94 , 181–187. Martin, W.H. and Creutz, C.E. (1987). Chromobindin A: a Ca2+ and ATP regulated chromaffin granule binding protein. J. Biol. Chem. , 262 , 2803–2810. Mehra, V., Sweetser, D. and Young, R.A. (1986). Efficient mapping of protein antigenic determinants. Proc. Natl. Acad. Sci. USA , 83 , 7013–7017. Melki, R. and Cowan, N.J. (1994). Facilitated folding of actins and tubulins occurs via a nucleotide-dependent interaction between cytoplasmic chaperonin and distinctive folding intermediates. Mol. Cell Biol. , 14 , 2895–2904. Melki, R., Rommelaere, H., Leguy, R., Vandekerckhove, J. and Ampe, C. (1996) Cofactor A Is a Molecular Chaperone Required for -Tubulin Folding: Functional and Structural Characterization. Biochemistry , 35(32), 10422–10435. Miklos, D., Caplan, S., Martens, D., Hynes, G., Pitluk, Z., Brown, C., Barrell, B., Horwich, A.L. and Willison, K. (1994). Primary structure and function of a second essential member of heterooligomeric TCP1 chaperonin complex of yeast, TCP1 . Proc. Natl. Acad. Sci. USA , 91 , 2743–2747. Mori, M., Murata, K., Kubota, H., Yamamoto, A., Matsushiro, A. and Morita, T. (1992). Cloning of a cDNA encoding the Tcp-1 (t complex polypeptide 1) homologue of Arabidopsis thaliana. Gene , 122 , 381–382. Moudjou, M., Bordes, N., Paintrand, M. and Bornens, M. (1996). -Tubulin in mammalian cells: the centrosomal and the cytosolic forms. J. of Cell Science , 109 , 875–887. Nitsch, M., Klumpp, M., Lupas, A. and Baumeister, W. (1997) The Thermosome: Alternating a and -Subunits Within the Chaperonin of the Archaeon Thermoplasma acidophilum. J. Mol. Biol , 267 , 142–149. Phipps, B., Hoffmann A., Stetter K.O. and Baumeister, W. (1991). A novel ATPase complex selectively accumulated upon heat shock is a major cellular component of thermophilic archaebacteria. EMBO J. , 10 , 1711–1722. Phipps, B.M., Typke, D., Hegerl, R., Volker, S., Hoffmann, A., Stetter, K.O. and Baumeister, W. (1993). Structure of a molecular chaperone from a thermophilic archaebacterium. Nature , 361 , 475–477. Reading, D.A., Hallberg, R.L. and Myers, A.M. (1989). Characterization of the yeast HSP60 gene coding for a mitochondrial assembly factor. Nature , 337 , 655–659. Rommelaere, H., van Troys, M., Gao, Y., Melki, R., Cowan, N.J., Vandekerckhove, J. and Ampe, C. (1993). Eukaryotic cytosolic chaperonin contains t-complex polypeptide 1 and seven related subunits. Proc. Natn. Acad. Sci. USA , 90 , 11975–11979. Roobol, A. and Garden, M.J. (1993). Identification of chaperonin particles in mammalian brain cytosol and t-complex polypeptide 1 as one of their components. J. Neurochem. , 60 , 2327–2330. Roobol, A., Holmes, F.E., Hayes, N.V.L., Baines, A.J. and Garden, A.J. (1995) Cytoplasmic chaperonin complex enter neurites developing in vitro and differ in subunit composition within single cells. J. Cell Sci . 107 , 1477–1488. Saibil, H.R. (1996) Chaperonin Structure and Conformational Changes. In The Chaperonins , Academic Press Inc. 107–135 Saitou, N. and Nei, M. (1987). Neighbor-joining method: A new method for


