Medicine, Faculty of
Medical Genetics, Department of
DSpace
UBCV
Kumar, Arun
Doctor of Philosophy - PhD
University of British Columbia
Proteostasis is critical for cell survival. Exposure to stressors such as DNA damage, heat, mutations, or ageing can accumulate proteins in non-native conformations. The accumulation of these misfolded proteins, through either gain- or loss-of-function, disrupts vital cellular processes. This is notably evident in neurodegenerative diseases such as Alzheimer’s, and Parkinson's which are linked to protein aggregation pathology. Therefore, cells strive to maintain a healthy proteome which is achieved through a complex network of protein quality control (PQC) circuits. Traditionally, PQC was thought to rely on two major pathways: molecular chaperones for refolding and proteolytic systems for degradation. However, this perspective has been challenged with the discovery of protein sequestration. Under severe stress conditions that overwhelm PQC systems, misfolded proteins are spatially sequestered to specialized and distinct membrane-less inclusions within the cell. This spatial compartmentalization represents a crucial aspect of PQC, facilitating the storage of disassembled or non-native proteins until they can be either refolded or degraded in a regulated manner. The model organism Saccharomyces cerevisiae or budding yeast has three well-established sequestration sites – the Insoluble PrOtein Deposit (IPOD), the JUxtaNuclear Quality control (JUNQ) site and the IntraNuclear Quality control (INQ) site. Although substantial efforts have contributed to identifying key proteins involved in the formation and dissolution of these sequestration sites, our understanding of the regulation and dynamics of INQ remains incomplete. In my thesis, I provide a comprehensive analysis of all proteins that have been found at INQ. Furthermore, I explore different pathways and stressors to expand our knowledge of the factors governing INQ formation. To do so, I first characterize Rpd3, a histone deacetylase, as a novel INQ marker and use its localization as a proxy for INQ levels. Through microscopy and genetic approaches, I demonstrate that INQ formation of Rpd3 is a general response to DNA damage. Furthermore, I explore the role of the post-translational modification (PTM) SUMOylation at INQ and document the DNA damage induced SUMOylation of chaperone Btn2 and Hsp42. Additionally, I engineer a non-SUMOylatable Btn2 and dissect its role in INQ clearance, placing it at the nexus of protein degradation and refolding at INQ.
DNA damage induced SUMOylation regulates intranuclear protein quality control in Saccharomyces cerevisiae by ARUN KUMAR B.Tech., Manipal Institute of Technology, 2017 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (MEDICAL GENETICS) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2023 © Arun Kumar, 2023 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: DNA damage induced SUMOylation regulates intranuclear protein quality control in Saccharomyces cerevisiae submitted by Arun Kumar in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Medical Genetics Examining Committee: Peter Stirling, Associate Professor, Department of Medical Genetics, UBC Supervisor Elizabeth Conibear, Professor, Department of Medical Genetics, UBC Supervisory Committee Member Pamela Hoodless, Professor, Department of Medical Genetics, UBC University Examiner Corey Nislow, Professor, Faculty of Pharmaceutical Sciences, UBC University Examiner Additional Supervisory Committee Members: Thibault Mayor, Professor, Department of Biochemistry and Molecular Biology, UBC Supervisory Committee Member Philip Hieter, Professor, Department of Medical Genetics, UBC Supervisory Committee Member iii Abstract Proteostasis is critical for cell survival. Exposure to stressors such as DNA damage, heat, mutations, or ageing can accumulate proteins in non-native conformations. The accumulation of these misfolded proteins, through either gain- or loss-of-function, disrupts vital cellular processes. This is notably evident in neurodegenerative diseases such as Alzheimer’s, and Parkinson's which are linked to protein aggregation pathology. Therefore, cells strive to maintain a healthy proteome which is achieved through a complex network of protein quality control (PQC) circuits. Traditionally, PQC was thought to rely on two major pathways: molecular chaperones for refolding and proteolytic systems for degradation. However, this perspective has been challenged with the discovery of protein sequestration. Under severe stress conditions that overwhelm PQC systems, misfolded proteins are spatially sequestered to specialized and distinct membrane-less inclusions within the cell. This spatial compartmentalization represents a crucial aspect of PQC, facilitating the storage of disassembled or non-native proteins until they can be either refolded or degraded in a regulated manner. The model organism Saccharomyces cerevisiae or budding yeast has three well-established sequestration sites – the Insoluble PrOtein Deposit (IPOD), the JUxtaNuclear Quality control (JUNQ) site and the IntraNuclear Quality control (INQ) site. Although substantial efforts have contributed to identifying key proteins involved in the formation and dissolution of these sequestration sites, our understanding of the regulation and dynamics of INQ remains incomplete. iv In my thesis, I provide a comprehensive analysis of all proteins that have been found at INQ. Furthermore, I explore different pathways and stressors to expand our knowledge of the factors governing INQ formation. To do so, I first characterize Rpd3, a histone deacetylase, as a novel INQ marker and use its localization as a proxy for INQ levels. Through microscopy and genetic approaches, I demonstrate that INQ formation of Rpd3 is a general response to DNA damage. Furthermore, I explore the role of the post-translational modification (PTM) SUMOylation at INQ and document the DNA damage induced SUMOylation of chaperone Btn2 and Hsp42. Additionally, I engineer a non-SUMOylatable Btn2 and dissect its role in INQ clearance, placing it at the nexus of protein degradation and refolding at INQ. v Lay Summary Proteins in our cells form in a chain that must fold into specific conformations to perform their functions. When this process goes awry, or cells are under stress, proteins tend to misfold and stick to each other. The resultant aggregates have been linked to aging, neurodegenerative diseases, and cancer. To prevent formation of these toxic aggregates of misfolded proteins, the cell employs various protein quality control (PQC) mechanisms. I used the model organism of budding yeast to investigate one such PQC mechanism called sequestration wherein misfolded proteins are trafficked to specialized compartments within the cell. Specifically, I explore the role of DNA damage as a stress and other genetic pathways in the formation of these compartments. In addition, I focus on a chemical tag called SUMOylation and its role in sequestration. Overall, this research expands our understanding of one of the most poorly understood PQC mechanisms. vi Preface This dissertation is the original intellectual product of Arun Kumar, unless otherwise noted below. A version of Chapter 1 has been published as a review article, written collaboratively by Veena Mathew, Peter C. Stirling and myself as Kumar*, A., Mathew, V. & Stirling, P.C. Nuclear protein quality control in yeast: The latest INQuiries. (2022) J Biol. Chem. 298(8) 102199. Additional text presented in Chapter 1 was published as a review article, written collaboratively by Louis-Alexandre Fournier, Peter C. Stirling and myself as Fournier*, L.-A.,Kumar*,A. & Stirling, P.C. Chromatin as a Platform for Modulating the Replicating Stress Response. (2018) Genes (Basel) 9, E622. *Designates co-first authorship. A version of Chapter 2, 3 and 4 formed the basis of a manuscript that has been submitted for peer-review, titled “DNA damage induced SUMOylation of Btn2 regulates nuclear protein quality control”. I designed, performed, and analyzed all the experiments with assistance in method development by Veena Mathew, and wrote the whole manuscript under the guidance of P. Stirling. Ethics approval for this research was obtained using the following certificate: Biosafety protocol B22-0122, entitled Exploring genome instability in eukaryotes. Published work that are not presented in this thesis are listed below: vii Kumar, A.*, Fournier, L-A.*, & Stirling, P.C. Integrative analysis and prediction of human R-loop binding proteins. (2022) G3 Genes|Genomes|Genet. 12(8), jkac142. *Designates co-first authorship. Mathew, V., Kumar, A., Jiang, Y.K., West, K., Tam, A.S., Stirling, P.C. Cdc48 regulates intranuclear quality control sequestration of the Hsh155 splicing factor in budding yeast. (2020) J Cell Sci 133 (23):jcs252551. Chang, E.Y-C., Tsai, S., Aristizabal, M.J., Wells, J.P., Coulombe, Y., Busatto, F.F., Chan, Y.A., Kumar, A., Zhu, Y.D., Wang, A.Y-H., Fournier, L-A., Hieter, P., Kobor, M.S., Masson, J-Y., Stirling, P.C. MRE11-RAD50-NBS1 promotes Fanconi Anemia R-loop suppression at transcription-replication conflicts. (2019) Nat Commun 10, 4265. Tam, A.S., Tsai, S., Chang, E.Y-C., Mathew, V., Shanks, A., Docking, T.R., Kumar, A., Bernard, D.G., Karsan, A., Stirling, P.C. DYNLL1 mis-splicing is associated with replicative genome instability in SF3B1 mutant cells. (2021). bioRxiv 2021.05.26.445839 viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables .............................................................................................................................. xiii List of Figures ............................................................................................................................. xiv List of abbreviations ................................................................................................................. xvii Acknowledgements ......................................................................................................................xx Dedication .................................................................................................................................. xxii Chapter 1: Introduction ................................................................................................................1 1.1 Proteostasis, misfolding and aggregation ....................................................................... 1 1.1.1 Co-translational folding of nascent polypeptides ....................................................... 3 1.1.2 Molecular chaperones and the respective families ..................................................... 4 1.1.2.1 The Hsp40/Hsp70/Hsp110 families .................................................................... 6 1.1.2.2 The Hsp90 family ............................................................................................... 9 1.1.2.3 Hsp104/AAA+ ATPases ................................................................................... 10 1.1.2.4 Chaperonins ...................................................................................................... 11 1.1.2.5 Small heat shock proteins (sHsps) .................................................................... 11 1.2 The three pillars of protein quality control ................................................................... 12 1.2.1 Disaggregation and protein refolding ....................................................................... 14 ix 1.2.2 Protein degradation ................................................................................................... 16 1.2.2.1 The Ubiquitin-Proteasome System ................................................................... 16 1.2.2.2 Ubiquitin ........................................................................................................... 17 1.2.2.3 SUMOylation .................................................................................................... 20 1.2.2.4 Autophagy ......................................................................................................... 22 1.2.3 Sequestration ............................................................................................................. 24 1.2.3.1 Intranuclear Quality Control: the nuclear protein sequestration structure ........ 26 1.2.3.1.1 The INQ proteome ...................................................................................... 29 1.2.3.1.2 Protein refolding at INQ .............................................................................. 31 1.2.3.1.3 Protein degradation at INQ ......................................................................... 34 1.2.3.1.4 Posttranslational modifications at INQ ....................................................... 37 1.2.3.1.5 Nucleophagy and the fate of INQ ............................................................... 39 1.2.3.2 Juxtanuclear Quality control site (JUNQ) and the Insoluble Protein Deposit (IPOD)…........................................................................................................................... 40 1.2.3.3 MitoStores and Stress Granules ........................................................................ 41 1.2.3.4 Aggresomes....................................................................................................... 41 1.3 DNA Replication stress and response ........................................................................... 43 1.3.1 Double strand breaks and DNA repair ...................................................................... 44 1.3.2 Chromatin organization during replication stress ..................................................... 45 1.3.2.1 Histone acetylation............................................................................................ 47 1.4 Thesis objectives ........................................................................................................... 49 Chapter 2: Rpd3 is a novel INQ marker. ..................................................................................51 2.1 Background ................................................................................................................... 51 x 2.2 Materials and Methods .................................................................................................. 53 2.2.1 Experimental Model and Subject Details ................................................................. 53 2.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods .................. 53 2.2.3 Western Blotting and protein stability time course................................................... 54 2.2.4 Fractionation of the insoluble pellet ......................................................................... 55 2.3 Results ........................................................................................................................... 55 2.3.1 Rpd3 is sequestered at INQ during replication stress. .............................................. 55 2.3.2 Sequestration to INQ is mediated by small-heat shock proteins. ............................. 57 2.3.3 Sequestration of Rpd3 to INQ is a DNA damage response. ..................................... 61 2.3.4 INQ turnover for Rpd3 is mediated by chaperones. ................................................. 64 2.3.4.1 Defining regions of Apj1 involved in INQ clearance. ...................................... 64 2.3.4.2 Nucleophagy affects INQ turnover. .................................................................. 67 2.4 Discussion ..................................................................................................................... 69 Chapter 3: DNA damage induces SUMOylation of sHsps and regulates INQ formation. ...71 3.1 Background ................................................................................................................... 71 3.2 Materials and Methods .................................................................................................. 72 3.2.1 Experimental Model and Subject Details ................................................................. 72 3.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods .................. 73 3.2.3 Western Blotting and protein stability time course................................................... 74 3.2.4 Fractionation of the insoluble pellet ......................................................................... 74 3.2.5 Ni-NTA pulldown of SUMOylated proteins ............................................................ 75 3.3 Results ........................................................................................................................... 76 3.3.1 Perturbation of SUMOylation reduces INQ formation. ............................................ 76 xi 3.3.2 DNA damage induces SUMOylation of sequestrase chaperones. ............................ 79 3.3.3 Identification of the SUMOylated Btn2 domain ....................................................... 83 3.3.4 Mapping the modified lysines in the Btn2 C-terminal domain................................. 86 3.4 Discussion ..................................................................................................................... 91 Chapter 4: Btn2 SUMOylation regulates INQ clearance. ........................................................93 4.1 Background ................................................................................................................... 93 4.2 Material and Methods ................................................................................................... 94 4.2.1 Experimental model and Subject details ................................................................... 94 4.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods .................. 95 4.2.3 Western Blotting and protein stability time course................................................... 96 4.2.4 Fractionation of the insoluble pellet ......................................................................... 96 4.3 Results ........................................................................................................................... 96 4.3.1 Non-SUMOylatable Btn2 is a functional sequestrase. ............................................. 96 4.3.2 Btn2 localization is unaffected by its SUMOylation status. ..................................... 98 4.3.3 Probing the role of Btn2 SUMOylation in its degradation. ...................................... 99 4.3.4 Loss of SUMOylation affects Btn2 solubility. ....................................................... 101 4.3.5 Btn2-SUMO restrains INQ accumulation. .............................................................. 104 4.3.6 Btn2 SUMOylation mediates INQ clearance. ......................................................... 106 4.3.7 Btn2-SUMO regulates ubiquitin levels in sequestration compartments. ................ 