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

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

Molecular chaperones and folding catalysts

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

Assisted protein folding

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

Molecular chaperones and folding catalysts

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 folding

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,

Molecular chaperones and folding catalysts

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

?

Molecular chaperones and folding catalysts

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

[Chuang et al., 1993]

Molecular chaperones and folding catalysts

16

1873

al., 1987]

H 034.3 39.8 HslK

49

?

334

1901.2– 1904.2

[Chuang et al., 1993]

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

Autoregulation of the heat

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

[Chuang et al., 1993]

94.8 HslX

51

?

652

4430.8– 4433.4

[Chuang et al., 1993]

94.8 HslY

45

652

4430.8– 4433.4

[Chuang et al., 1993]

94.8 HslZ

37

652

4430.8– 4433.4

[Chuang et al., 1993]

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

[Chuang et al., 1993]

29.7 HslF

51

?

260

1388.8– 1409.9

[Chuang et al., 1993]

Molecular chaperones and folding catalysts 29.7 HslG

18

41

?

260

1388.8– 1409.9

[Chuang et al., 1993]

36

?

265

1448.9– 1454.5

[Chuang et al., 1993]

30.6 HslJ

14

?

265

1448.9– 1454.5

[Chuang et al., 1993]

69.2 HslM

31

?

520

3331.7– 3350.3

[Chuang et al., 1993]

75.0 HslQ

24

?

620, 621 3549.6– 3552.9

[Chuang et al., 1993]

75.0 HslR

18

?

620, 621 3549.6– 3552.9

[Chuang et al., 1993]

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

Autoregulation of the heat

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

Molecular chaperones and folding catalysts

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

Autoregulation of the heat

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).

Molecular chaperones and folding catalysts

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

Autoregulation of the heat

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

Molecular chaperones and folding catalysts

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.

Autoregulation of the heat

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

Autoregulation of the heat

27

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

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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; 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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USA , 90 , 11019–11023. Liberek, K., Wall, D. and Georgopolous, C. (1995). The DnaJ chaperone catalytically activates the DnaK chaperone to preferentially bind the sigma-32 heat shock transcriptional regulator. Proc. Natl. Acad. Sci , 92 , 6224–6228. Lipinska, B., King, J., Ang, D. and Georgopolous, C. (1988). Sequence analysis and transcriptional regulation of the Escherichia coli grpE gene encoding a heat shock protein. Nuc’eic Acids Res. , 16 , 7545–7562. Lipinska, B., Sharma, S. and Georgopolous, C. (1988). Sequence analysis and regulation of the htrA gene of Escherichia coli: a sigma 32-independent mechanism of heatinducible transcription. Nucleic Acids Res. , 16 , 10053–10067. Lonetto, M.A., Brown, K.L., Rudd, K.E. and Buttner, M.J. (1994). Analysis of the Streptomyces coelicolor sigE gene reveals the existence of a subfamily of eubacterial RNA polymerase sigma factors involved in the regulation of extracytoplasmic functions . Proc. Natl. Acad. Sci. USA , 91 , 7573–7577. Martin, D.W., Schurr, M.J., Mudd, M.H., Govan, J.R., Holloway, B.W. and Deretic, V. (1993). Mechanism of conversion to mucoidy in Pseudomonas aeruginosa infecting cystic fibrosis patients. Proc. Natl. Acad. Sci. USA , 90 , 8377–8381. Martin, D.W., Schurr, M.J., Yu, H. and Deretic, V. (1994). Analysis of promoters controlled by the putative sigma factor AlgU regulating conversion to mucoidy in Pseudomonas aeruginosa: relationship to sigma E and stress response. J. Bacteriol. , 176 , 6688–6696. Maurizi, M.R., Clark, W.P., Katayama, V., Rudikoff, S., Pumphrey, J., Bowers, B. and Gottesman, S. (1990). Sequence and structure of ClpP, the proteolytic component of the ATP-dependent Clp protease of Escherichia coli. J. Biol. Chem. , 265 , 12536– 12545. McCarty, J.S., Rudiger, S., Schonfeld, H.J., Schneider-Mergener, J., Nakahigushi, K., Yura, T. and Bukau, B. (1996). Regulatory region C of the E. coli heat shock transcription factor, sigma32, constitutes a DnaK binding site and is conserved among eubacteria. J. Mol. Biol. , 256 , 829–837. Mecsas, J., Rouvière, P.E., Erickson, J.W., Donohue, T.J. and Gross, C.A. (1993). The activity of sigma E, an Escherichia coli heat-inducible sigma-factor, is modulated by expression of outer membrane proteins. Genes Dev. , 7 , 2618–2628. Missiakas, D., Betton, J.-M. and Raina, S. (1996). New components of protein folding in extracytoplasmic compartments of Escherichia coli SurA, FkpA and Skp/OmpH. Mol. Micro. , 21 , 871–884. Missiakas, D., Schwager, F., Betton, J.M., Georgopoulos, C. and Raina, S. (1996). Identification and characterization of HslV HslU (ClpQ ClpY) proteins involved in overall proteolysis of misfolded proteins in Escherichia coli. EMBO J . 15 , 6899– 6909. Missiakas, D., Mayer, M.P., Lemaire, M., Georgopoulos, C. and Raina, S. (1997). Modulation of the Escherichia coli (RpoE) heat-shock transcription-factor activity by the RseA, RseB and RseC proteins. Mol. Microbiol . 24 , 355–371. Mori, K., Ma, W., Gething, M.J. and Sambrook, J. (1993). A transmembrane protein with a cdc2+/ CDC28-related kinase activity is required for signaling from the ER to the nucleus. Cell , 74 , 743–756. Naczynski, Z.M., Mueller, C. and Kropinski, A.M. (1995). Cloning the gene for the heat shock response positive regulator (sigma 32 homolog). from Pseudomonas aeruginosa. Can. J. Microbiol. , 41 , 75–87. Nagai, H., Yuzawa, H., Kanemori, M. and Yura, T. (1994). A distinct segment of the

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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

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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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Sorger, P.K, and Pelham, H.R.B. (1987). Purification and characterization of a heat shock element binding protein from yeast. EMBO J. , 6 , 3035–3041. Sorger, P.I., Lewis, M.J. and Pelham, H.R.B. (1987). Heat shock factor is regulated differently in yeast and HeLa cells . Nature , 329 , 81–84. Theodorakis, N.G., Zand, D.J., Kotzbauer, P.T., Williams, G.T. and Morimoto, R.I. (1989). Hemin-induced transcriptional activation of the hsp70 gene during erythroid maturation in K562 cells is due to a heat shock factor-mediated stress response. Mol. Cell. Biol. , 9 , 3166–3173. Ubeda, M., Wang, X.-Z., Zinszner, H., Wu, I., Habener, J. and Ron, D. (1996). Stressinduced binding of transcription factor CHOP to a novel DNA-control element. Mol. Cell. Biol. , 16 , 1479–1489. Vuister, G.W., Kim, S.J., Wu, C. and Bax, A. (1994a). NMR evidence for similarities between the DNA binding regions of Drosophila melanogaster heat shock factor and the helix-turn-helix and HNF-3/forkhead families of transcription factors. Biochemistry , 33 , 10–16. Vuister, G.W., Kim, S.J., Orosz, A., Marquardt, J., Wu, C. and Bax, A. (1994b). Solution structure of the DNA binding domain of Drosohpila heat shock transcription factor. Nature Struct. Biol. , 1 , 605–614. Wang, X.-Z. and Ron, D. (1996). Stress0induced phosphorylation and activation of the transcription factor CHOP (GADD153). by p38 MAP -kinase. Science , 272 , 1347– 1349. Watowich, S.S. and Morimoto, R.I (1988). Complex regulation of heat shock and glucose responsive genes in human cells. Mol Cell. Biol. , 8 , 393–405. Welihinda, A.A. and Kaufman, R.J. (1996). The unfolded protein response pathway in saccharomyces cerevisiae. J. Biol. Chem. , 271 , 18181–18187. Westwood, J.T., Clos, J. and Wu, C. (1991). Stress-induced oligomerization and chromosomal relocalization of heat-shock factor. Nature , 353 , 822–827. Westwood, J.T. and Wu, C. (1993). Activation of Drosophila heat shock factor: conformational change associated with a monomer-to-trimer transition. Mol. Cell. Biol. , 13 , 3481–3486. Wiederrecht, G., Seto, D. and Parker, C.S. (1988). Isolation of the gene encoding the S. cerevisiae heat shock transcription factor. Cell , 54 , 841–853. Williams, G.T. and Morimoto, R.I. (1990). Maximal stress-induced transcription from the human hsp70 promoter requires interactions with the basal promoter elements independent of rotational alignment . Mol. Cell. Biol. , 10 , 3125–3136. Wisniewski, J., Orosz, A., Allada, R. and Wu, C. (1996). The C-terminal region of drosophila heat shock factor (HSF). contains a constitutively functional transactivation domain. Nucleic Acids Res. , 24 , 367–374. Wu, C. (1980). The 5’ ends of Drosophila heat shock genes in chromatin are hypersensitive to DNase1. Nature , 286 , 854–860. Wu, C., Wilson, S., Walker, B., David, I., Paisley, T., Zimarino, V. and Ueda, H. (1987). Purification and properties of Drosophila heat shock activator protein. Science , 238 , 1247–1253. Wu, C. (1984). Two protein-binding sites in chromatin implicated in the activation of heat-shock genes. Nature , 309 , 229–234. Wu, C. (1995). Heat shock transcription factors: structure and regulation. Annu. Rev. Cell Dev. Biol. , 11 , 441–469. Wu. C. (1995). Heat stress transcription factors. Annu. Rev. Cell Biol. , 11 , 441–469. Xiao, H. and Lis, J.T. (1988). Germline transformation used to define key features of the

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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

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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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C/EBP-homologous protein (CHOP/GADD153). Mol. Cell. Biol. , 16 , 4273–4280. Wang, X.Z. and Ron, D. (1996). Stress-induced phosphorylation and activation of the transcription factor CHOP (GADD153) by p38 MAP kinase. Science , 272 , 1347– 1349. Weeks, J.R., Hardin, S.E., Shen, J., Lee, J.M. and Greenleaf, A. (1993). Locus-specific variation in phosphorylation state of RNA polymerase II in vivo: correlations with gene activity and transcript processing. Genes & Dev. , 7 , 2329–2344. Welch, W.J. (1992). Mammalian stress response: cell physiology, structure/function of stress proteins, and implications for medicine and disease. Physiol. Rev. , 72 , 1063– 1081. Welihinda, A.A. Tirasophon, W., Green, S.R. and Kaufman, R.J. (1998). Protein serine/threonine phosphatase Ptc2P negatively regulates the unfolded-protein response by dephosphorylating Ire1p kinase. Mol. Cell Biol. , 18 , 1967–1977. Xia, Z., Dickens, M., Raingeaud, J., Davis, R.J. and Greenberg, M.E. (1995). Opposing effects of ERK and JNK-p38 MAP kinases on apoptosis. Science, 270 , 1326–1331. Xing, J., Ginty, D.D. and Greenberg, M.E. (1996). Coupling of the RAS-MAPK pathway to gene activation by RSK2, a growth factor-regulated CREB kinase. Science , 273 , 959–963. Yang, D., Tournier, C., Wysk, M., Lu, H.T., Xu, J., Davis, R.J. and Flavell, R.A. (1997). Targeted disruption of the MKK4 gene causes embryonic death, inhibition of c-Jun NH2-terminal kinase activation, and defects in AP-1 transcriptional activity. Proc. Natl. Acad, Sci. USA , 94 , 3004–3009. Yueh, A. and Schneider, R.J. (1996). Selective translation initiation by ribosome jumping in adenovirus-infected and heat-shocked cells. Genes & Dev. , 10 , 1557–1567. Zanke, B.W., Boudreau, K., Rubie, E., Winnett, E., Tibbles, L.A., Zon, L., Kyriakis, J., Liu, F.-F. and Woodgett, J.R. (1996). The stress-activated protein kinase pathway mediates cell death following injury induced by cis-platinum, UV irradiation or heat. Curr. Biol. , 6 , 606–613. Zarzov, P., Mazzoni, C. and Mann, C. (1996). The SLT2 (MPK1) MAP kinase is activated during periods of polarized cell growth in yeast. EMBO J. , 15 , 83–91. Zhou, M., Lambert, H. and Landry, J. (1993). Transient activation of a distinct serine protein kinase is responsible for 27-kDa heat shock protein phosphorylation in mitogen-stimulated and heat-shocked cells. J. Biol. Chem. , 268 , 35–43. Zigmond, S.H. (1996). Signal transduction and actin filament organization. Curr. Op. in Cell Biol. , 8 , 66–73. Zu, Y.L., Wu, F., Gilchrist, A., Ai, Y., Lbadia, M.E. and Huang, C.-K. (1994). The primary structure of a human MAP kinase activated protein kinase 2. Biochem. Biophys. Res. Commun. , 200 , 1118–1124.

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

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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

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89

(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).

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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 (p<0.01). On the other hand, the differences among Bgl, Rat-1, Sma, C-S, S-C and MV6 cells are not statistically significant (p-values around 0.4).

Rat-1 fibroblasts expressing a cloned human hsp70 gene, designated M21 (Li et al., 1992a). The constitutive expression of the human hsp70 gene in Rat-1 cells confers heat resistance as evidenced by the enhanced survival of heat-treated cells and resistance against heat-induced translational inhibition (termed translational tolerance) (Liu et al., 1992). In addition, after a 45°C, 25-minute heat treatment, the time required for RNA and protein synthesis to recover was considerably shorter in M21 cells than in control Rat-1 cells. These data demonstrate that the expression of human hsp70 in Rat-1 cells confers heat resistance and translational tolerance, and enhances the ability of cells to recover from heat-induced translational and transcriptional inhibition (Table 3). The effects of mutant human Hsp70 on the cellular heat sensitivity, transcriptional/translational activity after heat shock were studied (Li et al., 1995; Table 3). Neither a 4-bp out-of-frame deletion ( 21) nor an in-frame deletion of the nucleolar localization domain ( Sma) of human Hsp70 significantly affected the cells’ intrinsic heat sensitivity, heat-induced transcriptional/translational inhibition, or the subsequent recovery at 37°C. In contrast, cells expressing a mutant human Hsp70 missing the ATPbinding domain ( Bgl) were found to be resistant to both heat killing and translational inhibition. However, when the recovery kinetics of protein synthesis were evaluated, no significant differences were observed between Rat-1 cells and cells expressing any of these mutated forms of Hsp70. These studies provide strong support for a link between the expression of functional Hsp70 and protection of cellular procedures at elevated temperature. They further suggest that ATP-binding and/or hydrolysis by Hsp70 are dispensable in the Hsp70-mediated protection against thermal killing and translational inhibition. Perhaps Hsp70 lacking its ATP-binding domain can still bind to cellular proteins, stabilizing them and preventing their aggregation at elevated temperatures. On the other hand, because the mutant Hsp70 Bgl does not facilitate cells’ recovery from translational inhibition, ATP-binding and/or hydrolysis may be important in enabling the dissociation of Hsp70 from its substrates, or in facilitating the dissociation of aggregated protein complexes to restore their functional integrity. The ability of Hsp70 to protect cellular processes and enzymatic activities against heat

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stress has been demonstrated in a series of experiments. For example, in vitro studies by Skowyra et al., (Skowyra et al., 1990) have shown that the E. coli Hsp70 homologue, DnaK, can protect RNA polymerase from heat-induced inactivation, and that thermally inactivated RNA polymerase can be reactivated by DnaK in a process that requires ATP hydrolysis. Furthermore, the thermal protection efficiency of DnaK is enhanced by the action of its partner proteins DnaJ and GrpE (Hendrick et al., 1993). Similarly, mitochondrial Hsp60 has been shown to have thermal protective functions through its ability to prevent the thermal inactivation of dihydrofolate reductase imported into the mitochondria (Martin et al., 1992). It was also demonstrated that the in vivo enzymatic activities of luciferase and bgalactosidase were protected in thermotolerant mouse and Drosophila cells against a heat challenge (Nguyen et al., 1989). To extend these studies further to test the possibility that the intranuclear microenvironment may affect the stability of proteins under heat shock, Michels et al., (Michels et al., 1995) compared the in situ thermal stability of a reporter protein localized in the nucleus or in the cytoplasm, using recombinant firefly luciferase targeted to the cytoplasm (cyt-luciferase) or the nucleus (nuc-luciferase), respectively. In each case, decreased luciferase activity and solubility were found in lysates from heat-shocked cells, indicating thermal denaturation in situ. However, nucluciferase was more susceptible suggesting that the microenvironment of an intracellular compartment may modulate the thermal stability of proteins. In thermotolerant cells, the thermal inactivation of the recombinant luciferases by a second heat shock occurred at a slower rate with a lesser effect for the nuc-luciferase (threefold) than for the cytluciferase (sevenfold). The heat-inactivated luciferases were partially reactivated during recovery after stress, suggesting the capacity of both the cytoplasmic and nuclear compartments to reassemble proteins from an aggregated state. To further test the hypothesis that Hsp70 plays a role in protecting cells from thermal stress, the reporter gene firefly luciferase was constitutively expressed in mammalian Rat-1 cells or M21 cells (Rat-1 cells expressing high level of human Hsp70 via stable transfection). The effects of 43°C heat shock on luciferase activity in these cells were determined; in parallel, the solubility of luciferase in heat-shocked cells were monitored by immunoblot analysis of the soluble and insoluble protein fractions using antisera against luciferase, respectively. Our data demonstrate that in cells expressing a high level of intact human Hsp70, the heat inactivation of luciferase enzyme activity is attenuated (Li et al., 1992b). Similarly, a decrease in the insoluble fraction of luciferase molecules in these cells is observed (Li et al., 1992b). These data support the hypothesis that Hsp70 protects cells either by stabilization and prevention of thermal denaturation of normal proteins, and/or by facilitating the dissociation of protein aggregates. 3.2. Heat Shock Proteins and RNA Splicing Heat shock proteins have also been implicated in protecting the splicing machinery from disruption by heat shock (Yost et al., 1986; Yost et al., 1988; Yost et al., 1991): high temperatures are found to result in the accumulation of mRNA precursors due to the block in splicing. However, in thermotolerant cells, splicing is not disrupted and mature mRNAs accumulate. The mechanism by which this process is protected in thermotolerant

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cells is unclear. 3.3. Heat Shock Proteins and Apoptosis Heat induces cell death both in terms of loss of clonogenicity and apoptosis. Recently, it has been shown that thermotolerance protects against heat-induced apoptosis in human acute lymphoblastic leukemic T-cells (Mosser et al., 1992), as evidenced by a reduction in DNA fragmentation and in the fraction of cells with morphological appearance of apoptosis. However, whether heat shock proteins have a direct role in the protection against apoptosis is yet unknown. We studied heat-induced apoptosis and loss of clonogenicity in Rat-1 fibroblasts, thermotolerant Rat-1 (TT Rat-1), Rat-1 transfected with the human Hsp70 gene (M21) and Rat-1 transfected with the human c-myc protooncogene (Rat-1: myc). A significant fraction of 44°C heat-treated rat fibroblasts exhibited nuclear chromatin condensation, cytoplasmic contraction, loss of adhesion, dye exclusion, and DNA laddering (Figure 3, A-D). Relative to Rat-1, TT Rat-1 and M21 cells are heat-resistant, but Rat-1: myc cells are heat-sensitive, both in terms of apoptosis induction and clonogenic survival (Figure 3, E and F). This study suggests that apoptosis is an important mechanism of heat-induced cell killing in some cell lines, which is suppressed by an overexpression of the human HspTO gene, but enhanced by c-myc expression. These data also suggest that the effects of these genes on thermal sensitivity is in part due to their modulation of apoptosis; although, other factors as yet unknown may also contribute. Vidair and Dewey reported that heated cells died by two distinct modes (Dewey 1989, Vidair et al., 1988). In the “slow” mode, postulated to be associated with chromosome aberration, multinucleation, and aberrant mitotic division, cells maintain their physiological activity for several days prior to the loss of reproductive capability

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Figure 3 Induction of apoptosis in Rat-1, thermotolerant Rat-1 (TT Rat-1), and Rat-1 cells transfected with human Hsp70 (M21). or c-myc protooncogene (Rat-1: myc). In (A) and (B), cells were heated at 44°C for 30 min and studied 12 hours after heating. Cells were stained with 10 mg/ml acridine orange and studied under fluorescence microscope at 200X. (A) Detached cells: most cells are apoptotic as evidenced by chromatin condensation or nuclear fragmentation. (B) Attached cells: most cells are non-apoptotic. (C) DNA gel electrophoresis. Lane 1: Rat-1: myc, unheated control; lane 2: Rat-1: myc, attached cells, 12 hours after heating at 44°C for 30 min; and lane 3: Rat-1: myc, detached cells, same heat treatment as for lane 2. Equal amount of DNA (3 mg). was loaded per lane. (D) DNA gel electrophoresis of control and heated (24 hours after heating at 44°C for 40 min). Rat-1, TT Rat-1 and M21 cells. Lanes 1 and 2: heated M21 and Rat-1 cells; lanes 3 and 4: control M21 and Rat-1 cells; lanes 5 and 6: heated Rat1 and TT Rat-1 cells; lane 7: control TT Rat-1 cells. The attached and detached cells were pooled together for DNA gel electrophoresis

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analysis (3 mg DNA was loaded per lane). (E) Surviving fraction of Rat-1, TT Rat-1, M21 and Rat-1: myc cells heated for various periods at 44°C. □-, Rat-1; -●-, TT Rat-1; -■-, M21; and -O-, Rat-1: myc. (F) Dose response of heat-induced apoptosis assayed 24 hours after heating for Rat-1, TT Rat-1, M21 and Rat-1: myc cells. -□-, Rat-1; -●-, TT Rat1; -■-, M21; and -O-, Rat-1: myc. As in (D), cells were pooled and used for the analysis.