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reconstructing phylogenetic trees. Mol Biol. Evol. , 4 , 406–425. Soares, H., Penque, D., Mouta, C. and Rodrigues-Pousada, C. (1994). A Tetrahymena orthologue of the mouse chaperonin subunit CCT and its coexpression with tubulin during cilia recovery. J. Biol. Chem. , 269 , 29299–29307. Sternlicht, H., Farr, G.W., Sternlicht, M.L., Driscoll, J.K., Willison, K. and Yaffe, M.B. (1993). The tcomplex polypeptide 1 complex is a chaperonin for tubulin and actin in vivo. Proc. Natl. Acad. Sci. USA , 90 , 9422–9426. Stoldt, V., Rademacher, F., Kehren, V., Ernst, J.F., Pearce, D.A. and Sherman, F. Review (1996): The Cct Eukaryotic Chaperonin Subunits of Saccharomyces cerevisiae and Other Yeasts. Yeast , 12 , 523–529. Sun, H.B., Neff, A.W., Mescher, A.L. and Malacinski, G.M. (1995). Expression of the axolotl homologue of mouse chaperonin t-complex protein 1 during early development. Biochem. Biophys. Acta , 1260 , 157–166. Tian, G., Lewis, S.A., Feierbach, B., Stearns, T., Rommelaere, H., Ampe, C. and Cowan, N.J. (1997). Tubulin subunits exist in an activated conformational state generated and maintained by protein cofactors. J. Cell Biol. , 138 , 821–832. Tian, G., Vainberg, I.E., Tap, W.D., Lewis, S.A. and Cowan, N.J. (1995a). Specificity in chaperonin-mediated protein folding. Nature , 375 , 250–253. Tian, G., Vainberg, I.E., Tap, W.D., Lewis, S.A. and Cowan, N.J. (1995b). Quasi-native chaperonin-bound intermediates in facilitated protein folding. J. Biol. Chem. , 270 , 23910–23913. Tian, G., Huang, Y., Rommelaere, H., Vandekerckhove, J., Ampe, C. and Cowan, N.J. (1996). Pathway Leading to Correctly Folded -Tubulin. Cell 86 , 287–296. Trent, J.D., Nimmesgern, E., Wall, J.S., Hartl, F.-U. and Horwich, A.L. (1991). A molecular chaperone from a thermophilic archaebacterium is related to the eukaryotic protein t-complex polypeptide-1. Nature , 354 , 490–493. Trent, J.D., Gabrielsen, M., Jensen, B., Neuhard, J. and Olsen, J. (1994). Acquired thermotolerance and heat shock proteins in thermophiles from the three phylogenetic domains. J. Bacteriol. , 176 , 6148–6152. Ursic, D. and Ganetzky, B. (1988). A Drosophila melanogaster gene encodes a protein homologus to the mouse t complex polypeptide 1. Gene , 68 , 267–274. Ursic, D. and Culbertson, M.R. (1991). The yeast homolog to mouse Tcp-1 affects microtubule-mediated processes. Mol. Cell Biol. , 11 , 2629–2640. Ursic, D., Sedbrook, J.C., Himmel, K.L. and Culbertson, M.R. (1994). The essential yeast Tcp1 affects actin and microtubules. Mol. Biol. Cell , 5 , 1065–1080. Venner, T.J. and Gupta, R.S. (1990). Nucleotide sequence of mouse HSP60 (chaperonin, GroEL homolog) cDNA. Biophys. Acta , 1087. Vinh, D.B.-N. and Drubin, D.G. (1994). A yeast TCP-1 like protein is required for actin function in vivo. Proc. Natl. Acad. Sci. USA , 91 , 9116–9120. Waldmann, T., Nimmesgern, E., Nitsch, M., Peters, J., Pfeifer, G., Muller, S., Kellermann, J., Engel, A., Hartl, F.-U. and Baumeister, W. (1995a). The thermosome of Thermoplasma acidophilum and its relationship to the eukaryotic chaperonin TRiC. Eur. J. Biochem. , 227 , 848–856. Waldmann, T., Lupas, A., Kellerman, J., Peters, J. and Baumeister, W. (1995b). Primary structure of the thermosome from Thermoplasma acidophilum. Biol. Chem. Hoppe-


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Seyler , 376 , 119–126. Willison, K.R. and Kubota, H. (1994). The structure, function and genetics of the chaperonin containing TCP-1 (CCT) in eukaryotic cytosol. In The Biology of Heat Shock Proteins and Molecular Chaperones. R. I.Morimoto, A.Tissieres and C.Georgopoulos eds., (Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press), pp. 299–312. Willison, K.R. and Horwich, A.L. (1996). Structure and Function of Chaperonins in Archaebacteria and Eukaryotic Cytosol. In The Chaperonins , Academic Press Inc. 107–135. Wu-Baer, F., Lane, W.S. and Gaynor, R.B. (1996) Identification of a group of cellular cofactors that stimulate the binding of RNA polymerase II and TRP-185 to human immunodeficiency virus 1 TAR RNA. J. Biol. Chem. , 271 , 4201–4208. Xu, Z., Horwich, A.L. and Sigler, P.B. (1997) The crystal structure of the asymmetric GroEL-GroES-(ADP) 7 chaperonin complex. Nature , 388 , 741–750. Yaffe, M.B., Farr, G.W., Miklos, D., Horwich, A.L., Sternlicht, M.L. and Sternlicht, H. (1992). TCP1 complex is a molecular chaperone in tubulin biogenesis. Nature , 358 , 245–248.

