Regulation of Human 69-Kda Choline Acetyltransferase Protein Stability and Function by Molecular Chaperones and the Ubiquitin- Proteasome System

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Regulation of Human 69-kDa Choline Acetyltransferase Protein Stability and Function by Molecular Chaperones and the Ubiquitin- Proteasome System

Trevor M. Morey The University of Western Ontario

Supervisor Rylett, R. Jane The University of Western Ontario

Graduate Program in Physiology A thesis submitted in partial fulfillment of the equirr ements for the degree in Doctor of Philosophy © Trevor M. Morey 2017

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Recommended Citation Morey, Trevor M., "Regulation of Human 69-kDa Choline Acetyltransferase Protein Stability and Function by Molecular Chaperones and the Ubiquitin-Proteasome System" (2017). Electronic Thesis and Dissertation Repository. 5218. https://ir.lib.uwo.ca/etd/5218

This Dissertation/Thesis is brought to you for free and open access by Scholarship@Western. It has been accepted for inclusion in Electronic Thesis and Dissertation Repository by an authorized administrator of Scholarship@Western. For more information, please contact [email protected]. ABSTRACT:

The enzyme choline acetyltransferase (ChAT) mediates synthesis of the neurotransmitter acetylcholine required for cholinergic neurotransmission. ChAT mutations are linked to congenital myasthenic syndrome (CMS), a rare neuromuscular disorder. One CMS-related mutation, V18M, reduces ChAT enzyme activity and cellular protein levels, and is located within a highly-conserved N-terminal proline-rich motif at residues 14PKLPVPP20. It is currently unknown if this motif regulates ChAT function. In this thesis, I demonstrate that disruption of this proline-rich motif in mouse cholinergic SN56 cells reduces both the protein levels and cellular enzymatic activity of mutated P17A/P19A- and V18M-ChAT. The cellular loss of mutant ChAT protein appears to be a result of increased proteasome- dependent degradation due to enhanced ChAT ubiquitination. Using a novel fluorescent- biorthogonal pulse-chase protocol, I determined that the cellular protein half-life of P17A/P19A-ChAT (2.2 h) is substantially reduced compared to wild-type ChAT (19.7 h), and that proteasome inhibition by MG132 treatment increases the half-life and steady-state levels of ChAT protein. By proximity-dependent biotin identification (BioID), co- immunoprecipitation, and in situ proximity-ligation assay (PLA), I identified the heat shock proteins HSC/HSP70 and HSP90 as novel ChAT protein-interactors that are enriched in cells expressing mutant P17A/P19A-ChAT. Pharmacological inhibition of these HSPs by treatment with the HSC/HSP70 inhibitors 2-phenylethynesulfonamide (PES) or VER- 155008, or the HSP90 inhibitor 17-AAG reduced cellular ChAT activity and solubility, and enhanced ubiquitination and proteasomal loss of ChAT protein. The effects of HSP inhibition were greatest for mutant P17A/P19A- and V18M-ChAT. While I observed that ChAT interacts in situ with the HSP-associated E3 ubiquitin ligase CHIP, siRNA-mediated knock-down of CHIP had no effect on ChAT protein levels. Inhibition of HSC/HSP70 by

PES treatment sensitized ChAT to H2O2-induced insolubilization, and ChAT ubiquitination was enhanced following H2O2 treatment. Lastly, inhibition of the endoplasmic reticulum (ER)- and HSP-associated co-chaperone Cdc48/p97/Valosin-containing protein (VCP) prevented the degradation of ubiquitinated ChAT. Together my results identify novel mechanisms for the functional regulation of wild-type and CMS-related mutant ChAT by

multiple molecular chaperones and the ubiquitin-proteasome system that, importantly, may have broader implications for ChAT function during cellular stress and disease.

KEYWORDS:

Choline acetyltransferase (ChAT), molecular chaperones, protein degradation, protein quality control, proteasome, ubiquitination, click chemistry, BioID, proximity-ligation assay (PLA), HSC/HSP70, HSP90, CHIP/Stub1, Cdc48/p97/VCP, oxidative stress.

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CO-AUTHORSHIP STATEMENT

The studies detailed within this thesis were completed by Trevor M. Morey in the laboratory of Dr. R. Jane Rylett with the following co-author assistance:

1. Shawn Albers, M.Sc., assisted with cellular ChAT activity assays.

2. Dr. Brian Shilton, Department of Biochemistry (UWO), provided resources for, assisted with, and provided expertise for studies utilizing bacterially-expressed and purified recombinant ChAT protein.

3. Claudia Seah (Dr. Pasternak, UWO), assisted with in situ PLA experiments by performing confocal microscopy.

4. Dr. Warren Winick-Ng assisted with in situ PLA experiments by performing confocal microscopy and assisting with PLA optimization.

Initial experiments related to the effects of H2O2-induced oxidative stress on ChAT protein function were designed, completed, and analyzed previously by Shawn Albers, M.Sc.1

All other experiments were designed, and data was analyzed and interpreted by Trevor M. Morey and Dr. R. Jane Rylett. Experiments were performed by Trevor M. Morey in collaboration with the above co-authors. All authors contributed to the preparation of manuscripts for publication. This thesis was written by Trevor M. Morey.

1Albers, Shawn. “The role of choline acetyltransferase variants in Alzheimer’s disease models” (2013).

Electronic Thesis and Dissertation Repository. 1844. http://ir.lib.uwo.ca/etd/1844.

PUBLICATION STATEMENT

This thesis contains text, data, and figures that have been published in the followed peer- reviewed first-author publications:

1. Morey, T.M., Winick-Ng, W., Seah, C., and Rylett, R.J. (2017). Chaperone-mediated regulation of choline acetyltransferase protein stability and activity by HSC/HSP70, HSP90 and p97/VCP. Front Mol Neurosci. 10:415. doi: 10.3389/fnmol.2017.00415.

2. Morey, T.M., Albers, S., Shilton, B.H., and Rylett, R.J. (2016). Enhanced ubiquitination and proteasomal-degradation of catalytically-deficient human choline acetyltransferase mutants. J. Neurochem. 137, 630-646.

ACKNOWLEDGEMENTS

I am extremely grateful for the opportunity to complete this Ph.D thesis under the supervision of Dr. R. Jane Rylett. Her continued support, expertise, insight, and guidance throughout my studies provided me with the opportunity to learn, develop critical research skills, and most importantly to be successful during my time in her laboratory. I wish Jane all the best in her future research and look forward to future collaborations.

I would also like to thank my advisory committee members, Drs. Lina Dagnino, John Di Guglielmo, Susan Meakin, and Brian Shilton for providing invaluable critique, direction, and suggestions throughout my degree. Special thanks are extended to Dr. John Di Guglielmo for his generous read-through of my preliminary thesis and for providing his thoughts and expertise. To Drs. Brian Shilton and Martin Duennwald, thank you for providing me with your scientific and technical expertise that ultimately helped me design, implement, and interpret certain experiments completed within this thesis.

To Dr. Warren Winick-Ng, my colleague and friend, thank you for the many stimulating conversations, both research and non-research related, as well as for the laughs and good times while at work. I wish you all the best and success on your post-doctoral adventures in Berlin and beyond. In addition, I thank all my peers, friends, and colleagues for fostering a joyful, successful, and memorable learning and working experience.

To my parents Debi McKellar and Gary Morey - thank you for always being there to lend an ear, for your love and guidance, and for your belief that anything is possible.

Lastly, to my wife Suze, thank you for joining me on this incredible journey and for always motivating me to better myself, to take chances, and to always give my 100% in all things. Without your unwavering love and support, your patience and understanding, and your faith in me, this study would not have been possible. I hope one day, in your future endeavours, to extend the same supports as that have given to me throughout my studies.

Funding for this study was provided to R.J.R. by the Canadian Institutes of Health Research. T.M.M is a two-time recipient of the Ontario Graduate Scholarship.

TABLE OF CONTENTS

Abstract…………………………………………………………….……………………...ii

Keywords…………………………………………………………………………………iii

Co-authorship Statement………………………………………………………………….iv

Publication Statement………………………………………………………………….…..v

Acknowledgements…………………………………………………………………….....vi

Table of Contents…………………………………………………………………………vii

List of Figures……………………………………………………………………….……xii

List of Abbreviations……………………………………………………………...…….xvii

Chapter 1: Introduction……………………………………………………….………...….1

1.1: Proteostasis and protein quality control mechanisms……………….…………..1

1.2: The ubiquitin-proteasome system…………………..……………….…………..3

1.2.1: The proteasome…………………………………………..…….……….3

1.2.2: Ubiquitin…………………………….………………………………….3

1.2.3: Ubiquitination mechanisms……………………………………….……5

1.2.4: The ubiquitin code………………………………………..…………….8

1.2.5: Autophagy-lysosomal degradation………………………...………….10

1.3: Protein folding and molecular chaperones…………………….………….……11

1.3.1: Molecular chaperones overview………………....……………………11

1.3.2: Heat shock proteins and the heat shock response…………..…………..13

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1.3.3: The HSP chaperone network……………………………….…...……..14

1.3.4: HSC/HSP70………………………………………….………………..17

1.3.5: HSP90…………………………………………………………………19

1.3.6: Ubiquitin E3 ligase CHIP……………………………….………..……23

1.4: Molecular chaperones in human disease…………………………..….………..25

1.4.1: HSPs in Cancer………………………………………...……………...25

1.4.2: HSPs in aging and neurodegeneration…………..…….……………….26

1.4.3: HSPs and oxidative stress……………………………....……………..27

1.5: Cholinergic neurobiology……………………………………….……………..28

1.5.1: The cholinergic synapse……………………………….………………28

1.5.2: Choline acetyltransferase (ChAT)…………………………...………..31

1.5.3: ChAT in neurodegenerative diseases…………...……….…………….33

1.5.4: Congenial myasthenic syndrome (CMS) …………………..………….34

1.5.5: V18M-ChAT and the ChAT N-terminus…………..………..…………35

1.6: Study rationale, aims, and hypothesis…………………..…………....………40

Chapter 2: Materials and Methods…………………………………………….….………42

2.1: Protein expression plasmids……………………………………….…………..42

2.2: Cell culture, experimental treatments, and cell lysis…………………….……..42

2.3: Immunoprecipitation (IP) to assess ChAT ubiquitination……….…..…………45

2.4: Co-immunoprecipitation (co-IP) of ChAT with CHIP or HSPs………..………45

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2.5: SDS-PAGE and immunoblotting analysis…………………….………….……46

2.6: Purification of recombinant ChAT……………………………….……………47

2.7: Circular dichroism (CD) analysis and thermal denaturation…………..……….47

2.8: Strain-promoted alkyne-azide cycloaddition (SPAAC) pulse-chase….….……48

2.9: Soluble/insoluble Triton X-100 cellular fractionation and semi- denaturing detergent agarose gel electrophoresis (SDD-AGE)…….……….…50

2.10: Proximity-dependent biotin identification (BioID)…………….…………….51

2.11: Mass spectrometry………………………………………….…….…………..52

2.12: In situ proximity ligation assay (PLA)…………………………..……………53

2.13: Small-interfering RNA (siRNA)…………………….……………………….54

2.14: Statistical analysis……………………………………………………………54

Chapter 3: Results………………………..………………………………………….……56

3.1: ChAT mutations reduce steady-state protein levels and cellular enzymatic activity of ChAT…………………………………………...………56

3.2: In vitro characterization of recombinant proline-mutant ChAT…………..……61

3.3: Proteasome inhibition increases ChAT steady-state protein levels and cellular ChAT enzyme activity………………………………….………...64

3.4: Mutation of the N-terminal proline-rich motif enhances ChAT ubiquitination……………………………………………………..……67

3.5: ChAT protein can be targeted for proteasomal degradation by both K48-dependent and K48-independent polyubiquitination……….…...…..75

3.6: ChAT protein half-life is reduced upon mutation of the N-terminal proline-rich motif and can be increased by proteasome inhibition……….....….78

3.7: Identification of the heat shock proteins HSP70 and HSP90 as putative ChAT protein interactors………………………………...…….……..84

3.8: Co-immunoprecipitation of ChAT with HSPs is altered by mutation of N-terminal proline-rich motif of ChAT………………………..….93

3.9: Inhibition of HSC/HSP70-substrate protein interactions promotes accumulation of high molecular mass Triton-insoluble ChAT aggregates…...101

3.10: Inhibition of HSC/HSP70 or HSP90 ATPase activity reduces ChAT steady-state protein levels…………………………………………………..104

3.11: Proteasome inhibition prevents loss of ChAT protein following HSC/HSP70 and HSP90 inhibition………………………………….……...108

3.12: Inhibition of HSC/HSP70 and HSP90 activity enhances ChAT ubiquitination and reduces ChAT enzymatic activity…………….……...…111

3.13: ChAT interacts with the HSP-associated E3 ubiquitin ligase CHIP…...... …..115

3.14: CHIP co-expression promotes proteasomal degradation and insolubilization of ChAT……………………………………………..…….115

3.15: Oxidative stress by exposure of cells to H2O2 enhances ChAT ubiquitination…………………………………….………………….122

3.16: Oxidative stress by H2O2 treatment promotes the insolubilization of ChAT protein…………………………………………………....……….126

3.17: Co-immunoprecipitation of ChAT with HSC70 is altered

following exposure of cells to H2O2……………………………….…….….129

3.18: Inhibition of the co-chaperone Cdc48/p97/VCP prevents the degradation of ubiquitinated ChAT……..……………………………...…..131

Chapter 4: Discussion…………………………………………………….……………..135

4.1: Significant Findings……………………………………………….…………135

4.2: General Discussion……………………………………………….…………..138

4.3: Study Limitations and Alternative Approaches……………………..……..…149

4.4: Future Directions………………………………………………………....…..155

4.5: Significance and Concluding Remarks…………….…………………………160

References……………………………………………..………………………………..162

Appendices …………………………………………………...……………..………….215

Appendix A: Optimization of BioID in multiple mammalian cell lines………..215

Appendix B: Optimization of siRNA-mediated knock-down of endogenous HSC/HSP70 and HSP90………………...….……….217

John Wiley and Sons (Journal of Neurochemistry)…………………...…………218

Frontiers (Frontiers in Molecular Neuroscience)………………………………..223

Curriculum Vitae………………………………………………………………………..226

LIST OF FIGURES

Figure 1.1: Protein quality control overview………………………………..……………..2

Figure 1.2: The ubiquitin-proteasome system……………………………………………..4

Figure 1.3: The ubiquitination pathway…………………………………….……………..6

Figure 1.4: The ubiquitin code……………………………………………………….…....9

Figure 1.5: Molecular chaperones regulate cellular proteostasis………………...………12

Figure 1.6: The HSP chaperone network………………………………………..……….16

Figure 1.7: The HSP90 chaperone cycle………………………………………..………..22

Figure 1.8: The cholinergic synapse…………………………………………………...…30

Figure 1.9: Missense mutations within human ChAT are associated with development of the neuromuscular disorder congenital myasthenic syndrome (CMS)………………………………………………………….....36

Figure 1.10: Comparative sequence alignment of the highly-conserved N-terminal proline-rich motif of ChAT from multiple divergent animal species…….….38

Figure 1.11: Three-dimensional crystal structure of human 69-kDa ChAT……………..39

Figure 3.1: CMS-related ChAT mutants demonstrate reduced steady-state protein levels………………………………………………………...………57

Figure 3.2: N-terminal truncation of ChAT reduces ChAT steady-state proteins levels and cellular ChAT activity……………………………………………59

Figure 3.3: Mutation of conserved N-terminal proline-rich motif of ChAT reduces ChAT steady-state proteins levels and cellular ChAT activity……………...60

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Figure 3.4: Purification of bacterially-expressed recombinant wild-type and proline-mutant ChAT……………………………………………………..…62

Figure 3.5: In vitro analysis of recombinant wild-type and proline-mutant ChAT……...63

Figure 3.6: Proteasome inhibition increases ChAT steady-state protein levels and cellular ChAT activity…………………………………………………..65

Figure 3.7: Proteasome inhibition partially restores the steady-state protein levels and cellular activity of CMS-related ChAT mutants….…………...…68

Figure 3.8: Proteasome inhibition reveals the presence of multiple higher molecular mass ChAT proteins……………………………..……………….69

Figure 3.9: ChAT ubiquitination is increased upon mutation of the N-terminal proline-rich motif and in cases of CMS-related mutations…………….……71

Figure 3.10: Anti-ChAT IP reveals enhanced ubiquitination of mutant P17A/P19A-ChAT………………………………………………………....73

Figure 3.11: Wild-type and P17A/P19A-ChAT can be targeted for proteasomal degradation by both K48-dependent and K48-independent polyubiquitination………………………………………………………….76

Figure 3.12: Schematic overview of biorthogonal strain-promoted alkyne-azide cycloaddition (SPAAC) pulse-chase for fluorescent determination of protein half-life………………………………………………………….79

Figure 3.13: SPAAC pulse-chase reveals that ChAT protein half-life is reduced upon mutation of the N-terminal proline-rich motif……………….………81

Figure 3.14: Validation of SPAAC pulse-chase by cycloheximide (CHX)-chase assay...... 82

Figure 3.15: Proteasome inhibition increase ChAT protein half-life………….…………83

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Figure 3.16: Model for the regulation of ChAT protein stability by the ubiquitin- proteasome system……………………………………………………..…..86

Figure 3.17: Proximity-dependent biotin identification (BioID) for the determination of novel ChAT protein-protein interactions……….………..87

Figure 3.18: Identification of the heat shock proteins HSP70 and HSP90 as novel ChAT protein-interactions by BioID………………………………..90

Figure 3.19: MALDI-TOF-MS data for HSP70 following BioID…………...…………..91

Figure 3.20: MALDI-TOF-MS data for ChAT-BirA* fusion protein following BioID.…………………………………………………………...92

Figure 3.21: LC-ESI-MS/MS data for HSP90 following BioID…………..……………..94

Figure 3.22: LC-ESI-MS/MS data for HSP70 following BioID…………..……………..95

Figure 3.23: Immunoblot verification of putative ChAT interacting proteins following BioID………………………………………………...………….96

Figure 3.24: Co-immunoprecipitation of ChAT with HSC/HSP70 and HSP90 from HEK293 cells.…………………………………………………....…..97

Figure 3.25: Co-immunoprecipitation of ChAT with HSC70 and HSP90 from mouse cholinergic SN56 cells…………………………………………..….99

Figure 3.26: In situ interaction of ChAT with endogenous HSC70 and HSP90……..…100

Figure 3.27: Inhibition of HSC/HSP70-substrate protein interaction promotes accumulation of high molecular mass Triton-insoluble ChAT aggregates……………………………………………………….....102

Figure 3.28: Inhibition of HSC/HSP70 ATPase activity reduces ChAT steady-state protein levels in a dose-related manner……………………...105

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Figure 3.29: Inhibition of HSP90 by 17-AAG specifically reduces steady-state protein levels of mutant ChAT………………………………………..…..107

Figure 3.30: siRNA-mediated knock-down of HSP70 fails to sensitize wild-type ChAT to 17-AAG-induced loss of ChAT protein………………………...109

Figure 3.31: Proteasome inhibition by MG132 treatment prevents loss of ChAT protein following inhibition of HSC/HSP70 or HSP90 activity………….110

Figure 3.32: Inhibition of HSC/HSP70 and HSP90 activity enhances ChAT ubiquitination…………………………………………………...…112

Figure 3.33: Inhibition of HSC/HSP70 and HSP90 activity reduces cellular enzymatic activity of wild-type ChAT………………………………...….114

Figure 3.34: Co-immunoprecipitation of ChAT with HSP-associated E3 ubiquitin ligase C-terminus of HSC70-interaction protein (CHIP)………116

Figure 3.35: In situ interaction of ChAT with endogenous CHIP…………………..…..117

Figure 3.36: CHIP co-expression promotes proteasomal degradation of ChAT…….....119

Figure 3.37: The TPR domains of CHIP are critical for ChAT-CHIP protein interaction and promote CHIP-mediated ChAT insolubilization……..….120

Figure 3.38: siRNA knock-down of endogenous CHIP has no effect on the steady-state protein levels of wild-type or mutant ChAT……...…………123

Figure 3.39: Model for the regulation of ChAT protein stability by the heat-shock family of molecular chaperones……………...……………….124

Figure 3.40: H2O2 treatment promotes the generation of redox-dependent higher molecular mass ChAT proteins and decreases cellular ChAT activity…...125

Figure 3.41: Oxidative stress by H2O2 treatment enhances ChAT ubiquitination…………………………...…………………………...……127

Figure 3.42: Oxidative stress by H2O2 treatment promotes ChAT insolubilization…....128

Figure 3.43: HSC/HSP70 inhibition by PES treatment sensitizes ChAT to

H2O2-induced insolubilization……………………………………………130

Figure 3.44: Co-IP of ChAT with endogenous HSC70 is enhanced following

exposure of cells to H2O2……………………………………………...….132

Figure 3.45: Inhibition of the co-chaperone Cdc48/p97/Valosin-containing protein (VCP) prevents degradation of ubiquitinated ChAT……………..133

Figure 4.1: Model for the dynamic regulation of ChAT protein stability and function by molecular chaperones and the ubiquitin-proteasome system….139

Figure 4.2: Hypothetical model for the regulation of ChAT protein stability by the endoplasmic reticulum-associated degradation (ERAD) system…...158

Appendix A: Optimization of BioID in multiple mammalian cell lines………..…..…..215

Appendix B: Optimization of siRNA-mediated knock-down of endogenous HSC/HSP70 and HSP90…………………………………………...…….217

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LIST OF ABBREVIATIONS

15-DSG: 15-deoxyspergualin

17-AAG: 17-allylamino-17-demethoxygeldanamycin

AAA-ATPase: ATPases associated with diverse cellular activities

Aβ: β-amyloid

Acetyl-CoA; AcCoA: acetyl coenzyme A

ACh: acetylcholine

AChE: acetylcholinesterase

AD: Alzheimer’s disease

Aha1: activator of 90 kDa heat shock protein ATPase homology 1

AHA: L-azidohomoalanine

ALS: amyotrophic lateral sclerosis

ANOVA: analysis of variance

APC: anaphase-promoting complex

APP: amyloid precursor protein

ATP: adenosine triphosphate

BAG-1: BAG family molecular chaperone regulator 1

BCA: bicinchoninic acid

BH3: Bcl-2 homology region 3

BioID: proximity-dependent biotin identification

CaMKII: calcium/calmodulin-dependent kinase II

CD: circular dichroism

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CFTR: cystic fibrosis transmembrane conductance regulator

ChAT: choline acetyltransferase

CHIP: C-terminus of HSC70-interacting protein

Chol: choline

CHT: high-affinity choline transporter

CHX: cycloheximide

CK2: casein kinase II

CMS: congenital myasthenic syndrome

Co-IP: co-immunoprecipitation

CTab: anti-ChAT carboxyl-terminal peptide antibody

CTD: C-terminal domain

DAPI: 4’,6-diamidino-2-phenylindole

DIBO: 4-dibenzocyclooctynol

DMEM: Dulbecco’s modified eagle’s medium

DUB: deubiquitinating enzyme

E6-AP: E6-associated protein

Eer1: Eeyarestatin I

EGFR: Epidermal growth factor receptor eNOS: endothelial nitric oxide synthase

ER: endoplasmic reticulum

ERAD: endoplasmic reticulum-associated protein degradation

ESI-MS: electrospray ionization mass spectrometry

FBS: fetal bovine serum

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GAPDH: glyceraldehyde-3-phosphate dehydrogenase

GRP-78: 78-kDa glucose-regulated protein

HA: human influenza hemagglutinin

HBSS: Hank’s balanced salt solution

HECT: Homologous to the E6-AP carboxy terminus

HEK293: human embryonic kidney cells 293

HIF-1α: hypoxia-inducible factor 1-alpha

His6: hexa-polyhistidine-tag

Hop: HSP70-HSP90 organizing protein

HSC70: heat shock cognate 71 kDa protein

HSF1: heat shock factor 1

HSP: heat shock protein

HSP70: heat shock 70 kDa protein

HSP90: heat shock protein 90

HspBP1: HSP70-binding protein 1

HtpG: high-temperature protein G

IP: immunoprecipitation

IPSC: Induced pluripotent stem cell

LC3B-II: microtubule-associated protein 1A/1B-light chain 3 mAChR: muscarinic acetylcholine receptor

MAPK: mitogen-activated protein kinase

MD: Middle domain

MRE: mean residue ellipticity

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mtHSP70: mitochondrial HSP70 / mortalin nAChR: nicotinic acetylcholine receptor

NBD: nucleotide-binding domain

NEF: nucleotide exchange factor

NEM: N-ethylmaleimide nNOS: neuronal nitric oxide synthase

NTD: N-terminal domain

PBS: phosphate-buffered saline

PD: pull-down

PES: 2-phenylethynesulfonamide

PKC: protein kinase C

PLA: proximity ligation assay

Poly-Ub: polyubiquitin

PP5: serine/threonine-protein phosphatase type 5

PVDF: polyvinylidene fluoride

RING: Really interesting new gene

RIPA: radioimmune precipitation buffer

RNAi: RNA interference

ROS: reactive oxygen species

SBD: substrate-binding domain

SDD-AGE: semi-denaturing detergent agarose gel electrophoresis

SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis

SH3: Src-homology 3

siRNA: small-interfering RNA

SNP: single-nucleotide polymorphism

SPAAC: strain-promoted alkyne-azide cycloaddition

TAE: Tris-acetate-EDTA buffer

TAMRA: tetramethylrhodamine

TBS: Tris-buffered saline

TCEP: tris(2-carboxyethyl)phosphine hydrochloride

Tm: thermal denaturation midpoint

TRAP-1: tumor necrosis factor receptor-associated protein 1

TPR: tetratricopeptide repeat

Ub: ubiquitin

UBD: ubiquitin-binding domains

VAChT: vesicular acetylcholine transporter

VCP: Cdc48/p97/Valosin-containing protein

VGCC: voltage-gated calcium channels

WT: wild-type

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CHAPTER 1: INTRODUCTION

1.1: Proteostasis and protein quality control mechanisms

Preservation of protein homeostasis (i.e. proteostasis) is critical for the maintenance of cellular function under both physiological conditions and during cellular stress, whereas dysregulation of proteostatic mechanisms is a hallmark of many diseases. Regulation of cellular proteostasis relies on a complex interconnected network of cellular factors that collectively function as protein quality control mechanisms to promote protein function. These factors can be separated into four broadly defined systems (Figure 1.1); first, factors related to mRNA synthesis, processing, and stability, along with ribosome biogenesis regulate the synthesis of nascent proteins [1-3]. Second, nascent proteins are folded into mature, functional, and active proteins by various endoplasmic reticulum (ER)-associated and/or cytoplasmic molecular chaperones, such as heat shock proteins (HSPs), that cooperate with other cellular factors that regulate disulfide-bond formation, glycosylation, and protein phosphorylation [4-6]. Third, molecular chaperones play a critical role in the maintenance of protein function in cells exposed to various proteotoxic conditions, such as hyperthermia and oxidative stress, by preventing stress-induced protein misfolding and aggregation [7-10]. Fourth, and lastly, the proteolytic degradation of cellular proteins is essential for the timely turnover and attenuation of signalling proteins (e.g. transcription factors), and for the removal of excess, stress-damaged, misfolded, and dysfunctional proteins. Proteolysis of proteins is regulated through two major distinct cellular pathways, these being the autophagy-lysosome and the ubiquitin-proteasome systems [11-13], and, importantly, dysregulation of these proteolytic systems is known to be a critical feature of cancer and of various human neurodegenerative diseases [14-15]. Of note, many of these distinct protein quality control mechanisms not only work in tandem, but can also directly interact with one another, such as in the case for degradation of misfolded proteins by ER- associated protein degradation (ERAD) via the ubiquitin-proteasome system [16]. This thesis will hereafter explore in detail cellular mechanisms related to the ubiquitin- proteasome system and protein degradation, as well as various molecular chaperones that promote protein folding and stability and that target misfolded proteins for degradation.

Figure 1.1: Protein quality control overview. Regulation of protein homeostasis (i.e. proteostasis) is a critical process required for the maintenance of cellular function. Cellular proteostasis relies on protein quality control mechanisms that involve a complex interconnected network of cellular factors that regulate protein synthesis (e.g. ribosome biogenesis), maturation (e.g. endoplasmic reticulum (ER)-associated and cytoplasmic protein chaperones), protein maintenance during cellular stress, and protein degradation through, for example, the ubiquitin-proteasome system (UPS). Of note, many of these distinct cellular mechanisms not only work in tandem, but also interact with one another, such as in the case for ER-associated protein degradation (ERAD) of misfolded proteins via the ubiquitin-proteasome system.

1.2: The ubiquitin-proteasome system

1.2.1: The proteasome

The ubiquitin-proteasome system is one of two major cellular pathways responsible for the highly selective degradation of intracellular proteins in eukaryotic cells [11-12]. This system relies on two core components, the proteolytic 26S proteasome, and the small 8.5 kDa signalling protein ubiquitin (Figure 1.2). The eukaryotic 26S proteasome is a multi- subunit protein complex that consists of a hollow catalytic 20S core particle that is sandwiched between two ATP-dependent 19S regulatory particles [17-19]. The 20S core itself contains four stacked heptagonal rings consisting of two structural α-subunits that interact with the 19S regulatory particle and that are flank by two central β-subunits that impart threonine-dependent proteolytic activity to the 26S proteasome [20-22]. Multiple small molecule inhibitors that target the 20S proteasome core have been identified that are commonly used in cellular and molecular studies, such as MG132 and lactacystin [23-26], or that have been shown to have in vivo anti-tumorigenic properties such as bortezomib [27-28]. Substrate-specificity and activation of the 26S proteasome in regulated by the ATP-dependent 19S regulatory particles. These protein complexes contain multiple Rpt proteins, members of the ATPases associated with diverse cellular activities (AAA- ATPase) family, and non-ATPase Rpn proteins that together form a “molecular lid” that gates entry of substrates into 20S core particle for proteolytic breakdown [22, 29-32]. Recognition of proteasome substrates is mediated through interaction of Rpn proteins within the 19S regulatory particle, specifically Rpn10 and Rpn13, with the small protein ubiquitin when it is covalently attached to cellular proteins destined for proteolysis [33-36].

1.2.2: Ubiquitin

Ubiquitin is a highly-conserved 76 amino acid (8.5 kDa) regulatory protein expressed ubiquitously across eukaryota [37-40]. In mammals, ubiquitin is expressed from four different genes as precursor proteins that are either bound to the ribosomal biogenic

Figure 1.2: The ubiquitin-proteasome system. Cellular proteins are selectively targeted for proteasomal degradation by ATP-dependent post-translational modification with the small 8.5 kDa protein ubiquitin (Ub). Construction of substrate-bound polyubiquitin chains targets ubiquitinated proteins for proteolytic degradation by the 26S proteasome, a large multimeric protein complex consisting of a hollow catalytic 20S core particle sandwiched between two 19S regulatory particles. Binding of polyubiquitin chains to the Rpn proteins Rpn10 and Rpn13 located within the 19S regulatory particles promotes the deubiquitination, unfolding, and uptake of proteasome-targeted proteins into the 20S core where they are subsequently degraded in an ATP-dependent manner. Small molecule inhibitors of the proteasome, such as MG132 and lactacystin, interfere with the 20S core β-subunits, leading to accumulation of ubiquitinated proteasome substrates [36].

proteins L40 or S27 [41-42] or are expressed as tandem repeats that are post-translationally processed into free monomeric ubiquitin by deubiquitinating enzymes (DUBs) [43]. In the cell ubiquitin can most often be found either in its free and unbound monomeric state or covalently bound to other cellular proteins as either monomers or as a polyubiquitin (poly- Ub) chain [43-44]. Canonically, ubiquitination of cellular proteins occurs through the formation of a covalent isopeptide bond between residue Gly76 at the extreme C-terminus

of ubiquitin and the ε-amino group of a targeted lysine residue located either on a substrate protein or on a substrate-conjugated ubiquitin molecule [44-46]. Alternatively, protein ubiquitination has also been shown to occur on serine and threonine residues, as well as on N-terminal methionine residues [47-49]. Ubiquitination has been shown to have multiple diverse effects on protein function, including regulation of enzyme activity and subcellular trafficking, though is known foremost to promote the proteasome-dependent degradation of ubiquitinated proteins [44, 50-51].

1.2.3: Ubiquitination mechanisms

Conjugation of ubiquitin onto cellular proteins is a well-characterized multistep process that involves three distinct classes of enzymes in addition to ubiquitin (Figure 1.3). First, in an ATP-dependent manner monomeric free ubiquitin binds to and forms a thioester bond between the C-terminal residue Gly76 of ubiquitin and the catalytic cysteine residue of an E1 ubiquitin-activating enzyme. Second, the activated ubiquitin is transferred to the active site cysteine of an E2 ubiquitin-conjugating enzyme. Third, E3 ubiquitin ligases facilitate transfer of the active ubiquitin molecule onto a protein substrate by formation of a covalent isopeptide bond between reside Gly76 of ubiquitin and the ε-amino group of a substrate lysine residue. Conjugation of a single ubiquitin molecule on a protein substrate (i.e. monoubiquitination) can then serve as a primer for the construction of larger proteolytic poly-Ub chains by isopeptide attachment of free monomeric ubiquitin onto lysine residues within substrate-bound ubiquitin molecules. Alternatively, poly-Ub chains can be deconstructed and removed from substrate proteins by various DUBs [34, 44, 52-54]. While this process is known to exhibit a high degree of substrate specificity, it is believed that over 800 distinct cellular proteins are involved directly in the regulation of protein ubiquitination. Specifically, in humans two genes encode for two different E1 activating-- enzymes (UBA1 and UBA6) that are solely responsible for ubiquitin activation [55]. Conversely, it is estimated that approximately 40 different E2 conjugating-enzymes, over 600 unique E3 ubiquitin ligases, and roughly 100 distinct DUBs are expressed in human

Figure 1.3: The ubiquitination pathway. Ubiquitination of cellular proteins involves three distinct classes of enzymes. First, monomeric free ubiquitin (Ub) covalently binds to the catalytic cysteine residue of an E1 ubiquitin-activating enzyme in an ATP-dependent manner. Second, the activated ubiquitin molecule is transferred to the active site cysteine of an E2 ubiquitin-conjugating enzyme. Third, E3 ubiquitin ligases facilitate transfer of the active ubiquitin molecule onto protein substrates by promoting the formation of a covalent isopeptide bond between the C-terminal residue Gly76 of ubiquitin and the ε-amino group of a lysine residues on the substrate protein. Attachment of a single ubiquitin molecule (i.e. monoubiquitination) can then serve as a primer for the construction of larger proteolytic polyubiquitin (poly-Ub) chains by isopeptide attachment of free monomeric ubiquitin onto lysine residues located within substrate-bound ubiquitin molecules. Alternatively, poly-Ub chains can be deconstructed and monomeric ubiquitin can be removed from substrate proteins by various deubiquitinating enzymes (DUBs) [40, 43-44, 52, 61].

cells [56-59]. Subsequently, due to the substantial number of and their role in directly interacting with protein substrates, E3 ubiquitin ligases are thought to provide the majority of substrate specificity to the cellular ubiquitination system [44, 60-61].

E3 ubiquitin ligases can be categorized into two major families defined by their sequence/structural homology and the mechanism by which they transfer ubiquitin from E2 conjugating-enzymes onto their substrates. First, members of the homologous to the E6- AP carboxy terminus (HECT) family are characterized by their ability to first accept ubiquitin from E2 enzymes by formation of a covalent intermediate thioester bond via a conserved catalytic cysteine within the E3 HECT domain, then subsequently covalently transfer ubiquitin onto their substrates [62-63]. Of the over 600 estimated E3 ubiquitin ligases, to date only 28 HECT family members have been identified based on homology to the E3 ligase E6-associated protein (E6-AP) [57], with the WW-domain containing NEDD4 [64-66] and the BH3-domain containing HUWE1/Mule [67-68] representing two well-characterized HECT E3 ligases. Conversely, the large majority of E3 ubiquitin ligases belong to the really interesting new gene (RING) family that are characterized for their ability to facilitate direct conjugation of ubiquitin from E2 enzymes onto their substrates without forming covalent intermediates [69-70]. One notable feature of the RING family is the tendency of these E3 ubiquitin ligases to form large multi-protein heterocomplexes, particularly in collaboration with scaffolding Cullin proteins, with two well-documented examples being the Skp1-Cul1-F-box (SCF) complex and the anaphase-promoting complex (APC) [71-72]. Lastly, a smaller third family of RING-like E3 ubiquitin ligases has been identified that contain a conserved catalytic U-box domain [73-75], with the HSP- associated C-terminus of HSC70-interacting protein (CHIP/Stub1) being a notable U-box E3 ubiquitin ligase critical to the chaperone-mediated degradation of misfolded proteins [76-78]. It is important to note that E3 ubiquitin ligases can contain aspects from both HECT and RING ligases, such as Parkin [79], and that members from all E3 families are capable of both mono- and polyubiquitination of their protein substrates [44, 59-60].

1.2.4: The ubiquitin code

Protein ubiquitination has been shown to have both proteolytic and non-proteolytic functions. For example, studies have shown that monoubiquitination can regulate various cellular processes including cell cycle progression, mitosis/meiosis, and transcriptional activity [80-81], as well as endocytosis of plasma membrane-bound receptors and channels [82-84] and viral budding [85-87]. Conversely, the construction of substrate-bound poly- Ub chains canonically promotes the proteasome-dependent degradation of ubiquitinated cellular proteins, though polyubiquitination has also been shown to have non-proteolytic functions. The function of poly-Ub chains is dependent on the “lysine-linkages” between individual ubiquitin molecules within a substrate-bound poly-Ub chain, where ubiquitin contains seven lysine residues at positions K6, 11, 27, 29, 33, 48, and 63 that can be used to form differential ubiquitin-ubiquitin isopeptide linkages to construct poly-Ub chains (Figure 1.4). These different linkages produce structurally-distinct poly-Ub chains, such as for K48- and K63-linked chains, that allow for differential recognition by ubiquitin-binding proteins containing ubiquitin-binding domains (UBD) [88-90]. Classically, the proteolytic features of ubiquitination have been best characterized for K48-linked chains where modification of cellular proteins with poly-Ub chains containing at least four K48-linked ubiquitin molecules targets these polyubiquitinated proteins for proteasome-dependent degradation [44-45, 54, 60, 91-93]. Though, more recently K11-linked chains have also been shown to target substrates for proteasomal degradation and have been shown to regulate cellular processes such as cell cycle progression by the APC [94-96] or ERAD through interaction of K11-linked chains with the ER-associated AAA-ATPase cdc48/p97/Valosin-containing protein (VCP) [50, 97-98]. In contrast, K63-linked chains largely perform non-proteolytic functions and have been shown to regulate endocytic protein trafficking, DNA repair mechanisms, kinase activation, and ribosomal function [99- 102]. Interestingly, multiple studies have also documented the degradation of K63-linked substrates in a proteasome-independent nature via the autophagy-lysosome system [103- 105]. The remaining poly-Ub linkages (K6-, K27-, K29-, and K33-linked chains) have been less characterized and have unclear and/or unknown functions [44, 54, 60, 106].