(Dewey, 1989; Vidair and Dewey, 1988; Vidair et al., 1991). In the “rapid” mode, the underlying mechanism which is as yet unknown, cell death occurs within a few days after heating and is characterized by cell detachment from the culture surface. That the similar time course and the same loss of adhesion are observed for both this “rapid mode” and heat-induced apoptosis suggests further investigation to assess whether a common mechanism underlies these two phenomena. Recently, Strasser and Anderson have shown that both Bcl-2 gene expression and thermotolerance protect cells from heat-induced apoptosis and promote cell proliferation after thermal stress (Strasser et al., 1995). Based on these results, the authors suggested that heat shock induces apoptosis by two independent pathways, one inhibited by Bcl-2 and the other by the heat shock proteins. Our finding that Hsp70 overexpression reduces heat-induced apoptosis complements their findings. It has been reported that Hsp70 can interact with and alter the conformation of p53 protein (Hainaut et al., 1992), although the biological significance of p53-Hsp70 complexes is far from understood. Given the current belief that the induction of apoptosis requires p53 (Lowe et al., 1993, Lowe et al., 1993), the putative inactivation of p53 by Hsp70 may underly the observed decrease in apoptosis in M21 and IT Rat-1 cells. The protein product of the c-myc proto-oncogene is a transcription factor which has strong mitogenic effect. The induction of apoptosis by cmyc is believed to be due to a conflict of growth signals centered upon the G1/S transition (Evan et al., 1992; Shi et al., 1992), although the exact mechanism is largely unknown. It has also been reported that overexpression of c-myc gene enhances the induction of apoptosis by other insults, e.g. IL-3 deprivation or ionizing radiation (Askew et al., 1991; Clarke et al., 1993). Thus, our observation of the effect of cmyc in enhancing heat-induced apoptosis is consistent with the previous findings from other laboratories. 3.4. Thermotolerance and Heat Shock Protein Hsp27 In addition to Hsp70, the small heat shock protein, Hsp27, has been shown to have thermal protective functions. Landry and co-workers have overexpressed Hsp27 from various species into different cell lines and have observed that resistance to heat shock correlates with levels of Hsp27 (Huot et al., 1991; Lavoie et al., 1993). In addition it appears that phosphorylation of Hsp27 may play an important role in its thermal protective function. In support of this, Crete and Landry (Crete and Landry, 1990) observed that chemical agents such as cycloheximide, A23187 and EGTA, which induce phosphorylation but not accumulation of Hsp27, resulted in a significant degree of thermal protection. Furthermore, phosphorylation mutants of Hsp27 failed to protect cells from heat stress (Landry et al., 1993). However, this issue remains controversial because of recent evidence that the chaperone properties of the small heat shock protein

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contributes to the increased cellular thermoresistance in a phosphorylation-independent manner (Knauf et al., 1994). This study suggests that the phosphorylation dependent function of Hsp27 is distinct from its thermoresistance-mediating functions.

4. REGULATION OF HEAT SHOCK RESPONSE: POSSIBLE INVOLVEMENT OF KU AUTOANTIGEN Although the importance of the heat shock transcription factor HSF1 in the regulation of mammalian heat shock genes is well-established (Morimoto et al., 1990; see chapter by R.Morimoto), recent data indicate that activation of HSF1, by itself, is not sufficient for the induction of Hsp70 mRNA synthesis, suggesting the existence of additional regulatory factors (Jurivich et al., 1992; Liu et al., 1993; Mathur et al., 1994). A constitutive heat shock element binding factor (CHBF) was recently characterized in rodent cells that exhibits a tight inverse correlation with the HSE-binding activity of HSF1 in vitro (Kim et al., 1995; Liu et al., 1993). Upon heat shock, a rapid increase in the level of HSF1-HSE binding activity correlates with a rapid decrease in CHBF-HSE binding activity. During post-heat shock-recovery, as rates of heat shock gene transcription return to their pre-heat-shock levels, the disappearance of HSF1-HSE binding activity parallels the reappearance of CHBF-HSE binding activity. Additionally, HSF1-HSE binding is not sufficient for heat shock gene induction; agents such as sodium salicylate (Jurivich et al., 1992) and arsenite (Liu et al., 1993) elicit considerable HSF1HSE binding activity, yet do not result in significant induction of Hsp70 mRNA synthesis. Interestingly, unlike heat shock stimuli, these agents do not inactivate the CHBF activity, which remains at the high levels observed in untreated cells (Liu et al., 1993). These findings suggest that CHBF may be involved in the regulation of heat shock gene expression, possibly by acting as a represser (Liu et al., 1993). Further biochemical analysis of CHBF revealed that it is identical or closely related to the Ku autoantigen (Kim et al., 1995). The Ku autoantigen is a heterodimer of 70-kDa (Ku70) and 86-kDa (Ku80) polypeptides (Mimori et al., 1981, Reeves et al., 1989). In vitro studies have shown that this abundant nuclear protein binds to the termini of doublestranded DNA and DNA ending in stem-loop structures (Mimori et al., 1986). By virtue of its DNA binding activity, Ku serves as a regulatory component of the mammalian DNA-dependent protein kinase, DNA-PK, which is activated by DNA ends (Gottlieb et al., 1993). The other component of DNA-PK is a 460-kDa catalytic subunit (DNA-PKcs). Various cellular roles for Ku have been suggested, including a role in transcription, recombination, replication, and DNA repair (Amabis et al., 1990; Bannister et al., 1993; Dvir et al., 1993; Falzon et al., 1993; Gottlieb and Jackson, 1993; Lees-Miller et al., 1991; Lees-Miller et al., 1990; Reeves, 1992; Yaneva et al., 1986). However, until recently there is a paucity of experimental data on the in vivo function of Ku. With the finding that Ku80 complements the DNA double-strand break rejoining deficiency and V (D)J recombination defect in radiation-sensitive CHO mutant cell lines, a role for Ku in the repair of DNA double strand breaks and V(D)J rearrangement is now established (Smider et al., 1994, Taccioli et al., 1994). In another report, the 460-kDa catalytic subunit of DNA-PK was linked to the V(D)J recombination and DNA repair machinery,

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and shown to be the product of the murine gene SCID (Kirchgessner et al., 1995). To test whether the Ku-protein plays a role in the heat shock response, the intracellular concentration of Ku was modulated through the use of retroviral-mediated gene transfer technique. Coexpression of human Ku70 and Ku80 or expression of only the Ku70 subunit in rodent cells specifically inhibits the induction of Hsp70 upon heat shock (Li et al., 1995; Yang et al., 1996) (Figure 4). On the other hand, expression of human Ku80 alone does not have this effect. Thermal induction of other heat shock proteins in all of the Ku-overexpressing cell lines appears not to be significantly affected (Figure 4), nor the state of phosphorylation or the DNA-binding ability of HSF1 (Figure 5) (Yang et al., 1996). While Ku80 is involved in DNA repair, the 70-kDa component of the mammalian Ku-protein, but not the Ku80 subunit, appears to be involved in the regulation of Hsp70 gene expression. Thus, different subunits of Ku or different Ku-containing complexes may be involved in distinct biological processes. The fact that Ku is involved in gene regulation is not without precedent. Both the 70and 86-kDa subunits of Ku contain leucine/serine repeats which are reminiscent of similar “zipper” motifs found in a variety of transcription factors. Furthermore, Ku is a component of DNA-PK which phosphorylates several transcription factors, such as SP1 (Gottlieb and Jackson 1993), c-Jun (Bannister et al., 1993), p53 (Lees-Miller et al., 1990), c-Myc, Oct-1, and Oct-2 (Lees-Miller and Anderson 1991) in vitro. Studies also indicate that the Ku/DNA-PKcs complex modulates RNA polymerase I mediated transcription (Hoff et al., 1993; Knuth et al., 1990; Kuhn et al., 1995; Kuhn et al., 1993). Additionally, Ku has been found to be localized on certain transcriptionally active loci of chromosomal DNA (Amabis et al., 1990; Reeves, 1992). These data support a role for DNA-PK/Ku in transcription; specific genes regulated by DNA-PK/Ku need yet to be identified.

5. REQUIREMENT FOR KU80 IN GROWTH AND V(D)J RECOMBINATION It is well established that DNA double-strand break repair is important in maintaining cells’ genomic integrity. To determine the role of Ku80 in vivo, Ku80 was targeted in mice (Nussenzweig et al., 1996). Ku80 was inactivated by deleting 3.4 kilobases of the mouse Ku80 locus, including 100 base pairs of the promoter and the first two exons. Independently targeted embryonic stem (ES) cell clones were used to generate Ku80mutant mice. Heterozygous Ku80+/- mice were indistinguishable from their wild type littermates in all aspects, and 25% of the offspring born from Ku80+/- x Ku80+/- crosses were Ku80-/-. In contrast, new born Ku80-/- mice were significantly smaller than their wild type and heterozygous littermates (Figure 6). The growth retardation, as determined by weight difference, was first noted at embryonic day 15.5 (e15.5), and became statistically significant at e17.5. This difference increased to 40% at birth. During a 6month observation period, Ku80- /- mice grew and maintained body weights at 40–60% of controls, remaining proportional dwarfs. The reduced body weight in Ku80-/- mice does not appear to be a consequence of cell size, since comparisons of cell dimensions in kidney, muscle and liver sections showed no significant differences. Consistent with this

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growth defect, fibroblasts derived from Ku80-/- embryos showed an early loss of proliferating cells, a longer doubling time, and intact cell-cycle checkpoints that prevented

Figure 4 (left) Overexpression of Ku70 or Ku70 and Ku80 jointly specifically

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suppresses heat-induced Hsp70 expression. (A) Northern analysis of Hsp70 mRNA during post-heat-shock recovery at 37°C. Rat-1 cells, R7080– 6 cells (overexpressing human Ku-70 and Ku-80 protein jointly), and R70–15 cells (overexpressing human Ku-70 protein), were exposed to 45°C for 15 min, and returned to 37°C for 2–10 hr. Equal amounts of total RNA (10 mg). were loaded per lane, size fractionated on an agarose gel, transferred to Hybond-N membrane (Amersham), and probed with the human Hsp70 gene. C: no heat shock; 2, 4, 6, 8, 10, recovery time (hr). at 37°C. The heat-inducible Hsp70 mRNA is indicated by an arrowhead on the right margin. Note that the level of Hsp70 mRNA in heat-shocked R7080–6 and R70–15 cells is significantly reduced relative to that in Rat-1 cells. The same membrane was subsequently probed with a -actin gene. The levels of actin mRNA at all time points are not significantly different for all cell lines (see (B) below). The patterns of thermal induction of Hsp70 mRNA in R80–1 cells (overexpressing the human Ku-80 protein). are similar to the Rat-1 cells (data not presented). (B) Northern analysis of Hsc70, Hsp90, and Hsp27 mRNAs during post-heat-shock recovery. Rat-1 cells and R7080–6 cells were exposed to 45°C for 15 min, and returned to 37°C. Equal amounts of total RNA (10mg). were loaded per lane, size fractionated, and transferred to membranes as described in (A). The same membranes were probed sequentially with Hsp70, Hsc70, Hsp27, and Hsp90 and b-actin probes. C: no heat shock; H: heat-shocked. Total RNAs from control and heat-shocked cells (extracted 6–8 hr after heat shock), were shown. The levels of actin mRNA, used as a control, remain relatively constant after heat shock. The heat-induction of Hsc70 mRNA, Hsp27 mRNA and Hsp90 mRNA are clearly shown for both Rat-1 cells and R7080–6 cells. In contrast, the induction of Hsp70 mRNA in R7080–6 cells is significantly repressed.

cells with damaged DNA from entering the cell-cycle. This observation is in agreement with the correlation between senescence and defective DNA repair (Weirich-Schwaiger et al., 1994). Histologically, with the exception of lymphoid organs, Ku80−/− mice appear to be normal. However, spleen and lymph nodes were 5–10 fold smaller than controls and were devoid of lymphocytes. The thymus was also disproportionally small and had no corticalmedullary boundaries. Consistent with the histology, there was a complete absence of mature T and B cells. Both T and B lymphocyte development are arrested at early progenitor stages. Transient transfection experiments in mutant cell lines have shown that Ku80 is required for the formation of coding and signal joint during V(D)J recombination (Smider et al., 1994; Taccioli et al., 1994). To determine whether the KuSO−/− mutation affects rearrangements of antigen-receptor gene segments in T and B lymphocytes in vivo, immunoglobulin and T-cell antigen receptor (TCR) rearrangements were measured by polymerase chain reaction (PCR) (Carroll et al., 1991; Carroll et al., 1993; Costa et al., 1992; Schlissel et al., 1993). This analysis revealed that there is a profound deficiency in V(D)J rearrangement at either immunoglobulin or TCR loci in KuSO−/−

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mice.

Figure 5 Analysis of the HSF1-HSE binding activity and HSF1 phosphorylation in Ku-overexpressing cells. Monolayers of Rat-1 cells and R7080–6 cells were exposed to 45°C for 15 min, and returned to 37°C for 0–10 hr. Equal amounts of cell extracts were subjected to gel mobility shift analysis (upper panel), or immunoblot analysis (lower panel), using HSF1-specific antiserum. Upper panel: The position of HSF1-HSE binding complex is indicated by an arrow (HSF1). The CHBF/Ku binding activity is indicated by an arrowhead, and is much greater in cells overexpressing human Ku-protein. Lower panel: the level of phosphorylation of HSF1 during heat shock and subsequent recovery was examined by Western analysis with antiserum specific to HSF1, taking advantage of the known reduction in the electrophoretic mobility of HSF1 in SDS-polyacrylamide gel upon its phosphorylation (Sarge et al., 1993). The molecular sizes are indicated in kilodaltons. The heat-induced HSE binding activity and hyperphosphorylation of HSF1 are unaffected by the expression of the human Ku protein.

6. CONCLUSION Research from the past decade has made it clear that heat shock proteins are required for the development of thermotolerance. The fact that mammalian cells become permanently thermoresistant when transfected with Hsp70, and conversely, that cells become thermal

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sensitive when Hsp70 levels are reduced, directly demonstrates that Hsp70 plays a vital role in thermo tolerance. Hsp70 acts by stabilizing and preventing thermal denaturation of proteins, and by facilitating the dissociation of protein aggregates that are formed during conditions of stress. Other HSPs, such

Figure 6 Ku80 deficiency alters both pre- and post- natal growth (A) Photograph of 3 week old Ku80 −/− and Ku80 +/− mice (there is no significant difference in size between Ku80 +/− and Ku8 +/+ mice, not shown). (B) Growth of KuSO /− and Ku80 +/− embryos, as measured by weight. Pregnant females from KuSO +/− intercrosses were sacrificed at e13.5, e15.5, and e17.5 and 5–10 embryos of each genotype were weighed. The average weights of wild type (filled bars), heterozygous (hatched bars), and homozygous (white bars) embryos, and newborn mice are plotted. Error bars represent the standard deviation from the mean for each genotype. Statistical analysis using Student’s t test (p<0.05). showed significant differences in mean values between the three genotypes starting at e17.5, but not at earlier stages. (C) Post-natal growth of three female Ku80 +/− and three female Ku80−/− littermates. Weights of individual animals are plotted vs. age (in days). (D) Growth kinetics of primary fibroblasts. 105 passage two MEFs were plated in

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replicate 60 mm dishes and individual dishes trypsinized and counted every 24 hours as indicated. Doubling time was calculated to be 41 hours for Ku80 /− compared to 23 hours for Ku80 +/−, based on the exponential portion of the growth curve. Ku80 −/− MEF’s reached a four-fold lower saturation density than controls (2.66×104/cm2 for Ku80 /− compared to 1.1×105/cm2 for Ku80 +/−. Wild type and Ku80 +/− MEFs were indistinguishable (not shown). Data redrawn from Nussenzweig et al., (Nussenzweig et al., 1996)

as Hsp27, may also have thermal protective functions, and there may even be cooperative action among members of the heat shock protein family during thermotolerance development, as has been shown for the Hsp70 and Hsp60 chaperones during the folding of denatured proteins (Hendrick and Hartl, 1993). Studies both in vitro and in vivo have shown that chaperones have the ability to recognize structures that are exposed in unfolded proteins, protect polypeptides from aggregation or premature folding, and promote the refolding of proteins after denaturation. These folding and unfolding reactions are critical both during normal cellular biogenesis and during metabolic stress. Under various conditions of stress the increased need to repair denatured proteins and protect vital structures is fulfilled by the enhanced and preferential synthesis of chaperones. The importance of heat shock factor HSF1 in the regulation of mammalian heat shock gene expression is well-established. However, recent data indicate that activation of HSF1, by itself, is not sufficient for the induction of Hsp70mRNA synthesis. We have previously proposed a dual control mechanism for the regulation of Hsp70 gene expression in mammalian cells: a positive control mechanism mediated by HSF1, and a negative control mechanism mediated by CHEF. CHBF was found to be identical to the Ku-autoantigen. In rodent cell lines that stably and constitutively overexpress the human Ku subunits Ku70 and Ku80 jointly, or Ku 70 alone, the induction of Hsp70 upon heat shock is specifically suppressed (Li et al., 1995; Yang et al., 1996). Thermal induction of the other heat shock proteins appears not to be affected, nor is the state of phosphorylation or the DNA-binding ability of HSF1 affected. These data suggest a role for Ku in regulating Hsp70 expression. Accumulating experimental evidence implicates that Ku plays an important role in modulating cellular response to stresses, e.g., heat shock and ionizing radiation. Under normal growth conditions, Ku is linked to V(D)J recombination, T and B lymphocyte development and growth control. A challenge for future studies will be to elucidate how Ku is functionally involved in these processes.

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III. CELLULAR FUNCTIONS

6. GENETIC EVIDENCE FOR THE ROLES OF MOLECULAR CHAPERONES IN ESCHERICHIA COLI METABOLISM WILLIAM F.BURKHOLDER and MAX E.GOTTESMAN* Department of Biochemistry and Molecular Biophysics and Institute of Cancer Research, Columbia University, New York, NY 10032

1. Introduction 2. Genetic Analysis of Chaperone Function: The Case of dnaK 2.1. dnaK, dnaJ, and grpE Mutations Affect Gene Expression 2.2. E. coli Homologs of DnaK and DnaJ 2.3. Dominant dnaK Mutants and the Requirement for grpE 3. Genetic Evidence for the Roles of Chaperones in E. coli Metabolism 3.1. Protein Folding 3.2. Ribosomes and RNA Metabolism 3.3. Protein Export 3.4. Proteolysis 3.5. DNA Replication 3.6. Cell Division 3.7. Acquired Stress Tolerance and Survival during Starvation 4. The Orphaned Chaperones of E. coli: Homologs in Search of Function 4.1. HtpG (Hsp90) 4.2. IbpA and IbpB (Small Hsps) 5. Chaperones and Adaptation: Comparing Genomes 6. Acknowledgments 7. References *Corresponding author

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1. INTRODUCTION As the key regulated modulators of protein conformation in the cell, chaperones are likely to be involved in every aspect of cellular metabolism. A combination of genetic and biochemical analyses of chaperone function in E. coli has suggested possible roles for chaperones in a diverse range of cellular processes. Some of these roles are now fairly well understood on a mechanistic level, while others are still poorly explained. Efforts to unravel the involvement of chaperones in cellular metabolism are broadly motivated by three sets of goals: 1. to identify the cellular processes that chaperones participate in directly, the polypeptide substrates acted on, and the mechanisms of action of the chaperones involved, 2. to determine the ways in which chaperones are specialized, the extent to which their activities overlap, and how chaperones cooperate with each other, and 3. to characterize the regulatory effects of chaperones and any roles they may have in coordinating adaptive responses of the cell to changing environmental conditions. Genetic approaches can be very useful for revealing cellular processes influenced by chaperones and interactions between chaperones and their substrates. However, the road from a mutant phenotype to its underlying mechanism can be long and difficult: 1. A mutant phenotype may result from the disruption of a single cellular process, or from the simultaneous disruption of many processes. If several processes are affected, it may be difficult to identify specific mechanisms underlying the phenotype. 2. A chaperone mutation may affect a cellular process indirectly by altering gene regulation or protein expression. Most notable is the induction of heat shock proteins (hsps), many of which are chaperones or proteases, by mutations in dnaK, dnaJ, or grpE. 3. The role of a chaperone in a particular cellular process may go undetected because of the redundant or overlapping activity of other chaperones. 4. On the other hand, chaperones that appear to have redundant activities may be specialized in vivo due to different patterns of expression, interactions with different cofactors, or distinct substrate specificities. 5. Mutation or loss of one member of a chaperone complex may result in altered interactions of the complex with substrates, not just loss of complex function. Such mutations may therefore have a dominant phenotype. Section 2 of this review focuses on dnaK to illustrate the issues of regulatory effects of chaperone mutations, redundancy, and dominance, since these are particularly well understood in the case of dnaK These issues will be returned to with groEL in section 3.1 as we move on to survey the ways in which chaperones may participate in cellular metabolism. This review only covers the cytoplasmic chaperones of E. coli. Periplasmic chaperones and the specialized chaperones of pilus assembly are reviewed by Missiakis and Raina, and by Hultgren, et al. in this volume. Many chaperones, including DnaK and GroEL, function as dynamic, multimeric complexes with their cochaperones. The biochemistry of these complexes is reviewed in several chapters in this volume, by

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Ranson and Clarke, Burston and Saibil, and Buchberger et al. For other recent reviews of the genetics of molecular chaperones in E. coli see Georgopoulos et al. (1994) and Gross (1996). Useful earlier reviews are cited therein.

2. GENETIC ANALYSIS OF CHAPERONE FUNCTION: THE CASE OF DNAK 2.1. dnaK, dnaJ, and grpE Mutations Affect Gene Expression A central role of DnaK and its cochaperones DnaJ and GrpE is to down-regulate the dependent heat shock response, , encoded by the rpoH gene, is one of the two principal a factors regulating heat shock gene expression in E. coli (reviewed by Connolly et al. in this volume; Bukau, 1993; Yura et al., 1996). -dependent gene expression is induced by a variety of stresses, including misfolded cytoplasmic proteins. The second factor, , is induced by extreme heat shock and misfolded proteins in the extracytoplasmic space, and may mediate the heat shock response for that compartment (reviewed by Connolly et al., and Missiakas and Raina, in this volume; Missiakas and Raina, 1997). Cellular levels of are limiting, and increasing the level of up-regulates hsp expression (Grossman et al., 1987; Straus et al., 1987; Tilly et al., 1989). Mutations in dnaK, dnaJ, or grpE elevate basal hsp levels and eliminate the down-regulation of hsp synthesis during recovery from heat shock (Tilly et al., 1983; Straus et al., 1990). Since dnaK, dnaJ and grpE are positively regulated by , they form part of a homeostatic negative-feedback loop that maintains hsp synthesis at levels sufficient to cope with the concentrations of unfolded proteins in the cytoplasm (Craig and Gross, 1991). The failure of dnaK, dnaJ, and grpE mutants to down-regulate hsp expression is harmful to cells. Null mutants of dnaK are inviable at 16° or 40°C and grow slowly and filament at 30°C (Pack and Walker, 1987; Bukau and Walker, 1989a; Bukau and Walker, 1989b; Kang and Craig, 1990; McCarty and Walker, 1994). Mutations in rpoH (sidB alleles) that reduce -dependent gene expression fully suppress the slow growth and filamentation phenotypes of a dnaK52 mutant at 30°C and partially suppress cold sensitivity (Bukau and Walker, 1990). Rapid growth at 16°C and growth at 42°C can be restored by additional uncharacterized suppressor mutations (Bukau and Walker, 1990; Kang and Craig, 1990). Although dispensable for growth, dnaK appears to be active in many cellular processes over a range of growth temperatures, and it is likely that loss of dnaK is compensated for by other chaperones. A redundant chaperone might possibly be induced by the sidB mutations. In addition to inducing hsp expression, mutations in dnaK up- or down-regulate the expression of many other cellular proteins (Spence et al., 1990). Hence, the slow growth of unsuppressed dnaK, dnaJ, or grpE mutants could reflect either the reduced activity of specific proteins required for normal growth or the harmful effects of induced proteins. Some proteins are degraded at faster rates in dnaK mutants. This is due to the induction of proteases regulated by and perhaps also to the loss of dnaK chaperone activity (Gottesman, 1996; see section 3.4).

