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COFACTORS AND CO-CHAPERONES OF THE CHAPERONIN CCT: MECHANISTIC INSIGHTS AND IN VIVO RELEVANCE Peter C. Stirling B.Sc., University of Victoria, 2002 THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY In the Department of Molecular Biology and Biochemistry 0 Peter C. Stirling 2007 SIMON FRASER UNlVERSITY Spring 2007 All rights resewed. This work may not be reproduced in whole or in part, by photocopy or other means. without permission of the author

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Page 1: COFACTORS AND CO-CHAPERONES OF THE ...summit.sfu.ca/system/files/iritems1/2550/etd2862.pdfCOFACTORS AND CO-CHAPERONES OF THE CHAPERONIN CCT: MECHANISTIC INSIGHTS AND IN VIVO RELEVANCE

COFACTORS AND CO-CHAPERONES OF THE CHAPERONIN CCT:

MECHANISTIC INSIGHTS AND IN VIVO RELEVANCE

Peter C. Stirling B.Sc., University of Victoria, 2002

THESIS SUBMITTED IN PARTIAL FULFILLMENT O F THE REQUIREMENTS FOR THE DEGREE O F

DOCTOR O F PHILOSOPHY

In the Department of

Molecular Biology and Biochemistry

0 Peter C. Stirling 2007

SIMON FRASER UNlVERSITY

Spring 2007

All rights resewed. This work may not be reproduced in whole or in part, by photocopy

or other means. without permission of the author

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APPROVAL

Name:

Degree:

Title of Thesis:

Peter C. Stirling

Doctor of Philosophy

Cofactors and co-chaperones of the chaperonin CCT: mechanistic insights and in vivo relevance

Examining Committee:

Chair: Dr. William S. Davidson Psofcssos, Dcpartmcnt of Molccular Biology and Biochemistry

Dr. Michel R. Leroux Scnior Supcrvisor Associatc Profcssor, Dcpnrt~ncnt of Molccular Biology and Biochcmistry

Dr. Christopher T. Beh Supcrvisor Assistant Profcssor, Dcpartmcnt of Molccular Biology and Biochcmistry

Dr. Mark W. Paetzel Supcrvisor Assistant Profcssor, Dcpartmcnt of Molccular Biology and Biochcmistry

Dr. Bruce P. Brandhorst Intcsnal Examincr Profcssor, Dcpart~ncllt of Molccular Biology and Biochcmistry

Dr. \Valid Houry Extcrnal Examincr Associatc Profcssor, Dcpartmcnt of Riochcmistry, Univcsisty of Toronto

Date of Defense: April 3"l, 2007

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SIMON FRASER UNIVERSITYI i brary

DECLARATION OF PARTIAL COPYRIGHT LICENCE

The author, whose copyright is declared on the title page of this work, has granted to Simon Fraser University the right to lend this thesis, project or extended essay to users of the Simon Fraser University Library, and to make partial or single copies only for such users or in response to a request from the library of any other university, or other educational institution, on its own behalf or for one of its users.

The author has further granted permission to Simon Fraser University to keep or make a digital copy for use in its circulating collection (currently available to the public at the "Institutional Repository" link of the SFU Library website <www.lib.sfu.ca> at: ~http:llir.lib.sfu.calhandlell8921112~) and, without changing the content, to translate the thesislproject or extended essays, if technically possible, to any medium or format for the purpose of preservation of the digital work.

The author has further agreed that permission for multiple copying of this work for scholarly purposes may be granted by either the author or the Dean of Graduate Studies.

It is understood that copying or publication of this work for financial gain shall not be allowed without the author's written permission.

Permission for public performance, or limited permission for private scholarly use, of any multimedia materials forming part of this work, may have been granted by the author. This information may be found on the separately catalogued multimedia material and in the signed Partial Copyright Licence.

The original Partial Copyright Licence attesting to these terms, and signed by this author, may be found in the original bound copy of this work, retained in the Simon Fraser University Archive.

Simon Fraser University Library Burnaby, BC, Canada

Revised: Spring 2007

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ABSTRACT

Protein folding is the csscntial process by which a linear chain of amino acids

folds upon itself to adopt a defined three-dimensional structure. All proteins must

undergo folding to be functional and while the amino acid sequence dictates the tertiary

structure, in the crowded cellular environment the folding process is not always

spontaneous. To circumvent this problem proteins called molecular chaperones have

evolved to stabilize non-folded polypeptides and facilitate their transformation to the

folded state. This thesis revolves around a n~olecular chaperone called CCT. which uses a

barrel-shaped structure to bind non-native proteins and sequester them in a protected

environment to allow folding. My work has focused on two proteins that co-operate with

CCT in the cell to promote efficient folding of its substrates. The first, called prefoldin

(PFD), is another inolecular chaperone that binds non-native polypeptides and delivers

them to CCT for completion of folding. The second is a family of proteins called

phosducin-like proteins (PhLPs), which bind CCT and affect its ability to fold substrates.

Here we show that PFD uses long coiled-coil tentacles to grasp substrates using

interhelical-hydrophobic residues at the veiy tips. We find that the gencral properties of a

coiled-coil are sufficient to confer some chaperone activity indicating the importance of

this super-secondary structure to PFD function. We also find that archaeal PFD can alter

its shape to acconmodate substrates of different sizes, but that in most cases a large

proportion of the substrate protrudes from the PFD cavity. We also show that the

nlechanism of PhLP-mediated CCT regulation involves PhLP binding to CCT-substrate

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complexes and slowing of ATP hydrolysis. In yeast the PhLP homologues Plp 1 p and

Plp2p both affect cytoskeletal function but Plp2p, which is essential, also appears to

affect the cell cycle. Finally, we use a genomic approach to suggest novel cellular roles

for the chaperonin CC'T in pathways such as septin ring assembly. Altogether these

studies illuminate the role of C'C'T co-chaperones (PFD) and cofactors (PhLP) in

modulating the chaperonin's function and open up new research prospects by identifying

novel genetic interactors of CCT.

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DEDICATION

To my family. To my fathcr and his for fostering an inquisitive love of nature that

motivates me to ask questions about the natural world and to my mother for instilling a

love of language which allows me to discuss what I find.

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QUOTATIONS

"The surest way to corrupt a youth is to instruct him to hold in higher esteem those who

think alike than those who think differently." - Friedrich Nietzsche, The Duwn

"The sweetest and most inoffensive path of life leads through the avenues of science and

learning; and whoever can either remove any obstructions in this way or open up any new

prospect ought so far to be esteemed a benefactor to mankind." - David Hume, 1748 in

Ail Euyuiry Coi~c~c.iwing H L I / I I L ~ ~ I U / I L / ~ ~ : S ' ~ N I ~ C / ~ ~ I ~

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ACKNOWLEDGEMENTS

I would like to acknowledge the work of the undergraduate studcnts who hclped

me at various stages with the pro-ject, including Jennifer Obst, Jill Mwenifumbo, Melissa

Wong, Braydon Burgess, Naomi Farrell, Matthew Nesbitt, Andrea Feigl, Karam Takhar,

Clairessa Keays and Jayden Yamakaze. I want to thank Junchul Kim, Oliver Blacque,

Muneer Esmail, Nathan Bialas, Michael Healey, Michael Kennedy, Nicholas Inglis, and

especially Victor Lundin for reagents and helpful discussions about my research over the

ycars. I would also like to acknowledge the experimental and intellectual contributions of

our collaborators Drs. Ronald Melki, Jose Valpuesta, Masafumi Yohda, Tomatsu Zako,

Martin Srayko, Keith Willison, Charles Boone, Christopher Beh and members of their

laboratories. I would like to thank the support of my supervisory committee, Dr.

Christopher Bell, Dr. Mark Paetzel and my senior supervisor Dr. Michel Leroux without

wllonl my doctoral work would not have been possible.

v i i

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TABLE OF CONTENTS

.. Approval ............................................................................................................................ 11

... Abstract ............................................................................................................................. 111

........................................................................................................................... Dedication v

Quotations .................................................................................................................. vi

Acknowledgements ......................................................................................................... vii ... Table of Contents ......................................................................................................... VIII

.. List of Figures .................................................................................................................. XII

List of Tables ........................................................................................................... siv

Chapter 1 General introduction and research objectives ........................................ 1 1 .1 Historicalperspectives ........................................................................................ 2

........................................................................................ 1.2 Protein folding in vitrao 3 1.3

. . ......................................................................................... Protein folding r r ~ vwo 5 1.3.1 Cellular functions of inolecular chaperones ................................................ 5 1.3.2 Molecular strategies for binding non-native proteins .................................. 8

1.4 The eukaryotic type 11 chaperonin CCT ............................................................ 12 1.4.1 General features of chapcronins ................................................................. 12 1.4.2 Structure and niechanism of CCT ............................................................. 14 1.4.3 CCT substrate repertoire ............................................................................ 15 1.4.4 The actin and tubulin folding pathways ..................................................... 16

1.5 C'C'T cofactors .................................................................................................. 19 1.5.1 Prefoldin ..................................................................................................... I9 1 S .2 Phosducin-like proteins ............................................................................ I

1.6 Research ob-jectives ........................................................................................... 23 1.7 Figures ............................................................................................................... 25 1.8 Tables ................................................................................................................ 34

..................................................................................................... 1.9 Reference list 35

Chapter 2 Mutagenesis and electron microscopy characterize archaeal prefoldin as a molecular clamp with hydrophobic coiled-coil binding sites .............. 46

2.1 Abstract ............................................................................................................. 47 2.2 Introduction ....................................................................................................... 47 2.3 Mcthods ............................................................................................................. 50

2.3.1 Preparation of constructs ............................................................................ 50 2.3.2 Protein expression and purification .......................................................... I 2.3.3 Characterization of PFD variants ............................................................. 51

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.................................................. 2.3.4 Prevention of protein aggregation assays 52 ............................... 2.3.5 Formation and analysis of PFD-substrate con~plexes 52

2.3.6 Electron microscopy .................................................................................. 53 ........................... 2.3.7 Image processing and three-dimensional reconstruction 54

2.3.8 Miscellaneous ............................................................................................ 55 ..................................................................................... 2.4 Results and discussion -55

2.4.1 Properties of PFD coiled coils .................................................................. 55 .................................................. 2.4.2 Cavity surface forr~ied by the coiled coils 56

............................................... 2.4.3 Intrinsic properties of the coiled-coil motif 58 .................................................. 2.4.4 Hydrophobic interface of the coiled coils I

......................................................... 2.4.5 PFD functions as a molecular clamp 63 2.4.6 The interaction of PhPFD with unfolded proteins ..................................... 66

............ 2.4.7 Con~parison of archaeal and eukaryotic PFD binding mechanism 70 ..................................................................................................... 2.5 C:onclusion 72

............................................................................................................... 2.6 Figures 76 2.7 Tables ................................................................................................................ 91

..................................................................................................... 2.8 Reference list 92

Chapter 3 In vitru and in vivu analyses identify phosducin-like protein 3 ................................................................... as a novel cofactor of the chaperonin CCT 95

........................................................................................................... 3.1 Abstract 96 ....................................................................................................... 3.2 Introduction 96

3.3 Methods ........................................................................................................... 100 3.3.1 Purification of PhLP3 and CCT ............................................................... 100 3.3.2 Cell culture ............................................................................................... 100 3.3.3 It1 vilro translation, folding assays, and GST pull-downs ........................ 101

................ 3.3.4 Verification of actin and tubulin folding inhibition by PhLP3 I01 ................................................................ 3.3.5 ATPase activity measurements 102

................... 3.3.6 Purification of tubulin and microtubule-associated proteins 102 3.3.7 Co-sedimentation assay ........................................................................... 103 3.3.8 Sedimentation velocity measurements ..................................................... 103

........................................... 3.3.9 Sample preparation for electron microscopy 103 3.3.10 Electron microscopy and image processing ............................................. 104

.................................................... 3.3.1 1 Yeast strains, growth, and n~icroscopy 104 .................................................................................... 3.4 Results and discussion 105

3.4.1 Native PhLP3 associates with CCT likely as a monomer using ............................................................................... Both N and C termini 105

............. 3.4.2 PhLP3 forms terna~y conlplexes with CCT and actin or tubulin 107 ........................ 3.4.3 Excess PhLP3 inhibits actin and tubulin folding in vifro 109

........... 3.4.4 PhLP3 inhibits the ATPase activity of CCT bound to a substrate 110 3.4.5 Synthetic interactions of PLPI and prefoldin reveal links to tubulin

and actin function in vivo ......................................................................... 112 ........... 3.4.6 C'ellular defects in yucIOA yeast are enhanced by PLPI deletion 115

3.5 Conclusion ...................................................................................................... 119 ............................................................................................................. 3.6 Figures 121 .............................................................................................................. 3.7 Tables 132

....................................................... 3.8 Reference list ............................................ 134

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Chapter 4 Functional interaction between phosducin-like protein 2 and cytosolic chaperonin is essential for cytoskeletal protein function and cell cycle progression ............................................................................................................ 139

4.1 Abstract ........................................................................................................... 140 4.2 lntroduction ..................................................................................................... 140 4.3 Methods ........................................................................................................... 143

4.3.1 Purification of GST-PhLP2A and CCT ................................................... 143 4.3.2 In vitro translation and folding assays ..................................................... 143 4.3.3 Yeast strains, media and growth assays ................................................... 144

....................................................... 4.3.4 Plp2p, CCT co-immunoprecipitation 144 4.3.5 Drug and mating factor sensitivity assays ............................................... 144

.............................. 4.3.6 Generation of temperature-sensitive alleles of PLP2 145 4.3.7 Microscopy .............................................................................................. 145 4.3.8 High-copy suppression screen ................................................................. 146 4.3.9 Cell synchronization ................................................................................ 146

.................................................................................... 4.4 Results and discussion 147 .............................................. 4.4.1 Plp2p is an essential CCT-binding protein 147

4.4.2 Generation of PLP2 temperature-sensitive alleles ................................... 149 4.4.3 plp2-ts alleles exhibit cytoskeletal but not G-protein-related defects ...... 149 4.4.4 Microtubule and nuclear defects in plp2-ts alleles .................................. 151 4.4.5 Actin polarization defects in plp2-ts alleles ............................................. 153 4.4.6 Mammalian PhLP2A inhibits actin folding in vitro and binds CCT-

actin coniplexes ........................................................................................ 154 4.4.7 High-copy suppression ofplp2-ts alleles reveals links to the Gl/S

phase transition ........................................................................................ 156 4.5 Conclusion ...................................................................................................... 159

....... 4.5.1 PLP2 function is essential for viability but not G-protein signaling 160 ............................ 4.5.2 Cytoskeletal phenotypes in plp2 loss of function cells 160

............................... 4.5.3 Cell-cycle phenotypes in plp2 loss of function cells 162 4.5.4 Models of Plp2p-cell cycle connection ................................................... 162 4.5.5 Perspectives ............................................................................................. 164

4.6 Figures ............................................................................................................. 165 4.7 Tables .............................................................................................................. 180 4.8 Reference list ................................................................................................... 189

Chapter 5 Genetic interactors of yeast CCT and a novel role for the chaperonin in septin ring function ............................................................................... 194

5.1 Abstract ........................................................................................................... 195 5.2 Introduction ..................................................................................................... 195 5.3 Methods ........................................................................................................... 197

5.3.1 Yeast strains and manipulations ............................................................... 197 5.3.2 Microscopy .................................................................................... 1 9 7

5.4 Results and discussion ............................................................................... 197 ................ 5.4.1 Synthetic genetic array of a temperature sensitive CCT allele 197

......... 5.4.2 Analysis of septin function in yeast bearing mutant CCT subunits 199 ...................................................................................................... 5.5 Conclusion 202

5.6 Figures ............................................................................................................. 206

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5.7 Tables .............................................................................................................. 209 5.8 Reference list ................................................................................................... 215

Chapter 6 General Conclusions .......................................................................... 218 6.1 Reference list ................................................................................................... 222

Chapter 7 Appendices .............................................................................................. 224 7.1 Appendix I : An in v i tm expression cloning screen for novel CCT-

interacting proteins .......................................................................................... 224 7.1.1 Tables ....................................................................................................... 226 7.1.2 Appendix 1 reference list ......................................................................... 227

7.2 Appendix 2: Synthetic genetic array of ylplA cells ........................................ 228 7.2.1 Tables ....................................................................................................... 230 7.2.2 Appendix 2 reference list ..................................................................... 231

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LIST OF FIGURES

Figure 1-1 Folding pathways of engrailed homeodomain and egg-white lysozyme ...................................................................................................... 25

Figure 1-2 Structural featurcs of clamp-like chaperones ............................................. 26

Figure 1-3 Alternative structural strategies of molecular chaperones for stabilizing non-native substrates ............................................................... 27

Figure 1-4 Structural and functional features of chaperonins .................................... 29 Figure 1-5 Cellular pathways of folding by CCT ........................................................ 31 Figure 1-6 Structural features of prefoldin. phosducin-like protein and their

complexes with CCT .................................................................................. 32 Figure 2-1 Prefoldin (PFD) constructs used in this study ............................................ 76

.................... Figure 2-2 Multiple sequence alignment of archaeal prefoldin subunits 78 Figure 2-3 Architecture and surface conservation of archaeal prefoldin ................... 81

Figure 2-4 Chaperone activity of intradomain swap (switch) mutant complexes .................................................................................................... 82

Figure 2-5 Chaperone activity of chimeric complexes .................................................. 83

Figure 2-6 Hydrophobic a/d coiled-coil residues are required for chaperone activity ........................................................................................................ 84

Figure 2-7 Substrate binding occurs near the ends of flexible coiled coils ................. 86 Figure 2-8 The three-dimensional reconstruction of the complex between

PhPFD and several unfolded proteins .................................................... 88

Figure 2-9 The role of PhPFDa and PhPFDP subunits in the interaction with unfolded substrates .................................................................................... 89

Figure 2-10 Localization of the unfolded substrates in archaeal and eukaryotic PFDs ......................................................................................... 90

Figure 3-1 PhLP3 interacts with CCT in vivo ............................................................. 121 Figure 3-2 PhLP3 does not form binary complexes with unfolded actin.

tubulin o r different forms of native tubulin .......................................... 123

Figure 3-3 PhLP3 interacts with actin and tubulins in a ternary complex with CCT ........................................................................................................... 125

Figure 3-4 Electron microscopy of PhLP3-CCT and PhLP3-CCT-tubulin complexes .................................................................................................. 126

Figure 3-5 PhLP3 affects the folding of nascent actin and tubulin in vitro .............. 127

Figure 3-6 PhLP3 inhibits the ATPase activity of CCT in the presence of an actin o r tubulin substrate ........................................................................ 128

xii

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Figure 3-7 Synthetic effects of PLPl deletion in pacl0A yeast ................................... 129

Figure 3-8 puclOA cellular defects are enhanced by PLPI deletion .......................... 131 Figure 4-1 PhLP2lPlp2p is an essential CCT binding protein ................................... 165 Figure 4-2 Generation of temperature sensitiveplp2 alleles ...................................... 167

Figure 4-3 PLP2 loss of function does not impact pheromone sensitivity ................ 168

Figure 4-4 p@2-ts cells a re large and have increased sensitivity to cytoskeletal-destabilizing drugs .............................................................. 169

Figure 4-Splj~2-ts cells exhibit aberrant nuclear segregation and spindle orientation ................................................................................................. 170

Figure 4-6 Aberrant chitin levels and localization in plp2-ts and cct-ts cells ............ 172 Figure 4-7 Actin filament organization defects in plp2-ts cells .................................. 174 Figure 4-8 Mammalian PhLP2A binds CCT and modulates its activity in

vitrn ............................................................................................................ 175 Figure 4-9 High-copy suppressors ofplp2-1 indicate a role for PLP2 in cell

...................................................................................... cycle progression 177

Figure 4-1 0 Delayed rebudding in a-factor synchronized plp2-ts cells ..................... 179 Figure 5-1 Septin localization in CCT mutant alleles ................................................. 206 Figure 5-2 Relative growth defects of CCT-ts and cs mutants ................................... 208

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LIST OF TABLES

Table 1-1 List of known CCT substrates ....................................................................... 34

Table 2-1 Thermal stability of prefoldin variants monitored by circular dichroism ellipticity a t 222 nm .................................................................. 91

Table 3-1 Yeast strains used in this chapter ................................................................ 132

Table 3-2 Statistics of cellular yeast phenotypes ......................................................... 133

Table 4-1 Yeast strains used in this chapter ............................................................ 180

Table 4-2 Plasmids used in this study .......................................................................... 181

Table 4-3 Multiple buds in plp2-ts mutants ................................................................. 182

Table 4-4 Thickening of the bud neck junction in plp2-ts mutants ........................... 183

Table 4-5 Nuclear defects in plp2-ts cells ............................................................... 184

Table 4-6 Anaphase entry defects in plp2-ts mutants ................................................. 185

Table 4-7 Spindle misorientation in plp2-ts mutants ................................................. 186 .. 1 able 4-8 Actin organization defects in plp2-ts cells ................................................... 187

Table 4-9 Budding index of plp2-ts cells a t high temperature ................................... 188

Table 5-1 Yeast strains used in this chapter ................................................................ 209

Table 5-2 Verified genetic interactors of cctl-2 by synthetic genetic array ............. 210

Table 5-3 Septin localization in budded cells ........................................................... 213

Table 5-4 Septin localization in unbudded cells .......................................................... 214

Table 7-1 Strong. reproducible CCT co-migrating proteins ..................................... 226

Table 7-2 Less certain CCT co-migrating proteins .................................................... 226

Table 7-3 Synthetic genetic interactions ofplplA ....................................................... 230

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CHAPTER 1 GENERAL INTRODUCTION AND RESEARCH OBJECTIVES

Note regarding contributions: Portions of the following chapter were published as review articles in EMBO

~ - q m - / . s and Nu/zii.t. S/~.IIC/LII.LI/ on(/ MoIemIur BioIogy (Sections 1.2 and 1.3 and Figures 1 - 1 , 1-2 and 1-3). The authors of these articles are listed below.

Article 1 Stirling, P.C.*, Lundin, V.F.*, and Leroux, M.R. (2003). EMBO Reports 4, 565-570.

*These authors contributed equally

Article 2 Stirling, P.C., Bakhoum, S.F., Feigl, A.B., and Leroux, M.R. (2006). Nature Structural and Molecular Biology 13, 865-870.

I contributed approxin~ately 1/3 of the writing for the EMBO repoi./.s article with V.F.L. and M.R.L. contributed the remainder. I contributed nearly all of the writing for thc & l ~ ~ / ~ ~ r ~ S / IWO/~II -LI / lid I M O I C ~ L ' L I I U ~ BioIog~. article with editorial and conceptual contributions from S.F.B., A.B.F. and M.R.L.

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1. I Historical perspectives

Since Antinsen's observations in 1973 that denatured Ribonuclease A could

refold spontaneously upon dilution into a nativc buffer. it has been widely acknowledged

that the primary sequence of a polypeptide encodes all the infolmation needed to dictate

thc final tl~rcc-dimensional structure of thc protein (Anfinsen, 1973). This is important

because when a newly-made polypeptide emerges from the ribosome i t must adopt a

folded three-dimensional struchlre to perform its cellular functions. Folded proteins are

necessaly for all cellular functions, for examplc, to give cells their shape or for enzymes

to recognize their substrates. Moreover, non-folded proteins can be harmful to cells

because of their propensity to aggregate (Kopito, 2000). For many proteins the folding

process is essentially spontaneous although in the crowded environment of the cytosol

certain proteins require assistance to efficiently reach the native state (Hartl and Hayer-

Hartl, 2002). This requil-enient is fulfilled by molecular chaperones - proteins that bind

and stabilize non-nntive polypeptide species and facilitate their transition to the native

state.

The notion of a broad requirement in cells for molecular chaperones was first

posited in I986 by Hugh Pelham (Pelhani, 1986) although specific cases of chapcrones

had been identified earlier (Reviewed in Ellis, 1993). In the past 20 years the field has

grown immensely and literature searches for the term 'molecular chaperone' now yields

more than 20,000 articles (www.pubmed.coni). Genes for chaperones have been found

ubiquitously in all prokaryotic and eukaryotic genomes sequenced to date and many

chaperone genes are essential for life (Kapatai et al., 2006; Ursic and Culbertson, 199 1 ;

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Trott et a]., 2005). Chaperones and protein folding have also been found to modulate

aging in diverse organisms and diseases such as cancer. diabetes, and a host of

neurodegenerative diseases like Alzheimer's, Parkinson's and Huntington's (Behrends et

al., 2006; Kenyon, 2005; Kitamura et al., 2006: True. 2006; Tam et al., 2006; Whitesell

and Lindquist, 2005).

I t can be argued that the cellular folding apparatus ought to be considered an

extension of the central dogma of molecular biology. Protein coding genes are

transcribed and translated to produce a polypeptide, but without proper folding the

genetic information encoded by a gene could not be reflected in an organism's cellular

biology. This chapter will discuss some of the features of protein folding in vilro and in

vivo, focussing on the cellular roles for molecular chaperones. In addition, this chapter

will introduce the central players in this thesis, namely, the eukaryotic chaperonin CCT,

and its cofactors prefoldin and phosducin-like proteins.

1.2 Protein folding in vitro

All protcins are synthesized as linear polypeptide chains that are gradually

extruded from the ribosome. To become functional, these nascent proteins must shield

their exposed hydrophobic rcsidues and adopt a precise tertiary structure. I t has long been

known that primary amino-acid sequences dictate the tertiary structures of proteins, but

how folding into the native state occurs is still the subject of intense investigation. The

folding process is now seen as the downward path that an unstructured polypeptide takes

on a funnel-like free-energy surface representing the stcadily decreasing number of

conformations available to i t as i t reaches its native state (Dinner et a].. 2000).

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For small proteins, such as the engrailed homcodomain protein (7.5 kDa), folding

is thought to begin with a few native-like contacts that, once formed, promote a rapid

transition to the native state (Figure 1-1). In this two-step model, there arc no long-lived

intermediate states, and the seconda~y and tertiary structures form virtually

simultaneously (Daggett and Fersht, 2003).

The folding of larger proteins, such as lysozyme (Figure I-I), is more complex,

and usually involves transition states and intermediates that are represented as peaks and

valleys on the energy landscape. One theory is that on the initiation of folding,

secondary-structure elements form autonomously and collide to produce the tcrtiary

structure (Daggett and Fersht, 2003). De t?ovo, co-translational protein-folding in the cell

probably follows this type of pathway (Hartl and Hayer-Hart], 2002). Another hypothesis

is that the propensity of hydrophobic residues to associate stimulates the collapse of a

protein into a 'niol ten-globulet-like, conlpact state that has some non-na tive contacts. In

this scenario, the folding bottleneck for proteins is the reorganization of such incorrect

associations. A syncrgistic view of these two mechanisms, called nucleation-collapse,

probably explains the folding of most proteins ir? vi/t"o (Daggett and Fersht, 2003).

Folding intermediates, and non-native protein species in general, are usually

aggregation-prone. both in v i / m and in the crowded cellular environment (in vivo protein

concentration is -200mglmL; Siegers et a]., 1999). Therefore, it? vivo, they must bc

stabilized and ushered to their appropriate fate. be it biogenesis (folding, assembly and

transport), degradation, or sequestration into aggregated forms if they cannot reach their

native state or be disposed of.

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1.3 Protein folding in vivo

1.3.1 Cellular functions of molecular chaperones

A diverse and ubiquitous class of proteins, known as niolecular chaperones, has

evolved to transiently stabilize cxposed hydrophobic residues in non-native proteins.

Cellular functions for chaperones include assisting in biogenesis, modulation of protein

conformation and activity, disaggregation and refolding of proteins aftcr cellular stress

and, perhaps unexpectedly, the disassen~bly and unfolding of proteins for subsequent

degradation (Leroux and Hartl, 2000a). Although they are not historically classified as

chaperones, protcins that catalyse the cix-11-LIHS isomerization of proline residues

(peptidyl-prolyl isomerases) and the proper formation of disulphide bonds (protein

disulphide isomerases) often bind and stabilize non-native proteins (Leroux, 200 1). We

now consider some of the best-characterized chaperones, which we classify into three

broad functional categories: holding, folding and unfolding.

Hol~i'ing - Several molecular chaperones seem to have little more than a

stabilizing effect on non-native proteins, and usually require the participation of other

chaperones to assist with, for example, folding. These chaperones typically lack the

ability to undergo ATP-dependent conformational changes.

For example. heat-shock protein 40 (Hsp40) can prevent protein aggregation in

vilro, but requires the ATP-hydrolysing chaperone Hsp70 to fold substrates (Mayer et a].,

2000). Eukaryotic prefoldin, a hetero-hexameric chaperone complex, interacts with

nascent actin and tubulin chains and assists in their biogenesis by docking onto and

delivering substrates to the chaperonin-containi~ig TCP 1 (CCT), which is a cylindrical

'folding machine' (Martin-Benito et al., 2002). Archaeal prefoldin interacts

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indiscriminately with non-native proteins in vitro, suggesting that it boasts a wider rangc

of substrates than its eukaryotic counterpart (Lcroux et al. 1999; Siegert et a]., 2000).

Sniall heat-shock proteins (sHsps) belong to a diverse family of protein complexes that

trap denatured proteins on their surfaces (van Montfort et al.. 2001). I t is controversial as

to whether sHsps can use ATP to directly assist the refolding of their substrates, or

whether they are strictly reliant on ATP-dependent chaperones, such as Hsp70 or

chaperonins, to perform this task (Leroux, 2001). SecB, a bacterial chaperone, recognizes

and maintains newly synthesized precursor proteins in forms that are competent for

translocation by SecA, a chaperone ATPase that threads proteins into the SecYEG

translocon channel (Xu et a]., 2000). Lastly, chaperones such as the periplasmic protein

PapD can stabilize non-native proteins-in this case, unassembled pilus subunits-by

transiently 'donating' a structural element that is lacking in the substrate's tertiary

structure, but that is present (complemented) in the assembled quaternaly form (Sauer et

al.. 2002).

Folding - Chaperones that assist in protein folding often couple a holding (or

capturing) function with the ability to release the folded substrate in an ATP-dependent

manner. Hsp70 has a multitude of functions, two of which are stabilizing nascent

polypeptides and promoting the folding of non-native proteins through rounds of ATP-

dependent binding and release (Mayer et al., 2000). In bacteria, the function of the Hsp70

homologue, DnaK, partially overlaps with that of Trigger Factor (TF), a chaperone and

prolyl isonierase that is located near the ribosomal polypeptide exit tunnel (Teter et al.,

1999). The chaperonin (Hsp60) family of chaperones assists the folding of newly

translated proteins by sequestering aggregation-prone inter~nediates in a hydrophilic

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cavity and releasing them after ATP hydrolysis. Hsp90 is an interesting case of a highly

abundant chaperone that requires the cooperation of Hsp70 and other cofactors to

facilitate conformational switching between active and inactive client proteins despite its

ability to hydrolyse ATP (Pearl and Prodromou, 2002). The bacterial GroEL and

eukaryotic cytosolic CCT chaperonins interact with a range of substrates, although the

latter probably has specialized functions in folding actin and tubulin (Leroux and Hartl,

2000b).

An emerging concept is that chaperones cooperate extensively, sometimes

forming multi-chaperone systems that work sequentially or sinlultaneously lo ensure the

efficient biogenesis of cellular proteins in their respective cellular compartments (Leroux

and Hartl, 2000a). For example, sequential interactions with cytosolic Hsp70, prefoldin

and CCT are probably needed to assist actin and tubulin folding: at least in the case of

actin, prefoldin and CCT also cooperate closely to help i t rcach its native state (Siegers et

a]., 1999; Martin-Benito et al.. 2002). In mitochondria, several newly imported substrates

have been shown to interact first with Hsp70, and then with Hsp60, for productive

folding (Manning-Krieg et al., 199 1 ). As a final example, several endoplasmic reticulum

(ER) chaperones, such as calnexin, calreticulin, Hsp70 and Hsp90 homologues, Erp57

and UDP-gl~~cose:glycoprotein transferase (UGGT) work together or successively to

ensure the correct biogenesis of glycosylated proteins (Parodi, 2000).

Some pre-proteins contain sequences that fulfill the criteria for a chaperone, as

their cleaved pre-domains assist folding without being part of the final stnlchlre. The pro-

peptide of the protease subtilisin E encodes such an intramolecular chaperone (Shinde et

a]., 1993). An autotransporter such as BrkA provides a second example of an

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intramolecular chaperone. This bacterial protein contains a P-domain that fonns a P-

barrel channel, which is required for the proper biogenesis (secretion) of its a-domain

and is subsequently removed by proteolytic cleavage (Oliver et al., 2003).

UnfOlu'irig - Chaperones of the Hsp 1 00IC'lplAAA ' ( ATPase associated with

various cellular activities) ATPase family are ubiquitous ring structures that mediatc

protein unfolding and disassembly. Their activities depend on interactions with several

regions within their substrates and on the convcrsion of the energy gained from ATP

hydrolysis into conformational changes that exert a molecular 'crowbar' effect (Honvich

et al., 1999; Lee et at., 2003b; Leroux, 2001). Most AAA' ATPases typically cooperate

with other chaperones for folding or with proteases for degradation. Yeast Hsp104 and

bacterial C'lpB, for example, can disassemble aggregated proteins (or potentially. can

unfolded kinetically-trapped folding intermediates) and thus allow their reactivation in

conjunction with an Hsp701DnaK chaperone (Glover and Lindquist, 1998; Lee et al.,

2003b). When bound to the ends of proteases such as the eukaryotic and archaeal

proteasomes, and the structurally rclated bacterial HslV, the respective A A A ' ATPase

chaperones (Rpt subunits, PAN and HslU) promote the unfolding of their substrates and

their subsequent threading into the proteolytic chamber (Leroux, 2001).

1.3.2 iMolecular strategies for binding non-native proteins

The strategies used by molecular chaperones to stabilize non-native proteins,

which have been elucidated mainly by X-ray crystallography and cryoelectron

microscopy (c~yo-EM) studies, can be grouped broadly into one of four classes: the use

of clamps, cavities, specialized surfaces, and structural complementation.

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Clcrrnps - Recent crystallographic advances have shown that clamp-like stnictures

are one of the most common strategies employed by chaperones to stabilize non-nativc

proteins (Reviewed in Stirling et al., 2006b). The cradle-like structures of Trigger Factor

(TF) and SurA, the prong-like Hsp40 dimer, the jaws of Hsp70, the octopus-like

stn~ctures of prefoldin, Skp and Tin19.Tin110 or the dimeric Hsp90 structure all reveal

architectures designed to grasp substrate proteins in some manner. Indeed, this strategy

may be optimal for stabilizing niost non-native proteins (Figure 1-2; Ali et al., 2006:

Bitto and Mackay, 2002; Ferbitz et al., 2004; Harris et al., 2004; Huai et a]., 2005;

Korndorfer et al., 2004; Li et al., 2000; Ludlum et al., 2004; Meyer et al., 2003a; Sha et

al.. 2000; Siegert et al., 2000; Walton et al., 2004 Webb et a]., 2006; Zhu et al., 1996).

Mechanistically, the clamp-likc chaperones function in one of two different

manners, some using ATP hydrolysis to drive an active clamping process, and others

sin~ply having a structure that ii~nately grips and stabilizes non-native proteins. This sets

Hsp7O and Hsp90 apart from the other chaperones as they have defined nucleotide

hydrolysis-dependent reaction cycles with a limited time of interaction with substrate

proteins. The 'holdase' type chaperones do not have a regulated reaction cycle and

stabilize their substrates until some event causes release - for example, continued

translation of a doniain for ribosome-bound-TF, docking with other molecular

chaperones for PFD and Hsp40, or docking with membrane-bound receptors for Skp and

Tim9.TimlO. Interestingly. other than Hsp70, TF and SurA, which form clamp-like

features as mononiers, all of these chaperones need to oligomerize to function.

Oligornerization provides a way for two or more similar binding sites to be arrayed

around a space or cavity in which a non-native substrate protein can be partially

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sequestered. For example, Hsp40 proteins form a simplc two-pronged pincher to hold

substrate proteins while PFD displays six binding sites around a large central cavity

(Reviewed in Stirling ct al., 2006b).

CUL'III~S - Chaperonins consist of two stacked, oligomeric rings that form

chambers used to sequester non-native proteins and assist in their folding (Holwich et al.,

1999). Individual subunits are composed of a substrate-binding (apical) domain, which

lines the opening of the cavity and includes essential hydrophobic residues (nlostly

leucines, valines and tyrosines), an intermediatellinker domain, and an equatorial ATPasc

domain. The multivalent binding of substrates to the homo-oligomeric bacterial protein,

GroEL, was shown elegantly by sequentially mutating the substrate-binding sites of a

single polypeptide that encodes a conlplete heptameric ring (Farr et a]., 2000). Unlike

GroEL, the eukaryotic chaperonin CCT has eight unique binding sites per hetero-

oligomeric ring that are less hydrophobic in character. These probably evolved to bind a

wide spectrum of substrates while retaining some specificity; consistent with these

results, cryo-EM images show that actin and tubulin interact with two or more specific

subunits of CCT (Figure 1-3; Leroux and Hartl, 2000b: Llorca et al., 2000). During

chaperonin-assisted folding cycles, the substrates of GroEL and CCT are either

encapsulated by a cofactor (GroES) or an iris-like structure contained within CCT itself,

respectively (Hartl and Hayer-Hartl, 2002).

A A A ' ATPases form large, hexameric toroids, which probably also bind

substrates in a multivalent manner. The exact position and nature of their binding sites is

poorly understood, but they are probably adapted to the specific functions of the AAA'

ATPase in question. The function of these chaperones in unfolding or disassembly scenls

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to involve coupling substrate intcractions with large, nucleotide-dependent

conformational changes (Homicli et a]., 1999). As shown for the bacterial HslUV

chaperone-protease coniplex (Figure 1-3); many AAA' ATPases associate coaxially

with proteases, and are poised to untangle and unfold polypeptides and thread them into

the central proteolytic chamber (Bochtler et al., 2000).

Sur.Jircw - To prevent inappropriate interactions between non-native proteins and

components of the bulk cytosol, many chaperones use a relatively flat or corrugated

surface to bind exposed, unstable polypeptide regions. While using a surface often

suggests a more specific interaction than takes place in a clamp or cavity, this is not

always the case, as for the general secretion chaperone SecB. The crystal structure of

bacterial SecB ( 1 7 kDa) shows a dimer of dimers that has two hydrophobic grooves on

opposite faces of the niolecule (Figure 1-3). It has been suggested that a linear

polypeptide can wrap around a SecB tetranier, contacting both groovcs simultaneously

(Randall and Hardy, 2002). Unlike SecB the P-tubulin-specific chaperone cofactor A,

which is dimeric in yeast (Steinbacher, 1999; Figure 1-3), but apparently monomeric in

humans (Guasch et al., 2002), stabilizes its partially folded substrate on a primarily

hydrophilic surface.

A unicjue mechanism for stabilizing non-native proteins is used by sHsps.

~ M c ~ t l ~ a n o c ~ o c . c ~ w . j ~ ~ ~ ~ i ~ ~ ~ . s c ~ l ~ i i Hsp 16.5 is a spherical conlplex that is assembled from 12

dinieric building blocks (Figure 1-3; Kim et a]., 1998). van Montfort and colleagues

(200 1) found that subunits of a wheat sHsp (Hsp 16.9) form a smaller, cylindrical,

dodecameric complex. They suggested, from the structure and from otlier studies, that

sub-assembled species (dimers) dissociate to partition their exposed hydrophobic residues

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betwccn substrate b~nding and higher-order assen~bly. On oligomerization, i t is likely that

non-native proteins are not contained within the cavity of the less ordered conlplexes, but

are instead held on the outside surface, awaiting refolding by othcr chaperone systems.

Strticttird c ~ o r ~ ~ p l ~ ~ i ~ i ~ i i t ~ ~ t i o t t - Structural complementation is an exceptiorlal case,

in which a chaperonc contributes specific structural information to stabilize a non-native

substrate. The interaction between Pap pilus subunits and the periplasrnic chaperonc,

PapD, from uropathogenic E.vchcr.ic~l~iu coli illustrates the use of this strategy. The pilus

subunit contains an in~m~inoglobulin fold that lacks a P-strand, which, in the assembled

pilus structure, is provided by the neighbouring pilus subun~t. Before assembly. PapD

comple~nents, and thus stabilizes, a pilus subunit by donating a P-strand in an analogous

manner (Figure 1-3: Sauer et a]., 2002).

1.4 The eukaryotic type I1 chaperonin CCT

1.4.1 General features of chaperonins

Chaperonins are one of only a few ubiquitously conserved families of molecular

chaperone. Even the otherwise ubiquitous small heat shock protein is absent in certain

pathogenic mycobacteria (Kappe et al., 2002). The importance of chaperonins for cellular

viability is illustrated by their being encoded by essential genes in archaea, bacteria and

eukaryotes (Kapatai et al., 2006; Tilly et a]., 1981: Ursic and Culbertson, 1991).