113 4.3.8 Slx5/8 activity on the MMS-induced insoluble proteome. ..................................... 116 4.3.9 Discussion ............................................................................................................... 122 Chapter 5: Conclusion ...............................................................................................................124 5.1 Summary of data ......................................................................................................... 124 xii 5.2 DNA damage causes INQ formation. ......................................................................... 128 5.3 The significance of the INQ proteome. ....................................................................... 129 5.4 Importance of sequestration in humans, SUMO, and PML bodies ............................ 130 5.5 Limitations and caveats............................................................................................... 131 5.6 Future directions ......................................................................................................... 132 5.6.1 Hsp42 SUMOylation .............................................................................................. 132 5.6.2 Identification of the structural component of INQ deposition ................................ 133 5.6.3 Why sequester Rpd3? ............................................................................................. 134 5.6.4 The role of phase separation at INQ. ...................................................................... 135 Bibliography ...............................................................................................................................136 Appendices ..................................................................................................................................168 Appendix A ............................................................................................................................. 168 A.1 Yeast strains used in this study. .............................................................................. 168 A.2 Primers used in this study. ...................................................................................... 175 A.3 Plasmids used in this study. .................................................................................... 176 xiii List of Tables Table 1.1 The INQ Proteome. ....................................................................................................... 27 xiv List of Figures Figure 1.1 The protein folding landscape. ...................................................................................... 1 Figure 1.2 The Hsp70 chaperone cycle. .......................................................................................... 8 Figure 1.3 The triage decisions carried out by the proteostasis network. ..................................... 13 Figure 1.4 Major protein quality control sites in yeast. ................................................................ 25 Figure 1.5 Chaperones at INQ. ..................................................................................................... 32 Figure 1.6 Players and pathways for INQ refolding or degradation. ............................................ 37 Figure 2.1 Rpd3 is sequestered to INQ upon replication stress. ................................................... 56 Figure 2.2 Sequestration of Rpd3 to INQ is mediated by sHsps Btn2 and Hsp42. ...................... 58 Figure 2.3 Rpd3 complex partners do not relocalize to INQ. ....................................................... 60 Figure 2.4 INQ formation for Rpd3 following different genotoxic treatments. ........................... 61 Figure 2.5 DNA damage regulates INQ formation for Rpd3. ...................................................... 63 Figure 2.6 Apj1 affects turnover of Rpd3 at INQ. ........................................................................ 65 Figure 2.7 Cdc48 and nucleophagy regulate INQ turnover. ......................................................... 67 Figure 3.1 Schematic for the E1, E2, E3 cascade mediated SUMOylation. ................................. 76 Figure 3.2 Mutations in the SUMOylation cascade reduce INQ formation. ................................ 77 Figure 3.3 PolySUMOylation is involved in INQ turnover. ......................................................... 79 Figure 3.4 Schematic of the SUMOylation mini-screen utilizing GFP-constructs of substrates and chaperones found at INQ. ............................................................................................................. 80 Figure 3.5 INQ substrates are not SUMOylated upon replicative stress. ..................................... 81 Figure 3.6 DNA damage induces SUMOylation of sHsps Btn2 and Hsp42. ............................... 82 xv Figure 3.7 Schematic and NiPDs of GFP-Btn2 constructs used to determine the SUMOylated region. ........................................................................................................................................... 83 Figure 3.8 Btn2 is SUMOylated at its C-terminal domain. .......................................................... 84 Figure 3.9 Rescue experiments in a btn2Δ background using different domain constructs. ........ 85 Figure 3.10 Mutating all CTD lysines to arginine abolishes Btn2 SUMOylation. ....................... 87 Figure 3.11 Btn2 SUMOylation is lost in a Btn2-6KR construct. ................................................ 88 Figure 3.12 Induction, clearance, and stability of Btn2-6KR appears similar to Btn2-WT. ........ 89 Figure 3.13 Btn2 SUMOylation is still lost in Btn2-6KR when integrated into the genome. ...... 90 Figure 4.1 Loss of SUMOylation does not induce MMS sensitivity or growth defects. .............. 98 Figure 4.2 Btn2 localizes to the nucleus irrespective of its SUMOylation status......................... 99 Figure 4.3 Btn2 SUMOylation might regulate its degradation. .................................................. 100 Figure 4.4 SUMOylation regulates Btn2 solubility. ................................................................... 102 Figure 4.5 Computationally predicted soluble regions of Btn2. ................................................. 103 Figure 4.6 Elevated sequestration in Btn2-6KR cells. ................................................................ 105 Figure 4.7 Temperature sensitivity of fes1Δhsp104Δ cells is rescued by Btn2-6KR. ................ 106 Figure 4.8 Btn2-6KR is independent of Apj1-degradation and epistatic with Hsp104-refolding...................................................................................................................................................... 108 Figure 4.9 Rpd3 INQ measurements reflect Btn2 SUMO-Hsp104 epistasis.............................. 109 Figure 4.10 Clearance of cytoplasmic inclusions is unaffected by Btn2 SUMOylation. ........... 110 Figure 4.11 Btn2-6KR cells have an increased frequency of cells with >2 CytoQ foci. ............ 111 Figure 4.12 Btn2-SUMO does not affect luciferase refolding. ................................................... 112 Figure 4.13 Btn2 SUMOylation impacts ubiquitin and SUMOylation status of sequestration inclusions. ................................................................................................................................... 114 xvi Figure 4.14 Btn2 SUMOylation and turnover is controlled by Slx5/8. ...................................... 117 Figure 4.15 Slx5 and Apj1 regulate Btn2 SUMOylation levels. ................................................ 118 Figure 4.16 Slx5/8 recruitment to INQ is not dependent on Btn2 SUMOylation. ..................... 119 Figure 4.17 Slx5 is the E3 ligase responsible for depositing K48-Ub chains in the MMS-induced insoluble proteome. ..................................................................................................................... 120 Figure 4.18 INQ clearance is blocked in slx5Δ Btn2-6KR cells. ............................................... 122 Figure 5.1 Btn2 SUMOylation regulates INQ dynamics. ........................................................... 127 xvii List of abbreviations ACD Alpha crystallin domain aCLD Alpha crystallin like domain ADP Adenosine diphosphate ALS Amyotrophic lateral sclerosis ANOVA Analysis of variance ASF1 Anti-silencing function 1 ATM Ataxia-telangiectasia mutated protein ATP Adenosine triphosphate ATR Ataxia-telangiectasia and Rad3 related protein CAF Chromatin assembly factor 1 CCT Chaperonin containing CHX Cycloheximide CMA Chaperone mediated autophagy CMG Cdc45-MCM-GINS complex CMOS Complementary metal oxide semiconductor CP Core particle CPT Camptothecin CTD C terminal domain DAPI 4',6-diamidino-2-phenylindole DD Dimerization domain DDC DNA damage checkpoint DIC Differential interference contrast DNA Deoxyribonucleic acid DSB Double strand break DUB Deubiquitinase enzyme E1,2,3 Enzyme 1,2,3 EDTA Ethylenediaminetetraacetic acid EMS Ethyl methanesulfonate ESCRT Endosomal sorting complexes required for transport FACT Facilitates chromatin transcription FFL Firefly luciferase G/F Glycine/phenylalanine region G/M Glycine/methionine rich domain GFP Green fluorescent protein H2O2 Hydrogen peroxide HAT Histone acetyltransferase HDAC Histone deacetylase xviii HPD Histidine-Proline-Aspartic acid HR Homologous recombination Hsp Heat shock protein HU Hydroxyurea IDD Intrinsically disordered domain IDR Intrinsically disordered region INQ Intranuclear quality control IPOD Insoluble protein deposit JASSA Joint advanced SUMOylation site and SIM analyser JUNQ Juxtanuclear quality control K48 Lysine residue 48' K63 Lysine residue 63 KAT Lysine acetyltransferase KDAC Lysine deacetylase LIR LC3-interacting region LLPS Liquid-liquid phase separation LUBAC Linear ubiquitin chain assembly complex MCM2 Minichromosome maintenance 2 MD Middle domain MMS Methyl methanesulfonate NAC Nascent-chain associated complex NBD Nucleotide binding domain NEF Nucleotide exchange factor NER Nucleotide excision repair NHEJ Non-homologous end joining NLS Nuclear localization sequence NTA Nitrilotriacetic acid NTD N terminal domain NTP Nucleoside triphosphate NVJ Nucleolar vacuolar junction OD600 Optical density at 600nm PB1 Proteasome binding domain PCNA Proliferating cell nuclear antigen PD Pulldown PML Promyeloid leukemia PQC Protein quality control PrLD Prion like domain PTM Post-translational modification RAC Ribosome associated complex xix RNA Ribonucleic acid RP Regulatory particle RPA Replication protein A SARS Severe acute respiratory syndrome SBD Substrate binding domain SC Synthetic complete SDS Sodium dodecyl sulfate SEM Standard error or mean sHsp Small heat shock protein SIM SUMO interaction motif SNARE SNAP receptors snRNP Small nuclear ribonucleoproteins SUMO Small ubiquitin like modifier SWI/SNF Switching deficient/Sucrose-non-fermenting TCA Trichloroacetic acid Ub Ubiquitin UBD Ubiquitin binding domain UNT Untreated UPS Ubiquitin proteasome system UV Ultraviolet VCP Valosin-containing protein VHL Von Hippel-Lindau WT Wild type YPD Yeast peptone dextrose ZBD Zinc binding domain ZFLR Zinc finger like region xx Acknowledgements I would like to first express my heartfelt gratitude to my supervisor, Dr. Peter Stirling, for welcoming me into his lab and for his invaluable guidance and mentorship. Your support, encouragement, and excitement for science have helped me grow as a scientist and this work would not be possible without you. I would also like to thank my committee and their lab members - Drs. Elizabeth Conibear, Thibault Mayor and Phil Hieter for their expert insights and mentorship whenever I needed them. I am extremely grateful to the brilliant minds at work in the Stirling lab. I’d especially like to thank Alynn for her patience and support every single time I needed it. Sean, for our never-ending discussions on music and life. Katie, Leticia, Elizabeth, Hilary, and Cathy, for inspiring me with your drive to succeed. James, Justin and Louis for your friendship, camaraderie, and support in and outside the lab. I’ll forever cherish the game nights, lunch breaks and lab retreats we’ve all shared together. I would also like to thank all the lab alumni that have been instrumental in my growth, Karissa, Emily and Shuhe for their mentorship, and special thanks to Veena and Annie, for their constant support and friendship. I would also like to acknowledge the people in the TFL for all the conversations and discussions that have shaped my PhD experience. I’ve been fortunate to be surrounded by some amazing friends outside of work. To start, I would like to thank Ankush, Apte, Kushal, Kshitij, Patel, PD, Palash, and Gerry for their friendship. Keeping undergrad aside, our zoom calls every Saturday during lockdown is something that kept me going. To Shivangini, Srishti, Aditi, Kabbe, and Anirudh, thank you for your constant encouragement and for bearing with my not-so-often long calls. Special thanks to my Vancouver xxi family that made this city feel like home. To Johanna, Lydia, Mina and Dana, thank you for all the unconditional support every Friday night along with our favorite trash TV shows. To Himant, Rithika, Sai, and Maschanka, thank you for your company, for all the hikes and tennis sessions, for the movie and game nights, and for the concerts, which made my whole graduate experience a breeze. I would also like to express my gratitude to those whose names might not be mentioned, but whose friendship have been integral to my personal growth. Thank you all for your endless patience, kindness, and encouragement throughout this process. My journey would not be the same without you. Lastly, I would like to thank my family. To my parents, thank you for your unwavering love and encouragement through every step. I cannot fathom the sacrifices you have endured to put me where I am today, and for that I will be eternally grateful. To my brother and sister-in-law, Aravind and Prasanthi, thank you for being my second set of parents, for all your support, and for always looking out for me. I draw so much strength and courage knowing I have all of you by my side. Thank you for everything. xxii Dedication To the boy who dreamed. 1 Chapter 1: Introduction 1.1 Proteostasis, misfolding and aggregation Proteins are fundamental building blocks of life, essential for all processes occurring in a cell. They play constitutive roles in activating, executing, and maintaining integral pathways from DNA replication, cell differentiation, cargo trafficking, to even immune responses and oxygen transport. It is therefore in the interest of the cell to maintain protein homeostasis, or proteostasis, where the proteins are present in their respective folded conformations. Figure 1.1 The protein folding landscape. Schematic of a funnel-shaped free-energy landscape that proteins traverse as they sample different configurations during folding, increasing intramolecular contacts as they move towards a thermodynamically stable, native state. The figure was created using BioRender. 2 Proteins are synthesized as a linear chain of amino acids that must fold and in general adopt a three-dimensional structure before performing their cellular function. The process of folding has been described as a search across multiple conformations on a potential free energy landscape such that the properly folded state is one with minimum entropy and free energy. While it was thought that a functional protein always acquires a tertiary three-dimensional structure, recent advancements in determining protein structure have revealed the existence of intrinsically disordered regions (IDRs) (Oldfield & Dunker, 2014). These regions do not form rigid three-dimensional structures but instead exhibit flexibility and undergo conformational change often to perform different functions depending on the IDR conformation, protein-protein interactions, and the environment. Therefore, while a three-dimensional structure is not necessary, a proper conformation dictates protein function. As cells encounter different environmental insults, undergo ageing, or acquire mutations, proteins tend to misfold into non-native conformations. These misfolded proteins feature short stretches of hydrophobic regions which are typically buried inside their native conformation. These exposed regions are inherently “sticky” and if not acted on, could lead to accumulation of toxic protein aggregates. Aggregated misfolded proteins present two obvious problems. They could result in a gain-of-function phenotype by acting as a physical roadblock and interfering with protein trafficking (Chou et al., 2018; Woerner et al., 2016). Alternatively, they present a loss-of-function since their constituents can no longer function in their respective pathways (Winklhofer et al., 2008). This broad interference in cellular function is evident in 3 neurodegenerative diseases, cancer, and in aged cells (Ross & Poirier, 2004; Soto & Pritzkow, 2018). This chapter will discuss the concepts of protein folding, misfolding and the need for molecular chaperones in yeast. Additionally, I will introduce the three pillars of protein quality control: protein refolding, degradation, and sequestration with a stronger focus on the latter with respect to the nuclear sequestration site INQ. 1.1.1 Co-translational folding of nascent polypeptides Protein synthesis is carried out by the ribosomal machinery which consists of two major subunits: the small subunit that reads the mRNA template, and the large subunit that polymerizes the amino acids to create peptides. Since ribosomes control all protein production, mutations affecting protein synthesis rates, ribosome formation or elongation are associated with a heterogeneous class of diseases called ribosomopathies and also cancer (Klinge & Woolford, 2019). Translation occurs in a sequential manner. The mRNA template is fed to the ribosome and is read one codon at a time. Consequently, the peptide chain emerges in a vectorial fashion, starting at the N-terminus and extending as the ribosome moves along the mRNA. Synthesis is complete when the ribosome reads the stop codon, following which the peptide is released. Interestingly, this process of elongation is relatively slow with a rate of ~6 amino acids per second in eukaryotes, suggesting that synthesis of even small domains will require a couple of seconds (Kaiser & Liu, 2018). Protein folding on the other hand has been shown to be spontaneous and 4 occurs in the millisecond range. Therefore, it was posited and proven that ribosome bound-peptide folding can begin before synthesis is completed. One can imagine a situation where a translated N-terminal domain containing an alpha helix can take shape before the rest of the protein is translated. Folding, in turn, can be affected by elongation rates as some codons can be translated faster than the others. Additionally, ribosomal pausing due to mRNA structure (J.