Assembly and Substrate Recognition Properties of Human CCT
Subunits of the TRiC Chaperonin
by
Oksana A. Sergeeva
B.S. Chemistry/Biology
Harvey Mudd College, 2009
SUBMITTED TO THE DEPARTMENT OF BIOLOGY
IN PARTIAL FULFILLMENT OF THE REQUIREMENT
FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
AT THE
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
SEPTEMBER 2014
© Massachusetts Institute of Technology. All rights reserved.
Signature of Author: ____________________________________________________________
Department of Biology
May 2014
Certified by: __________________________________________________________________
Jonathan A. King
Professor of Molecular Biology, Thesis Supervisor
Accepted by: _________________________________________________________________
Amy E. Keating
Co-chair, Department Committee on Graduate Students
1
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Assembly and Substrate Recognition Properties of Human CCT
Subunits of the TRiC Chaperonin
by
Oksana A. Sergeeva
Submitted to the Department of Biology
at the Massachusetts Institute of Technology
on May 30th, 2014 in the Partial Fulfillment of the
Requirements for the Degree of
Doctor of Philosophy in Biology
ABSTRACT
Group II chaperonins are large multi-subunit complexes that fold cytosolic proteins to
their native structures. They are composed of two back-to-back rings of 7-9 subunits. The
eukaryotic cytosolic type II chaperonin Tailless Complex Polypeptide-1 (TCP-1) Ring Complex
(TRiC) consists of eight different subunits identified as Chaperonin Containing TCP-1 (CCT) α
(1) - θ (8). TRiC is necessary for folding about 10% of newly synthesized proteins and is
essential for folding actin and tubulin. Most of the research on TRiC in the last 20 years has
focused on yeast and bovine TRiC. However, recently, there has been inquiry into TRiC as a
target for disease therapy for Huntington’s disease, cataract, and some cancers. Consequently,
to understand human TRiC, we purified endogenous TRiC from HeLa cells for characterization.
These complexes contained all eight of the CCT subunits as determined by immunoblot. The
structures were well organized as double-rings of eight subunits each, using negative stain
electron microscopy (EM). Human TRiC was active in suppressing aggregation and refolding
two different substrates: luciferase (a model substrate) and human γD-crystallin (HγD-Crys; a
physiological substrate found in the eye lens).
To further understand human TRiC, we expressed all of the human CCT subunits, one
at a time in E. coli. This was done so that the subunit specificities of the CCT subunits could be
studied and so we could have a system where these proteins could be genetically manipulated.
Theoretically, all eight subunits in the mature TRiC-complex are needed to successfully
recognize all substrates that need to be folded in the cell. We found that two CCT subunits,
CCT4 and CCT5, but not the others, formed TRiC-like homo-oligomeric rings in the absence of
the other CCT subunits. Purification of these complexes and subsequent structural assays by
negative stain and cryo-EM showed that they formed double rings of eight subunits per ring.
Biochemically, we found that CCT4 and CCT5 hydrolyzed ATP at the same rate as human
TRiC, could refold luciferase to the same level as human TRiC, and suppressed aggregation of
HγD-Crys. The homo-oligomeric complexes also assisted the refolding of HγD-Crys, a property
not observed in the lens specific α-crystallin chaperone. On the substrates studied, CCT4 and
CCT5 homo-oligomers worked as efficiently as hetero-oligomeric TRiC. More stringent
substrates such as actin and tubulin need to be studied to further understand CCT specificity.
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Two mutations, one in CCT4 (C450Y) and one in CCT5 (H147R), have been implicated
in hereditary sensory neuropathy. In order to study the defective mutant proteins, we introduced
these mutations into the CCT4 and CCT5 constructs. We found that for CCT4, the newly
translated mutant polypeptide chains aggregated much more than wild-type (WT) CCT4. While
the mutant formed some rings in the E. coli lysate, as assayed by sucrose ultracentrifugation
gradients and negative stain EM, they were not stable throughout the purification and the final
purified sample contained few homo-oligomers. The mutant CCT5 polypeptide chains were
properly folded and assembled in homo-oligomers. H147R CCT5 was able to hydrolyze ATP at
a similar rate as WT CCT5. However, in the HγD-Crys aggregation suppression and refolding
assay, mutant huntingtin aggregation suppression assay, and actin refolding assay, mutant
CCT5 was not as efficient in suppression or refolding as WT CCT5. Therefore, the H147R
mutation in CCT5 led to a chaperoning defect while the C450Y mutation in CCT4 led to a
folding/stability defect.
In order to understand features of partially folded intermediates that group II chaperonins
recognize in a substrate, we investigated whether the archaeal group II chaperonin from
Methanococcus maripaludis (Mm-Cpn) could recognize a variety of HγD-Crys mutants. These
mutations were in regions of the protein that could act as recognition signals of substrate –
unpaired aromatics, domain interface, and buried core residues. We found that Mm-Cpn was
able to recognize all of these mutants, better than it recognized WT HγD-Crys. In addition, MmCpn could refold most of the mutants to levels higher than WT HγD-Crys. Therefore, we
concluded that Mm-Cpn doesn’t recognize any of the proposed recognition signals but
recognizes some β-sheet interface exposed in these mutants.
These studies further our knowledge of group II chaperonins and specifically human
TRiC, and open up some new avenues for the investigation of the folding, assembly and
function of this eukaryotic protein essential for the reproduction of all cells.
Thesis Supervisor: Jonathan A. King
Title: Professor of Molecular Biology
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ACKNOWLEDGEMENTS
There are many people who have helped me get to this point of finally writing up this
thesis and graduating. First and foremost, I want to thank my advisor Jonathan King. I definitely
could not have made it through the last four years without his support, encouragement, and
guidance. I have learned a great deal from him as a scientist, mentor, and person. Jon, thank
you for believing in me when I did not and supporting me through everything.
I need to thank all of the people in the King lab that I have interacted with and learned
from in the last four or five years: Ligia Acosta-Sampson, Jeannie Chew, Dan Goulet, Cammie
Haase-Pettingell, Althea Hill, Fan Kong, Kelly Knee, Kate Moreau, Jacqueline Piret, Liliana
Quintinar, Dessy Raytcheva, Nathaniel Schafheimer, Meme Tran, Takumi Takata, Cindy
Wooley, and Ginger Yang. Cammie: thank you for helping me in every random way you could:
running sucrose gradient and gels, making buffers (and pH-ing everything – what will I do
without you?), growing cells, and taking care of things when I had to run off to class. Cindy:
thank you for always being there to chat about random current events and taking care of all the
financial/grant issues. You both have made my life in lab less stressful and more entertaining.
Kate, Kelly, and Dessy: I am so glad that I met each one of you in lab and that we’re still friends
years later. I learned so much from each one of you in lab and life, and I know you all will go on
to great things. Nathaniel: you have kept me sane and grounded for the last few years. I
thoroughly enjoyed our lunches, venting sessions, and good times in lab. Thank you for always
being there for me and making me feel less alone. Meme: you were the best undergrad anyone
could ever ask for. Thank you for being hard working, amazing, and a joy to be around. I think
you have a bright future ahead of you! Eugene: I feel like I not only failed you with everything I
forgot to teach you, but also because I am leaving you all alone. However, I know you will be
just fine and go on to awesome things!
I would like to thank my committee: Amy Keating, Thomas Schwartz, and Susan
Lindquist. They have attended many meetings and offered valuable advice on my project, my
progress in the program, and my future. I especially want to thank Amy (who I also TAed for) for
always being encouraging and looking out for me and my training as a scientist. I also want to
acknowledge Frank Solomon for giving me advice and taking an interest in my project.
Next, I need to thank Wah Chiu. I am so lucky to have been part of the Nanomedicine
consortium and interacted with Wah and his lab over the last few years. He has always been
excited about my projects and about moving research along. Wah: thank you for letting me
come to BCM so often and giving me the opportunity to learn about cryo-EM from both the
technical and computational sides. At BCM, there are many colleagues I would like to
acknoweldge: Steve Lutdke, David Tweardy, Sarah Shahomoradian, Bo Chen, Michele Darrow,
Corey Hecksel, Rebecca Dillard, Soung-Hun Roh, Moses Kasembeli, Yao Cong, and Zhao
Wang. Thank you for putting up with all of my questions and helping out on all of my many cryoEM projects. The Nanomedicine consortium has also given me the opportunity to interact with
many professors and students in the chaperonin field. I would like to specifically thank Bill
Mobley, David Housman, and Chengbiao Wu for stimulating conversations; and, Koning Shen,
Tom Lopez, Em Sontag, Ryan McAndrew, and Henrique Pereira for sharing reagents, protocols,
and expertise in the chaperonin field. In addition, I want to acknowledge Zach Crook, the only
other grad student from MIT in our Nanomedicine group. Zach: thank you for answering all of
my huntingtin questions and giving me great feedback on all of my work and concerns. You are
one of the most hard-working people I know.
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I would like to thank my neighbors across the hall: the Gilbert lab. Boris, MK, Josh,
Thomas, Kristen, Pavan, Audra, Maria, Julia, and Wendy: thank you for always letting me
bother you and hang out in Starbucks+ with you. I have always been amazed by how well you
get along and how fun your lab atmosphere is. To my neighbors in the same hall, the Sinskey
lab (JQ, Jingnan, Chris, Tony, and Claudia): thanks for being so welcoming to me, inviting me to
Friday wine and cheese, and always having that random chemical I didn’t think actually existed.
I need to acknowledge my biology classmates in the entering class of 2009 for being an
exceptional group and somehow still remaining friends so many years later. I am so glad our
class data clubs, building 68 lunches, and random Muddy hang-outs are still going on. More
specifically, I would like to thank Paul Fields, Glenna Foight, Genny Gould, Heather Hoke, David
Kern, Katie Mattaini, and Boris Zinshteyn for all our coffee dates, trivia nights, knitting & game
nights, and in general for keeping me sane. You are all amazingly intelligent and dedicated, and
have made my last five years fun and engaging. I also want to acknowledge MIT classmates
outside of the biology department, in particular: Aimee Gillespie and Bridget Wall. Thanks for all
of the road trips, coffee dates, articles clubs, and dinners. I am incredibly grateful we met and
have become great friends! To my college friends in Boston/Cambridge (Nadia Abuelezam,
Hallie Kuhn, Christina Snyder, Terence Wong, Ken Loh, and Trevor Ashley): thank you for
making this transition to grad school easier, always being supportive, and strengthening our
friendship over these years. Finally, to my BFFs from middle school (Olga Obraztsova, Sakina
Palida, and Mika Wilbur): thanks so much for being there for me all of these years in every way.
I have enjoyed all of our random traveling to see each other, and all the letters, postcards, and
phone calls. I can’t believe we’ve been friends for more than half our lifetimes, and I can’t wait to
see what the next decade or two will bring for us all!
Last, but not least, I would like to thank my family. All four of my grandparents and both
my parents received PhDs in the sciences, so they have always encouraged me to pursue my
education. I thank them for instilling a love for science and learning in me. A big thank you (even
though I’m sure he doesn't want it) goes to my brother, Ivan. He started MIT as an undergrad a
year before I got here, and we had a great time both living in Cambridge and hanging out
together for the four years we overlapped. He was the one who gave me the courage to move
across the country (into the cold) to go to grad school here. Finally, I want to thank my husband,
Nate Jones and our son Chase. Nate somehow endured all of my grad school frustrations and
still respected me as a scientist and a person. Nate: thank you for always believing in me and
always being on my side. I can’t imagine these last five years of my life without you. Chase:
thanks for giving me the easiest pregnancy ever. You better be cute! [Edit: You are super cute
and the easiest baby! Thanks for your continued cooperation in letting me finish this thesis!]
This research was supported by National Institutes of Health grants (R01EY015834,
P41GM103832, and Common Fund Roadmap PN2EY016525). The Biophysical Instrumentation
Facility for the Study of Complex Macromolecular Systems (NSF-007031) is gratefully
acknowledged.
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BIOGRAPHICAL NOTE
EDUCATION
Ph.D.
Massachusetts Institute of Technology, Cambridge, MA
Expected 2014
Department of Biology
Chemical Biology Interface Training Program Student
B.S.
Harvey Mudd College, Claremont, CA
May 2009
Joint Chemistry/Biology with Psychology minor
Graduated with Distinction and Biology Department Honors
RESEARCH EXPERIENCE
2010-2014
Graduate Research Assistant with Professor Jonathan King
Biology Department, MIT, Cambridge, MA
2005-2009
Undergraduate Researcher with Professor David Asai
Biology Department, Harvey Mudd College, Claremont, CA
Summer 2008
SURF Student with Professor Seth Darst
Biophysics Department, Rockefeller University, New York, NY
Summer 2007
SURF Student with Professor Paul Insel
Pharmacology Dept., U. of California, San Diego, La Jolla, CA
Summer 2006
REU Student with Professors Bogdan Olenyuk and Katrina Miranda
Chemistry Department, University of Arizona, Tucson, AZ
2001-2004
NewBiotics, Inc. and ThioPharma, Inc., San Diego, CA
TEACHING EXPERIENCE
Spring 2013
Teaching Assistant, MIT, Cambridge, MA
7.41: Topics in Chemical Biology
Spring 2011
Teaching Assistant, MIT, Cambridge, MA
7.10/20.111: Physical Chemistry of Biomolecular Systems
Fall 2008
Teaching Assistant, Harvey Mudd College, Claremont, CA
Chem 24: Chemistry Laboratory
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PUBLICATIONS
Sergeeva OA, Tran MT, Haase-Pettingell C, King JA (2014). “Biochemical Characterization of
Mutants in Chaperonin Proteins CCT4 and CCT5 Associated with Hereditary Sensory
Neuropathy.” J. Biol. Chem. Submitted.
Sergeeva OA, Yang J, King JA, Knee KM (2014). “Group II archaeal chaperonin recognition of
partially folded human γD-crystallin mutants.” Protein Sci. 23: 693-702.
Sergeeva OA, Chen B, Haase-Pettingell C, Lutdke SJ, Chiu W, King JA (2013). “Human CCT4
and CCT5 chaperonin subunits expressed in E. coli form biologically active homo-oligomers.” J.
Biol. Chem. 288:17734-17744.
Knee KM, Sergeeva OA, King JA (2013). “Human TRiC complex purified from HeLa cells
contains all eight CCT subunits and is active in vitro.” Cell Stress and Chaperones 18:137-144.
Wilkes DE, et al. (2009). “Identification and characterization of dynein genes in Tetrahymena.”
Methods Cell Biol. 92:11-30.
Sergeeva OA, Khambatta HG, Cathers BE, Sergeeva MV (2003). “Kinetic properties of human
thymidylate synthase, an anticancer drug target.” Biochem. Biophys. Res. Commun. 307:297300.
8
TABLE OF CONTENTS
PREFATORY MATERIAL
Cover Page ................................................................................................................................... 1 Abstract ......................................................................................................................................... 3 Acknowledgements ....................................................................................................................... 5 Biographical Note .......................................................................................................................... 7 Table of Contents .......................................................................................................................... 9 List of Figures .............................................................................................................................. 13 List of Tables ............................................................................................................................... 15 List of Abbreviations .................................................................................................................... 17 CHAPTER 1:
Introduction
Protein Folding and Aggregation ................................................................................................. 20 ATP-Dependent Chaperones ...................................................................................................... 22 Hsp70 ...................................................................................................................................... 22 Hsp90 ...................................................................................................................................... 25 Chaperonins ............................................................................................................................ 27 Group I Chaperonins ................................................................................................................... 28 History ..................................................................................................................................... 28 Structure and Function ............................................................................................................ 31 Group II Chaperonins .................................................................................................................. 34 History ..................................................................................................................................... 34 Structure and Function ............................................................................................................ 37 Substrate Recognition by Group II Chaperonins ......................................................................... 39 Chaperonin Subunits/Domains Involved in Recognition ......................................................... 39 Features of the Substrate Recognized .................................................................................... 42 Chaperonin Complex Evolution ................................................................................................... 43 Homo-oligomeric Chaperonins ................................................................................................ 43 Hetero-oligomeric Chaperonins .............................................................................................. 43 Arrangement of CCT Subunits in TRiC ................................................................................... 43 Role of Chaperonins in Human Disease ..................................................................................... 46 Mutations in Human Chaperonin Genes ................................................................................. 46 Using TRiC to Ameliorate Diseases ........................................................................................ 46 Thesis Context ............................................................................................................................ 49 CHAPTER 2:
Human TRiC Complex Purified from HeLa Cells Contains All Eight CCT Subunits and is
Active In Vitro
Abstract ....................................................................................................................................... 52 Introduction ................................................................................................................................. 53 9
Materials and Methods ................................................................................................................ 55 TRiC Purification from HeLa Cells .......................................................................................... 55 SDS-PAGE and Immunoblots ................................................................................................. 56 Electron Microscopy ................................................................................................................ 56 Luciferase Refolding Assay ..................................................................................................... 56 Human γD-Crystallin Aggregation Suppression Assay ........................................................... 57 Results ........................................................................................................................................ 58 Purification .............................................................................................................................. 58 Structure .................................................................................................................................. 58 Activity ..................................................................................................................................... 64 Discussion ................................................................................................................................... 67 CHAPTER 3:
Human CCT4 and CCT5 Chaperonin Subunits Expressed in E. coli Form Biologically
Active Homo-oligomers Abstract ....................................................................................................................................... 70 Introduction ................................................................................................................................. 71 Materials and Methods ................................................................................................................ 73 CCT Subunit Expression ......................................................................................................... 73 CCT Subunit Purification ......................................................................................................... 73 Human TRiC and Mm-Cpn Purification ................................................................................... 74 Sucrose Gradient Sedimentation ............................................................................................ 74 SDS-PAGE and Immunoblots ................................................................................................. 74 Electron Microscopy ................................................................................................................ 74 Cryo-Electron Microscopy ....................................................................................................... 75 Thermal Denaturation by Circular Dichroism .......................................................................... 76 ATP Hydrolysis Assay ............................................................................................................. 76 Luciferase and Human γD-Crystallin Refolding Assays .......................................................... 76 Results ........................................................................................................................................ 78 Expression and Purification of CCT Subunits ......................................................................... 78 Structural Characterization of the CCT4 and CCT5 Homo-oligomers .................................... 83 Functional Characterization of the CCT4 and CCT5 Homo-oligomers ................................... 88 Discussion ................................................................................................................................... 95 CHAPTER 4:
Biochemical Characterization of Mutants in Chaperonin Proteins CCT4 and CCT5
Associated with Hereditary Sensory Neuropathy
Abstract ....................................................................................................................................... 98 Introduction ................................................................................................................................. 99 Materials and Methods .............................................................................................................. 104 Mutagenesis and Expression ................................................................................................ 104 Long-term Lysate Supernatant/Pellet Separation ................................................................. 104 Sucrose Gradient Sedimentation .......................................................................................... 104 SDS-PAGE and Immunoblots ............................................................................................... 104 CCT Subunit Purification ....................................................................................................... 105 10
Electron Microscopy and Circular Dichroism ........................................................................ 105 Native Gel Electrophoresis .................................................................................................... 106 ATP Hydrolysis and Human γD-Crystallin Refolding Assays ................................................ 106 Mutant Huntingtin Aggregation Suppression Assay .............................................................. 106 Actin Refolding Assay ........................................................................................................... 106 Results ...................................................................................................................................... 108 Mutant Protein Expression and Stability ............................................................................... 108 Mutant Protein Sedimentation ............................................................................................... 110 Mutant Protein Purification .................................................................................................... 113 Mutant Protein Structure ....................................................................................................... 115 CCT5 Mutant Activity ............................................................................................................ 119 Discussion ................................................................................................................................. 128 CHAPTER 5:
Group II Archaeal Chaperonin Recognition of Partially Folded Human γD-Crystallin
Mutants
Abstract ..................................................................................................................................... 132 Introduction ............................................................................................................................... 133 Materials & Methods ................................................................................................................. 136 Purification of HγD-Crys and Mm-Cpn .................................................................................. 136 Thermal Denaturation by Circular Dichroism ........................................................................ 136 Aggregation Suppression of HγD-Crys by Mm-Cpn .............................................................. 137 Quantification of Refolded HγD-Crys .................................................................................... 137 Results ...................................................................................................................................... 138 Buried Aromatic Pairs ........................................................................................................... 138 Domain Interface Residues ................................................................................................... 142 Buried Core Hydrophobic Mutants ........................................................................................ 146 Discussion ................................................................................................................................. 150 CHAPTER 6:
Final Discussion and Future Directions
Final Discussion ........................................................................................................................ 154 Future Directions ....................................................................................................................... 159 CHAPTER 7:
References ............................................................................................................................... 161 CHAPTER 8:
APPENDIX A:
Co-expression of CCT Subunits to Explore Subunit Assembly
Abstract ..................................................................................................................................... 184 Introduction ............................................................................................................................... 185 Materials and Methods .............................................................................................................. 189 Plasmid Construction ............................................................................................................ 189 Expression and Lysis ............................................................................................................ 189 11
Sucrose Gradient Sedimentation .......................................................................................... 189 SDS-PAGE and Immunoblots ............................................................................................... 189 Quantification ........................................................................................................................ 190 Results ...................................................................................................................................... 192 Summary of Each CCT Profile .............................................................................................. 195 Effect of Homo-oligomers on Full-length CCT Subunits and Their Fragments ..................... 199 Discussion ................................................................................................................................. 204 APPENDIX B:
Aggregation Suppression of Mutant Huntingtin by Chaperonins
Abstract ..................................................................................................................................... 208 Introduction ............................................................................................................................... 209 Materials and Methods .............................................................................................................. 211 Mutant Huntingtin Aggregation Suppression Assay .............................................................. 211 Results & Discussion ................................................................................................................ 212 12
LIST OF FIGURES
Figure 1-1: Effect of chaperones on protein folding and aggregation energy landscape ........... 21 Figure 1-2: Hsp70 structure and states ...................................................................................... 24 Figure 1-3: Hsp90 structure and states ...................................................................................... 26 Figure 1-4: Group I chaperonin structure and mechanism ......................................................... 33 Figure 1-5: Group II chaperonin structure and mechanism ........................................................ 38 Figure 1-6: Alignment of apical domains of CCT subunits ......................................................... 40 Figure 1-7: TRiC subunit arrangement differs between laboratories and methods .................... 45
Figure 2-1: Human TRiC primarily limited to the cytoplasmic fraction of HeLa cells .................. 59 Figure 2-2: Human TRiC purified by size exclusion chromatography ........................................ 60 Figure 2-3: Hsp70 and Hsp90 co-purified with TRiC when heparin affinity chromatography was
omitted ........................................................................................................................................ 61 Figure 2-4: All eight subunits present in purified human TRiC ................................................... 62 Figure 2-5: Negative stain TEM of purified human TRiC reveal double rings ............................ 63 Figure 2-6: Purified human TRiC active in refolding luciferase .................................................. 65 Figure 2-7: Purified human TRiC suppression of HγD-Crys aggregation and HγD-Crys nativelike state refolding ....................................................................................................................... 66
Figure 3-1: Expression of human CCT subunits in BL21 (DE3) RIL E. coli cells ....................... 79 Figure 3-2: Sucrose ultracentrifugation gradients of CCT subunits ........................................... 81 Figure 3-3: CCT5 purified by size exclusion chromatography as a 1 MDa complex.................. 82 Figure 3-4: Negative stain TEM of purified CCT4 and CCT5 homo-oligomers showed
morphology similar to human TRiC, and distinct from GroEL/ES ............................................... 84 Figure 3-5: Raw cryo-EM images of CCT5 homo-oligomers and 2D class averages indicated
two rings of eight subunits per ring ............................................................................................. 85 Figure 3-6: Cryo-EM reconstructions of CCT5 homo-oligomers suggested TRiC-like structures
.................................................................................................................................................... 87 Figure 3-7: Human TRiC is more stable than CCT4 and CCT5 homo-oligomers by thermal
denaturation using CD ................................................................................................................ 89 Figure 3-8: CCT4 and CCT5 homo-oligomers hydrolyze ATP at a similar rate to human TRiC 90 Figure 3-9: CCT4 and CCT5 homo-oligomers were active in refolding luciferase ..................... 92 Figure 3-10: CCT4 and CCT5 homo-oligomers suppressed aggregation of partially folded HγDCrys and promoted HγD-Crys native-like state refolding ............................................................ 94
Figure 4-1: Location of neuropathy mutations in CCT4 and CCT5 .......................................... 102 Figure 4-2: Expression levels of CCT4, CCT5, and their neuropathy mutants ........................ 109 Figure 4-3: Long-term lysate incubation of CCT4, CCT5, and their neuropathy mutants ........ 111 13
Figure 4-4: Sucrose ultracentrifugation gradients of CCT4, CCT5, and their neuropathy mutants
.................................................................................................................................................. 112 Figure 4-5: CCT4 and CCT5 purification off of the Co-NTA column ........................................ 114 Figure 4-6: Negative stain transmission electron micrographs of CCT4, CCT5, and their
neuropathy mutants .................................................................................................................. 116 Figure 4-7: Native gel electrophoresis of CCT5 and its neuropathy mutant ............................ 117 Figure 4-8: Far-UV circular dichroism scans and thermal denaturation of CCT5 and its
neuropathy mutant .................................................................................................................... 118 Figure 4-9: ATP hydrolysis of CCT5 and its neuropathy mutant .............................................. 120 Figure 4-10: Aggregation suppression of HγD-Crys by CCT5 and its neuropathy mutant....... 121 Figure 4-11: SDS-PAGE and quantification of HγD-Crys refolded by CCT5 and its neuropathy
mutant ....................................................................................................................................... 123 Figure 4-12: Mutant huntingtin aggregation suppression by CCT5 and its neuropathy mutant
.................................................................................................................................................. 125 Figure 4-13: Quantification of β-actin refolded by CCT5 and its neuropathy mutant ............... 126 Figure 4-14: Variations in protein concentration and ionic strength of β-actin refolded by CCT5
and its neuropathy mutant......................................................................................................... 127
Figure 5-1: HγD-Crys mutants chosen fall into three sets ........................................................ 135 Figure 5-2: HγD-Crys aromatic pair mutants suppressed by Mm-Cpn .................................... 140 Figure 5-3: HγD-Crys aromatic pair mutants refolded to native-like state by Mm-Cpn ............ 143 Figure 5-4: HγD-Crys mutants refolded by Mm-Cpn have native-like fluorescence ................ 144 Figure 5-5: HγD-Crys interface pair mutants suppressed and refolded to native-like state by
Mm-Cpn .................................................................................................................................... 145 Figure 5-6: HγD-Crys hydrophobic core mutants suppressed and refolded to native-like state by
Mm-Cpn .................................................................................................................................... 147 Figure 5-7: Most HγD-Crys mutants refolded to higher levels than WT HγD-Crys .................. 148
Figure 8-1: Immunoblot SDS-PAGE of sucrose gradient ultracentrifugation of CCT1-CCT4 .. 193 Figure 8-2: Immunoblot SDS-PAGE of sucrose gradient ultracentrifugation of CCT5-CCT8 .. 194 Figure 8-3: Quantified densities of full-length CCT species for each set of sucrose
ultracentrifugation gradients ...................................................................................................... 198 Figure 8-4: Quantified densities of fragmented CCT species for each set of sucrose
ultracentrifugation gradients ...................................................................................................... 201 Figure 8-5: Heat maps of CCT subunit complex formation alone, with Mm-Cpn, CCT4, or CCT5
.................................................................................................................................................. 202 Figure 8-6: Possible models for TRiC formation assuming assembly is started from CCT4 or
CCT5 homo-oligomers .............................................................................................................. 206 Figure 8-7: CCT5 and human TRiC suppress aggregation of mutant huntingtin while CCT4 and
Mm-Cpn do not ......................................................................................................................... 213 14
LIST OF TABLES
Table 1-1: CCT subunits implicated in substrate binding varies for different substrates ............ 41 Table 1-2: Mutations in chaperonin subunits lead to human disease......................................... 48
Table 4-1: Mutations in chaperonin genes leading to neuropathy diseases............................. 101
Table 5-1: All HγD-Crys mutants are destabilized compared to WT HγD-Crys ........................ 139 Table 5-2: Kinetics of Mm-Cpn suppression of HγD-Crys aggregation vary by mutant ........... 141
Table 8-1: Human CCT subunit expressed from eight different chromosomes........................ 186 Table 8-2: Antibodies against the CCT subunits ...................................................................... 191 Table 8-3: Summary table of full-length CCT subunits co-expressed with homo-oligomers .... 203 15
16
LIST OF ABBREVIATIONS
(in alphabetical order)
ANC: actin non-complementing
AP: alkaline phosphate
BBS: Bardet-Biedl syndrome
BCA: bicinchoninic acid assay
BIN: binucleated
BME: β-mercaptoethanol
CD: circular dichroism
CCT: chaperonin containing TCP-1
DHFR: dihydrofolate reductase
DTT: dithiothreitol
E. coli: Escherichia coli
EDTA: ethylenediaminetetraacetic acid
FSC: Fourier shell correlation
GdnHCl: guanidine hydrochloride
HγD-Crys: human γD-crystallin
Hsc: heat shock cognate
Hsp: heat shock protein
HSPD1: human mitochondrial Hsp60
HtpG: high temperature protein G
Htt: huntingtin
IP: immunoprecipitation
IPTG: isopropyl-β-thiogalactoside
mHtt: mutant huntingtin
Mm-Cpn: Methanococcus maripaludis chaperonin
NAC: nascent-chain associated complex
NaPi: sodium phosphate
NEF: nucleotide exchange factor
OE: overexpression
PDB: protein data bank
PEI: polyethelenimine
PVDF: polyvinylidene difluoride
pVHL: Von-Hippel Lindau tumor suppression protein
17
RuBisCO: ribulose-biphosphate carboxylase
sHSP: small heat shock proteins
SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis
SEC: size exclusion chromatography
TAP: TCP-1 associated proteins
TCP-1: Tailless Complex Polypeptide-1
TEM: transmission electron microscopy
TLC: thin layer chromatography
TPR: tetratricopeptide repeat
TRiC: TCP-1 Ring Complex
Tris: Tris(hydroxymethyl)-amino methane
WD40: 40 amino acid sequences ending in tryptophan and aspartate residues
WT: wild-type
18
CHAPTER 1:
Introduction
19
Protein Folding and Aggregation
The cell cytoplasm is a crowded space with a high concentration of macromolecules,
organelles, and small molecules. In this environment, not all newly synthesized proteins can fold
up into their native conformations. Roughly 30% of the proteins in both prokaryotic and
eukaryotic cells misfold or are degraded after translation, making protein misfolding a serious
obstacle for cell viability and reproduction (Schubert et al. 2000; Hartl and Hayer-Hartl 2009).
This failure to assume native state often leads to partially folded states that can result in
aggregation or the formation of toxic species (van den Berg et al. 1999; Slavotinek and
Biesecker 2001; Ellis 2003). These toxic species can be amyloid in nature, such as in prion
disease and other neurodegenerative disease (Berthelot et al. 2013; Olanow and Brundin 2013;
van der Putten and Lotz 2013). On the other hand, the toxic species may also be non-amyloid
(frequently termed “amorphous”) aggregation as in cataract (Wang and King 2010; Moreau and
King 2012). There are also cases where the oligomer species is toxic, as in many of the
neurodegenerative diseases such as Alzheimer’s Disease, Huntington’s Disease, and
Parkinson’s Disease (Frid et al. 2007; Berthelot et al. 2013; Denny et al. 2013; Margulis et al.
2013). Whatever the toxic conformation, many cell types have systems in place to decrease or
eliminate this species.
Molecular chaperones in the cell guide nascent and misfolded proteins to their native
states or protect misfolded proteins from aggregation (Figure 1-1) (Feldman and Frydman 2000;
Frydman 2001; Slavotinek and Biesecker 2001; Lee and Tsai 2005; McClellan et al. 2005; Ellis
2006; Broadley and Hartl 2009; Chen et al. 2011; Hartl et al. 2011). Direct experiments suggest
that up to 50% of newly synthesized proteins in both yeast and E. coli utilize chaperone
assistance (Teter et al. 1999; Feldman and Frydman 2000; Hartl and Hayer-Hartl 2009; Hartl et
al. 2011). Not only do chaperones act on nascent proteins, but they can also recognize and act
on proteins that become unfolded due to cellular stress or inherent destabilization from
mutations (Gregersen and Bross 2010). Chaperones can also pass substrates that are unable
to fold onto the proteasome system, therefore ridding the cell of these species that have
potential to aggregate (Chen et al. 2011; Hartl et al. 2011). Chaperones are divided into two
classes: ATP-dependent chaperones (termed foldases) and ATP-independent chaperones (also
termed small heat shock proteins or holdases) (Frydman 2001; Hartl et al. 2011). The ATPdependent chaperones are actually able to bind to partially unfolded or misfolded substrates
and help them in folding to more native-like states.
20
Unfolded State
Energy
Chaperones
Partially
Folded
States
Oligomers
Native
State
Non-amyloid
Aggregation
Amyloid
Intermolecular contacts
Intramolecular contacts
Figure 1-1: Effect of chaperones on protein folding and aggregation energy landscape
Protein conformations from the unfolded state (top) are shown as either having intermolecular
(left, orange) or intramolecular contacts (right, blue). Intramolecular contacts are shown as
partially folded states and the native state. Intermolecular contacts are shown as amyloid,
oligomers, and non-amyloid aggregation. Chaperones assist (green arrow) in driving proteins
from partially folded states into their native states, and inhibit (red bar-headed arrow) proteins
from going from the partially folded state to amyloid, oligomers, and non-amyloid aggregation.
Figure based off of Hartl et al. reviews (Hartl and Hayer-Hartl 2009; Hartl et al. 2011).
21
ATP-Dependent Chaperones
There are three main classes of chaperones that use ATP to drive proteins from partially
folded states to their native states. These are heat shock protein (Hsp) 70, Hsp90, and Hsp60
(chaperonins). They act in the cell at different times in the folding pathways, with Hsp70 acting
as the first chaperone to encounter a protein off of the ribosome, and the chaperonins and
Hsp90 acting downstream of Hsp70 (Hartl et al. 2011).
Hsp70
The first chaperones to bind to newly synthesized proteins and influence their folding in
an ATP-dependent manner are part of the Hsp70 family (DnaK in prokaryotes) (Frydman 2001;
Hartl et al. 2011; Mayer 2013). In eukaryotes, the nascent-chain associated complex (NAC), a
cluster of Hsp70-family chaperones at the ribosome exit site, greets the polypeptide chain (Hartl
et al. 2011). On the other hand, in prokaryotes, a protein called trigger factor binds to almost all
chains coming out of the ribosome, before these substrates can be transferred to DnaK (Teter
et al. 1999; Hartl et al. 2011). Hsp70 that is constitutively expressed is termed Heat shock
cognate (Hsc) 70 (Saibil 2013). However, other isoforms of Hsp70 can be induced under stress
conditions (Saibil 2013). Many human cancers overexpress Hsp70 family proteins and this
overexpression is linked to poor prognosis (Murphy 2013). Therefore, Hsp70 has been a recent
therapeutic target (Assimon et al. 2013).
Hsp70 has two domains: the N-terminal nucleotide-binding domain (43 kDa) and the Cterminal substrate-binding domain (27 kDa) (Figure 1-2) (Mayer 2013). Using ATP binding and
hydrolysis, it cycles between two conformational states: an ATP-bound low-affinity open state
where the substrate has high association and dissociation rates in the substrate-binding
domain, and a high-affinity closed state after ATP is hydrolyzed where the substrate has low on
and off rates (Mayer 2013). In general, Hsp70s have low ATP hydrolysis and release rates, but
with the help of a nucleotide exchange factors (NEFs; specifically GrpE in prokaryotes), the
ADP can be released, opening the Hsp70 and releasing the substrate (Mayer 2013). In addition
to NEFs, Hsp70s are assisted by other co-chaperones such as Hsp40 (DnaJ in prokaryotes),
which help bind substrates, bring them to Hsp70 and increase the rate of ATP hydrolysis (Hartl
et al. 2011). Hsp40 has a conserved J-domain, that interacts with the nucleotide-binding domain
of Hsp70 (Clare and Saibil 2013).