The overall structure of all chaperonins resen~bles a barrel with cavities at each

open end of the cylindrical structure (Figure 1-4A). The structure is formed by two rings

of -60kDa subunits stacked on top of one another in a back-to-back fashion. Each

subunit consists of three major domains: the eq~~atorial domain which comprises the

intcr-ring interface and is responsible for ATP binding and hydrolysis, the apical domain

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which is responsible for contacting substrate proteins, and the intermediate domain which

transmits allosteric signals between the equatorial and apical domains during a folding

reaction. Archaeal and euka~yotic chaperonins, tcrmed type 11 chaperonins, also have a

unique helical insertion in their apical domains which acts to enclose a bound substrate

protein in the cavity during the ATPase-driven folding cycle (Leroux and Hartl, 2000).

Bacterial chaperonins (also called Type I chaperonins) are found in eubactcrial

cytosols as well as in niitochondria, chloroplasts and in some archaea, which carry both

type 1 and I1 systems (Klunker et al., 2003). Type I chaperonins differ from their archaeal

and eukaryotic counterparts in several ways. First, bacterial chaperonins always have

only 7 subunits per ring, forming a 14-subunit complex canonically represented by E, coli

GroEL (Figure 1-4A). Secondly, since type I chaperonins do not have a helical extension

in their apical domains to encapsulate substrates they co-operate with a~iotlier heptanieric

protein complex called GroESICpn 10. GroES acts as a dissociable lid for GroEL by

docking to GroEL-substrate conlplexes and e.jecting the substrate inward to the aqueous

GroEL cavity for folding. A niodcl of the reaction cycle of type I chaperonins is shown in

Figure I-4B (See Figure Legend; Farr et al., 2003; Martin and Hartl, 1997). While Type

I chaperonins like GroEL bind non-native polypeptides proniiscuously in vitro, the in

vivo substrate repertoire of E. c-oli GroEL I-e\~eals only -250 strongly interacting proteins,

perhaps only -85 of which critically require GroEL f~~nctioii for folding (Houry et al.,

1999; Kerner et a]., 2005).

Archaeal genomes generally encodc 1, 2 or 3 type I 1 chaperonin subunits which

can assemble into con-~plexes with two stacked (usually) 8 membered rings (Kapatai et

a]., 2006; Martin-benito et a]., 2007). These complexes are often called the therniosome

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because they were initially isolated from organisms with very high optimal growth

tcmperaturcs and rcprescnt one of the major heat shock proteins in archaea (Trent et al.,

I99 1). Thermosomes have helical extensions likc CCT which closc in rcsponse to thc

nuclcotide status of the chaperonin (Gutsche ct a]., 2000; Meycr et a]., 2003b). The

structure of an archaeal thermosome is shown in Figure 1-4A and highlights the overall

similarity among all chaperonins. In general, type I 1 chaperonins arc less wcll

characterized than GroEL but recently, significant progress has becn made in

understanding structural and functional details of the eukaryotic chaperonin CCT, which

is discussed in greater detail below.

1.4.2 Structure and mechanism of C C T

The eukaryotic cytosolic chaperonin homolog is called CCT (Chaperonin

Containing Tailless complex polypeptide 1) or TRiC (TC'PI -Ring Complex) and is

found in the cytosol of all eukaryotes sequenced so far. Its three dimensional structure is

superficially similar to the thermosome (Figure 1-4) although it exhibits a more complex

subunit composition of 8 unique but related subunits arranged in two rings to form a 16

subunit oligomer (compared to 1-3 types of subunits in the thermosome) (Leroux and

Hartl, 2000; Llorca et al., 1999). CCT also has more hydrophilic substrate binding sites in

its apical domains than either GroEL or thermosome, likely because of its (relatively)

small and specific substrate repertoire (see 1.4.3 and Table 1-1; Spiess et al., 2004).

A simplified model of a CCT-mediated folding reaction is shown in Figure I-4D.

ADP-bound CC'T binds to an unfolded protein, either directly or after delivery by other

chaperones such as Hsp70 or prefoldin (Siegers et a]., 2003; Vainberg et al., 19%). CCT

then undergoes nucleotide exchange of ADP for ATP. The transition state of nucleotide

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hydrolysis has been shown to close thc lid forming a folding chamber in which thc

substrate is encapsulated (Meyer et a]., 2003b). Inorganic phosphatc and a possibly now-

native substrate protein are released together, restoring the initial ADP-CCT complex

(Figure 1-4). Mechanistically, CCT appears to utilize positive intra-ring co-operativity

and negative inter-ring co-operativity (Kafri and Horovitz, 2003; Kafri et al., 2001). In

other words, when one ring is occupied with nucleotidc. the affinity of the othcs ring for

nucleotide is reduced; the result of which is an innately asymmetric reaction mechanism

which keeps the two rings in different states of the folding cycle.

1.4.3 CCT substrate repertoire

The best characterized substrates of CCT are the cytoskeletal proteins actin, cx-

tubulin and P-tubulin. Indeed, along with y-tubulin, these substrates were long thought to

be the only substrates of CCT. CCT is likely to fold most actin and tubulin related

proteins; more recently, however, i t has become apparent than CCT achlally has many

additional substratcs, and several genome-wide studies are underway to try and extend

the known repertoire (Siegers et a]., 2003; Thulasiraman et a]., 1999). A review by Spiess

et al., (2004) summarizes the known CCT substrates (see also Table 1-1).

Aside from the commonalities between actin and its related proteins and between

a, P and y tubulin, CCT substrates also appear to be proteins that assemble into

oligomeric complexes (Table 1-1). CCT is also involved in the folding of several cell

cycle regulatory proteins (Camasses et al., 2003; Siegers et al, 2003). 111 yeast, CCT was

shown to be important for the biogenesis of the Anaphase Promoting Complex (APC:)

regulators Cdc20p and Cdh I p (Camasses et al., 2003). CC'T has independently been

shown to assist the biogenesis of the cell-cycle phosphatase regulator Cdc55p as well as

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the secretory protein Scc27p and the peroxisonml protein Pcx7p (Siegers ct a].. 2003).

Interestingly. the common features of these substrates seem to be a sequence motif called

a WD repeat. These structures f o m a propeller-likc structure out of repeating P-sheet

motifs (Figure 1-5A). Certain so-called j3-propeller proteins are thought to have

difficulty folding because their N- and C-termini must meet in order to form a stably

folded tertiary structure. The cylindrical CCT cavity apparently provides an environment

in which the topological requirements of these proteins have time to be met. Indeed,

proteomic studies reveal that numerous proteins with WD-repeats associate with CCT,

most of which are likely to be substrates (Cavin et al., 2006; Ho et al., 2002; Krogan et

al., 2006; Valpuesta et al., 2002).

Another better-characterized substrate of CCT is the tumour suppressor protein

VHL (Von Hippel Lindau). VHL is part of an E3-ubiquitin ligase con~plex whose

assembly with its subunits ElonginB and Elongin C is facilitated by CCT (Feldman et a].,

1999). Moreover, disease causing mutations in VHL have been shown to disrupt the

normal interaction of nascent VHL with CCT, highlighting the importance of the

chaperonin in the progression of VHL-related cancers (Feldman et al., 1999; Feldman et

a]., 2003).

1.4.4 The actin and tubulin folding pathways

The folding of cytoskeletal proteins places a unique demand on the folding

machinery both because of the abundance of these proteins and the need to tightly

regulate their levels. Protein folding is key to controlling the dynamic behaviour of

polymerizing microfilaments and microtubules. The activities of both the actin and

tubulin cytoskeleton arc dependent on proper monomer folding.

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Actin folding is somewhat simpler than tubulin in that it requires only the co-

chaperone prefoldin and the chaperonin CCT to reach thc native state, although this

process is also regulated by phosducin-like proteins (Mclaughlin et a]., 2002; Siegers et

a]., 1999; Stirling et a]., 2006a) (Figure 1-5B). CCT is critically required for this process

and prefoldin seems to greatly increase the speed and yield of actin folding (Siegers et al.,

1999). It is known that actin binds to both CCT and PFD in a quasi-native state and thc

structure of this quasi-native state has been modeled extensively (Llorca et a]., 1999;

Martin-benito et a]., 2002; McCormack et a]., 2001). Moreover, the folding of kinetically

trapped folding intermediates has been recently examined in vitro using purified

components (Pappenberger et al., 2006). The current model characterizes the folding-

competent intermediate as an extended form of actin moving around a hinge region at the

base of its nucleotide-binding cleft. CCT contacts actin at the two regions apical to this

hinge and somehow actively closes the conformation to make globular actin (Figure 1-

3A; Llorca et a].. 200 1 ; McCormack et a1.,200 1 ; Pappenberger et a]., 2006). Whether this

happens in one step or in multiple rounds of folding and release by CCT is contentious

and support exists for both models (Farr et a]., 1997; Siegers et a]., 1999; Pappenberger et

nl.. 2006). In yeast. actin filaments form patches and cables mediated by the Arp213

complex and the formins, respectively (Evangelista et a]., 2003). The polarization of actin

patches to the growing bud is essential for proper budding and for endocytosis. Actin

cables play a large role in organelle transport and other polarization processes during

budding. Actin also affects cell wall functions, such as the deposition of chitin.

In addidon to the CCT and PFD complexes, tubulin foldjng requires at least 7

additional proteins to effectively reach its native state. The need to assemble two

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polypeptide chains, a- and j3-tubulin, is likely the main reason for this added complexity.

When a tubulin chain is produced, i t is probably escorted to the CCT complex by PFD

(Geissler et al., 1998; Vainberg et a]., 1998). Interaction with CCT allows both a- and P-

tubulin to reach a quasi-native form that can go on to interact with other folding cofactors

(Figure I-SC). More recent studies, including some presented in this thesis, have also

shown that another CCT cofactor called Phosducin-like protein 3 modulates CCT-

mediated tubulin folding at the level of the CCT-substrate coniplex (Lacefield and

Solomon, 2003; Stirling et al., 2006n). Aftcr release fiom CCT, a-tubulin interacts with

either cofactor B (COB) or cofactor E (CoE) and j3-tubulin interacts with either cofactor A

(CoA) or cofactor D (COD) (Lopcz-fanarragga et al., 2001) (Figure 1-SC). Cofactors A

and B act as sinks for excess quasi-native a and P-tubulin respectively, and prevent the

aggregation of tubulin until i t can be released to CoE (for a ) or COD (for P). The COD+-

tubulin and CoE-a-tubulin complexes come together in a multinieric complex with

Cofactor C (CoC). This complex facilitates the dimeriyation of tubulin and releases a

soluble GDP-bound aij3 heterodimer from the cofactors (Figure 1-5C). Additionally, a

protein related to ADP-ribosylation factors called Arl2iCin4, in its GDP-bound form

regulates the interaction of COD with j3-tubulin and in this way further regulates the

pathway (Figure 1-SC). Finally, following nucleotide exchange of GDP for GTP, the

heterodimer is competcnt to assemble into growing niicrotubules. Microtubulcs are

essential for many intracellular transport processes and for cytokinesis. However, the

most well known role for microtubules lics in aligning and then segregating

chromosomes to the two spindle poles during mitosis.

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1.5 CCT cofactors

1.5.1 Prefoldin

In 1998 two groups rcported the discovcry of a novcl co-chaperone for C'C'T; one

group characterized the genes as involved in tubulin biogenesis in S. ccwvisitrc. (Geisslcr

et a]., 1998) and another group purified the co-chaperone as novel actin-binding activity

in rabbit reticulocyte lysate (Vainberg et al., 1998). The chaperone was called GiniC

(Genes Involved in Microtubule biogenesis Coniplex) i n yeast and prefoldin in mamnials

(Geissler et a]., 1998; Vainberg et al., 1998). Thus, GiniCIPrefoldin is a conserved co-

chaperone involved in cytoskeletal biogenesis.

Prefoldin (PFD) honiologues are found in all studied eukaryotes and archaea and

have been shown to co-operate with their chaperonins, CCT and thermosome,

respectively (Leroux et al., 1999; Okochi et a]., 2002; Vainberg et a]., 1998). In

eukaryotes, PFD consists of six unique but related subunits which assemble to form a

hexamer. In archaea, PFD is typically encoded by one a-type and one P-type subunit

which form an a 2 P 4 hexamer. The atomic structure of archaeal PFD reveals a jellytish-

shaped oligorner with six long coiled-coil tentacles protruding from a double P-barrel

base (Figure 1-6; Siegert et al., 2000). Eukaryotic PFD, whose structure is only known

by electron microscopy, adopts an essentially identical configuration to its archaeal

counterpart except for a kink which may exist in its outer P-type tentacles (Martin-benito

et al., 2002). Coiled coils are made up of two or more amphipathic a-helices which wrap

around one another and are composed of heptad repeats (abcdefg) with hydrophobic

residues at the a and d positions (Lupas, 1996). The hydrophobic residues interlock like

knobs-into-holes to form a supercoiled structure (Lupas, 1996).

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Substrate binding by thc PFD complex takcs place within the tips of the tentacle-

like coiled coils. For eukaryotic PFD the substrate is likely to be completely encapsulated

by approxiniately the distal 113 of the coiled coils, as has been observed for actin (Martin-

benito et al., 2002). For archaeal PFD, a more general type of substrate binding by

hydrophobic patches within the very distal tips of PFD has been shown (See Chapter 2:

Lundin et al., 2004; Martin-benito et a]., 2007). Unlike CCT, eukaryotic PFD appears to

be very specific for actins and tubulins and niay even recognize a coninion sequence

motif in the two unrelated protein families (Roninielaere et al., 2001).

PFD can be crosslinked to nascent polypeptide chains and may act in proximity to

the ribosome to stabilize (recruit) actin and tubulin chains for delivery to the chaperonin

CCT (Hansen et a]., 1999). It has also been established that PFD delivers nascent chains

directly to chaperonins by physically contacting the top of the cavity near the apical

domains (Figure 1-6; Martin-benito et al. 2002; Martin-benito et a]., 2007; Okochi et al.,

2004; Zako et al.. 2005). In the archaenl system, the substrate binding site overlaps with

the chaperonin binding site and chaperonin binding facilitates the release of substrate

protein from PFD (Okochi et al., 2004; Zako et al., 2005). Together these data sub, luest a

co-operative handoff mechanism for substrate transfer from PFD to chaperonin in which

chaperonin binding induces release of substrates from the PFD cavity to the receptive

chaperonin cavity. While it is likely that PFD acts to deliver newly made polypeptides to

a chaperonin, i t may also serve a 'quality control' function by binding not-yet-native

species of polypeptide that have been released from CCT after an unproductive folding

cycle. In this instance, PFD would re-deliver the substrate to CCT for another round of

folding.

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Loss of a PFD subunit in ycast leads to an array of cytoskeletal defects bascd on

aberrant actin and tubulin folding (Geissler et al., 1998; Vainbcrg et al., 1998). Deletions

of more than one PFD subunit does not have additional detectable defects and loss of all

six PFD genes y~elds an essentially identical phenotype to loss of one subunit (Siegers et

al., 1999; Siegers et a]., 2003). PFD is critical for maintaining an efficient speed and yield

of actin folding in ycast (Siegers et al., 1999). While loss of PFD function also rcduccs

the yield of tubulin folding, i t seems to affect the production of a-tubulin more than that

of P-tubulin (Alvarez et al., 1998: Lacefield and Solomon, 2003; Siegers et al., 1999).

The resultant imbalance is highly toxic to cells since excess P-tubulin interferes directly

with normal microtubule function. Interestingly, in the absence of certain Hsp70

homologues, PFD has been shown to interact with and assist the folding of substrates

other than actin or tubulin that i t does not nornially bind or has a lower affinity for

(Siegers et a]., 2003). This suggests that under abnormal conditions PFD can take on a

broader cellular role than i t normally serves. PFD function has also recently been

characterized in the model nematode C. elepns (Lundin et al.. 2007, submitted). In

worms, the PFD complex seems to have a greater role in tubulin function than in actin

function although these actin-related phenotypes may be more subtle in the worm. One

important difference between the yeast and the worm system is that PFD function 1s

essential in C. p I c y p ~ ~ ~ likely because of the more complex requirements for cytoskeletal

function in the multicellular eukaryote (Lundin et al., 2007, submitted).

1 S .2 Phosducin-like proteins

In 2002 the Phosducin-like Protein 1 (PhLP 1) was shown to interact with CCT

and interfere with thc folding of actin. PhLP1 bound CCT in a native state and was

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clearly behaving differently than a substrate protein (McLaughlin et a]., 2002). This led

the authors to propose a regulatory role for Phosducin-like Proteins, which has sincc been

supported by several studies and by work presented here in chapters 3 and 4 (Lacefield

and Solomon, 2003; Lukov et al., 2005; Lukov et a]., 2006; McLaughlin et a]., 2002;

Ogawa et a]., 2004; Stirling et a]., 2006a).

PliLPs arc sn~all protcins (-25-35kDa) that consist of an N-terminal helical

domain, a thioredoxin-like fold and short charged C'-terminal extension (Figure 1-6).

Three subfanlilies of PhLPs exist, namely PhLPI, PliLP2 and PhLP3 (Blaauw et al.,

2003). PhLPl proteins have n role in CC'T-mediated folding of heterotrilneric G-protein

subunits and actually contact the GP subunits directly, possibly as a nieans to facilitate

assenlbly with the Gy subunit (Blaauw et a]., 2003; McLaughlin et al., 2002; Lukov et al.,

2005; Lukov et al., 2006). PhLP3 proteins havc a role in tubulin folding in yeast and C',

e1cgcm.s and we show in Chapter 3 that they regulate actin and tubulin folding by C'C'T it1

vitr.o and iri vivo by contacting CCT-substrate coniplexes and slowing ATP hydrolysis

(Lacefield and Solomon, 2003; Ogawa et al., 2004; Stirling et al., 2006a). PhLP2 proteins

are not well-characterized although i t is known that they are essential in yeast and

Dic~tyostelium d i ~ ~ ~ o i ~ I ~ z i r ? l and that yeast PhLP2 physically interacts wi tli CCT (Blaauw et

al., 2003; Flanary et al.. 2000; Gavin et al., 2006).

PhLPs contact C'CT above the opening of one cavity spanning between apical

domains and inducing conformational changes in both the c0i.s and tr.trt7.s-rings of the

chaperonin (Figure 1-6; Martin-benito et a]., 2004; Stirling et al., 2006a). How the

allosteric signals are transmitted and whether the substrate binds to C'CT in the same ring

as PhLP or in the opposite ring is unknown. I t is clear that there would be sufficient room

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for a substrate to occupy the CCT cavity undesneath the lid created by the PhLP and

possibly also the CCT apical cxtensions. The study of PhLPs as they relate to chaperonin

function is in its infancy and Chapters 3 and 4 will show how our work has opened up

some ncw prospects in this rcgard.

1.6 Research objectives

Molecular chaperones serve a variety of essential cellular functions and,

chaperonins, unlike most other chaperones, are ubiquitously conserved and ubiquitously

essential (Kapatai et al., 2006; Tilly et a]., 198 1 ; Ursic and Culbertson, 199 1). CC'T

function serves a critical role in the biogenesis of the eukaryotic cytoskeleton but also

affects numerous other processes including the progression of the cell cycle. In recent

years mutations in the chaperonin have been implicated in several neuropathics and in

neuml development (Bouhouche et al., 2006: Lee et a]., 2003a; Matsuda and Mishina,

2004). Morcover CC'T is responsible for folding the VHL tumour suppressor which,

when mutated, loses its interaction with CCT and results In disease (Feldn~an et a].,

2003). Most recently. CCT has recently been shown by three different groups to

modulate the folded state of the polyglutamine-expanded proteins responsible for

diseases like Huntington's (Behrends et al., 2006; Kitamura et a]., 2006; Tam et a].,

2006). A basic understanding of the CCT folding machine can therefore help understand

cytoskeletal evolution and biogenesis, numerous disease states, and the cell cycle.

The goal of my doctoral research has bcen to gain novel mechanistic details of

how the CC'T cofactors PFD and PhLP(s) affect folding. Understanding how cofactors

like PFD or PhLP regulate CCT function in the cell will help to integrate the poorly

understood chaperone in to the greater cellular milieu. Cofactors that regulate substrate

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load~ng and nucleotide hydrolysis are found frequently for other molecular chapcrones

but were conspicuously absent in C'C'T research until relatively recently. Understanding

the effects of PFD and PhLP on the CCT reaction cycle and substrate recognition by C'C'T

will help define a more conlplete picture of CCT f~~nct ion in vivo.

Chapter 2 integrates two collaborative studies aimed at determ~ning the substrate

binding mechanism of archaeal PFD using a con~bination of biochemical techniques and

clectron microscopy. Chapter 3 describes the characterization of PhLP3 function in vitiw

and in the yeast S. cser-evisiuc as i t pertains to CCT and PFD function, respectively.

Chapter 4 presents our initial characterization of PhLP2 proteins with respect to CCT

function, primarily in yeast but also using inaninialian PhLP2. Finally, Chapter 5

describes a collaborative effort aimed at gaining new inforniation about CCT in vivo by

screening for synthetic genetic interactions. Our genomic data, combincd with that of a

collaborator, suggest a novel role for CCT in septin ring function.

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1.7 Figures

Figure 1-1 Folding pathways of engrailed homeodomain and egg-white lysozyme

(A) The cngrailed homeodomain (En-HD) folds rapidly, in nanoseconds to microseconds

(Mayor et a]., 2003). (B) Lysozyme has two significantly populated intcrmcdiatcs and

folds more slowly, with a timescale of n~illiseconds in vifro. The majority (70%) of the

lysozyme protein population folds relatively quickly into the a-domain intermediate, but

is slow to reach the near-native short-live UP-intermediate. Another 20% rapidly forms

the ap-intermediate directly. The a-domain is shown in red, and the P-domain in yellow.

(Taken from Stirling et al.. 2003).

Unfolded , En-HD

1 very fast

1 very fast

1 Unfolded 20% 70% lysozyme

slow I

very fast

/7 J r i b intermediate

Native

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Figure 1-2 Structural features of clamp-like chaperones

Surface representations of (A) Trigger Factor (TF, PDB ID: 1 W26), (B) yeast Hsp40

substrate-binding domain (Sis 1, I Cl3C;), (C') Prefoldin ( 1 FXK), (D) Skp ( I U2M), (E)

Tim9aTim I0 (2BSK), (F) the Hsp70 substrate (DnaK, I DXK), (G) Hsp90 (2CG9). For

TF (A) the ribosome-binding site is in green, the substrate-binding domains are in light

and dark yellow and the peptidyl prolyl isomcrasc domain is in bluc. For Hsp40,

prefoldin and Hsp90 (B, C and G) known or suspected substrate-binding sites are

coloured in red. For (E) Tim9 is light green and Tim 10 is dark green. For Hsp70 (F), a

peptide is shown within the clamp as orange spheres. (Taken from Stirling et a]., 2006b).

Trigger Factor -3

Prefoldin

Hsp40 (Sisl)

Hsp70 (DnaK)

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Figure 1-3 Alternative structural strategies of molecular chaperones for stabilizing non-native substrates

( A ) EM reconstructions of CCT-actin (left) and CCT-tubulin (centre) complexes and a

ribbon model of the atomic structure of HslU-HslV ( 1 (331) (right). Actin and tubulin are

shown in red. (B) Atomic surface representations of SccB ( 1 FX3) (left), Cofactor A

( I QSD) (centre) and small heat shock protein 16.5 (sHSP 16.5, I SHS) (right). For SecB

and Cofactor A putative substrate-binding residues are coloured red and for sHSP16.5

dimer subunits are coloured uniquely.

Cavities

SecB

Surfaces

Cofactor A Small HSP 16.5

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(C) Ribbon model of the PapD-PapK ( I PDK) complex showing strand complementation

(red strand) of the PapD chaperone (grey) to the PapK pilus subunit (gold). (Taken from

Stirling et a]., 2003)

Structural Complemenation

PapD-PapK

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Figure 1-4 Structural and functional features of chaperonins

(A) Crystal structures of GroELIES complex (left) and archaeal thermosome in the closed

conformation (right) (PDB ID'S: I SVT and I A6D respectively) (B) Reaction cycle of

GroELIES system (Farr et al., 2003; Martin and Hartl, 1997). A non-native protein binds

to the t i m s ring (upper left). Next, GroES and ATP join the substsate-bound ring,

ejecting ADP and GroES from the oppositc ring and encapsulating thc substrate (uppcr

right). The time needed for ATP hydrolysis allows the substrate to fold within the cavity

(lower right). Finally, ATP and GroES bind the 1)-crns ring and eject GI-oES, nucleotide

and the folded substrate protein (lower left).

cis

7 ATP n =

ADP

ATP I-lydrolys~s 10- 15 seconds

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(C) Doniain structure of type 11 chaperonins. Structural domains are labeled on the left

and functional features of each domain are noted on the right. (D) Reaction cycle of

euka~yotic CCT. ADP-CCT binds to a substrate and ADP is ejected (left3top). CCT-

substrate is loaded with ATP (top+right) and the transition state of hydrolysis closes the

lid (rigl~t+bottom). Folded substrates are released along with inorganic phosphate

restoring the ADP-CCT complex (bottom+left).

C Structural Domains Functional Features

Helcal protruston - -rorrns l ~ d of fold~ng charriber

Ap~cal dornam - -Substrate b~ndmg s~ te

lnterrnedmte domain - , T r a n s ~ ~ ~ ~ t s alloste~~c s~gnal from ATPnsc to substrate b~ndmg s~te L

Equator~al dornam - I I -ATP hydrolys~s rite

- Nalive substrate

Lid Closure

t13

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Figure 1-5 Cellular pathways of folding by CCT

(A) A ribbon n~odel of a WD-repeat protein likely to be a CCT substrate, (P-transducin,

PDB ID: ITBG; Sondek et a]., 1996). Modulators in the pathway of (A) actin and (B)

tubulin folding (see main text for description; Lopez-Fanarraga et al., 2001).

Phosducin-like protein

4 Native Actin

- -

Non-native actin- folding intermediate

CCT

Ccfactor E Cofactor B

Cofactor A

Cofaclor C - L>\ Microtubules

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Figure 1-6 Structural features of prefoldin, phosducin-like protein and their

complexes with CCT

(A) Backbone trace of archaeal PFD (PDB code: 1 FXK). Individual a or P subunits are

shown (left, top and bottom) to highlight the coiled-coils and P-hairpins. The backbone

trace of the complete hexanier is shown (centre and right) without amino acid sidechains

for clarity. (B) Atomic structure of phosducin removed from the coniplcx with GPy (PDB

code: 2TRC; Gaudet et a]., 1996). The N-terminal helical domain is coloured green and

the C-terminal tliioredoxin-like domain is coloured red.

A Front view

ti-subunit

Mew mto cavity tx , B

Phosducin

I-ielical domain

Thioredoxin-like domain

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(C) Three-dimensional electron n~icroscopy reconstructions of apo-CC'T (left), PFD-

CCT (centre), and PhLPl-CCT (right) (Martin-benito et a]., 2002; Martin-benito et al.,

2004). PFD and PhLP I are coloured red in the centre and right panels, respectively.

CCT-PFD

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1.8 Tables

Table 1 -1 List of known CCT substrates (Reproduced from Spiess et al., 2004)

Part of an WD Molecular oligomeric repeat

Protein Weight (kDa) complex a actin, p actin 42.1,41.7 Yes? a tubulin P tubulin 7 tubulin 50.2, 49.8, 51.2 Yes

Myosin heavy chain Luciferin 4- monooxygenase Ga-Transducin Von Hippel-Lindau disease tumour suppressor GI-S specific cyclin E l Cofilin Actin-depolymerizing factor 1 Actin-related protein V (Centractin) Hepatitis B virus capsid protein EBNA-3 nuclear protein Gag polyprotein of M- PMV Histone deacetylase 3 SET domain protein 3 Probably histone deacetylase HOS2 Cell division control protein 20 (Cdc20) Cell division control protein 15 (Cdhl) Protein phosphatase PP2A regulatory subunit B (Cdc55) Peroxisomal targeting signal 2 receptor (Pex7) Pre-mRNA splicing factor PRP46 Coatomer p' subunit (Sec27) Guanine-nucleotide binding protein P subunit (Ste4)

Yes

N 0

Yes

Yes Yes No

No

Yes

Yes No?

Yes Yes Yes

Yes

Yes

Yes

Yes

N 0

Yes

Yes

Yes

motif N 0

N 0

No

N 0

N 0

No N 0

No

No

N 0

N 0

N 0

No No No

No

Yes

Yes

Yes

Yes

Yes

Yes

Yes

References Gao et al., 1992 Yaffe et al., 1992; Melki et al., 1997 Srikakulam and Winkelmann, 1999

Frydman et al., 1994 Farr et al.. 1997

Feldman et al., 1999 Won et al., 1998 Melki et al., 1997

Melki et al., 1997

Melki et a1.,1993

Lingappa et al., 1994 Kashuba et al., 1999

Hong et al., 2001 Guenther et al., 2002 Pijnappel et at., 2001

Pijnappel et al., 2001 Camasses et al., 2003 Camasses et al., 2003

Siegers et al., 2003

Siegers et al., 2003

Siegers et al., 2003

Siegers et al., 2003

Siegers et al., 2003

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1.9 Reference list

Ali. M.M., Roe, S.M., Vaughan, C.K., Meyer, P., Panaretou, B., Piper, P.W., Prodromou, C., and Pearl, L.H. (2006). C~ystal stn~cture of an Hsp90-nucleotide-p23lSba1 closed chaperone complex. Nature 440, 10 13- 10 17.

Alvarez, P., Smith, A., Fleming, J., and Solon~on, F. (1998). Modulation of tubulin polypeptide ratios by the yeast protein Pac 1 Op. Genetics 149, 857-864.

Anfinsen, C.B. ( 1973). Principles that govern the folding of protein chains. Science 1 X I , 223-230.

Behrends, C., Langer, C.A., Boteva, R., Bottcher, U.M., Stemp, M.J., Schaffar, G., Rao, B.V., Giese, A., Kretzschmar, H., Siegers, K., and Hartl, F.U. (2006). Chaperonin TRiC promotes the assembly of polyQ expansion proteins into nontoxic oligomers. Mol Cell 23, 887-897.

Bitto, E., and McKay, D.B. (2002). Crystallographic structure of SurA, a molecular chaperone that facilitates folding of outer membrane porins. Structure lo1 1489- 1498.

Blaauw, M., Knol, J.C., Kortholt, A., Roelofs, J., Ruchira, Postma, M., Visser, A.J., van Haastert, P.J. (2003). Phosducin-like proteins in Dictyostelimn discvideuni: implications for the phosducin family of proteins. EMBO J 22, 5047-5057.

Bochtler, M., Hartmann, C., Song, H.K., Bourenkov, G.P., Bartunik, H.D., and Huber, R. (2000) The structures of HsIU and the ATP-dependent protease HsIU-HslV. Nature 403, 800-805.

Bouhouche, A., Benomar, A., Bouslam, N., Chkili, T., and Yahyaoui, M. (2006). Mutation in the epsilon subunit of the cytosolic chaperonin-containing t-complex peptide- 1 (CctS) gene causes autosomal recessive mutilating sensoly neuropatliy w-ith spastic paraplegia. J Med Genet 43,44 1-443.

Camasscs, A., Bogdanova, A., Shevchenko, A., and Zachariae, W. (2003). The CCT chaperonin promotes activation of the anaphase-promoting complex through the generation of functional Cdc20. Mol Cell 12, 87-1 00.

Daggett, V., and Fersht A.R. (2003). Is there a unifying mechanism for protein folding'? Trends Biochenl Sci 28, 18-25.

Dinner, A.R., Sali, A,, Smith, L.J., Dobson, C.M., and Karplus, M. (2000). Understanding protein folding via free-energy surfaces from theory and experiment. Trends Biochem Sci 25,33 1-339.

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Valpuesta, J.M., Martin-Benito, J., Gomcz-Puertas, P., Carrascosa, J.L., and Willison, K.R. (2002). Structure and function of a protein folding machine: the eukaryotic cytosolic chaperonin CCT. FEBS Lett 529, 1 1 - 16.

van Montfort, R.L.M., Basha, E., Friedrich, K.L., Slingsby, C., and Vierling, E. (2001). Crystal structure and assembly of a eukaryotic small heat shock protein. Nature Struct Biol 8. 1025-1 030.

Walton, T.A., and Sousa, M.C. (2004). Crystal structure of Skp, a prefoldin-like chaperone that protects soluble and membrane proteins from aggregation. Mol Cell 15. 367-374.

Webb, C.T., Gorman, M.A., Lazarou, M., Ryan, M.T., and Gulbis, J.M. (2006). Crystal structure of the rnitochondrial chaperone TIM9.10 reveals a six-bladed alpha- propeller. Mol Cell 2 1 , 123- 133.

Wells, C.A., Dingus, J., and Hildebrandt, J.D. (2006). Role of the chaperonin CCT/TRiC7 complex in G protein betagamma-dimer assembly. J Biol Chem 28 I , 2022 1-20232.

Whitesell, L., and Lindquist, S.L. (2005). HSP90 and the chaperoning of cancer. Nat Rev Cancer 5, 76 1-772.

Won, K.A., Schumacher, R.J., Farr, G.W., Honvich, A.L., and Reed, S.I. (1998). Maturation of human cyclin E requires the function of eukaryotic chaperonin CCT Mol Cell Biol 18, 7584-7589.

Xu, Z., Knafels, J.D., and Yoshino, K. (2000). Crystal structure of the bacterial protein export chaperone SecB. Nature Struct Biol 7, 1 172-1 177.

Yaffe, M.B., Fan-, G.W., Miklos, D., Horwich, A.L., Sternlicht, M.L., and Sternlicht, H. ( 1992). TCPI complex is a niolecular chaperonc in tubulin biogenesis. Nature 358, 245-248.

Zako, T., lizuka. R., Okochi, M., Nomura, T., Ueno, T., Tadakuma, H., Yohda, M., and Funatsu, T. (2005). Facilitated release of substrate protein from prefoldin by chaperonin. FEBS lett 579, 37 18-3724.

Zhu, X., Zhao, X., Burkholder, W.F., Gragerov, A,, Ogata, C.M., Gottesman, M.E., and Hendrickson, W.A. (1996). Structural analysis of substrate binding by the molecular chaperone DnaK. Science 272, 1606- 16 14.

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CHAPTER 2 MUTAGENESIS AND ELECTRON MICROSCOPY CHARACTERIZE ARCHAEAL PREFOLDIN AS A MOLECULAR CLAMP WITH HYDROPHOBIC COILED-COIL BINDING SITES

Note regarding contributions: The following chapter is a composite of two papers published in ThP PI-occcu'i~gs

oftkc. Nntiorld Accru'v~i~y qf'Scienct. USA (2004) and in Stnictzrrr (2007), respectively. The authors of these studies are listed below.

Article I Lundin, V. F.*, Stirling, P. C.", Gomez-Reino, J., Mwenifumbo, J. C., Obst, J. M., Valpuesta, J. M., and Leroux, M. R. (2004). Molecular clamp mechanism of substrate binding by hydrophobic coiled coil residues in the archaeal chaperone, prefoldin. Proc Natl Acad Sci USA 10 1,4367-4372.

t

V.F.L. and P.C.S. contributed equally to this work.

Article 2 Martin-Benito, J., Gomez-Reino, J., Stirling, P.C., Lundin, V.F., Gomcz-Puertas, P., Boskovic. J., Chacon, P., Fernandez, J.J., Berenguer, J., Leroux, M.R., and Valpuesta. J.M. (2007). Divergent Substrate-Binding Mechanisms Reveal an Evolutionary Specialization of Eukaryotic Prefoldin Compared to Its Archaeal Counterpart. Structure 1 5 , 1 0 1 - 1 1 0. Reprodrrced with perrtzissiorz. from Elsevier.

As a co-first author on article # I I generated the data for Table 2-1, Figures 2-1, 2-2,2-4 and 2-9A, C equally with V.F.L. V.F.L. generated Figure 2-5 and I generated Figures 2-3, 2-6 and 2-7A, D. The group of Jose M. Valpuesta (particularly J.G.R. and J.M.B.) generated the data for Figure 2-7B,C, 2-8, 2-9B and 2-10. I wrote approximately 113 of the PrVAS article along with V.F.L. and M.R.L. (up to 2.4.5 inclusive) and the group of Jose M. Valpuesta wrote nearly all of the Strtictui-c article (2.4.6, 2.4.7). To article #2 I contributed purified proteins, the data shown in Figure 2-9A, as well as preliminary data with the three substrates studied and intellectual contributions to developing the story.

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2.1 Abstract

Prefoldin (PFD) is a jellyfish-shaped molecular chaperone that has becn proposed

to play a general role in de novo protein folding in archaea and is known to assist the

biogcnesis of actins, tubulins, and potentially other proteins in eukaryotes. Using point

mutants, chimeras, and intradomain swap variants, we show that the six coiled-coil

tentacles of archaeal PFD act in concert to bind and stabilize non-native proteins near the

opening of the cavity they form. Importantly, the interaction between chaperone and

substrate depends on the mostly buried interhelical hydrophobic residues of the coiled

coils. We also show, by electron microscopy (EM), that the tentacles can undergo

inovement to acconmodate an unfolded substrate. By performing three-dimensional EM

reconstnictions of PFD in complex with three substrates of different sizes we confirm

that the chaperone moves to accommodate larger substrates and that larger substrates

contact more PFD subunits. Analysis of PFD truncations both functionally and by EM

show that one subunit type can compensate for loss of the other in the context of the

intact hexamer. Finally, a conlparison of eukaiyotic PFD and archaeal PFD binding to the

same substrate, actin. shows that the two chaperones employ different modes of binding.

This observation likely reflects the substrate specificity of euka~yotic PFD compared to

its more general archaeal counterpart.

2.2 Introduction

Coiled coils consist of two or more parallel or antiparallel aniphipathic a-helices

that twist around one another to form supercoils (Lupas, 1996). The primary sequences of

the helices display a heptad repeat (abcdefg), where apolar residues are found

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preferentially in the first (a) and fourth (d) positions. Although the knobs-into-holes

packing of the hydrophobic residues is the predominant stabilizing force for a coiled coil,

inter- and intrahelical ionic interactions can act to further stabilize or destabilize its

supersecondary stn~cture (Lupas, 1996). Coiled coils are found in several molecular

chaperones: a diverse family of proteins whose collective cellular role is to ensure the

quality control (e.g., folding, assembly, and transport) of non-native proteins (Hart1 and

Hayer-Hartl, 2002; Stirling et a]., 2003). Archaeal prefoldin (PFD) is a chaperone that

contains six canonical antiparallel coiled coils whose N- and C-terminal helices project

outward from a double P-barrel oligomerization domain; the overall shape of the

hexameric protein complex, assembled from two PFDa and four PFDP subunits (a2P4),

resembles a jellyfish with six tentacles (Siegert et a].. 2000). In solution, its tentacles are

likely to be fully solvated and independently mobile (Siegert et a]., 2000). A lower-

resolution electron microscope image of recombinant human PFD, which consists of six

different proteins ( t ~ l o a class and four P class subunits), rcvcals that i t posscsses the

same overall structure (Martin-benito et a]., 2002).

Like other chaperones, archaeal PFD can selectively interact with and stabilize

non-native (unfolded) polypeptides that exposc hydrophobic surfaces in v i t r~ ) , helping to

prevent their aggregation (Siegert et al., 2000; Leroux et a]., 1999; Okochi et a]., 2002).

Preliminary studies have shown that deletion of the distal coiled-coil regions in either the

a or p subunit abrogates chaperone activity it? vitr-o, implying that PFD grasps its

substrates in a multivalent manner (Sicgert et a]., 2000). Similarly, the eukaryotic PFD-

actin complex recently visualized by EM shows that non-native actin, one of its

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substrates, makes multiple contacts with the distal regions of the tentacles (Martin-benito

et a]., 2002).

In the crowded cellular environment, eukaryotic PFD 1s likely to transiently

stabilize ribosome-bound nascent polypeptides (Hanscn et a]., 1999) before shuttling

them to a chaperonin (an ATP-dependent cylindrical chaperone) for completion of

folding (Vainbcrg et al., 1998). Functional cooperation between PFD and eukaryotic

cytosolic Chaperonin Containing TCP-I (CCT) likely arises through a transient ternary

complex with substrate that accelerates folding and prevents aggregation (Martin-benito

et al., 2002; Vainberg et al., 1998; Geissler et a]., 1998; Siegers et a]., 1999). The range of

substrates bound by eukaryotic PFD overlaps at least in part with CCT. because it

includes actins and tubulins (Vainberg et a]., 1998; Geissler et a]., 1998; Siegers et a].,

1999; Leroux and Hartl, 2000; Rommelacrc et al., 2001).

Although the in vivo substrates of archaeal PFD are not known, its ability to

stabilize a wide array of unfolded proteins in vitro (e.g., rhodanesc, actin, lysozyme,

firefly luciferase, and GFP) suggests that i t performs a general role in recognizing and

assisting the biogenesis (folding) of non-nativc proteins in the archaeal cytosol (Lerous et

al., 1999; Leroux, 2000; Okochi et a]., 2002). Archaeal PFD seems to function as an

ATP-independent holdase for non-native protans before passing on the substrate to an

ATP-dependent chaperonin, much like its eukaryotic counterpart (Stirling et a]., 2003;

Siegert et a]., 2000; Leroux et a]., 1999; Okochi et a]., 2002; Okochi et a]., 2004;

Vainberg ct a]., 1998; Zako et a]., 2005). How PFD binds and stabilizes non-native

proteins at the molecular level therefore represents a fundamental question that needs to

be explored.