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Transcriptional regulators other than are regulated by DnaK Consequently, a variety of adaptive cellular responses are impaired in dnaK mutants. DnaK is required for induction of the stationary phase-specific factor, (Muffler et al., 1997; Rockabrand et al., 1998; see section 3.7). DnaK both increases the rate of translation and decreases its rate of degradation. One of the positive regulators of the capsular polysaccharide biosynthetic (cps) genes, RcsA, aggregates in dnaK and dnaJ mutants (Jubete et al. 1996; Jubete, Maurizi, and Gottesman, manuscript in preparation). The tendency of RcsA to aggregate in the absence of DnaK may explain the decrease in cps gene expression that occurs with increasing temperature. Titration of DnaK and DnaJ by competing substrates at higher temperatures may lead to the formation of inactive RcsA aggregates. As with the cps genes, flagellar gene expression is reduced in dnaK, dnaJ, and grpE mutants. The effect is due to reduced expression of the positive regulators (encoded by fliA) and FhlD/C, resulting in a nonmotile phenotype (Shi et al., 1992). Gene expression in dnaK mutants may also be affected by competition between and other subunits for RNA polymerase, although this does not appear to be the case with (Muffler et al., 1997). 2.2. E. coli Homologs of DnaK and DnaJ Though DnaK, DnaJ, and GrpE act together as a complex, the phenotypes of null mutations in their corresponding genes differ in severity. grpE null mutants are inviable at all temperatures in an otherwise wild-type strain background (Ang and Georgopoulos, 1989). dnaK null mutants are inviable below 16°C and above 40°C (Bukau and Walker, 1989a; Kang and Craig, 1990; McCarty and Walker, 1994). At 30°C, dnaK mutants grow slowly, have poor viability, frequently form long filaments, and are defective in chromosome segregation (Paek and Walker, 1987; Bukau and Walker, 1989a; Bukau and Walker, 1989b; Kang and Craig, 1990). dnaJ null mutants have the mildest phenotypes of the three. They are not cold sensitive and are inviable only at 43°C and above (Sell et al., 1990). Like dnaK mutants, dnaJ mutants grow slowly and filament at 30°C, though less markedly than dnaK mutants. In the absence of GrpE, DnaK and DnaJ may be toxic to cells (see section 2.3), and this may explain why grpE is an essential gene. On the other hand, dnaJ mutants may have less severe phenotypes than dnaK mutants because of the activity of a redundant dnaJ/homolog, cbpA. The phenotypes of cbpA dnaJ double mutants are similar to those of dnaK mutants (Ueguchi et al., 1994; Ueguchi et al., 1995; Wegrzyn et al., 1996). A cbpA null mutation confers no apparent phenotype in a dnaJ + strain. Overexpression of CbpA suppresses the phenotypes of dnaJ null and missense mutants. Two lines of genetic evidence suggest that CbpA replaces DnaJ in the complex with DnaK rather than acting in a parallel bypass pathway: (1) overexpression of CbpA does not rescue the growth defects of dnaK, grpE, groEL, groES, or rpoH missense mutants, and, (2) the phenotype of a dnaK mutant is not exacerbated by a second mutation in cbpA (Ueguchi et al., 1994). CbpA and DnaK have not yet been shown to interact functionally in vitro. Expression of cbpA is regulated by the stationary phase-specific sigma factor and is induced when cells enter stationary phase or during phosphate limitation (Yamashino et al., 1994; see section 3.7). Unlike dnaJ, cbpA is not induced by heat shock. CbpA is 39%

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identical to DnaJ but lacks the DnaJ cysteine rich region (Ueguchi et al., 1994). This region appears to be required for interactions of DnaJ with some, but not all, polypeptide substrates, and for promoting the formation of stable DnaK-substrate complexes (Szabo et al., 1996; Banecki et al., 1996). In addition to cbpA, E. coli has two other dnaJ homologs, djlA and hscB (yfhE) (Clarke et al., 1996; Seaton and Vickery, 1994). DjlA and the protein encoded by hscB, Hsc20, are similar to DnaJ only in the conserved J-domain, which is required for the binding of DnaJ homologs to the specific hsp70s with which they function (Wall et al., 1994; Schlenstedt et al., 1995). DjlA is an integral membrane protein, with the J-domain at its cytoplasmic C-terminus (Clarke et al., 1996). DjlA thus belongs to a growing family of Jdomain-containing integral and membrane-associated proteins that are known or presumed to target hsp70s for membrane localized functions (Cyr et al., 1994). Overexpression of DjlA induces expression of the capsular polysaccharide biosynthetic (cps) genes (Kelley and Georgopoulos, 1997; Clarke et al., 1997). This phenotype requires specific sequences in the transmembrane domain of DjlA, a functional ‘Jdomain’, DnaK, and GrpE. The phenotype may thus result from DjlA acting as a cochaperone with DnaK and GrpE on a component of the signaling pathway regulating cps expression, such as the integral membrane sensor kinase RcsC. In this model, the transmembrane domain of DjlA may participate in substrate recognition (Clarke et al., 1997). Deletion of djlA has a modest effect on the timing of cps gene expression (Kelley and Georgopoulos, 1997). E. coli also has a second dnaK homolog, Hsc66, that is 42% identical to DnaK (Seaton and Vickery, 1994). The gene encoding Hsc66, hscA, is in the same operon as the dnaJ homolog hscB, encoding Hsc20. Hsc66 and Hsc20 appear to functionally interact in a manner analogous to DnaK and DnaJ (Vickery et al., 1997). A possible target of Hsc66 action is the E. coli nucleoid-binding protein H-NS (Kawula and Lelivelt, 1994), which modulates the expression of other environmentally regulated genes, including the stationary sigma factor, (Barth et al., 1995). Hsc66 may antagonize H-NS activity, since inactivation of hscA suppresses a putative hns promoter-down mutation, but has no effect on a hns mutant (Kawula and Lelivelt, 1994). The pattern of protein expression following cold shock is altered in hscA null mutants (Lelivelt and Kawula, 1995). This altered expression pattern maybe related to H-NS function, since H-NS is induced by cold shock. 2.3. Dominant dnaK Mutants and the Requirement for grpE The analysis of dominant negative dnaK mutants has helped clarify the basis of dnaK mutant phenotypes and provided insights into the mechanism of dnaK action. Of the dnaK mutations selected as failing to down-regulate heat shock gene expression, roughly 20% are dominant (Wild et al., 1992b). The two dnaK mutants exhibiting the strongest dominant phenotypes, GD229 and GD341, encode proteins defective in ATP-binding (Wild et al., 1992b; mutations are designated by single letter amino acid code and residue number). ATP binding is required for release of both bound substrate and GrpE from DnaK (Palleros et al., 1993; Zylicz et al., 1987) and is stimulated by GrpE-catalyzed nucleotide exchange (Liberek et al., 1991). This suggests two complementary

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mechanisms of dominance: 1. The DnaK mutants bind polypeptide substrates but are slow to dissociate, thus blocking the spontaneous or chaperone-assisted folding or function of these substrates. Dominant DnaK mutations may therefore interfere with cellular processes unaffected by DnaK loss-of-function mutations (see section 3.3). 2. The mutant proteins are defective in releasing bound GrpE, and thus titrate GrpE away from wild-type DnaK. Without GrpE-catalyzed nucleotide exchange, wild-type DnaK would also be slow to dissociate from substrates. Consistent with the suggestion that defects in substrate release can result in a dominant negative phenotype, the dominant EK171 mutant is defective in the ATP-dependent release of peptide substrates (Buchberger et al., 1994). Similarly, a C-terminal fragment of DnaK (CTF) containing only the substrate binding domain is toxic when overexpressed in wild-type cells (Burkholder et al., 1996; see Buchberger et al., in this volume, for a description of the domain structure of DnaK). CTF binds and releases substrates in a manner similar to ADP-bound full-length DnaK (Gragerov et al., 1994). Mutations in the CTF that reduce substrate binding relieve toxicity (Burkholder et al., 1996). DnaK mutants may also inhibit cell growth by titrating GrpE, as suggested by mutants of an N-terminal fragment (NTF) of DnaK comprising only the ATP binding domain (Burkholder and Gottesman, unpublished). Three NTF mutants were isolated that were defective in the ATP-dependent release of GrpE, GE228, GS229 and GD341. The mutants are toxic to cells when overexpressed, but coexpression of GrpE suppressed the lethality of GE228. This suggests that the mutants inhibit E. coli growth by titrating GrpE. The mutations carried by these NTFs are similar or identical to previously identified dominant mutants of full-length DnaK (Wild et al., 1992b). The full-length DnaK mutants EK171 and DN201 are also defective in ATP-dependent release of GrpE (Kamath-Loeb et al., 1995). The GD229 and EK171 dominant negative mutants of DnaK impair cellular growth at 30°C much more severely when coexpressed with wild-type DnaK than when expressed by themselves. GD229 has the stronger synthetic phenotype. Expression of GD229 from a single chromosomal copy is lethal to duaK + merodiploids (Wild et al., 1992b), whereas EK171 only retards growth when overexpressed from a multicopy plasmid (Buchberger et al., 1994). Neither of the two models of dominance outlined above explain why the toxicity of GD229 and EK171 requires wild-type dnaK. It is possible that the mutant proteins titrate GrpE but are not lethal because they fail to form stable complexes with polypeptide substrates. Complex formation between DnaK and polypeptide substrates is stimulated by ATP in the presence of DnaJ (cf. Wawrzynów et al., 1995a). GD229 and EK171 have been shown to be defective in either ATP-binding or ATP hydrolysis stimulated by substrates and DnaJ (Wild et al., 1992b; Buchberger et al., 1994; KamathLoeb et al., 1995). These mutants may, therefore, be unable to form stable ATPdependent complexes. EK171 and DN201 have wild-type affinities for peptide substrates in the absence of ATP, but these interactions may be qualitatively different than forming stable ATP- and DnaJ-dependent complexes. Extragenic suppressor mutations that permit the deletion of grpE in a dnaK + strain

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have been reported (Ang and Georgopoulos, 1989). grpE can also be deleted in cells expressing DnaK mutants that appear to bind substrates less stably. One such mutant, DnaK332, has an increased basal rate of nucleotide exchange. A carboxyterminal truncation of DnaK also permits deletion of grpE (described in Georgopoulos et al., 1994). The effect of the truncation may be to remove the lid that locks bound substrate into ADP-bound DnaK while preserving the substrate binding cleft itself (see Zhu et al., 1996). These suppressor mutations of dnaK lend further support to the model that GrpE is essential to dissociate stable complexes formed between DnaK and substrate polypeptides.

3. GENETIC EVIDENCE FOR THE ROLES OF CHAPERONES IN E. COLI METABOLISM 3.1. Protein Folding 3.1.1. Cooperative Action of the DnaK and GroEL Complexes In this section, we focus on genetic evidence for the involvement of chaperones in protein folding in E. coli. Other evidence for the role of chaperones in protein folding is reviewed extensively in other chapters of this volume. The cellular requirement for chaperone function increases with increasing temperature. This is due in part to the increased rates of growth and protein synthesis that occur at higher temperatures and also to the temperature sensitivity of the folding intermediates of many proteins (for a recent review, see King et al., 1996). Cells lacking rpoH are unable to induce hsp expression and cannot grow above 20°C (Zhou et al., 1988). Shifting to nonpermissive temperature does not immediately halt protein synthesis in rpoH mutants, but most newly synthesized polypeptides aggregate and form cytoplasmic inclusion bodies (Gragerov et al., 1991; Kucharczyk et al., 1991; Gragerov et al., 1992). Overexpression of either GroEL/ES or DnaK/ J raises the permissive temperature of rpoH null mutants and suppresses the aggregation of newly synthesized polypeptides (Kusukawa and Yura, 1988; Gragerov et al., 1992). However, the defects in growth and protein aggregation can be suppressed with much lower levels of these chaperones if both pairs are coexpressed. Similarly, dnaK756 groEL100 or dnaK756 groES30 double mutants display extensive aggregation at 42°C whereas single mutants do not (Gragerov et al., 1992). Thus, aggregation is more efficiently suppressed by the cooperative action of GroEL/ES and DnaK/J. How do GroEL/ES and DnaK/J function cooperatively? NMR studies indicate that DnaK binds unfolded peptide substrates, whereas GroEL binds substrates that have some secondary structure (Landry et al., 1992). These results are supported by two lines of evidence. First, the crystal structure of the substrate binding domain of DnaK shows that substrate polypeptides must contain fully unfolded regions to be encapsidated in the DnaK binding cleft (Zhu et al., 1996; see Ha et al., and Buchberger et al., this volume). Second, biochemical analyses reveal interactions of GroEL with partially folded and even fully folded substrate proteins (see Burston and Saibil, this volume). These data, together

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with studies of protein folding in mitochondria and in reconstituted in vitro reactions, suggest that DnaK/J and GroEL/ES act sequentially in the folding pathway. In this model, DnaK and DnaJ bind nascent unfolded polypeptides emerging from the ribosome to block premature folding and aggregation, and perhaps facilitate polypeptide translocation (see Nelson et al., 1992; Welch et al., this volume). Then full-length unfolded or partially folded polypeptides are transferred to GroEL/ES where they fold to the native state in one or more cycles of binding and release. Consistent with a sequential folding pathway, DnaK and DnaJ, but not GroEL, are found to bind ribosome-associated nascent polypeptides (Gaitanaris et al., 1994). Further, pulse-chase experiments indicate that newly synthesized full-length CI associates transiently with GroEL/ES before entering the soluble fraction (Gaitanaris et al., 1994). The vectorial model has its limitations, however. If DnaK and GroEL work at separate stages of folding, how is it that each can substitute for the other to suppress aggregation when overexpressed? Along the same lines, why is a dnaK strain viable at 30°C? It is possible that other chaperones may substitute for DnaK or GroEL at either early or late stages of protein folding. It is also possible, however, that the pathway of chaperoneassisted folding is more dynamic and flexible, with substrates passing back and forth between different chaperones until properly folded or degraded (see, for example Buchberger et al., 1996; Burston et al., 1996; reviewed in Bukau et al., 1996; Gragerov, 1997). Although no global aggregation of mature proteins is seen in rpoH mutants at nonpermissive temperature (Gragerov et al., 1991), temperature sensitive proteins may still require chaperones to prevent aggregation at high temperature and to refold correctly when returned to permissive temperature. Thus, firefly luciferase heterologously expressed in E. coli is inactivated when cells are shifted to high temperature. Luciferase activity is restored in the absence of new protein synthesis upon returning cells to permissive temperature (Schröder et al., 1993). Activity is not recovered in dnaK, dnaJ, or grpE mutants, although wild-type levels of the luciferase protein are still present. The chaperone mutants are, therefore, defective in protecting luciferase from irreversible heat inactivation. In vitro, DnaK, DnaJ and GrpE are required to refold denatured luciferase at 30°C (Schröder et al., 1993; Szabo et al., 1994). Whereas denatured full-length luciferase requires DnaK for refolding in vitro and in vivo, newly synthesized luciferase folds readily into active form at 30°C in a dnaK52 strain (Hesterkamp and Bukau, personal communication). The nascent polypeptide folding pathway for luciferase thus appears to be distinct from the pathway stabilizing and refolding the heat-inactivated protein. 3.1.2. A Central Role for GroEL/ES groEL and groES are essential for growth at all temperatures (Fayet et al., 1989). Consistent with the homeostatic model of hsp regulation (Craig and Gross, 1991; Bukau, 1993), GroEL/ES is expressed at levels just sufficient to meet the needs of the cell. Reducing GroEL/ES expression to 25% of normal levels slows cell growth and induces -dependent hsps at 37°C (Kanemori et al., 1994). Without sufficient GroEL/ES hsp expression is presumably stimulated by accumulated unfolded polypeptides. When strains carrying the groEL201 (EK461) mutation are shifted to 37°C, the concentrations of

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roughly 50% of the newly synthesized soluble proteins are reduced (Horwich et al., 1993). Many of these proteins aggregate, and the rest may become susceptible to proteolysis. Other proteins appear to be inactivated while remaining stable and soluble. Unlike rpoH mutants, massive aggregation of nascent polypeptides and formation of inclusion bodies is not seen in the groEL201 mutant. The protein species affected by groEL201 may indicate which proteins interact with GroEL to fold at 37°C. Roughly 50% of E. coli cytoplasmic proteins, when diluted from denaturant, bind GroEL (Viitanen et al., 1992; Horwich et al., 1993). The ability of several proteins to interact with GroEL in vitro correlates with their defective folding in groEL201 in vivo (Horwich et al., 1993). However, estimates of the proportion of nascent polypeptides that GroEL could interact with during growth at 37°C, given the abundance of GroEL, the rate of protein synthesis, and the rates of substrate binding and release by GroEL in vitro, suggest that GroEL may only be involved in the folding of 5–30% of cellular polypeptides (Lorimer, 1996). Efforts to identify particular in vivo substrates of GroEL/ES are discussed in the following sections. The first identified substrates of GroEL/ES are the capsid head proteins of bacteriophages and T4 and the tail of bacteriophage T5 (for review, see Georgopoulos et al., 1990; Zeilstra-Ryalls et al., 1991; Georgopoulos et al., 1994). groEL and groES mutants are resistant to infection, growth in the mutants is blocked at an early step of phage head morphogenesis, at the point waen the B protein assembles into the homomeric preconnector ring. Assembly of the bacteriophage T4 head and bacteriophage T5 tail are also blocked at early stages. GroEL appears to play a similar role in the assembly of certain endogenous protein complexes. Mutations in groEL and groES block the anaerobic processing and assembly of one of the three NiFe hydrogenases in E. coli, HYD3, at a step sensitive to the availability of Ni2+ (Rodrigue et al., 1996). The mutations also interfere with the activity of a second NiFe hydrogenease. Ni2+ appears to be incorporated into HYD3 during folding, since it is buried in the native protein structure of a related hydrogenase. GroEL may stabilize HYD3 until Ni2+ is bound and folding can proceed. GroEL binds the HYD3 precursor in vitro, consistent with a direct role in HYD3 processing and assembly. In what is probably a related reaction, precursors of the MoFe nitrogenase complex of Klebsiella pneumoniae transiently associate with GroEL during assembly in vivo (Govezensky et al., 1991). E. coli GroEL facilitates assembly of the complex when it is expressed in E. coli or when E. coli GroEL is overexpressed in Klebsiella pneumoniae. The requirement of chaperones for the assembly of cofactor-dependent complexes may turn out to be a general rule. Interestingly, the assembly of a molybdenum-dependent nitrate reductase complex in E. coli appears to depend in part on a dedicated chaperone, NarJ (Liu and DeMoss, 1997; Blasco et al., 1998). GroEL may play a direct role in gene regulation by promoting the folding of transcriptional activators. GroEL is required in vivo for activation of both the NodD and the NifA transcriptional activators from Rhizobium meliloti and Klebsiella pneumoniae, respectively. It physically interacts with both in vitro, supporting a direct role for the chaperone in their activation (Ogawa and Long, 1995; Govezensky et al., 1991; Govezensky et al., 1994). Since groEL and groES mutations are pleiotropic, it is difficult to identify specific

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substrates of GroEL/ES that might be affected by the mutations. The problem has been approached genetically by identifying allele-specific interactions between mutations in groEL or groES and the gene encoding a putative substrate. For instance, single-stranded DNA binding protein (SSB) was identified as a possible target for GroEL when groEL411 was isolated as an allele-specific, dominant suppressor of the ssb-1 mutant allele (Ruben et al., 1988). Another ssb mutation, ssb-13, is specifically suppressed by the groEL46 mutation (Laine and Meyer, 1992). Importantly, neither ssb mutation is suppressed by overexpressing wild-type GroEL. Allele-specific interactions and suppressor analysis have also been used to define sites of interaction between GroEL, GroES, and substrates such as DnaA46 (Zeilstra-Ryalls et al., 1993; Zeilstra-Ryalls et al., 1994; Zeilstra-Ryalls et al., 1996). The systematic structure/function analyses of GroEL/ES in vitro have made rapid progress on the same questions (Fenton et al., 1994; see Burston and Saibil, this volume). A change in GroEL substrate specificity occurs following infection with bacteriophage T4. Phage T4 encodes a functional homolog of GroES, gp31, which interacts with GroEL and can replace GroES for many functions in E. coli (van der Vies et al., 1994; for a review of early work, see Zeilstra-Ryalls et al., 1991). gp31 and GroEL are essential for T4 growth, whereas GroES is dispensable. This suggests that the GroEL-gp31 complex uniquely supports the folding of the T4 substrate polypeptide, the capsid head protein gp23, while still retaining activity towards endogenous bacterial substrates. The threedimensional structure of gp31 is similar to that of GroES, but gp31 complexed to GroEL forms a larger and more hydrophilic cavity encapsulating protein folding intermediates than is formed by the GroEL-GroES complex (Hunt et al., 1997). 3.2. Ribosomes and RNA Metabolism A major E. coli protein associated with nascent polypeptide chains is trigger factor. Trigger factor is a ribosome-associated peptidyl prolyl cis-trans isomerase that can be cross-linked to nascent chains of both cytosolic and secreted polypeptides (Lill et al., 1988; Stoller et al., 1995; Valent et al., 1995; Hesterkamp et al., 1996; reviewed in Hesterkamp and Bukau, 1996; see Fischer and Schmid, this volume). Trigger factor stabilizes the outer membrane precursor protein, proOmpA, in a translocation competent state in vitro, suggesting that it can act as a chaperone (Crooke and Wickner, 1987; Crooke et al., 1988). However, depletion of trigger factor in vivo has no effect on proOmpA export, even in a secB null mutant (section 3.3). Trigger factor also associates with GroEL and stimulates its binding to denatured proteins, the unstable fusion protein CRAG, and other model polypeptide substrates (Kandror et al., 1995; Kandror et al., 1997; see section 3.4). Trigger factor is induced by cold shock and appears to increase cell viability at low temperatures (Kandror and Goldberg, 1997). Underexpression or overexpression of trigger factor induces cell filamentation (Guthrie and Wickner, 1990). It has not been demonstrated that the peptidyl prolyl isomerase and chaperone-like activities of trigger factor are related. However, isomerization of prolyl peptide bonds is a significant rate-limiting step for the folding of some proteins (reviewed in Galat and Metcalfe, 1995; Schmid, 1995). In addition to trigger factor, a growing list of cytosolic and periplasmic peptidyl prolyl isomerases has been identified in E. coli. These include a