The substrate-binding domain of Hsp70 specifically binds stretches of five hydrophobic
amino acids with positive amino acids on either side, which occur on average every 40 amino
acids in many globular proteins (Mayer 2013). These recognition elements are most likely
22
buried in the hydrophobic core of folded proteins. Therefore, Hsp70 only acts on unfolded or
partially folded chains. By binding to these substrates, Hsp70 decreases their potential to
aggregate in the cell. When released, the substrate is more competent to fold by burying the
bound hydrophobic patches into its core (Hartl et al. 2011). If the substrate cannot fold, it may
be bound again by Hsp70, passed on to the other chaperones in the cell, or targeted for
degradation (Hartl et al. 2011; Saibil 2013).
23
Figure 1-2: Hsp70 structure and states
Hsp70 has two domains: the substrate binding domain (cyan) and the nucleotide binding
domain (magenta). It cycles between a low-affinity ATP-bound state where the substrate binding
site is exposed and there are high rates of substrate association and dissociation (A) and a
high-affinity state where the substrate binding site is closed and therefore the substrate is tightly
bound (B). The substrate and nucleotide binding sites of both states are labeled. Low-affinity
state: PDB: 4B9Q; high-affinity state: PDB: 2KHO.
24
Hsp90
Hsp90 acts on substrates (specifically termed clients) to regulate their conformations
and mature them (Hartl et al. 2011). These clients are involved in crucial cell processes such as
signal transduction, innate and adaptive immunity, and protein trafficking (Picard 2008; Taipale
et al. 2010). Because of its central role in important cell functions, Hsp90 seems to act as a
buffer for protein evolution, allowing destabilized proteins to fold and mature (Rutherford and
Lindquist 1998; Lindquist 2009). Like Hsp70, Hsp90 is also overexpressed in most human
cancers, making it a robust therapeutic target (Whitesell and Lindquist 2005). Interestingly, while
Hsp90 is essential in eukaryotic cells, the bacterial homolog high temperature protein G (HtpG)
is not essential and Archaea do not possess any Hsp90 homologs (Taipale et al. 2010).
Hsp90 is made up of three domains: a N-terminal ATP binding domain, a middle domain,
and a C-terminal dimerization domain (Figure 1-3) (Taipale et al. 2010; Clare and Saibil 2013). It
functions as a homo-dimer and cycles through two conformations: an open conformation where
only the C-terminal domains interact and a closed ATP-bound conformation in which both the Cterminal and N-terminal domains are interacting and the N-terminal domains are twisted relative
to each other (Clare and Saibil 2013). Clients on their own seem to bind to any of the three
domains, but co-chaperones preferentially bind to the C-terminal domain (Clare and Saibil
2013). These co-chaperones have tetratricopeptide repeat (TPR) domains that bind to the
MEEDV sequence in Hsp90 C-terminus (Taipale et al. 2010; Hartl et al. 2011). The cochaperones are specific for their clients, increasing the diversity of interactions by Hsp90
(Taipale et al. 2010). These co-chaperones also regulate the ATP hydrolysis cycle of Hsp90,
allowing clients to undergo various conformational transitions that lead to their maturation (Hartl
et al. 2011).
25
Figure 1-3: Hsp90 structure and states
Hsp90 has three domains: the N-terminal ATP-binding domain (orange), the middle domain
(blue), and the C-terminal dimerization domain (green). It cycles between an open state (A)
where substrates can easily associate and dissociate and an ATP-bound closed state where
substrates are tightly bound (B). Open state: PDB: 2IOQ; closed state: PDB: 2CG9.
26
Chaperonins
Like Hsp70, chaperonins bind partially folded substrates and through conformational
changes, induced by ATP-binding and hydrolysis, release a more native-like substrate
(Frydman 2001). However, unlike Hsp70, substrate folding by chaperonins occurs in a cavity
within the chaperonin where the substrate is sequestered, in whole or in part, away from the
environment of the cell (Tang et al. 2006). Since substrates up to 60 kDa can fold in this cavity,
whole proteins and domains can be recognized and encapsulated (Xu et al. 1997). Little is
understood about how exactly the substrate achieves a more native-like conformation within this
cavity (Gershenson and Gierasch 2011).
Chaperonins are composed of two back-to-back rings with seven to nine subunits each
(Horwich et al. 2007). Each subunit is divided into three domains: equatorial, intermediate, and
apical (Braig et al. 1994). ATP hydrolysis and inter-ring negative allostery occur at the equatorial
domain (Spiess et al. 2004). The hinge-like intermediate domain connects the equatorial and
apical domains (Braig et al. 1994). Substrates are recognized via the apical domains, taken into
the cavity, folded, and then released through the same opening (Horwich et al. 2007). The
chaperonins are divided into two groups: group I (found in prokaryotes and in the chloroplasts
and mitochondria of eukaryotes) and group II (found in archaeal and eukaryotic cytosols)
(Frydman 2001). Group I chaperonins bind approximately 12% and group II chaperonins bind 915% of newly synthesized proteins in their respective cytosols (Ewalt et al. 1997; Thulasiraman
et al. 1999; Frydman 2001). Many features of the structure and mechanism of group I
chaperonins have been elucidated; however, studies of group II chaperonins are more limited
due to their complexity.
27
Group I Chaperonins
History
In the late 1960s and early 1970s, many laboratories were investigating phage mutations
that led to defects in lysogenic or lytic propagation or recombination (Georgopoulos 2006).
Costa Georgopoulos and Ira Herskowitz decided to look at mutations in E. coli that would hinder
the development of λ phage (Georgopoulos 1971). They used C600 E. coli strain containing a
supE amber nonsense suppressor gene (Georgopoulos 1971). After mutagenesis, the bacteria
were spread on LB agar plates with λcI- and 434cI- phage (Georgopoulos 1971). These phage
were chosen because they do not lysogenize so their infection would absolutely result in death
of the host, and they have distinct host-surface receptor attachment so host surface receptor
mutations would only be observed if both surface receptor genes contained mutants. The
concentration of phage to apply to the bacteria was very carefully chosen as to get enough
bacterial colonies lysed by the phage (rather than just “nibbled”), but not so many that all of the
bacterial colonies are killed (Georgopoulos 1971; Georgopoulos 2006). At the correct
conditions, most of the bacteria were killed by the phage, but every thousandth colony was large
and unaffected by the phage. The bacteria in these colonies had mutations that blocked phage
growth and were therefore named gro mutants (Georgopoulos 1971). Two of these mutants,
gro15 and groC3, were studied further and eventually identified as part of two distinct
chaperone systems (Georgopoulos 1971).
Georgopoulos observed that most λcI- phage did not form plaques on the Gro15 E. coli
mutant, but with a frequency of about 10-7, plaques did form (1971). Therefore, the phage that
formed these plaques were able to compensate for the Gro15 mutation. The phage mutants
were purified and tested on a variety of other amber-suppressing and non-suppressing E. coli
strains (Georgopoulos 1971). About 20% of these λ phage mutants could not kill the nonsuppressing wild-type (WT) E. coli strains but did form plaques on the amber-suppressing
bacterial strains. This lead to the conclusion that the compensatory mutants found in the λ
phage had a suppressible amber mutation in an essential phage gene (Georgopoulos 1971).
Herskowitz tested a collection of essential λ phage amber mutants for complementation with the
λ phage Gro15 compensatory mutations, and found that the λ phage Gro15 compensatory
mutations mapped to gene P of λ phage (Georgopoulos 1971; Georgopoulos 2006). This gene
was known to be responsible for λDNA replication. Two other λ phage mutants that possessed
mutations in gene P, λPam3 and λPam80, were shown to also form plaques on C600 supE
gro15 mutant E. coli (Georgopoulos 1971). However, many of the isolated λ phage Gro15
compensatory mutations and the known gene P λ phage mutations were unique. Now that the λ
28
phage compensatory mutants were mapped, the mutations in E. coli could further be studied.
Herskowitz showed that most of the Gro15 E. coli mutant (and a few other mutants with the
same phenotypes named groP mutants) mapped to the dnaB gene (Georgopoulos 1971).
Georgopoulos and Herskowitz reasoned that for λDNA replication to occur, the λP protein must
interact with DnaB protein of E. coli (Georgopoulos 1971). As DnaB is a helicase, it is not
surprising that they found this replication-associated protein in their genetic screen (Zylicz et al.
1984).
There was one groP mutant (groPAB756) that did not map to DnaB, but instead mapped
near the thr locus of E. coli and was renamed groPC756 (Georgopoulos 1977). Georgopoulos
went on to show that this mutant was responsible for both phage growth defects and host
temperature sensitivity, therefore concluding that this GroPC protein forms a complex with the
λP protein (1977). Concurrently, Michael Feiss’s laboratory isolated an E. coli mutant defective
in λ phage growth and bacterial growth temperature sensitivity (Sunshine et al. 1977). They
showed that this mutant (groPC259) was closely linked to groPAB756 (Sunshine et al. 1977).
Around the same time, Saito and Uchida were isolating E. coli mutants (named grp for groPlike), which interfere with λDNA replication and are temperature sensitive (1977). As with
Georgopoulos and Herskowitz, some of their mutants mapped to DnaB, but one class of
mutants (grpC) fell near groPC756 and groPC259 (Saito and Uchida 1977). In collaboration, it
was discovered that the groPC and grpC mutants fell into two distinct complementation classes:
groPC756 and groPC259, which Saito and Uchida renamed dnaK and dnaJ, respectively
(Yochem et al. 1978). Saito and Uchida went on to isolate the essential nucleotide exchange
factor grpE from another class of their mutants, therefore rounding out the DnaJ/DnaK/GrpE
chaperone system (1978).
The other original gro mutant identified by Georgopoulos and Herskowitz was groC3.
Georgopoulos et al. isolated compensatory λ phage mutants, about 30% of which had amber
suppressing mutations or temperature-sensitive mutations in gene E of λ phage (1973). The
gene encoded the capsid of the phage and was referred to as λε, therefore the mutant was
renamed GroEAC3 (and similar mutants were designated GroE mutants) (Georgopoulos et al.
1973). The GroE mutants were split into two classes: GroEA on which λεA plaques but not λεB
plaques formed, and GroEB on which λεB plaques but not λεA plaques formed (Georgopoulos
et al. 1973). Both λεA and λεB phage were isolated as compensatory mutants of GroE bacterial
mutants. Both bacterial mutant classes had problems growing at high temperatures. While all of
λεA phage mutants mapped to the E gene, the λεB phage mutants fell in both the E and B
genes (Georgopoulos et al. 1973). Transmission electron micrographs (TEMs) of groE mutant
29
bacteria infected with WT λ phage showed that the λ capsid (E gene) was incorrectly
assembled, showing that groE bacteria affected the λB protein function (Georgopoulos et al.
1973).
One groE bacterial mutant, GroEA44, not only resisted λ phage growth, but also T4
phage growth. As before, T4 plaque forming phage mutations were present at a frequency of
10-7, and referred to as T4ε. One of these mutants, T4ε1, could no longer grow on groE mutants
(groEB515),
which
propagated
WT
T4
phage
(Georgopoulos
et
al.
1972).
Using
complementation tests with known T4 amber suppressing mutants, this phage mutation was
mapped to gene 31 of T4 (which makes Gp31) (Georgopoulos et al. 1972). Therefore, it was
concluded that Gp31 of T4 phage interacted with the groEA44 gene product (Georgopoulos
1971). Concurrently, Ulrich Laemmli et al. showed that Gp31-defective T4 phage had no heads
and that without Gp31, the T4 capsid protein, p23, aggregated into insoluble clumps (1970).
Employing λgroE+-infected cells, which could overexpress the GroE protein, Georgopoulos and
colleagues found that GroE was a protein of approximately 60 kDa, had a tetradecameric
structure and could hydrolyze ATP (Georgopoulos and Hohn 1978; Hendrix 1979).
Further investigation of groE mutants showed that they fell into two complementation
groups (Tilly et al. 1981). These were renamed GroEL (large 60 kDa product) and GroES (small
15 kDa product) (Tilly et al. 1981). The GroES product was found to be a co-chaperone of
GroEL, and their interaction was ATP-dependent (Chandrasekhar et al. 1986). The T4 phage
protein Gp31 is another co-chaperone essential for Gp23 capsid formation and takes over the
function of GroES (Ang et al. 2000). The Gp31 co-chaperone is approximately the same size as
GroES, but its interaction with GroEL creates a larger cage in which Gp23 can fit and fold
(Bakkes et al. 2005; Clare et al. 2006). Interestingly, while λ phage, and obviously E. coli,
require the GroES co-chaperone, T4 phage does not (Ang et al. 2000). Therefore, isolation of
host mutants by T4 infection would not have identified the crucial GroES co-chaperone. As the
temperature sensitive properties of GroEL/GroES were part of the criteria of their discovery, it
was not until almost ten years later that their requirement for E. coli cell growth at all
temperatures was verified (Fayet et al. 1989).
While Georgopoulos was investigating the effect of bacterial mutation on phage
assembly, the laboratory of R. John Ellis was researching a protein that bound and helped
assemble Ribulose-biphosphate carboxylase (RuBisCO), a protein complex involved in carbon
fixation. RuBisCO subunit binding protein was isolated in a much more biochemical way than
GroEL. RuBisCO subunit binding protein was found to bind to RuBisCO in isolated pea
chloroplasts, but not be part of RuBisCO itself (Hemmingsen et al. 1988). Biochemical studies
30
showed that it was made up of 60 kDa and 61 kDa subunits making an approximately 700 kDa
complex with ATPase activity (Hemmingsen and Ellis 1986). With the help of protein
sequences, Ellis and Georgopoulos assembled their findings to conclude that GroEL and
RuBisCO subunit binding protein were members of the same family (Hemmingsen et al. 1988).
Around the same time, McMullin and Hallberg had concluded that the protein they found in the
mitochondria of Tetrahymena thermophila was homologous to GroEL (1987; 1988). Ellis,
Georgopoulos, and colleagues coined the term “chaperonin” for this family of proteins, derived
from the term “molecular chaperone” that Laskey used for nucleoplasmin, a protein that bound
histones and helped assemble them onto DNA (Laskey et al. 1978). The term “chaperonin” was
quickly accepted as work from the Hartl and Horwich groups on the mitochondrial Hsp60 in
yeast employed that term in publication shortly afterwards (Cheng et al. 1989).
Structure and Function
The most studied of the bacterial group I chaperonins, GroEL from E. coli, consists of
two back-to-back rings of seven 57 kDa identical subunits each with an inner cavity volume of
85,000 Å3 (Figure 1-4A) (Braig et al. 1994; Fenton and Horwich 1997). GroEL is a stressinduced chaperone like Hsp70 and Hsp90 (Horwich et al. 2007). GroEL requires the cofactor
GroES, or Hsp10, which acts as a lid to close the cavity when a substrate protein is bound and
encapsulated (Hunt et al. 1996). GroES is a dome-shaped structure composed of seven
individual subunits (Chen et al. 1994; Fenton and Horwich 1997). When GroES binds to GroEL,
the volume of the cavity expands to about 175,000 Å3 (Figure 1-4B), therefore allowing for
larger proteins to fit in the cavity (Xu et al. 1997). Substrate binding occurs when exposed
hydrophobic residues in the substrate folding intermediate make contacts with the hydrophobic
residues in the apical domain of the subunits of one ring of GroEL (cis ring; Figure 1-4C, 2) (Lin
and Rye 2006). When ATP binds to the cis ring, the substrate experiences global stretching and
local segmental tightening (Figure 1-4C, 3) (Sharma et al. 2008; Kim et al. 2010). GroES can
bind to the hydrophobic residues of the apical domains where the substrate is bound, pushing
the substrate into the cavity of the ring (Figure 1-4C, 4) (Frydman 2001). The substrate is
encapsulated in the cavity and refolded for approximately 15 seconds as the ATP is hydrolyzed
(Figure 1-4C, 5) (Tang et al. 2006). ATP and substrate binding to the trans ring cause GroES
dissociation and substrate release (Figure 1-4C, 6) (Lin and Rye 2006).
The released, more native-like, substrate may spontaneously assume its native state or
may need another round of encapsulation (Weissman et al. 1994). Due to its proximity to the
apical domain when released, the substrate can easily rebind to the cis ring of the chaperonin
31
when the trans ring completes its cycle (Weissman et al. 1994). The actual substrate folding
mechanism inside the chaperonin is unknown, but there is some evidence that sequestration of
the substrate from the environment produces an Anfinsen cage, where the substrate can freely
fold (Ellis 2003; Apetri and Horwich 2008; Horwich et al. 2009). Other studies have shown that
positively charged residues lining the cavity of GroEL play an active role in substrate folding,
especially for larger proteins that directly contact these residues (Xu et al. 1997; Tang et al.
2006). Studies of GroEL substrates show that GroEL prefers substrates with multiple α/β
domains with buried hydrophobic β-sheets (Houry et al. 1999; Kerner et al. 2005; Hirtreiter et al.
2009). These architectures are slow to fold and are prone to aggregation (Kerner et al. 2005;
Fujiwara et al. 2010; Raineri et al. 2010).
32
Figure 1-4: Group I chaperonin structure and mechanism
Each subunit has equatorial (blue), intermediate (green), and apical domains (purple). The
cavity of the group I chaperonin GroEL expands from the open state (A; PDB: 1OEL) to the
closed state (B; PDB: 1AON) in complex with the GroES lid (red). The mechanism of a group I
chaperonin involves an ATP hydrolysis transition state in which the substrate is folded (C, see
text for details).
33
Group II Chaperonins
History
Genetically unique tailless mice mutants have been studied since the early 1930s
(Chesley and Dunn 1936). Two mutations have been isolated from the work: the dominant Tlocus mutation and the recessive lethal t allele mutants (Bennett 1975). WT mice have normal
length tails, whereas the double heterozygous T/t mice have the tailless phenotype. Further, T/+
mice have short tail but t/+ mice have normal tails. Additionally, tx/ty males were sterile and the
sperm transmission ratios of tx/+ males resulted in as many as 99% of the progeny received the
t haplotype (Bennett 1975). The double heterozygous T/t mice bred true since both T/T and t/t
mice died as embryos (Bennett 1975). This was the first balanced lethal mammalian system
(Waelsch 1989). Therefore, the tailless phenotype seems to arise from the interaction of the Tlocus and the t allele (Bennett 1975).
It was not until the 1970s that the mapping of the t complex could be achieved. The t
chromosome 17 is different than the WT chromosome 17 in a region of inversions (named the t
complex) spanning 30 Mbs (Bennett 1975). This region contains around 100 genes (Bennett
1975). As defects were seen in spermatogenesis of male mice, Silver et al. isolated
spermatogenic cells from WT and t haplotype mice (1979). Cells were labeled with
35
S-
methionine in vitro, the protein fraction was isolated, and the proteins were separated using twodimensional isoelectric focusing and gel electrophoresis (Silver et al. 1979). They found that a
single abundant protein spot, p63/6.9, differed between WT and t haplotype mice. In WT mice,
p63 was a bit more acidic, therefore labeled p63/6.9a, whereas in t haplotype mice, the protein
was labeled p63/6.9b (Silver et al. 1979). They extended their work to other cell types
(splenocytes, thymocytes, and two carcinoma cell lines) to find that these cell types only
contained the WT p63/6.9b (Silver et al. 1979). This protein was coined Tailless Complex
Polypeptide 1 (TCP-1) (Silver 1985). Although research continued on the t complex in terms of
its implication in development (Waelsch 1989) and transmission ratio (Willison 1986), how
exactly it caused the tailless phenotype and whether TCP-1 actively played a role is still unclear.
To better understand t haplotype chromosome evolution, Keith Willison and colleagues
cloned TCP-1b (WT) and part of TCP-1a (t haplotype) genes. They found no sequence similarity
to known proteins and at least six nucleotide differences between the two genes (Willison et al.
1986). Willison’s laboratory went on to study TCP-1 in the cell and found (due to nonspecific
antibodies (Lewis et al. 1992)) that it associated with the cytoplasmic part of the Golgi
membrane (Willison et al. 1989). Therefore, they concluded that TCP-1 plays a role in transport
of proteins through the exocytic pathway (Willison et al. 1989). Working on microtubules in
34
yeast, Ursic and Culbertson found that an essential gene in yeast shared sequence identity with
mouse TCP-1 (Ursic and Culbertson). They isolated and characterized a cold sensitive yeast
TCP-1 mutant and concluded that TCP-1 affected the microtubules responsible for spindle pole
body positioning (Ursic and Culbertson 1991). Despite their logical conclusions based on their
data, both groups were misled. At this time, the publication by Georgopoulos and Ellis outlining
the chaperonins gave both groups the direction they needed to continue studying the actual role
of TCP-1 in the cell (Hemmingsen et al. 1988). Comparing the protein sequence of TCP-1 to
those of the rest of the chaperonins, Ellis speculated that TCP-1 was the eukaryotic cytosolic
chaperonin (1990). Unpublished data mentioned in his 1990 review bolsters this theory,
because the TCP-1 antibody recognized proteins in the crude extracts of pea leaves but not
chloroplasts or mitochondria (Ellis 1990).
While neither Georgopoulos nor Ellis crossed over to the group II chaperonin field, the
team of Ulrich Hartl and Arthur Horwich who got their start studying mitochondrial Hsp60 in
yeast, stumbled upon the connection between the archaeal thermosome and TCP-1 (Trent et al.
1991). They reasoned that thermophilic factor 55 (TF55) was part of the chaperonin family by
showing structural similarity to GroEL via electron microscopy and functional similarity to GroEL
as both chaperonins form a complex with a substrate,
35
S-methionine labeled Su9-DHFR (part
of subunit 9 of F0-ATPase fused to dihydrofolate reductase), diluted out of denaturant (Trent et
al. 1991). Additionally, protein sequence similarity was shown to be strongest between TF55
and TCP-1 than any other chaperonins, suggesting that they form a subclass of chaperonins,
possibly specialized in cytoskeletal assembly (Trent et al. 1991). This paper propelled the study
of TCP-1 as a chaperonin, although its chaperoning function was only speculative at this time.
In the summer of 1992, two papers published back-to-back in Nature, verified the
chaperoning function of TCP-1. The first paper, from the tubulin-focused Sternlicht laboratory in
collaboration with Horwich, reported that in rabbit reticulocytes, newly made tubulin subunits
enter a 900K complex before being competent to assemble into microtubules (Yaffe et al. 1992).
This complex consisted of a set of polypeptides between 55-60K and one of which reacted with
the TCP-1 antibody (Yaffe et al. 1992). Therefore, the authors concluded that tubulin interacted
with the TCP-1 complex to acquire a more assembly-competent form (Yaffe et al. 1992). This
was concurrently shown for actin assembly in rabbit reticulocytes by Nicholas Cowan’s
laboratory (Gao et al. 1992). The second article in the same issue was a report from the Willison
lab preliminarily characterizing human and mouse TCP-1 (Lewis et al. 1992). They found that in
both species, TCP-1 made a complex of approximately 900K (Lewis et al. 1992). Additionally,
TCP-1 associated with four to six unidentified polypeptides and two Hsp70 homologs, coined
35
TCP-1 associated proteins (TAPs) (Lewis et al. 1992). They speculated that the unidentified
polypeptides may be TCP-1 related proteins which were just being found in a variety of
organisms by sequence similarity (Lewis et al. 1992). Due to this heterogeneity of the TCP-1
complex, and the observation that TCP-1 levels were not increased in response to stress, the
Willison lab concluded that TCP-1 was unique from GroEL (Lewis et al. 1992).
At this time, there was only indirect evidence of the chaperoning properties of TCP-1
complex. Judith Frydman in the Hartl Lab reported that TCP-1, renamed TCP-1 Ring Complex
(TRiC), refolded unfolded substrates in vitro and definitively showed that TRiC did not need the
Hsp10 co-chaperonin required for GroEL-assisted protein folding (1992). Frydman et al.
demonstrated that purified bovine TRiC associated with at least five other polypeptides which
were sequenced to show at least 40% identity with TCP-1, suggesting that the TRiC is made up
of a number of homologous proteins (Frydman et al. 1992). Additionally, Frydman et al. showed
that TRiC can bind to and refold firefly luciferase from denaturant whereas GroEL can bind
denatured luciferase but not refold it (1992). This was the first direct evidence verifying that
TRiC functions as a chaperonin (Frydman et al. 1992). While this showed that TRiC had the
ability to fold substrates other than actin and tubulin, there was still only evidence of TRiCassisted actin and tubulin folding in the cell (Sternlicht et al. 1993).
The eight subunits of TRiC were first identified by Rommelaere, et al. and subsequently
sequenced and mapped by the Hartl and Willison laboratories (Rommelaere et al. 1993; Kubota
et al. 1994; Li et al. 1994). Rommelaere et al. also showed that bovine TRiC was structurally
consistent and functionally identical with rabbit reticulocyte TRiC (1993). All eight human
subunits were sequenced by the end of 1994 (Kubota et al. 1995). The Willison lab renamed
TRiC to CCT (Chaperonin Containing TCP-1) (Kubota et al. 1994). Although the term TRiC is
more widely used for the complex, CCTx is commonly used to designate subunit x of the
complex.
While TCP-1 (now CCT1) was identified from its difference between t haplotype and WT
mice, other subunits were also found concurrently through genetic screens. The group of
Huffaker was searching for mutants of tubulin that lead to binucleated cells (cells with two nuclei
produced by defective spindles) (Chen et al. 1994). Two of their mutants, BIN2 and BIN3
(binucleated), were further characterized, and mapped to CCT3 and CCT2, respectively (Chen
et al. 1994). In the meantime, while searching for temperature sensitive mutations of actin in S.
cerevisiae, the Drubin laboratory found non-complementing extragenic mutants of actininteracting proteins (Vinh et al. 1993). One of their mutants, ANC2 (actin non-complementing 2),
was mapped to CCT4 (Vinh and Drubin 1994). Recently, the actual ANC2 mutant was identified
36
to be CCT4 G345 and characterized to abolish ATP-induced allostery of TRiC (Shimon et al.
2008).
Structure and Function
The group II chaperonin mechanism is different from the group I chaperonin mechanism
due to the structural differences of the two groups. The archaeal group II chaperonins consist of
two 7-9 subunit rings that have 1-3 different subunits, while the eukaryotic group II chaperonin
Tailless Complex Polypeptide-1 (TCP-1) Ring Complex (TRiC) consists of two identical rings,
each with eight different subunits (Bigotti and Clarke 2008). While the archaeal chaperonins can
be stress induced like the group I chaperonins, the eukaryotic TRiC is not a stress-inducible
chaperone (Horwich et al. 2007). Overall, the subunits have the same domain organization as
those of GroEL, but since group II chaperonins do not require a cofactor like GroES, the apical
domain has a helical protrusion which acts as a built-in lid (Figure 1-5A) (Reissmann et al.
2007). Unlike expanding in the group I chaperonins, the volume of the cavity of the group II
chaperonins contracts from about 350,000 Å3 in the open state to 130,000 Å3 in the closed state
(Figure 1-5B) (Huo et al. 2010; Pereira et al. 2010). Although group II chaperonins do not have
a lid-like co-chaperone, they do interact with prefoldin, a co-chaperone that binds some
substrates and brings them to the apical domain (Gutsche et al. 1999; Martín-Benito et al. 2002;
Sahlan et al. 2010). Most of the research on prefoldin structure and function has employed the
archaeal group II chaperonins, so little is known about this co-chaperone in eukaryotic cells.
The homo-oligomeric archaeal Methanococcus maripaludis chaperonin (Mm-Cpn) has
provided a useful group II chaperonin model because it allows for recombinant expression of
site-directed mutations (Spiess et al. 2004). Recent studies using a variety of Mm-Cpn mutants
have revealed the mechanism of substrate folding differs from that of the group I chaperonin
(Douglas et al. 2011). The substrate folding intermediate binds to the cis ring of the chaperonin
at the apical domains (Figure 1-5C, 2). ATP binds to the cis ring and the lid begins to close
(Figure 1-5C, 3) (Douglas et al. 2011). The ATP hydrolysis transition state (Figure 1-5C, 4)
precedes substrate release into the cavity, which is caused by scraping the substrate from the
apical domain via lid closing (Figure 1-5C, 5) (Douglas et al. 2011). Once the substrate is
released into the cavity of the chaperonin, it folds into a more native-like conformation (Figure 15C, 6) (Douglas et al. 2011). The cause of lid opening is unknown but it may be substrate
binding at the trans ring (Figure 1-5C, 7).
37
Figure 1-5: Group II chaperonin structure and mechanism
Each subunit has equatorial (blue), intermediate (green), and apical domains (purple) along with
a built-in lid (red). The cavity of the group II chaperonin narrows from the open state (A; PDB:
3KO1) to the closed state (B; PDB: 3KFB). The mechanism of a group II chaperonin involves an
ATP hydrolysis transition state which precedes substrate release into the cavity for folding (C,
see text for details).
38
Substrate Recognition by Group II Chaperonins
Chaperonin Subunits/Domains Involved in Recognition
The CCT subunits have a 30% sequence identity but their largest divergence is in the
apical substrate-recognition domain (Figure 1-6). This suggests that the different subunits of
TRiC may have evolved distinctive subunit specificities (Kim et al. 1994; Frydman 2001; Spiess
et al. 2006). Evolutionarily, this may be due to eukaryotic substrates being more difficult to fold
because of their increased complexity, therefore needing more motifs for substrate recognition
(Frydman 2001). In addition, by having very distinct apical domains but still having the CCT
subunits form one structure, many substrates can be folded in the same space.
Although only a few substrates have been studied, it is clear that not all CCT subunits
are implicated in binding of non-native-state substrates to TRiC (Table 1-1) (Llorca et al. 1999;
Hynes and Willison 2000; Llorca et al. 2000; Spiess et al. 2006; Tam et al. 2006). Many
substrates appear to bind across the ring, thus contacting subunits on either side of the ring
(Llorca et al. 2000; Martín-Benito et al. 2004). There are two alternate binding models
suggested for TRiC binding to actin and tubulin due to subunit arrangement geometry (Llorca et
al. 2000). In both TRiC-actin models, actin binds to five different CCT subunits (CCT1-3, 7, 8 or
CCT 2, 4-6, 8) while both TRiC-tubulin models suggest that tubulin binds to two different CCT
subunits (CCT2 then CCT4 or CCT5 then CCT4). The lack of high-resolution TRiC-actin or
TRiC-tubulin structures along with inconsistencies in subunit assignment have made it difficult to
distinguish the correct model with the methods employed. Other substrates such as huntingtin
and Von Hippel Lindau tumor suppressor protein (pVHL) have been more clearly studied using
crosslinking and overexpression methods. The apical domains of CCT1 and CCT4 have been
implicated in TRiC binding to polyglutamines such as those in the pathogenic exon one of
huntingtin (Tam et al. 2006). Spiess et al. demonstrated that TRiC subunits CCT1 and CCT7
bind pVHL (2006). While these studies show that not all CCT subunits are required to bind a
variety of substrates to TRiC, we cannot firmly conclude whether the eight CCT subunits are
specific for binding these substrates. For example, it may be that CCT1 preferentially binds
actin, although in the absence of CCT1 another subunit could perform this function, albeit less
efficiently.
39
Figure 1-6: Alignment of apical domains of CCT subunits
The apical domains of all eight CCT subunits were aligned. The blue residues are conserved in
6 or more of the CCT subunits while the red residues are conserved in all 8 CCT subunits.
While overall alignment is reasonable, there is considerable variation in the apical domains
between the CCT subunits.
40
Table 1-1: CCT subunits implicated in substrate binding varies for different substrates
Substrate
Function
CCT Subunit Bound
Method
Reference
Tubulin
Cytoskeleton
1-3, 7, 8 or 2, 4-6, 8
TEM reconstruction, IP
a
Actin
Cytoskeleton
2 and 4 or 4 and 5
TEM reconstruction, IP
b
Huntingtin
Scaffolding
1 and 4
OE in yeast & neurons
c
pVHL
Cell cycle control
1 and 7
Crosslinking, co-IP
d
TEM = transmission electron microscopy
IP = immunoprecipitation
OE = overexpression
pVHL = protein von Hippel-Lindau
a = (Llorca et al. 2000; Llorca et al. 2001)
b = (Llorca et al. 1999; Llorca et al. 2000)
c = (Tam et al. 2006)
d = (Feldman et al. 2003)
41
Features of the Substrate Recognized
GroEL has been studied extensively for its substrate recognition properties (GómezPuertas et al. 2004; Yébenes et al. 2011). Most substrate proteins of GroEL have been
identified by mass spectrometry, in vivo GroEL interaction, and bioinformatics (Kerner et al.
2005; Fujiwara et al. 2010; Tartaglia et al. 2010). These proteins contain aggregation-prone
folds and have an enrichment of alanines and glycines (Fujiwara et al. 2010). GroEL can bind
unfolded substrates with exposed hydrophobics through hydrophobic patches on its subunits
(Horwich et al. 2007). More specifically, NMR studies show helices 8 and 9 of GroEL bind to an
amphipathic helix of a substrate peptide, suggesting that GroEL can recognize amphipathic
elements in substrates (Li et al. 2009). These helices were earlier identified as crucial for both
substrate binding and for binding of the co-chaperone GroES, by studying point mutations in
that region (Fenton et al. 1994).
The group II chaperonin TRiC prefers substrates with extended β-sheets, whose folds
contain hydrophobic patches and are slow to fold (Yam et al. 2008). The interactions of TRiC
with pVHL demonstrated that TRiC recognizes two hydrophobic β-sheets termed Box 1 and Box
2 (Feldman et al. 2003). These motifs, which are buried in the native state, bind through
hydrophobic interactions to TRiC (Feldman et al. 2003). TRiC has been shown to recognize
delineated hydrophobic sections of actin and tubulin (Rommelaere et al. 1999). Further
molecular dynamics simulations between CCT3 and β-tubulin shows that TRiC recognizes this
substrate through hydrophobic and electrostatic interactions, particularly via a salt bridge
network between tubulin and CCT3 (Jayasinghe et al. 2010). In addition, TRiC recognizes βpropeller proteins, specifically the hydrophobic third β-strand of the second WD40 (40 aminoacid stretch ending with tryptophan and aspartate residues) repeat of G protein β and WD40
repeats 3-5 in Cdc20 (Camasses et al. 2003; Kubota et al. 2006). β-propellers are extremely βsheet rich and fold slowly, making them ideal group II chaperonin substrates (Gromiha and
Selvaraj 2004; Smith 2008).
42
Chaperonin Complex Evolution
Homo-oligomeric Chaperonins
All prokaryotic group I chaperonins are homo-oligomeric in structure (Horwich and
Willison 1993). Many archaeal group II chaperonins are also homo-oligomeric (Large et al.
2009). Due to the fact that group I chaperonins have the lid-like GroES co-chaperone, it seems
that this may have evolved in the prokaryotic lineage, rather than being lost in the archaeal
lineage (Dekker et al. 2011). Therefore, the group I and group II chaperonins might have had a
common ancestral chaperonin origin, from which they diverged and evolved (Dekker et al.
2011). Within eukaryotic mitochondria, Hsp60 forms homo-oligomeric chaperonins (Cheng et al.
