Molecular Chaperones of the Endoplasmic Reticulum [1 ed.] 9781616684594, 9781616681739

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Copyright © 2010. Nova Science Publishers, Incorporated. All rights reserved. Molecular Chaperones of the Endoplasmic Reticulum, Nova Science Publishers, Incorporated, 2010. ProQuest Ebook Central,

Copyright © 2010. Nova Science Publishers, Incorporated. All rights reserved. Molecular Chaperones of the Endoplasmic Reticulum, Nova Science Publishers, Incorporated, 2010. ProQuest Ebook Central,

PROTEIN SCIENCE AND ENGINEERING

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MOLECULAR CHAPERONES OF THE ENDOPLASMIC RETICULUM

No part of this digital document may be reproduced, stored in a retrieval system or transmitted in any form or by any means. The publisher has taken reasonable care in the preparation of this digital document, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information Molecular Chaperones of the Endoplasmic Reticulum, Nova Science Publishers, Incorporated, 2010. ProQuest Ebook contained herein. This digital document is sold with the clear understanding that the publisher is not engaged in

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Molecular Chaperones of the Endoplasmic Reticulum Martin Schröder 2010. ISBN: 978-1-61668-173-9 Molecular Chaperones of the Endoplasmic Reticulum Martin Schröder 2010. ISBN: 978-1-61668-459-4 (E-book) Insulin Receptor Substrate Signaling in the Central Nervous System Susanna Freude and Markus Schubert 2010. ISBN: 978-1-61668-255-2 Insulin Receptor Substrate Signaling in the Central Nervous System Susanna Freude and Markus Schubert 2010. ISBN: 978-1-61668-615-4 (E-book)

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PROTEIN SCIENCE AND ENGINEERING

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MOLECULAR CHAPERONES OF THE ENDOPLASMIC RETICULUM

MARTIN SCHRÖDER

Nova Science Publishers, Inc. New York

Molecular Chaperones of the Endoplasmic Reticulum, Nova Science Publishers, Incorporated, 2010. ProQuest Ebook

Copyright © 2010 by Nova Science Publishers, Inc. All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers’ use of, or reliance upon, this material.

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Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. LIBRARY OF CONGRESS CATALOGING-IN-PUBLICATION DATA Molecular chaperones of the endoplasmic reticulum / Martin Schrvder. p. ; cm. Includes bibliographical references and index. ISBN H%RRN 1. Molecular chaperones. 2. Endoplasmic reticulum. I. Title. [DNLM: 1. Endoplasmic Reticulum--physiology. 2. Molecular Chaperones--physiology. 3. Protein Folding. QU 350 S381m 2010] QP552.M64S37 2010 571.6'5--dc22 2010003161

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CONTENTS

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Preface

vii

Chapter 1

Introduction

1

Chapter 2

Physicochemical Principles Governing Protein Folding

9

Chapter 3

The ER

15

Chapter 4

Molecular Chaperones of the ER

23

Chapter 5

Conclusion

67

Acknowledgements

69

References

71

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PREFACE An important function of the endoplasmic reticulum (ER) is the folding of newly synthesized polypeptide chains. Several general and specialized chaperones operate in the ER to assist protein folding. The HSP70 class chaperones BiP and GRP170, their DnaJ and GrpE co-chaperones, the HSP90 class chaperone GRP94, and the lectin chaperones calnexin, calmegin, and calreticulin are the major general chaperones of the ER. HSP47 is an example of a specialized chaperone required for collagen folding. Protein foldases such as protein disulfide isomerases and cis-trans peptidyl prolyl isomerases catalyze two otherwise ratelimiting steps in protein folding, formation and isomerization of disulfide bonds and the cis-trans isomerization of peptidyl-prolyl bonds. Chaperone-assisted folding of the polypeptide chain, disulfide bond formation and posttranslational modification of the polypeptide chain, most notably by addition of large oligosaccharides to exposed asparagine residues by oligosaccharyl transferase interact in the ER in a complex manner to produce properly folded proteins. In addition to functioning in protein folding chaperones participate in other vital functions of the ER, such as import of polypeptide chains by translocation, quality control of protein folding status and targeting of permanently unfolded proteins for retrograde translocation and proteasomal degradation, and regulation of the activity of ER Ca2+ channels. In this book I review our current knowledge of protein folding in the ER and of the functions of ER-resident molecular chaperones in protein folding in the ER.

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Chapter 1

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INTRODUCTION Molecular chaperones are a ubiquitous class of proteins that play important roles in protein folding and protection of cells from several stresses associated with disruption of the native three dimensional structures of proteins [1-3]. Protein folding is governed by the hydrophobic effect resulting in burial of hydrophobic amino acid side chains in the center of the protein [46]. Exposure of these hydrophobic stretches at the surface of a protein is highly toxic to cells as it leads to aggregation of proteins within a cell, aberrant protein-protein interactions that inactivate essential components of the cellular machinery, and disruption of phospholipid bilayers [7]. Molecular chaperones detect these hydrophobic surface areas and shield them from interaction with other proteins and biological membranes [8-10]. Constitutive expression of molecular chaperones in the two major protein folding compartments of an eukaryotic cell, the cytosol and the endoplasmic reticulum (ER), reflects their function in folding of newly synthesized polypeptide chains and translocation of polypeptide chains across lipid bilayers. Many molecular chaperones have been identified as proteins induced upon heat stress, a condition leading to protein unfolding, and have originally been classified as heat shock proteins (HSPs) [11]. Several classes of molecular chaperones or HSPs have been distinguished based on the molecular masses of their founding members, for example small HSPs (sHSPs, IbpA/IbpB in Escherichia coli), HSP60/HSP10 (GroEL/GroES in E. coli), HSP70/HSP40 (DnaK/DnaJ in E. coli), HSP90 (HtpG in E. coli), and HSP100/CLP (ClpB in E. coli). Heat shock also induces expression of several proteases which by degrading unfolded proteins protect cells from accumulation of unfolded proteins. Induction of chaperones and proteases by heat shock has been termed the heat shock response. Other

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stresses, such as oxidative stress or viral infections associated with immense production of viral proteins, also cause a heat shock response [12, 13]. The heat shock response is ubiquitous and affects all organelles of an eukaryotic cell. However, organelle-specific heat shock responses, such as a mitochondrial heat shock or unfolded protein response (UPR), or the UPR of the ER, selectively induce chaperones and proteases of these organelles. The ER is a major protein folding compartment in eukaryotic cells second only to the cytosol. Its folding clients are secretory proteins, transmembrane proteins of the plasma membrane, and proteins residing in the secretory pathway. Not surprisingly, the ER harbors many chaperone classes that are also found in the cytosol, for example the HSP70 chaperones heavy chain binding protein (BiP) [14]/glucose-regulated protein of 78 kDa (GRP78) [1517]/karyogamy 2 protein (Kar2p) [18], luminal HSP seventy 1 protein (Lhs1p) [19]/chaperone in the ER 1 protein (Cer1p) [20]/Ssi1p [21]/oxygen-regulated protein of 150 kDa (ORP150) [22-24]/GRP170 [25-28], the HSP90 chaperone GRP94 [15, 16, 29]/adenotin [30, 31]/endoplasmin [32]/tumor rejection antigen glycoprotein of 96 kDa (gp96) [33, 34]/ER protein of 99 kDa (ERp99) [35]/HSP108 [36]/Ca2+-binding protein 4 [37]/protein kinase of 80 kDa [38], and the HSP40 chaperones ER DnaJ 1 (ERdj1)/murine DnaJ-like protein 1 (MTJ1) [39, 40], ERdj2/secretory 63 protein (Sec63p) [41-44], ERdj3 [45, 46]/human ER associated DnaJ (HEDJ) [47]/Saccharomyces cerevisiae DnaJ 1 protein (Scj1p) [48, 49], ERdj4 [50]/microvascular endothelial differentiation gene 1 (MDG1) [51-53], ERdj5/Erj5p/JPDI [54-57], DnaJ-like protein of the ER membrane (Jem1p) [58], and p58IPK/DnaJc3 [59, 60]. sHSPs or HSP60s of the ER are currently unknown. Two major differences between protein folding in the ER and the cytosol exist. First, many proteins targeted to the ER undergo several post-translational modifications, of which N-linked glycosylation plays important roles in protein folding in the ER. Therefore, the ER contains specialized lectin chaperones – calnexin [61]/p88 [62, 63], calreticulin [64, 65]/Ca2+-binding protein of 63 kDa (CAB-63) [66]/calregulin [67]/Ca2+-regulated protein of 55 kDa (CRP55) [65, 68]/high affinity Ca2+binding protein (HACBP) [69]/Ca2+-binding protein 3 (CaBP3) [70], and the testis-specific lectin chaperone calmegin [71-73]/calnexin-t [74, 75] – that recognize the N-linked glycosylation status of proteins folding in the ER. Second, disulfide formation between two cysteine thiol groups occurs in the ER, a protein folding reaction catalyzed by a large family of protein disulfide isomerases (PDIs, Figure 1) [76, 77]. PDIs and cis-trans peptidyl prolyl isomerases (PPIs), a ubiquitous family of proteins whose members catalyze the cis-trans isomerization of peptidyl-prolyl bonds (Figure 2), are often called

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Introduction

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protein foldases. Conformational, chaperone-assisted, and oxidative folding intersect [78], which is reflected by several PDIs possessing chaperone activity. These ER luminal chaperones and foldases also play important roles in cellular processes closely associated with protein folding: Translocation of newly synthesized polypeptide chains into the ER [79, 80], recognition of unfolded polypeptide chains for retrograde translocation out of the ER and degradation by the proteasome in a process called ER associated protein degradation (ERAD) [81-83], and activation of signaling pathways termed the UPR that coordinate protein folding demand and capacity of the ER (Figure 3) [84, 85]. Here I will review how molecular chaperones of the ER assist protein folding and detect unfolded proteins in the ER.

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Introduction

Figure 1. Oxidative protein folding in the ER. (A) Disulfide bond formation and isomerization reactions catalyzed by prototypic PDI. (B) PDI families in

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Saccharomyces cerevisiae (top), Homo sapiens (middle), and Arabidopsis thaliana (bottom). Thioredoxin-like domains are indicated by white or hatched rectangles. Catalytic centers are indicated in single letter amino acid code in the a and a’ domains. The b and b’ domains are indicated by hatched rectangles. Black rectangles represent transmembrane domains, and black rectangles with the lettering ‘SP’ signal peptides. ER retrieval sequences are indicated at the C-terminus in single letter amino acid code. Numbers represent the lengths of the proproteins. (C) Rooted, phylogenetic tree of A. thaliana (At), H. sapiens (Hs), and S. cerevisiae (Sc) PDIs generated with PHYLIP 3.68. The GenBank accession numbers are: AtPDI1 (CAB41088), AtPDI2 (BAB09837), AtPDI3 (AAG51554), AtPDI4 (BAB02677), AtPDI5 (AAD41430), AtPDI6 (NM_106400), AtPDI7 (Q9T042), AtPDI8 (AAK62431), AtPDI9 (AAB91984), AtPDI10 (AAF40463), AtPDI11 (NP_001078074), AtPDI12 (NP_566664), Eps1p (P40557), ERdj5 (Q8IXB1), ERp18 (O95881), ERp27 (Q96DN0), ERp28 (P30040), ERp44 (Q9BS26), ERp46 (Q8NBS9), ERp57 (P30101), ERp72 (P13667), Eug1p (P32474), Mpd1p (Q12404), Mpd2p (Q99316), P5 (Q15084), PDI (P07237), Pdi1p (P17967), PDILT (Q8N807), PDIp (Q13087), PDIr (Q14554), TMX (Q9H3N1), TMX2 (Q9Y320), and TMX3 (Q96JJ7). A. thaliana PDIs are labeled according to [554].