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cytosolic FK506-binding protein homolog, SlyD, mutants of which affect the growth of E. coli and the function of a phage encoded lysis protein (Roof et al., 1996; see Missiakas and Raina, this volume, for a review of the role of periplasmic peptidyl prolyl isomerases). A role for hsps in ribosome function is suggested by the activation of the ribosomeassociated enzyme RelA by dnaK and dna/mutations at nonpermissive temperature. RelA is a guanosine tetraphosphate (ppGpp) synthetase that is activated when the proportion of charged tRNAs is growth-limiting. High levels of ppGpp block stable RNA synthesis via the stringent response pathway (for a recent review, see Cashel et al., 1996). Heat shock raises ppGpp concentrations in dnaK and dnaJ mutants (Tanaka et al., 1985; Itikawa et al., 1986). Correspondingly, when dnaK, dnaJ, or grpE mutants are shifted to nonpermissive temperature, RNA synthesis rapidly shuts off while DNA and protein synthesis continue for some time (Itikawa and Ryu, 1979; Wada et al., 1982; Ang et al., 1986). The accumulation of ppGpp and the rapid inhibition of RNA synthesis are relAdependent (Tanaka et al., 1985; Itikawa et al., 1986). A transient burst of ppGpp synthesis also occurs in some, but not all, wild-type control strains following heat shock. In these cells, however, ppGpp concentrations decline to new basal levels with kinetics similar to the shutoff of hsp induction. It is unclear why ppGpp concentrations increase during heat shock and why dnaK and dnaJ mutants are defective in down-regulating ppGpp synthesis following heat shock. Charged tRNA levels may become limiting after temperature upshift as a result of an increased rate of polypeptide synthesis. Alternatively, an aminoacyl-tRNA synthetic pathway could be inhibited by heat shock. For instance, the metA gene product, homoserine transsuccinylase, an essential enzyme in the methionine biosynthesis pathway, is unstable at 42°C (reviewed in Ron et al., 1990). Phosphorylation of the glutaminyl- and threonyl-tRNA synthetases is reduced in dnaK and dnaJ mutants (Wada et al., 1986), although it has not been proven that the activity of the enzymes is affected by phosphorylation. Ribosome function might also be directly affected following heat shock. A role for hsp70s in ribosomal function in S. cerevisiae has been proposed (Nelson et al., 1992), and recent in vitro evidence suggests that chaperone-mediated folding of nascent polypeptides may be coupled to translation termination and polypeptide release in E. coli (Kudlicki et al., 1994). DnaK may participate in the assembly of 30S and 50S ribosomal subunits. Ribosomal precursors accumulate in dnaK missense mutants upon shift to nonpermissive temperature (Alix and Guérin, 1993). The accumulated precursors remain competent for assembly, and chase into mature ribosomes when the cells are returned to permissive temperature. The assembly defects of dnaK mutants are not suppressed by introducing rpoH or relA mutations, indicating that neither hsp overexpression nor induction of the stringent response is responsible for the defects. The pattern of precursor accumulation varies among different mutant alleles of dnaK, presumably reflecting differences in behavior of the mutant DnaK proteins. GroEL has been suggested to regulate ribosomal RNA processing and mRNA turnover. When a groEL mutant is shifted to nonpermissive temperature, the 9S precursor to ribosomal p5S RNA accumulates, consistent with reduced RNase E processing (Sohlberg et al., 1993). Conversely, overexpression of a fragment of GroEL suppresses a

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temperature sensitive ams (RNase E) mutation (Chanda et al., 1985). GroEL copurifies with RNase E and binds a mutant RNase E protein, but direct binding to wild-type RNase E has not been observed (Sohlberg et al., 1993; Miczak et al., 1996). GroEL also forms stable complexes with polynucleotide phosphorylase, which itself associates with RNase E, suggesting that GroEL may be a component of protein complexes that process and degrade RNA (Ybarra and Horowitz, 1996). Recent work implies that a form of GroEL may possess an opposing activity, binding RNA and protecting it from degradation (Georgellis et al., 1995; Georgellis, Vytvytska, Täuber, Rezazadek, Brandsch, Hartl, and von Gabain, manuscript in preparation). A fraction of GroEL copurifies with an RNA binding activity that inhibits RNase digestion of 9S RNA in vitro. The binding activity is induced by anaerobic growth or phosphate limitation, conditions under which OmpA mRNA is stabilized in vivo. An unphosphorylated monomeric form of GroEL also predominates under these conditions, and may correspond to the RNA binding activity. These data suggest that GroEL may not only play a role in protein folding, but also in mRNA stability and in the regulation of translation. 3.3. Protein Export Most proteins destined for the periplasmic space or the outer membrane are translocated across the inner membrane by a machinery encoded by the sec genes (for reviews, see Pugsley, 1993; den Blaauwen and Driessen, 1996; Murphy and Beckwith, 1996). Once in the periplasmic space, other general or specialized chaperone activities are required for protein folding and transport (see Missiakis and Raina, this volume). The insertion of proteins into the inner membrane is less well understood, and the role of the sec-encoded export machinery in this process is not defined. Though some cotranslational protein export occurs in E. coli, most proteins are exported posttranslationally. These proteins are presumably maintained in a transport-competent unfolded or loosely folded conformation. SecB is a cytosolic chaperone that binds proteins destined for export and prevents their folding, aggregation, or degradation (for recent reviews, see Collier, 1993; Randall and Hardy, 1995; Welch et al., this volume). In addition, SecB may facilitate the targeting of precursor proteins to the translocation apparatus by binding to SecA, a cytosolic factor that cooperates directly with the SecY inner membrane translocase during export. Null secB mutants are defective in exporting some, but not all, proteins. Thus, efficient targeting of SecB-independent proteins to the secretory apparatus either does not require a chaperone or utilizes a chaperone other than SecB. The chaperone function of SecB appears to be partially redundant with DnaK. secB null mutants fail to grow in rich medium below 42°C and are defective for protein export in minimal medium (Kumamoto and Beckwith, 1985). These phenotypes are suppressed by overexpression of (Altman et al., 1991). Overexpression of DnaK and DnaJ is sufficient to suppress the growth and export defects of a secB null mutant, whereas overexpression of DnaK or GroEL/ES alone does not (Altman et al., 1991; Wild et al., 1992a). Conversely, reducing hsp expression prevents the growth of a secB null mutant, suggesting that -dependent genes compensate for the absence of secB (Altman et al., 1991). DnaK is necessary for this compensation, since the growth of a secB null mutant is

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blocked by strongly defective dnaJ or dnaK mutations or by depletion of DnaK, DnaJ, or GrpE (Wild et al., 1992a; Wild et al., 1996). During depletion, the export of SecBindependent proteins is impaired and the export of SecB-dependent proteins is reduced below the levels seen in a secB single mutant. Since the export of SecB-independent proteins is only affected in the absence of both SecB and DnaK, the two chaperones appear to be fully redundant for exporting this class of proteins. However, one of the SecB-independent proteins affected by DnaK depletion, ribose binding protein (RBP), is not a substrate for SecB (reviewed in Collier, 1993). SecB is thought to recognize its substrates in part through kinetic partitioning, binding to slow folding substrates in preference to substrates that fold quickly (reviewed in Randall and Hardy, 1995). RBP folds rapidly in vitro, consistent with its not being a substrate of SecB. How, then, does RBP export become dependent on SecB in the absence of DnaK? If the folding of RBP is slower in vivo in the absence of DnaK, then depletion of DnaK may make RBP a suitable substrate for SecB. The export of slow-folding mutants of RBP is, in fact, SecB dependent (Teschke et al., 1991; Kim et al., 1992). Interactions between DnaK and specific exported proteins are suggested by the allelespecific effects of dnaK and dnaJ mutations on PhoA translocation (Wild et al., 1992a). Translocation of PhoA to the periplasmic space is SecB-dependent at 30°C but SecBindependent at higher temperatures. Two dominant mutations of dnaK, EK171 and GD229, and a dnaJ mutation, HQ33, inhibit PhoA export at 30°C without affecting export of several other SecB-dependent and SecB-independent proteins. The defect is not due to loss of functional DnaK, since depletion of DnaK, DnaJ, or GrpE does not inhibit PhoA export in secB + cells (Wild et al., 1996). The mutant proteins may instead interfere with export by binding irreversibly to PhoA precursors or some other factor specifically required for PhoA export (see section 2.3). Doubt is cast on this interpretation, however, by the finding that GrpE depletion, which globally stabilizes DnaK-substrate complexes, does not inhibit PhoA export. GroEL/ES may also participate in the export of particular proteins. Pre- -lactamase export is SecB-independent and is impaired in groEL or groES mutants (Kusukawa et al., 1989; Laminet et al., 1991). Consistent with a direct role for GroEL/ES in this reaction, GroEL interacts with pre- -lactamase in vitro to suppress aggregation, maintain translocation competence, or promote either folding or unfolding (Bochkareva et al., 1988; Laminet et al., 1990; Zahn and Plückthun, 1992). Pre- -lactamase export is also reduced by depleting DnaK, DnaJ, or GrpE in a secB null mutant, although not as dramatically as by groEL or groES mutations (Wild et al., 1992a; Wild et al., 1996). This suggests that GroEL/ES plays a primary role in pre- -lactamase export whereas DnaK and SecB play secondary roles. GroEL/ES also maintains newly translated lactose permease, an integral inner membrane protein, in a conformation competent for insertion into the cytosolic face of membrane vesicles in vitro (Bochkareva et al., 1996). Overexpression of GroEL/ES or DnaK/J facilitates the export of certain heterologously expressed proteins (see, for example, Phillips and Silhavy, 1990; Bergès et al., 1996). In general, however, chaperone overexpression has had only mixed success in promoting protein folding and secretion (for recent reviews, see Makrides, 1996; Thomas et al., 1997). Furthermore, suppression of protein export defects by increasing chaperone levels may be unrelated to chaperone function. Thus, the growth and export defects of several

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cold-sensitive secretory pathway mutants are suppressed by overexpressing a variety of proteins, including GroEL/ ES or even inactive truncation mutants of GroEL (Danese et al., 1995). Similarly, suppression of a secY24 mutant by overexpressing HtpG may be a nonspecific effect of reduced growth rate (Shirai et al., 1996). Slow growth and lower rates of protein synthesis, conferred by mutations or by protein synthesis inhibitors, also suppress sec mutants (Lee and Beckwith, 1986; Schatz and Beckwith, 1990). 3.4. Proteolysis Proteases, like chaperones, adapt cells to changing environmental conditions and protect them from stress. Proteases down-regulate unneeded or inappropriate cell functions during adaptation. Proteins induced by carbon starvation, for example, are degraded after starving cells are resupplemented with glucose (Damerau and St. John, 1993; for reviews, see Gottesman, 1996; Sherman and Goldberg, 1996). Several E. coli ATP-dependent proteases are induced by heat shock. They presumably protect the cell by degrading misfolded or damaged proteins. To be accessible to proteases, substrates must be soluble. By blocking aggregation and perhaps by promoting unfolding, chaperones aid in the presentation of substrates to proteases (reviewed by Maurizi et al., this volume). Chaperone-assisted proteolysis of a substrate may compete with aggregation and chaperone-mediated protein folding. This may account for the varied effects of chaperone mutations on the degradation of specific proteins in vivo. Mutations in dnaK, dnaJ, grpE, or groEL stabilize some rapidly degraded proteins, such as unfolded or misfolded polypeptides induced by puromycin or canavinine, the unstable regulatory factor RcsA, and the mutant protein PhoA61 (Keller and Simon, 1988; Straus et al., 1988; Sherman and Goldberg, 1992; Jubete et al. 1996). Mutations in dnaK, dnaJ, grpE, or groEL may increase the rate of proteolysis of other proteins, however, including a temperaturesensitive mutant of the CI represser, the C protein of bacteriophage Mu, and the E. coli UmuC protein (Keller and Simon, 1988; Sand et al., 1995; Donnelly and Walker, 1989). UmuC, which forms a heterodimer with UmuD’, may be stabilized by GroEL/ES until the heterodimer is formed (Donnelly and Walker, 1992). The UmuC-UmuD’ complex is required for the error-prone repair of DNA damage. groEL and groES mutants are defective in UV-induced mutagenesis and the reactivation of UV-damaged bacteriophage, perhaps as a result of lowered UmuC levels (Donnelly and Walker, 1989; Liu and Tessman, 1990). The association of unstable proteins with chaperones may increase their rates of degradation. Missense mutations of dnaK or grpE that stimulate the binding of PhoA61 to DnaK enhance PhoA61 degradation. Conversely, a loss-of-function mutation in dnaJ that decreases the amount of PhoA61 associated with DnaK reduces PhoA61 degradation (Sherman and Goldberg, 1992). Similarly, conditions that enhance the association of the unstable fusion protein CRAG with GroEL increase its rate of degradation by the protease ClpP (Kandror et al., 1994). The chaperone trigger factor facilitates the binding of CRAG to GroEL and stimulates CRAG degradation (Kandror et al., 1995). The importance of chaperone activities for proteolysis is underscored by the recent finding that the ATPase subunits of the ClpP protease, ClpA and ClpX, are chaperones in their own right (Wickner et al., 1994; Levchenko et al., 1995; Wawrzynów et al., 1995b;

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Kruklitis et al., 1996; reviewed in Schirmer et al., 1996; Wawrzynów et al., 1996; Gottesman et al., 1997; Maurizi et al., this volume). In vivo evidence for the ClpPindependent chaperone activity of ClpX is discussed in section 3.5. ClpA and ClpX, which are members of the hsp100 family of chaperones, were first identified as factors that target specific polypeptide substrates to the ClpP protease for degradation. The ATPase subunits are now thought to not only recognize and target substrates to the ClpP protease but also to actively unfold them for presen-tation to ClpP. Recognition of substrates by ClpA and ClpX is not sufficient for proteolysis, however, since a stable protein tagged with a sequence recognized and bound by ClpX is not degraded by ClpP (Laachouch et al., 1996). Another member of the E. coli hsp100 family, ClpB, solubilizes protein aggregates, which are then rapidly degraded (Inoue and Rechsteiner, 1994; Laskowska et al., 1996a). ClpB is the nearest E. coli homolog to the S. cerevisiae, chaperone Hsp104, which also solubilizes protein aggregates in vivo (Parsell et al., 1994). ClpB is not required to target solubilized proteins for degradation, since the soluble fraction of aggregation-prone proteins is degraded at the same rate in clpB and wild-type cells (Inoue and Rechsteiner, 1994). Mutants of clpB, the protease Lon, and the protease subunits ClpA and ClpP are all impaired in the rapid removal of protein aggregates that form transiently in heat shocked cells (Laskowska et al., 1996a). 3.5. DNA Replication Much of the insight into the mechanism of DnaK chaperone function has come from studying the role of DnaK in the initiation of DNA replication. DnaK is required for efficient initiation of replication by bacteriophage A, and plasmids P1 and F, and perhaps in the initiation of E. coli chromosome replication (for a recent review, see Chattoraj, 1995). E. coli strains carrying missense mutations in dnaK, dnaJ, and grpE were isolated originally as resistant to killing by bacteriophage (Georgopoulos and Herskowitz, 1971; Sunshine et al., 1977; Saito and Uchida, 1977). growth is blocked in these mutants at the stage of DNA replication initiation (Georgopoulos and Herskowitz, 1971; Georgopoulos, 1977; Sunshine et al., 1977; Saito and Uchida, 1977; Saito and Uchida, 1978). Specific mutations in the P protein, a bacteriophage-encoded replication initiation factor, restore growth on the mutant strains. DnaK, DnaJ, and GrpE proteins are required in vitro to remove P from the replication initiation complex assembled at the origin of replication (reviewed by Zylicz et al., this volume). Removal of P from the complex permits subsequent DNA unwinding by DnaB helicase, which is initially recruited to the origin by P. The P mutants that bypass the requirement for wild-type DnaK, DnaJ, and GrpE activities form less stable complexes with DnaB helicase than wild-type P (Konieczny and Marszalek, 1995). A similar role for the ClpX chaperone during bacteriophage replication has recently been shown. ClpX, which is required for bacteriophage Mu replication in vivo, modifies the conformation of the replication initiation complex in vitro to permit subsequent partial disassembly of the complex and initiation of replication (Mhammedi-Alaoui et al., 1994; Levchenko et al., 1995; Kruklitis et al., 1996). In addition to blocking replication, mutations in dnaK, dnaJ, and grpE reduce the

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efficiency of replication of plasmids carrying a minimal P1 origin of replication (Tilly and Yarmolinsky, 1989; Bukau and Walker, 1989b) and interfere with the autoregulation of the P1 replication initiation protein, RepA (Tilly et al., 1990). DnaK and DnaJ activate RepA for DNA binding by dissociating inactive RepA dimers into monomers (Wickner et al., 1991a; Wickner et al., 1991b; Wickner et al., 1992). RepA mutants with increased activity in dnaK, dnaJ, and grpE mutant hosts appear to form less stable dimers in vitro (Mukhopadhyay et al., 1994). It has been suggested that DnaK not only dissociates RepA into monomers but also activates misfolded monomeric RepA (DasGupta et al., 1993). DnaK, DnaJ, and GrpE are required for mini-F plasmid replication in vivo, as well (Ezaki et al., 1989; Kawasaki et al., 1990). DnaK disassembles dimers of the F plasmid replication factor RepE into active monomers (Ishiai et al., 1994), analogous to its action on RepA. Unlike RepA dimers, RepE dimers bind DNA, but recognize different sites than those bound by the monomer. Like the corresponding RepA mutants, DnaKindependent RepE mutants are defective in dimerization in vitro. When mutants of dnaK or dnaJ are shifted to nonpermissive temperature, DNA synthesis declines after a short lag, whereas protein synthesis continues for a longer time (Saito and Uchida, 1977; Saito and Uchida, 1978; Itikawa and Ryu, 1979; Wada et al., 1982; Sakakibara, 1988). It was on the basis of this replication phenotype that the genes were designated dna, grouping them together with other genes required for host DNA synthesis. DNA synthesis is also reduced relative to protein synthesis in a grpE280 mutant at nonpermissive temperature (Ang et al., 1986). Replication of the E. coli chromosome is initiated at a single origin, oriC, by the ATPbound form of the initiation factor, DnaA. When dnaK mutants are shifted to nonpermissive temperature, chromosome replication continues to completion and then ceases (Sakakibara, 1988). Upon return to permissive temperature, a burst of replication at oriC occurs, indicating that the mutants had been blocked at the stage of replication initiation (Sakakibara, 1988; Ohki and Smith, 1989). Consistent with this interpretation, DNA replication persists for several hours at nonpermissive temperature in a dnaK rnh double mutant, in which replication initiation is DnaA-independent. The rnh mutation does not suppress the temperature sensitivity of dnaK mutants, however (Sakakibara, 1988; Bukau and Walker, 1989b), indicating that other essential cellular functions are defective as well (see section 3.6). DnaK may only be required for DnaA-dependent replication at elevated temperatures, since at 30°C a naK52 rpoH mutant grows well (Bukau and Walker, 1990; see section 2.1) and an onC-dependent plasmid is able to replicate in a dnaK756 mutant (Malki et al., 1991). In vitro evidence supports a role for DnaK in DnaA-dependent replication. DnaA forms inactive aggregates containing phospholipids that can be dissociated into active form by incubation with DnaK or phospholipase A2 (Hwang et al., 1990). In vitro, DnaK stimulates replication initiation at oriC by wild-type or mutant DnaA and may protect DnaA from heat inactivation (Zylicz et al., 1985; Hwang and Kaguni, 1991; Malki et al., 1991; Hupp and Kaguni, 1993a; Hupp and Kaguni, 1993b). GroEL/ES may also play a role in E. coli DNA replication. When temperature sensitive groEL and groES mutants are shifted to nonpermissive temperature, DNA synthesis ceases earlier than protein synthesis (Wada and Itikawa, 1984). Genetic evidence is consistent with mutant DnaA being a substrate of GroEL/ES. Two classes of

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thermosensitive dnaA mutants have been identified. The first class, including dnaA46, recovers activity when returned to permissive temperature. The second is irreversibly inactivated. Overexpression of GroEL/ES suppresses the thermosensitivity of the first but not the second class of mutants (Jenkins et al., 1986; Fayet et al., 1986). Curiously, overexpressing GroEL/ES in either a dnaA46 mutant strain or a dnaA46/dnaA + merodiploid strain confers a cold-sensitive phenotype that is not seen when GroEL/ES is overexpressed in a dnaA + strain (Hansen et al., 1984, Fayet et al., 1986;Jenkins et al., 1986; Katayama and Nagata, 1991). Though the genetic data suggests a role for GroEL/ES in activating a DnaA mutant, this may simply represent the ability of the chaperone to bind improperly folded proteins rather than indicate a role for GroEL/ES in the function of wild-type DnaA. 3.6. Cell Division Mutants of dnaK, dnaJ, and grpE filament without septa at permissive temperatures. Filamentation is more extensive when the mutants are shifted to nonpermissive temperatures (Georgopoulos and Herskowitz, 1971; Paek and Walker, 1987; Bukau and Walker, 1989a; Kang and Craig, 1990; Sell et al., 1990). groEL and groES mutants also filament without septa at nonpermissive temperature (Georgopoulos and Eisen, 1974; Wada and Itikawa, 1984; Horwich et al., 1993), as do cells with limiting or overproduced trigger factor (Guthrie and Wickner, 1990). Since mutations in a large number of other genes confer filamentation phenotypes, it has been difficult to determine the cellular targets affected by the chaperone mutations. Filamentation of dnaK52 mutants can be fully suppressed by the sidB mutant alleles of rpoH, which lower hsp expression (see section 2.1). This suggests that filamentation results from the overexpression of hsps or the consequent down-regulation of other proteins. It also implies that dnaK either does not play a direct role in cell division or is compensated for by a redundant activity. It should be noted that rpoH mutations that lower hsp expression more than the sidB alleles induce filamentation at nonpermissive temperature (Tsuchido et al., 1986). Thus, inappropriately low levels of hsp expression also block cell division. Filamentation of dnaK mutants is also suppressed when cells are grown in minimal glucose medium instead of rich medium (Georgopoulos et al., 1994; Rockabrand et al., 1995). Perhaps as a consequence, dnaK mutants retain higher viability and are less likely to acquire suppressing mutations when cultured in minimal medium. A factor required for cell division may be limiting in dnaK mutants grown in rich medium due to faster growth or to differences in gene expression in rich vs. minimal media. Filamentation might result from induction of a specific inhibitor of cell division or from impaired chromosome partition or septum formation. The block in cell division in dnaK52 mutants does not result from induction of the cell division inhibitor SulA (Paek and Walker, 1987; Bukau and Walker, 1989a; Bukau and Walker, 1989b). Mutants of ftsZ, which encodes a major structural component of the septum, also filament without septa. One possibility is that FtsZ may be malfunctioning in dnaK52 mutants. Overexpressing FtsZ does suppress filamentation of a dnaK52 mutant at 30°C (Bukau and Walker, 1989a; Bukau and Walker, 1989b). The mutants still grow poorly, however, and acquire subpopulations of minicells and anucleate cells. Furthermore, ftsZmutants