-D. Wen et al., 2008), codon choice (Mohammad et al., 2019), or polyA tracts (Koutmou et al., 2015), while deleterious, could aid in providing extra time for the nascent polypeptide chain to take shape. Before the nascent chain emerges from the ribosome, it must cross a long exit tunnel in the large subunit. The positioning of this tunnel introduces a regulatory step as only limited folding can occur within it, therefore limiting tertiary structure formation. However, because this process occurs co-translationally, the extruded section of the N-terminus can acquire premature non-native conformations and aggregate while the rest of the protein is being translated. This is prevented by the concerted action of molecular machines called chaperones that bind to nascent polypeptides co-translationally and provide them with the proper environment to conform to its functional structure. Aside from protecting nascent proteins, chaperones also protect the proteome from stress, thus forming the foundation of every protein quality control (PQC) pathway. 1.1.2 Molecular chaperones and the respective families Pioneering research from Anfinsen revealed that small, denatured proteins can spontaneously refold in vitro, highlighting that all the information required for a three-dimensional structure is 5 encoded in the amino acid sequence (Anfinsen, 1973). While spontaneous folding might be easily achieved for certain proteins, large proteins, complexes, or proteins present in crowded environments require assistance from chaperones to reach their native state. Molecular chaperones are defined as proteins that interact with and aid in the assembly or folding of another protein without being part of its final structure (Kim et al., 2013). They do so by binding exposed hydrophobic regions on nascent polypeptides or misfolded proteins and maintaining them in a soluble, folding-competent state. This prevents inappropriate intra- and inter-molecular interactions and promotes folding into the native state. While prokaryotes have only one ribosome-associated chaperone called the Trigger Factor, eukaryotes have two major complexes (Pechmann et al., 2013). First, the nascent-chain associated complex (NAC) which has been suggested to sense nascent polypeptides in the ribosomal tunnel, is a heterodimeric complex comprised of an α- and β- subunit both of which appear to have binding capabilities to both the ribosome and the emerging nascent chain (Beatrix et al., 2000; Gamerdinger et al., 2019; Kirstein-Miles et al., 2013). Second, the ribosome-associated complex (RAC) which consists of a tripartite system between the Hsp70-like protein Ssz1, the Hsp40 zuotin (Hsp70L1 and MPP11 in mammals), and Hsp70s Ssb1/Ssb2 (Otto et al., 2005; Zhang et al., 2017, 2020). Both complexes have been shown to compete for ribosome binding but also are functionally connected to each other, as simultaneous deletion of genes functioning in both pathways caused synthetic sickness. Remarkably, deleting both NAC and RAC in yeast resulted in decreased levels of ribosomal subunits and translating ribosomes indicating both complexes have a key role in determining the protein folding and production capacity of the cell (Koplin et al., 2010). 6 1.1.2.1 The Hsp40/Hsp70/Hsp110 families The Hsp70 family (the number represents molecular weights which forms the basis for classification of heat shock response (Hsp) proteins) of chaperones are one of the most versatile and ubiquitous chaperones, present in different cellular compartments. Hsp70s control a multitude of PQC pathways ranging from de novo protein folding, refolding, and disaggregation to crosstalk with other chaperone systems and proteolysis. Humans contain at least 13 HSP70 proteins (Radons, 2016) while yeast code for at least 11 – constitutive and cytosolic Ssa1 and Ssa2, heat-inducible Ssa3 and Ssa4, paralogs Ssb1 and Ssb2, ribosome-associated Ssz1, mitochondria-specific Ssc1, Ssq1, and Ecm10, and endoplasmic reticulum Kar2 (Lotz et al., 2019). Structurally, Hsp70s consist of two highly conserved domains, namely the N-terminal nucleotide bindin domain (NBD) and the substrate binding domain (SBD) that are joined by a flexible linker (Saibil, 2013). Additionally, they can act as monomers or form homodimers for efficient binding to its partners and subsequent peptide capture (Morgner et al., 2015; Nitika et al., 2022; Trcka et al., 2019). Hsp70s function in a tripartite system, with the help of two other co-chaperones namely, an Hsp40 and a nucleotide exchange factor (NEF; belonging to the Hsp110 family). Together, they bind short hydrophobic sequence motifs practically present almost all partially folded proteins, allowing them to bind substrates independent of size and function. With respect to ATP, Hsp70s are present in two forms: ATP-bound and ADP-bound states. The ATP-bound Hsp70 has a low substrate affinity and a faster exchange rate as opposed to the ADP-bound Hsp70 that has a higher substrate affinity and slower exchange rate. This has to do with 7 the position of the C-terminus which acts as a lid over the SBD in an ADP- bound Hsp70, thus trapping the substrate (Mayer & Gierasch, 2019). As discussed in detail later, Hsp40s bind to substrates and deliver them to an ATP-bound Hsp70, simultaneously stimulating their ATPase activity. ATP hydrolysis results in a structural change, trapping the client in an ADP-bound Hsp70 allowing them to fold. Consequently, NEFs promote the release of ADP and re-binding of ATP to facilitate substrate release and start a new cycle of substrate binding (Mayer & Gierasch, 2019). This repetitive process of cycles of substrate capture, and release form the basis of the Hsp40/Hsp70/NEF system which is represented in Figure 1.2. 8 Figure 1.2 The Hsp70 chaperone cycle. Nascent polypeptides are handed over to Hsp70s as they exit the ribosome, where they undergo multiple rounds of association and dissociation during chain elongation. Consequently, they can either fold spontaneously or bind to Hsp70s through Hsp40s, which ultimately stimulate their ATPase activity to fold peptides into a proper conformation. Hsp70s switch between high- and low-affinity states as they bind and hydrolyze ATP (Grey box). Once bound to a substrate, the C-terminal lid closes on the substrate binding domain, trapping the polypeptide, allowing it to fold into its native state. Upon the action of an NEF, Hsp70s release their substrate by binding to ATP and achieving a low-affinity ATP bound state. When encountering nascent polypeptides or misfolded proteins, Hsp70s can also enlist the help of other chaperones such as Hsp90s or the TRiC/CCT system for further folding or target misfolded proteins for degradation via the proteasome. Adapted and modified from (Rosenzweig et al., 2019) using BioRender. As mentioned above, Hsp40s bind and transfer substrate proteins to Hsp70s for folding. In doing so, Hsp40s control, direct and diversify Hsp70 substrate specificity thus acting as important regulators of protein folding. In line with that, the number of Hsp40 proteins far exceed that of Hsp70s. For instance, humans code for at least 49 Hsp40s while yeast code for 22 proteins (Kampinga & Craig, 2010; Piette et al., 2021). While some Hsp40s might act on the same substrate illustrating redundancy, most of them act on different groups of proteins and under different stress conditions. They can be further classified into three different classes: I, II, and III (Q. Liu et al., 2020). The classification is based on the position of their characteristic J domain, which stimulates the ATPase activity of Hsp70s. Class I Hsp40s generally have five domains namely the J domain, a glycine/phenylalanine rich domain, two carboxyl-terminal domains 9 CTD1 and CTD2, and a dimerization domain. Class II Hsp40s have a similar domain organization except that they do not contain a zinc finger-like region that is found in the Class I CTD1. Unlike Class I and II, the J domain in Class III Hsp40s is not restricted to the N-terminus and can be in any location (Q. Liu et al., 2020). While the role of domains like the zinc finger-like domain are still unclear, structural data has revealed that Hsp40s dimerize using their dimerization domain and capture substrates via their CTD domains. Once substrate-bound, they interact with their respective Hsp70 to perform distinct cellular functions. Most importantly, they stimulate the ATPase to Hsp70s through their conserved HIS-PRO-ASP (HPD) motif in J domain. Interestingly, diverse Hsp70-independent roles have been documented for Hsp40s by acting in concert with other proteins to promote motility of flagella, viral replication, pre-mRNA splicing and spliceosome disassembly (Ajit Tamadaddi & Sahi, 2016). 1.1.2.2 The Hsp90 family Downstream to the Hsp70/40 system lie the next class of chaperones that belong to the Hsp90/100 families. Similar to Hsp70 and Hsp40, dimerization is essential for Hsp90 function, and this is carried out through its CTD (Wayne & Bolon, 2007). Additionally, substrate transfer from Hsp70s to Hsp90s is bridged through the conserved co-chaperone HOP/Sti1 that also binds to the MEEVD motif in the CTD of the Hsp90 (Johnson et al., 1998; Wegele et al., 2006). Once bound, Hsp90s also utilize ATP-hydrolysis to control protein folding of its substrates by interacting with other co-chaperones like Aha1 or Sgt1 (Schopf et al., 2017). Despite similar roles, Hsp90 differs heavily from Hsp70 in that it specifically controls protein folding for a subset of the proteome while Hsp70s are constitutively binding to all proteins. Hsp90 facilitates assembly of multiprotein complexes such as the kinetochore, kinases, receptors (Boczek et al., 10 2015; Lorenz et al., 2014; Schopf et al., 2017) but also promotes ligand-protein interactions such as siRNA loading on the Argonaute 2 complex, and association of telomerase with DNA (DeZwaan et al., 2009; Iwasaki et al., 2015). Therefore, while Hsp90 is a key regulator of protein folding and proteostasis, it also has a profound effect on signalling cascades and networks. 1.1.2.3 Hsp104/AAA+ ATPases On the other hand, Hsp100s such as Hsp104 in yeast and ClpB in bacteria have the distinctive ability to resolubilize and refold aggregated proteins (Seraphim & Houry, 2020). Based on its structure and mode of action, Hsp104 also belongs to the AAA+ (ATPases Associated with diverse cellular Activities) protein superfamily, a diverse class of conserved enzymes that also utilize energy derived from ATP-dependent reactions to govern multiple processes including protein quality control. For example, Hsp104 can form hexamers and using an ATP-dependent threading mechanism, dissolve and resolubilize protein aggregates. In conjunction with the Hsp70 machinery, this forms the basis for refolding of non-native misfolded proteins (discussed in detail later in Section 1.2.1 titled “Disaggregation and protein refolding”). Similarly, another AAA+ protein called Cdc48 functions as a chaperone by recognizing and extracting ubiquitinated proteins from complexes or membranes and processes them for subsequent degradation and processing (Baek et al., 2013). Since other AAA+ proteins such as the initiator PCNA or helicase MCM are essential for DNA replication, these proteins are pivotal for maintaining homeostasis (Seraphim & Houry, 2020). 11 1.1.2.4 Chaperonins Chaperonins/Hsp60s form an essential class of chaperones ubiquitously conserved across different kinds of life. They are ATP-driven megastructures that function through oligomerization as opposed to most chaperones mentioned till now that function in dimers. Chaperonins can be classified into two classes: Class I which are found in prokaryotes, chloroplasts and mitochondria and Class II which are archaeal or cytosolic eukaryotic proteins (Horovitz et al., 2022). Both are well studied in that research on GroEL (Class I) and TRiC/CCT (Class II) have helped us develop an understanding of how this class of chaperones function. TRiC/CCT, for example, is made up of 8 unique subunits that bind each other in a stepwise manner to form a ring-like structure. Once formed, two of these rings are stacked on each other to form a barrel-shaped 16-subunit CCT complex that has open ends on each side with a cavity in the middle (Kalisman et al., 2012; Leitner et al., 2012). Chaperonins also work in concert with other co-chaperones. For CCT, Prefoldin is one such co-chaperone that helps bind nascent-polypeptides and delivers it to CCT for proper folding. Together, they appear to have a major role in facilitating the folding of cytoskeletal proteins such as actin, tubulin, actin-related proteins, cofilin and more that have essential roles in mitosis and cell polarity (Balchin et al., 2018; X. Chen et al., 1994; Horovitz et al., 2022; Melki et al., 1993). 1.1.2.5 Small heat shock proteins (sHsps) Unlike other chaperone families mentioned till now, sHsps are simpler ATP-independent chaperones that prevent irreversible aggregation of misfolded proteins by acting as “holdases”. Since they do not possess the ability to hydrolyze ATP, sHsps cannot affect protein folding, rather they bind hydrophobic stretches on non-native proteins and either maintain the sHsp-12 protein complex to avoid aggregation or sequester these misfolded proteins into specialized compartments where they can be acted on by other chaperones. They contain a characteristic alpha crystallin domain (ACD) which is flanked by two disordered regions, namely the NTD and the CTD (Haslbeck et al., 2019). While the length and functions of the NTD and CTD differ between sHsps, they bind substrates using their ACD. Like chaperonins, sHsps have also been found to oligomerize to carry out their function. Canonically, the IX(I/V) motif in the CTD plays a prominent role in oligomerization (Haslbeck et al., 2019). Proteins bound to sHsps are eventually refolded or degraded with assistance from other PQC machineries, however substrate transfer and choice are mechanisms that are yet to be discovered. The role of sHsps in sequestration will be discussed in Section 1.2.3.1. 1.2 The three pillars of protein quality control Irreversible protein aggregation has formed the hallmark of a group of diseases called proteinopathies which include neurodegenerative diseases like Huntington’s, Parkinsons, Amyloidosis and more. Misfolded proteins, therefore, are constantly recognized, sequestered, refolded, or degraded to prevent accumulation of aggregates. Moreover, as we age, the burden of misfolded proteins increases thus imposing a strain of chaperones to maintain proteostasis. In such cases, the cell must employ pathways specifically tasked with the role of clearing protein aggregates. Refolding, sequestration, and degradation represent the three fundamental triage decisions for non-native proteins and collectively define the idea of a PQC circuit. 13 Figure 1.3 The triage decisions carried out by the proteostasis network. Proteins in different conformational states are under the constant action of chaperones that strive to achieve proteostasis by refolding, degrading, or sequestering misfolded proteins while aiding in the formation of properly folded proteins. Adapted and modified from (Hipp et al., 2019) using BioRender. 14 1.2.1 Disaggregation and protein refolding An aggregate poses an interesting problem to chaperones. When nascent proteins are bound to chaperones, they are also maintained in a soluble state. Misfolding and consequent aggregation shift the proteins to an insoluble state, from which they must be resolubilized and disaggregated in order to be processed for refolding or degradation. The importance of this process became evident in yeast, where cells lacking the Hsp104 disaggregase could not recover or tolerate a severe heat shock, a stress condition which induces large levels of misfolding and aggregation (Sanchez & Lindquist, 1990). Our understanding of disaggregation is derived largely from studies on ClpB (bacterial disaggregase) and Hsp104 (Haslberger et al., 2008; Lum et al., 2004; Wendler et al., 2009). Metazoans, interestingly, have lost the ability to code for disaggregases at some point in evolution. However, the Hsp40/70/110 system appears to compensate for this loss by having disaggregase activities of their own (Gao et al., 2015; Kirstein et al., 2017; Nillegoda et al., 2015; Rampelt et al., 2012). Hsp104 and ClpB belong to the AAA+ ATPase and Hsp100 class of chaperones, that rely on ATP hydrolysis to disaggregate large complexes and refold substrates by threading them through their inner core. Structurally, both disaggregases contain an NTD, two distinct yet conserved nucleotide-binding domains NBD1/2 that are essential for ATP hydrolysis, and a middle domain (MD) that regulates ATPase activity and mediates Hsp70 interaction (Mogk et al., 2018; Yokom et al., 2016). Once assembled into hexamers, energy derived from ATP hydrolysis is suggested to transform into a pulling force through the inner core, ultimately threading large assemblies and misfolded proteins through a polypeptide loop extrusion mechanism (Avellaneda et al., 2020). 15 Studies in yeast revealed that Hsp104 on its own cannot promote refolding of firefly luciferase (Glover & Lindquist, 1998). Rather, it requires cooperation with the Hsp40/70 system for substrate transfer and stimulation of ATPase activity. The Hsp70/Hsp104 bipartite system works together to coat aggregates with Hsp70s that physically recruit Hsp104/ClpB molecules for disaggregation. Since Hsp70s bind to almost all polypeptides with the help of their Hsp40 co-chaperones, one could image stress induced Hsp40s regulate binding of Hsp70s specifically to aggregates thus regulating free Hsp70 levels under stress. Aside from delivering substrates, Hsp70s also stimulate the ATPase activity of Hsp104/ClpB by binding to its MD domain, and shifting the Hsp104/ClpB conformation to an active state (Mogk et al., 2018; Rosenzweig et al., 2013; Seyffer et al., 2012). Remarkably, this physical interaction is precisely regulated to avoid misdirection and binding of Hsp104 to nascent polypeptides. Firstly, interaction between the MD domain on a Hsp104/ClpB and an ATP-bound Hsp70 is prevented due to steric hindrance, thus ensuring interaction only with a substrate-bound Hsp70 (Hayashi et al., 2017). Secondly, two or more Hsp70 molecules are required to establish a physical interaction with Hsp104, which could be achieved in a situation where multiple Hsp70s are in close proximity to each other, bound to an aggregate (Oguchi et al., 2012). Ultimately, the solubilized unfolded protein that emerges from an Hsp70-Hsp104 complex can either undergo spontaneous refolding or bind to chaperones like Hsp70s or Hsp90s in the vicinity. Interestingly, refolding appears to be preferred upon stress recovery, suggesting that it is energetically favorable to refold important proteins instead of making them again. However, resolubilized proteins from aggregates can also undergo degradation via the proteasome or through autophagy. 