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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 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
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
SIMON FRASER UNIVERSITYI i brary
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Simon Fraser University Library Burnaby, BC, Canada
Revised: Spring 2007
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
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.
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.
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 ~
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
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
.................................................. 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
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
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
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
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
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
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.
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 ;
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).
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.
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
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
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
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.
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
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
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
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
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
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
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
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.
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
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.
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).
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.
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
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
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
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.
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
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)
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
(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
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
(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
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
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
(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
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
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Siegers et al., 2003
Siegers et al., 2003
Siegers et al., 2003
Siegers et al., 2003
Siegers et al., 2003
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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.
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
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
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.
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.
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
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
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
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).
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
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-
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
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
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
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
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
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
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
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
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
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-
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
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
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
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
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).
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.
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
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
(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.
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
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 ~
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-
(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
Figure 2-2 continued
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
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)
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)
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.
(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
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 - + -
(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
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.]
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
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
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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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.
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
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),
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
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
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
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
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.
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
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
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,
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).
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
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
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).
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
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.
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,
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
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
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
~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
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
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
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
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.
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
(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
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%)
(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
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
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]
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
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)
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
(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
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
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
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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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).
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).
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
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
(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).
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
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
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
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
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.
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
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
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
( 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
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
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"-
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
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
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
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
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.
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
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.
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.
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.
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.
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
(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.
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
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
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
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.
(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
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.
(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 ~ ,
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
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
(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
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
(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.
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
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
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
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
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
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
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
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
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
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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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).
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.
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.
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
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).
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
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-
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
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
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
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
interaction can be considered more dircct or more indirect remains to bc determined by
specific experiments with each interactor.
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
(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
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).
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
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
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
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
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
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.
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.
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.
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
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
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).
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.
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.
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.
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
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.
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
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.
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
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.
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,
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.