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In this study, we characterize the function of archaeal PFD complexes using a

panel of a and 13 subunit variants assen~blcd in different con~binations and use EM to

visualize the interaction between the chapcrone and several non-native protein. Our data

show that archaeal PFD utilizes, in a concerted manner, partially buried hydrophobic

residues in the tips of flexible coiled coils to interact with and prevent the aggregation of

its non-native substrate. Three dimensional reconstructions of PFD in complex with

substrates of different size and shape, including actin, confirm our biochemical data and a

comparison with eukaryotic PFD-actin complexes suggests a fascinating evolutionary

specialization of the two chaperones.

2.3 Methods

2.3.1 Preparation of constructs

PCR-based mutagenesis was used to create point mutant, intradomain swapped.

and chimeric prefoldin constructs of both the a and subunits. For mutations near the N

and C termini, one pair of mutagenic primers was used to amplify the product. For

mutations further from the termini, two or three sets of nested mutagenic primers were

used in successive PC'R reactions. PCR products were subcloned into pRSET6a at NdeI

and BamHl sites, and the constructs were verified by DNA sequencing (see Figure 2-1

for the amino acid sequences of constructs from Lundin et al, 2004). P. koi.iko.shii PFD

truncations corrcspond to amino acids 13- 104 for PIIPFDP~' and 15- 130 for P ~ P F D ~ ~ '

Figure 2-9.

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2.3.2 Protein expression and purification

Wild-type or mutant Py~.oc.oc~c.r/.s hoi-ikoshii or ~ ~ ~ ~ / / ~ L I ~ ~ ~ / / I ~ ~ I ' I ~ I o / ~ c I L ' / ~ I -

//7rr.1tloul//otl*op/lic~11~s PFD subunits were produced in Esc~l7c~ric~l~iu c d i strain

BL2 l(DE3)pLysS. purified, and assembled into a2P4 complexes as described (Siegert et

a1,2000). Purified protein complexes (stored frozen in 25% glycerol) were dialyzed

against buffer A (20 mM sodium phosphate, pH 8.011 00 mM NaCI) at 4 O C ' and

concentrated with Centriprep Y M- I 0 or Ultra- I 5 centrifugal filter units (Millipore)

before analysis. Protein concentrations were determined by quantitative amino acid

analyses (Alberta Peptide Institute, Edmonton, AB) and Bradford protein assays (Bio-

Rad). The observed molecular weights o f recombinant PFD subunits, determined by

matrix-assisted laser desorption ionization time-of-tlight mass spectrometry, were as

predicted; in the case MtPCePFD6, the initiating methionine was absent. Recombinant

polyhistidine-tagged GFP (F99SIM 153TlV 163A) was expressed and purified as

described (Sakikawa et al., 1999).

2.3.3 Characterization of PFD variants

All PFD complexes were characterized by analytical size-exclusion

chromatography (SEC) and circular dichroism (CD). For SEC, samples were run on a

Superdex S200HR PC 3.2130 column (Amersham Pharmacia) equilibrated in buffer A.

Far-UV C'D spectra were recorded on a Jasco (Easton, MD) 7 10 spectropolarimeter by

using I0 accumulations from 260 to 190 nm at room temperature. Protein samples were

diluted to 0.4 mglml in buffer A. Thermal denaturation experiments were performed

essentially as described (Fandrich et al., 2000). Stability of PFD variants was examined

by monitoring the CD ellipticity at 222 nm as a function of temperature with a heating

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rate of 1.3"C/niin in buffer A and a path length of 20 mm. Melting temperatures (Tm)

reported in Table 2-1 correspond to the dissociation of the hexamer containing the

indicated PFD variant in a complex with the other wild-type subunit (Fhdrich et a].,

2000).

2.3.4 Prevention of protein aggregation assays

In vitr-o chaperone activity of PFD variants was determined essentially as

described (Leroux et a]., 1999). Briefly, hen egg-white lysozyme (Sigma) was dissolved

in denaturing buffer (6 M guanidine-HCIII 00 1nM NaC1120 mM sodium phosphate, pH

8.0150 mM DTT) and then diluted 1 0 0 ~ to a final concentration of 2 pM into buffer A

alone or containing various concentrations of either wild-type or mutant PFD complexes.

Aggregation of substrate was monitored spectrophotonietrically at 360 nm (which detects

light scattering by the aggregates) for 10 min at 2S•‹C. Raw absorbance data were

normalized, and relative aggregation was defined as the fraction of the final absorbance

value observed in the buffer A alone control. Conalbumin aggregation assays were

performed as above, except the protein was diluted to a final concentration of 0.75 p M .

Each sample was run at least in duplicate for a given experiment, and each experiment

was repeated at least twice on separate occasions. The data presented are representative

trials.

2.3.5 Formation and analysis of PFD-substrate complexes

Native GFP in 20 mM TrisC1/100 mM NaC'III mM DTT, pH 8.0 (buffer B) was

acid-denatured by adding HC'I to a final concentration of 12.5 mM. For SEC analysis,

1 16 p M denatured GFP was diluted into buffer A or buffer A containing I I pM prefoldin

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to a final concentration of I 1 pM and mixed rapidly. The mixture was incubated on ice

for 1 hour, centrifuged for 5 min at 16,100 x g, and 50 p1 was analyzed by SEC as

described above. Peptide backbone absorbance was monitored at 222 nm and GFP

excitation at 396 nm. Fractions analyzed by SDSIPAGE were PFD Peak A, 1.25- to 1.5-

1111 elution volume, and GFP Peak B, 1.60- to 1 .S5-ml elution volurnc. Before EM, PFD

was dialyzed against buffer B and mixed with acid-denatured GFP as above without

column purification. For the formation of actin-, lysozyme- and conalbumin-PFD

complexes the substrates were diluted fifty fold from 6M Guanidine-HCl buffer into a

solution containing P~~r.oc~occ*r/.r hoi-koshii PFD (PhPFD) at a I : I final ratio (5pM each).

2.3.6 Electron microscopy

Aliquots of substrate-free PFD or PFD-GFP con~plexes were applied to carbon

grids and negatively stained with 2% uranyl acetate. Images were taken under low-dose

conditions at a ~ 6 0 , 0 0 0 nominal magnification in a JEOL I200EX-I1 electron microscope

operated at I00 kV and recorded on Kodak SO- 163 film. For image processing, 1,265

substrate-free and 1,926 GFP-bound PFD particles displaying U-shaped side views were

selected from independent samples.

For Figure 2-7B U-shaped views were selected among other views (i.e., W-

shaped views) essentially as described (Martin-benito et al., 2002). The presence or

absence of GFP in the sample did not affect the distribution of the two PFD views.

Particles were centered and aligned by using a fi-ee-pattern algorithm (Penczek et al.,

1992) and subsequently subjected to a neural network classification procedure (Marabini

and Carazo, 1994). This procedure served to discriminate, when analyzing the putative

PFD!GFP complexes, between particles containing a stain-excluding region between the

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tentacles (i.e., GFP-bound particles; 20% of the population) and those possessing a stain-

penetrating rcgion (i.c., the substrate-free particles; 80% of the population). The

hon~ogeneous populations from the two independent samplcs were subsequently

processed and averaged.

2.3.7 Image processing and three-dimensional reconstruction

Micrographs were digitized in a Zeiss SC'AI scanner with a sampling window

corresponding to 2.8 AIpixeI. The three-dimensional reconstruction of PhPFD was

generatcd from negatively-stained, randomly oriented particles, using the EMAN

package for single-particle three-dimensional reconstruction (Ludtke et al., 1999). The

initial volume was generated by the common line procedure included in the EMAN

package, using the average classes obtained after multivariate statistical analysis. A 2-

fold symmetrization was imposed on the volumes generated throughout the iterative

process. The final resolution was estimated to be 19 with the 0.5 criterion for the

Fourier shell correlation coefficient between two independent reconstructions. For the

three-dimensional reconstri~ctions of the PhPFD:lysozyme, ~ h ~ ~ ~ ~ " : l ~ s o z ~ m e ,

PhPFD:GFP, PhPFD:conalbumin and PhPFD:actin complexes, the corresponding

particles were subjected to thc reconstruction procedure described above, except that the

volume of the unconiplexed PhPFD was ~ ~ s c d as the reference volume and that no

symmetry was imposed throughout the reconstruction process. The final resolution for

the PhPFD:lysozyme, ~ h ~ ~ ~ ~ ~ ' : l ~ s o z ~ r n e , PhPFDGFP, PhPFD:conalbumin and

PhPFD:actin complexes was 20, 20, 21. 22 and 19 A, respectively. Visualization of the

volumes was carried out using AMIRA (http://anlira.zeb.de).

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2.3.8 Miscellaneous

Molar aniounts refer to licxamers for PFD co~npleses (84 kDa) and monomers for

lysozyine ( I4 kDa), co~ialbumin (75 kDa), and GFP (27 kDa). Multiplc sequence

alignments were performed by using clustalx (ftp://ftp-igbn1c.u-strasbgti-/p~ib/clustalx)

software followcd by manual editing. Silnilarity scores were assigned by using a Gomet

PAM250 similarity matrix. Molecular graphics of the PFD crystal structure (1 FXK) were

prepared using Pyniol (Delano Scientific).

2.4 Results and discussion

2.4.1 Properties of PFD coiled coils

In an attempt to understand the attributes of the coiled coils that confer the ability

of PFD to interact with and stabilize non-native proteins, we evaluated the amino acid

sequence conservation of the coiled coils in relation to the 3D structure of the chaperone.

We constructed multiple sequence alignments of 1 I PFD a and [3 subunits from different

genera and assigned a score for the degree of conservation at each residue (Figure 2-2).

We noted, based on the crystal structure, that the N-terminal helices of the archaeal a and

p subunit coiled coils face into the rectangular cavity, ostensibly forming the binding

surface for non-native proteins; in contrast, C-terniinal helices localize mainly to the

outside surface of the cavity (Figure 2-3). It was thus surprising to find that the primary

sequences of the N-terminal helices were not more conserved on the whole than those of

the C-terminal helices if they indeed contained the substrate-binding site (Figure 2-2).

More importantly, when we mapped the amino acid conservation onto the structure of

PFD, there were few highly conserved surface-exposed residues that were preferentially

found inside the cavity (Figure 2-3). A large proportion of the conserved residues are

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within the partially buried hydrophobic core (a/d residues) of the coiled coils. Together,

these observations led us to hypothesize that the basis of action of PFD likely depcnds on

the unique spatial arrangement, and intrinsic properties of the coiled coils rather than on

the presence of conserved patches of substrate binding surface-exposed residues.

2.4.2 Cavity surface formed by the coiled coils

To test for the potential presence of an essential solvcnt-exposed binding site on

the cavity surface of PFD, we designed mutants in which the C-terminal helix coniprises

the cavity surface and the N-terminal helix faces the external solvent. To this end, the

amino acid sequences of the distal N- and C-terminal coiled-coil helices were switched

relative to the wild-type subunits (Figures 2-1 and 2-4A). We chose a crossover point

near the middle of the coiled coils, because removal of protein sequences beyond this

point (i.e., the distal region) negates substrate binding by PFD (Siegert et al., 2000).

Therefore, the proximal-to-distal sequence of side chains in both of the switched N- and

C-terminal helices is identical to that of the wild-type subunits. Although the backbone

polarity within the swapped helical regions is reversed relative to the wild-type sequence,

the overall structural properties of the switched coiled coils appear to be essentially

unchanged (sec below).

The recombinant a and P switch (SW) mutant subunits (aS'" and psi') were

assembled with each other or with wild-type subunits (a or P). To assess the structural

integrity of these complexes, we performed analytical SEC', far-UV CD. and measured

thernial stability by CD, as described (Data not shown; Fandrich et al., 2000 Siegert et

al., 2000; Table 2-1). SEC indicated that the mutant and wild-type complexes had an

identical Stoke's radius. implying tlie same overall shape and coiled-coil length. The far-

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UV CD spectra revealed that the secondary structure content of the mutant complexes

was indistinguishable from wild type, confirming thcrc were no gross structural dcfects.

We also monitored thermal stability by CD at 222 nm and found that the complexes

possess melting temperatures (Tm) comparable to their wild-type counterpart, which

denatures at 61•‹C (Table 2-1). Therefore, the switch mutations alter the nature of the

cavity surface without affecting the amino acid composition or significantly altering thc

structure or stability of the PFD hexamer.

Each PFD complex containing PFDa and -P variants was tested for chaperone

activity in a standard prevention-of-aggregation assay by using lysozynre as a model

substrate (Siegert et a]., 2000; Leroux et al., 1 999). Importantly, the aggregation of

lysozyme is not affected by the presence of irrelevant proteins, includmg aldolase and

native lysozyn~e, even at elevated concentrations; moreover, inactive PFD variants have

no effect on the aggregation of denatured proteins (Siegert et a]., 2000). Last, the assay is

performed at a temperature (25•‹C) well below the melting points of the as"- and pF\\'-

containing complexes (59•‹C and 53"C, respectively; Table 2-1).

A PFD complex assembled from a switch and wild-type P subunits (aS"P)

prevented the aggregation of denatured lysozyme as efficiently as wild-type PFD (Figure

2-4B). The P switch mutant, when combined with wild-type a subunit (a~"'), displayed

reduced yet significant activity at the same 1 : 1 molar ratio over denatured lysozynie and

almost full activity at a 4: 1 ratio ovcr substrate (Figure 2-4B). Finally, a complex

',\\' S\V containing both a and P switch mutants (a P ), in which all six coiled coils are

switchcd, still had detectable chaperone activity at a 2: 1 molar ratio over substrate and

significant activity at a 4: 1 ratio. This concentration-dependent prevention of aggregation

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activity shows that the switch mutations do not abolish all PFD activity, although their

activities are significantly reduced (Figure 2-4B).

Because of the a& stoichiornetry of the PFD hexanier, the as'\'13, aps\\', and

us\\Yp'\\ variants represent complexes i n which increasing amounts of the cavity surface

(i.e., two, four, or six coiled-coil tentacles, respectively) are affected, and this coincides

with a gradual decrease in activity (Figure 2-4B). The additional loss in activity of

aS\4'p"4\elativc to upS\'; appears to reveal a defect in thc a"" variant and a contribution

from both subunit types toward chaperone act~vity. I t is therefore possible that some of

the residues that normally face into the cavity do in fact contribute to substrate binding.

These residues do not appear to be essential for interaction with a non-native protein to

occur, although their absence may reduce the substrate-binding affinity of the chaperone.

These data represent a previously undescribed and interesting finding that is consistent

with our observation that the interior N-terminal helix of PFD is not detectably more

conserved than the outward-facing C-terminal helix (Figure 2-2).

2.4.3 Intrinsic properties of the coiled-coil motif

Given both the relative paucity of conserved solvent-exposed residues in the

putative substrate-binding site and the fact that the switch mutations rctain partial

chaperone activity, we hypothesized that PFD function may depend to a large degree on

intrinsic properties of the coiled-coil motif. We therefore predicted that heterologous

coiled-coil sequences should at least partially support the chaperone activity of the

complex.

To test our hypothesis, we engineered PFD chimeras in which coiled-coil regions

of the myosin I1 heavy chain, Rad5O zinc hook domain, and Ccrcnorli~~hc/iti. r l epns

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PFD6 (one of the four eukaryotic 13 class PFD subunits) were fused to shortened coiled

coi l s of the PFDP ( P ~ ~ Y ~ ~ N ~ ~ , 3 H ; ~ 1 S 0 , and P ~ ' " ' ~ ' ~ ( ~ ), and a similar rcgion of Rad5O was

fused to the PFDa subunit (Figure 2-5A). The heterologous sequences have very

low sequence identity (9-1 I %) and similarity (35-36%) to the corresponding archaeal

sequences and were designed to form a coiled coil with the same number and register of

heptad repeats as that of wild-type PFD. As with the switch mutant complexes, the

chimeric coniplexes were found to be indistinguishable from wild-type PFD hexamers by

SEC and far-UV CDI eliminating the possibility of major structural perturbations.

Thermal denaturation of the chimeras revealed similar stabilities to wild-type PFD, with

the exception of the a p c'cPI:Do chimera, which melted at a lower temperature (Tm = 43•‹C').

This behavior is consistent with the low optimal growth temperature of C. c.kCgm~s ( 1 5-

20•‹C) but is still significantly above the assay temperature used (Table 2-1).

When tested for chaperone activity, the aRadSO~ chimeric complex displayed near-

wild-type activity (Figure2-SB). PFD complexes of the wild-type a subunit with

chimeric p subunits ( a p C'cPI:DO a p M y s i ~ ~ , and apl'"d"') were also able to bind and stabilize

denatured lysozyme (Figure2-SB), although these variants had a range of intermediate

chaperone activities relative to wild-type. As with the PFD switch mutants, that a PFD

conlplex in which only the a subunit is chimeric has more activity than P subunit

k1d50. chimeras (i.e., aH'"""p VS. a p , Figure2-SB) is consistent with the presence of four

potential binding sites for the P subunits and only two for the a subunits. Therefore, even

though both subunit types niay be similarly compromised, the defect is expected to be

more apparent in the P chimeras. Importantly, the activities of the chimeras are greater

than that observed when the corresponding subunit is truncated, even though the

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truncations remove less of the coiled coil than is replaced in the chimeras (Siegert et al.,

2000). This partial rescue of activity shows that thc chimeric coilcd-coil regions, which

are devoid of archaeal PFD sequence, can contribute to the chapcrone activity of the PFD

complex.

f~n t150 Remarkably, the a and pR'"'50 chimeric subunits, which arc partially activc in

a wild-type background, had no measurable chaperone activity when assembled together

(Figure2-SB). Thus, either wild-type subunit confcrs a significant level of activity in the

context of a mutation in the other subunit, whereas the same mutations in all six subunits

seem to eliminate function. This cooperation between a and P subunits is an important

concept for PFD function and reflects its ability to bind substrates multivalently (Siegert

et al., 2000; Simons et al., 2004). Altogether, these results are consistent with the notion

that prcfoldin coiled coils have evolved specific features particular to their chaperone

function, which do not occur to the same extent in any heterologous coiled-coil sequence.

C'cl'l I)h In this respect, i t is notable that up . which contains a eukaryotic P class subunit,

MYONII was more efficient at stabilizing denatured lysozyme than the other chimeras (up . or

upRadS0) (Figure2-SB), despite the lack of consenred sequence (Figure 2-1) or amino

acid composition. This suggests that certain unique properties of the coiled coils are

likely shared by eukaryotic and archaeal PFD subunits. Indeed, archaeal PFDP can

partially complement the cytoskeletal phenotype caused by the lack of the

S t r c . c * J l c r i - o v cc~rc~visitrc p class (giml or gitr14) genes (Leroux ct al., 1999).

The loss of chaperone activity in the chimeric complexes could in principle be

accounted for by the absence of specific solvent-exposed residues that normally

contribute to substrate binding in wild-type PFD. Alternatively, or in addition, the

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chimeras could be deficient in subtle properties that affect the accessibility of their

interhelical hydrophobic residues. The second possibility led us to hypothesize that the

common ald hydrophobic residues may, in large part, form the intrinsic property of coiled

coils involved in substrate recognition and binding by PFD.

2.4.4 Hydrophobic interface of the coiled coils

Archaeal PFD, like some well-characterized nlolecular chaperones that bind non-

native proteins promiscuously (for example, Hsp7O and the bacterial chaperonin GroEL;

H a d and Hayer-Hartl, 2002; Bukau and Ho~wich, 1998) likely recognizes its substrates

mainly because they expose normally buried hydrophobic surfaces (Leroux et al., 1999).

Unlike other chnperones, however, the surface displayed within the cavity of PFD, where

substrate binding is expected to occur, is almost entirely devoid of distinctly solvent-

exposed hydrophobic residues (Siegert et al., 2000). The apolar interhelical interface of

the coiled coils may therefore be directly responsible for interactions with substrates.

To test this hypothesis, we engineered scrine-substituted PFD variants at one to

four pairs of the predominantly hydrophobic ald heptad repeat residues in the distal ends

of the u and p subunit coiled coils. Serine was chosen because of its relatively sn~all size,

polar character, and tolerance for inclusion into a-helices (Lawrence and Johnson, 2002;

Myszka and Chaiken, 1994). The intended effect of the sequential substitutions is to

gradually remove the potential hydrophobic-binding site while retaining intact helices

and thc overall rod-like struchlre of the tentacle. PFDu subunits mutated in up to three

a/d pairs (aH"', aHM2, and atmu), and p subunits mutated in up to four aid pairs (P"~" ,

pHM2, PHM3, and pHM4 ) were constructed (Figure 2-6A; HM, hydrophobic mutant). The

overall stiuctures and stabilities of these mutants were essentially identical to wild-type

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PFD, as judged by SEC, far-UV CD, and thennal denaturation studies (Table 2-1 and

data not shown).

PFD complexes containing a wild-type subunit and any hydrophobic point mutant

wcre found to have near-wild-type activities at a 1 : 1 ratio of chaperone hesanier to

denatured lysozyme (Figure 2-6B, Left). To test for functional cooperation between the

a and p subunits, we assayed the activity of a complex containing three substituted pairs

Hh13 H M J in the a subunit and four substitutions in the p subunit ( a p ). Remarkably, this

PFD variant was unable to prevent the aggregation of denatured lysozyme (Figure 2-6B,

Right) or conalbumin (Figure 2-7D). We verified the structural integrity of this inactive

complex as described above and also examined i t by EM (Dr. .lose M. Valpuesta data not

shown). The latter analysis showed that the ultrastructural features of the mutant were

indistinguishable from wild-type archaeal (e.g., see Figure 2-7B) and eukaryotic PFD

(Martin-benito et al., 2002).

Compared to the inactive c ~ ~ ~ " ' l ~ ~ ~ " % o m ~ l e x , the presence of two additional pairs

of aid residues in the a subunit (aH"IpHM4) partially restored the activity of the con~plex

(Figure 2-6B Right). As might be expected, con~plexes assembled from less severely

HM2 H M 3 mutated variants, lacking 3 a/d pairs in PFDP, and two or three pairs in PFDa (a p a H M 3 H M 3 p - ) showed intermediate activities compared to wild-type PFD (Figure 2-6B

Right). PFD complexes with the same PFDa backgrounds as above but lacking only two

HMZ HMZ aid pairs in PFDB (a and aH"3p'1ML) showed nearly full activity (Figure 2-6B

Right).

Altogether, these results demonstrate three important properties of the coiled coils

in archaeal PFD. First, the partially buried hydrophobic interface between the

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amphipathic helices is required for effective interaction and stabilization of a non-native

substrate by the chaperone. We also observed this effect in the partially activc P 11:1d 5 0

chimera; replacing the first four pairs of aid residues with serine (p R,1d~OIliL13 ) impaired its

ability to function in the complex (Data not shown). I t is notable that the residues

comprising the binding site are only partially exposed and are at the apex of the coiled-

coil tentacles (Figure 2-6C), where increased flexibility (indicated by higher B factors in

the tip regions; see Protein Data Bank ID I FXK) may facilitate the cxposure of

interhelical apolar residues. Indeed, such increased exposure may result from the partial

unwinding of the coiled coils, as suggested by Siegert et al. (2000). Second, the binding

site appears to be diffuse because therc is progressive loss of function as more apolar

residues are substituted with serine; the binding site could therefore cxtend somewhat

beyond the residues mutated in both the a and fi subunits. Last, the coiled coils act in a

concerted or multivalent fashion to stabilize a non-native protein, because alterations in

either subunit alone have much less profound effects on chaperone activity than

mutations in both subunits. An alternate intrcpretation of our results would bc that the

HM mutations disrupted the structure of the tips enough to re-orient the true substrate in

such a way as to inactivate it. We favour the notion that the interhelical hydrophobic

residues themselves become more exposed to substrate upon binding. However, the role

of electrostatic or backbone contacts cannot be wholly discounted. I t remains possible

that some fully exposed surface residues in this region assist in substrate binding.

2.4.5 PFD functions as a molecular clamp

Our lnutational analyses of PFD suggest that hydrophobic residues at the core of

each coiled coil conlprise the niajor substrate-binding surface. To analyze the manner in

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which PFD interacts with a substrate, we generated a stable PFD-substrate complex and

visualized i t by EM.

By mixing acid-denatured GFP (27 kDa) with PFD from P. /?or-ikoshii (PhPFD),

we could observe a complex by SEC (Figure 2-7A). Because PhPFD binds GFP with a

somewhat higher aftinity than I~ / h e ~ r n o ~ ~ ~ ~ / o t r ~ o p / ~ i ~ ~ z ~ . s PFD (all other biochemical assays

were performed on MtPFD), we used PhPFD for subsequent EM studies. EM images

were obtained by negative staining of two independent samples, namely substrate-frec

PhPFD and PhPFD-GFP con~plexes. For each sample, U-shaped views were selected

among other views ( r .g W-shaped views) as described (Martin-benito et a]., 2002)

(Figure 2-7A-2 and A-4). This view permitted the unambiguous identification,

processing, and averaging of substrate-free PFD and, after classitication of the PFDIGFP

complexes according to the absence or presence of stain in the intertentacle area, of the

GFP-bound prefoldin (20% of the total population).

The processed EM image of substrate-free PFD (Figure 2-7B-2) appcars identical

in overall structure and geometry to the molecular surface of the PFD crystal structure

(Figure 2-7B-1). When comparing substrate-free PFD to substrate-bound PFD, a stain-

excluding region representing the bound GFP molecule at the distal tips of the coiled

coils is immediately apparent (Figure 2-7B-4). The location of the GFP confirms the

distal coiled-coil regions as the substrate-binding site and explains the sensitivity of the

chaperone to mutagenesis in this region.

In addition, the EM images show that the PFD tentacles have flexed outward to

accommodate the non-native GFP (Figure 2-7B-3 and B-4). An overlay c o n t o ~ ~ r map

shows that compared to the unbound state, the observed expansion of the cavity

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corresponds to an outward motion of - 12@ for the tentacles (Figure 2-7B-3). This

confornlational change appears to represent an m bloc movement of the coiled coil. We

suggest that a hinge rcgion, consisting of the loops connecting the coiled coils to the P-

barrel oligomerization domain, could be responsible for the observed flexibility and

outward nloven~ent of the a and p supercoils (Figure 2-7C). Interestingly, the recent EM

image reconstruction of actin within the cavity of eukaryotic PFD showed no apparent

movemcnt of the coiled coils to accommodate this larger (45-kDa) non-nativc protein

(Martin-benito et al., 2002). In this reconstn~ction, actin appears to be in a nonglobular

conformation that spans the entire opening of the cavity. Together, these observations

suggest that the PFD tentacles can move independently to create the cavity shape needed

for efficient interaction with substrates of different conformations andior sizes.

Archaeal PFD is known to interact with proteins as small as 14 kDa (lysozyme)

and as large as 62 kDa (firetly luciferase) (Siegert et al., 2000; Leroux et al., 1999).

Figure 2-7D shows that both PhPFD and MtPFD are able to completely prevent the

aggregation of the 75-kDa protein conalbumin at a 5-fold molar exccss over thc substrate.

By comparison, MtPFD lacking its distal aid residues is essentially inactive at the same

concentration (Figure 2-7D). This finding extends the upper size limit of proteins known

to interact with PFD. Furthermore, the EM results, which show that PFD tentacles are

tlexible, may explain how the chaperone can interact with proteins of such diverse sizes.

The wide range of substrate sizes bound by archaeal PFD is consistent with an in

vivo role in binding and stabilizing a large repertoire of nascent proteins. Indeed, during

synthesis, many polypeptides must be stabilized cotranslationally before spontaneous, or

chaperone-assisted, folding (Hart1 and Hayer-Hartl, 2002). In bacteria, the chaperones

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DnaK (an Hsp70 honiolog) and trigger factor cooperate to perform this general

stabilizing function (Teter et a]., 1999; Deucrling et al., 1999). I t has been suggested that

prefoldin could functionally replace these chaperones in archaea, where trigger factor,

and often Hsp70, arc conspicuously absent (Leroux et a]., 1999; Leroux, 2000). If this is

the case, i t is not surprising that PFD displays a general ability to recognize non-native

proteins, and that those proteins can vary greatly in size and shape, as thcy would in vivo.

2.4.6 The interaction of PhPFD with unfolded proteins

The bioche~nical studies performed with several archaeal PFDs suggest for these

chaperones a promiscuous role in the protection and delive~y of unfolded proteins to their

corresponding therniosomcs (Leroux et a]., 1999; Okochi ct a]., 2002). Unlikc eukaryotic

PFD, which has only been shown to interact directly with non-native actin and tubulin.

archaeal PFDs appear to bind denatured proteins indiscriminately. We therefore sought to

visualize directly how PhPFD could interact with substrates of different size and

structure. We chose as substrates denatured fonns of the mostly cx-helical lysozynic (14

kDa), the medium-size green fluorescent protein (GFP; 27 kDa) that is composed mostly

of P-strands, and the large, alp protein conalbun~in (75 kDa). The three proteins wcre

chemically denatured and independently incubated with PhPFD. Aliquots of the

complexes formed were subsequently stained with 2% uranyl acetate and particles were

sclected and used for the three-dimensional reconstruction of the PhPFD:lysozynie,

PhPFD:GFP and PhPFD:conalbumin complexes (3 158, 324 1 and 3 173 particles,

respectively). In all three cases (Figure 2-8), the volumcs generated reveal the typical

structure of PFD obtained so far, a structure with six tentacles hanging from a rectangular

base. However, unlikc the three-dimensional reconstruction of the apo-PhPFD (Figure 2-

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HA), the volunies of tlie PhPFD:lysozyme (Figure 2-8B), PhPFD:GFP (Figure 2-8C) and

PhPFD:conalbuniin (Figure 2-8D) complexes rcveal a stain-excluding mass interacting

with the tip of some of the PliPFD tentacles. The masses of each unfolded protein

protrude from the PhPFD cavity and their sizes are consistent with that of their

corresponding native structures (see atomic structures in Figure 2-8). The volunles

reconstructed also reveal that the number of PhPFD subunits involved in the interaction

with the unfolded substrates increases with the size of the denatured protein (see the

bottoni views for each of the three-din~ensional reconstructions). Accordingly, lysozyme

interacts with a pair of PhPFDP subunits (Figure 2-8B), GFP binds to a pair of PhPFDP

subunits plus one of the PliPFDa subunits (Figure 2-8C) and the largest protein,

conalbumin, interacts with all six PhPFD subunits (Figure 2-8D). The arrangement of the

tentacles in the PhPFD:substrate complexes (Figure 2-8) seems to deviate from the

position of the apo-PhPFD tentacles (Figure 2-8A), which suggests a flexing of tlie

coiled coils to accommodate tlie interaction with substrates of different size and shape

consistent with ous previous obseniations (Figure 2-7).

To confirm biochemically these structural results we generated PhPFD mutants

with truncations of the N- and C- termini for both PhPFDa and PhPFDP subunits, which

correspond topologically to the tips of the chaperone tentacles, and subsequently tested

their intcraction with different substrates. For GFP, it was previously shown by truncation

analysis that the PhPFDP subunits are important for substrate binding activity, whereas

the PhPFDa subunits are less critical (Okochi et al., 2004), which agrees with the

stn~ctural data shown hcrc. To further dissect the relative contribution of each tentacle to

binding different proteins, we used lysozynie and conalbumin as substrates. The two

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proteins were chemically denatured and their aggregation upon dilution into non-

denaturing buffer was assayed in the absence or presence of eithcr wild-type PhPFD, a

dcletion mutant with a truncation in the PhPFDa subunits ( P ~ P F D ~ ~ ' ~ ) , another mutant

with the same type of truncation in the PliPFDP subunits ( P I I P F D ~ P ~ ' ) or a mutant with

Tr Tr truncations in both subunits (PhPFDa P ) (Figure 2-9A and C). The results obtained

show that in the casc of lysozymc. renioval of the PhPFDu tips results in a small decrease

in the prevention of aggregation, as compared to wild-type PIiPFD (Figure 2-9A),

consistent with our demonstration that only PhPFDP subunits are involved in the

interaction with lysozyme (Figure 2-86). Unexpectedly however, the activity of the

chaperone with truncated PhPFDP subunits is not completely abolished (Figure 2-9A).

This apparent paradox could be explained if the PhPFDa subunits substitute for the

PhPFDP ones in the stabilization of the unfolded protein, once the tips of thc latter are

removed. Indeed, this is what happens, as revealed by a three-dimensional reconstruction

of the coniplex formed between P ~ P F D P ' ' and unfolded lysozyme (2729 particles

analyzed; Figure 2-9B). Strikingly, the volunie reconstructed shows the unfolded

lysozyme interacting with the pair of central PhPFDa subunits conipared with the

PhPFDP subunit in the wild-type complex (Figure 2-86). At this stage, it is unclear

whether the different shapes of the unfolded lysozyme bound to PhPFD (Figure 2-86) or

P ~ P F D P ~ ' (Figure 2-9B) stems from the relatively low resolving power of the three-

dimensional recolistructions or reflects the stabilization of an alternate form of unfolded

lysozyme between the two types of PhPFD subunits.

When the same prevention-of-aggregation experiments were performed with

denatured co~ialbun~in (Figure 2-9C), we obscrved that removal of the PhPFDu tips only

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slightly reduces the ability of the chaperone to prevent aggregation, and truncation of the

PhPFDp tips resulted in only a further small increase in the aggregation of the non-native

protein. Only the tnlncation of both PhPFDa and PhPFDP tips abolish PhPFD protection

of conalbumin aggregation (Figure 2-9C). These data clearly indicates that all six PhPFD

subunits are used in the stabilization of unfolded conalbumin, a finding consistent with

mutagenesis data showing that archaeal PFDa and PFDP coiled coil tentacles act

synergistically to stabilize non-native proteins (Figure 2-4. 2-5,2-6 and 2-7). In addition,

the results confirm our observation that a11 PFD subunit tentacles are engaged in the

P11PFD:conalbumin complex observed by electron microscopy (Figure 2-8D).

Archaeal PFDs have been shown to interact with a wide range of substrates,

protecting them from unwanted interactions and delivering them into the chaperonin

cavity (Lcroux ct aI., 1999; Zako et al., 2005). The results described here confilm the

pronliscuity of this chaperone in the interaction with unfolded proteins, since a stable

interaction takes place between PhPFD and three proteins of different size and secondary

structure: lysozyme, a small protein (14 kDa) of mostly a-helical nature; GFP, a protein

of medium size (27 kDa) that forms a j3-barrel in its native conformation, and

conalbumin, an a/j3 protein of large size (75 kDa). In all three cases, the unfolded

proteins seem to have reached a degree of compactness before interacting with PhPFD.

Curiously enough, and despite the fact that the tips of both PhPFDa and PhPFDP

subunits contain a large number of hydrophobic residues in their inner surface. the tips of

one of the PhPFDP pairs are always the ones to recognize and bind the unfolded protein.

A small protein like lysozyme (14 kDa) only requires such an interaction (Figure 2-8A)

but larger ones require binding to additional PhPFD subunits (Figure 2-8C and D). Our

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reconstructions also reveal an inherent plasticity in the PhPFD con~plex allowing it to

interact with substrates of vastly differcnt s i x , consistent with the two-dimensional

imaging (Figure 2-7). Finally, the observation that the PhPFDa subunit can functionally

complement for PhPFDP when i t is truncated (Figure 2-9) is consistcnt with the intrinsic

properties of coiled coils being most important for archaeal PFD function ( i .0. based on

their hydrophobic interhelical core: Figure 2-4, 2-5 and 2-6).

2.4.7 Comparison of archaeal and eukaryotic PFD binding mechanism

The three-dimensional structures of PhPFD complexed to three unfolded proteins

reveal a stsuctural plasticity of the archaeal chaperone, since its tentacles deviate from the

structure obtained in the apo-PhPFD to accomniodate the denatured proteins (Figure 2-

8). This finding is remarkable because it shows for the first time that the jellyfish-like

architecture and flexibility of archaeal PFD is ideally suited for interacting with a diverse

array of non-native proteins with different sizes and shapes. reflecting the likcly gencral

function of the chaperone in assisting de m v o protein folding. Even more interesting is

our finding that the three unfolded proteins are not confined in the cavity formcd by the

PhPFD tentacles but rather protrude fi-on1 it (Figure 2-lUA, 2-IOB and 2-10C). This is a

surprising result, given the fact that in the three-dimensional reconstruction of the

eukalyotic PFD-unfolded actin complex (Martin-benito et al., 2002), the cytoskeletal

protein is found almost entirely encapsulated in the chaperone cavity (Figure 2-1 OD).

The difference in localization cannot bc ascribed to substrate sizes, as actin has a

molecular mass (42 kDa) intermediate between that of GFP (27 kDa) and conalbumin (75

kDa), both of which are nearly excluded from the archaeal PFD cavity (Figure 2-8). In

fact, a three-dimensional reconstruction carried out with 28 12 particles of a complex

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between PhPFD and unfolded actin shows the cytoskcletal protein not encapsulated in the

chaperone cavity but interacting with the tips of the PhPFD tentacles (Figure 2-1OE).

The cylindrical shape of the unfolded actin, is similar however to that shown to be

interacting with the eukaryotic PFD (Martin-benito et a]., 2002) or with the chaperonin

CCT (Llorca et al., 1999), which strengthens the notion of actin reaching a ccrtain degree

of sccondary structurc by itsclf, bcfore interacting with the chapcrones (Schiiler et al.,

2000).

The difference between substrate interaction in the case of the archaeal PFD and

encaps~~lation in the eukaryotic one suggests a distinct role for the two types of PFDs that

might have originated when the simpler archaeal-like PFD evolved towards a structure

with a more complex subunit composition and function in the ancestral eukaryotic

chaperone. The evolution of PFDs correlates with the evolution of the Group 11-type

chaperonins that they serve (Leroux and Hartl. 2000). Whereas archaeal chaperonins and

PFDs arc composed respectively of 1-3 and 2 types of subunits, the eukaryotic cytosolic

chaperonin CC'T and PFD are coinposed respectively of 8 and 6 different subunits. This

co-evolution towards a higher complexity correlates with a specialization in the function

of both cliaperonins and PFDs, so whereas the archaeal PFDs and chaperonins seem to

act on a variety of substrates (Gu tsche et al., 1 999; Leroux et a]., 1 999; Leroux and Hartl,

2000), the eukaryotic PFD and C'CT has been shown to be mostly involved in the folding

of a more limited set of substrates, including two (actins and tubulins) that are restricted

to the eukaryal domain (Geissler et al., 1998; Siegers et al., 2003; Vainberg et al., 1998;

Valpuesta et a]., 2005).

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The evolution of PFDs i n terms of structure and specialization seems to be

associated with a change in their function, fro111 an archaeal chaperone that traps and thus

stabilizes unfolded proteins until their transfer to the thermosome, to an eukalyotic onc

that recognizes a certain set of unfolded proteins (i.e. actins and tubulins) and shields

them in its cavity ~mtil their transfer to K T . This protective role of the euka~yotic PFD is

so important that its presence increases by at least 5 fold the amount of actin folded by

CCT iri vivo (Siegers et al., 1999). The change in the role of PFD, from a stabilizer and

carrier in the archaeal PFDs to a more conlplex, protective role in the eukaryotic ones,

must be accompanied by changes in the mechanism of substrate recognition and

interaction. Therefore, whereas the recognition mechanism in archaeal PFDs relies on

non-specific, hydrophobic interactions (Okochi et al., 2004; Siegert et al., 2000), the

eukaryotic PFDs have evolved more specific interactions based on particular sequences

in the chaperone and the unfolded protein. T h ~ s has been shown for the cytoskeletal

proteins b-actin, a-, P-, y-tubulin, and actin-related protein ARP- I (Rommelaere et al.,

200 I), which seem to possess at least two conserved PED-binding sites in their sequence.

Likewise, truncation experin~ents in the subunits of human PFD reveal specific domains

for interaction with tubulin and actin (Sirnons et al., 2004).

2.5 Conclusion

In the present study of the archaeal molecular chaperone prefoldin, we uncovered

a singular ability of its coiled coils to interact with and stabilize non-native proteins. The

substrate-binding mechanism of PFD appears to depend on at least three distinct

properties of the coiled coils.

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I. A flexible molecular clamp-likc motion, apparently as a means to grip

substrates of varying shapes and/or sizes.

. . 1 1 . Interlielical hydrophobic residues at the distal tips that are likely to directly

contact exposed apolar patches in non-native substrates.

. . . 111. A concerted action of multiple weak binding sites, where the four outer P

subunits appear to contribute more to binding than the two central a

subunits.