1989). However, in chloroplasts of algae and plants, the Hsp60 group I chaperonin seems to
have multiple chaperonin subunits (Nishio et al. 1999; Hill and Hemmingsen 2001).
Hetero-oligomeric Chaperonins
Hetero-oligomeric chaperonins have evolved presumably from homo-oligomeric
ancestors, especially in the archaeal lineage where some chaperonins have two or three
subunits (Archibald et al. 1999; Large et al. 2009). The archaea Thermoplasma acidophilum has
alternating α and β subunits, while Haloferax volcanii has three subunits denoted cct1-3 (Large
et al. 2009). Most extreme is the eukaryotic group II chaperonin TRiC which has eight different
subunits (Hartl et al. 2011). While the number of different subunits has increased, the number of
subunits per ring has stayed fairly constant with 7 for the group I chaperonins and 8 or 9 for the
group II chaperonins (Archibald et al. 1999). The evolution of hetero-oligomerization in the
archaeal chaperonins is postulated to have happened due to gene duplication and then
specificity to a class of substrates via divergent mutations in the paralogs (Archibald et al.
1999). Along with evolution of substrate specificity, there also evolved a preference or an ability
to hetero-oligomerize rather than homo-oligomerize in the cell (Archibald et al. 1999). Therefore,
different subunits can more strongly interact with each other than themselves. Hsp60 in
chloroplasts, mentioned above, may have evolved similarly (Nishio et al. 1999). The eukaryotic
TRiC may be an extreme version of the evolution events seen in archaeal chaperonins. In fact,
phylogenetic studies of the CCT subunits show that their divergence was due to positive
selection after duplication events (Fares and Wolfe 2003).
Arrangement of CCT Subunits in TRiC
Studies of TRiC have been limited due to its multiple subunit species. TRiC subunit
arrangement has been a source of controversy in the field, because different methods and
43
laboratories generated varying models (Figure 1-7) (Liou and Willison 1997; Martín-Benito et al.
2007; Cong et al. 2010; Dekker et al. 2011). Even the same laboratories have obtained different
structures when employing different methods. One issue is that the structures generated for
identifying the arrangement are too low in resolution to distinguish the subunits, further
complicating identification of CCT-substrate interactions.
The first proposed arrangement, from biochemical studies in the Willison group on
bovine TRiC, showed this order: CCT 1-5-6-2-3-8-4-7 (Liou and Willison 1997). Working
together, the Willison and Valpuesta groups employed immunogold negative stain EM with
antibodies against specific subunits (CCT1, CCT4, CCT7 and CCT8) to obtain the same
arrangement of bovine TRiC (Llorca et al. 2000). These groups then furthered these studies by
using cryo-EM of bovine TRiC with surface-specific antibodies against CCT4, allowing them to
solve the register of the two rings (Martín-Benito et al. 2007).
In the meantime, the Frydman and Chiu groups used high resolution cryo-EM to obtain a
structure of bovine TRiC where the slight differences in structure between the subunits could be
resolved, giving the arrangement: CCT1-7-5-4-8-3-2-6 (Cong et al. 2010). The Willison group
obtained a crystal structure of rabbit α-actin bound to yeast TRiC, and found that their previous
arrangement docked well into the electron density (Dekker et al. 2011). They did note that the
register of the crystal structure differed by one subunit counter-clockwise as compared to the
earlier EM studies (Dekker et al. 2011). Most recently, two cross-linking and mass spectrometry
were in agreement about a new arrangement: CCT1-3-6-8-7-5-2-4. One study was performed in
the Levitt group on bovine TRiC; the other by the groups of Frydman, Chiu, Hartl, and
Aebersold, who used both bovine and yeast TRiC to obtain the same arrangement (Kalisman et
al. 2012; Leitner et al. 2012). The authors of the latter work note that their new arrangement has
a better fit in the yeast TRiC crystal structure than the one used by the Willison group (Leitner et
al. 2012).
44
Figure 1-7: TRiC subunit arrangement differs between laboratories and methods
Arrangements of the CCT subunits have been suggested around the ring (shown in color on
top) and between the two rings (shown in numbers below with the homotypic inter-ring
interactions indicated). A: Crosslinking and mass spectroscopy of bovine and yeast TRiC
(Kalisman et al. 2012; Leitner et al. 2012) B: Cryo-EM reconstruction of bovine TRiC (Cong et
al. 2010) C: Cryo-EM reconstruction and crosslinking studies of bovine TRiC (Liou and Willison
1997; Llorca et al. 2000; Martín-Benito et al. 2007) D: Crystal structure of yeast TRiC (Dekker et
al. 2011).
45
Role of Chaperonins in Human Disease
Mutations in Human Chaperonin Genes
Sensory neuropathies are diseases affecting the degeneration of the nerve fibers of
sensory neurons, leading to ulceration and inability to feel pain (Cavanagh et al. 1979; Thomas
et al. 1994). Single point mutations in the DNA that result in CCT4 (C450Y) and CCT5 (H147R)
mutations have been implicated to lead to hereditary sensory neuropathy in a stock of SpragueDawley rats and in a Moroccan family, respectively (Lee et al. 2003; Bouhouche et al. 2006).
Similarly, a mitochondrial Hsp60 (V98I) mutation was identified in a French family with
neuropathy (Bross et al. 2008). In vitro studies showed that this mutant affected the group I
chaperonin’s ability to bind and refold its substrates (Bross et al. 2008). Therefore, it is likely that
the CCT mutations affect the substrate binding abilities of CCT4 and CCT5, therefore affecting
TRiC function.
In addition to neuropathy, ciliopathy is also related to mutations in the human chaperonin
genes (Table 1-2). Human Bardet-Biedl syndrome (BBS) is a ciliopathy primarily affecting the
renal cells (Marion et al. 2011). Three BBS genes: BBS6 (also called MKKS for McKusickKaufman Syndrome), BBS10, and BBS12 share sequence identity with the CCT genes
(Mukherjee et al. 2010). In fact, TRiC seems to interact with these three proteins as part of the
BBS-chaperonin complex, which is required for BBSome (made up of the remaining BBS genes:
1, 2, 4, 5, 7, 8, and 9) assembly (Zhang et al. 2012). Mutations in these genes have been
implicated in BBS (Nakane and Biesecker 2005; Stoetzel et al. 2006; Stoetzel et al. 2007;
Billingsley et al. 2010). Due to the high conservation in sequence between the BBS and CCT
genes and the large physiological affects of this disease, the mutated sites might be crucial to
TRiC function, as well.
Using TRiC to Ameliorate Diseases
Since TRiC can recognize and help misfolded species, it has been postulated that it
could be targeted in various diseases for therapy (Slavotinek and Biesecker 2001; Broadley and
Hartl 2009). As with Hsp70 and Hsp90, CCT subunits are also overexpressed in many cancers
(Yokota et al. 2001; Boudiaf-Benmammar et al. 2013). The devastating effects of expanded
polyglutamine stretches, as found in Huntington’s disease, have been ameliorated in cells
where CCT subunits are upregulated (Kitamura et al. 2006; Tam et al. 2006). Huntington’s
disease is marked by an expansion in the first exon of the protein huntingtin, leading to
psychological and physical ailments such as depression and chorea (Walker 2007). Both
Kitamura et al. and Tam et al. demonstrated that knocking down CCT subunits increased
46
mutant Htt aggregates, while overexpressing CCT subunits lead to inhibited neuronal cell death
(2006; 2006). Using cryo-EM, not only has TRiC been shown to suppress in vitro Htt
aggregation, but an interaction between Htt and TRiC was revealed (Shahmoradian et al. 2013).
However, not all of TRiC is needed for these beneficial effects of the chaperonin. Just the CCT1
apical domain has been shown to enter cells, decrease huntingtin aggregation and increase cell
viability (Sontag et al. 2013). While this is promising, the apical domain is still 20 kDa in size, so
it would prove difficult to develop into a pharmaceutical therapy. Therefore, small molecules for
inhibition or activation of TRiC or chaperonin function have also been investigated (Bergeron et
al. 2009). Finding a small molecule that could enhance interaction between TRiC and a diseasecausing substrate, while not significantly sacrificing TRiC function with the rest of its substrates
would be ideal.
47
Table 1-2: Mutations in chaperonin subunits lead to human disease
Mutation
Gene
Conservation
Domain
Result
Reference
C450Y
CCT4 rat
Only CCT4
Equatorial
Neuropathy
a
H147R
CCT5 human
Only CCT5
Equatorial
Neuropathy
b
V98I
HSPD1
None
Equatorial
Neuropathy
c
G41R
BBS6 human
All but CCT3
Equatorial
BBS
d
A242S
BBS6 human
CCT1, 5, 7, & 8
Apical
BBS
e
L55P
BBS10 human
All
Equatorial
BBS
d
L414S
BBS10 human
CCT1-3, 5, & 7
Intermediate
BBS
f
T501M
BBS12 human
All but CCT1
Intermediate
BBS
g
G539D
BBS12 human
All but CCT6
Equatorial
BBS
d
HSPD1 = human mitochondrial Hsp60
BBS = Bardet-Biedl syndrome, a ciliopathy
a = (Lee et al. 2003)
b = (Bouhouche et al. 2006)
c = (Hansen et al. 2002)
d = (Billingsley et al. 2010)
e = (Nakane and Biesecker 2005)
f = (Stoetzel et al. 2006)
g = (Stoetzel et al. 2007)
48
Thesis Context
This thesis aims to understand the properties of human TRiC. Is human TRiC similar to
the bovine and yeast TRiC that has been studied? Are CCT subunits specific or redundant for
substrates? Are all eight CCT subunits needed to assemble into chaperonin rings? Do the
neuropathy mutations in the CCT subunits affect TRiC function? And lastly, how is heterooligomeric TRiC assembled from the CCT subunits?
In Chapter 2, I present the purification and characterization of human TRiC from HeLa
cells. This material is consistent with TRiC that has been purified from bovine and yeast
sources. However, due to its endogenous nature, its yield is low and it cannot be genetically
manipulated. Therefore, I move on to purify the CCT subunits one at a time in Chapter 3.
Surprisingly, two of the CCT subunits, CCT4 and CCT5, but none of the others, formed TRiClike homo-oligomeric rings. I went on to characterize their structure and function with a number
of substrates. Coincidentally, CCT4 and CCT5 are the loci of point mutations implicated in
neuropathy diseases. My homo-oligomeric system was ideal to make and study these mutations
for defects in chaperonin structure and function. The characterization of these mutants in this
system is outlined in Chapter 4. Since I have been able to purify these homo-oligomers, a
natural progression is using these as starting points for hetero-oligomeric TRiC assembly. To
that end, Appendix A of Chapter 8, I describe the formation of hetero-oligomeric rings between
each CCT subunit and the homo-oligomeric CCT species. While this data is still preliminary, this
gives us some initial information about TRiC assembly in the cell. Appendix B shows the first
steps toward assessing whether the CCT subunits are specific for substrates as it shows that
CCT5 can efficiently suppress mutant huntingtin aggregation while CCT4 cannot.
49
50
CHAPTER 2:
Human TRiC Complex Purified from HeLa Cells
Contains All Eight CCT Subunits and is Active In Vitro*
* This research was originally published in Cell Stress and Chaperones and has been adapted
for presentation here.
Kelly M. Knee, Oksana A. Sergeeva, and Jonathan A. King (2013). “Human TRiC Complex
Purified from HeLa Cells Contains All Eight CCT Subunits and is Active In Vitro.” Cell Stress
and Chaperones 18:137-144. doi: 10.1007/s12192-012-0357-z © Springer.
KMK initiated the research and performed some experiments; OAS performed most
experiments and wrote the manuscript; JAK supervised the research and edited the manuscript.
51
Abstract
Archaeal and eukaryotic cytosols contain group II chaperonins, which have a double
barrel structure and fold proteins inside a cavity in an ATP-dependent manner. The most
complex of the chaperonins, the eukaryotic TCP-1 Ring Complex (TRiC), has eight different
subunits, Chaperone Containing TCP-1 (CCT1-8), that are arranged so that there is one of each
subunit per ring. Aspects of the structure and function of the bovine and yeast TRiC have been
characterized, but studies of human TRiC are very limited.
We have isolated and purified endogenous human TRiC from HeLa suspension cells.
This purified human TRiC contained all eight CCT subunits organized into double barrel rings,
consistent with what has been found for bovine and yeast TRiC. The purified human TRiC is
active as demonstrated by the luciferase refolding assay. As a more stringent test, we examined
human TRiC’s interaction with the physiological substrate human γD-crystallin. In addition to
suppressing off-pathway aggregation, TRiC was able to assist the refolding of the crystalline
molecules, an activity not found with the lens chaperone, α-crystallin. Additionally, we show that
human TRiC associates with the heat shock protein 70 (Hsp70) and heat shock protein 90
(Hsp90) chaperones. Purification of human endogenous TRiC from HeLa cells will enable
further characterization of this key chaperonin, required for the reproduction of all human cells.
52
Introduction
TRiC was first identified as a specific chaperone for actin and tubulin. In rabbit
reticulocytes, it was found that newly made tubulin subunits entered a 900-kDa complex before
becoming competent to assemble into microtubules (Yaffe et al. 1992). Concurrently,
preliminary characterization of human and mouse TCP-1 found that in both species, TCP-1
made a complex of approximately 900 kDa and that TCP-1 associated with four to six
unidentified polypeptides and two Hsp70 homologs (Lewis et al. 1992). Another simultaneous
study showed that TRiC was capable of binding and folding proteins to their native states
(Frydman et al. 1992). Structures of TRiC with tubulin and actin have since been resolved and
show that TRiC recognizes and binds these essential protein substrates that first led to its
discovery (Hynes and Willison 2000; Llorca et al. 2001; Neirynck et al. 2006; Muñoz et al.
2011). However, TRiC does not only bind actin and tubulin; Thulasiraman et al. demonstrated
that TRiC binds 9-15% of newly synthesized proteins in [35S]-methionine pulse labeled baby
hamster kidney cells (1999).
The mechanism and high-resolution structure of group II chaperonins has been
elucidated using the archaeal chaperonin of Methanococcus maripaludis, Mm-Cpn (Pereira et
al. 2010; Zhang et al. 2010; Douglas et al. 2011; Pereira et al. 2012). Due to the increased
complexity of TRiC much of the structural and biochemical research on group II chaperonins
was carried out with less complex archaeal chaperonins, such as Mm-Cpn. The most common
preparation of TRiC for scientific research is the purification of endogenous TRiC from bovine
testes tissue (Frydman et al. 1992; Ferreyra and Frydman 2000; Feldman et al. 2003).
Purification of endogenous TRiC from rabbit reticulocytes has also been effective (Gao et al.
1993; Nimmesgern and Hartl 1993; Frydman et al. 1994; Norcum 1996). More recently,
purification of exogenously tagged yeast TRiC in yeast has been developed (Pappenberger et
al. 2006; Dekker et al. 2011), along with purification of endogenous yeast TRiC by exogenously
tagging an interacting protein (Leitner et al. 2012). Co-expression of all eight human CCT
subunits in baby hamster kidney cells has been attempted but resulted in very low yields
(Machida et al. 2012). While these purifications have increased the opportunities to study TRiC,
research of human TRiC is still lacking.
Investigations of the arrangement of the CCT subunits in TRiC purified from different
species have given conflicting results (Cong et al. 2010; Dekker et al. 2011). However, a recent
novel mass spectrometry method established that bovine TRiC purified from testes tissue and
yeast TRiC purified via an interacting protein had the same arrangement with, CCT2 and CCT6
forming the homotypic contacts (Leitner et al. 2012). This does not rule out that other CCT
53
subunit arrangement of TRiC may exist, especially in different tissues and at different
developmental stages. Previous research has shown that TRiC complexes containing specific
subunits may have different roles (Roobol et al. 1995) and even that the CCT subunits may
have functions in the cell independent of the TRiC complex (Roobol and Carden 1999).
The largest divergence of sequence between the CCT subunits is in the apical
substrate-recognition domain, suggesting that the different subunits of TRiC may have evolved
distinctive subunit specificities (Kim et al. 1994; Frydman 2001; Spiess et al. 2006). Although
only a few substrates have been studied, it is clear that not all CCT subunits are involved in
binding of non-native-state substrates to TRiC (Hynes and Willison 2000; Llorca et al. 2001;
Feldman et al. 2003; Spiess et al. 2006). The apical domains of CCT1 and CCT4 have been
implicated in TRiC binding to exon one of the huntingtin protein (Tam et al. 2006), while Spiess
et al. demonstrated that TRiC subunits CCT1 and CCT7 bind pVHL (2006). While these studies
show that not all CCT subunits are required to bind a substrate to TRiC, we cannot firmly
conclude whether only specific CCT subunits can bind particular substrates.
While there is some, albeit far from complete, knowledge of the CCT subunit
arrangement and the recognition of substrates by specific CCT subunits, there has been little
study on the assembly of TRiC from the CCT subunits. With the eight CCT subunits expressed
from seven different genes, the assembly of TRiC must be very finely regulated (Kubota et al.
1999). This regulation may mean that the TRiC rings can contain a different arrangement and
ratio of CCT subunits at some point in their lifetime. To investigate whether human TRiC found
in epithelial cells is the same as bovine TRiC from testes tissue, we purified endogenous TRiC
from an epithelial cell derivative line, HeLa. HeLa cells have previously been shown to express
all eight CCT proteins at high levels (Fountoulakis et al. 2004), making this an ideal cell line for
endogenous TRiC purification.
54
Materials and Methods
TRiC Purification from HeLa Cells
A starter culture of HeLa suspension cells (HeLa-S3; ATCC) was grown in S-MEM
(Sigma) supplemented with 10% fetal bovine serum (FBS), 1% l-Glu, and 1% Penicillin and
Streptomycin. From this starter culture, Cell Essentials, Inc. (Boston, MA) grew a 20 L
suspension culture of HeLa cells, resulting in a cell pellet of approximately 100 g.
All of the following steps were preformed at 4 °C. The HeLa cells were lysed following
the HeLa nuclear extraction protocol (Tran et al. 2001). Briefly, the pellet was washed twice with
iced phosphate buffer (137 mM NaCl, 2.68 mM KCl, 4.29 mM Na2HPO4, 1.47 mM KH2PO4). The
packed cell volume (PCV) of the pellet was determined and the pellet was resuspended in two
PCVs of hypotonic buffer A (10 mM Tris, pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT) and
mixed thoroughly. The HeLa cells were dounced 35 times with pestle B. The dounced cells were
centrifuged at 2,500 × g for 15 minutes, resulting in three layers. The top cytoplasmic layer
contained TRiC and was therefore supplemented with 1 mM ATP and used in the subsequent
purification.
The human TRiC purification hereafter loosely follows the bovine TRiC purification
described by Ferreyra & Frydman (2000). Two ammonium sulfate precipitations (25% then
55%) were preformed on the cytosolic fraction isolated above. Human TRiC was found in the
supernatant of the 25% ammonium sulfate cut and the pellet of the 55% ammonium sulfate cut.
This pellet was dissolved in a minimal volume of MQ-A (20 mM HEPES/KOH, pH 7.4, 50 mM
NaCl, 5 mM MgCl2, 10% glycerol, 1 mM DTT, 0.1 mM PMSF, 0.1 mM EDTA, 1 mM ATP) and
placed in 50-kDa MWCO dialysis tubing (SpectraPor) and dialyzed twice (2 hours to overnight)
against MQ-A at 4 °C.
The dialyzed sample was centrifuged at 15,000 × g to remove aggregates and passed
over a HiLoad 26/10 Q sepharose column (GE Healthcare). Human TRiC was eluted off of this
column by 40% MQ-B (MQ-A with 1 M NaCl). The fractions containing TRiC were pooled,
diluted in half by MQ-A, and applied to a Heparin HiTrap HP 5x5 mL column (GE Healthcare).
Human TRiC eluted during a 14 column volume gradient from 20% to 65% MQ-B. The fractions
containing TRiC were pooled and concentrated down to 1 mL using Vivaspin ultraconcentrators
(Satorius Stedim). This sample was loaded on a Superose 6 10/300 GL size exclusion column
(GE Healthcare). Human TRiC eluted by MQ-A around 12-14.5 mL of the size exclusion
column, consistent with that of a 1 MDa complex. These fractions were pooled, concentrated,
and the protein concentration was measured using the BCA assay (Pierce) with BSA as the
standard.
55
SDS-PAGE and Immunoblots
Proteins were separated by SDS-PAGE (14%) at 165 V for 1 h after boiling in reducing
buffer (60 mM Tris, pH 6.8, 2% SDS, 5% β-mercaptoethanol, 10% glycerol, bromophenol blue
for color) for 5 min. The gels were stained with Coomassie blue or Krypton (Pierce). Transfer
was conducted for 1.5 h at 300 mA in transfer buffer (10% methanol, 25 mM Tris, 192 mM
glycine) onto 0.45 µm polyvinylidene difluoride (PVDF) membranes (Millipore).
The primary antibodies used for CCT1-8 were from Santa Cruz Biotechnology: CCT1,
sc-53454; CCT2, sc-28556; CCT3, sc-33145; CCT4, sc-58865; CCT5, sc-13886; CCT6, sc100958; CCT7, sc-130441; and CCT8, sc-13891. The Hsp70 and Hsp90 antibodies were from
Enzo Life Sciences: Hsc70/Hsp70, SPA-820; and, Hsp90a, SPA-840. The secondary antibodies
were Alkaline Phosphatase (AP)-conjugated (Millipore) and the membranes were visualized
using AP-conjugate substrate kit (BioRad).
Electron Microscopy
Copper grids with Formvar carbon coating (400 mesh, Ted Pella) were glow discharged
for 20 s and 5 µL of purified human TRiC was placed on the grids for 5 min. Excess sample on
the grids was blotted off using filter paper and the grids were floated onto a drop of filtered 1.5%
uranyl acetate (Sigma-Aldrich) for 45 s. Grids were visualized under a JEOL 1200 SX
transmission electron microscope (TEM), and digital micrographs were taken using an AMT
16000S camera system.
Luciferase Refolding Assay
The luciferase refolding assay was preformed as described in Thulasiraman et al.
(2000). Briefly, 8.2 µM of luciferase (Promega) was unfolded in unfolding buffer (6 M guanidine
hydrochloride, 25 mM HEPES/KOH pH 7.4, 50 mM KOAc, and 5 mM DTT) at room temperature
for 1 hour with mixing. The unfolded luciferase was diluted 1:40 (205 nM) in unfolding buffer and
then further diluted 1:25 (8.2 nM) into refolding buffer (25 mM HEPES/KOH, pH 7.4, 100 mM
KOAc, 10 mM Mg(OAc)2, 2 mM DTT, 1 mM ATP, 10 mM creatine phosphate, 40 U/mL creatine
kinase, 2% DMSO) with or without 400 nM of purified human TRiC. At various time points, an
aliquot of the refolding reaction was diluted 1:25 into Steady-Glo Assay Reagent buffer
(Promega) and luminescence was measured on a FLUOstar Optima plate reader (BMG
Labtech) with FLUOstar Optima software.
56
Human γD-Crystallin Aggregation Suppression Assay
The aggregation suppression assay is described in detail in Knee et al. (2011). Briefly,
23 µM human γD-crystallin was unfolded overnight at 37 °C in unfolding buffer (5.5 M guanidine
hydrochloride, 50 mM Tris-HCl, pH 7.5, and 5 mM DTT). To initiate aggregation the unfolded
protein was diluted 1:10 (2.3 µM) into refolding buffer (50 mM Tris-HCl, pH 7.5, 1 mM DTT, 50
mM KCl, 5 mM MgCl2, 1 mM ATP) with or without 2.3 µM purified human TRiC. Aggregation
kinetics were measured at 350 nm on a Cary UV/Vis spectrophotometer (Varian) using the
Varian Kinetics program.
57
Results
Purification
The first step in purifying endogenous human TRiC from HeLa cells was separating the
cytoplasmic fraction of the cells from the nuclei, because TRiC is a cytoplasmic chaperonin.
This was accomplished using a HeLa nuclear extraction protocol (Tran et al. 2001) and verified
by immunoblots probed with the CCT1 primary antibody (Figure 2-1). Next, a series of
ammonium sulfate cuts further purified TRiC from other HeLa proteins. The resuspended pellet
was passed over three chromatography steps: anion exchange (Q sepharose), Heparin affinity,
and Superose-6 size exclusion chromatography. The elution peak from the size exclusion
column was between 12 and 14.5 mL (Figure 2-2). This was consistent with other TRiC
purifications and the purification of Mm-Cpn (Frydman et al. 1994; Reissmann et al. 2007). The
average yield of this purification from a 100 g HeLa cell pellet was 5 mg of purified human TRiC
with ~90% purity.
When Heparin affinity chromatography was omitted from the purification, Hsp70 and
Hsp90 co-purified with human TRiC (Figure 2-3). This interaction may be due to human TRiC
binding to Hsp70 and Hsp90 in HeLa cells, while exchanging substrates between the
chaperones. When Heparin affinity chromatography was utilized, Hsp70 and Hsp90 were not
seen in the purified human TRiC sample, demonstrating that while this interaction was quite
robust, it could be eliminated.
Structure
All eight CCT subunits were present in the purified human TRiC sample as seen by
immunoblots probed with antibodies against each of the eight subunits (Figure 2-4). They were
all present in roughly equal stoichiometry. By negative stain TEM, purified human TRiC
appeared as two back-to-back rings approximately 185 Å in height and 165 Å in diameter
(Figure 2-5). The morphology of purified human TRiC was consistent with that of purified TRiC
reported in the literature (Cong et al. 2010; Dekker et al. 2011).
58
Figure 2-1: Human TRiC primarily limited to the cytoplasmic fraction of HeLa cells
Immunoblot probed with CCT1 of three layers seen after the lysis: cytoplasmic layer (C), middle
layer (M), and nuclei layer (N). Most of the CCT1 (labeled), and therefore TRiC, was found in
the cytoplasm of the lysed cells.
59
Figure 2-2: Human TRiC purified by size exclusion chromatography
The input (inp) and elution volumes (7.5-14.5 mL) are shown on Coomassie-stained 14% SDSPAGE. TRiC (labeled) appears as a series of bands ~60 kDa in size that are eluted in volumes
of 12-14.5 mL consistent with a 1 MDa complex.
60
Figure 2-3: Hsp70 and Hsp90 co-purified with TRiC when heparin affinity chromatography was
omitted
Immunoblots probed with antibodies against CCT1, Hsp70, and Hsp90 clearly show Hsp70 and
Hsp90 present in the purified human TRiC sample when heparin affinity chromatography was
not used.
61
Figure 2-4: All eight subunits present in purified human TRiC
A. A series of bands consistent with TRiC were present in the final purified human TRiC sample
as shown on Coomassie-stained 14% SDS-PAGE. B. Immunoblots probed with all 8 CCT
primary antibodies show that the purified human TRiC sample contains all eight CCT subunits in
approximately equal ratios. There are no degradation products of the subunits in the purified
human TRiC sample.
62
Figure 2-5: Negative stain TEM of purified human TRiC reveal double rings
The morphology of human TRiC is consistent with that of TRiC purified from other species. The
complexes were ~165 Å in diameter and ~185 Å in height and shown here at 150 K
magnification. Scale bar: 100 nm.
63
Activity
Human TRiC purified from HeLa cells was active in the luciferase refolding assay, which
has been previously used to test activity of TRiC purified from bovine testes (Frydman et al.
1992) and rabbit reticulocytes (Nimmesgern and Hartl 1993; Frydman et al. 1994). In the assay,
luciferase was unfolded and then diluted into buffer with purified human TRiC (Thulasiraman et
al. 2000). The presence of refolded luciferase in the mixture was assayed by addition of luciferin
and subsequent luminescence production monitoring. Purified human TRiC refolded luciferase
for over two hours at room temperature (Figure 2-6).
Though luciferase has frequently been used to assay the refolding activity of a variety of
chaperonins, it is not an authentic substrate for human TRiC. A human protein whose folding
and competing aggregation has been systematically studied is human γD-crystallin (HγD-Crys)
(Kosinski-Collins and King 2003; Flaugh et al. 2005). TRiC is almost certainly present in lens
epithelium cells and primary lens fibers (Hoehenwarter et al. 2008), making the interaction
between HγD-Crys and TRiC an authentic one. In characterizing the activity of the archaeal
chaperonin Mm-Cpn, Knee et al. found that Mm-Cpn both suppressed the aggregation of HγDCrys, but also enhanced its refolding in vitro (2011). The major lens chaperone, α-crystallin, is
holdase so it can suppress aggregation of substrates, but not refold them (Moreau and King
2012). Likewise, in vitro, α-crystallin can suppress HγD-Crys aggregation, but cannot refold the
molecules (Acosta-Sampson and King 2010; Moreau and King 2012).
Therefore, we decided to assess whether human TRiC was active with respect to the
HγD-Crys substrate. In this assay, when unfolded HγD-Crys was diluted from denaturant into
buffer at concentrations of 50 µg/mL, partially folded intermediates partitioned between
productive refolding and off-pathway aggregation. This aggregation was monitored by sample
turbidity. When purified human TRiC was added to the buffer, aggregation was significantly
suppressed (Figure 2-7A). Furthermore, native-like HγD-Crys could be detected when the
filtered sample was assessed on SDS-PAGE (Figure 2-7B), suggesting HγD-Crys was refolded
by human TRiC.
In summary, human TRiC was purified from HeLa cells by first extracting the cytoplasmic
layer of the cells and then performing three chromatography steps. The purified material
contained all eight CCT subunits in approximately equal ratios. TRiC could be effectively
separated from other chaperones in the cells by Heparin affinity chromatography. Purified
human TRiC possessed the back-to-back ring morphology that defines the structure of
chaperonins. Our purified human TRiC was not only active in refolding the model substrate
luciferase, but also suppressed the aggregation and refolded the authentic substrate HγD-Crys.
64
Figure 2-6: Purified human TRiC active in refolding luciferase
Human TRiC (blue) refolds luciferase more efficiently than the BSA (green) or water (red)
controls. Human TRiC is active over two hours at room temperature.
65
Figure 2-7: Purified human TRiC suppression of HγD-Crys aggregation and HγD-Crys nativelike state refolding
A. Aggregation of HγD-Crys (red) can be suppressed by the addition of human TRiC (blue) by
approximately 80% after fifteen minutes at 37 °C. B. When filtered, the aggregation suppression
samples were observed on Krypton-stained 14% SDS-PAGE. The sample with purified human
TRiC showed a band corresponding to HγD-Crys indicating that human TRiC can refold HγDCrys to a native-like state.
66
Discussion
While TRiC has been readily purified endogenously from bovine testes (Frydman et al.
1992; Leitner et al. 2012) and pseudo-exogenously from yeast (Pappenberger et al. 2006;
Leitner et al. 2012), purification of human TRiC has been limited. Expression of all eight
subunits exogenously from the cloned genes is difficult. The direct approach of growing a large
amount of human epithelial cells and purifying endogenous TRiC has been successful in
producing the authentic human chaperonin.
Molecular chaperones have been postulated to be viable therapeutic targets (Almeida et
al. 2011). There has been evidence that TRiC may play a role in suppressing Huntington’s
disease by decreasing huntingtin aggregate formation (Kitamura et al. 2006; Tam et al. 2009). If
TRiC is to be used as a therapeutic in the clinic, it will be necessary to study human TRiC to
further understand the chaperonin function inside human cells. While the TRiC isolated from
bovine testes may overall be similar to the human version, there are differences in all eight
subunits between bovine and human TRiC, let alone any type of arrangement or assembly
differences that have yet to be elucidated.
Human TRiC activity in assisting the refolding of firefly luciferase corresponds to
activities reported for other mammalian TRiC complexes (Frydman et al. 1992; Nimmesgern
and Hartl 1993; Frydman et al. 1994). The lens crystallins represent a more rigorous test for the
activity of human TRiC. The lens crystallins must remain stable and folded throughout life for the
aggregated state results in the lens disease cataract (Moreau and King 2012). The α-crystallin
chaperone present at high concentrations in the lens suppresses the aggregation of HγD-Crys,
but cannot refold it (Horwitz 1992; Evans et al. 2008; Acosta-Sampson and King 2010). The
results showed that human TRiC both suppressed the aggregation of partially folded HγD-Crys
and, in the presence of ATP, was able to refold the chains. Human TRiC may in fact play a role
in protecting the lens fibers from cortical cataract (Mitchell et al. 1997).
We found that Hsp70 and Hsp90 bound to purified human TRiC if one of the
chromatography steps is omitted. This is not surprising for it has been shown that TRiC copurifies with Hsp70 and Hsp90 in rabbit reticulocytes (Nimmesgern and Hartl 1993; Frydman et
al. 1994). However, while the shuttling of substrates between Hsp70 and TRiC has been widely
studied (Kabir et al. 2011), substrate exchange from TRiC to Hsp90 is much less understood.
The substantial amounts of Hsp90 that co-purified with TRiC from HeLa cells may make this a
good system for further studying the TRiC-Hsp90 interaction.
Other further directions with human TRiC will be to attempt high-resolution structural
studies for comparison to yeast and bovine TRiC. The arrangement of human TRiC purified
67
from HeLa cells may be different than that of bovine TRiC purified from testes, not only because
of the differences in species as mentioned above but also due to differences between the tissue
and the cells in culture. Furthermore, it will be interesting to see whether purified human TRiC
can refold actin and tubulin, the two largest substrates of TRiC, as efficiently as bovine TRiC.
Consequently, we plan to study whether each CCT subunit is needed to recognize and refold
particular substrates, such as actin and tubulin.
It has been postulated that the eight different CCT subunits of TRiC are needed to
recognize a variety of substrates (Llorca et al. 2001; Feldman et al. 2003). The CCT subunits
may recognize different types of proteins e.g., CCT2 may recognize beta-propeller proteins,
while CCT8 may recognize hydrophobic beta sheets. Even more specifically the CCT subunits
may recognize different proteins e.g., CCT1 may bind huntingtin (Tam et al. 2006) while CCT7
recognizes pVHL (Spiess et al. 2006). However, with our limited knowledge on the substrate
recognition of TRiC, it is unknown if different CCT subunits specifically or redundantly recognize
these various substrates. It may be that while CCT1 recognizes huntingtin with the highest
efficiency, CCT4 or CCT7 can bind it when CCT1 is not present or defective.
The arrangement of TRiC has been determined for TRiC purified from bovine testes and
from yeast, but it is unknown how this arrangement varies among tissues or at different
developmental stages. Also unknown, as alluded to above, is how TRiC can assemble into this
arrangement. As with other large complexes, it is likely that an extremely regulated sequence of
events is needed for the final arrangement. It has recently been shown that chaperonin-like
Bardet-Biedl syndrome (BBS) subunits assemble into the final BBSome complex by sequential
addition of each subunit (Zhang et al. 2012). Such fine sequential assembly is possible for TRiC
as well, therefore requiring more research about this complex chaperonin. Our purified TRiC
material is an important first step for furthering the knowledge on this crucial human chaperonin.
68
CHAPTER 3:
Human CCT4 and CCT5 Chaperonin Subunits
Expressed in E. coli Form Biologically Active Homo-oligomers*
* This research was originally published in the Journal of Biological Chemistry and has been
adapted for presentation here.
Oksana A. Sergeeva, Bo Chen, Cameron Haase-Pettingell, Steven J. Lutdke, Wah Chiu, and
Jonathan A. King (2013). “Human CCT4 and CCT5 Chaperonin Subunits Expressed in E. coli
Form
Biologically
Active
Homo-oligomers.”
J.
Biol.
Chem.
288:17734-17744.
doi:
10.1074/jbc.M112.443929 © The American Society for Biochemistry and Molecular Biology.