Figure 2. Cis-trans peptidyl prolyl bond isomerization. For most peptide bonds the cis configuration is disfavored because of steric hindrance between the two Cα atoms. If the C-terminal amino acid is proline, this energetic difference between the cis and trans configurations is largely eliminated. Therefore, peptidyl-prolyl bonds exist both in the trans and cis configuration. The partially double bonded character of the peptide bond hinders free rotation around the carbonyl C – N bond (EA – activation energy, t1/2 – half life). Catalysis by cis-trans peptidyl prolyl isomerases decreases EA and increases the rate of conversion of the trans into the cis configuration and vice versa.

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Introduction

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Figure 3. Biological functions of molecular chaperones in the ER.

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Chapter 2

PHYSICOCHEMICAL PRINCIPLES GOVERNING PROTEIN FOLDING

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As for any other chemical reaction protein folding reactions have to be energetically favorable under biochemical conditions, i.e. characterized by a negative free Gibbs enthalpy, and have to be reasonably fast to be productive in the life time of a cell. Thermodynamic and kinetic implications on protein folding in a cellular environment are summarized in the following paragraphs.

2.1. THERMODYNAMICS OF PROTEIN FOLDING Each conformation of a given polypeptide chain is characterized by a certain free Gibbs enthalpy. In this way the primary structure of a protein determines the protein’s folding landscape in which the free enthalpy is plotted against each conformation, characterized by the fractions of native and nonnative interactions of its amino acid side chains [5]. In an aqueous environment the Gibbs free energy of the water-protein system is largely determined by the number of hydrophobic amino acid residues exposed at the surface of the protein [86, 87]. The more hydrophobic residues are exposed at the surface, the higher the Gibbs free energy of the water-protein system. This effect – also known as the hydrophobic effect - is largely an entropic effect, because minimization of the surface between hydrophobic molecules and the bulk water phase liberates water molecules from structured ‘cages’ formed at this surface to maximize hydrogen bonding between water molecules close to the surface. With increasing temperature the entropic consequences of the

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hydrophobic effect become less pronounced, because water molecules become more mobile as hydrogen bonds between water molecules are broken. The cellular consequence of elevated temperature is the heat shock response caused by entropic destabilization of protein structure. Exposure of polar groups at the surface, on the other hand, results in a much less pronounced decrease in the Gibbs free energy of the system, because the energetic and entropic differences between interactions of a water molecule with other water molecules or hydrophilic amino acid side chains are small. To be energetically favorable, protein folding hides as many hydrophobic amino acids in the center of the protein as possible, thereby decreasing the surface between hydrophobic amino acid side chains and the surrounding water and, in turn, the Gibbs free energy of the water-protein system by increasing its entropy. On an energy landscape a polypeptide chain folds along pathways leading to ever decreasing free energies until the conformation with the lowest free energy is reached. Theoretical calculations support the idea that this conformation is the native conformation for most proteins [88, 89]. These biophysical considerations on how the thermodynamics of the water-protein system energetically govern protein folding are summarized by Anfinsen’s dogma, stating that the information for the native conformation of a protein is encoded by its primary structure [90].

2.2. KINETICS OF PROTEIN FOLDING Thermodynamic considerations do not inform on the kinetics of protein folding. Levinthal’s paradox states that even for a protein consisting of only 100 amino acids too many conformations are possible if a random sampling mechanism probing each conformation for only 10-11 s is considered to allow the protein to fold in the life time of the universe [91-93]. A solution to Levinthal’s paradox is constriction of the folding choices of the unfolded protein by assuming that proteins fold along discrete pathways very much as small molecule reactions proceed along a few pathways. However, the complex nature of proteins and the similar, but not identical, physiochemical properties of their constituents, make such an easy picture unlikely. On the energy landscape discussed above a hydrophobic collapse initiates protein folding, especially in larger proteins. By progressing toward conformations with decreasing free energies sufficient conformational constraints are retained to solve Levinthals’ paradox. In the hydrophobic collapse hydrophobic side chains are buried in the center of the protein shielding them from surrounding

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Physicochemical Principles Governing Protein Folding

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water [6]. An important feature of the hydrophobic collapse is burial of charged and hydrophilic amino acids in the hydrophobic core of the initial structure formed by the hydrophobic collapse. These charged and hydrophilic residues provide energy signatures to individual conformations and influence the directionality of further folding of this intermediate. Without formation of salt bridges, disulfide bonds, or hydrogen bonds in the hydrophobic core energetic differences between individual conformations would be small and a thermodynamic driving force and directionality of further folding would not exist [6]. Secondary structure elements in proteins fold very fast. For example, βturns or α-helices are formed within 0.1-10 µs [94]. Structurally simple and small proteins fold in less than 50 µs [95-97], whereas more complex structures, e.g. β-sheets, fold more slowly [98]. Modules or domains of larger proteins fold independently of each other into near native structures [99]. In a final cooperative folding event water is excluded from the protein core and the native structure formed [100]. Translation of mRNAs proceeds at ~2-8 amino acid residues per second [101-106] which is slow compared to formation of secondary structure elements. To form secondary and tertiary structural elements in which residues far apart in the amino acid sequence interact, e.g. β-sheets or disulfide bonds, the preceding residues must be maintained in a folding competent state until the interacting partners are added to the polypeptide chain. The high protein concentration in vivo exacerbates this problem. The protein concentration in the ER reaches ~100 g/l (~2 mM). Assembly of IgG heavy and light chains present in the ER of a plasma cell at concentrations of ~4-6 µM can, in principle, be a diffusion-controlled reaction [6]. The hydrophobic effect is not affected by structural differences in different polypeptide chains and does not distinguish between interactions of hydrophobic amino acid side chains with other hydrophobic amino acid side chains or lipids. As a consequence, aggregation triggered by aberrant hydrophobic interactions between two polypeptide chains and disruption of phospholipid bilayers by unfolded proteins is a significant problem for all cells. To solve these problems cells invented molecular chaperones, classes of proteins that interact with unfolded proteins and shield unfolded proteins from interaction with hydrophobic interaction partners, i.e. other proteins and phospholipid bilayers (Figure 4).

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Figure 4. Competition between folding, chaperone interaction and aggregation of an unfolded protein. An unfolded protein folds through several intermediates into its native state. Aggregation kinetically competes with productive folding, because protein folding is governed by the hydrophobic effect. Interaction with chaperones stabilizes the unfolded conformation and kinetically partitions the unfolded protein toward productive folding. Chaperones recognize hydrophobic surface patches on unfolded proteins and, therefore, efficiently compete with aggregation. Chaperones exist in a high and low affinity state for the substrate. In HSP70 chaperone foldases the high affinity state is terminated by ATP for ADP exchange, resulting in release of the substrate from the chaperone. This allows the unfolded substrate, which is conformationally locked when bound to the chaperone, to continue on its folding pathway.

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2.3. TOXICITY OF PROTEIN AGGREGATES Protein aggregates or partially folded folding intermediates are extremely toxic to cells. Several mechanisms of protein aggregate toxicity are recognized. Protein aggregates may interact with other proteins whose biological function is to interact with other proteins. For example, huntingtin protein interacts with the transcriptional co-activator CBP through glutaminerich stretches present in both proteins, leading to downregulation of transcription of genes controlled by cAMP response elements (CREs) [7, 107112]. Sequestration and inactivation of other transcription factors by polyglutamine stretches has also been reported for other polyglutamine diseases, such as spinocerebellar ataxias 1 and 3, dentatorubral pallidoluysian atrophy, and Kennedy disease [7, 113-115]. A second class of proteins affected in this way are molecular chaperones and proteins of the protein folding quality control machinery, for example the proteasome. HSP70 chaperones are associated with huntingtin [116-118], Lewy bodies formed by aggregated α-synuclein in Parkinson’s disease [7, 119], and intracellular amyloid β (Aβ) aggregates [120]. Protein aggregate toxicity may be mediated by sequestration and inactivation of molecular chaperones. Depletion of molecular chaperones in turn impairs folding of essential, metastable proteins [121]. Consistent with this view is that overexpression of chaperones mitigates protein aggregate toxicity [7, 122]. In addition, one would expect that anterograde and retrograde protein translocation mechanisms at the ER and the mitochondria are affected, because of the involvement of cytosolic chaperones in these processes. Second, proteasomal function is impaired by Lewy bodies and α-synuclein protofibrils [123, 124], in Alzheimer’s disease by Aβ and phosphorylated tau [125-128], Parkinson’s disease [129], and by polyglutamine proteins [130-133], which can at least in part be attributed to direct inhibition of proteasome activity by aggregated proteins. Inhibition of the proteasome abolishes ERAD. Likewise, depletion of cytosolic chaperones may interfere with retrotranslocation of unfolded proteins out of the ER and extraction of stalled, translocating polypeptide chains out of the translocation channel in a process called pre-emptive quality control [134], again inhibiting ERAD and causing ER stress.

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Chapter 3

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THE ER Protein folding in the ER is more complex than in the cytosol mainly because the ER is a separate metabolic compartment surrounded by a phospholipid bilayer [135]. Newly synthesized proteins are transported into the ER by translocation across a phospholipid bilayer and co- and posttranslationally modified by addition of oligosaccharides to asparagine residues in consensus Asn-X-Ser/Thr (X = any amino acid except proline) motifs and by oxidation of cysteine thiol groups to form disulfide bonds. Disulfide bond formation increases the structural rigidity of extracellular proteins and increases their half-life, whereas glycosylation significantly increases the hydrophilicity of proteins. These modifications affect the conformational folding pathway and folding choices of the polypeptide backbone of the protein.

3.1. TOPOLOGY OF THE ER The luminal space of the ER is topologically equivalent to the extracellular space. A hydrophobic signal sequence targets proteins to the ER membrane for co-translational crossing of the ER membrane through the Sec61p complex [136-139]. Posttranslational entry into the ER, once the complete polypeptide chain has been synthesized, is an alternative mechanism to enter the ER. Signal peptidase co-translationally cleaves off the signal peptide. Bacterial signal peptidases process their substrates after translation of ~80% of the polypeptide chain [140, 141], suggesting that the signal peptide is present during initial folding steps of the protein. Indeed, the signal sequence influences the timing of N-linked glycosylation and signal sequence cleavage

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[142]. Inefficient cleavage can result in prolonged interaction of the protein with ER chaperones [143].

