Molecular Chaperons

MOLECULAR CHAPERONES AND FOLDING CATALYSTS

MOLECULAR CHAPERONES AND FOLDING CATALYSTSRegulation, Cellular Function and MechanismsEdited 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 572.645 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 203206, 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 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 III. CELLULAR FUNCTIONS 13 39 3

59 85

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.Jakbkiewicz , 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 IV. MECHANISMS 196 226 260

291 310 325

346

365 384 421

A. Overview 19. Spontaneous versus assisted protein folding R.Jaenicke and R.Seckler 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 479 448

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 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 625 663 537 570 605

693 722

747

PREFACEOne 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 Mller for expert help in preparation of the manuscripts.

CONTRIBUTORSBogdan Banecki Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 Gdansk, Kladki 24 Poland Piotr Barski Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 Gdansk, Kladki 24 Poland Olivier Bensaude Unit de Gntique Molculaire Dpartement de Biologie cole Normale Suprieure 46 rue dUlm 75230 Paris Cedex 05 France Adam Blaszczak Polish Academy of Science Institute of Biochemistry and Biophysics Laboratory of Molecular Biology University of Gdansk 80822 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 fr Biophysik und Physikalische Biochemie Universitt Regensburg Universittsstr. 31 D-93040 Regensburg Germany Bernd Bukau Institut fr Biochemie und Molekularbiologie Universitt 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 fr Biochemie und Molekularbiologie Universitt Freiburg Hermann-Herder-Str. 7 D-79104 Freiburg Germany Marlena Duchniewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 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 fr Biophysik und Physikalische Biochemie Universitt Regensburg Universittsstr. 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 80822 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 943055400 USA Ingrid G.Haas Institut fr Biochemie I Universitt 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 fr Biophysik und Physikalische Biochemie Universitt Regensburg D-93040 Regensburg

Germany Joanna Jakbkiewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 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 943055400 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 80822 Gdansk, Kladki 24 Poland Joanna Krzewska Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 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 80822 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 80822 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 943055400 USA Dominique Missiakas Centre National de Recherche Scientifique LIDSM-CBBM 31 Chemin Joseph-Aiguier 13402 Marseille Cedex 20 France Michel Morange Unit de Gntique Molculaire Dpartement de Biologie cole Normale Suprieure 46 rue dUlm 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-Universitt Am Botanischen Garten 19 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 fr Biochemie und Molekularbiologie Universitt Freiburg Hermann-Herder-Str. 7 D-79104 Freiburg Germany

Joanna Puzewicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 Gdansk, Kladki 24 Poland Satish Raina Centre Medical Universitaire Dpartement de Biochimie Mdicale 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 fr 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 fr Biochemie Universitt Bayreuth D-95440 Bayreuth Germany Robert Seckler Institut fr Biophysik und Physikalische Biochemie Universitt Regensburg D-93040 Regensburg Germany Jrgen Soll Botanisches Institut Christian-Albrechts-Universitt Am Botanischen Garten 19 D-24118 Kiel Germany Marcelo C.Sousa Department of Structural Biology Stanford University School of Medicine Stanford, CA 943055400 USA Shigeki Takeda Department of Structural Biology Stanford University School of Medicine Stanford, CA 943055400 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 80822 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 943055400 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 Universitt des Saarlandes D-66421 Homburg Germany Maciej Zylicz Department of Molecular and Cellular Biology Faculty of Biotechnology University of Gdansk 80822 Gdansk, Kladki 24 Poland

I. INTRODUCTION

1. ASSISTED PROTEIN FOLDINGB.BUKAU1, * , F.X.SCHMID2 and J.BUCHNER31 Institut

fr Biochemie und Molekularbiologie, Universitt Freiburg, HermannHerder-Str. 7, D-79104 Freiburg, Germany 2 Laboratorium fr Biochemie, Universitt Bayreuth, D-95440 Bayreuth, Germany 3 Institut fr Biophysik and Physikalische Biochemie, Universitt 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

Molecular chaperones and folding catalystsfolding enzymes that act very early in the life spans of proteins. 2.2. Molecular Chaperones

6

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 MembersHsp100 ClpA, ClpB, ClpX, ClpY Hsp104, Hsp78

Functions

Book Chapters 2

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) prevention of aggregation and Bose et al; assistance of refolding of misfolded Toft proteins; regulation of activity of kinases and steroid hormone receptors 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 co-chaperone of Hsp70 Buchberger et al.