Figure 1.4: The ubiquitin code. (A) Three-dimensional (3D) ribbon diagram of the crystal structure of human erythrocytic ubiquitin (PDB: 1UBQ) [107]. Ubiquitin contains seven lysine residues (shown in green) at positions K6, 11, 27, 29, 33, 48, and 63 that can all be used for the extension of poly-ubiquitin chains by formation of covalent isopeptide linkages between the C-terminal Gly76 residue (red) of a free ubiquitin molecule and lysine residues within substrate-bound ubiquitin. (B) Cellular proteins targeted for degradation are first monoubiquitinated by covalent attachment of monomeric ubiquitin (Ub) onto substrate lysine residues by E3 ubiquitin ligases. Monoubiquitination can then either alter substrate protein function (i.e. subcellular trafficking and/or protein signalling) or can serve as a primer for the construction of larger proteolytic poly-Ub chains. Classically, proteasome- dependent degradation of cellular proteins requires recognition of substrate-bound K48- linked poly-Ub chains by the 26S proteasome. Alternatively, K11-linked poly-Ub chains have also been implicated in the proteasomal degradation of cell cycle modulators by the anaphase-promoting complex (APC) and of endoplasmic-reticulum-associated protein degradation (ERAD) substrates. Conversely, K63-linked poly-Ub chains largely perform non-proteolytic functions (i.e. regulation of endosomal trafficking, kinase activation, and DNA repair mechanisms), though can also promote the lysosomal-dependent degradation of K63-linked proteins. K6-, 27-, 29-, and 33-linked poly-Ub chains have unclear and/or unknown functions [44, 50-51, 54, 60, 87].

1.2.5: Autophagy-lysosomal degradation

In addition to the ubiquitin-proteasome system, the autophagy-lysosome system is another distinct cellular pathway that mediates degradation of intracellular proteins. Cytoplasmic materials destined for lysosomal degradation, such as damaged organelles and excess, foreign, insoluble, and/or aggregated proteins, are first enveloped into autophagosome vesicles through a process called autophagy [108-109]. Subsequently, autophagosomes fuse with the lysosome, a membrane-bound acidic organelle that contains various pH- sensitive proteases with broad proteolytic activity, resulting in the gross proteolysis of cytoplasmic materials within autophagosomes [110-111]. Though multiple mechanisms

can direct cellular proteins to the lysosome, ubiquitination of lysosomal targets with K63- linked poly-Ub chains is a well documented signal for lysosomal proteolysis [51, 103-105, 112-113]. In addition, alternative poly-Ub chains, such as K27- or K29-linked chains, have been shown to target cellular proteins for lysosomal degradation [114-115]. Lastly, important to the molecular study of lysosome function, multiple lysosomal inhibitors have been identified, such as the vacuolar-type H+-ATPase inhibitor bafilomycin-A1 [116] and the lysosomotropic agent chloroquine [117].

1.3: Protein folding and molecular chaperones

1.3.1: Molecular chaperones overview

Cellular proteolytic pathways are essential to protein quality control and the maintenance of proteome integrity through the break-down of excessive, foreign, misfolded, and dysfunctional proteins, as well as for the turn-over of short-lived signaling proteins. To balance these pro-degradative pathways, cells also rely on opposing molecular mechanisms that promote the stabilization of cellular protein through protein folding, maturation, and maintenance. Proper and completed protein folding of not only newly synthesized nascent proteins, but also of misfolded proteins during cellular stress, represents a critical proteostatic mechanism as the stability and function of most cellular proteins is dependent on obtaining specific three-dimensionally folded structures. The cellular mechanisms that oversee protein folding are extensively varied, though central to this process are molecular chaperones that not only interact with and promote the folding of nascent and misfolded/aggregated proteins but can also “triage” terminally misfolded proteins for proteolytic degradation (Figure 1.5) [118-121]. As such, dysregulation of protein chaperones has been associated with the accumulation of mutated, misfolded, and/or aggregated proteins in several human diseases, such as cancer and age-related neurodegeneration [122-125].

Figure 1.5: Molecular chaperones regulate cellular proteostasis. Molecular chaperones interact with and promote the folding and stabilization of non-native nascent and semi- folded proteins. Additionally, molecular chaperones assist in the refolding and disaggregation of misfolded and/or aggregated proteins following cellular stress (e.g. hyperthermia and oxidative stress) or in the case of mutated or disease-related aggregate- prone proteins (e.g. poly-Q-expanded huntingtin, β-amyloid (Aβ), and α-synuclein). Lastly, molecular chaperones can promote ubiquitination and degradation of terminally misfolded and/or aggregated proteins through either the ubiquitin-proteasome system or the autophagy-lysosome system [119-121].

The term protein chaperone was first introduced in 1978 to describe the ability of the nuclear protein nucleoplasmin to facilitate assembly of nucleosomes, the core subunits of chromatin [126-127]. Later, this term was extended to encompass both prokaryotic and eukaryotic proteins that actively promote the folding of nascent proteins and the assembly of oligomeric protein complexes [128-130]. Today, this classification has expanded to cover over 200 different molecular chaperones that are found throughout the cell, many of which are localized to specific organelles or subcellular regions, and that collectively regulate an exhaustive array of cellular process. Examples of these molecular chaperones include ribosome-binding chaperones (e.g. nascent-chain-associated and ribosome- associated complexes) [131], ER-associated chaperones (e.g. 78 kDa glucose-regulated protein (GRP-78) and protein disulfide isomerase) [132-133], cytoplasmic chaperones (e.g. heat shock proteins (HSPs) and chaperonins) [134-135], mitochondrial chaperones (e.g. Ssc1 and Ssq1) [136], and the aforementioned nuclear chaperone nucleoplasmin [119-120]. Of these, HSPs represent a diverse family of molecular chaperones containing members that are localized throughout the cell and that are involved in the protein folding and stabilization of a multitude of differing cellular proteins [118-120, 135, 137-138].

1.3.2: Heat shock proteins and the heat shock response

HSPs belong to a ubiquitous family of molecular chaperones that are involved in many diverse protein quality control functions, including nascent protein folding, refolding of stress-misfolded and/or aggregated proteins, subcellular trafficking, and in the degradation of terminally misfolded proteins [118, 120, 135]. Interestingly, HSPs were inadvertently discovered in 1962 when Dr. Ferruccio Ritossa first observed chromosome puffing in Drosophila larvae following exposure to elevated temperatures, suggesting the presence of heat-induced changes in gene transcription [139]. This was later confirmed in 1974 by Tissières et al. when Drosophila chromosome puffing was correlated with heat-induced protein expression [140], a response that has since been shown to be a universal response to acute heat shock from bacteria to humans [141-143]. To date numerous HSP family members have been identified that can be separated into six major subfamilies based on

function, homology, and molecular mass, including small HSPs such as HSP27 and HSP33, HSP40/DnaJs, HSP60s, HSP70s, HSP90s, and HSP100s [8, 144-145]. Importantly, though first discovered in the context of acute heat stress, many HSP are now known to be constitutively expressed under non-stressed physiological conditions, such as heat shock cognate 71 kDa protein (HSC70) and HSP90-β, whereas others, such as HSP70 and HSP90- α, are expressed when cells are exposed to protein-misfolding stresses such as hyperthermia and oxidative stress [8, 135, 146].

Elevated expression of stress-inducible HSPs during cellular stress is a critical event required to prevent stress-induced misfolding and aggregation of cellular proteins [8, 135, 144]. The stress-induced expression of HSPs in mammalian cells, i.e. the heat shock response, is regulated by the transcription factor heat shock factor 1 (HSF1) which, under non-stressed conditions, is maintained as an inactive cytosolic monomer by interaction with various constitutively-expressed HSPs [147-149]. The accumulation of stress-denatured proteins following acute proteotoxic stress competes with HSF1 for interaction with these constitutively-expressed HSPs, thus relieving the HSP-mediated sequestration of HSF1 and allowing for trimerization and nuclear import of HSF1 and subsequent stress-induced expression of HSP70 and HSP90-α [150-153]. In turn, HSF1-mediated expression of HSPs protects cells from stress-induced accumulation of misfolded proteins, cellular dysfunction, and apoptosis by chaperoning stress-denatured proteins [8, 154-158]. Lastly, once cellular stress has been relieved HSPs can bind to, sequester, and inactivate HSF1, resulting in abrogation of the heat shock response and restoration of cellular proteostasis [159-161]. Importantly, while many HSP subfamilies contain both constitutive and stress-expressed members, many of these, such as inducible HSP90-α and constitutive HSP90-β, share a high degree of sequence, structural, mechanistic, and substrate homology.

1.3.3: The HSP chaperone network

Despite their name, most members of the HSP family are constitutively-expressed under non-stressed physiological conditions, such as HSP40/DnaJ, HSC70, and HSP90-β that act

together to promote the folding of nascent proteins and to prevent aggregation of misfolded and/or mutant proteins [120, 146, 162]. These HSPs have been shown to chaperone substrate proteins in a cooperative and processive manner (Figure 1.6) where non-native nascent proteins are first recognized by and bind to the co-chaperone HSP40 that then interacts with HSC/HSP70 to form a ternary chaperone-substrate complex [163-165]. Affinity of these HSPs for their protein substrates is mediated through interaction of their substrate-binding domains with exposed peptide sequences rich in hydrophobic amino acids that are located within non-native substrate proteins [166-169]. Next, HSP40 stimulates hydrolysis of HSC/HSP70-bound ATP, causing HSC/HSP70 to adopt an ADP- bound closed-conformation that secures the non-native substrate protein and promotes protein folding [164, 170-171]. Lastly, nucleotide exchange factors (NEF), such as BAG family molecular chaperone regulator 1 (BAG-1) [172] and HSP70-binding protein 1 (HspBP1) [173], promote ADP-to-ATP exchange, causing HSC/HSP70 to adopt an ATP- bound open-conformation that releases the folded substrate protein [174-175]. Though, if a native fully-folded state is not obtained, semi-folded HSP substrates can re-enter this chaperone cycle through repeated interaction with HSP40 and HSC/HSP70 [176-177]. Alternatively, semi-folded HSC/HSP70 substrates can be transferred to HSP90 for further processing by the HSP70-HSP90 organizing protein (HOP/STI1) that reversibly links and mediates transfer of proteins between these molecular chaperones [179-180]. In addition to nascent proteins, misfolded and aggregated proteins can also enter the HSP chaperone network through interaction with the disaggregases HSP104 and HSP110 in cooperation with HSP40 and HSC/HSP70 [181-183]. Lastly, if the native-folded state of a substrate protein cannot be obtained, HSC/HSP70 and HSP90 can triage terminally misfolded proteins for proteasome-dependent degradation through interaction with the HSP- associated E3 ubiquitin ligase C-terminus of HSC70-interacting protein (CHIP) [76-78, 184]. Though many HSPs regulate protein folding, both HSC/HSP70 and HSP90 are central to the maintenance of proteostasis, and as such this thesis will continue to review these HSPs with a focus on their dual roles in protein folding and promoting degradation of misfolded proteins.

Figure 1.6: The HSP chaperone network. (1) Non-native nascent and misfolded proteins bind via exposed hydrophobic regions to the co-chaperone HSP40/DnaJ that then interacts with HSC/HSP70 to form a ternary chaperone-substrate complex. (2) HSP40 stimulates hydrolysis of HSC/HSP70-bound ATP, causing HSC/HSP70 to adopt an ADP-bound closed-conformation that secures non-native protein substrates and promotes protein folding. (3) Nucleotide exchange factors (NEF), such as BAG-1 and HspBP1, promote ADP-to-ATP exchange of HSC/HSP70 (4), causing HSC/HSP70 to adopt an ATP-bound open-conformation that releases the folded substrate protein (5a). If folding is incomplete (5b), non-native semi-folded substrates can re-enter this chaperone cycle through repeated interaction with HSP40 and HSC/HSP70, or (6) can be transferred to HSP90 for further processing by the HSP70-HSP90 organizing protein (HOP/STI1), a tetratricopeptide repeat (TPR)-containing co-chaperone that reversibly links and mediates transfer of substrate proteins between HSC/HSP70 and HSP90. (7) Aggregated proteins can also enter the HSP chaperone network via interaction with the disaggregases HSP104/110 and HSP110 in cooperation with HSP40 and HSC/HSP70. (8) Lastly, HSC/HSP70 and HSP90 can alternatively triage terminally misfolded proteins for proteasome-dependent degradation through interaction with the HSP-associated and TPR-containing E3 ubiquitin ligase C- terminus of HSC70-interacting protein (CHIP) [6, 118, 137, 176].

1.3.4: HSC/HSP70

The HSC/HSP70 family is an extensively studied chaperone subfamily that consists of multiple highly-related proteins found through both prokaryota and eukaryota that range in molecular mass from 66 kDa to 78 kDa. While best know for their role in the co- or post- translational folding of approximately 20% of all nascent proteins [185-187], HSC/HSP70 proteins can also regulate the subcellular trafficking of substrate proteins [188-189] and the degradation of misfolded proteins [78, 184, 190]. Multiple genes in eukaryotes encode for HSC/HSP70 proteins that differ in their molecular mass, expression profile, and subcellular localization [146, 191-193]. For example, the prototypical 70-kDa stress-inducible HSP70- 1a/b (collectively termed HSP70) [191, 193] and the constitutively-expressed HSP70-8 (i.e.

HSC70) are primarily localized to the cell cytoplasm [194-195]. In comparison, the constitutively-expressed HSP70-5 and HSP70-9, known respectively as 78-kDa glucose- regulated protein (GRP-78/BiP) [196-197] and mortalin (mtHSP70/GRP-75) [198], are localized to the ER lumen or mitochondrial matrix respectively. This contrasts with prokaryotes that express only three HSC/HSP70 homologs, including DnaK, Hsc66, and Hsc62 [199-200]. Of note, many eukaryotic HSC/HSP70 proteins have overlapping functions and share similar substrates, though also have distinct functions typically determined by their subcellular localization [146].

Though differences in expression profile and subcellular localization exist, all known members of the HSC/HSP70 family share highly homologous amino acid sequences and conserved domain structures. These domains include an N-terminal nucleotide-binding domain (NBD) with weak ATPase activity, a hydrophobic linker region, a C-terminal substrate-binding domain (SBD), and an extreme C-terminal EEVD-motif that facilitates interaction of HSC/HSP70 proteins with various tetratricopeptide repeat (TPR)-containing co-chaperones such as HSP40, BAG-1, HspBP1, HOP/STI1, and CHIP [6, 146, 176]. The C-terminal SBD can be further subdivided into an α-helical lid that sits adjacent to a substrate-binding β-sandwich that recognizes label and solvent-exposed 4-7 amino acid sequences within substrate proteins enriched in hydrophobic residues [166-169, 201-202]. Mechanistically, ATP-ADP binding to the N-terminal NBD alters affinity of HSC/HSP70 for non-native protein substrates by allosterically altering the C-terminal SBD. Specifically, cycling of HSC/HSP70 between a low-affinity ATP-bound state and a high-affinity ADP- bound state by HSP40 and various NEFs is responsible, respectively, for the allosteric opening or closing of the C-terminal α-lid domain onto protein substrates bound to the C- terminal substrate-binding β-sandwich [203-206]. Accordingly, HSC/HSP70-mediated protein folding is believed to function through a process of kinetic partitioning where non- native hydrophobic regions within protein substrates transiently interact with the C- terminal SBD of HSC/HSP70, thus preventing substrate aggregation by allowing for the unobstructed folding of hydrophilic domains and the subsequent burial of hydrophobic residues/domains upon substrate release [4, 6, 8, 176]. Importantly, the availability of exposed hydrophobic regions within HSC/HSP70 substrates can promote repeated HSP-

substrate binding and folding either until a fully-folded native state is achieved or, if not, the non-native substrate is transferred to HSP90 for further processing or is targeted for proteolytic degradation [118, 137, 176].

Multiple small molecule inhibitors of HSC/HSP70 have been discovered and characterized that can bind to and alter the activity and function of HSC/HSP70 [195, 207- 208]. Due to the ATP-dependent nature of HSC/HSP70-mediated protein folding, inhibition of the N-terminal NBD has proven to be a successful route for the discovery and design of HSC/HSP70 inhibitors, including 3’-sulfogalactolipids [209], the allosteric small molecules MAL3-101 and MKT-007 [210-211], and the ATP mimetic VER-155008 [212- 214]. Alternatively, compounds that interfere with the C-terminal SBD of HSC/HSP70 have also been characterized, including neutralizing peptide aptamers [215], and the small molecules 2-phenylethynesulfonamide (PES) and the modified PES-Cl [216-218]. Lastly, the small molecule 15-deoxyspergualin (15-DSG) and its associated derivatives have been shown to act through multiple means by inhibiting ATPase activity, substrate binding, and by blocking the extreme C-terminal EEVD-motif required for co-chaperone interaction with HSC/HSP70 [210, 219-220]. Importantly, inhibition of HSC/HSP70 substrate binding (e.g. PES) has been shown to result in the accumulation of misfolded and aggregated HSC/HSP70 substrates [216-217], while inhibition of HSC/HSP70 ATPase activity (e.g. VER-155008) results in enhanced degradation of HSC/HSP70 substrates [213-214, 221]. Altogether, the variety of available HSC/HSP70 inhibitors has greatly enhanced the molecular study of HSC/HSP70 function and of their protein substrates, and subsequently has impacted research into human diseases associated with dysregulation of HSPs and proteostasis [218, 222-225].

1.3.5: HSP90

The HSP90 family is another well-conserved subfamily of both constitutive and stress- induced molecular chaperones that make up an estimated 1-2% of all intracellular proteins under physiological conditions [162, 226-227]. Like HSC/HSP70, HSP90 proteins play a

critical role in both the folding and stabilization of nascent and stress-damaged proteins [149, 228-229], as well as in the degradation of misfolded proteins [184, 230-231]. HSP90s are believed to interact with and regulate the function of at least 200 different cellular proteins [232], though, despite this apparent promiscuity, HSP90 is thought to contain a greater substrate specificity compared to that of HSC/HSP70 which, respectively, interacts with a diverse and extensive range of non-native proteins [233-235]. Specifically, in the current model (Figure 1.6) HSP90 acts downstream of HSC/HSP70 to interact with semi- or pre-folded proteins and appears to play a notable role in the structural maturation and conformational activation of signalling proteins such as steroid receptors and kinases [236- 238]. Subsequently, a high proportion of identified HSP90 substrates are either enzymes and/or kinases involved in the regulation of cell cycle control [239-240], cell survival [241], epigenetics [242-243], or in hormone receptor and cell signalling [244-247]. Thus, HSP90 functions as a critical regulator of a subset of the proteome that integrates together several cellular signalling pathways through its chaperoning function [232, 235, 248].

Similar to HSC/HSP70, multiple genes in eukaryotes encode for HSP90 proteins that differ in their molecular mass, expression profile, and subcellular localization [249- 250]. Specifically, three cytoplasmic isoforms of the prototypical 90-kDa HSP90 have been identified including the constitutively-expressed HSP90-β and the stress-inducible HSP90- α1/α2 [251-253]. Additionally, ER- and mitochondrial-localized isoforms of HSP90 have been identified, including ERp99/GRP-94 [254-255] and tumor necrosis factor receptor- associated protein 1 (TRAP-1) respectively [256]. In contrast, prokaryotes express a single HSP90 homolog, high-temperature protein G (HtpG) [257].

All known members of the eukaryotic HSP90 family share highly conserved domains that consist of a nucleotide-binding N-terminal domain (NTD) with ATPase activity, a charged linker region, a substrate-binding middle domain (MD), and a C- terminal domain (CTD) responsible for HSP90 dimerization [4, 162, 227, 235]. Furthermore, HSP90s contain an extreme C-terminal MEEVD-motif that allows for interaction with various TPR-containing co-chaperones such as HOP/STI1 [258-260], peptidyl-prolyl-isomerases (e.g. cyclophilin 40) [261], and serine/threonine-protein phosphatase type 5 (PP5) [262]. Mechanistically, HSP90 functions as a C-terminally-bound

homodimer where ATP-ADP cycling at the NTD causes allosteric conformational changes to the CTD that facilitates substrate folding (Figure 1.7). Specifically, HSP90 in the absence of nucleotide binding forms an early-stage “apo-open” complex that binds to semi-folded protein substrates in complex with HSC/HSP70 and HOP/STI1 [178-180, 263-264]. Like other HSPs, binding of non-native proteins to HSP90 is promoted by the availability of exposed hydrophobic sequences within protein substrates [234, 246. 265]. Next, ATP binding to the NTDs of HSP90, and subsequent ATP hydrolysis, causes dimerization of the NTDs with concomitant compaction of the HSP90 dimer into a “closed” conformation, securing and promoting the conformational folding of protein substrates [266-268]. Following ATP hydrolysis, the NTDs separate, release ADP, and the folded protein substrate is released, thus returning HSP90 to the apo-open state ready for new or repeated substrate binding [263, 268-269]. Multiple co-chaperones can directly influence the HSP90 chaperone cycle, including HOP/STI1 and Cdc37/p50 which both stabilize HSP90 in its open conformation and inhibit ATP hydrolysis [270-271], while HOP/STI1 additionally facilitates substrate transfer from HSC/HSP70 to HSP90 [178-180]. Furthermore, following ATP and protein substrate binding to HSP90, two molecules of p23 can bind to the ATP-bound NTDs and stabilize the substrate-bound HSP90 dimer prior to ATP hydrolysis [272-273]. Conversely, the co-chaperone activator of 90 kDa heat shock protein ATPase homology 1 (Aha1) can stimulate the inherent ATPase activity of HSP90 by binding to the MD of HSP90, causing transition of HSP90 to its closed, pro-folding conformation [271, 274]. In addition, various TPR-containing co-chaperones can interact with HSP90 through its C-terminal MEEVD-motif, such as peptidyl-prolyl-isomerases, to assist in the final conformation folding and activation of HSP90 substrates [261-262]. Lastly, HSP90 can interact with the E3 ubiquitin ligase CHIP in a TPR-mediated fashion that promotes degradation of terminally misfolded protein substrates [230, 275-276].

Various small molecules have been discovered and characterized that can bind to and alter the activity and function of HSP90 [277-278]. Most of these inhibitors target the NTDs of HSP90 and their associated ATPase activity, such as the natural antibiotics radicicol [279-280] and geldanamycin, of which was instrumental in deciphering the structural and molecular mechanisms of the HSP90-ATP chaperone cycle [279-282].

Figure 1.7: The HSP90 chaperone cycle: (1) In the absence of HSP90 nucleotide-binding semi-folded non-native protein substrates, in complex with HSC/HSP70 and HOP/STI1, interact with a HSP90 homodimer bound together by the C-terminal domains (CTD) of HSP90. The co-chaperone Cdc37 prevents binding of ATP to the N-terminal domains (NTD) of HSP90 and maintains an early-stage “apo-open” complex. (2) ATP binds to and induce dimerization of the HSP90 NTDs. The co-chaperone p23 stabilizes an “intermediate” substrate-bound HSP90 dimer prior to ATP hydrolysis by binding to the ATP-bound NTDs. (3) Binding of the co-chaperone Aha1 to the middle domain (MD) of HSP90 promotes ATP hydrolysis, causing an allosteric compaction of the HSP90 dimer into a “closed” conformation, securing and promoting conformational folding of protein substrates. Various tetratricopeptide repeat (TPR)-containing co-chaperones, such as peptidyl-prolyl-isomerases (PPI) and serine/threonine-protein phosphatase type 5 (PP5), interact with the HSP90 C-terminal MEEVD-motif to aid in substrate folding. Following ATP hydrolysis, the HSP90 NTDs separate, release ADP, and the folded protein substrate is released (4), returning HSP90 to the apo-open state where it is ready for new or (5a) repeated protein substrate binding. Alternatively, (5b) HSP90 can interact with the TPR- containing E3 ubiquitin ligase CHIP to promote degradation of terminally misfolded HSP90 substrates [162, 227, 235].

Subsequently, a geldanamycin derivative, 17-allylamino-17-demethoxygeldanamycin (17- AAG), has been shown to be a potent inhibitor of HSP90 activity [283-284], resulting in enhanced proteasomal degradation of HSP90 substrates [230, 285-287]. Lastly, coumarin antibiotics, such as novobiocin, have been shown to interfere with the dimerization function of the HSP90 CTD, resulting in proteolytic loss of cellular HSP90 substrates [288-289].

1.3.6: Ubiquitin E3 ligase CHIP

Degradation of dysfunctional, mutated, and misfolded/aggregated proteins is a critical event required to maintain cellular proteostasis. Importantly, both HSC/HSP70 and HSP90

can triage misfolded proteins for proteolytic degradation through interaction with the E3 ubiquitin ligase CHIP/Stub1 [184, 276, 290]. CHIP is a cytosolic member of the U-box family of RING-like E3 ubiquitin ligases that contains a C-terminal U-box domain with E3 ligase activity [77, 291-292] and three tandem N-terminal TPR domains that facilitate interaction of CHIP with the C-terminal EEDV-motifs of HSC/HSP70 and HSP90 [76-78, 293]. CHIP has been shown to regulate diverse cellular functions due to its capacity to ubiquitinate a multitude of HSC/HSP70 and HSP90 substrates, including the glucocorticoid receptor [184, 294], hypoxia-inducible factor 1-alpha (HIF-1α) [295], the receptor tyrosine kinase ErbB2 [230, 285], the transcription factors Foxp3 and RFX1 [296-297], and the cystic fibrosis transmembrane conductance regulator (CFTR) [298-299]. Importantly, CHIP can target its protein substrates to distinct proteolytic pathways through the synthesis of either K48- or K63-linked poly-Ub chains. Specifically, interaction of CHIP with the E2 ubiquitin-conjugating enzyme UbcH5 promotes K48-linked polyubiquitination and proteasome-dependent degradation of CHIP substrates [292, 300-301], whereas interaction with the E2 enzyme Ubc13 has been shown to promote K63-linked polyubiquitination and endocytosis of growth hormone receptors [302-303]. Owing to its regulatory roles in a range of diverse cellular processes, CHIP mutations are associated with development of hereditary cerebellar ataxias, including Gordon Holms spinocerebellar ataxia syndrome [304-305], as well as widespread multisystemic neurodegeneration [306].

The substrate specificity of CHIP is dependent on interaction with HSPs where it preferentially ubiquitinates HSP-bound misfolded proteins [77, 184, 291]. Of note, various HSP-interacting co-chaperones have also been shown to affect CHIP function, including BAG-1 which cooperates with CHIP to target HSC/HSP70 substrates for degradation [307- 308], as well as BAG-2 and HspBP1 which, conversely, inhibit CHIP ligase activity [299, 309-310]. To date no small molecule inhibitors of CHIP have been identified, though siRNA-mediated knock-down of CHIP has proven to be an effective means for investigating CHIP-substrate dynamics [296, 311-312]. In addition to HSP-bound protein substrates, CHIP can also directly ubiquitinate and regulate both the activity and the degradation of HSC/HSP70 and HSP90 under non-stressed conditions [291, 313-315], and furthermore plays a key role in restoring cellular proteostasis following cellular stress by

degrading stress-induced HSPs [316-317]. Though CHIP is well-known for its role in chaperone-mediated proteolysis, one intriguing and yet unclear question is which factors regulate the triage decisioning between folding or degradation of HSP substrates. One leading theory stipulates that CHIP can compete through its tandem N-terminal TPR domains with other pro-folding TPR-containing co-chaperones for binding to the C- terminal EEVD-motif of HSC/HSP70 and HSP90. In support, CHIP has been shown to compete with HOP/STI1 for binding to C-terminus of HSP70 and HSP90, and subsequently that C-terminal phosphorylation of these HSPs regulates the alternate binding of pro- folding HOP/STI1 and pro-degradative CHIP [276, 315, 318].

1.4: Molecular chaperones in human disease

1.4.1: HSPs in Cancer

Provided their essential role in maintaining cellular proteostasis, HSPs have been implicated in the pathogenesis of many human cancers and neurodegenerative diseases. In respect to cancers, one of the defining features and drivers of tumorigenesis is the build-up of mutated and dysregulated proteins, especially those associated with regulation of cell cycle and cell survival mechanisms [319-321]. Importantly, HSPs such as HSC/HSP70 and HSP90 have been shown to chaperone and stabilize mutant and dysregulated cancer-related proteins, such as mutated and inactive p53, thereby promoting cell cycle dysregulation and tumorigenesis [322-323]. Additionally, HSPs play a critical anti-apoptotic role during tumorigenesis by preventing stress-induced apoptosis due to hypoxic build-up of oxidative and proteotoxic stress within the microtumor environment [324-325]. Consequently, dysregulated and heightened expression of both constitutive and stress-induced HSPs is a common feature of many cancers which often correlates with poorer clinical prognosis and can promote resistance to chemotherapy [323, 326]. Therefore, an area of intense study for cancer therapy is the development of clinically-relevant inhibitors of HSPs, such as the HSP90 inhibitor 17-AAG that has shown promising results in clinical trial [327-328].

1.4.2: HSPs in aging and neurodegeneration

The functions of HSPs have also been implicated during the aging process and in the pathology of human neurodegenerative diseases [10, 329]. One significant feature of cellular aging is a decline in both the constitutive and stress-inducible expression of HSPs, resulting in a diminished capacity of aging cells to chaperone non-native proteins, as well as respond to cellular and proteotoxic stress [330-332]. In turn, this age-related decline in chaperoning capacity can lead to the accumulation of misfolded aggregate-prone proteins, loss of cellular proteostasis, and accelerated cell death [332-333]. Importantly, increased aberrant protein misfolding and accumulation of protein aggregates during aging has been associated with the development of type 2 diabetes mellitus [334] as well as various neurodegenerative diseases, including Alzheimer’s disease (AD), Parkinson’s and Huntington’s disease, and amyotrophic lateral sclerosis (ALS) [4, 6, 10]. Although the mechanisms responsible for these different age-related diseases are varied, one common pathological feature is the accumulation of cytotoxic amyloid or amyloid-like oligomers and aggregates [335-336].

The cytotoxic properties of these age-related misfolded and aggregated proteins have been foremost attributed to their ability to directly impair organelle function, such as mitochondria [337], the ER [338], and autophagosomes [339]. To exacerbate this issue, protein aggregates can inadvertently interact with exposed hydrophobic regions found within non-native nascent proteins and in numerous structurally-labile proteins, such as enzymes, kinases, and transcription factors, thereby co-aggregating with and sequestering these cellular proteins from performing their normal functions, resulting in multifactorial cytotoxicity [122, 340-341]. Lastly, protein aggregates can directly impair activity of the ubiquitin-proteasome system, leading to the establishment of an aggregate-driven positive feedback system that promotes the build-up of pathogenic protein aggregates, resulting in gross proteostatic dysregulation, induction of proteotoxic cellular stress, and enhanced cell death [4, 124, 342-343]. Of note, multiple studies have demonstrated that HSPs have cytoprotective properties in multiple neurological protein misfolded diseases [10, 344- 345]. Specifically, in the case of Huntington’s disease, HSP70 and HSP40 have been shown to chaperone and prevent the assembly of mutant polyglutamine-expanded proteins, such

as huntingtin, into cytotoxic aggregates [346-347]. Additionally, HSPs can interfere with the toxic accumulation of AD-associated hyperphosphorylated tau protein or β-amyloid (Aβ) [348-349], or with Parkinson’s-associated α-synuclein [350]. Consequently, pharmacological upregulation of HSP expression or chaperone activity [351-352], enhancement of proteasome activity [353-354], or stimulation of the ubiquitination of misfolded and aggregated protein [15, 355-356] represent exciting new avenues for the treatment of protein misfolding neurodegenerative diseases.

1.4.3: HSPs and oxidative stress

Common to both cancer and neurodegenerative diseases is the build-up of oxidative stress and concomitant acceleration of protein misfolding and cell death [357-358]. A powerful and significant source of cellular oxidative stress is the generation of free reactive oxygen species (ROS) that have the capacity to oxidize cellular components, including cellular proteins, lipids, and DNA [359-361]. For example, ROS-mediated oxidation of thiol- containing cysteine residues within cellular proteins can either promote the formation of aberrant intra- and intermolecular disulfide bonds or can stimulate modification of oxidized proteins with various protective antioxidant modifications such as glutathione or thioredoxin [362-364]. Though the formation of disulfide bonds during the synthesis of nascent proteins is often required for proper protein function, the formation of stress- induced disulfide bonds during oxidative stress can induce protein misfolding with severe and damaging functional consequences [361, 364-365]. Therefore, provided the critical role of cysteine residues in disulfide bond formation and protein folding, it is reasonable to hypothesize that cysteine-rich proteins may be more susceptible to oxidative misfolding and dysfunction. One such protein, the disease-associated E3 ubiquitin ligase Parkin, has a higher-than-average cysteine content of 7.5% compared to 2.26% for the global human proteome [366] or 1.6% for intracellular mammalian proteins [367]. Importantly, studies have demonstrated that exposure of Parkin to H2O2, a potent inducer of cellular oxidative damage, can induce misfolding, inactivation, and aggregation of Parkin [368-370].

Interestingly, HSPs play a key role in mitigating the cytotoxic and pro-apoptotic effects of oxidative stress by chaperoning stress-damaged and aggregate-prone proteins [7, 371]. Of note, prolonged cellular exposure to oxidative stress can lead to the induction of the heat shock response through oxidative depletion of glutathione, an antioxidant and redox modulator that can compete with and prevent the formation of aberrant oxidative disulfide bonds [372-374]. Oxidative depletion of glutathione in turn promotes the oxidation of cellular proteins and formation of non-native disulfide bonds, causing gross cellular protein misfolding that subsequently stimulates activation of HSF-1 and the heat shock response [375-377]. Consequently, stress-induced HSP70 and HSP90 can either bind to and chaperone oxidized proteins to maintain and/or restore protein function, or alternatively can promote proteasomal degradation of oxidatively damaged proteins [371, 378-379]. In addition to their stress-induced functions, HSPs have also been found to constitutively maintain an available pool of reduced glutathione that can buffer against acute oxidative stress [380-381]. Lastly, in the case of neurodegenerative diseases various HSPs have been shown to suppress the oxidative properties of mutant and misfolded huntingtin, Aβ1-42, and the ALS-associated DNA-binding protein TDP-43 [382-384]. Taken together, these studies establish a critical role for HSPs not only in the regulation of nascent protein folding, but also in the maintenance of cellular proteostasis during exposure to cellular stress and in human disease.

1.5: Cholinergic neurobiology

1.5.1: The cholinergic synapse

Build-up of oxidative stress and dysregulation of cellular proteostasis are common cellular events observed during aging and in many age-related neurodegenerative diseases, including AD, and may represent mechanisms that are critical to the pathogenesis of these diseases. Work in the Rylett Laboratory is primarily focused on the role of cholinergic neurons in the brain during aging and AD [385-386] as these neurons appear to be particularly sensitive to oxidative stress and to AD-related dysfunction and degeneration

[387-388]. Cholinergic neurons are critical components of the central, autonomic, and the somatic nervous system where they are essential to the regulation of bodily homeostasis through their ability to synthesize and release the neurotransmitter acetylcholine (ACh) [389-390]. In turn, ACh regulates diverse processes throughout the human body, including cognitive functions such as behaviour and memory, as well as autonomic (e.g. heart rate, pupil dilation, and gastric function) and neuromuscular processes. Consequently, dysregulation of ACh synthesis and/or neurotransmission has been observed in multiple neurodegenerative and neuromuscular disorders [388, 391].

Cholinergic neurotransmission has been extensively studied over the past fifty-plus years and is known now to involve the cooperative actions of multiple presynaptic proteins that oversee the synthesis and release of ACh, as well as postsynaptic ACh receptors that relay cholinergic neural activity into discreet post-synaptic signalling events (Figure 1.8). Briefly, in this system ACh is first produced in the presynaptic terminals of cholinergic neurons by the cytoplasmic enzyme choline acetyltransferase (ChAT) using the substrates choline and acetyl coenzyme A (acetyl-CoA) [392]. ACh is then transported and stored in presynaptic lipid-bound vesicles by the vesicular acetylcholine transporter (VAChT) [393]. Depolarization and activation of cholinergic neurons induces calcium-dependent fusion of ACh-containing vesicles with the presynaptic terminal plasma membrane, resulting in release of ACh into the synaptic cleft [394-395]. Subsequently, released ACh then binds to postsynaptic membrane-bound receptors, such as nicotinic and muscarinic ACh receptors, that then initiate downstream postsynaptic signalling events [396-397]. To attenuate cholinergic signalling, ACh is broken down into choline and acetate by the enzyme acetylcholinesterase (AChE) located within the synaptic cleft [398]. Lastly, extracellular choline is transported back into the presynaptic terminals of cholinergic neurons by the high-affinity choline transporter (CHT) where it can be used by ChAT for the renewed synthesis of ACh [390, 399]. Importantly, many of these cholinergic proteins have been found to be dysregulated in neurodegenerative and neuromuscular diseases [400-402], prompting researchers to develop treatment-oriented cholinergic modulators, such as the acetylcholinesterase inhibitors donepezil and rivastigmine for the treatment of AD [403].