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3.2. CHEMICAL COMPOSITION OF THE ER The pH in the ER is near neutral [144] and comparable to the cytosol. In mammalian cells the ER or its specialized counterpart, the sarcoplasmic reticulum (SR) in muscle cells, is the major site for Ca2+ storage. ER luminal free Ca2+ concentrations reach 1-5 mM, compared to 0.1 µM in the cytosol [145, 146]. ER luminal Ca2+ concentrations rapidly and frequently fluctuate as the ER Ca2+ pool is mobilized during intracellular signaling (Figure 5) [147]. Ca2+ release and entry channels in the ER membrane control the ER luminal Ca2+ concentration. Ca2+ release channels are the ryanodine receptors (RyR) [148-151] and the inositol 1,4,5-trisphosphate receptors (IP3R) [152-154]. IP3Rs and RyRs can coexist in the same cell [155]. Three isoforms (RyR1, RyR2, and RyR3) [156-160] and several splice variants of the RyR are found in many tissues. RyRs function as homotetramers [161]. and show highest expression in striated muscle [149, 157, 162-170]. In skeletal muscle RyR1 is the predominant isoform in mammals [166, 171-173], in cardiac muscle RyR2 is predominant. Excitation-contraction coupling through interaction with voltage-dependent Ca2+ channels in the t-tubule membrane, i.e. the dihydropyridine receptors (DHPRs) [174-177], activate RyR in the SR. Depolarization of the t-tubule membrane induces a conformational change in the DHPR that activates RyRs stimulating RyR-mediated Ca2+ release from the SR [149]. IP3R also exist in three isoforms with one to two alternative splice variants per isoform and also form heterotetramers [178]. Binding of the second messenger inositol 1,4,5-trisphosphate, which is generated at the plasma membrane, opens the IP3Rs to release Ca2+ from the ER. The action of the Ca2+ release channels is countered by ATP-dependent Ca2+ pumps in the ER membrane, the sarco/endoplasmic reticulum Ca2+ ATPases (SERCA) 1-3 [179-181]. SERCA pumps mediate most of the Ca2+ influx into the ER. ATP hydrolysis is coupled to Ca2+ transport by these pumps because SERCA pumps transport Ca2+ against a Ca2+ gradient from the cytosol into the ER [179-181]. SERCAs can operate in reverse mode in which they synthesize ATP and contribute to Ca2+ leakage from the ER [182]. Three SERCA genes are found in humans, ATP2A1 encoding SERCA1, ATP2A2 encoding SERCA2, and ATP2A3 encoding SERCA3 [180]. All genes possess several splice variants with characteristic tissue-specific expression patterns [180].

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The ER

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SERCA2b and SERCA3 are nearly ubiquitously expressed [180]. Binding of calreticulin [183] or calnexin [184] to a 12 amino acid luminal extension tail specific to SERCA2b inhibits SERCA2b when the ER luminal Ca2+ store is full [185]. In addition to Ca2+ channels and pumps in the ER membrane Ca2+binding proteins of the ER increase the capacity of the ER luminal Ca2+ store allowing to store up to 20 mM Ca2+ [186, 187] while maintaining a free Ca2+ concentration of 1-5 mM [188]. The most abundant Ca2+-binding protein in the SR is calsequestrin [186], followed by the 165 kDa histidine-rich Ca2+-binding protein (HRC) [186, 189, 190] and sarcalumenin [186, 190, 191].

Figure 5. ER Ca2+ homeostasis. Abbreviations: CRT – calreticulin, CSQ – calsequestrin, DAG – diacylglycerol, HRC – 165 kDa histidine-rich Ca2+-binding protein, IP3 – inositol 1,4,5-trisphosphate, PI4,5P2 – phosphatidylinositol 4,5bisphosphate, ROC – receptor-operated channel, ROS – reactive oxygen species, SLN – sarcalumenin, SOC – store-operated channel, VOC – voltage-operated channel.

Calsequestrin has ~50 Ca2+-binding sites and the Kd for Ca2+ is ~1 mM [192-194]. Calsequestrin is anchored via the transmembrane proteins triadin

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[195-198] and junctin [199, 200] to the RyR. At low (< 10 mM) Ca2+ concentrations calsequestrin inhibits the RyR [186, 201]. Ca2+ homeostasis of the ER is important for protein folding in the ER because many chaperones of the ER are Ca2+ binding proteins. In fact, many ER luminal chaperones have first been identified as Ca2+-binding proteins [37, 68, 70, 202, 203]. Further, the activity of several chaperones is regulated by alterations in Ca2+ concentration. Depletion of Ca2+ dissociated BiP from Tcell antigen receptor α chain [204]. Lectin chaperones bind to oligosaccharides at high Ca2+ concentrations, but not at Ca2+ concentrations representing a depleted ER Ca2+ store (~100 μM) [205]. These examples suggest that depletion of ER luminal Ca2+ inactivates ER chaperones and causes accumulation of unfolded proteins in the ER. This conclusion is supported by strong induction of the UPR by the irreversible SERCA inhibitor thapsigargin [206-209]. Ca2+ dissociates oligomeric calnexin [203], whereas low Ca2+ concentrations promote interaction of calreticulin with PDI [210, 211]. Likewise, interaction of calreticulin with ERp57 is affected by changes in the Ca2+ concentration [210, 212]. It is likely that fluctuations in ER luminal Ca2+ concentration also affect protein-protein interactions in other chaperone complexes [213-217] and regulate interaction of chaperones with SERCA pumps and Ca2+ release channels. The physiological consequences of regulation of chaperone interactions by Ca2+ remain to be elucidated. Ca2+ also participates in protein folding reactions. Ca2+ can engage in formation of salt bridges in proteins affecting protein folding in a protein specific manner [214, 218]. The folding of several proteins, for example apo-α-lactalbumin, requires Ca2+ [219].

3.3. REDOX BUFFERING OF THE ER Two redox buffer systems exist in the ER, the reduced/oxidized glutathione redox pair and the redox pairs of reduced and oxidized nicotinamide dinucleotide (NADH/NAD+) and NADPH/NADP+. These redox pairs are separate from their counterparts in the cytosol, and can exist in a different redox status. For example, the ratio of reduced (GSH) to oxidized (GSSG) glutathione is 30:1 to 100:1 in the cytosol, whereas in the ER this ratio is 1:1 to 3:1 [220] making this redox pair more oxidizing in the ER. Generation of one molecule H2O2 per formed disulfide bond [221] may contribute to the oxidizing character of the ER [222]. Fenton decay of H2O2 at

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cytochromes and other metal ion-containing proteins of the ER membrane generates the highly oxidizing and toxic hydroxyl radical [223, 224]. Thioredoxin reductase [225] and glutathione reductase [226] appear to be absent from the ER, indicating that the glutathione and nicotinamide dinucleotide redox pairs are uncoupled in the ER. Net reduction of cortisone to cortisol by 11β-hydroxysteroid dehydrogenase 1 (11βHSD1), which catalyzes a reversible reaction, indicates that the redox potential of the NAD(P)H/NAD(P)+ redox pairs in the ER is reducing. Only the GSH/GSSG redox pair is linked to oxidative protein folding. The ER membrane is virtually impermeable for GSSG, but possesses saturable transport systems for GSH [227], indicating that maintaining the luminal GSH/GSSG ratio requires additional redox systems. The Foyer-Halliwell-Asada cycle (Figure 6) connects the GSH/GSSG redox pair to the ascorbate/dehydroascorbate redox pair. Ascorbate is synthesized in the ER lumen by a membrane-bound flavoprotein, gulonolactone oxidase, in most animals, but not humans [228, 229]. A byproduct of this reaction is H2O2 [230]. The ER membrane is permeable for dehydroascorbate, but not ascorbate [231]. Ingested ascorbate is oxidized by ascorbate oxidases allowing uptake of dehydroascorbate by the ER. Dehydroascorbate is an electron acceptor in oxidative protein folding and accepts electrons from PDI [232] and may also function in maintaining the GSH/GSSG redox buffer of the ER. Ascorbate is also a cofactor for luminal enzymes such as prolyl- and lysyl hydroxylases [233, 234], which play important roles in collagen synthesis. Lipid-soluble electron carriers, especially the tocopherol/tocopheryl radical redox system may relay dehydroascorbate into the ER lumen [235, 236]. The vitamin K redox cycle (Figure 7) participates in γ-carboxylation of glutamate side chains in secretory proteins [237]. Vitamin K epoxide reductase (VKORC1) regenerates vitamin K from vitamin K 2,3 epoxide. VKORC1 contains a thioredoxin-like CXXC center and forms a stable complex with PDI [238], suggesting that VKORC1 accepts electrons from cysteinyl thiols of folding cargo proteins of the ER.

3.4. N-LINKED GLYCOSYLATION IN THE ER N-linked glycosylation is initiated by transfer of a core oligosaccharide from a membrane-bound dolichol phosphate anchor to consensus Asn-XSer/Thr (X ≠ Pro) residues in the polypeptide chain (Figure 8) [239, 240]. Glycosylation serves several purposes in protein folding. First, due to the

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hydrophilic nature of carbohydrates, glycosylation increases the solubility of glycoproteins and defines the attachment area for the surface of the protein. Second, due to their large hydrated volume oligosaccharides shield the attachment area from surrounding proteins and thus act as a chaperone.

Figure 6. Foyer-Halliwell-Asada cycles in the ER contribute to disulfide bond formation and may contribute to maintaining the redox potential of the GSH/GSSG redox pair in the ER because the ER membrane is nearly impermeable for GSSG.

Figure 7. Coupling of L-glutamate γ-carboxylation to oxidative protein folding in the endoplasmic reticulum. Molecular Chaperones of the Endoplasmic Reticulum, Nova Science Publishers, Incorporated, 2010. ProQuest Ebook

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Third, oligosaccharides interact with the peptide backbone and stabilize its conformation [241]. Fourth, sequential trimming of sugar residues is monitored by the lectin chaperone machinery to report on the folding status of the protein (Figure 3; [242, 243]) and will be discussed in detail below.

Figure 8. The pathway of N-linked glycosylation.

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Chapter 4

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MOLECULAR CHAPERONES OF THE ER Two classes of proteins assist protein folding in the ER: molecular chaperones and protein foldases. Foldases catalyze protein folding reactions that otherwise would be too slow to allow for productive protein folding in a cellular environment. Prominent examples for foldases are peptidyl-prolyl cistrans isomerases (Figure 2) and PDIs (Figure 1). Molecular chaperones facilitate protein folding by shielding unfolded hydrophobic regions from surrounding proteins and are further divided in chaperone foldases and chaperone holdases.

4.1. CHAPERONE FOLDASES Chaperone foldases consist of at least two domains, a substrate binding domain (SBD) and a nucleotide binding domain (NBD). They employ ATP hydrolysis cycles to cycle through ATP- and ADP-bound states that in- or decrease their affinities for unfolded substrates by altering the conformation of their SBD [244, 245]. In the high affinity state the unfolded substrate is bound to the chaperone and conformationally locked because of the interaction of the unfolded domain with the chaperone. The low affinity state terminates the substrate-chaperone interaction (Figure 9). Released substrates kinetically partition between folding to the native state, aggregation, and rebinding to the chaperone (Figure 4). Preferential interaction of the chaperone with the unfolded protein provides directionality to the folding of the substrate. ATP hydrolysis and ADP/ATP exchange provide the energy to drive conformational changes in the SBD.