25. STRUCTURE AND MECHANISM OF HSP70 PROTEINS JEUNG-HOI HA, ERIC R.JOHNSON, DAVID B.McKAY*, MARCELO C.SOUSA, SHIGEKI TAKEDA and SIGURD M.WILBANKS Department of Structural Biology, Stanford University School of Medicine, Stanford, California 94305–5400, USA

1. Introduction 2. Structure 3. ATPase Activity 4. Coupling of the ATPase Activity to the Peptide Binding Actitivity 5. Modulation of Hsp70 Activity 5.1. Self-association 5.2. Posttranslational Modification 5.3. Accessory Proteins 6. Epilogue 7. References 1. INTRODUCTION The 70 kDa heat shock related proteins (referred to generically as “Hsp70s” in this chapter—taken to include the endoplasmic reticulum-resident BiPs, cytoplasmic/ nuclear Hsc70s, prokaryotic DnaKs and others) comprise a family of molecular chaperones that are essential for cell viability. Their biochemical and biological functions are discussed in depth in several chapters of this volume, and hence they will not be elaborated extensively here. In brief, the Hsp70s cyclically bind and release “unstructured” segments of polypeptides; the binding presumably suppresses aggregation or misfolding of the peptide segments, while their release gives them the opportunity to coalesce into a correctly folded tertiary structure. The kinetics of the binding and release cycle is modulated by ATP/ADP; in the presence of ATP, peptide release is relatively rapid, while in the presence of ADP, Hsp70-peptide complexes are stable and long-lived. In vivo, unstructured segments of polypeptide may present themselves in many different contexts ranging from nascent polypeptides emerging from ribosomes during translation (Beckmann et al., 1990; Hartl, 1996; Welch et al., this volume), *Corresponding author


626

polypeptides translocating across membranes into intracellular organelles (Scherer et al., 1990; Schatz and Dobberstein, 1996; chapter in this volume by Welch et al.; Haas and Zimmermann; Dekker and Pfanner; Muckel and Soll) and protein denaturation (e.g. heatdenatured proteins arising from thermal stress of cells (Skowyra et al., 1990; Li et al., this volume). The focus of this chapter is the structure and biochemical mechanism of the Hsp70 proteins. In this context, it is convenient to delineate the in vitro activities of the Hsp70 proteins as (i) an ATPase activity; (ii) a peptide binding activity (discussed in detail by Buchberger et al. in this volume); (iii) a mechanism of coupling binding and release of peptides to the enzymatic ATPase cycle; and (iv) mechanisms of modulating the Hsp70 activities through interaction through self-association, posttranslational modification and interaction with accessory proteins. For a detailed description of the functional cycle of the procaryotic DnaK homolog, and in particular the role of the accessory proteins including DnaJ the reader is referred to the chapter by Buchberger et al. 2. STRUCTURE Approximately 100 amino acid sequences (derived from DNA sequences) are currently known for Hsp70-related proteins (Swiss-prot release 34.0, 4/96). The mature proteins are typically ~640–650 amino acid residues in length. The sequences show a high degree of homology, with the most widely separated prokaryotic and eukaryotic sequences sharing ~50% amino acid identity. The ATPase activity is localized in the amino-terminal ~385 residues, while the peptide binding activity resides in ~160 residues following the ATPase domain (Figure 1). The carboxy terminal regions of the proteins are the most variable in sequence; the eukaryotic Hsp70s have a glycine, proline rich segment of variable length and with an approximate (GGMP)n repeat whose function is still unknown. Additionally, the non-organellar eukaryotic Hsp70s end with a conserved EEVD sequence whose presence is important for stable binding of model denatured polypeptide substrates by Hsp70s (Freeman et al., 1995). The X-ray crystallographic structure of the ATPase domain of bovine Hsc70 is known (Flaherty et al., 1990) (Figure 2). It consists of two lobes with a cleft between them; nucleotide is bound at the base of the cleft and is almost entirely inaccessible to solvent. The tertiary structure of the ATPase domain is highly similar to that of DnaK (Harrison, et al., 1997) and human Hsp70 (Sriram, et al., 1997). The first available structure of an Hsp70 ATPase domain clearly showed a striking similarity to that of actin

Figure 1 Schematic delineation of functional domains within the primary

Structure and mechanism of Hsp70 proteins

627

sequence of Hsp70 proteins.

Figure 2 Structure of the ATPase domain, (a) Schematic drawing of the crystallographic structure of bovine Hsc70 ATPase domain with MgADP, Pi and two K+ ions. Separate structural domains are shown


628

in separate colors. Mg2+ ion is yellow, K+ ions are blue. Drawing made with Molscript (Kraulis, 1991). (b) Folding topology, with domains in same relative orientation as in (a) Secondary structure assignment determined with the program DSSP (Kabsch et al., 1983); numbering referenced to bovine Hsc70 primary sequence.