Hsp90

HtpG

Hsp90, Grp94, ERp99, endoplasmin, Hsp108, gp96 Hsp70, Hsc70, Ssa14, Ssb1, 2, Ssc, Ssh1, Lhs1, Kar2, BiP, Grp78

Hsp70

DnaK, HscA (Hsc66)

DnaJ3

DnaJ, DjlA, CbpA, HscB

Hsp40, Ydj1, Sec63, Auxilin, CSPs, Mdj1, Hdj1, Hdj2 Mge1p

GrpE

GrpE

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

Buchberger et al.

Folding systemsHSP

Prokaryotic MembersIbpA, IbpB

Eukaryotic Members

Functions

Book Chapters2

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

Assisted protein foldingPapD SecB PapD SecB

7Hultgren et al.

assembly of bacterial pili

prevention of folding and Welch et al. targeting of precursor proteins to translocase (bacteria) folding and assembly of collagen folding of proteins in the ER folding of proteins in the ER Bose et al. Haas and Zimmermann Haas and Zimmermann

Hsp47 Calnexin

Hsp47 Calnexin Calreticulin

Calreticulin 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) co-chaperone of GroEL Burston and Saibil; Ranson and Clarke Willison

Hsp10

GroES, gp31

Hsp10, Cpn10

CCT

TF55

TRiC

folding of newly synthesized and misfolded proteins (eukaryotic cytosol)

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

Assisted protein foldingmotifs for chaperones.

9

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, 1019. 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, 480486. Ehrnsperger, M., Grber, 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, 221229. Ellis, J. (1987). Proteins as molecular chaperones. Nature (London), 328, 378379. Ellis, R.J., and Hemmingsen, S.M. (1989). Molecular chaperones: proteins essential for the biogenesis of some macromolecular structures. Trends Biochem. Sci. , 14, 33942. 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, 439449. 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, 499510. Johnson, J.L., and Craig, E.A. (1997). Protein folding in vivo: Unraveling complex pathways. Cell , 90, 201204. Lindquist, S. (1997). Mad cows meet Psi-chotic yeast: the expansion of the prion disease. Cell , 89, 495498. Privalov, P.L. (1979). Stability of proteins. Adv. Protein Chem. , 33, 167241. Prusiner, S.B. (1997). Prion diseases and the BSE crisis. Science , 278, 245251. Thomas, P.J., Qu, B.-H., and Pedersen, P.L. (1995). Defective protein folding as a basis of human disease. Trends Biochem. Sci. , 20, 456459. Wetzel, R. (1996). For protein misassembly, its the I decade. Cell , 86, 699702.

II. REGULATION

2. AUTOREGULATION OF THE HEAT SHOCK RESPONSE IN PROCARYOTESLYNN CONNOLLY1, TAKASHI YURA2 and CAROL A.GROSS3, * 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 941431 Department

1. Introduction 2. Regulation of the 2.1. Discovery of 2.2. How does 2.4. Regulation of 2.5. Regulation of 3. Regulation of the 3.1. Discovery of 3.2. What is the nature of the signal inducing 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

heat shock response regulate the response to temperature shift? stability activity heat shock regulon? (24) heat shock response activity? ?

2.3. Translational regulation of

2.6. What are the signals governing expression of the


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 25C and 40C, deletion of the gene encoding restricts growth to temperatures below 20C (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; Rouvire 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 designationRegulon .3 .3 .3 HtpY DnaK DnaJ B 066.0 H 036.5 H 094.0 H 094.1 10.0 ClpP 10.0 ClpX 10.0 HslA 10.8 HtpG 19.2 HslC 39.3 GapA I 033.5 C 062.5 F 021.5

Molec. Function Weight

Kohara Physical Reference(s) Clones Map

21 69 39 89

? chaperone chaperone protease

?

[Missiakas et al., 1993]

101, 102 11.715.5 [Bardwell et al., 1984] 101, 102 11.715.5 [Bardwell et al., 1986] 148 464.1 468.4 [Gayda et al., 1985]

10.0 Lon

24(22) 46 65 70 80 35.5

protease chaperone ? chaperone ?