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Figure 9. HSP70 chaperone ATPase cycle. HSP70 chaperones, such as BiP, cycle between an ADP- and ATP-bound state. Chaperone-bound ATP is hydrolyzed by the ATPase activity of the chaperone. Binding of DnaJ proteins to the HSP70 chaperone stimulates the ATPase activity of the HSP70 chaperone. GrpE co-chaperones stimulate the nucleotide exchange reaction. The ATP-bound state is characterized by low affinity for the substrate and fast exchange of substrates on the chaperone. The ADP-bound state is characterized by high substrate affinity and slow exchange of substrates.

The only clear example for a chaperone foldase with an ATP hydrolysis cycle in the ER is the HSP70 class chaperone BiP. The HSP90 class chaperone GRP94 has been suggested to be a chaperone holdase due to its barely detectable ATPase activity, but elucidation of its three dimensional structure together with clear demonstration of ATPase activity for GRP94 have now shifted the discussion toward GRP94 being a chaperone foldase [246-248]. The lectin chaperones calnexin, calreticulin, and calmegin may also be considered to be chaperone foldases in which substrate recognition and ATP consumption cycles have been uncoupled (Figure 10). The function of chaperone foldases is tightly regulated by co-chaperones [249-251]. In the case of HSP70 class chaperones HSP40 or DnaJ co-chaperones stimulate the ATPase activity of HSP70 class chaperones and load substrates onto their

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HSP70 partners [251], whereas GrpE co-chaperones are nucleotide exchange factors for HSP70 class chaperones [250].

Figure 10. The calnexin cycle. Abbreviations: CNX – calnexin, EDEM – ER degradation enhancer, mannosidase α-like, G – D-glucose, M – D-mannose, and UGGT - uridine diphosphate (UDP)-glucose:glycoprotein glucosyl transferase. Reprinted from Mutation Research, volume 569, authors M. Schröder and R. J. Kaufman, title “ER stress and the unfolded protein response”, pages 29-63, Copyright (2004), with permission from Elsevier.

For cytosolic HSP70s several DnaJ co-chaperones and nucleotide exchange factors have been well characterized. For their ER luminal counterparts at least six DnaJ co-chaperones (ERdj1/MTJ1 [39, 40], ERdj2/Sec63p [41-44], ERdj3/HEDJ/Scj1p [45-49], ERdj4 [50]/MDG1 [5053], ERdj5/Erj5p [54, 55], Jem1p [58], and p58IPK/DnaJc3 [59, 60]) and one nucleotide exchange factor for BiP (suppressor of Ire1/Lhs1 synthetic lethality 1 protein (Sil1p)/synthetic lethal with the 7S RNA mutation 1 protein (Sls1p)/BiP-associated protein (BAP) [252-257]) have been identified.

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Further, the HSP70 chaperone Lhs1p has been shown to function as a nucleotide exchange factor for BiP [258]. In addition, several auxiliary cochaperones for cytosolic HSP70s have been described for which ER luminal counterparts have not been identified [259-264]. For HSP90 chaperones the functional diversity of co-chaperones is even more complex. HSP90 cochaperones that stimulate the ATPase activity of HSP90 chaperones, cochaperones that are PPIs, or that may be involved in targeting proteins for degradation, such as the ubiquitin ligase carboxyl terminus of heat shockcognate (Hsc) protein of 70 kDa-interacting protein (CHIP), or in bridging the HSP70 and HSP90 chaperone systems have been described [265]. GRP94 cochaperones remain unknown.

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4.2. CHAPERONE HOLDASES Chaperone holdases do not employ ATP hydrolysis cycles. Examples for chaperone holdases are sHSPs, HSP33, DnaJ proteins, and the chaperone activity of PDI. Chaperone holdases interact with substrate proteins through hydrophobic interactions [266-268] but do not support folding of the substrate to its native state [267, 269-272]. Structural maturation of the substrate may terminate the substrate-holdase interaction [273]. Some holdases provide a chaperone buffer to chaperone foldases and closely cooperate with certain chaperone foldases [270, 274-276]. Under conditions unfavorable for protein folding the substrate is transferred from the foldase to the holdase to which they remain bound until conditions become favorable for folding again. Transfer of the substrate back to the foldase reinitiates folding of the substrate thus preventing futile, ATP-consuming folding attempts under stress conditions. Nevertheless, chaperone holdases may also cycle between a high and low affinity conformation, in which case the transition is triggered by changes in the redox status of the cell or other environmental factors such as temperature [273]. In the following chapters the molecular function of the major ER luminal chaperones will be discussed in the context of the biochemical features of the ER. Because of the more advanced state of research into cytosolic chaperones information on the function and regulation of these chaperones will be incorporated into these discussions to highlight potential gaps in knowledge and mechanistic understanding of chaperonemediated protein folding in the ER.

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4.3 THE HSP70 CLASS MOLECULAR CHAPERONES BIP AND LHS1P

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BiP accounts for ~5% of the soluble, luminal protein of the ER. It belongs to the well-characterized class of HSP70 chaperones which are found in all eukaryotic organelles and the cytosol of bacteria (Figure 11) [277, 278].

Figure 11. Phylogenetic tree of the HSP70 superfamily created with PHYLIP 3.68. Abbreviations: B. t. - Bos taurus, C. e. - Caenorhabditis elegans, C. g. - Cricetulus griseus, D. m. - Drosophila melanogaster, E. c. - Escherichia coli, H. s. - Homo sapiens, M. m. - Mus musculus, N. c. - Neurospora crassa, R. n. - Rattus norvegicus, S. c. - Saccharomyces cerevisiae, S. f. - Strongylocentrotus franciscanus, S. p. Schizosaccharomyces pombe. Genbank accesion numbers: 14G8.3. C. e. (CAA91809), APG-1 M. m. (NP_035150), APG-2 M. m. (Q61316), BiP S. c. (NP_012500), BiP H. s. (CAA61201), BiP M. m. (NP_071705), DnaK E. c. (NP_285706), GRP170 C. e. (CEESE89FB), GRP170 C. g. (Q60432), GRP170 H. s. (NP_006380), GRP170 R. n. (AAB05672), HSC70 B. t. (P19120), HSP70 B. t. (P34933), HSP88 N. c. (AAC23862), HSP105α H. s. (NP_006635), HSP105α M. m. (BAA11035), HSP110 C. g. (Q60446), HSP110 D. m. (CAB38172), HSP110 S. f. (AAB09038), Lhs1p S. c. (NP_012850), Pss1p S. p. (AAC18441), Ssa1p S. c. (NP_009396), Sse1p S. c. (P32589), Sse2p S. c. (P32590), and Yam6p S. p. (Q10061).

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Figure 12. Domain architecture of the HSP70 family showing the relative arrangement of the nucleotide-binding domain (NBD), the β-sandwich and α-helical lid of the substrate binding domain (SBD), the C-terminal extension (CTE) characteristic to HSP110 and GRP170/Lhs1p family members, and a long acidic loop (LAL) inserted between the β-sandwich and α-helical lid of the substrate binding domain present in some HSP110 and GRP170 chaperones. For ER luminal HSP70s the C-terminal ER retrieval sequence is shown. Kar2p is shown as an example for an archetypal HSP70 and Sse1p as an example for a HSP110 family member. The drawing is not to scale.

Much of what we know about BiP has been learnt from work on cytosolic HSP70s and its bacterial ortholog, DnaK. BiP was originally identified as GRP78 [15-17], a protein induced by glucose starvation and, independently, as the immunoglobulin heavy chain-binding protein [14]. BiP is a chaperone foldase of broad substrate specificity and assists in the folding of newly synthesized protein in the ER [279, 280]. BiP also has important functions in co- and posttranslational translocation of nascent polypeptide chains across the ER membrane [79, 138], sealing the luminal end of the translocon [281], recognition of damaged or misfolded proteins for retrograde translocation and degradation [282, 283], and regulation of unfolded protein response signaling (Figure 3) [284-287]. A common denominator in most of its biological functions is preferential interaction with unfolded protein substrates and prevention of aggregation of these unfolded proteins. The budding yeast ortholog, Kar2p, is required for karyogamy, the fusion of the two nuclei of haploid a and α cells during mating of haploid yeast cells [18, 288, 289]. The second ER luminal HSP70 protein, GRP170/Lhs1p [19-21], belongs to the HSP70 subfamily of HSP110/GRP170 chaperones (Figure 11). These chaperones stimulate the ATPase activity of other HSP70 chaperones [20, 255, 258, 290-300]. A mammalian ortholog of Lhs1p has been identified as a

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protein induced by glucose starvation (GRP170, [25-28]) and hypoxia (ORP150, [22-24]).

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4.3.1 Structure and Function of the NBD BiP follows the structural blueprint of all HSP70 chaperones. They consist of a conserved 44 kDa N-terminal NBD and a 25 kDa C-terminal domain divided into a conserved SBD of 15 kDa followed by a 10 kDa immediate Cterminal domain (Figure 12) [8-10, 278, 301]. Conformational coupling of both domains by a flexible hinge region connecting both domains is the basis for ATP hydrolysis driven cycling of HSP70 chaperones through two alternating states of high (ADP-bound) and low (ATP-bound) affinity for unfolded substrates [302-307]. X-ray crystallography has provided insight into the three dimensional structures of the NBD of the 70 kDa bovine heat shock cognate protein (HSC70) [308-310] and the SBD of DnaK [311]. The structure of the NBD of DnaK closely resembles the structure of the NBD of HSC70 [312]. The N-terminal domain of BiP shares 66% sequence identity and 81% homology with HSC70 [278] and can easily be modeled onto the HSC70 structure [313], suggesting that structural lessons learnt from HSC70 and DnaK are valid to understand BiP. The KM of HSP70 chaperones for ATP is ~1-2 μM and is well below the cytosolic ATP concentration of 1-2 mM [249, 302, 305, 314-322]. For ER luminal HSP70s the ATP concentration in the ER lumen is of consequence, which remains unknown. ATP is transported into the ER by an ADP/AMP antiport mechanism [323]. Therefore, it is possible that the protein folding capacity of the ER is limited by ATP import, especially under conditions of high secretory activity or ER stress. The NBD consists of two large, globular lobes (I and II) each containing two subdomains (A and B). The two lobes are connected by two crossed αhelices and are separated by a deep cleft to which ATP binds and in which ATP hydrolysis takes place (Figure 13) [308]. Both lobes and the connecting helices contribute to forming the binding pocket for ATP, one Mg2+ and two K+ ions at the bottom of the cleft [308, 324]. ATP is positioned through a hydrophobic adenosine binding pocket [308] and interactions of one Mg2+ and two K+ ions with the triphosphate group [324]. The carboxyl groups of three aspartates (D10, D199, and D206 in bovine HSC70) are involved in coordination of the metal ions [324, 325].

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Figure 13. Crystal structure of the ATPase domain of bovine HSC70 (PDB entry no 1ba0). The two lobes are colored red and blue, ADP yellow, Pi orange, and several residues involved in MgADP coordination (D10 and D199) and catalysis (K71) are colored white. The N- and C-termini are labelled ‘N’ and ‘C’, respectively. The numbering of the two lobes of the ATPase domain is indicated [555].