(Kabsch et al., 1990); further, the structural domains of Hsc70 and actin that are involved in nucleotide binding have identical folding topology and a similar tertiary structure to the equivalent regions in hexokinase (Fletterick et al., 1975) and glycerol kinase (Hurley et al., 1993). Although the overall sequence similarity between actin and Hsc70 is relatively low, many of the residues that interact with MgADP and MgATP are identical, raising the possibility of parallels between the two families of proteins in the manner in which they utilize ATP in their respective functions (Flaherty et al., 1991). A proposal was made several years ago, based on a weak amino acid sequence similarity, that the structure of the peptide binding domain of Hsp70s would be similar to that of the peptide binding portion of major histocompatibiltiy complex (MHC) proteins (Flajnik et al., 1991; Rippmann et al., 1991). This model was appealing since it rationalized the interaction of Hsp70 proteins with unstructured segments of polypeptides. A shadow of doubt first fell on this model when NMR studies of an Hsc70 peptide binding domain led to a secondary structure assignment and folding topology that was fundamentally different from that of MHC peptide binding domains (Morshauser et al., 1995). Recently, the X-ray crystallographic structure of the DnaK peptide binding domain (residues 389–607) complexed with a seven residue peptide (Asn-Arg-Leu-LeuLeu-Thr-Gly), has revealed a novel tertiary fold that is unrelated to the MHC structure (Zhu et al., 1996) (Figure 3). The peptide binding moiety is composed of an 8-stranded antiparallel structure which forms a crevice with a single, deep pocket that accommodates the central leucine of the bound peptide. Although the secondary structure diagram depicts a pair of four-stranded sheets, only one of these segments (strands 3 and 6–8, Figure 3) forms a continuous sheet. In the other segment, the first two strands (strands 1 and 2, Figure 3) arc over the top of the peptide, while the other two strands (strands 4 and 5, Figure 3) run under it. Although biochemical studies suggested that a minimum of seven residues was required for optimal peptide affinity (Flynn et al., 1991), only the central five residues of the peptide make substantial interaction with the peptide binding domain in the crystallographic structure. One residue appears to be “buried” in a binding pocket of the protein; it is the central leucine of the peptide, which is sandwiched between the two pairs of beta strands, and whose side chain protrudes downward into the crevice between the strands in the view shown in Figure 3. A long alpha helix lies across the crevice, but does not actually contact the peptide; it has been suggested that this helix acts as a “latch” to stabilize peptide complexes and to control the kinetics of peptide binding and release. This suggestion implies that the crystal structure shows the stable peptide binding form and that the transient binding form has yet to be observed. The functional imperatives of Hsp70 proteins to recognize a diversity of unstructured polypeptides and to modulate their peptide binding and release rates is consonant with the structure of the DnaK peptide binding domain. The hydrophobic binding pocket for the peptide rationalizes the preference for aliphatic hydrophobic amino acids in peptides bound by Hsp70s (Flynn et al., 1991; Rüdiger et al., 1997a, b; see chapter Buchberger et


629

al., this volume). The long helix that lies over the top of the peptide binding crevice suggests a mechanism whereby peptide binding and release would be relatively slow under circumstances where the helix

Figure 3 Structure of the peptide binding domain, (a) Schematic Molscript drawing of the crystallographic structure of E. coli; DnaK peptide binding domain. Peptide is shown as a ball-and-stick model, with


630

oxygen, red; nitrogen, blue; carbon, gray, (b) Folding topology. Secondary structure assignment determined with the program DSSP; numbering referenced to E. coli DnaK primary sequence.

was latched over the crevice, but relatively rapid if the helix swung outward to allow unhindered access to the crevice. The helix is bound to the peptide binding groove through the polypeptide backbone at the N-terminus and by salt bridges at the Cterminus. Disruption of one of these salt bridges destabilizes peptide binding (Ha, et al., 1997), supporting the hypothesis that the helix serves as a latch controlling access of the peptide to the crevice. 3. ATPase ACTIVITY The steady state ATPase turnover rate (Table 1) and nucleotide affinity (Table 2) have been reported for several Hsp70-related proteins, as well as for the aminoterminal ATPase fragment of bovine and rat Hsc70 (Table 3). The general consensus that emerges from these studies is, (i) the steady state ATPase turnover rate is relatively slow; reported kcat values range 0.02 min-1 to 1.0 min-1, with the faster rates generally measured under conditions of peptide stimulation of the ATPase activity; steady state Km values for MgATP range ~0.1–1 M; (ii) within the enzymatic cycle, ATP hydrolysis is also relatively slow and under many conditions, it appears to be the rate-limiting step of the cycle (Gao et al., 1993; McCarty et al., 1995; Theyssen et al., 1996) (although, by contrast, Pi release is rate-limiting for the isolated ATPase domain of Hsc70 (Ha et al., 1994)); (iii) affinities of both MgATP and MgADP for Hsp70 proteins is submicomolar, with reported values ranging 0.01–1 M; in absence of divalent ion, the affinities are significantly weaker, ~1–10 M; (iv) following ATP hydrolysis, at least for Hsc70 product release is ordered, with Pi preceeding ADP (Ha et al., 1994). The ATPase activity requires Mg2+; the K0.5 for the Mg2+-dependence of the activity is ~1–10 M (Wilbanks et al., 1994). The ATPase activity is also dependent on monovalent ion with a K0.5 20–50 M; K+ (ionic radius=1.33 Å) induces maximal activity, while significantly smaller (e.g. Na+ or Li+ with ionic radii 0.97 Å and 0.68 Å respectively) or larger (e.g. Cs+, ionic radius=1.67 Å) monovalent ions give a substantially lower ATPase activity (O’Brien et al., 1995). A pathway for chemical hydrolysis of ATP has been suggested (Figure 4), based on both the effects of mutagenesis of residues in the nucleotide binding cleft (Table 3) and the comparison of crystallographic structures of the Hsc70 ATPase fragment that have ATP or slowly hydrolyzable analogs bound (AMPPNP bound to wild type protein; ATP bound to several different mutant proteins) to those that have ADP bound (Flaherty et al., 1994). In the suggested scenario, MgATP would initially bind in a conformation that was not correctly aligned for hydrolysis, possibly with an H2O molecule intercalated between the Mg2+ ion and the phosphate of the nucleotide, as observed crystallographically. Then, the phosphate would rearrange to form a “tight” , -bidentate complex with the Mg2+ ion. This is proposed to align the terminal phosphate of the nucleotide for in-line attack by an H2O molecule or OH ion, ultimately yielding ADP and Pi as products. Model-building of a hypothetical transition state of the hydrolysis reaction suggested an