Our three diniensional electron microscopy studies (Figure 2-8, 2-9 and 2-10)

confirm the flexible nature of archaeal PFD and the localization of the substrate binding

site to the very tips of the coiled coil tentacles. Moreover, these structural studies reveal

an increase in the number of subunits employed in substrate binding for substrates of

increasing size. The EM reconstructions also show that much of the substrate protrudes

froni the PFD cavity. even for small substrates (Figure 2-10). As predicted by our

biochemical studies, the a-subunits can functionally replace the P-subunits for lysozyme

binding, supporting the notion that both subunit-types rely on similarly non-specific

hydrophobic coiled-coil properties to bind substrates (Figure 2-9).

In contrast to archaeal PFD, i t has been suggested that eukaryotic PFD may

interact specifically with a limited number of substrates given that it has evolved six

divergent and potentially specialized subunits (Geissler et al., 1998; Hansen et al., 1999;

Leroux et al., 1999; Siegers et al., 1999; Vainberg et al., 1998) however, it is conceivable

that some of its six subunits rely on the same properties as PFD to bind exposed

hydrophobic patches on its substrates. Indeed. the C. elcog~~ns PFD6 coiled coil

complemented the function of the chimeric archaeal chaperone more efficiently than

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otlicr exogenous coiled coils (Figure 2-5). Moreover, because of their comparable

quaternary structures (Martin-benito et al., 2002; Siegert et a]., 2000), it is possible that

eukaryotic PFD could alter its cavity shape to acconiniodate different substrates, in the

same manner as the archaeal chaperone. While the scope of eukaryotic PFD function is

unknown our studies have revealed a distinct mechanisni of substrate-binding for the

archaeal and eukaryotic cliapcrones. Eukaryotic PFD seems to totally encapsulate its

substrate(s), contacting a large surface area with its largely hydrophobic cavity surface

(Martin-Benito et al., 2002; Figure 2-10). This is in contrast to archaeal PFD which, even

when binding the same denatured-actin substrate, binds using only with the hydrophobic

patches at its tentacle tips, leaving much of the substrate protruding from its cavity

(Figure 2-10).

Coiled coils are highly abundant in the proteomes of all organisms, accounting for

an estimated 2-3% of all protein residues (Wolf ct a]., 1997). Therefore, in addition to

shedding light into the chaperone function of archaeal, and potentially eukaryotic, PFD,

our findings are of particular significance because all coiled coils share a nonpolar core.

The binding property observed for this region in archaeal PFD could extend to other

coiled-coil-containing proteins, including molecular chaperones or those that interact

with any molecule exposing an apolar surface. Cofactor A, for example, is a tubulin-

specific three-stranded coiled-coil cliapernne that appears to stabilize quasi~iative P-

tubulin on its niostly hydrophilic surface (Steinbacher, 1999). The contribution of the

interhelical apnlar residues may play an unrecognized yet important role in substrate

binding. The five-stranded coiled-coil protein, COMPcc, is an interesting case of a

nonchaperone protein that binds vitamin D within the network of apolar ald residues

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(Ozbek ct a]., 2002). Although this latter intcraction is highly specific, other coiled coils

could conceivably bind a range of hydrophobic n~olecules using their interhelical

hydrophobic residues. In conclusion, our findings provide significant insight into the

mechanism of the molecular chaperone function of PFD and extend the known functions

of coiled coils to include molecular recognition via their common hydrophobic interface.

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2.6 Figures

Figure 2-1 Prefoldin (PFD) constructs used in this study

Primary amino acid sequences of iWethuno/l7c~1n10h~1c*/e/. /hei.i~~o~~z~/oti~opI~ic'z~.s and

Pyr-ococcw.s horikoshii PFDa and PFDP variants. Secondary structures are indicated as an

h for a-helix and an s for P-strand. In this study, wild-type P subunit (P) refers to the

fully active f01-111 used in Siegert et a]. (Siegert et a]., 2000). Predicted hydrophobic a/d

residues in the heptad repeat of the coiled coil are indicated by a or d and shown in bold

in the sequence. Switched N- and C-terminal regions are underlined. Exogenous coiled-

coil sequences are shaded gray, and serine substitutions are highlighted in black.

3 subunits

2 s t r u c t . hhhhhhhhhkhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhh

C o i l a d c o i l ( d ) a d a d a d a d a d a d

ME~RLEEIVNQLNIYQSQVELIOQQMEAVRATISELEILEKTLSDIQG--KDCS~LVPVGAGSFII Phu M I R M A Q m K E L E K L A Y E Y Q v L Q A Q A Q I W I Q N L E L L N L P I G A G s F L l - I RLEEIWQLNIYQSQVELIOQOMU\VRATISELEILEKTLSDIQG--KDGSETLVPVGAGSFIl

C I EIWQLNIYQSQVELI~EAVRATISELEILEKTLSDIQG--KDGSETLVPVGAGSFII I M QLNIYQSQVELIOOQHEAVRATISELEILEKTLSDI~--KDGS~LVPVGAGSFII I EAVRATISELEILEKTLSDIQG- -KDGSETLVPVGAGSFII

MgL~IBggRIW~SQVELI~EAVW\TISELEILEKTLSDIQc--KDGSETLVPVGAGSFII

[I subunits

2 utruct.

coiled coil

M E L P O N V Q H Q L A Q P O Q L P O Q A Q A I S V Q A ~ M Q I N E T Q K K S S G N I L I R V A MQNIP~VQAM~LDTYOOQLQLVIgQKQKVQADLNEAKKALEEIETLPDDAQIYKTVGTLIVK~

LAQFQQWAQAISVQKQTVEMQINmKALEOLSRAADDAFWKSSGNILIRVA eAQAISVQKQTVEMQINETQKALEELSRAADDAEVYKSSGNILIRVA

QTVEMQINETQKALEELSRAADDAENYKSSCNILIRVA RQTVP(QINETOKALEELSWDAWYKSSGNIL1RVA

Mlunumm TVEMQINETQKALEELSRAADDAEWKSSGNILIRVA

+ ~ I ~ O P I X H ~ ~ ~ Q N E M Q ~ N ~ K A L E E L S R R A D D A E W K S S G N I LIRVA

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Figure 2-1 continued

assso sees hhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhh

~ELKDTSEVIMSVGAGVAIKKNPEDA#ESIKSQICNELESTLQWENLRKITDIHMKLSPpAEELLKKVRGSGE {GVIVDKNNAIVSVGSGYAVERSIDEAISFLEKRLKEYDEAIKKTQGALAELEKRIGEVARKAQWQOKQS~SFKVKK CAELKDTSEVIHSVGAGVAIKKNFEDAMESIKSQKNELEST~KHGENLRKITDIHnKLS K KVRGSG E CAELKDTSEVIMSVGAGVAIKKNFEDAMESIKSQKNELESTMKHGENLRKITDI~KLS KKVRGSGE ~ELKDTSEVI~SVGAGVAIKKNFEDAMESIKSOKNELES~LQKMGENLRKITDII.IM~ KKVRGSGE EAELKDTSEVIMSVGAGVAI KKNFEDAMESI KSQKNELESTLQKLlLMplLEVOSOY ( A E L K D T S E V I H S V G A G V A I K K N S E D A M E S I K S Q K N E L E S T L Q K M G E N L R K I T D ~ ~ ~ ~ Q C V T P

hhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhhh

KDELTEEMEKLETLOLRSKTIERQSERVMKKMEMQVNIQUIMKGA KEKAVQELKEKIETLEVRLNALNRQEQKINEKVKELTQKIQAALRPPTAG KDELTEELQEKLETLQLREKTIERQEERVHKKLQEMQVNIQEAMK KDELTEELQEKLETLQLREKTIERQEERVHKKLQMQVNIQ KDELTEELQEKLETLQLREKTIERQEERVHKKLQMQ KDELTEELQEKLETLQLREKTI ERQEERVMKKLQ*

m E L T E E r X ) E K L E T m L R E K n r r w I m I ~ ~ p Q K D E L T E E L Q E K L ~ T L Q L R E ~ P ~

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Figure 2-2 multiple sequence alignment of archaeal prcfoldin subunits

(A) Column score output of a Gonnet PAM 250 similarity matrix. Grey bars represent ultl

residues at the interhelical interface of the coiled coils, and secondary (2") structures are

indicated as a-helix, cylinder, and P-strand, arrow.

2" struc. ( I . subunits - QQQ-

2' struc.

[I subunits - 00-

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(B) Multiple sequence alignmcnts of PFDa and -P subunits froin I I archacal spccics,

with L I / L / hydrophobic residues marked at the top of the alignment. Residues similar at

1011 1 residues are colored yellow, or light blue if they are at UIL/ hydrupl~obic core

positions. Residues identical at 1011 1 positions arc colored purple. Mt,

~ M c ~ / l i ~ ~ r ~ o / l i c r ~ ~ ~ ~ o l ? ~ ~ c / e ~ - / /~~ i~ inouz i /~ /~~oyh ic~z~ . s ; Ap, Aei.opjv-zir11 yc.inix; Af, Ai.c~/~~~c~oglohzi.\

1; subunits

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Figure 2-2 continued

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Figure 2-3 Architecture and surface conservation of archaeal prefoldin

(A) Space tilling representation of prefoldin ( 1 FXK) shown looking down into the cavity

(Left) and from inside, viewing the cavity surface of three subunits (Right). The N-

terminal helix (a4-29; P516-29) is blue, and the C-terminal helix (a1 1 1- 141; P88- I I I) is

orange. (B) Conservation of the prefoldin coiled-coil regions inside vs. outside the cavity

across 1 1 archaeal species. Residues are colored as in the alignment in Figure 2-2. Two

similar amino acids at the N terminus of the P-subunit (residues 3 and 4) are not shown

because they were not resolved in the crystal structure.

top vlew of cav~ty

inside view

ins~de view

outs~de view

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Figure 2-4 Chaperone activity of intradomain swap (switch) mutant complexes

(A) Scheniatic representations of a and P switch mutants. For the wild-type subunits, the

N- and C-terminal helices are colored white and gray, respectively; the interhelical aid

rcsidues are represented as dark ovals and correspond to thosc shown in the PFD subunit

alignments (Figure 2-2). For tlie switch mutants, tlie numbering scheme and colors used

correspond to the wild-type sequences and show where the crossover points occur. (B)

Effect of PFD switch mutants on the aggregation of denatured lysozynic. Relativc

aggregation of 2 pM lysozymc (monitored at 360 nm) during I0 min in buffer alone or in

the presence of wild-type or prefoldin variants (as shown on the right of each curce). PFD

complexes were at 2 pM unless otherwise indicated. [NOTE: This data was generated

by V.F.L. and myself. j

., , buffer Am'

0 1 2 3 4 5 6 7 8 9 1 0 ' A p Time (minutes)

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Figure 2-5 Chaperone activity of chimeric complexes

(A) Schematic representations of chimeric PFD subunits. The exogenous coiled-coil

region is colored black, and thc n~imbers adjacent to the helical regions refer to the amino

acid position at which the fusion has taken place. (B) Effect of chimeric PFD mutants on

the aggregation of 2 pM denatured lysozyme. PFD con~plexes (as shown on the right of

each curve) were at 2 pM unless otherwise indicated. [NOTE: This data was generated

by V.F.L.]

Time (minutes)

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Figure 2-6 Hydrophobic ald coiled-coil residues are required for chaperone activity

(A) Schematic representations of the ald residue point mutants. Hydrophobic residues

bctween the coiled coils are dark ovals; residues mutated to serine are colorcd whitc.

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(B) Effect of hydrophobic a/d point mutants on the aggregation of 2 p M denat~ired

lysozyme. Each PFD complex, shown on the right, was tested at 2 pM. (C) Mutated

hydrophobic rcsidues shown on the PFD crystal stnicture ( I FXK). PFD is shown looking

into the cavity (Left), and from inside, viewing the cavity surface (Right). Residues that

were mutatcd are colored green. Not shown is one pair of hydrophobic residues (L3 and

A1 13) in thep subunit termini and one amino acid (D.3) in the N terminus of the a

subunit, which were not resolved in the c~ystal structure but may also form part of the

coiled coil (4, 6)

Time (minutes) Time (minutes)

buffer U,Hlv12~p14

top view of cavity inside view

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Figure 2-7 Substrate binding occurs near the ends of flexible coiled coils

(A) Coelution of PFD and GFP on a superdex 200 size-exclusion column (Left). The

green dashed line is PFD (PhPFD) plus denatured GFP monitored at 222 nrn, the thick

blue line is GFP alone monitored at its excitation lnaxiinuin (396 nm), and the red line is

PFD plus denatured GFP monitored at 396 nni. Coelution was also demonstrated by

SDSIPAGE analysis of peak A (1.25-1 3 0 in]) with and without PIIPFD and of peak B

without PhPFD (1.65-1.80 ml) (Right). (B) Interaction of unfolded GFP with PFD. ( I )

Molecular surface of PFD crystal structure; negatively stained and avcraged EM images

of substrate-free prefoldin (2), and GFP-bound PFD (4). (3 Upper) Mergcd contour maps

(blue, PFD alone; red, PFD + GFP). (Lower) The approximate t i l t angle change (-1 2"

opening) of the substrate-bound PFD subunits (red) relative to that of PFD alone was

estimated manually from B3 using Adobe Illustrator C S (Adobe) (the contour map is

shaded bluc).

A

I peaks A 6 300 n n

. PFD+GFP , (222 nm) -15 % 5 GFP o

Volume (ml)

+ GFP

+ PhPFDu 15-

10- + PhPFDll

GFP + + + PFD - + -

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(C') The putative hinge domains connecting PFD coiled coils and the fi-barrel domain are

shown with arrows (I FXK). Gray dashed lines indicate the (approximate) 12" opening

motion, and the P-barrel oligomerization domain regions are circled with narrow black

H M 3 HM4 dashed lines. (D) Effect of wild-type (Ph and Mt) and Mta P PFD conlplexes on the

aggregation of denatured conalbumin (75 kDa). Aggregation assays were performed as

for denatured lysozyme except conalbumin was 0.75 pM and PFD or its variants (Right)

were added at a 5: 1 ratio over substrate (3.75 pM). [NOTE: Data in B and C were

generated by J.G.R. and J.IM.VI

Time (minutes)

buffer

(XHM3 HM4 P

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Figure 2-8 The three-dimensional reconstruction of the complex between PhPFD and several unfolded proteins

A) Three orthogonal views of thc three-dimensional reconstruction of apo-PhPFD. B-D) The

same views of the three-dimensional reconstructions of PhPFD complexed to unfolded

lysozyme (B), GFP (C) and conalbumin (D). The bottom images correspond respectively to the

atomic structures of lysozynie, GFP and conalbumin, at the same scale. Bar represents 50 A.

[NOTE: J.M.B., J.G.R., P.G.P., J.Boskovic., P.C., J.J.F., J.Berenguer and J.M.V.

generated this data. We generated recombinant prefoldin and demonstrated PhPFD

binding to the substrates.]

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Figure 2-9 The role of PhPFDa and PhPFDP subunits in the interaction with unfolded substrates

A) Effect of truncation of the tips of PhPFDa and PhPFDlJ subunits in the prevention of

lysozyme aggregation. B) Two orthogonal views of the three-dimensional reconstruction of the

conlplex betwcen P ~ P F D ~ ~ " and unfolded lysozyme. C) Effect of truncation of the tips of

PhPFDa and PhPFDP subunits in thc prevention of conalbumin aggregation. T~uncations are

described in section 2.3.1. [NOTE: J.M.B., J.C.R., P.C.P., J.Boskovic., P.C., J.J.F.,

J.Berenguer and J.M.V. generated the data in B. V.F.L generated the data in C.1

. . . a41Tr

rn . rn . rn AlM.(ype PFD

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Figure 2-10 Localization of the unfolded substrates in archaeal and eukaryotic

PFDs.

A-C) Two orthogonal vicws of the three-dimensional reconstruction of the complex

between PhPFD and unfolded lysozyme (A), GFP (B) and conalbumin (C). D) The same

two views of the three-dimensional reconstruction of the complex between human PFD

and unfolded actin. In red the mass corresponding to the unfolded substrate. E) The same

two views of the three-dimensional reconstruction of the complex between PhPFD and

unfolded actin. In all cases the unfolded protein is depicted in red, except in (D, bottom),

which is colored in light red to indicate that the mass of the unfolded actin is enclosed in

the chaperone cavity. [NOTE: J.M.B., J.C.R., P.C.P., J.Boskovic., P.C., J.J.F.,

J.Berenguer and J.M.V. generated this data. The experiments with archaeal

prefoldin and actin were our own idea.]

Lysozyme GFP Conalbumin Human PFD + PhPFD + Actin Actin

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2.7 Tables

Table 2-1 Thermal stability of prefoldin variants monitored by circular dichroism ellipticity at 222 nm

Construct T"I, c* Construct T,,l, O C*

M t up conlplex 6 1 p~~, l t~so~h,~4 5 8

Ph a p complex >80 u l ~ h 4 I 6 1 5 \\I a 5 9 C I ~ ~ 2 60

pSW 5 3 u H M l 5 9

uR:'t150 6 1 pHM1 5 8

p('cP1~"h 43 pHM2 54

P ~ ~ ~ ~ ' ~ ~ ' ~ 48 pH"' 57

~ ~ ~ ' ~ l ~ ~ 69 pHM4 53 *Melting temperature (T,,,) reported is the dissociation of the prefoldin complex

containing the indicated subunit in a wild-type background (Fandrich et al., 2000).

PhPFD truncations did not melt under the assay conditions as shown for wildtype PhPFD.

[NOTE: This data was generated equally by V.F.L. and mysc1f.l

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2.8 Reference list

Bukau, B., and Honvich, A. L. (1998). The Hsp70 and Hsp60 chaperone machines. Cell 92.35 1-366.

Deuerling, E., Schulze-Specking, A., Tonioyasu, T.. Mogk, A., and Bukau, B. (1999). Trigger factor and DnaK coopcrate in folding of newly synthesized proteins. Nature 400,693-696.

Fandrich, M., Tito, M. A., Leroux, M. R., Rostom, A. A., Hartl, F. U., Dobson, C. M., and Robinson, C. V. (2000). Observation of the noncovalent assembly and disassembly pathways of the chaperone coniplex MtGimC by mass spectrometry. Proc Natl Acad Sci USA 97, 141 5 1-141 55.

Geissler, S., Siegers, K., and Schiebel, E. (1 998). A novel protein complex promoting formation of functional alpha- and gamma-tubulin. EMBO J 17, 952-966.

Gutsche, I., Essen, L. O., and Baunieister, W. (1999). Group I1 chaperonins: new TRIC:(k)s and turns of a protein folding niachine. J Mol Biol 293, 295-3 12.

Hansen, W. J., Cowan, N. J., and Welch, W. J. (1999). Prefoldin-nascent chain complexes in the folding of cytoskeletal proteins. J Cell Biol 145,265-277.

Hartl, F. U., and Hayer-Hartl, M. (2002). Molecular chaperones in the cytosol: from nascent chain to folded protein. Science 295, 1852-1 858.

Lawrence, J. R., and Johnson, W. C'. (2002). Lifson-Roig nucleation for alpha-helices i n tritluoroethanol: context has a strong effect on the helical propensity of amino acids. Biophys Cheni 10 1-1 02,375-385.

Leroux. M. R., Fandrich. M., Klunker, D., Siegers, K., Lupas, A. N., Brown, J. R., Schiebel, E., Dobson, C. M., and Hartl, F. U. (1999). MtGimC, a novel archaeal chaperone related to the eukaryotic chaperonin cofactor GimCIprefoldin. EMBO J 18.6730-6743.

Leroux, M. R., and Hartl, F. U. (2000). Protein folding: Versatility of the cytosolic chaperonin TRiCICCT. Clurr Biol 10, R260-264.

Llorca, O., McCormack, E. A., Hynes, G., Grantham, J., Cordell, J., Carrascosa, J. L., NJillison, K. R., Fernandez, J. J., and Valpuesta, J. M. (1999). Eukaryotic type I1 chaperonin CCT interacts with actin through specific subunits. Nature 402, 693- 696.

Ludtke S. J., Baldwin, P. R., and Cliiu, W. (1999). EMAN: semi automated software for high-resolution single-particle reconstructions. J Struct Biol 128, 82-97.

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Lundin, V. F., Stirling, P. C., Gomcz-Reino, J., Mwenifi~mbo, J. C., Obst, J. M., Valpuesta. J . M., and Leroux, M. R. (2004). Molecular clamp mechanism of substrate binding by hydrophobic coiled coil residues in the archaeal chaperone. prefoldin. Proc Natl Acad Sci USA 101,4367-4372.

Lupas, A. (1 996). Coiled coils: new structures and new functions. Trends Biochem Sci 2 1,375-382.

Marabini, R.. and Carazo, J. M. (1 994). Pattern recognition and classification of images of biological macromolecules using artificial neural networks. Biophys J 66, 1804- 1814.

Martin-Benito, J., Boskovic, J., Gomez-Puertas, P., Carrascosa. J. L., Simons, C'., Lewis, S. A., Bartolini, F., Cowan, N.C., and Valpuesta, J. M. (2002). Structure of eukaryotic prefoldin and of its complexes with unfolded actin and the cytosolic chaperonin CCT. EMBO J 2 1, 6377-6386.

Myszka, D. G., and Chaiken, I . M. ( 1994). Design and characterization of an intramolecular antiparallel coiled coil peptide. Biochemistry 33, 2363-2372.

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Schiiler, H., Lindberg, U., Scliutt, C. E., and Karlsson, R. (2000). Thermal unfolding of G-actin nionitored with the DNase I-inhibition assay. Eur J Biocliem 267,476-486.

Siegers, K., Waldniann, T., Leroux, M. R.. Grein, K., Shevchenko, A., Schiebcl, E., and Hartl, F.U. (1999). Compartmentation of protein folding in vivo: sequestration of non-native polypeptide by the chapcronin-GinK system. EMBO J 18, 75-84.

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CHAPTER 3 IN VITRO AND IN VIVO ANALYSES IDENTIFY PHOSDUCIN-LIKE PROTEIN 3 AS A NOVEL COFACTOR OF THE CHAPERONIN CCT

Note regarding contributions: The following chapter was published in the Jozrrnal ~fBiologicnl Cl~cmistry. The

a~~tl iors of the study are listed below.

Stirling, P.C., Cuellar, J . , Alfaro, G.A., El Khadali, F., Beh, C.T., Valpuesta, J.M., Melki, R., and Leroux, M.R. (2006). .J Biol Chem 28 1 70 12-702 1 .

As the first author I contributed most of the data and wrote the article. J. Cuellar and J.M. Valpuesta contributed the electron microscopy in Figure 3-4. G.A. Alfaro and C.T. Beh provided strains, technical advice and provided suggestions to improve the manuscript. F. El Khadali and R. Melki did the ATPase assays (Figure 3-6) and the sedimentation experiments in Figure3-2B, C and D.

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3.1 Abstract

Many ATP-dependent molecular chaperones, includmg Hsp70, Hsp90, and the

chaperonins GroELIHsp60, require cofactor proteins to regulate their ATPase activities

and thus folding f~~nctions it1 vivo. One conspicuous exception has been the eukaryotic

chaperonin CCT. for which no regulator of its ATPase activity, other than non-native

substrate proteins, is known. We identify the evolutionarily conserved PhLP3

(phosducin-like protein 3) as a modulator of CCT function it1 vi/t.o and in vivo. PhLP3

binds CCT, spanning the cylindrical chaperonin cavity and contacting at least two

subunits. When present in a ternary complex with CCT and an actin or tubulin substrate,

PhLP3 signitkantly diminishes the chaperonin ATPase activity, and accordingly, excess

PliLP3 perturbs actin or tubulin folding in vilr-o. Most interestingly, however, the

S L I C , C - ~ L ~ I . O ~ ~ ! I C * C . C ~ I ' C V ~ S ~ L ~ ~ PhLP3 homologue is required for proper actin and tubulin

function. This cellular role of PhLP3 is most apparent in a strain that also lacks prefoldin,

a chaperone that facilitates CCT-mediated actm and tubulin folding. We propose that the

antagonistic actions of PhLP3 and prefoldin serve to modulate CCT activity and play a

key role in establishing a functional cytoskeleton in vivo.

3.2 Introduction

A significant proportion of proteins requires the assistance of molecular

chaperones to ensure proper biogenesis during and following translation on ribosomes

(Hart1 and Hayer-Hartl, 2002; Stirling et al., 2003). Most chaperones that actively fold

polypeptides depend on ATP hydrolysis, a process often regulated by cofactor proteins.

For example, Hsp40 modulates the ATPase function of Hsp70, whereas factors such as

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GrpE or Bag1 control nucleotide exchange (Bimston et al., 19%; Szabo et a]., 1994).

Hsp9O is also regulated by many cofactors, for example p23 and C'dc37 (Siligardi et al.,

2004; Young and Hartl, 2000). One class of toroid-shaped chaperones, termed

chaperonins, are conserved across all domains of life and assist the folding of many

cytosolic proteins (Thulasiraman et a]., 1999; Kerncr et al., 2005). In eukaryotes, the

chaperonin containing TCP-I (CCT, also termed TRiC or c-cpn) consists of two stacked

rings each formed by eight related subunits (Hartl and Hayes-Hartl, 2002; Stirling et a].,

2003; Spiess et a]., 2004). C'CT binds substrate proteins and, through ATP-dependent

confor~national changes, encapsulates them in a central cavity. Upon release into the

cytosol, the substrates may have folded to the native state or may require additional

rounds of CCT binding and release (Farr ct al., 1997; Siegers et a]., 1999). CCT is

required for folding nascent actin and tubulin and has been shown by

immunoprccipitation studies to interact with a wide range (1 0%) of polypeptides. many

of which may be substrates (Thulasiraman et a].. 1999). Unlike other ATP-dependent

chaperones, there is no evidence that C'C'T cooperates with protein cofC~ctors to modulate

its ATP hydrolysis. Hop/p60. a cofactor of Hsp70 and Hsp90, pron~otes nucleotide

exchange by CCT in vilrw, but the significance of these findings is unknown (Gebauer et

al., 1998).

CCT differs from its bacterial chaperonin counterpart, GroEL, in that i t does not

encapsulate its substrates with a GroES-like cofactor that tits over the chaperonin cavity

(Hartl and Hayes-Hartl, 2002). Instead, built-in protrusions within the apical regions of

CCT close the central cavity during folding (Spiess et al., 2004; Meyer et al., 2003). At

least for actins and tubulins, another chaperone, prefoldin (PFD, also named GimC),

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participates in CC'T-mediated folding. PFD is a heterohexan~eric complex that uses its

octopus-like structure to clamp onto non-nativc proteins (Siegert et al., 2000; Martin-

Bcnito et a]., 2002; Lundin et al., 2004). In eukaryotes, PFD interacts with nascent chains

and facilitates their transfer to CCT via direct interactions with the chaperonin (Siegers ct

a]., 1999; Martin-Bcnito et al., 2002; Geissler et al., 1998; Vainberg et a]., 1998; Hansen

et al., 1999). PFD also promotes the efficient release of native actin from CC'T (Siegers et

a]., 1999) by a n~echanisn~ that is not understood.

Recently, PhLPl (phosducin-like protein I) was shown to inhibit CCT-mediated

folding, and a regulatory interaction was proposed (Mclaughlin et a]., 2002). Phosducin,

which, ~ ~ n l i k e PhLPI, does not interact with CCT, modulates retinal phototransduction by

binding Gp subunits of transducin and preventing their reassociation with G a following a

signaling event (Mclaughlin et al., 2002; Yoshida et al., 1994). The three known families

of phosducin-like proteins likely participate in G-protein signaling, but they have also

been implicated in other processes.

PhLPI, a close relative of phosducin, binds newly made GP and assists in the

assen~bly of a GPy complex (Lukov et al., 2005). A recent electron microscopy

reconstruction of a mamnialian C'CT-PhLP I complex shows that PhLP I binds to the

apical domains of multiple chaperonin subunits, siniultaneously occluding the cavity of

the cis ring and altering the conformation of the trvrns ring (Martin-benito et a]., 2004).

However, the biological consequence of PhLPl binding to the chaperonin is not

completely understood. PhLP2 has an unknown but essential function in yeast and

Dic~tyostclirrtn, where it has been implicated in cell cycle progression and G-protein

signaling (Blaauw et al., 2003; Flanary et al., 2000). PhLP3 (called APACD or TXNDC9

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in manimals) has been linked to G-protein signaling in yeast but also influences tubulin

fiinction in both yeast and Ctrenol.l~trhtli/iss deg~rn.s (Flanary et al., 2000; Lacefield and

Solomon, 2003, Ogawa et al.. 2004). Deletion of the yeast PhLP3 gene orthologue,

PLPI, rescues the benomyl supersensiti\~ity of strains such asp~lcIOA (a PFD subunit) or

luh3A (an a-tubulin variant) that have an excess of undinierized j3-tubulin (Lacefield and

Solomon, 2003). Folded but undimerized 13-tubulin is thought to interfere with normal

microtubule asseniblylf~inction and is toxic when not associated with a-tubulin. Plp I p

does not affect the levels of p-tubulin but rather its folded state, so that in the absence of

Plp l p some 13-tubulin appears in nontoxic aggregates. Aside from the presence of

aggregates, the deletion of PLPI in yeast has not been shown to possess obvious defects,

consistent with the wild-type (WT) phenotype of PltLP3 knockouts in Dic.fyo.s/elizm~

disc~oitkw~n (Blaauw et al., 2003). Unlike yeast and Dic*/jm/~lilim, however, C. c1cgt1n.s

PhLP3 is essential as its disruption by RNA interference results in a failure of the first

embryonic cell division. The arrested emblyos possess short astral microtubules

compared with control embryos, suggesting that PhLP3 plays a role in niicrotubule

organization (Ogawa et a]., 2004).

In this study, we identified human PhLP3 as a novel CCT-binding protein. We

show that PhLP3 forms ternary complexes with CCT and cither actin or tubulin and

negatively impacts their folding. Functional assays suggest that this occurs by slowing

the ATPase activity of the chaperonin and not through direct competition with substrates.

In vivo, yeast PhLP3 appears to coordinate the proper biogenesis of actin and t~ibulin with

PFD. Our results idcntifj, PhLP3 as a novel CCT cofactor and suggest that the balance of

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PhLP3 and PFD activities helps CCT modulate the levels of folded actin and tubulin, an

essential process for maintenance o f a f~~nctional cytoskeleton.

3.3 Methods

3.3.1 Purification of PhLP3 and C C T

Hunian PhLP3 (full-length; GenBankTM accession number NM 005783) and a

truncated version (tPhLP3; residues 1-1 93), cloned into pRSET6a, or GST-tagged

variants (Figure 3-1 F) cloned into pGEX-6p, were produced in BL2 1 [DE3]pLysS as

described (Lundin et al., 2004). Inclusion bodies were washed in 50 mM Tris-CI, pH 8.0,

I mM EDTA, 0.75% Triton X- 100, resuspended in 20 mM Tris-CI, 8 M urea, pH 8.0,

filtered, and passed over a Q-Sepharose column (Aniershani Biosciences). Fractions

containing pure PliLP3 or tPhLP3 were refolded by dialysis against 20 mM sodium

phosphate, 100 mM NaCI, and 1 mM DTT, pH 8.0. GST-tagged proteins were purified

with glutathione-Sepharose 4B as per the manufacturer's instructions (Aniersliani

Biosciences). CCT was purified from rabbit reticulocyte lysatc as described (Melki et a].,

1997) and was shown to be f~inctional by its ability to refold denatured actin in v i / m

(Melki et al., 1997).

3.3.2 Cell culture

Hunian embryonic kidney (HEK) 293T cells were transfected with pCMV-n~~vc-

PhLP3 or empty vector with PolyFect reagent (Qiagen). For imniunoprecipitations, HEK

cells were lysed mechanically in 1P buffer (25 mM Tris-C'1, 100 mM KC], 2 1i1M EDTA,

I niM DTT, and 0. I mM phenylmethylsulfony1 fluoride, pH 7.4), and cell debris was

removed by centrifi~gation at 15,600 x g for 10 niin. The supernatant was incubated with

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anti-tiiyc~ beads (Covance) at 4 O C ' for 60 mln. Beads werc washed three times in IP buffcr

and analyzed by SDS-PAGE and Western blot. TCP I was detected with a rat anti-TCPI

antibody (StressGen Biotechnology). Cell lysatc supernatants were fractionated on a

Superose 6 column PC3.2130 (Amersham Biosciences). 52-pl fractions were collected,

and I0 pI of each was loaded on an SDS-polyacrylamide gel and Western-blotted for nlyc

or TCP 1 . Separately run size standards were blue dextran (2 MDa), thyroglobulin (669

kDa), aldolase ( I50 kDa), serum albumin (69 kDa), and carbonic anhydrase (29 kDa).

3.3.3 In vitro translation, folding assays, and GST pull-downs

Actin and tubulin were translated with the T7 quick-coupled translation kit

(Promega) in the presence of 2 pM recombinant PhLP3 or tPhLP3 (Cowan, 1998;

Leroux, 2000). At time points, aliquots of the reaction were frozen in native gel running

buffer and then thawed on ice and analyzed by native gel electrophoresis (Leroux, 2000).

GST pull-downs were done according to the manufacturer's instructions by using

glutathione-Sepharose-4B (Aniersham Bioscicnces). CCT ( 1 00 nM) was incubated with

I0 times molar excesses of GST, GST-PhLP3, or GST-PhLP3 and 30 timcs excess

untagged PhLP3. GST-fused trimcations of PhLP3 were incubated in 10% reticulocyte

lysate (Proniega) for 30 minutes on ice and precipitated according to n~anufacturer's

instructions.

3.3.4 Verification of actin and tubulin folding inhibition by PhLP3

We verified that PhLP3 inhibited native actin production by the addition of

DNase I , which interacts only with native actin. In the absence of full-length PhLP3, a

new band corresponding to a DNase I-actin complex appeared, but when PhLP3 was

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present. no such shift was observed because of the lack of native actin (Leroux, 2000). To

positively identify the lowest band in the tubulin-folding assay, we added taxol to the

reactions to stabilize microtubules, thus rcrnoving alp-heterodimers from thc material

loaded on the gel. The intensity of the lowest band was diminished in the prcsence of

taxol, indicating that hctcrodimers were present. Howevcr, the band did not disappear

completely, leaving the possibility that some p-tubulin-cofactor A complex is present.

"s -~ct in and s3'-P-tubulin were expressed and purified as described (Leroux, 2000).

3.3.5 ATPase activity measurements

ATP hydrolysis at 30 OC was measured in folding buffer containing 2 mM [.j-

3 ' ~ ] ~ ~ ~ and either I pM CCT alone, CCT with denatured client proteins (0.1 mglml

actin or P-tubulin), CC'T with PhLP3 (0.9 mglrnl) or CCT with denatured client proteins

and either full-length or truncated PhLP3 (tPhLP3 at 1.1 mglml), by extraction of the

[ 3 2 ~ ] phosphomolybdate complex formed in 1 N HCI as described (Melki, et a]., 1990).

3.3.6 Purification of tubulin and microtubule-associated proteins

Dinieric alp-tubulin was purified from pig brain by three polymerization cycles

followed by phosphocellulose chromatography and stored at -80 O C in buffer D (0.05 M

PIPES, pH 6.9, 0.5 mM EGTA, 0.25 mM MgC'I2, 3.4 M glycerol. and 200 pM GTP)

(Melki et al., 1996). Microtubule associated proteins (MAPS) were isolated from

niicroh~bules by phosphocellulose and DEAE-Sephadex chromatography (Kuznetsov et

a]., 198 1 ) and stored at -80 OC in buffer D.

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3.3.7 Co-sedimentation assay

Microtubules (50 uM tubulin) were assembled in buffer E (0.1 M KjPOq, pH 7.5,

0.5 mM EGTA, 0.5 niM MgC1:) supplemented with 3 mM MgC12, I mM GTP, and a 2

M excess of taxol. After 15 min at 37 "C, the microtubule solution was supplc~nented

with PhLP3 (to 50 pM) or MAPS (to 0.5 mglml) and incubated at 37 "C for 30 min before

spinning at 200,000 x g at 37 "C for 10 niin in a TL1 00-Tabletop ultracentrifuge

(Beckman). Supernatant and pellet fractions were then analyzed on SDS-polyacrylamide

gels.

3.3.8 Sedimentation velocity measurements

Sedimentation velocity experiments were carried out with a Beckman Optima

XL-A analytical ultracentrifuge equipped with a 60 Ti four-hole rotor and cells with two-

channel 12-mm path length centerpieces. Measurements were made at 45,000 rpm and 15

"C using tubulin and PhLP3 (0.3 and 0.44 mglnil, respectively) in buffer E. The apparent

distributions of sedinienta tion coefficients were obtained with the program DCDT

(Stafford, 1 992).

3.3.9 Sample preparation for electron microscopy

CCT was purified from bovine testis as described previously (Martin-benito et al.,

2002). The CC'T-PhLP3 complexes were formed by incubating CCT and PhLP3 in a 1 : 10

molar ratio for 30 min at 25 "C. The CCT-tubulin complexes were formed by denaturing

bovine brain tubulin (Cytoskeleton, Inc.) in 6 M guanidine hydrochloride and subsequent

100-fold dilution in buffer (20 mM HEPES, pH 7.4, 50 mM KC1, 5 mM MgC'I2, 1 mM

EDTA, 2 mM DTT) containing 0.9 pM purified CCT (chaperonin:tubulin molar ratio of

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1 : 12). For the ternary con~plex between CC'T, PhLP3, and unfolded tubulin, denatured

tubulin was diluted 100 times in buffer containing PhLP3. After 5 niin, CCT was added

so that the CCT:PhLP3:tubulin molar ratio was I : 10: 10.

3.3.10 Electron microscopy and image processing

For electron microscopy of the various CC'T coniplexes, 5-p1 aliquots were

applied to glow-discharged carbon grids for I min and then stained for I min with 2%

uranyl acetate. Images were recorded at 0'-tilt in a JEOL 1200EX-I1 electron nlicroscope

operated at 100 kV and recorded at ~ 6 0 , 0 0 0 noniinal magnification. Micrographs were

digitized in a Zeiss SCAI scanner with a sampling window corresponding to 3.5 Mpixel.

For two-dimensional classification and averaging, top views of CCT particles were

selected, classified, and averaged using a maximunl-likelihood multireference refinement

algorithm (Scheres, et al., 2005) included in the XMIPP software package (Marabini and

Carazo, 1994).

3.3.1 1 Yeast strains, growth, and microscopy

For yeast strains, see Table 3-1. Singly deleted yeast strains were obtained from

the knockout collection (s288c background) and other mutants were made in house.

Yeasts were grown on YPD or YPD + 200 pg/ml geneticin sulfate (Invitrogen; Adams et

al., 1997). Matings and tetrad dissections were perfomled as described (Adams et al..

1997). Lat~unculin B (Sigma) sensitivity assays were performed according to (Ayscough

et al., 1997). For microscopy, cells were grown to mid-log phase beforc live imaging or

formaldehyde or methanol:acetic acid fixing and staining with TRITC-con-jugated

plialloidin (rho-phalloidin) or 4', 6'-diamidino-2-phenylindole (DAPI) (Sigma) according

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to manufacturer's instructions (Adams et al., 1997). For cell sizing, cell diameters were

measured perpendicular to the mother-bud axis at the largest point. Cellular defects were

statistically validated using either an independent variable t test or 2 analysis as

appropriate. The p-values associated with particular statistical tests are shown for each

phenotype scored in Table 3-2.

3.4 Results and discussion

3.4.1 Native PhLP3 associates with CCT likely as a monomer using Both N and C termini

Based on literature showing that PhLPl interacts with CC'T, and that yeast and C.

t.Iqans PhLP3 are implicated in P-tubulin folding and microtubule dynamics,

respectively (Mclauglilin et al., 2002; Lacefield and Solomon, 2003; Ogawa et al., 2004),

we investigated whether mammalian PhLP3 also interacts with CCT. I t could not be

assumed that PhLP3 would bind CCT, as i t has only 15% identity and 37% similarity Lo

human PhLPl, whereas phosducin, which does not bind CCT, is 38% identical and 64%

siniilar to PhLPI. However, production of PhLP3 in rabbit reticulocyte lysate followed

by native gel analysis showed that the radiolabeled PhLP3 co-migrates with the position

of CCT similar to what is seen for nascent actin (Figure 3-1A). To observe thc

association of PhLP3 and CCT in vivo, we expressed inyc epitope-tagged PhLP3 in HEK

cells and itnmunopl-ecipitated PhLP3 using a monoclonal antibody specific for the n?-vcc

epitope. CCT was efficiently co-immunoprecipitated from nlyc--PhLP3-expressing cells

but not froni cells transfected with vector alone (Figure 3-IB), suggesting that PhLP3

interacts physiologically with CCT. To assess whether the interaction is direct or indirect,

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wc expressed and purified recornbinant GST-tagged PliLP3 from Escherichi~~ cdi, and

we found that GST-PhLP3, but not GST alone; precipitated purified CCT complex in

pull-down assays. indicating that CCT directly interacts with PhLP3 (Figure 3-1C).