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Mutagenesis of any of these aspartates to asparagine or serine decreases kcat to 1-10% of the wild-type enzyme [322, 326]. A conserved lysine (K71) and a glutamate (D175) are positioned opposite to the γ-phosphate group of ATP in the crystal structure of HSC70 [308]. Mutation of K71 to a neutral amino acid completely destroys ATPase activity [325], suggesting that this lysine positions a H2O molecule or OH- anion for in-line nucleophilic attack on the γ-phosphate group or acts as a base to increase the nucleophilicity of the nucleophile [325]. In contrast, the effect of mutation of E175, which may participate in correct positioning of the water molecule for nucleophilic attack on the γ-phosphate group, to glutamine or serine is not that dramatic. Taken together, these data suggest the following mechanism for ATP hydrolysis (Figure 14) [310, 324-326]: The MgATP complex binds initially through coordination of the Mg2+ ion with the carboxyl groups of D10 and D199 and a non-bonded oxygen atom of the β-phosphate group of ATP. Rearrangement leads to formation of a β,γ-bidentate complex in which the Mg2+ ion is coordinated by the carboxyl groups of D10 and D199 and one oxygen atom of the β- and γ-phosphate groups of ATP. The two K+ ions stabilize the β,γbidentated complex, explaining the absolute dependence of ATP hydrolysis on the presence of K+ ions [327]. The β,γ-bidentated complex aligns the terminal phosphate group for in-line nucleophilic attack by a water molecule or OHanion. Lysine 71 participates in the nucleophilic attack by stabilizing or accepting a proton from the OH- anion or water molecule (Figure 14). The rate-limiting step of ATP hydrolysis is the chemical, hydrolytic step [317].

4.3.2. Structure and Function of the SBD The SBD is formed by two structural units, a β-sandwich consisting of two sheets of four antiparallel β-strands with a hydrophobic groove on its upper side and a lid formed by four α-helices located above the hydrophobic groove. The two units are connected via an additional α-helix (Figure 15). The α-helices are making direct contacts to the substrate. The hinged lid has been proposed to gate access to the hydrophobic groove. ATP binding to the NBD opens the peptide binding pocket, whereas ATP hydrolysis closes the pocket to increase the affinity of the chaperone for its substrates. Two loops emanating from the top sheet of the β-sandwich form the hydrophobic pocket in DnaK. The hydrophobic pocket consists of a floor of hydrophobic amino acid residues surmounted by an hydrophobic arch.

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Figure 14. Simplified model of the β,γ-bidentated complex in the catalytic center of the NBD of bovine HSC70.

Figure 15. Crystal structure of the SBD of DnaK (PDB entry no. 1dkx). The βsandwich and hydrophobic groove are colored red, the α-helical lid is shown in blue, and the peptide in white. The N- and C-termini are labelled ‘N’ and ‘C’, respectively, the α-helices 1 to 5 [311].

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Both, the hydrophobic floor and arch engage in hydrophobic interactions with the substrate and determine substrate specificity. The hydrophobic pocket forms a channel with a cross section of ~5 x 7 Å. In DnaK negatively charged residues flank the ends of the hydrophobic channel thus favoring the presence of basic amino acids at the end positions of interacting peptides. The main chain of the bound peptide also forms several hydrogen bounds with the main chain of the chaperone. This geometry and enclosure of the bound substrate by lid closure requires that the remainder of the substrate protein is separated by ~10 Å from its region interacting with the chaperone, suggesting that DnaK substrates are considerably unfolded. Analysis of peptide libraries has identified consensus motifs for interaction with DnaK [328-330] and BiP [303, 304]. The consensus peptide interacting with DnaK consists of 4-5 hydrophobic residues, preferably leucines, flanked by basic residues. Acidic residues are not present in the peptide core, and disfavored in the flanking regions, which is in good agreeance with the structure of the DnaK peptide binding pocket. These sequences are often buried in the hydrophobic core of a protein and found in β-sheets. Similar approaches with BiP have established that BiP displays selectivity for hydrophobic peptides with the consensus sequence HyXHyXHyXHy (Hy = large hydrophobic amino acid, X = any amino acid). High KM values (1-100 mM) [303, 304] of HSP70 chaperones suggest a wide substrate spectrum. Protein aggregation occurs on a time scale of seconds. If binding of unfolded substrates to HSP70 chaperones is to compete successfully with their aggregation, unfolded substrates must interact with HSP70 chaperones on a similar, if not faster, timescale as protein aggregation. The rate constant for association of nucleotide-free HSP70 chaperones with peptide substrates are ~10-100 M-1s-1 [331]. At this rate, complex formation would take at least several minutes. ATP binding accelerates this rate constant at least ~10 fold, but substrate binding requires conversion of the ATP to the ADP-bound state. The turnover number for unstimulated ATP hydrolysis by HSP70 chaperones is ~0.1 min-1 [249, 302, 305, 317-322, 332], making ATP hydrolysis the ratelimiting step for the HSP70 chaperone substrate interaction. Substrate binding to HSP70 chaperones stimulates the ATPase activity of HSP70 chaperones [302, 303], but is not considered to sufficiently elevate the rate of ATP hydrolysis to allow HSP70 chaperones to kinetically compete with aggregation of the substrate. To make HSP70 chaperones productive chaperones in vivo, ATP hydrolysis is stimulated by substrate binding [302, 303] and specialized co-chaperones of the HSP40 family.

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Figure 16. Crystal structure of amino acids 1-554 of bovine HSC70 (1yuw) [333]. Coloring: NBD – blue, SBD – red, linker – yellow, helix 1 of the SBD – orange, and portion of the SBD mimicking a substrate – white.

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4.3.3. Communication between the SBD and NBD in BiP The crystal structure of the SBD suggests that lid closure induced by ATP hydrolysis in the NBD locks the substrate in the substrate binding pocket and increases the affinity of HSP70 chaperones for their substrates. Insight into interdomain communication between the NBD and SBD has been provided by the 2.6 Å structure of bovine HSC70 in the nucleotide-free state lacking ~10 kDa at the C-terminus (Figure 16) [333]. This structure encompasses the βsandwich forming the SBD and the α-helix connecting the hinged lid to the bottom of the substrate binding pocket. The two domains are connected by a flexible linker of 10 aa. The hinge helix of the SBD contacts a β-strand and a loop in the groove opposite to the ATP binding cleft of the NBD. Closing of the ATP binding cleft upon nucleotide binding/release or ATP hydrolysis impinges on the groove interacting with the hinge helix of the SBD, transducing ATP hydrolysis cycle induced movements in the ATP binding cleft to the lid of the SBD (Figure 16). Intrusion of the linker into the interdomain space may result in considerable conformational changes in this region, which may lead to differential movement of the helical lid and the βsandwich of the SBD. An important role of linker movement in communication between both domains is supported by biochemical and crystallographic data [311, 333-338]. A conserved arginine, R151 in E. coli DnaK, exposed at the surface of this groove is involved in a hydrogen bond relay originating from the ATP binding cleft and may contribute to conformational responses of the groove to the nature of the bound nucleotide [339]. Two additional surface exposed, positively charged residues in the NBD, K155 and R167, and one negatively charged residue in the linker, D393, are involved in communication between the NBD and SBD [340]. Therefore, the linker appears to be the lever that is wielded to induce the ATP hydrolysis competent conformation in the NBD. A smaller interaction surface formed by a loop of the SBD and a loop in the NBD is also important for conformational communication between both domains [341].

4.3.4. HSP40/DnaJ Co-Chaperones for BiP Without stimulation of ATP hydrolysis and ADP-ATP exchange HSP70 chaperones cannot kinetically compete with protein aggregation. HSP40 cochaperones stimulate the ATPase activity of HSP70 chaperones, whereas GrpE co-chaperones are adenine nucleotide exchange factors for HSP70s (Figure

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17). HSP40 co-chaperones are characterized by a conserved J domain of ~70 amino acids. The J domain is critical for stimulation of HSP70 ATPase activity. The J domain is formed by four α-helices connected via three loops (Figure 18). The J domain is stabilized by hydrophobic interactions between amino acid residues protruding from helices 2 and 3. The loop between helices 2 and 3 contains a conserved HPD sequence motif distinguishing J proteins from J-like proteins. The HPD motif is important for interaction of J proteins with HSP70 chaperones and stimulation of their ATPase activity. The J domain contacts the groove between lobes IA and IIA of the HSC70 NBD. Stimulation of ATPase activity is achieved by directing the linker toward a hydrophobic patch on the surface of the NBD. The SBD is displaced upon J protein binding from the NBD, suggesting that the SBD gains conformational freedom to capture substrates presented to the HSP70 chaperone by the J protein [342, 343]. Interaction of DnaJ co-chaperones also increases the stability of the chaperone-substrate complex [344] and affects the substrate specificity of HSP70 class chaperones [344, 345]. J proteins are classified into three categories based on the nature of domains they share with E. coli DnaJ, the founding member of this protein family (Figure 19) [346]. Class I J proteins contain a glycine and/or phenylalanine rich region and a Zn2+-finger in addition to the J domain, class II J proteins only the glycine/phenylalanine rich region, and class III J proteins only a J domain. Class I J proteins, such as ER luminal Scj1p [48] and HEDJ [347], interact with unfolded proteins through their Zn2+-fingers [348, 349]. This function may have been taken on by other domains such a glutamine-rich regions or tetratricopeptide binding motif repeats (TPR) in some class III J proteins, for example p58IPK (Figure 19) [350, 351]. Class II J proteins also interact with unfolded proteins [56, 352]. As a consequence class I and II J proteins load HSP70 substrates onto HSP70 chaperone foldases (Figure 17). DnaJ proteins associate with these substrates fast enough to kinetically compete with aggregation successfully [331, 353, 354] and act catalytically in loading substrates onto HSP70 chaperones [355, 356], suggesting that the majority of HSP70 substrate interactions are mediated by DnaJ proteins. The cytosolic J protein TPR2/TTC2/DJC7 can offload unfolded polypeptide chains from the HSP90 foldase machinery onto the HSP70 foldase machinery [357]. Affinity panning has revealed that the class I HSP40 Ydj1p binds peptides harboring hydrophobic stretches [358]. Both, class I and II HSP40s, function as dimers [353]. The crystal structure of a peptide-binding fragment of Sis1p, a class II HSP40, has revealed a dimeric structure with a large cleft (Figure 19C) [359].

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Figure 17. Revised HSP70 ATPase cycle incorporating loading of substrates onto HSP70 chaperones by DnaJ proteins. The HSP70 chaperone is shown in black. J protein-mediated substrate loading overcomes the slow association kinetics of HSP70 chaperones with their substrates in vitro and enables HSP70 chaperones to kinetically compete with protein aggregation.

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Figure 18. NMR solution structure of the J domain of E. coli DnaJ (residues 1-78) (PDB entry no. 1bqz) [556]. The N- and C-termini are labeled ‘N’ and ‘C’, respectively. The four α-helices are labeled 1 to 4. The HPD motif is indicated as a ball and stick model.

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Figure 19. DnaJ proteins. (A) Domain structure of class I, II, and III J proteins. (B) J proteins of the ER. The J domain of Hlj1p is localized in the cytosol. Abbreviations and symbols: C – L-cysteine, D – L-aspartate, E – L-glutamate, F – L-phenylalanine, G – glycine, J - J domain, Q – L-glutamine, SANT - SANT domain, I to IV - thioredoxinlike domains, 1 to 9 - TPR repeats. Black boxes in the middle of the protein represent transmembrane segments, directly at the N-terminus signal peptides. The drawings are not to scale. (C) Crystal structure of the protein-binding domain of the DnaJ cochaperone Ydj1p (PDB entry 1nlt, [360]. Ydj1p is shown in blue, the bound substrate GWLYEIS in red. Only the monomeric form of Ydj1p is shown. (D) Model for substrate transfer from DnaJ co-chaperones onto HSP70 chaperones based on the crystal structure of dimeric protein binding domains of Sis1p (PDB entry 1c3g, [359]).