631

experimentally observed H2O molecule that is hydrogen bonded to the -amino group of the side chain of Lys71 in the Hsc70 ATPase fragment as a candidate catalytic H2O molecule. Lys71 could then act either to accept a proton from the H2O molecule or to stabilize an OHTable 1 Kinetic parameters of unstimulated and substrate-stimulated steady state ATPase activities of wild-type Hsp70s derived from data in the literature. Sequence of peptide A: KRQIYTDLEMNRLGK: a-pp peptide: CALLOSRLLLSAPRAAATARA: peptide C: KLIGVLSSLFRPK: peptide A7: RRLIEDAETAARG: S peptide: KETAAAKFEROHMDSSTSAA: cytochrome C peptide: IFAGIKKKAERADLIAYLKQATAK; f-AF1 fluorescen-FYQLALT: Mpp: MALLQSRLLLSAPRAAATARA.

# Protein

Ligand (conc.) (1)

k cat K M ( Assay (min-1) M) condition

1 Bovine Hsc70 natural

0.5 mg/ml triton-washed clathrin cages

1

0.7

20 mM (Braell et al., HEPES 1984) 25 mM KC1 10 mM (NH4) 2SO4 2mMMg (OAc)2 0.8 mM DTT pH 7.0, 37°C


vesicular stomatitis virus glycoprotein peptides

»0.2

nd

(Flynn et al., 20 mM 1989) HEPES 20 mM NaCl 2 mM MgC12 1% sodium cholate 8 M ATP pH 7.0, 37°C


clathrin cages & various peptides derived from clathrin

1

nd

40 mM HEPES 75 mM KC1 4.5 mM Mg (OAc)2 550 M ATP pH 7.0, 37°C

4 Bovine Hsc70

apo cyt C:

natural

0

g/ml

Reference

(DeLuca-Flaherty et al., 1990)

20 mM (Sadis et al., HEPES-KOH 1992) 0.075

1.37

25 mM KC1 10 mM (NH4) 2SO4

Molecular chaperones and folding catalysts 400

# Protein

Ligand (conc.) (1)

5 Bovine Hsc70

peptide C:

natural

0.19

g/ml

k cat (min1)

1.44

K M( M)

632

2mMMg (OAc)2 0.1 mM EDTA 1mM DTT pH 7.0, 37°C

Assay condition

Reference

Reportedly same as (Wang et al., 1993)

no peptide

0.028

nd

1.2mM

0.29

nd


none

0.037±0.002 nd

20 mM imidazole (Gao et al., 25 mM KCl 1994) 10 mM (NH4)2SO4 2 mM Mg(OAc)2 1mM DTT 0.1mM ATP pH 7.0, 25°C

7 Bovine Hsc70

no peptide

»0.02

20 mM imidazole

natural



nd

#3 above.

1 mM of:

25 mM KCl

cyt C peptide

»0.12

10 mM (NH4)2SO4

peptide C

»0.08

2mM Mg (OAc)2

S peptide

»0.04

1mM DTT

KFERQ

»0.03

0.2 mM ATP pH 7.0, 25°C

faf1

nd

10 mM MOPS

no peptide

0.027±0.013

150 mM KCl 4.5 mM Mg(OAc)2

83 M

0.051±0.007 Vmax

pH 7.0, 25°C

faf1

nd

10 mM MOPS

no peptide

0.32±0.02


83 M

0.20±0.08 Vmax

pH 7.0, 25°C 1:1 cysteine string

(Greene et al., 1995)

(Braun et al., 1996)

(Braun, et al., 1996)


633

protein

# Protein

Ligand (conc.) (1)

10 Bovine Hsc70

faf1

natural


12 Bovine Hsc70

nd

(Ha, et al., 1997)


80 M

0.2±0.04

50 g/ml bovine serum albumin

152 M

0.29±0.01 khyd, single turnover

2.3 M Hsc70 10 nM ATP pH 7.0, 37°C

faf1

nd

40 mM HEPES

(Ha et al., 1997)

no peptide

0.029±0.0006

150 mM NaCl 4.5mM Mg (OAc)2

152 M


50 g/ml bovine serum albumin 2.3 M Hsc70 10 nM ATP pH 7.0, 37°C

faf1

Ligand (conc.) (1)

nd

40 mM HEPES

(Ha et al., 1997)