Figure 1.8: The cholinergic synapse. (1) In the presynaptic terminals of cholinergic neurons, the enzyme choline acetyltransferase (ChAT) synthesizes the neurotransmitter acetylcholine (ACh) using the substrates choline and acetyl-CoA. (2) ACh is transported into and stored in lipid-bound synaptic vesicles by the vesicular acetylcholine transporter (VAChT). (3) Depolarization and activation of cholinergic neurons induces Ca2+ influx into the presynaptic terminals through voltage-gated calcium channels (VGCCs), resulting in fusion of ACh-containing vesicles with the terminal plasma membrane and release of ACh into the synaptic cleft. (4) ACh binds to postsynaptic membrane-bound nicotinic and/or muscarinic acetylcholine receptors (n/mAChRs), resulting in the opening of ligand-gated ion channels (nicotinic) and/or activation of G-coupled protein signalling (muscarinic) that alters postsynaptic ion dynamics and downstream subcellular signalling. (5) Cholinergic signalling is attenuated by breakdown of ACh in the synaptic cleft into choline and acetate by the enzyme acetylcholinesterase (AChE). (6) Lastly, free choline is transported back into cholinergic presynaptic terminals by the high-affinity choline transporter (CHT) where it can be used again as a substrate for ChAT in the repeated synthesis of ACh [389, 392].

1.5.2: Choline acetyltransferase (ChAT)

The enzyme ChAT holds a particularly vital role in cholinergic neural function as it is the sole protein responsible for the synthesis of ACh. ChAT is a member of the CoA-dependent superfamily of acyltransferases that includes, for example, carnitine and chloramphenicol acetyltransferases [404-405], and is a phenotypic marker of central, autonomic, and somatic cholinergic neurons [389-390]. To date, multiple ChAT transcripts have been identified in both human and rodents that are formed by alternative splicing and differential use of the non-coding exons R, N, M, S and H [392, 406-408]. All transcripts encode a common 69- kDa ChAT enzyme that is localized predominantly to the neuronal cytoplasm where it generates ACh, while the spinal cord-isolated S-transcript also encodes for a 74-kDa isoform [407, 409-411]. Additionally, unique to humans and non-human primates the M- transcript also encodes for an 82-kDa ChAT protein in addition to 69-kDa ChAT due to the presence of a unique in-frame translation initiation codon located 5’ to the common

translation initiation site for 69-kDa ChAT [407, 411, 412-413]. This primate-specific 82- kDa ChAT protein contains a nuclear localization signal (NLS) located within its extended 118 amino acid N-terminus that directs it to the nuclei in human brain and spinal cord cholinergic neurons [385, 414-415]. While our laboratory has shown that 82-kDa ChAT can synthesize ACh [386, 414-415], the function/s of this nuclear ChAT isoform remain under investigation, though more recent studies have gathered evidence that 82-kDa ChAT may be involved in regulating chromatin organization and the gene expression of important amyloid precursor protein (APP) processing proteins such as GGA3 [386, 416-417].

Though different molecular mass ChAT protein isoforms exist, the cytoplasmic 69- kDa variant is considered the canonical ChAT enzyme responsible for ACh synthesis due to its shared expression from the aforementioned mRNA transcripts in multiple animal species. Consequently, provided its essential role in cholinergic neurotransmission, multiple studies have investigated the regulation of ChAT enzyme activity with a focus on the structural and/or molecular and biochemical properties of 69-kDa ChAT. Importantly, the crystal structures of both rat [418-419] and human 69-kDa ChAT [420] have been solved. Structurally, human ChAT contains two functional domains, a substrate-binding domain that binds to its two substrates choline and acetyl-CoA, and a catalytic domain with an interfacial active site tunnel that runs between these two domains [420]. An active site histidine residue, His324, catalyzes transfer of the acetyl group from acetyl-CoA to choline in a sequential reaction resulting in ACh synthesis [420-422]. Impaired in vitro activity of purified recombinant ChAT has been demonstrated under oxidative conditions and when ChAT is exposed to either sulfhydryl-reactive or arginine-modifying reagents [423-424]. Subsequently, the arginine residues Arg442 and Arg443 have been shown to be indirectly involved in CoA-binding through interactions with a surface “P-loop” that stabilizes acetyl- CoA within the active site [420, 425]. In addition to these structural determinants of ChAT activity, post-translational phosphorylation of ChAT has also been shown to regulate its enzymatic activity where phosphorylation by protein kinase C (PKC) and casein kinase II (PK2) increases the in vitro activity of recombinant human ChAT [426]. Importantly, in human HEK293 or IMR32 neuroblastoma cells residues Ser440 and Ser476 of human 69-kDa ChAT have been shown to be phosphorylated by PKC in a hierarchical fashion such that

phosphorylation of Ser476 regulates the basal activity of ChAT and where PKC activation promotes phosphorylation of residue Ser440, subsequently enhancing ChAT activity [410, 427]. Interestingly, mutation of residue Arg442 not only prevents PKC-mediated phosphorylation of ChAT at residue Ser440 but also dramatically reduces basal ChAT enzymatic activity [427]. Analysis of the human ChAT crystal structure reveals that these residues, Ser440 and Arg442, can form hydrogen bonds between each other [420], suggesting a potential biochemical link between ChAT phosphorylation and protein structural changes that may influence acetyl-CoA binding and subsequent enzymatic activity of human ChAT.

1.5.3: ChAT in neurodegenerative diseases

Alterations in ChAT activity have been found in multiple neurological disorders, including AD [428-429], ALS [430] and in Huntington’s disease [431]. Of significance, impairment of ChAT activity [432-435] and reduced ChAT mRNA expression [436-437] have been associated with the clinical progression of AD. Furthermore, exposure of cells to AD- related and cytotoxic Aβ1-42 oligomers results in calcium/calmodulin-dependent kinase II (CaMKII)-mediated phosphorylation of ChAT on residue Thr456, altering ChAT activity, and promoting interaction of ChAT with the ER- and HSP-associated co-chaperone Cdc48/p97/VCP [438-440]. VCP is a critical mediator for the proteasome-dependent degradation of ubiquitinated ERAD substrates [50, 97, 441-442], as well as for the maturation of ubiquitin-containing autophagosomes that are critical to the autophagy- lysosome system [443-445]. While it is currently unknown whether ChAT protein stability and function is regulated by either the ubiquitin-proteasome and/or autophagy-lysosome system, these studies collectively suggest that ChAT ubiquitination and protein stability may be altered following exposure to cytotoxic Aβ stress. Lastly, aggregated forms of Aβ have been found to reduce ChAT activity in vivo in the striatum of aged rats [446]. These studies detailing alteration in ChAT enzyme activity during AD or following exposure to cytotoxic Aβ, together with the observation that significant loss of cholinergic neurons occurs in patients with AD [447-449], forms the basis for the cholinergic hypothesis of AD and suggest a critical role for ChAT during AD [450-451]. Subsequently, studies have

investigated whether coding and non-coding single-nucleotide polymorphisms (SNPs) within the human ChAT gene are genetically linked to the development of AD, though comparison of these studies reveals variable results [452-456].

1.5.4: Congenial myasthenic syndrome (CMS)

Though alterations in ChAT activity and mRNA expression have been correlated with the progression of AD and other neurodegenerative diseases, direct and conclusive genetic linkage of ChAT missense mutations with these neurodegenerative diseases has yet to be observed. Conversely, ChAT mutations have been causatively-linked to the development of the rare neuromuscular disorder congenital myasthenic syndrome (CMS) [411, 457- 458]. CMS is an inheritable and heterogeneous disorder with an early-childhood onset in which neuromuscular synaptic mechanisms are compromised, resulting in chronic muscle weakness (i.e. myasthenia), difficulties in movement, eating, and breathing, and in severe cases of CMS periods of episodic apnea that can be fatal without medical intervention [402, 459]. Of note, CMS differs from other myasthenic diseases, such as myasthenia gravis and Lambert-Eaton syndrome, due to the genetic/mutational origins of CMS compared to the non-genetic autoimmune features of these other myasthenic diseases [402, 460-461]. Myasthenic syndromes, including CMS, are often highly-treatable disease, though treatment efficacy specifically for CMS patients is highly dependent on the underlying genetic cause for a patients CMS diagnosis. Importantly, to date mutations in no fewer than 20 different neuromuscular proteins, including cholinergic proteins, have been causatively- linked to CMS development [402, 462-463].

While mutations in various proteins localized either pre- and/or postsynaptically at neuromuscular junctions are linked to CMS [402, 459], early in vitro electrophysiological studies of CMS patients revealed that ACh synthesis can be directly impaired, suggesting a role for ChAT in the pathogenesis of CMS [464-466]. Subsequently, multiple studies have identified over 40 unique CMS-related missense mutations in the human 69-kDa ChAT protein, many of which have been biochemically characterized [411, 457-458, 467-

473]. The severity of CMS-related ChAT mutations has been associated with the position of the mutated residues within the three-dimensional structure of ChAT where, in general, mutations involving the ChAT active site can severely impair enzymatic activity, often through alterations in acetyl-CoA binding. Furthermore, these active site mutations often display a concomitant reduction in ChAT protein levels, resulting in a synergistic loss in ACh synthesis associated with development of severe CMS (Figure 1.9) [411, 457-458]. One CMS-related mutations located within the ChAT active site tunnel is an arginine-to- histidine mutation of residue Arg442, a residue our laboratory has shown to regulate both basal ChAT activity and the PKC-mediated phosphorylation of ChAT on residue Ser440 [420, 427]. Residue Arg442 is hypothesized to indirectly stabilize acetyl-CoA binding to ChAT, and while the CMS-related R442H mutation does not appear to affect ChAT protein levels in cells [411, 427], this mutation does dramatically reduce the in vitro affinity of recombinant R442H-ChAT for acetyl-CoA, causing a 99% loss in catalytic activity [411].

1.5.5: V18M-ChAT and the ChAT N-terminus

Interestingly, several catalytically-deficient CMS-related ChAT mutations have been found that are positioned distal to the ChAT active site and are hypothesized to work through allosteric mechanisms, though these mutations tend to produce less severe catalytic defects and often do not affect protein expression of ChAT [411, 457-458]. One notable exception is a valine-to-methionine mutation of the N-terminal residue Val18 that was identified in four unrelated CMS patients with severe myasthenic symptoms [457-458]. Biochemical characterization of this mutation not only revealed a notable decrease in the in vitro affinity of recombinant V18M-ChAT for acetyl-CoA, leading to a 30-75% loss in overall catalytic efficiency, but also a remarkable 30-40% reduction in the steady-state protein levels of mutant V18M-ChAT compared to wild-type ChAT when transiently expressed in either BOSC 23 or HEK293 cells (Figure 1.9) [457-458]. Furthermore, this V18M mutation was found to have negligible effects on the in vitro thermal stability of the enzyme [457], suggesting that the loss of V18M-ChAT catalytic activity and cellular protein expression is not likely due to gross alterations in ChAT protein folding.

Figure 1.9: Missense mutations within human ChAT are associated with development of the neuromuscular disorder congenital myasthenic syndrome (CMS). CMS-related ChAT mutations disrupt ChAT function by (A) reducing the steady-state protein levels of mutant ChAT in cells and/or (B) by altering choline (chol) and/or acetyl-CoA (AcCoA) binding. In general, mutations involving the ChAT active site (red asterisks) severely impair ChAT enzymatic activity, frequently due to a loss in AcCoA affinity, and are often accompanied by a concomitant reduction in ChAT protein levels resulting in a synergistic loss in ACh synthesis that is associated with development of severe CMS. Interestingly a valine-to-methionine mutation of the N-terminal residue Val18 has been identified in four unrelated CMS patients with severe myasthenic symptoms. This residue is positioned distal to the ChAT active site where mutation of Val18 has been found to reduce both the cellular steady-state levels protein levels of and the in vitro AcCoA affinity of mutant V18M-ChAT (green asterisks; D and E). Shown are anti-ChAT immunoblot from transfected COS cells (A), in vitro kinetic parameters (B), and mutation summaries (C, D, and E) for various CMS-related ChAT mutants collated from three independent studies completed previously by other laboratories [A-C, 411; D, 457; E, 458]. Amino acid numbering corresponds to the human 82-kDa ChAT protein sequence (+118 amino acids vs. 69-kDa ChAT).

The mechanisms by which this CMS-related V18M mutation enacts its effects on cellular ChAT function remains to be elucidated. One hypothesis provided by the authors of the above studies is that mutation of residue Val18 may disrupt a hydrogen-bond network that involves other active site-localized residues that contribute to acetyl-CoA binding, such as residue Ser440 [457-458]. Importantly, in silico analysis of the human 69-kDa ChAT amino acid sequence revealed that residue Val18 is positioned within a highly-conserved N- terminal proline-rich motif with sequence 14PKLPVPP20 (Figure 1.10). This motif is positioned distal to the ChAT active site, is surface-exposed, and forms part of a stably- folded domain (Figure 1.11) where neither Val18 nor any other residue in this motif participates directly in acetyl-CoA binding [420, 457-458]. Interestingly, this highly- conserved proline-rich motif of ChAT shares structural and sequence homology with the core PxxP-binding motif for Src homology 3 (SH3)-binding domains [420, 474-475].

Figure 1.10: Comparative sequence alignment of the highly-conserved N-terminal proline-rich motif of ChAT from multiple divergent animal species. An in silico NCBI Blastp search with downstream sequence alignment using Clustal Omega (EMBL-EBI) was completed to compare the first 50 N-terminal amino acids of human 69-kDa ChAT to the indicated animal species. The sequence for the N-terminal proline-rich motif of human ChAT (14PKLPVPP20), including the CMS-related residue Val18 (i.e. V18M), is indicated. Sequence alignment revealed either complete (green) or a high degree (yellow) of amino acid conservation within the ChAT proline-rich motif between multiple species, with residues differing from the human 69-kDa ChAT sequence highlighted in purple.

Figure 1.11: Three-dimensional crystal structure of human 69-kDa ChAT. Shown is a space-fill 3D model of the human 69-kDa ChAT structure, bound to acetyl-CoA, that was solved previously in the Rylett Laboratory (PDB: 2FY4) [420]. ChAT contains a highly- conserved N-terminal proline-rich motif at residues 14PKLPVPP20 (green) that shares homology with the core PxxP-binding motif for Src homology 3 (SH3)-binding domains. SH3-mediated protein-protein interactions are abundant within intracellular signalling networks and regulate diverse protein and cellular functions [474-479]. A CMS-related valine-to-methionine mutations of residue Val18 (red) has been identified from four unrelated CMS patients with severe myasthenia. This mutation is located within the ChAT N-terminal proline-rich motif and has been shown previously to reduce the cellular steady- state protein levels and in vitro catalytic activity of mutant V18M-ChAT [457-458].

Protein-protein interactions between SH3-motifs and SH3-domain-containing proteins are abundant within intracellular signalling networks and have been shown to regulate the subcellular trafficking, phosphorylation, activity, and stability of multiple cellular proteins [475-479]. To date only a handful of protein-protein interaction involving human ChAT has been identified, including the protein kinases PKC and CaMKII and the co-chaperone VCP [438]. As such, it is currently unknown whether ChAT engages in SH3-mediated protein-protein interactions that may alter ChAT function, or furthermore if the ChAT N- terminal proline-rich motif may play a functional role in the regulation of ChAT enzyme activity and/or protein stability.

1.6: Study rationale, aims, and hypothesis

Maintenance of cellular proteostasis represents an intricate, ongoing, and critical process required for the preservation of cellular function and cell survival. To accomplish this, cells utilize a complex network of pro-stabilizing molecular chaperones, such as HSPs, and pro- degradation proteins, such as those involved with the ubiquitin-proteasome system, to efficiently triage cellular proteins between these two opposing cellular fates. In addition, HSPs play a significant role in the protection against cellular proteotoxic stress (e.g. hyperthermia and oxidative stress) by either chaperoning stress-misfolded proteins to promote their folding and stability or, conversely, to triage terminally misfolded proteins for proteolytic degradation, thereby maintaining proteostasis and preventing stress-induced cell death. Chronic exposure to oxidative stress, and the associated dysregulation of molecular chaperones and cellular proteostasis is a common factor in the pathogenesis of multiple human diseases, including cancer and many neurodegenerative diseases such as AD. Work in our laboratory is focused on the regulation of cholinergic neural function during aging as cholinergic neurons appear to be especially sensitive to oxidative stress and AD-related dysfunction. The cholinergic enzyme ChAT is responsible for the synthesis of the neurotransmitter ACh and is essential to cholinergic neural function. Missense mutations found within human 69-kDa ChAT protein have been causatively-linked to the rare inheritable neuromuscular disorder CMS. One CMS-related ChAT mutation, V18M,

reduces both in vitro enzyme activity and cellular steady-state protein levels, and is positioned within a highly-conserved, surface-exposed, N-terminal proline-rich motif at residues 14PKLPVPP20 that share homology with SH3-binding motifs. It is currently unknown what cellular factors regulate ChAT protein stability, and whether the N-terminal proline-rich motif of ChAT regulates ChAT function.

The specific aims of my present study were to two-fold. First, I was to assess the role of the ChAT N-terminus, and in particular the proline-rich motif surrounding the CMS- related residue Val18, in the regulation of ChAT protein function. Second, and in conjunction with my first aim, I was to determine cellular mechanisms involved in the regulation of ChAT protein stability, potentially through alteration of the ChAT N- terminus. Thus, I hypothesized that ChAT protein stability and enzyme activity are regulated by the highly-conserved N-terminal proline-rich motif of ChAT, and furthermore that ChAT protein stability is regulated by molecular chaperones in collaboration with the ubiquitin-proteasome system.

Briefly, to evaluate these hypothesises I addressed the cellular protein expression, enzymatic activity, and the degradation of both wild-type and CMS-mutant ChAT protein in transiently transfected mouse cholinergic SN56 neural cells. To assess the role of the proline-rich motif on ChAT protein function, CMS-related V18M-ChAT was compared to an engineered proline-to-alanine ChAT mutant, P17A/P19A-ChAT. Initial assessment was carried out by anti-ChAT immunoblotting and by radioenzymatic assay to assess cellular ChAT protein levels and activity respectively. Additionally, mechanisms that regulate the degradation of ChAT were determined using the proteasome inhibitor MG132 and the lysosome inhibitor chloroquine, and subsequently ChAT ubiquitination was tested by anti- ubiquitin immunoblotting from anti-ChAT immunoprecipitation protein samples. Together these experiments formed the basis for further experimentation into the regulation of ChAT protein stability and function.

CHAPTER 2: MATERIALS AND METHODS

2.1: Protein expression plasmids

For expression in mammalian cells, cDNA encoding wild-type human 69-kDa ChAT, ligated to pcDNA3.1+ vector [440], was used as a template for preparation of N-terminally truncated, proline-mutant, and CMS-mutant ChAT. Multiple N-terminally truncated ChAT isoforms (1-5, 1-10, 1-15, 1-20 and 1-30) were prepared by PCR amplification and ligated to pcDNA3.1+ vector. Additionally, multiple proline-to-alanine ChAT mutants (P14A, P17A, P19A, P20A and P17A/P19A) and CMS-mutant ChAT (V18M, A513T and T490N) [457-458] were prepared by site-directed mutagenesis according to the QuikChange II Kit (Agilent/Stratagene). For bacterial expression, cDNA encoding wild- type human 69-kDa ChAT, ligated to pProEXHTa vector [480], was used as a template for preparation of P17A-, P19A- and P17A/P19A-ChAT mutants by site-directed mutagenesis. Plasmids encoding wild-type and mutant HA-tagged ubiquitin [481-482] were purchased from Addgene: wild-type (#17608), K0 (#17603), K11 (#22901), K48 (#17605), K48R (#17604), and K63 (#17606). For BioID experiments, pcDNA3.1/MCS-BirA(R118G)-HA (BirA*; Addgene #36047) [483] served as a template for the ligation of PCR-amplified wild-type and P17A/P19A-ChAT cDNA. For the heterologous expression of wild-type and CHIP mutants (ΔTPR and ΔU-box), plasmids were kindly provided by Dr. Yosef Shaul (Weizmann Institute of Science) [484] with permission by Dr. Cam Patterson (UNC School of Medicine). CHIP cDNA was PCR amplified with the addition of a C-terminal FLAG- tag and ligated to pcDNA3.1+ vector. Expression plasmids were validated by full-length DNA sequencing prior to use for mammalian/bacterial expression.

2.2: Cell culture, experimental treatments, and cell lysis

Mouse cholinergic SN56 neural cells (gift from Dr. J.K. Blusztajn, Boston University) [485] or human HEK293 cells (ATCC) were grown as monolayers in DMEM supplemented with 5-10% FBS (Invitrogen) and 1% Pen-Strep at 37C with 5% CO2. Prior

to experimental treatments, cells were transiently transfected with protein expression plasmids (detailed above) by either calcium phosphate [486] or using Lipofectamine 2000/3000 (Invitrogen) at ~50% confluence for 18-24 h at 37C. All laboratory procedures were conducted with Biosafety Approval BIO-RRI-0022.

For determination of ChAT steady-state protein levels and cellular ChAT activity of wild-type or mutant ChAT, ChAT-expressing SN56 cells were collected and lysed on ice in ChAT assay lysis buffer (10 mM Na2HPO4; pH 7.4, 0.87 mM EDTA, 0.5% Triton X- 100 and 1.5 mM eserine sulphate) supplemented with mammalian protease inhibitor cocktail (Sigma), phosphatase inhibitor cocktail (10 mM NaF, 1 mM Na3VO4, 20 mM

Na2HPO4, 3 mM -glycerolphosphate, 5 mM sodium pyrophosphate), and 50 µM MG132 (Enzo Life Sciences). Lysates were centrifuged for 10 min at 21,000 g at 4C and protein concentrations were measured by Bradford Protein Assay (Bio-Rad). Lysate supernatants were prepared for analysis of ChAT protein levels by denaturing aliquots of cell lysate in 1x Laemmli sample buffer (63 mM Tris-HCl; pH 6.8, 10% glycerol, 2% SDS, 0.005% bromophenol blue, 2.5% 2-mercaptoethanol) at 95C for 10 min followed by anti-ChAT immunoblotting. Additionally, cell lysates were used to determine cellular ChAT activity by modified radioenzymatic assay [487-488]. To investigate the effect of proteasome and/or lysosome inhibitors on ChAT steady-state protein levels and cellular ChAT activity, SN56 cells transiently expressing wild-type, P17A/P19A-ChAT, or CMS-mutant ChAT were treated with 5 µM MG132, 25 µM chloroquine (Sigma), or vehicle (DMSO or water respectively) for 18 h at 37C prior to immunoblotting or radioenzymatic assay.

The impact of inhibition of HSC/HSP70, HSP90, or p97/VCP function on ChAT steady-state protein levels was determined in SN56 cells expressing either wild-type or mutant ChAT proteins that were treated with varying concentrations of the HSC/HSP70 inhibitor VER-155008 (5 – 50 µM; Sigma), the HSP90 inhibitor 17-AAG (0.5 – 2 µM; StressMarq Biosciences), or the p97/VCP inhibitor Eeyarestatin-I (Eer1; 5 – 10 µM; Sigma) for 18 – 24 h at 37°C. Cells were collected and lysed on ice in RIPA buffer (50 mM Tris-HCl; pH 8.0, 150 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS) supplemented with protease/phosphatase inhibitors (above), 50 µM MG132, 10 mM N-

ethylmaleimide (NEM; Calbiochem), and 800 U/ml DNase I (Invitrogen). Lysates were centrifuged for 10 min at 21,000 g at 4C, protein concentrations were measured by BCA protein assay (Thermo), and lysate supernatants were denatured in 1x Laemmli sample buffer at 95C for 10 min then analyzed by anti-ChAT immunoblotting.

To measure cellular ChAT activity following HSC/HSP70 and/or HSP90 inhibition SN56 cells expressing wild-type or mutant ChAT proteins were treated with either 40 µM VER-155008 or 1 µM 17-AAG either alone or in combination at 37°C for 24 h. Cells were collected and lysed on ice in ChAT assay lysis buffer (above) supplemented with protease/phosphatase inhibitors and 50 µM MG132. Lysates were centrifuged for 10 min at 21,000 g at 4C and used for analysis of cellular ChAT activity as described above.

The effects of either proteasome or lysosome inhibition on ChAT steady-state protein levels during HSC/HSP70 and HSP90 inhibition were investigated in SN56 cells expressing wild-type or mutant ChAT. Cells were treated either alone with 40 µM VER- 155008 or 1 µM 17-AAG at 37°C for 24 h or co-treated for 18 h with either 5 µM MG132 or 50 µM chloroquine. To lyse cells and release aggregated proteins following HSP and proteasome inhibition, cells were boiled at 95°C for 5 min and sonicated in 1% SDS lysis buffer (50 mM Tris-HCl; pH 8.0, 150 mM NaCl, 1% SDS) supplemented with protease/phosphatase inhibitors, 50 µM MG132 and 10 mM NEM. Protein samples were prepared for anti-ChAT immunoblotting as above.

For ubiquitination studies, SN56 cells were transiently transfected to express wild- type, P17A/P19A-ChAT or CMS-mutant ChAT alone, or in combination, with HA-tagged wild-type, K0, K11, K48, K48R or K63-ubiquitin, then treated with either 5 µM MG132, 2.5 µM lactacystin (EMD), or 25 µM chloroquine for 18 h at 37C. For oxidative stress studies SN56 cells transiently co-expressing wild-type ChAT with HA-ubiquitin were pre- treated for 18 h with 5 µM MG132 then co-treated for an additional 30 or 60 min with 1 mM H2O2. Alternatively, SN56 cells transfected to express either wild-type or mutant ChAT were treated at 37°C with 40 µM VER-155008 for 24 h, 1 µM 17-AAG for either 8 or 24 h, or 10 µM Eer1 for 18 h to inhibit HSC/HSP70, HSP90, or p97/VCP respectively. To block degradation of ubiquitinated ChAT following HSP inhibition cells were co-treated

with 20 µM MG132 for 6 h prior to being collected; MG132 co-treatment was omitted when cells were treated with Eer1. Cells were collected and lysed on ice in RIPA buffer (above) or in RIPA buffer supplemented with 0.5% SDS. Lysates were sonicated to shear DNA, centrifuged for 10 min at 21,000 g at 4C, then either anti-HA or anti-ChAT immunoprecipitations were completed and analyzed by immunoblotting.

2.3: Immunoprecipitation (IP) to assess ChAT ubiquitination

For anti-HA IPs, 1 mg aliquots of cleared whole cell lysate were diluted to a final volume of 1 ml in supplemented RIPA buffer (above), pre-cleared by incubation with 30 µl of anti- mouse IgG-agarose beads (Sigma) for 1 h at 4C, then HA-tagged ubiquitinated proteins were immunocaptured onto 25 µl of monoclonal anti-HA-agarose beads (Sigma) for 3 h at 4C. For anti-ChAT IPs, 2-4 mg aliquots of cleared whole cell lysate were diluted to a final volume of 1 ml in supplemented RIPA buffer, incubated with 25 µg/mg cellular protein of anti-ChAT primary antibody (CTab) [440] at 4C for 18 h, then immune complexes were captured onto 50 µl of protein-G Dynabeads (Invitrogen) for 1 h at 4C. For both anti-HA and anti-ChAT IPs, immunocaptured samples were washed 4-6 times with ice-cold RIPA buffer and immune complexes were eluted into 50 µl of 2x Laemmli sample buffer (above) by heating samples at 85C for 15 min with intermittent mixing. IP samples were analyzed by immunoblotting with anti-ChAT, anti-HA, and anti-ubiquitin primary antibodies. Total protein samples were prepared by denaturing whole cell lysates in 1x Laemmli sample buffer at 95C for 5-10 min prior to immunoblotting.

2.4: Co-immunoprecipitation (co-IP) of ChAT with CHIP or HSPs

For co-IP of ChAT with endogenous HSPs, endogenous CHIP, or heterologous FLAG- tagged CHIP either HEK293 or SN56 cells were transfected to transiently express either wild-type or mutant ChAT protein alone or to be co-expressed with either FLAG-CHIP or with mutant FLAG-CHIP-ΔTPR or FLAG-CHIP-ΔU-box. For co-IP of ChAT with

endogenous HSPs following oxidative stress, SN56 cells transiently expressing wild-type

ChAT were treated with either 100 µM H2O2 for 1 – 8 h or with 1 mM H2O2 for 1 h at 37°C. Cells were collected on ice and lysed in co-IP lysis buffer (10 mM HEPES; pH 7.4,

5 mM MgCl2, 1 mM EGTA, 50 mM NaCl, 5% glycerol, 0.5% Triton X-100) supplemented with protease/phosphatase inhibitors, 50 µM MG132, and 10 mM NEM. Lysates were centrifuged for 15 min at 15,000 g at 4C, protein concentrations were measured by Bradford protein assay, then aliquots of cleared whole cell lysates containing either 2 mg (FLAG-CHIP) or 4 mg (endogenous proteins) of protein were incubated with CTab primary antibody at 4C for 18 h. Immune complexes were captured onto 50 µl of protein-G Dynabeads for 1 h at 4C, washed 4-6 times in co-IP lysis buffer, then eluted into 50 µl of 2x Laemmli sample buffer by heating samples at 85C for 15 min with intermittent mixing. Co-IP samples were analyzed by immunoblotting. Total protein samples were prepared by denaturing whole cell lysates in 1x Laemmli sample buffer at 95C for 5-10 min.

2.5: SDS-PAGE and immunoblotting analysis

Denatured protein samples from whole cell lysates and IP/co-IPs were resolved on 7.5, 10, 12, or 15% SDS-PAGE gels, then transferred to PVDF membranes (Bio-Rad) by semi-dry or wet-tank electroblotting. For immunoblotting, membranes were blocked for 1 h at 21°C in 5% non-fat milk powder in PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4; pH 7.4) containing 0.15% Triton X-100 (PBST), followed by incubation overnight at 4C with primary antibody. Probed membranes were washed with PBST, and primary antibodies were detected using 1:10,000 peroxidase-coupled secondary antibodies (Jackson ImmunoResearch) and Clarity Western ECL Substrate (Bio-Rad) on a ChemiDoc MP system (Bio-Rad). The following primary antibodies were used: 1:1000 ChAT (CTab) [440], either 1:1000 actin (Santa Cruz) or 1:10,000 β-actin (Sigma), 1:10,000 GAPDH (Cell Signalling), 1:1000 HSC70 (StressMarq), 1:2000 HSP70 (Thermo), 1:2000 HSP90 (StressMarq), 1:1000 CHIP (Santa Cruz), 1:500 HA (Roche), 1:1000 ubiquitin (Santa Cruz), 1:1000 FLAG-M2 (Sigma), and 1:1000 LC3B (Thermo). For anti-CHIP immunoblotting from anti-ChAT co-IP samples, 1:1000 VeriBlot IP secondary antibody

(Abcam) was used as a substitute for standard secondary antibodies to reduce IgG light- chain interference of endogenous 35-kDa CHIP detection.

2.6: Purification of recombinant ChAT

Chemically competent E. coli BL21 (DE3) cells were heat shock transformed at 42°C with pProEXHTa expression plasmids encoding N-terminal His6-tagged wild-type, P17A-, P19A- or P17A/P19A-ChAT and grown in auto-induction media (1% tryptone, 0.5% yeast extract, 50 mM Na2HPO4, 50 mM KH2PO4, 25 mM (NH4)2SO4, 0.5% glycerol, 0.05% glucose, and 0.2% lactose) at 18C on a shaker (225 rpm) for 48 h until OD600 = ~8.0. Cells were pelleted by centrifugation at 5,000 g for 20 min at 4C, resuspended in Ni2+-column buffer (50 mM Na2HPO4; pH 7.5, 1.0 M KCl, 10% glycerol) supplemented with 50 mM imidazole. Purification of His6-tagged ChAT was completed by Ni2+-affinity purification with follow-up cation exchange chromatography over either Ni2+-Chelating Sepharose Fast Flow or SP-Sepharose Fast Flow (GE Healthcare), respectively, as performed previously in our laboratory [480]. Following cation exchange chromatography, protein samples were analyzed on Coomassie-stained SDS-PAGE gels and eluent fractions containing purified recombinant ChAT were pooled, flash-frozen in liquid nitrogen, and stored at -80C. Final protein concentrations of pooled fractions containing purified recombinant ChAT protein were determined using a Nanodrop UV spectrometer (Thermo) with Abs0.1% = 0.789 at 280 nm. Samples of purified recombinant ChAT were denatured in 1x Laemmli sample buffer at 95C for 5 min, resolved on 10% SDS-PAGE gels, stained with Imperial Protein Stain (Thermo), and imaged using a ChemiDoc MP system. In vitro specific activity of purified recombinant ChAT proteins was assessed by radioenzymatic assay [487-488].

2.7: Circular dichroism (CD) analysis and thermal denaturation

Purified recombinant wild-type, P17A-, P17A/P19A- and P19A-ChAT were dialyzed overnight at 4C against cation exchange buffer without NaN3 (10 mM Na2HPO4; pH 7.0,

10% glycerol, 0.5 mM tris(2-carboxyethyl)phosphine hydrochloride (TCEP; Sigma)).

Proteins were diluted to 0.2 mg/ml in cation exchange buffer without NaN3 and CD spectra were generated using a Jasco J-810 spectropolarimeter at 4C in a 0.1 cm path length cuvette over a wavelength range of 195-260 nm at 0.5 nm intervals. A total of 20 reads were averaged per recombinant ChAT protein, and CD spectra were finalized by correcting for background buffer spectra and calculating mean residue ellipticity (MRE). Following

CD, the thermal denaturation midpoint (Tm) was determined for each recombinant ChAT protein by measuring the loss of helicity at 222 nm over a 4-95C range at 0.5C intervals.

Tm was calculated as previously described [489].

2.8: Strain-promoted alkyne-azide cycloaddition (SPAAC) pulse-chase

SN56 cells grown to ~90% confluence on 100 mm dishes were transfected by Lipofectamine 2000 for 6 h at 37C to mediate transient expression of wild-type or P17A/P19A-ChAT, then trypsinized and re-plated onto 60 mm dishes and grown overnight. SPAAC pulse-chase was initiated by first live-labeling (pulse) cells at 37C for 4 h in methionine-free DMEM (Invitrogen) supplemented with 5% dialyzed FBS (Invitrogen) and 50 μM Click-iT L-azidohomoalanine (AHA; Invitrogen). Control cells were grown for 4 h in methionine-containing DMEM with 5% FBS. AHA-labeled cells were then washed once with and subsequently incubated in methionine-containing DMEM with 5% FBS for 2 - 24 h at 37C (chase). For studies of ChAT protein half-life during proteasome inhibition cells were treated with 5 µM MG132 throughout both the 4 h pulse and 24 h chase period. Cells were collected on ice either immediately following the pulse period (control and 0 h) or following the chase periods (2, 4, 10, and 24 h). For fluorescent labeling of AHA-labeled proteins from whole cell lysates, collected cells were lysed on ice in 300 µl RIPA buffer supplemented with protease/phosphatase inhibitors, 50 µM MG132, 10 mM NEM, and 10 mM of freshly prepared iodoacetamide. Lysates were centrifuged for 10 min at 21,000 g at 4C and 50 µl aliquots of whole cell lysate were fluorescently-labelled by incubating with 10 μM Click-iT TAMRA-DIBO (Invitrogen) for 1 h at 21°C followed by denaturation in 1x Laemmli sample buffer at 95C for 5 min. For analysis, equal amounts of proteins (e.g.

25 µg) from each experimental sample were resolved on 7.5% SDS-PAGE gels and fluorescence was detected in-gel using a ChemiDoc MP system at an excitation/emission of 555/580 nm. Additionally, for downstream analysis anti-ChAT immunoblotting of AHA/TAMRA-labeled protein samples from whole cell lysates was also completed.

To determine ChAT protein half-life, anti-ChAT IPs were prepared from AHA- labeled cells by diluting 200 µl of cleared whole cell lysate, as prepared above, with 300 µl of supplemented RIPA buffer. Diluted lysates were incubated with 25 µg of anti-ChAT CTab primary antibody at 4C for 3 h followed by immunocapture onto 50 µl of protein-G Dynabeads for 1 h at 4C. Immunocaptured samples were washed four times with RIPA buffer, once with PBS to remove detergents, then subsequently reacted with 10 μM Click- iT TAMRA-DIBO in 1x PBS for 1 h at 21°C with gentle agitation. Samples were washed twice with RIPA buffer to remove excess TAMRA-DIBO and AHA/TAMRA-labeled ChAT was eluted into 50 µl of 2x Laemmli sample buffer at 85C for 15 min with intermittent mixing. For analysis, equal volumes of immunoprecipitated AHA/TAMRA- labeled ChAT (e.g. 20 µl) were resolved on 7.5% SDS-PAGE gels and in-gel fluorescence was detected as above. Additionally, for downstream analysis anti-ChAT immunoblotting of AHA/TAMRA-labeled protein samples from anti-ChAT IPs was also completed.

To calculate ChAT protein half-life, I assumed that the amount of fluorescent AHA/TAMRA-labeled ChAT, P(t), decays exponentially under first order kinetics

-t according to the equation, P(t) = Poe , where Po is the fluorescence intensity at t = 0. The slope of decay () was calculated by plotting fluorescence intensity of immunoprecipitated AHA/TAMRA-labeled ChAT, corrected for the levels of total immunoprecipitated ChAT as measured by parallel anti-ChAT immunoblotting, on a semi-logarithmic scale and performing linear regression. ChAT protein half-life, T1/2, was calculated according to first order kinetics where T1/2 = ln(2)/ [490].

Validation of SPAAC pulse-chase was performed using a cycloheximide (CHX)- chase assay. SN56 cells were transfected and plated as for SPAAC pulse-chase to transiently express wild-type or P17A/P19A-ChAT. Cells were treated with either 100 µg/ml CHX for 2, 4, 6 or 8 h, or with DMSO-control for 8 h. Cells were collected and lysed

on ice in supplemented RIPA buffer, lysates were centrifuged for 10 min at 21,000 g at 4C, and protein samples were prepared for anti-ChAT immunoblotting by denaturing cell lysates in 1x Laemmli sample buffer at 95C for 5 min. ChAT immunoreactive bands were quantified by densitometry, plotted on semi-logarithmic scale, and analyzed as above by linear regression to determine a slope of decay for wild-type and P17A/P19A-ChAT.