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differ from dnaK52 mutants in that they exhibit normal nucleoid segregation (Bukau and Walker, 1989b). Thus, although FtsZ may be malfunctioning in dnaK52 mutants, the mutants are probably also blocked at another stage of cell division. Donachie (1993) has noted that the nucleoid segregation defect of dnaK52 strains resembles that of mukB null and mukB106 missense mutants at nonpermissive temperature (Niki et al., 1991). MukB is a myosin-like protein that may play a role in chromosome segregation. The similarity of the dnaK and mukB phenotypes is strengthened by the finding that overexpressing DksA, which suppresses the temperaturesensitivity of a mukB106 mutant (Yamanaka et al., 1994), also partially suppresses the filamentation and temperature-sensitivity of dnaK, dnaJ, and grpE mutants (Kang and Craig, 1990).dksA is homologous to several other bacterial and bacteriophage genes of unknown function (Bass et al., 1996). In addition to suppressing mukB and dnaK mutations, dksA has been isolated as a multicopy or missense suppressor of two other kinds of mutations, both related to cell division: 1. Overexpressing DksA suppresses the temperature-sensitive and filamentation phenotypes of a null mutation in prc, which encodes a periplasmic protease that is involved in the proteolytic processing of peptidoglycan precursors and the essential cell division protein PBP3 (Bass et al., 1996). PBP3 is required for peptidoglycan synthesis at the septum. 2. A missense mutation in dksA suppresses the replication initiation defects of pSC101 plasmids carrying mutations at the origin of replication in the replication enhancer element (Ohkubo and Yamaguchi, 1997). The missense allele also suppresses the growth defects of a dnaK mutation. The allele appears to be a gain-of-function mutation, because a null mutation in dksA has no effect on pSC101 replication). The finding that dksA suppresses defects in replication initiation, chromosome segregation, and septation conferred by various mutations and also suppresses dnaK mutant growth phenotypes may indicate that all the mutations affect some common process at a key stage of cell division. Alternatively, dksA may be a general bypass suppressor of various cell division defects. This is unlikely, however, since overexpressing DksA has no effect on the growth and filamentation phenotypes of groEL or groES mutants (Kang and Craig, 1990). A null mutation in dksA has no apparent effect on cell growth in rich media but retards growth in minimal media (Kang and Craig, 1990). The regulation of peptidoglycan biosynthesis is intimately coupled with septum formation and cell division (Nanninga, 1991; Donachie, 1993). dksA establishes a tentative link between the cell division defects of dnaK mutants and the defects in peptidoglycan synthesis caused by prc mutants. A more direct connection between dnaK and peptidoglycan synthesis is made by a second multicopy suppressor of dnaK, dapE (msgB), which is an essential gene in the peptidoglycan biosynthetic pathway (Wu et al., 1992). Overexpressing DapE partially suppresses the growth defects of dnaK, dnaJ, and grpE mutants. As with dksA, dapE may compensate for a specific defect of dnaK or instead be a nonspecific bypass suppressor. Very little is known about the basis for the filamentation phenotypes of groEL or groES mutants. However, the role of GroEL/ES in activating the F plasmid encoded

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inhibitor of cell division, LetD, has been characterized in some detail. LetD (ccdB) and a second F plasmid encoded factor, LetA (ccdA), form a suicide/antidote system that blocks division of cells that have lost the plasmid (for review, see Jensen and Gerdes, 1995). LetD is a stable protein that blocks chromosome partitioning by binding and inhibiting DNA gyrase subunit A. LetA is an unstable protein, degraded by the lon protease (Van Melderen et al., 1994), that inhibits LetD activity by dissociating LetD from DNA gyrase. LetD activity is inhibited by mutations in groEL, groES, or two other host genes, tldD and tldE (Miki et al., 1988; Miki et al., 1992; Murayama et al., 1996). The activity of LetD is restored in these mutants by inactivation of csrA. CsrA, which is induced upon entry into stationary phase, regulates carbon storage by repressing glycogen biosynthesis and gluconeogenesis. The simplest model for these genetic data is that csrA or another gene under its control inhibits LetD activity but is itself inhibited by tldD/E and groEL/ES. The function of GroEL/ES might be to activate TldD or TldE. Since single and double null mutants of tldD and tldE have no discernible effect on growth in cells lacking the F plasmid, it is not clear what role they normally play. Interestingly, a subset of hsps that includes DnaK and GroEL is induced by LetD and other inhibitors of DNA gyrase, such as nalidixic acid and novobiocin (Krueger and Walker, 1984; VanBogelen et al., 1987b; Kaneko et al., 1996). 3.7. Acquired Stress Tolerance and Survival during Starvation Cells develop resistance to otherwise lethal exposures of stresses by at least two mechanisms, prior exposure to sublethal levels of certain stresses during exponential growth (acquired stress tolerance) and starvation-induced stress tolerance. The mechanism and regulation of acquired stress tolerance are still poorly understood (reviewed by Parsell et al., 1994; Li et al., this volume), whereas the regulation of starvation-induced stress tolerance is somewhat better understood (for recent reviews, see Hengge-Aronis, 1996; Loewen and Hengge-Aronis, 1994; Nyström, 1995). The response to starvation is mediated principally by the alternative sigma factor, , which initiates the transition into stationary phase. In response to induction, starved cells synthesize trehalose, which protects against heat and osmotic stress, and undergo other global changes in macromolecular synthesis and cell morphology that are likely to protect against stress. Since hsps are required for basal stress tolerance and are induced by starvation and other stresses, they are likely to be important for both acquired and starvation-induced stress tolerance, but their roles are not yet clear. The -dependent induction of hsps following heat shock is necessary for cells to acquire thermotolerance (Yamamori and Yura, 1982). In contrast, hsps induced by overexpression of from a heterologous promoter do not confer thermotolerance (VanBogelen et al., 1987a). The patterns of hsp induction when is overexpressed differs from those seen during heat shock. This raises the possibility that is not sufficient to induce one or more hsps required for thermotolerance. A comparison of the hsps induced by stresses that do or do not confer thermotolerance failed to identify a set of hsps uniquely associated with acquired thermotolerance (VanBogelen et al., 1987a). Thus, proteins other than hsps are likely to be necessary for the establishment of thermotolerance. These might include proteins under the regulation of , a subset of

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which are induced by heat shock (Muffler et al., 1997). Conflicting results have been reported, however, regarding the role of in acquired thermotolerance. Under similar conditions, has been found to be necessary (McCann et al., 1991) or unnecessary (Hengge-Aronis et al., 1991) for acquired thermotolerance. Deletion of the gene for , rpoE, has indicated that the regulon is not necessary for acquired thermotolerance (Rouvière et al., 1995). Conflicting results have also been reported for the role of dnaK in acquired stress tolerance. Mutants of dnaK that are temperature sensitive at 42°C fail to acquire thermotolerance induced by heat shock at 42°C (Taniguchi et al., 1989; Delaney, 1990; Wild et al., 1993; Rockabrand et al., 1995). In contrast, the dnaK756 mutant, which is temperature resistant below 43°C, still acquires thermotolerance following heat shock at 42°C (Ramsay, 1988). Perhaps these results are explained by the temperature sensitivities of the dnaK mutants during the initial heat shock. Some cellular function may be impaired in the more defective dnaK mutants at 42°C, blocking the acquisition of thermotolerance, though not reducing viability during the heat shock period. This would indicate some role for dnaK during the acquisition of thermotolerance. The role of dnaK in acquired resistance to other stresses, such as exposure to H2O2, is not clear. In one study, a nonsense mutant of dnaK did not develop H2O2-induced H2O2 resistance (Taniguchi et al., 1989), whereas a dnaK52 mutant did under similar conditions in another study (Rockabrand et al., 1995). In S. cerevisiae, Hsp104 plays a major role in mediating acquired stress tolerance (Sanchez and Lindquist, 1990; see Lindquist and Schirmer, this volume). ClpB is the E. coli member of the hsp100 family of chaperones most closely related to Hsp104. Like Hsp104, ClpB appears to solubilize cytosolic protein aggregates (see section 3.4). However, null mutants of clpB are not defective for acquired thermotolerance, although they grow more slowly at 44° C and lose viability somewhat faster than a wild-type strain at 50°C (Squires et al., 1991). A closely related clpB homolog in a strain of the cyanobacterium Synechococcus sp. contributes to acquired thermotolerance (Eriksson et al., 1996). Both Hsp104 and its mitochondrial homolog in S. cerevisiae, Hsp78, can partially compensate for reduced Hsp70 function (Sanchez et al., 1993; Schmitt et al., 1995). Whether a similar redundancy might exist in E. coli between DnaK and ClpB has not been explored. Small hsps also confer stress tolerance in some organisms. Two members of the small hsp family have been identified in E. coli, IbpA and IbpB, but no studies of their role in stress tolerance have been reported (see section 4.2). The regulation of starvation-induced stress tolerance is quite distinct from that of acquired stress tolerance, and the roles of hsps also appear to differ. -dependent induction of hsp expression is not required for short-term viability of cells during starvation or for development of starvation-induced thermotolerance and H2O2 resistance, although a subset of the -dependent hsps, including DnaK, GroEL/ ES, and HtpG, is induced by glucose starvation. These hsps are not induced by starvation in a rpoH mutant, but the mutant still acquires starvation-induced thermotolerance and H2O2 resistance. A rpoH mutant is fully viable for up to 20 days of glucose starvation, but thereafter loses viability more rapidly than its wild-type parent (Jenkins et al., 1991). In contrast to rpoH, dnaK mutations inhibit many of the adaptive changes that occur during starvation, including development of starvation-induced thermotolerance and

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resistance to H2O2 (Rockabrand et al., 1995). Starving dnaK52 mutants do not undergo the shift from a rod-like to a spherical cell morphology exhibited by starving wild-type cells. The mutant cells shorten in length, indicating arrest at an early stage of the transition. These phenotypes resemble those of strains deleted for rpoS, the gene encoding the stationary phase-specific factor, . dnaK52 mutants are, in fact, defective in the induction of and -dependent genes (Muffler et al., 1997; Rockabrand et al., 1998). dnaK appears to be required both for the increased translation and the proteolytic stabilization of as cells enter stationary phase. The effects of dnaK mutants during starvation appear to extend beyond their role in regulation, however. Recent evidence suggests that starvation-induced thermotolerance is mediated both by a -dependent pathway, which is impaired in dnaK52 mutants, and by a separate DnaK-dependent/ -independent pathway (Rockabrand et al., 1998). The starvation responses of dnaK and rpoS mutants also differ in that dnaK mutants retain viability during brief periods of glucose starvation under certain conditions (Rockabrand et al., 1995) whereas rpoS mutants rapidly lose viability. The resistance of dnaK52 mutants to brief periods of glucose starvation is similar to rpoH strains and contrasts to the sensitivity of rpoS strains. Evidently, dnaK mutants are not defective in all aspects of the starvation response. The sensitivities of dnaK mutants to glucose starvation do vary under different conditions, however. dnaK52 mutants remain viable in liquid minimal medium when starved for glucose for up to three days (Rockabrand et al., 1995), whereas dnaK103 rapidly loses viability on minimal plates when starved for the same length of time (Spence et al., 1990). This could reflect differences between the mutant alleles, but it is also possible that high cell densities or drying on plates sensitizes dnaK mutants to carbon starvation. The role of GroEL/ES during stationary phase has not been examined closely. A groEL mutant strain that expresses a truncated form of GroEL lacking the last 16 residues recovers slowly from stationary phase if starved at 42°C (McLennan et al., 1993). The mutant otherwise behaves like wild-type when screened for a range of phenotypes common to groEL mutants. This impairment of stationary phase survival may be allelespecific, or it may have been missed in more defective groEL mutants.

4. THE ORPHANED CHAPERONES OF E. COLI: HOMOLOGS IN SEARCH OF FUNCTION 4.1. HtpG (Hsp90) htpG encodes a protein highly homologous to the eucaryotic hsp90 chaperone family. Expression of htpG is -dependent and is induced under a wide range of stress conditions (Bardwell and Craig, 1987; for review see Buchner, 1996). Null strains of htpG grow nearly as well as wild-type, are mildly temperature sensitive (Bardwell and Craig, 1988), and retain full viability during starvation (Spence et al., 1990). The strongest evidence for HtpG having chaperone activity comes from in vitro studies, where HtpG, like its eucaryotic counterparts, suppresses the aggregation of unstable

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folding intermediates (Jakob et al., 1995). An in vivo role for htpG is suggested by its identification as a multicopy suppressor of the growth defects of strains with limiting FtsH (HflB) expression or function (Shirai et al., 1996). FtsH is an essential integral membrane ATP-dependent protease that degrades and thus negatively regulates both (Herman et al., 1995; Tomoyasu et al., 1995) and, under certain conditions, SecY, an integral membrane component of the protein export machinery (Kihara et al., 1995). Missense mutants of ftsH are defective in protein export and membrane protein localization (Akiyama et al., 1994). Overexpression of HtpG suppresses mislocalization, but has no effect on protein export. In contrast, overexpression of groEL/ES suppresses the export but not the mislocalization defects of ftsH mutants. The mechanisms underlying these phenotypes and the possible roles of HtpG and GroEL/ES remain to be determined. htpG was also previously identified as a multicopy suppressor of the temperature sensitivity of the secY24 mutation (Ueguchi and Ito, 1992), but this may have been an indirect effect (Shirai et al., 1996). The eucaryotic hsp90s act in concert with other cofactors to stabilize intermediates in signal transduction pathways and might participate with hsp70s in the general cytoplasmic folding pathway (for a recent reviews see Buchner, 1996; Johnson and Craig, 1997). HtpG might also work with yet-to-be-defined cofactors in similar roles. 4.2. IbpA and IbpB (Small Hsps) The 16 kd IbpA and IbpB proteins are members of the small hsp (hsp20) family. The two proteins are about 48% identical and share three conserved C-terminal motifs with eucaryotic small hsps (Allen et al., 1992; Chuang et al., 1993; Jakob and Buchner, 1994). IbpA and IbpB are coexpressed from a -dependent operon and are strongly induced by heat shock. They have recently been reported to accumulate in the outer membrane following heat shock (Laskowska et al., 1996b). Both associate with proteins that transiently aggregate in wild-type cells following heat shock and inclusion bodies formed by the overexpression of unstable proteins (Allen et al., 1992; Laskowska et al., 1996b). One role of IbpA/B maybe to protect cells during starvation. A small hsp induced during long-term starvation in Mycobacterium tuberculosis increases stationary-phase survival of M. tuberculosis (Yuan et al., 1996). An effect of starvation on IbpA/B expression in E. coli has not been reported.

5. Chaperones and adaptation: comparing genomes This review has focused only on the genetics of chaperones in E. coli, with little reference to studies in other bacteria. However, one way of trying to understand how the requirements for different chaperones may change with changing conditions is to ask how bacterial species that have adapted to very different lifestyles and growth conditions differ in the chaperones that they use and the ways that they use them. Among the first four bacterial genomes to be completely sequenced, only those of Haemophilus influenzae and Escherichia coli have representatives of all the major chaperone families (Fleischmann et al., 1995; Blattner et al., 1997). Mycoplasma

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genitalium lacks hsp90 and so far has no recognizable small hsps (Fraser et al., 1995; Mushegian and Koonin, 1996). The archaea Methanococcus jannaschii and Archaeoglobus fulgidus appear to lack an hsp70 complex entirely, as well as several other chaperone families (Bult et al., 1996; Klenk et al., 1997; see Koonin et al., 1997). This is not characteristic of the archaea generally, since homologs of dnaK have been identified in other archaebacterial species (Macario et al., 1991; Gupta and Singh, 1992; Gupta and Singh, 1994; Smith et al., 1997). The number of members of each chaperone family represented in an organism is also not conserved. E. coli and H. influenzae each have two hsp70 members, homologs of DnaK and Hsc66. By contrast, M. genitalium has only one hsp70. With respect to the hsp60 family, the three bacteria each have only a single homolog, while the nitrogen fixing bacteria Rhizobium meliloti (Rusanganwa and Gupta, 1993; Ogawa and Long, 1995) and Bradyrhizobium japonicum (Fischer et al., 1993) carry four or more groEL genes. The increased number of groEL/groES paralogs in the nitrogen fixing bacteria Rhizobium meliloti and Bradyrhizobium japonicum is matched by an increased specialization in their regulation or function. Expression of one of the groE operons in Bradyrhizobium japonicum is coordinately regulated with the genes involved in nitrogen fixation (Fischer et al., 1993). Similarly, high levels of expression from one of the groE operons of Rhizobium meliloti is required for nitrogen fixation and normal root nodule formation (section 3.1). Other examples of specialized regulation or function of the conserved chaperones are being found in different bacteria. The expression and cellular localization of DnaK and GroEL are regulated in a cell-cycle-dependent manner during the growth and differentiation of Caulobacter crescentus (Reuter and Shapiro, 1987; Gomes et al., 1990; Avedissian and Lopes Gomes, 1996). Several developmental responses of Myxococcus xanthus mediated by cell-cell signaling require a constitutively expressed dnaK homolog, sglK, including social motility and fruiting body formation (Yang et al., 1998). Finally, novel solutions to common problems are turning up in different bacteria. One example of this is the regulation of heat shock gene expression in the Gram-positive bacterium Bacillus subtilis. The expression of dnaK and several other heat shock genes in B. subtilis are principally regulated by a transcriptional repressor, HrcA, rather than by a transcriptional activator as in E. coli (reviewed in Hecker et al., 1996). GroEL promotes the formation of active HrcA and thus down-regulates the heat shock response of B. subtilis, much as DnaK negatively regulates the heat shock response in E. coli (Mogk et al., 1997). Clearly, comparative studies of chaperone regulation and function in divergent species of bacteria are beginning to provide a new and fruitful approach to understanding how chaperones are integrated into the metabolic and adaptive pathways of the cell.

6. ACKNOWLEDGMENTS We gratefully acknowledge former lab members George Gaitanaris and Alexander Gragerov for many years of fruitful and instructive collaboration and discussion. We thank Alexander Gragerov, Art Horwich, Susan Lindquist, Christos Panagiotidis, Justina

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Voulgaris, Randolph Watnick, and Joanne Burkholder for their comments on the manuscript, and Bernd Bukau, Alexander von Gabain, and Susan Gottesman for communicating results prior to publication. Work in our laboratory is supported by NIH grant GM37219.

