Assembly and Substrate Recognition Properties of Human CCT Subunits of the Tric Chaperonin

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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

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

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.

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, Mm- Cpn 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

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 cryo- EM 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.

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.

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

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. 307:297- 300.

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

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

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

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

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 native- like 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 HγD- Crys 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

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

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

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

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

CHAPTER 1:

Introduction

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 ATP- dependent chaperones are actually able to bind to partially unfolded or misfolded substrates and help them in folding to more native-like states.

Unfolded State

Chaperones

Partially Oligomers Energy Folded States 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).

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 C- terminal 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

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).

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.

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 C- terminal 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 co- chaperones 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).

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.

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 9- 15% 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.

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 non- suppressing 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

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 groP- like), 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

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

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 stress- induced 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

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).

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).

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 T- locus 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 T- locus 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 35S- methionine in vitro, the protein fraction was isolated, and the proteins were separated using two- dimensional 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

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, 35S-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

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 TRiC- assisted 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 actin- interacting 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

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 1- 5C, 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).

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).

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.

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.

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)

Features of the Substrate Recognized GroEL has been studied extensively for its substrate recognition properties (Gómez- Puertas 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 β- propeller proteins, specifically the hydrophobic third β-strand of the second WD40 (40 amino- acid 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 β- sheet rich and fold slowly, making them ideal group II chaperonin substrates (Gromiha and Selvaraj 2004; Smith 2008).

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

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).

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).

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 Sprague- Dawley 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 McKusick- Kaufman 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

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 disease- causing substrate, while not significantly sacrificing TRiC function with the rest of its substrates would be ideal.

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)

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 hetero- oligomeric 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 TRiC- like 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.

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.

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.

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

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.

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.

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, sc- 100958; 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.

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.

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).

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.

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% SDS- PAGE. 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.

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.

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.

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.

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 HγD- Crys, 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.

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.

Figure 2-7: Purified human TRiC suppression of HγD-Crys aggregation and HγD-Crys native- like 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 HγD- Crys to a native-like state.

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 co- purifies 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

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.

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.

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 γD- crystallin 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.

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-binding- deficient 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).

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 co- expression 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.

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

(Cambridge, MA). Peptides were identified by searching for hits in the Mascot database. N- terminal 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.

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 semi- automatically 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 low- resolution 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 sub- datasets, 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

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 &

King (2010) and Knee et al. (2011) and was modified in this study by use of a decreased chaperonin concentration of 145 nM.

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% SDS- PAGE 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.

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.

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.

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).

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.

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.

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.

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.

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.

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 homo- oligomers. 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.

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 3- 8).

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 wild- type nature of human TRiC. Adding ATP slightly decreased the denaturation midpoint of the chaperonins, primarily due to the increase in cooperativity.

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.

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).

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) homo- oligomers 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.

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 partially- folded 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 homo- oligomer 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).

Figure 3-10: CCT4 and CCT5 homo-oligomers suppressed aggregation of partially folded HγD- Crys 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.

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 homo- oligomeric 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 cryo- EM 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 homo- oligomers 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).

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 homo- oligomers. 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.

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.

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.

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

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 CCT1- 8) 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 Sprague- Dawley 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

aHSPD1: 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 anti- T7 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.

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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 solid- lined 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.

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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.

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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 4- 10B). 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.

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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 HγD- Crys 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.

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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 homo- oligomeric 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-

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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 homo- oligomer 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

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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 well- characterized 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 HγD- Crys 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

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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).

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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 HγD-

Crys 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 -1 -1 was determined by A280 using an extinction coefficient of 42,860 M cm . 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.

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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.

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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

1published in Flaugh, et al. 2006 2published in Moreau & King 2009 3published in Kong & King 2011

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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).

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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

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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% SDS- PAGE. 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 HγD- Crys: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 HγD- Crys 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).

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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.

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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).

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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 HγD- Crys 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.

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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 Mm- Cpn/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 SDS- PAGE 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.

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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: 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.

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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.

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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 HγD- Crys mutants to levels higher than WT.

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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 Mm- Cpn-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 Mm- Cpn 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

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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.

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Final Discussion and Future Directions

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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 Hsp70- TRiC 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

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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, homo- oligomers 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 HγD- Crys, 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 HγD- Crys, 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

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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

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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 β- sheet 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 substrate- binding domain recognizes stretches of five hydrophobic amino acids, flanked by positively- charged amino acids, normally buried in the native state (Mayer 2013). Small heat shock

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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; Martínez- Alonso 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 high- quantity 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.

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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 post- translational 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.

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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 hetero- oligomerizing 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.

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CHAPTER 7:

References

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CHAPTER 8: APPENDIX A:

Co-expression of CCT Subunits to Explore Subunit Assembly*

* Cameron Haase-Pettingell is acknowledged for her technical assistance.

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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

aAll information from National Center for Biotechnology Information

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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 homo- oligomeric 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

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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 assembly- competent forms.

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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 6x- Histag, 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

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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

aSCB = Santa Cruz Biotechnology bterm = 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.

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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.

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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,

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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 co- expression 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

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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 35- kDa fragment presence in a more soluble (14S) species, showing potential chaperoning of this fragment by CCT4.

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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.

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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 8- 3) and various fragment (Figure 8-4) bands. In summary, for the full-length CCT species, many of the CCT subunits (CCT1-2; CCT7- 8) 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 Mm- Cpn, 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 co- expressed 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 co- expressed 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 full- length species carry over to fragments, such as 42-kDa CCT1 having enrichment in the

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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.

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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.

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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 - + -

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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 hetero- oligomeric 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 hetero- oligomeric 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

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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.

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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.

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CHAPTER 8: APPENDIX B:

Aggregation Suppression of Mutant Huntingtin by Chaperonins

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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.

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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).

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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.

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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

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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.

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1.25 hTRiC CCT4 CCT5 MmCpn

1.00 * * 0.75 * * * 0.50

0.25 + Quantified Aggregation of Htt Aggregation Quantified

0.00 0x 1x 0x 1x 2x 0x 1x 2x 3x 0.5x 0.5x 0.5x 15x30x 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.

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