GST-tagged PhLP3 also selectively bound CC'T from either HEK cell or rabbit

reticulocyte lysates (Data not shown). Thc interaction of the purified components likely

involves native PliLP3 because addition of excess folded (native) untagged PliLP3 (sec

below) competes with GST-PhLP3 for CCT binding in solution (Figure 3-1 D). These

finding5 provide evidence that PhLP3 interacts with CCT it? vivo in a folded, native form,

rather than as a non-native substrate, siniilar to PhLPl (Mclaughlin, et al., 2002).

Untagged recombinant PhLP3 adopts a high degrec of secondary structure and is

thermally stable by circular dicliroism, two indications that i t is properly folded (Data not

shown). Moreover, PhLP3-CCT complexcs resemble native PhLP I -CCT coniplexes by

electron nlicroscopy, and PhLP3 does not behave like a non-native substrate in functional

assays with CCT (see Figures 3-4,3-5,3-6 below). Sedimentation velocity analytical

ultracentrifugation yielded measurenients of 2.7 S for PhLP3, consistent with its 26.5-

kDa monomeric size (Figure 3-2B). Additionally, when analyzed by size exclusion

chrornatography, myc-PhLP3 from HEK cell extracts eluted not only at the position

predicted for a PhLP3 monomer but also co-eluted with CCT (Figure 3-1 E). Our

observations are therefore consistent with the existence of cytosolic monomeric and

CCT-associated forms of PhLP3. Indeed, we also observed a cytosolic staining pattern

for my- and green fluorescent protein-tagged PhLP3 by ininiunocytochemistry as had

previously been published (Data not shown; Ogawa et al., 2004).

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Structurally. PliLP3 consists of a central domain with homology to tliioredoxin,

and flanking N- and C-terminal regions predicted to form a coiled coil and a potentially

disordered region, respectively (Gaudet et al., 1996). We created several truncations of

PhLP3 fused to GST and tested their ability to interact with CCT in 10% reticulocyte

lysate (Figure 3-1 F). Removal of 27 N-ter~ninal amino acids did not affect binding, but

removal of the cntire N-terminal region up to the thioredoxin domain (residues 65-226)

abrogated C'CT binding. Most interestingly, truncation of only 8 C-terminal amino acids

strongly diniinished CCT binding. Consistent with these findings, the thioredoxin domain

alone (residues 65-1 9 I), which is conserved in all phosducin and phosducin-like

proteins, is not sufficient for CCT binding (Figure 3-IF). Notably, both PhLP3 and

PhLPI seen1 to use similar regions for interaction with CCT. which suggests a similar

binding mechanism (Martin-benito et a]., 2004).

3.4.2 PhLP3 forms ternary complexes with CCT and actin o r tubulin

In light of the role for yeast PhLP3 (PLPI) in p-tubulin biogenesis (Lacefield and

Solomon, 2003), we hypothesized that PhLP3 may act as a molecular chaperone that

participates in CCT-mediated ti~bulin folding. We therefore investigated whether PhLP3

interacts directly with unfolded tubulin or actin, both of which are substrates of PFD and

CCT (Geissler et al., 1998; Vainberg et al., 1998). Our results indicate that in vitr-o,

PhLP3 does not appear to interact with non-native actin or tubulin in isolation in a native

gel shift experiment (Figure 3-2A), suggesting that i t may not act as a chaperone. To

assess whether PhLP3 may interact with other forms of tubulin, we tested for a potential

interaction with a- and P-tubulin monomers exchanging in and out of heterodimers or

with polymerized microtubules. PhLP3 was not found to interact with up-heterodimers

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undergoing exchange in solution (Figure 3-2B). PhLP3 also did not co-precipitate with

intact n~icrotubules after pelleting by ultracentrifugation (Figure 3-2C), although known

MAPS did (Figure 3-2D). Most interestingly, native PhLP3 did not interfere with the

ability of CCT to bind the denatured substrates even at a 10: 1 ratio (Figure 3-2A). This is

in contrast to the previous finding that the related protein PhLPI seems to directly

compcte with substrates for CCT binding (Mclaughlin et al., 2002).

Because CCT interacts with both PhLP3 and tubulin. we tested if PhLP3 could

affect tubulin biogenesis in a ternary complex with CCT and substrate. We translated a-

tubulin, P-tubulin, or actin separately in ['5~]n~ethionine-supplemented rabbit reticulocyte

lysate (which contains C'CT) and could precipitate all three radiolabeled polypeptides

with GST-PhLP3 (Figure 3-3A). This result shows that PhLP3 may not be specific for P-

tubulin, as suggested previously (Lacef eld and Solomon, 2003), and provides evidence

for a ternary interaction. If the con~plex between PhLP3 and the nascent protein is binary

instead of ternary, the interaction should take place in a heterologous E. coli in vitr-o

translation system, which lacks CCT. Figure 3-3B shows that E, coli lysate will not

support the interaction between PhLP3 and translated P-tubulin unless exogenous CCT is

addcd (Figure 3-3B). These data corroborate the notion that PhLP3 does not interact

directly with substrate proteins (Figure 3-2) and instead provide evidence that PhLP3

binds to CCT-substrate con~plexes, forming C'CT-PhLP3-substrate ternary con~plexes.

Such a conclusion is supported by electron microscopy experiments (see below) and by

f~~nctional assays w ~ t h CCT, PhLP3, and e~ther actin or tubulin (Figure 3-6).

To confirm the presence of ternary con~plexes, we compared negative-stained

electron microscopy images of CCT alone and CCT mixed with PhLP3, denatured

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tubulin, or both denatured tubulin and PhLP3. The average iniage of apoCCT (Figure 3-

4A) reveals an empty cavity, filled with stain and therefore darker, representing the

unoccupied substrate interaction site of the CCT complex (Llorca et a]., 2000). The

average image of the CCT-PhLP3 coniplex (Figure 3-4B) show^ a stain-excluding mass

crossing the chaperonin cavity, vely similar to the interaction between CCT and PhLP I

(Figure 3-4C) (Martin-benito et al., 2004), although in the casc of the comparativcly

smaller PhLP3 protein, the interaction secms to involve fewer CCT subunits. The averagc

image of the CCT-tubulin complex (Figure 3-4D), similar to that previously described

(Llorca et al., 2000), reveals a greater occlusion of the CCT cavity by the unfolded

tubulin compared with PhLP3. More importantly, the average image of CCT in complex

wit11 both PhLP3 and denatured tubulin (Figure 3-4E) differs from that obtained for the

C'CT-PhLP3 and the CCT-tubulin complexes, in that the stain-penetrating regions are

hardly noticeable (compare the chaperonin cavity in Figure 3-4E with those in Figure 3-

4B and 3-4D). Based on the images obtained, we favor the possibility that the substrate

and PhLP3 bind CCT in c ~ i s , possibly contacting one another in the context of the ternary

complex.

3.4.3 Excess PhLP3 inhibits actin and tubulin folding in vitro

To understand how PhLP3 might affect C'CT function, we carried out in vitro

actin and tubulin folding assays. Previous data showed that mammalian PhLPl inhibits

actin and luciferase folding by CCT (Mclaughlin et a]., 2002). Using G,a as a substrate,

the authors (Mclaughlin et al., 2002) proposed that PhLP 1 abrogated substrate binding to

CCT by direct competition, which does not seem to be the case for PhLP3 (Figures 3-2,

3-3,3-4).

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A time course experiment of newly translated actin or P-tubulin folding in rabbit

reticulocyte lysate was performed in the presence of either excess PliLP3 or a truncated

form of PhLP3 unable to bind CCT (tPhLP3; residues 1-193) (Figure 3-5A and B). For

both actin and tubulin, the three species v~sible on the native gels reprcscnt CCT-

substrate co~nplexes at the top, PFD-substrate cotnplexes in thc middle, and native actin

at the bottom or, in the case of tubulin, either ap-heterodimers or a co~nplex of quasi-

native P-tubulin and cofactor A (Vainberg et al., 1998; Leroux, 2000; Lopez-Fanarraga,

2001) (see Methods). In the presence of excess full-length PhLP3, native actin and

tubulin (lowest band) are produced vely inefficiently. The resulting unfolded nctin and

tubulin proteins seem to be captured by PFD, consistent with a role for PFD as a sink for

unfolded CCT substrates (Figure 3-SA and B) (Siegers et a]., 1999; Vainberg et a].,

1998). The effect of PhLP3 depends on its interaction with CCT. as an excess of tPhLP3

had no effect. similar to buffer-alone controls (Figure 3-SA and B).

Thus, aside from simply inhibiting folding, the presence of PhLP3 forces both

CCT substrates onto PFD. Whether the substrates on PFD in this case represent a pool

that is cycling through the chaperonin without folding productively or whether substrate

turnover at CCT is reduced, leading to an occupied population of chaperonin complexes

to which PFD cannot deliver substrates, is not known. Indeed, a possible mutual

exclusivity behveen PFD and PhLP3 binding to CCT remains to be explored.

3.4.4 PhLP3 inhibits the ATPase activity of'CCT bound to a substrate

Unlike the niodel proposed for PhLP I . the inhibition of actin or tubulin folding by

excess PhLP3 is probably not caused by direct competition with substrate, because

PhLP3 f o r m ternary complexes with CCT and substrate (Figures 3-3 and 3-4). Another

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possible inhibitory n~echanisn~ would be if PhLP3 alters the ability of CCT to hydrolyse

ATP. Figure 3-6A and B, shows the relative ATP hydrolysis activities of folding-

competent rabbit CCT with either denatured actin or tubulin alone, with PhLP3 alone, or

with a substrate and PhLP3. In these assays, PhLP3 alone had no effcct on the basal

ATPase activity of C'C'T (Figure 3-6A). As reported previously (Melki et al., I997), the

addition of either denatured actin or -tubulin significantly increased ATP hydrolysis by

CCT (Figure 3-6A and B). When either denatured actin or P-tubulin and PhLP3 were

both added to CCT, ATP hydrolysis decreased well below the basal ATP hydrolysis

levels of CCT alone (Figure 3-6A and B). As expected, truncated PhLP3 (tPhLP3),

which does not interact with CCT, did not produce an inhibitory effect. The data are

again consistent with PhLP3 acting in a terna~y complex with CCT and either actin or

tubulin rather than competing with them, because the ATPase activities of CCT-substrate

or CCT-PhLP3 differ from that of the CCT-PhLP3-substrate combination.

The substrate-dependent inhibition of chaperonin ATPase activity by PhLP3

provides a plausible explanation as to why actin or tubulin folding by CCT is inhibited by

PhLP3 (Figure 3-5A and B) and hints at a complex allosteric relationship between the

chaperonin, PhLP3, and substrate. It1 vivo, the concentration of PhLP3 is likcly to be

significantly less than that of CCT. However, the amount of PhLP3 may be sufficient to

slow the reaction cycle of a significant proportion of newly formed CCT-actin and CCT-

tubulin con~plexes. PhLP3 may recognize and act on particular CCT-substrate complexes

and modulate, rather than merely inhibit, CCT-mediated folding at physiologically

relevant concentrations.

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Given the prevalence of cofactors that nlodulate the ATPase of other chaperones,

the absence of similar cofactors for CCT has been conspicuous. Many other nucleotide-

dependent chaperones have at least one cofactor protein that intluences ATP hydrolysis;

Hsp70 has Hsp40 (Szabo et al., 1 994); Hsp90 has Aha I, p23, and Cdc37 (Siligardi et al.,

2004; Young and Hartl, 2000); and even the C'C'T hon~ologue GroEL has GroES

(Chandrasekhar ct al., 1986). For CCT, i t will be interesting to establish if other

phosducin-like proteins have similar or differcnt effects on its ATPase activity, and to

determine the significance of HopIp60 as a nucleotide exchange factor for CCT (Gebauer

et al., 1998). Direct effects of PFD on the CCT ATPase have not been tested. but because

substrates stimulate the ATPase and PFD promotes delivery of substrates to CCT, PFD

could have the indirect effect of stimulating the reaction cycle of CCT (Siegers et a].,

1999; Geissler ct al., 1998; Vainberg et al., 1998; Melki et a]., 1997). More importantly,

this function of PFD would antagonize the inhibitory action of PhLP3.

3.4.5 Synthetic interactions of PLPl and prefoldin reveal links to tubulin and actin function in vivo

To understand the relevance of our in viftu data to cytoskeletal function in vivo,

we turned to S. cScwvisi~w. Yeast lacking the PFD subunit PAC10 possess less folded

tubulin overall but more P-tubulin relative to a-tubulin, an imbalance that is not tolerated

by yeast and rcsults in supersensitivity to the niicrotubule-depolymerizing drug benomyl

(Geissler et al., 1998; Alvarez et al., 1998). In this context, the toxic levels of P-tubulin

can be reduced if the folding of P-tubulin into its functional form is inhibited. Thc present

model for yeast PhLP3 (PlpI p) function is that i t promotes the folding of P-tubulin

without influencing tubulin expression (Lacefield and Solomon, 2004). Accordingly,

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PLPl deletion suppresses PLIL'IOA benoniyl sensitivity because less folded P-tubulin is

produced (Lacefield and Solomon, 2004) (Figure 3-7A). We also found that at 20 O C ' , the

plplA mutant was itself more resistant to benomyl than WT (Figure 3-7A), suggesting

that even in WT cells a reduction in tolded P-tubulin also counters benomyl-induced

imbalances in alp-tubulin levels (Lacefield and Solomon, 2003).

On the surface, the apparently positive effects of PLPI deletion on p-tubulin

folding may seem to be in conflict with our in vi t m data showing that excess PliLP3

inhibits tubulin folding. However, as mentioned previously, the in vivo ratio of PhLP3 to

CCT is likely to be lower such that the modulatory effects of PhLP3 may actually be

helpful. An increase in the t h e CCT and tubulin are associated could allow for more

efficient folding and perhaps require fewer rounds of CCT binding and release. In the

case of the bacterial chaperonin GroEL, substrate proteins that normally required several

rounds of binding and release were shown to reach the native state while trapped in a

mutant chaperonin unable to release polypeptides (Weissman et a]., 1996). Similarly,

PhLP3 may increase substrate retention time, leading to a better folding yield within a

particular C'CT reaction cycle. I t is also possible that a delay helps the quasi-native

tubulin associate with downstream cofactors required for its assen~bly into heterodimers

(Lopez-Fanarraga et al., 200 1 ).

Six proteins (cofactors A-E and CIN4lARL2) work downstream of CC'T and PFD

to promote the formation of alp-tubulin heterodimers (Lopez-Fanarraga et al., 200 1).

Although PAC10 deletion in yeasts lacking any one cofactor leads to lethality or growth

defects, in plplA cells lacking either cofactor A(RBL2). C (CIIV~), D (CIIVI), or E

(PACZ), no synthetic growth defects were observed (Tong et a]., 2004). We surmise that

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because cofactor deletion strains have fewer polymerization-cumpctent tubulin

heterodimers, the deletion of PACIO, which Icads to an excess of quasi-native P-tubulin,

further interferes with niicrotubule assenibly and therefore results in sickness or Icthality.

On the other hand, PLPI deletion does not affect tubulin ratios, and this could be why i t

does not display synthetic interactions with cofactor deletions. Moreover, unlike the case

for ytrcl0A (Figure 3-7A), there was no effect of PLPI deletion on the bcnomyl

supersensitivity of the cofactor deletions (Figure 3-7B). This is consistent with the

different effects of PFD and cofactor deletions on the ratios of a- and P-tubulin; PLPI

deletion corrects thc excess of quasi-native P-tubulin responsible for benomyl sensitivity

in paclOA cells. although i t has no observable effect on cofactor deleted cells, which have

a norn~al ratio of a-to P-tubulin (Abruzzi et al., 2002; Hoyt et al., 1997).

The putative specificity of Plp I p for P-tubulin (Lacefield and Solonion, 2003)

implies that Plpl p acts on particular CCT-substrate coniplexes and/or may affect CCT-

substrate complexes differentially, i .c, positively, negatively, or not at all. In addition to

P-tubulin, i i ~ vitro data suggest that PhLP3 also impacts actin function (Figures 3-3 and

3-5). Furthermore, it is known that PFD assists actin folding and that PFD subunit

deletions are hypersensiti\~e to the drug latn~nculin (Geissler et al., 1998), which

specifically sequesters native actin monomers (Ayscough et al., 1997). Remarkably, we

found that yuc~lOAplplA yeast were resistant to latrunculin relative to ~ L I L - I O A ccIIs

(Figure 3-7C). As a control for specificity, we found that latrunculin did not affect any of

the tubulin cofactor deletions, consistent with literature showing no effect of latrunculin

on microtubule function (Ayscough et al., 1997). Latn~nculin resistance in~plies that more

folded actin is present in prrcIOAplplA cells than in pcrcal0A cells. In fact, rho-phalloidin

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staining of tilan~entous actin (F-actin) revealed lower staining intensity in p~iclOA cells

than in identically treated WT orplplA cells (Figure 3-7D). More importantly, deletion

of PLPI in the same haploid ytrc,lOA strain restored F-actin staining intensity, suggesting

that more F-actin is present in pciclOA plplA cells than in puclOA cells (Figure 3-7D).

These observations are consistent with published data showing that PFD deletions have

an 50% lower yield of actin folding (Siegers et al., 1999) and with our data showing

latrunculin resistance in p~ic l0A ylplA cells relative to ytrcI0A cells.

Aside from benomyl and latrunculin scnsitivity,pnclOA cells are sensitive to high

osmolari ty and low temperatures, likely representing actin and tubulin defects,

respectively (Figure 3-7E and F) (Geissler et al., 1998; Vainberg et al., 1998). Although

in each case plplA cells were comparable with WT, we found that on high osmolarity

media (1.5 M sorbitol (Figure 3-7E) or 1 M NaCI) or on media grown at low

temperatures (20 or 25 "C) puclOAplplA cells grew slower than puclOAcells (Figure 3-

7F). Additional evidence that Plplp and PFD work togcther to promotc cellular viability

was revealed in a large scale synthetic interaction study that showed that cells lacking

PLPI and any of five PFD subunits had a slow growth phenotype (Tong et al., 2004), a

finding we have independently confirn~ed (Appendix 2).

3.4.6 Cellular defects in pacl0A yeast a re enhanced by PLPI deletion

To understand further the relationship between PFD and Plplp, we examined the

known pncIOA cellular defects of increased ccll size, aberrant chromosome segregation,

and disorganization of cortical actin patches (Geissler et al., 1998; Vainberg et al., 1998)

in plplAand paclOA plpIA mutants. To determine the effects of PLPI deletion on the

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~wclOA phenotypes, we used differential interference contrast or fluorescence

niicroscopy of yeast stained with DAPI (to stain nuclei) and rho-phalloidin (to stain F-

actin). Although i n each case the plplAcells appeared WT, the deletion of PLPl in a

ptrcl0A strain led to exacerbation of nearly all phenotypes examined.

As expected (Geissler et a].. 1998), when visualized by differential interference

contrast, yuclOA cells had a larger diameter perpendicular to the mother-daughter axis

cornpared with WT cells (4.62 versus 4.40 pni WT, t = 4 .467 , p < 0.001). TheptrcIOA

plylA cells were even larger on avesage than pat-lOA cells (5.05 p n , t = -5.914, p <

0.001). A significant proportion ofynclOA ply1A cells also exhibited a thickening of the

bud neck junction between mother and daughter cells, a possible indication of actin

cytoskeleton defects (Table 3-2). DAPI staining revealed that pucl0A cells often failed to

segregate chroniosonies properly to the large bud (37% defect) as compared with WT

cells ( 1 5%). The pcrclOAplplA cells exhibited even more penetrant defects (53%).

Consistent with these observations, sonie p~rcI0A cells were anucleate and unbudded

(6%) or multinucleate and large budded (4.5%), two phenotypes not observed in WT

cells but exacerbated in ytrc-lOA ylylA cells (19% anucleate; 14.5% multinucleate)

(Figure 3-8A and Table 3-2). A significant increase in the proportion of large budded

cells was observed for ycrclOA yeast (43%) compared with WT yeast (35%), and a more

severe defect was observed in yuclOAplylA yeast (53.5%). The increase in large budded

cells suggests a G2lM cell cycle delay, potentially indicating a checkpoint response to the

defects in DNA segregation (Lew and Burke, 2003). On the whole, these phenotypes are

consistent with previously described microtubule and actin defects for PFD subunit

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deletions (Geissler ct al., 1998; Vainberg et a]., 1998) and with the idea that Plp l p works

in coordination with PFD to promote cytoskeletal function.

Rho-phalloidin staining revealed actin patch polarization dcfects in p c ~ l O A cells

that were exacerbated by PLPI deletion. In 20% of large budded p c l O A cells, cortical

actin was depolarized, whereas actm in 47%) o fy~ i~~1OAplp lA cells and only 3.6% of WT

cells was depolarized. As published for another PFD subunit deletion (Vainberg et al..

1998), i n p c l O A yeast there was an abundance of unbudded cells with diffuse cortical

actin patches (40%) compared with WT (8.5%) (Figure 3-8B). Unbuclded yaclOAplylA

cells were slightly more affected (57%). Actin polarization in s~nall budded yrrclOA cells

was comparable with WT; however, the deletion of PLPI in p~rclOA haploid cells led to

significant depolarization (14% versus 3% ofpuclOA cells) (Figure 3-8B and Table 3-

2). Finally, rho-phalloidin-stained actin filaments were visible in only 66% ofpc1OA

cells with daughter buds compared with 85% of WT budded cells. This phenotype was

not significantly altered inpnclOAplplA cells as 62% contained actin filaments,

implying that actin cable formation is not f~irtlier impaired by PLPI deletion in

ycrclOA cells (Table 3-2).

Although i t has been suggested otherwise (Lacefield and Solomon, 2003), in our

hands the function of PLPI appears to extend beyond tubulin to actin. Other genes whose

mutations lead to latrunculin resistance promote actin instability in WT yeast (Ayscough

et a]., 1997); thus, Plp I p may regulate the levels of folded monomeric actin and in this

way regulate filament stability. Plpl p also promotes microtubule instability, in this case

by facilitating toxic quasi-native P-tubulin production. Quasi-native tubulin is toxic

because i t interferes with normal microtubule function. Although PFD and Plpl p have

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opposing effects on niicrofilanient and microtubule stability, as rcvealed by drug

sensitivity, they clearly work together to promote cellular viability (Figures 3-7 and 3-8)

(Tong et al., 2004). Compared with PACIO or PLPI deletions. p ~ 1 c I 0 A p l p I A cells are

slow growing and exhibit more significant tubulin defects (cold sensitivity and abnonnal

DNA segregation) and actin defects (osmosensitivity and incorrect actin-

organizationlpolarization) (Figure 3-8). y l p l A cells are resistant to benomyl because a

proportion of both tubulins aggregate (potentially more P-tubulin), rcscuing strains that

have an imbalance, such as ptrcIOA, from the toxicity of excess P-tubulin (Lacefield and

Solomon, 2003). However, tubulin aggregation leads to fewer polymerization-competent

heterodimers; the net result is that in strains where tubulin is already compro~niscd,

~nicrotubule function is further reduced and gives rise to aggravated cold sensitivity and

chromoson~e segregation defects (Figures 3-7 and 3-8). The casc for actin may be

somewhat different; ~ L I c I O A cells have 50% the WT yield of folded actin (Siegers et a].,

1999), leading to latrunculin sensitivity and other phenotypes (Cieissler ct al., 1998;

Vainberg et a]., 1998). PLPI deletion in a pc . IOA strain restores latrunculin resistance

and the amount of F-actin visible by rho-phalloidin staining to lcvels comparable with

WT (Figure 3-7C and D). This suggests that Plp Ip inhibits actin folding in a p~lc.lOA

background, a notion consistent with our in vitro data (Figure 3-5). However, pucIOA

p l p l A cells are more sensitive to high osmolarity, are larger, and exhibit a greater number

of cells with disorganized cortical actin than p~rc10A cells.

I t is unclear why PLPI deletion ameliorates certain ycrclOA actin phenotypes and

exacerbates others. Although the in vitrw data and the close genetic relationship with PFD

suggest a direct effect on actin folding by Plplp, there may also be indirect effects that

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could explain why PLPI deletion gives rise to disparate actin phenotypes (Figures 3-7

and 3-8). Plpl p and/or PFD may affect other CCT substrates independent of actin

monomer production, and the effects on the substrate(s) could lead to enhanced dcfects i n

actin filament assembly or organization. The actin-related proteins (ARPs) represent

candidates responsible for such indirect effects. ARPs regulate actin filament nucleation

and organization, and some ARPs are known CCT and prefoldin substrates (Rommelaere

et a]., 2001). Alternatively, the speed and timing of actin production niay be dysregulated

in the absence of PFD and Plpl p, leading to defects in filament organization and

function. Further experiments are required to understand precisely the effects of PFD and

Plplp action at CCT on downstream actin folding and organization. Moreover, the impact

that the other phosducin honiologue in yeast, Plp2, has on CCT. actin, and tubulin is

unclear. A complete nlodel of how PhLPs regulate C'CT in yeast will require a better

understanding of Plp2 function. Although Plp2p may function somewhat like Plplp, i t

seems that the f~inctions will differ to some degree because P L P is essential and PLPI is

not. and PLPI overexpression cannot complement the loss of P L P (Flanary et a]., 2000).

3.5 Conclusion

In general, little is known about the biological functions of phosducin-like

proteins outside of their role in G-protein signaling (Yoshida et a]., 1994; Lukov et a].,

2005; Blaauw et al., 2003; Flanary et a]., 2000). The following two studies have provided

clues to a new cellular role for PhLP3: one in Strccl7trt-ot~z~vces cet-cvi.\itre, where Plp 1 p

was iniplicated in P-tubulin folding (Lacefield and Solomon, 2003); and the other in C.

C I ~ ~ I I S , showing PhLP3 is important for correct niicrotubule architecture (Ogawa et al.,

2004). Based on our findings, we propose that PhLP3 acts as a novel cofactor of CCT

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that niodulates its ability to fold substrate proteins. We show that PhLP3 binds to CCT,

spanning the central cavity perhaps at thc level of the apical domains above the substrate

cavity, as does PhLP I (Martin-benito et a]., 2004). We also show that PhLP3 can form

ternary complexes with CCT and actin or tubulin and affect the folding of the two

substrates in vitro. Finally, we show that PhLP3 slows the rate of ATP hydrolysis by

CCT when in the presence of an unfolded substrate protein. The mechanism of PhLP3

function seenis to be different from that proposed for PhLP 1 (Mclaughlin et a]., 2002). I t

is possible that the t ~ l o proteins work differently at the level of CCT; alternatively, they

could have different effects on different substrates. Indeed, the competition of PhLP 1 for

binding to CCT was shown with G,u and not with actin or tubulin (Mclaughlin et al.,

2002). Further experiments will be required to fully understand the similarities and

differences between PhLP isofornis.

Finally, we detnonstrate that PhLP3 iri vivo, in con-junction with PFD, is required

not only for tubulin biogenesis but also to regulate the formation of a functional actin

cytoskeleton. Earlier data have suggested that Plp Ip functions upstream of the tubulin

folding cofactors to modulate the folding of P-tubulin in vivo, and our findings are

consistent with this notion (Lacefield and Solomon, 2003). Together our data point to an

antagonistic relationship between PFD and PhLP3 at C'CT that when balanced leads to

optinlal cytoskeletal protein biogenesis and function. Overall, these studies establish

PhLP3 as a novel CCT cofactor, suggest a mechanism for the regulation of CCT function

by phosducin-like proteins, and reveal a new modulator of cytoskeletal protein

biogenesis.

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Figures

Figure 3-1 PhLP3 interacts with CCT i t 2 vivo

(A) Native gel analysis of ir7 vilro translated "S-labelled actin and PhLP3. CCT-actin

binary con~plexes and folded actin are indicated. (B) Immunoprecipitation (IP) using anti

n i y antibody of HEK cell lysates expressing r~1jx--PhLP3 or pCMV (empty vector).

Purified C'CT (right lane) or imrnunoprecipitated CCT was detected by Western Blot

(WB) using an anti-TCPI antibody. ( C ) Coomassie stained gel showing GST-pull-down

(IP) of the CCT complex by GST-PhLP3 but not GST alone (two right lanes). A sample

of the reaction mixture prior to the pull-down (On) was included to show that equal

amounts of CCT were used for each pull-down (two left lanes). (D) WB showing a GST-

pull-down of CCT incubated with GST, GST-PhLP3, or GST-PhLP3 and excess nativc

untagged PhLP3.

A

CCT-Actin -

Native - Actin

Native PAGE

Whole Cell Extracts anti-myc IP Purified CCT

pCMV myc-PhLP3 pCMV myc-PhLP3

WB: anti-TCPI

WB. anti-TCPI

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(E) Size exclusion chromatography of HEK ccll lysate expressing n1,vc-PhLP3. Fractions

were immunoblotted for CCT (TCPI) and t~1j .c . . Whole cell extract (WCE) represent

loading controls for the western blots (F) GST-fused PhLP3 truncations (schematics

shown on the left) were used in pull-down assays with CCT to determine binding regions

(right).

CCT

I

Helical Thioredoxin Acidic N-terminus Domain C-terminus Residues

I , V V &&V

1-218 WB: anti-TCP1

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Figure 3-2 PhLP3 does not form binary complexes with unfolded actin, tubulin or different forms of native tubulin

(A) Denatured '%labelled actin or tubulin (D*actin or D*P-tubulin, as shown) were

diluted 1 : 100 into buffer containing no addition (-), PhL133 ( 1.5 pM) (+), C'CT (0.15 pM)

or both and analyzed on a native gel. CCT-substrate complexes are indicated with an

arrow. The position where native PhLP3 normally runs is also indicated. (B)

Sedimentation coefficients for PhLP3, tubulin heterodimers or a mixture of the two.

Buffer CCT Buffer CCT -- -- - + + - + - + PhLP3

PhLP3 -P 1 c complex 1

Protein Sedimentation Coefficients

Tubulin Heterodimer 5.3 S -- -- - -- -

Tubulin Heterodimer 2.7 S (52%). + PhLP3 5.3 S (41%)

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(C. D) Microtubules assembled in v i k o were pelleted at high speed with PhLP3 (C') or

purified microtubule-associated proteins (MAPS) (D). The supernatant (S) and pellet (P)

fractions were separated by SDS-gel electrophoresis and coornassie stained. In (C), buffer

(S) corresponds to PhLP3 without microtubules added. The positions of a- and P-tubulin

(Tubulin), PhLP3 or MAPS are indicated. [NOTE: F.E.K. and R. M. completed the

experiments in B, C, and Dj

C PhLP3 - - + + Buffer

P S P S S ;;

+ MAPS kDa P s

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Figure 3-3 PhLP3 interacts with actin and tubulins in a ternary complex with CC'?'

(A) Actin. a- and P-tubulin were translated in reticulocyte lysate supplemented with CiST

or GST-PhLP3, precipitated, and analyzed by SDS-PAGE and autoradiography. The

lower pancl are control reactions showing total translated products. (B) P-tubulin was

translated in an E. c d i lysate GST or GST-PhLP3 with or without CCT and analyzed as

in (A).

Reticulocyte lysate

GST pull-down 1 - -I Translation

Actin P-tubulin rx-tubulin

E. coli lysate

GST pull-down

Translation l - - - w t - u

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Figurc 3-4 Electron microscopy of PhLP3-CCT and PhLP3-CCT-tubulin cornplexcs

(A-E) Two-dimensional averages of negative-stained electron microscopy images of

Apo-CCT (453 particles analyzed) (A), CCT-PhLP3 (847 particles) (B), C'CT-PhLPl

(Martin-benito et a]., 2004) (C), C'CT-tubulin (both a and P isoforms; 570 particles) (D)

and CCT-PhLP3-tubulin complexes (530 particles) (E). Scale bar indicates I00 A.

[NOTE: J. C. and J.M.V. completed these experiments]

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Figure 3-5 PhLP3 affects the folding of nascent actin and tubulin iiz vitro

Actin (A) or P-tubulin (B) was translated i11 the presence of excess PhLP3 (WT) or

tPhLP3 (t; residues 1 - 193) and time points (as indicated) were analyzcd on native gels

(lower panel). Thc upper panel shows SDS-PAGE analyzed translation products over

time for the reactions. Tubulin* refers to either ap-heterodirners of C'ofactor A-P-tubulin

con~plexes (refer to Methods).

I - -.- - - Translation

1 1 ----- 5 10 20 30 40 Tirne (minutes)

Actin folding

W

I .- a -- - - - I Translation

+ Folded Actin

20 30 40 50 60 Tirne (minutes) p-tubulin folding

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Figure 3-6 PhLP3 inhibits the ATPase activity of CCT in the presence of an actin or tubulin substrate

(A) ATP hydrolysis by CCT was nieasurcd over 60 minutes for CCT alone (+), CCT t

PhLP3 (n), CCT + actin done (x), C:CT + actin and tPhLP3 (0) or CCT + actin and fidl-

length PhLP3 (A). B-tubulin was also tested (B) with CCT alone (x), CCT + tPhLP3 (a),

and CCT + fill1 length PliLP3 (A).[NOTE: F.E.K. and R.M. completed these

experiments]

Time (minutes)

CCT + Actin

CCT + Actin + tPhLP3

CCT + PhLP3

CCT alone

CCT + Actin + PhLP3 1

CCT + P-tubulin

CCT + [j-tubulin + tPhLP3

CCT alone

CCT + P-tubulin + PhLP3

Time (minutes)

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Figure 3-7 Synthetic effects of PLPZ deletion in paclOA yeast

(A and B) Saturated cultures were serially diluted tenfold, plated on rich medium

containing benomyl (as indicated) and photographed after 3 days of growth at 20 "C or

30 "C (as indicated). Cclls grown on control plates lacking benomyl grew identically (sec

Figure 3-7C).

Benomyl 20l~g/mL 5!1g/rnL 12 5 !~g/rnL 30'C 20•‹C

Benomyl 20ug/mL 5gglmL 2.5pg/mL2.5pg/rnL 30•‹C

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(C) Latrunculin B (LatB) sensitivity of wild-type ( WT), plpl A, ptrc.lOA and puc-lOA

plplA strains were determined as described (Ayscough et al., 1997) and are shown

norn~alized to 1 .OO for wt cells. (D) Images of identically treated, equally exposed raw

images of each strain; right panels show the same images with equally increased contrasts

to illustrate the differences in staining intensity. ( E and F) Yeast werc treated as in (A).

plated on rich media with 1.5 M sorbitol (E) or no addition (F) and grown at the indicated

temperature for 3 days.

25'C 30•‹C

YPD + 1.5M Sorbitol

D l ncreased Raw Images

Rhodamine Phalloidin

YPD

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Figure 3-8 pacI0A cellular defects are enhanced by PLPI deletion

(A and B) Cells in mid-log phase were fixed and stained with DAPI (A) or rho-plialloidin

(B). Arrows indicated multinucleate or anucleate cells in (A) and aberrant actin patch

polarization in (B). Actin cables are denoted by a '*'.

A wt p ~ p ~ a

Rhodamine Phallo~din

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3.7 Tables

Table 3-1 Yeast strains used in this chapter

Strain Name Cienotype MLY I10 MLY 1 1 1 MLY 1 12 MLY l I 3 MLY 114 MLY 1 I5 MLY116 M L Y l l 7 MLY I18

MLY I19

MLY 120

MLY 121

MLY 122

Mat a 111~13-52, 1~112-3, -1 12, l1is3, 1~115 . Aplpl: :KanMX4 Mat a ur-03-52, le112-3, -112, his3, tnctI5, Apuc.lO::KanMX4 Mat a ~11.~13-52, 1~112-3, -112, hi.53, rnctl5, Acinl : :KanMX4 Mat a 111-u3-52, lcw2-3, -112, hi.r3, nletI5, Ac.i112::KanMX4 Mat a ZII-LI~-52, lm2-3 , -112, his3, me t l5 , Arhld::KanMX4 Mat a z1r~13-52, 1~112-3, -1 12, his3, nlctI5, Ap~1c,2::KanMX4 Mat a zrl.u3-52, 1~112-3, -1 12, his3, l~~.s2-XOl, Ap~1czlO::KanMX4 Mat a ~ 1 . ~ 3 - 5 2 , lezr2-3, - I 12, his3, ~lx2-h '0/ , Aplpl : :KanMX4 Mat a 11r-u3-52, lez12-3, -112, his.3, met/-5, lp2-$01. Aplpl : :KanMX4, Ap~lclO::KanMX4 Mat a zlr-u3-52, lezr2-3, -1 12, his3A1, Aplp/::KanMX4, Ac,inl::KanMX4 Mat a zrr-~13-52, le112-3, -1 12, hi.s3Al, metl5. Aplpl : :KanMX4. Ac,inZ::KanMX4 Mat a ul-u3-52, lcv12-3. -1 12, his3A1, nrc.115, Aplpl : : KanMX4, Apuc2::KanMX4 Mat a z1ru3-52, lezr2-3, -1 12, his3A1, lys2-801, Aplpl : :KanMX4, Al+hl2::KanMX4

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Table 3-2 Statistics of cellular yeast phenotypes

p-\due relati\le to p-value relali\e lo

Phenotype Strain n = I'ercentage wild-type p ~ 1 ~ ~ 1 O A p I p l A

\Y I 126 1 5. 0O'X N/A N/A Cllro~nosome

y l p l A segregation defect in large-budded W ' I o A

Multinucleate p u c I OA 309 4.50% p-;0.0 1 p 4 . 0 0 I large-budded cells p u ~ l 0 A p l p l A 228 14.50% p<0.00 1 NiA

u1 t 105 0.00% N/A N/A

plp I A 96 0.00% N/A N/A

Anucleale IXICIOA 100 6.00% p4 .025 p 4 . 0 1

unbudcled cells /XI(, IOApIp / A 156 19.20% pi 0.00 1 N/A

\vt 45 1 35.30% N/A N/A

p111 1 A Proportion of lotal

384 35.90% N/A N /A

cells that are pat, 1 OA 840 42.60% p 0 . 0 2 5 p.4M.N I

large-budded /)ma I OAplp I A 482 53.50% p<0.00 1 N /A

\v t 141 X5.200/;, N /A N/A

p/p I A 165 88.50% N /A N/A

Actin filaments in I ) ~ ~ ~ ~ I ~ ) ~ 125 65.60% p 4 . 0 0 1 N/A

budded cells ~ L I L , I OAplp I A 3 62 62.40% p 4 . 0 0 1 N/A

\\I t 114 8.50% N /A N /A

/I/]> I A Aberrant actin pol:iri~ation, p c I OA

pip I A Aberrant actin polarization, yuc 1 OA small-bucltled p u c l OAplp 1 A 124 14.30% pi0.00 1 NiA

wt 8 8 3.600/;, N/ A N/A

pip / A Aberrant actin polarization. pcrc 1 OA large-budded p c 1 ~ * 1 OApIp I A 97 46.70% p-4.001 N/A

MI t 174 1.10% N/A N /A

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Stafford, W. F., 111 ( 1992). Boundary analysis in sedimentation transport experiments: a procedure for obtaining sedimentation coefficient distributions using the time derivative of the concentration profile. Anal Biochem 203, 295-301.

Stirling, P. C., Lundin, V. F., and Leroux, M. R. (2003). Getting a grip on non-native proteins. EMBO Rep 4, 565-570.

Stirling, P.C., Cuellar, J., Alfaro. G.A., El Khadali, F., Beh, C.T., Valpuesta, J.M., Melki, R., and Leroux, M.R. (2006). PhLP3 niodulates CC'T-mediated actin and tubulin folding via ternary complexes with substrates. J Biol Chem 28 1, 70 12-702 1.

Szabo, A., Langer, T., Schroder, H., Flanagan, J., Bukau, B., and Hartl, F. U. (1994). The ATP hydrolysis-dependent reaction cycle of the Escherichitr coli Hsp70 system DnaK, DnaJ, and GrpE. Proc Natl Acad Sci USA 9 1, 10345-10349.

Thulasiranian, V., Yang, C. F., and Frydman, J. (1999). In vivo ncwly translated polypeptides are sequestered in a protccted folding environment. EMBO J. 18, 85- 95.

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Tong, A. H., Lesagc, G., Bader, G. D.. Ding, Id., Xu, H., Xin, X., Y o ~ ~ n g , J.. Bei-riz. G. F., Brost. R. L., Chang, M., et al. (2004). Cilobal mapping of the yeast genctic interaction network. Science 303, 808-8 13.

Vainberg, I . E., Lewis, S. A., Rommelaere, H., Ampe, C., Vandekerckhove, J . , Klein, H. L., and Cowan. N. J . ( 1998). Prefoldin, a chaperone that delivers unfolded proteins to cytosolic chaperonin. Cell 93, 863-873.

Weissman, J . S., Rye, H. S., Fenton, W. A., Beechem, J. M., and Horwich, A. L. (1996) . Characterization of the active intermediate of a GroEL-GroES-111ediated protein folding reaction. Cell 84, 48 1-490.

Yoshida, T., Willardson, B. M., Wilkins, J. F., Jensen, G. J., Thornton, B. D., and Bitensky, M. W. ( 1994). The phosphorylation state of phosducin detem~ines its ability to block transducin subunit interactions and inhibit transducin binding to activated rhodopsin. J Biol Chem 269. 24050-24057

Young, J. C., and Hartl, F. U. (2000). Polypeptide release by Hsp90 involves ATP hydrolysis and is enhanced by the co-chaperone p23. EMBO J 19,5930-5940.