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Van Melderen, L., Bernard, P. and Couturier, M. (1994). Lon-dependent proteolysis of CcdA is the key control for activation of CcdB in plasmid-free segregant bacteria. Mol Microbiol. , 11 , 1151–1157. VanBogelen, R.A., Acton, M.A. and Neidhardt, F.C. (1987a). Induction of the heat shock regulon does not produce thermotolerance in Escherichia coli . Genes Dev. , 1 , 525– 531. VanBogelen, R.A., Kelley, P.M. and Neidhardt, F.C. (1987b). Differential induction of heat shock, SOS, and oxidation stress regulons and accumulation of nucleotides in Escherichia coli. J. Bacteriol , 169 , 26–32. Vickery, L.E., Silberg, J.J. and Ta, D.T. (1997). Hsc66 and Hsc20, a new heat shock cognate molecular chaperone system from Escherichia coli. Protein Sci. , 6 , 1047– 1056. Viitanen, P.V., Gatenby, A.A. and Lorimer, G.H. (1992). Purified chaperonin 60 (groEL) interacts with the nonnative states of a multitude of Escherichia coli proteins. Protein Sci. , 1 , 363–369. Wada, M., Kadokami, Y. and Itikawa, H. (1982). Thermosensitive synthesis of DNA and RNA in dnaJ mutants of Escherichia coli K-12. Jpn. J. Genet. , 57 , 407–413. Wada, M. and Itikawa, H. (1984). Participation of Escherichia coliK-12 groE gene products in the synthesis of cellular DNA and RNA. J. Bacteriol. , 157 , 694–696. Wada, M., Sekine, K. and Itikawa, H. (1986). Participation of the dnaK and dnaJ gene products in phosphorylation of glutaminyl-tRNA synthetase and threonyl-tRNA synthetase of Escherichia coli K-12. J. Bacterial , 168 , 213–220. Wall, D., Zylicz, M. and Georgopoulos, C. (1994). The NH2-terminal 108 amino acids of the Escherichia coli DnaJ protein stimulate the ATPase activity of DnaK and are sufficient for replication. J. Biol Chem. , 269 , 5446–5451. Wawrzynow, A., Banecki, B., Wall, D., Liberek, K., Georgopoulos, C. and Zylicz, M. (1995a). ATP hydrolysis is required for the DnaJ-dependent activation of DnaK chaperone for binding to both native and denatured protein substrates. J. Biol Chem. , 270 , 19307–19311. Wawrzynow, A., Wojtkowiak, D., Marszalek, J., Banecki, B., Jonsen, M., Graves, B., Georgopoulos, C. and Zylicz, M. (1995b). 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. Wawrzynow, A., Banecki, B. and Zylicz, M. (1996). The Clp ATPases define a novel class of molecular chaperones. Mol Microbiol , 21 , 895–899. Wegrzyn, A., Taylor, K. and Wegrzyn, G. (1996). The cbpA chaperone gene function compensates for dnaJ in plasmid replication during amino acid starvation of Escherichia coli. J. Bacteriol. , 178 , 5847–5849. 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. (1991a). Function of DnaJ and DnaK as chaperones in originspecific DNA binding by RepA. Nature , 350 , 165–167. Wickner, S., Hoskins, J. and McKenney, K. (1991b). Monomerization of RepA dimers by heat shock proteins activates binding to DNA replication origin. Proc. Natl Acad. Sci. USA , 88 , 7903–7907. Wickner, S., Skowyra, D., Hoskins, J. and McKenney, K. (1992). DnaJ, DnaK, and GrpE heat shock proteins are required in onP1 DNA replication solely at the RepA

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7. GENETIC DISSECTION OF THE Hsp70 CHAPERONE SYSTEM OF YEAST ELIZABETH CRAIG * , WEI YAN and PHILIP JAMES Department of Biomolecular Chemistry, University of Wisconsin, Madison, WI, USA 53706

1. Introduction 2. The Hsp70 Class 2.1 Hsp70s of the Cytosol/Nucleus 2.1.1. The Ssa Hsp70s 2.1.2. The Ssb Hsp70s 2.1.3. Basis of the Functional Differences Between Ssa and Ssb Hsp70s 2.1.4. The Sse Class of Hsp70s 2.1.5. Pdr13 2.2. Hsp70s of the Mitochondria 2.3. Hsp70s of the ER 3. The DnaJ Multigene Family of Yeast 3.1. DnaJs of the Cytosol/Nucleus: Ydj1, Sis1 and Zuo1 3.1.1. Ydj1 3.1.2. Sis1 3.1.3. Zuo1 3.2. DnaJ of the Mitochondria 3.3. DnaJs of the ER: Sec63 and Scj1 3.4. Other DnaJs: Xdj1 and Caj1 4. MGE1, a GrpE-related Gene of the Mitochondrial Matrix 5. Functional Interactions Among Classes of Chaperones in Protein Folding 6. Conclusions 7. References *Corresponding

author

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1. INTRODUCTION It has been known for more than a decade that eucaryotes have multiple families of genes encoding molecular chaperones, particularly those of the Hsp70 and DnaJ classes. The completion of the sequence of the S. cerevisiae genome allows a unique opportunity to consider the entire array of molecular chaperones within a eucaryotic organism. Although some of the predicted proteins are yet to be analyzed, a great deal of useful information has been obtained for many members. Different, but related proteins are found in various cellular compartments: the cytosol/nucleus, ER and mitochondria. Each of these three cellular compartments has more than one type of Hsp70, while the ER and the cytosol have more than one DnaJ. The presence of multiple chaperones of both the Hsp70 and DnaJ classes raises the question of whether related proteins in the same compartment are functionally different. In addition, it will be important to dissect the interrelationship between the Hsp70s and proteins with which they interact, such as DnaJs, in order to understand the roles of chaperones in cellular processes. Below we outline the families of molecular chaperones and their functions within the cell as we understand them today, focusing on those of the Hsp70 class and the proteins that function with them.

2. THE Hsp70 CLASS Analysis of the complete genome sequence of S. cerevisiae revealed a total of 14 open reading frames predicted to encode proteins related to Hsp70s. Ten of these were identified and reported prior to the sequencing of the yeast genome. The sequence relationship among these 14 proteins is shown in Figure 1. The proteins in the top part of the tree, Ssa1 through Ssh1, share a high level of sequence identity with one another and with identified Hsp70s from other organisms. For example, the Ssa subfamily is most closely related to Hsp70s of the cytosol of other eucaryotes. Kar2 is most closely related to BiP of the mammalian ER, while Ssc1, Ssh1, and Ssj1 are most closely related to other mitochondrial Hsp70s and bacterial Hsp70s such as DnaK. The others, Pdr13, Ssi1, and Sse1, 2, are less related and may ultimately form individual Hsp70-related subgroups. The Sse1, 2 and Ssi1 proteins are considerably larger than members of the “core” Hsp70 group and are most obviously related to Hsp70s in the ATPase domain. Further functional information is required to determine whether these proteins have the characteristic ATPase and peptide binding activities of Hsp70s. Figure 1 also indicates that the cytosol, ER and mitochondria each contain more than one family of Hsp70. Our current state of knowledge is sufficient to establish that at least the Ssa and Ssb proteins are present in the cytosol, Kar2 and Ssi1 in the lumen of the ER and Ssc1, Ssh1 and Ssj1 in the matrix of the mitochondria. Below, the individual classes of Hsp70s are discussed, emphasizing the roles they play in cellular processes and the similarities and differences among them.

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2.1. Hsp70s of the Cytosol/Nucleus The yeast cytosol contains at least two Hsp70 subfamilies, the Ssas and Ssbs. Analysis of Hsp70s from a diverse set of organisms revealed that the SSA proteins are more closely related to the cytosolic Hsp70s from other organisms than to the SSB proteins (Boorstein et al., 1994). Therefore, the SSB proteins represent an ancient divergence from other known cytosolic Hsp70s. As discussed in more detail below, the SSA and SSB proteins are functionally distinct. The Ssa proteins function in the regulation of the heat shock response, the translocation of some proteins into organelles, and the folding of proteins in the cytosol. The Ssb proteins associate

Figure 1 Phylogenetic tree of the Saccharomyces cerevisiae Hsp70 proteins. Subcellular location and phenotypes of deletion mutants are indicated for each Hsp70 family. The tree was constructed using MegAlign software from DNASTAR (Madison, WI) and the clustal method. Amino acid sequences were truncated at the N-terminus to remove the leader sequences of Hsp70s localized in mitochondria or ER. The first amino acids included in the alignment are: 4 (Ssa1–4, Sse1–2), 9 (Ssb1–2), 32 (Ssc1), 52 (Kar2), 30 (Ssh1), 22 (Ssi1), 29 (Ssj1), 39 (Pdr13).

with translating ribosomes and ssb1 ssb2 mutants are sensitive to some translationinhibiting drugs. In the next three sections, we describe the Ssa and Ssb families independently, and then discuss experiments which delineate the functional differences between them. In addition, the Sse1, 2 and Pdr13 Hsp70s, which are

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discussed briefly below, likely also reside in the cytosolic compartment since they contain no obvious organellar targeting sequence. 2.1.1. The Ssa Hsp70s The Ssa subfamily is essential and is composed of four members that share between 84 and 97% identity (Werner-Washburne et al., 1987). Under optimal growth conditions, only SSA1 and SSA2 are expressed, with the more distantly-related SSA3 and SSA4 being induced in response to a temperature upshift or other stress. While expression of SSA2 remains unchanged during stress conditions, expression of SSA1 is increased. Although strains carrying mutations in different combinations of SSA genes have different phenotypes, this is likely due to differences in expression levels; a combination of genetic experiments have provided no evidence of functional differences among members of the SSA Hsp70 family (Craig et al., 1995). That is, cells with the same level of any of the four individual Ssa proteins or a combination of Ssa proteins appear phenotypically the same, at least under conditions tested thus far. The complexity and essentiality of the SSA family of genes illustrates the central role of this class of Hsp70s in cellular function. Analysis of strains containing mutations of SSA genes have been used to begin to dissect the multiple roles played by the Ssa proteins. Strains lacking both Ssa1 and Ssa2, the two constitutively expressed members of this family, have been particularly useful, ssa1 ssa2 strains grow more slowly than wild-type strains at temperatures up to 35°C and are unable to form colonies at 37°C (Craig and Jacobsen, 1984). The viability of cells lacking Ssa1 and Ssa2, along with their constitutive thermotolerance, is indicative of the role of Ssa proteins in regulating the heat shock response (see chapter by Morimoto). Like cytosolic Hsp70s of other eucaryotes, Ssa proteins negatively regulate the activity of Hsf, the heat shock transcription factor. In ssa1 ssa2 strains the expression of a variety of heat shock proteins is induced even at low temperatures (Craig and Jacobsen, 1984; Boorstein and Craig, 1990a, b). Among the genes whose expression is enhanced are SSA3 and SSA4; it is this expression that allows for viability of ssa1 ssa2 cells. However the expression of Ssa3 and Ssa4 does not reach the levels seen for Ssa1 and Ssa2 under normal conditions, explaining the failed regulation of heat-inducible genes in these strains. If overexpressed, Ssa3 or Ssa4 can restore the expression of heat shock proteins to normal levels. Consistent with the role of SSA proteins regulating Hsf activity, when Ssa1 is overexpressed the induction of the SSA3 and SSA4 genes in response to a heat shock is greatly reduced (Stone and Craig, 1990). Extragenic suppressors of the slow growth defect of an ssa1 ssa2 strain have been isolated (Nelson et al., 1992). Analysis of these suppressors has provided additional insights into the importance of the Ssa proteins in regulating the expression of heat shock-inducible proteins. One such suppressor, named EXA3, rescued the slow growth phenotype of ssa1 ssa2 strains at temperatures such as 23° and 30°C. EXA3 was subsequently shown to be an allele of HSF1, which encodes the Hsf transcription factor (Halladay and Craig, 1995). The initial expectation was that a mutation in HSF1 able to suppress the growth defect of ssa1 ssa2 cells would cause increased Hsf activity and thus increased Ssa3 and Ssa4 expression. However, the induction of Hsf-driven transcription

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in response to a heat shock is significantly delayed in an EXA3–1 strain compared to wild-type. In addition, the basal level of Hsf-driven expression at all temperatures was several fold lower in the mutant than in the wild-type strain. This decreased expression suggests that the EXA3–1 allele encodes an Hsf with reduced activity. Consistent with this idea, the EXA3–1 allele of Hsf contains a single point mutation which causes the change of a proline to a glutamine in the DNA binding domain of Hsf (Halladay and Craig, 1995). As a result, the DNA binding activity of the mutant Hsf was reduced about 50 fold in in vitro gel shift assays. The affected proline is at position 214, which is absolutely conserved in Hsf homologs from all organisms examined and produces a prominent kink (Harrison et al., 1994) near the end of the first helix of the helix-turnhelix DNA binding motif. How can these phenotypes be explained? Under normal conditions, the Ssa1 and Ssa2 proteins serve as negative regulators of Hsf, repressing the transcription of heat shock proteins. In an ssa1 ssa2 strain, Hsf activity is increased and a wide variety of heat shock proteins are constitutively expressed. The constitutive expression of the normally heatinducible Ssa3 and Ssa4 proteins allows the cells to survive in the absence of Ssa1 and Ssa2, but at the same time, the constitutive expression of other heat-inducible genes appears to be detrimental. Consistent with this idea is the observation that an additional copy of Hsf is deleterious to the growth of ssa1 ssa2 cells (Halladay and Craig, 1995). The reduced activity of the EXA3–1 allele of Hsf apparently aids the growth of ssa1 ssa2 cells by decreasing the expression of these detrimental proteins. Thus in an ssa1 ssa2 strain a delicate balance must be met to allow sufficient expression of Ssa proteins to perform functions other than heat shock regulation, while limiting the detrimental expression of other Hsps. Evidence has also been found for a similar situation in prokaryotes (Bukau and Walker, 1990). In E. coli, the Hsp70 DnaK plays an important role in regulating the heat shock response by modulating the level of the subunit of RNA polymerase (see Chapter by Connolly et al.). This subunit is required for the initiation of transcription of heat-inducible genes (Georgopoulos et al., 1994). Bukau and Walker (Bukau and Walker, 1990) selected suppressors of the slow growth phenotype of a dnaK deletion strain and identified mutants of . Interestingly, these mutants had either decreased activity or decreased stability of , leading to the hypothesis that increased expression of Hsps was in part responsible for the growth defects of dnaK mutants. Though Ssa proteins are critical for regulating the heat shock response, they carry out other functions as well. The initial indication that Ssa proteins are involved in the translocation of proteins into the ER and mitochondria came from studies of an ssa1 ssa2 ssa4 mutant which contained an SSA 1 gene under the control of the regulatable promoter GAL1 (Deshaies et al., 1988). Shift from galactose- to glucose-based media caused the repression of SSA1 transcription. By three hours after the shift to glucose the amount of Ssa1 present was significantly reduced and the precursor forms of several organellar proteins had accumulated in the cytosol. However the requirement for SSA function in protein translocation in vivo is certainly not universal. Of three mitochondrial proteins tested, the processing of only one, the subunit of the F1F0 ATPase, was dramatically affected. Of six proteins destined for the ER, the translocation of only prepro- -factor and proteinase A was inhibited.

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The time delay required for Ssa1 depletion in these experiments left open the possibility that the translocation defect could be an indirect effect of the loss of Ssa function. However, similar translocation defects were recently observed using a strain carrying a temperature-sensitive SSA1 allele, ssa1–45 ssa2 ssa3 ssa4 (Becker et al., 1996); in this case the inactivation of Ssa protein should be very rapid. The defect in prepro- -factor translocation appears within 2 minutes after shift to the nonpermissive temperature in the ssa1–45 strain, suggesting a direct requirement for Ssa in the process. Interestingly, in this mutant the block in translocation was not complete. More than 50% of the protein accumulated in the precursor form, with the remainder rapidly reaching the mature form. The accumulated precursor could not be chased through the secretory pathway however. This apparent dichotomy—either complete blockage or normal translocation—could be explained in two ways. It is likely that the translocation process utilizes the chaperone activity of Hsp70s, and that some protein precursors require an association with Hsp70 to prevent aggregation, which would block subsequent translocation. In one scenario, the ssa1–45 mutation could be leaky, providing sufficient activity for translocation of a portion of the affected precursors, but leaving the rest to aggregate. Consistent with this idea, Ssa has been found associated with prepro- -factor in in vitro translation extracts (Chirico, 1992). Alternatively, the affected proteins might enter the ER by two different pathways, one that is dependent on Ssa proteins, and thus is completely blocked in the mutant, and one that is Ssa-independent. If this is true, proteins that fail to complete the Ssa-dependent pathway must not be able to return to the Ssaindependent one, perhaps due to aggregation. Recently, evidence has been reported for two translocation pathways, one SRP-independent and one SRP-dependent (Ng et al., 1996). In addition to the Ssas, the DnaJ homolog Ydj1 is also involved in protein translocation, as a YDJ1 temperature sensitive mutant accumulates the precursor forms of certain proteins (Caplan et al., 1992). A comparison of the proteins whose translocation either was or was not affected in ssa1–45 and ydj1 mutant strains shows a striking similarity (Becker et al., 1996). This similarity suggests that Ydj1 and Ssa1 function together in the translocation process. Consistent with this idea, the ssa1–45 ssa2 ssa3 ssa4 mutations are synthetically lethal with a ydj1 null mutation. In addition, an ssa2 ssa3 ssa4 ydj1 strain contains a functional SSA1 gene, but is defective in prepro- -factor translocation under conditions where neither a ydjl nor an ssa2 ssa3 ssa4 strain show precursor accumulation. These results suggest a specific functional interaction between the Ssa proteins and Ydj1 in the translocation process. Significantly, ssb1 ssb2 ydj1 strains have no synthetic phenotypes. Finally, there is compelling in vitro evidence for a role of cytosolic Hsp70s in folding of denatured proteins (Levy et al., 1995; Freeman et al., 1995; Szabo et al., 1994). Ssa1 in combination with the DnaJ homologue Ydj1 facilitates the efficient refolding of denatured luciferase. This again appears to be an Ssa-specific function, since Ssb proteins do not function in this assay (Ziegelhoffer and Craig, unpublished result). Although strong in vivo evidence is yet to be obtained for a direct role in this process, Ssa proteins are likely to be intimately involved in protein folding in the cell.

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2.1.2. The Ssb Hsp70s The SSB family contains two proteins, which are 99.3% identical and are encoded by the SSB1 and SSB2 genes. Unlike other Hsp70s, their expression is dramatically decreased upon a temperature upshift (Craig and Jacobsen, 1985). The SSB genes are not essential; ssb1 ssb2 cells are cold-sensitive for growth. No phenotypes of single ssb1 or ssb2 mutants have been detected, suggesting that these closely related proteins are functionally identical. It has been hypothesized that SSB proteins are involved in protein translation (Nelson et al., 1992). In part, this idea is based upon the fact that ssb1 ssb2 cells are sensitive to certain drugs which inhibit protein synthesis, including verrucarin A and aminoglycosides such as hygromycin B. In addition, the bulk of Ssb protein is found to sediment with polysomes upon sucrose gradient fractionation of crude cell extracts. This interaction appears to be specific for actively translating ribosomes, as it is disrupted in the presence of puromycin, treatment which results in the release of nascent chains from ribosomes. Given the propensity of Hsp70s to bind to polypeptides which are in an extended conformation (Landry et al., 1992), it has been proposed that Ssb proteins bind to nascent chains as they emerge from the large ribosomal subunit, thereby preventing their premature folding (Nelson et al., 1992) (see chapter by Welch et al.). This hypothesis is supported by recent data demonstrating that Ssb can be crosslinked to the nascent chain on translating ribosomes (Pçund, Wiedmann and Craig, unpublished results). A genetic screen for multicopy suppressors of the cold-sensitive phenotype of ssb1 ssb2 strains resulted in the isolation of a novel homologue of the GTP-binding protein, EF1 , which catalyzes the binding of aminoacyl-transfer RNAs to the ribosome (Nelson et al., 1992). The function of this gene, called HBS1, is unknown; a strain lacking Hbs1 has no obvious phenotype. However, its similarity to EF1 is consistent with the role of Ssb proteins in translation. SSB1 itself has been isolated as a multicopy suppressor of two very different cellular defects. Increased expression of SSB1 suppresses a temperature-sensitive mutation in the Y7 subunit of the multicatalytic proteasome, which functions to degrade both ubiquitinconjugated and non-ubiquitinated substrates (Ohba, 1994). In addition, SSB1 has been identified as a multicopy suppressor of the temperature-sensitive dbf3–1 allele, which was originally characterized as having a cell cycle phenotype due to inhibition of DNA synthesis (Shea et al., 1994). DBF3 is also identical to PRP8, which encodes the U5 snRNP, an important component of the yeast splicosome. These genetic interactions are intriguing, but it is not clear whether they are indicative of specific Ssb functions in proteolysis and RNA splicing, or an indirect effect of the role of SSB proteins in translation. 2.1.3. Basis of the Functional Differences Between Ssa and Ssb Hsp 70s Because eucaryotes have such a large number of different Hsp70s, a basic question is whether these Hsp70s are functionally distinct or overlap in function. The Ssa and Ssb Hsp70s of the cytosol of S. cerevisiae have been studied most extensively. The SSA

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proteins are quite similar to one another in function since any of the four are able to allow wild-type or nearly wild-type growth at all temperatures if present in sufficient quantities (i.e., similar to the normal amount of total SSA protein) (Craig and Jacobsen, 1985; Craig et al., 1995). Similarly, the Ssb1 and Ssb2 proteins are interchangeable. However, it is clear that the Ssas and Ssbs are functionally distinct. Not only, as described above, are the phenotypes of SSA and SSB mutants very different, but even when differences in regulation are accounted for, members of one family can not rescue mutants of the other. Thus expression of Ssb1 under the control of an SSA promoter cannot rescue the phenotypes of an SSA mutant strain; similarly, Ssa1 under the control of an SSB promoter cannot rescue an SSB mutant strain. Differences have been observed between Ssa and Ssb proteins at the biochemical level as well. Ssa protein can facilitate the uncoating of clathrin-coated vesicles (see chapter by Eisenberg and Greene), while Ssbs cannot (Gao et al., 1991). Also, as noted above, Ssas in conjunction with Ydj1 can facilitate the refolding of denatured luciferase (Levy et al., 1995), while Ssbs cannot (Ziegelhoffer and Craig, unpublished result). The basis for the functional differences between the SSA and SSB proteins has been analyzed using chimeric proteins (James et al., 1997). Chimeras were constructed that contain all combinations of the 44 kDa ATPase, the 18 kDa peptide binding and the 10 kDa variable domains characteristic of Hsp70s (see chapter by Ha et al.). While no chimeras were able to rescue the growth phenotypes of SSA mutants, several chimeras allowed growth of ssb1 ssb2 cells at 18°C and/or in the presence of the aminoglycoside hygromycin B. It has been hypothesized that the functional differences between Hsp70s are determined by differences in peptide-binding specificity. However, a chimera that contains the peptide binding domain of Ssa1 was able to rescue both the cold-sensitive and the hygromycin B-sensitive phenotypes of an ssb1 ssb2 strain. Thus peptide binding specificity is not the determinant of the functional differences between these two Hsp70 families. If differences in peptide binding specificity do not explain the functional distinction between these two classes, then what does? An analysis of additional chimeras (James et al., 1997) revealed that rescue of the cold-sensitive phenotype required only that the chimera contain the Ssb1 ATPase domain, while rescue of the hygromycin B sensitivity was more complex (see Table 1). No single domain of Ssb was sufficient for hygromycin B rescue, but also no single domain was required. Rather, a chimera must have two Ssb1 domains to confer hygromycin B resistance, but any two will suffice. In addition, the data in Table 1 demonstrate that the cold-sensitive and hygromycin B-sensitive phenotypes of ssb1 ssb2 strains are separable, since the BAA chimera rescues cold sensitivity but not hygromycin B sensitivity while the ABB chimera rescues hygromycin B sensitivity but not cold sensitivity. Therefore, the SSB proteins carry out two different functions in the cell. Since the ability of a chimera to rescue hygromycin B sensitivity correlates with its ability to associate with ribosomes (with one exception; see Table 1), at least this phenotype is likely to represent a function of Ssb proteins in the translation process. Results from the chimeras demonstrate that the peptide binding domain is dispensible for two different fuctions of the Ssb proteins, and thus is not the source of functional specificity. Those same results suggest a model in which functional specificity depends on interactions between several Hsp70 domains and other components of the cellular

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machinery. The specific Hsp70 domains involved can vary from one function to another. According to this model, an interaction with the ATPase domain is required for the function involved in the rescue of cold sensitivity. On the other hand, all three domains of Ssb appear to interact with the ribosomes or other components of the translational apparatus. These interactions must be additive and sufficiently strong that the loss of any one domain by the

Table 1 Rescue of ssb1 ssb2 mutant phenotypes by Ssa/Ssb chimeric proteins

ssb1 ssb2 Phenotype a 44kDa ATPase Domain

18kDa Peptide Binding Domain

10 kDa Variable Domain

Cold Sensitivity

Hygromycin B Sensitivity

Polysome Association

B

B

B

+

+

+

B

A

A

+

–

–

A

B

A

–

–

–

A

A

B

–

–

–

A

B

B

–

+

+

B

A

B

+

+

+

B

B

A

+

+

–

A

A

A

–

–

–

a. Growth at 18°C or in the presence of 70 µg/ml hygromycin B.