This cleft has been proposed to accommodate the SBD of HSP70s to facilitate substrate transfer (Figure 19D). Unfolded substrates may interact with Sis1p monomers through interaction with the parallel β-sheet in domain I [359]. The crystal structure of the class I HSP40 Ydj1p in complex with the peptide GWLYEIS shows the peptide interacting with β-sheets in domain I [360]. Mutation of analogous residues in the predicted β-sheet in domain I of ER-localized ERdj3/HEDJ corrupted the ability of ERdj3 to interact with IgG heavy chains in vivo and denatured firefly luciferase in vitro [361], suggesting that ERdj3 and Ydj1p employ a similar substrate recognition mechanism. In vivo ERdj3 formed multimers [361], suggesting that ERdj3 may transfer substrates onto BiP in a similar way as Sis1p. Whereas the structure of Ydj1p shows a monomer, the protein forms a dimer in solution. Modeling supports that Ydj1p monomers can form a dimer with a three dimensional structure similar to the structure of Sis1p. Computational analysis suggests that Ydj1p recognizes peptides with the consensus sequence

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GX[LMQ](P)X(P)(CIMPVW), where [XY] denotes either X or Y and (XY) denotes neither X nor Y [362]. A J protein specific to the ER is Erj5p, which contains four thioredoxin-like domains and probably is involved in coordinating oxidative folding driven by disulfide bond formation and isomerization with conformational folding of the polypeptide backbone. A second function of J proteins is to target HSP70 chaperones to specific sites of action. For example, yeast Sec63p recruits BiP to the translocon to participate in translocation of nascent polypeptide chains into the ER [363]. Mammalian SEC63 and MTJ1 also function in translocation [364, 365]. Hlj1p, whose J domain is exposed to the cytosol, recruits the HSP70 chaperones Ssa1p and Ssa2p to the ER membrane to participate in ERAD. All mammalian ER luminal J proteins interact with BiP and stimulate its ATPase activity in vitro [40, 44, 46, 57, 347, 366, 367]. As J proteins are believed to form equimolar complexes with HSP70 chaperones the large structural diversity of J proteins suggests that these co-chaperones tether BiP to sub-ER localizations to participate in specific processes such as translocation, specify substrate selectivity of BiP, and the fate of the interacting substrate.

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4.3.5. Nucleotide Exchange Factors (NEFs) Nucleotide exchange factors are the second class of HSP70 cochaperones. Several protein families provide NEF function to HSP70 chaperones. The archetypal NEF is E. coli GrpE. Two other dedicated NEFs that are structurally different from GrpE, are the BAG-1 family of NEFs [368], and NEFs characterized by several α-helical armadillo repeats. The latter class includes cytosolic HSP70 binding protein 1 (HspBP1) [369] and ER luminal BAP [252]/Sil1p [255, 370]/Sls1p [253, 254, 257]. Finally, the HSP70 subfamilies of HSP110 and GRP170/Lhs1p chaperones provide NEF services to HSP70 chaperones. GrpE is a 197 aa protein consisting of a central αhelical dimerization domain and a C-terminal β-sheet domain (Figure 20) [249, 371, 372]. GrpE accelerates ADP release from DnaK-ADP ~5,000 fold by decreasing the affinity of DnaK for ADP ~200-fold [373]. Together, GrpE and DnaJ increase ATP hydrolysis by DnaK several hundred-fold [251], which makes HSP70 proteins efficient chaperones despite their intrinsic low turnover number for ATP. GrpE-catalyzed nucleotide release may be a two step process, in which facilitated ADP release precedes release of GrpE from the chaperone, which is triggered by ATP binding to the chaperone (Figure 17) [374, 375].

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Figure 20. Molecular action of NEFs on the NBD of HSP70 chaperones. (A) The βsheet of a GrpE dimer reaches deep into the nucleotide binding cleft between the two lobes of the NBD. A 14° rotation of subdomain IIb opens the cleft and facilitates nucleotide release (PDB entry 1dkg, [312]). (B) The three helix bundle of BAG-1 contacts subdomains Ib and IIb. In analogy to GrpE a 14° rotation of subdomain IIb opens the cleft and facilitates nucleotide release (PDB entry 1hx1, [368]). (C) The four armadillo repeats of HspBP1 contact subdomain IIb inducing vertical movement of subdomain IIb in relation to subdomain Ib to open the nucleotide binding cleft (PDB entry 1xqs, [369]).

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Structural analysis of three HSP70-NEF complexes, GrpE-DnaK [312], Bag-1-HSC70 [368], and HspBP1-HSP70 [369], has revealed diversity in how different NEFs catalyze ADP release from HSP70 chaperones (Figure 20). The β-sheet domain of one GrpE subunit contacts the nucleotide binding cleft of DnaK and induces a rotation of 14° of subdomain IIB of the DnaK NBD. BAG-1 also induces a 14° rotation of subdomain IIB, but by another mechanism. GrpE contacts the back of the NBD and reaches into the nucleotide binding cleft with its β-sheets to open subdomain IIB. In contrast, BAG-1 opens subdomain IIB by contacting both sides of the central cleft from above via α-helices. While different in structure and their mode of interaction with HSP70s, the mode of action of GrpE and BAG-1 is conserved. In contrast, HspBP1 uses a different mechanism. HspBP1 contacts subdomain IIB of HSP70 via its four α-helical armadillo-like domains. Sterical clashes with subdomain IB are resolved by movement of the two lobes of the NBD in relation to each other, leading to opening of the nucleotide binding cleft. The ER possesses one NEF, BAP [252]/Sil1p [255, 370]/Sls1p [253, 254, 257], which may adopt an HspBP1-like fold and, therefore, may open the nucleotide binding cleft of BiP in an analogous way to HspBP1. Mutations in SIL1 cause Marinesco-Sjögren syndrome, a neurodegenerative disease, in humans [376, 377] and ataxia in the woozy mouse [256]. Further, deletion of SIL1 in yeast is not lethal [255]. These observations suggest that additional NEFs are present in the ER [292]. Synthetic lethality of deletion of the HSP70 Lhs1p with SIL1 suggests that Lhs1p and Sil1p share a common function [255]. Indeed, Lhs1p stimulates nucleotide exchange on BiP in vitro [258] and is present in the ER in complexes with BiP [213]. Recent work has shown that NEF activity for the HSP70 chaperone Lhs1p reflects on a common function of HSP70 chaperones belonging to the GRP170/Lhs1p and HSP110 subfamilies. The HSP70 protein family can be divided further into two subfamilies, archetypal and atypical HSP70s (Figs. 11 and 12). Atypical HSP70s can be further divided into HSP110 and ER luminal GRP170/Lhs1p HSP70s [300, 378]. Atypical HSP70s are distinguished from archetypal HSP70s by the presence of a C-terminal extension and a long acidic loop insertion between the β-sandwich and αhelical lid of the SBD [300, 378]. The cytosolic HSP110 Sse1p and Lhs1p form 1:1 stoichiometric complexes with Ssa [293, 294, 379] and BiP [258], respectively. Complex formation with nanomolar affinity is independent of other co-chaperones and can be reconstituted in vitro [258, 291, 380]. The interaction is mediated by the NBDs of both chaperones [291, 379-382]. Like Lhs1p HSP110s are nucleotide exchange factors for their interacting HSP70

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partners [258, 293, 294, 379]. The cytosolic HSP110 Sse1p is a NEF for the cytosolic HSP70s Ssa1-4p and ribosome-associated Ssb chaperones [291, 380]. A C-terminal truncation of Sse1p did not complement growth phenotypes of a sse1Δ strain and was ineffective as a NEF in vitro, suggesting that the C-terminal extension mediates nucleotide exchange. Crystal structures for two HSP110·HSP70 complexes, a complex between yeast Sse1p and the NBD of human HSP70 [381] and between yeast Sse1p and the C-terminal ~10 kDa deletion of bovine HSC70 [382] have revealed extensive interactions between both NBDs (Figure 21). In these complexes extensive contacts between opposite lobes of both NBDs have been observed in H/D exchange experiments [383]. In addition, the C-terminal 3 helix bundle domain (3HBD) of Sse1p contacts subdomain II of the HSP70 NBD and tilts lobe II of the NBD in the HSP70 to open the nucleotide-binding cleft in the NBD [381-383]. As a consequence ADP can no longer make contact to both binding pockets for the adenine moiety and the diphosphate group, resulting in a decrease in affinity of ADP and dissociation of ADP from HSP70. Altogether, the HSP110 Sse1p appears to use a BAG-1 like NEF mechanism (Figure 20). The question then arises what, if any, is the role of the ATPase domain in HSP110 proteins? The interaction between HSP110 and HSP70 depends on the ATP-bound state of the HSP110 chaperone [293, 294, 379]. In Sse1p nucleotide binding induces conformational compaction in two steps. ADP binding induces a compacted, but still inactive conformation, even at large excess, while binding of ATP induces the active, compacted conformation capable to interact with HSP70s. ATPase-deficient K69M-Sse1p exhibits dissociation kinetics from Ssa1p that are indistinguishable from WT Sse1p [384] and fully complements a sse1Δ sse2Δ mutation in vivo [293, 385]. Mutations decreasing the affinity for nucleotide interfered with Sse1p function [385]. These data suggest that the ATPase activity of Sse1p is dispensable for function. ATP binding to Ssa1p terminates its interaction with Sse1p [384]. Stochastic ATP for ADP exchange driven by the cellular concentration ratio of ~10:1 for these nucleotides may contribute to release of Sse1p from Ssa1p by converting it into the inactive, ADP-bound conformation. Subtle differences in HSP110 NEF function may also exist. For example, binding of mammalian HSP105α to HSP70 stimulates its ATPase activity [299], suggesting that either HSP70 hands over substrates to HSP105α (Figure 22) or that stimulation of the ATPase activity of HSP105α by HSP70 is a mechanism to control the activity of HSP105α.

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Figure 21. Crystal structure of the HSP110-HSP70 complex (PDB entry 3c7w, [382]). Sse1p is shown in yellow to brown tones and HSC70 in blue tones. The SBDs and the two lobes of the NBDs of Sse1p and HSC70 are labeled.

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Figure 22. Model for function of the atypical HSP70 chaperones HSP110 and GRP170/Lhs1p as NEFs for archetypal HSP70 chaperones.