OAS initiated the research, performed most experiments, and wrote the manuscript; BC
performed some experiments and computational analysis, CHP performed some experiments,
SJL supervised the research and performed computational analysis; WC supervised the
research and edited the manuscript; JAK supervised the research and edited the manuscript.
69
Abstract
Chaperonins are a family of chaperones that encapsulate their substrates and assist
their folding in an ATP-dependent manner. The ubiquitous eukaryotic chaperonin, TCP-1 Ring
Complex (TRiC), is a hetero-oligomeric complex composed of two rings each formed from eight
different CCT (Chaperonin Containing TCP-1) subunits. Each CCT subunit may have distinct
substrate recognition and ATP-hydrolysis properties. We have expressed each human CCT
subunit individually in E. coli to investigate whether they form chaperonin-like double ring
complexes. CCT4 and CCT5, but not the other six CCT subunits, formed high molecular weight
complexes within the E. coli cells that sedimented about 20S in sucrose gradients.
When CCT4 and CCT5 were purified, they were both organized as two back-to-back
rings of eight subunits each, as seen by negative stain and cryo-electron microscopy. This
morphology is consistent with that of the hetero-oligomeric double-ring TRiC purified from
bovine testes and HeLa cells. Both CCT4 and CCT5 homo-oligomers hydrolyzed ATP at a rate
similar to human TRiC, and were active as assayed by luciferase refolding and human γDcrystallin aggregation suppression and refolding. Thus both CCT4 and CCT5 homo-oligomers
have the property of forming eight-fold double rings absent the other subunits, and these
complexes carry out chaperonin reactions without other partner subunits.
70
Introduction
The eukaryotic group II chaperonin TRiC consists of two identical rings, each with eight
different CCT subunits (Cong et al. 2010). Through a variety of structural, functional, and cell
biology methods, interactions between TRiC and its main substrates, actin and tubulin, have
been well characterized (Frydman et al. 1992; Lewis et al. 1992; Yaffe et al. 1992; Hynes and
Willison 2000; Llorca et al. 2001; Neirynck et al. 2006; Muñoz et al. 2011). However, TRiC
binding is not limited to actin and tubulin; TRiC binds 9-15% of newly synthesized proteins in
[35S]-methionine pulse labeled baby hamster kidney cells (Thulasiraman et al. 1999). Recent
research has focused on the arrangement of the eight CCT subunits in TRiC, the binding and
hydrolysis of ATP in TRiC, and the recognition of substrates by specific CCT subunits of TRiC.
The arrangement of CCT subunits in TRiC has been a source of controversy (Liou and
Willison 1997; Martín-Benito et al. 2007; Cong et al. 2010; Dekker et al. 2011). However,
recently, a novel method has established a consistent arrangement for bovine and yeast TRiC
with CCT2 and CCT6 making homo-typic contacts between the rings (Kalisman et al. 2012;
Leitner et al. 2012). This does not explicitly exclude the existence of other CCT subunit
arrangements. With the eight CCT subunits expressed from seven different genes, the
assembly of TRiC must be regulated to insure one of each subunit per mature ring (Kubota et
al. 1999). In fact, TRiC could contain a different arrangement and ratio of CCT subunits in
different tissues, or in different stages of embryonic development. Furthermore, there is
evidence that TRiC variants containing specific subunits may have different roles (Roobol et al.
1995) and that the CCT subunits may have additional functions in the cell independent of TRiC
chaperonin function (Roobol and Carden 1999).
It has recently been found that the different CCT subunits of TRiC bind ATP with
different affinities (Reissmann et al. 2012). In order for the TRiC chaperonin to close, every
subunit does not need to bind ATP, unlike the ATP binding mechanism in GroEL/ES (Horwich et
al. 2007), where every GroEL subunit has to bind an ATP for closure. Only four of the CCT
subunits (CCT1, CCT2, CCT4, and CCT5) seemed to bind ATP at physiological concentrations,
representing high ATP-affinity subunits (Reissmann et al. 2012). Introducing ATP-bindingdeficient and ATP-hydrolysis-deficient mutations into the other subunits (CCT3, CCT6, CCT7,
and CCT8) in yeast did not affect yeast growth (Reissmann et al. 2012). Combining this
information with the recent consistent arrangement of CCT subunits around TRiC (where the
high ATP-affinity subunits are located together on one half of the ring), suggests that the high
ATP-affinity subunits regulate an asymmetrical power stroke that drives ATP hydrolysis (Leitner
et al. 2012; Reissmann et al. 2012).
71
The apical substrate-recognition domain exhibits the largest divergence of sequence
among the CCT subunits, suggesting that this heterogeneity among CCT subunits evolved to
recognize and refold a variety of substrates in the eukaryotic cytosol (Kim et al. 1994; Frydman
2001; Spiess et al. 2006). Although only a limited number of substrates have been investigated,
binding of non-native-state substrates to TRiC may not involve all CCT subunits (Hynes and
Willison 2000; Llorca et al. 2001; Feldman et al. 2003; Spiess et al. 2006). Many substrates
appear to bind across the ring, thus contacting subunits on either side of the ring (Llorca et al.
2000; Martín-Benito et al. 2004). CCT1 and CCT7 bind pVHL (Spiess et al. 2006), while CCT1
and CCT4 bind polyglutamines such as those in exon one of the mutant huntingtin protein (Tam
et al. 2006; Sontag et al. 2013). Not all CCT subunits bind a substrate, but it is unknown
whether only specific CCT subunits can bind a particular substrate.
Eukaryotic TRiC has been purified from yeast (Pappenberger et al. 2006; Dekker et al.
2011; Leitner et al. 2012), from bovine (Frydman et al. 1992; Ferreyra and Frydman 2000;
Feldman et al. 2003) and mouse (Liou and Willison 1997; Llorca et al. 1999; Llorca et al. 2000)
testes, and more recently from HeLa cells (Knee et al. 2013). The potential of TRiC as a target
of therapeutic agent will benefit from access to human TRiC (Knee et al. 2013). However,
purification of human TRiC from HeLa cells is expensive (Knee et al. 2013) and recombinant coexpression of all eight CCT subunits has resulted in very low yields (Machida et al. 2012). In
order to understand how individual human CCT subunits function (both in terms of ATP binding
and hydrolysis, and substrate recognition and folding), we have successfully expressed single
subunits in E. coli. When we purified two of the CCT proteins, CCT4 and CCT5, to our surprise,
they were organized into chaperonin-like homo-oligomeric rings that exhibited chaperonin
activities.
72
Materials and Methods
CCT Subunit Expression
The pET21b vector was modified (pET21b*) to include a TEV protease cleavage site
between the end of the inserted gene and the C-terminal 6x-His tag. The human CCT genes
were synthesized by Genescript (Piscataway, NJ) and inserted into the pET21b* vector using
the following restriction sites per gene: CCT1, NdeI and NheI; CCT2, NdeI and BamHI; CCT3,
SacII and BamHI; CCT4, NdeI and BamHI; CCT5, NdeI and BamHI; CCT6, NdeI and BamHI;
CCT7, SacII and BamHI; CCT8, SacII and BamHI. Each plasmid was confirmed by sequencing
(Genewiz). Recombinant single CCT subunits were prepared by plasmid transformation into E.
coli BL21 (DE3) RIL cells. The cells were grown in Super Broth to OD 5.0 at 37 °C and then
shifted to 18 °C and induced with 0.5 mM IPTG. After an overnight induction, cultures were
pelleted by centrifugation for 15 min. The cells were resuspended in CCT-A (20 mM
HEPES/KOH pH 7.4, 300 mM NaCl, 10 mM MgCl2, 10% glycerol, 1 mM DTT, 1 mM ATP) with
addition of one EDTA-free Complete protease inhibitor (Roche) per L of culture.
CCT Subunit Purification
After the addition of 1 mM DTT, 5 mM MgCl2, and 5 µg/mL of DNase, the cells were
lysed via French Press at a pressure of 16,000 pounds per square inch. The lysate was
centrifuged at 20,000 x g for 45 min. The supernatant was removed by pipetting, 0.45 µm
filtered, and passed over a Ni-NTA column (Pierce). After loading, the column was first washed
with 100% CCT-A, then the CCT single subunit was eluted off of the column in a linear gradient
from 10 to 100% CCT-B (CCT-A but with 250 mM imidazole). The fractions containing the CCT
single subunit were combined and concentrated using Vivaspin ultraconcentrators (Satorius
Stedim). The protein was diluted with CCT-A down to 25 mM imidazole. After the addition of
TEV protease, the CCT single subunit was incubated over night at 4 °C with gentle rocking.
The His-tag-cleaved CCT single subunit was 0.45 µm filtered and applied again to the
Ni-NTA column, to which it no longer bound. The flow through fractions containing the CCT
single subunit were combined, further concentrated, and passed over a Superose 6 10/300 GL
size exclusion column (GE Healthcare). CCT4 and CCT5 single subunits eluted by CCT-SEC
(CCT-A but with 5% glycerol and no ATP) around 12-14.5 mL off of the size exclusion column,
consistent with that of a 1 MDa complex. These fractions were pooled, concentrated, and the
protein concentration was measured using the BCA assay (Pierce) with BSA as the standard.
The purified CCT subunit band was cut out, trypsin digested, and LC-MS/MS analysis was run
on a Qstar mass spectrometer by Biopolymer and Proteomics Core Facility at the Koch Institute
73
(Cambridge, MA). Peptides were identified by searching for hits in the Mascot database. Nterminal sequencing was conducted by Tufts Medical Core Facility (Boston, MA).
Human TRiC and Mm-Cpn Purification
The human TRiC control sample was purified as described at length in Knee et al.
(2013). Mm-Cpn was purified as described in Knee et al. with the slight variation that the protein
was grown up in Super Broth (2011).
Sucrose Gradient Sedimentation
Isokinetic 5-40% sucrose (in CCT-SEC buffer) gradients were prepared via the gradient
master (BioComp Instruments) and ultracentrifuged at 4 °C using a SW50 rotor for 18 h at
28,000 rpm (Beckman). Twenty-four fractions were collected using a gradient fractionator
(BioComp Instruments).
SDS-PAGE and Immunoblots
Proteins were separated by SDS-PAGE (14% or 10%) at 165 V for 1 h after boiling in
reducing buffer (60 mM Tris, pH 6.8, 2% SDS, 5% β-mercaptoethanol, 10% glycerol,
bromophenol blue for color) for 5 min. The gels were stained with Coomassie blue or Krypton
(Pierce). Transfer was conducted for 1.5 h at 300 mA in transfer buffer (10% methanol, 25 mM
Tris, 192 mM glycine) onto 0.45 µm polyvinylidene difluoride (PVDF) membranes (Millipore).
The primary antibodies used for CCT1-8 were from Santa Cruz Biotechnology: CCT1, sc-53454;
CCT2, sc-28556; CCT3, sc-33145; CCT4, sc-58865; CCT5, sc-13886; CCT6, sc-100958;
CCT7, sc-130441; and CCT8, sc-13891. The secondary antibodies were Alkaline Phosphatase
(AP)-conjugated (Millipore) and the membranes were visualized using the AP-conjugate
substrate kit (BioRad).
Electron Microscopy
Copper grids with Formvar carbon coating (400 mesh, Ted Pella) were glow discharged
for 20 s and 5 µL of purified chaperonin was placed on the grids for 5 min. Excess sample on
the grids was blotted off using filter paper and the grids were floated onto a drop of filtered 1.5%
uranyl acetate (Sigma-Aldrich) for 45 s. Grids were visualized under a JEOL 1200 SX
transmission electron microscope (TEM), and digital micrographs were taken using an AMT
16000S camera system.
74
Cryo-Electron Microscopy
For the apo state, 0.5 mg/mL CCT4 or CCT5 was used for cryo-EM sample preparation.
For the closed state, 0.35 mg/mL CCT5 was incubated for 30 min at 37 °C with 5 mM Al(NO3)3,
30 mM NaF, and 1 mM ATP. A volume of 2.5 µL was applied onto a plasma-cleaned grid
(R1.2/1.3, Quantifoil Micro Tools) and plunge-frozen into liquid ethane operated automatically by
Vitrobot Mark III device (FEI). Images for both states were taken at 71,361x detector
magnification on a JEM 2200FS microscope (JEOL) with Omega in-column energy filter (energy
slit = 20 eV) and recorded on a Gatan 4Kx4K CCD with a dose of 20 e/A2. A total of 54 (apo
state CCT5) and 160 (ATP-AlFx state CCT5) CCD frames were taken with defocuses range
from 2 mm - 3.5 mm.
A total of 5,000 particles (apo) or 6,307 particles (ATP-AlFx state) were boxed out semiautomatically by e2boxer.py (EMAN2) (Tang et al. 2007). The reference-free 2D class averages
were generated by using e2refine2d.py (EMAN2) to group 5,000 particles in each dataset into
100 classes. For 3D reconstruction, all the 6,307 particles in ATP-AlFx state were fit using the
automatic CTF fitting program fitctf.py (EMAN), and then manually examined and adjusted using
the EMAN program ctfit.py. A C8 symmetry-imposed multiple model refinement approach was
used initially to separate conformationally heterogeneous particles into different groups using
multirefine as previously described (Chen et al. 2006; Cong et al. 2011). The initial model was
generated from the dataset by startcsym.py (EMAN). Three initial models for refinement were
generated by adding different random noise at a level of σ=0.15 to the initial model and the
dataset is divided into three subclasses. After iterative refinements, the refined models
converged and one refined model was associated with 2,974 (~47%) particles. This subset of
particle images was re-processed from scratch without imposing any symmetry.
An initial asymmetric model was generated for this subset of particle images and no
symmetry was applied during the refinement process. When the refined map converged, a
Gaussian low-pass filter with cut-off frequency of 0.04 Å-1 (25Å) was applied to interpret the lowresolution features of the map. A rotational correlation plot was used to assess the symmetry of
the map, and a strong 8-fold symmetry was observed.
After C8 symmetry was observed in the map, it was imposed during another round of 3D
refinement (EMAN) providing a C8-symmetrized initial model. After refinement, the final
resolution was measured to be 22 Å (0.143 criterion (Rosenthal and Henderson 2003)). To
further validate the map, the dataset of 2,974 particles was divided into two independent subdatasets, with 1,487 particles each. Two phase-randomized symmetry-free models were used
as initial models for these two datasets. The phase-randomized models were generated as
75
follows: symmetry-free refined model was subject to a phase randomization process by using a
low-pass phase randomization filter in EMAN2 to randomize the Fourier phases below 33Å.
Fourier Shell Correlation (FSC) between these two template models demonstrated the expected
zero mean correlation beyond 33Å resolution. The choice of 33 Å was based on an expectation
that the final resolution would be better than this. The two independent datasets were then
refined independently from these 2 starting models. This “gold standard” resolution (Scheres
and Chen 2012) was measured as 22 Å based on the FSC=0.143 cut-off criterion (Rosenthal
and Henderson 2003).
Thermal Denaturation by Circular Dichroism
The secondary structure of the chaperonins was assayed at 100 µg/mL of protein in
filtered and degassed 10 mM Tris, 20 mM KCl. For protein in the closed state, 1 mM ATP-γS
was added to the buffer. Temperature was raised from 25 to 100 °C in 5 °C steps and
equilibrated for 5 min at each temperature. Far-UV circular dichroism (CD) spectra from 260 nm
to 195 nm were obtained for each chaperonin and the buffer using an AVIV Model 202 CD
spectrophotometer at each temperature. The buffer was subtracted at each temperature and the
signal at 227 nm was selected for thermal denaturation analysis. Transition midpoints were
determined using a two-state unfolding fit in Prism (GraphPad).
ATP Hydrolysis Assay
The ATP hydrolysis assays were preformed as described in Reissmann et al. (2007).
Briefly, 250 nM of chaperonin was incubated for 5 min at 30°C in 1.25x reaction mix. At time
zero, ATP was added to a final concentration of 2 mM with [α-32P]ATP (Perkin Elmer) at a
concentration of 0.002 µCi/µL, and the reaction proceeded at 30°C. At each indicated point, 2
µL of the sample was taken out of the reaction and spotted onto a polyethelenimine (PEI)cellulose thin layer chromatography (TLC) plate (Macherey-Nagel). The plates were run using a
mobile phase of 1 M LiCl and 2 M formic acid, air-dried, and exposed to a phosphorimager.
After 24 hours, the screen was scanned by a Typhoon imager (GE Healthcare), and the amount
of [α-32P]ADP was quantified using ImageJ.
Luciferase and Human γD-Crystallin Refolding Assays
The luciferase refolding assay was performed as described in Knee et al. (2013). The
human γD-crystallin aggregation suppression assay is described in detail in Acosta-Sampson &
76
King (2010) and Knee et al. (2011) and was modified in this study by use of a decreased
chaperonin concentration of 145 nM.
77
Results
Expression and Purification of CCT Subunits
The CCT subunits were successfully cloned into a modified pET21b vector that included
a TEV protease site before the C-terminal 6x-His tag. Due to the variations in DNA sequences
of the CCT subunits, two of four different restriction enzymes were used to insert each CCT
subunit DNA sequence into the vector. Of four different E. coli expression lines – (BL21 (DE3)
Gold, BL21 (DE3) pLysS, Rosetta (DE3) pLysS, BL21 (DE3) RIL) – BL21 (DE3) RIL, was found
to express full length CCT subunit protein to the highest level. The CCT subunit sequences
were non-optimized human sequences and the BL21 (DE3) RIL cell line is enhanced for
expressing human sequences in E. coli. Some of the CCT subunits accumulated at much higher
levels than other CCT subunits as seen by the cells lysates electrophoresed through 10% SDSPAGE and Coomassie stained (Figure 3-1A). The expression levels were verified by
immunoblots of the cell lysate proteins probed by each of the respective CCT antibodies (Figure
3-1B).
For a number of the subunits, lower molecular species were clearly visible in the
immunoblots. Four of these antibodies are monoclonal – CCT1, CCT4, CCT6, CCT7 –
indicating that the lower molecular weight species are CCT fragments produced by degradation.
These fragment levels were not sensitive to time of incubation of the lysates, suggesting that
proteolysis was happening within the expressing cells. Variation of temperature of cell growth,
conditions of induction, and treatment of the lysed cells, did not have a significant effect on the
differences in expression among the eight subunits. A 53-kDa fragment was present in the
CCT4 expression. Mass spectrometry and N-terminal sequencing identified the fragment to lack
the first 60 amino acids of CCT4. We considered that fragment might be the result of late
translation initiation, but mutations of the suspected methionine did not significantly decrease
the level of the fragment. Therefore, the fragment might be the result of a specific protease
acting within the cell.
In our attempts to express subunits without a His-tag, we had difficulty separating the
CCT subunits from the endogenous GroEL/S complexes. Though we cannot rule out differences
in transcription or translation, we suspect that the differences in subunit accumulation may
reflect whether or not the translated CCT chains are able to utilize the E. coli
chaperone/chaperonin apparatus to assist their folding and assembly.
78
Figure 3-1: Expression of human CCT subunits in BL21 (DE3) RIL E. coli cells
A. Cells expressing each of the eight subunits (CCT1-8), an Mm-Cpn control (Mm-Cpn), and an
uninduced control (UI) were electrophoresed through 10% SDS-PAGE and stained with
Coomassie blue. Arrows indicate the major overexpressed band in each lane that has an
induced plasmid. B. The same samples in A were separated by 10% SDS-PAGE, transferred,
and probed with each of the eight CCT antibodies. We saw no cross-reaction of each of the
CCT antibodies with any other CCT subunit. Arrows mark the antigenic band. Antibodies to
CCT2, CCT3, CCT5, and CCT8 were polyclonal, while antibodies against CCT1, CCT4, CCT6,
and CCT7 were monoclonal. Filled circles designate bands that are CCT subunits while open
circles designate bands that may not be CCT subunits.
79
To test whether the expressed CCT subunits formed higher order complexes, lysates of
cells expressing the CCT subunits were fractionated on sucrose gradients. Use of the lysates
rather than purified proteins allowed us to verify that these complexes are forming within the E.
coli cells and that that the C-terminal His-tag did not impede subunit assembly. The
conformation of the CCT subunits in the sucrose gradients was compared via immunoblots due
to the low expression of some of the subunits and the abundance of E. coli proteins present in
the lysate (Figure 3-2). While CCT4 and CCT5 formed higher order complexes of similar
sedimentation to human TRiC and Mm-Cpn, the other CCT subunits did not. CCT2 was the only
other CCT subunit possibly forming very low levels of a ~20S complex; however, we did not
observe rings in these samples by electron microscopy (EM). CCT1 and CCT6 subunits were
found throughout the sucrose gradients – these may represent aggregated states or subunits
associating with ribosome subunits. Mass spectrometry and EM of purified CCT1 showed that
CCT1 bound to ribosomes, consistent with its position in the sucrose gradients. The rest of the
subunits: CCT2, CCT3, CCT7 and CCT8, were recovered as slowly sedimenting species.
CCT4 and CCT5 were chosen for further purification due to their assembled state and
high expression level. The purification of the CCT subunits followed a standard 6x-His tagged
protein purification. The cells were lysed with a French Press; this was found to be most
effective in maximizing CCT subunit protein yield, compared to sonication or chemical lysis.
Supernatant/pellet separation of the lysed species did show that a fraction of the CCT protein
ended up in the pellet. We did not investigate the nature of these chains but believe that they
resided in inclusion bodies.
The lysates were passed over a Ni-NTA column and eluted off with a gradient of
imidazole concentrations from 25 to 250 mM. The protein was concentrated, diluted to a lower
imidazole concentration (25 mM), and incubated with TEV protease overnight. The TEV-cleaved
CCT subunit protein was passed again over the Ni-NTA column to which it no longer bound.
The resulting protein fractions were concentrated and passed over a size exclusion column. The
major protein peak eluted between 12 mL and 14.5 mL (CCT5 shown in Figure 3-3). This elution
was consistent with a 1 MDa complex and corresponding to the elution volume of both human
TRiC and Mm-Cpn (Knee et al. 2011; Knee et al. 2013). The symmetry of the distribution and
absence of a trailing edge indicates that the complexes were not dissociating into subunits
under the conditions of the fractionation. Sucrose gradients on purified protein confirmed the
existence of a complex with no dissociated monomers.
80
Figure 3-2: Sucrose ultracentrifugation gradients of CCT subunits
BL21 (DE3) RIL cells expressing each of the eight subunits (CCT1-8) were lysed, fractionated
through sucrose gradients, separated by 10% SDS-PAGE, transferred, and probed with their
respective antibodies. Fractions from the top (5%) through two-thirds (27%) of the sucrose
gradients are shown. For each CCT subunit, on the left, the immunoblot region between 75 kDa
(top line) and 50 kDa (bottom line). Some CCT subunits were sedimenting as soluble subunits
(subunits), others as complexes (complexes), and some were binding to ribosomes
(ribosomes).
81
Figure 3-3: CCT5 purified by size exclusion chromatography as a 1 MDa complex
The input (Input) and various elution volumes (10-16 mL) were electrophoresed through 14%
SDS-PAGE and stained with Coomassie blue. CCT5 appeared as a band ~60 kDa in size
eluted in volumes of 12-14.5 mL consistent with a 1 MDa complex.
82
The final yield was approximately 2 mg per liter of lysate for CCT4 and 5 mg per liter of
lysate for CCT5. Interestingly, CCT5 was concentrated up to 10 mg/mL without issues while
CCT4 tended to precipitate above 1.5 mg/mL. The purified CCT4 and CCT5 subunits were
verified by immunoblots and mass spectrometry. From the mass spectrometry, there was no
detectable GroE in the preparations of purified His-tag-cleaved CCT single subunits complexes.
Structural Characterization of the CCT4 and CCT5 Homo-oligomers
When viewed by negative stain EM, both CCT4 and CCT5 homo-oligomers formed rings
(Figure 3-4A and Figure 3-4B). These rings were approximately 160 Å wide and 180 Å tall,
consistent with that of other group II chaperonins, such as human TRiC and Mm-Cpn (Knee et
al. 2011; Knee et al. 2013). The rings seen for CCT4 and CCT5 homo-oligomers were similar to
that of human TRiC (Figure 3-4C), but distinctly different from those of GroEL/ES from E. coli
(Figure 3-4D). The difference was seen in not only the top views of the rings (eight in
CCT4/CCT5/human TRiC and seven in GroEL – shown as insets), but also in the side views (as
shown with open arrows). The CCT4 homo-oligomer structure looked hollow in the center, but
CCT5 seemed to contain extra density. This extra density was present throughout the three
steps of purification and persisted with or without ATP presence.
Additionally, we observed end-to-end homo-oligomer polymers in EM for CCT4 but not
CCT5 homo-oligomers. Trent et al. reported similar filaments formed by the chaperonin of the
archaea Sulfolobus shibatae and postulated cytoskeletal or regulatory roles for such polymers
(1997). While these were present at low level in negative stain EM, they were common when
CCT4 was viewed in cryo-electron microscopy (cryo-EM) (Figure 3-5A). The presence of these
polymers impeded structural study of CCT4 by cryo-EM.
83
Figure 3-4: Negative stain TEM of purified CCT4 and CCT5 homo-oligomers showed
morphology similar to human TRiC, and distinct from GroEL/ES
The morphology of CCT4 (A) and CCT5 (B) was consistent with that of group II chaperonins.
The complexes were ~160 Å in diameter and ~180 Å in height and shown here at 200 K
magnification. The morphology of human TRiC (C) and GroEL/ES (D) is shown as a control to
the CCT4/CCT5 morphology. These GroEL/ES complexes can be distinguished due to their
subunit per ring differences (seven for GroEL, eight for TRiC; shown as insets) and unique side
view morphology (shown by open arrows). The scale bar represents 100 nm.
84
Figure 3-5: Raw cryo-EM images of CCT5 homo-oligomers and 2D class averages indicated
two rings of eight subunits per ring
A. Raw cryo-EM image of CCT4 homo-oligomers end-on-end polymers. B. Raw cryo-EM image
of the apo/open state CCT5 homo-oligomer with an inset of the 2D classification in the top view,
showing eight subunits per ring. C. Raw cryo-EM image of the ATP-AlFx/closed state CCT5
homo-oligomer with an inset of the 2D classification of the top and side view, showing that the
CCT5 chaperonin complex consisted of two back-to-back rings with eight subunits per ring. The
scale bars represent 50 nm.
85
To further understand the quaternary structure of CCT5 homo-oligomers, we performed
cryo-EM of this complex in both the apo and ATP-AlFx states. In the apo state (Figure 3-5B), a
reference-free 2D class average approach was taken to demonstrate that the top view class
average (inset) displayed eight density blobs without imposing any assumption on the symmetry
in the analysis. The apo state resulted in preferred orientation of end-on views, which has been
encountered in the cryo-EM studies of TRiC or Mm-Cpn in their apo states (Zhang et al. 2010;
Cong et al. 2011). When incubated with ATP-AlFx, similar features were also seen in the raw
images and two orthogonal views of 2D class averages of CCT5 (Figure 3-5C and insets).
However, the density became more continuous from the end-on view of the 2D class average,
which was also observed with TRiC/CCT or Mm-Cpn in ATP-AlFx states compared to their apo
states (Zhang et al. 2010; Cong et al. 2011). This suggests that CCT5 homo-oligomer is
capable of hydrolyzing ATP and closing the complex. To carry out the 3-D reconstruction of
CCT5 homo-oligomers, we used the ATP-AlFx condition because it allowed us to obtain
sufficient number of particle images in different orientations needed for a 3D structure
determination.
In the image reconstruction of CCT5 particle images, we noted significant conformational
heterogeneity, not unusual for reconstructions of group II chaperonins. However ~47% of
particles could be sorted out computationally to have homologous conformation. This data
subset was reprocessed from scratch with a symmetry-free initial model (Figure 3-6A and
Figure 3-6B). A symmetry-free reconstruction of this subset of particle images clearly showed
that the CCT5 complex had similar quaternary structure as TRiC or Mm-Cpn (Figure 3-6C).
A rotational correlation analysis was carried out for the symmetry-free reconstructed
map and eight peaks were observed with approximately 45º spacing when the structure was
rotated along the central axis from 0º to 360º, indicating the presence of eight-fold symmetry in
the complex along the central axis (Figure 3-6D). With C8 symmetry imposed, the reconstructed
map (Figure 3-6E) further improved to 22 Å resolution based on phase randomized resolution
test with two independent data sets (Figure 3-6F). Interestingly, the CCT5 complex had a more
elongated conformation along the symmetry axis compared with TRiC and the two rings were
not exactly identical (i.e. lack of 2-fold symmetry). One possibility is that the heterogeneous
subunits of TRiC have stronger intra-ring interactions that could be conducive to a more
compact closed state.
86
Figure 3-6: Cryo-EM reconstructions of CCT5 homo-oligomers suggested TRiC-like structures
A. Symmetry-free initial template of CCT5 homo-oligomers in the ATP-AlFx state from three
different views (top, bottom and side). B. A rotational correlation analysis of the symmetry-free
reconstructed map along the central axis from 0º to 360º shows only C2 symmetry. C. Three
different views of the symmetry-free 3D reconstruction map of ATP-AlFx state of CCT5 homooligomers. D. A rotational correlation analysis of the symmetry-free reconstructed map along the
central axis from 0º to 360º shows eight peaks with approximately 45º spacing, suggesting
eight-fold symmetry of the reconstruction. E. 3D reconstruction of CCT5 homo-oligomers in the
ATP-AlFx/closed state with C8 symmetry imposed shows a TRiC-like structure in three views. F.
The resolution measured at 0.143 cutoff in the Fourier Shell Correlation between the two initial
models with random phase was ~33 Å (blue) while between the two C8 symmetry imposed final
maps was ~22 Å (red). The maps shown in A, C and E are radially colored.
87
To further investigate this, we performed thermal denaturation studies by circular
dichroism (CD) on human TRiC, CCT4, and CCT5. All three chaperonins had similar far UV CD
scans with human TRiC having a minimum at 225 nm while CCT4 and CCT5 had minima at 228
nm (Figure 3-7A). Occasionally, CCT4 and CCT5 had an unusual minimum at 247 nm,
attributed to ATP or ADP self-association (Heyn and Bretz 1975). This signal was reduced
during purification of CCT5; however, CCT4 samples retained this signal, but the stoichiometry
of the nucleotide was less than 1% of the protein chains (Heyn and Bretz 1975).
Both CCT4 and CCT5 melted at lower temperature than human TRiC (53 °C for CCT4;
60 °C for CCT5; 68 °C for TRiC) (Figure 3-7B). This suggested that subunit-subunit interactions
within TRiC stabilized its secondary structure more than the structures of CCT5 and CCT4. The
transition was much more cooperative for CCT4 and CCT5 than for human TRiC. While no
aggregation was visible upon heating with any of these chaperonins, the denaturation process
was not reversible and rings were not observed by EM after sample denaturation.
When ATP was added in the form of ATP-γS, we saw a very small decrease in melting
temperature (2° for CCT5, and 5° for human TRiC and CCT4), primarily attributed to increased
cooperativity of melting. While addition of ATP should not change the actual secondary
structure, and therefore melting temperature, our results are consistent with the loss of flexibility
in the apical domains of the closed structure, resulting in a more symmetric, uniform structure.
Functional Characterization of the CCT4 and CCT5 Homo-oligomers
The hydrolysis of ATP is an important functional characteristic of the chaperonins. The
ATP hydrolysis properties of CCT4 and CCT5 were assayed using radioactive ATP.
Surprisingly, CCT4 and CCT5 hydrolyzed ATP at a rate comparable to human TRiC (Figure 38).
88
Figure 3-7: Human TRiC is more stable than CCT4 and CCT5 homo-oligomers by thermal
denaturation using CD
A. CD scans of CCT4 (orange), CCT5 (green), and human TRiC (blue) from 260 nm to 200 nm.
CCT4 and CCT5 had minima at 228 nm while human TRiC had a minimum at 225 nm. B. CD
signal at 226 nm was monitored while CCT4 (orange), CCT5 (green), and human TRiC (blue)
were thermally denatured from 25 °C to 100 °C. The denaturation midpoint of CCT4 was 53 °C
and CCT5 was 60 °C while that of human TRiC was 68 °C, suggesting CCT4 and CCT5
complexes were less stable than that of human TRiC. CCT4 had the most cooperative
transition, followed by CCT5, and then human TRiC consistent with the hetero-oligomeric wildtype nature of human TRiC. Adding ATP slightly decreased the denaturation midpoint of the
chaperonins, primarily due to the increase in cooperativity.
89
Figure 3-8: CCT4 and CCT5 homo-oligomers hydrolyze ATP at a similar rate to human TRiC
The generation of [α-32P]ADP was quantified over time for 250 nM CCT4 (orange), CCT5
(green), human TRiC (blue), and BSA (magenta), and a water control (cyan). CCT4 and CCT5
show very similar ATP hydrolysis properties as human TRiC.
90
CCT4 and CCT5 homo-oligomers were assayed for refolding of luciferase (Thulasiraman
et al. 2000), which we previously used to test the substrate refolding activity of human TRiC
(Knee et al. 2013). In the experiment, unfolded luciferase was diluted into buffer with chaperonin
(Thulasiraman et al. 2000). Addition of luciferin and subsequent monitoring of luminescence
production assayed the presence of refolded luciferase in the mixture. At a concentration of 400
nM, human TRiC, and CCT4 and CCT5 homo-oligomers refolded luciferase to about the same
level, leveling off after two hours (Figure 3-9A). When the concentration of chaperonin was
varied, CCT4 and CCT5 homo-oligomers showed higher activity at higher concentrations, as
evidenced by higher luciferase activity (Figure 3-9B). While the range of chaperonin
concentrations (measured as a 16-mer) varied from 0 nM to 300 nM, luciferase concentration
was constant at approximately 10 nM. The luciferase refolding activity of CCT4 and CCT5
homo-oligomers over this range indicated that the folding could be directly attributed to these
two chaperonins and not to any buffer component.
While luciferase is a model substrate for the chaperonins, a more stringent human
substrate is human γD-crystallin (HγD-Crys) (Knee et al. 2011; Knee et al. 2013). HγD-Crys is
found in the lens of the eye and its damage or unfolding can lead to cataract (Moreau and King
2012). CCT subunits have been found in cataracts by proteomic studies and there is evidence
that TRiC interacts with HγD-Crys in the lens periphery, making HγD-Crys an authentic human
TRiC substrate (Hoehenwarter et al. 2008). Its folding and unfolding have been extensively
studied (Kosinski-Collins and King 2003; Flaugh et al. 2005). While some chaperones, such as
the major lens chaperone, α-crystallin, can only suppress HγD-Crys aggregation, group II
chaperonins have been shown to actively suppress and refold HγD-Crys molecules in vitro
(Acosta-Sampson and King 2010; Knee et al. 2011; Moreau and King 2012; Knee et al. 2013).
Knee et al. found that this suppression and refolding ability was strictly ATP-dependent (2011).
In this assay, when unfolded HγD-Crys was diluted from high concentration of guanidinium
hydrochloride (GdnHCl) into buffer at concentrations of 50 µg/mL, partially folded intermediates
partitioned between productive refolding and off-pathway aggregation. This aggregation was
monitored by sample turbidity (OD at 350 nm).
91
Figure 3-9: CCT4 and CCT5 homo-oligomers were active in refolding luciferase
A. CCT4 (orange), CCT5 (green), and human TRiC (blue) at 400 nM were active in refolding
luciferase as compared to the BSA (magenta) control for over two hours at room temperature.