2.9: Soluble/insoluble Triton X-100 cellular fractionation and semi- denaturing detergent agarose gel electrophoresis (SDD-AGE)

ChAT protein solubility was assessed in SN56 cells, transiently expressing either wild-type or mutant ChAT following treatment with 20 µM 2-phenylethynesulfonamide (PES; Sigma), a HSC/HSP70 inhibitor, at 37°C for 24 h. Alternatively, ChAT-expressing SN56 cells were treated with varying concentrations of H2O2 for either 24 h (100 nM – 200 µM) or for 1 h (1 µM – 2 mM) alone or were pre-treated with 20 µM PES for 24 h followed by co-treatment with 100 µM H2O2 for 1 – 8 h at 37°C. Cells were collected and lysed on ice in 0.1% Triton X-100 lysis buffer (50 mM Tris-HCL; pH 8.0, 150 mM NaCl, 0.1% Triton X-100) supplemented with protease/phosphatase inhibitors, 50 µM MG132 and 10 mM NEM. Lysates were centrifuged for 15 min at 15,000 g at 4C and aliquots of Triton-soluble supernatant were prepared for immunoblotting by denaturing in 1x Laemmli sample buffer at 95C for 10 min. Triton-insoluble proteins were prepared for immunoblotting by washing Triton-insoluble pellets once with ice-cold PBS, then denaturing the pellets in an equal volume of 2x Laemmli sample buffer with 5% 2-mercaptoethanol at 85C for 15 min prior to separation on SDS-PAGE gels. For analysis of ChAT solubility following co- expression of either FLAG-CHIP or with mutant FLAG-CHIP-ΔTPR or FLAG-CHIP-ΔU- box, cells were lysed in 0.5% Triton X-100 co-IP lysis buffer (above) and Triton- soluble/insoluble protein samples were prepared as above.

In some cases, following PES treatment Triton-insoluble proteins were further analyzed in parallel by semi-denaturing detergent agarose gel electrophoresis (SDD-AGE) [491] to determine if PES treatment results in the generation of SDS-resistant ChAT

aggregates. Briefly, Triton-insoluble protein samples were loaded onto 1.8% agarose gels containing 0.5% SDS and resolved by electrophoresis at 80V for ~2 h in 1x TAE buffer (40 mM Tris-HCl; pH 7.6, 20 mM acetic acid, 1 mM EDTA) supplemented with 0.1% SDS. Resolved proteins were transferred to PVDF membranes by overnight capillary transfer in 1x TBS (50 mM Tris-HCl; pH 7.6, 150 mM NaCl) with 10% methanol at 21°C. Membranes were blocked for 1 h at 21°C in 5% non-fat milk powder in PBST then immunoblotted for ChAT protein as described above.

2.10: Proximity-dependent biotin identification (BioID)

HEK293 cells expressing wild-type- or P17A/P19A-ChAT-BirA* fusion proteins were live-labeled with 50 µM biotin at 37°C for 24 h to allow proximity-dependent biotinylation of cellular proteins. Controls cells were transfected with either empty vector or plasmids encoding untagged ChAT or BirA* [483]. Cells were washed three times on ice with PBS to remove excess biotin, lysed in supplemented RIPA buffer (above), centrifuged for 10 min at 21,000 g at 4C, then aliquots of cleared whole cell lysate containing 5 mg protein were incubated with 50 µl of MyOne Streptavidin C1 Dynabeads (Invitrogen) at 4°C for 18 h. Streptavidin pull-down (PD) samples were washed once with 1% SDS lysis buffer (50 mM Tris-HCl; pH 8.0, 150 mM NaCl, 1% SDS), 3-4 times with RIPA buffer, then biotinylated proteins were eluted into 50 µl of 2x Laemmli sample buffer at 85C for 15 min. Biotinylated proteins were resolved on SDS-PAGE gels, transferred to PVDF membranes, and membranes were blocked overnight at 4°C in PBST with 5% BSA prior to incubation with, and subsequent blotting using 1:20,000 Pierce High Sensitivity Streptavidin-HRP (Thermo) for 1 h at 21°C. Total protein samples were prepared by denaturing whole cell lysates in 1x Laemmli sample buffer at 95C for 10 min prior to immunoblotting. For large-scale BioID to identify novel ChAT protein-interactors, HEK293 cells expressing either untagged ChAT (control) or ChAT-BirA* fusion proteins were live labeled with 50 µM biotin then collected, washed, lysed and centrifuged as above. Aliquots of cleared whole cell lysate containing 25 mg protein were incubated with 250 µl of MyOne Streptavidin C1 Dynabeads at 4°C for 18 h. Streptavidin PD samples were

washed as described above, eluted into 100 µl of 2x Laemmli sample buffer, and resolved on 10% SDS-PAGE gels. To visualize biotinylated proteins, the SDS-PAGE gel was stained using Pierce Silver Stain for Mass Spectrometry (Thermo).

2.11: Mass spectrometry

Following large-scale BioID proteins of interest were excised from silver-stained SDS- PAGE gels using an Ettan Spot Picker (GE Healthcare Life Sciences). In-gel digestion was performed at the London Regional Proteomics Center - Functional Proteomics Facility (FPF; Western University) using a MassPREP automated digester station (PerkinElmer) where gel pieces were first de-stained (50 mM Na2S2O3 and 15 mM K3Fe(CN)6), followed by protein reduction (10 mM dithiothreitol), alkylation (55 mM iodoacetamide), and tryptic digestion. Digested peptides were extracted in 1% formic acid with 2% acetonitrile, lyophilized, then dissolved in 10% acetonitrile containing 0.1% trifluoroacetic acid prior to mass spectrometry by either matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS) or by liquid chromatography-electrospray ionization- tandem mass spectrometry (LC-ESI-MS/MS).

For MALDI-TOF-MS tryptic-digested peptides were mixed 1:1 (v/v) with a 5 mg/mL –cyano-4–hydroxycinnamic acid MALDI matrix that was prepared in 6 mM

NH4H2PO4, 50% acetonitrile, and 0.1% trifluoroacetic acid, then submitted to the London Regional Proteomics Center - FPF for analysis. Mass spectrum were obtained using a TOF/TOF 5800 System (Applied Biosystems, Sciex) equipped with a 349 nm OptiBeam On-Axis laser using a laser pulse-rate of 400 Hz. Reflectron positive mode was used and was externally calibrated at 50 ppm mass tolerance and internally at 10 ppm. Each mass spectrum was collected as a sum of 500 shots. Data acquisition and processing were completed using a TOF/TOF Series Explorer and Data Explorer respectively (Applied Biosystems, Sciex). The resulting data were analyzed using the online Mascot Server (Matrix Science) against the NCBI Human protein database with the following selected as

variable modifications; biotin (K), carbamidomethyl (C), NEM (C), oxidation (M). Peptide tolerance was set at ± 50 ppm for protein identification.

For LC-ESI-MS/MS, tryptic-digested peptides were submitted for protein identification by the London Regional Proteomics Center – Biological Mass Spectrometry Laboratory (BMSL; Western University). Briefly, digested peptides were injected into and analyzed using a LC-ESI-MS/MS system comprised of a nanoAcquity UPLC (Waters) followed by an in-line QExactive Hybrid Quadrupole-Orbitrap mass spectrometer (Thermo) using a top-12 double-play method. For LC, peptides were trapped onto a reverse-phase Acquity Symmetry C18 column, 5 µm, 180 µm x 200 mm (Waters), and subsequently were separated/eluted using a H2O-acetonitrile with 0.1% formic acid solvent gradient applied to a Acquity Peptide BEH C18 column, 1.7 um, 75 um x 250 mm (Waters). Resulting data were analyzed using Peaks 7.5 software (Bioinformatics Solutions Inc.) against the NCBI Human protein database allowing for variable carbamidomethylation or modification of cysteine residues with NEM with filter criteria set to FDR = 1% and UP = 1 (unique protein) to show protein identification.

2.12: In situ proximity ligation assay (PLA)

To evaluate in situ ChAT protein-interactors, SN56 cells were plated on 35 mm glass- bottom confocal dishes and transiently transfected to express wild-type 69-kDa ChAT. Cells were washed with HBSS, formalin-fixed (4% paraformaldehyde in HBSS) for 15 min, permeabilized with 0.1% Triton X-100, blocked for 1 h in HBSS supplemented with 3% donkey serum, then finally incubated for 1 h with primary antibodies targeting ChAT (1:100; Chemicon, goat primary) together with either endogenous HSC70 (1:100; StressMarq, mouse primary), HSP90 (1:200; StressMarq, mouse primary) or CHIP (1:200; Santa Cruz, rabbit primary); all steps were performed at 21°C. Controls cells were transfected with empty vector or were not incubated with primary antibodies. DuoLink PLA technology (Sigma) was utilized to visualize in situ interactions by incubating cells with Duolink oligonucleotide-linked secondary antibodies (anti-goat PLUS with either

anti-mouse MINUS or anti-rabbit MINUS) for 1 h at 37°C. Cells were washed with Duolink In Situ Wash Buffer A, then incubated with 1 U/µL DNA ligase for 30 min followed by 10 U/µL DNA polymerase for 2 h at 37°C according to manufacturer’s instructions. Finally, cells were washed with Duolink In Situ Wash Buffer B and were overlaid with Duolink In Situ Mounting Medium with DAPI. Images were acquired using a Leica True Confocal Scanner (TCS) 8 SpectroPhotometer (SP8; Leica Microsystems Inc.) confocal laser microscope with a 63x-magnification oil-immersion objective (NA = 1.4) and processed using ImageJ 1.50b software (NIH).

2.13: Small-interfering RNA (siRNA)

SN56 cells were incubated for a total of 72 h at 37°C with 25 nM of siRNA targeting either mouse Stub1/CHIP (Silencer Select ID: s80537; Thermo) or mouse HSP70 (sc-35605; Santa Cruz) following transfection with siLentFect Lipid Reagent (Bio-Rad). As a control, cells were either mock transfected with siLentFect alone or transfected with 25 nM of scramble-control siRNA (Silencer Select #4390843; Thermo). After 24 h of endogenous CHIP/HSP70 knock-down, cells were transfected with Lipofectamine 3000 to transiently express either heterologous wild-type or mutant ChAT for an additional 48 h at 37°C. Additionally, cells incubated with anti-HSP70 siRNA were treated with 1 µM 17-AAG for the final 24 h of HSP70 knock-down at 37°C. Cells were then collected and lysed on ice in RIPA buffer containing 0.5% SDS, protease/phosphatase inhibitors, 50 µM MG132 and 10 mM NEM. Lysates were sonicated, centrifuged for 10 min at 21,000 g at 4C, denatured in 1x Laemmli sample buffer at 95C for 10 min, then analyzed by immunoblotting.

2.14: Statistical analysis

For statistical analysis of steady-state proteins levels from immunoblots prepared using whole cell lysates, immunoreactive protein levels were measured by densitometry using ImageLab 5.0 software (Bio-Rad), normalized to either β-actin or GAPDH protein levels,

and graphed as mean ± SEM from individual independent replicate experiments (n). For analysis of anti-ChAT co-IP immunoblots, immunoreactive bands for each co- immunoprecipitated protein (i.e. wild-type and mutant ChAT proteins, HSC70/HSP70, HSP90 or FLAG-CHIP) were separately measured by densitometry then internally normalized to the total densitometric value for that protein from within their respective immunoblots to obtain the relative levels of each co-immunoprecipitated protein. These relative levels of HSPs or FLAG-ChAT were then expressed as a function of the relative level of co-immunoprecipitated ChAT, and data was graphed as mean ± SEM from n number of independent replicate co-IP experiments. Statistical analysis for experiments completed within this thesis was performed by Student’s t-test, one-way ANOVA with either Tukey’s or Dunnett’s post-hoc test, or by two-way ANOVA with Bonferroni’s post- hoc test using GraphPad Prism software. Statistical significance was set at p≤0.05.

CHAPTER 3: RESULTS

3.1: ChAT mutations reduce steady-state protein levels and cellular enzymatic activity of ChAT

Multiple missense mutations have been identified in the human 69-kDa ChAT protein that have a causative association with development of CMS, with several of these mutations reducing both the in vitro enzyme activity of purified recombinant ChAT and the cellular steady-state protein levels of mutant ChAT when expressed in COS, BOSC 23, or HEK293 cells (Figure 1.9) [411, 457-458]. Two such catalytically-deficient mutations, V18M and A513T, are positioned distal from the ChAT active site and reduce ChAT protein levels and enzymatic activity through unknown mechanism [457]. As these earlier studies were performed in non-neural cells, it is currently unknown if these CMS-related mutations also reduce ChAT steady-state protein levels in cholinergic cells. Thus, I generated expression plasmids encoding for three different catalytically-deficient CMS-related ChAT mutations (V18M, A513T, and T490N) that were then transiently expressed in mouse cholinergic SN56 neural cells; T490N-ChAT differs from V18M and A513T as this mutation does not reduce steady-state ChAT protein levels when expressed in human kidney BOSC 23 cells [457]. Following anti-ChAT immunoblotting (Figure 3.1a) I confirmed that the steady-state protein levels of both V18M- (p≤0.001) and A513T-ChAT (p≤0.05) are reduced compared to wild-type ChAT in cholinergic SN56 cells, whereas the protein levels of T490N-ChAT were unaffected as anticipated (Figure 3.1b). As introduced earlier, residue Val18 is located within a highly-conserved and surface-exposed N-terminal proline-rich motif at residues 14PKLPVPP20 that shares homology with the core PxxP-binding motif for SH3-domain- containg proteins (Figs 1.10 and 1.11) [420, 474]. Therefore, to test whether residues within this proline-rich motif beyond residue Val18 may regulate ChAT steady-state protein levels, I also generated an expression plasmid encoding for ChAT with proline-to-alanine mutation of residues Pro17 together with Pro19 (P17A/P19A). When transiently expressed in SN56 and analyzed by anti-ChAT immunoblotting, I observed that the steady-state protein levels of P17A/P19A-ChAT is also reduced compared to wild-type ChAT (p≤0.001; Figure 3.1).

Figure 3.1: CMS-related ChAT mutants demonstrate reduced steady-state protein levels. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type (WT), P17A/P19A-, or CMS-related mutant ChAT (V18M-, A513T-, and T490N-ChAT). Control cells were transfected with empty pcDNA3.1+ vector. Anti-actin immunoblotting was completed as a loading control. (B) Steady-state protein levels of P17A/P19A- (***p≤0.001), V18M- (***p≤0.001), or A513T-ChAT (*p≤0.05) are reduced compared to wild-type ChAT (one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=5). In agreement with previously reported results from human kidney BOSC 23 cells transiently expressing mutant T490N-ChAT [457], no significant changes in the steady-state protein levels of T490N-ChAT were observed in SN56 cells as compared to wild-type ChAT.

Both the P17A/P19A and CMS-related V18M mutations suggest a critical role for the ChAT N-terminus in regulating ChAT protein function. The crystal structure of human 69-kDa ChAT suggests that residues Met1 to Ser11 are unstructured, while Gly12 through Gln30 are structured and surface-exposed [420]. Therefore, to first locate which regions within the ChAT N-terminus are functionally critical I generated expression plasmids encoding for ChAT with progressive truncation of the N-terminus up to and including residue Gln30 (1-5, 1-10, 1-15, 1-20 and 1-30). These N-terminally truncated ChAT mutants were then transiently expressed in SN56 cells and cell lysates were analyzed for

both steady-state protein levels, by anti-ChAT immunoblotting, and for cellular ChAT activity. When compared to wild-type ChAT, I found that truncation of the ChAT N- terminus up to and including residue Glu10 (1-5 and Δ1-10) had no effect on either ChAT steady-state protein levels (Figures 3.2a and 3.2b) or cellular ChAT activity (Figure 3.2c). However, deletion of residues including Lys15 and up-to Gln30 (1-15, 1-20 and 1-30) dramatically reduced steady-state protein levels of these truncated ChAT mutants in SN56 cells and abolished their cellular catalytic activity (p≤0.001).

Residue Lys15, like Val18, is located within the highly-conserved N-terminal proline- rich motif of ChAT the encompasses residues 14PKLPVPP20. Proline residues impart structural rigidity to polypeptides, and proline-rich motifs are often involved in cellular signalling processes [474, 492-493]. On this basis, I further investigated the conserved N- terminal proline-rich motif to find whether mutation of this motif beyond the P17A/P19A and CMS-related V18M mutations might also affect ChAT activity and cellular ChAT protein levels. Plasmids encoding for ChAT with various N-terminal proline-to-alanine mutations (P14A, P17A, P19A, and P20A) were generated and subsequently assessed in SN56 cells alongside of P17A/P19A-ChAT. As illustrated in Figures 3.3a and 3.3b, mutation of any proline residue within this proline-rich motif reduced ChAT steady-state protein levels when compared to wild-type ChAT, with P17A/P19A-ChAT yielding the greatest reduction (80%; p≤0.01). Moreover, mutation of these proline residues reduces cellular ChAT activity, with P17A/P19A-ChAT being catalytically inactive (p≤0.001; Figure 3.3c). Interestingly, though statistical differences in the steady-state protein levels between individual proline mutants were not observed, mutation of residue Pro17 alone did result in a greater reduction in cellular ChAT activity when compared to mutation of either residues Pro14 (p≤0.05) or Pro20 (p≤0.01) alone. Collectively, these results, by disruption, removal, or mutation of the ChAT N-terminal proline-rich motif, suggest a critical role for this motif on ChAT function.

Figure 3.2: N-terminal truncation of ChAT reduces ChAT steady-state proteins levels and cellular ChAT activity. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing either full-length wild-type (WT) or N-terminally truncated ChAT mutants. Control cells were transfected with empty pcDNA3.1+ vector. Anti-actin immunoblotting was completed as a loading control. Truncation up-to and including residue Glu10 (1-5 and Δ1-10) had no effect on either steady-state protein levels (B) or cellular ChAT activity (C) when compared to wild-type ChAT. Conversely, truncation of residues including Lys15 and up-to Gln30 (1-15, 1-20 and 1-30) reduces steady-state protein levels and cellular ChAT activity compared to wild-type ChAT (***p≤0.001, one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=3).

Figure 3.3: Mutation of conserved N-terminal proline-rich motif of ChAT reduces ChAT steady-state proteins levels and cellular ChAT activity. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type (WT) or various proline-to- alanine ChAT mutants. Control cells were transfected with empty pcDNA3.1+ vector. Anti-actin immunoblotting was completed as a loading control. (B) Proline-to-alanine mutation reduces the steady-state protein levels of proline-mutant ChAT compared to wild- type ChAT with P17A/P19A-ChAT yielding the greatest reduction (**p≤0.01, one-way ANOVA with Tukey’s post-hoc test, mean ± SEM, n=5). (C) Proline-to-alanine mutation reduces cellular ChAT activity of proline-mutant ChAT compared to wild-type ChAT with P17A/P19A-ChAT yielding the greatest reduction (***p≤0.001). Cellular activity of mutant P17A-ChAT is reduced compared to either P14A- (*p≤0.05) or P20A-ChAT (**p≤0.01, one-way ANOVA with Tukey’s post-hoc test, mean ± SEM, n=5).

3.2: In vitro characterization of recombinant proline-mutant ChAT

The introduction of proline-to-alanine mutations within the N-terminal proline-rich motif of ChAT could potentially alter ChAT protein secondary structure, thereby resulting in reduced enzyme activity, gross structural instability, protein misfolding, and increased protein degradation. To determine the structural stability of mutant ChAT, I purified bacterially-expressed recombinant wild-type and proline-mutant ChAT proteins (Figure 3.4). I made the important observation that, in contrast to my results in mouse cholinergic SN56 cells, expression levels of proline-mutant ChAT proteins (P17A-, P17A/P19A-, and P19A-ChAT) in E. coli BL21 cells did not differ from that of wild-type ChAT protein (data not shown). Moreover, the proline mutants behaved similarly to wild-type ChAT during purification, yielding stable and pure recombinant protein preparations each with an approximate molecular mass of 65-kDa in agreement with previously published results from our laboratory (Figure 3.4a) [480]. Using these recombinant ChAT proteins, I first determined whether proline-mutant ChAT displayed reduced in vitro enzymatic activity similar to that observed in cells. One consideration when measuring ChAT activity from cell lysates is that both ChAT protein expression and innate enzymatic activity factor into these measurements, and therefore loss of cellular ChAT protein, such as that observed for proline-mutant ChAT (Figures 3.2 and 3.3), can synergize with catalytic defects. Thus, using equal amounts of purified recombinant wild-type and proline-mutant ChAT, I measured the in vitro specific activity of recombinant ChAT by radioenzymatic assay (Figure 3.4b). As anticipated, proline-to-alanine mutation of residue Pro17 either alone or in combination with Pro19 significantly reduced in vitro specific activity of these proline- mutant ChAT proteins when compared to wild-type ChAT, with P17A/P19A-ChAT yielding the greatest reduction (76%; p≤0.05). Surprisingly, unlike in SN56 cells, the in vitro specific activity of P19A-ChAT did not differ from that of wild-type ChAT.

Given the surface-exposed positions of residues Pro17 and Pro19 within the human 69-kDa ChAT structure [420] it is unlikely that observed differences in in vitro specific activity of recombinant P17A- or P17A/P19A-ChAT are due to substantial changes in protein secondary structure or stability. To confirm this, I completed circular dichroism (CD) analysis on purified recombinant ChAT and, importantly, observed no significant

Figure 3.4: Purification of bacterially-expressed recombinant wild-type and proline- mutant ChAT. (A) Representative Imperial Protein-stained SDS-PAGE gel loaded with 1.0 or 2.0 µg of purified bacterially-expressed recombinant wild-type (WT) and proline- mutant ChAT. (B) Proline-to-alanine mutation of residue Pro17 either alone or in combination with Pro19 (i.e. P17A/P19A) reduces the in vitro specific activity of purified recombinant mutant ChAT when compared to wild-type ChAT (***p≤0.001), with P17A/P19A-ChAT yielding the greatest reduction compared to either P17A- (*p≤0.05) or P19A-ChAT (***p≤0.001). No significant change in in vitro specific activity was observed for mutant P19A-ChAT when compared to wild-type ChAT (one-way ANOVA with Tukey’s post-hoc test, mean ± SEM, n=8).

differences in the gross protein secondary structure between the different recombinant ChAT proteins tested (i.e. wild-type, P17A, P17A/P19A and P19A) (Figure 3.5a). To assess the in vitro stability of recombinant wild-type and proline-mutant ChAT I performed a thermal denaturation assay by tracking loss of helicity at 222 nm during a thermal melt from 4 to 95C (Figure 3.5b). In support of my CD results showing the absence of gross structural changes in the recombinant proline-mutant ChAT proteins, the denaturation

Figure 3.5: In vitro analysis of recombinant wild-type and proline-mutant ChAT. (A) Circular dichroism analysis of purified bacterially-expressed recombinant ChAT reveals that both wild-type (WT) and proline-mutant ChAT are well-structured, primarily alpha helical, and that there are no overall changes in gross protein secondary structure of proline- mutant ChAT as compared to that for wild-type ChAT. Mean residue ellipticity (MRE) was measured over a wavelength range of 195-260 nm at 0.5 nm intervals. (B) Proline-to- alanine mutation does not significantly affect the in vitro denaturation midpoint (Tm) of recombinant P17A- (50.65 ± 0.081C), P19A- (49.45 ± 0.07C), or P17A/P19A-ChAT

(50.35 ± 0.07C) as compared to wild-type ChAT (51.15 ± 0.03C). Tm was determined by tracking loss of helicity at 222 nm from 4-95°C.

midpoint (Tm) of ChAT was not significantly altered by mutations involving the N-terminal proline-rich motif as follows; wild-type (51.15 ± 0.03C), P17A-ChAT (50.65 ± 0.081C), P17A/P19A-ChAT (50.35 ± 0.07C) and P19A-ChAT (49.45 ± 0.07C).

3.3: Proteasome inhibition increases ChAT steady-state protein levels and cellular ChAT enzyme activity

My studies using purified recombinant ChAT suggest that mutation of the N-terminal proline-rich motif can result in loss of enzymatic activity though does not result in gross changes in protein secondary structure or thermal stability. Interestingly, in mammalian SN56 cells N-terminal mutation of ChAT, including P17A/P19A and CMS-related V18M, results in reduced ChAT protein levels (Figures 3.1 – 3.3), though the cellular mechanisms responsible for this reduction in mutant ChAT protein levels have not been defined. Loss of cellular protein levels can be due to enhanced protein degradation that is mediated through either proteasomal or lysosomal proteolysis; it is currently unknown whether ChAT is degraded by either of these cellular proteolytic pathways. Thus, I tested whether treatment with either the proteasome inhibitor MG132 [26] or the lysosome inhibitor chloroquine [117] could alter the steady-state protein levels and/or cellular activity of ChAT in SN56 cells transiently expressing either wild-type (Figure 3.6a) or P17A/P19A- ChAT (Figure 3.6c). Importantly, following analysis of cell lysates by anti-ChAT immunoblotting, I observed an increase in the steady-state protein levels for both wild-type (p≤0.01; Figure 3.6b) and P17A/P19A-ChAT (p≤0.001; Figure 3.6d) in cells treated with 5 µM MG132 for 18 h when compared to cells treated with 25 µM chloroquine or to DMSO-treated control cells. Additionally, by radioenzymatic assay MG132 treatment was found to significantly increased the cellular activity of wild-type ChAT as compared to treatment with either chloroquine or DMSO-control (p≤0.00; Figure 3.6e). In contrast, treatment with either MG132 or chloroquine failed to increase the cellular activity of P17A/P19A-ChAT (Figure 3.6e), suggesting that this mutant enzyme maintains impaired cellular catalytic activity despite increased protein levels following MG132 treatment. Proteasomal inhibition with MG132 was verified by accumulation of ubiquitinated

Figure 3.6: Proteasome inhibition increases ChAT steady-state protein levels and cellular ChAT activity. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type (WT) ChAT that were treated for 18 h with 5 µM MG132 or 25 μM chloroquine (CQ). Proteasome inhibition by MG132 treatment or lysosome inhibition by CQ treatment was validated by immunoblotting for the accumulation of either ubiquitinated cellular proteins or for the lysosome-associated protein LC3B-II, respectively. Anti-actin immunoblotting was completed as a loading control. (B) Steady-state protein levels of wild- type ChAT are increased in MG132-treated cells compared to either CQ-treated cells or DMSO-treated control cells (**p≤0.01, one-way ANOVA with Tukey’s post-hoc test, mean ± SEM, n=3). (C) Anti-ChAT immunoblots from SN56 cells transiently expressing mutant P17A/P19A-ChAT that were treated and assessed as in (A). (D) Steady-state protein levels of P17A/P19A-ChAT are increased in MG132-treated cells compared to CQ-treated cells or DMSO-treated control cells (***p≤0.001, one-way ANOVA with Tukey’s post- hoc test, mean ± SEM, n=3). (E) Cellular ChAT activity is increased in wild-type ChAT expressing cells treated with MG132 as compared to either CQ-treated cells or DMSO- treated control cells (***p≤0.001, two-way ANOVA with Bonferroni’s post-hoc test, mean ± SEM, n=3). Neither MG132 nor CQ treatment increased the cellular activity of mutant P17A/P19A-ChAT.

proteins (Figure 3.6a), whereas lysosomal inhibition with chloroquine was verified by accumulation of a lysosome-associated isoform of microtubule-associated protein 1A/1B- light chain 3 (LC3B-II; Figure 3.6c) [494].

Together these results suggest that wild-type and mutant P17A/P19A-ChAT may be targeted for proteasomal degradation. To next determine whether CMS-related mutant ChAT may also be regulated by the proteasome I tested whether proteasomal inhibition could increase the steady-state protein levels of CMS-mutant ChAT, and if this may correlate with increased cellular ChAT activity. Thus, SN56 cells transiently expressing wild-type or CMS-related V18M-, A513T-, or T490N-ChAT were treated for 18 h with 5 µM MG132 and steady-state protein levels and cellular ChAT activity were measured. As

shown in Figures 3.7a and 3.7b, proteasome inhibition significantly increased the steady- state protein levels of both A513T- (p≤0.05) and T490N-ChAT (p≤0.01) as compared to DMSO-treated control cells. Though not statistically significant, I did observe a trend towards increased steady-state protein levels of V18M-ChAT with MG132 treatment when compared to DMSO-control (p=0.114). Lastly, MG132 treatment also increased the cellular activity of both A513T- (p≤0.01) and T490N-ChAT (p≤0.001) as compared to DMSO-treated control cells, but, like P17A/P19A-ChAT, had no effect on the cellular activity of V18M-ChAT (Figure 3.7c).

3.4: Mutation of the N-terminal proline-rich motif enhances ChAT ubiquitination

My results with MG132 suggest that cellular stability of ChAT protein is regulated through proteasome-dependent degradation. Canonically, post-translational ubiquitination of cellular proteins by covalent attachment of poly-Ub chains targets proteins for proteasomal degradation [44, 60, 93]. Ubiquitination of ChAT protein has not been reported previously, and therefore as a preliminary experiment I treated SN56 cells transient expressing either wild-type or P17A/P19A with either 5 µM MG132 or 25 µM chloroquine for 18 h. Cells were subsequently collected, lysed, and assayed by anti-ChAT immunoblotting for the presence of higher molecular mass ChAT proteins that may be representative of ubiquitinated forms of ChAT (Figure 3.8). As anticipated, I observed accumulation of higher molecular mass ChAT proteins following treatment with 5 µM MG132 that were not present in cells treated with either DMSO-control (Figure 3.8a, top panel, anti-ChAT long-exposure) or in cells treated with 25 µM chloroquine (Figure 3.8b). Importantly, by comparing the levels of higher molecular mass ChAT protein following MG132 treatment, relative to total ChAT protein levels, I observed a greater abundance of higher molecular mass P17A/P19A-ChAT as compared to that for wild-type ChAT (p≤0.01; Figure 3.8c).

The presence of higher molecular mass ChAT immunoreactive proteins following proteasome inhibition suggests that ChAT may be ubiquitinated in cells. Therefore, I

Figure 3.7: Proteasome inhibition partially restores the steady-state protein levels and cellular activity of CMS-related ChAT mutants. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type (WT), V18M-, A513T-, or T490N-ChAT that were treated with 5 μM MG132 for 18 h. Proteasome inhibition by MG132 treatment was validated by immunoblotting for the accumulation of ubiquitinated cellular proteins. Anti- actin immunoblotting was completed as a loading control. (B) Steady-state protein levels of A513T- (*p≤0.05) and T490N-ChAT (**p≤0.01) are increased in MG132-treated cells as compared to DMSO-treated control cells. Though not statistically significant, a trend towards increased steady-state protein levels of V18M-ChAT with MG132 treatment was observed when compared to DMSO-control (p=0.114, two-way ANOVA with Bonferroni’s post-hoc test, mean ± SEM, n=4). (C) Cellular ChAT activity of wild-type (***p≤0.001), A513T- (**p≤0.01), and T490N-ChAT (***p≤0.001) are increased in MG132-treated cells compared to DMSO-treated control cells (two-way ANOVA with Bonferroni’s post-hoc test, mean ± SEM, n=4).

Figure 3.8: Proteasome inhibition reveals the presence of multiple higher molecular mass ChAT proteins. Anti-ChAT immunoblots from SN56 cells transiently expressing either wild-type (WT) or P17A/P19A-ChAT that were treated with either (A) 5 µM MG132 or (B) 25 µM chloroquine (CQ) for 18 h. Control cells were transfected with empty pcDNA3.1+ vector. Proteasome inhibition by MG132 treatment or lysosome inhibition by CQ treatment was validated by immunoblotting for the accumulation of either ubiquitinated cellular proteins or the lysosome-associated protein LC3B-II, respectively. Anti-actin immunoblotting was completed as a loading control. Treatment with MG132, but not CQ or DMSO-control, resulted in the accumulation of multiple higher molecular mass immunoreactive ChAT proteins in cells expressing either wild-type or P17A/P19A-ChAT. (C) The levels of higher molecular mass ChAT proteins following MG132 treatment (A), relative to total ChAT, is greater in cells expressing P17A/P19A-ChAT compared to wild- type ChAT (**p≤0.01, Student’s t-test, mean ± SEM, n=4).

co-expressed wild-type or P17A/P19A-ChAT with HA-tagged ubiquitin in SN56 cells and completed anti-HA IPs to isolate for ubiquitinated cellular proteins and to test for the presence of ChAT protein ubiquitination. Following anti-ChAT immunoblotting from anti- HA IP samples, I observed that both wild-type and P17A/P19A-ChAT are ubiquitinated, and furthermore that treatment of cells with 5 µM MG132 for 18 h increased the recovery of ubiquitinated ChAT compared to DMSO-treated control cells (Figure 3.9a). Importantly, the abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, is greater for P17A/P19A-ChAT. Treatment of cells with 25 µM chloroquine had negligible effect on the recovery of ubiquitinated wild-type or P17A/P19A-ChAT (Figure 3.9b). To determine whether ubiquitination of CMS-mutant ChAT is also altered, I completed anti-HA IPs from SN56 cells transiently co-expressing wild-type or CMS- mutant ChAT with HA-tagged ubiquitin that were treated with 5 µM MG132 for 18 h. As anticipated, by anti-ChAT immunoblotting from anti-HA IP samples I observed increased recovery of ubiquitinated wild-type and CMS-mutant ChAT proteins following proteasome inhibition by MG132 treatment. Importantly, the abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, was greater for both V18M- and A513T-ChAT as compared to wild-type ChAT (Figure 3.9c). In support of Figure 3.1, the ubiquitination of T490N-ChAT appears to be unchanged from that of wild-type ChAT.

To validate my results regarding enhanced ubiquitination of P17A/P19A-ChAT, I performed anti-ChAT IPs from SN56 cells transiently co-expressing wild-type or P17A/P19A-ChAT with HA-tagged ubiquitin that were treated for 18 h with either 5µM MG132, 25 µM chloroquine, or with 2.5 µM lactacystin, another proteasome inhibitor [23, 25] (Figure 3.10a). Following anti-HA immunoblotting from anti-ChAT IP samples, I observed increased recovery of ubiquitinated ChAT following proteasome inhibition by treatment with either MG132 or lactacystin compared to DMSO-treated control cells. The abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, was greater for P17A/P19A-ChAT when compared to wild-type ChAT. Treatment with chloroquine had no effect on recovery of ubiquitinated ChAT by anti-ChAT IP.

These initial experiments investigating ChAT ubiquitination were performed in cells transiently overexpressing HA-tagged ubiquitin, and therefore I next wanted to

Figure 3.9: ChAT ubiquitination is increased upon mutation of the N-terminal proline-rich motif and in cases of CMS-related mutations. (A) Representative anti- ChAT and anti-HA immunoblots demonstrating that treatment of SN56 cells with 5 μM MG132 for 18 h increases recovery of ubiquitinated wild-type (WT) and P17A/P19A- ChAT by anti-HA IP from cells co-expressing HA-tagged ubiquitin (HA-Ub) as compared to DMSO-treated control cells. The abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, is greater with P17A/P19A-ChAT as analyzed by anti-ChAT immunoblotting. Control cells were transfected to transiently express ChAT without HA-Ub or were transfected with empty pcDNA3.1+ vector. Proteasome inhibition by MG132 treatment was validated by anti-HA immunoblotting for the accumulation of ubiquitinated cellular proteins from whole cell lysates. Anti-actin immunoblotting was completed as a loading control (n=5). (B) Treatment of SN56 cells with 25 μM chloroquine (CQ) for 18 h had negligible effect on recovery of ubiquitinated wild-type or P17A/P19A- ChAT by anti-HA IPs from cells co-expressing HA-Ub as compared to DMSO-treated control cells. Lysosome inhibition by CQ treatment was validated by immunoblotting for the accumulation of the lysosome-associated protein LC3B-II (n=3). (C) Treatment of SN56 cells with 5 μM MG132 for 18 h enhances recovery of ubiquitinated wild-type and CMS-mutant ChAT by anti-HA IPs from cells co-expressing HA-tagged ubiquitin as compared to DMSO-treated control cells. The abundance of ubiquitinated ChAT, relative to total ChAT protein levels, is greater with mutant P17A/P19A-, V18M-, and A513T- ChAT compared to wild-type ChAT as evaluated by anti-ChAT immunoblotting (n=5).

confirm that ChAT is ubiquitinated in cells under conditions of endogenously-expressed ubiquitin. Thus, I completed anti-ChAT IPs from SN56 cells transiently expressing only either wild-type or P17A/P19A-ChAT and that were treated for 18 h with either 5 µM MG132 or 25 µM chloroquine (Figure 3.10b). Following anti-ubiquitin immunoblotting from anti-ChAT IP samples, I observed that both wild-type and mutant P17A/P19A-ChAT are ubiquitinated in SN56 cells with endogenous ubiquitin. In addition, recovery of ubiquitinated ChAT was again increased following proteasome inhibition by MG132, and

Figure 3.10: Anti-ChAT IP reveals enhanced ubiquitination of mutant P17A/P19A- ChAT. (A) Representative anti-ChAT and anti-HA immunoblots demonstrating that proteasome inhibition by treatment of SN56 cells with 5 μM MG132 or 2.5 μM lactacystin (Lac) for 18 h increases recovery of ubiquitinated wild-type (WT) and P17A/P19A-ChAT following anti-ChAT IPs from cells co-expressing HA-tagged ubiquitin (HA-Ub) as compared to DMSO-treated control cells. Distinct bands in anti-HA immunoblots corresponding to different molecular mass species of ubiquitinated ChAT are indicated by red arrows. The abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, is greater with P17A/P19A-ChAT compared to wild-type ChAT. Treatment with 25 μM CQ for 18 h had no effect on recovery of ubiquitinated ChAT. Proteasome inhibition by MG132 treatment or lysosome inhibition by CQ treatment was validated by immunoblotting from whole cell lysates for the accumulation of either ubiquitinated cellular proteins (i.e. HA-Ub) or for the lysosome-associated protein LC3B- II, respectively. Anti-actin immunoblotting was completed as a loading control (n=5). (B) Ubiquitination of P17A/P19A-ChAT with endogenous ubiquitin is enhanced compared to wild-type ChAT in SN56 cells transfected to transiently express only either wild-type or P17A/P19A-ChAT. Ubiquitinated ChAT was analyzed by anti-ubiquitin immunoblotting from anti-ChAT IPs samples. Proteasome inhibition by 5 µM MG132 treatment for 18 h increases recovery of ubiquitinated ChAT compared to DMSO-control, whereas treatment with 25 μM CQ for 18 h had no effect on recovery of ubiquitinated ChAT. Distinct bands in anti-ubiquitin immunoblots corresponding to different molecular mass species of ubiquitinated ChAT are indicated by red arrows. Control cells were transfected with empty pcDNA3.1+ vector (n=4).

furthermore that the abundance of ubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, is greater for P17A/P19A-ChAT when compared to wild-type ChAT. Of importance, these results with endogenously-expressed ubiquitin reflect my earlier results using transient co-expression of ChAT with HA-tagged ubiquitin.