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In yeast, BiP/Kar2p is present in ~2500 fold excess over Lhs1p [386]. Further, the yeast ER possesses a ~20 fold excess of Sil1p over Lhs1p [386], suggesting that Lhs1p participates only in a fraction of the chaperoning activities of BiP. Deletion of LHS1 and SIL1 is synthetic lethal, showing that these two NEFs are the only NEFs in a yet to be identified specific chaperoning activity of BiP and that they can substitute for each other in this chaperoning activity. In translocation of nascent polypeptide chains across the ER the activities of Sil1p and Lhs1p can be separated. Sil1p functions in cotranslational translocation [254, 255], whereas Lhs1p appears to function only in posttranslational translocation of polypeptide chains into the ER [19, 21, 290, 387]. In this regard viability of sil1Δ and lhs1Δ strains suggests the existence of additional NEFs in the yeast ER that substitute for Lhs1p and/or Sil1p in translocation. Nucleotide exchange releases substrates from HSP70 chaperones. Therefore, stimulation of nucleotide exchange on BiP by Lhs1p may simply facilitate substrate release from BiP (Figure 22). Alternatively, but non-mutually exclusively, Lhs1p may take over substrates from BiP, or both chaperones may co-ordinate their chaperoning activities on a particular substrate molecule (Figure 22). Intriguingly, the relationship between BiP and Lhs1p is reciprocal in that BiP stimulates the ATPase activity of Lhs1p [258]. Action of BiP on Lhs1p in a J protein-like fashion suggests that BiP may by transferring substrates onto Lhs1p. Lhs1p, therefore, may be involved in folding of partially folded substrates, a subset of substrates, or in targeting of terminally misfolded substrates toward degradation.

4.3.6 Regulation of BiP by Posttranslational Modification In an inactive state, HSP70 chaperones, including BiP, form stable dimeric and oligomeric complexes [305, 314, 388, 389]. In the oligomeric state, HSP70 chaperones are bound to ADP, phosphorylated in their SBDs [314, 388-394], and ADP-ribosylated [389, 390, 395-397]. These oligomeric forms do not interact with unfolded substrates and have been proposed to function as a reserve pool. In in vitro experiments both peptides [305] and ATP [388] dissociate the oligomeric form of BiP. The monomeric and oligomeric BiP pools are readily interconvertible. In vivo, conversion of BiP to the monomeric, unmodified form precedes induction of expression of its gene [389, 398]. However, it is currently unknown if these posttranslational modifications are required for BiP function, or are solely coinciding. Autophosphorylation of purified BiP preparations was suggested [392, 393],

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but phosphorylation due to a contaminating kinase was not convincingly ruled out as a source for phosphate incorporation into BiP.

4.4. THE HSP90 CLASS ER-RESIDENT MOLECULAR CHAPERONE GRP94 GRP94 is a member of the HSP90 class of chaperones and consists of an N-terminal NBD (N domain), followed by a charged region (M domain), a dimerization domain (C domain) and C-terminal ER retention motif (Figure 23A) [399]. HSP90 chaperones are obligate dimeric chaperones and belong to the GHKL superfamily of ATPases and histidine kinases [400].

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4.4.1. The GHKL Superfamily of ATPases and the Bergerat Fold GHKL protein family members are DNA topoisomerases of the gyrase class, MutL-like DNA mismatch repair proteins, histidine kinases of the HisAsp signal transduction system, and HSP90 chaperones. These proteins share a unique nucleotide binding domain which is characterized by the so called Bergerat fold [401-407]. ATP is bound to the Bergerat fold in an unusual, kinked conformation. The Bergerat fold is characterized by four conserved motifs, motif I (also called ‘N box’, uubEuuuaNouDA), II (G1 box, uxuxDNGxGuxbaauxxuu), III (G2 box, uGxxGxouxSxxxuoxbuTuxT), and IV (G3 box, TxnGT, a – acidic residue, b – basic residue, o – small residue, u – bulky hydrophobic residue) [400]. The Bergerat fold consists of a sandwich formed by a bottom of four-stranded antiparallel β-sheets and a top formed by three α-helices [400] (Figure 24A). Helices α1 and α3 are oriented parallel to the strands of the β-sheet, whereas helix α2 is perpendicular to the strands of the β-sheet. A loop (loop 4) between helices α2 and α3 functions as a flexible lid that can close over the ATP binding pocket. The N box lies at the Cterminus of helix α1. Its conserved asparagine (N107 in canine/human GRP94) co-ordinates an octahedrally co-ordinated Mg2+ ion that contacts all phosphate groups of the bound nucleotide through solvent-mediated hydrogen bonds. The G1 box lies in loop 3 connecting the β2 sheet and helix α2. The conserved aspartate (D149) of the G1 box forms a hydrogen bond with the N6amine of the adenine moiety. The conserved glycines of the G1 and G2 box form flexible hinges for the ATP lid.

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Figure 23. Structure of GRP94. (A) Domain structure of GRP94. The boundaries of the N, M, and C domains are indicated. KEKE – potential protein-protein interaction motif, L ZIP – leucine zipper, NBD – nucleotide binding domain, and SP – signal peptide. (B) Crystal structure of the open conformation of GRP94 (PDB entry 2o1u, [246]). (C) Crystal structure of the closed, twisted V conformation of yeast HSP82 (PDB entry 2cg9, [411]. In (B) and (C) the two protomers of a dimer are shown in blue and red, respectively. The N domain is displayed in lighter and the M and C domains in darker colors. The bound nucleotide is shown in yellow and R448 in GRP94 (R380 in yeast HSP82) in white. Movement of this arginine into proximity of the phosphate groups of the nucleotide can be seen by comparing its position in relation to the nucleotide in the left (blue) protomers in parts (B) and (C).

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Figure 24. Structure of the NBD of GRP94. (A) Schematic of the Bergerat ATPbinding fold. (B) Two views of the crystal structure of canine GRP94 (PDB no. 1tc0 [557]). α-helices and β-strands of the Bergerat fold are shown in blue and red, respectively. Amino acids making contacts with bound nucleotide and bound ADP are shown in a ball and stick model and are labeled. The Mg2+ ion is shown in yellow.

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The ATP lid in HSP90 chaperones consists of two helices and one loop. The conserved glycine of the G1 box and the threonine/serine (S213) at the end of strand β3 make solvent-mediated contacts with the N1 atom and the N6-amine of the adenine moiety (Figure 24B). A conserved lysine (K168) and asparagine (N162) in the ATP lid contact the β-phosphate of ATP and the adenine and ribose moieties, respectively. Glutamate 103 in the N box is involved in nucleophilic attack of a water molecule on the γ-phosphate group of ATP [401, 408-410]. It acts as a base that positions the water molecule in line for attack and participates in abstracting a proton from the water molecule. An important feature of all GHKL ATPases is that a second conserved lysine or arginine residue residing in a domain immediately adjacent to the NBD, the M domain in HSP90 proteins, also hydrogen bonds to this water molecule in the hydrolytically active state. In yeast Hsp82 this is R380 [411], in human HSP90α2 R400, and in GRP94 R448 [246]. Mutation of this basic residue in E. coli GyrB [25], a Drosophila homolog of DNA gyrase [412], MutL [402], yeast Hsp82 [411], and GRP94 [246] decreases or completely destroys ATPase activity. A consequence of this ‘split ATPase’ arrangement is that substantial conformational rearrangements are necessary to bring this basic residue in line for formation of a hydrolysis competent ATPase domain.

4.4.2. The ATP Hydrolysis Cycle of HSP90 Chaperones In cytosolic HSP90s and bacterial HtpG these domain movements are initiated upon ATP binding (Figure 25). In the nucleotide free, apo form HSP90 chaperones assume an open V-shaped conformation (Figure 23B). ATP binding triggers closure of the ATP lid. Lid closure facilitates dimerization of the NBD domain by inducing conformational changes in a short N-terminal segment [413]. These conformational changes in the N domain trigger further substantial changes in the rest of the HSP90 dimer including a ~90° rotation of the N and M domains. This brings the catalytic arginine in the M domain in proximity of the catalytic aspartate and activates the water molecule for in line attack on the γ-phosphate group of ATP. Overall, the ATPase competent chaperone assumes a closed, twisted V conformation (Figure 23C). This conformation is metastable and relaxes upon ATP hydrolysis into the open, V-shaped conformation (Figure 25).

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Figure 25. ATPase cycle of the cytosolic chaperone HSP90. ATP binding to the N domain (N) induces conformational changes in the N domain leading to lid closure and dimerization of the two N domains of both protomers in a HSP90 dimer. This brings R380 in the middle (M) domain in proximity of the γ-phosphate group of ATP generating active ATPase centers. Upon ATP hydrolysis the strained ‘twisted V’ conformation relaxes into the ‘open V’ conformation and releases Pi and ADP. The C domain (C) mediates dimer formation of HSP90. GRP94 undergoes a similar ATPase cycle as HSP90 in vitro, but the importance of this ATPase cycle for the in vivo function of GRP94 is not clear. Cytosolic HSP90 co-chaperones such as Aha1p, Cdc37p, Sba1p, and Sti1p regulate conformational changes in the HSP90 dimer. Similar GRP94 co-chaperones are currently unknown.

ATP binding and hydrolysis are faster than conformational changes leading to the closed, ATP hydrolysis competent ‘twisted V’ conformation after nucleotide binding (Figure 25) [414]. These rate-limiting conformational changes are regulated by several co-chaperones, such as Cdc37p [415-420], Hop/Sti1p [421-423], Aha1p [424-427], and p23/Sba1p [428, 429], and Hch1p [430]. In the presence of ATP the activation energy of two conformational changes, one associated with the closed and one associated with the open state, is decreased [431]. A likely explanation is that hydrolysis of the γ-phosphate group accelerates conformational changes in the closed state, whereas dissociation of ADP or inorganic phosphate accelerates the conformational

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change in the open state. The energetic differences in four conformations observed in single molecule fluorescence resonance energy transfer (FRET) experiments are small [431], suggesting that all conformations can be stochastically sampled even in the absence of nucleotide. This conclusion is supported by the observation of several conformations for HSP90 in vitro [432, 433]. The ATPase activity of yeast HSP90 is required for viability [408, 409], suggesting that imposition of directionality of the ATPase cycle by virtually irreversible ATP hydrolysis or acceleration of discrete conformational changes in the ATPase are required for function of HSP90. Identification of the CDK4-HSP90 interaction surface couples the ATP hydrolysis cycle of HSP90 chaperones to substrate recognition and interaction. HSP90 chaperones, including GRP94, have low ATP turnover numbers allowing them to hydrolyze one molecule of ATP every 1-2 min in case of yeast Hsp82 [409, 434] and GRP94 [246, 247] and every ~20 min in case of human HSP90 [435]. This low ATPase activity is essential for function of yeast HSP90 [408, 409]. The extensive conformational rearrangements triggered by ATP binding are slower than ATP hydrolysis. Along the same argumentation as for HSP70 chaperones, HSP90 chaperones by themselves should not be able to compete kinetically with substrate aggregation in vivo if these conformational changes control substrate affinity. Therefore, several cochaperones for cytosolic HSP90s exist that regulate their ATPase activities, their conformational transitions, and load substrates onto cytosolic HSP90s, thus bridging the HSP70 and HSP90 machineries [436-438]. No cochaperones for GRP94 have been identified. Sequences involved in cochaperone interactions in cytosolic HSP90s are not conserved in GRP94, suggesting at the very least that if GRP94 co-chaperones exist that these will not simply be ER luminal paralogs of cytosolic HSP90 co-chaperones. GRP94 is insensitive to the nature of the bound adenine nucleotide and assumes an open, V-shaped conformation in the apo form, bound to ADP or nonhydrolyzable 5'-adenylyl-β,γ-imidodiphosphate. The crystal structure of full length GRP94 reveals that the γ-phosphate group of bound 5'-adenylyl-β,γimidodiphosphate makes no direct or water-mediated contacts with any parts of GRP94 [246]. Point mutations in the NBD inactivate GRP94 resulting in defects in galactosyl transferase activity in Leishmania donovani [439] and surface expression of toll-like receptors 1, 2, and 4 [440]. However, mutations that diminish nucleotide binding more severely compromise GRP94 function than point mutations predicted to render the NBD inactive [440]. GRP94 possesses a low ATPase activity in vitro that is comparable to the ATPase activity of other HSP90 chaperones [247, 413, 441-443]. In contrast to yeast