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CHAPTER 4 FUNCTIONAL INTERACTION BETWEEN PHOSDUCIN-LIKE PROTEIN 2 AND CYTOSOLIC CHAPERONIN IS ESSENTIAL FOR CYTOSKELETAL PROTEIN FUNCTION AND CELL CYCLE PROGRESSION

Note regarding contributions: The following chapter has been accepted for publication to Molecz~lrr~ Biology o/'

tht. Cdl. The authors of the study are listed below.

Stirling, P.C., Srayko, M., Takhar, K.S., Pozniakovsky, A., Hyman, A.A., and Leroux, M.R. (2007). Functional interaction between phosducin-like protein 2 and cytosolic chaperonin is essential for cytoskeletal protein function and cell cycle progression. Accepted to Moleculw B i o h ~ 7 ) (?/'the Cell. (Copyright, ASCB, 2007)

As the first author I contributed most of the data and wrote nearly all of the article, along with significant editorial guidance from M.R. Leroux. K.S. Takhar performed the high copy suppression screen (Figure 4-9). M. Srayko, A. Pozniakovsky and A.A. Hyman, generated the temperature-sensitive alleles of PLP2, showed co- immunoprecipitation of Plp2p with CCT and performed the FACS analysis on synclvonized plp2-ts cells (Figure 4-1 and 4-9).

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4.1 Abstract

The chaperonin C'CT maintains cellular protein folding homeostasis in the

eukaryotic cytosol by assisting the biogenesis of many proteins, including actins,

tubulins, and regulators of the cell cycle. Here, we demonstrate that the essential and

conscrved eukaryotic phosducin-like protein 2 (PhLP2lPLP2) physically interacts with

CCT and modulates its folding activity. Consistent with this functional interaction.

temperature-sensitive alleles of S. cw-evisi~w PLP2 exhibit cytoskcletal and cell cyclc

defects. We uncovered several high-copy suppressors of the ply2 alleles, all of which are

associated with G 11s cell cycle progression but which do not appreciably affect

cytoskeletal protein function or fully rescue the growth defects. Our data support a model

in which Plp2p modulates the biogenesis of several CCT substrates relating to cell cycle

and cytoskeletal function. which together contribute to the essential function of PLP2.

4.2 Introduction

Phosducin-like proteins (PhLPs) are a conserved family of small thioredoxin-like

proteins that were originally identified as modulators of heterotrimeric Ci-protein

signaling in the retina (Schsoder and Lohse, 1996). Subsequently, they have been shown

to have roles in G-protein signaling in other cell types as well as having G-protein

independent functions (Blauuw et al., 2003; Flanary et al., 2000). One of the other

functions of PhLPs seems to be the regulation of the eukaryotic protein-folding machine

known as CCT (chaperonin Containing Tcpl; also called TRiC for TCP 1 containing

Ring Complex) (Lukov et al., 2005; Lukov et al., 2006; Martin-benito et al., 2004; -

McLaughlin et al., 2002; Stirling et al., 2006).

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Chaperonins arc oligomeric ~no lec~~la r chaperones that bind non-native proteins

and facilitate their transition to the native state (Hart1 and Haycr-Hartl, 2002). These

barrel-shaped molecular machines undergo large conformational changes to encapsulate

and release bound substrate proteins during thcir folding cycle (Spiess et al., 2004). In the

eukaryotic cytosol, the chaperonin CCT ensures the correct folding and assen~bly of a

wide variety of proteins. The best-characterized substrates of CCT are actins and

tubulins, although scveral recent studies have extended the number of known CC'T

substrates (Camasses et al., 2003; Siegers et al., 2003; Spiess et a]., 2004; Thulasiranian

et al., 1999). CCT cooperates with another chaperone called prefoldin (PFD) in the

folding of actins and tubulins (Geissler et al., 1998; Vainberg et al., 1998). PFD uses six

long coiled-coil 'tentacles' to stabilize substrate proteins at the opening of its jelly-fish

shaped cavity (Lundin et a]., 2004; Siegert et a]., 2000; Chapter 2). Together, PFD and

CCT compose a folding pathway for cytoskelctal proteins which, along with PhLPs,

control the folding of actin and tubulin (Lacefield and Solomon, 2003; Siegers et al.,

1999; Stirling et al., 2006).

PhLPs can be subdivided into three ho~nologous families, called PhLPI, PhLP2

and PhLP3, that share an N-terniinal helical domain, a central thioredoxin-like fold and a

charged C'-terminal extension (Blaauw et al., 2003). PhLPl proteins are the best

cl~aracterized and function both in G-protein signaling and the regulation of CCT (Lukov

et al., 2005; Lukov et al., 2006; Martin-benito et al., 2004; McLaughlin et al., 2002;

Schroder and Lohse, 1996). PhLP3 proteins act as modulators of C'CT and have a role in

actin and tubulin biogenesis in yeast and microhlbule function in C. e/egm~.s (Lacefield

and Solomon, 2003; Ogawa et al., 2004; Stirling et al., 2006). PhLP2 proteins arc the

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least characterized PhLP isofor~ns but are essential in Dic.l_)~osteli~~m c/'i.sc.oidcwn~ and

S~~c*c~hu t -o r?~~~c~ . \ . . w w i s i ~ ~ e (Blaauw et a]., 2003; Flanary et a]., 2000). PhLP2 may act as a

regulator of apoptosis in nian~malian cells, but the significance and mechanism of this

function is unclear (Wilkinson et a]., 2004).

Yeast possess honiologues of PhLP3 and PliLP2, which are encoded by PLPI and

PLP.2, respectively (Blaauw et a]., 2003). PLPI/PIILP.? has been implicated in CC'T-

mediated folding of actin and tubulin and is thought to work at the level of CCT to

regulate the ATP-hydrolysis dependent turnover of CCT-substrate complexes (Lacefield

and Solomon, 2003; Stirling et a]., 2006; Chapter 3). Although PLPI plays an important

function in regulating actin and tubulin folding, plplA cells have no apparent growth

defects. Indeed, most plp lA phenotypes are only detected in strains that also lack a

functional PFD complex (Lacefield and Solomon, 2003; StirIing et a]., 2006).

Conversely, PLPZ is essential for growth but its function is not well understood (Flanary

ct a]., 2000; Lopez et al., 2003).

To elucidate the essential function of I'LP2, we undertook studies of yeast and

human PhLP2 homologues. Yeast Plp2p and the homologous human PhLP2A bind to

CCT as suggested by proteonie-wide studies in yeast (Gavin et a]., 2006). In this regard,

we show that ii? v im) human PhLP2A inhibits actin folding and forms ternary complexes

with CCT and actin. We also show in yeast that teniperah~re sensitive (IS) alleles of PLP2

are defective in CCT regulated processes such as actin and tubulin function, and cell

cycle progression, plp2-IS alleles do not however exhibit altered sensitivity or resistance

to a-factor, supporting the notion that regulating G-protein signaling is not part of the

essential function of Plp2p, as night be thc case for certain nianimalian phosducins

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(Schroder and Lohse, 1996). Finally, we idenlify high-copy suppressors ofplp2-ts alleles

that indicate an essential function for P L P during G 11s-phase cell cyclc progression.

Our data support a model in which Plp2p modulates the biogenesis of sevcral C'CT

substrates, which together contribute to tlie essential function of PLP2.

4.3 Methods

4.3.1 Purification of' GST-PhLP2A and CCT

Human PhLP2A (Accession: AF 1 1 05 l I) and PliLP3 (Accession: NM - 005783)

were cloned into tlie GST fusion vector pGEX-6p, and expressed in BL21 [DE3]pLysS as

described (Stirling et al., 2006). GST-fusion proteins were purifled with glutathione-

sepharose 4B as per manufacturer's instructions (Amcrsham). GST alone, used as a

control, was expressed and purified exactly as for GST-PIiLP2A. CCT was purified from

rabbit reticulocyte lysate as described (Gao et al., 1 992; Melki et al., 1 997).

4.3.2 In vitro translation and folding assays

Actin and tubulin were translated in tlie T7 quick coupled translation reaction

(Promega) according to manufacturer's instructions. Actin was also translated in E. coli

lysate using the EcoPro system (Novagen) according to manufacturer's instl-uctions.

Recombinant GST or GST-PhLP2A was added at approximately 100 times excess to

endogenous levels of CCT prior to translation (Cowan, 1998). At various time points

aliquots of the reaction were frozen in native gel riming buffer then thawed on ice and

analyzed by native gel electophorcsis (Leroux, 2000). GST-pulldowns were perfornicd

with glutathione sepharose 4B beads according to ~iianufacturer's instructions

(Aniersham).

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4.3.3 Yeast strains, media and growth assays

For yeast strains and plasmids see Tables 4-1 and 4-2. Yeast werc grown on

YEPD, or synthetic complete mcdias as required (Adams et al, 1997). 5'-Fluoroorotic

acid platcs were also made as described (Ada~ns et a]., 1997). For temperature-sensitive

growth assays, log-phase cells were serially diluted by 10-fold and spotted on thc

appropriate media and cultured at the temperatures indicated. To assess the reversibility

of the ts-a1 leks, the indicated strains were cultured at the restrictive temperature (37•‹C)

for the times shown and plating efficiency was assessed directly under the

microdissection nlicroscope as described (Amberg et a]., 2005).

4.3.4 Plp2p, CCT co-immunoprecipitation

For im~nunoprecipitatio~~-i~nn~i~~~oblotting experiments, a strain bearing integrated

twit -tagged Plp2 and HA-tagged CC'T2 (AHY 994), or control strains lack~ng the tagged

Plp2, or both tagged proteins, were grown to mid-log phase. Extracts were prepared as

described (Zachariac ct a]., 1998) from 2x 10" cclls in 0.4 mL of buffer B70 (number

~ndicates mill~molar potasslum acetate). Cleared extracts (0.35 mL or 4 mg) were

incubated for 60 min with antibodies, which were captured with 40 pI of protein A-

Sepharose for 60 mi11 as previously descr~bed (Camasses et a]., 2003). Beads wcre

washed with the buffers B70 plus BSA (I mg/mL), B70, B 150, B200, and B70 followed

by ~mmunoblot analysis.

4.3.5 Drug and mating factor sensitivity assays

Latrunculin B (LatB; Sigma) sensitivity assays were performed as described

(Asycough et al., 1997). Benomyl or mating fiictor sensitivity assays were perfonned

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essentially as described for LatB. Brietly. sterilc filter papcr disks werc soaked with the

drug or pheromone at the concentration indicatcd and placcd on a soft agar overlay

containing a particular yeast strain. The radius of clearance around the disk was measured

aftcr two days of growth at 30•‹C. Congenic control strains carrying wild-type copies of

thc genes under examination wcre included for each experiment.

4.3.6 Generation of temperature-sensitive alleles of PLPZ

A ply2 null strain was made by HIS3 gene insertion into the PLP2 coding region.

To generate the ylp2-ts mutants, the PLPZ gene (including 1 kb of the upstream promoter

and 120 bp after the stop codon) was cloned into the pRS405 Leu vector. ply2 mutations

were generated within the coding region by error-prone PCR [0.3 niM MnC12,0.5 mM

dCTP, 0.5 mM dTTP, 0.1 niM dGTP, 0.1 mM dATP, wild-type Taq polymerase for 28

cycles] and the wild-type PLP2 ORF was replaced via gap-repair. Mutagenized plasmids

able to rescue the p1p2 11~111 strain were then screened for temperature sensitivity.

Plasniids from temperature sensitive transformants were sequenced, retransformed and

integrated into AHY955 with a LEU2 marker before counterselection of the URA3-

marked PLPZ plasniid with S'fluoroorotic acid (Table 4-1).

4.3.7 Microscopy

Yeast strains were grown to mid-log phase in YPD or SC media before direct

imaging or fixing and staining. Tetramethylrhodamine isothiocyanate-conjugated

phalloidin (Sigma) and DAPl were used, according to manufacturers instructions, to stain

foniialdehyde fixed yeast cells as described (Adams et al, 1997). C'alcofluor white

staining was carried out as in Amberg et al. (2005). Actin immunofluorescence was

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performed as described by A d a m et al. ( 1 997). Bricfly, formaldehyde fixed yeast cclls

were digested with Zymolyasc 20T, bound to Teflon masked slides and fixed

successively in methanol and acetone prior to actin antibody staining. For cell sizing, at

least one hundred cell diameters were measurcd in Openlab 5.0.2 (lmprovision) for each

strain perpendicular to the mothcr-bud axis at the largest point. Cellular defects were

statistically validated using an independent variable t-test or Chi square analysis, as

appropriate.

4.3.8 High-copy suppression screen

A Y Ep24-based library of URAS-marked plasmids containing -10kb fragments of

S. cet-evisiue genomic DNA (Carlson and Botstein, 1982) was transformed into plp2-1

cells. Half of the cells were plated at a permissive temperature (29•‹C) to calculate

transformation efficiency and half were plated at the non-permissive temperature (37•‹C)

to identify suppressing clones. Colonies which grew at 3 i • ‹C after 48 hours were regrown

and the plasmid DNA was isolated and screened by PCR to determine whether it

contained a copy of PLP2. Plasmid inserts lacking PLP2 were sequenced at both ends

and genes therein were identified using the yeast genome database

(www.yeastgenome.org). Plasmids bearing some of the individual genes identified were a

kind gift from Dr. .loaquin Arifio (Mufioz et a]., 2003).

4.3.9 Cell synchronization

For FACS analysis, cells were synchronized as small G 1 cells by centrifugal

elutriation as described by Schwob and Nasmyth (1993). After elutriation, san~ples were

incubated at 37•‹C and, at 15 minute intervals, stained for DNA content with propidium

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iodide. All samples were analyzed on a Bccton Dickinson FACScan (San Josc, CA) as

described (Epstein and Cross, 1992). For rebudding analyses, cells were synchronized

with a-factor according to the low pH method (Amberg et a]., 2005). Synchro~iizcd cells

were shifted to 3 7 T for I hour prior to releasc into 37OC media. Time points were taken

after release and the budding index of the culture scored.

4.4 Results and discussion

4.4.1 Plp2p is an essential CCT-binding protein

Previously we found that Plp lp, the yeast PhLP3 honiologue, interacts with CCT

and cooperates with prefoldin (PFD) to modulate actin and tubulin function in vivo

(Stirling et al., 2006). We initially hypothesized that the only other yeast phosducin-like

protein, Plp2p, might function similarly to modulate CCT activity. Howevcr, several lines

of evidence suggest that the Plpl and Plp2 proteins are unlikely to have identical

functions in vivo. Yeast lacking PLPl are viable whereas yeast lacking PhLP2 (PLP2) do

not survive. In addition, overexpression of PLPl cannot complement the deletion of

PLPb (Flanary et al., 2000).

To determine whether S, ce~wi . r i~~c . Plp2p influences CCT-mediated protein

folding, we first tested if Plp2p interacts with CCT in vivo. When ni,vc-tagged Plp2p was

co-expressed with HA-tagged CCT. CCT co-precipitated with lnyc-Plp2p using an anti-

~ I J K * antibody, indicating a potentially direct physical interaction between the two proteins

(Figure 4-I A). This result is consistent W I th previous proteome-scale TAP-tagging

studies that also identified an interaction between CCT subunits and Plp2p (Ciavin et al..

2006). To assess whether this interaction is conserved in mammalian cells, we purified a

GST-tagged human PhLP2A fusion protein for use in in vitro experiments. Similar to

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PhLP3, purified CCT co-precipitated with the GST-fused PhLP2A, but not GST alone,

indicating that PhLP2A and C'C'T form a coniplex (Figure 4-1B; Stirling ct a]., 2006).

Given that both native PhLP3 and PhLPI form con~plexes with CCT (Stirling et a].,

2006; McLaughlin et al., 2002), our results with Plp2p/PhLP2A now confirm that all

phosducin-like proteins, but not phosducin itself (Martin-benito et al., 2004; McLaughlin

et a]., 2002), intcract with CCT, not as substrates but rather as native binding partners.

There has been some dispute as to whether PLP2 is truly essential for growth or is

instead required only for spore germination (Flanary et a]., 2000; lope^ et a]., 2003).

Flanary et al. (2000) showed that no viable pIp2A spores were isolated from heterozygous

diploids (PLP2/pIp2A) that were sporulated and dissected, a finding we independently

confirmed. Lopez et a]., (2003) showed that a complementing inainmalian PhLP2 bearing

plasmid rescued the lethality ofplp2A cells but. after long term culturing in non-selective

medium, plp2A cells do not retain the plasmid, suggesting to the authors that the gene is

not essential. Unfortunately, this experinicnt allows for the accumulation of secondary

suppressors and is not conclusive. To clarify this discrepancy, we generated a haploid

yeast strain lacking the chromosonial copy of PLP2 and balanced with a URA3-marked

plasmid containing PLP2. When cultured on iiiediuni containing the drug S'fluoroorotic

acid. which selects against cells carrying a functional copy of the URA3 gene, no growth

was obscrved (Figure 4-1C). While i t is still possible that PLP2 is essential in only somc

strain backgrounds, this experiment denlonstrates that some yeast cannot survive without

PLP2.

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4.4.2 Generation of PLPZ temperature-sensitive alleles

In order to study the effects of P L P loss-of-function we generated temperature-

sensitive allelcs of I'LP2 by error-prone PCR mutagenesis of the wild-typc gene. Two of

the alleles identified werc chosen for further characterization, namely pIp2-I, the most

severe allelc. and pl/?2-2, a less severe allele (Figure 4-2A). The pIp2-l mutant cxhibits

growth defccts at 30•‹C, while plp2-2 cells are only slightly impaired for growth at 34•‹C'.

Both mutant alleles cause lethality at 37OC. The two alleles contain multiple sequence

changes (Figure 4-2B), but interestingly, both are mutated at Q91, which aligns to the

region at the C-terminus of helix 3 between the N-terminal and the thioredoxin domains

(Blaauw et al., 2003; Gaudet et al., 1996). To detern~ine whether or not the t.s-phenotype

was reversible, we cultured wild-type, plp2-I, and plp2-2 cells at 37OC and then

microdissected arrested cells onto solid medium and incubated them at 25OC, a

pernlissive temperature of growth for all strains. Figure 4-2C shows that the temperature

sensitivity is not reversible forplp2-1 cells after 4 hours at 37OC, since more than 50% of

the cells are inviable. On the other hand, the tenlperat~lre sensitivity ofplp2-2 cells is

largely reversible since even after 8 hours at 37OC, 70% of the cells form colonies after

shifting to the pel-missive temperature (Figure 4-2C). Thus, phenotypes observed in

plp2-2 cells at 37OC do not reflect general cell death defects, but rather indicate specific

cellular defects caused by PLP2 loss-of-function.

4.4.3 plp2-1s alleles exhibit cytoskeletal but not G-protein-related defects

Given the putative role of yeast phosducin-like proteins as negative regulators of

heterotrimeric C;-protein signaling, we tested the sensitivity ofplp2-IS strains to mating

pheromone, which is a read-out of heterotrimeric yeast G-protein activation. Yeast ~~ t i l i ze

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a receptor-coupled heterotrimeric G-protcin signaling cascade to sense mating

plicronione In the environmcnt. When this pathway is stimulated, yeast undergo a cell

cycle arrest which can be overcome by the deletion of downstream signaling components

such as STE7, STEII or STEIS. However, consistent with the inability of a STE7 deletion

to rescue loss of PLPS (Flanary et al., 2000), we saw no additional sensitivity to

pheromone as compared to yeast carrying wild-type PLP.? (Figure 4-3). In contrast, thc

sst2A control strain (Sst2p nornially desensitizes cells to the pheromone signal) was

highly sensitive to pheromone, as expected (C'han and Otte, 1982; Figure 4-3). These

data therefore support an essential role for Plp2p outside of mating pheromone signaling

and possibly relating to CCT function (Flanary et al., 2000).

While CCT impacts a wide variety of substrate proteins, its most abundant and

best characterized substrates arc the cytoskeletal proteins actin and tubulin and,

accordingly. yeast t.s alleles of CCT display various actin- and tubulin-related cytoskeletal

defects (Gao et a]., 1992; Siegers et al., 1999; Spiess et a]., 2004; Thulasiraman, et a].,

1999: Ursic et al., 1994; Vinh and Drubin, 1994). We therefore predicted that if Plp2p

modulates the function of CCT by way of direct physical interaction (Figure 4-1A and 4-

1 B), the yIp2-ts alleles might also exhibit cytoskeletal anoniaIies. To investigate this

possibility, we first testcd the sensitivity ofylp2-ts allelcs to actin and microtubule-

disrupting drugs. Figure 4-4A shows that ply2-ts alleles exhibit sensitivity to both

latrunculin and benomyl, indicating niicrofilanient and niicrotubule defects, respectively.

Similar phenotypes are also observable in strains carrying t s or cold-sensitive (cs) alleles

of several CCT genes (Data not shown; Geissler et a]., 1998; Ursic et al., 1994; Vinh and

Drubin, 1994). The latrunculin sensitivity phenotype of the yIp2-t.s alleles are also

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consistent with synthetic genetic interactions between a doxycycline-repressible copy of

PLP2 and mutant alleles of hnil and q 1 2 (Mnainineh ct at., 2004: Davienvala et al.,

2005). Bni I p (formin) and the actin-related Arp2p protein are involved in thc forniation

of actin cables and actin patches, respectively (Reviewed in Evangelists et al., 2003).

Under the microscope, plp2-fs cells also exhibited morphological phenotypes

consistcnt with actin and tubulin cytoskeleton defects. When cultured at the non-

permissive tenlperature of 37OC, p/p2-/.v cells became larger (Figure 4-4B), a possible

indication of defects in actin filament organization (Drubin et a]., 1993). Indeed, ylp2-I

cclls were significantly larger than their wild-type counterparts even at perrnissivc

temperatures (Figure 4-4B). Moreover. after 8 hours of growth at 37OC. the pIp2-/.s cells

exhibited budding defects such as multiple buds (0% wild-type, 7%plp2-I, 30% ,71172-2;

p<O.O 1 ; Table 4-3) and a thickening of the bud-neck junction between mother and

daughter cells (0% wild-type, 1 1% plp2-I, 26% plp2-2; p<O.OOl, Table 4-4). These types

of budding defects are also consistent with disrupted actin function or organization

(Drubin et al., 1993).

4.4.4 Microtubulc and nuclear defects in plp2-ts alleles

Microtubules are essential for orienting the mitotic spindle and for proper

segregation of cliron~osomes during anaphase. The improper segregation of a nucleus to

the daughter bud, therefore, can be an indication of niicrotubule defects. Such a

phenotype can be observed, for example. by abrogating prefoldin, PLPI, or CCT function

in yeast (Lacefield and Solon~on, 2003; Stirling et a]., 2006; Ursic et al., 1994). We

stained nuclei in yly2-fs cells with DAPI to examine possible defects in nuclear

segregation. Compared to wild-type cells, a significant (p<O.OI) number of unbudded

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( I I %), small-budded ( l I YO) and large-budded (22%) pIp2- 1 cells contained multiple

nuclei when incubated at the restrictive temperature for 4 hours (Figurc 4-SA and Table

5). Siniilarly, the niultinucleate plicnotype was found to be significant ( p 4 . 0 0 I) in sn~all-

and large-budded pIp2-2 cells (I 6% and 23% sespectivcly) at 37OC for 4 hours. A

statistically signif'icant number of unbudded pIp2-l cells were also anucleatc at both

permissive and non-permissive ternperaturcs although the defect was more penetrant at

high temperature (6% and 2896, respectivcly) (Figure 4-5A and Table 4-5).

Quantification of the number of cells going through anaphase revealed that both plp2-I

and plp2-2 had significantly fewer cells in anaphase than wildtype at the non-permissive

temperature (37% wild-type, 22% ylp2-l and 17% plp2-2 large-budded cclls in anaphase;

Table 4-6). This type of defect is consistent with the activation of a cell cycle checkpoint,

possibly in response to incorrect anaphase spindle positioning (Lew and Burke, 2003).

One possible explanation for the observed benomyl sensitivity and DNA

segregation defects of the plp2-/s alleles is a deficiency in tubulin cytoskeleton function.

We thcrefore expressed a CiFP-a-tubulin (Tublp) fusion protein in wild-type, plp2-1 and

plp2-2 cells to assess the integrity of microtubules. Intcrestingly, we observed

superficially normal microtubules i n the ply2 mutant cells incubated at the non-

permissive temperature of 37•‹C for 4 hours (Data not shown). Considering that aberrant

spindle postioning could also lead to segregation defects, and that BNII (formin) and

P L P interact genetically (Davie~wala et al., 2005), we speculated that the benomyl

sensitivity and defects in nuclear segregation may result from errors in spindle

positioning, and not spindle assembly or tubulin defects y c ~ s c ~ (Davie~wala et a]., 2005).

When we examincd the orientation of spindles with respect to the mother-bud axis, we

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found that 45% and 42% ( ~ ~ 0 . 0 0 1 ) of pIp2-I and pIp2-2 cells, respectively, had

obviously mis-oricnted mitotic spindles (Figure 4-5B and Table 4-7). Sincc actin

function is required for establishing the correct orientation of the mitotic spindle in thc

GI -phase of the cell cycle (Theesfeld et al., 1999), spindle mis-orientation could in

principle explain both the latrunculin and benomyl sensitivity observed (Figure 4-4A).

Howcver, whether these defects relate solely to actin dysfunction or whether tubulin also

plays a role remains unclear.

4.4.5 Actin polarization defects in pIp2-ts alleles

To better understand the actin cytoskeletal defects in cells with defects in PLP2

function, we stained the actin cytoskeleton with rhoda~ninc-phalloidin. Remarkably, the

plp2-ts cells were refractory to standard phalloidin staining protocols, especially after

growth at high temperatures: we surmise, based on calcofluor staining of cell ~ i a l l chitin,

that this was likely due to cell wall defects (Figure 4-6A). Chitin defects have been

obsei-ved before for specific actin mutants (Drubin et a]., 1993), and wc observed

n~islocalized chitin patches in both temperature-sensitive PLP.? and CCT mutants at non-

permissive temperatures (Figure 4-6). Interestingly, no such defects were observed in

deletions of either the prefoldin subunit gene PACIO or the phosducin-like gene PLPI

(Figure 4-6B). We therefore removed the cell wall in order to visualize actin filaments

clearly with anti-actin antibodies (See Materials and Methods; Figure 4-7). When

examined by i~nrnunofluorescencc, actin localization in the pIp2-ts mutants was

essentially normal at 25OC, but after shifting to the restrictive temperature of 37•‹C for 4

hours, sevcre defects were observed in both ylp2-I and pIp2-2 cells (Figure 4-7 and

Table 4-8). After incubation at 37OC, actin cables were absent in a very large proportion

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of unbudded plp2-I (93%) and plp.2-2 cells (86%), small-budded plp2-I (72Y0) and plp2-

2 cells (94%) and large-budded plp2-l(90%) and plp2-2 cells (91 %) (Figure 4-7 and

Table 4-8). In plp2-1 and plp2-2 cells, the polarization of the actin cytoskeleton was also

defective (Figure 4-7 and Table 4-8). In wild-type cclls, cortical actin patches are

polarized to the bud site i11 unbudded cells and toward the bud tip in small-budded cells.

Howcvcr, at 37OC, normal actin patch polarization in plp2-l cells was observed in only

12% unbudded, 36% small-budded, and 109'0 large budded cells. In ~1172-2 cells at 37OC,

normal actin patch polarization was observed in only 6% unbudded, 30% small budded,

and 7% large-budded cells. A significant proportion of unbudded plp2-I cells also

exhibited abnormal actin patch polarization even at 2S•‹C (Table 4-8). These data are

consistent with the genetic interactions reportcd behveen doxycycline-repressible PLP2

alleles and mutant alleles of BNIl and ARP2, which are normally required for the

formation of actin cables and patches, respectively (Davienvala et al., 2005; Evangelists

ct a]., 2003). Importantly, these data also support our observed functional cooperation

between Plp2p and CCT in actin folding, since mutations in CC'T or PFD subunits show

similar actin organization defects (Geissler et a]., 1998; Ursic et a].. 1994; Vainberg eta]. ,

1999; Vinh and Drubin, 1994).

4.4.6 Mammalian PhLP2A inhibits actin folding in vilro and binds CCT-actin complexes

Because disruption of PLP2 function appears to strongly affect the actin

cytosketeton irt vivo, we exploited the robust in vitro actin folding system developed

using reticulocyte lysate (Cowan. 1998; Leroux, 2000; Vainberg et a]., 2000) to exarninc

whether PhLP2 n~odulates the folding function of CCT. In this system, nascent s3"-

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methionine radiolabelled actin is produced and actin folding is examined on a native

polyacrylamide gel (Leroux, 2000). Since marnmalian actin, CCT, and other mammalian

components have been well-tested in this system, we used purified GST-fused human

PhLP2A protein to test for its effect on actin folding. Figure 4-8A shows that in the

presence of excess GST-PhLP2A, the production of native actin was greatly inhibited at

all time points assaycd. The presence of GST-PhLP2A also led to an increase in the

amount of radiolabelled actin associated with both CCT and PFD complexes (Figure 4-

8A), whose positions on native gels and actin-binding activities are well established

(Cowan, 1998; Leroux, 2000; Vainberg et al., 2000). This result is consistent with

PhLP2A slowing the reaction cycle or inhibiting the activity of CCT; this inhibition could

potentially be taking place in a ternary complex with CCT and substrate, as reported for

PhLP3. leading to a backlog of unfolded substrate that ends up bound to PFD (Figure 4-

8A; Stirling et al., 2006; Lukov et al., 2006).

We therefore examined whether, similar to what was observed for PhLP3,

PhLP2A could form ternary complexes with C'CT and substrate (c.g., ('CT-PhLP3-actin

or CCT-PhLP3-tubulin; Stirling et a]., 2006). To this end, we performed GST-pulldowns

of in virt-o translation reactions of actin or tubulin containing exogenous GST-PhLP2A or

GST alone as a negative control. Figure 4-8B shows in vilro translations of actin and P-

tubulin in reticulocyte lysate, which contains endogenous CCT, that have been

precipitated with glutathione conjugated beads. GST-PhLP2A effectively precipitates

both actin and P-tubulin while GST alone has no such effect. To determine whether the

interaction between PhLP2A and a cytoskeletal protein (actin) depends on CCT (as

opposed to PhLP2A binding the substrate directly), we performed a similar experiment

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with an E. coli lysate translation system, which lacks CCT. Wc found that GST-PhLP2A

only precipitates actin when exogenous, purified CCT was added (Figure 4-8C).

Altogether, these data suggest that PhLP2A negatively nlodulates CCT function in a

manner that may be similar to that of the mammalian PhLP3 protein (Stirling et al.,

2006). This functional mechanism may also extend to PhLP1 proteins, which have

recently been shown to form ternary complexes with CCT and GP to facilitate

heterotrimcric G-protein assembly (Lukov et al., 2006).

4.4.7 High-copy suppression ofp/j12-ts alleles reveals links to the G11S phase transition

Our finding that Plp2p influences actin niicrofilanient and tubulin niicrotubule

function in S. iwevisiue is consistent with the fact that Plp2p associates with and affects

the function of CCT, a chaperone essential for actin and tubulin folding. However, CCT

has been implicated in the biogenesis of several other proteins, some of which are linked

to the cell cyclc. In order to take an unbiased approach to understanding the essential

function(s) of PLP2, which may be CCT-dependent or (potentially) independent, we

executed a high-copy suppression screen of the yly2-ts alleles. A high-copy plasn~id

library carrying -10 kB fragments of yeast genomic DNA (C'arlson and Botstein, 1982)

was transfol-nied into theplp2-1 strain, and colonies were plated at 37OC. Plasniids

isolated from colonies that grew well at 37•‹C were screened by PCR to exclude those

carrying the PLP2 gene, a predictable suppressor. Plasmids that did not contain PLPZ

were sequenced to identify the gene(s) present in the suppressing plasmid. One high-copy

fragment we isolated contained PLCI, a yeast phospholipase C homologue. We

confirmed that PLCI was a suppressor becausc a galactose-inducible PLCI construct

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could partially suppress both pIp2-l and pIp2-2 alleles (Figure 4-9E). Another

suppressing plasmid cncoded two genes, VHSI and PAMI. Thcse were expressed

individually in yIp2-I cells and, surprisingly, both were found to partially suppress the

temperature-sensitive defect (Figure 4-9A). PA IWI was previously identified as a high-

copy suppressor of the lethality associated with complete loss of protein phosphatase 2A

function (Hu and Ronne, 1994). VHSI was previously identified in a screen for

suppression of the lethality associated with loss of the SIT4 protein phosphataae and

phosphatase regulator HL4L3 (Mufioz et a]., 2003). In the same screen from which VHSI

was previously identified, several other genes, ~ncluding protein phosphatase subunits

and other genes active in the G 1 IS phase transition, were found to suppress loss of SIT4

and HAL3. To try and identify additional plp2-/ suppressors, we tested some of the sit4-

htrl3 suppressors found in the screen by Muiioz et al. (2003; Figure 4-9B). Remarkably,

the genes YHS2, VHS3, PTC2, PTC'.?, YAP7. HAL3, HAL5 and CLN3 were also able to

partially suppress temperature-sensitive growth defects of ylp2-l cells. This apparent lack

of specificity suggests that driving cells to continue through the cell cycle helps to

somewhat alleviate thcplp-7-based tenlpcrahlre sensitivity. The extensive overlap

behveen suppressors of plp2-ts alleles and suppressors of sit4-htrl3 mutants (Muiioz et a].,

2003) suggests either that Plp2p directly effects Sit4p or Hal3p or that Plp2p works in a

parallel pathway to that of Slt4p and Hal3p.

While the partial suppressors ofplp2-I have somc divcrse functions, many are

involved in protein phosphorylationi'dephosphorylation and they all play roles in

promoting the GI IS phase transition of the cell cycle (Muiioz ct al., 2003). These data

therefore suggest that PLP2 also has a rolc in G 1-progression. Indeed, we observed an

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accumulation of unbudded pIp2-l cells aftcr 2 hours at 37OC (50% unbuddcd, p.-0.00 1;

Table 4-9), although this could also be related to thc actin defects reported abovc. Also,

the observation that plp2-ts cells become larger than wild-type cells at 3 7 O C ' , is consistent

with a pause in progression from GI to S phase. To test these suppositions, we examined

the entry of synchronized pIp2-l cells into S-phase. Synclironizcd I N cell populations

werc prepared by clutriation, grown at 37OC, staincd at various time points for DNA

content and subjected to fluorescence activated cell sorting (FACS). Figure 4-9C shows

that plp2-I cells are delayed in their entry into S-phase compared to their wild-type

counterparts. Even after 4 hours, there are still a greater proportion of ylp2-l cells in CJ I

than G2, while approxiniately half of the wild-type cells are found in G2 (Figure 4-YC).

This delay in cell cycle progression is consistent with the G I IS function of the ylp2-l

suppressors we identified, and the accumulation of unbudded cells in asynchronous plp2-

/ cell populations at restrictive temperatures (Table 4-9). We also observed a delay in

budding when a-factor-synchronizcd plp2-t.s cells were releascd in 37OC media (Figure

4-10). Importantly, entry into S-phase seems to be indepvm'eni of actin function

(McMillan et al., 1998) as we found that the actin cytoskeleton appears siniilarly

defective in plp-3-I and plp2-2 cells carrying high-copy empty, PTC2- or Y'4P7-

containing plasmids (Data not shown). Our data therefore suggest an additional, actin-

independent role for PLP2 in the cell cycle.

To explore the possibility of allele-specific suppression, we tested two of the

stronger suppressors of plp2- 1 ( YL4P7 and PTC2) in plp2-2 cells. While high-copy

production of YAP7 suppresses y1p2-2 temperature sensitivity, PTCZ overexpression

actually inhibits the growth ofylp2-2 cells (Figure 4-YD). Also consistent with

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functional differences between thc ply2 alleles, unbudded cells did not accumulate at the

restrictive tempcrature for plp2-2 as they did for plp2-I (Table 4-9). Finally, wc observcd

that heterozygous diploids carryingplp2-I but not ylp2-2 retained some temperaturc

sensitive growth defects at 37OC (Figure 4-9F). Homozygous diploids of each plp2-/.s

allele were very sensitive to high temperatures and the presence of a low copy plasmid

completely rescued this sensitivity as seen for haploids (Figure 4-9F). Thesc data suggest

that plp2-I has semi-dominant affects on the cell because the single copy of PLP2 in the

heterozygotes does not completely rescue the growth defects whereas the prescncc of

plasmid in homozygotes, which introduces several copies of PLP2, does rescue the

growth defects. Together, the observed allele specificity supports a role for PLP2 in the

cell cycle because the cytoskeletal phenotypes ofplp2-l and plp2-2 are nearly idcntical

(Figures 4-7). Indeed, if the observed cell cycle defects were somehow an indirect effect

of cytoskeletal defccts, thc two ply2 alleles would be expected to behave similarly.

4.5 Conclusion

In recent years, the phosducin-like proteins PhLPl and PhLP3 have emerged as

modulators of cytosolic chaperonin function in euka~yotes (McLaughlin et a]., 2002;

Lacefield and Solomon, 2003; Martin-benito ct a]., 2004; Lukov et a]., 2006; Stirling et

al., 2006). We now report in vitro and in viva studies demonstrating that the PhLP2

subgroup also n~odulates CCT function. PhLP2 exerts similar effects on CC'T as does

PhLP3, in that PhLP2 binds CCT, and when in excess, slows protein folding in a tcmary

conlplex with substrates (Figures 4-1 and 4-8). Using loss-of-function alleles of PLP2,

we uncovcred a role for Plp2p in both cytoskeletal protein function and the cell cycle.

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4.5.1 PLP2 function is essential for viability but not G-protein signaling

Although pliosducin-like proteins havc gcncrally been implicated in

hetcrotrimeric G-protein signaling (Schroder and Lohse, 1996), here we establisli that

P L P is unlikely to play a critical role in heterotrimeric G-protein signaling (Figure 4-3).

I t is possible that thc weak interaction between Plp2p and Ste4p (GP) reported by Flanary

et al. (2000) is due to a ternary CCT-Ste4p-Plp2p complex that forms during Ste4p

folding, as shown for the mammalian proteins GP (STE4 homologue), CCT and PhLPI

(Lukov et a]., 2006) and not a binary Plp2p-Ste4p complex as the authors suggest

(Flanary ct a].. 2000). The function of Plp2p appears closely aligned with that of CCT;

Plp2p binds CCT in yeast and ts-alleles of PLP2 and CC'T subunits exhibit similar

phenotypes, consistent with a functional cooperation between the CCT chapcronin and

Plp2p (Figures 4-1 through 4-10; Camasses et al., 2003; Siegers et al., 1999; Ursic et a].,

1994; Vinh and Drubin, 1994). Importantly, and similar to CCT (Ursic et a]., 1994; Vinh

and Drubin, 1994), we conclusively demonstrate that PLP2 is essential for viability in S.

cwoviar (Figure 4-1).

4.5.2 Cytoskeletal phenotypes inp/p2 loss of function cells

The reason for the essential nature of PLP2 is not entirely clear, although we

establish cytoskeletal phenotypes as a profound cellular defect of cells lacking functional

Plp2p. Cells carrying plp2-ts alleles are sensitive to the drugs latrunculin and benoniyl.

which disrupt actin and tubulin filaments. respectively (Figure 4-4). The ylp2-ts strains

also have aberrant budding morphology, become larger and accumulate multinucleate

and anucleate cells, all of which can be indirect indicators of cytoskeletal defects (Figure

4-4 and 4-5 and Table 4-5). Moreover, the mitotic (microtubule) spindles ofylp2-l and

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p/p2-2 cells bcconle niis-oriented with respect to the mother-daughter axis when the cells

are cultured at high temperature (Figure 4-5B). The p/p2-ts strains also show weakened

polarization of actin patches and a nearly complete loss of actin cables (Figure 4-7).

Together, these data support a role for Plp2p in actin and t~ibulin function, and arc

consistent with previous findings that a reduction in PLP2 expression using a

doxycycline repressible promoter allele lcads to synthetic growth defccts in the prescnce

of BN/I or ARP2 mutants (Davierwala et a]., 2005). These genetic interactions support

the aberrant actin polarization and spindle nis-orientation observed in both y/p2-/A

alleles. Furthermore, the interaction of Plp2p with CCT strongly supports a role for Plp2p

in the production of actin and tubulin. Interestingly, our data also suggest that actin-based

fi~nctions may be more sensitive to Plp2p disruption than are tubulin-based functions.

This may reflect that the cellular demand for properly folded actin in the budding cell is

higher than it is for tubulin, and that actin-based phenotypes manifest before tubulin-

specific defects can be detected. In conclusion, our stlldies provide unequivocal evidence

that both actin and ti~bulin functions are compromised when Plp2p function is perturbed.