3.5: ChAT protein can be targeted for proteasomal degradation by both K48-dependent and K48-independent polyubiquitination

My results with the proteasome inhibitors MG132 and lactacystin and either HA-tagged or endogenously-expressed ubiquitin suggest that ChAT protein stability is regulated by the ubiquitin-proteasome system. As illustrated in Figure 1.4, proteins targeted for proteasome- dependent degradation are modified by E3 ubiquitin ligases through covalent attachment of monomeric ubiquitin onto lysine residues that then serves as a primer for construction of K48-linked poly-Ub chains by addition of free ubiquitin onto residue Lys48 of substrate- bound ubiquitin. Subsequently, K48-linked poly-Ub chains are directly recognized by the 26S-proteasome, resulting in proteolysis of these polyubiquitinated proteins [44, 60, 93]. In addition to Lys48, ubiquitin contains six other lysine residues (K6, 11, 27, 29, 33 and 63) that can be used for formation of poly-Ub chains that differ from K48-linked chains [44, 50]. These alternative poly-Ub chains can have non-proteolytic roles, with K63-linked chains best known to regulate endosomal substrate trafficking, receptor endocytosis, kinase activation and DNA repair [44, 51].

My initial work investigating ChAT ubiquitination was performed under conditions of proteasome inhibition by treatment of cells with either MG132 or lactacystin, and thus I wanted to further characterize ChAT ubiquitination in SN56 cells having active and functional proteasomes. To this end, SN56 cells transiently expressing either wild-type (Figure 3.11a) or P17A/P19A-ChAT (Figure 3.11b) were co-transfected to express HA- tagged wild-type ubiquitin or different HA-tagged ubiquitin mutants that either cannot form poly-Ub chains (K0), that can only form K48-linked chains (K48), or that can only form non-K48-linked poly-Ub chains (i.e. K11, K48R, or K63) and anti-HA IPs were completed to isolate ubiquitinated cellular proteins. Following anti-ChAT immunoblotting from anti- HA IP samples, I observed that recovery of polyubiquitinated wild-type and P17A/P19A- ChAT was reduced in SN56 cells that co-express K48-ubiqutin as compared to wild-type- ubiquitin. Alternatively, prevention of K48-linked polyubiquitination of ChAT by co- expression of either K0-, K11- (only K11-linked chains), or K48R-ubiquitin increased recovery of polyubiquitinated wild-type and P17A/P19A-ChAT in anti-HA IPs. Co-

Figure 3.11: Wild-type and P17A/P19A-ChAT can be targeted for proteasomal degradation by both K48-dependent and K48-independent polyubiquitination. Recovery of ubiquitinated wild-type (WT) ChAT (A) (n=3) and P17A/P19A-ChAT (B) (n=3) is reduced in SN56 cells co-expressing HA-tagged mutant ubiquitin (Ub)-K48 (only K48-linked polyubiquitin chains) compared to wild-type HA-Ub as assessed by anti-ChAT immunoblotting following anti-HA IPs. Conversely, recovery of wild-type and mutant P17A/P19A-ChAT is increased in cells with co-expression of HA-tagged Ub-K0 (no polyubiquitin linkages) or either Ub-K11 or Ub-K48R that are unable to form K48-linked polyubiquitin chains. Control cells were transfected with empty pcDNA3.1+ vector. Anti- actin immunoblotting from whole cell lysates was completed as a loading control. (C) Treatment with 5 μM MG132 for 18 h increases recovery of both K48-linked and non-K48- linked (K48R) polyubiquitinated wild-type ChAT and P17A/P19A-ChAT in SN56 cells co-expressing either HA-Ub-K48 or HA-Ub-K48R compared to DMSO-treated control cells. Cellular ChAT ubiquitination was assessed by anti-ChAT immunoblotting following anti-HA IPs. Proteasome inhibition by MG132 treatment was validated by anti-HA immunoblotting (i.e. HA-Ub) for the accumulation of ubiquitinated cellular proteins from whole cell lysates (n=4).

-expression with K63-ubiquitin (only K63-linked chains) had no effect on the recovery of polyubiquitinated wild-type or P17A/P19A-ChAT as compared to co-expression with wild- type-ubiquitin. Closer examination of ubiquitinated P17A/P19A-ChAT from anti-HA IP samples (Figure 3.11b, top panel, anti-ChAT immunoblot) revealed an additional higher molecular mass ubiquitinated ChAT species in samples co-expressing K0, K11 or K48R- ubiquitin when compared to cells co-expressing wild-type ubiquitin. This additional ubiquitinated ChAT species suggests that increased P17A/P19A-ChAT ubiquitination may be due to multi-ubiquitination of additional and/or alternative lysine residues within mutant ChAT that may differ from those utilized for the basal ubiquitination of wild-type ChAT.

These results suggest that ChAT protein degradation by the proteasome is regulated through K48-linked polyubiquitination. To confirm this, we treated cells for 18 h with 5

µM MG132 that were transiently co-expressing either wild-type or P17A/P19A-ChAT with HA-tagged wild-type, K48-, or K48R-ubiquitin (Figure 3.11c). As anticipated, proteasome inhibition increased the recovery of K48-linked polyubiquitinated ChAT. Interestingly, MG132 treatment also enhanced the recovery of non-K48-linked polyubiquitinated ChAT when ChAT was co-expressed with K48R-ubiquitin. The abundance of both K48-linked and non-K48-linked polyubiquitinated ChAT, relative to total ChAT protein levels from whole cell lysates, appears greater for P17A/P19A-ChAT compared to wild-type ChAT.

3.6: ChAT protein half-life is reduced upon mutation of the N-terminal proline-rich motif and can be increased by proteasome inhibition

My assessment of ChAT ubiquitination suggests that enhanced ubiquitination, and resultant enhanced proteasome-dependent degradation, may be responsible for the observed reduction in the steady-state protein levels of P17A/P19A-ChAT. To confirm that the rate of degradation of P17A/P19A-ChAT is increased, I investigated whether the protein half- life of P17A/P19A-ChAT is reduced in SN56 cells when proteasomes are functional. To measure protein half-life, I developed a novel fluorescent strain-promoted alkyne-azide cycloaddition (SPAAC) pulse-chase procedure that utilizes non-radioactive labeling and detection reagents (Figure 3.12). This protocol is an adaptation on classical 35S-methionine pulse-chase labeling and is based on metal-free “click” chemistry whereby azide-labeled compounds react specifically and efficiently with strained cyclooctynes, by 1,3-dipolar cycloaddition, to form stable 1,2,3-triazole conjugates without the need of a metal catalyst [495-496]. While SPAAC reactions have been used previously both in vitro and in vivo to measure global changes in protein synthesis [497-500], my study is the first to apply the use of SPAAC reactions to measure half-life of a specific cellular protein. Briefly, in my protocol proteins are live-labeled (pulsed) with L-azidohomoalanine (AHA), a biorthogonal methionine analog that contains a reactive azide moiety, under methionine- free conditions and then chased with excess methionine. AHA-labeled proteins are then detected by reaction with 4-dibenzocyclooctynol (DIBO), a strained cyclooctyne, that

Figure 3.12: Schematic overview of biorthogonal strain-promoted alkyne-azide cycloaddition (SPAAC) pulse-chase for fluorescent determination of protein half-life. (1) Briefly, proteins are live-labeled (pulsed) with L-azidohomoalanine (AHA), a biorthogonal methionine analog that contains a reactive azide moiety, under methionine- free conditions and then (2) chased with medium containing methionine for desired times (e.g. up-to 24 hours). (3) Cell are then collected, lysed, and (4) immunoprecipitation for a protein-of-interest (e.g. ChAT) is completed. (5) AHA-labeled proteins are reacted with the strained cyclooctyne 4-dibenzocyclooctynol (DIBO) that is modified with a fluorescent tetramethylrhodamine (TAMRA) probe to form stable fluorescent triazole conjugates. (6) AHA/TAMRA-labeled proteins are resolved on SDS-PAGE gels, and (7a) TAMRA fluorescence is detected in-gel at an excitation/emission of 555/580 nm. In addition, (7b) AHA/TAMRA-labeled proteins can be transferred to PVDF membranes and standard immunoblotting can be completed for downstream analysis.

is modified with a fluorescent tetramethylrhodamine (TAMRA) probe. Importantly, treatment of HEK293 cells with AHA has been shown to be non-toxic, resulting in highly- specific labeling of newly-synthesized proteins that does not affect ubiquitination or protein degradation of these AHA-labeled proteins [498].

Using this pulse-chase protocol in SN56 cells transiently expressing either wild-type or P17A/P19A-ChAT, I detected the global progressive loss of fluorescent AHA/TAMRA- labeled proteins from whole cell lysates during the 0-24 h chase period in the absence of changes in total protein levels as measured in parallel by anti-actin immunoblotting (Figure 3.13a). Additionally, following anti-ChAT IP from ChAT-expressing SN56 cells live- labeled with AHA, I also observed progressive loss of fluorescent AHA/TAMRA-labeled ChAT during the 0-24 h chase period. Importantly, the decline in protein levels of immunoprecipitated AHA/TAMRA-labeled P17A/P19A-ChAT is more rapid than that of wild-type ChAT. By quantifying the fluorescence intensities of AHA/TAMRA-labeled ChAT, I determined that the protein half-life of P17A/P19A-ChAT (2.2 h) is significantly reduced by ~10-fold compared to wild-type ChAT (19.7 h; F(1,41)=110.043, p≤0.0001; Figure 3.13b). A 24 h time point was not included for the quantification of P17A/P19A- ChAT half-life as the fluorescence intensity was indistinguishable from background.

To validate the results of my novel SPAAC pulse-chase assay, I performed a cycloheximide (CHX)-chase assay on SN56 cells transiently expressing either wild-type or P17A/P19A-ChAT. CHX is a potent, though toxic, inhibitor of protein biosynthesis commonly used for the molecular determination of protein half-life [501]. Thus, by anti- ChAT immunoblotting from ChAT-expressing SN56 cells that were treated with 100 µg/ml CHX for 2 – 8 h (Figure 3.14a) I determined that the cellular half-life of P17A/P19A-ChAT (4 h) is again ~10-fold shorter than that of wild-type ChAT (45 h; F(1,46)=36.802, p≤0.0001; Figure 3.14b). Although the absolute half-life values determined for wild-type ChAT and P17A/P19A-ChAT by these two methods differ, importantly they both revealed that the protein half-life of P17A/P19A-ChAT is ~10% that of wild-type ChAT.

My earlier results show that proteasome inhibition by MG132 treatment resulted in increased steady-state protein levels of both wild-type and P17A/P19A-ChAT (Figure 3.6).

Figure 3.13: SPAAC pulse-chase reveals that ChAT protein half-life is reduced upon mutation of the N-terminal proline-rich motif. (A) Fluorescent detection of immunoprecipitated (IP) AHA/TAMRA-labeled wild-type (WT) and P17A/P19A-ChAT following SPAAC pulse-chase (Figure 3.12) from transiently transfected SN56 cells. Control cells were incubated without AHA (i.e. unlabeled). Fluorescent AHA/TAMRA- labeled proteins from either whole cell lysates or anti-ChAT IPs were detected in resolved SDS-PAGE gels at an Ex/Em of 555/580 nm. Anti-ChAT and anti-actin immunoblotting was completed on AHA/TAMRA-labeled samples for downstream data analysis and as loading controls. (B) Protein half-life of P17A/P17A-ChAT (2.2 h) is reduced compared to wild-type ChAT (19.7 h). ChAT fluorescence intensities from (A) were plotted on a semi- logarithmic scale and linear regression was completed. The slopes of decay for wild-type and P17A/P19A-ChAT were significantly different: F(1,41)=110.043, p≤0.0001, n=5.

Figure 3.14: Validation of SPAAC pulse-chase by cycloheximide (CHX)-chase assay. (A) SN56 cells, transiently expressing either wild-type (WT) or P17A/P19A-ChAT, were treated with 100 μg/ml CHX for indicated time points (hours) and anti-ChAT immunoblotting was completed. Anti-actin immunoblotting was completed as a loading control. (B) Protein half-life of P17A/P17A-ChAT (4 h) is reduced as compared to wild- type ChAT (45 h). Steady-state ChAT protein levels from (A) were plotted on a semi- logarithmic scale and linear regression completed. The slopes of decay for wild-type and P17A/P19A-ChAT were significantly different: F(1,46) = 36.802, p≤0.0001, n=5.

Therefore, I tested next whether proteasome inhibition would correlate with an increase in ChAT protein half-life. To determine ChAT protein half-life during proteasome inhibition, I performed SPAAC pulse-chase on SN56 cells transiently expressing either wild-type (Figure 3.15a) or P17A/P19A-ChAT (Figure 3.15b) that were treated with 5 µM MG132 throughout both the 4 h pulse and 24 h chase periods. Following anti-ChAT IP from AHA- labeled SN56 cells and quantification of fluorescence intensities of immunoprecipitated AHA/TAMRA-label ChAT, I observed that proteasome inhibition by MG132 treatment prevented the decay of wild-type ChAT protein as compared to DMSO-treated control cells (half-life = 22.5 h; Figure 3.15c). Additionally, the protein half-life of P17A/P19A-ChAT was also increased during MG132 treatment (16.8 h) as compared to

Figure 3.15: Proteasome inhibition increase ChAT protein half-life. Fluorescent detection of immunoprecipitated (IP) AHA/TAMRA-labeled wild-type (WT; A) or P17A/P19A-ChAT (B) following SPAAC pulse-chase from transiently transfected SN56 cells that were treated with either DMSO-control or 5 µM MG132 throughout both the 4 h AHA-labeling and the 24 h chase periods. Control cells were incubated without AHA (i.e. unlabeled). Fluorescent AHA/TAMRA-labeled proteins were detected in resolved SDS- PAGE gels at an Ex/Em of 555/580 nm. Anti-ChAT and anti-actin immunoblotting was completed on AHA/TAMRA-labeled samples for downstream data analysis and as loading controls. (C) Proteasome inhibition by MG132 treatment increased the protein half-life of wild-type ChAT (no protein decay) as compared to DMSO-treated control cells (22.5 h). Wild-type ChAT fluorescence intensities from (A) were plotted on a semi-logarithmic scale and linear regression was completed. The slopes of decay for DMSO- and MG132-treated cells were significantly different from each other: F(1,46) = 4.79241, p≤0.05, n=5. (D) Proteasome inhibition by MG132 treatment increased the protein half-life of mutant P17A/P19A-ChAT (16.6 h) as compared to DMSO-control (2.2 h). ChAT fluorescence intensities from (B) were plotted on a semi-logarithmic scale and linear regression was completed. The slopes of decay for DMSO and MG132-treated cells were significantly different from each other: F(1,41)=18.9864, p≤0.0001, n=5.

DMSO-control (2.2 h; Figure 3.15d). No 24 h time point was included for protein half-life quantification from DMSO-treated control cells expressing P17A/P19A-ChAT as the fluorescence intensity was indistinguishable from background.

3.7: Identification of the heat shock proteins HSP70 and HSP90 as putative ChAT protein interactors

To summarize my findings thus far, I initially sought to identify a functional role of the N- terminal proline-rich motif of ChAT and have shown that both cellular ChAT protein levels and enzyme activity are dramatically reduced following alteration of this motif, particularly

for the P17A/P19A and CMS-related V18M mutations. Furthermore, I also demonstrated for the first time that cellular degradation of ChAT protein is modulated through the ubiquitin-proteasome system, and that ubiquitination of mutant ChAT is enhanced. The mechanism/s responsible for basal ChAT ubiquitination or enhanced ubiquitination of catalytically-deficient ChAT have not been elucidated (Figure 3.16) [502].

Regulation of protein stability through the ubiquitin-proteasome system involves a complex network of protein chaperones, E3 ubiquitin ligases, and other cellular proteostatic factors. Therefore, to identify putative protein-interactors that may regulate ChAT protein stability I utilized the BioID proteomics approach [483] to screen for direct and proximal ChAT-interacting proteins. Briefly, in the BioID protocol cells expressing fusions proteins consisting of a protein-of-interest (e.g. ChAT) fused to the 35-kDa HA-tagged promiscuous biotin ligase BirA-R118G-HA (BirA*) are treated with exogenous biotin to allow for proximity-dependent biotinylation of proximal and directly-interacting proteins. Cells are then lysed, streptavidin pull-downs (PDs) are completed, and biotinylated proteins can be analyzed by SDS-PAGE and identified by mass spectrometry (Figure 3.17a). To optimize this protocol for my studies I chose to compare wild-type ChAT to P17A/P19A-ChAT as this mutation had the greatest impact on ChAT ubiquitination and degradation following modification of the proline-rich motif [502]. Thus, HEK293 cells transiently expressing either wild-type- or P17A/P19A-ChAT-BirA* fusion proteins were treated with either vehicle-control (water) or 50 µM biotin for 24 h to allow proximity-dependent biotinylation of ChAT-interacting proteins. Subsequent analysis of protein samples from whole cell lysates by anti-ChAT and anti-HA immunoblotting revealed expression of ChAT-BirA* fusion proteins with an apparent molecular mass of ~100 kDa as compared to untagged ChAT proteins at ~70 kDa (Figure 3.17b, top panels). The steady-state protein levels of P17A/P19A-ChAT-BirA* was less than that of wild-type-ChAT-BirA* in agreement with my previous results demonstrating enhanced ubiquitination and degradation of mutant P17A/P19A-ChAT (Figures 3.10 and 3.13) [502]. Furthermore, the BioID-dependent biotinylation of cellular proteins was analyzed by blotting of protein samples obtained from streptavidin PDs with HRP-conjugated streptavidin (Strep-HRP). As anticipated, I detected biotinylation of multiple cellular proteins in samples from biotin-treated HEK293 cells

Figure 3.16: Model for the regulation of ChAT protein stability by the ubiquitin- proteasome system. ChAT synthesizes the neurotransmitter acetylcholine using acetyl- CoA (AcCoA) and choline as substrates in cholinergic neurons. Steady-state protein levels of wild-type (WT) ChAT are regulated by ubiquitination (Ub) and subsequent degradation by the 26S-proteasome. Ubiquitination and proteasomal degradation of ChAT is enhanced upon mutation of the N-terminal proline-rich motif of ChAT (i.e. P17A/P19A) as well as for the CMS-related V18M- and A513T-ChAT mutants. These mutations also reduce the cellular catalytic activity of mutant ChAT, thereby resulting in a synergistic impairment in cellular acetylcholine production (red hashed arrow).

Figure 3.17: Proximity-dependent biotin identification (BioID) for the determination of novel ChAT protein-protein interactions. (A) Overview of BioID protocol. Briefly, a protein of interest (e.g. ChAT), fused to the promiscuous biotin ligase BirA-R118G-HA (BirA*), is expressed in mammalian cells (e.g. HEK293 cells). Upon the addition of exogenous biotin to live-cell growth media, proteins that are either proximal-to or directly bound-to BirA* fusion protein will be live-labeled with biotin. Cells are then lysed, and biotinylated proteins are subsequently purified by streptavidin pull-down, resolved on SDS-PAGE gels, and identified by mass spectrometry. (B) Optimization of BioID in HEK293 cells transiently expressing wild-type (WT) or P17A/P19A-ChAT fused to HA- tagged BirA*. Control cells were transfected with empty pcDNA3.1+ vector or plasmids encoding either untagged ChAT or unfused BirA*. Cells were treated for 24 h with 50 µM biotin to induce proximity-dependent biotinylation of ChAT-interacting cellular proteins or with vehicle-control (water). Protein samples were prepared from both whole cell lysates and from streptavidin pull-downs (PD: Strep), resolved on SDS-PAGE gels, and transferred to PVDF membranes. Biotinylated cellular proteins were detected by blotting with HRP- conjugated streptavidin (Strep-HRP). ChAT-BirA* fusion proteins were detected by anti- ChAT and anti-HA immunoblotting. Anti-actin immunoblotting from whole cell lysates was completed as a loading control (n=2).

that expressed ChAT-BirA* fusion proteins, but not in control cells grown in the absence of biotin (vehicle) or in cells expressing untagged ChAT (Figure 3.17b). Importantly, the abundance of several of these biotinylated proteins was different in cells expressing P17A/P19A-ChAT-BirA* as compared to wild-type-ChAT-BirA*; an ~70 kDa protein was notably enriched in cells expressing P17A/P19A-ChAT-BirA*.

My initial BioID experiments suggest that multiple proteins interact with and may differentially regulate P17A/P19A-ChAT as compared to wild-type ChAT. My primary goal for using BioID was to identify putative ChAT protein-interactors that may be responsible for the enhanced degradation of mutant ChAT observed earlier, and thus biotinylated proteins enriched in cells expressing P17A/P19A-ChAT-BirA* were of prime

interest to identify. Therefore, I completed a large-scale BioID experiment in HEK293 cells expressing either wild-type-ChAT-BirA* or P17A/P19A-ChAT-BirA* and, following streptavidin PDs, biotinylated proteins were visualized on a silver stained SDS-PAGE gel (Figure 3.18). Two proteins of interest that were enriched in cells expressing P17A/P19A- ChAT-BirA* with apparent molecular masses of ~70 kDa and ~90 kDa were excised from the gel, along with an intense band at ~100 kDa correlating with the estimated molecular mass of the putative P17A/P19A-ChAT-BirA* fusion protein. In-gel protein samples were digested with trypsin and identified by either matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS) and/or by liquid chromatography- electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Initial identification by MALDI-TOF-MS revealed the ~70 kDa protein as the heat shock protein HSP70 and, as anticipated, the ~100 kDa protein as ChAT-BirA* (Figure 3.18). By comparing the peptides masses generated by MALDI-TOF-MS to the NCBI protein database I obtained a 30% sequence coverage for human HSP70 (20 peptides; Figure 3.19). The ~100 kDa protein analyzed from cells expressing P17A/P19A-ChAT-BirA* was confirmed as ChAT- BirA* with a sequence coverage for human 69-kDa ChAT and bacterial BirA of 31% (20 peptides) and 42% (16 peptides) respectively (Figure 3.20). My data did not contain peptides with masses that matched tryptic peptides that would cover the mutated residues in the P17A/P19A-ChAT sequence. Identification of the BioID-enriched ~90 kDa protein could not be achieved by MALDI-TOF-MS due to low sample yield, though subsequently by using the more sensitive LC-ESI-MS/MS approach I was able to identify this protein as HSP90 with a 16% sequence coverage (10 peptides; Figure 3.21). Furthermore, by using LC-ESI-MS/MS I obtained a greater sequence coverage for HSP70 of 34% (25 peptides; Figure 3.22). Importantly, tryptic peptides with masses indicating the addition of biotin were detected for both HSP70 and ChAT-BirA* by MALDI-TOF-MS; these peptides contained lysine residue that could be sites for biotinylation by BirA*.

To verify these proteins identified by MALDI and LC-ESI-MS/MS following BioID, I performed anti-HSP70 and anti-HSP90 immunoblotting on biotinylated protein samples obtained by streptavidin PDs from biotin-treated HEK293 cells expressing either wild-type-ChAT-BirA* or P17A/P19A-ChAT-BirA* (Figure 3.23). Importantly, as

Figure 3.18: Identification of the heat shock proteins HSP70 and HSP90 as novel ChAT protein-interactions by BioID. Streptavidin pull-down samples prepared from biotin-treated HEK293 cells expressing either wild-type (WT)-ChAT-BirA* or P17A/P19A-ChAT-BirA* fusion proteins that were resolved and visualized on a silver- stained SDS-PAGE gels. Two biotinylated proteins (~70 kDa and ~90 kDa) that were enriched in samples expressing P17A/P19A-ChAT-BirA* were identified by MALDI- TOF-MS or LC-ESI-MS/MS as HSP70 and HSP90, respectively. An ~100 kDa protein correlating with the estimated molecular mass of the P17A/P19A-ChAT-BirA* fusion protein was confirmed by MALDI-TOF-MS as ChAT-BirA*. Biotin-treated control cells were transfected to express untagged wild-type ChAT (n=1).

Figure 3.19: MALDI-TOF-MS data for HSP70 following BioID. (A) Peptide coverage map of human heat shock 70 kDa protein 1B (HSP70; NCBIprot database, RefSeq: NP_005337.2) obtained by MALDI-TOF-MS from P17A/P19A-ChAT-BirA* BioID samples (Figure 3.18). A total sequence coverage of 30% was obtained. (B) Peptide list of 20 unique peptides with indicated amino acid modifications and relevant MALDI-TOF- MA data for HSP70. One peptide (bordered) was identified with a mass-shift correlating with biotinylation of residue Lys507 of human HSP70.

Figure 3.20: MALDI-TOF-MS data for ChAT-BirA* fusion protein following BioID. (A) Peptide coverage map of human 69-kDa choline acetyltransferase (ChAT; NCBIprot database; GenBank: AAI30618.1) obtained by MALDI-TOF-MS from P17A/P19A- ChAT-BirA* BioID samples (Figure 3.18). A total sequence coverage of 33% was obtained for human 69-kDa ChAT. (C) Peptide coverage map of bacterial BirA (NCBIprot database; GenBank: AEP13897.1) obtained by MALDI-TOF-MS from P17A/P19A-ChAT-BirA* BioID samples. A total sequence coverage of 42% was obtained for bacterial BirA. Peptide lists of either 23 (B; ChAT) or 16 (D; BirA) unique peptides with indicated modifications and relevant MALDI-TOF-MA data. One ChAT peptide (B; bordered) was identified with a mass-shift correlating with biotinylation of residue Lys4 of human ChAT.

anticipated I observed PD of endogenous HSP70 and HSP90 from biotin-treated cells expressing P17A/P19A-ChAT-BirA*; streptavidin PD of these HSPs was not consistently observed from cells expressing wild-type-ChAT-BirA* (2 of 4 trials). As controls, neither HSP70 nor HSP90 were visualized in immunoblots prepared from streptavidin PDs from biotin-treated control cells that expressed untagged ChAT. In addition, anti-ChAT and anti- HA immunoblots revealed recovery of ChAT-BirA*, but not untagged ChAT, following streptavidin PD, suggesting that ChAT-BirA* can undergo self-biotinylation. This has been reported in other BioID studies for BirA*-tagged proteins [483] and confirms my MALDI- TOF-MS analysis of biotinylated P17A/P19A-ChAT-BirA* (Figure 3.20).

3.8: Co-immunoprecipitation of ChAT with HSPs is altered by mutation of N-terminal proline-rich motif of ChAT

HSPs, such as HSP70 and HSP90, are well-characterized molecular chaperones capable of performing protein triage decisions by either promoting the folding and stabilization of cellular proteins or by targeting terminally misfolded proteins for degradation [118, 135]. As such, these HSPs may represent molecular chaperones involved in the stabilization of ChAT and/or that may promote degradation of mutant or misfolded ChAT. Therefore, to

Figure 3.21: LC-ESI-MS/MS data for HSP90 following BioID. (A) Peptide coverage map of human heat shock protein HSP 90-beta isoform c (HSP90; NCBInr_new database, RefSeq: NP_001258901.1) obtained by LC-ESI-MS/MS from P17A/P19A-ChAT-BirA* BioID samples (Figure 3.18). A total sequence coverage of 16% with 10 unique peptides with indicated modifications was obtained. (B) LC-ESI-MS/MS peptide list for human HSP90 with relevant MS data following ChAT-BirA* BioID.

to verify these putative BioID interactions between ChAT and endogenous HSP70 and HSP90, I performed co-IP assays from HEK293 cells expressing either untagged wild-type or P17A/P19A-ChAT (Figure 3.24a). Interestingly, I was able to observe both HSP70 and HSP90 in anti-ChAT co-IP samples from cells expressing either wild-type or P17A/P19A- ChAT. Furthermore, in addition to HSP70 and HSP90, I also found that endogenous HSC70, a constitutively-expressed HSP that has analogous chaperone function and shares similar substrate proteins to that of HSP70 [194], can co-IP with ChAT (Figure 3.24a). Importantly, by comparing the levels of ChAT protein to that of HSPs in anti-ChAT co-IP

Figure 3.22: LC-ESI-MS/MS data for HSP70 following BioID. (A) Peptide coverage map of human heat shock 70 kDa protein 1B (HSP70; NCBInr_new database, RefSeq: NP_005337.2) obtained by LC-ESI-MS/MS from P17A/P19A-ChAT-BirA* BioID samples (Figure 3.18). A total sequence coverage of 34% with 25 unique peptides with indicated modifications was obtained. (B) LC-ESI-MS/MS peptide list for human HSP70 with relevant MS data following ChAT-BirA* BioID.

Figure 3.23: Immunoblot verification of putative ChAT interacting proteins following BioID. Confirmation of endogenous HSP70 and HSP90 as putative ChAT-interacting proteins by immunoblotting of protein samples following streptavidin pull-down (PD: Strep). HEK293 cells, transiently expressing HA-tagged wild-type (WT)-ChAT-BirA* or P17A/P19A-ChAT-BirA*, were treated for 24 h with 50 µM of exogenous biotin. Biotin- treated control cells were transfected with empty vector or with vectors encoding for untagged ChAT proteins. Biotinylated cellular proteins from whole cell lysates were detected with HRP-conjugated streptavidin (Strep-HRP). ChAT-BirA* fusion proteins were detected by anti-ChAT and anti-HA immunoblotting. Anti-actin immunoblotting from whole cell lysates was completed as a loading control (n=4).

Figure 3.24: Co-immunoprecipitation of ChAT with HSC/HSP70 and HSP90 from HEK293 cells. (A) Immunoblots showing co-immunoprecipitation (co-IP) of ChAT with endogenous HSC70, HSP70, and HSP90 following anti-ChAT co-IP from HEK293 cells transiently expressing either wild-type (WT) or P17A/P19A-ChAT. Control cells were transfected with empty pcDNA3.1+ vector. Anti-actin immunoblotting from whole cell lysates was completed as a loading control. Co-IP of P17A/P19A-ChAT with HSP70 (B), HSP90 (C), and HSC70 (D) is greater than that of wild-type ChAT (***p≤0.001, Student’s t-test, mean ± SEM, n=4).

samples, I found enhanced association of mutant P17A/P19A-ChAT with endogenous HSP70 (p≤0.001; Figure 3.24b), HSP90 (p≤0.001; Figure 3.24c) and HSC70 (p≤0.001; Figure 3.24d) relative to wild-type ChAT.

My initial BioID and co-IP experiments were performed in HEK293 cells as these cells are easily transfected to transiently express heterologous proteins. As my earlier experiments assessing ChAT ubiquitination and degradation were carried out with mouse cholinergic SN56 cells, I next confirmed my novel results by performing anti-ChAT co-IPs from SN56 neural cells expressing either wild-type or P17A/P19A-ChAT. Furthermore, to assess whether enhanced association of ChAT with HSC/HSP70 and HSP90 would also be seen in selected CMS-related ChAT mutant proteins, I also evaluated anti-ChAT co-IPs from SN56 cells expressing either V18M- or A513T-ChAT. These two catalytically- deficient CMS-related mutations were selected as they were shown earlier in my study to enhance the ubiquitination and degradation of these ChAT mutant proteins [Figures 3.1, 3.7, and 3.9]. Importantly, I detected both endogenous HSC70 and HSP90 in anti-ChAT co-IP samples from SN56 cells expressing either wild-type or mutant ChAT proteins (Figure 3.25a). Unlike HEK293 cells, I did not observe constitutive expression of HSP70 in whole cell lysates from SN56 cells. By comparing the relative levels of ChAT proteins to that of endogenous HSC70 in anti-ChAT co-IP samples, I found enhanced association of both P17A/P19A- (p≤0.001) and V18M-ChAT (p≤0.05), but not A513T-ChAT, with HSC70 as compared to wild-type ChAT (Figure 3.25b). Interestingly, there were no significant differences in association of mutant ChAT with HSP90, but there was a trend towards enhanced association of HSP90 with P17A/P17A-ChAT (p=0.09; Figure 3.25c).

Lastly, to confirm whether endogenous HSC70 and HSP90 interact with ChAT in situ, I performed a proximity ligation assay (PLA) in SN56 cells transiently expressing wild-type ChAT. As predicted, I observed in situ interaction of wild-type ChAT with endogenous HSC70 and HSP90 in cells (Figure 3.26). Confocal images show that the ChAT-HSP PLA complexes are predominantly cytoplasmic in agreement with the previously reported subcellular distribution of human 69-kDa ChAT [414]. As controls, PLA complexes were not observed in SN56 cells that were transfected with empty vector

Figure 3.25: Co-immunoprecipitation of ChAT with HSC70 and HSP90 from mouse cholinergic SN56 cells. (A) Immunoblots showing co-immunoprecipitation (co-IP) of ChAT with endogenous HSC70 and HSP90 following anti-ChAT co-IP from mouse cholinergic SN56 cells expressing wild-type (WT), P17A/P19A-ChAT, or the CMS-related mutants V18M- or A513T-ChAT. Control cells were transfected with empty pcDNA3.1+ vector. Anti-actin immunoblotting from whole cell lysates was completed as a loading control. (B) Co-IP of P17A/P19A-ChAT (***p≤0.001) and V18M-ChAT (*p≤0.05), but not A531T-ChAT, with HSC70 is greater than that of wild-type ChAT (mean ± SEM, n=5). (C) While there was a trend toward increased HSP90 interaction with P17A/P19A-ChAT (p=0.09), no significant differences were observed for HSP90 interaction with mutant ChAT as compared to wild-type ChAT in SN56 cells (mean ± SEM, n=5). Statistical analysis for (B) and (C) was performed by one-way ANOVA with Dunnett’s post-hoc test.

Figure 3.26: In situ interaction of ChAT with endogenous HSC70 and HSP90. Detection of in situ ChAT-HSP interactions with HSC70 and HSP90 by proximity ligation assay (PLA) in SN56 cells transiently expressing heterologous wild-type (WT) ChAT. Formalin-fixed cells were first co-labeling with goat anti-ChAT together with either mouse anti-HSC70 or mouse anti-HSP90 primary antibodies, then incubated with oligonucleotide- linked secondary antibodies. Following DNA ligation and DNA amplification using the Duolink In Situ Orange Kit (Sigma), in situ ChAT-HSP interactions were imaged by confocal microscopy at an excitation/emission of 554/576 nm. Positive in situ ChAT-HSP interactions where visualized and are presented as fluorescent red dots while nuclei were stained with DAPI (blue). Control cells were either transfected with empty pcDNA3.1+ vector or had primary antibodies omitted from the assay (No 1o antibodies). Images are representative of 3 independent experiments; scale bars are 10 µm.

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and incubated with primary antibodies, or when primary antibodies recognizing ChAT, HSC70, or HSP90 were omitted from the assay.

3.9: Inhibition of HSC/HSP70-substrate protein interactions promotes accumulation of high molecular mass Triton-insoluble ChAT aggregates

Functionally, HSPs aid in the stabilization of cellular proteins by interacting with exposed peptide sequences rich in hydrophobic amino acids, thereby preventing unintended protein aggregation and promoting protein folding and solubility [118, 135, 202, 503]. Therefore, to test whether HSC/HSP70 may regulate the folding of ChAT protein, I treated SN56 cells, transiently expressing wild-type or mutant ChAT, with the HSC/HSP70 inhibitor 2- phenylethynesulfonamide (PES), a small-molecule that interferes with HSC/HSP70- substrate interactions leading to substrate insolubilization and the subsequent accumulation of misfolded and aggregation-prone proteins [216-217]. Cells were treated with 20 µM PES for 24 h, then levels of Triton-soluble and insoluble ChAT protein were assessed by immunoblotting to determine levels of misfolded and/or aggregated ChAT (Figure 3.27a). Importantly, I found that PES treatment of cells did not alter the Triton-soluble levels of either wild-type or mutant ChAT proteins (Figure 3.27b), but levels of Triton-insoluble wild-type (p≤0.01), P17A/P19A- (p≤0.001), V18M- (p≤0.01) and A513T-ChAT proteins (p≤0.001) were significantly increased compared to DMSO-treated control cells (Figure 3.27c). Moreover, immunoblots of Triton-insoluble samples from PES-treated cells revealed accumulation of high molecular mass ChAT-immunoreactive proteins and the presence of insoluble ChAT that was retained in the loading pockets of the stacking layer of SDS-PAGE gels (Figure 3.27a, top panel, anti-ChAT long-exposure). The accumulation of high molecular mass Triton-insoluble ChAT was greater for mutant ChAT than for wild- type ChAT, with levels of Triton-insoluble P17A/P19A-ChAT having been the greatest. Interestingly, high molecular mass Triton-insoluble P17A/P19A-ChAT was also observed in DMSO-treated cells, though at a lower abundance than following PES treatment.

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Figure 3.27: Inhibition of HSC/HSP70-substrate protein interaction promotes accumulation of high molecular mass Triton-insoluble ChAT aggregates. (A) Anti- ChAT immunoblots from SN56 cells expressing wild-type (WT) or mutant ChAT that were treated for 24 h with either 20 μM PES, a HSC/HSP70 inhibitor, or DMSO-vehicle. Cells were lysed in buffer containing 0.1% Triton X-100 and fractionated into soluble and insoluble proteins. Triton X-100 insoluble proteins were solubilized prior to SDS-PAGE by denaturing in 2x Laemmli sample buffer with 5% 2-mercaptoethanol. PES treatment resulted in production of high molecular mass Triton-insoluble ChAT-immunoreactive proteins that accumulated in the loading pockets of the SDS-PAGE gels (top panel, anti- ChAT long-exposure). The abundance of insoluble ChAT in the loading pockets appears to be greater for mutant ChAT as compared to wild-type ChAT, with levels of Triton- insoluble P17A/P19A-ChAT appearing the greatest. Anti-actin immunoblotting from both Triton-soluble and -insoluble samples was completed as a loading control. (B) Treatment with 20 μM PES had no effect on the total levels of Triton-soluble ChAT protein. (C) PES treatment significantly increased the levels of Triton-insoluble wild-type (**p≤0.01), P17A/P19A- (***p≤0.001), V18M- (**p≤0.01) and A513T-ChAT (***p≤0.001) proteins compared to vehicle-control (two-way ANOVA with Bonferroni’s post-hoc test, mean ± SEM, n=6). (D) Triton-insoluble protein samples from (A) were analyzed by semi- denaturing detergent agarose gel electrophoresis (SDD-AGE) revealing the presence of insoluble SDS-resistant ChAT aggregates in PES-treated cells. The abundance of these SDS-resistant ChAT aggregates is greater for mutant ChAT compared to wild-type ChAT, with P17A/P19A-ChAT being the greatest (n=6).