16 1.2.2 Protein degradation The rules and guidelines followed by PQC pathways that decide between refolding or degradation of a misfolded substrate remains elusive. From a thermodynamic perspective, refolding would be prioritized over degradation since both protein production and degradation are energetically expensive. However, under conditions of stress, cells need to respond quickly by regulating splicing and suppressing production of ribosomes (Gasch et al., 2000; Mathew et al., 2017). Additionally, this evokes a stress response that includes expression of stress response genes such as chaperones and therefore, degradation of terminally misfolded proteins could aid in supplying the building blocks for other proteins. Degradation is primarily carried out by two major pathways: the ubiquitin-proteasome system (UPS) or through autophagy (Rousseau & Bertolotti, 2018). While autophagy helps degrade proteins, large complexes, and insoluble species especially under stress, genes involved in this process are not essential in normal conditions. On the other hand, most of the UPS genes are essential indicating an important and constitutive role for the UPS in degrading proteins. Indeed, inhibiting the proteasome results in cell death through dysfunction of amino acid homeostasis suggesting its involvement as the main pathway in turnover of proteins in basal conditions (Suraweera et al., 2012). 1.2.2.1 The Ubiquitin-Proteasome System The eukaryotic 26S proteasome is a conserved 2.5MDa multisubunit protease that regulates the degradation of ~80% of the cellular proteome from short lived and regulatory proteins to terminally misfolded proteins thus controlling a variety of processes. It is comprised of the 20S 17 core particle (CP) flanked by one or two 19S regulatory particle (RP) both of which are made up of multiple subunits themselves (Rousseau & Bertolotti, 2018). In brief, the RP is comprised of multiple AAA+ ATPases responsible for substrate recognition, unfolding and translocation into the CP core which contains proteolytic peptidases for degradation. Due to its complexity and the sheer number of subunits required, production and assembly of a functional proteasome is energetically costly. As a result, expression of proteasomal subunits must be tightly regulated, to ensure efficient and stoichiometric production which is in part achieved by having a common transcription factor like Rpn4. Remarkably, Rpn4 has a very short half-life and is degraded by the proteasome (Xie & Varshavsky, 2001). Consequently, Rpn4 abundance increases when the proteasome is compromised, thus increasing proteasome subunit production. 1.2.2.2 Ubiquitin An assembled and functional proteasome primarily recognizes its substrates via the post-translational modification (PTM), ubiquitin. Ubiquitin is a highly conserved 76-amino acid polypeptide that is covalently attached to a protein via Lys-linked isopeptide bonds or Cys/Ser/Thr-linked esterification in a three-step manner: First, ubiquitin activating E1 enzymes catalyze the adenylation of the terminal glycine in ubiquitin resulting in a high energy thioester bond between the two (Finley et al., 2012; Pohl & Dikic, 2019). Next, the activated ubiquitin is conjugated onto the amide group on E2 conjugating enzymes by transesterification and finally, is transferred onto substrates via a third group of proteins called E3 ubiquitin ligases. The number of E3 ligases (>60 in yeast, >500 in humans) outnumber E2 conjugases (11 in yeast, >50 in humans) by a margin across species indicating that E3 ligases control substrate specificity (Finley et al., 2012; Pohl & Dikic, 2019). Additionally, E3 ligases share many substrates 18 indicating redundancy in case other parallel pathways are disrupted. Importantly, some E2 enzymes can modify substrates on their own, circumventing the need for an E3 ligase, thus adding another layer of regulation (Hoeller et al., 2007). Once modified, a ubiquitinated protein can be acted on by proteases called deubiquitinases (DUBs) that cleave the ubiquitin moiety and recycle it for further rounds of modification. Ubiquitination of a protein has been shown to regulate multiple facets of a protein’s function including activation, localization, interactions, stability, and degradation (Pohl & Dikic, 2019). This level of versatility is achieved due to its intrinsic ability to be ubiquitinated itself, therefore forming chains of varying lengths and topologies. The ubiquitin moiety contains 7 lysines that can be modified – K6, K11, K27, K29, K33, K48 and K63 – either simultaneously, giving rise to a multi-monoubiquitinated substrate, or to form chains on a certain lysine residue. These chains can either be homotypic (same linkage) or heterotypic (different linkages; branched vs non-branched) thus further expanding the ubiquitin code and its complexity (Swatek & Komander, 2016). K48-, K29- and K11- chains act as strong signals for proteasomal degradation, whereas K63- chains are involved in many proteasome-independent pathways including inflammatory signalling, endocytosis, and the DNA damage response. However, K63- chains have also been shown to regulate proteasomal degradation by acting as a seed to form a branched K48-/K63- heterotypic ubiquitin chain (Ohtake et al., 2018). The remaining linkages have been less studied comparatively, which could be in part due to their low abundance (P. Xu et al., 2009). In addition to lysine linkages, linear methionine (M)1-linked ubiquitin chains can also be assembled by a 19 single E3, linear ubiquitin chain assembly complex (LUBAC) and have established roles in tumor necrosis factor signalling, along with K63-Ub chains. Substrates can also be modified at multiple lysines simultaneously across the protein, resulting in a multimonoubiquitinated substrate, that can act as an underappreciated degradation signal (Dimova et al., 2012). Processing of ubiquitinated substrates by the proteasome or DUBs requires cleavage of the polyUb chain, which results in the formation of unanchored polyUb chains and free ubiquitin that is necessary to maintain Ub homeostasis. Initially thought to be toxic and detrimental, there is a growing appreciation for unanchored Ub chains as key regulators for degradation and signalling (Blount et al., 2020; Hao et al., 2013; Xia et al., 2009). Moreover, advances in proteomics, gene editing and spectroscopy have expanded our knowledge of non-Lys ubiquitination. Remarkably, the bacterial pathogen Legionella pneumophila was found to employ its own E1 and E2 independent mechanism for ubiquitination to Arg residues instead of Lys (Qiu et al., 2016). Also, in line with viruses disrupting host Ub homeostasis to increase pathogenicity, the SARS coronavirus was recently shown to express a papain-like protease that specifically cleaves K48-Ub linkages, thus exhibiting DUB activity (Békés et al., 2016). Overall, ubiquitin is a central player in regulating not only protein degradation but various other processes in the cell. The ubiquitin code represents a complex set of pathways, with multiple layers of regulation. With new tailored E3 linkages, improved proteomics, and editing tools, our understanding of these pathways will only grow over time (Wegmann et al., 2022). 20 1.2.2.3 SUMOylation SUMO, or small ubiquitin-like modifier, is a PTM that is part of a class of proteins called Ubiquitin-like modifiers alongside NEDD8, ISG15 and more that are structurally similar to Ub and are covalently added onto substrates but differ in sequence and charge. SUMO shares remarkable similarity to ubiquitin in that it also requires an E1-E2-E3 cascade for a substrate to be SUMOylated (Vertegaal, 2022). Moreover, canonical SUMOylation occurs on a lysine, and like ubiquitin, SUMO can be modified itself, allowing linkage or chain formation to produce polySUMO chains. While they share similarities, they do have some obvious differences. Unlike ubiquitin, SUMO needs to be activated in order to be conjugated onto substrates, a process which is carried out by SUMO proteases. Also, SUMOylation is regulated by a single E2 enzyme, as compared to ubiquitin which has 11 E2s in yeast and more than 50 in humans. Finally, while ubiquitin sequence is strongly conserved from yeast to humans, SUMO has five different isoforms in humans (SUMO1-5), eight in plants while only Smt3 exists in yeast (Vertegaal, 2022). Despite these differences, SUMOylation is also essential for cellular maintenance as evidenced by its roles in gene expression, DNA damage response, proteostasis, and cell cycle progression (Han et al., 2018). Alongside degradation, SUMOylation exerts a range of functions from controlling protein stability to localization (Psakhye et al., 2019; Sarangi & Zhao, 2015; D. Wen et al., 2017). Since SUMOylation also occurs at lysines, it antagonistically competes with ubiquitination and decides on the fate of a large list of substrates. In yeast, SUMO/Smt3 contains three lysines (K11,15, and 19) in its NTD that can be further modified for polySUMO chain formation. Whether SUMO chains form homo- and heterotypic chains similar to ubiquitin 21 remains to be explored. However, SUMO chains have a well-documented role in protein degradation. SUMO-targeting Ubiquitin Ligases (StUbLs) are a class of E3 ligases that ubiquitinate polySUMOylated proteins for degradation. These proteins contain multiple SUMO-interaction motifs (SIMs) in tandem that bind to a polySUMOylated substrate and using the RING E3 ligase domain, catalyze the ubiquitination of their substrate (Jansen & Vertegaal, 2021). Yeast (Slx5/8, Uls1, Rad18) and humans (RNF4, RNF111) both code for StUbLs. They are essential in maintaining genome integrity as they are involved in timely degradation of cell cycle progression and DNA replication proteins (Psakhye et al., 2019; Talhaoui et al., 2018; Thu et al., 2016). Additionally, they are critical for SUMO homeostasis since they regulate the degradation of the SUMO conjugation machinery itself (R. Kumar et al., 2017). Degradation-independent roles have also been noted, for example, RNF111 works in concert with E2s Ubc13-Mms2 to K63-polyubiquitinate SUMOylated DNA damage response protein xeroderma pigmentosum C (XPC) amongst others to promote efficient nucleotide exchange repair (NER) upon exposure to ultraviolet (UV) radiation (Poulsen et al., 2013). The addition of SUMO and Ub, together or separately, also affects protein solubility. This is an important feature to note since misfolded proteins can be refolded or degraded only when they are present in a soluble-competent state. Essentially, while these PTMs modulate the fate of a protein, polymeric chain formation of either can maintain a soluble state that could be worked on by chaperones. Even though we know these PTMs impart solubility, their roles, and the extent to which they affect protein, refolding, degradation, and sequestration remains to be tested. 22 1.2.2.4 Autophagy Contrary to the UPS that deals with spontaneous, small misfolded proteins, autophagy is preferred for larger aggregates, damaged organelles and lipid droplets which would otherwise over burden or exceed the capacity of the proteasome. Through this process, substrates are engulfed by the autophagic machinery and delivered to the lysosomes for degradation. Overall, autophagy can be classified into three different classes based on the delivery method and the cargo constituents themselves (Dikic, 2017). First, is the most well studied and overarching pathway called macroautophagy that utilizes a double membraned organelle called the autophagosome to sequester cytoplasmic aggregates and organelles and delivers them the lysosomes for bulk degradation. Second, is microautophagy where proteins are directly engulfed or invaginated into lysosomes. And third, chaperone-mediated autophagy (CMA) (only in mammals) involves the selective recruitment of misfolded cytoplasmic proteins featuring a KFERQ-like motif through via Hsp70s HSPA8/HSC70 which recognize, unfold, and transfer substrates to lysosomes for degradation (Dikic, 2017). The different classes of autophagy can be further broken into two classes: bulk vs selective. While bulk autophagy degrades substrates indiscriminately, selective autophagy is specialized degradation of cargo mediated by receptors. The importance of selective autophagy spans all organelles as most have their own specialized autophagic clearance mechanism, such as nucleophagy (nucleus), proteophagy/aggrephagy (proteasomes/aggregates), mitophagy (mitochondria) and many more (Zaffagnini & Martens, 2016). These processes are essential in maintaining organelle-homeostasis by aiding in degradation of harmful species, controlling amino acid flux and recycling of nutrients under stress such as nitrogen starvation or DNA 23 damage. Interestingly, nucleophagy also falls under micronucleophagy as it involves invagination of a portion of the nucleus at the nucleolar-vacuolar junction (NVJ) directly into lytic vacuoles. This process is governed by two proteins NVJ1 and VAC8 that mark the NVJ site in the nucleus and is involved in the clearance of nuclear envelope components, nuclear lamina, chromatin and most recently, misfolded proteins (Papandreou & Tavernarakis, 2019; Sontag et al., 2023). Despite their mechanistic differences, there is strong evidence of crosstalk between the UPS and autophagy pathways. This is in part due to ubiquitin, as both pathways use ubiquitin as a degradation signal. Additionally, Hsp70s bound to a ubiquitinated misfolded protein could either deliver the substrate to the proteasome or the CMA machinery, thus adding another layer of regulation. Similarly, Ub-receptor proteins like p62/SQSTM1 contain a ubiquitin-binding domain (UBD), a proteasome-binding domain (PB1), and a domain to bind autophagic membranes (LIR domain), and can thus target substrates to both the autophagic machinery and the proteasome (Dikic, 2017). Overall, maintaining chaperone and ubiquitin concentration appears to be crucial for protein degradation. Consequently, blocking UPS via proteasome inhibition increases autophagic capacity (Ding et al., 2004; Pandey et al., 2007; D. Wang et al., 2019). On the other hand, the compensation for autophagic inhibition by the UPS remains unclear. While there are reports of increased proteasomal capacity via the upregulation of proteasomal, inhibition of autophagy also appears to inhibit the UPS (Dikic, 2017; X. J. Wang et al., 2013). The latter could be explained by proteophagy wherein proteasomes themselves can be ubiquitinated and degraded via autophagy (Besche et al., 2014; Cohen-Kaplan et al., 2016). Ubiquitinated proteasomes could compete with functional proteasomes and affect the proteomic 24 flux. Therefore, while UPS and autophagy are distinct mechanisms, they appear to work together in a tightly regulated manner and control the degradation of unwanted proteins in a cell to achieve proteostasis. 1.2.3 Sequestration When PQC homeostasis is first overwhelmed during stress, cells acutely sequester and organize non-native proteins into specialized compartments/aggregates/inclusions. This pillar of PQC is tightly regulated by chaperones that bind to misfolded proteins and sequester them at specific sites in the cell to facilitate refolding or degradation. This process of compartmentalization has several proposed advantages. Spatially organizing misfolded proteins not only reduces their cytotoxicity but also reduces the burden on other PQC pathways (Ungelenk et al., 2016; Wolfe et al., 2013). This in turn prevents PQC collapse and promotes stress recovery (Escusa-Toret et al., 2013; Ho et al., 2019). Furthermore, recruitment of degradation and refolding machineries to these sites could facilitate the clearance of misfolded proteins (explained later in Section 1.2.3.1.2). Additionally, sequestration causes asymmetric inheritance of damaged proteins, resulting in inclusion-free daughter cells but contributing to replicative aging in mother cells (Hill et al., 2016; Zhou et al., 2014). In yeast, various PQC deposition sites have been described including three major inclusions: the insoluble protein deposit (IPOD), the juxtanuclear quality control site (JUNQ), and the intranuclear quality control site (INQ) (Kumar et al., 2022). While the IPOD, JUNQ reside in the cytoplasm, the INQ is present inside the nucleus and adjacent to the nucleolus. Like JUNQ, the INQ is highly dynamic and contains the proteasome, disaggregation and degradation 25 machineries, and is formed upon proteasome inhibition (Miller et al., 2015; Tkach et al., 2012). Since both the INQ and the JUNQ were discovered using the same model substrates, there has been an ongoing discussion about whether they represent the same or independent deposition sites. However, it was recently confirmed that the INQ and JUNQ are separated by the nuclear envelope (Sontag et al., 2023). Alongside this distinction, other differences have also been noted. Genotoxic stress induced via the alkylating agent methyl methanesulfonate (MMS) or oxidative damaging agent hydrogen peroxide induces the relocalization of several endogenous proteins to the INQ, whereas robust JUNQ formation is not seen (Mathew et al., 2017). This DNA damage–dependent relocalization to the INQ is not affected by actin or microtubule depolymerizing drugs like latrunculin B or nocodazole, suggesting that an intact cytoskeleton might not be a prerequisite for INQ deposition (Gallina et al., 2015). Figure 1.4 Major protein quality control sites in yeast. Shown are nuclear or cytoplasmic misfolded proteins during stress responses (lightning bolts) being recruited to the INQ, JUNQ, or IPOD through the action of sequestrases Hsp42 or Btn2. 26 The localization of proteasomes to the INQ and the JUNQ are highlighted. IPOD, insoluble protein deposit; JUNQ, juxtanuclear quality control site. Adapted from (Kumar et al., 2022). 1.2.3.1 Intranuclear Quality Control: the nuclear protein sequestration structure Similar to other PQC sites, the INQ was first discovered using aggregation prone reporter proteins like the small ubiquitin-like modifier (SUMO)–conjugating E2 enzyme temperature-sensitive (ts) allele Ubc9-ts-GFP and von Hippen Lindau protein (VHL) and was shown to reside inside the nucleus using immunoelectron microscopy (Miller et al., 2015). However, recent studies have found that many endogenous proteins are sequestered to the INQ upon genotoxic stress (Table 1). Its formation can be induced by various DNA damaging agents such as MMS, ethyl methanesulfonate, hydroxyurea, hydrogen peroxide, UV radiation, and even upon heat shock coupled with proteasome inhibition (Mathew et al., 2017; Saugar et al., 2017). Various efforts have gone into cataloging the proteins that relocalize to the INQ using mass spectrometric approaches. To date, 37 proteins have been found to relocalize to the INQ. These proteins, or the INQ substrates, perform a diverse set of functions such as chromatin remodeling and transcription (Rpd3, Hos2, Cmr1), RNA splicing (Hsh155, Cdc40), or replication fork structure cleavage (Mus81-Mms4 and Slx1-Slx4). The INQ also harbors various chaperones and degradation machinery, including the AAA ATPase Cdc48, the disaggregase Hsp104, the sHsps Hsp42 and Btn2, the Hsp40 J-domain protein Apj1, the Slx5/8 ubiquitin ligase, and the proteasome, that can together decide the fate of the INQ substrates (Table 1). 