B. When the chaperonin concentration was varied, CCT4 (orange) and CCT5 (green) homooligomers are more active in refolding luciferase with increasing concentration. The luciferase
concentration was constant at 10 nM. For this experiment, n = 3, and the error bars shown are
standard error of the mean.
92
Under the conditions of this assay, containing residual 0.55 M GdnHCl, both CCT4 and
CCT5
homo-oligomers
exhibited
slow
polymerization
by
themselves.
Therefore,
the
concentration of the chaperonin was decreased (16-fold) to 145 nM, as compared to the 2.3 µM
used in previous studies, but the concentration of HγD-Crys was unchanged (Knee et al. 2011).
When CCT4 or CCT5 homo-oligomers were added to the reaction, aggregation of partiallyfolded HγD-Crys was significantly suppressed (Figure 3-10A). While turbidity in the HγD-Crys
aggregation suppression by CCT4 homo-oligomer reached a plateau, HγD-Crys aggregation
suppression by CCT5 homo-oligomer showed continuing increase in turbidity. We attributed this
to CCT5 homo-oligomer polymerization. Previous studies showed that Mm-Cpn and human
TRiC suppress HγD-Crys aggregation by 60-80% (Knee et al. 2011; Knee et al. 2013). At the
significantly reduced concentrations used in this study, both CCT4 and CCT5 homo-oligomers
still suppressed HγD-Crys aggregation by approximately 50%. When CCT5 homo-oligomer was
assayed without ATP, there was less HγD-Crys aggregation suppression and the CCT5 homooligomer polymerization was even more distinct. The initial curve of HγD-Crys aggregation
suppression by CCT5 homo-oligomer without ATP was consistent with that seen for HγD-Crys
aggregation suppression by Mm-Cpn without ATP (Knee et al. 2011). At the conclusion of the
assay, the samples were filtered to remove large aggregates and electrophoresed through 14%
SDS-PAGE. A 20-kDa band consistent with HγD-Crys was seen in the sample with CCT4 and
CCT5 homo-oligomer, but not in the HγD-Crys alone or BSA control samples, indicating that a
fraction of the partially folded HγD-Crys was refolded to native-like state specifically by the
chaperonins (Figure 3-10B; only CCT4 and controls are shown for clarity). The activity of human
TRiC to refold HγD-Crys is reported in Knee et al. and is consistent in levels seen here with
CCT4 and CCT5 (2013).
93
Figure 3-10: CCT4 and CCT5 homo-oligomers suppressed aggregation of partially folded HγDCrys and promoted HγD-Crys native-like state refolding
A. Aggregation of HγD-Crys (blue) was suppressed by the addition of human CCT4 (orange) or
CCT5 (green) by approximately 50% after 15 min at 37 °C. CCT5 tended to self-polymerize
showing a higher turbidity. Without ATP (magenta), HγD-Crys aggregation suppression by
CCT5 was decreased and the CCT5 polymerization was seen more clearly. Curves shown are
representative; the assay was repeated 3-5 times for each chaperonin. B. After filtering, the
samples of HγD-Crys with or without chaperonins were separated by 14% SDS-PAGE and
stained by Krypton. Without chaperonin (---) and with BSA (BSA), no HγD-Crys band was
present, but with Mm-Cpn (Mm-Cpn) and CCT4 (CCT4), HγD-Crys was seen, indicating that it
was refolded to native-like state.
94
Discussion
To our surprise, CCT4 and CCT5 subunits purified out of E. coli formed homo-oligomeric
chaperonin-like complexes. These novel complexes not only possessed morphology consistent
with human TRiC, but were also active in refolding two different substrates. Since these homooligomeric complexes lack many of the wild type subunit/subunit interactions, and are less
stable than the complete endogenous complex, they may not have the structural integrity of the
complete complexes. However, they are clearly active double eight-fold barrels.
The differential expression levels of CCT subunits have been observed in fibroblasts and
mouse tissues (Kubota et al. 1999; Satish et al. 2011), but may be present in many other
tissues and cell types. The expression differences we saw for the CCT subunits in E. coli cells
may reflect differential folding efficiency or stability of the CCT subunits in these cells. The
expression of CCT4 and CCT5 may have been robust in part because the CCT subunit proteins
folded successfully and were assembled into rings inside the cells and were therefore resistant
to degradation. Cheng et al. showed that the folding and assembly of Hsp60 after import into
mitochondria depended on the existence of pre-assembled Hsp60 complexes (Cheng et al.
1990). That result implied that folding of Hsp60 depends on Hsp60 chaperonin function. The
human CCT subunits may also require chaperone or chaperonin assistance in their folding, at
least within the E. coli cytoplasm.
Most of the CCT5 particles contained density within the chaperonin. Although there are
minor contaminating bands seen by SDS-PAGE, no one impurity could account for the density
seen within most of the particles. One hypothesis is that many different newly synthesized E.
coli proteins may be recognized and bound by the CCT5 homo-oligomers. Another explanation
for that density is that CCT5 chains synthesized within the E. coli cells or damaged during the
purification are recognized by the CCT5 homo-oligomeric chaperonin and bound. Further cryoEM and mass spectroscopy studies may distinguish between these hypotheses.
The ATP hydrolysis properties of CCT4 and CCT5 were surprising, because recently
Reissmann et al. revealed that CCT4 and CCT5 have higher affinity for ATP than the other CCT
subunits (2012). The authors postulated that CCT4 and CCT5 – which in the latest consistent
TRiC arrangement are on one side of the ring (Leitner et al. 2012) – drive an asymmetrical
power stroke of ATP hydrolysis that pushes the folding cycle. For CCT4 and CCT5 homooligomers to have similar ATP hydrolysis properties as human TRiC may mean that in these
complexes, each subunit binds ATP as in GroEL/ES or that the identical subunits take turns
hydrolyzing ATP as seen with the ClpX protease rings (Horwich et al. 2007; Glynn et al. 2009).
95
The ATP hydrolysis properties of the homo-oligomers should be further explored to better
understand the mechanism involved in folding cycle in these chaperonins.
In response to the model that specific CCT subunits recognize particular substrates, we
were interested to see whether one CCT subunit homo-oligomer but not the other could
recognize and refold our tested substrates. However, both CCT4 and CCT5 homo-oligomers
recognized and refolded both luciferase and human γD-crystallin. In order to accurately study
the specificity or redundancy of the CCT subunits, we plan to study substrates that are
proposed to only interact with some of the subunits such as actin, tubulin, huntingtin and pVHL
(Llorca et al. 2001; Spiess et al. 2006; Tam et al. 2006).
All reported structures of TRiC purified from tissues describe rings of eight different
subunits. It had been assumed that all eight were obligatory for assembly. Machida et al. have
co-expressed all eight subunits in baby hamster kidney cells and showed that they formed a
TRiC-like complex (2012). While they showed that the CCT subunits were in equal
stoichiometry, the complexes they observed could have contained some CCT4 or CCT5 homooligomers.
The results reported here raise the question of what prevents homo-oligomers from
forming in cells? There is no evidence for regulation at the level of transcription or translation
that would prevent this. Recently, work with the V0 ring of the V-ATPase showed that the
specific arrangement of subunits evolved due to mutations in interfaces between subunits,
rather than evolution of subunit function (Finnigan et al. 2012). In light of that work, it may be
that CCT4 and CCT5 retained the ability to form homo-oligomer contacts but the rest of the CCT
subunits did not. Therefore, TRiC may only be regulated at the level of assembly, as in
bacteriophage, where the interactions of soluble subunits with growing complexes drive the
specificity of association (Kikuchi and King 1975). More recently, Zhang et al. showed that the
chaperonin-like Bardet-Biedl syndrome (BBS) subunits help assemble the final BBSome
complex by sequential addition of each subunit (2012). This needs to be explored for TRiC
through direct in vitro dissociation and re-assembly experiments.
In summary, we have successfully purified CCT homo-oligomer subunit complexes from
E. coli. These subunits have TRiC-like morphology and are active in refolding two substrates.
This novel system will be employed to further study the subunit specificity and redundancy of
the CCT subunits within TRiC and to further provide insight into the assembly of TRiC in the
cell.
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CHAPTER 4:
Biochemical Characterization of Mutants in Chaperonin Proteins CCT4 and CCT5
Associated with Hereditary Sensory Neuropathy*
* This research was submitted to Journal of Biological Chemistry and has been adapted for
presentation here.
Oksana A. Sergeeva, Meme T. Tran, Cameron Haase-Pettingell, and Jonathan A. King (2014).
“Biochemical Characterization of Mutants in Chaperonin Proteins CCT4 and CCT5 Associated
with Hereditary Sensory Neuropathy.” J. Biol. Chem. Submitted.
OAS initiated the research, performed most experiments, and wrote the manuscript; MTT
performed some experiments; CHP performed some experiments and edited the manuscript;
JAK supervised the research and edited the manuscript.
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Abstract
Hereditary sensory neuropathies are a class of disorders marked by degeneration of the
nerve fibers in the sensory periphery neurons. Recently, two mutations were identified in the
subunits of the eukaryotic cytosolic chaperonin, TRiC, a protein machine responsible for folding
actin and tubulin the cell. C450Y CCT4 was identified in a stock of Sprague-Dawley rats, while
H147R CCT5 was found in a human Moroccan family. As with many genetically identified
mutations associated with neuropathies, the underlying molecular basis of the mutants was not
defined. We investigated the biochemical properties of these mutants using an expression
system in E. coli that produces homo-oligomeric rings of CCT4 and CCT5.
Full-length versions of both mutant protein chains were expressed in E. coli at levels
approaching that of the wild-type (WT) chains. Sucrose gradient centrifugation revealed
chaperonin-sized complexes of both WT and mutant chaperonins, but with reduced recovery of
C450Y CCT4 soluble subunits. Electron microscopy of negatively stained samples of C450Y
CCT4 revealed few ring-shaped species, while WT CCT4, H147R CCT5, and WT CCT5
revealed similar ring structures. CCT5 complexes were assayed for their ability to suppress
aggregation of and refold the model substrate γD-crystallin, suppress aggregation of mutant
huntingtin, and refold the physiological substrate β-actin in vitro. H147R CCT5 was not as
efficient in chaperoning these substrates as WT CCT5. The subtle effects of these mutations is
consistent with the homozygous disease phenotype, in which most functions are carried out
during development and adulthood, but some selective function is lost or reduced.
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Introduction
Sensory neurons are nerve cells that convert external stimuli from the environment into
internal stimuli. A rare group of disorders, hereditary sensory neuropathies (HSNs), affect
sensory neurons resulting in a range of clinical symptoms (Auer-Grumbach 2008). These
disorders are marked by the degeneration of the myelinated nerve fibers in the peripheral
sensory neurons and the autonomic neurons that control the involuntary nervous system
(Thomas et al. 1994; Auer-Grumbach 2008). These defects may manifest as ulceration of the
feet, inability to feel pain (especially in the lower limbs), and severe pains in the distal limbs
(Thomas et al. 1994; Auer-Grumbach 2008). Genetic screening of many neuropathy families
has led to the discovery of several mutated genes associated with HSNs and other related
neuropathy diseases. These neuropathies may be inherited through autosomal dominant or
autosomal recessive forms, and are heterogeneous in their pathological and behavioral
symptoms (Cavanagh et al. 1979; Thomas et al. 1994; Rotthier et al. 2009). While age of onset
is variable, severe instances of this disease can have both onset and death within childhood
(Thomas et al. 1994).
Point mutations in three chaperonin genes have been implicated in this class of
neuropathies (Table 4-1) (Hansen et al. 2002; Lee et al. 2003; Bouhouche et al. 2006). While
only two are true HSNs, the other, hereditary spastic paraplegia (HSP), has some important
phenotypic overlaps with HSNs (Timmerman et al. 2013). One of these HSNs is actually
characterized as being a HSN with spastic paraplegia, even further showing the phenotypic
heterogeneity of these disorders (Bouhouche et al. 2006). Two of these have been found in
human populations, making their study potentially valuable for understanding and eventually
treating human neuropathy diseases. How these mutations lead to the disease phenotypes is
still unknown (Auer-Grumbach 2008).
Chaperonins are ATP-dependent chaperones that assist in folding substrate proteins
inside a cavity. They are made of back-to-back rings of 7-9 subunits each (Hartl et al. 2011).
Chaperonins are divided into two classes: type I, found in bacteria, chloroplasts, and
mitochondria; and type II, found in archaeal and eukaryotic cytosols (Hartl et al. 2011). While
there are structural and functional differences between the two classes, they share the same
domain architecture: an equatorial domain making subunit-subunit contacts and forming the
ATP binding and hydrolysis site; an apical domain recognizing substrate to be brought into the
cavity; and an intermediate domain acting as a hinge-like region between the other two domains
(Hartl et al. 2011). The eukaryotic cytosolic chaperonin, TRiC, is involved in the folding and
assembly of dozens of essential eukaryotic proteins (Frydman 2001; Hartl et al. 2011). The
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most important proteins it folds are tubulin and actin, which are especially crucial in neurons
(Lundin et al. 2010). Unlike most of the type I and some of the archaeal type II chaperonins,
which contain identical subunits in both rings, TRiC contains 8 different subunits (termed CCT18) in each of its two rings (Frydman 2001).
Two of the identified HSNs have mutations in two of the CCT genes: CCT4 and CCT5. A
point mutation in the CCT5 gene, A492G, has been associated with human hereditary sensory
neuropathy in a Moroccan family (Bouhouche et al. 2006). These patients are homozygous
recessive for this mutation in exon 4 of the CCT5, which translates to H147R in the protein
(Bouhouche et al. 2006). Hereditary sensory neuropathy has also been identified in a SpragueDawley rat strain, associated with a single point mutation in the CCT4 gene: G1349A (Lee et al.
2003). The affected rats are homozygous recessive for this mutation in CCT4, resulting in the
mutant C450Y in the protein (Lee et al. 2003). Both H147 in CCT5 and C450 in CCT4 are well
conserved in a variety of species (Lee et al. 2003; Bouhouche et al. 2006). Both mutant amino
acid replacements are in the equatorial domain of the CCT subunit, therefore possibly affecting
intra- or inter-ring formation in the chaperonin complex, or ATP hydrolysis activity (Figure 4-1).
However, the actual molecular basis has not been investigated.
The other chaperonin mutation leading to neuropathy was V98I in the mitochondrial
Hsp60 (HSPD1 gene), identified in a French family with HSP (Hansen et al. 2002). While this is
in a type I chaperonin, unlike the type II chaperonin CCT mutations, the two chaperonins have
similar functions, and may therefore share a molecular defect in order to manifest similar
disease phenotypes. This mutant protein was studied biochemically and within bacterial cells. In
vitro studies showed that this substitution affected both ATP hydrolysis and chaperoning
(aggregation suppression and refolding) ability as a homo-oligomer (Bross et al. 2008). In vivo
studies showed that the ATP hydrolysis defect was ameliorated when only a few of the mutated
subunits were in the chaperonin rings. However, the chaperoning defect, while slight, was
enough to cause problems with protein folding (Bross et al. 2008). Having a subtle defect in
these diseases is not too surprising because these patients do live to adulthood, so the
chaperonins have to be functional, albeit slightly suppressed, through their lifetimes.
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Table 4-1: Mutations in chaperonin genes leading to neuropathy diseases
Protein
Mutation
Domain
Inheritance
Identified
Disease
CCT4
C450Y
Equatorial
Recessive
Sprague-Dawley rats
Hereditary sensory neuropathya
CCT5
H147R
Equatorial
Recessive
Moroccan family
Mutilating sensory neuropathyb
HSPD1a
V98I
Equatorial
Dominant
French family
Hereditary spastic paraplegiac
a
HSPD1: human mitochondrial Hsp60
b
(Lee et al. 2003)
c
(Bouhouche et al. 2006)
d
(Hansen et al. 2002)
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Figure 4-1: Location of neuropathy mutations in CCT4 and CCT5
Location of C450Y in CCT4 (A) and H147R in CCT5 (B) are shown in yellow with black arrows
pointing to them. The equatorial domains of the subunits are shown in magenta, the
intermediate domains in green, and apical domain in cyan. PDB: 3P9D.
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Human TRiC expressed in HeLa cells is assembled from eight different protein subunits
(Knee et al. 2013) and has not been amenable to efficient genetic manipulation. However,
CCT4 subunits and CCT5 subunits form homo-oligomeric TRiC-like rings when expressed in E.
coli (Sergeeva et al. 2013). These rings have eight subunits per ring and are active in
hydrolyzing ATP, suppressing aggregation, and refolding a variety of substrates (Sergeeva et al.
2013). Therefore, we have used expression of the single CCT4 and CCT5 subunits as an
experimental system to study the biochemical basis of the CCT4 and CCT5 mutants associated
with hereditary sensory neuropathies.
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Materials and Methods
Mutagenesis and Expression
Wild-type plasmids were previously constructed by modifying the pET21b vector to
contain a TEV protease cleavage site between the end of the inserted gene (CCT4 or CCT5)
and the C-terminal 6x-His-tag (Sergeeva et al. 2013). Site-directed mutagenesis was used to
introduce the neuropathy mutations (G1349A to make C450Y in CCT4; A440G to make H147R
in CCT5) into the plasmids. Mutations were confirmed by sequencing (Genewiz). Plasmids were
transformed into E. coli BL21 (DE3) RIL cells. Proteins were expressed as before (Sergeeva et
al. 2013). Briefly, the cells were grown in Super Broth to OD 5.0 at 37 °C and then shifted to 18
°C and induced with 0.5 mM IPTG. After the overnight induction, cultures were pelleted by
centrifugation for 15 min, and the cells were resuspended in CCT-A (20 mM HEPES/KOH pH
7.4, 300 mM NaCl, 10 mM MgCl2, 10% glycerol, 1 mM DTT, 1 mM ATP) with addition of one
EDTA-free Complete protease inhibitor (Roche) per L of culture.
Long-term Lysate Supernatant/Pellet Separation
E. coli expressing WT and mutant CCT4 and CCT5 were grown and expressed as
above but without the addition of protease inhibitors. The cells were lysed via French Press and
incubated at 4 °C. At specified time points (0, 4, 7, 11, and 14 days), 200 µL aliquots were taken
from the lysates and spun down at 11,500 x g for 30 minutes. The supernatant was extracted
and the pellets were resuspended in CCT-A. SDS-PAGE loading dye (see below) was added to
both the supernatant and pellet, and samples were boiled for 10 minutes, and then frozen at -20
°C until all samples were collected.
Sucrose Gradient Sedimentation
Using CCT-A buffer, 5-40% sucrose gradients were prepared by the gradient master
(BioComp Instruments). Lysates (100 µL) were added carefully to the top and gradients were
ultracentrifuged at 4 °C for 18 h at 37,000 rpm using a SW41 rotor (Beckman). Twenty fractions
were collected using a gradient fractionator (BioComp Instruments), and one bottom fraction
was collected from the remaining gradient.
SDS-PAGE and Immunoblots
Proteins were separated by SDS-PAGE (10%, 12%, or 14%) at 165 V for 1 h after
boiling in reducing buffer (60 mM Tris, pH 6.8, 2% SDS, 5% β-mercaptoethanol, 10% glycerol,
bromophenol blue for color) for 5 min. The gels were stained with Coomassie blue. Transfer
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was conducted for 1.5 h at 300 mA in transfer buffer (10% methanol, 25 mM Tris, 192 mM
glycine) onto 0.45 µm polyvinylidene difluoride (PVDF) membranes (Millipore). The primary
antibodies for the CCT subunits were from Santa Cruz Biotechnology: CCT4, sc-48865; and
CCT5, sc-13886. The secondary antibodies were Alkaline Phosphatase (AP)-conjugated
(Millipore) and the membranes were visualized using the AP-conjugate substrate kit (BioRad).
Band quantification was done using ImageJ.
CCT Subunit Purification
Purification was carried out as previously published (Sergeeva et al. 2013) with a few
slight differences outlined below. Briefly, after lysis via French Press, the lysate was centrifuged,
and the supernatant was removed by pipetting. The supernatant was passed through a 0.45 µm
filter and loaded over a Co-NTA column (Pierce). After loading, the column was first washed
with 100% CCT-A, then 5% CCT-B (CCT-A but with 250 mM imidazole), the CCT single subunit
was eluted off of the column in a linear gradient from 5 to 100% CCT-B. CCT4 protein was
washed with more column volumes of 5% CCT-B than CCT5 protein, due to the presence of a
53-kDa fragment that could be decreased by more thorough washing at that percentage of
imidazole. The fractions containing the CCT single subunit were combined and concentrated,
and then diluted with CCT-A down to 25 mM imidazole. The His-tag was cleaved by TEV
protease and the protein was applied again to the Co-NTA column, to which it no longer bound.
The fractions containing the CCT single subunit were combined, further concentrated, and
passed over a Superose 6 10/300 GL size exclusion column (GE Healthcare). CCT single
subunits were eluted by CCT-SEC (CCT-A but with 5% glycerol and no ATP) off of the size
exclusion column. These fractions were pooled, concentrated, and the protein concentration
was measured using the Bradford assay (BioRad) with BSA as the standard.
Electron Microscopy and Circular Dichroism
Negative stain transmission electron microscopy was carried out as published previously
(Sergeeva et al. 2013). The secondary structure of the chaperonins was assayed by far-UV
circular dichroism at 100 µg/mL of protein in filtered and degassed 10 mM Tris, 20 mM KCl.
Spectra from 260 nm to 195 nm were obtained for each chaperonin and the buffer using an
AVIV Model 202 CD spectrophotometer. Thermal denaturation was carried out by increasing the
temperature in 5 °C increments from 25 °C to 100 °C, with a 5 min incubation before each
spectra was measured. Mean molar ellipticity at 227 nm was used as the metric for protein
folded percentage. Points were fit to a two-state denaturation curve in Prism.
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Native Gel Electrophoresis
CCT5 and its mutant were diluted to 0.5 mg/mL and mixed 2:1 with Bio-Rad Native
Sample Buffer (161-0738). Samples were loaded on Bio-Rad Criterion XT 3-8% Tris-Acetate
gels (345-0131) with 100 mM tricine and 100 mM Tris base running buffer (the cathode buffer
contained 0.02% Coomassie blue G 250). Gels were run at 4 °C either for 3 hours at 150 V or
overnight at 10 mA, and stained with Coomassie blue.
ATP Hydrolysis and Human γD-Crystallin Refolding Assays
The ATP hydrolysis assay was first described in Reissmann et al. (2007) and repeated
with slight modifications in Sergeeva at al. (2013). The human γD-crystallin aggregation
suppression assay is described in detail in Acosta-Sampson & King (2010) and Knee et al.
(2011) and was modified in Sergeeva et al. (2013) to the conditions used in this study. Refolding
percentages were calculated as in Sergeeva et al. (2014) with the same mutant (Y92A/Y97A)
human γD-crystallin protein purification outlined there.
Mutant Huntingtin Aggregation Suppression Assay
Mutant huntingtin (mHtt) aggregation suppression assay was modified from Tam et al.
(2006). Briefly, GST-, His-, and S-tagged exon 1 of Htt with 53 poly glutamines, and containing
a TEV protease cleavage site between the GST-tag and the rest of construct, was purified using
a Co-NTA column, followed by a glutathione agarose column (Pierce). To initiate an aggregation
suppression reaction, 5 µM of the mHtt protein in a buffer (20 mM Tris, 50 mM KCl, 5 mM
MgCl2, 5 mM DTT, and 1 mM ATP) containing various concentrations of chaperonin was
cleaved with 0.1 mM TEV protease. This reaction was left at 30 °C for 16 hours. The reaction
was stopped by equal volume addition of 4% SDS, boiled for 10 minutes, and filtered through
0.22 µm cellulose acetate membrane (GE Healthcare). The membrane was washed and
blocked using 5% milk in TBS. An AP-conjugated antibody against the S-tag (EMD Millipore)
was used to detect amount of mHtt trapped in the membrane. Ovalbumin was used as a control
and concentration of CCT5 was calculated as in the HγD-Crys assay. Quantification was done
in ImageJ where suppression was calculated as decrease from the ovalbumin control.
Actin Refolding Assay
Actin refolding assay was modified from Machida et al. (2012). Briefly, pET28a
containing T7- and His-tagged β-actin was translated using New England Biolabs PURExpress
In Vitro Synthesis kit (E6800S) for 2 h at 37 °C. The translated actin was diluted by half into an
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equal mix of buffer (100 mM HEPES/KOH pH 7.5, 300 mM KCl, 10 mM MgCl2, and 1 mM ATP)
and 4 mg/mL chaperonin or BSA, and actin was allowed to be refolded for 2 h at 37 °C.
Variations of ionic strength (changing KCl concentration to 100 and 500 mM) and concentration
(changing chaperonin concentration to 1 and 2 mg/mL) were also carried out. Trypsin was
added to a final concentration of 20 ng/µL for 15 min at 32 °C to degrade all non-native actin.
SDS-PAGE loading dye (see above) was added to the samples and samples were boiled for 10
minutes. Samples were run on 12% SDS-PAGE, transferred to PVDF, and probed with an antiT7 antibody (Novagen 69522-3). Quantification was done using ImageJ, with ratios taken for
each in vitro actin experiment and normalized to 1000 for WT CCT5.
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Results
Mutant Protein Expression and Stability
Each neuropathy mutation was introduced into the plasmid constructs containing the
CCT4 or CCT5 WT sequences using site-directed mutagenesis. Both full-length mutant proteins
were expressed in E. coli at, at sufficiently high levels to be directly identifiable using Coomassie
stain (Figure 4-2). The mutant expression level was divided by the WT expression levels for
each dilution to quantify how much less of the mutant was expressed. For C450Y CCT4, the
levels monitored by Coomassie stain were reduced to about 80% as compared to WT CCT4 in
both the supernatant and pellet. By Coomassie stain, CCT5 expression levels were comparable
for both the WT and the H147R mutant in the supernatant. In the pellet, the levels of the mutant
were slightly lower than those of the WT, at about 80% of the WT accumulation.
To increase the sensitivity of detection of the truncated chains, the same gels were
probed with a CCT4 and CCT5 antibodies, respectively. With the increased sensitivity of
immunoblotting, a shorter fragment of 53 kDa was clearly detected, for both WT and C450Y
CCT4 chains. This truncated product was previously shown to be missing the first 60 amino
acids of the protein either due to a delayed translation start or a specific protease in the E. coli
lysate (Sergeeva et al. 2013). The shortened mutant chain was present at higher levels in the
pellet than the supernatant. This suggests association into an inclusion body, common for
misfolded or incomplete polypeptide chains.
A more significant difference was seen in the recovery of C450Y CCT4 as compared to
WT CCT4. The mutant chains accumulated to about 30% of the level of the WT in the
supernatant, and 60% of the level of the WT in the pellet. This presumably represents reduced
efficiency in the partial refolding of the chains during the transfer out of SDS to the membrane in
the immunoblot procedure. This is consistent with increased fractionation of the mutant chains
into the pellet fractions. In the immunoblot assays using the CCT5 antibody, H147R CCT5 was
not significantly decreased from WT CCT5 in either the supernatant or pellet. Therefore, the
expression and recovery of the H147R CCT5 mutant in E. coli did not differ from expression and
recovery of WT CCT5.
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Figure 4-2: Expression levels of CCT4, CCT5, and their neuropathy mutants
Supernatant (left) and pellet (right) of E.coli cells expressing CCT4 or CCT5 were diluted by two
from 1/25 to 1/800; solid arrows point to full-length CCT protein, dashed arrows point to CCT4
fragment of 53 kDa. The expression levels were quantified via ImageJ and calculated for the
mutants as mutant level divided by WT level for each dilution. For the Coomassie-stained gels
(top half), the expression levels were almost the same in the WT and mutant. The immunoblots
(against CCT4 or CCT5, respectively) shows a decreased recovery of antigenic C450Y CCT4
as compared to WT CCT4 in the supernatant.
109
To understand the fate of both WT and mutant chains, we incubated the lysate at 4 °C
without protease inhibitors for up to 2 weeks, taking samples for pellet/supernatant separations
every 3-4 days. Over time, both WT and mutant proteins accumulated in the pellet, suggesting
that they became aggregated rather than becoming susceptible to proteases and being
degraded in the lysates (Figure 4-3). This was especially true for C450Y CCT4, which was
mostly in the pellet fraction by about day 7 by both Coomassie-stained gel and immunoblot. WT
CCT4 had a much higher level in the pellet initially than C450Y CCT4 as seen by immunoblot,
especially for the 53-kDa fragment. However, by looking at the Coomassie-stained gel, we see
that WT CCT4 also accumulated in the pellet over time, however slower than C450Y CCT4. For
CCT5, by Coomassie-stained gel and immunoblot, both WT and mutant levels in the pellet
increased from day 0 to day 11, suggesting a fraction of chains aggregated. However, overall,
the amount in the pellet and supernatant of CCT5 had much smaller changes over time than
those for CCT4. In general, CCT5 was more stable than CCT4 in the lysate over the period
assayed, with C450Y CCT4 being the least stable subunit of the four tested. The loss of soluble
chains looks to be due to aggregation rather than proteolysis for all four proteins.
Mutant Protein Sedimentation
The supernatants of the E. coli lysates expressing both WT and mutant chaperonins
were assayed by sucrose gradient ultracentrifugation, to assess whether they were organized
into high molecular weight complexes (Figure 4-4). The sedimentation patterns for both WT
CCT5 and H147R CCT5 were similar, with a distinct species in the 18S complex region and
some presence of soluble subunits at the top of the gradient. For CCT4, WT CCT4 exhibited a
distinct 22S complex species composed of both full length and truncated CCT4 chains. For the
C450Y CCT4 lysate, recovery of unassembled subunits was sharply reduced compared to the
WT control. The majority of C450Y subunits recovered sedimented at the 22S region of the
gradient, but the mutant species seemed to sediment slightly faster and more broadly than the
WT species. The rapidly sedimenting chains to the right of the 22S peak may represent
aggregated chains, corresponding to the increased recovery in the pellets from Figure 4-2. The
mutant fragments behaved similarly as the mutant full-length chains. This overall pattern is
consistent with misfolding and loss of mutant soluble subunits – either through degradation or
inclusion body formation, but with some successful assembly of the remaining subunits. The two
CCT oligomer species, identified here as 18S and 22S, sediment similarly to the 20S
sedimentation seen for the endogenous WT TRiC isolated from HeLa cells (Knee et al. 2013).
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Figure 4-3: Long-term lysate incubation of CCT4, CCT5, and their neuropathy mutants
Lysates of CCT4 (A) and CCT5 (B) and their neuropathy mutant were incubated for 0, 4, 7, or
11 days and then underwent pellet/supernatant separations. Both coomassie and immunoblot
SDS-PAGE is shown with full-length CCT4 or CCT5 in between dotted lines, respectively. Two
E.coli fragments that accumulate in the pellet are indicated with asterisks, while two CCT4
fragments that accumulate in the pellet are indicated with +-signs. The full-length proteins are
quantified to the right of each gel with WT in blue and Mutant in magenta, and pellet in solidlined circles and supernatant in dashed-lined squares. Both CCT4 and CCT5 and their
neuropathy mutants (especially C450Y CCT4) accumulate in the pellet over time, suggesting
aggregation of the full-length species.
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Figure 4-4: Sucrose ultracentrifugation gradients of CCT4, CCT5, and their neuropathy mutants
Centrifuged lysates were immunoblotted for CCT4 (top) and CCT5 (bottom), respectively; solid
arrows point to full-length CCT protein, dashed arrows point to CCT4 fragment of 53 kDa.
C450Y CCT4 showed a distinctly different sedimentation pattern (no soluble monomer species
and a more broad 22S species, possibly slightly faster sedimenting) as compared to WT CCT4.
H147R CCT5 and WT CCT5 had very similar sedimentation patterns. The WT sedimentation
patterns shown here are consistent with those published in Sergeeva et al. (2013), but have
been more specifically labeled as 22S and 18S for CCT4 and CCT5, respectively.
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Mutant Protein Purification
The CCT chaperonins and their neuropathy mutants were purified from the lysates by
cobalt affinity chromatography (Figure 4-5). The elution profiles of WT and H147R CCT5 were
similar, both proteins eluting off of the cobalt affinity column in approximately the same
amounts. For CCT4, the pattern of elution of full-length WT chains differed from that of the
fragment, suggesting that they were not in a complex with each other under these conditions.
Note that the fragments also bind to the cobalt column indicating that they carry the C-terminal
His-tags. In order to decrease the amount of CCT4 fragment eluting with full-length CCT4 off of
the cobalt column, a longer 5% CCT-B wash was used. Therefore, the WT CCT4 protein
partitioned between weakly bound chains eluting at low imidazole and tightly bound chains
eluting at higher concentrations. C450Y CCT4 – both full-length and fragment - was recovered
from the column at significantly lower levels than WT CCT4. This suggested that the
conformation and stability of the mutant CCT4 subunits was altered, so that it was either
aggregating, or that it no longer efficiently bound to the cobalt column. This may be because the
His-tag was buried or otherwise inaccessible for binding.
Both WT and mutant proteins were further purified by TEV protease cleavage to remove
the His-tag, followed by size exclusion chromatography. Due to the low concentration off of the
cobalt column, the CCT4 C450Y mutant protein was much less pure and at a significantly lower
yield than the WT CCT4. However, it did elute off the size-exclusion column at the same place
as WT CCT4, suggesting that some proportion of ring-like complexes were assembled, but they
were not stable or sufficient enough for a large sample to be purified. This limited our ability to
assay its properties compared to WT CCT4. The neuropathy mutant of CCT5, on the other
hand, was successfully purified with the His-tag removed to levels similar to those of WT CCT5.
113
Figure 4-5: CCT4 and CCT5 purification off of the Co-NTA column
Fractions of CCT4 (left; WT, top and C450Y, bottom) and CCT5 (right; WT, top and H147R,
bottom) from the 5% wash (5%) and elution (arrow to 50%) off of the Co-NTA column were run
on 10% Coomassie-stained SDS-PAGE; solid arrows point to full-length CCT protein, dashed
arrows point to CCT4 fragment of 53 kDa. WT CCT4 had significantly more protein eluting off of
the column than C450Y CCT4, even with the difference in expression levels taken into account.
There was no significant difference between the elution of WT CCT5 and H147R CCT5.
114
Mutant Protein Structure
Final purified samples, off of the size exclusion column, were examined by negative
stain transmission electron microscopy (TEM). WT CCT4, WT CCT5, and H147R CCT5 all had
similar morphology (Fig. 5) appearing as well formed rings oriented along the beam axis. C450Y
CCT4 had few to no rings and for the most part appeared as aggregated species by negative
stain TEM. The lack of ring species at the end of the mutant CCT4 purification suggests that the
mutant CCT4 protein may be unstable, even in the multimeric state. Fractions of mutant CCT4
off of the cobalt column did show a few rings by negative stain EM, and the size exclusion
elution volume and lysate sucrose ultracentrifugation gradients did suggest a chaperonin-sized
species. While C450Y CCT4 may be capable of forming rings, they did not persist throughout
the purification, possibly succumbing to aggregation or dissociation. The experiments in Figures
4-2 though 4-6 taken together indicate that the defect in C450Y is one of subunit folding and
stability.
For WT and H147R CCT5, purified samples could be obtained and were run on native
gel electrophoresis. H147R CCT5 repeatedly ran slightly slower than WT CCT5, suggesting that
its charge difference was on the surface of the protein, therefore altering its running properties
on a native gel (Figure 4-7). Additionally, both WT and mutant CCT5 were well-formed
complexes of approximately 1 MDa with no smear of degraded subunits or monomer subunits.
This assay also verified that the protein purified was indeed mutant CCT5.