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and human HSP90 and the mitochondrial HSP90, TRAP1 (tumor necrosis factor receptor-associated protein 1), GRP94 remains largely (~97%) [247] in the open state after ATP binding. Trapping of ATP by closure of the ATPase domain (cf. above) is required for ATP hydrolysis. In contrast to yeast and human HSP90 ATP binding does not commit GRP94 to ATP hydrolysis [247]. These biochemical data are in agreement with mutational analysis of GRP94, which shows that mutations in amino acids involved in nucleotide binding have more severe consequences than mutations in residues involved in catalysis of ATP hydrolysis [440]. One explanation may be that nucleotide binding is simply required for stabilization of GRP94. GRP94 already denatures slightly at 30°C [247], whereas other HSP90s start to denature at 4055°C [247, 441]. GRP94 is required for secretion of insulin-like growth factor (IGF)-II [444]. grp94-/- MEFs, while viable in the presence of serum, are hypersensitive to serum starvation, because of their inability to produce IGF-II [444]. Neither GRP94 mutants in which residues involved in nucleotide binding or hydrolysis of the γ-phosphate group fully complement the hypersensitivity of grp94-/- cells to serum deprivation [445]. These experiments, for the first time, reveal a requirement for nucleotide hydrolysis by GRP94 in vivo. It, however, remains to be established what this function of ATP hydrolysis is and how the GRP94 ATP hydrolysis cycle is controlled in vivo. Maybe its own substrates function as co-chaperones to regulate its conformational transitions and its ATPase activity.

4.4.3. Substrate Recognition A second important question is how HSP90 class chaperones interact with their substrates. A protein kinase client, CDK4, binds to the concave surface of the closed, twisted V conformation [446]. Several hydrophobic amino acid residues may be involved in substrate recognition. A methionine pair at the top of the C domain (M658 and M663 in human GRP94) [447], surface-exposed aromatic side chains of Y652, W654 and Y678 in GRP94, and W300 in yeast Hsp82 (W320 in human HSP90α2) [448] may be involved in making hydrophobic contacts to unfolded substrates. W300 is replaced in GRP94 by a hydrophilic amino acid. A stretch of alternating basic and acidic residues, called a KEKE motif [449], may also be involved in protein-protein interactions. These first insights into the substrate-HSP90 chaperone interaction require extension to further substrates, but already indicate that HSP90 class chaperones use an extensive interaction surface to recognize their

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substrates. As a consequence it becomes clear why HSP90 class chaperones prefer partially folded substrates over completely unfolded substrates and hierarchically act after HSP70 chaperones in protein folding. Whatever the substrate recognition mechanism for GRP94 is, its in vivo substrates appear to be limited. GRP94 appears to possess a limited substrate spectrum including immunoglobulin G molecules [450, 451], several integrins [440] and toll-like receptors [452], plant CLAVATA proteins [453], and IGF-II [444]. This may explain why GRP94 is not found in all eukaryotes and why grp94-/- mouse embryonic fibroblasts are viable [454].

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4.5. THE LECTIN CHAPERONES CALNEXIN, CALRETICULIN, AND CALMEGIN Proteins entering the ER are modified co-translationally by transfer of the oligosaccharide Glc3Man9GlcNAc2 (Glc - D-glucose, GlcNAc – 2-Nacetylamide-D-glucose, Man – D-mannose) from dolicholpyrophosphateGlcNAc2Man9Glc3 located in the ER membrane to consensus Asn-X-Ser/Thr residues by oligosaccharyltransferase (OST, Figure 8) [239, 240]. The two terminal D-glucose moieties are rapidly (t1/2 < 3 min [455]) removed by sequential action of α-glucosidases I and II [456]. The monoglucosylated form interacts with the lectin chaperones calnexin, calreticulin, and the testisspecific calnexin homolog calmegin/calnexin-t (Figure 26) [457]. Removal of this glucose by α-glucosidase II releases the unfolded substrate from calnexin/calreticulin/calmegin (Figure 10) [458]. If still unfolded, the enzyme UDP-glucose:glycoprotein glucosyltransferase (UGGT) reglucosylates the Man9GlcNAc2 oligosaccharide. This triggers a second round of interaction with the lectin chaperones [242, 243, 459]. This glycoprotein folding cycle is known as the calnexin/calreticulin cycle [456, 458-460]. The only component of this cycle for which evidence is available that it recognizes hydrophobic amino acid stretches is UGGT [461, 462]. Demannosylation events catalyzed by ER resident α-mannosidases and α-mannosidases residing in the ER Golgi intermediate compartment and the early cisternae of the Golgi complex are critical for extraction of permanently misfolded proteins from the calnexin cycle. In this way the glycosylation status of a newly synthesized protein functions as a molecular clock determining the fate of the polypeptide chain to which the oligosaccharide is attached.

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Figure 26. (A) Domain structures for calreticulin, calnexin, and calmegin. Disulfide bonds are indicated by -S-S-. Amino acid sequences at the C-termini represent ER retention sequences. Circles labeled A represent the acidic repeat sequence PXXIXDPDAXKPEDWDE. Squares labeled B represent the proline-rich repeat sequence GXWXPPXIXNPXYX. (B) Crystal structure of the N and P domains of calnexin (PDB entry 1jhn [465]). The P domain is shown in red, the N domain in blue, and amino acid residues involved in carbohydrate binding in white.

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4.5.1 Structure and Function of Calnexin and Calreticulin Calreticulin, calnexin, and calmegin display a high degree of functional and structural similarity (Figure 26). Calreticulin and calnexin are chaperones for ~100 newly synthesized glycoproteins. Their substrate spectra are overlapping, but not identical [463]. Several viral glycoproteins that preferably interact with calnexin displayed increased interactions with calreticulin in calnexin knock-out cells [22], providing evidence for redundancy between calreticulin and calnexin. However, cellular glycoproteins did not switch over from calnexin to calreticulin and instead were found to be associated with BiP [464]. Calreticulin is a soluble, 46 kDa Ca2+-binding protein of the ER, whereas calnexin and calmegin are 90 kDa and 70 kDa type I transmembrane proteins of the ER, respectively (Figure 26). Both proteins consist of an Nterminal N domain, a middle proline-rich P domain, and a C-terminal C domain. The NMR structures of the P domain of calreticulin and the crystal structure of the N and P domains of calnexin have been solved [465-468]. The N domain consists of a globular β-sandwich formed by eight antiparallel βstrands, has similarity to legume lectin, and functions as a lectin recognizing the monoglucosylated N-linked oligosaccharide GlcMan9GlcNAc2 [469, 470]. In both proteins several residues have been identified that participate in binding to the D-glucose moiety. In calreticulin these are Y109, D135 [470], and possibly K111, Y128, and D317 [471, 472]. In calnexin Y164, K166, Y185, M188, E216, and E425 are important for oligosaccharide binding [473]. Oligosaccharide binding to the N domain induces conformational changes in these chaperones which are important for substrate binding [474]. In addition to recognizing monoglucosylated oligosaccharides calnexin and calreticulin also recognize non-native conformers via the polypeptide backbone of the unfolded protein [458, 475-477]. In vitro calnexin and calreticulin suppress the aggregation of glycosylated, but also of nonglycosylated proteins [474, 478-480], demonstrating a lectin-independent chaperone function. Physiological ER Ca2+ and millimolar ATP concentrations potentiate this chaperone activity [481]. In vivo, lectin-deficient calnexin and calreticulin interact with a broad spectrum of proteins, including heavy chains of major histocompatibility complex (MHC) class I molecules [473, 482]. Screening of peptide libraries has revealed that a stretch of at least five hydrophobic amino acids is a prerequisite for peptide-binding to calreticulin [483, 484]. The polypeptide-interacting region has been mapped to the globular N and the extended P domains of calnexin and calreticulin [469, 470, 485-487]. The P domain is characterized by two repeat sequences (repeat A,

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IXDPXA/DXKPEDWDX, and repeat B, GXWXPPXIXNPXYX, X = any amino acid). Calnexin harbors four [61] and calreticulin three repeats (Figure 26) [64, 65]. The P domain forms an extended structure stabilized by three antiparallel β-sheets [465-468]. Calnexin and calreticulin closely co-operate with the PDI ERp57 in protein folding [469, 488]. The P domain mediates the interaction with ERp57 [467, 485, 489, 490]. Mutagenesis experiments have confirmed that the extreme tip of the P domain is important for interaction of calreticulin with ERp57 [485]. Both the N and P domains bind Ca2+, but the major Ca2+ binding domain in calreticulin is its C domain. The C domain of calreticulin binds >50% of ER luminal Ca2+ with low affinity (Kd ~ 2 mM), but high capacity (25 mol Ca2+/mol of protein) [486]. In contrast, the C domain of calnexin consists of a transmembrane domain, followed by stretches of negatively charged amino acids capable of binding Ca2+ and a C-terminal ER retention motif [61, 491]. Ca2+ binding to and phosphorylation of the C domain of calnexin may regulate interaction of calnexin with other proteins [492-494]. For example, interaction of calnexin with SERCA2b may be regulated by phosphorylation of S562 and contribute to coupling of ER luminal chaperone function to Ca2+ signaling. Phosphorylation of S554 and S564 regulates interaction with the cytosolic sorting protein phosphofurin acidic cluster sorting protein 2 (PACS2), which localizes to ER-mitochondrial junctions [495].

4.5.2. α-Glucosidase II Entry of substrates into the calnexin/calreticulin cycle is controlled by αglucosidase II, a soluble, heterodimeric protein of the ER. The α-subunit carries the catalytic activity and is retained in the ER through interaction with the β-subunit ending in a KDEL ER retention sequence [496-498]. The βsubunit is characterized by stretches with high homology to the mannose binding domain of the mannose 6-phosphate receptor [499]. The mannose 6phosphate receptor homologous domain interacts with mannose residues in the B and C arms of the oligosaccharide enhancing deglucosylation of the interacting oligosaccharide by the catalytic α subunit [500, 501]. Cleavage of the two D-glucose moieties from the oligosaccharide by α-glucosidase II is sequential and may involve release of the substrate from the enzyme after the first hydrolytic event and reassociation of the substrate with the enzyme before the D-glucose moiety is released by α-glucosidase II [502], because of

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different orientations of the two α-glycosidic bonds in space. This transient release of α-glucosidase II from its substrate may allow calnexin/calreticulin to bind to the monoglucosylated form. In this model the affinities and association kinetics of α-glucosidase II, calnexin, and calreticulin determine how monoglucosylated substrates are partitioned between deglucosylation and entry into the calnexin/calreticulin cycle at the ER luminal side of the ER membrane or in the lumen of the ER. α-Glucosidase II displays 20% activity with GlcMan8GlcNAc2 and