27 Proteins Function References Rpd3, Hos2, Arp6 Chromatin remodelling (Gallina et al., 2015) Mrc1, Cmr1*, Mus81, Mms4, Slx1, Slx4, Rad1 Genome integrity (Gallina et al., 2015; Saugar et al., 2017) Apc4*, Cdc20*, Cdc27 Anaphase-promoting complex (Gallina et al., 2015) Hsh155, Cdc40* Splicing (Gallina et al., 2015; Mathew et al., 2017) Pph3, Pph22, Fig4 Phosphatases (Gallina et al., 2015) Btn2, Apj1, Cdc48, Sis1, Hsp104, Slx8, Rpn11, Cct6 Chaperones and degradation machinery (Gallina et al., 2015; Ho et al., 2019; Mathew et al., 2020) Gln1, Dug2* Glutamine metabolism (Gallina et al., 2015) Mkt1, Lst8, Emg1, Ylr126c, Rbd2, Dus3, Pal2, Qcr6, Mdh2 Other (Gallina et al., 2015) *Proteins containing one or more WD-40 repeats Table 1.1 The INQ Proteome. INQ formation is governed by two sequestrases, Btn2 and Hsp42, that increase in abundance upon DNA damage in an Hsf1-dependent manner (Solís et al., 2016). Sequestrases, usually sHsps, are ATP-independent chaperones that bind to the exposed hydrophobic regions of misfolded proteins and sequester them to specific deposition sites. As explained earlier, once 28 bound to an sHsp, a misfolded protein is protected from aggregation but does not actively refold. Hence, sHsps are also termed holdases or aggregases as they hold misfolded proteins and are acted upon by other ATP-dependent chaperones to refold or degrade the misfolded protein. Each sHsp contains a characteristic alpha crystallin domain that is made up of antiparallel beta strands that form a beta sandwich (Haslbeck et al., 2019). The alpha crystallin domain is flanked by two disordered regions, namely the N-terminal domain (NTD) and the C-terminal domain (CTD). Functional diversity amongst sHsps is determined by the NTD and CTD as differing length and amino acid makeup of the NTD and CTD can control substrate specificity (Specht et al., 2011). The NTD of Hsp42 is vital for its aggregate sorting function as Hsp42 mutants deleted for its NTD do not restore formation of VHL aggregates upon heat shock and proteasomal impairment using MG132. Interestingly, swapping the Hsp42-NTD with Hsp26 to create a Hsp42NTD-Hsp26 chimera could efficiently sort VHL into aggregates, indicating that the NTD of Hsp42 determines its substrate specificity. This NTD was further dissected to reveal a prion-like domain that is crucial for cytoplasmic sequestration and acts as the major substrate-binding site (Grousl et al., 2018). While these studies focused on Hsp42 domains and cytoplasmic sequestration, we can assume the same regions could be involved in the INQ sorting. In comparison, Btn2 is a v-SNARE-binding protein with standalone sequestrase activity and sHsp-like features (Ho et al., 2019; Kama et al., 2007). It mediates late endosome-golgi recycling but has an established role as a crucial sequestrase for deposition of misfolded proteins at the INQ. Dissection of Btn2 domains revealed similarity with sHsps in that Btn2 contains an alpha-crystallin–like domain (aCLD) flanked by a disordered NTD and CTD. Unlike Hsp42, Btn2’s NTD is not involved in the INQ 29 formation but rather acts as a recruitment domain for disaggregases that aid in the INQ solubilization. Instead, both the aCLD and CTD are essential for the INQ formation with the aCLD acting as the substrate-binding site showcasing the flexibility between various sequestrase domains in carrying out their activity (Ho et al., 2019). 1.2.3.1.1 The INQ proteome Why do only certain proteins get sequestered to the INQ? Of the 37 endogenous proteins so far found at the INQ, there do not appear to be any major functional enrichments. With respect to common possible motifs, only six INQ proteins: Cmr1, Cdc20, Apc4, Lst8, and Dug2, contain at least one WD40 repeat. WD40 repeat domains are β-sheet enriched structural motifs that act as protein scaffolds and aid in protein–protein interactions (Xu & Min, 2011). It has been shown that Cmr1’s WD40 domain is critical for its localization to the INQ. Non-native WD repeat proteins could explain why the chaperonin CCT, which is required for WD protein biogenesis, is also found at the INQ. Therefore, while sequences within the WD40 repeats could be involved in chaperone recognition, there does not appear to be a single domain or motif shared amongst all the INQ substrates. On average, the INQ substrates are long, stable, hydrophobic, enriched for charged amino acids, abundant, and soluble as the nuclear proteome, indicating the absence of a unique feature (Kumar et al., 2022). While some of the INQ substrate proteins are likely misfolded and therefore sequestered, recent studies have also addressed the idea that sequestration to the INQ has specific regulatory roles during stress. To cope with DNA damage, a transcriptional response represses ribosomal gene production and activates chaperones. This ensures reduced and controlled production of new 30 proteins that help with stress recovery and preserving amino acid and energy availability. For example, the U2 spliceosome protein Hsh155 disassociates from its partners upon DNA damage and only Hsh155 is sequestered to the INQ (Mathew et al., 2017). Since ribosomal transcripts are the major source of splicing flux in yeast, it is proposed that Hsh155 sequestration and the associated splicing defect, served an additional posttranscriptional role in slowing ribosome production during stress. Consistent with this, in BTN2-deleted cells Hsh155 is not sequestered to the INQ and splicing is restored even during DNA damage. In another study focusing on the structure-specific endonucleases Mus81-Mms4 which relocalize to the INQ as a complex, the authors found that Mus81 retention in the INQ is determined by the phosphorylation state of Mms4 and that, when phosphorylated, the Mus81-Mms4 is activated and released from the INQ (Saugar et al., 2017). Accordingly, deleting Mms4 or prematurely phosphorylating it, retained Mus81 at the INQ or released Mus81 from the INQ, respectively. Furthermore, genetic perturbations that increased or decreased recombination intermediates that need to be resolved by Mus81-Mms4 greatly affected the release and persistence of this complex at the INQ. These studies reveal that nuclear functions are regulated by the INQ sequestration. It is therefore hypothesized that alongside its role as a PQC site, the INQ also acts as a “storage locker” for endogenous proteins that are sequestered to restrict or enhance their activity. These proteins are not necessarily misfolded but could be actively recruited to the INQ as part of stress responses. Another recent study on the INQ protein Mrc1 supports this model. The authors find that Mrc1 disassociates from the replication fork upon DNA damage and is sequestered at the INQ (Colding et al., 2021). Upon stress recovery, Mrc1 is released from the INQ and rejoins the replication fork for efficient restart of DNA replication. Together, this suggests the INQ is a 31 unique kind of stress-responsive quality control compartment with protein triage and pathway regulatory functions. 1.2.3.1.2 Protein refolding at INQ Once sequestered to the INQ compartment, substrate proteins are protected until stresses are removed or cells begin to recover. This reversible, and stress-induced formation is further evidenced by the rapid dissipation of INQ when the stressor such as MMS is washed out (Mathew et al., 2017). The fate of most INQ substrates is not known, but they must be either reactivated or degraded in concert with the disappearance of the INQ. As noted previously, DNA replication and repair proteins like Mus81-Mms4 or Mrc1 seem capable of returning to their cellular functions to help restart the cell cycle (Colding et al., 2021; Saugar et al., 2017). Whether this is generalizable is unclear, however, there is evidence of an active refolding machinery at the INQ. First, the complement of chaperones that localize to the INQ include the Hsp104 disaggregase, which is associated with protein refolding in partnership with the Hsp40/70/110 system. Indeed, the Hsp110/NEF Sse1 is also involved in the INQ clearance, and loss of either Hsp104 or Sse1 leads to a dramatic increase in the INQ foci during stress (Colding et al., 2021; Mathew et al., 2017). The direct effect of the Hsp40 chaperone Sis1 on the INQ is not clear; however, by measuring nuclear luciferase activity after heat stress, it has been shown to work together with Hsp70/Hsp104 to help in the INQ protein refolding. In fact, Btn2 is involved in recruiting Sis1 to the INQ and consequently forms a part of the Hsp70/104 refolding pathway (Ho et al., 2019). 32 Figure 1.5 Chaperones at INQ. A, domain architecture of five chaperones that have been found at INQ. NTD = N-terminal domain; αCLD = alpha-crystallin–like domain; CTD = C-terminal domain; PrLD = Prion-like domain; IDD = intrinsically disordered domain; CLD = alpha crystallin domain; J = J-domain; G/F = glycine/phenylalanine rich domain; G/M = glycine/methionine rich domain; D = dimerization domain; ZBD = zinc-binding domain; NBD1/2 = nucleotide-binding domain 1 or 2; MD = middle domain. B, competition-based model between Hsp40/70-sHsp proteins decides INQ substrate fate. The model shows that an exchangeable shell of sHsps enables access of Hsp40/70 chaperones to non-native substrates ready to enter the refolding or degradation pathway. Adapted from (Kumar et al., 2022). 33 Sequestrases/sHsps prevent irreversible aggregation by binding to misfolded proteins to form solubilized assemblies. These assemblies form two substructures—a dynamic outer shell with continuous binding and release of sHsps and an inner stable core, which contains fewer sHsps bound to their substrates (Żwirowski et al., 2017). Hsp70s are recruited to these assemblies through their respective Hsp40s. A competition-based model suggests that Hsp70s displace sHsps in the outer shell by competing for binding to misfolded substrates in the inner core. This in turn excludes new sHsps from re-entering and reassociating with these assemblies as Hsp70s can only be disassociated after interacting with NEFs that release ADP, allowing ATP to rebind to Hsp70 and release substrates. Hsp70s bound to substrates further recruit and activate the disaggregase Hsp104, which can refold misfolded proteins through its central channel (J. Lee et al., 2013, p. 104). While presumably the mechanisms defining the sHsp/Hsp70/Hsp104 system are at play during the reactivation of some of the INQ proteins, there are not yet studies that directly test reactivation of the endogenous INQ substrates by this pathway. A recent study showcased a negative genetic interaction at high temperatures between Btn2 or Hsp42 and a background lacking both the NEF Fes1 and Hsp104 (Ho et al., 2019). Deleting FES1 and HSP104 together results in a stabilized Hsp70–substrate interaction, thus reducing the pool of active Hsp70 and hindering the overall activity of this pathway. This temperature sensitivity was neither seen with cells lacking Hsp26 nor upon inhibiting Hsp90 activity, indicating that this defect is specific to the INQ sequestrases and low Hsp70 activity and that sequestrase activity and therefore INQ formation becomes crucial when Hsp70 activity is limited. This finding was supported by a phenotypic rescue upon expressing Sse1, another NEF largely excluded from the nucleus, with a nuclear localization sequence (NLS). Expressing 34 Hsp42 with a NLS in a btn2Δfes1Δhsp104Δ or expressing BTN2 lacking its NLS in a hsp42Δfes1Δhsp104Δ background also rescued the temperature sensitivity, indicating that the site of sequestration is interchangeable, and as long as sequestration occurs, the cells can cope with stress (Ho et al., 2019). Surprisingly, a BTN2 construct without its NTD (Btn2-ΔNTD-NLS) not only rescued the growth defect in a btn2Δfes1Δhsp104Δ background but also grew better than fes1Δhsp104Δ at high temperatures. This gain of function was explained by the Btn2-NTD mediating recruitment of disaggregases to the INQ for INQ solubilization (Ho et al., 2019). Btn2 NTD recruits Sis1 and consequently Hsp70 to the INQ, which in turn recruits Hsp104. Therefore, in a Btn2-ΔNTD-NLS fes1Δhsp104Δ background, the Hsp70/Hsp104 bichaperone system is not recruited to the INQ. This slightly increases Hsp70 availability throughout the cell and simultaneously leads to INQ persistence since the INQ would prevent irreversible aggregation of misfolded proteins. Interestingly, similar luciferase assays using another Hsp40 chaperone Apj1 did not show a function in refolding but degradation of nuclear aggregates instead, thus indicating some misfolded INQ aggregates are destined for proteasomal degradation rather than refolding and reactivation (den Brave et al., 2020). 1.2.3.1.3 Protein degradation at INQ Direct evidence of UPS proteins at the INQ is limited, but our observations and others suggest that by inhibiting the proteasome using MG132, INQ substrates aggregate more aggressively under stress. This implies that the proteasome and degradation of its substrates is essential for maintaining INQ homeostasis, which is supported by the enrichment of the proteasomal subunit 35 Rpn11 at the INQ (Gallina et al., 2015). Surprisingly, immunostaining of the INQ revealed the absence of ubiquitin, which was further tested using two misfolded substrates tGnd1-GFP and ΔΔCPY∗ that can be ubiquitinated by E3 ligases San1 and Ubr1 (Miller et al., 2015). Their sequestration to the INQ was unaffected by San1 or Ubr1 status thus arguing against the role of ubiquitination as a sorting signal in the INQ. This was further supported for the splicing factor Hsh155 whose localization at the INQ was unaffected by SAN1 deletion (Mathew et al., 2017). In contrast, cells lacking SAN1 showed delayed clearance of Mrc1 from the INQ under stress (Colding et al., 2021). Furthermore, in the same study, Cdc48 was identified as a regulator of Mrc1 retention in the INQ, raising questions about whether a Cdc48-San1 degradation pathway may exist for some of the INQ substrate proteins. The Cdc48 ATPase complex is a highly conserved chaperone that functions in protein complex disassembly, and degradation. Cdc48 functions as a chaperone in processes like endoplasmic reticulum associated degradation, membrane fusion, autophagy, cell cycle regulation, and replication stress (Baek et al., 2013). Misfolded Cdc48 substrates are ubiquitinated, bound by Cdc48, and delivered to the proteasome for degradation (Elsasser & Finley, 2005). Importantly, Cdc48 localizes to the INQ during cellular stress and suppresses the INQ substrate retention under both stressed and unstressed conditions (Mathew et al., 2020). To date, Cdc48 has been shown to regulate the localization and turnover of the two INQ substrates, Hsh155 and Mrc1. The Hsh155 INQ aggregation was significantly increased in CDC48 ts mutants (Mathew et al., 2020). The ATPase dead allele of Cdc48 could not rescue the cdc48-ts phenotype compared to the WT CDC48 plasmid, indicating that the ATPase activity is essential to retain Hsh155 aggregates in the INQ. Hsh155 interacts directly with Cdc48, whereas Mrc1 interacts with Cdc48 36 cofactors like Otu1, Ubx5, and Ufd1 as shown by bimolecular fluorescence complementation (Colding et al., 2021; Mathew et al., 2020). Cdc48 binding and interaction with these INQ substrates also influences their function in splicing (Hsh155) and replication (Mrc1). Thus, Cdc48 is likely an important chaperone adaptor for the degradation of at least a subset of the INQ proteins. Hsp40 chaperones like Ydj1 and Sis1 together with E3 ligases and Hsp70 help to maintain protein stability and support protein degradation pathways in general. Although Mrc1 showed Cdc48-dependent interactions with Ydj1 and Sis1 (Colding et al., 2021), a direct involvement of an Hsp40 chaperone in the INQ substrate degradation pathway was not clear until recent descriptions of the protein Apj1, which localizes to the INQ under stress and affects the distribution and sequestration of the INQ substrates (den Brave et al., 2020). Using heat-aggregated nuclear luciferase (LuciDM-NLS-GFP) and measuring the recovery of the luciferase activity after heat-induced inactivation, apj1Δ cells showed increased protein recovery and stability. These authors suggest that Apj1, along with Hsp70 and Hsp110, supports nuclear protein disaggregation and solubility independently of Hsp104. Moreover, Apj1 was found to promote degradation of aggregated nuclear proteins and compete with Hsp104 that drives refolding (den Brave et al., 2020). This suggests that two independent outcomes exist as a protective mechanism in the nucleus to eliminate stress-induced toxic protein aggregates: an Hsp104-dependent pathway for refolding and an Apj1-dependent pathway for degradation. 37 Figure 1.6 Players and pathways for INQ refolding or degradation. The left half shows potential regulators of the INQ substrate fate involving protein refolding/reactivation. The right half shows the INQ substrate fates involving proteasomal degradation (and potentially nucleophagy). DNA damage is indicated as a strong stress for endogenous protein aggregation, but other stresses can also induce INQ. Adapted from (Kumar et al., 2022). 