To assess the conformation of the mutant CCT5 subunits, far-UV circular dichroism (CD)
scans of WT and H147R CCT5 were obtained, along with thermal melts of both proteins as
tracked by CD. They exhibited very similar spectra with minima at 227 nm and a very similar
thermal denaturation midpoint of approximately 60 °C (Figure 4-8A). The denaturation of the
mutant was less cooperative than the denaturation of the WT, possibly pointing to some
difference in subunit contacts within or between the rings (Figure 4-8B). However, in general,
the H147R mutation in CCT5 did not disrupt subunit structure or complex assembly.
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Figure 4-6: Negative stain transmission electron micrographs of CCT4, CCT5, and their
neuropathy mutants
WT CCT4 (top, left), WT CCT5 (bottom, left) and H147R CCT5 (bottom, right) formed TRiC-like
rings of approximately the same size that were visualized here after a full purification and elution
off of the size exclusion column. At the end of the purification, C450Y CCT4 (top, right)
contained more aggregates and did not display rings by TEM. Scale bars, 100 nm. WT CCT4
and WT CCT5 rings are consistent with those published in Sergeeva et al. (2013).
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Figure 4-7: Native gel electrophoresis of CCT5 and its neuropathy mutant
Mm-Cpn (control), WT CCT5, and H147R CCT5 were run on native gel electrophoresis. Vertical
lines for visual comparison designate the chaperonin complexes in each lane. The H147R
CCT5 mutant runs slightly slower than WT CCT5, suggesting that the mutation alters the outer
charge of the mutant chaperonin.
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Figure 4-8: Far-UV circular dichroism scans and thermal denaturation of CCT5 and its
neuropathy mutant
A. CD scans of WT CCT5 (blue) and H147R CCT5 (magenta) showed similar spectra from 260
nm to 195 nm; the minima are approximately 227 nm. B. Thermal denaturation of WT CCT5
(blue) and H147R CCT5 (magenta) by CD had approximately the same midpoint of 60 °C,
although the profiles were slightly different in terms of cooperativity. The mean molar ellipticity at
227 was used as the proxy for protein folding percentage. The WT CCT5 scan and thermal melt
are consistent with those published in Sergeeva et al. (2013).
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CCT5 Mutant Activity
In order to investigate how the mutation may lead to neuropathy, chaperonin activity
assays were performed. Due to the position of the mutation in the equatorial domain, one likely
defect might be in ATP hydrolysis of the mutant chaperonin. The purified CCT5 and H147R
CCT5 complexes were therefore assayed for their ability to hydrolyze ATP. As shown in Figure
4-9, the hydrolysis rates were very similar between WT and mutant CCT5.
The critical functions of group II chaperonins are believed to be suppressing the
intracellular aggregation of partially folded intermediates, and assisting the folding to the native
state. We therefore assayed CCT5 and H147R CCT5 for suppression of off-pathway
aggregation, and refolding in vitro to the native state. The substrate used in these experiments
was human γD crystallin (HγD-Crys), whose off-pathway aggregation and productive refolding
has been systematically studied (Kosinski-Collins and King 2003; Kosinski-Collins et al. 2004;
Flaugh et al. 2005; Flaugh et al. 2005; Chen et al. 2006; Flaugh et al. 2006; Moreau and King
2009; Acosta-Sampson and King 2010; Kong and King 2011). Endogenous human TRiC
purified from HeLa cells and WT CCT4 and CCT5 homo-oligomers are active in both assays
(Knee et al. 2013; Sergeeva et al. 2013).
As can be seen in Figure 4-10, the HγD chains aggregated to high molecular weight
complexes after dilution out of denaturant (Kosinski-Collins and King 2003). When WT CCT5
was added, the aggregation of WT HγD-Crys was suppressed. This is consistent with what was
seen previously for CCT5 suppression of WT HγD-Crys aggregation (Sergeeva et al. 2013).
H147R CCT5 was able to suppress mutant aggregation at first, but showed an increase in
turbidity that was similar to WT HγD-Crys alone at the end of the reaction (Figure 4-10A).
Therefore, the mutant protein appears to have an altered reaction with the substrate in this
reaction compared to WT CCT5. A potentially more stringent substrate was also assayed with
WT and H147R CCT5. In this case, the aggregating protein was HγD-Crys carrying a double
alanine substitution of tyrosines, Y92A/Y97A (Kong and King 2011; Sergeeva et al. 2014).
Suppression of aggregation by WT CCT5 was similar to that found with WT HγD-Crys (Figure 410B). The H147R CCT5 protein had an altered interaction compared to WT CCT5, mimicking
the results seen for WT HγD-Crys. For both HγD-Crys substrates, H147R CCT5 less efficiently
suppressed aggregation than WT CCT5.
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Figure 4-9: ATP hydrolysis of CCT5 and its neuropathy mutant
WT CCT5 (blue) and H147R CCT5 (magenta) showed similar rates of ATP hydrolysis as
measured by quantified generation of [α-32P]ADP over time. The values shown for WT CCT5
were previously published in Sergeeva et al. (2013).
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Figure 4-10: Aggregation suppression of HγD-Crys by CCT5 and its neuropathy mutant
Aggregation of WT (orange, A) or Y92A/Y97A (orange, B) HγD-Crys was suppressed more
efficiently by WT CCT5 (blue) than H147R CCT5 (magenta). Without HγD-Crys, WT CCT5
(purple) and H147R CCT5 (green) did not show any self-polymerization. The curves are
representative; the assays were repeated three to five times and showed the same trends.
121
Previously, we showed that CCT5 had an increase in turbidity throughout the assay,
which we attributed to self-polymerization. However, when WT and H147R CCT5 were added to
the assay without HγD-Crys (Figure 4-10), we did not see an increase in turbidity, suggesting
that it was not self-polymerization but rather polymerization or aggregation of the complex
between CCT5 and HγD-Crys that was causing the increase in turbidity throughout the assay.
We cannot exclude that the decrease in aggregation suppression of HγD-Crys by H147R CCT5
may be due to increased aggregation of the H147R CCT5/HγD-Crys complex.
Along with aggregation suppression, we can also assay the amount of HγD-Crys
refolded by the chaperonin. Residual background refolding is present but is significantly less
than the amount of HγD-Crys actively refolded by the chaperonins (Figure 4-11A). When we
quantified the amount of WT and Y92A/Y97A HγD-Crys refolded by WT and H147R CCT5, we
observed a significant decrease in the amount refolded by H147R CCT5 as compared to WT
CCT5 (Figure 4-11B). This decrease was approximately 30% for WT HγD-Crys and 20% for
Y92A/Y97A HγD-Crys, but this amount of refolded HγD-Crys by H147R CCT5 was not
significantly different than background refolding in both cases. In general, Y92A/Y97A HγD-Crys
was refolded to lower levels than WT HγD-Crys, contrary to what was seen for the archaeal
Mm-Cpn chaperonin previously (Sergeeva et al. 2014).
While HγD-Crys is an authentic substrate of TRiC in the periphery of the eye lens, its
value is limited when surveying how H147R CCT5 may lead to neuropathy. Therefore, we also
challenged mutant CCT5 to two other human substrates associated with the brain. The first is
huntingtin (Htt), a very large, 3144 amino acid (348 kDa), soluble cytoplasmic protein. Although
it is ubiquitously expressed, it is found at high levels in the central nervous system and the
testes (Wetzel 2012). WT Htt has various functions in cells such as acting as a scaffold protein,
and playing a role in neuronal gene transcription, and axonal and vesicular transport (Bates
2005). Htt in its pathological form contains an expanded repeat of CAG resulting in 36+
polyglutamines (Walker 2007). Aggregates of mutant Htt (mHtt) have been found in patient
brains, consistent with the idea that aggregation of the pathological protein is part of the disease
(Arrasate and Finkbeiner 2012; Clabough 2013). These aggregates contain fragments of the
mHtt protein, the shortest of which includes only the first exon of Htt wherein the polyglutamine
region is located (Wetzel 2012).
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Figure 4-11: SDS-PAGE and quantification of HγD-Crys refolded by CCT5 and its neuropathy
mutant
A. 14% Coomassie-stained SDS-PAGE of either WT (left) or Y92A/Y97A (right) refolded HγDCrys alone (---), with WT (WT), or with H147R (H147R) CCT5 is shown. Some residual
background refolding can be seen, but there is significantly more refolding by the chaperonins.
B. WT CCT5 (blue) refolded significantly more WT (left) or Y92A/Y97A (right) HγD-Crys than
H147R CCT5 (magenta). Both chaperonins refolded more WT than Y92A/Y97A HγD-Crys. Error
bars are SEM from 3 independent quantifications; single asterisks denote significance at p <
0.05 by t-test, double asterisks denote significance at p < 0.01 by t-test.
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Previous studies have shown that TRiC interacts with mHtt and decreases its
aggregation (Tam et al. 2006; Tam et al. 2009; Shahmoradian et al. 2013; Sontag et al. 2013).
We assayed both WT and H147R CCT5 for their ability to suppress mHtt. For this assay, we
used an mHtt protein that was GST-tagged and contained at TEV protease site. When we
added TEV protease to the reaction containing mHtt and either WT or H147R CCT5, the mHtt
would aggregate. We were able to see how much aggregation was suppressed by the
chaperonins by using a filter trap assay and probing with an antibody against the mHtt
construct. While both were able to suppress mHtt, WT CCT5 was more efficient in at least one
concentration than H147R CCT5 (Figure 4-12).
The second more neuropathy-related substrate we assayed is highly expressed in
neurons and is one of TRiC’s major substrates: β-actin (Lundin et al. 2010). For this assay, we
synthesized T7-tagged β-actin in vitro and allowed WT or H147R CCT5 to fold it to native state
(with BSA as a control). The samples were cleaved with trypsin so only the native β-actin
persisted, run on SDS-PAGE, transferred to immunoblot, and probed with an anti-T7 antibody.
We found that H147R CCT5 folded significantly less β-actin than WT CCT5 (Figure 4-13).
However, in this assay, unlike the HγD-Crys refolding assay, the background folding of β-actin,
as seen by the BSA negative control, was minimal, so the amount folded by H147R CCT5 was
still significant. To further investigate the actin refolding properties, we varied both concentration
of chaperonin and ionic strength of the buffer (Figure 4-14). For each of these conditions, the
mutant CCT5 did not refold as much as WT CCT5. Interestingly, while we saw a concentration
dependence when we varied concentration, we were able to confirm that the concentration of
KCl in the buffer we used above was the optimal concentration for actin refolding.
Overall, WT CCT5 was more efficient at suppressing HγD-Crys aggregation, refolding
HγD-Crys (by about 30%), suppressing mHtt aggregation (by about 40%), and folding β-actin
than H147R CCT5 (by about 20%). This suggests that the defect in H147R CCT5 is that of
chaperonin function.
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Figure 4-12: Mutant huntingtin aggregation suppression by CCT5 and its neuropathy mutant
A. Representative filter trap samples probed with an antibody to mHtt; ratios are mHtt: CCT5. B.
Quantifications of multiple experiments as in A. WT CCT5 suppressed mHtt more efficiently
than H147R CCT5 at all ratios, but only significantly at the 1:1 ratio. Data normalized to
ovalbumin control (1.0); Error bars are SEM from 2 independent quantification; double asterisks
denote significance at p < 0.01 by t-test.
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Figure 4-13: Quantification of β-actin refolded by CCT5 and its neuropathy mutant
A. Representative 12% SDS-PAGE immunoblot probed with anti-T7 antibody of refolded actin in
the presence of BSA, WT CCT5, or H147R CCT5 is shown. The arrow points to β-actin. B.
Quantification of multiple experiments as in A. H147R CCT5 refolded significantly less actin
than WT CCT5. WT CCT5 refolded intensity was normalized to 1000; Error bars are SEM from
4 independent quantifications; single asterisks denote significance at p < 0.05 by t-test, triple
asterisks denote significance at p < 0.001 by t-test.
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Figure 4-14: Variations in protein concentration and ionic strength of β-actin refolded by CCT5
and its neuropathy mutant
Same assay as in Fig. 12 but with variations in protein concentration (A) and ionic strength of
the buffer (B). H147R CCT5 refolded significantly less actin than WT CCT5 in all variations.
There was a protein concentration dependence (A) and the optimal ionic strength was 300 mM
KCl (B). Conditions used in Fig. 12 were normalized to 1000; Error bars are SEM from 3
independent quantifications; double asterisks denote significance at p < 0.01 by t-test, triple
asterisks denote significance at p < 0.001 by t-test.
127
Discussion
Human CCT4 and CCT5 subunits of the TRiC group II chaperonins assemble into
double barrel TRiC-like rings in the absence of the other seven CCT subunits (Sergeeva et al.
2013). We have used this homo-oligomerization of CCT4 and CCT5 subunits to investigate two
neuropathy mutations identified in these chaperonin subunits. Based on the in vitro work on
V98I Hsp60, and the fact that these patients survive to adulthood, we were expecting a only
subtle differences in the function of these mutated subunits (Bross et al. 2008).
The H147R CCT5 mutant subunits assembled into oligomeric rings with similar
efficiency as WT CCT5 subunits. The melting temperature for the mutant rings was similar to
that for the WT CCT5 indicating that the H147R substitution did not cause a major defect in
chaperonin structure. These chaperonin-like complexes hydrolyzed ATP with similar efficiency
as WT CCT5 complexes. However, when assayed for the ability to suppress in vitro aggregation
of HγD-Crys, their efficiency was reduced. The ability of the mutant complexes to chaperone the
refolding of HγD-Crys back to the native-like state was also significantly reduced. Additionally,
H147R CCT5 folded significantly less β-actin than WT CCT5. Note however, that in most of
these assays the mutant complexes exhibited substantial levels of activity, with respect to WT
CCT5 and negative controls. Our experiments do not distinguish a reduction in the initial
efficiency of recognizing and binding partially folded substrates, from an actual alteration of the
chaperoning reaction that proceeds within the lumen of the complex.
The HγD-Crys aggregation suppression and refolding assay used in this study has been
used for many other chaperonins (Knee et al. 2011; Knee et al. 2013; Sergeeva et al. 2013;
Sergeeva et al. 2014). Interestingly, the crystallin mutant used herein, Y92A/Y97A, was refolded
to higher levels by the archaeal chaperonin Mm-Cpn (Sergeeva et al. 2014). Here both WT
CCT5 and H147R CCT5 refolded the mutant substrate chains to levels of about a third of those
of WT HγD-Crys chains. H147R CCT5 refolded less of both WT and Y92A/Y97A HγD-Crys than
WT CCT5, showing that the refolded amount is even worse when mutant substrates are
chaperoned by this mutant chaperonin. The β-actin assay was used before in various iterations
(Llorca et al. 1999; Pappenberger et al. 2006; Machida et al. 2012), but is novel for these homooligomeric complexes. Seeing a significant difference between our WT and mutant CCT5 with
this stringent substrate bolstered the theory that this mutant was responsible for neuropathy due
to its decreased chaperoning ability.
Another discrepancy worth noting is that of mHtt and CCT5. It was previously shown that
only CCT1 and CCT4 could suppress aggregation of mHtt (Tam et al. 2006). Here we show that
CCT5 is capable of also suppressing mHtt aggregation. In that study, CCT subunits were co-
128
overexpressed in yeast with mHtt constructs. The conformation of the CCT subunits in these
over-expressed cells is unknown so they could have been misfolded or aggregated, therefore
not showing any efficacy. These latest results of CCT5 being able to suppress mHtt aggregation
may allow the study of CCT subunits other than CCT1 in modulating mHtt aggregation.
The H147R substitution introduces a charge change into the CCT5 subunits. The
guanido group of arginine is generally found at the surface of soluble proteins. Direct evidence
of this change was seen by native gel electrophoresis. Such increased charge density might
reduce chaperonin activity both in suppressing aggregation and refolding, due to electrostatic
effects.
The defect in C450Y CCT4 was in the folding and stability of the mutant subunit itself,
which may affect the complex formation ability. Compared to WT CCT4 chains, a larger fraction
accumulated aggregated in the pellet fraction of cells, accumulation of soluble subunits was
reduced, and formation of organized rings was sharply reduced.
Due to the homo-oligomer nature of our system, it is hard to assess whether a normal
TRiC ring with seven other CCT subunits would be equally disrupted. That may depend on how
TRiC is assembled in the cell. If CCT4 is one of the last subunits added, even an unstable
CCT4 subunit may be incorporated and function sufficiently as part of the full ring. However, if
CCT4 needs to form homo-oligomeric rings on the way to the mature TRiC complex, the
mutation may results in a more defective phenotype. In either case, if C450Y CCT4 mutant folds
less efficiently in the cytoplasm, or is subject to increased aggregation, it could reduce levels of
functional TRiC, thus affecting folding of any of the numerous TRiC substrates. These are likely
to have differential importance in different cell types.
Another feature of C450Y CCT4 mutant subunit is that the amino acid change itself is
from a cysteine to a tyrosine, which may be easily post-translationally modified by a kinase.
Either the loss of the cysteine or the gain of the tyrosine could affect post-translational
modifications for downstream signaling (Abe et al. 2009). If C450Y CCT4 does incorporate itself
as part of TRiC, it may be modified with respect to WT subunits. Unfortunately, very little is
known of the control of chaperonin activity by post-translational modifications of TRiC.
It was encouraging to see a similar chaperoning defect in H147R CCT5 as seen for V98I
Hsp60 (Bross et al. 2008). While in our system, the defect was exaggerated due to the homooligomer nature of the chaperonins, any decrease in protein folding function of TRiC will
negatively affect many essential substrate proteins, including tubulin and actin. Since neurons
contain a high abundance of microtubules, tubulin is a good candidate for a substrate that may
be most affected (Lundin et al. 2010). There have been reports of sensory neuropathy induced
129
by taxanes (paclitaxel and docetaxel; anti-cancer drugs used in chemotherapy), where it is
postulated that the taxanes promote microtubule aggregation, specifically in neurons (Hagiwara
and Sunada 2004). Therefore, the H147R CCT5 hereditary sensory neuropathy may very well
be working through the same mechanism.
By studying purified human C450Y CCT4 and H147R CCT5 expressed in E. coli, we
have found very subtle biochemical defects in these neuropathy-associated mutants as
compared to WT. Whether these defects are exactly the issues contributing to neuropathy within
the Moroccan family or the Sprague-Dawley rat strain remains to be seen. To further investigate
chaperonin activity of the CCT5 neuropathy mutant, it will be crucial to use more physiological
neuronal substrates in these aggregation and refolding assays. β-Actin is a good first candidate,
but others will need to be tested. However, sorting out which substrates are predominantly
affected by the CCT mutant substitutions will require characterizing the substrates associated
with TRiC within human neuronal cells expressing the neuropathy mutations. The use of patient
or rat cell lines of these neuropathies would be ideal in being able to investigate these mutants
in the disease context.
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CHAPTER 5:
Group II Archaeal Chaperonin Recognition of
Partially Folded Human γD-Crystallin Mutants*
* This research was originally published in Protein Science and has been adapted for
presentation here.
Oksana A. Sergeeva, Jingkun Yang, Jonathan A. King and Kelly M. Knee (2014). “Group II
archaeal chaperonin recognition of partially folded human γD-crystallin mutants.” Protein
Science 23: 693-702. doi: 10.1002/pro.2452 © The Protein Society.
OAS performed most experiments and wrote the manuscript; JY performed some experiments;
JAK supervised the research and edited the manuscript; KMK initiated the research and
performed some experiments.
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Abstract
The features in partially folded intermediates that allow the group II chaperonins to
distinguish partially folded from native states remain unclear. The archaeal group II chaperonin
from Methanococcus maripaludis (Mm-Cpn) assists the in vitro refolding of the wellcharacterized β-sheet lens protein human γD-crystallin (HγD-Crys). The buried cores of this
Greek key conformation and the domain interface include a variety of side chains, which might
be exposed in partially folded intermediates. We sought to assess whether particular features
buried in the native state, but absent from the native protein surface, might be serving as
recognition signals. The features tested were a) paired aromatic side chains, b) side chains in
the interface between the duplicated domains of HγD-Crys, and c) side chains in the buried core
which result in congenital cataract when substituted. We tested the Mm-Cpn suppression of
aggregation of these HγD-Crys mutants refolding upon dilution out of denaturant.
Mm-Cpn was capable of suppressing the off-pathway aggregation of the three classes of
mutants indicating that the buried residues were not recognition signals. In fact, Mm-Cpn
recognized the HγD-Crys mutants better than wild-type (WT) and refolded most mutant HγDCrys to levels twice that of WT HγD-Crys. This presumably represents the increased population
or longer lifetimes of the partially folded intermediates of the destabilized mutant proteins. The
results suggest that Mm-Cpn does not recognize the features of HγD-Crys tested – paired
aromatic residues, exposed domain interface, or destabilized core – but rather recognizes other
features of the partially folded β-sheet conformation that are absent or inaccessible in the native
state of HγD-Crys.
Note: This research was done chronologically before the other chapters of the thesis and that is
why Mm-Cpn rather than CCT/TRiC was used as the chaperonin.
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Introduction
Group II chaperonins are found in the cytosol of eukaryotes and archaea, and fold their
substrates by encapsulating them inside a cavity away from other proteins and macromolecules.
They are composed of two back-to-back rings of one to eight different subunits (Hartl et al.
2011). The mechanism and high-resolution structures of group II chaperonins have been
elucidated using the archaeal chaperonin from Methanococcus maripaludis (Mm-Cpn) (Pereira
et al. 2010; Zhang et al. 2010; Douglas et al. 2011; Zhang et al. 2011; Pereira et al. 2012).
However, it is still largely unknown how exactly group II chaperonins recognize their substrates.
While only a few studies of substrate recognition by group II chaperones have been carried out,
but there are a few specific substrate examples. The eukaryotic group II chaperonin, TRiC,
prefers substrates with extended β-sheets, whose folds contain hydrophobic patches and are
slow to fold (Yam et al. 2008). TRiC recognizes two hydrophobic β-sheets termed Box 1 and
Box 2 on pVHL (Feldman et al. 2003), and specific parts of two different WD40 proteins: the
hydrophobic third β-strand of the second WD40 repeat of G protein β and WD40 repeats 3-5 in
Cdc20 (Camasses et al. 2003; Kubota et al. 2006). Among these important examples, it is still
unclear which feature of the hydrophobic β-sheets the group II chaperonin recognizes.
To better understand the recognition of β-sheet proteins by group II chaperonins, we
have used the β-sheet protein human γD-crystallin (HγD-Crys) as a substrate (Basak et al.
2003). This two-domain/four-Greek-key protein, found in the eye lens where there is no protein
turnover, must maintain its native structure for the human lifetime. Deleterious modification and
damage can accumulate over time, inducing partial unfolding of HγD-Crys, which can lead to
cataract (Wang and King 2010). HγD-Crys is a genuine substrate for chaperones; it interacts
with the small heat shock chaperone α-crystallin in the lens nucleus and may interact with TRiC
in the epithelial cells of the lens periphery (Hoehenwarter et al. 2008; Acosta-Sampson and King
2010). The folded state of the protein can be assayed using four buried tryptophans located in
each quadrant of the protein, which are quenched in the native state (Kosinski-Collins and King
2003; Kosinski-Collins et al. 2004). Using this fluorescence property, the folding pathway of
HγD-Crys has been elucidated and involves an intermediate that has a folded C-terminus and
unfolded N-terminus (Kosinski-Collins et al. 2004).
The in vitro folding, unfolding, and off-pathway aggregation of HγD-Crys has been well
characterized, including the effects of diverse amino acid substitutions (Kosinski-Collins et al.
2004; Flaugh et al. 2005; Flaugh et al. 2005; Chen et al. 2006; Flaugh et al. 2006; Mills et al.
2007; Moreau and King 2009; Kong and King 2011). Many of these substitutions induce a
partial unfolding of the native structure, which is thought to be key in substrate recognition by
133
the chaperones (Hartl et al. 2011). Such mutations in the crystallins have been previously used
to study substrate-chaperone interactions. Another crystallin protein, βB2-crystallin (βB2-Crys)
has been characterized as a substrate for the α-crystallin lens chaperone. The substrate
proteins carried domain interface mutations of glutamine to glutamate, to simulate deamidation
in the lens (Takata et al. 2009; Takata et al. 2010). α-Crystallin could only partially rescue the
aggregation of these deamidated mutants, primarily because their aggregation involved
intermediates that were not as readily recognized by α-crystallin (Michiel et al. 2010).
Furthermore, the aggregation of the double deamidated mutant was found to be less efficiently
suppressed by α-crystallin than the aggregation of wild-type (WT) βB2-Crys, due to the fact that
this mutant aggregated faster than WT, not allowing α-crystallin enough time to recognize and
bind it (Lampi et al. 2012). We sought to understand similar interactions between the group II
chaperonins and a β-sheet substrate, HγD-Crys.
Earlier research showed that the group II chaperonin Mm-Cpn can recognize and refold
partially folded, but not native, β-sheet rich WT HγD-Crys (Knee et al. 2011). While HγD-Crys is
not a genuine substrate for the archaeal chaperonin, its extensive characterization makes it an
ideal model substrate for study. Candidates for features recognized by the chaperonin include
amino acid side chain conformations buried in the native state, but exposed in partially folded
intermediates. These include the unpaired domain interface (based on the βB2-Crys example),
exposed or unpaired aromatics, or exposed hydrophobic core residues (Figure 5-1). HγD-Crys
has fourteen tyrosines, six phenylalanines, and four tryptophans that make a substantial
contribution to the buried β-sheet cores. Many of these occur in pairs or clusters. Though these
residues contribute to overall stability, polypeptide chains with alanine substitutions are
generally able to fold to the native state (Knee et al. 2011). As part of the effort to identify what
features Mm-Cpn recognizes in its substrates, we also examined Mm-Cpn interactions with
mutant HγD-Crys altered in the buried core (Moreau and King 2009).
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Figure 5-1: HγD-Crys mutants chosen fall into three sets
Crystal structure (PDB: 1HK0) of HγD-Crys with mutant residues highlighted in red (aromatic
pair Y92/Y97), orange (aromatic pair Y133/Y138), blue (interface pair Q54/Q143), and green
(core residues L5, V75, and I90) in front view (A) and down view (B).
135
Materials & Methods
Purification of HγD-Crys and Mm-Cpn
WT and mutant HγD-Crys were expressed and purified as published (Kosinski-Collins et
al. 2004). Briefly, pQE.1 plasmid containing the HγD-Crys gene was transformed into M15 E.
coli cells. Cells were grown up to OD600 1.0 and induced with 1 mM IPTG for 3 hours at 37 °C.
Cells were spun down and resuspended in lysis buffer (50 mM NaPi, 300 mM NaCl, 15 mM
imidazole, pH 8.0). Cells were lysed by sonication and cell debris was pelleted at 11,500 x g for
45 minutes. The lysate was passed over a Ni-NTA column (Qiagen) after filtering. The HγDCrys protein was eluted using a linear gradient to 100% B (50 mM NaPi, 300 mM NaCl, 250 mM
imidazole, pH 8.0). The fractions containing the protein were identified by SDS-PAGE and then
dialyzed three times against 10 mM ammonium acetate, pH 7.0. The concentration of HγD-Crys
was determined by A280 using an extinction coefficient of 42,860 M-1cm-1.
Mm-Cpn was expressed and purified as published (Knee et al. 2011). Briefly, pET21a
plasmid containing the Mm-Cpn gene was transformed into BL21 (DE3) Rosetta E. coli cells.
The cells were grown to OD600 0.6 and moved to 18°C and induced with 1 mM IPTG overnight
(~16 hours). Cells were pelleted and resuspended in MQ-A buffer (20 mM HEPES-KOH, pH
7.4, 50 mM NaCl, 5 mM MgCl2, 0.1 mM EDTA, 10% glycerol, 1 mM DTT, 0.1 mM PMSF, 1 mM
ATP). Cells were lysed via French Press and debris was pelleted at 11,500 x g for 45 minutes.
An ammonium sulfate cut of 55% was performed on the lysate, and the supernatant of the cut
was dialyzed against MQ-A (without ATP) overnight. After filtering, the sample was applied to a
Q Sepharose FF column (GE Healthcare). The sample eluted on a linear gradient to MQ-B (20
mM HEPES-KOH, pH 7.4, 1 M NaCl, 5 mM MgCl2, 0.1 mM EDTA, 10% glycerol, 1 mM DTT, 0.1
mM PMSF, 1 mM ATP). The fractions containing Mm-Cpn were identified by SDS-PAGE and
then concentrated with Vivaspin ultraconcentrators (Sartorius Stedim). The concentrated
sample was loaded onto a Superose 6 GL column (GE Healthcare) and eluted from the column
using MQ-A. The fractions containing Mm-Cpn were identified by SDS-PAGE, pooled,
concentrated, and buffer exchanged into MQ-A without ATP. The concentration of Mm-Cpn was
determined using the BCA Assay (Pierce) with BSA as a standard.
Thermal Denaturation by Circular Dichroism
Circular dichroism of HγD-Crys mutants at 100 µg/mL in 10 mM NaPi, pH 7.0 was
measured. Signal at 218 nm was monitored as the temperature was increased by 1 °C from 25
to 90 °C. Two-state fitting was done in MATLAB on three independent measurements to
calculate the midpoint of thermal denaturation.
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Aggregation Suppression of HγD-Crys by Mm-Cpn
The HγD-Crys aggregation suppression assay was performed just as in Knee et al
(2011). Briefly, 22 µM HγD-Crys (or 50 µg/mL) was unfolded overnight at 37 °C in unfolding
buffer (5.5 M GdnHCl, 50 mM Tris-HCl, pH 7.5, 5 mM DTT). Other concentrations (100 µg/mL,
150 µg/mL, and 200 µg/mL) were used to establish concentration curves for the mutants and
WT. The unfolded HγD-Crys was diluted 1:10 into refolding buffer (50 mM Tris-HCl, pH 7.5, 50
mM KCl, 5 mM MgCl2, 1 mM DTT, 1 mM ATP) with or without 22 µM Mm-Cpn to start the
aggregation reaction. The kinetics of the reaction were monitored as solution turbidity at 350 nm
in a Varian Cary 50 UV/Vis spectrophotometer, using the Varian Kinetics program. A two-state
kinetic fit was performed in MATLAB on six to nine independent aggregation experiments. The
change in absorbance due to light scattering (ΔA) was calculated by subtracting the minimum
value (usually time 0) from the fit maximum value.
After the aggregation reaction, the samples were 0.22 µm filtered, loaded on a Superose
6 GL column (GE Healthcare) and eluted from the column using MQ-A (without ATP). Fractions
containing refolded HγD-Crys were identified by SDS-PAGE with Krypton staining (Pierce), and
concentrated to 0.5 mL. The fluorescence of the refolded HγD-Crys was measured on a Hitachi
F-4500 Fluorimeter using unfolded HγD-Crys and native HγD-Crys as controls.
Quantification of Refolded HγD-Crys
To quantify the amount of HγD-Crys refolded, three independent 50 µL aggregation
suppression experiments were set up as outlined above. The samples were 0.22 µm filtered
and the residual GdnHCl was removed using the SDS-PAGE Sample Prep Kit (Pierce).
Samples were run on a 14% SDS-PAGE with HγD-Crys standards of known concentrations.
ImageJ was used to quantify amount of HγD-Crys refolded as compared to the standard curve
of HγD-Crys standards.
137
Results
The mutants we selected (Figure 5-1) fell into three classes: buried aromatic pairs,
domain interface residues, and buried hydrophobic core mutants. While all of the mutants were
slightly destabilized from WT (Table 5-1), they were all still very stable and could easily fold up
when expressed in E. coli. To assay for chaperonin recognition of HγD-Crys intermediates, we
initiated refolding by dilution out of denaturant. The protein concentration after dilution was
selected so that aggregation of partially folded intermediates predominated over spontaneous
refolding (Acosta-Sampson and King 2010). Aggregation of the HγD-Crys mutants in the
presence and absence of Mm-Cpn was measured by monitoring turbidity at 37 °C over 30
minutes (representative curves in Figures 5-2, 5-5A, and 5-6A). When the mutant HγD-Crys
proteins alone were tested for their aggregation in this reaction, their changes in light scattering
were not statistically different from the change in light scattering for WT (ΔA = 0.3-0.5) (Knee et
al. 2011). Although this could suggest that their aggregation mechanism is that of WT, the
changes in light scattering as a function of concentration suggested that the pathways of
aggregation for the mutants differed from that of WT. Therefore, it is likely that different
intermediate states are populated for different lengths of time between the mutants and WT.
Buried Aromatic Pairs
Paired aromatic residues in the partially folded state of HγD-Crys may constitute a signal
of recognition by the chaperonin. Paired tyrosine residues in HγD-Crys are highly conserved.
We examined two of these pairs in the C-terminal domain: Y97/Y92 in one of the Greek keys
and Y138/Y133 in the other Greek key. Mutant proteins with only one member of the Y92/Y97
and Y133/Y138 pairs replaced by alanine, and double mutant proteins Y92A/Y97A and
Y133A/Y138A with both tyrosines substituted were used as substrates for Mm-Cpn function.
Figure 5-2A shows that Mm-Cpn suppressed the off-pathway aggregation of Y92A, Y97A, and
Y92A/Y97A proteins by about 70% (Table 5-2). The aggregation of Y138A, Y133A and
Y133A/Y138 was also efficiently suppressed by incubation with Mm-Cpn (by about 50%; Figure
5-2B and Table 5-2), though not to the same extent as the other tyrosine pair.
138
Table 5-1: All HγD-Crys mutants are destabilized compared to WT HγD-Crys
HγD-Crys
Thermal Denaturation (°C)
WT
83.8 ± 1.31
L5S
73.0 ± 0.22
V75D
71.7 ± 0.22
I90F
74.8 ± 0.42
Q54A
80.1 ± 0.1
Q143A
78.6 ± 0.2
Q54AQ143A
77.8 ± 0.1
Y92A
77.0 ± 0.23
Y97A
79.0 ± 0.23
Y92AY97A
75.6 ± 0.2
Y133A
72.1 ± 0.23
Y138A
71.3 ± 0.63
Y133AY138A
73.5 ± 0.3
1
published in Flaugh, et al. 2006
2
published in Moreau & King 2009
3
published in Kong & King 2011
139
Figure 5-2: HγD-Crys aromatic pair mutants suppressed by Mm-Cpn
Representative curves of aggregation suppression of Y92A, Y97A, and Y92A/Y97A (A) and
Y133A, Y138A, and Y133A/Y138A (B) HγD-Crys mutants are shown in lighter colors alone and
in darker colors with Mm-Cpn. While all mutants are suppressed by Mm-Cpn, the mutants of the
first aromatic set (Y92/Y97) seem to be slightly better suppressed than those of the second
aromatic set (Y133/Y138).