substitution of Ssa sequences does not disrupt the functional interaction. In this model the peptide binding activity of Hsp70 is considered to be a “generic” activity, much like the ATPase activity, which is ferried to particular locations by specific interactions involving other parts of the protein. Such interactions may serve to target an Hsp70 to, for example, the ribosome in the case of Ssb, or to associate with cohort proteins such as DnaJs. 2.1.4. The Sse Class of Hsp70s The Sse1 and Sse2 proteins are quite distantly related to other Hsp70s (approximately 13 to 28% identical) but are 76% identical to each other. Another protein distantly related to Hsp70s is the mammalian heat-inducible protein, Hsp110 (Subjeck et al., 1983). Hsp110 is present at basal levels in mammalian cells and is induced upon a heat shock. A portion of Hsp110 has been found associated with the nucleolus, but the function of Hsp110 is unknown. The Sse proteins and Hsp110 are at least as similar to each other as either is to yeast or mammalian Hsp70s (Lee-Yoon et al., 1995). The subcellular localization of the Sses has not been determined. However, their lack of obvious targeting sequences

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suggests that they may be localized to the cytosol, or perhaps the nucleus. The Sse proteins were originally isolated based upon their calmodulin binding activity (Mukai et al., 1993). In addition, SSE1 was identified as a multicopy suppressor of the heat shock sensitive phenotype of an ira1 mutation, which causes hyperactivation of the RAS-cAMP pathway (Shirayama et al., 1993). The single sse2 mutant has no observable phenotype while sse1 mutant cells have a slow growth phenotype. The phenotype of the double mutant is indistinguishable from that of the sse1 mutant. Like Hsp110, there is no understanding of the functional role which Sses play in the cell. 2.1.5. Pdr13 A recently discovered Hsp70 encoded by PDR13 has been implicated in the regulation of genes involved in pleiotropic drug resistance. PDR13 was isolated in a screen for multicopy suppressors which elevates the resistance of cells to the mitochodrial ATPase inhibitor oligomycin; in addition, overexpression of PDR13 renders cells resistant to cycloheximide (Katzmann et al., 1995). The same phenotype results from an overexpression of YOR1, a gene encoding an ABC transporter. Interestingly, increased copy number of PDR13 leads to elevated expression of YOR1 and other genes involved in pleiotropic drug resistance (Hallstrom et al., 1998). These Pdr1 3 effects require the presence of the zinc finger transcription factor Pdr1, which is necessary for both expression of YOR1 (Balzi and Goffeau, 1995) and oligomycin resistance. Strains that lack Pdr1 are cold-sensitive for growth and hypersensitive to drugs. Together these results suggest that one function of Pdr1 3 may be to positively regulate the transcription factor Pdr1, and thus the expression of a set of genes involved in drug resistance, including ABC transporters. 2.2. Hsp70s of the Mitochondria The matrix of the yeast mitochondria contains three Hsp70s. The most abundant is the well-characterized SSC1 protein which is essential for translocation of many proteins across the mitochondrial membranes. The mechanism of action of this Hsp70 in the translocation process has been studied extensively. Ssc1 is tethered to the inner membrane via its interaction with Tim44, a component of the translocation complex. It binds to the mature part of the translocating polypeptide as well as the presequence. The mechanism by which Ssc1 provides the directionality to the movement of the polypeptide chain is a matter of debate. Ssc1 may harness the energy of Brownian motion by preventing “backsliding” of the polypeptide chain. On the other hand, Ssc1 may act as a force generating motor, with energy derived from a conformational change in Ssc1 as a result of the binding/hydrolysis of ATP. The chapter by Dekker and Pfanner discusses in detail the function of Ssc1 in translocation. Consistent with the apparent multiple roles of the cytosolic Ssa proteins, analysis of conditional SSC1 mutants has provided evidence that Ssc1 is involved in several other mitochondrial processes. These additional functions include the folding of both mitochondrially-encoded (Hermann et al., 1994) and translocated polypeptides. Degradation of matrix proteins by the PIM1 pro tease is also hampered in SSC1 mutants

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suggesting a role for Ssc1 in presenting proteins to the proteolytic apparatus in a conformation permissive for degradation (Wagner et al., 1994). In all of these roles Ssc1 is presumed to bind to partially unfolded protein substrates, acting in cycles of binding and release as do other members of the Hsp70 family. In a seemingly different capacity, Ssc1 is the noncatalytic subunit of the mitochondrial site-specific endonuclease Endo.Sce1 (Morishima et al., 1990), and is required for activity of the catalytic subunit. SSH1 and SSJ1, two genes uncovered during the genome sequencing project, have recently begun to be examined. Only a preliminary examination of SSJ1 has been done at this time. Ssj1 is 80% identical to Ssc1 and contains an apparent mitochondrial leader sequence. However, mitochondrial localization has not been demonstrated at this time. No phenotype of a disruption mutant has been observed (Forster, Schilke, Davis and Craig, unpublished result). Since Ssj1 appears to be expressed at much lower levels than Ssc1, and since the proteins are about 80% identical, one might expect that overexpression of Ssj1 could compensate for lack of Ssc1. However preliminary experiments in which expression of Ssj1 was driven by the SSC1 promoter, ssc1 mutant phenotypes were not suppressed. An understanding of the role of this Hsp70 therefore awaits more extensive investigation. Unlike SSJ1, disruption of SSH1, which encodes a protein 53% identical to Ssc1, is detrimental to yeast cells. SSH1 is not an essential gene; however, cells lacking Ssh1 are cold-sensitive, growing very poorly at temperatures of 23°C and below (Schilke et al., 1996). The functional relationship between Ssc1 and Ssh1 is complex. The level of Ssc1 in the matrix is much higher than that of Ssh1, perhaps as much as a hundred times. However, overexpression of SSC1 can suppress the cold-sensitive phenotype of a ssh1 deletion. In contrast, a fusion which expresses Ssh1 under the control of the SSC1 promoter does not suppress the phenotypes of an SSC1 temperature-sensitive mutation. These results suggest that Ssh1 is unable to carry out the essential roles of Ssc1 when expressed at the same levels as Ssc1. However, Ssc1 can carry out the functions of Ssh1 when present in several hundred fold excess. The function of Ssh1 in mitochondria is not known. However clues have come from the analysis of suppressors of the cold-sensitive phenotype (Schilke et al., 1996). These suppressors arose spontaneously at a high frequency. All were found to be petite mutants, that is, they were respiratory incompetent. 15 of the 16 analyzed were rho0 mutants that lacked mitochondrial DNA (mtDNA) completely, while the 16th contained reduced amounts of mtDNA. Therefore, absence or reduction in the amount of mtDNA overcomes the deleterious effects of the absence of Ssh1. This result suggests two possible roles for Ssh1 in mitochondrial physiology. First, Ssh1 might be involved in the folding of mitochondrially-encoded proteins. In this case the synthesis of proteins in the mitochondria might be deleterious because these proteins would be likely to misfold and aggregate, interfering with other mitochondrial functions. Elimination of the information encoding these proteins would eliminate this problem. However, mitochondrially encoded proteins synthesized in the absence of Ssh1 appear to be soluble. A second hypothesis is that Ssh1 is involved in mtDNA replication and that the absence of Ssh1 results in alterations in the DNA replication process that are deleterious to the cell. In this model, also, the absence of mtDNA would eliminate the problem. In theory these two models could be distinguished by the analysis of rho− mutants of

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SSH1. rho− mutants have deletions of mtDNA which eliminate protein synthesis; however, the remaining mtDNA is amplified such that rho− cells have the same total amount of mtDNA as do wild-type rho+ cells (Dujon, 1981). If the deleterious effect caused by the absence of Ssh1 is due to its role in protein folding, rho− cells should be suppressed, since no mitochondrial protein synthesis occurs. However, if DNA replication is defective in ssh1 cells, rho− and rho+ cells should have the same phenotype, since both contain the same amount of mitochondrial DNA. However, even though rho− cells arise at a rate of 1–5% in a normal population, no rho− ssh1 cells were obtained upon analysis of 4500 cells. The failure to obtain rho− ssh1 cells suggests that the presence of rho- DNA in an ssh1 background is deleterious. These studies cannot distinguish definitively between the two most obvious roles for Ssh1. However, the failure to isolate rho- cells and the fact that the growth of ssh1 cells does not correlate with the amount of mitochondrial proteins synthesized, suggest an involvement of Ssh1 in DNA replication. Further studies will be required to define the cellular role of Ssh1 and to determine the basis of the functional differences between Ssh1 and Ssc1. 2.3. Hsp70s of theER The lumen of the ER of S. cerevisiae contains two members of the Hsp70 family, the essential KAR2 protein, which is the homologue of mammalian BiP, and a recently identified protein Ssil/Lhs1/Cer1 (Craven et al., 1996; Baxter et al., 1996; Hamilton and Flynn, 1996) which is 25% identical in sequence to Kar2 as well as other Hsp70s. Kar2 is important in the translocation of proteins from the cytosol into the lumen of the ER, as shown by both in vivo and in vitro experiments (Rose et al., 1989; Vogel et al., 1990). Kar2 interacts directly with translocating polypeptides and has been proposed to provide a driving force for the unidirectional movement of the polypeptide across the ER membrane. This role is analogous to that carried out by Ssc1 of the mitochondrial matrix to drive translocation into that organelle. This function of Kar2 and its homologue BiP in the translocation process is described in more detail in the chapter by Haas and Zimmermann. Kar2 appears to also be involved in the folding of proteins after their translocation. In a KAR2 mutant the vacuolar protein carboxypeptidase Y aggregates after translocation into the ER, and fails to be transported into the vacuole (Simons et al., 1995). Unlike KAR2, SSI1/LHS1/CER1 is not an essential gene. Strains lacking Ssi1 are coldsensitive for growth (Baxter et al., 1996; Craven et al., 1996; Hamilton and Flynn, 1996). However the severity of the defect varies with the strain background. ssi1 strains accumulate precursor forms of several proteins which are normally translocated into the ER. In one case the precursor has been shown to be on the cytosolic side of the ER membrane indicating a defect in translocation. However, consistent with the nonessential nature of Ssil, the block is not complete. Overexpression of Scj1, a DnaJ of the ER, suppresses the translocation defect of ssi1. In addition to the presumptive role in translocation, it has been suggested that Ssi1 functions in the folding of proteins in the lumen. This suggestion is based upon the fact that the expression of Ssil, like Kar2, is induced by the presence of incorrectly folded proteins in the ER. In addition, an SSI1 deletion displays synthetic lethality with a mutation in IRE1, which encodes a protein

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required for the transcriptional response to an increase in unfolded protein in the ER (Craven et al., 1996). Thus, like Kar2, Ssil is thought to be involved in both the translocation and subsequent folding of proteins in the ER lumen. Ssil has also been implicated in the refolding of denatured proteins. The ER has an ATP-dependent, heat-resistant machinery which is able to refold thermally denatured proteins in cells (Jamsa et al., 1995). In the absence of Ssi1, an in vivo heat-denatured and aggregated reporter enzyme failed to be resolubilized and reactivated, as in wild-type cells. Instead, it was slowly degraded (Saris et al., 1997). Genetic interactions between KAR2 and SSI1 provide some clues as to the relationship between these related proteins (Baxter et al., 1996; Craven et al., 1996; Hamilton and Flynn, 1996). A number of conditional KAR2 mutants exhibit different synthetic growth defects in the presence of the ssi1 deletion mutation. Conditional KAR2 alleles have been grouped into classes based on their phenotypes (Rose, Vogel and Scidmore, personal communication). Class I alleles such as kar2–159 cause a block in translocation at the nonpermissive temperature and rapid loss of viability. Class II alleles, which fail to grow but remain viable at the nonpermissive temperature, do not cause a block in translocation. However, cells having alleles of this class, such as kar2–1, have defects in a number of posttranslocational events. Class III alleles such as kar2–191 show both translocational and posttranslocational defects. The genetic interactions of an ssi1 deletion with the various KAR2 mutants can be grouped into the same classes (Baxter et al., 1996). Class I alleles show synthetic lethality with ssi1. For example, a ssi1 kar2–159 is inviable at all temperatures, including those at which both ssi1 and kar2–159 strains are viable. Class II alleles are not synthetically lethal but show impaired growth compared to both parental strains. On the other hand, a Class III allele, kar2–191, shows better growth in the presence of the SSI1 mutation than in its absence. In addition, overexpression of Ssi1 is deleterious in a kar2–191 background, but not in Class I or Class II kar2 strains tested. What is the explanation of these complex genetic interactions? Since ssi1 cells are impaired in the translocation process, as well as protein folding events, it is likely that Ssi1 can partially substitute for KAR2 function, thus explaining the synthetic lethality or growth defect of Class I and Class II mutant strains, respectively. The synthetic growth enhancement of the Class III allele is more complicated to explain. It has been previously shown that this allele is synthetically lethal with sec63–1, a missense mutation in the gene encoding a component of the translocation machinery homologous to DnaJ (see chapter, Haas and Zimmermann), even in the presence of wild-type KAR2 (Scidmore et al., 1993), suggesting dominant effects of this mutation. Perhaps kar2–191 protein interacts with the translocation apparatus in an inhibitory fashion and Ssi1 enhances this interaction. Such an interaction would lead to improved growth in the absence of Ssi1. Consistent with this idea, increased expression of Ssi1 in this genetic background is deleterious. More thorough analysis of the various alleles of KAR2 should prove very helpful in understanding the relationship between these two Hsp70s of the ER lumen.

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3. THE DNAJ MULTIGENE FAMILY OF YEAST DnaJ has the ability to bind unfolded polypeptides and prevent their aggregation and therefore can be considered a chaperone in its own right. DnaJ, however, also interacts with DnaK, stimulating the hydrolysis of ATP (see chapter by Buchberger et al.). Using the defining motif of DnaJ-related proteins, the “J” domain, 16 potential DnaJs have been identified through screening of the yeast genome, only nine of which have been reported in the literature. Ydj1 is the best studied DnaJ-related protein of S. cerevisiae and has many of the same properties as E. coli DnaJ. Although the studies of other yeast DnaJs is not as advanced, it is likely that all interact with one or more Hsp70s and some, particularly those with an obvious cysteine-rich motif able to form a “zinc finger”, also bind unfolded proteins. Below the best studied DnaJs are discussed. 3.1. DnaJs of the Cytosol/Nucleus: Ydj1, Sis1 and Zuol 3.1.1. Ydj1 The best studied DnaJ of yeast is Ydj1 (Mas5). Ydj1 was identified independently by two groups based on its association with membranes (Caplan and Douglas, 1991) and its role in the facilitation of translocation into mitochondria (Atencio and Yaffe, 1992). Ydj1 is not essential, but cells lacking it grow very poorly at 23°C and are unable to form colonies at 37°C. The structural organization of Ydj1 is similar to that of DnaJ of E.coli, containing glycine- and cysteine-rich regions in addition to the “J” domain (Figure 2) (see chapter by Buchberger et al., for structure/function aspects of DnaJ proteins). In fact, expression of DnaJ can overcome some of the

Figure 2 Schematic diagram of DnaJ homologs from Saccharomyces

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cerevisiae. The protein structure of the eight DnaJ homologs of S. cerevisiae that have been studied to date are shown. Protein motifs discussed in the text are shown. Subcellular location and phenotypes of deletion mutants are indicated.

defects caused by the lack of Ydj1. Unlike other DnaJs of S. cerevisiae, Ydj1 contains a “CAAX” box at its C-terminus and is farnesylated (Caplan et al., 1992), explaining the observed association with membranes. While at 23°C about 15% of Ydj1 was found associated with the membrane fraction, after a shift to 37°C for 1 hour, more than 65% was found membrane-associated. Cells carrying a mutant form of Ydj1 having a substitution for the conserved cysteine in the “CAAX” box are temperature-sensitive for growth and have increased amounts of Ydj1 in the cytosol. Together these results suggest that a membrane-associated function of Ydj1 may be particularly important at high temperatures. What functions does Ydj1 carry out in the cell? Analysis of the temperature-sensitive ydj1-151 mutant strongly indicates that Ydj1 functions in the translocation of some proteins from the cytosol to the ER and mitochondria (Caplan et al., 1992). The ydj1-151 mutant is defective in translocation of prepro- -factor but not three other ER proteins tested, and in the translocation of the F1 , and subunits of the F1F0 ATPase into mitochondria. As discussed in more detail in section 6, Ydj1 has also been implicated in the folding and activation of several heterologous proteins which require Hsp90 function. Although there is no genetic evidence for the function of Ydj1 in the general protein folding pathway, biochemical experiments strongly suggest that possibility. Like DnaJ acting with DnaK, Ydj1 acting with Ssa1 can faciltate the refolding of denatured luciferase (Levy et al., 1995). Furthermore, by itself, Ydj1 can bind to denatured rhodanase and prevent its aggregation (Cyr, 1995). 3.1.2. Sis1 Sis1 is an essential DnaJ of the yeast cytosol (Luke et al., 1991). While Sis1 contains the “J” domain and the glycine-rich repeat, it does not possess the cysteine-rich region found in Ydj1 and DnaJ. However, it does possess a Met-rich region. Whether this region is analogous to the cysteine-rich region in allowing binding to polypeptide substrates is unknown. Compelling genetic and physiological experiments indicate a role for Sis1 in protein synthesis, specifically initiation (Zhong and Arndt, 1993). First, Sis1 comigrates with polysomes and is specifically enriched in fractions that contain only a few ribosomes per messenger. This biased distribution toward small polysomes suggests that Sis1 associates with ribosomes only during and shortly after initiation. In addition, 80S monosomes rapidly accumulate upon temperature upshift of a temperature-sensitive sis1 mutant, indicating a defect in initiation in the absence of functional Sis1. Extragenic suppressors of sis1 ts mutations were found to be in genes that encode ribosomal proteins of the 60S subunit. Although this result underscores a role for Sis1 in translation, how it might function in the initiation process remains unknown. It is likely that Sis1 is involved in either assembly or dissembly of a complex rather than in general protein folding, since it is apparently present only in the very early stages of translation. It should also be

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remembered that Sis1 was first isolated as a multicopy suppressor of mutations in SIT4, which encodes a serine/threonine phosphatase (Luke et al., 1991). Whether this suppressing ability is an indirect effect of its role in translation or reflects an additional function remains to be defined. 3.1.3. Zuo1 Zuo1 was originally detected in nuclear extracts in an assay designed to isolate Z-DNAbinding proteins (Zhang et al., 1992). Independently, Zuo1 was isolated as a transfer RNA binding protein (Wilhelm et al., 1994). Isolation by these two independent nucleic acid binding assays strongly suggests that Zuo1 functions by binding either DNA or RNA, or both. Zuo1 is not an essential protein. However, zou1 strains grow slowly compared to wild-type strains. Unlike Ydj1, Zuo1 contains no cysteine rich domain. Zuo1 has yet to be purified in sufficient quantities to determine whether it can interact with any Hsp70s, prevent protein aggregation, or facilitate protein folding. 3.2. DnaJ of the Mitochondria A single DnaJ, Mdj1, has been identified in the matrix of the mitochondria (Rowley et al., 1994). Mdj1 is not an essential protein, however mdj1 mutants are unable to grow at 37°C. Interestingly, mdj1 mutants are unable to maintain mitochondrial DNA, that is they are rho0. It is unclear at this time whether Mdj1 is actually involved in DNA replication, since many mutations which disrupt the structure or function of mitochondria fail to maintain normal mitochondrial DNA (Piskur, 1994). Further analysis of strains lacking Mdj1 has indicated a requirement for Mdj1 in the efficient folding of proteins imported into mitochondria, as well as to prevent their aggregation (Rowley et al., 1994). Luciferase which has been imported into mitochondria aggregates after partial unfolding in response to a shift to 40°C. This aggregation occurs significantly more rapidly in mdj1 mitochondria compared to wild-type mitochondria (Prip-Buus et al., 1996). These results with Mdj1 are reminiscent of those obtained with DnaJ of E. coli. For example, DnaJ is able to maintain the soluble state of denatured luciferase (Szabo et al., 1994; Schröder et al., 1993). Perhaps these similarities are not surprising considering the symbiotic origin of mitochondria. However, more direct comparisons of DnaJ and Mdj1 await purification of the mitochondrial DnaJ. 3.3. DnaJs of the ER: Sec63, Scj1 and Jem1 Two DnaJs of the ER have been extensively studied. Sec63, an essential integral membrane protein with its “J” domain exposed to the lumenal side of the membrane and Scj1, a soluble protein of the lumen. Recently a third DnaJ of the ER, Jem1, has been identified (Nishikawa and Endo, 1997). Like Sec63, Jem1 is localized to the ER membrane facing the lumen. jem1 cells are defective in karyogamy, as are kar2 mutants, suggesting that Jem1 may function together with Kar2 in nuclear fusion. jem1 cells are temperature-sensitive in the presence of an scj1 deletion. Sec63 was originally identified in a screen for mutants defective in translocation of

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proteins from the cytosol across the ER membrane (Sadler et al., 1989; Sanders et al., 1992). Sec63 is related to DnaJ only in a region of about 70 amino acids. This “J” domain is situated between two membrane spanning domains and is exposed to the ER lumen. The Kar2:Sec63 interaction was established through analysis of Kar2-Sec63 complexes isolated from detergent-treated microsomes (Brodsky and Schekman, 1993; Scidmore et al., 1993). A mutation resulting in the alteration of a highly conserved alanine of the “J” domain to a threonine resulted in an unstable complex. In addition, the wild-type complex could be disrupted by the addition of an ATP analogue, indicative of a dynamic interaction modulated by nucleotide binding or hydrolysis. That the interaction between Sec63 and Kar2 is important is underscored by the finding that kar2–1 and sec63–1 mutants are synthetically lethal and that second-site suppressors of sec63–1 map to KAR2 (Scidmore et al., 1993). Scj1, which was initially believed to be a mitochondrial protein, is found almost exclusively in the lumen of the ER (Schlenstedt et al., 1995). Unlike Sec63, whose only homology with DnaJ is the “J” domain, Scj1 contains a cysteine-rich region and a glycine-rich region as well as a “J” domain. Scj1 was originally isolated as a gene which in multiple copies causes mislocalization of a protein that should have been targeted to the nucleus (Blumberg and Silver, 1991). However, strains lacking Scj1 grow as well as wild-type strains under a number of conditions tested. Additional genetic studies suggest that Scj1 interacts functionally with Kar2 (Schlenstedt et al., 1995). kar2–159 is synthetically lethal with scj1; interestingly, kar2–1 is synthetically lethal with sec63–1 but is viable in combination with an SCJ1 deletion. As a test of whether Scj1 interacts directly with Kar2, the “J” domain of Scj1 was swapped for the Sec63 “J” domain. The chimeric Sec63-Scj1 protein could substitute for the wild-type Sec63, maintaining viability of a SEC63 deletion strain. These results suggest that Kar2 interacts with two different DnaJ-type proteins in the ER. Proteins being translocated into the ER may first interact with Kar2 and Sec63, and then, after release from the translocation machinery, with Kar2 and Scj1. This analysis of DnaJ chimeras was extended by substituting the “J” domain of mitochondrial Mdj1 and cytosolic Sis1 into Sec63 (Schlenstedt et al., 1995). Neither chimeric protein was able to substitute for Sec63. However, changing only 3 nonconserved amino acids of the Sis1 “J” domain to those found in Sec63 allowed the Sis1:Sec63 chimera to function. The ability of the Scj1 “J” domain to function in place of that of Sec63, while the “J” domains of Sis1 and Mdj1 cannot, may reflect an evolution in the specificity of Hsp70/DnaJ pairs. The interactions between Hsp70s and DnaJs are believed to be specific, such that each Hsp70 will function only with a specific subset of DnaJs. It is likely that Hsp70/DnaJ pairs have co-evolved to maintain important functional interactions, while diverging from family members that are separated by cellular localization or functional differences. Analysis of the numerous DnaJs which have been identified in the genome, but have yet to be functionally analyzed may be required before the full range of interactions between Hsp70s and DnaJs is appreciated. 3.4. Other DnaJs: Xdj1 and Caj1 The basic characterization of two other genes encoding proteins with significant

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similarity to the “J” domain, XDJ1 and CAJ1, has been reported. XDJ1, identified based on its sequence similarity to DnaJ, encodes a protein with both a glycine-rich and a cysteine-rich domain as well as the characteristic “J” domain (Schwarz et al., 1994). Strains lacking Xdj1 have no obvious phenotype and expression from XDJ1 was not detected at either the RNA or protein level. Therefore, it is likely that XDJ1 is expressed at an extremely low level or under a particular condition which was not analyzed in the study. Caj1 was isolated due to its propensity to bind calmodulin (Mukai et al., 1993); Ssb1, Sse1 and Sse2 were isolated in the same study. Caj1 contains no glycine-rich domain, nor a cysteine-rich domain. However, the middle portion of the protein contains a periodic repeat of leucine residues, potentially a leucine zipper. The growth properties of a CAJ1 disruption mutant were indistinguishable from wild-type.