1.2.3.1.4 Posttranslational modifications at INQ To date, two PTMs, namely ubiquitination and SUMOylation have been implicated in the INQ formation and dissolution. Both have established roles in protein sorting, degradation, transport, and signaling, but their roles at the INQ are yet to be elucidated (Gill, 2004). Additionally, protein phosphatases, Pph21 and Pph22, and lysine deacetylases, Hos2 and Rpd3, localize to the 38 INQ, raising the possibility that removal of these modifications could be important for the INQ resolution (Table 1). Ubiquitination is critical for localization of misfolded proteins to JUNQ; however, a general connection between the INQ and ubiquitination has not been established (Kaganovich et al., 2008). The case for the role of ubiquitin is clearer for some of the INQ substrates, such as Mrc1, which is ubiquitinated by the E3 ligase Dia2. Polyubiquitination of Mrc1 releases it from the replication fork and may serve as a recruitment signal to the INQ since it fails to relocalize to the INQ in a dia2Δ strain (Gallina et al., 2015; Mimura et al., 2009). Interestingly, overexpression of BTN2 in a dia2Δ strain partially rescues the Mrc1 INQ localization, indicating that factors other than ubiquitination could govern INQ formation (Colding et al., 2021). Ubiquitination also controls the recruitment of Mus81-Mms4 to the INQ as deletion of the E3 ligase complex members Rad6 or Bre1 abrogates their sequestration (Saugar et al., 2017). Whether this is a direct effect or an indirect result of altered DNA damage signaling is unknown. SUMOylation on the other hand is suggested to be involved in turnover of proteins at the INQ. SUMO (Smt3) colocalizes with Cmr1 and Slx5 upon DNA damage confirming its presence at the INQ (Gallina et al., 2015). Interestingly, loss of APJ1 shows synthetic lethal interactions with slx5Δ, which is a subunit of the yeast StUbL Slx5/8 (Sahi et al., 2013). While it is unclear whether degradation of the INQ substrates by Apj1 depends on StUbLs or SUMO, the majority of StUbL substrates are nuclear proteins including transcription factors, DNA replication and repair proteins (Garza & Pillus, 2013). Sequestration of Hsh155 to the INQ upon DNA damage is also dependent on SUMO conjugase and E3 ligase Ubc9 and Siz1, respectively. This implied 39 that the INQ substrates could be SUMOylated upon DNA damage and that this could act as a recruitment signal for Btn2 and Hsp42 to sequester proteins to the INQ. Indeed, the SUMO-binding molecular chaperone Cdc48/VCP is also recruited to the INQ where it functions to counteract INQ persistence (Mathew et al., 2020). Further studies focusing on the INQ and SUMOylation should reveal the SUMOylated proteome at the INQ, its role in the formation, and consequently clearance of the INQ. 1.2.3.1.5 Nucleophagy and the fate of INQ It was hypothesized and recently proven that the INQ is spatially located at the nucleolar-vacuolar junction (NVJ), adjacent to the nucleolus (Sontag et al., 2023). Apart from refolding and degradation within the nucleus, this pathway adds an additional layer for clearance of misfolded proteins wherein INQ can be extruded into a lytic vacuole and degraded therein. Using a combination of cryogenic soft x-ray tomography and super resolution microscopy, authors showed that INQ and JUNQ form on opposite sides of the nuclear envelope and home to the same site, at the NVJ (Sontag et al., 2023). Genetic manipulations revealed that the homing signal, and therefore trafficking to the NVJ was mediated by the endosomal sorting complexes required for transport (ESCRT) machinery, specifically the proteins Chm7 and Vps23. Additionally, the extrusion of INQ to the vacuole was found to be mediated by another ESCRT protein, namely Vps4 (Sontag et al., 2023). This location of this process presents an exciting possibility for the exchange of misfolded proteins between JUNQ and INQ through nuclear pores or diffusion through the nuclear membrane. Moreover, the fate of proteins at INQ remains elusive. Perhaps, if refolding and UPS-mediated degradation pathways are insufficient to handle 40 the load of non-native proteins, the INQ could be subject to nucleophagy. Further research will be required to explore this possibility. 1.2.3.2 Juxtanuclear Quality control site (JUNQ) and the Insoluble Protein Deposit (IPOD) Similar to INQ, which is the sequestration site in the nucleus, JUNQ and IPOD are sites found in the cytoplasm. The IPOD is perivacuolar, adjacent to the preautophagosomal structure, and primarily made up of amyloidogenic proteins that are terminally aggregated, immobile and show a slow rate of diffusion with the cytosol (Kaganovich et al., 2008). On the other hand, the JUNQ is associated with the nuclear envelope but is peripheral to it and resides outside the nucleus. Contrary to the IPOD, the JUNQ is highly dynamic in nature, soluble, and concentrates the 26S proteasome and disaggregation machinery and therefore acts as an active site for protein degradation and refolding. Partitioning between these two sites depends on the misfolded protein’s ubiquitination state as blocking ubiquitination causes proteins to localize to IPOD instead of JUNQ and genetically engineering a ubiquitinated Rnq1-GFP localized to both the IPOD and JUNQ instead of only the IPOD (Kaganovich et al., 2008). Other sites such as Q bodies, stress foci, and peripheral aggregates have also been found but have since been grouped together as CytoQ bodies that could potentially represent that same deposition (Escusa-Toret et al., 2013; Specht et al., 2011; Spokoini et al., 2012). CytoQ bodies contain soluble misfolded proteins and undergo ATP-dependent coalescence that could represent the pathway that ultimately leads to JUNQ formation (Sontag et al., 2017). 41 1.2.3.3 MitoStores and Stress Granules In addition to INQ, JUNQ and IPOD, which have been the main focus of sequestration studies, there are other compartments that are being discovered. MitoStores, are cytoplasmic compartments that are distinct to JUNQ and IPOD as they are in close proximity to the mitochondria (Krämer et al., 2023). Upon mitochondrial import stress, proteins are sequestered to MitoStores through Hsp42, again showcasing its pleiotropic role in maintaining proteostasis through sequestration. Unlike other compartments, however, proteins that localize to MitoStores appear to carry a mitochondrial targeting sequence in their NTD suggesting that these proteins are mitochondrial precursors that are sequestered under stress. While it was thought that sequestration is random, MitoStores represent an ideal example to show that it is a strictly controlled process. Researchers utilized the split-GFP collection to pinpoint proteins interacting with Hsp104 at MitoStores (Krämer et al., 2023). Interestingly, while Hsp104 interacts with many mitochondrial proteins under stress, when the UPS is perturbed via deletion of RPN4, Hsp104 interacts with new, and more mitochondrial proteins. This suggests that if present, the UPS can degrade certain precursors but, in its absence, these precursors are sequestered to MitoStores to be acted on later. Similarly, stress granules, previously known to contain only mRNA and RNA binding proteins, have been found to sequester misfolded proteins, which in turn recruit chaperones including HSP70s for turnover (Mateju et al., 2017). 1.2.3.4 Aggresomes In line with the previous compartments mentioned, aggresomes represent juxtanuclear cytoplasmic cytoprotective compartments that harbor ubiquitinated misfolded proteins in human cells (Johnston & Samant, 2021). However, unlike in yeast where binding and sequestration is 42 mediated by sHsps, HDAC6, a cytoplasmic deacetylase is responsible for aggresomes formation. It does so by binding to polyubiquitin chains on misfolded proteins via its Zinc Finger Ubiquitin Binding Protease domain and interacting with the dynein-dynactin system through its CTD to translocate its cargo to aggresomes (Boyault et al., 2006; Kawaguchi et al., 2003). While chain specificity remains contested, HDAC6 appears to bind strongly to K63-polyUb chains compared to K48-polyUb chains, suggesting an important role for K63- chains in aggresome formation (Olzmann et al., 2007). With respect to the proteome at aggresomes, expected PQC factors have been documented at these inclusions, such as the HSP40/70/90 system, CCT, ubiquitin adaptors like p62/SQSTM1, sHsps, p97/Cdc48/VCP, many E3 ligases and DUBS. Since many PQC proteins involved in removal of misfolded proteins relocalize to aggresomes, clearance from these inclusions seems to be through both proteasome-mediated degradation and autophagy. However, one proposed mechanism is that the proteasome DUB Poh1 produces unanchored K63-polyUb chains which bind and activate HDAC6 to facilitate clearance via autophagy when the proteasome is burdened or challenged, thus linking the two clearance routes (Hao et al., 2013). In conclusion, the three pillars of protein quality control: refolding, degradation and sequestration mediate and control a multitude of processes essential for cell survival. One such process is regulating the process of DNA replication during cell division which requires the assembly of multiple protein complexes, chromatin remodelling, post translational modifications and ensuring the expression, localization and proper folding of all proteins that execute the DNA damage response. Additionally, stresses such as reactive oxygen species and chemical agents like methyl methanesulfonate cause DNA replication stress that evoke a dynamic proteomic response 43 to activate the DNA damage response that is essential for survival. A few of these pathways are discussed in detail below. 1.3 DNA Replication stress and response During cell division, the entire genome needs to be replicated accurately to ensure faithful inheritance of information. DNA replication is carried out by the replisome, a highly complex structures, consisting of a multimeric helicase complex termed CMG (Cdc45–MCM–GINS), multi-subunit leading and lagging strand polymerases, a processivity factor called PCNA (proliferating cell nuclear antigen), a trimeric single-stranded DNA (ssDNA) binding protein RPA (Replication Protein A) and numerous other factors. The progression of the replisome can be impaired by various physical barriers. Such barriers include DNA–protein and DNA–DNA crosslinks, DNA lesions such as thymidine dimers, DNA secondary structures such as G-quadruplexes, transcription-replication conflicts associated with DNA:RNA hybrid structures called R-loops, topological stresses, depletion of nucleotide pools and other conditions (Zeman & Cimprich, 2014). Paused replisomes result in a structure with the replisome, a leading strand and a lagging strand, called a replication fork. When replication fork slowing and stalling occur frequently during S-phase, cells experience replication stress. Replication stress notably occurs in cancer where it is likely to drive mutational processes that underlie genome instability phenotypes associated with the disease. Indeed, oncogene activation is sufficient to induce DNA replication stress (Kotsantis et al., 2018). Stalled replication forks must be restarted or rescued by a converging fork in order to prevent under-replicated DNA entering mitosis. Stalled forks first expose ssDNA that is coated with 44 RPA. RPA stimulates the activation of the Ataxia-telangiectasia and Rad3 related protein (ATR) kinase which initiates a cascade of signalling events that can stall the cell cycle and recruit various effectors to the stalled fork (Zeman & Cimprich, 2014). Replication stress is often induced in vitro by damaging DNA with alkylating agents like methyl methanesulfonate (MMS), by creating protein–DNA crosslinks with the topoisomerase I inhibitor camptothecin (CPT), by depleting nucleotide pools with hydroxyurea (HU), through reactive oxygen species using hydrogen peroxide, or by directly inhibiting DNA replication with aphidicolin. Depending on the nature and timing of the fork stalling event, replication forks can be restarted via lesion bypass mechanisms such as translesion synthesis, repriming of replication after the lesion, or template switching (Zeman & Cimprich, 2014). Alternatively, the fork can be cleaved to create a broken end that can lead to fork restart by homologous recombination (HR) (Pepe & West, 2014). Cellular resilience to fork stalling is a complex process that remains an active area of investigation. 1.3.1 Double strand breaks and DNA repair A stalled fork can eventually collapse and result in the formation of a double strand break (DSB). DSBs, or damaged DNA, need to be repaired immediately as unrepaired breaks can drive mutagenesis. DSBs are sensed and acted on by first responders such as the MRE11-RAD50-NBS1 (MRN complex; MRX in Yeast) nuclease that activate the ataxia-telangiectasia mutated (ATM) kinase and ataxia-telangiectasia and Rad3 related (ATR) kinase which in turn activate and control different repair mechanisms (Duursma et al., 2013; J.-H. Lee & Paull, 2005). With respect to DSB repair, two major mechanisms have been described: the homologous recombination (HR) and the non-homologous end-joining (NHEJ) pathway. 45 HR and NHEJ operate to preserve genome integrity but with distinct cell cycle profiles. NHEJ is available to cells throughout the cell cycle and essentially consists of religating the broken DNA ends. It is a quick but error-prone pathway that results in the formation of small insertions or deletions at the DSB site (Mao et al., 2008; Shrivastav et al., 2008). On the other hand, HR requires the presence of a sister chromatid that can be used as a template to repair the DSB. Therefore, it is only activated during S/G2 phases of the cell cycle and is preferred over NHEJ since it is error-free in nature (Mao et al., 2008; Shrivastav et al., 2008). Additionally, HR requires processing of the DSB ends by nucleases such as MRN that can resect the break, generating ssDNA overhangs. These overhangs are coated by RAD51 which can then invade the homologous sister chromatid that can be used as a template for DNA polymerases which extend the invading strand and repair the DSB. While they have their own machinery, the choice between HR and NHEJ depends on various factors including the cell cycle stage, availability of necessary repair factors and the accessibility of the damaged DNA (Shrivastav et al., 2008). The latter is heavily influenced by the epigenetic state and how chromatin is organized upon replication stress. 1.3.2 Chromatin organization during replication stress Nucleosomes are the basis of chromatin organization in eukaryotes. Nucleosomes are assembled by the sequential association of two histone H3–H4 dimers and two histone H2A–H2B to form an octameric protein complex that is wrapped by 147 base-pairs of DNA (Luger et al., 1997). Nucleosome assembly requires the aid of protein chaperones that function in the storage, transport, modification, or deposition of histones onto DNA. Packaging of DNA with 46 nucleosomes to form chromatin is essential for genome integrity. Histone modifications and dynamics regulate not only chromatin compaction and gene expression but also DNA repair, DNA replication and mitosis (Hauer & Gasser, 2017). Perturbation of any of these processes can lead to DNA damage and mutations. The process of chromatin replication presents additional challenges to cells. Nucleosomes must be removed ahead of advancing replication forks and be recycled behind the forks onto the nascent sister chromatids to preserve the epigenetic state of the replicated locus. Briefly, during replication, newly synthesized nucleosome core particles histone H3 and H4 can be bound by the histone chaperone anti-silencing function 1 (ASF1), which associates ahead of the replication machinery with the MCM2 (MiniChromosome Maintenance 2) subunit of the core helicase complex (Alabert & Groth, 2012; Clément & Almouzni, 2015). ASF1 may also coordinate the first steps of histone recycling at the leading edge through the deposition of histone “pre-disposition marks,” which signals for the disassembled and nascent histone complexes to be re-incorporated into the newly synthesized DNA by other histone chaperones, like the chromatin assembly factor 1 (CAF-1) (Das et al., 2009). CAF-1 is recruited to sites of DNA synthesis through interactions with PCNA (Yang et al., 2016). Histone pre-disposition marks include the di-acetylation of H4 at lysines 5 and 12 (H4K5K12) by the H4K5K12 lysine acetyltransferase RBAP46/HAT1 in human cells and the acetylation of H3K56ac and H3K27ac by Rtt109 in budding yeast (H3K14ac and H3K18ac in humans) prior to deposition into newly synthesized DNA (Alabert & Groth, 2012). 47 While ASF1-CAF-1 driven recycling of H3–H4 is important for replication progression, more recent studies reveal crucial intrinsic roles for the replication machinery itself as a histone chaperone. Core subunits of the leading strand polymerase (Polε), POLE3-POLE4, have been demonstrated to have histone chaperone activity and promote histone recycling (Bellelli et al., 2018). The ssDNA-binding protein complex RPA is also present at every replisome and has been ascribed a role in histone deposition. RPA binds directly to H3–H4 complexes and facilitates their deposition into adjacent double stranded DNA, likely in concert with histone chaperones in the cell (Liu et al., 2017). Finally, the MCM2 subunit of the replicative helicase also binds directly to histones and works with the FACT complex to promote histone recycling during replication. More recently, the histone recycling function of MCM2 was shown to be essential to maintain epigenetic inheritance of histone post-translational modifications (Foltman et al., 2013; Petryk et al., 2018). Thus, the FACT-MCM2 pathway may provide most of the essential histone recycling activity in normal cells. Moreover, the action of various chromatin remodelling complexes such as SWI/SNF (Switching deficient/Sucrose-Non-Fermenting), Imitation Switch (ISWI) and INO80 (Inositol requiring 80) is important for normal replication (Vincent et al., 2008). Thus, core replisome components like Polε, MCM2 and RPA, along with regulation by dedicated histone chaperones and chromatin remodellers together ensure histone deposition and recycling in replication, which in turn ensures epigenetic stability during cell division. 1.3.2.1 Histone acetylation Once formed, histones can undergo frequent cycles of post-translational modification. PTMs on histones have been shown to control functional changes in chromatin structure, gene expression, protein-protein interactions, and transgenerational epigenetic inheritance. One of the most 48 heavily studied histone PTMs is the acetylation of lysines, largely in the amino-terminal tail of histones. Similar to ubiquitination and SUMOylation discussed earlier, histone acetylation is highly reversible and is modulated by a number of enzymes. The histone acetyl mark is laid down by a class of proteins call the histone/lysine acetyltransferases (HATs/KATs) and removed by histone/lysine deacetylases (HDACs/KDACs) (Seto & Yoshida, 2014). Usually co-expressed, these enzymes possess substrate specificity but also showcase redundancy. Interestingly, there is growing evidence that HDACs can also deacetylate non-histone proteins and have acetylation-independent roles such as HDAC6’s role in aggresomes formation and clearance. Human HDACs specifically are grouped into four classes based on their sequence similarity to yeast HDACs. Class I HDACs have sequence similarity to the yeast protein Rpd3 which has important roles in regulating transcription, DNA replication through origin firing and the DNA damage response (He et al., 2022; Ortega et al., 2019; Sharma et al., 2007). Research aimed at understanding regulators of DNA replication stress is growing dramatically. Much of this work has naturally focused on understanding the DNA transactions that promote lesion bypass and fork restart, or fork cleavage. However, replication stress signalling, fork reversal, lesion bypass and fork cleavage all appear to be influenced by changes to cellular proteome through the action of chromatin remodellers and chaperones. Just as chromatin is recognized as a key regulator of DNA DSB repair, it will be important that models of replication stress tolerance be understood in the context of protein quality control. This is because there is great current interest in the anti-cancer therapeutic potential of both replication stress targeting agents such as ATR inhibitors, and in hijacking the PQC system for targeted protein degradation 49 (Nalawansha & Crews, 2020; Pilié et al., 2019). The intersection of these fields presents new opportunities to understand a complex biological system and improve human health. 