Furthernlore, the effect of the phosducin-like protein is likely mediated via a direct

functional interaction with CCT alone, or with CCT associated with an actin/tubulin

substrate protein (Figures 4-1 and 4-8). Given that disrupting prefoldin function in

S. L ' ~ I ' c L ' ~ . \ ~ L I ~ specifically r e s~~ l t s in severe actin and tubulin cytoskeletal defects

comparable to those of the severe plp2-t.r allele, we hypothesize that at least one CCT-

dependent (or potentially independent) Plp2p function is unrelated to the cytoskeleton but

is cssential for cell growth and survival.

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4.5.3 Cell-cyclc phenotypes in p1p2 loss of function cells

Whilc our characterization of the ylp2-ts alleles showed many cytoskeletal

phenotypes consistcnt with an interaction with CCT. an unbiascd high-copy suppression

screen retcaled a link to the cell cycle, especially the progression out of G 1 phase into S

phase. When we examined cell cycle progression by DNA content in plp2-l cells, we

found a delay in DNA replication (Figure 4-9). Moreover, we obsenled an excess of

unbudded cells when the mutant strains were cultured at high temperature (Table 4-9).

While i t is possible that some of these defects relate to actin dysfunction, which can delay

budding (as we observed in Figure 4-10), it is likely that Plp2p also affects one or more

non-cytoskeletal, cell cycle-related CCT substrate(s), as we elaborate below.

4.5.4 Models of Plp2p-cell cycle connection

Other groups have shown that CCT has an important role in cell cycle progression

bccause of its effect on the biogenesis of the anaphase pronioting complex (APC)

regulators Cdc20p and Cdh I p and the protein phosphatase subunit Cdc55p, any one of

which could be regulated by Plp2p (Caniasses et al., 2003; Siegers et al., 2003). Since

these particular CCT substrates contain WD-repeats, a role for Plp2p seems plausible in

light of the known role for other phosducin-like proteins in WD-repeat folding/assembly

by C'CT (Lukov et al., 2005; Lukot et al., 2006). For example, impaired CCT function

leads to precocious cntly into S-phase because of loss of Cdh I -APC activity, which

nor~nally inhib~ts entry into S-phase (Camasses et al.. 2003; Harper et al., 2002). If Plp2p

were actlng to negatively regulate thc CCT-mediated Cdh I p-APC assembly, thcn ylp2-l

alleles may accunlulate excess Cdh Ip-APC, thus slowing entry into S-phase.

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An alternative connection supported by the litcrature 1s one between C'CT and

type 2A protein phosphatases (PP2As). CCT is known to assist the folding of the ycast

PP2A rcgulator Cdc55p, and physical interact~ons have been reported between CCT and

the yeast phosphatase components Sit4p, Pph2 l p, Pph22p, Pph3p, and Tap42p (Gavin et

al., 2006; Ho et a]., 2002; Siegers et a]., 2003). The finding that PAMI, which bypasses a

loss of PP2A activity (Hu and Rome, 1994), can weakly suppress pIp2-I and pIp2-2

alleles (Figure 4-9A) is consistent with a role for Plp2p and CCT in the folding of PP2A

components. Indeed, all the suppressors we identified were also identified in a screen for

suppression of lethality of strains lacking the phosphatase SIT4 and the phosphatase

regulator HAL3 (Muiioz ct a]., 2003). Moreover, the toxicity associated with

overexpression of SIT4 and SAP155 (a Sit4p rcgi~latory protein) is suppressed by

increased CCT6 copy number in a manner that is not understood (Kabir et a]., 2005).

Importantly, Sit4p and other PP2As play roles in both GI progression and cytoskelctal

organization (Muiioz et a]., 2003; Stark, 1996).

The aforementioned models of PLPZ function, while speculative, could help

explain the diverse and weak G 1 -related suppressors we identified as well as the

polari7atio11, cytoskeletal and cell cycle defects resulting from plp2 mutations. Also of

importance is the finding that, in mammalian cells, CCT itself was shown to be

upregulated at the G 11s phase transition and C'CT depletion caused arrest of cells at the

same cell cycle stagc (Grantham et a]., 2006; Yokota et a]., 1999). These obsesvations

suggest an increased folding requirement for some CCT substrates at this cell cycle stagc,

consistent with our findings.

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4.5.5 Pcrspectives

Given the literature and the data presented here, a picture of Plp2p activity

emerges in which it functions to regulatc the folding of several CCT substrates that

together control cytoskeletal morphogenesis and cell cycle progression. Based on thc

phenotypes we observed in plp2-bs alleles, candidates for these critical substrates include

actin and actin-related proteins, tubulin. Cdc20p, Cdc55p, Cdh I p and/or as-yet-

unconfirmed CCT substrate(s) such as Sit4p. While some of these substrates are essential

and others arc dispensable, we propose that the additive effects of their altered folding by

CCT contributes to the complex phenotypes and eventual lethality associated with loss of

PLP2 function. Alternatively, there could be one ci-itical substrate whose biogenesis

wholely depends on Plp2p, although we favour the former option because of the

phenotypes of PLP2 mutations. Actin seems to be particularly affected by PLP2

mutations, suggesting an important role in controlling actin biogenesis andlor the

function of an actin regulator like Bni I plArp2p. However, we cannot rule out a role for

Plp2p in tubulin biogenesis although severe effects, such as those seen in prefoldin

mutants, were not observed (Geissler et a!., 1998). The conserved nature of PhLP-CCT

cooperation and the diversity of substrates likely to be impacted by PhLPs reveals thc

importance of understanding PhLPs for a con~plete picture of chaperonin function in

eukaryotes.

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Figures

Figure 4-1 PhLP2IPlp2p is an essential CCT binding protein

(A) Cct2p-HA is imlnunoprecipitated using an anti-lnyc antibody from S. cerevisicrc cells

co-expressing Cct2p-HA and Plp2p-1y1c but not from control wild-type (WT) or Cct2p-

HA-cxpressing cells. HA-tagged Cct2p is detected by Western blot analysis using an

anti-HA antibody. [NOTE: M.S. and A.P. generated the data in Al. (B) Western blot

showing CCT co-precipitating in vitro with GST-PhLP2A or GST-PhLP3 in a GST

pulldown, but not the GST-alone control. Purified CCT was run as a control to show that

the antibody recognizes CCT (left panel).

Western Blot

Y & *

Anti-myc lmmunoprecitation

anti-CCT I Western Blot

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(C) Counterselection of a URA3-marked plasmid carrying PLP.? ([pURAJ P L P J ] ) in

pIp2A cells with 5'-fluoroorotic acid (5'FOA) shows that PLP2 is an essential gem. Cells

prototrophic (PLP.? URA3) and auxotrophic (PLP2 I I I Y L ? - ~ ~ ) for LITLIC~I (UTLI) were

included as controls.

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Figure 4-2 Generation of' temperature sensitivepIp2 alleles

(A) Temperature sensi tive alleles of PLP2, namely plp2- I and pIp2-2, wcrc spotted on

YPD media and grown at the temperatures shown for 48 hours. p1p2-I displays a more

severe /s phenotype than pIp2-2. (B) Table showing the amino acid mutations present in

twoplp2-/s alleles. (C') Reversibility of thepk)2-/.~ alleles was assessed following growth

at 37•‹C' for thc times shown. Shifted cells were platcd at a pcrniissive temperature and the

number of niicrocolony forming cells was assessed under a microdissection microscope.

plp2-2 temperature sensitivity is largely reversible compared to the more severe plp2-I

allele. [NOTE: The temperature-sensitive strains were generated by M.S. and A.P.

in the lab of A.A.H.1

Mutations in plp2-ts alleles

Hours at 37•‹C

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Figure 4-3 PLP2 loss of function does not impact pheromone sensitivity

Relative sensitivities to mating pheromo~~e-i11c1~1ced death f'orplplA and pIp2-t.s cells as

assessed by the radius of clearance caused by a pheromone-soaked disk. sst2A was

included as a pheromone-sensitive control. Radius of clearance is indicated in

millimetres.

Sensitivity to Mating Pheromone

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Figure 4-4 plp2-ts cells are large and have increased sensitivity to cytoskeletal- destabilizing drugs.

(A) Benomyl and latrunculin sensitivity ofyly2-l and ylp2-2 mutants relative to wild-

type (PLP2) cells, as determined by relative clearance caused by drug-inoculated paper

discs. (B) Cell sizes of the indicated strains wcrc measured perpendicular to the mother-

daughter axis for at least 100 cells. An asterisk (*) indicates that, relative to wild-type, the

cells were significantly larger (p<0.0 1 ) as determined by an independent variable /-test.

A

O Wildtype

1 0 plp2- 1

2 4 Hours at 37•‹C

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Figure 4-SpIp2-ts cells exhibit aberrant nuclear segregation and spindle orientation

(A) Images of DAPI-stained PLPZ, pIp2-1 and yIp2-2 cells grown at permissive (23•‹C)

and non-permissive temperatures (37•‹C'). A I T ~ Y S indicate multinucleate cells. Scale bars

indicate 1 0 pm.

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(B) GFP-a- t~~bul in expressing PLP2, plp2-I and plp2-2 cells were visualized and scored

for spindle orientation with respect to the mother-daughter axls. Cells are outlined in

white and the pcrccntages of mis-or~ented spmdles are ind~cated bclow each panel.

Representative images are shown of norlnal spindles for w~ld-type and plp2-/ and plp2-2

cclls at 23OC and of mis-oriented spindles in plp2-I and plp2-2 at 37OC. The imagcs are

of equal scale, plp2-1.v cells are sin~ply larger at high temperature.

070 rn~s-or~enrea

nln 3- I

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Figure 4-6 Aberrant chitin levels and localization in plp2-ts and cct-ts cclls

(A) Mid-log phase PLPZ. plp2-I or ylp2-2 cells were grown at 2S•‹C' or 37OC for four

hours before staining with calcofluor white, which stains the chitin-containing cell wall.

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(B) Mid-log phase/)//)/A, ~ L K . / O A . c.ct/-2 and crt4-/ cells were grown at 37OC for four

hours and stained with calcofluor white. Cells grown at 2S•‹C appeared wild-type (not

shown) while C'CT mutants accumulated cxcess chitin (ccbtl-2) or mislocalizcd chitin

patches (cc.14-1). Arrows show abberant chitinous patches in mother cells and daughter

buds and erroneously thick cell wall chitin in cctl-2 mutants. Scale bar indicates I0 p n ~ ,

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Figure 4-7 Actin filament organization defects inpl1~2-ts cells

( A ) PLPZ, p1p2-l and p1p2-2 cells grown at permissive temperature (25•‹C') or non-

permissive temperature (37•‹C') were stained with anti-actin antibodies. Arrows indicatc

cells without actin cables and with poorly polarized patches. Scale bar indicates 10 pm.

25•‹C 37•‹C

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Figure 4-8 Mammalian PhLP2A binds CCT and modulates its activity in vitro

(A) In vitr-o folding reactions of nascent s3'-labclled actin in the prescnce of GST or

GST-PhLP2A. CCT:actin and PFD:actin binary complexes. as well as native actin, are

indicated. The identity of the fast-migrating band as native monomeric actin was verified

by DnaseI-shifting on the native gel (right panel). An SDS-gel illustrating the relative

amounts of translation products is shown (lower panel).

Native Gel

Translation

t Native Actin

----- 10 20 30 40 50 Dnase I control

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(B) Co-precipitation of ~ ~ ~ - 1 a b e l l e d actin or P-tubulin with GST-PhLP2A in a

reticulocyte translation rcaction. The lower panel shows that the lcvels of translation wcrc

comparable. (C') C'o-precipitation of s"-labelled actin produced in E. coli lysate takes

place only in the presence of exogenously added CCT. Actin cDNA was translated in

E . c d i lysate with or without the addition of purified rabbit CCT and the relative levels of

the translation products are shown on the left. GST-PhLP2A co-precipitated the newly

made actin only in reactions to which CCT had been added whereas GST alone had no

such affect as shown on the right.

Reticulocyte Lysate Translation

C )

GST pulldown

E. coli Lysate Translation

@ * Translation

Actin 13-tubulin

V '

On < < 6 p$

5 c: material 0 0

CCT - + - - + +

Actin

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Figure 4-9 High-copy suppressors of plp2-1 indicate a role for PLP2 in cell cycle progression

Partial suppressors ofp lp2- l temperature sensitivity identified i n a high-copy screen (A)

and suppressors identified in the literature (B). pIp2-I cells carrying the indicated

plasmids were grown to log-phase and serially-diluted on Sc-Ura media and grown for 2-

3 days at permissive (25•‹C) or restrictive (37•‹C) temperatures before imaging. (C) A

delay in DNA replication is observed in synchronized plp2-I cells. Graphs indicate

fluorescence activated cell sorting (FACS) analyis of cells stained for DNA content over

time. [NOTE: Figure 9-C was completed by IM.S., A.P. in the lab of A.A.H. The

suppressors shown in 9A were identified by K.S.T. and preliminary testing of the

suppressors in 9B was done by K.S.T.1

A

Wildtype

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(D) Serially dilutcd p/p2-2 cultures carrying the indicated plasmids wcre grown for 2-3

days on Sc-Ure as in (A) and (B) before imaging. (E) Serially diluted PLPZ, pIp.7-I and

p/p2-2 cells carrying a galactose-inducible copy of PLCI were grown on glucosc

containg (repressing PLCl expression) and galactose/raffinose containing (inducing

PLCI expression) nicdia at 25•‹C and 37•‹C. (F) Diploid yeast plated as in (A) and grown

at 25•‹C and 37•‹C' for 3 days before imaging. ** Indicates where ylp2-llPLP2

heterozygous diploids reveal growth defects not seen in p/p2-2/PLPZ cclls.

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Figure 4-10 Delayed rebudding in a-factor synchronized plp2-ts cells

PLPZ (e), pIp2-/ ( x ) and pIp2-2 ( A ) cclls were synchronized with a-factor by the low-

pH method according to Amberg et al. (2005). Cells were shifted to 37OC for I hour prior

to release into 37OC media. Cells tixed at the time points indicated werc scored for the

presence of a daughter bud. (B) n-values and percentage budded cells for the data shown

in (A).

B

Time (Minutes)

0 30 45 60 7 5 90 120 150

NT PLPZ n = % budded

15.60% 15.20% 68.80% 90.10% 90.50% 89.90% 77.00% 81 10%

Minutes

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4.7 Tables

Table 4-1 Yeast strains used in tflis cflapter

Strain name Genotype Source MLY IOO MLY 110

MLYII1

MLY 150

AHY95 1

AHY955

AHY988

AHY994

AHY997

AHY999

DUYSSX

DUY559

DUY560

DUY 56 1

DDY 299 MLY 151

MLY 152

MLY 153 MLY 154 MLY 155

Mat a ~ ( 1 3 - 5 2 , 1 ~ ~ 2 - 3 , -1 12. his3, t1lctl5 Mat a ~ 1 ~ 3 - 5 2 , 14112-3, -112, h i d , mrtl.5. Aplyl:: KanMX4 Mat a ut~13-52, 1 ~ ~ 2 - 3 , -112, his.3, 111ctl5, ApclO::KanMX4 Mat a r/t173-52, 1~112-3, -112, his3, nlctl5, Avst2:: KanMX4 Mat a ULI'LJZ-1, l~~.s.?-~YOl, ut.cr3-52, 1 ~ ~ 2 - 3 , 112, 17is3- 11 , trpl-1 ylp2A::Hihs3 [yRS416 PLP2 URA31 Mat a ade2-1, ~~:s2-HOl, lrvcr3-52, ti-pl-1, lez12-3,- 112, his3-1 I , plp2A::Hi.s3. GFP-TUB1 ::His3 bRS4lCi PLP2 UR/43] Mat a udc2- I , lys2-801. 1/1-03-52, ler12-3, 112, l1is3- I I , try1 -1, PLP2-t17ycI):; His3 Mat a (&2-1, I):s~-SOI, r/t.u3-52, lc112-3, 112, l7is3- 11, trpl-I, PLP2-n7jx~Y::Hi.s3. CCT2-HA::KI Ttpl , yep4A:: His3 Mat a ude2-I, l~s2-~SO/ , 1/tu3-52, ttlyl-I, ler12-3,- 112, l7is3-11, ylp2A::His3. GFP-TUB I::Hi.r3, ,91122- I :: Lm2 Mat a m'~.2-1, bs2-801, 11r~73-52, t i p / - / , 1~112-3,- 112, 17is3-I I , y/y2A::Hi.s3, GFP-TUB1 ::Hi.s3, p1112-2:: Lcw2 Mat a ut.u3-52, 1 . ~ 2 - 3 , -1 12, ttpl-7, cc-tlA::Ler/2 [CCTI TRP I CEN YCpMS381 Mat a ut-(13-52, ler12-3, -1 12, ttpl-7, cctlA::Lru2 [cc.tl-1 TRPI CEN Y C ~ I M S ~ S ] Mat a z1t~13-52, lc.112-3, -1 12, ttpl-7, ct.flA::Lcu2 [ c ~ ~ t l - 2 TRP I CEN YCpMS381 Mat a wu3-52, lc.112-3, -112, ttpl-7, cctIA::Lclr,' [cctl-3 TRPI CEN YCpiVfS381 Mat a wa3-52, lcu2-3, -1 12, ~ t 4 - 1 Mat a l a p1/12-l::Lc~/2/ply2-1::Lcu2 *relevant genotype Mat a l a plp2-2::Le1/2/ylp2-2::Lc.1/2 *relevant genotype Mat a l a plp2-2::Leu2/PLP2 *relevant genotype Mat a l a plp2-l::Ler/2/PLP2 "relevant genotype Mat a l a PLP2/PLP2 *relevant genotype

Y KO collection Y KO collection

YKO collection

Y KO collection

This study

This study

This study

This study

This study

This study

Doris Ursic

Doris Ursic

Doris Ursic

Doris Ursic

David Drubin This study

This study

This study This study This study

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Table 4-2 Plasmids used in this study

Relevant features Empty Vector PLP2 PTC2 VHSI VHSZ VHS3 PTC2 PTC3 YAP7 HAL5 CLlV3 HAL3

Genotype

URA3 2 p URA3 C'EN PLP2 URA3 211 PAM I URA3 2p VHS I URA3 2 p VHS2 URA3 211 VHS3 URA3 211 PTC2 URA3 21.1 PTC'3 URA3 211 YAP7 URA3 211 HAL5 URA3 2 p CLN3 URA3 2 p HAL3

Source Dr. Christopher Bell This study Muiioz et a]., 2003 Muiioz et a]., 2003 Mrlfioz et al., 2003 M ~ f i o z ct al., 2003 Muiioz et al., 2003 Muiioz et al., 2003 Muiioz et al., 2003 Muiioz et al., 2003 Muiioz et al., 2003 Muiioz et al., 2003

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Table 4-3 Mr~ltiple buds in plp2-ts mutants

(y' 0 slllgly . .

budded Strain n= cell

WT (2S•‹C) 101 100% WT (37OC) 1 16 100%

p11,-1 (25•‹C) 1 13 99%

plp2- 1 (37•‹C) 1 1 5 93% plp2-2 (25•‹C) 123 100% pIp2-2 (37•‹C) 1 04 70%

'%, multiple budded cell

0% 0'% 1 % 7% 1%

30%

p-value vs. WT at same

concljtions NIA N/A N.S.

p a 0 1 N.S.

p<0.00 1

NIA = Not applicable N.S. = Not signifmnt

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Table 4-4 Thickening of the bud neck junction in pfp2-ts mutants

% normal Strain n= bud neck

% thick bud neck p-value

2%) N/A 0% N/A 2% N.S. 12% p<o.oo 1 1% N.S.

26% p<O.OOI

N/A = Not applicable N.S. = Not significant

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Tablc 4-5 Nuclear defects in plp2-ts cclls

Strain n= multinucleate p-value n= anucleate p-value

Unbudded cells

WT(25OC) 106 0% NIA 106 0% N/A WT(37"C) 102 1 % N/A 102 0% N/A

plp2-l (25•‹C) 107 4% p<0.05 101 6% ~ ~ 0 . 0 2 5 pIp2- 1 (37•‹C) 10 1 11% p<O.01 73 28% pi0.00 1 pl@-2 (25•‹C) 104 0% N.S. 100 0'30 N.S. plp2-2 (37•‹C) 100 4% N.S. 94 6% p<0.025

Small-budded

WT (25•‹C) 108 0 N/A 108 0% N/ A

WT (37•‹C) 88 0% N/A 88 0% N/A plp2-1 (25•‹C) $2 1 '% N.S. 81 1 % N. S. plp2- 1 (37•‹C) 7 1 11% p<0.001 70 1% N.S. plp2-2 (25•‹C) 1 00 0% N.S. 100 0% N.S. ~1,172-2 (37•‹C) 105 16% p<0.001 102 3% N.S.

Large-budded

WT (25•‹C) 1 10 0% N/A 110 0% N/A WT (37OC) 102 0% N/A 102 0% N/ A

plp2-1 (25•‹C:) 100 4% pi0.05 103 0% N.S. ylp2-1 (37•‹C) 103 22% p<0.001 100 0% N.S. p1p2-2 (25•‹C) 104 0% N.S. 100 0% N.S. plp2-2 (3 7•‹C) 1 03 23% p<0.001 101 2% N.S.

N/A = Not applicable N.S. = Not significant

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Table 4-6 Anaphase entry defects in plp2-ts mutants

96 large- budded cells

Strain n= in anaphase WT (25•‹C) 110 36% NIA WT (37•‹C) 102 37% N.S.

plp2-l (25•‹C) 100 3 1% N.S. ,~1/12-l (37•‹C) 103 22% pi0.025 ~ 4 ~ 2 - 2 (25•‹C') 104 38% N S . plp2-2 (37•‹C) 103 17% p<O.O I

N/A = Not applicable N.S. = Not significant

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Table 4-7 Spindle misorientation in plp2-ts mutants

% correct Strain n= orienatjon

[Yo incorrect

orientation p-value 9% N/A 7 (Yo NIA 8 (Yo N.S.

45% p<O.OOl 1 2 % N. S.

42(%1 p<O.OO 1

NIA = Not applicable N.S. = Not significant

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Table 4-8 Actin organization defects in plp2-ts cells

Strain Normal Aberrant Visible p-valuc (Growth Budding actin patch nctin patch actin rclativc

Temperature) stage n= polarization polarization cables to WT WT (25•‹C) unbudded

small large

WT (37• ‹C) unbudded small large

yIp2-l (25•‹C) unbudded small large

ylp2- l (37•‹C) unbudded small large

plp2-2 (25•‹C) unbudded small large

plp2-2 (37•‹C) unbudded small large

N/A N/A N/A N/A NIA N / A

p<O.OO 1 N.S. N.S.

p<o.oo 1 p<o.oo 1 p<o.oo 1

N.S. N.S. N.S.

p.cO.00 1 p<o.oo 1 p<0.00 1

N/A = Not applicable N.S. = Not significant

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Table 4-9 Budding index ofplp2-ts cells at high temperature

37•‹C 37•‹C 37•‹C 25•‹C 2 hours 4 hours 6 hours

n= 1 1 1 120 115 114 WT unbudded 32% 30% 24% 33%

small 32% 36% 29% 27% large 35% 34% 47% 39%

p/p2- 1 11 = 122 161 140 132 unbudded 38% 50% " 44% * 52% *

small 3 1% 29% 27% 17% large 3 I (% 22% 29% 32%)

-- --

p/p2-2 n= 117 131 113 113 unbudded 30% 21% 3 0% 36%

small 31% 40% 37% 3 6% large 3 9% 39% 33% 27%

* Significantly different from WT at the same conditions (pCO.0 1 , ~ 0 . 0 I and c-0.025)

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4.8 Reference list

Adams, A., Gottschling, D.E., Kaiser, C.A., and Stearns, T. (1 997). Methods in Yeast Genetics: A Cold Spring Ilarbour Laboratoiy Course Manual, Cold Spring Harbour: Cold Spring Harbour Press.

Amberg, D.C., Burke, D.J. and Strathern, J.N. (2005). Methods in Yeast Genetics, 2005 edition. Cold Spring Harbor: Cold Spring Harbor Laboratory Press.

Ayscough, K.R., Stryker, J., Pokala, N., Sanders, M., Crews, P., and Drubin, D.G. (1 997). High rates of actin filament turnover in budding yeast and roles for actin in establishment and maintenance of cell polarity revealed using the actin inhibitor latrunculin-A. J Cell Biol 137, 399-416.

Blaauw, M., k lo l , J.C., Kortholt, A., Roelofs, J., Ruchira, Postma, M., Visser, A.J., van Haastert, P.J. (2003). Phosducin-like proteins in Dictyosieli~lm cliscoitle~~m: inlplications for the phosducin fanlily of proteins. EMBO J 22, 5047-5057

Camasses, A., Bogdanova, A., Shevchenko, A., and Zachariae, W. (2003). The CCT chaperonin promotes activation of the anaphase-promoting con~plex through the generation of functional Cdc20. Mol Cell 12, 87- 100.

Carlson, M., and Botstein, D. (1 982). TWO differentially regulated mRNAs with different 5' ends encode secreted with intracellular forms of yeast invertase. Cell 28, 145- 154.

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CHAPTER 5 GENETIC INTERACTORS OF YEAST CCT AND A NOVEL ROLE FOR THE CHAPERONIN IN SEPTIN RING FUNCTION

Note regarding contributions: Data presented in this chapter represent part of an article in preparation from a

collaborative effort between Michel Leroux's lab, and the groups of Dr. Keith Willison (Chester Beatty Laboratories at The Institute of Cancer Research, London England) and Dr. Charles Bootie (Banting and Best Department of Medical Research, University of Toronto, Toronto Canada).

Dr. Boone's group generated the ir.11-2::NatMX strain used to initiate the synthetic genetic array (SGA) screen and performed the preliminary SGA screen itself (Table 5-1 and 5-2). Dr. Willison uncovered physical interactions of CC'T with septin subunits. Aside from writing this chapter I validated the SCiA screen by randoni spore analysis (Table 5-2) and examined the localization of the septins (Cdc3p and Cdc lop) in ccv-1.r1c.s and other mutant strains (Figures 5-1 and 5-2 and Tables 5-3 and 5-4).

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5.1 Abstract

One of the important qucstions regarding CCT function in the cell is the nature of

its substrate repertoire. While a handfid of CCT substrates have been identified, lnrgc

scale studies indicate that many more substrates are likely to be found. A genomic

approach was undertaken by our group to identify novel cellular processes in which CCT

participates and ultimately with the goal of finding novel substrates that require CCT for

folding. Coniplementing this study was thc identification by our collaborator that several

septin subunits physically interact with CCT. As a proof-of-principle, we show that CCT

function is critical for the normal localization of septin rings iri vivo as predicted by our

collaborative data.

5.2 Introduction

The chaperonin CCT has long been known to be critical for the biogenesis of

actins and tubulins (Sternlicht et al., 1993). Accordingly, loss of CC'T function leads to

actin and tubulin cytoskeletal defects in tissue culture cells. S. ccrwisiue, and C'. c~1~g~ln.s

(Grantham et al., 2006; Lundin et a]., 2007, submitted; Ursic and Culbertson, 1991 : Ursic

et al., 1994; Vinh and Drubin, 1994). I t had been thought that actins and tubulins

represented the only CCT substrates though other proccsses might be dcfective these

losses of function are masked by the pervasive and lethal cytoskeletal defects. In recent

years, a handful of additional CCT substrates were identified that suggested a broader

cellular role for the chaperonin than previously thought (Canlasses et al., 2003, Feld~nan

ct al., 1999; Siegers et al., 2003). However, the true scope of the CCT substrate repertoire

remains uncertain.

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Large-scale, unbiased proteomic analyses in yeast have revealed a large nuinber

of physical intcractions for CC'T, niany of which could represent substrate protcins

(Gavin et al.. 2006; Ho et a]., 2002; Krogan et a].. 2006). One problem with this type of

analysis is that the N- and C'-termini of CC'T are buried within its cavity, mcaning that

any terminal fusion could be disruptive to the 16 subunit hetero-oligonieric CCT

coniplcx, and that affinity purification of thcse CCT fusions is highly inefficient duc to a

largely inaccessible affinity tag. This problem was circumvented in 2006 when

Pappenberger et a]., (2006) described tlie cngineering of a calmodulili-bindi~ig peptide

(CBP) into a surface exposed loop of the Cct3p subunit (C'BP-C'CT3). This tag allowed

purification of functional yeast CCT for the first time, a reagent the authors used to

carefully analyze tlie folding kinetics of actin in vitro (Pappenberger et al., 2006).

In this study we use the synthetic genetic array (SGA) technique to assess the

genetic interaction network of the temperature sensitive c(V1-2 allele of CCT. %A

combines the allele of interest with all viable single deletions in the yeast genome and

scores tliese double mutants for fitness (Tong et a]., 2001). In this way we identified 72

lion-essential genes W ~ O S C deletion exacerbates the temperature sensitive phenotype of

cr t l -2 cells. At the same time our collaborator. Dr. Keith Willison, indicated that the

CBP-CCT3 fusion interacts with several septin ring subunits (personal communicat~on).

Based on these data we showed that CCT is likely to have a role in septin function

independent of its role in actin and tubulin folding. Together tliese types of analyses

provide a powerfill tool for generating new liypotlieses about CCT function in vivo and

should ultimately allow the identification of novel CCT substrates.

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5.3 Methods

5.3.1 Yeast strains and manipulations

Yeast strains used are listed in Table 5-1. Yeast plasmid transformations were

performed as described (Amberg et a]., 2005). The SGA experiments were performed

exactly as described three times and only those genes appearing in at least two screens

were included in the final list (Tong and Boone, 2006). Random spore analysis was done

as described (Tong and Boone, 2006) at 30•‹C, which is a normally permissive

temperature for cut 1-2.

5.3.2 Microscopy

GFP fusion plasmids containing CDC3 and CDC10 were a kind gift of Dr.

Christopher Beh. Cells were grown to log phase before shifting to the non-permissive

temperature, 15•‹C for cold-sensitive cells or 37•‹C for heat-sensitive cells, and grown for

16 or 4 hours respectively before live cell imaging on a Leica DM 6000 epifluorescence

microscopc with the appropriate fluorescence filter. Images were analyzed in Openlab

5.0.2 (Improvision). Cellular defects were scorcd manually and chi square analysis was

used to determine the significance of any differences from wildtype cells under the same

conditions.

5.4 Results and discussion

5.4.1 Synthetic genetic array of a temperature sensitive CCT allele

Genetic interactions can imply that the interacting genes work in the same, or

parallel, pathways in the cell to execute a common function. This information can be

extremely usefi.11 for placing a gene in a biochemical pathway or identifying redundancies

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it7 vivo. In recent years synthetic genetic array (SGA) technology has allowed the crcation

of extensive genetic interaction networks both on a genon~e-wide scalc (Davierwala et al.,

2005; Tong ct al., 200 I; Tong et al., 2004) and with respect to specific target proccsscs

(Drccs et al.. 200 1 ; Zhao et al.. 2005). The technical details of this procedure are

described at length in Tong and Boone (2006). The resultant data have becn cxtrernely

valuable for assigning functions to novel genes and enabling researchers to gencrate

testable l~ypotheses about their system of interest.

To try to expand the known scope of C'CT function in the cell we executed an

SGA scrcen using the ccfl-2 temperature sensitive allele to screen the viable deletion

collection. Interactions with 72 non-redundant gene deletions were validated by random

spore analysis and found to grow more slowly at 30•‹C (a normally pem~issive

temperature for ccfl-2) when in combination with the ccfl-2 allele (Table 5-2). Based on

the gene ontology (GO) annotation of the SGA genetic interactions appeared to be biased

toward cytoskeletal and chromatin remodeling functions (Table 5-2). The cytoskeleton-

related interactions were predicted because of the known effects of CCT on actin and

tubulin folding. We identified several prefoldin subunits (PACIO, YKE2, GIM-i), a

phosducin-like protein (PLPI) and the tubulin folding cofactors CIh'l (cofactor D) and

CIN2 (cofactor C) as genetic interactors of ccfl-2. We also identified downstream genes

which rcgulate actin during polarized ccll growth, cytokinesis and endocytosis (ARCIH,

BEMI, CLA4, pRKI and SLA I ; Table 5-2). Perhaps also expected to come along with

actin defccts wc identified some gcncs involved in vesicular transport and ccll wall

biogenesis (e.g. RUD3, ECM.33; sce Table 5-2).

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Thc bias towards chromatin remodeling coniponents may be a non-specific

phenomenon, indeed sonie of the genes identit'icd haw more than 100 known synthetic

genetic interaction!, (LGEI, BREI, CDC73, HTZI, SkVRI; Tong et a]., 2004). However,

the sheer number of chromatin-related interactions (25 of 72 or -35%) combined with the

knowledge that CCT likely assists the biogenesis of several histone deacetylase

complexes makes these currently tenuous connections an interesting area for future

investigation (Ciuenther, et a]., 2002; Pi-jnappel et a]., 2001). CCT has becn ascribed a

direct functional rolc in the folding of a n~anlnlalian SET3 histone deacetylase (HDAC)

homologue (Guenther, et a]., 2002) and in the biogenesis of two yeast HDAC's (Set3p and

Hos2p) (Pijnappel et a]., 200 1 ). I t may be that CCT dysfunction leads to misfolding of a

few HDAC components which in turn explains the large number of genetic interactions

between ccrl-2 and chroniatin modifying genes.

Interestingly, a collaborator, Dr. Keith Willison, identified physical interactions

with three core components of the septin ring (Cdc3p, Cdcl Op, CdcI 2p) and with a septin

kinase (Gin4p; K. Willison, personal comniunication). This data is supported by the

genetic interaction between cctl-2 and the septin kinase CLA4 which we may otherwise

have considered a non-specific effect of actin defects. Cla4p is a septin regulatoly kinase

involved in assembly of the septin filaments (Versele and Thorner, 2005). We chose to

examine the potential role for CCT in septin function in greater detail.

5.4.2 Analysis of septin function in yeast bearing mutant CCT subunits

The identification of physical interactions between C'C'T, three septin subunits

(Cdc3p, C'dc 1 Op, Cdc 12p) and one septin kinase (Gin4p), in conlbination with the genetic

interaction between L'ILI~A and cc.11-2 suggest a novel role for CCT in modulating septin

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assembly or function. The septin ring forms in the G 1 -phase as a patch at the incipient

bud site before expanding into a ring or disk during bud emergence (Vcrscle and Thorner,

2005). As the bud emerges septins re-organize into an hourglass-shaped collar around the

bud neck which is distributed cvenly between mother and daughter cells (Versele and

Thorner, 2005). During M-phase the septin collar splits, in a protein phosphatase-

dependant fashion, to form a pair of split rings before cytokinesis and septin disassembly

(Versele and Thorner. 2005).

To explore a possible role for CC'T in septin function we examined the

organization of Cdc3p- and Cdc lop-GFP fusions in cells bearing cold- and heat-sensitive

alleles of CCTI, CCT2, CCT3 and CCT4. The localization of Cdc3p and Cdc 1 Op is

normally confined to a single patch in unbuddcd cells, an hourglass-shaped ring in the

bud neck of small and large-budded cells and a split ring in very large-budded cells as

described above (Versele and Thorner, 2005). We found that the pattern of septin

localization was altered for both Cdc3p and Cdc10p in cold-sensitive (Y*/ I - I and in heat

sensitive c~4/4-1 cells (Figure 5-1 and Table 5-3). In budded cells the morphology of the

septin collar was found to be disrupted in a significant percentage of cc./I-I and cc.14-I

cells at the non-permissive temperatures (Table 5-3). Moreover, we saw aberrant cortical

GFP patches in the mutant cells, which were not found in wildtype (Figure 5-1A and

Table 5-3). Strangely. these phenotypes were also observed in the cold sensitive c.c/I-3

allele for the localization of Cdc10p but not Cdc3p (Table 5-3). Finally, in unbudded

cca/4-I cells we observed septin patches that seemed to be retained following cell division

while a new septin structure was assembled on the opposite side of the cell (Figure 5-1 B

and Table 5-4). In~portantly, all of these phenotypes were not observed in ccl2-4 or cc'/3-

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I alleles or in deletions of the CCT co-factors PLPI and the prefoldin subunit PAC'IO

(Table 5-4). Even cells lacking both PLPI and PAC10 had largely wild-type septin

localization although a small but significant percentage of septin collars wcre slightly

abnormal in these cells (Table 5-3).

I t is possible that the reason septin defects are only scen in some CCT mutant

strains relates to the relative severity of the alleles. If the ccfl-1 and c.c.14-1 alleles werc

more penetrant than the other alleles tested, they may be revealing a general septin defect

causcd by a dysfunctional CCT holo-complex. In support of the specific nature of the

defects we observed that the 1s growth defects in ccfl-2 cells was at least as severe as

those of c(8/4-/ cells (Figure 5-2). Also, c.c#/l-l and cc//-3 cells have similarly severe

growth defects at 25OC and all of the cs-alleles tested show strong growth defects when

grown at 20•‹C (Figure 5-2). Therefore at the non-permissive temperatures used in our

septin localization expcriments (15OC and 37•‹C') all of the 1s- and cs-alleles should have

shown defects if the septin mislocalization observed were a non-specific result of

abrogating CCT function.

Altogether the preceding data are consistent with the newly discovered physical

association of CCT with septins and suggest a direct role for the chaperonin in regulating

septin ring structure and function (Figure 5-1; K. Willison, personal communication).

Moreover, the fact that certain alleles of CCT and deletion of a PFD subunit do not affect

septins suggests that the defects i11 c r t 1-1 and (13/4- 1 cells are not secondary effects of

actin/tubulin dysfunction. Our data do not seem to suggest a role for CCT in septin

assenibly/folding because in imst of the defective cells the septins are still localized

approxiinately correctly at the bud neck. Instead, we see additional septin structures or

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misshapen septin rings in some CCT mutant alleles (Figure 5-1). Therefore. our data

point toward a role for CCT in the niaintenancc of proper scptin filament structure and

possibly a role in septin disassembly. This latter proposition stems largely from the

additional septin patches in unbudded ccf4-1 cells, shown in Figure 5-lB, which may

represent septin rings that failed to completely disasse~llble after the previous cytokinesis.

The presence of two such septin structures in unbudded cells is strikingly similar to the

phenotype of CDC3 mutants that block Cdc3p pl~osphorylation (Tang and Reed, 2002).

Cdc3p phosphorylation by the cell cycle kinasc Cdc28p, along with GI cyclins and likely

other factors, regulates septin disasse~nbly in early GI prior to the for~iiation of a new

septin cap at the presumptive bud site (Tang and Reed, 2002). If CCT were promoting

Cdc3p phosphorylation in G I it could explain some of the phenotypes we observed. In

support of this possibility, proteomic studies suggest CCT can interact with both Cdc28p

and C'dc3p (Ho et al., 2002; K. Willison personal conimunication) and several recent

studies suggest that it has a role in GI progression (Chapter 4, Stirling et a]., 2007,

accepted; Grantliani et a]., 2006: Yokota et a]., 1999). A highly speculative model could

be envisioned in which CCT could facilitate the interaction between Cdc28p and Cdc3p

in very early G 1 to promote septin disassembly by bringing the kinase (C'dc28p) to its

substrate (Cdc3p) .

5.5 Conclusion

Our knowledge of CC'T function in vivo is continually expanding beyond the

canonical roles in actin and tubulin biogenesis. These studies aimed to rapidly implicate

CCT in new cellular processes using a genomic approach. We identified 72 genes that

genetically i~iteract ~ ~ i t h a temperature sensitive CCTI allele. Integrating this data set

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with known literature revealed plausible roles for CCT in chroniatin remodelling and, as

expected. cytoskelctal function. In the casc of chromatin rcmodeling, CCT has becn

shown to assist the folding of some HDAC's (Gucnther et al.. 2002; P~jnappel et a].,

2001). I t must be noted that we identified many genetic interactions with c.c./I-2 not

relating to either chromatin or the cytoskeleton directly. Some of these interactions secm

to point to specific biological processes, for example NUPI 70, NUPISH, iMOGI and

NEiMI (Table 5-2) all relate to nuclear-cytoplasmic shuttling of proteins and RNAs. The

nicchanism behind these genetic interactions rcmains an interesting avenue of future

experiment.

The nun~ber of potentially interesting interactions with CCT is very high, even

within the published literature. For example, Sda I p is a nuclear protein involved in actin

filanient organization, cell cycle progression through G I and ribosonie biogencsis

(Buscemi et al., 2000; Dez et al., 2006); all phenotypes that could plausibly relatc to

CCT's roles in cytoskeletal and cell-cycle protein biogenesis. Moreover, Sda I p has also

recently been shown to physically associate with the Cct4p and Cct8p subunits (Krogan

et al., 2006). These phenotypic similarities and the physical interactions strongly suggest

a heretofore unrecognized relationship between the CCT complex and SdaI p that remains

to be explored.