4. Mge1, A GrpE-RELATED GENE OF THE MITOCHONDRIAL MATRIX In addition to its interaction with DnaJ, DnaK physically interacts with GrpE, a nucleotide exchange factor that promotes the release of bound ATP or ADP (Johnson et al., 1989; Buchberger et al., 1994) (see chapter by Buchberger et al.). Functionally, the ability of GrpE to promote ADP release is more important, as it allows the protein to exchange ADP for ATP, which is present in higher concentrations in the cell. Although there have been thorough searches for GrpE homologues in all cellular compartments of eucaryotes, such proteins have only been found in the matrix of mitochondria. It is possible that the fundamental properties of the cytosolic and ER Hsp70s obviate the need for such release factors. There are several pieces of evidence that suggest cytosolic Hsp70s may in fact be biochemically different from procaryotic or mitochondrial Hsp70s. For example, although the intrinsic ATPase activities of the cytosolic yeast Ssa1 and E. coli DnaK are similar, the ability of Ydj1 to stimulate the Ssa1-ATPase activity is significantly greater (Cyr and Douglas, 1994; Ziegelhoffer et al., 1995) than that reported for the DnaK-DnaJ system, in which GrpE is also required for maximal stimulation (Liberek et al., 1991). Furthermore, polypeptide substrates decrease the stability of nucleotide binding to cytosolic Hsp70 (Sadis and Hightower, 1992; Ziegelhoffer et al., 1995), an effect not observed with DnaK Interestingly, a protein termed Hip, which actually stabilizes the Hsc70-ADP complex, has been isolated in mammalian cells (Höhfeld et al., 1995; see chapters by Ha et al., and Buchberger et al.). Most compelling is the fact that cytosolic Hsp70s require only an appropriate DnaJ homologue for refolding activity, while DnaK requires both DnaJ and GrpE. Together this evidence supports a circumstantial case for the absence of a nucleotide exchange factor for Hsp70s in the cytosol. Alternatively, however, nucleotide exchange factors may exist, but be unrelated in sequence to GrpE, and thus have escaped detection up to this point. The best characterized mitochondrial GrpE-related protein is that of S. cerevisiae, called Mge1 or Yge1. MGE1 was first identified on the basis of its similarity in sequence to E. coli grpE (Bolliger et al., 1994; Laloraya et al., 1994), as well as its ability, in multiple copies, to promote staurosporine resistance (Ikeda et al., 1994). MGE1 is an essential gene encoding a protein that is 34% identical to GrpE. Temperature-sensitive

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mutations in MGE1 result in delayed translocation of precursors as well as impaired maturation of imported proteins (Laloraya et al., 1995; Westerman et al., 1995). It is quite certain that Mge1 and Ssc1 function together in the translocation process. Mge1 can be quantitatively co-immunoprecipitated with Ssc1 in isolated mitochondria (Ungermann et al., 1994), and both Mge1 and Ssc1 can be co-immunoprecipitated with a precursor protein that is trapped at the site of import (Voos et al., 1994). The role of Mge1 in this process is discussed in more detail in the chapter by Dekker and Pfanner. Using purified components, Mge1 has been shown to stimulate the release of both ATP and ADP from Ssc1 (Miao et al., 1996). Like GrpE’s interaction with DnaK, Mge1 stably associates with Ssc1 in vitro and this interaction is resistant to high salt, but disrupted by the addition of ATP. The temperature sensitive mutant ssc1–3 produces a protein that fails to stably interact with Mge1, and Mge1 cannot stimulate the release of ATP from this mutant protein. The change in the ssc1–3 protein lies very close to the position of an analogous loop structure on the surface of DnaK which has been implicated in its interaction with GrpE (Buchberger et al., 1994). Deletion of this loop in Ssc1 eliminates detectable interaction with Mge1 (Miao et al., 1996). Interestingly, two mutants with a single amino acid change in this loop do not eliminate the stable binding with Mge1, yet do not stimulate the release of nucleotide. Therefore this loop is likely to be important not only for binding to Mge1 but also for generating a conformational change necessary for Mge1-induced nucleotide release.

5. FUNCTIONAL INTERACTIONS AMONG CLASSES OF CHAPERONES IN PROTEIN FOLDING As discussed in some detail above, Hsp70s and DnaJs function together. Recent data support the idea that these two chaperones also function with other classes of chaperones as well. For example, both Hsp70 and DnaJ function with chaperones of the Hsp90 class (Bohen et al., 1995). It has been known for some time that in mammalian cells Hsp90s function in the maturation of transcription factors of the steroid hormone receptor class. In addition, Hsp90 associates with the oncogenic tyrosine kinase p60v-src during its maturation. More recently, both Hsp70s and DnaJs have also been shown to associate with steroid hormone receptors during the maturation process (Smith and Toft, 1993). Maturation of both receptors and p60v-src have been studied in a heterologous yeast system. Expression of active receptors in the heterologous yeast system requires the endogenous Hsp90 as well as Ydj1 (Picard et al., 1990; Bohen and Yamamoto, 1993). Also, expression of p60s-src in yeast cells results in lethality (Brugge et al., 1983; Dey et al., 1996; Kimura et al., 1995), but both HSP90 and YDJ1 mutants are resistant to the toxic effects of p60v-src (Xu and Lindquist, 1993). Genetic evidence indicates an overlap in function between the SSA proteins and Hsp104, a molecular chaperone of the Clp/Hsp100 class (see chapters by Maurizi et al., and Lindquist et al.). Hsp104 has been shown to be critical for the reactivation of denatured luciferase in vivo (Parsell et al., 1994), while Ssa proteins can carry out this function in vitro. Phenotypic analysis of strains containing mutations in both SSA genes and HSP104 have also been informative. For example, overexpression of Ssa1 in cells

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lacking Hsp104 partially restores thermotolerance to hsp104 cells, while the survival of ssa1 ssa3 ssa4 hsp104 cells is 100 fold less than hsp104 cells after exposure to high temperatures (Sanchez et al., 1993). In addition, ssa1 ssa2 cells grow more slowly than wild-type but reasonably well at 30°C, while hsp104 mutants show no detectable growth defect. However, ssa1 ssa2 hsp104 cells are barely viable under the same conditions. A more detailed understanding of the functional relationship between the Ssas and Hsp104 awaits further experimentation.

6. CONCLUSIONS As described in this chapter, we have learned a great deal about the function of molecular chaperones of the Hsp70 and DnaJ classes since their first identification. We now know that the Hsp70s of yeast fall into multigene families, that each well-studied family participates in multiple cellular functions, and that they function with other classes of chaperone. Both biochemical and genetic experiments indicate that many of the Hsp70 and DnaJ chaperones within a cell are functionally distinct. The recent completion of the sequence of the yeast genome has allowed the identification of all members of the established classes of molecular chaperones in this organism. This information should provide the opportunity in the coming years to map out the complex web of proteinprotein interactions, cooperative roles, and cellular functions among the full complement of molecular chaperones in a single eukaryotic cell. Only then will we truly begin to understand the intricacies of the roles of chaperones in cell physiology.

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8. FUNCTIONS IN DEVELOPMENT MICHEL MORANGE Unité de Génétique Moléculaire, Département de Biologie, Ecole Normale Supérieure, 46 rue d’Ulm, 75230 Paris Cedex 05, France

1. Introduction 2. The Stress-inducibility of Chaperones During Development 3. The Different Experimental Systems Used to Follow Chaperone Expression During Embryogenesis 4. Chaperones as actors in development 5. Mechanisms Controlling the Developmental Expression of Chaperones 6. Research Perspectives 7. Acknowledgements 8. References

1. INTRODUCTION The study of the variations in expression of chaperones during development, and the role of these variations, is at the same time trivial and difficult. Trivial because it is well known that the expression of chaperones differs widely from one adult tissue to another (Tanguay et al., 1993) and that these differences are established during development. Moreover, it has been extensively documented that the synthesis of chaperones is dependent upon metabolic conditions (Lanks, 1986) and is not uniform during the different phases of the cell cycle (Wu and Morimoto, 1985; Milarski and Morimoto, 1985; Jérome et al., 1993). Since the embryo is submitted during development to dramatic variations in metabolism, and given that the duration of the different cell cycle phases varies widely in embryonic cells, it can be expected that the expression of chaperones will vary during development. If one considers the functions of chaperones in protein folding (Hartl, 1996), translocation across membranes but also protein degradation (Hayes and Dice, 1996), it is also obvious that the concentrations of chaperones and heat shock proteins (HSPs) will vary during development, in parallel with protein synthesis, cell remodeling or secretion (as during the formation of the extracellular matrix). As we shall briefly show below, a large number of observations in different organisms provides evidence for variations in chaperone levels and synthesis during embryogenesis (Heikkila, 1993; Morange, 1997). However these observations do not satisfy

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developmental biologists, who are not interested in variations in the levels of expression of housekeeping genes, but are looking for genes specifically involved in crucial steps of embryogenesis. As we shall see, only now are we beginning to have data clearly demonstrating that chaperones play a major structural and/or regulatory role at some specific steps of development. The numerous early studies describing the expression of chaperones during development were necessary steps to point out these crucial functions. These studies required the preliminary characterization of the chaperone-encoding genes in the different organisms studied. Due to the complexity of the genetic organization of heat shock genes, this has been a long and tedious task which is still not fully completed. We will rapidly present the main developmental systems used, outlining the specific interest of each and the most important observations which have emerged. We will then focus on three different systems which are closest to providing data showing the crucial requirement of chaperones in development. We will conclude with a few words about the mechanisms which might be responsible for the specific expression of chaperones during embryogenesis and the developments we can expect in the near future.

2. THE STRESS-INDUCIBILITY OF CHAPERONES DURING DEVELOPMENT The most meaningful observations for developmental biologists concern the spontaneous expression of chaperones during embryogenesis. However, chaperones are also stress proteins, able to participate in the repair of cell damage (Parsell and Lindquist, 1993; Freeman and Morimoto, 1996). Two major arguments support the study of variations in chaperone (HSP) inducibility during development: 1) Stress treatments lead to uneven alteration of normal embryonic development. Some tissues and embryological processes are more sensitive to stress than others: for instance, in mammals, neural tube closure can be perturbed by many different chemical or physical stresses, leading to serious abnormalities (Walsh et al., 1991; Walsh et al., 1997). This sensitivity is probably due to the perturbation in relative amounts of important regulatory proteins which follows stress (Welte et al., 1995). 2) The inducibility of the stress proteins is different in the different tissues of an adult, but also in the different tissues of an embryo and at different stages of development. As a general rule, the early embryo is poorly responsive to stress. This is true from plants (reviewed in Vierling, 1991) and Drosophila to mammals. In mammals, the fully grown oocyte is unresponsive to stress and heat shock (Curci et al., 1991), as is the early embryo before the blastocyst stage (Morange et al., 1984). The uninducibility of heat shock genes is also observed in some differentiated cell lines where it might be linked with methylation (Gorzowski et al., 1995). One major goal of embryologists is to try to relate these two groups of observations and to elucidate the role of induced HSPs and chaperones in the protection of the embryo. The answer is not straightforward. The presence of a strong HSP induction in adult or embryonic tissues does not prevent these tissues from being a target for the stress: everything happens as if the intensity of HSP induction was more a reflection of the damages supported by the cells than an adaptive response of these cells (Brown, 1994). In

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some cases, such as in mammalian testis, it has even been suggested that the accumulation of HSPs might have deleterious effects, blocking cell division or differentiation (Sarge, 1995).

3. THE DIFFERENT EXPERIMENTAL SYSTEMS USED TO FOLLOW CHAPERONE EXPRESSION DURING EMBRYOGENESIS We will only briefly discuss HSP and chaperone expression during plant development. The abundance and diversity of small HSPs, but also of other stress proteins such as GRP78, the presence of members of each HSP family in the different cell compartments (Boston et al., 1996), the occurrence of different heat shock transcription factors, some of which are themselves HSPs (Nover et al., 1996), make this study particularly difficult. However, it is obvious that the expression of the different HSPs, as well as their intracellular distribution, both in the presence and absence of a stress, is highly variable during development and in the different tissues of the plant. It is remarkable how diverse physiological conditions (stress?) are able to alter HSP synthesis: light, dehydration or starvation, addition of plant hormones (abscisic and gibberellic acids), mechanical wounds, as well as the circadian rhythm. The mechanisms involved in these variations of expression appear as diverse as the conditions leading to them (regulation by translational control or modification of protein stability as well as by transcription). Even if it clearly appears that HSP synthesis is “part of the developmental program” of plants (Wehmeyer et al., 1996), it has not yet been demonstrated that HSPs are essential components of this developmental program. In animals, the best studied system is, for historical reasons, Drosophila. In this organism, there exist four genes coding for the small HSPs, whose inducibility and spontaneous expression vary dramatically during development (Haass et al., 1990). For instance, HSP23 is specifically expressed during embryogenesis in glial cells identified as Midline glial cells (Tanguay, 1989). Similar observations have been reported for the three other small hsps which are expressed in characteristic structures of the larva and adult, mainly in the central nervous system and gonads (Arrigo and Mehlen, 1994; Michaud et al., 1997). The most puzzling observations concern the localization of hsp83 mRNA during oogenesis and early embryogenesis. This mRNA is transferred from nurse cells to the oocyte. It is evenly distributed throughout the embryo during the initial divisions, and then remains concentrated at the posterior pole by a selective stabilization process. It is then taken up by the pole cells. hsp83 gene is also transcribed from the zygotic genome at the syncitial blastoderm stage in the anterior third of the embryo (Ding et al., 1993). Both maternal and zygotic expressions are under the strict control of important developmental genes of Drosophila, oskar for maternal gene expression and bicoid for the zygotic one, suggesting that the control of HSP83 synthesis is critically important at these early stages of embryogenesis. Another developmental system in which the expression of chaperones has been studied at the very early stages is the amphibian embryo (Heikkila, 1993; Herberts et al., 1993; Heikkila et al., 1997). The need to isolate specific probes and to prepare antibodies has so far prevented a full use of this system, which is particularly well suited to the study of the

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first steps of embryogenesis. It has been clearly demonstrated that the expression and inducibility of heat shock genes is controlled during oogenesis and early embryonic development by chromatin state, as well as by specific activators (Landsberger et al., 1995a and b; Ali et al., 1993). Stabilization of mRNAs might also be important for the progressive appearance of the heat-inducible expression of the small HSPs (Ohan and Heikkila, 1995). One recent observation has underlined the spontaneous variations in the level of one small HSP (HSP30) (Helbing et al., 1996) which occur during development. Mouse embryo has also been studied at very early stages for a differential induced or spontaneous expression of HSPs and chaperones (Mezger et al., 1991). The preimplantation phase has been most closely examined and data on the expression of the major HSPs and chaperones during mouse post-implantation development is still incomplete. One of the earliest events associated with the onset of zygotic genome transcription (Schultz, 1993) is the transient expression of the hsp70 gene (Bensaude et al., 1983; Manejwala et al., 1991; Christians et al., 1997a). Both maternal and paternal genomes contribute to this synthesis as demonstrated with parthenogenotes and androgenotes (Barra and Renard, 1988). The hsc70 gene is also expressed but, in contrast to hsp70, its expression remains high during later development (Morange et al., 1984; Giebel et al., 1988). Transgenic animals harboring a hybrid hsp70.1-luciferasegene reproduce this transient expression (Christians et al., 1995), definitely demonstrating that this expression is not an artefact of manipulation. Two mechanisms have been proposed for this transient expression: activation by the maternal HSF1 factor, which is abundant and localized in the nucleus at this stage of development (Christians et al., 1997b), or the action of the basal promoter elements (Bevilacqua et al., 1995; Bevilacqua et al., 1997). The physiological significance of this expression remains unknown and the generalization of this phenomenon to other mammals (and vertebrates) is problematic. hsp70 is also spontaneously expressed later in development, first in the extra-embryonic tissues, and then in specific tissues of the embryo (Kothary et al., 1987; Loones et al., 1997). The precise nature of the tissues or cells concerned, and the meaning of these expressions, await further studies. The expression in these tissues contrasts with the very low constitutive expression of HSP70 in most cells. A transient expression of HSP70 during in vitro differentiation of the human cell line K562 has been described (Theodorakis et al., 1989; Sistonen et al., 1992 and 1994) and shown to also occur during normal hematopoiesis (Singh andYu, 1984). The generalization of this observation to other mammals (in particular mouse) is once again problematic. Both HSC70 and HSP90 are ubiquitously expressed, in the adult as well as in embryonic tissues of mice. These two proteins, as well as HSP90 and HSP60, are also expressed at a very high level at the blastocyst stage and during the period immediately following implantation of the embryo (Bensaude and Morange, 1983). A remarkably high level of HSC70 and HSP90 can be found during mid-gestation in the nervous system and in neural crest cells, as well as in the hypertrophic chondrocytes during bone morphogenesis (Loones, Figure 1; Loones et al., 1997). In the first case, this high expression persists in the adult (Itoh et al., 1993; Izumoto

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Figure 1 HSP distribution in the ulna cartilage primordium of a day 15.5 mouse embryo, revealed with peroxidase-coupled second antibody, a) HSP90 is expressed all along the chondrocyte differentiation pathway, in particular in the hypertrophic chondrocytes. b) HSC70 expression during chondrogenesis is limited to hypertrophic chondrocytes. scale bar: 100 m

and Herbert, 1993) and might play a crucial role in the protection of the nervous system against aggressions (Nowak et al., 1994). Three observations on the mammalian embryo deserve further study and will be discussed below. 1) The existence of a Heat Shock Transcription Factor (HSF2) specifically expressed during most of embryogenesis, as well as during hematopoiesis or spermatogenesis. 2) The very specific expression of the small HSP/ -crystallins in the heart, muscle and some neural structures in the brain during development. 3) The highly specific expression of minor members of the HSP/HSC70 family during spermatogenesis. The mouse embryo surely has further surprises and discoveries in store since the potential of this developmental system (existence of multiple mutations in the developmental genes, possibility of gene disruption by homologous recombination, existence of many in vitro differentiation systems) is far from being fully exploited.

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The last system to have provided some clues on the role of chaperones and HSPs during development is the zebrafish. This organism is presently the best adapted to the combined use of genetic and developmental techniques, a sort of hybrid of what is possible in the mouse and in the amphibian. The work is still in its initial phases and the characterization of the different HSP and chaperone genes is not yet complete. For instance, nothing is presently known about the structure and expression of the small HSPs in this organism. However by whole mount in situ hybridization analysis, Sass et al. (1996) have revealed the very specific localization of zebrafish hsp90 mRNA in muscles during development (see below). hsp90 mRNA does not show the same distribution, but is detectable within the central nervous system at control temperatures. Finally, despite its interest for developmental biology, C. elegans has not so far furnished outstanding results concerning the role of chaperones and HSPs in development. However, it has confirmed the heat-inducible tissue specific expression of small HSPs during development (Stringham et al., 1992), quite reminiscent of what is observed in Drosophila.

4. CHAPERONES AS ACTORS IN DEVELOPMENT Previous data clearly demonstrate the variations in the levels of HSP and chaperone gene expression during embryogenesis. We would now like to describe in more detail three selected situations in which additional experimental data support the hypothesis that HSPs and chaperones are not only parts of the developmental program, but important actors in these programs. The first concerns the expression of hsp90during zebrafish development (Sass et al., 1996). The level of expression of this gene is weak and limited to a small subset of cells within the pre-somitic mesoderm, later in the somites and pectoral fin buds of the developing embryo. The expression of the gene coding for one of the Myogenic Regulatory Factors (MyoD) coincides with the expression of hsp90 in the somites and pectoral fin buds. The expressions of hsp90 and myoD are down-regulated in adult muscle. HSP90 has been shown to enhance the DNA-binding activity of murine MyoD synthesized in vitro (Shaknovich et al., 1992), as well as of the MyoD/E12 heterodim