140
Table 5-2: Kinetics of Mm-Cpn suppression of HγD-Crys aggregation vary by mutant
Half-Time
Change in Light Scattering (Final – Initial)
HγD-Crys
only
+ Mm-Cpn
only
+ Mm-Cpn
Suppression %
WT*
48 ± 1
398 ± 122
0.30 ± 0.01
0.10 ± 0.02
62 ± 15
L5S
58 ± 8
811 ± 344
0.36 ± 0.03
0.15 ± 0.05
58 ± 20
V75D
42 ± 5
488 ± 44
0.29 ± 0.01
0.15 ± 0.03
47 ± 10
I90F
63 ± 6
1000 ± 139
0.52 ± 0.04
0.27 ± 0.03
49 ± 7
Q54A
75 ± 11
986 ± 353
0.25 ± 0.04
0.12 ± 0.02
52 ± 11
Q143A
66 ± 6
828 ± 87
0.35 ± 0.02
0.19 ± 0.02
47 ± 6
Q54AQ143A
98 ± 5
327 ± 86
0.42 ± 0.04
0.19 ± 0.03
55 ± 9
Y92A
47 ± 9
867 ± 206
0.35 ± 0.06
0.13 ± 0.02
63 ± 14
Y97A
47 ± 5
1078 ± 155
0.44 ± 0.04
0.12 ± 0.01
73 ± 10
Y92AY97A
50 ± 5
755 ± 79
0.41 ± 0.03
0.12 ± 0.01
69 ± 5
Y133A
71 ± 2
367 ± 13
0.51 ± 0.01
0.24 ± 0.02
52 ± 3
Y138A
70 ± 1
290 ± 13
0.55 ± 0.01
0.35 ± 0.02
37 ± 3
Y133AY138A
67 ± 1
321 ± 14
0.55 ± 0.03
0.24 ± 0.02
56 ± 6
*published in Knee, et al. 2010
141
After the aggregation suppression reaction, samples were filtered through 0.22 µm
membranes to remove high molecular weight aggregates and fractionated by size exclusion
chromatography. As shown in Figures 5-3A and 5-3B, three peaks of protein were recovered:
the small peak of transient Mm-Cpn/HγD-Crys complex eluting earliest, the central peak
representing free chaperonin, followed by the refolded substrate eluting last in the series.
Fluorescence emission measurements of the refolded HγD-Crys species confirmed that they
were in a native-like conformation (Figure 5-4). This confirmed that the mutant folding
intermediates were successfully recognized, bound to, and refolded by Mm-Cpn.
After filtering, the aggregation suppression reactions were also run out on 14% SDSPAGE. A band corresponding to native-like HγD-Crys was seen for all mutants, indicating that
Mm-Cpn was efficiently refolding the mutant proteins. For WT, about 20% of HγD-Crys was
refolded by Mm-Cpn when the aggregation suppression assay was carried out at a 1:1 HγDCrys:Mm-Cpn ratio. Interestingly, while Mm-Cpn refolded twice as much of the Y92A/Y97A
mutant pair, it only refolded half as much of the Y133A/Y138A mutant pair (Figure 5-7). This
may be because: the Y133A/Y138A mutants aggregated more than the Y92A/Y97A mutant pair,
the Y133A/Y138A mutants were more destabilized than the Y92A/Y97A mutant pair (Table 5-1),
or the intermediate species populated by Y133A/Y138A were less recognized by Mm-Cpn than
the Y92A/Y97A intermediates.
Domain Interface Residues
All known γ- and β-crystallins have duplicated domains, and thus contain a distinctive
domain interface. A feature of the interface is a pair of interacting glutamines, one from each
domain (Flaugh et al. 2005; Flaugh et al. 2005). Studies of the folding and unfolding of HγDCrys reveal the presence of a relatively long-lived intermediate with the C-terminal domain
folded, and the N-terminal domain disordered (Kosinski-Collins and King 2003). Thus, in this
species, the C-terminal face of the domain interface is presumably exposed. If this is
recognized, altering residues in the interface might affect chaperonin recognition or binding. We
therefore examined as substrates single replacements of each glutamine (Q54A and Q143A)
and the double mutant Q54A/Q143A (Flaugh et al. 2005). In the presence of Mm-Cpn, their
aggregation was suppressed with efficiencies of about 50% (Figure 5-5A and Table 5-2).
142
Figure 5-3: HγD-Crys aromatic pair mutants refolded to native-like state by Mm-Cpn
Size exclusion chromatography of Y92A, Y97A, and Y92A/Y97A (A) and Y133A, Y138A, and
Y133A/Y138A (B) HγD-Crys mutants show three distinct peaks: Mm-Cpn-HγD-Crys complex
(~12 mL), Mm-Cpn only (~16 mL), and refolded HγD-Crys (~21 mL). All mutants are refolded to
native-like HγD-Crys by Mm-Cpn.
143
Figure 5-4: HγD-Crys mutants refolded by Mm-Cpn have native-like fluorescence
Fluorescence measurements of two representative aromatic mutants (Y97A in purple and
Y138A in red) show that the refolded species have native-like fluorescence when compared to
native WT HγD-Crys (blue) and unfolded WT HγD-Crys (orange).
144
Figure 5-5: HγD-Crys interface pair mutants suppressed and refolded to native-like state by
Mm-Cpn
A. Representative curves of aggregation suppression of Q54A, Q143A, and Q54A/Q143A HγDCrys mutants are shown in lighter colors alone and in darker colors with Mm-Cpn. B. Size
exclusion chromatography of Q54A, Q143A, and Q54A/Q143A HγD-Crys mutants show three
distinct peaks: Mm-Cpn-HγD-Crys complex (~12 mL), Mm-Cpn only (~16 mL), and refolded
HγD-Crys (~21 mL). All mutants are refolded to native-like HγD-Crys by Mm-Cpn.
145
These reactions were filtered to removed high molecular weight aggregates and
analyzed by size exclusion chromatography for molecules refolded back to the native state
(Figure 5-5B). The irregularity in the monomer peak for the double mutant might represent the
presence of a perturbed folded conformation for the double mutant, given its reduced stability
compared to WT (Table 5-1) (Flaugh et al. 2005). The decreased recovery of the MmCpn/substrate complex is consistent with the increased yield of refolded native-like HγD-Crys
molecules. A significant amount of refolded monomers was recovered from all three mutant
proteins, as seen by fluorescence of the refolded HγD-Crys peak from size exclusion and SDSPAGE of filtered aggregation suppression reactions. As with the Y92A/Y97A set of aromatic
mutants, the yield of refolded molecules carrying the interface mutants was higher than that of
WT HγD-Crys (Figure 5-7).
Buried Core Hydrophobic Mutants
Mutants at three buried core sites in γ-crystallins result in congenital cataracts in mice:
L5S, V75D, I90F (Sinha et al. 2001; Graw et al. 2002; Graw et al. 2004). When these
substitutions were made in HγD-Crys, they resulted in significant destabilization of the protein
(Moreau and King 2009). The off-pathway aggregation of these mutants was monitored by
turbidity as above, in the presence or absence of Mm-Cpn. As can be seen in Figure 5-6A, all
three mutant chains were suppressed by Mm-Cpn to about 50% (Table 5-2). The aggregation of
I90F during refolding was distinctively slowed upon incubation with chaperonin, but turbidity
continued to increase at a slow rate during the course of the incubation.
To assess whether the mutant proteins were refolded by Mm-Cpn during the reaction,
the reactions were filtered, and analyzed by size exclusion chromatography. The early eluting
HγD-Crys/Mm-Cpn complex was recovered in all three reaction mixtures. A significant yield of
refolded HγD-Crys proteins were recovered for all congenital mutants by fluorescence
measurement of the HγD-Crys peak from size exclusion chromatography and SDS-PAGE of
filtered aggregation suppression reactions. The two mutant proteins that were not better
refolded than WT were V75D and I90F (Figure 5-7). Moreau and King showed that V75D
exhibited an aggregation pathway from the native state, and that the aggregation intermediates
were poorly recognized by the lens chaperone αB-crystallin (2012). Similar phenomena may be
operating with Mm-Cpn.
146
Figure 5-6: HγD-Crys hydrophobic core mutants suppressed and refolded to native-like state by
Mm-Cpn
A. Representative curves of aggregation suppression of L5S, V75D, and I90F HγD-Crys
mutants are shown in lighter colors alone and in darker colors with Mm-Cpn. B. Size exclusion
chromatography of L5S, V75D, and I90F HγD-Crys mutants show three distinct peaks: MmCpn-HγD-Crys complex (~12 mL), Mm-Cpn only (~16 mL), and refolded HγD-Crys (~21 mL). All
mutants are refolded to native-like HγD-Crys by Mm-Cpn.
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Figure 5-7: Most HγD-Crys mutants refolded to higher levels than WT HγD-Crys
Amount refolded relative to WT (of which 20% is refolded) is shown with aromatic mutants in red
and orange, interface mutants in blue, and core mutants in green. Most mutants were refolded
to twice the level of WT, except for the more destabilized Y133A/Y138A aromatic pair mutants
and the potentially harder to recognize V75D and I90F core mutants.
148
In general, although Mm-Cpn did not suppress aggregation of HγD-Crys mutants
differentially from aggregation of WT HγD-Crys, Mm-Cpn better recognized partially folded
intermediates of most HγD-Crys mutants than WT, and also successfully refolded most HγDCrys mutants to levels higher than WT.
149
Discussion
Investigations of the substrate recognition by both group I and group II chaperonins
indicate that they recognize partially folded conformations of their target proteins. For most
substrates binding to the native state is weak or undetectable. In general, it has not been
possible to image, at high resolution, the bound conformation or the substrate, so direct
assessment of the binding or recognition sites on the substrate are limited (Dekker et al. 2011).
Genetic studies have often revealed residues involved, such as the Box 1 and Box 2 residues in
VHL protein, recognized by TRiC (Feldman et al. 2003). However, the conformation of these
residues in the bound state remains unclear.
The ability to refold HγD-Crys in vitro under physiological conditions – pH 7 and 37 °C –
led to the identification of a partially folded intermediate with the N-terminus disordered and the
C-terminus folded. As the concentration of protein increased, off-pathway aggregation
competed with productive refolding. These reactions have provided a substrate for the lens
chaperone α-crystallin, and for the group II chaperonins Mm-Cpn and TRiC (Acosta-Sampson
and King 2010; Knee et al. 2011; Sergeeva et al. 2013)
Characterization of the recognition by the lens chaperone α-crystallin indicated that the
conformation recognized was a slightly earlier intermediate with the C-terminus not fully folded
(Acosta-Sampson and King 2010). The contributions of a large set of sites to the folding and
stability of HγD-Crys have been systematically examined (Kosinski-Collins and King 2003;
Kosinski-Collins et al. 2004; Flaugh et al. 2005; Chen et al. 2006; Kong and King 2011). By
studying the ability of Mm-Cpn to suppress the aggregation and/or enhance the refolding of
mutants crystallins, we hoped to identify specific resides that were signals for substrates
involved in group II chaperonin recognition.
Though the off-pathway aggregation of some mutants were suppressed slightly better
(Y97A) than others (L5S), aggregation of all mutants was sufficiently suppressed by Mm-Cpn.
Similarly all the mutant proteins were aided in their refolding to the native state by incubation
with Mm-Cpn. Refolding is likely to be a more stringent test than aggregation suppression by the
group II chaperonins. These results suggests that the features we examined as possible MmCpn-recognition signals – aromatic pairs, interface contacts, and selected buried core residues
– were actually not signals for the chaperonin. The aggregation half-time of all mutants in the
absence of Mm-Cpn was about the same as the aggregation half-time of WT HγD-Crys (~50 s)
(Knee et al. 2011). However, the aggregation half-time of most mutants in the presence of MmCpn was longer than the aggregation half-time of WT HγD-Crys (for example, 1062 ± 349
seconds for Y97A vs. 398 ± 122 seconds for WT) (Knee et al. 2011), suggesting Mm-Cpn
150
recognized the mutant intermediates more efficiently than it recognized WT HγD-Crys folding
intermediates.
An interesting result from this study was the increased refolding of HγD-Crys mutants
upon incubation with Mm-Cpn as compared to WT HγD-Crys. Mm-Cpn actually seemed to be
recognizing these mutant intermediates better than WT. Many of these mutants slow the rate of
refolding when examined by kinetic refolding experiments (Kosinski-Collins and King 2003;
Flaugh et al. 2006; Kong and King 2011). It seems likely that the partially folded intermediates
that are the chaperonin substrates have longer lifetimes, or increased populations, which would
result in better recognition by the chaperonin. The recovery of substrate/chaperonin complexes
via size exclusion chromatography supports this interpretation. However, we cannot rule out the
possibility that the altered amino acids increase exposure or availability of the actual recognition
signals, for example due to the substitution of bulky tyrosines and glutamines with smaller
alanines. Additional experiments on the nature of the substrate/Mm-Cpn complexes will be
needed to resolve these two models.
Presumably Mm-Cpn is recognizing some feature of the β-sheet that is exposed in a
variety of folding intermediates. Due to the β-sheet-rich structure of HγD-Crys, this feature could
potentially be a β-sheet interface between the Greek keys of a domain, or an exposed surface
of a β-strand. This is especially likely due to the evidence that group II chaperonins
preferentially recognize and refold β-sheet-rich proteins and recognize hydrophobic β-sheets
(Camasses et al. 2003; Yam et al. 2008). How exactly Mm-Cpn can recognize a β-strand is still
unknown. Hsp70 chaperones recognize their substrates through hydrophobic residues and a
similar interaction has been theorized to occur in the group II chaperonins (Frydman 2001;
Feldman et al. 2003). Since one side of a β-strand can be quite hydrophobic, Mm-Cpn can
potentially recognize this feature and bind the partially folded protein. On the other hand, if the
chaperonin recognized features of the β-sheet backbone, this might be only mildly affected by
the side chain composition.
It has long been known that chaperones such as Hsp90 act as buffers for mutant
substrate proteins (Rutherford and Lindquist 1998). This buffering allows proteins to sample
possible beneficial mutations without sacrificing folding ability, therefore assisting protein
evolution (Rutherford and Lindquist 1998). The results in this work provide an additional
example of this phenomenon with chaperonins. A variety of mutants, all somewhat destabilized
in the native state, were successfully chaperoned during folding by the group II chaperonin.
151
152
CHAPTER 6:
Final Discussion and Future Directions
153
Final Discussion
As far as we currently understand, all human cell types require functional TRiC
chaperonin for cell division and growth. While there has been a great deal of research on
archaeal group II chaperonins, and bovine and yeast TRiC, studies of human TRiC have been
limited. This is in large part due to the fact that bovine tissues and yeast cultures are easier to
obtain than human tissues or large-scale cultures of human cells. However, as more findings
emerge of TRiC’s interaction with protein substrates implicated in diseases, characterization of
human TRiC will increase in importance. To that end, this thesis investigated structure and
function of endogenous human TRiC, the properties of individual human CCT subunits,
mutations in two human CCT subunits implicated in disease, substrate recognition signals for
group II chaperonins, and how human CCT subunits may assemble into mature TRiC.
In Chapter 2, I report characterization of the human TRiC protein purified from HeLa
cells. This material was well organized, containing all eight CCT subunits. By electron
microscopy of negatively stained samples, the complexes appeared as back-to-back rings with
eight subunits each. This is consistent with the studies of bovine and yeast TRiC (Frydman et al.
1992; Liou and Willison 1997; Pappenberger et al. 2006; Cong et al. 2010; Cong et al. 2011;
Dekker et al. 2011). Human TRiC was active with two different substrates previously used in
chaperonin assays: luciferase and human HγD-Crys (Frydman et al. 1992; Knee et al. 2011).
Interestingly, Hsp90 and Hsp70 associated with human TRiC through many steps of the
purification, only to be finally disrupted by heparin chromatography. This interaction suggests
that there are ways the cell shuttles substrates between the chaperone systems. While Hsp70TRiC interaction has been previously characterized, the Hsp90-TRiC interaction remains to be
characterized (Hartl et al. 2011). In general, human TRiC was similar to the other TRiC species
studied previously.
In Chapter 3, I report studies of the individual human CCT subunits. I expressed each of
the eight CCT subunits one at a time in E. coli and was able to recover full-length chains. By
sucrose gradient ultracentrifugation, these chains were predominantly either soluble monomer
or dimer species (CCT2, CCT3, CCT7, and CCT8), formed 20S complexes (CCT4 and CCT5),
or seemed to stick to ribosomes or may have aggregated through the gradient (CCT1 and
CCT6). I was particularly interested in the two species that formed rings absent from the other
subunits – CCT4 and CCT5. Further purification of these species and subsequent structural
characterization by negative-stain and cryo-EM showed that they were double rings of eight
subunits per ring. Activity assays showed they were active in hydrolyzing ATP, refolding
luciferase, and suppressing and refolding HγD-Crys. The fact that these complexes formed and
154
were fully active was surprising. Some archaeal chaperonins can form homo-oligomeric rings
even if they possess more than one chaperonin subunit (Sahlan et al. 2009). However, homooligomers of the CCT subunits have never been postulated or reported. General consensus in
the field is that TRiC only forms hetero-oligomers of eight subunits per ring. Since I was able to
show the presence of active homo-oligomers, it may be that there are species of CCT/TRiC not
limited to canonical TRiC. In Appendix B of Chapter 8, I assay the two homo-oligomers CCT4
and CCT5, along with human TRiC and archaael Mm-Cpn, for their ability to suppress
aggregation of mHtt. While CCT5 can suppress its aggregation as well as human TRiC, both
CCT4 and Mm-Cpn fail to adequately suppress mHtt aggregation. Therefore, we show mHtt
recognition and binding is not only specific to the eukaryotic chaperonin, but is more efficient
with CCT5 than CCT4. This is the first evidence that points to the CCT subunits interacting with
particular substrates.
In Chapter 4, I report studies of two CCT mutants that have been postulated to be
causally associated with neuropathies (Lee et al. 2003; Bouhouche et al. 2006). These
mutations have only been found in CCT4 and CCT5, the two subunits that homo-oligomerize in
E. coli. These two mutations, both in the equatorial domain, lead to different defects when
studied in our homo-oligomer system. C450Y CCT4 mutant was defective in folding and
assembly, while H147R CCT5 mutant was unable to suppress aggregation of and refold HγDCrys, suppress aggregation of mHtt, and refold β-actin to the same level as WT CCT5. While it
is premature to speculate that these defects are what cause neuropathy in the patients, finding
a clear difference between WT and mutant at the biochemical level was encouraging.
Presumably, the defects are exaggerated in our homo-oligomer system, so they are potentially
subtler when these mutated subunits are part of mature TRiC. This is not surprising, however,
because these patients do live to adulthood, so the defect cannot be too extreme.
In Chapter 5, I diverge from the human CCT theme to explore substrate signals for
chaperonin recognition. Mm-Cpn was used as the chaperonin in these studies due to its ease of
purification and its characterized aggregation suppression of a substrate, HγD-Crys (Knee et al.
2011). Mm-Cpn was challenged with mutants of potential substrate recognition signals in HγDCrys, to see whether it could still recognize this substrate when it was missing something
crucial. These signals were in the unpaired aromatics, domain interface, and hydrophobic core.
Mm-Cpn not only still recognized all of these mutants, it actually recognized them better than
WT and refolded them to higher levels than it refolded WT. This suggested that what Mm-Cpn
was actually recognizing was a β-sheet interface that was more exposed when we replaced
some of the bulkier side-chains with smaller alanines. This is actually not too surprising because
155
group II chaperonin substrates have little in common other than being more β-sheet-rich than
other proteins in the cytoplasm (Yam et al. 2008). By recognizing a more general feature of the
substrate, the group II chaperonins expand the number and types of substrates that they can
assist.
Finally, in Appendix A of Chapter 8, I returned to the question of how human CCT
subunits are assembled into mature TRiC. The CCT subunits are expressed from seven
different chromosomes, indicating a requirement for the regulation of their complex assembly. In
order to study potential interactions on the way to TRiC, I co-expressed each CCT subunit with
a homo-oligomerizing chaperonin subunit (either CCT4, CCT5, or Mm-Cpn). CCT5 was the
most efficient in driving CCT subunits into chaperonin-sized 20S complexes, as surveyed by
sucrose ultracentrifugation gradients, affecting all subunits but CCT6. CCT4, on the other hand,
only pushed CCT8 into 20S species. Mm-Cpn, surprisingly, interacted intermediately between
CCT5 and CCT4. I was expecting this ancestral subunit to interact with all of the CCT subunits,
but it may be that many of the CCT subunits diverged too far and can no longer interact with it.
A limitations of these experiments was the difficulty in distinguishing whether the assisting
species acted at the level of folding, generating subunits competent for assembly, or whether
the effect was specifically at the level of subunit assembly. While these co-expression results
are still preliminary, and I cannot be sure these 20S complexes are true hetero-oligomers, the
selectivity of the interactions gives us a starting place for thinking about TRiC assembly. One
model, consistent with the experiments, is that the CCT4 and CCT5 homo-oligomers are
starting templates due to their increased stability in complexes in comparison to monomer
subunits.
Although investigations in the last twenty years have expanded our knowledge of the
eukaryotic chaperonin, investigations on TRiC have focused on its ATP hydrolysis and substrate
interaction properties (Willison et al. 1989; Llorca et al. 1999; Llorca et al. 2001; Feldman et al.
2003; Spiess et al. 2006; Tam et al. 2006; Reissmann et al. 2007; Booth et al. 2008; Yam et al.
2008; Zhang et al. 2010; Dekker et al. 2011; Douglas et al. 2011; Jiang et al. 2011; Pereira et al.
2012; Reissmann et al. 2012). The surprising finding that homo-oligomeric CCT subunits can be
as active as hetero-oligomeric TRiC (at least for the substrates investigated herein), suggests
that the CCT subunits of TRiC have more dynamic roles than just being part of a static complex.
More work will need to be done to better understand if TRiC is made up of different subunits
during development or in tissues where it would be beneficial to have more of one CCT subunit
than another. It’s possible that tissue and substrate-specific TRiC rings exits but have not been
isolated or reported due to the fact that TRiC research has been limited to specific bovine and
156
yeast materials, and the technology to discern these small structural differences has not yet
been developed.
The interaction between HγD-Crys substrate and the group II chaperonins Mm-Cpn and
TRiC is presumably through β-sheet or strand recognition, making it an representative substrate
for chaperonin studies. This human substrate of the eye lens most likely interacts with TRiC in
the lens periphery during the development of the eye lens. The interaction between Mm-Cpn
and HγD-Crys is more artificial, but important chaperonin recognition information can be
gathered from that substrate-chaperonin combination. In the lens, the main small heat shock
chaperone is α-crystallin (Moreau and King 2012). In vitro, α-crystallin can suppress
aggregation of HγD-Crys by binding and holding the partially folded species, but cannot refold
the chains to native-like state (Acosta-Sampson and King 2010; Moreau and King 2012). On the
other hand, TRiC (and CCT4 and CCT5) and Mm-Cpn can suppress aggregation of HγD-Crys
and refold this substrate to native-like state. This is a special property of the chaperonins that
the small heat shock chaperones do not possess.
In addition, we found that Mm-Cpn and CCT5 were able to differentially refold some
mutants of HγD-Crys. Mm-Cpn refolded a wide range of mutants to levels higher than that of
WT, suggesting that it was preferentially helping the folding of more destabilized chains. While
other chaperones have been implicated in protein evolution by assisting mutant proteins in
folding, therefore allowing them to adapt potentially beneficial mutations, there has been little
study of this for the chaperonins (Rutherford and Lindquist 1998; Lindquist 2009). Interestingly,
one mutant of the paired aromatics, Y92A/Y97A HγD-Crys, was refolded to twice the level of
WT HγD-Crys by Mm-Cpn, but only to half the level of WT HγD-Crys by CCT5. This may mean
that CCT5 is less efficient at recognizing destabilized mutants than Mm-Cpn or that it cannot
assist them as well as Mm-Cpn can. It may also be that CCT5 has trouble recognizing this
mutant but that another CCT subunit may better recognize it or that CCT5 may be more efficient
at recognizing and refolding different HγD-Crys mutants.
While studies of substrate recognition by TRiC and other group II chaperonins are
limited, specific recognition signals on the substrate have been studied for other chaperones
and complexes. In general, we found that the group II chaperonin potentially recognizes the βsheet interface rather than a specific feature in the substrate sequence. Three other examples
where substrates are recognized in a non-specific way are: Hsp70, small heat shock proteins
such as α-crystallin, and the major histocompatibility complex (MHC). For Hsp70, the substratebinding domain recognizes stretches of five hydrophobic amino acids, flanked by positivelycharged amino acids, normally buried in the native state (Mayer 2013). Small heat shock
157
proteins such as α-crystallin recognize their substrates also through hydrophobic interactions,
but due to their assemblies, the interaction is not limited to linear stretches of amino acids (Clark
et al. 2012). The MHC displays peptides on the cell surface for other immune cells to recognize.
These peptides, usually 8-12 amino acids in length, are bound to the MHC in a well-regulated
manner (Purcell 2000; Cresswell et al. 2005). However, while there is some specificity in
peptides for specific MHCs, overall, the recognition of peptides is quite degenerate to allow
presentation of a wide variety of antigens (Rothbard and Gefter 1991). In light of these
examples, it is quite feasible that group II chaperonins also interact with substrates in a general
way, using common features of the substrate such as β-sheet interfaces.
Being able to make large quantities of natively-folded proteins in E. coli is important for
the biotechnology sector. While expression of human or other potential therapeutic proteins in
E. coli can be optimized, not all proteins are able to fold in bacteria and end up degraded or in
inclusion bodies. One way to increase the fraction of folded exogenous chains in E. coli is the
addition of molecular chaperones to the cells (Ito and Wagner 2004; de Marco 2007; MartínezAlonso et al. 2009). However, most of the chaperones that have been added in such protocols
have been DnaK, DnaJ, and GroE: all bacterial components. Due to our work of being able to
form structural and functional human CCT subunits in E. coli, co-expression of our CCT4 and
CCT5 chains with difficult-to-fold human proteins may be an ideal strategy for producing highquantity folded proteins. If human proteins need to be expressed to high levels in E. coli, there is
a good chance that they need the human chaperonin (or at least a subset of it) to fold correctly
in bacterial cells.
158
Future Directions
One of my main interests in studying CCT subunits individually was to determine if
different CCT subunits are needed to recognize and bind specific substrates. While this
question was partly answered by the model substrates in this thesis, more stringent substrates
such as actin and tubulin would have to be used to fully address this model. Therefore, one of
the most important next steps is addressing substrate specificity with actin and tubulin. As of
right now, it looks like CCT4 and CCT5 can function just as well as human TRiC on their own.
However, this would be at odds with the argument that TRiC evolved eight different CCT
subunits to recognize and bind different kinds of substrates (Kim et al. 1994). One of the
reasons I might be seeing no specificity in the substrates that I tested is that these are not
obligate substrates of TRiC and therefore any CCT subunit is sufficient to recognize and bind
them. Without testing actin and tubulin (and possibly other more strict substrates), I will not
know for sure about the substrate specificity properties of the CCT subunits.
Another important direction of study with the homo-oligomeric CCT subunits is the
question of ATP hydrolysis. Recent work from the Frydman lab showed that TRiC has an
asymmetrical power stroke driven by the subunits that bind and hydrolyze ATP most efficiently –
CCT4 and CCT5 (Reissmann et al. 2012). However, I saw that the ATP hydrolysis of rate of
CCT4 and CCT5 was the same as that of human TRiC. Therefore, in the homo-oligomeric
complexes, there must be another mechanism for regulating ATP hydrolysis rates. It is unknown
whether that is by only allowing some subunits to bind and hydrolyze ATP or letting all subunits
bind and hydrolyze ATP but somehow decreasing the rate. In order to explore this, I can use our
system to genetically engineering homo-oligomers with mutations that inhibit ATP hydrolysis or
binding and assay their ATP hydrolysis rates.
Due to the limited study of human TRiC and its regulation, the extent of how posttranslational modifications affect TRiC structure or function is not yet known. One example is
that in response to growth factors or tumor promoters, CCT2 is phosphorylated at serine 260,
leading to cell proliferation (Abe et al. 2009). The CCT4 neuropathy mutation may be another
example where the regulation of the mutation may also be at a post-translational level – with a
cysteine being replaced by a tyrosine. Both these residues may be post-translationally modified,
so either the presence or absence of such modification may lead to a regulation defect in the
mutant. Therefore, one subsequent experiment is identifying whether there are modifications of
endogenous human TRiC. Since I already have the endogenous material from HeLa cells, it
would only require some high-resolution mass spectrometry to attempt to identify various
modified sites in TRiC.
159
The interactions between CCT subunits were a good way to study hetero-oligomer
formation on the way to mature TRiC. However, I cannot be sure that the 20S complexes of
sucrose ultracentrifugation gradients are indeed hetero-oligomers between two different
subunits or the homo-oligomer chaperoning the chains of the other subunits. More experiments
need to be done to verify and address this. One way to check this is to use negative stain EM
on CCT4 homo-oligomers alone and CCT4-CCT8 complexes. We know that CCT4 does not
contain anything in the cavity like CCT5 sometimes does, so if we see clear rings with nothing in
the cavity, it suggests that CCT4 and CCT8 are forming homo-oligomers. If, on the other hand,
we see a significantly higher proportion of rings with filled cavities in the purified sample where
CCT4 and CCT8 are co-expressed, it suggests that CCT4 is chaperoning rather than heterooligomerizing with CCT8. With CCT5, parsing out chaperoning versus hetero-oligomerizing is
harder since it does seem to sometimes contain its own chains inside its cavity. For that, native
gels may prove valuable. Eventually, it would be ideal to be able to sequentially add each
subunit one-by-one to make mature TRiC, following some of our proposed assembly schemes.
In order to do this, we would need to purify each subunit as monomers, which has not yet been
successful. Varying the purification conditions and possibly adding co-chaperones (Mm-Cpn
specifically) may allow us to purify some of the CCT subunits as monomers, in vitro add them to
CCT4 and CCT5, and assay whether hetero-oligomerization occurs.
One other more eventual future direction is verifying whether the defects seen for CCT4
and CCT5 neuropathy mutants are what is causing neuropathy in the patients. To do this, we
would need patient or rat cell lines carrying these homozygous mutations. Since these mutants
may be part of TRiC, we can purify TRiC from these cells and assay its stability and
chaperoning abilities. Based on our data, we would expect TRiC from the C450Y CCT4 mutant
to be less stable, possibly melting sooner by thermal denaturation. For H147R CCT5, we would
expect chaperoning ability to be hindered in the TRiC purified from the patient cells, possibly
inhibiting its ability to refold HγD-Crys and β-actin in vitro as compared to control TRiC. These
experiments, while not simple, will be the best way to show that our results are consistent with
the disease pathology.
160
CHAPTER 7:
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182
CHAPTER 8:
APPENDIX A:
Co-expression of CCT Subunits
to Explore Subunit Assembly*
* Cameron Haase-Pettingell is acknowledged for her technical assistance.
183
Abstract
The eight CCT subunits of TRiC are expressed from seven different chromosomes in the
cell. In order to assemble into mature TRiC, which contains one of each of these subunits, they
must be translated, correctly folded and assembled into the TRiC complex. Previous studies
showed that two of the subunits, CCT4 and CCT5, could form TRiC-like homo-oligomeric rings
absent of the other CCT subunits, while none of the other subunits formed such complexes. To
explore potential subunit-subunit, we co-expressed the homo-oligomerizing chaperonin CCT4
and CCT5 subunits with CCT1-8 one at a time in E. coli.
We found that CCT5 drove all of the CCT subunits but CCT6 into 20S complexes, while
CCT4 only interacted with CCT8 to push it into chaperonin rings. We hypothesize that these
specific interactions may be due to the formation of hetero-oligomers in E. coli, although more
work needs to verify this assumption. Models of TRiC assembly have been proposed based on
this hetero-oligomer data that rely on the CCT4 and/or CCT5 homo-oligomers as starting
complexes. Eventually, analysis of CCT arrangement in various tissues and at different
developmental times may provide additional information on TRiC assembly and CCT subunit
composition.
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Introduction
The eukaryotic chaperonin TRiC is made up of eight different subunits, designated
CCT1-CCT8. These subunits are expressed off of seven different chromosomes, but CCT6 has
two isoforms that are expressed from two different chromosomes, totaling nine subunits
expressed off of eight chromosomes (Table 8-1; NCBI). Most of the CCT subunits have various
characterized splice isoforms, but neither those nor the two different CCT subunits have been
studied in any rigorous manner. The CCT6 subunit used most frequently experimentally has
been CCT6A. Expression levels of individual CCT subunits indicate that the levels of each
subunit relative to every other is about equal, but these levels as a whole vary between different
tissues, and can be up or down regulated in cancer (Kubota et al. 1999; Yokota et al. 2001;
Boudiaf-Benmammar et al. 2013; Finka and Goloubinoff 2013). This may suggest that the levels
are finely regulated so that the subunits can form a complex with each subunit appearing once,
but there is also data to show that the subunits may have alternate functions apart from TRiC
(Roobol and Carden 1999).
While it is generally accepted that mature TRIC is made up of two rings of eight different
subunits, the arrangement of these subunits is still debated. Overall, the sequence identity
between CCT subunits is about 30%, making them structurally very similar (Horwich et al.
2007). Therefore, their individual conformations within TRiC have been difficult to resolve via
conventional negative-stain electron microscopy or low-resolution x-ray crystallography (Bigotti
and Clarke 2008). Over the years, many other methods have been used to determine the
arrangement of primarily bovine TRiC, but more recently, also yeast TRiC.
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Table 8-1: Human CCT subunit expressed from eight different chromosomes
Subunit
Chromosome
Location
Accession Numbera
1
(α) Alpha
6
6q25.3-q26
NM_030752.2
2
(β) Beta
12
12q15
NM_006431.2
3
(γ) Gamma
1
1q23
NM_005998.4
4
(δ) Delta
2
2p15
NM_006430.3
5
(ε) Epsilon
5
5p15.2
NM_012073.3
6A
(ζ) Zeta A
7
7p11.2
NM_001762.3
6B
(ζ) Zeta B
17
17q12
NM_006584.3
7
(η) Eta
2
2p13.2
NM_006429.3
8
(θ) Theta
21
21q22.11
NM_006585.3
a
All information from National Center for Biotechnology Information
186
The first proposed arrangement, from biochemical studies by the Willison group on
bovine TRiC, showed this order: CCT 1-5-6-2-3-8-4-7 (Liou and Willison 1997). Later, the
Frydman and Chiu groups used high resolution cryo-EM to obtain a structure of bovine TRiC
where the slight differences in structure between the subunits could be resolved, giving the
arrangement: CCT1-7-5-4-8-3-2-6 (Cong et al. 2010). The Willison group obtained a crystal
structure of yeast TRiC binding rabbit α-actin, and found that their previous arrangement docked
well into the electron density (Dekker et al. 2011). They did note that the register of the crystal
structure differed by one subunit counter-clockwise as compared to the earlier EM studies
(Dekker et al. 2011). Most recently, two cross-linking and mass spectrometry were in agreement
with a new arrangement: CCT1-3-6-8-7-5-2-4 (Kalisman et al. 2012; Leitner et al. 2012). The
authors of the latter work note that their new arrangement has a better fit in the yeast TRiC
crystal structure than the one used by the Willison group (Leitner et al. 2012).
Due to the controversy surrounding the “correct” mature TRiC arrangement, it is possible
that more than one arrangement may occur in TRiC purifications in these studies. Even with the
high-resolution methods, many particles, and therefore potential arrangements are discarded
during the processing stages. What may be even more likely is that different arrangement would
exist in various tissues or at distinctive developmental stages. The finding reported in Chapter 3
that two of the CCT subunits (CCT4 and CCT5) form TRiC-like homo-oligomers on their own,
suggests they might have particular competence for subunit assembly. We also previously
showed that all of the other CCT subunits did not form homo-oligomers when expressed in E.
coli.
Therefore, to explore the interactions between CCT4 and CCT5 and other CCT subunits
we co-expressed each CCT subunit one at a time with CCT4 or with CCT5. Regardless of which
of the models discussed above is correct, in all cases CCT4 and CCT5 would interact in TRiC
with three or four other subunits – one on either side within the ring and one or two (depending
on exact register) in the other ring. We also included in the experiment co-expression with the
archaeal chaperonin subunit of Methanococcus maripaludis (Mm-Cpn), which forms a homooligomeric 16-subunit chaperonin when expressed in E. coli (Reissmann et al. 2007; Douglas et
al. 2011; Knee et al. 2011). The archaeal Mm-Cpn was used as an evolutionary control, due to
the fact that the archaeal thermosome chaperonin genes are the presumed ancestors of the
eukaryotic chaperonin genes (Archibald et al. 1999; Horwich et al. 2007; Dekker et al. 2011;
Yébenes et al. 2011). These homo-oligomer subunits may drive the CCT subunits into
chaperonin complexes, showing a specific interaction possibly leading to hetero-oligomerization.