To assess whether high molecular mass Triton-insoluble ChAT observed following PES treatment may consist of detergent-resistant ChAT aggregates, I performed semi- denaturing detergent agarose gel electrophoresis (SDD-AGE) [491, 504] on Triton- insoluble protein samples assessed in Figure 3.27a. Using anti-ChAT immunoblotting, I observed accumulation of high molecular mass SDS-resistant ChAT protein aggregates from lysates of PES-treated cells, but not generally from DMSO-treated control cells (Figure 3.27d). The abundance of these insoluble SDS-resistant ChAT aggregates was

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greater for mutant ChAT as compared to wild-type ChAT, with P17A/P19A-ChAT having been the greatest. Moreover, insoluble SDS-resistant aggregates of P17A/P17A-ChAT were found in DMSO-treated cells, suggesting that this mutation may promote ChAT insolubility and aggregation under basal conditions.

3.10: Inhibition of HSC/HSP70 or HSP90 ATPase activity reduces ChAT steady-state protein levels

In addition to interaction with substrate proteins the protein-chaperoning function of HSC/HSP70 requires activity of its N-terminal ATPase domain, and inhibition of this domain by the small-molecule VER-155008 has been shown to promote the degradation of HSC/HSP70 clients [213-214]. Thus, to determine if HSC/HSP70 activity is required for ChAT protein stability, SN56 cells expressing either wild-type (Figure 3.28a), P17A/P19A- (Figure 3.28b), V18M- (Figure 3.28c) or A513T-ChAT (Figure 3.28d) were treated with varying concentrations (5-50 µM) of VER-155008 for 24 h. When analyzed by anti-ChAT immunoblotting, I observed dose-related reductions in the steady-state protein levels for both wild-type and mutant ChAT in VER-155008 treated cells. Interestingly, mutant ChAT appears more sensitive than wild-type ChAT to HSC/HSP70 inhibition. Specifically, a significant decrease in wild-type ChAT steady-state protein level was observed when cells were treated with at least 20 µM VER-155008 (p≤0.05; Figure 3.28e), while lower concentrations of VER-155008 were sufficient to significantly reduce protein levels of P17A/P19A- (5 µM; p≤0.001; Figure 3.28f), V18M- (5 µM; p≤0.001; Figure 3.28g), or A513T-ChAT (10 µM; p≤0.001; Figure 3.28h), as compared to DMSO-treated control cells. Furthermore, treatment of ChAT-expressing cells with 50 µM VER-155008 reduced ChAT steady-state proteins levels, compared to DMSO-treated control cells, as follows: wild-type ChAT (38.6 ± 4.97%), P17A/P19A-ChAT (61.0 ± 4.17%), V18M-ChAT (75.7 ± 2.82%) and A513T-ChAT (53.6 ± 2.69%).

Like HSC/HSP70, HSP90 has an N-terminal ATP-binding domain with ATPase activity that is required for HSP90 chaperoning function, and pharmacological inhibition

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Figure 3.28: Inhibition of HSC/HSP70 ATPase activity reduces ChAT steady-state protein levels in a dose-related manner. Anti-ChAT immunoblots from SN56 cells expressing wild-type (WT; A), P17A/P19A- (B), V18M- (C) or A513T-ChAT (D) that were treated for 24 h with 5-50 µM VER-155008, an inhibitor of HSC/HSP70 activity. Control cells were treated with DMSO-vehicle. Anti-actin immunoblotting was completed as a loading control. Steady-state protein levels of wild-type (E), P17A/P19A- (F), V18M- (G) and A513T-ChAT (H) are reduced following treatment with VER-155008 as compared to DMSO-control (**p≤0.01, ***p≤0.001, one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=4). Mutant ChAT appears more sensitive to HSC/HSP70 inhibition where treatment with 50 µM VER-155008 reduced ChAT steady-state proteins levels, compared to vehicle-control, as follows; wild-type ChAT (38.63% ± 4.97), P17A/P19A-ChAT (61.04% ± 4.17), V18M-ChAT (75.71% ± 2.82), and A513T-ChAT (53.60% ± 2.69).

of this domain by the small molecule 17-allylamino-17-demethoxygeldanamycin (17- AAG) has been shown to promote proteasomal degradation of HSP90 clients [286-287]. Therefore, to determine whether HSP90 activity may also promote ChAT protein stabilization, I treated ChAT-expressing SN56 cells with 0.5-2 µM 17-AAG for 24 h (Figure 3.29). Analysis by anti-ChAT immunoblotting showed that treatment of cells with concentrations of 17-AAG up to 2 µM had no effect on the steady-state levels of wild-type ChAT protein (Figures 3.29a and 3.29e). Importantly though, treatment with 0.5-2 µM 17- AAG did significantly reduce the steady-state levels of mutant P17A/P19A- (p≤0.001; Figures 3.29b and 3.29f), V18M- (p≤0.001; Figures 3.29c and 3.29g), and A513T-ChAT proteins (p≤0.001; Figures 3.29d and 3.29h) as compared to DMSO-treated control cells.

Overall, these data using the HSP inhibitors VER-155008 and 17-AAG suggest that while the ATPase activity of HSC/HSP70 is necessary for both wild-type and mutant ChAT protein stability, the ATPase activity of HSP90 may be uniquely critical for stabilization of mutant ChAT, but not of wild-type ChAT protein. Interestingly, previous studies have shown that treatment of cells with 17-AAG can lead to the expression of stress-inducible HSP70 [287, 505-506]. Therefore, to assess whether 17-AAG-induced HSP70 may have a

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Figure 3.29: Inhibition of HSP90 by 17-AAG specifically reduces steady-state protein levels of mutant ChAT. Anti-ChAT immunoblots from SN56 cells expressing wild-type (WT; A), P17A/P19A- (B), V18M- (C), or A513T-ChAT (D) that were treated for 24 h with 0.5-2 µM with 17-AAG, an inhibitor of HSP90 activity. Control cells were treated with DMSO-vehicle. Anti-GAPDH immunoblotting was completed as a loading control. (E) Treatment of cells with 17-AAG at concentrations up to 2 µM has no effect on the steady-state levels of wild-type ChAT protein. By comparison, steady-state protein levels of P17A/P19A- (F), V18M- (G), and A513T-ChAT (H) are reduced following treatment of cells with 17-AAG as compared to DMSO-control (***p≤0.001, one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=4).

protective role towards ChAT protein stability during HSP90 inhibition I performed siRNA-mediated knock-down of HSP70 in SN56 cells, transiently expressing wild-type or mutant ChAT, that were treated with 1 µM 17-AAG for 24 h (Figure 3.30). In support of Figure 3.29, treatment with 17-AAG once again reduced the proteins levels of mutant P17A/P19A-, V18M-, and A513T-ChAT but had no effect on the steady-state protein levels of wild-type ChAT as compared to DMSO-treated control cells. While I found that 17- AAG treatment did induce HSP70 protein expression in SN56 cells, siRNA knock-down of HSP70 had no additional effect on the loss of steady-state protein levels of mutant ChAT following 17-AAG treatment and, furthermore, failed to sensitize wild-type ChAT to 17- AAG-induced loss of ChAT protein.

3.11: Proteasome inhibition prevents loss of ChAT protein following HSC/HSP70 and HSP90 inhibition

My earlier results suggest that ChAT protein degradation is regulated by the proteasome (Figures 3.6, 3.7, 3.15). Therefore, to test whether the loss of ChAT protein following inhibition of either HSC/HSP70 or HSP90 was due to proteasome-dependent degradation of ChAT, I co-treated ChAT-expressing SN56 cells with either VER-155008 or 17-AAG

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Figure 3.30: siRNA-mediated knock-down of HSP70 fails to sensitize wild-type ChAT to 17-AAG-induced loss of ChAT protein. Anti-ChAT immunoblots from SN56 cells transiently expressing either wild-type (WT) or mutant ChAT that were co-transfected with 25 nM of either anti-HSP70 siRNA or scramble-control siRNA for 72 h. In addition, cells were treated for the final 24 h of siRNA incubation with either 1 µM 17-AAG or with DMSO-vehicle control. 17-AAG-induced expression of and subsequent siRNA knock- down of HSP70 was assessed by anti-HSP70 immunoblotting. Anti-actin immunoblotting was completed as a loading control (n=3).

along with the proteasome inhibitor MG132. Importantly, I observed that treatment of cells expressing either wild-type and mutant ChAT with 40 µM VER-155008 for 24 h reduced ChAT steady-state protein levels, and that co-treatment of cells with 5 µM MG132 for 18 h attenuated this effect (Figure 3.31a). In addition, treatment of ChAT-expressing cells with 1 µM 17-AAG alone for 24 h reduced the steady-state protein levels of mutant ChAT only,

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Figure 3.31: Proteasome inhibition by MG132 treatment prevents loss of ChAT protein following inhibition of HSC/HSP70 or HSP90 activity. Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type (WT) or mutant ChAT that were co-treated with either 40 µM VER-155008 (A) or 1 µM 17-AAG (B) for 24 h and with 5 µM MG132 for the final 18 h of HSP inhibition (n=4). Lysosomal inhibition by co- treatment of ChAT-expressing cells with 50 µM chloroquine (CQ) for 18 h did not prevent the effects of 24 h treatment with either VER-155008 (C) or 17-AAG (D) on ChAT steady- state protein levels (n=3). Control cells were treated with DMSO-vehicle or alone with either VER-155008 or 17-AAG. Proteasome inhibition by MG132 treatment or lysosome inhibition by CQ treatment was validated by immunoblotting for the accumulation of either ubiquitinated cellular proteins or for the lysosome-associated protein LC3B-II, respectively. Anti-actin immunoblotting was completed as a loading control.

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while co-treatment with 5 µM MG132 for 18 h also prevented this effect (Figure 3.31b). As anticipated, and in support of my earlier findings, co-treatment of ChAT-expressing SN56 cells for 18 h with 50 µM chloroquine to inhibit lysosomal proteolysis did not prevent the effects of either 40 µM VER-155008 (Figure 3.31c) or 1 µM 17-AAG (Figure 3.31d) on the steady-state levels of either wild-type or mutant ChAT protein.

3.12: Inhibition of HSC/HSP70 and HSP90 activity enhances ChAT ubiquitination and reduces ChAT enzymatic activity

In addition to protein folding, HSPs perform essential protein quality control functions by triaging stress-damaged or terminally misfolded proteins for degradation through the ubiquitin-proteasome system [118, 135]. Therefore, to determine if inhibition of either HSC/HSP70 or HSP90 may promote degradation of ChAT through enhanced ChAT ubiquitination, I treated ChAT-expressing SN56 cells with either 40 µM VER-155008 for 24 h or 1 µM 17-AAG for either 8 or 24 h, along with 20 µM MG132 during the final 6 h of HSP inhibition to prevent the degradation of ubiquitinated ChAT. By anti-ubiquitin immunoblotting from anti-ChAT IPs, I observed that treatment of cells with VER-155008 resulted in enhanced ubiquitination of both wild-type and mutant ChAT as compared to DMSO-treated control cells (Figure 3.32a). Furthermore, ubiquitination of mutant ChAT, in particular P17A/P19A-ChAT, was greater than that of wild-type ChAT following VER- 155008 treatment when compared to DMSO-treated control cells. Surprisingly, while treatment of cells with 17-AAG for 24 h had no effect on the ubiquitination of either wild- type or mutant ChAT proteins, treatment for 8 h did enhance ChAT ubiquitination, with this effect being greater for mutant than for wild-type ChAT, particularly for P17A/P19A- ChAT, when compared to DMSO-control (Figure 3.32b).

Canonically, ubiquitination targets proteins for proteasomal and/or lysosomal degradation, but it can also regulate subcellular trafficking, receptor endocytosis, and enzymatic activity of various proteins [44]. Therefore, to determine if HSP inhibition may also affect ChAT enzymatic activity, I treated SN56 cells transiently expressing

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Figure 3.32: Inhibition of HSC/HSP70 and HSP90 activity enhances ChAT ubiquitination. (A) Representative anti-ChAT and anti-ubiquitin immunoblots from SN56 cells expressing wild-type (WT) or mutant ChAT that were treated for 24 h with 40 μM VER-155008. To inhibit degradation of ubiquitinated ChAT cells were co-treated for the final 6 h of VER-155008 treatment with 20 μM MG132. ChAT ubiquitination was then assessed by anti-ubiquitin immunoblotting following anti-ChAT immunoprecipitation (IP). Inhibition of HSC/HSP70 by VER-155008 treatment enhanced ChAT ubiquitination, where levels of ubiquitinated mutant ChAT, particularly P17A/P19A-ChAT, are greater than that of wild-type ChAT. Control cells were transfected with empty pcDNA3.1+ vector and/or were treated with DMSO-vehicle. Proteasome inhibition by MG132 treatment was validated by immunoblotting from whole cell lysates for the accumulation of ubiquitinated cellular proteins. Anti-actin immunoblotting from whole cell lysates was completed as a loading control (n=3). (B) Immunoblots following anti-ChAT IP from ChAT-expressing SN56 cells that were co-treated for either 8 or 24 h with 1 μM 17-AAG and for the final 6 h with 20 μM MG132. Inhibition of HSP90 by treatment with 1 μM 17-AAG for 8 h, but not for 24 h, enhanced ubiquitination of wild-type and mutant ChAT; the ubiquitination of ChAT appears to be greater for mutant ChAT than for wild-type ChAT following 17-AAG treatment, with levels of ubiquitinated P17A/P19A-ChAT being the greatest (n=3).

heterologous wild-type ChAT with either 40 µM VER-155008 or 1 µM 17-AAG alone or in combination for 24 h, then analyzed cellular ChAT activity by radioenzymatic assay. Interestingly, I found that the cellular activity of wild-type ChAT is reduced following 24 h treatment of cells with either 40 μM VER-155008, 1 µM 17-AAG, or a combination of the two as compared to DMSO-treated control cells (p≤0.01; Figure 3.33). While treatment with VER-155008 alone trended towards a greater reduction in cellular ChAT activity when compared to 17-AAG alone (p=0.058), co-treatment of cells with VER-155008 and 17-AAG did lead to a greater reduction in wild-type ChAT activity as compared to 17- AAG alone (p≤0.01). Importantly, these data show that while inhibition of HSP90 by 17- AAG treatment had no effect on the steady-state protein levels of wild-type ChAT (Figures

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Figure 3.33: Inhibition of HSC/HSP70 and HSP90 activity reduces cellular enzymatic activity of wild-type ChAT. Cellular enzymatic activity of heterologously-expressed wild- type ChAT is reduced following treatment of SN56 cells for 24 h with either 40 μM VER- 155008 or 1 µM 17-AAG alone or a combination of the two as compared to DMSO-control treated cells (**p≤0.01). While treatment with VER-155008 trended towards a greater reduction in cellular ChAT activity compared to 17-AAG (p=0.058), co-treatment with these inhibitors did result in a greater reduction in ChAT activity as compared to 17-AAG alone (**p≤0.01, one-way ANOVA with Tukey’s post-hoc test, mean ± SEM, n=3).

3.29 and 3.31), this treatment did significantly reduce the cellular enzymatic activity of wild-type ChAT like that observed with HSC/HSP70 inhibition by VER-155008 treatment, suggesting that the folding activity of both HSC/HSP70 and HSP90 are necessary in order to generate fully-functional and active ChAT enzyme.

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3.13: ChAT interacts with the HSP-associated E3 ubiquitin ligase CHIP

The E3 ubiquitin ligase CHIP/Stub1 ubiquitinates proteins in an HSP-dependent manner through direct interaction with either HSC/HSP70 or HSP90 and their bound substrate proteins [76-78]. My results demonstrating enhanced ubiquitination and proteasomal degradation of ChAT protein following HSC/HSP70 and HSP90 inhibition suggest that ubiquitination of ChAT may be regulated by CHIP. Therefore, to first determine if ChAT may interact with CHIP, I performed anti-ChAT co-IPs from SN56 cells co-expressing either wild-type or mutant ChAT with FLAG-tagged CHIP (Figure 3.34a). As anticipated, FLAG-CHIP was detected in anti-ChAT co-IP samples from both wild-type and mutant ChAT-expressing cells by anti-FLAG immunoblotting. By comparing the levels of immunoprecipitated FLAG-CHIP to that of ChAT protein, I observed enhanced co-IP of FLAG-CHIP with mutant P17A/P19A- (p≤0.001), V18M- (p≤0.001) and A513T-ChAT (p≤0.05) as compared to wild-type ChAT (Figure 3.34b). Subsequently, association of wild-type and mutant ChAT with endogenous CHIP was confirmed by anti-ChAT co-IP from ChAT-expressing SN56 cells with follow-up anti-CHIP immunoblotting (Figure 3.34c). Lastly, to assess whether ChAT can interact with CHIP in situ I also performed PLA assays to visualize and confirm, by confocal microscopy, the presence of in situ interactions between wild-type ChAT and endogenous CHIP in SN56 cells (Figure 3.35). As predicted, I observed predominantly cytoplasmic in situ interaction of wild-type ChAT with endogenous CHIP in cells. As controls, I did not observe in situ interactions in control cells that were either transfected with either empty vector or when primary antibodies towards ChAT and CHIP were omitted from the assay.

3.14: CHIP co-expression promotes proteasomal degradation and insolubilization of ChAT

Interaction of ChAT with CHIP may promote the ubiquitination and proteasome-dependent degradation of ChAT. Therefore, to determine if ChAT is a substrate for CHIP, I first tested if co-expression of FLAG-CHIP promotes loss of ChAT protein in SN56 cells transiently

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Figure 3.34: Co-immunoprecipitation of ChAT with HSP-associated E3 ubiquitin ligase C-terminus of HSC70-interaction protein (CHIP). (A) Immunoblots showing co- immunoprecipitation (co-IP) of ChAT with FLAG-CHIP from SN56 cells co-expressing either wild-type (WT) or mutant ChAT with FLAG-tagged CHIP. Control cells were transfected with empty pcDNA3.1+ vector or to express FLAG-CHIP alone. Anti-FLAG and anti-actin immunoblotting from whole cell lysates was completed as loading controls. (B) Co-IP of ChAT with FLAG-CHIP is enhanced for P17A/P19A- (***p≤0.001), V18M- (***p≤0.001), and A513T-ChAT (*p≤0.05) as compared to wild-type ChAT (one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=5). (C) Co-IP of heterologously- expressed wild-type and mutant ChAT with endogenous CHIP following anti-ChAT co-IP from SN56 cells. Control cells were transfected with empty vector (n=3).

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Figure 3.35: In situ interaction of ChAT with endogenous CHIP. Detection of in situ ChAT-CHIP interactions by proximity ligation assay (PLA) in SN56 cells transiently expressing heterologous wild-type ChAT. Formalin-fixed cells were first co-labeling with goat anti-ChAT together with rabbit anti-CHIP primary antibodies, then incubated with oligonucleotide-linked secondary antibodies. Following DNA ligation and DNA amplification using the Duolink In Situ Orange Kit (Sigma), in situ ChAT-CHIP interactions were imaged by confocal microscopy at an excitation/emission of 554/576 nm. Positive in situ ChAT-CHIP interactions were visualized and presented as fluorescent red dots while nuclei were stained with DAPI (blue). Control cells were either transfected with empty pcDNA3.1+ vector or primary antibodies omitted from the assay (No 1°). Images are representative of 4 independent experiments; scale bars are 10 µm.

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expressing wild-type or mutant ChAT. By anti-ChAT immunoblotting (Figure 3.36a) I observed that co-expression of FLAG-CHIP significantly reduced the steady-state protein levels of wild-type (p≤0.001), P17A/P19A- (p≤0.05), V18M- (p≤0.001), and A513T-ChAT (p≤0.001) as compared to cells co-transfected with equal amounts (µg) of empty vector (Figure 3.36b). To assess whether the effects of FLAG-CHIP co-expression on reduced ChAT protein levels were proteasome dependent, SN56 cells, transiently co-expressing wild-type ChAT with FLAG-CHIP, were treated for 18 h with either 5 µM MG132 or 50 µM chloroquine alone or with these two inhibitors together in combination. Following anti- ChAT immunoblotting I found that proteasome inhibition by treatment with either MG132 alone or together with chloroquine partially ameliorated the FLAG-CHIP-induced loss of wild-type ChAT protein (Figure 3.36c). As anticipated, treatment with chloroquine alone failed to prevent the loss of wild-type ChAT protein in cells co-expressing FLAG-CHIP.

CHIP has two distinct functional protein domains, a C-terminal U-box domain that contains E3 ubiquitin ligase activity and three N-terminal TPR domains that allow for and facilitate interaction of CHIP with HSPs and their substrates (Figure 3.37a) [73, 76, 276, 315]. To determine if the E3 ligase activity of CHIP is required to promote CHIP-associated loss of ChAT protein, and furthermore whether the TPR domains of CHIP are required for ChAT-CHIP interaction, SN56 cells were transiently transfected to co-express wild-type ChAT with either full-length wild-type FLAG-CHIP or mutant FLAG-CHIP missing either the N-terminal TPR domains (ΔTPR) or the C-terminal U-box domain (ΔU-box). Cells were collected, lysed in buffer containing 0.5% Triton X-100, and lysates were used for completion of both anti-ChAT co-IPs and for the immunoblot assessment of Triton-soluble and -insoluble ChAT proteins (Figure 3.37b). By anti-FLAG immunoblotting from anti- ChAT co-IP samples I observed that wild-type ChAT can co-IP with both full-length FLAG-CHIP or FLAG-CHIP-ΔU-box but was unable to co-IP with FLAG-CHIP-ΔTPR. Surprisingly, co-expression of either full-length FLAG-CHIP or FLAG-CHIP-ΔU-box reduced the Triton-soluble levels of ChAT protein and led to a reciprocal increase in the Triton-insoluble levels of ChAT. These effects on ChAT solubility were not observed when ChAT was co-expressed with FLAG-CHIP-ΔTPR, suggesting that the TRP

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Figure 3.36: CHIP co-expression promotes proteasomal degradation of ChAT. (A) Anti-ChAT immunoblots from SN56 cells co-expressing wild-type (WT) or mutant ChAT with FLAG-tagged CHIP. Anti-FLAG and anti-actin immunoblotting was completed as loading controls. (B) The steady-state proteins levels of wild-type (***p≤0.001), P17A/P19A- (*p≤0.05), V18M- (***p≤0.001), and A513T-ChAT (*p≤0.05) are reduced following co-expression of FLAG-CHIP as compared to control cells co-transfected with empty pcDNA 3.1+ vector (two-way ANOVA with Bonferroni’s post-hoc test, mean ± SEM, n=4). (C) Proteasome inhibition partially attenuates loss of ChAT protein following co-expression of FLAG-CHIP. Immunoblots from SN56 cells co-expressing wild-type ChAT with FLAG-tagged CHIP that were treated with either 5 µM MG132 or 50 µM chloroquine (CQ) alone or in combination for 18 h. Control cells were treated with DMSO- vehicle and/or transfected with either empty pcDNA 3.1+ vector or to express wild-type ChAT alone. Proteasome inhibition by MG132 treatment was validated by immunoblotting for the accumulation of ubiquitinated cellular proteins (n=6).

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Figure 3.37: The TPR domains of CHIP are critical for ChAT-CHIP protein interaction and promote CHIP-mediated ChAT insolubilization. (A) Schematic overview of the functional protein domains of the E3 ubiquitin ligase CHIP/Stub1. CHIP contains three N-terminal tetracopeptide repeat (TPR) domains that facilitate interaction of CHIP with HSPs and their substrates. Additionally, a C-terminal U-box domain imparts E3 ligase activity to CHIP. (B) The TPR domains, but not the U-box domain, of CHIP are necessary for ChAT-CHIP protein interaction. Additionally, co-expression of either full- length FLAG-CHIP or FLAG-CHIP-ΔU-box increased Triton-insoluble levels of wild-type (WT) ChAT protein while ChAT co-expression with FLAG-CHIP-ΔTPR had no effect on ChAT solubility. Shown are representative anti-ChAT and anti-FLAG immunoblots following anti-ChAT co-immunoprecipitation (co-IP) from SN56 cells transiently co- expressing wild-type ChAT with either full-length FLAG-tagged CHIP or with mutant FLAG-CHIP missing either its TPR domains (ΔTPR) or U-box domain (ΔU-box). Control cells were transfected with empty pcDNA3.1+ vector or to express either wild-type ChAT or full-length FLAG-CHIP alone. Cells were lysed in 0.5% Triton X-100, fractionated into Triton-soluble and -insoluble fractions, and anti-ChAT co-IPs were completed from the soluble fractions. Triton X-100 insoluble proteins were solubilized prior to SDS-PAGE by denaturing in 2x Laemmli sample buffer. Anti-actin immunoblotting from Triton-soluble and -insoluble proteins samples was completed as loading controls (n=5).

domains of CHIP are critical for not only ChAT-CHIP association, but also for the effects of CHIP co-expression on ChAT protein solubility and degradation.

These initial studies investigating the role of CHIP on the regulation of ChAT protein degradation and stability were completed in cells co-expressing ChAT with heterologous FLAG-tagged CHIP. Though I was able to show that ChAT can co-IP and interact in situ with endogenous CHIP (Figures 3.34c and 3.35), it is unknown whether ChAT may be a substrate for degradation by endogenous CHIP. Thus, to determine if endogenous CHIP regulates ChAT protein stability I monitored ChAT steady-state protein levels from SN56 cells, transiently expressing wild-type or mutant ChAT, that were co-

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transfected with siRNA targeting endogenous mouse CHIP (Figure 3.38). Surprisingly, as assessed by anti-ChAT and anti-CHIP immunoblotting, while I was able to achieve a greater-than 95% CHIP knock-down in cells, siRNA-mediated knock-down of endogenous CHIP had no effect on the steady-state levels of either wild-type or mutant ChAT protein.

3.15: Oxidative stress by exposure of cells to H2O2 enhances ChAT ubiquitination

In summary of my results investigating the regulation of ChAT by HSPs, I have shown that ChAT interacts with and is a substrate for HSC/HSP70 and HSP90, where inhibition of HSC/HSP70 protein interactions results in accumulation of insoluble ChAT protein, and inhibition of HSC/HSP70 and HSP90 activity led to reduced cellular ChAT activity, along with enhanced ChAT ubiquitination and proteasome-dependent loss of ChAT protein. Furthermore, the effects of HSP inhibition appear greater for mutant ChAT (P17A/P19A-, V18M-, and A513T-ChAT) than for wild-type ChAT (Figure 3.39).

HSPs, such as HSC/HSP70 and HSP90, have been best characterized for their role in promoting the folding and stabilization of nascent proteins, as well as in refolding of misfolding-prone and aggregated proteins during proteotoxic stress such as hyperthermia [118, 135, 154, 371]. Oxidation of cellular proteins and subsequent formation of aberrant stress-induced disulfide bonds during oxidative stress can induce protein misfolding with severe functional consequences [361, 364-365]. Interestingly, ChAT has a higher-than- average cysteine content of 3.2% compared to 1.6% for intracellular mammalian proteins [367] which, like the cysteine-rich E3 ubiquitin ligase Parkin, may sensitize ChAT to oxidative damage [370]. Importantly, work performed in our laboratory by Shawn Albers, M.Sc., has demonstrated that treatment of ChAT-expressing SN56 cells with 50 µM – 2 mM of H2O2, a potent inducer of cellular oxidative stress, results in the dose- and redox- dependent accumulation of higher molecular mass wild-type ChAT proteins that can be observed by anti-ChAT immunoblotting specifically in non-reducing conditions (Figure 3.40a). Additionally, Shawn Albers has shown that treatment of ChAT-expressing SN56

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Figure 3.38: siRNA knock-down of endogenous CHIP has no effect on the steady-state protein levels of wild-type or mutant ChAT. Anti-ChAT immunoblots from ChAT- expressing SN56 cells that were transfected with 25 nM of either anti-CHIP siRNA or scramble-control siRNA for 72 h. Cells were co-transfected to express wild-type (WT) or mutant ChAT for the final 48 h of siRNA incubation. Control cells were mock-transfected with siLentFect transfection reagent. Knock-down of CHIP was assessed by anti-CHIP immunoblotting. Anti-actin immunoblotting was completed as a loading control (n=4).

cells for up-to 4 h with 1 mM H2O2 reduced the cellular activity of wild-type ChAT when compared to vehicle-treated control cells (p<0.001; Figure 3.40b) [507]. Lastly, Nunes- Tavares and colleagues [439] have demonstrated that cellular ChAT activity is reduced following exposure of neural cells to Aβ oligomers, a hallmark of AD and stimulator of cellular oxidative stress [429, 508-509].

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Figure 3.39: Model for the regulation of ChAT protein stability by the heat-shock family of molecular chaperones. Under physiological conditions ChAT protein folding is promoted by the molecular chaperoning-function of the heat shock proteins HSC/HSP70 and HSP90, resulting in stable and active ChAT enzyme. Though, if ChAT protein folding becomes compromised, potentially in case of ChAT mutations (e.g. P17A/P19A, V18M, and A513T) or following HSP inhibition (e.g. PES, VER-155008, and 17-AAG), misfolded and/or insoluble ChAT is targeted for proteasomal degradation via enhanced ubiquitination by a yet-to-be identified chaperone-associated E3 ubiquitin ligase. It is currently unknown whether cellular stresses, such as oxidative stress, can induce ChAT dysfunction through enhanced misfolding and subsequent degradation of ChAT, and/or if HSPs may assist in the stabilization of ChAT protein during cellular stress.

The aforementioned studies suggest that ChAT protein function, and potentially protein stability, may be altered during cellular exposure to oxidative stress. Additional studies have demonstrated that the ubiquitination, and subsequent proteasomal degradation, of various cellular proteins is altered during oxidative stress [510-512]. Importantly, as presented earlier in this thesis, I have demonstrated that ChAT protein stability is regulated by the ubiquitin-proteasome system, though it is unknown whether ChAT ubiquitination is altered during oxidative stress. Therefore, ChAT ubiquitination was monitored in SN56 cells, transiently co-expressing wild-type ChAT with HA-tagged ubiquitin, that were

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Figure 3.40: H2O2 treatment promotes the generation of redox-dependent higher molecular mass ChAT proteins and decreases cellular ChAT activity. (A) Anti-Myc immunoblots from SN56 cells, transiently expressing Myc-tagged 69-kDa wild-type

ChAT, that were treated for 1 hour with 1 mM H2O2. Control cells were transfected with empty pcDNA3.1+ vector or were treated with vehicle-control (0 µM; water). Treatment of ChAT-expressing cells with H2O2 generated higher molecular mass ChAT proteins that were observed on anti-Myc immunoblots only under non-reducing conditions. Anti-actin immunoblotting was completed as a loading control (n=3). (B) Cellular ChAT activity is reduced in SN56 cells following treatment with 1 mM H2O2 for 240 min as compared to vehicle-treated cells (***p≤0.001, one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM, n=4). Experiments were completed previously by Shawn Albers, M.Sc. [507].

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treated for 30 – 60 min with 1 mM H2O2 to induce cellular oxidative stress. To prevent degradation of ubiquitinated ChAT cells were co-treated for 18 h with 5 µM MG132 prior- to and during treatment with H2O2. Following anti-HA IP and anti-ChAT immunoblotting I observed that ChAT ubiquitination is enhanced following treatment of cells with 1 mM

H2O2 for either 30 or 60 min as compared to DMSO-treated control cells (Figure 3.41). No differences in the abundance of ubiquitinated ChAT were detected when comparing the 30 min H2O2-treated cells to the 60 min-treated cells.

3.16: Oxidative stress by H2O2 treatment promotes the insolubilization of ChAT protein

These preliminary ubiquitination experiments following H2O2 treatment suggest that ChAT protein stability may be negatively affected by oxidative damage. Furthermore, work by Shawn Albers showing the appearance of higher molecular mass ChAT proteins under non- reducing conditions following H2O2 exposure suggests that ChAT may be susceptible to oxidative misfolding that is dependent on cysteine oxidation and formation of aberrant disulfide bonds. Therefore, to first assess whether ChAT solubility, and potentially ChAT protein folding, is altered during oxidative stress I treated SN56 cells transiently expressing wild-type ChAT with varying concentrations of H2O2 for either 24 h (100 nM – 200 µM) or for 1 h (1 µM – 2 mM). Cells were lysed with buffer containing 0.1% Triton X-100 and Triton-soluble and -insoluble ChAT proteins were fractionated. Following anti-ChAT immunoblotting I observed that treatment of ChAT-expressing cells for 24 h with 100 nM

– 200 µM H2O2 had no significant effect on the overall solubility of ChAT protein (Figure 3.42a). Interestingly, though no changes in Triton-soluble levels of ChAT were detected, enhanced levels of Triton-insoluble ChAT were observed following 1 h treatment of cells specifically with 0.5-2 mM H2O2 compared to vehicle-treated control cells (Figure 3.42b).

HSPs, such as HSC/HSP70 and HSP90, have been shown to bind to and chaperone oxidized proteins to maintain and/or restore protein function during oxidative conditions [371]. My earlier studies using the HSC/HSP70 inhibitor PES suggest that the folding and

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Figure 3.41: Oxidative stress by H2O2 treatment enhances ChAT ubiquitination. Representative anti-ChAT and anti-HA immunoblots from SN56 cells transiently co- expressing wild-type ChAT with HA-tagged ubiquitin (HA-Ub) that were treated with 1 mM H2O2 for either 30 or 60 min. Levels of ubiquitinated ChAT protein were assessed by anti-ChAT immunoblotting from anti-HA IP samples. Cells were treated with 5 µM

MG132 for 18 h prior-to and during H2O2 treatment to prevent proteasomal degradation of ubiquitinated proteins. Control cells were either transfected with empty pcDNA3.1+ vector or to express wild-type ChAT or HA-Ub alone, or were treated with vehicle-control (water). Proteasome inhibition by MG132 treatment was validated by anti-HA immunoblotting from whole cell lysates for the accumulation of ubiquitinated cellular proteins. Anti-actin immunoblotting from whole cell lysates was completed as a loading control (n=3).

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Figure 3.42: Oxidative stress by H2O2 treatment promotes ChAT insolubilization. Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type ChAT that were treated with either 0.1 – 200 µM H2O2 for 24 h (A) or with 1 µM – 2 mM H2O2 for 1h (B). Control cells were treated with water-vehicle. Cells were lysed in 0.1% Triton X-100 and Triton-soluble and -insoluble proteins were fractionated. Triton X-100 insoluble proteins were solubilized prior to SDS-PAGE by denaturing in 2x Laemmli sample buffer with 5% 2-mercaptoethanol. No differences in ChAT protein solubility was detected following 24 h of 0.1 – 200 µM H2O2 treatment (A; n=3). While no changes in the Triton-soluble levels of

ChAT were detected following 1h H2O2 treatment, levels of Triton-insoluble ChAT protein were increased in cells specifically treated for 1 h with 0.5 – 2 mM H2O2 (B; n=3). Anti- actin immunoblotting was completed as loading controls.

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solubility of ChAT protein may be promoted by interaction of ChAT with HSC/HSP70 (Figure 3.27). Therefore, to determine whether HSC/HSP70 may play a protective role during acute oxidative stress by preventing oxidative insolubilization of ChAT, I co-treated wild-type ChAT-expressing SN56 cells with 1-8 h of 100 µM H2O2 that had been pre- treated for 24 h with 20 µM PES. Following anti-ChAT immunoblotting, I again observed that treatment of cells with 20 µM PES increased the levels of higher molecular mass Triton-insoluble ChAT protein as compared to DMSO-treated control cells (Figure 3.43). Furthermore, PES treatment again led to the accumulation of Triton-insoluble ChAT aggregates retained in the loading pockets of the stacking layer of SDS-PAGE gels. Importantly, I observed that pre-treatment of cells with 20 µM PES sensitized ChAT to

H2O2-induced insolubilization such that treatment of ChAT-expressing cells for 4 or 6 h with 100 µM H2O2 resulted in an enhanced accumulation of higher molecular mass Triton- insoluble ChAT proteins as compared to PES or H2O2 treatment alone. Lastly, greater levels of Triton-insoluble ChAT aggregates were observed in the loading pockets of SDS-PAGE gels when cells treated with H2O2 for 4 or 6 h were first pre-treated with 20 µM PES.

3.17: Co-immunoprecipitation of ChAT with HSC70 is altered following exposure of cells to H2O2

Protein misfolding stresses, such as oxidative stress, can promote the formation of aberrant disulfide-bonds and exposure of hydrophobic regions within various cellular proteins, increasing the affinity of HSPs for these misfolded proteins [246, 365, 371]. My ChAT solubility experiments with H2O2 and PES treatment suggest that ChAT protein folding may be negatively impacted following exposure to acute elevated levels of oxidative stress

(i.e. 1 mM H2O2), though under more moderate oxidative conditions (i.e. 100 µM H2O2) that interaction of ChAT with HSC/HSP70 may protect ChAT from oxidative misfolding. Therefore, I next assessed by co-IP assay whether interaction of ChAT with either HSC70 or HSP90 is enhanced in ChAT-expressing SN56 cells treated for 1 - 8 h with 100 µM

H2O2 or for 1 h with 1 mM H2O2. Following anti-HSC70 and anti-HSP90 immunoblotting from anti-ChAT co-IP samples, I observed that co-IP of wild-type ChAT with endogenous

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Figure 3.43: HSC/HSP70 inhibition by PES treatment sensitizes ChAT to H2O2- induced insolubilization. Anti-ChAT immunoblots from SN56 cells transiently expressing wild-type ChAT that were pre-treated for 24 h with 20 µM PES then co-treated for an additional 1-8 h with 100 µM H2O2. Control cells were treated with DMSO- or water- vehicle (Veh) for PES or H2O2 treatment respectively. Cells were lysed in 0.1% Triton X- 100 and Triton-soluble and -insoluble proteins were fractionated. Triton X-100 insoluble proteins were solubilized prior to SDS-PAGE by denaturing in 2x Laemmli sample buffer with 5% 2-mercaptoethanol. Co-treatment of cells with PES and H2O2 had no effect on the total levels of Triton-soluble ChAT protein. However, co-treatment with PES and 4-6 h of

H2O2 resulted in greater accumulation of higher molecular mass Triton-insoluble ChAT proteins as compared to cells treated with PES or H2O2 alone. Deposition of Triton- insoluble ChAT aggregates could be observed in the loading pockets of the SDS-PAGE gels following co-treatment of cells with PES and 4-6 h H2O2 (top panel, anti-ChAT long- exposure). Anti-actin immunoblotting was completed as loading controls (n=3).