148 148

464.1 468.4 464.1 468.4

[Maurizi et al., 1990] [Gottesman et al., 1993] [Chuang et al., 1993]

152 212

501.5 504.2 921.6 936.7

[Bardwell et al., 1987] [Chuang et al., 1993] [Charpentier et

dehydrogenase 330, 331 1872


16al., 1987]

1873 H 034.3 39.8 HslK 40.3 HtpX 56.0 ClpB F 084.1 E 072.0 B 025.3 B 082.0 49 32 84 ? ? chaperone 334 ? 437 1901.2 1904.2 ? 2741.2 2743.7

[Chuang et al., 1993] [Kornitzer et al., 1991] [Kitagawa et al., 1991; Squires et al., 1992] [Lipinska et al., 1988] [Burton et al., 1981] [Herman et al., 1995; Tomoyasu et al., 1993] [Herman et al., 1995; Tomoyasu et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993]

56.8 GrpE 67.0 69.2 FtsJ

26 70 26

nucleotide 438, 439 2757.7 exchange factor 2763.6 sigma factor 509 520 3233.0 3236.2 3331.7 3350.3 3331.7 3350.3

69.2 HflB

70

protease

520

75.0 HslO 75.0 HslP 81.2 HtrM (RfaD) 83.0 IbpB (HtpE, HslS) 83.0 IbpA (HtpN, HslT) 89.0 ClpY (HtpI, HslU)) 89.0 HslV (HtpO) 90.0 HtrC 94.2 GroEL B 056.5 C 014.7

33 30 34 16.3

? ? epimerase chaperone

620, 621 3549.6 3552.0

575, 576 3815.3 3816.4 566, 567 3889.9 3892.7 566, 567 3889.9 3892.7 538, 539 4149.5 4151.7 538, 539 4149.5 4151.7

[Raina et al., 1991] [Allen et al., 1992; Chuang et al., 1993] [Allen et al., 1992; Chuang et al., 1993] [Chuang et al., 1993, Missiakas et al., 1996] [Chuang et al., 1993, Missiakas et al., 1996] [Raina et al., 1990]

G 013.5

15.8

chaperone

D 048.5

49

chaperone

G 021.0

21

protease

21 60

? chaperone 648, 649 4400.5

[Hemmingsen et


174405.7 al., 1988] [Hemmingsen et al., 1988] [Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Aa]

94.2 GroES 94.2 HslW 94.8 HslX 94.8 HslY 94.8 HslZ HtpK

C 015.4

16 22 51 45 37

chaperone ? ?

648, 649 4400.5 4405.7 648, 649 4400.5 4405.7 652 652 652 4430.8 4433.4 4430.8 4433.4 4430.8 4433.4

F 010.1

10

Min Protein Alphanumeric designationRegulon HtpT Regulon: 3.9 DegP (HtrA) A 039.5

Molec. Function Weight

Kohara Physical Reference(s) Clones Map

40

[Aa]

50

protease

117, 118 181182

[Lipinska et al., 1988; Strauch et al., 1989] [Lonetto et al., 1994; Nashimoto 1993; Raina et al., 1995] [Landick et al., 1984; Yura et al., 1984] [Danese et al., 1997]

55.5

sigma factor

435

2718

77.5

F 033.4

sigma factor PPlase

613

3614 3625

74.9 tkpA Others: 29.2 PspA E 026.0 28

625, 626

257, 258 1374 1378 ? ? 260 260 1388.8 1409.9 1388.8 1409.9

[Lipinska et al., 1988; Yamamori et al., 1982] [Chuang et al., 1993] [Chuang et al., 1993]

29.7 HslE 29.7 HslF

60 51

Molecular chaperones and folding catalysts29.7 HslG 30.6 HslI (HtpH) 30.6 HslJ 69.2 HslM 75.0 HslQ 75.0 HslR 93.5 LysU D 060.5 D 033.4 41 36 14 31 24 18 60 ? ? ? ? ? ? LysyltRNA synthetase 260 265 265 520

18[Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Chuang et al., 1993] [Lveque et al., 1990]

1388.8 1409.9 1448.9 1454.5 1448.9 1454.5 3331.7 3350.3

620, 621 3549.6 3552.9 620, 621 3549.6 3552.9 646, 647 4381.8 4383.2

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 30C to 42C, 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 T 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 42C to 30C, 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 30C (Straus et al., 1989; Taura et al., 1989). By one to two doublings after downshift, the cell gradually resumes the 30C 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 620), 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 110210, 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 364433; amino acid 122144) 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 (T=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 (T=18 minutes) is much slower than the in vivo T 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 (T>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 inputsexpression 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.


25

3. REGULATION OF THE

(

) 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 30C, but drives over 90% at the lethal temperature of 50C (Erickson et al., 1987). The continued production of at 50C 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; Rouvire 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 (Rouvire 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 (Rouvire 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 (Rouvire et al., 1996).


26

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; Rouvire 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 (Rouvire 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; Rouvire 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 Peas, 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; Rouvire 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 18C, and cells lacking rapidly accumulate a suppressor of this lethality (De Las Peas et al., 1997b). Cells lacking and containing this suppressor form colonies at 42C to 43C 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; Rouvire 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; Rouvire 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


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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; Rouvire 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 50C 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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