The expectation would be that the archaeal Mm-Cpn would hetero-oligomerize with the most
187
CCT subunits, while CCT4 and CCT5 would only hetero-oligomerize with specific subunits they
interact with on the way to becoming mature TRiC rings. Alternatively, Mm-Cpn, CCT4, or CCT5
may also act as chaperones, allowing the other CCT subunits to fold to more assemblycompetent forms.
188
Materials and Methods
Plasmid Construction
Two chaperonin subunits at a time were inserted in the pETDuet plasmid (Novagen).
The CCT subunit genes (1-8) contained a c-terminal TEV protease cleavage site and a 6xHistag, whereas the Mm-Cpn gene was inserted unmodified. Multiple cloning site one (MCS2)
contained CCT4, CCT5, Mm-Cpn or nothing, while MCS1 contained CCT1-8, for a total of 32
plasmids. The restriction enzymes for CCT1-8 in MCS1 were SpeI and AscI, where SpeI was
inserted into the plasmid via mutagenesis. For MCS2, the restriction enzymes for CCT4 and
CCT5 were NdeI and KpnI, while for Mm-Cpn they were NdeI and BamHI. All plasmids were
confirmed by sequencing (Genewiz).
Expression and Lysis
Plasmids were transformed into E. coli BL21 (DE3) RIL cells. Proteins were expressed
as before (Sergeeva et al. 2013). Briefly, cells were grown in Super Broth to OD 5.0 at 37 °C
and then shifted to 18 °C and induced with 0.5 mM IPTG. After the overnight induction, cultures
were pelleted by centrifugation for 15 min, and the cells were resuspended in CCT-A (20 mM
HEPES/KOH pH 7.4, 300 mM NaCl, 10 mM MgCl2, 10% glycerol, 1 mM DTT, 1 mM ATP) with
addition of one EDTA-free Complete protease inhibitor (Roche) per L of culture. To lyse the
cells, 1 mM DTT, 5 mM MgCl2, and 2.5 mg/mL lysozyme, and 10 µg/mL DNase was added to
the pellets. After an incubation with shaking at approximately 12 °C, the cells were lysed via
French Press. Debris was spun down at 11,500 x g and supernatant was isolated by pipetting.
Pellet was resuspended in CCT-A buffer.
Sucrose Gradient Sedimentation
Using CCT-A buffer, 5-40% sucrose gradients were prepared by the gradient master
(BioComp Instruments). Lysates (100 µL) were added carefully to the top and gradients were
ultracentrifuged at 4 °C for 18 h at 28,000 rpm using a SW50 rotor (Beckman). Nineteen or
twenty fractions were collected using a gradient fractionator (BioComp Instruments), and one
bottom fraction was collected from the leftover gradient.
SDS-PAGE and Immunoblots
Proteins were separated by SDS-PAGE (10%) at 165 V for 1 h after boiling in reducing
buffer (60 mM Tris, pH 6.8, 2% SDS, 5% β-mercaptoethanol, 10% glycerol, bromophenol blue
for color) for 5 min. The gels were stained with Coomassie blue. Transfer was conducted for 1.5
189
h at 300 mA in transfer buffer (10% methanol, 25 mM Tris, 192 mM glycine) onto 0.45 µm
polyvinylidene difluoride (PVDF) membranes (Millipore). The primary antibodies were from
Santa Cruz Biotechnology (Table 8-2). An anti-His(C-term)-AP antibody (Life technologies) was
sometimes used for further verification. The secondary antibodies were Alkaline Phosphatase
(AP)-conjugated (Millipore) and the membranes were visualized using the AP-conjugate
substrate kit (BioRad).
Quantification
Band quantification was done using ImageJ for both full-length and fragments of the
CCT subunits. The band densities of all 19 fractions (20 for CCT1) were summed and each
fraction was divided by the total to calculate a percentage of the total density in the gradient in
each fraction. Likelihood of complex formation was calculated by adding the values in the 20S
region (fractions 9-12 for CCT2-8, fractions 11-14 for CCT1), subtracting all of the values for
each CCT subunit alone, and then normalizing within Mm-Cpn, CCT4, or CCT5 categories. All
graphs and the nuanced heat map were created in Microsoft Excel.
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Table 8-2: Antibodies against the CCT subunits
Subunit
SCBa Accession #
Clone
Host
Clonality
Epitopeb
1
(α) Alpha
sc-53454
91A
Rat
Monoclonal
C-term half
2
(β) Beta
sc-28556
H-80
Rabbit
Polyclonal
C-term (454-535)
3
(γ) Gamma
sc-33145
H-300
Rabbit
Polyclonal
Internal (101-400)
4
(δ) Delta
sc-137092
H-1
Mouse
Monoclonal
Internal (176-400)
5
(ε) Epsilon
sc-13886
C-15
Goat
Polyclonal
C-term
6
(ζ) Zeta
sc-100958
G-06
Mouse
Monoclonal
Non-specific
7
(η) Eta
sc-13887
N-18
Goat
Polyclonal
N-term
8
(θ) Theta
sc-13891
N-18
Goat
Polyclonal
N-term
a
SCB = Santa Cruz Biotechnology
b
term = terminus; numbers denote amino acids
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Results
In order to investigate how TRiC is assembled, we utilized the properties of CCT4,
CCT5, and the archaeal Mm-Cpn to assemble homo-oligomers in E. coli (Sergeeva et al. 2013).
In order to do this, the pETDuet plasmid, which contains two multiple cloning sites, was used. In
one multiple cloning site, CCT4, CCT5, Mm-Cpn, or no sequence, was inserted. In the other
site, we inserted CCT1-8, one at a time. Therefore, we had a repository of 32 plasmids to study
pair-wise chaperonin-subunit interactions. The plasmids were transformed one at a time into
BL21 (DE3) RIL cells, grown and expressed in 1 L cultures, harvested, and lysed via French
Press (Sergeeva et al. 2013). The debris was spun down, and 100 µL of the supernatant was
applied to 5-40% isokinetic sucrose gradients and centrifuged in a swinging bucket rotor.
Gradients were fractionated and the fractions were run on SDS-PAGE, transferred to
membranes, and probed with the appropriate CCT subunit corresponding to CCT1-8.
All CCT subunit had a species corresponding to the approximately 60-kDa full-length
CCT species, and all CCT subunits but CCT6 had various shorter and longer fragments
(Figures 8-1 and 8-2). These fragments included mid-length fragments, such as the CCT4
fragment of 53 kDa that has been previously observed and preliminarily characterized
(Sergeeva et al. 2013). Other mid-length fragments included 42-kDa species for CCT1 and
CCT3, 40-kDa species for CCT5, and a 36-kDa species for CCT2. All CCT subunits with
fragments also had smaller-sized fragments of approximately 35 kDa and/or 27-30 kDa. Some
CCT subunits had both 35 kDa and 27 kDa fragments, but only the strongest one was included
for further analysis due to their similarity. Interestingly, CCT5-8 had larger fragments of
approximately 75 kDa. Due to the fact that all of these fragments were identified by immunoblot,
these 75-kDa species reacted with CCT antibodies, suggesting that there may have been some
transcriptional disregulation to cause larger proteins to be translated.
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Figure 8-1: Immunoblot SDS-PAGE of sucrose gradient ultracentrifugation of CCT1-CCT4
Lysed cells expressing pETDuet plasmids with each CCT subunit alone, with Mm-Cpn, with
CCT4, or with CCT5 were applied to sucrose gradients and then fractionated. The fractions
were run on 10% SDS-PAGE, transferred, probed with the corresponding CCT antibody, and
shown here (top, 5%, to 40%, and one bottom fraction, B). Full-length CCT proteins are
indicated with solid arrows and various fragments are labeled and indicated with dashed arrows.
Fractions are separated by dotted vertical lines for easier visual comparison and the fractions
corresponding to approximately the 20S region of the gradient are outlined in solid vertical lines
and labeled below each set of gels.
193
Figure 8-2: Immunoblot SDS-PAGE of sucrose gradient ultracentrifugation of CCT5-CCT8
Lysed cells expressing pETDuet plasmids with each CCT subunit alone, with Mm-Cpn, with
CCT4, or with CCT5 were applied to sucrose gradients and then fractionated. The fractions
were run on 10% SDS-PAGE, transferred, probed with the corresponding CCT antibody, and
shown here (top, 5%, to 40%, and one bottom fraction, B). Full-length CCT proteins are
indicated with solid arrows and various fragments are labeled and indicated with dashed arrows.
Fractions are separated by dotted vertical lines for easier visual comparison and the fractions
corresponding to approximately the 20S region of the gradient are outlined in solid vertical lines
and labeled below each set of gels.
194
Summary of Each CCT Profile
Each CCT subunit expressed alone from the pETDuet plasmid had a similar
sedimentation pattern as that subunit expressed from the pET21a plasmid previously (Sergeeva
et al. 2013). For CCT1, the full-length chains accumulated as a slowly sedimenting species and
species sedimenting faster than 20S. These chains may be aggregated as inclusion bodies or
associated with ribosome subunits, present in this region of the gradient. In the presence of
Mm-Cpn, the soluble CCT1 full-length subunits sedimented somewhat slower than 20S, at
approximately 14S. We suspect that the CCT1 chains may have been chaperoned in the cell by
Mm-Cpn. The co-expression of CCT4 with CCT1 drew most CCT1 chains into very soluble
species (monomers or dimers). When CCT5 was co-expressed with CCT1, although the
expression of full-length CCT1 was lower than full-length CCT1 expressed alone, most of the
full-length CCT1 was in the 20S region where CCT5 sediments as a homo-oligomer. This may
mean that CCT1 and CCT5 are co-assembling in the cell or that CCT5 is chaperoning CCT1
chains inside its cavity. The two fragments of CCT1 (30 and 42 kDa) sedimented in the 20S
region where Mm-Cpn sediments. These chains are most likely associated with the Mm-Cpn
rings or perhaps incorporated into them. Both fragments were also affected by the presence of
CCT4, sedimenting as soluble subunits in that 14S region, showing that there is some
interaction between CCT4 and the fragments of CCT1. Co-expression of CCT5 affected only the
30-kDa fragment, driving it into very soluble species.
For CCT2, the expression of full-length chains alone was low, even when co-expressed
with CCT4 and CCT5; but, when co-expressed with Mm-Cpn, the yield of full-length chains
increased, especially at 20S. This suggests that Mm-Cpn may be chaperoning the folding of
CCT2 subunits, and that these remain associated with the ring complex, or are incorporated.
CCT5 co-expression also pushed some of the full-length chains into sedimentation at 20S,
implying that CCT5 may co-assemble or chaperone CCT2. The major CCT2 fragment
expressed (30 kDa) sedimented slowly as soluble subunits, alone, or in the presence of CCT4
and CCT5. With Mm-Cpn, it sedimented in the 20S region, once again suggesting Mm-Cpn may
be chaperoning or binding this CCT fragment. The minor CCT2 fragment (36 kDa) sedimented
further down the gradient than 20S alone, or when co-expressed with any of the homo-oligomer
subunits, showing that it was not affected by chaperonin co-expression.
CCT3 expression was robust, as the full-length chains were the major species
accumulating. The majority of the full-length species was slowly sedimenting soluble subunit
forms, as well as chains sedimenting faster than 20S. The 20S species were most prominent
when co-expressed with Mm-Cpn and CCT5, suggesting these homo-oligomers associate,
195
chaperone, or co-assemble with CCT3. One of the minor fragments of CCT3 (42 kDa) was
predominantly found in the 20S region when CCT3 was co-expressed with Mm-Cpn. This CCT3
fragment was also found as more soluble species in the presence of CCT5. This suggests a
chaperoning role for this fragment by both Mm-Cpn and CCT5. The other fragment (27 kDa),
sedimenting as soluble (monomers or dimers) and faster than 20S species, showed no changes
with any of the co-expressed homo-oligomer subunits.
Since CCT4 and CCT5 already have a dominant presence at 20S as homo-oligomeric
rings, it was more difficult to interpret the changes in sedimentation patterns when the other
homo-oligomer subunits were co-expressed. We did find that there were more full-length 20S
species when the dose of CCT4 or CCT5 was doubled, by expressing each from both MCSI and
MCSII of the plasmid, which was encouraging. One interesting outcome was that co-expression
of Mm-Cpn disrupted full-length CCT4 20S species and instead full-length CCT4 populated a
species of more soluble subunits. Co-expression of Mm-Cpn brought CCT4 53-kDa fragment
into the 20S species and also increased the proportion of soluble subunits sedimenting at 14S,
therefore probably chaperoning these chains. Other than that interaction, homo-oligomer coexpression did not affect the other CCT4 fragments (35 kDa, 75 kDa, or 53 kDa with CCT4 or
CCT5), which were primarily found as soluble species or species sedimenting faster than 20S.
For CCT5, the majority of the fragments (27 kDa, 40 kDa, or 75 kDa) were once again
unaffected by co-expression of the other homo-oligomer subunits, but Mm-Cpn co-expression
did drive some of the 27-kDa fragments into 20S subunits, effectively chaperoning them.
The CCT6 full-length chains formed broad distributions in all four lysates, indicative of
poorly folded, misfolded or aggregating chains. There was little evidence of preferential
incorporation into rings in the presence of the CCT4 or CCT5 homo-oligomeric rings. Mm-Cpn
co-expression did drive some of the CCT6 full-length subunits into 20S ring or soluble subunit
(14S) species. CCT4 co-expression seemed to increase the full-length CCT6 chains that were
very soluble (monomers or dimers), as seen for co-expression of CCT4 with both CCT1 and
CCT2. CCT6 was the only species without detectable fragments by immunoblot.
CCT7 full-length chains accumulated as both soluble subunits and faster sedimenting
species when expressed alone. Their expression level was robust as they were the major
species accumulating in the lysates. In the presence of CCT5, the full-length chains sedimenting
at 20S increased, indicating complex formation or hetero-oligomer assembly. Co-expression
with Mm-Cpn seemed to drive some of the CCT7 full-length subunits into 20S species, but most
were found in more soluble species at 14S. Both fragments of CCT7 (35 and 75 kDa) were
196
unaffected by the presence of the homo-oligomer subunits, primarily sedimenting faster than
20S.
CCT8 full-length chains alone accumulated predominantly as faster sedimenting forms
with some soluble subunits. However, chains sedimenting at 20S were increased in the lysates
co-expressing CCT4 and CCT5 rings. Mm-Cpn co-expression pushed the CCT8 full-length
subunits into more soluble species of 14S, suggesting that it was chaperoning these chains.
The fragments of CCT8 (35 and 75 kDa), generally sedimented faster than 20S, showed no
change overall in the presence of the homo-oligomer subunits, but CCT4 did increase CCT8 35kDa fragment presence in a more soluble (14S) species, showing potential chaperoning of this
fragment by CCT4.
197
Figure 8-3: Quantified densities of full-length CCT species for each set of sucrose
ultracentrifugation gradients
Full-length CCT species from Figures 8-1 and 8-2 were quantified using ImageJ. The band
density in each fraction was divided by the total density of all fractions in a given gradient and is
plotted here. Lines correspond to either each CCT subunit: alone, red; with Mm-Cpn, blue; with
CCT4, orange; or, with CCT5, green. Dashed vertical lines correspond to approximately 20S
complexes sedimenting in the gradient. Asterisks indicate areas where there was unique
enrichment in species and are colored in agreement with the lines.
198
Effect of Homo-oligomers on Full-length CCT Subunits and Their Fragments
To help visually interpret the complex sucrose gradient patterns, the full-length and
fragment CCT species were quantified to analyze the changes in sedimentation patterns
between the different conditions, and therefore the effect of co-expression with CCT4, CCT5, or
Mm-Cpn. Quantifications are shown as plots of fraction of band densities of full-length (Figure 83) and various fragment (Figure 8-4) bands.
In summary, for the full-length CCT species, many of the CCT subunits (CCT1-2; CCT78) alone show unique enrichment in a species sedimenting faster than 20S. This is a region of
the gradients where ribosomes sediment, so these species may either be specifically interacting
with the ribosome to help fold proteins in E. coli or may be stuck on the ribosome because these
subunits alone have trouble folding themselves. This faster sedimenting species is significantly
decreased or eliminated in the presence of CCT4, CCT5, or Mm-Cpn, suggesting that these
homo-oligomeric chaperonins may interact with these particular CCT subunits to drive them
away from the ribosomes.
One overarching pattern seen in the gradients was that when co-expressed with MmCpn, almost all the full-length CCT subunits (all but CCT2) are found in soluble species,
sedimenting slower than 20S (approximately 14S). These are most likely some kind of
oligomers (but not 16-mers), possibly between both the CCT subunit and Mm-Cpn. When coexpressed with CCT4, many full-length CCT subunits (CCT1-2, 6) are enriched in a very small
and soluble species. This position in the gradient corresponds to CCT monomers or dimers,
implying that CCT4 may drive these CCT subunits into dissociating into stable smaller species.
Overall, the fragments of the CCT subunits had fewer pattern changes when coexpressed with homo-oligomeric subunits (Figure 8-4). Particularly, the larger sized fragments
(75 kDa) showed no change at all and had very similar patterns when comparing the different
CCT subunits to each other, specifically CCT5, CCT7, and CCT8. For the small- and mid-sized
fragments, there were a few specific enrichments; most notably Mm-Cpn driving the 42-kDa
CCT1, 30-kDa CCT1, 30-kDa CCT2, 42-kDa CCT3, 53-kDa CCT4, and 27-kDa CCT5
fragments to 20S complexes. CCT5 enriched a few species (42-kDa CCT1 and 42-kDa CCT3)
in small monomer/dimer soluble subunits, as CCT4 had done for many full-length species.
CCT4, on the other hand encouraged several of the mid- to small-sized fragments (42-kDa
CCT1, 30-kDa CCT1, and 35-kDa CCT8) to become part of the 14S complex that we observed
for Mm-Cpn co-expression with the full-length species. Some of the observations for the fulllength species carry over to fragments, such as 42-kDa CCT1 having enrichment in the
199
ribosomal region of the gradient, and Mm-Cpn making the 53-kDa CCT4 fragment more soluble,
driving it to sediment at 14S.
Most interesting for the investigation of CCT interactions on the way to TRiC assembly is
formation of 20S complexes when the CCT subunits are co-expressed with homo-oligomeric
subunits (Figure 8-5; Table 8-3). In terms of that, CCT5 was most efficient at driving CCT
subunits into 20S sedimentation, showing enrichment for all CCT subunits but CCT6. Mm-Cpn
was second most effective, pushing CCT2-3, CCT6, and possibly CCT7 into 20S complexes.
CCT4 was least effective, only showing any 20S interaction with CCT8. We went on to further
quantify the interaction of the full-length CCT subunits with the homo-oligomeric subunits in the
20S region. Therefore, we calculated the likelihood of the homo-oligomers having an effect on
the CCT subunits. We did this by adding the densities for the 20S fractions, subtracting out the
control (CCT1-8 only), and normalizing within each homo-oligomeric subunit. CCT4 and CCT5
were exempt from this normalization because we have shown that they form rings on their own.
In general, the two heat maps have good agreement between each other.
200
Figure 8-4: Quantified densities of fragmented CCT species for each set of sucrose
ultracentrifugation gradients
Fragment CCT species from Figures 8-1 and 8-2 were quantified using ImageJ. The density in
each fraction was divided by the total density of all fractions in a given gradient and is plotted
here for mid length fragments (36-53 kDa), short fragments (27-35 kDa), and long fragments (75
kDa). Lines correspond to either each CCT subunit: alone, red; with Mm-Cpn, blue; with CCT4,
orange; or, with CCT5, green. Dashed vertical lines correspond to approximately 20S
complexes sedimenting in the gradient. Asterisks indicate areas where there was unique
enrichment in species and are colored in agreement with the lines.
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Figure 8-5: Heat maps of CCT subunit complex formation alone, with Mm-Cpn, CCT4, or CCT5
A. A binary heat map based on whether a 20S complex species is present (blue) or is not
present (red) qualitatively for each CCT subunit in the data in Figure 8-3. A 0 (red) or 1 (blue)
scale is shown on the right. B. A more nuanced heat map based on how much of a 20S
complex species is present for each CCT subunit in the data. A nuanced scale is shown on the
right, corresponding to the likelihood of a 20S species formed under the specific conditions in
the map. All CCT4 and CCT5 interactions are shown in blue (most likely formation) because
these form established 20S species. See Materials and Methods for calculations.
202
Table 8-3: Summary table of full-length CCT subunits co-expressed with homo-oligomers
Subunit Pair
Soluble subunits
20S Species
Faster Sedimenting
CCT1 alone
-
-
+
CCT1 + Mm-Cpn
+
-
-
CCT1 + CCT4
+
-
+
CCT1 + CCT5
+
-
+
CCT2 alone
+
-
+
CCT2 + Mm-Cpn
-
+
-
CCT2 + CCT4
+
-
-
CCT2 + CCT5
+
+
-
CCT3 alone
+
-
-
CCT3 + Mm-Cpn
+
+
-
CCT3 + CCT4
+
-
-
CCT3 + CCT5
+
+
-
CCT4 alone
+
+
-
CCT4 + Mm-Cpn
+
-
-
CCT4 + CCT4
+
+
-
CCT4 + CCT5
+
+
+
CCT5 alone
-
+
-
CCT5 + Mm-Cpn
+
+
-
CCT5 + CCT4
-
+
-
CCT5 + CCT5
-
+
-
CCT6 alone
-
-
+
CCT6 + Mm-Cpn
+
+
-
CCT6 + CCT4
+
-
+
CCT6 + CCT5
-
-
+
CCT7 alone
+
-
+
CCT7 + Mm-Cpn
+
-
-
CCT7 + CCT4
+
-
+
CCT7 + CCT5
+
+
-
CCT8 alone
-
-
+
CCT8 + Mm-Cpn
+
-
-
CCT8 + CCT4
-
+
-
CCT8 + CCT5
-
+
-
203
Discussion
The experiments herein are the first step in understanding specific interactions between
CCT subunits, possibly for formation of hetero-oligomeric TRiC. Our data shows that CCT5
interacts with most CCT subunits (all but CCT6) by driving them into 20S complexes. These
effects could be due to a) the CCT5 homo-oligomers performing an active chaperonin function
within E. coli for the other CCT subunits being expressed, b) the other CCT subunits being
incorporated into the CCT5 ring complexes, or c) a combination of both a and b. Interestingly,
CCT5 did not push any of the CCT fragments into 20S complexes, meaning that it wasn’t
actively interacting with or binding to any of the CCT fragments. This may indicate that we are
may be seeing some hetero-oligomeric interactions with the full-length CCT subunits. Some of
the subunit fragments (CCT1 and CCT3) were found to sediment as more soluble species,
showing that CCT5 did possibly display some transient chaperone function.
We were expecting the archaeal chaperonin Mm-Cpn to be better at interacting with the
CCT subunits, due to its evolutionary role, but it fell intermediate between CCT5 and CCT4 in
number of CCT subunits it drew into 20S complexes. We did see that Mm-Cpn was efficient at
chaperoning both full-length and fragment CCT subunits, due to the sedimentation of all the
CCT full-length subunits and many fragments (CCT1-5) in 20S complexes or more soluble 14S
species in the presence of Mm-Cpn. This chaperoning of both CCT full-length and fragments
chains suggests that Mm-Cpn can recognize the CCT subunits in E. coli and interact with them
– either in the chaperonin complex (20S) or transiently to make them more stable and soluble
subunits (14S).
CCT4, surprisingly, only interacted with full-length CCT8 to push it into 20S complexes.
It did interact with full-length CCT1, CCT2, and CCT6, so that these subunits sedimented as
very soluble species, and interacted with the fragments of CCT1 and CCT8 in a way that these
fragments were sedimenting as soluble 14S species. Although we see no interaction with the
fragments in the 20S complex sedimentation, it does seem that CCT4 is capable of chaperoning
some of the CCT subunits. However, as with CCT5, it only interacts with the full-length CCT
subunits to push them into the 20S complexes, implying that it may be incorporating these CCT
subunits into its ring complexes.
The most crucial next step is to learn whether these interactions are bona fide heterooligomeric interactions or whether the homo-oligomers are actually chaperoning the CCT
subunits they interact with. For now, we will assume that these interactions are true heterooligomeric interactions, due to the fact that they are very specific (especially for CCT5 that
doesn't chaperone any fragments) and we don’t just see CCT4 or CCT5 interacting with all of
204
the CCT subunits (as we see for Mm-Cpn), possibly suggesting a more probable chaperoning
effect.
If our assumption that these are hetero-oligomers holds, we can use our data to propose
some theories of how TRiC may assemble inside the cell. The homo-oligomer state of CCT4
and CCT5 is much more stable than any CCT subunit on its own. Therefore, we will start with
the assumption that CCT4 and/or CCT5 homo-oligomers are formed first. One possible model is
a sequential model where each CCT subunit gets added one at a time (Figure 8-6A). Since
CCT4 only interacts with CCT8, if we started with just CCT4, then CCT8 would be the first to be
add into the ring, then CCT5, and the subunits that interact with CCT5 from most to least
strongly: CCT2, CCT7, CCT1, and CCT3. Finally, CCT6 would be added because it did not
interact with CCT4 nor CCT5. Another model would have CCT4 and CCT5 each starting on
their own with one other subunit as hetero-oligomers, and then coming together, and having
each other subunit added on sequentially (Figure 8-6B). Finally, subunits may be added
sequentially to CCT5, until CCT4 and CCT8 are added in, and then CCT6 is added on last
(Figure 8-6C).
For these models we are assuming that a mature TRiC complex with each subunit
appearing once per ring is made every time TRiC is assembled. However, this has not yet been
definitively proven. It would be probable that in cells that need to fold one substrate more so
than any other, the subunit that best recognizes this substrate is preferred over other subunits.
This may make it so that there are rings that hypothetically have two CCT4s or two CCT1s, and
no CCT8s or CCT3s. By isolating TRiC from more sources and tissues, we will be able to better
understand not only the subunit assembly but also the subunit arrangement. Methods such as
native mass spectrometry would be best to address these questions, once this type of
endogenous TRiC material is successfully isolated.
205
Figure 8-6: Possible models for TRiC formation assuming assembly is started from CCT4 or
CCT5 homo-oligomers
A. A sequential model for TRiC formation starting with CCT4. B. A model of TRiC formation
starting with both CCT4 and CCT5, then sequential assembly. C. A model for TRiC assembly
ending with CCT6 added last and CCT4 just interacting with CCT8. All models are based on
data from the heat map in Figure 8-5B.
206
CHAPTER 8:
APPENDIX B:
Aggregation Suppression of Mutant Huntingtin
by Chaperonins
207
Abstract
Huntingtin is a scaffolding protein in the brain that in its pathological form is responsible
for Huntington’s Disease. Suppression of mutant huntingtin by TRiC or CCT subunits has been
previously studied. Research showed that CCT1 and CCT4 were the optimal subunits for
huntingtin aggregation suppression. In order to attempt to treat huntingtin by modulating the
huntingtin-CCT interaction, more needs to be known about how exactly the CCT subunit
suppress this protein. We assayed huntingtin aggregation suppression by full human TRiC,
homo-oligomeric CCT4 and CCT5, and the archaeal chaperonin Mm-Cpn. While human TRiC
and CCT5 significantly decreased aggregation of huntingtin, CCT4 and Mm-Cpn (even at much
higher concentrations) failed to significantly decrease huntingtin. Therefore, the interaction
between huntingtin and the CCT subunits is very specific to the eukaryotic chaperonin, and
more specifically CCT5 rather than CCT4. This also is the first evidence of the CCT subunits
being specific for a substrate, with CCT5 showing more efficiency than CCT4.
208
Introduction
Huntington’s disease is an autosomal dominant gain of function disease with genetic
anticipation (Bates 2005). It affects about 5-7 of every 100,000 people (Bates 2005; Walker
2007). The symptoms of Huntington’s disease include physical traits such as chorea and
psychological traits such as apathy, irritability, anxiety and dysphoria (Walker 2007). The
disease primarily affects the striatum inside the basal ganglia and the cerebral cortex (Walker
2007). Huntington’s disease is caused by a pathological version of the protein huntingtin, in
which a section of the protein has an increased number of CAG repeats (Bates 2005; Walker
2007). Wild-type huntingtin protein has between 10-35 CAG repeats, and therefore
polyglutamines, whereas mutant huntingtin (mHtt) has at least 40 polyglutamines (Bates 2005).
The longer the stretch of polyglutamines in the mHtt protein, the earlier onset of symptoms
occurs (Walker 2007; Wetzel 2012), consistent with diseases displaying genetic anticipation.
The progression of the disease to death after onset is about 15-20 years (Bates 2005; Walker
2007).
Huntingtin is a very large, 3144 amino acid (348 kDa) soluble cytoplasmic protein.
Although it is ubiquitously expressed, it is found at high levels in the central nervous system and
the testes (Wetzel 2012). Wild-type huntingtin has various functions in cells such as acting as a
scaffold protein, and playing a role in neuronal gene transcription, and axonal and vesicular
transport (Bates 2005). Aggregates of mHtt have been found in patient brains, consistent with
the idea that aggregation of the pathological protein is part of the disease (Arrasate and
Finkbeiner 2012; Clabough 2013). These aggregates contain fragments of the mHtt protein, the
shortest of which includes only the first exon of huntingtin wherein the polyQ region is located
(Wetzel 2012). These aggregates can be part of inclusion bodies or amyloidogenic fibrils of the
mHtt protein (Wetzel 2012). It is unknown, however, whether the aggregation is a mechanism of
disease suppression or part of the progression of the disease, due to the fact that there is still
uncertainty surrounding the question of what the toxic species of this disease is. Multiple reports
have implicated soluble oligomers of mHtt as the toxic species (Margulis et al. 2013; van der
Putten and Lotz 2013). Even if aggregates are the cell’s way of suppressing the deleterious
effects of mHtt, these inclusion bodies contain other important cellular proteins, therefore
resulting in transcription disregulation of the cell (Arrasate and Finkbeiner 2012; Clabough
2013). Being able to target these aggregates by molecular chaperones would give an
opportunity to ameliorate the cell’s burden (Arrasate and Finkbeiner 2012; Margulis et al. 2013;
van der Putten and Lotz 2013).
209
TRiC has been shown to decrease mHtt aggregation in vitro, in yeast cells, and in cell
culture. When an exon 1 construct of mHtt was induced to aggregate, a 1:1 ratio of TRiC to
mHtt completely suppressed its aggregation in vitro (Tam et al. 2006). Additionally, each CCT
subunit was co-expressed with a construct of exon 1 of mHtt, and CCT1 and CCT4 significantly
diffused huntingtin aggregates (Tam et al. 2006). This was consistent with CCT1 and CCT4
recognizing mHtt specifically and actively disaggregating the huntingtin aggregates. By using a
UV-inducible cross linker, it was found that TRiC binds directly to and sequesters the N-terminal
seventeen residues immediately preceding the polyglutamine region of mHtt (Tam et al. 2009).
Structural research using cryo-EM has also shown that TRiC suppress mHtt fibrils and binds to
mHtt oligomers (Shahmoradian et al. 2013). More recently, a construct with just the apical
domain of CCT1 was shown to decrease huntingtin aggregation when applied exogenously to
PC12 cells (Sontag et al. 2013). This construct also decreased toxicity of striatal cells derived
from Huntington’s Disease model mice, showing efficacy in treating Huntington’s Disease by a
TRiC-like construct (Sontag et al. 2013).
Based on this research, TRiC is an ideal target for treatment of huntington’s disease.
Although increasing TRiC expression and function may be detrimental to the cell, finding a way
to selectively increase interaction between TRiC and mHtt would be promising. The most direct
method would be through a small molecule that directly promoted the interaction between TRiC
and mHtt. Knowledge of the structure of the TRiC/mHtt interaction would be crucial in
understanding what kind of small molecule would need to be designed.
210
Materials and Methods
Mutant Huntingtin Aggregation Suppression Assay
mHtt aggregation suppression assay was modified from Tam et al. (2006). Briefly, GST-,
His-, and S-tagged exon 1 of Htt with 53 poly glutamines, and containing a TEV protease
cleavage site between the GST-tag and the rest of construct, was purified using a Co-NTA
column, followed by a glutathione agarose column (Pierce). To initiate an aggregation
suppression reaction, 5 µM of the mHtt protein in a buffer (20 mM Tris, 50 mM KCl, 5 mM
MgCl2, 5 mM DTT, and 1 mM ATP) containing various concentrations of chaperonin was
cleaved with 0.1 mM TEV protease. This reaction was left at 30 °C for 16 hours. The reaction
was stopped by equal volume addition of 4% SDS, boiled for 10 minutes, and filtered through
0.22 µm cellulose acetate membrane (GE Healthcare). The membrane was washed and
blocked using 5% milk in TBS. An AP-conjugated antibody against the S-tag (EMD Millipore)
was used to detect amount of mHtt trapped in the membrane. Ovalbumin was used as a control
and concentration of CCT5 was calculated as in the HγD-Crys assay. Quantification was done
in ImageJ where suppression was calculated as decrease from the ovalbumin control.
211
Results & Discussion
In order to study whether the chaperonins suppressed mHtt aggregation in vitro, we
quantified the aggregation by using an exon 1 construct of mHtt (containing 51 polyglutamines)
in the presence of the chaperonins. CCT5 significantly suppressed aggregation of huntingtin, as
did human TRiC (Knee et al. 2013), but not the archaeal chaperonin Mm-Cpn or CCT4 (Figure
8-7). This means that the interaction between huntintin and TRiC is specific for the eukaryotic
chaperonin and is more dependent on CCT5 than CCT4.
We can use structural studies such as cryo-EM to better assess the interaction between
mHtt and CCT5. Having a specific CCT subunit able to interact with and suppress mHtt can
allow us to target small molecules to this interface. Further, because CCT5 can suppress
aggregation much more efficiently than CCT4, this is the best example of a CCT subunit specific
for a substrate. This gives credibility to the theory that there are 8 CCT subunits so that they can
each recognize and bind different classes of substrates (Kim et al. 1994). More substrates (such
as actin, tubulin, and pVHL) will have to be tested to further substantiate this theory.
212
1.25
Quantified Aggregation of Htt
hTRiC
CCT4
CCT5
MmCpn
1.00
*
0.75
*
*
*
*
0.50
+
0.25
0
0. x
5x
1x
0
0. x
5x
1x
2x
0
0. x
5x
1x
2x
3
15 x
30x
x
0.00
Concentration of Chaperonin
Figure 8-7: CCT5 and human TRiC suppress aggregation of mutant huntingtin while CCT4 and
Mm-Cpn do not
Concentration of human TRiC, CCT4 and CCT5 homo-oligomer, and the archaeal chaperonin
MmCpn calculated so that 1x would be one 60 kDa chaperonin subunit per one 20 kDa
huntingtin construct. TRiC and CCT5 homo-oligomer suppressed huntingtin aggregation
significantly (* is p < 0.05; + is p < 0.01) while CCT4 and Mm-Cpn did not significantly suppress
huntingtin aggregation even with Mm-Cpn at a 30x concentration of chaperonin as compared to
mHtt. 0x refers to the ovalbumin control. Error bars shown are SEM of 2-5 repeats.
213
214