We did verify that CC'T plays a role in organimtion of the septin ring in vivo. This

function seems to bc independent of the effects of CCT on actin and tubulin since

deletions of a PFD subunit or certain alleles of CCT, which do have actin and tubulin

defects, do not phenocopy the CCT mutants with septin defects (Table 5-3; Ckissler et

al.. 1998; Stirling et al., 2006; Ursic et al., 1994; Vinh and Drubin, 1994). Moreover, the

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physical interactions idcntified between CCT and CdcSp, Cdcl Op and Cdc l2p and the

allele specificity suggest a direct rolc for the C'C'T complex in septin finction. Wc

propose that CCT is involved in the maintenance andlor disasscrnbly of septin structures

because even in CCT mutants the septins are produced normally and localize to

approxin~ately the correct region of the cell. If CCT were required for folding septins

themselves wc would predict either rcduced an~ounts of GFP-septin being produced or an

amorphous aggregate to appear, which was not observed. Finally, the ability to express

properly folded septins in bactcria (unlike the critical CCT substrates actin and tubulin)

suggest that CCT is not crucial for dc. now folding of septin subunits (Versele et al.,

2004). Precisely where CCT is involved in septin organization or disassembly remains to

be elucidated. Interestingly, the yeast casein kinase homologues, Yck 1 p and Yck2p, are

known to affect septin biogenesis and are proposed to phosphorylate the Cct6p and Cct7p

subunits, respectively (Ptacek et al., 2005; Robinson et al,. 1999). The importance of

CCT phosphorylation for its iii vivo function is totally unknown but could help to regulate

the assembly of certain CC'T-interacting proteins such as thc septins. Indeed, examination

of casein kinase consensus phosphorylation sites in the amino acid sequence Cct6p and

Cct7p suggests that the modification is more likely to occur near or within the apical

domains and thus may be more likely to affect protein binding than ATP hydrolysis.

Together these studies create a number of novel avenues for future experiment

that should help to explain some of the in vivo functions of the CCT complex. Whether

the septins reprcscnt substrate proteins or proteins which CCT binds in a native state for

some unknown reason is currently unclear. Similarly the degree to which each genetic

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interaction can be considered more dircct or more indirect remains to bc determined by

specific experiments with each interactor.

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5.6 Figures

Figure 5-1 Septin localization in CCT mutant alleles

Localization of GFP-tagged septins in wildtype, c c ~ I - I (cold-sensitive) and c 'c~4-I (heat

sensitive) alleles. ( A ) Cdc3p-GFP localization in budded cells at permissive (30•‹C for

cr11- I and 25•‹C' for ~ ~ 1 4 - I ) and non-permissive temperatures ( 1 5•‹C' for cec/ I- I and 37•‹C'

for cc1.4-I). CCTI cells are shown at 30•‹C (permissive) and 37•‹C (non-permissive) but

were also normal at 25OC and 15•‹C (see Table 5-3)

Permissive

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(B) Localization of Cdc3p-GFP and Cdc lop-GFP in unbudded wildtype (DUYSSX) and

c c ~ 4 - I (DDY299) cells at 25•‹C (permissive temperature) and 37•‹C (non-permissive

temperature). For (A) and (B) representative images are shown of normal cells or of

defective cells where relevant. Scale bars indicate 10 pm.

Wildtype CnC3-GFP

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Figure 5-2 Relative growth defects of CCT-ts and c.s mutants

Heat sensitive (A) or cold-sensitive (B) C'CT mutant strains were grown to log phase.

scrially diluted, spotted on rich n~edia and grown for 48 hours or 72 hours in the case of

the 20•‹C' panel in (B).

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5.7 Tables

Table 5-1 Yeast strains used in this chapter

Strain # Genotype MLY 100 M L Y 1 10 M L Y I I I MLY l I8

M L Y 133

Pappenberger et a]., 2006 DUY55X DUY559

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Tablc 5-2 Verified genetic interactors of cctl-2 by synthetic genetic array (NOTE: Generated in collaboration with the group of Dr. Charles Boone)

YLROISW

YLRl l0C I--

I YFKOIYW

Y N L 153C'

BREI

C'INI

C'IN2

C%,4 4

C'TK I

D E P l

NOC'I

HOS2

Gene Function . .

transc~minasc a c t ~ v ~ t y

molecular function unknown

structural constituenl of cytoskeleton

ubiquitin-protein ligase activity

transcription regulalor activity

niolecular fi~nction i~nknown

RNA polymerase 11 lranscriplion eloneation factor activitv

beta-tubulin binding

molecular t'unction unknown

protein scrinelthreonine kinase activity . .

protcin kinase a c t ~ v ~ t y

transcription regulator activity

DNA replication origin binding

molecular function unknown

C'-8 sterol isomerase activity

1 -phosphatidylinositol-3-phosphate 5- kinase activity

I,3-beta-glucan synthase activity

FAD transnorler aclivilv

lubulin binding

histone deacetvlase activitv

histone cleacctylase activity

~ lpha - 1 ,h-~n;~~ino\ylll.;~nsferiibe activity

NAD-dependent litstone deacetyliw lctlvlty

Bio-process

biological proccss unknown

mRNA esport from nnclcus

actin filament organi~nlion

est;~blishment of cell polarity (sensu Fungi)

chromatin silencing at lelomere

telomere maintenance

cell wall organization and biogenesis

leloniere maintenance

post-choperonin tubulin folding pat hwny

microti~bule-based process

C'ytokinesis, apical bud growh, rho signaling

telornere ~naintcnance

telotnere maintenance

invasive growth (sensi~ Saccharomyces)

cell wall organi~ation and biogenesis

ergosterol biosynlhesis

response to stress

cell wall organization and biogenesis

FAD transnort

tubulin folding

regi~lalion of transcription, DNA-dependent

regu1;ition of transcription. DNA-dependent

cell wall mannoprotein biosynthesis

regulation of transcr~ption. DNA-dependent

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regulation of transcriplion from RNA nolvmer:ise I 1 nromoter chromatin hindine

positive regulation of lranscription from RSA polymerase I I promoters

Dublous ORF. o\ erlaps w ~ t h YO111 36u (IDH2)

cell wall organization and biogenesis

meiosis

protein import into nucleus

sporulation (sensu Fungi)

mRNA export from nucleus

mRNA exuort from nucleus

structural constituent of cell \ v d l

molecular function unknown

Ran CiTPase binding

molecular function imknown

structi~ral ~nolecule activity

structural molecule activitv

KREI

LGEI

1LIO G I

NEIMI

NUPI 7(

moleci~lar fimction unknown biolorrical orocess unknown

P,I C'l 0

PHO2.I

tubulin foldine

histone deacetvlase activitv chromatin modification

CiTPase inhibitor ac~ivitv beta-tubulin folding

protein amino acid phosphory lation urotein serinelthreonine kinase activ itv

Phosphatidylserine decarboxylase activily

protein binding

PSL) I

RC'YI

phosphatidylcholine hiosynthesis

endocy tosis

invasive growth (sensu Saccharomyces) rnolecular function i~nknown

cell wall biosynthesis (sensu Fungi) molecular function unknown

invasive growth (sensu Saccharornyces)

biological process unknown

molecular functwn unLnou n

Protein reqi~~red for sporulat~on

mitochondrial genomc maintenance DNA-directed RNA polymerase activity

RNA polymerase 11 transcription doneation hclor activilv lelomere maintenance

lranscription terminalion from Pol l l promoter. RNA aolymerase(A) coupled molecular fimction unknown

Eli to golgi vesicle-mediated Iransport molecular function ~~riknown YOR2 I OC'

~nvasive growth (sensu Saccharomy ces)

.elomere maintenance

;phingolipid metabolism

molecular function unknown

histone deacetylase activity

Sphingolipid alpha-hydroxylase

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histone lysine N-methyltrar~sli-rrise activilv (1-13-K4 snecific)

NAD-dependent histone deacelylase ;~ctivily

transcription corepressor activity

protein binding, bndging

phospliatidylinosibl-3A-bisphosphate binding

molecular function unknown

transcription cofiictor activity

transcription cohclor aclivily

histone lysine N-methyltransferasc activity ( M 3 - K 4 specific)

pl-otein-cysleine S- palmitoleyltransferase :ictiv~ty

helicase activity

protein transporter activity

ubiquitin-protein ligase activity

transcription corepressor activity

nucleosome binding

histone binding

tubulin binding

glucose-6-pI1osph:ite I -dehydrogenase activitv

niolecular tiinction unknown

Dubious O K F , overlaps with YDR456w (NIIXI)

molecular filnclion ilnlinown

chromatin silencing ul telo~nere

histone deacety lation

chromatin silencing at telomere

cell \vall organization and biogenesis

niicroautophagy

telomere maintenance

sporulafion (sensu Fungi)

histone acetylation

chromatin silencing :it telomerc

teloniere maintenance

protein amino x i d palmitoylation

chromatin rc~nodeling

protein target in,^ to membrane

protein monoubiquitination

telomere maintenance

lelomere maintenance

lelomere maintenance

protein folding

pentose phosphate shunt. oxidative branch

lelomere maintenance

biological process unknown

biological process unknown

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Table 5-4 Septin localization in unbudded cells

hannal Single Opposlng extra Total # Cell\ Strarn Genotype Temp. Patch patch analy~ed

C'DC3 C'DCIO CDC3 CDCIO CDC3 CDClO

MLY 100 \VT 3 0 100.0% 1 O0.O0/u 0.0'%, 0.0% 77 57 DUY55S ('('7'1 2 5 I . 93.8% 8.9% 6.2%) 79 8 1

37 95.4% 94.9% 4.6% 5.1% 65 SO DDY299 ~ ~ 1 4 - / 25 100.0% 100.0?/0 0.0% 0.0'%;, 54 95

37 80.7%' 79.7% 19.3% 20.3% 57 59 I Bold numbers indicate a p-value <0.01 relative to wildtype cells under the same conditions

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5.8 Reference list

Amberg, D.C., Burke, D.J. and Strathcrn. J.N. (2005). Methods in Yeast Genetics, 2005 edition. Cold Spring Harbor: Cold Spring Harbor Laboratory Press.

Buscemi, G., Saracino, F., Masnada, D., and Carbone, M.L. (2000). The S~/c~c~l?~/~.ot~;vc'c~~v c~ct.evi.ri~/c~ SDA 1 gene is recpired for actin cytoskeleton organization and cell cycle progression. J Cell Sci 1 13, 1 199- 12 1 1 .

Caniasses, A., Bogdanova, A., Shevchenko, A,, and Zachariae, W. (2003). Thc CCT chaperonin promotes activation of the anaphase-promoting complex through the generation of functional Cdc20. Mol Cell 12, 87- 100.

Davienvala, A.P., Haynes, J., Li, Z., Brost, R.L., Robinson, M.D.. Yu, L., Mnaimneli, S., Ding, H., Zhu. H.. Chen, Y.. et al. (2005). The synthetic genetic interaction spectrum of essential genes. Nat Genet 37, 1 147- 1 152.

Dez, C., Houseley, J., and Tollenfey, D. (2006). Surveillance of nuclear-restricted pre- ribosomes within a subnucleolar region of'S~~c~c~l?ut~otnj~c'es cct.evisicre. EMBO J 25, 1534- 1546.

Drees, B.L., Sundin, B., Brazeau, E., Caviston, J.P., Chen, G.C., Guo, W., Kozminski, K.G., Lau, M.W., Moskow, J.J., Tong, A.. et al. (2001). A protein interaction niap for cell polarity development. J Cell Biol 154, 549-57 1 .

Feldman, D.E., Tliulasiraman, V., Ferreyra, R.G., and Fryd~nan, J. (1 999). Formation of the VHL-elongin BC tumor suppressor complex is mediated by the chaperonin TRiC. Mol Cell 4, 105 1 - 106 1.

Gavin, A.C., Aloy, P., Grandi, P.. Krause, R., Boesche, M., Marzioch, M., Rau, C., Jensen, L.J., Bastuck, S., Dumpelfeld, B., et al. (2006). Proteome survey reveals modularity of the yeast cell machine~y. Nature 440. 63 1-636.

Geissler, S., Siegers, K., and Schiebel, E. (1 998). A novel protein coniplex promoting forniation of functional a- and y-tubulin. EMBO J 17, 952-966.

Grantham. J., Brackley, K.I., and Willison, K.R. (2006). Substantial CCT activity is required for cell cycle progression and cytoskeletal organization in ~iiammalian cells. Exp Cell Res 3 12, 2309-2324.

Guenther, M.G.. Yu, .I., Kao, G.D., Yen. T.J., and Lazar, M.A. (2002). Assembly of the SMRT-histone deacetylase 3 repression complex requires the TCP- 1 ring complex. Genes Dev 16, 3 130-3 135.

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Hartl, F.U., and Hayer-Hartl, M. (2002). Molecular chaperones in the cytosol: from nascent chain to folded protein. Science 295, 1852- 1858.

Ho, Y., Gruhler, A.. Heilbut, A., Bader, G.D., Moore, L., Adams, S.L., Millar, A., Taylor, P.. Bennett, K., Boutilier, K., et al. (2002). Systcinatic identification of protein conlplexes in S~~c.c. /~~i-onqw.s cwevisiw by mass spectrometry. Nature 4 15, 1 80- 183.

Krogan, N.J., Cagney, G., Yu, H., Zhong, G., Guo, X., Ignatchenko, A., Li, J., Pu, S.. Datta, N. , Tikuisis, A.P. et al. (2006). Global landscape of protein complexes in the yeast S ~ ~ c c h ~ ~ i w i ~ q r ' c . ~ C ' C I ~ V ~ S ~ N C . Nature 440. 637-43.

Lundin, V.F., Srayko, M., Hyman, A.A., and Leroux, M.R. (2007). Efficient chaperone- mediated tubulin folding is required for cell division and cell migration in C. e1egun.s. Submitted to Devclopmentul Biologs.

Pappenberger, G.. McCormack, E.A., and Willison, K.R. (2006). Quantitative actin folding reactions using yeast CCT purified via an internal tag in the C'C1T3/gamma subunit. J Mol Biol 360,484-496.

Pijnappel, W.W., Schaft, D., Roguev, A., Shevchenko, A., Tekotte, H.. Wilm, M., Rigaut, G., Seraphin, B., Aasland, R., and Stewart, A.F. (200 1). The S. cer.evi.sitrc SET3 coniplex includes two histone deacetylases, Hos2 and Hstl, and is a nieiotic- specific repressor of the sporulation gene program. Genes Dev 15, 299 1-3004.

Ptacek, J., Devgan, G., Michaud, G., Zhu, H., Zhu, X., Fasolo, J . , Guo, H., Jona, G., Breitkreutz, A., Sopko, R., et al. (2005). Global analysis of protein phosphorylation in yeast. Nature 438,679-684.

Robinson, L.C., Bradley, C., Bryan, J.D., Jerome, A., Kweon, Y., and Panek, H.R. (1 999). The Yck2 yeast casein I<inase 1 isoform shows cell cycle-specifk localization to sites of polarized growth and is required for proper septin organization. Mol Biol Cell 10, 1077- 1092.

Siegers, K., Bolter, B., Schwarz, J.P., Bottcher, U.M., Guha, S., and Hartl, F.U. (2003). TRiCICCT cooperates with different upstream chaperones in the folding of distinct protein classes. EMBO J 22, 5230-5240.

Sternlicht, H., Farr, G.W., Sternlicht, M.L., Driscoll, J.K., Willison, K., and Yaffe, M.B. (1993). The t-complex polypeptide 1 cornplex is a chaperonin for tubulin and actin in vivo. Proc Natl Acad Sci USA 90, 9422-9426.

Stirling, P.C., C'uellar, J., Alfaro, G.A., El Khadali, F., Beh, C.T., Valpuesta, J.M., Melki, R., and Leroux, M.R. (2006). PhLP3 niodulates CCT-mediated actin and tubulin folding via ternary conlplexes with substrates. J Biol Chem 28 1 . 70 12-702 1.

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Stirling, P.C., Srayko, M., Takhar, K.S., Pozniakovsky, A., Hyman, A.A., and Leroux. M.R. (2007). Phosducin-likc Protein 2 is Required for Multiple Functions of CCT. Accepted to ~Mo/~'cw/(r~- Bio/ogv o j /lie C'c.ll.

Tang, C.S., and Reed. S.I. (2002) Phosphorylation of the septin cdc3 in g I by thc cdc28 kinasc is cssential for efficient septin ring disassembly. Cell Cyclc 1, 42-49.

Tong. A.H., Evangelists, M., Parsons, A.B., Xu. H., Bader, G.D.. Page, N., Robinson. M., Raghibizadeh, S., Hogue, C.W., Busscy, H., et al. (2001). Systematic genetic analysis with ordered arrays of yeast deletion mutants. Science 294. 2364-2368.

Tong, A.H.. Lesage, G., Bader. G.D., Ding, H., Xu, H., Xin, X., Young, J., Berriz, G.F., Brost, R.L., Chang, M.. et al. (2004). Global mapping of thc yeast gcnetic interaction network. Scicnce 303, 808-8 13.

Tong, A.H., and Boone, C. (2006). Synthetic genetic array analysis in Strc~c~hcwoni)~c~c~.~ cwcwisicrc~. Methods Mol Biol 3 13. 17 1 - 192.

Ursic, D., Sedbrook. J.C., Himmel, K.L., and Culbertson, M.R. (1 994). The essential yeast Tcpl protein affects actin and microtubules. Mol Biol Cell 5, 1065- 1080.

Ursic, D., and Culbertson, M.R. (1 99 1). The yeast homolog to mouse Tcp- 1 affects microtubule-mediated processes. Mol Cell Biol 1 1 , 2629-2640.

Vcrsele, M., Gullbrand, B., Shulewitz, M.J, Cid, V.J., Bahmanyar, S., Chen, R.E., Barth, P., Alber, T., and Thorner, J. (2004). Protein-protein interactions governing septin heteropentamer assembly and septin filament organization in Scrc~chcrt-on~~~c~c~.~ cwVvisicre. Mol Biol Cell 15, 4568-4583.

Versele. M., and Thorner, J. (2005). Some asse~nbly required: yeast septins provide the instruction manual. Trends Cell Biol 15,4 14-424,

Vinh, D.B. and Drubin, D.G. (1 994). A yeast TCP- l -like protcin is requircd for actin function in vivo. Proc Natl Acad Sci USA 9 1 , 9 1 16-9 120.

Yokota, S., Yanagi, H., Yura, T., and Kubota, H. (1999). Cytosolic chaperonin is up- regulated during cell growth. Preferential expression and binding to tubulin at G(I)/S transition through early S phase. J Biol Chem 274, 37070-37078.

Zhao, R.. Davey, M., Hsu, Y.C., Kaplanek, P., Tong, A., Parsons, A.B., Krogan, N., Cagney, G., Mai, D., Greenblatt. J.. et al. (2005). Navigating the chaperone network: an integrative map of physical and genetic interactions mediated by the hsp9O chapcrone. Cell 120, 7 15-727.

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CHAPTER 6 GENERAL CONCLUSIONS

Molecular chaperones have an essential role to play in all cclls and understanding

their in vivo functions at a basic level is critical to our fundamental understanding of

cellular function. The findings presented in this thesis provide important new information

about the cellular roles of CCT and its cofactors. While the thesis is unified by our

interest in understanding the chaperonin CCT, the work can be broken into three discrcte

subcategories: First, to understand the substrate binding site and mechanism of the CCT

co-chaperone PFD; second, to understand the mechanism and in vivo significancc of CCT

regulation by phosducin-like proteins, particularly the PhLP2 and PhLP3 families; third

to expand the known cellular rolcs for C'CT beyond cytoskeletal protein folding.

Previous studies had implicated the distal half of the PFD coiled-coils as

important for substrate binding (Sicgert ct al., 2000). Our work (Chapter 2), idcntificd

the hydrophobic interhelical residues at the very distal tips of the coiled coils as the

substrate binding site (Lundin et a]., 2004). These residues are an inherent part of all

coiled coils and PFD mutants were able to retain partial chaperone function when we

replaced the tentacles with irrelevant coiled coils sequences. Whether these coils partially

unwind to expose a great surface area to substrates remains unknown. Our collaborative

efforts with Jose Valpuesta's group provided excellent structural support for our model of

substrate binding as well as significant new insights (Martin-benito et a].. 2007; Lundin et

a]., 2004). Archaeal PFD can alter its conformation to accon~n~odate substrates of

different shapes and sizes which would be predicted given the large size range of

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substrates (14-75kDa). Moreover, archaeal PFD appears to grip only part of the substrate:

leaving much of the lion-native protein protruding from its clamp-like cavity (Martin-

Bcnito et al., 2007). These lattcr two points are in stark contrast to eukaryotic PFD which

does not appear to alter its shape when bound to non-native actin and also co~nplctely

envelopes the actin, leaving very little or no part protruding from its cavity (Martin-

bcnito et a]., 2002). These findings are consistent with a highly specific hand-in-glovc

interaction between eukaryotic PFD and its substrates compared to the much more

promiscuous substrate-binding behaviour of archaeal PFD. All together this work

explores a novel mode of n~olecular recognition by hydrophobic sequences within coiled

coils and greatly improves our knowledge of substrate binding by archaeal PFDs. The

work also suggests that eukaryotic prefoldin has gained over its archaeal counterpart a

distinct evolutionary specialization in the manner i t interacts with and stabilizes actin, a

protein only present in eukalyotes (Martin-benito et a]., 2007).

While phosducin-like prote~ns had previously been shown to regulate CCT our

specifc aims were ( I ) to gain mechanistic information about how they regulated folding

by CCT, (2) to characterize the PhLPs which had yeast homologues to enable facile in

vivo assessment of PhLP function. The interaction between PliLP3 and CCT was initially

discovered as part of an in vitro expression screen for novel C'CT substrates (Appendix

I , Chapter 7.1). Once focussed on PhLP3 (Chapter 3) we showed that i t forms a

coniplex with C'CT in vivo and that, when present in excess, PhLP3 inhibits actin and

tubulin folding i r~ vitrv. This inhibition does not take place through direct competition

with substrate protein for CC'T binding, as had been suggested (McLaughlin et al., 2002).

Instead PhLP3 forms tcrnaiy cotnplexes with CC'T and substrate and slows the turnover

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of ATP to ADP in the context of this complex (Stirling et al., 2006). Whether PhLP3 acts

prior to hydrolysis or affects nucleotide exchange is uncertain; however, the effect on the

nucleotide cycle provides a rational explanation for why excess PhLP3 inhibits protein

folding in vitr-o. We showed that in ~*ivo , yeast PhLP3. PlpI p works with PFD to

modulate actin and tubulin biogenesis (Lacefield and Solomon, 2003; Stirling et a].,

2006). This latter data is supportcd strongly by unpublished synthetic gcnctic array

(SGA) data probing the genetic interactions of PLPI (Appendix 2, Chapter 7.2).

The other yeast PhLP homologue belongs to the class I 1 family and is called PLP2

in yeast (Flanary et a]., 2000; Blaauw et a]., 2003). We contirnied that PLP2 is an

essential gene and that the mammalian honiologue, PhLP2A, behaves much like PhLP3

i17 vitw (Chapter 4) . Collaborators showed that Plp2p binds CCT in vivo and generated

temperature sensitive alleles of PLPZ which we characterized in a number of ways.

Consistent with a role in modulating CCT, we found a variety of cytoskeletal defects i n

yIp2-t.r cells, including sensitivity to latrunculin and benomyl, mitotic spindle

misorientation, actin cable disruption and depolurization of actin patches (Stirling et al.,

2007, Accepted). We also identified high-copy suppressors ofplp2-1.1' alleles which

suggested a role in GI /S phase progression. This finding was supported by an observed

cell cycle delay in synchronized mutant cells as well as an accumulation of unbudded

cells in asynchronous populations (Stirling et a]., 2007, Accepted). Since CCT is known

to fold both regulators of the cytoskeleton and the cell cycle these data are entirely

supportive of a model in which Plp2p regulates the biogenesis of several CCT substrates,

the identity of which remains to be determined (Camasses et a]., 2003; Siegers et a].,

2003).

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Finally, my doctoral work aimed to idcntify novel cellular roles for the

chaperonin using yeast as a model system (Chapter 5). Wc undertook synthetic genetic

array analysis of one temperature sensitive CCT allele in collaboration with Dr. Charles

Boone's group. Meanwhile a collaborator, Dr. Keith Willison identified novel physical

interactions between CCT and septin subunits. In order to validate and explore these

findings we examincd septin localization in sevcral strains bcaring mutations in CC'T

subunits. Consistent with the results of our genomic experiments, CC'T alleles exhibited a

previously unidentified septin architecture defect (Chapter 5). This was not likely to be a

seconda~y effect of actin defects since PFD deletions and certain alleles of CCT did not

exhibit septin defects in spite of their known actin defects (Ursic et al., 1994: Stirling et

al., 2006; Vainberg et al., 1998; Vinh and Drubin, 1994).

In combination with recent literature, work presented here has helped define the

mechanism of archaeal PFD fiinction (Lundin et al., 2004; Okochi et al., 2004). We also

defined a novel mechanism for PhLP-mediated CCT regulation and suggested a CCT-

modulatory function for the essential PhLP homologue, PLP2 (Stirling et al.. 2006;

Stirling et a]., 2007, Accepted). Finally genomic approaches to understand CCT function

in the cell have suggested a role for the chaperonin in septin filnction and open up many

prospects for future research.

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6.1 Reference list

Blaauw. M., Knol, J.C., Kortholt, A., Roelofs, J., Ruchira, Postma, M., Visser. A.J., van Haastert, P.J. (2003). Phosducin-like proteins in L)ic/yos/eli~m~ u'iscoiu'wn~: implications for the phosducin family of proteins. EMBO J 22, 5047-5057.

Camasscs, A., Bogdanova, A., Slievchenko, A., and Zachariae, W. (2003). The CCT chaperonin promotes activation of the anaphase-promoting complex though the generation of functional Cdc20. Mol Cell 12, 87- 100.

Flanary, P.L., DiBello, P.R., Estrada, P., and Dohlman, H.G. (2000). Functional analysis of Plp 1 and Plp2, two homologues of phosducin in yeast. J Biol Chem 275, 18462- 18469.

Laccfield, S., and Solomon, F. (2003). A novel step in beta-tubulin folding is important for heterodimer formation in Strc~c.hcr~.o~?~j)~'c~~s cwrvisitrc. Genetics 165, 53 1-54 1 .

Lundin, V.F., Stirling, P.C., Gomez-Reino, J., Mwenifimbo, J.C., Obst, J.M., Valpuesta, J.M., and Leroux, M.R. (2004). Molecular clamp mechanism of substrate binding by hydrophobic coiled-coil residues of the archaeal chaperone prefoldin. Proc Natl Acad Sci USA 10 1,4367-4372.

Martin-Benito, J., Boskovic, J., Goniez-Puertas, P., Carrascosa, J. L., Sinions, C., Lewis, S. A., Bartolini, F., Cowan, N.C., and Valpuesta, J. M. (2002). Structure of eukaryotic prefoldin and of its complexes with unfoldcd actin and the cytvsolic chaperonin CCT. EMBO J 2 1,6377-6386.

Martin-Benito, J., Gomez-Reino, J., Stirling, P.C., Lundin, V.F., Gomez-Puertas, P., Boskovic, J., Chacon, P., Fernandez, .I.J., Berenguer, J., Leroux, M.R., and Valpuesta, J.M. (2007). Divergent Substrate-Binding Mechanisms Reveal an Evolutionary Specialization of Eukaryotic Prefoldin Compared to Its Archaeal Counterpart. Stn~cture 15, I0 I - 1 10.

McLaughlin, J. N., Thulin, C. D., Hart, S. J., Resing, K. A., Ahn, N. G., and Willardson, B. M. (2002). Regulatory interaction of phosducin-like protein with the cytosolic chaperonin complex. Proc Natl Acad Sci USA 99, 7962-7967.

Okochi, M., Nomura, T., Zako, T., Arakawa, T., Iizuka, R., Ueda, H., Funatsu, T., Leroux, M., and Yohda, M. (2004). Kinetics and binding sites for interaction of prefoldin with group 11 chaperonin: contiguous non-native substrate and chaperonin binding sites in archaeal prefoldin. J Biol Chenl 279, 3 1788-3 1795.

Siegers, K., Bolter, B.. Schwarz, J.P., Bottcher, U.M., Guha, S., and Hartl, F.U. (2003). TRiCICCT cooperates with different upstream chaperones in the folding of distinct protein classes. EMBO J 22, 5230-5240.

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Siegcst, R., Lcroux, M. R., Scheufler. C., Hartl, F. U., and Moarefi, I. (2000). Structure of the molecular chaperone prefoldin. Unique interaction of multiple coiled coil tentacles with unfolded proteins. Cell 103. 62 1-632.

Stisling, P.C., Cuellas, .I., Alfaro, G.A., El Khadali, F., Beh, C.T., Valpuesta, J.M., Melki, R., and Lesoux, M.R. (2006). PhLP3 modulates CCT-mediated actin and tubulin folding via ternary complexes with substrates. J Biol Chem 28 1 , 70 12-702 1.

Stirling, P.C., Srayko, M., Takhar, K.S., Pozniakovsky, A,, Hyman, A.A., and Lcroux, M.R. (2007). Phosducin-like Protein 2 is Required for Multiple Functions of CCT. Accepted to Molccul~r~ Biolog~, o/ /he Cell.

Ursic, D., Sedbrook, J.C'., Himmel, K.L., and Culbertson, M.R. (1994). The essential yeast Tcp 1 protein affects actin and microtubules. Mol Biol Cell 5, 1065- 1080.

Vainberg, I.E., Lewis, S.A., Romn~elacre, H., Ampe, C., Vandekerckhove, J., Klein, H.L., and C'owan, N.J. (1998). Prefoldin, a chaperone that delivers unfolded proteins to cytosolic chaperonin. Cell 93, 863-873.

Vinh, D.B. and Drubin, D.G. (1994). A yeast TC'P-I-like protein is required for actin function in vivo. Proc Natl Acad Sci USA '1 1. 9 1 16-9 120.

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CHAPTER 7 APPENDICES

7.1 Appendix 1 : An in vitro expression cloning screen for novel CCT- interacting proteins

Prior to identifying PhLP3 as a CCT-interacting protein I was in thc process of

executing an in vitro expression screen to identify novel CCT binding proteins being

produced in reticulocyte lysate. The approach involved translating individual cDNAs in

v i tm and looking for a native gel migration pattern similar to that of the known CCT

substrates actin and tubulin (Leroux, 2000). Initially I translated 134 cDNAs individually

in reticulocyte lysate and examined the migration of the radiolabelled protein products on

a native gel (Cowan, 1998; Leroux, 2000). This type of analysis is useful because known

CCT-substrate complexes have a discrete migration pattern on these gels, regardless of

the nature of the substrate. Some cDNAs were collected from members of our lab but the

DNAs werc pri~narily a kind gift of Dr. Takahiro Nagase at the Kazusa DNA research

institute in Japan (Kikuno et al., 2002). 1 also added PhLP2A and PhLP3 cDNAs to the

screen as I predicted that they would interact with CCT since, at the time, only PhLPl

was known to interact (McLaughlin et al., 2002).

The resultant screen yielded 9 proteins which strongly and reproducibly co-

migrated exactly with the position of CCT-actin or CC'T-tubulin co~nplexes suggesting

that they may be binding to CCT (Table 7-1). The screen also revealed a further 18

proteins which appeared to weakly co-migrate with CCT (Table 7-2). These latter I X

proteins had a variety of problems leading to uncertainty regarding their migration, either

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weak translation, a smearcd band or faint, not-always-reproducible banding around the

position of CCT.

These interactions, whether strong or suspect, needed validation and to this end

many were cloned for expression in tissue culhire. Our plan was to examine the co-

precipitation of our candidate proteins with CCT from cells and to assess their

localization in cells with RNA interference-induccd decreases in the levels of functional

CCT. However, we became aware of another group doing a larger scale vcrsion of our

screen that had already identified more than 80 polypeptides which co-precipitated

directly with CCT. This knowledge, combined with a growing interest in PhLP3 and

PhLP2A as modulators of CCT, led to dropping the screen as a primary investigation in

favour of the phosducin-like proteins (See Chapter 3 and 4). Whether these CCT co-

migrating proteins, other than PhLP2A and PhLP3. are truly CCT-interacting proteins is

not known and remains to be validated. One might speculate that the Kelch-repeat

proteins (Kelch-like Protein 1 and 4; Table 7-1 and 7-2) are good candidates because

they are structurally related to the WD repeats in GP subunits (Harashinla and Heitman,

2002). Unfortunately, many of the genes identified do not have clear yeast homologues

which may hamper detailed assessment of their requirement for CCT in the future.

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Table 7-1 Strong, reproducible CCT co-migrating proteins

Table 7-2 Less certain CCT co-migrating proteins

Accession #* Q9259S Q9COHh Q9UIF8 Q9BYZ6 Q92620 Q 14203 QXlY47 Q9LlM54 Q9UPY3 Q9Y2G2 Q 15034 Q 15027 P53992

**KIAA clone information can be accessed at \vwm~.kaz~~sa.or.ip/I~utle/

K I A A #** KIAA0201 KlAA1687 KIAA 1476 KIAA0717 KIAA0224

Q96Q07 Q9P286 Q9Y2A7 0 6 0 2 16 Q96RK4

Name MSP I05 Kelch like 4 Bromodomain adjaccnt to Zn finger domain 2 B Rho related BTB containing protein 2 PRP 16

KIAA0385 KIAA 1489 KIAA0389 KIAA0928 KlAA0055 KIAA0032 KlAA0050 KlAA0079

X-linked mental retardation candidate BTB and kclch domain containing protein Myosin6 Endoribon~~clease Dicer TLiCAN HER3 C'enlaurin Beta 1 -

Sec24C

*Query accession # at h t t ~ : / / w w w . n c b i . n l n ~ . n i h . ~ o v / e n t r e z / c i

KIAA I SXO K1AA 1264 KIAA0587 KIAA00721, N/ A

BTB/PO% domam containing protein 9 Serinelthreon~ne-protein kinase PAK 7 Nck associated protein 1 Hum;in rad2 1 homolog Bardet-Biedl syndron~e protein 4

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7.1.2 Appendix 1 reference list

Cowan, N.J. ( 1998). Manimalian cytosolic chaperonin. Methods Enzyniol 290, 230-241.

Harashinla, T., and Heitman, J. (2002). The Galpha protein Gpa2 controls yenst differentiation by interacting with kelch repcat proteins that niiniic Gbeta subunits, Mol Cell 10, 163- 173.

Kikuno, R., Nagase, T., Waki. M., and Ohara, 0. (2002). HUGE: a database for human large proteins identified in the Kazusa cDNA sequencing project. Nucleic Acids Res 30, 166- 168.

Leroux, M.R. (2000). Analysis of eukaryotic molecular chaperone coniplexes involved in actin folding. Methods Mol Biol 140, 195-206.

McLaughlin. J. N., Thulin, C. D., Hart, S. J., Resing, K. A., Ahn, N. G., and Willardson, B. M. (2002). Regulatory interaction of phosducin-like protein with the cytosolic chaperonin complex. Proc Natl Acad Sci USA 99, 7962-7967.

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7.2 Appendix 2: Synthetic genetic array ofp lp ld cells

Wc collaborated with the lab of Dr. Charles Boone (University of Toronto) to

execute a synthetic genetic array (SGA) screen oEylylA cells in an effort to gencrate a

list of genes which interacted genetically with PLPI. Because PLPI was not used as a

bait in any of the previous screens the only genetic interactions known previously were

with four of the six prefoldin subunits (Tong et al., 2004). The generation of an unbiased

list of genetic interactions with PLPI would help to understand the biological pathways

in which i t participates. Given the genetic interactions with prefoldin and our previous

data (Stirling et al., 2006; Chapter 3), we predicted a list enriched for cytoskeletal and/or

CCT-related genes.

The lab of Dr. Charles Boone executed the SGA screen as described (Tong and

Boone, 2006) and generated an electronically scored list of 95 unvalidatcd gcnes whose

deletions were deleterious in a ylplA background (Mat a cunIA::MFA/pr-HIS3 1 ~ y l A

11ru3A0 Icu2AO his3AI mc.tl5A11 ~~~IA::IVCI/IMX). I validated this list by random spore

analysis of the individual interactions and came up with 25 clear positives (Table 7-3).

Thc reniaindcr of the initial interactions were apparently false positives or interactions

which were too subtle to detect with random spore analysis and the human eye (Table 7-

3). This list may not be saturating but, as mentioned, the subtlety of some of the

interactions made confirmation difficult and this list is comprised of strong true positive

genetic interactions.

The interactions we identified are not surprising giving our previous tindings that

Plp I p modulates CCT and prefoldin mediated actin and tubulin folding (Lacefield and

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Solomon, 2003: Stirling et al., 2006). As such most of the interactions identified relate to

cell polarity, microtubule or actin function. vesicular transport and protein targeting. As

an internal control we found four of the previously known genetic interactions with

prefoldin subunits (GIIW. GlhI5, P,4CIO and YKE2; Tong et al., 2004; Stirling et a].,

2006). We also found direct reniodellers of actin and tubulin such as the Arp213

component ARCIS or the kinesin-associated protein CIKI . Further removed from these

direct cytoskeletal effectors we find genes such as the kinetochore component (NKP2)

and the endocytic regulator (RVS167) whose absence in combination with cytoskeletal

dcfects could lead to the obscrved sickness. Importantly, these interactions support the

notion that there really are subtle cytoskeletal biogenesis defects in plp1A cells that

bcconie apparent in certain gcnetic backgrounds, even though we failed to detect them in

previous work with the PLPI single deletion (Stirling et a]., 2006 Chapter 3).

These results came too late to be included in Stirling et al. (2006) and do not stand

alone as a publishable article. The data are included here only because they are highly

supportive of our findings in Chapters 3 and Chapter 5 and further extend our

understanding of Plp I p function in vivo.

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Table 7-3 Synthetic genetic interactions of pll)lA

Orf'Narne YLR370C'

YER155C'

Y FRO36 W YMRI98W

YPROI7C

YNL136W YDR3XSW

YNL 1 S3C

Y ML094W Y ERO92W YCiL236C YLR315W YKR082W

YCiR078C YGL023C'

Y P R I 9 l W

YCjR I X3C

YCiR25XC Y PRO43 W YCiL252C' YDR3XtlW - Y LR268W

YBRI71W

Y DL033C

Y LR200W

*[The data University of Toronto]

Gene Name ARC'IS

BEM2

C'DC'26 C'lh'l

DSS4

EAF7 EFT2

GI IW

G'l1tf.5

IES5 MTOI

iVl(r2 1VLlP133

PAC10 P I B

QC'R2

QC'R'I

R A D RPL43A RTG.2 RVSl67 SEC'ZZ

SEC66

SLAL?

YKE2

presented in this

Gene Function Structural constituent of cytoskeleton

Signal transducer activity

Protein binding Microtubule motor activity Guanyl-nucleotidc exchange factor activity

Molecular fi~nction unknown TI-anslation elongation factor activity

Tubulin binding. C'CT co-chaperone

Tubulin binding; C'C'T co-chaperone Molecular fimction u n k n o \ \ ~ ~ Molecular fi~nction unknown Molecular function unknown Structural molecule activity

Tubulin binding. CC'T co-chaperone Phosphaticiylinositol binding U bicluinol-cytochro~ne-c reductase activity LJbiquinol-cytochrome-c reductase nctivity

Single-stranded DNA specific endodeoxyribonuclease activity Stluchrral constituent of ribosome Transcription regulator activity C'ytoskeletal protein binding \;-SNARE activity

Protein lransporter activily t KNA (5-lnetby laminornethyl-2- thiouridylale)-1ne11.iyItr:11isferase activity

Tubulin binding list was generated in collaboration

Bio-process Actin filament organization

Cell wall organization and biogenesis

Ubiquitin-dependent protein catabolism Meiosis

Secretory pathway

Regulation of transcription from Pol 11 promoter Translational elongation Prefoldin complex, tubulin folding Prefoldin complex, tubulin folding Biological process unknown Protein biosynthesis Biological process unknown mRNA-nucleus export Prefoldin complex, tubulin folding Vesicle-mediated transport

Aerobic respiration

Aerobic respiration

Nucleotide-excision repair. DNA incision, 3 '40 lesion Protein biosynthesis Intracellular signaling cascade Endocytosis ER to Ciolgi transport

Posttranslational prolein- membrane targeting

Biological PI-ocess lrnknown Prefoldin complex, tubulin folding

with Dr. Charles Boone,

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7.2.2 Appendix 2 reference list

Lacefield, S., and Solomon, F. (2003). A novel step in beta-tubulin folding is important for heterodimer formation in S t r ~ d m w n i y m cwc.vivicre. Genetics 165, 53 1-54 1.

Stirling, P.C., C'uellar, J.. Alfaro, G.A., El Khadali, F., Heh, C.T.. Valpuesta, J.M., Melki. R., and Leroux, M.R. (2006). PhLP3 modulates CCT-mcdiated actin and tubulin folding via ternary complexes with substrates. J Biol Chem 281, 70 12-702 1 .

Tong, A.H., Lesage, G., Bader, G.D., Ding, H., Xu, H., Xin, X., Young, J., Berriz, G.F., Brost, R.L., Chang, M., et al. (2004). Global mapping of the yeast genetic interaction network. Science 303, 808-8 13.

Tong, A.H., and Boone, C. (2006). Synthetic genetic array analysis in S~rcc.h~~~-oi~~vc*e.v cewvisicre. Methods Mol Biol 3 13, 1 7 1 - 192.