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HSC70 is enhanced following acute exposure of cells to 100 µM H2O2 for either 1 or 2 h as compared to vehicle-treated control cells (Figure 3.44). Interestingly, exposure of ChAT- expressing cells with 1 mM H2O2 for 1 h or to 100 µM H2O2 for extended periods of time (4 – 8 h) had no effect on ChAT-HSC70 co-IP. Lastly, co-IP of ChAT with endogenous

HSP90 appears to be unaffected by treatment with either 100 µM or 1 mM H2O2.

3.18: Inhibition of the co-chaperone Cdc48/p97/VCP prevents the degradation of ubiquitinated ChAT

Previously in our laboratory we have shown that the co-chaperone Cdc48/p97/Valosin- containing protein (VCP) can stably interact with ChAT in IMR32 neural cells exposed to oxidative Aβ [440, 508]. Interestingly, VCP has also been documented to modulate HSP substrates through interaction with HSC/HSP70 [513] and HSP90 [514], and additionally can regulate the subcellular trafficking and degradation of ubiquitinated proteins [98, 441- 442]. It is currently unknown if VCP may regulate ChAT ubiquitination and degradation under basal conditions, and therefore to examine if ChAT protein stability is regulated by VCP I treated SN56 cells, transiently expressing either wild-type or mutant ChAT, for 18 h with 5-10 µM Eeyarestatin I (Eer1), a small-molecule inhibitor of VCP [515-516]. Following anti-ChAT immunoblotting, I observed accumulation of high molecular mass wild-type and mutant ChAT-immunoreactive proteins following cell treatment with 10 µM Eer1, but not in control cells treated with DMSO-vehicle (Figure 3.45a). The abundance of high molecular mass ChAT was greater for mutant than for wild-type ChAT, with levels of P17A/P19A-ChAT being the greatest. Additionally, Eer1 treatment led to accumulation of insoluble high molecular mass ChAT proteins for both wild-type and mutant ChAT that were retained in the loading pockets of the stacking layer of SDS-PAGE gels.

VCP can promote the proteasome-dependent degradation of ubiquitinated proteins from both the ER and the cytoplasm [441-442], and furthermore inhibition of VCP by Eer1 treatment has been shown to prevent the degradation of ubiquitinated VCP substrates [515- 516]. Thus, to determine if the higher molecular mass ChAT proteins observed following

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Figure 3.44: Co-IP of ChAT with endogenous HSC70 is enhanced following exposure of cells to H2O2. Immunoblots showing co-immunoprecipitation (co-IP) of ChAT with endogenous HSC70 and HSP90 from SN56 cells transiently expressing wild-type ChAT that were treated with 100 µM H2O2 for 1-8 h or with 1 mM H2O2 for 1 h. Controls cells were either treated with water-vehicle (Veh) or were transfected with empty pcDNA3.1+ vector. As assessed by anti-HSC70 and anti-HSP90 immunoblotting, co-IP of wild-type

ChAT with endogenous HSC70 is enhanced in cells treated with 100 µM H2O2 for 1-2 h, whereas co-IP of ChAT with endogenous HSP90 is unaffected by H2O2 treatment. Anti- actin immunoblotting from whole cell lysates was completed as a loading control (n=4).

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Figure 3.45: Inhibition of the co-chaperone Cdc48/p97/Valosin-containing protein (VCP) prevents degradation of ubiquitinated ChAT. (A) Anti-ChAT immunoblots from SN56 cells transiently expressing either wild-type (WT) or mutant ChAT that were treated for 18 h with 5-10 μM Eeyarestatin I (Eer1), a VCP inhibitor, or with DMSO-vehicle control. Inhibition of VCP with 10 µM Eer1 resulted in the generation of high molecular mass ChAT-immunoreactive proteins and the presence of insoluble ChAT retained in the loading pockets of SDS-PAGE gels (top panel, anti-ChAT long-exposure). Accumulation of high molecular mass ChAT appears greater for mutant ChAT compared to wild-type ChAT, with levels of P17A/P19A-ChAT being the greatest. Anti-actin immunoblotting was completed as a loading control (n=3). (B) Inhibition of VCP attenuates the degradation of ubiquitinated ChAT. ChAT ubiquitination was analyzed by anti-ubiquitin immunoblotting following anti-ChAT IP from ChAT-expressing SN56 cells that were treated for 18 h with 10 μM Eer1. Control cells were transfected with empty pcDNA31+ vector and/or treated with DMSO-vehicle. Proteasome function was maintained to allow for the degradation of ubiquitinated ChAT in DMSO-treated control conditions. The abundance of ubiquitinated ChAT following VCP inhibition appears to be greater for mutant ChAT, with levels of ubiquitinated P17A/P19A-ChAT being the greatest. Anti-actin and anti-ubiquitin immunoblotting from whole cell lysates were completed as loading controls (n=3).

VCP inhibition are ubiquitinated forms of ChAT, and therefore whether ubiquitinated ChAT may be degraded in a VCP-dependent manner, I completed anti-ChAT IPs from ChAT-expressing SN56 cells that were treated with 10 µM Eer1 for 18 h. Importantly, to maintain proteasome function and thus to allow for the degradation of ChAT in cells treated with DMSO-control I omitted the addition of MG132. By anti-ubiquitin immunoblotting of anti-ChAT IP samples, I confirmed that VCP inhibition prevented the degradation of ubiquitinated wild-type and mutant ChAT when compared to DMSO-treated control cells (Figure 3.45b). The abundance of ubiquitinated ChAT following VCP inhibition was greater for mutant ChAT when compared to wild-type ChAT, with levels of ubiquitinated P17A/P19A-ChAT being the greatest. Lastly, higher molecular mass ubiquitinated ChAT was observed in the loading pockets of SDS-PAGE gels following Eer1 treatment.

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CHAPTER 4: DISCUSSION

4.1: Significant Findings

Maintenance of ChAT protein stability and activity are essential for cholinergic neural function, and thus are critical for the regulation of both central and autonomic neural processes as well as somatic neuromuscular functions. Unfortunately, little is known regarding the cellular factors that either stabilize ChAT protein or, conversely, that promote the degradation of ChAT. Alterations in ChAT protein stability and enzymatic activity have been documented and correlated with the pathogenesis of the neuromuscular disease CMS. Previous studies from other laboratories have identified and characterized multiple CMS- related ChAT mutations that result in reduced in vitro ChAT activity as well as reduced steady-state protein levels when expressed in mammalian cells [411, 457-458]. One of these CMS-related mutations, V18M, is positioned distal to the ChAT active site and is located within a highly-conserved and surface-exposed proline-rich motif at residues 14PKLPVPP20 that shares homology with PxxP SH3-binding motifs [420, 457-458, 474]. The cellular mechanisms underlying the dysfunction of mutant V18M-ChAT protein are still unknown, though mutation of residue Val18 could potentially result in disruption of the ChAT N-terminal proline-rich motif; it is currently unknown whether this proline-rich motif regulates ChAT function. Thus, the goal of my thesis was to first investigate a functional role of the ChAT N-terminus on human 69-kDa ChAT protein, with a focus on the highly-conserved proline-rich motif that surrounds the CMS-related residue Val18. Second, and in conjunction with my first aim, I sought to determine cellular mechanisms that regulate the basal stabilization and/or degradation of wild-type ChAT protein and/or that regulate mutant ChAT protein stability.

In summary, this thesis details the characterization and establishment of a role for the conserved N-terminal proline-rich motif in regulating the protein stability and activity of human 69-kDa ChAT. To begin, I found that disruption of the ChAT proline-rich motif by either N-terminal truncation or proline-to-alanine mutation, or in relation to the CMS- related V18M mutation, reduces both the steady-state protein levels and cellular activity of mutant ChAT when transiently expressed in mouse cholinergic SN56 cells. Interestingly,

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proline-to-alanine mutation of residues Pro17 and Pro19 together (P17A/P19A) rendered the mutant enzyme inactive in cellular lysates and yielded the greatest loss of ChAT protein in cells similar to that observed for N-terminal truncations that involved partial or complete removal of the ChAT proline-rich motif. The in vitro catalytic activity of purified bacterially-expressed recombinant P17A/P19A-ChAT is impaired, though this does not appear to be due to substantial changes in either gross protein secondary structure or in vitro thermal stability of mutant ChAT. Importantly, I demonstrated for the first time that ChAT is ubiquitinated in cells and that the ubiquitination of mutant P17A/P19A- and V18M-ChAT is enhanced, resulting in increased proteasome-dependent degradation and loss of mutant ChAT protein in SN56 cells. The cellular loss of P17A/P19A-ChAT protein correlated with a reduction in ChAT protein half-life as measured using a novel fluorescent- biorthogonal pulse-chase protocol. Proteasome inhibition prevented the degradation of ubiquitinated wild-type and mutant ChAT, increasing the steady-state protein levels and protein half-life of ChAT. Lastly, I observed that ChAT can undergo K48-dependent and K48-independent polyubiquitination and proteasomal degradation. These data have been peer reviewed and published in the Journal of Neurochemistry [502].

Of significance, these initial studies demonstrated that ChAT protein stability is regulated by the ubiquitin-proteasome system, and that ubiquitination of catalytically- deficient mutant ChAT is enhanced, thus identifying a mechanism that may be responsible for the cellular loss of CMS-related ChAT mutant protein. The molecular mechanisms that regulate basal ChAT ubiquitination and/or that promote enhanced ubiquitination of mutant ChAT remained unknown following the aforementioned experiments, and thus I sought to identify novel cellular mechanisms that regulate ChAT protein stability. By BioID, co- immunoprecipitation, and in situ PLA, I identified the heat shock proteins HSC/HSP70 and HSP90 as novel ChAT protein-interactors in mammalian cells. Both of these molecular chaperones are well-known for their role in promoting the folding and stabilization of cellular proteins. Thus, by pharmacological means I found that inhibition of HSC/HSP70- substrate interactions by treatment of SN56 cells with the HSC/HSP70 inhibitor PES resulted in the accumulation of Triton-insoluble and SDS-resistant wild-type and mutant ChAT protein aggregates. Furthermore, inhibition of HSC/HSP70 and HSP90 activity by

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treatment of cells with VER-155008 or 17-AAG, respectively, led to reduced cellular ChAT activity, enhanced ChAT ubiquitination, and increased proteasome-dependent loss of ChAT protein. Importantly, the effects of HSP inhibition were greater for mutant ChAT (P17A/P19A-, V18M-, and A513T-ChAT) as compared to wild-type ChAT. HSPs can alternatively promote the ubiquitination and degradation of misfolded proteins through cooperative interaction with the E3 ubiquitin ligase CHIP/Stub1, and while I show that ChAT interacts with CHIP by both co-IP assays and in situ PLA, siRNA-mediated knock- down of CHIP had no effect on either wild-type or mutant ChAT protein levels. However, inhibition of the ER- and HSP-associated co-chaperone Cdc48/p97/VCP prevented the degradation of ubiquitinated ChAT. Together, these results identify novel mechanisms for the functional regulation of both wild-type and CMS-related mutant ChAT by pro- stabilizing HSPs and the pro-degradative co-chaperone p97/VCP which may have broader implications for ChAT function during cellular stress and disease. These data have been peer reviewed and published in Frontiers in Molecular Neuroscience [517]

An important function of HSPs is to chaperone cellular proteins when cells are exposed to acute proteotoxic stresses, such as hyperthermia and oxidative stress, thereby maintaining protein stability, solubility, and function. Earlier studies in our laboratory by Shawn Albers, M.Sc., showed that cellular ChAT activity is decreased in cells exposed to oxidative stress by H2O2 treatment, and furthermore that ChAT forms redox-dependent higher molecular mass oligomers following H2O2 treatment [507]. In my work, I found that the solubility of ChAT protein is decreased, and that ChAT ubiquitination is enhanced in cells treated with H2O2. Furthermore, inhibition of HSC/HSP70-substrate interactions by

PES treatment further sensitized ChAT to H2O2-induced insolubilization and formation of higher molecular mass ChAT protein aggregates. Lastly, as assessed by co-IP assays, the association of ChAT with HSC70, but not HSP90, is enhanced following H2O2-treatment, suggesting that HSC70 may help to prevent oxidative dysfunction of ChAT by chaperoning ChAT protein and preventing ChAT misfolding during exposure to oxidative stress. These initial experiments investigating the effects of oxidative stress on ChAT protein stability, solubility, and function will help to inform and provide rationale for future work within the Rylett Laboratory as will be discussed later in this thesis.

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Taken altogether, my results detail within this thesis support a model (Figure 4.1) where the heat shock proteins HSC/HSP70 and HSP90 promote the folding, stabilization, and catalytic activity of human 69-kDa ChAT in a cooperative manner, thereby ultimately regulating ACh synthesis and cholinergic neural function. Furthermore, the basal turnover of ChAT protein is regulated by the ubiquitin-proteasome system where the co-chaperone p97/VCP promotes the degradation of ubiquitinated ChAT. I propose that ChAT mutations that impair ChAT enzymatic activity, such as P17A/P19A and CMS-related V18M, do so by interfering with the correct/complete conformational folding of ChAT in cells, thus leading to ChAT destabilization and the generation of misfolded mutant ChAT protein. Additionally, under cellular conditions that can promote protein misfolding, such as during exposure to oxidative stress or when HSP functions are inhibited, ChAT protein folding becomes compromised, resulting in the formation of misfolded and insoluble ChAT protein aggregates. Subsequently, misfolded and dysfunctional ChAT protein is targeted for cellular clearance via enhanced ubiquitination and proteasome-dependent degradation, potentially through chaperone-mediated mechanisms. The identity of the E3 ubiquitin ligase responsible for ChAT ubiquitination under either basal conditions or that promotes enhanced ubiquitination of mutant and/or misfolded ChAT remains to be elucidated.

4.2: General Discussion

Canonically, HSPs such as HSC/HSP70 and HSP90 have been characterized for their role in the folding and stabilization of nascent proteins, as well as refolding of aggregation- and misfolding-prone proteins during proteotoxic stress [118, 135, 154-155]. In the present studies I show that inhibition of HSC/HSP70 led to the insolubilization, destabilization, and degradation of both wild-type and mutant ChAT, and while I found that inhibition of HSP90 reduced cellular ChAT activity, this had no effect on the steady-state protein levels of wild-type ChAT. In the currently accepted model (Figure 1.6) multiple HSPs can act together to chaperone cellular proteins in a cooperative and processive manner, with initial recognition and folding of non-native nascent proteins completed by a HSP40-HSC/HSP70 complex, followed by transfer of semi-folded HSP substrates to HSP90 by HSP70-HSP90

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Figure 4.1: Model for the dynamic regulation of ChAT protein stability and function by molecular chaperones and the ubiquitin-proteasome system. (1) The heat shock proteins HSC/HSP70 and HSP90 act in a cooperative and processive manner to promote the folding, stabilization, and catalytic activity of human 69-kDa ChAT protein. (2) Under basal conditions the co-chaperone Cdc48/p97/Valosin-conaining protein (VCP) regulates the turnover of ChAT protein where inhibition of VCP by Eeyarestatin I (Eer1) treatment prevents the proteasomal degradation of ubiquitinated ChAT. (3) ChAT protein folding can become compromised following exposure of cells to either oxidative stress (i.e. H2O2) or to HSP inhibition (i.e. PES/VER-155008 and 17-AAG), as well as in cases where mutation of ChAT impairs ChAT enzymatic activity, such as with P17A/P19A and CMS-related V18M, leading to the generation of misfolded ChAT protein. (4) Misfolded ChAT is then targeted for cellular clearance through enhanced ubiquitination and proteasome-dependent degradation, potentially through chaperone-mediated mechanisms. The E3 ubiquitin ligase responsible for ChAT ubiquitination under either basal conditions or that promotes the enhanced ubiquitination of mutant and/or misfolded ChAT remains to be elucidated.

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organizing protein (Hop) and subsequent final folding by HSP90 [178-180]. Thus, HSP90 works downstream of HSC/HSP70 to interact with semi- or pre-folded proteins where formation of stable HSP90-substrate interactions is promoted by availability of labile solvent-accessible hydrophobic domains, such as those found in ligand-binding domains and enzyme active sites [234-236, 238, 246, 503]. As such, HSP90 plays an essential role in the structural maturation and conformational activation of various cellular kinases and enzymes involved in a wide array of cell signaling pathways, such as endothelial nitric oxide synthase (eNOS) [518], the cell cycle-associated tyrosine kinase Wee1 [519], steroid hormone receptors [244, 520], and the receptor tyrosine kinase ErbB2 [245, 503]. This contrasts with HSC/HSP70 that interacts with a diverse and extensive range of nascent non- native proteins (20% of all nascent proteins) that minimally contain solvent-exposed regions enriched with 4-7 hydrophobic residues [167-168, 187, 202]. Thus, I propose that ChAT protein folding, stability, and cellular activity is regulated in a similar processive manner by HSC/HSP70 and HSP90 respectively. It is important to note that inhibition of the ATPase activity of either HSC/HSP70 or HSP90 resulted in the loss of cellular ChAT activity, suggesting that cooperation of these HSPs is essential for the formation of natively-folded and active ChAT enzyme, and thus that these HSPs may be critical to cholinergic neural function. Interestingly though, inhibition of HSC/HSP70 and/or HSP90 did not fully abolish ChAT enzymatic activity, nor did it result in complete loss of ChAT protein in cells suggesting that these HSPs may work in collaboration with other cellular molecular chaperones to promote ChAT protein folding, stability, and function.

Multiple co-chaperones regulate the function of HSPs, including HSP40, Hop, and CHIP. Previous studies from our laboratory have shown that the co-chaperone Cdc48/p97/VCP can interact with ChAT in IMR-32 neural cells exposed to Aβ [440]. VCP is a member of the AAA-ATPase family that has diverse cellular functions, including the ability to interact with and modulate HSC/HSP70 and HSP90 substrates [513-514], as well as to interact with ubiquitinated proteins to regulate their subcellular trafficking and degradation [97, 441-442]. Importantly, in the present studies I not only demonstrate for the first time that ChAT is ubiquitinated in cells and that the degradation of ubiquitinated ChAT is proteasome dependent, but also show that pharmacological inhibition of VCP led

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to the accumulation of ubiquitinated ChAT and prevented its degradation in cells with functional proteasomes. Related to its proteolytic functions, studies have demonstrated a role for VCP in the handling and degradation of protein aggregates [521], as well as in the degradation of ubiquitinated proteins by the autophagy-lysosomal system [443-444]. While the autophagy-lysosomal system is one possible mechanism involved in the VCP-mediated degradation of ubiquitinated ChAT, my studies show that ChAT protein degradation is largely unaffected by pharmacological inhibition of the lysosome and, conversely, is regulated chiefly by the proteasome as assessed using the proteasome inhibitors MG132 and lactacystin. Interestingly, VCP is best known for its role in promoting the proteasome- dependent degradation of ERAD substrates [441-442], where VCP is critical for the ER- to-cytoplasm transfer of proteins ubiquitinated in the ER, thereby directly mediating degradation of ERAD substrates by the cytosolic 26S-proteasome [522]. Furthermore, VCP can directly influence the ubiquitination of ERAD substrates by interacting with the ER- associated E3 ubiquitin ligases gp78/AMFR and Hrd1/SYVN1 [522-524] that act downstream of 78-kDa glucose-regulated protein (GRP-78), an ER-associated HSP known to promote protein folding and stability of nascent proteins in a fashion similar to that of HSC/HSP70 [525]. Therefore, an alternative possibility by which VCP regulates the degradation of ubiquitinated ChAT may involve ERAD-associated mechanisms in addition to those related to cytosolic HSPs, and thus I hypothesize that ChAT may be regulated by various cytosolic and/or ER-associated chaperones that act cooperatively to promote ChAT protein function. Importantly, while my data suggest that the proteasomal degradation of ubiquitinated ChAT is promoted by VCP, it is unclear whether this pro-degradative function of VCP is primarily HSP- or ER-associated. Lastly, though the E3 ubiquitin ligase/s responsible for the ubiquitination of ChAT protein have yet to be identified, my results establish the possibility that ChAT ubiquitination may be regulated by the E3 ubiquitin ligases gp78 and/or Hrd1; this requires further investigation.

Many neurological disorders, including AD, Parkinson’s, and Huntington’s disease exhibit dysfunction of HSPs, accumulation of misfolded and aggregate-prone proteins, and build-up of oxidative stress [10, 526]. Protein oxidation and subsequent formation of aberrant stress-induced disulfide bonds during cellular exposure to oxidative stress can

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result in gross misfolding of cellular proteins with severe functional consequences [361, 365]. As an example, the cysteine-rich (7.5%) E3 ubiquitin ligase Parkin has been shown to misfold, inactivate, and aggregate following exposure to H2O2 [368-370]. Of significance to my studies, human 69-kDa ChAT also has a higher-than-average cysteine content of 3.2% compared to 1.6% for intracellular mammalian proteins [367] which may sensitize ChAT to oxidative damage. In support, previous studies from both our laboratory and by others [439, 507] have demonstrated that cellular ChAT activity is reduced following exposure of neural cells to oligomeric Aβ1-42, a molecular hallmark of AD and stimulator of cellular oxidative stress [429, 508]. Additionally, our laboratory has shown that Aβ1-42 treatment results in stable interaction of ChAT with VCP [440] and, together with my results establishing a novel role for VCP in the degradation of ubiquitinated ChAT, suggest that ChAT ubiquitination and subsequent protein stability may be altered during cellular stress. Importantly, work in this thesis shows that acute exposure of ChAT- expressing cells to H2O2-induced oxidative stress results in enhanced insolubilization and ubiquitination of ChAT protein. One well-known function for HSPs, such as HSC/HSP70 and HSP90, is to maintain proteostasis during acute exposure to cellular stress [118, 135, 154-155], and as such during oxidative conditions these HSPs have been shown to chaperone oxidized proteins in order to maintain and/or restore protein function [371]. My results using pharmacological inhibitors of HSC/HSP70 and HSP90 suggest that these HSPs regulate basal ChAT protein stability in non-stressed cells, and therefore it is likely that they may also chaperone ChAT during oxidative stress. In support, by co-IP assay I found that interaction of ChAT with HSC/HSP70 appears enhanced in H2O2-treated cells, and furthermore that HSC/HSP70 inhibition promoted the H2O2-induced insolubilization of ChAT protein. It is currently unknown whether exposure to oxidative Aβ results in alterations in ChAT ubiquitination or solubility, or whether ChAT protein stability is altered following exposure to other cellular stresses, such as hyperthermia, or during human protein misfolding diseases such as AD.

In addition to their role as cellular protein chaperones that promote protein folding and stabilization, HSC/HSP70 and HSP90 are also well-known for their ability to triage mutated, misfolded, and aggregated proteins for degradation through coordination with the

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cytosolic U-box-containing and HSP-associated E3 ubiquitin ligase CHIP/Stub1 [76-78, 184]. In fact, HSP-mediated protein degradation often requires the formation of pro- degradative trimeric complexes between substrate-bound HSC/HSP70 or HSP90 and the N-terminal TPR domains of CHIP [76, 78, 293]. As such, CHIP has been shown to regulate a diverse range of cellular functions through CHIP-mediated ubiquitination of HSC/HSP70 and HSP90 substrate proteins, including but not limited to the glucocorticoid receptor [184, 294], hypoxia-inducible factor 1-alpha (HIF-1α) [295], and the receptor tyrosine kinase ErbB2 [230, 285]. In my present studies, I demonstrate that inhibition of HSC/HSP70 and HSP90 enhances the ubiquitination of ChAT protein, and subsequently show that ChAT interacts in situ with endogenous CHIP in SN56 cells. Furthermore, though overexpression of CHIP promoted the proteasomal degradation of ChAT, siRNA-mediated knock-down of CHIP failed to stabilize and prevent the degradation of ChAT protein. While unexpected, one plausible reason for this observation is that other HSP-associated E3 ubiquitin ligases may regulate ChAT ubiquitination as previous studies have demonstrated functional redundancy for CHIP-mediated degradation of HSP clients in CHIP-/- mouse fibroblasts [527]. As such, the E3 ubiquitin ligase Parkin has been shown to associate with both HSPs and CHIP to co-regulate the chaperone-mediated degradation of HSP clients such as neuronal nitric oxide synthase (nNOS) and various polyglutamine-expanded proteins [527- 528]. In addition, other cytosolic E3 ubiquitin ligases can also regulate the ubiquitination of nascent and misfolded HSP clients, including E6-associated protein (E6-AP) and the N- end rule E3 ligase Ubr1 [529-530]. Interestingly, in my studies I found that the HSP- binding TPR domains of CHIP, but not the catalytic U-box domain, are required for both the interaction of CHIP with ChAT as well as for the loss of ChAT protein following CHIP overexpression. Importantly, CHIP has been previously shown to compete with other pro- folding TPR-containing co-chaperones, such as HOP, for binding to the C-terminal EEVD- motif of HSC/HSP70 and HSP90 thus promoting the destabilization of HSP substrates [276, 318]. Thus, I hypothesize that alternative HSP-related E3 ubiquitin ligases, such as Parkin or E6-AP, may regulate the ubiquitination of ChAT protein, potentially in cooperation with CHIP, whereas CHIP overexpression likely promotes ChAT protein degradation by interfering with the pro-folding functions of HSPs that stabilize ChAT.

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To my knowledge, work in this thesis demonstrates for the first time the ubiquitination of human 69-kDa ChAT protein in mammalian cells. Subsequent characterization of ChAT ubiquitination revealed that ubiquitinated ChAT is degraded in a proteasome-dependent manner, and furthermore that proteasome inhibition by MG132 treatment prevents degradation of both K48-linked and non-K48-linked polyubiquitinated ChAT. Canonically, cellular proteins targeted for proteasomal degradation are done so through covalent modification with poly-Ub chains containing at least four K48-linked ubiquitin molecules [44, 54, 93]. However, studies investigating either the APC, a RING- domain E3 ubiquitin ligase involved in cell cycle regulation, or the ERAD-associated co- chaperone p97/VCP have identified K11-linked poly-Ub chains as an alternative signal for proteasomal degradation [94-95, 97-98]. These studies, together with my results, suggest that ChAT degradation may be regulated by K48- and/or K11-linked polyubiquitination. Interestingly, in cells with functional proteasomes where ChAT ubiquitination was limited to modification with proteolytic K11-linked poly-Ub chains I observed impaired degradation of polyubiquitinated ChAT. Therefore, I hypothesize that K48-mediated proteasomal degradation represents the dominant mechanism for degradation of ChAT protein in cells, with K11-mediated degradation potentially representing either a low abundance or compensatory mechanism for ChAT protein degradation when K48-mediated degradation is prevented or impaired. Furthermore, given that VCP can interact with K11- linked polyubiquitinated proteins [98], and that interaction of ChAT with VCP in enhanced in neural cells exposed to Aβ1-42 [440], ubiquitination of ChAT with alternative K11-linked poly-Ub chains may represent a unique modification of ChAT during cellular stress, though this requires future confirmation.

The initial goal of my thesis was to determine a functional role for the highly- conserved N-terminal proline-rich motif of human 69-kDa ChAT at residues 14PKLPVPP20. Subsequently, my studies have demonstrated that this motif is essential for the catalytic activity of ChAT both in vitro and in cells, as well as for the cellular stability of ChAT protein. Briefly, I show that the ubiquitination and proteasomal degradation of mutant P17A/P19A-ChAT is enhanced compared to wild-type human 69-kDa ChAT, correlating with a loss in cellular steady-state protein levels and a reduction in the protein half-life of

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P17A/P19A-ChAT. In studies performed by others, disease-related ChAT mutations have been identified in CMS patients that reduce both in vitro enzymatic activity and cellular steady-state protein levels of ChAT when expressed in mammalian cells [411, 457-458], though the mechanisms underlying the loss of CMS-mutant ChAT protein in cells remains unknown. One of these catalytically-deficient CMS-related mutations, V18M, is located within the N-terminal proline-rich motif of ChAT and was found previously to reduce both the in vitro catalytic efficiency and the steady-state protein levels of the mutant enzyme [457-458]. Importantly, my results show that the ubiquitination and proteasomal degradation of mutant V18M-ChAT is increased when compared to wild-type ChAT, paralleling my observations made for catalytically-inactive P17A/P19A-ChAT. Moreover, I observed increased ubiquitination of an additional non-active site and catalytically- deficient CMS-related mutant, A513T-ChAT, that has been previously shown to also display reduced steady-state ChAT protein levels in cells [457]. Given these data, ChAT mutations positioned either within or outside of the N-terminal proline-rich motif that reduce ChAT catalytic activity may also promote enhanced ubiquitination and proteasomal degradation of mutant ChAT, thereby synergistically enhancing the catalytic defects of these mutants in vivo. Thus, I propose that enhanced ubiquitination and subsequent proteolytic degradation may be a common feature among various CMS-related mutant ChAT proteins, suggesting that prevention of proteasomal degradation may enhance the cellular activity of some CMS-related ChAT mutants. In support of this, I have shown that proteasome inhibition increases the steady-state protein levels and cellular enzymatic activity of both wild-type and the CMS-related ChAT mutants A513T- and T490N-ChAT. Conversely, while proteasome inhibition did increase the steady-state protein levels of mutant P17A/P19A- and CMS-related V18M-ChAT, these mutants remained catalytically impaired in cells. Thus, these results suggest that inhibition of ChAT ubiquitination and/or proteasomal degradation may represent a novel avenue for the in vivo stabilization and synergistic enhancement of the enzymatic activity of certain CMS-related ChAT mutants.

While my results show that mutation of the N-terminal proline-rich motif enhances ChAT ubiquitination and degradation in cells, the cellular mechanisms responsible for these effects remain to be elucidated. Interestingly, the N-terminal proline-rich motif of

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ChAT shares structural and sequence homology with the core “PxxP” SH3 binding motif [420, 474-477]. Interactions between SH3 motif- and SH3 domain-containing proteins have been shown to regulate the subcellular trafficking, activity, and stability of multiple cellular proteins, most often by modulating changes in protein phosphorylation through SH3 domain-containing adaptor proteins, such as Grb2 and Nck, and various cellular protein tyrosine kinases [476-479]. Prior to work completed in this thesis, only a handful of protein- protein interaction involving human ChAT had been identified, of which include the protein kinases PKC and CaMKII and the co-chaperone p97/VCP [438]. Importantly, work in our laboratory has shown that hierarchical phosphorylation of ChAT by either PKC or CaMKII alters the cellular activity of ChAT, indicating that ChAT enzymatic function is influenced by dynamic phosphorylation [410, 426-427, 440]. Furthermore, studies by others have demonstrated that phosphorylation can influence the ubiquitination and degradation of multiples members involved within the EGFR/MAPK signalling network through various phospho-binding E3 ubiquitin ligases, such as members of the Cbl family [531-533]. It is currently unknown whether ChAT engages in SH3-dependent protein-protein interactions mediated through its N-terminal proline-rich motif. Furthermore, it is unknown if ChAT mutations, specifically those found within the N-terminal proline rich motif (i.e. V18M), alter the phosphorylation of ChAT, and whether this may correlate with loss of cellular activity and protein stability of mutant ChAT.

As discussed above, in this thesis I found that the ubiquitination and degradation of specific CMS-related ChAT mutants (i.e. V18M- and A513T-ChAT), as well as mutant P17A/P19A-ChAT, is enhanced in cells. All three of these mutations are positioned distal to the ChAT active site [420, 457], and while the mechanisms responsible for the loss of mutant ChAT protein in mammalian cells are not yet fully clear, one possibility is that these mutations may produce local labile changes to the ChAT protein secondary structure that are then responsible for both the catalytic impairment of ChAT and that target these ChAT mutants for chaperone-mediated degradation. Importantly, using purified bacterially- expressed recombinant ChAT proteins I found that mutation of the ChAT N-terminal proline-rich motif does not produce substantial changes to the overall gross protein secondary structure or the in vitro thermal stability of purified recombinant ChAT as

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measured by circular dichroism analysis. As such, it is not obvious how these mutations destabilize ChAT protein in vivo, and thus by BioID I sought to identify ChAT protein- interactors responsible for the stabilization and/or enhanced degradation of mutant ChAT in mammalian cells. Though I did not detect any SH3-mediated ChAT protein-protein interactions by BioID, I did identify the heat shock proteins HSC/HSP70 and HSP90 as novel ChAT protein-chaperones and observed by co-IP assay enhanced interaction of mutant ChAT with these HSPs as compared to that seen for wild-type ChAT. Furthermore, I show that various ChAT mutants (i.e. P17A/P19A, V18M, and A513T) have increased sensitivity to pharmacological inhibition of both HSC/HSP70 and HSP90 where, importantly, HSP90 inhibition resulted in the proteasomal loss of mutant but not of wild- type ChAT. In general, HSPs aid in the stabilization of their substrates through a process of kinetic partitioning, whereby they interact with exposed hydrophobic domains within cellular proteins that is thought to prevent client protein misfolding and aggregation [6, 167-168, 503]. In support of this, I found that inhibition of HSC/HSP70-substrate interactions by PES treatment resulted in loss of ChAT protein solubility and the generation of Triton-insoluble and SDS-resistant ChAT aggregates, with the abundance of these PES- induced insoluble ChAT proteins being greatest for mutant ChAT. Interestingly, Triton- insoluble aggregates of mutant P17A/P19A-ChAT protein was observed in cells with functional HSPs, suggesting that the folding of this mutant ChAT protein may be compromised under basal conditions, resulting in the production of catalytically-impaired ChAT that may be susceptible to misfolding and inactivation. While my results suggest that HSC/HSP70 and HSP90 are critical for the stabilization of mutant ChAT, these HSPs are also known to promote the degradation of misfolded proteins through the ubiquitin- proteasome system [6, 76-78, 118]. Importantly, I observed enhanced ubiquitination of both wild-type and mutant ChAT following HSP inhibition, suggesting that impairment of HSP- mediated folding of ChAT protein may promote the ubiquitination and degradation of misfolded ChAT. Therefore, taken together my data suggest that mutations, such as CMS- related V18M and A513T, may compromise ChAT protein folding and the conformational activation of ChAT in vivo, leading to enhanced chaperone-mediated ChAT degradation and/or enhanced dependence of mutant ChAT proteins on HSPs to maintain mutant ChAT stability and function. It is important to note that my HSP observations related to mutant

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ChAT are significant as protein misfolding and enhanced functional dependence on molecular chaperones is frequently observed for mutated and disease-relevant proteins [125, 534]. Thus, my observations may represent a general HSP-related mechanism for the regulation of ChAT protein stability that may apply to not only various CMS-related ChAT mutants beyond those studied presently, but also to the maintenance of ChAT function during cellular stress and disease, such as that observed in this thesis for ChAT-expressing cells exposed to oxidative stress by H2O2 treatment.

An important outcome of my studies was the development of a novel approach to the study of cellular protein half-life. The biochemical determination of protein half-life can provide insight into the stability, regulation, and function of cellular proteins. Two differing methods are commonly used to determine protein half-life; ribosomal inhibition by CHX treatment, a potent yet cytotoxic inhibitor of protein synthesis [501], or pulse-chase experiments that utilize radioactive 35S-methionine live-labeling [535-537]. While 35S- methionine labeling is held as the gold standard for pulse-chase analysis, metabolic radiolabeling has been found to induce cellular damage and reduce cell survival, suggesting that care is needed when interpreting results gathered by this approach [538-539]. In response to these concerns, I have developed an alternative and novel protocol for the biorthogonal determination of protein half-life by fluorescent labeling of nascent proteins using “click chemistry” [502]. My protocol is a direct adaptation on 35S-methionine pulse- chase labeling whereby newly synthesized nascent proteins are first live-labeled in culture with the biorthogonal amino acid L-azidohomoalanine (AHA), a non-radioactive and azide- containing methionine analog, that are then reacted with fluorescently-tagged strained cyclooctynes to form stable fluorescent triazole conjugates using metal-free SPAAC reactions [495-496]. Importantly, using this novel protocol I not only determined the protein half-life of human 69-kDa ChAT, but have also shown that ChAT protein half-life is reduced following mutation of ChAT N-terminal proline-rich motif (i.e. P17A/P19A) or that it is increased following cellular proteasome inhibition by MG132 treatment. While SPAAC reactions have been used previously both in vitro and in vivo to measure global changes in protein synthesis [497-500], to my knowledge my studies are the first to use SPAAC reactions to measure the protein half-life of a specific cellular protein of interest.

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My protocol has several advantages over the more standardized CHX or 35S-methionine pulse-chase assays, foremost being that AHA live-labeling of HEK293 cells has been previously shown to be highly specific for newly-synthesized proteins and, importantly, is non-toxic to cells [498]. Second, AHA-labeled proteins can be used for both fluorescent and immunoblotting analysis from the same sample preparation, thereby eliminating the need to prepare both labelled and non-labelled protein samples when performing similar 35S-methionine pulse-chase experiments. Lastly, an additional advantage of my protocol is the ever-growing selection of commercially available biorthogonal amino acids and complementary fluorescent or chemiluminescent probes that can be used in copper-free SPAAC reactions. This advantage allows for increased flexibility when designing pulse- chase experiments and increases the compatibility of my protocol with commonly-used laboratory equipment and procedures without the reliance on radioactive 35S-methionine or cytotoxic CHX. Overall, I propose that my SPAAC pulse-chase protocol represents a novel, non-toxic, and non-radioactive alternative/replacement to classical CHX or 35S-methionine pulse-chase methods, and furthermore is an important example of the utility for click chemistry in the safe and effective alteration of existing and well-practiced methods.