0.016 to 0.072

150 mM KCl

0.1 to 0.52

4.5 mM Mg (OAc)2

khyd, single turnover

50 g/ml bovine serum albumin 2.3 M Hsc70 10 nM ATP pH 7.0, 37°C

kcat (min- 1 )

13 Bovine Hsc70 faf1 no peptide

Reference

40 mM HEPES

0.03±0.006

152 M

recombinant

K M ( Assay condition M)

no peptide

recombinant no peptide

# Protein

k cat (min- 1 )

K M ( Assay M) condition nd

0.006±0.002

40 mM HEPES 150 mM NaCl 4.5 mM Mg (OAc)2

Reference (Ha et al., 1997)

Molecular chaperones and folding catalysts 152 M

14 Rat Hsc70 recombinant


S-peptide:

634

50 g/ml BSA 2.3 M Hsc70 10 nM ATP pH 7.0, 37°C nd

40 mM HEPES

(Wang et al., 1993)

no peptide

0.16

75 mM KCl 4.5 mM Mg (OAc)2

1.2 M

0.38

pH 7.0, 37°C

15 Canine BiP natural

none

0.34

0.11

50 mM MES (Kassenbrock et 25 mM NaCl al., 1989) 1 mM MgCl2 1 mg/ml BSA pH 6.0, 37°C

16 Bovine BiP

no peptide

0.02

nd

20 mM HEPES

natural

20 mM NaCl various peptides

17 Bov. BiP natural

(Flynn et al., 1989)

0.08–0.16

peptide A:

2 mM MgCl2 1 % sodium cholate 8 M ATP pH 7.0, 37°C 20 mM HEPES

no peptide

0.027

20 mM KCl 10 mM (NH4)2SO4

1 mM

0.15

2 mM Mg (OAc)2 0.5 mM DTT 0.25 mM PMSF pH 7.0, 37°C

# Protein

Ligand kcat (min- K M ( (conc.) (1) 1 ) M)

Assay condition

18 Murine BiP

peptide A

20 mM HEPES

(Blond-Elguindi et al., 1993)

Reference (Blond-Elguindi et al., 1993)

Structure and mechanism of Hsp70 proteins recombinant

no peptide: 0.1

0.4

635

20 mM KCl 10 mM (NH4) 2SO4 2 mM Mg(OAc)2 0.5 mM DTT 0.25 mM PMSF pH 7.0, 37°C

1 mM

0.15

19 Hamster BiP recombinant

none

0.4±0.1

1.5±0.2

20 mM HEPES 25 mM KCl 2 mM MgCl2 0.1 mM EDTA 0.5 mM DTT pH 7.0, 37°C

(Wei et al., 1995b)

20 E. coli DnaK overproduced

none

0.15

nd

100 mM HEPES 10 % (v/v) glycerol 35 mM KCl 5 mM MgCl2

(McCarty et al., 1991)

5 mM mercaptoethanol 70 M ATP pH 8.1, 37°C 21 E. coli DnaK overproduced

none

1

200

50 mM Tris-HCl 2 mM MgCl2 0.2 mg/ml BSA 5 mM mercaptoethanol 5% (v/v) glycerol pH 8.0, 30°C

(Zylicz et al, 1983)

# Protein

Ligand (conc.) (1)

K M( kcat 1 (min- ) M)

Assay condition

22 E. coli DnaK

none

0.23

20

30 mM HEPES (Liberek et al., 40 mM KCl 1991a) 50 mM NaCl 7mM Mg (OAc)2 2mM DTT 0.29 mg/ml BSA pH 7.6, 30°C

23 E. coli DnaK

a-pp peptide:

nd

25 mM HEPES (Schmid et al., 1994)

Reference


636

overproduced

no peptide

0.13

100 mM KCl 5 M DnaK

5 M

0.13

24 E. coli DnaK

peptide C:

overproduced

no peptide

0.035

50 mM KCl 11 mM Mg (OAc)2

500 mM

0.24

pH 7.6, 30°C

pH 7.0, 30°C 3.9±0.7

25 mM HEPES-KOH

25 E. coli DnaK

peptide C:

overproduced

no peptide

0.045

50 mM KCl 10mM Mg (OAc)2

200 M

0.41

200 M ATP pH 7.6, 30°C

no peptide

0.03


# Protein

nd

25 mM HEPES-KOH

nd

50 mM HEPES-KOH 50 mM KCl

0.5 mM pep. 0.12 A

10 mM Mg (OAc)2 0.2 mM ATP

0.5 mM pep. 0.21 C

pH 7.6, 30°C

Ligand kcat (min- 1 ) (conc.)