4.3: Study Limitations and Alternative Approaches

The major limitation for any study is the choice of biological model/s and the associated techniques used to assess study objectives. For my studies, to determine the effects of ChAT mutations on the regulation of ChAT protein stability and function I used transiently transfected cells lines due to their availability, ease of use, and speed by which I was able assess my initial experimental aims centered around mutated forms of human ChAT protein. Given that ChAT is a defining cholinergic protein, the use of cholinergic neural cells in my studies was imperative to minimalize the acquisition of misleading results associated with studying neural proteins in non-neural cells. In addition, as transient transfection was to be my primary means to express wild-type and mutant ChAT protein in cells, transfection efficiency and the ability to express heterologous proteins were major considerations for an appropriate cell line for my study. Many well-documented neural cell

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lines exist, including human neuroblastoma SH-SY5Y cells [540-541], a cell line that has been extensively used for the study of cholinergic molecular mechanisms related to neurogenerative diseases [542-544]. Though useful for the study of either endogenous neural proteins or for the generation of stable cell lines, SH-SY5Y cells are notoriously difficult to transfect, transiently express heterologous proteins at low levels relative to other mammalian cell lines, and often experience transfection-related cytotoxicity. Therefore, to circumvent these limitations, for the majority of my studies I used mouse cholinergic SN56 cells [485, 545], a neural cell line that has been used by other groups to study both cholinergic and ChAT function in cell culture [546-550]. Ultimately, my choice for using mouse SN56 cells was four-fold, being that these cells are easy to transfect by multiple methods (e.g. calcium phosphate and Lipofectamine), express relatively higher levels of heterologous protein, are resistant to transfection-induced cytotoxicity, and in culture require minimal serum (2 – 5%) and grow quickly with a doubling-time of ~24 hours. Importantly, SN56 cells also have an undifferentiated cholinergic phenotype [485, 547], whereas SH-SY5Y cells require differentiation to attain this phenotype [544, 551]. These attributes together make SN56 cells an excellent and effective option for the molecular study of cholinergic processes in cell culture that require transection-based methods.

Though SN56 cells proved useful throughout my thesis, there were limitations to my study of human ChAT protein function in these cells. First, my studies were completed in undifferentiated SN56 cells, and while these cells do have a basal cholinergic phenotype, differentiation using retinoic acid and dibutyryl-cAMP has been shown to enhance the cholinergic properties of SN56 cells, correlating with enhanced endogenous ChAT activity and ACh content, reduced cell division, and neurite outgrowth [485, 547]. It is currently unknown what impact differentiation would have had on my results, though I hypothesize that any effects may have been negligible as the mechanisms found to regulate ChAT protein stability in this thesis, i.e. the ubiquitin-proteasome system and HSPs, are known to be well-conserved between many different tissue types. A second limitation to my use of SN56 cells is that I studied a human protein in mouse cells. While this is common when studying disease-related human proteins in mouse models [552-553], this limitation is often avoidable in cell culture studies due to the variety of available human cell lines. As

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discussed above, mouse SN56 cells are an excellent option for cholinergic studies requiring transfection, and while the mouse proteome does differ from that found in human cells, a high degree of compositional and function proteomic similarity has been observed between cells from these two species [554-555]. Interestingly, in my studies I observed results that differed in some respects when analyzing anti-ChAT co-IPs completed from either human kidney HEK293 or mouse SN56 cells. Specifically, unique to HEK293 cells I observed co- IP of ChAT with stress-inducible HSP70 and saw enhanced interaction of P17A/P19A- ChAT with HSP90; co-IP of ChAT with HSC70 and HSP90 was observed in both HEK293 and SN56 cells. Of note, HEK293 cells have elevated constitutive expression of HSP70 due to the E1A-mediated immortalization of these cells [556-557], whereas in SN56 cells the stress-induced expression of HSP70 was limited to cells exposed to either hyperthermia (data not shown) or to the HSP90 inhibitor 17-AAG (Figure 3.30). Therefore, observational differences made between these two cell lines are likely multifactorial (i.e. species/tissue type and HSP expression) and may be unique to HEK293 cells. It is unknown if my results would have also differed if assessed in human cholinergic cells, such as SH-SY5Y cells.

As is common for cell-based studies, transient transfections were used throughout my experiments as I required straightforward, routine, effective, and consistent heterologous expression of multiple proteins of interest including engineered and CMS-related ChAT mutants. A limitation of this approach is the typical overexpression of heterologous protein observed when using expression vectors, such as pcDNA3.1, that contain strong constitutive promoters [558]. As it is often unknown whether overexpressed heterologous proteins behave like or are regulated in a similar fashion to their endogenous counterparts, studies that can wholly or partially investigate endogenous proteins have an advantage by limiting the potential for unintended effects due to protein overexpression. Importantly, though my studies relied on heterologous expression of wild-type or mutant ChAT, my experiments focused on ChAT function as it was related to various endogenous proteins, including ubiquitin, HSC/HSP70, HSP90, CHIP, and p97/VCP. It is currently unknown whether endogenous ChAT protein is regulated likewise to that observed in my studies. One alternative to transient heterologous protein expression is the generation of cells stably expressing a heterologous protein of interest [559], though this carries risks associated with

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random genome integration of plasmid cDNA and unintended cellular consequences due to long-term constitutive protein overexpression. To address these issues, modern techniques have been developed that allow for guided genomic integration of inducible cDNA (e.g. Flp-In T-Rex System; Thermo) or for genomic editing of endogenous proteins using CRISPR-Cas9 technology [560-561], though transient transfection remains the most standard and simplest approach for the expression of heterologous proteins in cell lines.

The most significant limitation of my study was the extensive use of immortalized cell lines to study ChAT protein stability, and while the use of cell lines remains a standard approach for the molecular study of protein function, care must be taken when interpreting results generated from such studies. Specifically, most immortalized cell lines are derived from genetically unstable tumorigenic cells, including SN56 cells [485, 545, 562-563], and thus a major concern is that these cell likely harbour tumour-promoting proteomic changes that could have an unforeseen or deleterious impact on experimental results. While this is a well-known limitation, the availability and accessibility of immortalized cell lines has made them an effective molecular tool, particularly for exploratory studies such as mine. Alternatively, the use of non-tumorigenic primary cells or induced pluripotent stem cells (IPSCs) has gained popularity due to the development of effective transfection techniques for primary cells [564], as well as the creation of protocols for the growth, differentiation, and use of IPSCs. While proposed to be a more physiologically-relevant tool for the study of biological processes, the use of primary cells or IPSCs remains today both costly and challenging, particularly due to difficulties in the isolation, culturing, and transfection of these cells. Though these constraints may reduce the experimental flexibility of these cells as compared to immortalized cell lines, I propose that it will be important to assess whether select results from my thesis are reproducible in either primary neurons or IPSCs.

In addition to these study-wide limitations, experiment-specific limitations also need to be addressed. First, to characterizing ChAT ubiquitination in SN56 cells I overexpressed various ubiquitin mutants over-top of an endogenous ubiquitin background (Figure 3.11). Unexpectedly, I observed enhanced recovery of ubiquitinated ChAT in MG132-treated cells expressing heterologous K48R-ubiquitin that is unable to form proteolytic K48-linked poly-Ub chains (Figure 3.11c). While this result may suggest that ChAT can be targeted

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for proteasomal degradation by non-classical polyubiquitin chains, such as K11-linked poly-Ub chains, it is more likely that under these experimental conditions small quantities of ChAT protein could be degraded through ubiquitination with endogenous K48-linked poly-Ub chains. To address this limitation in future experiments, ChAT polyubiquitination can be further characterized following anti-ChAT IPs using linkage-specific anti-ubiquitin antibodies (e.g. K11, K48, and K63 linkages) in substitution of total-ubiquitin antibodies that were used earlier in my studies (Figure 3.10).

A second experimental limitation is that my BioID experiments were completed in human HEK293 cells, and not in cholinergic SN56 cells, primarily due to the superior transfectability and transient expression of heterologous proteins typically seen in HEK293 cells. In support of my use of HEK293 cells, I found in initial optimization experiments that the transient expression of multiple ChAT-BirA* fusion proteins was the greatest in HEK293 cells as compared to either SN56 or SH-SY5Y cells, and that this correlated with a greater abundance in the total amount of biotinylated cellular proteins that could be used downstream for BioID (Appendix A). Additionally, another limitation of my BioID studies is that my large-scale BioID experiment with follow-up mass spectrometry was completed once without repetition (n=1), though this did result in the identification of HSP70 and HSP90 as potential ChAT-interacting proteins (Figures 3.18 – 3.22). It is important to note that while my large-scale BioID experiment was completed only once, biotinylated proteins correlating with the molecular mass of HSP70 were observed in repeated optimization experiments on streptavidin-HRP blots (Figure 3.17; n=2), and furthermore that HSP70 and HSP90 were directly confirmed by immunoblotting from streptavidin PDs from follow-up small-scale BioID experiments (Figure 3.23; n=4). While there are limitations to my use of HEK293 cells for BioID, the goal of these experiments was ultimately to identify potential ChAT-interacting proteins that could then later be confirmed by alternative methods, such as by co-IP and in situ PLA assays, in cholinergic cells. As such, and importantly, my BioID results were largely confirmed and expanded upon by follow-up studies in cholinergic SN56 cells, though I propose that any future BioID experiments investigating ChAT protein-interactions should be completed in cholinergic cells where feasible.

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Following BioID I used in situ PLA assays to test and confirm the presence of ChAT protein-interactions with HSC70 and HSP90 in SN56 cells. One limitation to my approach of these experiments was that cells were not co-stained with a second alternative anti-ChAT antibody that could be used to verify that PLA-positive cells were expressing heterologous human ChAT protein. The stringent nature of the in situ PLA technique is based on the observation that, to generate a positive PLA reaction, two antibody-labeled proteins must be no more than 40 nm apart from each other [565-567], and therefore it is unlikely to generate PLA signals unless the tested proteins in questions are in direct contact. Importantly, as no positive PLA signals was generated in control cells transfected with empty vector that were incubated with anti-ChAT and either anti-HSP or anti-CHIP antibodies (Figures 3.26 and 3.35), I propose that this limitation does not significantly impact my results. However, in future PLA experiments involving heterologous expression of human ChAT protein it will be important that this limitation be addressed.

Lastly, a fourth experimental limitation to my studies is the use of small molecule inhibitors for the study of HSC/HSP70, HSP90, and p97/VCP function. While this is a common approach to the study of molecular pathways, and provided that the inhibitors used within my studies, including PES and VER-155008 (HSC/HSP70), 17-AAG (HSP90), and Eer1 (VCP), have all been previously shown to be selective for and efficacious against their intended targets [213, 217, 516, 568], off-target effects of these small molecules must always be considered. The most common alternatives to small molecule inhibitors typically involve reducing (knock-down) or eliminating (knockout) the endogenous expression of a protein of interest. While knockout studies that do not require the generation of transgenic animals are becoming more common and accessible due to the advent of CRISP-Cas9 technology [560-561], RNAi-mediated knock-down remains the standard approach for cell-based studies that offers a high degree of experimental flexibility for the study of endogenous proteins in cells. Unfortunately, in preliminary experiments I found that siRNA-mediated knock-down of HSC/HSP70 reduced the cellular viability of SN56 cells by 70-90% as compared to cells transfected with scramble-control siRNA when assessed by alamar blue assay (Appendix B). Furthermore, while I was able to obtain an ~50% knock-down of HSP90 in SN56 cells, this is most likely insufficient to prevent the cellular

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chaperoning functions of HSP90 due to the high endogenous levels of HSP90 (1-2% of all cellular protein) [162, 227]. In relation to VCP inhibition, other studies have previously shown comparable results between Eer1 treatment and siRNA-mediated knock-down of VCP [515-516]. Though knock-down studies have been used to investigate the function of many cellular proteins, one advantage of pharmacological inhibition is the possibility to target different functional domains within a protein of interest. In my studies, this advantage was most evident when using the HSC/HSP70 inhibitors PES (substrate binding) and VER- 155008 (ATPase activity) that produced differing though complementary results towards ChAT protein solubility and stability in SN56 cells. Taken together, I propose that the use of small molecule inhibitors within this thesis was a safe and effective approach to studying the cellular mechanisms that regulate ChAT protein function.

4.4: Future Directions

Future work related to my studies can be separated into two categories involving either short-term follow-up experiments or more long-term projects. To begin, in the short-term it will be important to follow-up on select observations from my thesis given the experimental limitations to my studies as discussed above. Subsequently, I propose that an immediate follow-up study should be to test whether my core observations from SN56 cells are reproducible in alternative cholinergic cells, such as differentiated human SH-SY5Y cells and/or primary mouse neuron cultures. Importantly, these cell models can be used to confirm that (a) P17A/P19A- and CMS-mutant ChAT are targeted for proteasome- dependent degradation through enhanced ubiquitination, (b) that wild-type and mutant ChAT interact with HSC/HSP70 and HSP90, (c) that HSP inhibition promotes the insolubilization, ubiquitination, and degradation of ChAT protein, and (d) that the solubility and ubiquitination of ChAT is altered following exposure to cellular oxidative stress. Most if not all these follow-up experiments in alternative cholinergic cells will need to be performed using heterologous-expressed wild-type and/or mutant ChAT protein, and while this poses a potential setback due to transfection difficulties, recent studies have detailed new and enhanced transfection techniques designed for difficult-to-transfect cell

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lines (i.e. SH-SY5Y cells) and primary neuron cultures [564, 569-570]. Furthermore, alongside of these conformational studies using either human cholinergic cells lines or primary neuron cultures it may be useful to also assess whether (a) endogenous ChAT is targeted for proteasomal degradation, and (b) if HSP inhibition promotes degradation and/or reduces the enzymatic activity of endogenous ChAT in cells. Unfortunately, as tested previously in our laboratory, the relative protein expression of endogenous 69-kDa ChAT in cholinergic cells is very low when compared to cells transiently transfected to express heterologous ChAT (data not shown), making the molecular study of endogenous ChAT protein very challenging. As such, these follow-up experiments investigating endogenous ChAT are a possibility, though are not a priority.

In addition to these alternative cell model studies, the direct follow-up of specific experiments from this thesis in SN56 cells would be to useful to further build on my studies and expand our understanding of the mechanisms that regulate ChAT protein stability. These include, for example, (a) the use of chain-specific anti-ubiquitin antibodies to characterize ChAT polyubiquitination, (b) to assess whether inhibition of HSC/HSP70 by PES treatment results in enhanced ChAT ubiquitination in addition to reducing the solubility of ChAT protein as observed above (Figures 3.27 and 3.43), (c) to test if VCP inhibition using Eer1 also promotes ChAT insolubilization, and (d) to investigate if ChAT interacts with VCP under non-stressed conditions using co-IP and/or in situ PLA assays. In addition, further experimentation related to my novel SPAAC pulse-chase protocol is needed to build on and provide more evidence for the applicability, flexibility, and accuracy of this novel technique as compared to more standard CHX or 35S-methionine pulse-chase assays. To this end it will be important to perform SPAAC pulse-chase in alternative cell lines, potentially including primary cells, using either heterologous or endogenous proteins that have either an unknown (e.g. 82-kDa ChAT) or well-defined protein half-life (e.g. p53) [571-572]. Additionally, given the wide-array of commercially available click chemistry reagents, these follow-up SPAAC pulse-chase experiments could use alternative probes, such as various DIBO-conjugated Alexa Fluors, or Biotin-DIBO (Thermo) that would enable determination of protein half-life by streptavidin-HRP blotting without the need of a fluorescence detector or the use of radioactivity. It is important to note that, aside from

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alternative reagents, all the above immediate follow-up studies to my experimental work would utilize similar or exact techniques that were used throughout my thesis, allowing for the quick and straightforward assessment of these experiments.

Results from my thesis support further long-term investigation into alternative mechanisms beyond cytosolic HSPs that may regulate ChAT protein stability and function under physiological conditions and/or during cell stress and disease (Figure 4.2). First, in relation to my results detailing the effects of VCP inhibition, the stability and function of ChAT protein could be assessed in relation to ERAD-associated mechanisms. As discussed above, VCP is best known for its critical role in mediating ER-to-cytoplasm transfer of proteins ubiquitinated in the ER [441-442, 522], and furthermore inhibition of VCP by Eer1 treatment has been previously shown to inhibit the degradation of ubiquitinated VCP and ERAD substrates [515-516]. Thus, as in my studies I showed that VCP inhibition prevented the degradation of ubiquitinated ChAT, it is possible that the basal ubiquitination of ChAT may be regulated through ERAD-related mechanisms. Though I found that the E3 ubiquitin ligase CHIP interacts with ChAT in situ, siRNA-mediated knock-down of CHIP had no effect on ChAT protein levels, and thus the E3 ligase/s responsible for ChAT ubiquitination remain to be elucidated. Ubiquitination of mammalian ERAD substrates is regulated by two differing ER-bound E3 ubiquitin ligases, gp78/AMFR and Hrd1/SYVN1 [573-575]. Importantly, these E3 ubiquitin ligases and their ubiquitinated substrates can interact with VCP through the ER-localized adaptor protein Ubx2 [576-577], acting together to mediate the retrotranslocation of ERAD substrates into the cytoplasm for degradation by the 26S proteasome [522-524]. Though pharmacological inhibitors of these E3 ubiquitin ligases are not commercially available, siRNA-mediated knock-down has been used in the past as an effective tool to study the functions of gp78 and Hrd1 and of their cellular substrates [578- 580]. Interestingly, gp78, Hrd1, and VCP act together downstream of GRP-78 and GRP- 94, two ER-localized molecular chaperones that are related-to and that share homologous functions to HSC/HSP70 and HSP90 respectively [525, 555, 575, 581-582]. Thus, given that my studies have demonstrated a role for the cytoplasmic chaperones HSC/HSP70 and HSP90 on ChAT protein stability and function, it is possible that the homologous and ER- associated GRP-78 and/or GRP-94 may also chaperone nascent ChAT protein. Like gp78

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Figure 4.2: Hypothetical model for the regulation of ChAT protein stability by the endoplasmic reticulum-associated degradation (ERAD) system. (1) Nascent ChAT protein interacts with and is chaperoned by the heat shock proteins GRP-78 and GRP-94 in the ER lumen. (2) Once localized to the cell cytoplasm, semi-folded ChAT protein interacts with HSC/HSP70 and HSP90 in order to produce fully-folded and conformationally-active ChAT enzyme. (3) If initial folding of nascent ChAT protein in the ER by GRP-78 and/or GRP-94 is unsuccessful, ChAT may be ubiquitinated by the ER-localized E3 ubiquitin ligases gp78/AMFR and/or Hrd1/SYVN1. (4) p97/VCP, linked to the ER by the adaptor protein Ubx2, regulates retrotranslocation of ER-ubiquitinated ChAT into the cytoplasm for degradation by the cytosolic 26S proteasome. VCP inhibition by Eeyarestatin I (Eer1) treatment prevents the degradation of ubiquitinated ChAT. (5) Cellular stresses, such as ER and/or oxidative stress (e.g. H2O2, Aβ), may promote the misfolding of ChAT protein and subsequent degradation of ChAT through ERAD and/or cytosolic HSP mechanisms.

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and Hrd1, knock-down studies have been used in the past to elucidate the cellular functions and substrates of either GRP-78 [583-584] or GRP-94 [254, 585]. Therefore, alongside siRNA knock-down studies, I propose that co-IP and in situ PLA assays could be used to quickly and effectively assess whether ChAT may interact with various ERAD-associated chaperones and E3 ubiquitin ligases, and if so to determine if interaction with these ER proteins regulates ChAT protein stability in a manner consistent with their known ERAD- related functions. These initial studies can then be expanded upon by investigating ChAT ubiquitination and degradation under conditions of ERAD impairment (i.e. knock-down) or in cells exposed to various ER stress-inducing small molecules, including tunicamycin, thapsigargin, and brefeldin A [586-587]. Lastly, one of the most well-studied roles for ERAD is as a central hub for multiple protein quality control mechanisms that act together to triage misfolded and mutated proteins for proteasomal degradation [588-589], and as such ERAD may represent a cellular pathway responsible for the enhanced ubiquitination and degradation of various CMS-related ChAT mutants.

Lastly, another important avenue for future investigation will be to identify the cellular mechanisms that may promote ChAT stability and function during cellular stress and disease. Results from my studies related to oxidative stress suggest that cytosolic HSPs, particularly HSC/HSP70, may chaperone ChAT during cellular stress so to prevent stress- induced misfolding, loss of solubility, enhanced ubiquitination, and subsequent degradation of ChAT protein. Though these preliminary studies were completed using H2O2, a potent inducer of oxidative stress [590-591], alternative studies have demonstrated that ChAT activity is altered in cells acutely exposed to AD-related and oxidative Aβ1-42 oligomers [438-440]. Additionally, impairment of ChAT activity has been associated with the pathogenesis of AD [432-435], an age-related neurodegenerative disease characterized partly by the dysregulation of molecular chaperones, including HSPs, and various protein quality control mechanisms [10, 329]. As such, an important question that will need addressing is whether acute and/or prolonged cellular exposure to oxidative Aβ1-42 may promote the insolubilization and ubiquitination of ChAT protein like that observed in my studies following H2O2 treatment. Furthermore, a potential protective role for HSC/HSP70 and/or HSP90 towards ChAT function during Aβ-stress could be assessed by co-treatment

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of ChAT-expressing cells with Aβ1-42 in combination with inhibitors of either HSC/HSP70 (e.g. PES and VER-155008) or HSP90 (e.g. 17-AAG). Following these treatments, the solubility, ubiquitination, and enzymatic activity of ChAT protein could be determined using similar techniques to those used in the thesis, including but not limited to Triton X- 100 fractionation, IP/co-IP assays, and radioenzymatic assay to determine cellular ChAT activity. Interestingly, multiple studies have reported on the potential therapeutic potential for either activation of HSF1 (i.e. the heat shock response) or enhancement of HSC/HSP70 activity on the pathogenesis of protein misfolding neurodegenerative diseases, including AD [592-594], and as such it would be interesting to test if these strategies may help to ameliorate ChAT dysfunction during cellular stress and disease. It is important to note that, in addition to oxidative stress, another key feature of many age-related neurodegenerative diseases is heightened ER stress and ER dysfunction due to the accumulation of misfolded proteins [595-596], and subsequently studies have shown that cellular exposure to cytotoxic Aβ can directly induce ER stress [597-598]. Of significance, previous work from our laboratory has shown that interaction of ChAT with the ER-associated co-chaperone p97/VCP is enhanced in neural cells treated with Aβ1-42 [440], suggesting that the regulation of ChAT ubiquitination and function may be altered following exposure to ER and/or oxidative stress. Therefore, these proposed studies investigating the effects of Aβ- stress on ChAT protein stability may be intimately linked with my other proposed studies assessing a potential role for ER-associated chaperones and ERAD-related mechanisms on the regulation of ChAT protein function (Figure 4.2).

4.5: Significance and Concluding Remarks

In summary, for this thesis I initially set out to determine a functional role for the highly- conserved N-terminal proline-rich motif of human 69-kDa ChAT and found that both the cellular protein stability and enzyme activity of ChAT protein is dramatically reduced following alteration of this motif. In addition, I demonstrated for the first time that ChAT protein levels are modulated through the ubiquitin-proteasome system, and furthermore that the ubiquitination of catalytically-deficient ChAT, such as CMS-related V18M-ChAT,

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is increased. In search for cellular mechanisms that may regulate ChAT protein stability and/or that are responsible for enhanced ubiquitination of mutant ChAT, I found evidence that a network of cytoplasmic and ER-associated chaperones, which included the heat shock proteins HSC/HSP70 and HSP90 along with the co-chaperone Cdc48/p97/VCP, regulates ChAT protein stability and function. The regulation of specific catalytically- deficient ChAT mutants (i.e. P17A/P19A, V18M, and A513T-ChAT) by these chaperones is altered, suggesting a mechanism that may be relevant to other CMS-related ChAT mutants or to ChAT protein during cellular stress and disease. Subsequently, in preliminary experiments I observed evidence that HSC/HSP70 may play a role in the maintenance of ChAT protein solubility, stability, and function during cellular exposure to oxidative stress. For future studies, this thesis supports research into the mechanisms that regulate ChAT ubiquitination and the role that ubiquitination may play in relation to ChAT dysfunction during cellular stress. Lastly, my results merit additional assessment of the roles for various cytosolic and ER-associated HSPs, along with the co-chaperone VCP, on ChAT stability and function under both physiological conditions and during cellular stress and disease.

In conclusion, this thesis has established and characterized the roles of various molecular chaperones and the ubiquitin-proteasome system on the regulation of human 69- kDa ChAT protein stability and function. It is important to note that to date ChAT is the sole enzyme known to synthesize the neurotransmitter ACh, and thus is both critical to and necessary for the function of cholinergic neurons [390, 392, 599]. These neurons can be found throughout the central, peripheral, and somatic nervous systems, where they are involved in the regulation of numerous bodily functions. These processes include cognitive functions such as behaviour, attention, and learning/memory, as well as various autonomic (e.g. pupil dilation, heart rate, and gastric and renal function) and neuromuscular processes (i.e. movement and motor control) [389-390, 392, 599-600]. Consequently, dysregulation of ChAT function, ACh synthesis, and cholinergic neurotransmission has been observed in multiple neurodegenerative and neuromuscular disorders including AD and CMS [387- 388, 411, 457-458]. Thus, by gaining a deeper understanding of the factors that regulate ChAT protein stability and activity, the work presented within this thesis may have broader implications to the study of ChAT protein function during cellular stress and in disease.

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APPENDICES

Appendix A: Optimization of BioID in multiple mammalian cell lines. Comparison of ChAT-BirA* fusion protein expression and the levels of biotinylated cellular proteins in human HEK293 as compared to either human SH-SY5Y (A; n=2) or mouse SN56 cells (B; n=2) following BioID. Cells were transfected to transiently express either 69-kDa (A and B) or 82-kDa (B) ChAT-BirA* fusion proteins using either Lipofectamine 2000 (A) or 3000 (A and B). Cells were treated for 24 h with 50 µM biotin to induce proximity- dependent biotinylation of ChAT-interacting proteins prior to collection and lysis. Biotin- treated control cells were transfected with empty pcDNA3.1+ vector or vectors encoding for untagged 69- or 82-kDa ChAT proteins. Altogether, transfection of human HEK293 cells using Lipofectamine 3000 was found to be superior for both the transient expression of ChAT-BirA* and for the accumulation and detection of cellular biotinylated proteins as compared to either SH-SY5Y or SN56 cells. As such, this method was used to perform my BioID experiments as detailed earlier (Figures 3.17 – 3.23). Biotinylated cellular proteins from whole cell lysates were detected by blotting with HRP-conjugated streptavidin (Strep- HRP). Relative expression of ChAT-BirA* fusion proteins and of untagged ChAT proteins were assessed by anti-ChAT immunoblotting. Anti-actin immunoblotting from whole cell lysates was completed as a loading control. Optimization experiments for BioID were completed by Sabin Rajkarnikar, B.Sc.

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Appendix B: Optimization of siRNA-mediated knock-down of endogenous HSC/HSP70 and HSP90. (A) Anti-HSC70 and anti-HSP90 immunoblots from SN56 cells that were transfected for 72 h with 10-100 nM of either anti-HSC70 or anti-HSP90 siRNA. Control cells were mock-transfected or transfected with 50 nM of scramble-control siRNA. Following incubation with siRNAs an ~50% knock-down of either HSC70 and HSP90 was observed. Anti-actin immunoblotting was completed as a loading control (n=2). (B) siRNA knock-down of HSC70 promotes the stress-induced expression of both HSP70 and HSP90 in SN56 cells. Accumulation of stress-induced HSP70 was preventable by co-transfecting cells with anti-HSP70 siRNA (n=2). (C) Knock-down of HSC70 either alone or together with stress-induced HSP70 reduces the viability of SN56 cells as compared to scramble- control transfected cells (***p≤0.001; one-way ANOVA with Dunnett’s post-hoc test, mean ± SEM). Cell viability was measured by alamar blue assay after 72 h of siRNA incubation (n=2).

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v1.10 Last updated September 2015 Questions? [email protected] or +1-855-239-3415 (toll free in the US) or +1-978-646-2777.

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Dear Dr. Morey,

Thank you for your email.

I would like to confirm that under the Frontiers Terms and Conditions, authors retain the copyright to their work. All Frontiers articles are Open Access and distributed under the terms of the Creative Commons Attribution License (CC-BY 3.0), which permits the re-use, distribution and reproduction of material from published articles, provided the original authors and source are credited.

Please keep in mind that if anything in the paper, such as a figure, was already under copyright restriction from any other third-party, then you would have to seek permission to reuse the item.

I hope I have addressed your question. Please let me know if you have any other questions or concerns.

Best regards,

Iacopo

Iacopo Marcon, PhD

Journal Development Specialist

Frontiers | Editorial Office - Journal Development Team

Journal Manager: Stéphanie Maret, PhD

------Forwarded message ------From: Frontiers Editorial Office Date: Tue, Feb 27, 2018 at 9:22 AM Subject: Fwd: Permission to Use Copyrighted Material in a Doctoral/Master’s Thesis To: Frontiers Research Topics Neuroscience

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------Forwarded message ------From: Trevor Morey Date: Mon, Feb 26, 2018 at 5:57 PM Subject: Permission to Use Copyrighted Material in a Doctoral/Master’s Thesis To: [email protected]

Hello,

I am a University of Western Ontario graduate student completing my Doctoral / Master’s thesis entitled “Regulation of Human 69-kDa Choline Acetyltransferase Protein Stability and Function by Molecular Chaperones and the Ubiquitin-Proteasome System”. My thesis will be available in full-text on the internet for reference, study and / or copy. Except in situations where a thesis is under embargo or restriction, the electronic version will be accessible through the Western Libraries web pages, the Library’s web catalogue, and also through web search engines. I will also be granting Library and Archives Canada and ProQuest/UMI a non-exclusive license to reproduce, loan, distribute, or sell single copies of my thesis by any means and in any form or format. These rights will in no way restrict republication of the material in any other form by you or by others authorized by you.

I would like permission to allow inclusion of figures and text from the following publication; the materials will be attributed through citation:

Morey, T.M., Winick-Ng, W., Seah, C., and Rylett, R.J. (2017). Chaperone-mediated regulation of choline acetyltransferase protein stability and activity by HSC/HSP70, HSP90 and p97/VCP. Front Mol Neurosci. 10:415. doi: 10.3389/fnmol.2017.00415.

Please confirm in writing or by email that these arrangements meet with your approval.

Sincerely Trevor Morey

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CURRICULUM VITAE

Trevor M. Morey PubMed ID: “Morey TM”

Education:

• Ph.D Physiology; Western University: 2012 – 2018. o Supervisor: Dr. R. Jane Rylett o Thesis: Regulation of Human 69-kDa Choline Acetyltransferase Protein Stability and Function by Molecular Chaperones and the Ubiquitin- Proteasome System – Defended January 2018

• M.Sc. Molecular and Cellular Biology; University of Guelph: 2009 – 2012. o Supervisor: Dr. Richard Mosser o Thesis: The Effect of Noxa serine-13 Phosphorylation on Hyperthermia- induced Apoptosis – Defended January 2012.

• B.Sc. Molecular Biology and Genetics; University of Guelph: 2005 – 2009.

Scholarships and Awards:

• Robarts Research Institute Trainee Award; awarded a platform presentation at Robarts Research Retreat – June 2016

• Queen Elizabeth II Scholarship – Ontario Graduate Scholarship; $15,000 awarded annually; Awarded 2x: September 2013–2014, September 2014–2015

Publications:

1. Morey, T.M., Winick-Ng, W., Seah, C., and Rylett, R.J. (2017). Chaperone-mediated regulation of choline acetyltransferase protein stability and activity by HSC/HSP70, HSP90 and p97/VCP. Front. Mol. Neurosci. 10:415. doi: 10.3389/fnmol.2017.00415.

2. Winick-Ng, W., Caetano, F.A., Winick-Ng, J., Morey, T.M., Heit, B., and Rylett, R.J. (2016). 82-kDa choline acetyltransferase and SATB1 localize to -amyloid induced matrix attachment regions. Sci. Rep. 6:23914. doi: 10.1038/srep23914.

3. Morey, T.M., Albers, S., Shilton, B.H., and Rylett, R.J. (2016). Enhanced ubiquitination and proteasomal-degradation of catalytically-deficient human choline acetyltransferase mutants. J. Neurochem. 137, 630-646.

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4. Morey, T.M., Roufayel, R., Johnston, D.S., and Mosser, D.D. (2015). Heat shock inhibition of CDK5 increases NOXA levels through miR-23a repression. J. Biol. Chem. 290, 11443-11454.

Scientific Meetings and Conferences:

• Robarts Research Retreat, London, Ontario, Canada; June 20, 2017 o Poster: Ubiquitination and Protein Stability of Catalytically-deficient Human Choline Acetyltransferase Mutants are regulated by the molecular chaperones HSC70 and HSP90s o Authors: Trevor M. Morey, Claudia Seah, and R. Jane Rylett o Scope: Institutional

• London Health Research Day; London, Ontario, Canada; March 28, 2017 o Poster: Ubiquitination and Protein Stability of Catalytically-deficient Human Choline Acetyltransferase Mutants are regulated by the molecular chaperones HSC/HSP70 and HSP90s o Authors: Trevor M. Morey, and R. Jane Rylett o Scope: Institutional o Facilitator for Salon B Platform Presentations, morning session

• Physiology and Pharmacology Annual Research Day; London, Ontario, Canada; November 1, 2016. o Poster: Regulation of Catalytically-deficient Human Choline Acetyltransferase Mutants by the Heat Shock Family of Molecular Chaperones and the E3 Ubiquitin Ligase CHIP o Authors: Trevor M. Morey, Warren Winick-Ng, and R. Jane Rylett o Scope: Departmental o Awarded first place prize - Neurosciences

• Robarts Research Retreat, London, Ontario, Canada; June 13, 2016 o Platform Presentation: Enhanced Ubiquitination and Proteasomal- degradation of Catalytically-deficient Human Choline Acetyltransferase Mutants o Authors: Trevor M. Morey, Shawn Albers, and R. Jane Rylett o Scope: Institutional

• London Health Research Day; London, Ontario, Canada; March 29, 2016. o Platform Presentation: Enhanced Ubiquitination and Proteasomal- degradation of Catalytically-deficient Human Choline Acetyltransferase Mutants o Authors: Trevor M. Morey, Shawn Albers, and R. Jane Rylett o Scope: Institutional

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• Physiology and Pharmacology Annual Research Day; London, Ontario, Canada; November 3, 2015. o Poster: Decreased Activity and Increased Ubiquitin-Mediated Degradation of Human Choline Acetyltransferase by Mutation of an N-terminal Proline-Rich Motif. o Authors: Trevor M. Morey, Shawn Albers, Brian H. Shilton, and R. Jane Rylett o Scope: Departmental o Awarded first place prize - Neurosciences

• Neuroscience 2015; Society for Neuroscience, Chicago, Illinois, USA; October 17-21, 2015. o Poster: Decreased Activity and Increased Ubiquitin-Mediated Degradation of Human Choline Acetyltransferase by Mutation of an N-terminal Proline-Rich Motif. o Authors: Trevor M. Morey, Shawn Albers, Brian H. Shilton, and R. Jane Rylett o Scope: International

• Robarts Research Retreat; London, Ontario, Canada; June 8, 2015. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Institutional

• London Health Research Day; London, Ontario, Canada; April 1, 2015. o Platform Presentation: Regulation of Human Choline Acetyltransferase Protein Stability and Enzyme Activity by an N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Institutional

• Physiology and Pharmacology Annual Research Day; London, Ontario, Canada; November 4, 2014. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Departmental o Awarded first place prize - Neurosciences

• Canadian Student Health Research Forum; Winnipeg, Manitoba, Canada; June 10-12, 2014. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: National; invite-only o Awarded an honorable mention

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• Robarts Research Retreat; London, Ontario, Canada; June 9, 2014. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Institutional

• Southern Ontario Neuroscience Association; London, Ontario, Canada; May 5, 2014. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Regional

• London Health Research Day; London, Ontario, Canada; March 18, 2014. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Institutional o Awarded first place prize – Neuroscience and Mental Health

• Physiology and Pharmacology Annual Research Day; London, Ontario, Canada; November 4, 2013. o Poster: Regulation of Human Choline Acetyltransferase Protein Stability by a N-terminal Proline-rich Motif. o Authors: Trevor M. Morey, R. Jane Rylett o Scope: Departmental

• Molecular and Cellular Biology Annual Research Day; Guelph, Ontario, Canada; April 2009. o Poster: The Effect of Noxa Serine-13 Phosphorylation on Hyperthermia- induced Apoptosis. o Authors: Trevor M. Morey, Richard Mosser o Scope: Departmental

Scientific and Technical Expertise:

• Mammalian cell culture: o Culture and maintenance of primary murine cells and mammalian cell lines. o Plasmid transfection of adherent and non-adherent human cells lines. o Generation of stably-expressing mammalian cells lines. o Pharmacological experimentation.

• Molecular techniques: o Plasmid design and generation: PCR (for both insertion and mutagenesis cloning), restriction digestion and purification, plasmid ligation.

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o Plasmid preparation: Bacterial transformation, plating, growth, and purification. o Immunoblotting: Mammalian cell lysis, protein sample preparation, 1D SDS- PAGE, electroblotting, and protein detection by chemiluminescence using film or ChemiDoc. o Immunoprecipitation, co-immunoprecipitation, and affinity purification. o Confocal microscopy o Advanced techniques: BioID, PLA, SDD-AGE. o Fluorescent and colorimetric plate-reader assays. o Radioenzymatic assays, 32P-orthophosphate live-labeling and detection. o Generation of novel assays (i.e. SPAAC pulse-chase).

• Biochemical techniques: o Recombinant protein purification: Expression in E. coli BL21, bacterial lysis, Ni2+ affinity purification, TEV digestion, dialysis, cation exchange purification. o Biochemical plate-reader assays: enzymatic assays. o Circular dichroism with tandem thermal denaturation for measurement of in vitro protein structure and thermal stability. o MALDI and ESI mass spectrometry to identify protein-protein interactors

Teaching Experience:

• Phys1020 – Human Physiology, Western University, Sept. 2015 – April 2016. o Supervisor: Christine Bell o Position: Teaching Assistant

• Phys1020 – Human Physiology, Western University, Sept. 2014 – April 2015. o Supervisor: Christine Bell o Position: Teaching Assistant

• Phys1020 – Human Physiology, Western University, Sept. 2013 – April 2014. o Supervisor: Tom Stavraky o Position: Teaching Assistant

• Phys3130y – Physiology Laboratory, Western University, Sept. 2013 – April 2013. o Supervisor: Tom Stavraky o Position: Teaching Assistant

• Biol1090 – Introduction to Molecular and Cellular Biology, University of Guelph, Jan. 2011 – April 2011. o Supervisor: Kim Kirby o Position: Teaching Assistant

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• MCB4010 – Advanced Cell Biology, University of Guelph, Jan. 2011 – April 2011; Jan. 2010 – April 2010. o Supervisor: Dr. Richard Mosser o Position: Teaching Assistant

• MBG2000 – Introductory Genetics, University of Guelph, Sept. 2010 – Dec. 2010; May 2010 – Aug. 2010; Sept. 2009 – Dec. 2009. o Supervisor: Christine Schisler o Position: Teaching Assistant

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