KM(

M)

(Buchberger et al., 1994b)

(McCarty et al., 1995)

(Kamath-Loeb et al., 1995)

Assay condition

Reference

(Theyssen et al., 1996)

(1)

27 E. coli DnaK

none

overproduced

28 E. coli DnaK

peptide Mpp

overproduced

no peptide

single turnover

0.073

50 mM Tris-HCl 100mM KCl

0.09±0.006

Kd

5 mM MgCl2

steady state

2 mM EDTA

0.09±0.01

2 mM dithioerythritol pH 7.5, 25°C nd

0.02 (NaCl) 0.05 (KCl)

25 mM HEPES 100 mM KCl or NaCl 5 mM MgCl2

(Feifel et al., 1996)

Structure and mechanism of Hsp70 proteins 50 M 29 E. coli DnaK overproduced


# Protein

0.02 (NaCl) 0.15 (KCl)

peptide Mpp

0.4 mM ATP pH 7.0, 25°C nd

25 mM HEPES

(Feifel et al., 1996)

none

0.018 -cofactor 0.036+grpE 0.19+DnaJ 2.9+DnaJ+grpE

50 M

0.03 -cofactor 0.048+grpE 0.20+DnaJ 2.9+DnaJ+grpE

none

0.019+0.001 steady state

0.019±0.002

0.018±0.004 khyd

0.001±0.0003 11mM Mg (OAc) Kd 2 50 g/ml BSA pH 7.6, 25°C

Ligand (conc.)

100 mM KCl orNaCl 5 mM MgCl2 1 M DnaK 0.1mM ATP pH 7.0, 25°C

25 mM HEPES 200 mM potassium glutamate

(Russell et al., 1998)

kcat (min1)

K M( M)

Assay condition

Reference

nd

25 mM HEPES 50 mM KCl 11 mM MgCl2, 5 mM mercaptoethanol pH 7.0, 37°C

(Pierpaoli et al., 1997)

50 mM Tris-HCl

(Ziegelhoffer et al., 1995)

(1)


none

0.084±0.006 khyd

32 S. cerevisiae

none

2.5 mM KCl

Ssa1P natural

637

0.028±0.003

4.2±1.0

various [KCl] 2 mM MgCl2

15 mM KCl

0.68 ± 0.006

5 mM mercaptoethaol

0.046±0.003

0.19 ± 0.07

pH 7.5, 33°C

200 mM KCl 0.045±0.007

Molecular chaperones and folding catalysts 33 S. cerevisiae Ssa1P natural

peptide A7:

50 mM Tris-HCl

no peptide 5 mM KCl

200 M

638 (Ziegelhoffer et al., 1995)

various [KCl]

0.01

2 mM MgCl2

50 mM KCl

5 mM mercaptoethaol

0.04

10

5 mM KCl

pH 7.5, 33°C

M ATP

0.01 50 mM KCl 0.1 (1)Sequence

of peptide A: KRQIYTDLEMNRLGK; a-pp peptide: CALLQSRLLLSAPRAAATARA; peptide C: KLIGVLSSLFRPK; peptide A7: RRLIEDAETAARG; S peptide: KETAAAKFERQHMDSSTSAA; cytochrome C peptide: IFAGIKKKAERADLIAYLKQATAK.

Table 2 Binding affinities of adenine nucleotides for Hsp70 proteins

Protein

Nucleotide K d (

Bovine Hsc70 natural

ATP ADP

Bovine Hsc70 natural Bovine Hsc70 natural

M)

Assay conditions

Method/comments Reference

0.7 1.35

40 mM HEPES 75 mM KCl 4.5 mM Mg (OAc)2 0.8 mM DTT, 5% BSA pH 7.0, 37°C

Nucleotide binding proceeded for 5'; then, free nucleotide removed by charcoal.

ATP ADP

9.5±3.9 1.6±0.5

20 mM Na phosphate 200 mM KCl pH 6.5, 4°C

Equilibrium dialysis. (Palleros et al., 1991)

ATP ADP AMPPNP dATP

0.012 0.018 2.8 23

20 mM Imidazole, 25 mM KCl 10 mM (NH4) 2SO4 2 mM Mg (OAc)2 1 mM DTT

Equilibrium dialysis: (Gao et al., AMPPNP and dATP 1994) measured directly; ATP pH 7.0, 25°C and ADP measured by competition with AMPPNP.

(Schmid et al., 1985)


639

Bovine ATP Hsc70 ADP E543K recombinant

0.042±0.007 0.11±0.02

40 mM HEPES Equilibrium binding. (Ha et al., 75 mM KCl (by filter binding) 1994) 4.5 mM Mg (OAc)2 pH 7.0, 25°C

Bovine ATP Hsc70 E543K recombinant

Two-step binding; K1=1.5 M K2=0.03

40 mM HEPES 75 mM KCl 4.5 mM Mg (OAc)2 pH 7.0, 25°C

ADP

0.020±0.002

Kinetics of change in (Ha et al., tryptophan 1995) fluorescence when nucleotides bind.

Protein

Nucleotide K d (mM)

Assay conditions

Method/comments Referenc

Bovine Hsc70

ATP

20 mM HEPES

Kinetics of nucleotide

E543K recombinant

no peptide

0.011±0.005 150 mM KCl with peptide 3 mM Mg(OAc)2 0.022±0.001 pH 7.0, 25°C

(Takeda e al., 1996)

binding, (by flourescence).

ADP

no peptide (peptide=fluorescein 0.048±0.005 -FYQLALT) with peptide 0.099±0.014

Bovine Hsc70 44 kDa ATPase fragment

ADP

0.094±0.012 40 mM HEPES 75 mM KCl 4.5 mM Mg(OAc)2 pH 7.0, 25°C

Equilibrium binding (Ha et al. and kinetics of 1994) nucleotide binding.

Rat Hsc70 recombinant

ATP/ADP

0.2–0.3

25 mM Tris-HCl 120 mM NaCl 5 mM MgCl2 pH 7.0, room temp.

Binding of (Wang et nucleotide to al., 1993) protein immobilized on Sepharose.

Rat Hsc70 recombinant WT, D10N

ATP/ADP

0.3

40 mM HEPES 75 mM KCl 5 mM Mg (OAc)2 pH 7.0, 4°C

Binding of (Huang e nucleotide to al., 1993) protein immobilized on Sepharose.

1.1±0.1 1,400±700

20 mM HEPES Filter binding 25 mM KCl 10 mM (NH4)2SO4 1 mM Mg(OAc)2 pH 7.0, r. t.

Mung bean ATP site 1 Hsc70 ATP site 2 (pig brain was found to be indistinguishable)

Protein

Nucleotide K d (mM)

Assay conditions

(Buxbaum et al., 1996)


Molecular chaperones and folding catalysts E. coli DnaK MABAoverproduced ADP*

640

0.09±0.01

50 mM Tris/HCl Kinetics of change 100 mM KCl in MABA* flourescence;

AMP

17.6±2.5

5 mM MgCl2

Competition with

ADP

0.13±0.03

2 mM EDTA

MABA-ADP* at

AMPPNP

1.6±0.1

2 mM dithioerythritol

equilibrium

ATPgS

0.044±0.002

pH 7.5, 25°C

GDP

90±14

ATP

0.007

(Theyssen et al., 1996)

Kinteics of change in tryptophan fluoresence

E. coli DnaK MABAoverproduced ADP*

20

0 mM Tris/HCl 100 mM KCl 5 mM MgCl2 2 mM EDTA 2 mM dithioerythritol 1:1 grpE pH 7.5, 25°C

E. coli DnaK ATP

0.001±0.0003 25 mM HEPES

overproduced ADP

0.0012±0.0003 200 mM potassium glutamate 11 mM Mg (OAc)2 5 mM PO4 50 g/ml BSA pH 7.6, 25°C

Kinteics of change in MABA* flourescence

(Packschies et al., 1997)

Filter binding

(Russell et al., 1998)

*N8-[4-[(N’-methyl(anthraniloyl)-amino]butyl]-8-aminoadenosine 5’-diphosphate

Protein

Nucleotide K d (mM)

Assay conditions


E. coli DnaK

ATP

20 mM HEPES 100 mM NaCl 100 mM KCl ~10 nM Mg2+, controlled with 20 mM

Equilibrium dialysis.

0.5

(Skowyra et al., 1995)


641

EDTA 0.1 mg/ml BSA 5% glycerol 5 mM DTT pH 7.6, 4°C Hamster BiP

ATP

0.2±0.1

20 mM HEPES

recombinant ADP

0.29±0.12

25 mM KCl

T229G

ATP

0.11±0.1

2 mM MgCl2

ADP

0.15±0.11

0.1 mM EDTA

ATP

0.12±0.11

0.5 mM DTT

ADP

0.16±0.12

pH 7.0, 4°C

T37G

Equilibrium dialysis.

(Wei et al., 1995b)

G227, 226D ATP, ADP not detectable

Table 3 Mutants in the isolated ATPase domain of Hsc70 derived from data in the literature(1)

Protein Mutation E. coli k cat ( min−1 ) DnaK equivalent residue Bovine w.t.

0.8±0.1

Rat*

0.28

w.t.

k cat KM( (mutant) M) k cat (w.t.) 0.7±0.1

KM Reference (mutant) KM (w.t.) (O’Brien et al., 1993) (Huang et al., 1993)

Bovine w.t.

0.53±0.07

0.49±0.26

(Wilbanks et al., 1994)

Bovine w.t. (25° C)

0.14±0.012

0.37±0.09

(Ha et al., 1994)

Rat*

D10N

D8

0

0

(Huang et al., 1993)

Bovine D10N

D8

0.021±0.008

0.04 10.1±4.3

20 (Wilbanks et al., 1994)

Bovine D10S

D8

0.009±0.003

0.02 5.9±2.3

12 (Wilbanks et al., 1994)


642

Bovine C17K

C15

0.02±0.0003*

0.037 0.04±0.01

0.07 (Wilbanks et al., 1998)

Bovine K71A

K70

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