1.4 Thesis objectives Spatial compartmentalization is a key facet of protein quality control that serves to store disassembled or non-native proteins until triage to the refolding or degradation machinery can occur in a regulated manner. Yeast cells sequester nuclear proteins at intranuclear quality control bodies (INQ) in response to various stresses, although the regulation of this process remains poorly understood. The goal of my thesis is to investigate the role of the post-translational modification SUMOylation in mediating INQ formation and clearance. Thesis hypothesis: DNA damage induced SUMOylation regulates formation and clearance of proteins at INQ. The objective of Chapter 2 was to establish Rpd3, a histone deacetylase, as a new marker for INQ under DNA damage and assessing factors that influence its sequestration. This was accomplished by utilizing C-terminal GFP fusion constructs, monitoring protein localization using microscopy, assessing INQ formation, and by performing fractionation in various genetic backgrounds. The outcome of this work demonstrated that sequestration of Rpd3 to INQ is a general response to DNA damage and that this relocalization is governed by many proteins including sHsps Btn2 and Hsp42. 50 The objective of Chapter 3 was to investigate the role of SUMOylation in INQ formation. This was accomplished by combining microscopy with genetic perturbations in the SUMOylation cascade and performing immunoprecipitation to purify SUMOylated proteins under DNA damage. The outcome of this work showed that SUMOylation is involved in INQ formation while polySUMOylation has a role in INQ clearance. Additionally, DNA damage induces monoSUMOylation and diSUMOylation of the two sHsps Btn2 and Hsp42, respectively. Finally, efforts to map the modified lysine in Btn2 revealed that SUMOylation of Btn2 occurs at its C-terminus. The objective of Chapter 4 was to understand the role of Btn2 SUMOylation. This was accomplished by genomically integrating the non-SUMOylatable Btn2 construct from Chapter 3 and assessing its role in INQ formation and clearance. The outcome of this work revealed that Btn2 SUMOylation is not important for INQ formation but rather is at the nexus of protein refolding and degradation at INQ, thereby regulating INQ clearance. 51 Chapter 2: Rpd3 is a novel INQ marker. This chapter is based on an article currently under review: Kumar, A, Mathew, V., Stirling, P.C. “DNA damage induced SUMOylation of Btn2 regulates nuclear protein quality control”. (2023) This was a collaborative effort with Dr. Veena Mathew who aided in method development under the supervision of Dr. Peter Stirling. 2.1 Background Protein Quality Control (PQC) circuits are mediated and executed by molecular chaperones that either refold, degrade, or sequester misfolded proteins, forming the three pillars of PQC. Under severe stress conditions that overburden PQC systems, misfolded proteins are spatially sequestered to specialized and distinct membrane-less inclusions within the cell (Kaganovich et al., 2008; Miller et al., 2015). This controlled aggregation process includes relocalization of chaperones involved in refolding and degradation pathways suggesting that sequestration could help improve local concentration of PQC factors to promote stress recovery. In Saccharomyces cerevisiae, misfolded proteins have been found to partition into one of three inclusions: Nuclear misfolded proteins are sequestered to the peri-nucleolar intranuclear quality control site (INQ), whereas cytoplasmic proteins can be sequestered to two distinct inclusions based on their solubility. If soluble, cytoplasmic misfolded proteins are ubiquitinated and sequestered to the peri-nuclear juxtanuclear quality control site (JUNQ) while insoluble proteins are targeted to the peri-vacuolar insoluble protein deposit (IPOD) (Kumar et al., 2022; Sontag et al., 2017). 52 Multiple small cytoplasmic foci containing misfolded proteins have also been reported but are thought to coalesce to form the JUNQ (Escusa-Toret et al., 2013). While these inclusions are site-specific, there have been reports of interplay such that cytoplasmic misfolded proteins can be shuttled between the JUNQ to INQ. Furthermore, the INQ and JUNQ have been found to home to the nuclear-vacuolar junction supporting this idea that there can be transfer of misfolded proteins to and from different sequestration sites (Sontag et al., 2023). The spatiotemporal organization of misfolded proteins is orchestrated by small-heat shock proteins (sHsps) Btn2 and Hsp42 in S. cerevisiae (Miller et al., 2015). Studies utilizing terminally misfolded proteins (e.g. VHL, Ubc9-tsGFP) have uncovered site-specific roles of these sHsps such that Hsp42 targets proteins to cytoplasmic inclusions whereas Btn2 sequesters proteins to INQ (Grousl et al., 2018; Ho et al., 2019). Interestingly, endogenous proteins have also been found to relocalize to INQ under replication stress and heat shock coupled with proteasomal disruption (Gallina et al., 2015; Mathew et al., 2017, 2020; Saugar et al., 2017). An initial screen performed by our group, coupled with other studies revealed a catalogue of proteins that relocalize to INQ including Rpd3 and Hos2, two histone deacetylases, a U2 sn-RNP splicing component Hsh155, an Hsp40 chaperone Apj1 and a protein degradation machine Cdc48. While previous efforts by our group have helped characterized the sequestration of Hsh155 and Cdc48-mediated clearance at INQ, our understanding of factors that govern INQ formation and the reason for sequestering endogenous proteins remains poorly understood. Here, in Chapter 2, I will establish Rpd3 as a new INQ marker under replication stress and use it as my model substrate to explore the role of DNA damage in INQ formation. I show that 53 exposure to chemicals inducing DNA damage leads to Btn2- and Hsp42- dependent sequestration of Rpd3 to INQ. Moreover, only Rpd3 and not its complex partners get sequestered to INQ. I evaluate and assess different pathways that regulate INQ formation by Rpd3. Finally, I test different mutations and chemicals and show that sequestration of Rpd3 to INQ is a general response to DNA damage. 2.2 Materials and Methods 2.2.1 Experimental Model and Subject Details Yeast (Saccharomyces cerevisiae) strains used in this study (s288c), database IDs, genotypes, primers, and plasmids used in this study are listed in Appendix 1. All strains were grown under nutrient rich YPD medium or synthetic medium lacking amino acids for auxotrophic selection unless otherwise indicated. For all MMS treatment conditions, log cells were exposed to 0.05% MMS (∼99%; Sigma) in synthetic complete (SC) medium for 2 h unless otherwise noted. For growth curves and washouts, MMS was then washed out and replaced with fresh MMS-free medium. All other agents DNA damaging agents used were hydroxyurea (200mM), hydrogen peroxide (2mM), camptothecin (25μM), and ethyl methanesulfonate (0.5%) for 2 h in SC medium before imaging. 2.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods Imaging and subsequent analysis were performed as described previously (Mathew et al., 2017). Log-phase yeast were mounted on slides pre-treated with concanavalin A, in SC growth medium. Immobilized cells were imaged using an Objective HCX PL APO 1.40 NA oil immersion 100× objective on an inverted Leica DMi8 microscope with a motorized DIC (differential interference 54 contrast) turret (for DIC imaging) and a filter cube set for FITC/TRITC (for GFP and mCherry fluorescence imaging). The images were captured at room temperature using a scientific complementary metal oxide semiconductor (sCMOS) camera (ORCA Flash 4.0 V2; Hamamatsu), collected using MetaMorph Premier acquisition software and post processed [including gamma adjustments, counting of cells with/without foci, Z-stacking, projection of maximum intensity and foci intensity measurements] using ImageJ (National Institutes of Health). For all microscopy experiments, the significance of the differences was determined using Prism version 8 or higher (Graphpad). For foci and nuclear intensity measurements, samples were compared with t-tests or ANOVA; Prism performs F-tests for variance as part of this analysis. For comparisons of proportions, Fisher tests were used, and P-values were Holm–Bonferroni corrected in the event of multiple comparisons. Sample sizes and specific statistical details for each image analysis are listed in the figure legends. 2.2.3 Western Blotting and protein stability time course For western blotting, whole-cell extracts of logarithmic phase cells were prepared using trichloroacetic acid (TCA) extraction and blotted with experiment-specific antibodies. For the stability time course, overnight cultures of the indicated strains were diluted to below an OD600 nm of 0.2 and allowed to progress into the logarithmic phase before collection. The cells were treated with or without cycloheximide (CHX) at a concentration of 200 μg/ml for the indicated times (0–120 min). Final samples of 2×107 cells were collected by centrifugation, and whole-cell extracts were prepared by TCA extraction and used for immunoblotting. 55 2.2.4 Fractionation of the insoluble pellet Cellular fractionation was performed as described in. In brief, logarithmic cells were collected after or before MMS treatment. Frozen cells were resuspended in a fractionation buffer (100mM HEPES, 1% Triton X-100, 300mM NaCl, protease inhibitor cocktail) and lysed by bead beating. Lysates were precleared at 100g, 4℃ for 5 minutes. Supernatant (whole cell lysate) was recovered and fractionated at 16000g, at 4℃ for 10 minutes to separate into soluble (S) and insoluble (P) fractions. Equal amounts were loaded onto gels followed by immunoblotting. 2.3 Results 2.3.1 Rpd3 is sequestered at INQ during replication stress. Previous high-throughput microscopy studies have noted relocalization of the histone deacetylases Rpd3 and Hos2 to INQ (Gallina et al., 2015; Mathew et al., 2017). While the functional significance of sequestering these two HDACs specifically is still not understood, Hos2 fused with an mCherry tag has been used as INQ marker. While this construct marks INQ successfully, the fluorophore has low brightness and stability as it gets photobleached over long exposure times. In an attempt to create a better marker for INQ, I confirmed the relocalization of Rpd3 by using a C-terminal GFP fusion and monitored its sequestration using microscopy. To induce replication stress, I used the alkylating agent methyl methanesulfonate (MMS). This will be a recurring drug treatment throughout this thesis as MMS alkylates DNA bases to create guanine and adenine alkyl adducts that act as replication fork barriers. Fork pausing eventually leads to DSBs that cause DNA damage. MMS has been used as a replication stressor in multiple 56 INQ studies and has become an accepted form of treatment to induce INQ sequestration (Gallina et al., 2015; Mathew et al., 2017). Figure 2.1 Rpd3 is sequestered to INQ upon replication stress. a Rpd3 forms inclusions upon acute exposure to 0.05% Methyl methanesulfonate (MMS) which induces replicative stress. Right, representative image; Left, quantification of cells with Rpd3-GFP foci. UNT = untreated. b, c Representative images of colocalization of Rpd3-GFP with Hta2-mCherry and Hos2-mCherry respectively. All error bars represent means ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, Fisher’s test. Scale bars: 5µm. Following MMS exposure, Rpd3 formed inclusions within an hour (Figure 2.1a). While Rpd3 itself is canonically present in the nucleus, we used a Hta2-mCherry nuclear marker to confirm its localization (Figure 2.1b). Indeed, we found that Rpd3 formed both nuclear and cytoplasmic 57 inclusions upon MMS treatment. Interestingly, Rpd3 formed nuclear inclusions first, rapidly within an hour followed by a gradual increase in cytoplasmic inclusions. This suggests that since Rpd3 is nuclear, its sequestration upon DNA damage is first directed to nuclear inclusions and overtime gets shuttled to cytoplasmic inclusions as well. Additionally, to confirm the nature of these inclusions, we colocalized Rpd3-GFP with Hos2-mCherry to confirm that the Rpd3 nuclear inclusions were in fact INQ (Figure 2.1c). 2.3.2 Sequestration to INQ is mediated by small-heat shock proteins. As highlighted previously, sequestration to INQ and other inclusions is orchestrated by the chaperones Btn2 and Hsp42 (Ho et al., 2019; Miller et al., 2015). To this end, I retested Rpd3 INQ localization in btn2Δ and hsp42Δ cells. While deleting HSP42 abrogated all inclusions, deleting BTN2 removed INQ formation whereas cytoplasmic inclusions (hereby CytoQ) remained (Figure 2.2a). In line with this, cell fractionation revealed that Rpd3 was mostly soluble in unstressed cells but that induction of replication stress via MMS treatment caused a shift to the insoluble fraction (or pellet). Deleting BTN2 reduced Rpd3 protein levels in the insoluble pellet whereas an Hsp42 deletion completely blocked the accumulation of Rpd3 in the pellet (Figure 2.2b). While this supported the site-specific role of Btn2 targeting proteins to nuclear inclusions, it suggests that Hsp42 might also play a major role in sequestering proteins to both nuclear and cytoplasmic inclusions under DNA damage. Furthermore, the presence of some Rpd3 in the insoluble fraction in cells lacking Btn2 suggests that the pellet contained both INQ and CytoQ inclusions. It is important to note that loss of HSP42 appeared to affect Rpd3 protein levels in the whole cell 58 extract (down to ~50% of WT Rpd3) which restricted the conclusions I could draw for the role of Hsp42 in sequestering Rpd3 to different compartments. Figure 2.2 Sequestration of Rpd3 to INQ is mediated by sHsps Btn2 and Hsp42. a Deletions of compartment specific sequestrases Hsp42 and Btn2 results in a significant reduction of Rpd3 INQ foci in MMS treated cells. b Cellular fractionation supports imaging data showing Hsp42 and Btn2-dependent shift of Rpd3-GFP to the insoluble pellet under stress. W = whole cell extract, S = supernatant, P = insoluble pellet. All error bars represent means ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, Fisher’s test. Scale bars: 5µm. Normalized protein levels in different fractions was calculated from duplicate experiments. 59 Rpd3 forms three major protein complexes that control transcriptional regulation through their histone deacetylation activity (Baker et al., 2013; Carrozza et al., 2005; Keogh et al., 2005). The Rpd3L and Rpd3S complexes both have a common catalytic core complex made up of Rpd3-Sin3-Ume1 with additional subunits that dictate complex specificity (X.-F. Chen et al., 2012). Rpd3μ on the other hand is composed of Rpd3, Snt2 and Ecm5 and is important for oxidative stress response (Figure 2.3a). Since other INQ proteins can either be sequestered within complexes (e.g. Mus81-Mms4) or can disassociate from their complexes (e.g. Hsh155), we tested whether Rpd3 complex subunits also relocalize to INQ (Mathew et al., 2017; Saugar et al., 2017). I imaged C-terminal GFP fusions of ten Rpd3 complex subunits and found that none of the subunits tested relocalize to INQ upon replication stress (Figure 2.3a). Fractionation of Rpd3L (Pho23), Rpd3S (Rco1) and Rpd3μ(Snt2) subunits confirmed that, unlike Rpd3 itself, all remained in the soluble fraction after MMS treatment (Figure 2.3b). These data suggest that Rpd3 dissociates from its complexes or is sequestered to INQ as a nascent unassembled polypeptide by Btn2 and Hsp42 following MMS treatment. 60 Figure 2.3 Rpd3 complex partners do not relocalize to INQ. a Schematic of the various Rpd3-containing complexes and representative images of GFP-tagged Rpd3 complex subunits after MMS treatment. b Fractionation of Rpd3 complex subunits in both untreated and MMS-treated conditions followed by GFP western blots or an anti-Pgk1 loading control. UNT = untreated. W = whole cell extract; S = supernatant; P = pellet. Scale bars: 5µm. 61 2.3.3 Sequestration of Rpd3 to INQ is a DNA damage response. While the major mechanism of DNA damage via MMS treatment is through the alkylation of DNA bases resulting in replication fork stalling, blockage and double strand breaks, there have been reports of MMS alkylating RNA and peptides (Groth et al., 2010; Thomas et al., 2020; Yan et al., 2019). To understand whether INQ formation is a general response to DNA damage and not through proteotoxic artifacts introduced by MMS, I tested a panel of DNA damaging agents each having a different mechanism of action. I used hydrogen peroxide (oxidative damage), camptothecin (topoisomerase-poison), hydroxyurea (replication fork stalling), and ethyl methanesulfonate (alkylating damage) and found that all DNA damaging agents caused INQ formation for Rpd3 indicating that this is likely to be a general response to replication stress and not due to protein reactivity of MMS (Figure 2.4a). Figure 2.4 INQ formation for Rpd3 following different genotoxic treatments. a Rpd3-GFP foci levels following different genotoxic stressors. H2O2 (2mM), Camptothecin (CPT; 25uM), Hydroxyurea (HU; 200mM) and Ethyl methanesulfonate (EMS; 0.5%). b Representative images are shown. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, Fisher’s test. Scale bars: 5µm. 62 If INQ formation is a general response to replication stress, then genetic perturbations not requiring any exogenous stress should recapitulate the phenotype. To test this I used mutations that inactivate the DNA damage response kinases Mec1 and Tel1 (Gasch et al., 2001; Tkach et al., 2012; Zhao, 2001). I found that both mec1Δsml1Δ and tel1Δ cells have elevated INQ formation after MMS treatment, hinting that a combination of MMS and a higher internal rate of DNA damage caused due to these deletions resulted in a stronger response (Figure 2.5a). Remarkably, sml1Δ cells had a similar level of INQ formation both in unstressed and MMS treated cells when compared to mec1Δsml1Δ, tel1Δ, and mec1Δsml1Δtel1Δ cells (Figure 2.5a). Sml1 inhibits dNTP pool production and is degraded via Mec1 and Tel1-mediated phosphorylation. Consistent with my HU data (from Figure 2.4a), which also inhibits dNTP production, overexpression of SML1 caused INQ formation in WT cells (Figure 2.5b). Deletions of SML1, rather than impairing replication, increase dNTP pools, increase replication fork speed, and favour late replication origin firing (Poli et al., 2012). Since both increased expression and deletion of SML1 seem to promote INQ formation my data suggests that careful balancing of the nucleotide pool and replication dynamics are key signals for Rpd3 deposition at INQ. Importantly, a population of unstressed cells also formed INQ for Rpd3 in these mutants. Budding index analysis revealed that the cells undergoing spontaneous INQ formation were small budded cells and therefore likely to be in S-phase (Figure 2.5c). This aligns with my hypothesis that a higher burden of internal replication stress was sufficient for INQ formation as part of the DNA replication stress response. 63 To further delineate the DNA damage response arm that dictates INQ formation, I tested Rpd3-INQ formation in DNA damage checkpoint (DDC) (Rad9, Chk1) and DNA replication checkpoint (Mrc1, Tof1, Csm3) mutants. Only DDC mutants had increased INQ formation suggesting that a functioning DDC opposed INQ formation during replication stress (Figure 2.5d). Overall, I concluded that sequestration of Rpd3 to INQ is a general response to DNA replication perturbations. Figure 2.5 DNA damage regulates INQ formation for Rpd3. a Quantification of Rpd3 foci formation in sml1Δ, mec1Δsml1Δ, tel1Δ, and mec1Δsml1Δtel1Δ cells. b Rpd3-foci quantification showing SML1 overexpression rescues sml1Δ INQ phenotype 64 but also induces INQ formation in untreated cells. c Rpd3-GFP foci counts as a function of budding index for unstressed cells harboring sml1, mec1 and tel1 deletions. Representative images are shown at right. d Quantification of Rpd3 INQ formation in DNA repair deficient mutants. Mean percentage values of 2/3 replicates are shown below each dataset. All error bars represent ± SEM, n=2/3, >100 cells each. ****, p<0.0001, Fisher’s test. Scale bars: 5µm. 2.3.4 INQ turnover for Rpd3 is mediated by chaperones. Once misfolded proteins have been deposited at INQ, they either need to be refolded or degraded. Assuming Rpd3 is misfolded upon DNA damage (discussed in detail in Section 2.4), its turnover from INQ must be mediated by chaperones. Previous studies have suggested degradation-specific INQ chaperones such as the Hsp40 Apj1, the AAA+ ATPase Cdc48 and nucleophagy regulating proteins Nvj1 and Vac8 (den Brave et al., 2020; Mathew et al., 2020; Sontag et al., 2023). In order to understand whether Rpd3 is under constant action of these chaperones, I monitored INQ formation for Rpd3 in different genetic backgrounds. 2.3.4.1 Defining regions of Apj1 involved in INQ clearance. Cells lacking APJ1 had higher levels of INQ foci upon DNA damage compared to a WT strain, indicating that Apj1 is involved in clearance of Rpd3 from INQ (Figure 2.6b). This is in line with previous observations that Apj1 forms a degradation-dependent pathway that dictates the fate of proteins at INQ. Interestingly, untreated apj1Δ cells had spontaneous foci indicating that Rpd3 might be under a constant action of Apj1, such that blocking degradation results in accumulation of Rpd3 at INQ. 65 Figure 2.6 Apj1 affects turnover of Rpd3 at INQ. a Schematic representing domain organization in Apj1. b, c Rpd3-GFP foci levels in an apj1Δ background transformed with an empty vector, the J domain mutant H34Q, domain truncated mutants for ZnF and G/FS, and hydrophobic cleft point mutants. Mean percentage values of 2/3 replicates are shown below each dataset. All error bars represent ± SEM, n=2/3, >100 cells each. ****, p<0.0001, Fisher’s test. I next sought to understand which domains of Apj1 regulate INQ clearance. Apj1 belongs to the Class I family of Hsp40s with a J domain at its NTD followed by a glycine/phenylalanine region (G/FS), a Zinc-Finger domain (ZnF), a CTD and a dimerization domain (DD) (Figure 2.6a). Additionally, I also used a J domain mutant H34Q where the HIS-PRO-ASP conserved motif is mutated to PHE-PRO-ASP, thus perturbing the J domain activity resulting in reduced Hsp70 66 ATPase activity (Sahi et al., 2013). I transformed plasmids containing domain-specific mutants of Apj1 into an apj1Δ background in an attempt to rescue increased foci phenotype. As expected, the full-length wild type (wt) plasmid completely reduced foci and caused a reduction in INQ levels. Conversely, the J domain mutant. H34Q, only caused a partial rescue highlighting the importance of the Hsp40s in stimulating Hsp70 activity (Figure 2.6b). While the ZnF domain mutant achieved a full rescue, the G/FS domain mutant appeared to only achieve a partial rescue. Since the G/FS domain has been proposed to determine the client specificity for another Hsp40 Sis1, I hypothesized that a complete rescue was blocked by other Hsp40s that might share some substrate specificity at INQ. In addition to specific domains, Hsp40s bind peptides through hydrophobic clefts spanning the protein. To determine whether these clefts are essential for INQ turnover, I utilized point mutants where the residues were mutated to serine (Sahi et al., 2013). Out of the six constructs that were transformed, four of them, namely I179S, L198S, L200S, and L277S failed to rescue INQ foci levels (Figure 2.6c). Moreover, the L200S construct had almost no change in foci compared to apj1Δ cells transformed with an empty vector, indicating that hydrophobic cleft residues have a critical role in peptide binding and therefore in controlling the Apj1 interactome at INQ. Building on the notion that chaperones regulate INQ turnover, I tested Rpd3 foci levels in a Cdc48 ts allele cdc48-4601 that was previously used in our lab to test regulation of the splicing protein Hsh155 at INQ. Similar to Apj1, I noted spontaneous INQ formation for Rpd3 at a non-permissive temperature of 30°C (Figure 2.7a). Additionally, treating these cells with MMS only exacerbated the phenotype resulting in a dramatic increase in foci levels compared to the WT. 67 Figure 2.7 Cdc48 and nucleophagy regulate INQ turnover. a Quantification of Rpd3-GFP foci levels in a cdc48-4601 ts allele at a non-permissive temperature of 30°C. b Quantification of INQ and CytoQ foci for Rpd3-GFP in nucleophagy-deficient mutants. Mean percentage values of 2/3 replicates are shown below each dataset. All error bars represent ± SEM, n=2/3, >100 cells each. ****, p<0.0001, Fisher’s test. 2.3.4.2 Nucleophagy affects INQ turnover. With respect to its location in the nucleus, INQ has been found to localize to the nucleolar-vacuolar (NVJ) junction beside the nucleolus (Sontag et al., 2023). The localization itself suggests an interesting fate for INQ wherein the NVJ is invaginated into a lytic vacuole present in the cytoplasm for degradation of misfolded proteins. Since the NVJ is adjacent to the nucleolus, a hub for rDNA and ribosome biogenesis, studies have shown that rDNA is extruded out of the nucleolus before nucleophagy indicating that this process is tightly controlled and regulated (Mostofa et al., 2018). The sites are marked by the nuclear membrane protein NVJ1 68 and the vacuolar protein VAC8 both of which are essential for nucleophagy. Additionally, the transmembrane protein ATG39 acts as a nucleophagy receptor that controls the extrusion of the nucleus at the NVJ (Papandreou & Tavernarakis, 2019). Since Rpd3 is found at both INQ and JUNQ/CytoQ, I wondered whether nucleophagy plays any role in Rpd3 turnover from INQ. To this end, I monitored Rpd3 localization after MMS exposure in cells lacking Nvj1, Vac8 and Atg39. Loss of NVJ1 resulted in increased INQ levels for Rpd3 while simultaneously reducing CytoQ levels suggesting that Rpd3 is no longer able to relocate to CytoQ foci and is stuck at INQ (Figure 2.7b). This further supports the hypothesis that proteins first accumulate at INQ and then CytoQ. At first, this sequential sequestration was thought to be due to compartments reaching their maximum capacity, and therefore sequestration was targeted to a different site. However, my data suggests this translocation could be through nucleophagy and might be a normal route for degradation. Surprisingly, vac8Δ cells had reduced formation for CytoQ while INQ levels were unaffected compared to WT cells (Figure 2.7b). While this result needs to be investigated further, it might suggest an important role for the localization of the vacuole, JUNQ and therefore “marking” of INQ in its formation. Interestingly, loss of ATG39 increased both INQ and CytoQ levels indicating a constant role for nucleophagy in degrading misfolded proteins at both sites. The choices that dictate protein fate at INQ between refolding, and degradation, and even degradation through nucleophagy remain elusive and might be answered upon further study. 69 2.4 Discussion Rpd3 was previously identified as a candidate INQ resident protein in a large-scale screen, but this study presents the first direct tests of Rpd3 as an INQ resident protein. I show that Rpd3, but not its normal interaction partners, becomes insoluble and INQ-associated upon replication stress. This behaviour is regulated by chaperone networks previously linked to INQ. Interestingly, Rpd3 shares a lysine deacetylase activity with another INQ protein, Hos2, both of which are known histone deacetylases (Sharma et al., 2007). Why these deacetylases relocalize to INQ during stress remains unknown. Rpd3 and Hos2 have many substrates and large-scale effects on gene expression and on genome stability (Alejandro-Osorio et al., 2009; A. Wang et al., 2002). It is possible that the dramatic gene expression reprogramming that takes place upon MMS treatment requires immediate post-transcriptional control of HDAC activity that is achieved through sequestration. This would be similar to a model we previously presented for the INQ localization of a core splicing factor, Hsh155, whose sequestration could reduce the splicing flux of ribosomal protein genes during the stress response (Mathew et al., 2017). It is also possible that roles for Rpd3 in DNA replication must be regulated during the MMS response. Rpd3 has been implicated in delaying the firing of late replicating origins (Aparicio et al., 2004), and we note spontaneous recruitment of Rpd3 to INQ in sml1Δ cells that have faster replication velocity, and early firing of late replicating origins (Poli et al., 2012). Thus, Rpd3 sequestration at INQ could help rapidly change the chromatin state at late origins, promoting recovery from MMS. Indeed, this kind of adaptive sequestration to INQ has been proposed for Mus81 and other DNA endonucleases, where the authors suggested INQ as a regulatory hub for controlling the timing of nuclease activity late in the cell cycle (Saugar et al., 2017). Additional 70 work is required to understand the functional significance and substrate selection criteria for novel INQ proteins like Rpd3, and whether all proteins at INQ are misfolded or sequestered in their active forms as a form of PQC. 71 Chapter 3: DNA damage induces SUMOylation of sHsps and regulates INQ formation. This chapter is based on an article currently under review: Kumar, A, Mathew, V., Stirling, P.C. “DNA damage induced SUMOylation of Btn2 regulates nuclear protein quality control”. (2023) This was a collaborative effort with Dr. Veena Mathew who aided in method development under the supervision of Dr. Peter Stirling. Additionally, Figure 3.5b was published in the article Mathew, V., Kumar, A., Jiang, Y.K., West, K., Tam, A.S., Stirling, P.C. Cdc48 regulates intranuclear quality control sequestration of the Hsh155 splicing factor in budding yeast. (2020) J Cell Sci 133 (23):jcs252551 where I contributed to the conception, investigation and visualization of the study. 3.1 Background Having established a system to track INQ formation and clearance through Rpd3, I sought to understand the role of PTMs in INQ regulation. Studies have revealed that ubiquitination is required for JUNQ localization but dispensable for deposition at INQ under proteolytic stress (Kaganovich et al., 2008; Miller et al., 2015). In contrast, ubiquitination appears to be an important sorting signal to INQ under replication stress for certain endogenous proteins (Gallina et al., 2015; Saugar et al., 2017). In an effort to characterize the role of other PTMs at INQ, we 72 focused on the small ubiquitin-like modifier, SUMO. Similar to ubiquitination, SUMOylation has well-documented roles in regulating protein stability, degradation and subcellular localization (Psakhye et al., 2019; Sarangi & Zhao, 2015; D. Wen et al., 2017), and previous genetic and imaging data placing yeast Smt3 (SUMO) at INQ along with Cdc48 and the SUMO-targeting ubiquitin ligase (StUbL) Slx5, implicate SUMO in INQ regulation (Gallina et al., 2015; Mathew et al., 2020). Here in Chapter 3, using the established system from Chapter 2 to monitor INQ formation and clearance using Rpd3, I explore the role of SUMOylation at INQ. I show that perturbation of SUMOylation reduces INQ formation, while polySUMOylation plays a role in INQ clearance. In an attempt to determine whether INQ proteins are SUMOylated, I observed that DNA damage induces SUMOylation of both sHsps Btn2 and Hsp42 and that SUMOylation of Btn2 occurs at its C-terminus. I further map the modified lysines on Btn2 to create a non-SUMOylatable construct that I later use in Chapter 4 to understand the role of Btn2 SUMOylation in INQ regulation. 3.2 Materials and Methods 3.2.1 Experimental Model and Subject Details Yeast (Saccharomyces cerevisiae) strains used in this study (s288c), database IDs, genotypes, primers, and plasmids used in this study are listed in Appendix 1. All strains were grown under nutrient rich YPD medium or synthetic medium lacking amino acids for auxotrophic selection unless otherwise indicated. For all MMS treatment conditions, log cells were exposed to 0.05% MMS (∼99%; Sigma) in synthetic complete (SC) medium for 2 h unless otherwise noted. For 73 growth curves and washouts, MMS was then washed out and replaced with fresh MMS-free medium. Plasmid constructs were created using the pRS316 plasmid backbone and the NotI and Kpn1 restriction sites. Btn2 CTD lysine-to-arginine mutants were created using gene blocks that were purchased from Integrated DNA Technologies. Constructs included necessary fragments to stitch a full length Btn2 into pRS316 that were used for further experiments. Primers are listed in Appendix 1. 3.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods Imaging and subsequent analysis were performed as described previously (Mathew et al., 2017). Log-phase yeast were mounted on slides pre-treated with concanavalin A, in SC growth medium. Immobilized cells were imaged using an Objective HCX PL APO 1.40 NA oil immersion 100× objective on an inverted Leica DMi8 microscope with a motorized DIC (differential interference contrast) turret (for DIC imaging) and a filter cube set for FITC/TRITC (for GFP and mCherry fluorescence imaging). The images were captured at room temperature using a scientific complementary metal oxide semiconductor (sCMOS) camera (ORCA Flash 4.0 V2; Hamamatsu), collected using MetaMorph Premier acquisition software and post processed [including gamma adjustments, counting of cells with/without foci, Z-stacking, projection of maximum intensity and foci intensity measurements] using ImageJ (National Institutes of Health). For all microscopy experiments, the significance of the differences was determined using Prism version 8 or higher (Graphpad). For foci and nuclear intensity measurements, samples were compared with t-tests or ANOVA; Prism performs F-tests for variance as part of this analysis. For comparisons of proportions, Fisher tests were used, and P-values were Holm–74 Bonferroni corrected in the event of multiple comparisons. Sample sizes and specific statistical details for each image analysis are listed in the figure legends. 3.2.3 Western Blotting and protein stability time course For western blotting, whole-cell extracts of logarithmic phase cells were prepared using trichloroacetic acid (TCA) extraction and blotted with experiment-specific antibodies. For the stability time course, overnight cultures of the indicated strains were diluted to below an OD600 nm of 0.2 and allowed to progress into the logarithmic phase before collection. The cells were treated with or without cycloheximide (CHX) at a concentration of 200 μg/ml for the indicated times (0–120 min). Final samples of 2×107 cells were collected by centrifugation, and whole-cell extracts were prepared by TCA extraction and used for immunoblotting. 3.2.4 Fractionation of the insoluble pellet Cellular fractionation was performed as described in. In brief, logarithmic cells were collected after or before MMS treatment. Frozen cells were resuspended in a fractionation buffer (100mM HEPES, 1% Triton X-100, 300mM NaCl, protease inhibitor cocktail) and lysed by bead beating. Lysates were precleared at 100g, 4℃ for 5 minutes. Supernatant (whole cell lysate) was recovered and fractionated at 16000g, at 4℃ for 10 minutes to separate into soluble (S) and insoluble (P) fractions. Equal amounts were loaded onto gels followed by immunoblotting. 75 3.2.5 Ni-NTA pulldown of SUMOylated proteins SUMO-conjugated proteins were detected by performing protein pulldowns under denaturing conditions, as described previously (Ulrich & Davies, 2009). In brief, strains of interest were transformed with a plasmid containing His-tagged SUMO (Smt3–His×7; Addgene) under the control of a copper-inducible promoter. Logarithmically growing cells were harvested at an OD600 nm of 0.6–0.8, and a total of 109 cells were collected by centrifugation (4000 rpm, 3724 g; 5 min at 4°C). Pelleted cells were washed and resuspended in 5 ml of pre-chilled water. Cells were then lysed with 800 µl of 1.85 M NaOH containing 7.5% (v/v) 2-mercaptoethanol, followed by 20 min incubation on ice. Protein precipitation was carried out by adding 800 µl 55% TCA on ice for 20 min. Precipitates were collected by centrifugation (8000 g, 20 min, 4°C) and were resuspended in 1 ml Buffer A (6 M guanidine hydrochloride, 100 mM sodium phosphate and 10 mM Tris-HCl, pH 8.0) and incubated on a rotating block at room temperature for 1 h to solubilize the precipitate completely. The resulting solution was then centrifuged (16,000 g, 10 min, 4°C), and supernatant was transferred to tubes containing 30 µl precleared Ni-NTA agarose beads (Qiagen) in the presence of 0.05% Tween-20 and 15 mM imidazole. These were incubated overnight on a rotating block at room temperature. Following binding, the beads were washed twice with Buffer A containing 0.05% Tween-20 and four times with Buffer C (8 M urea, 100 mM sodium phosphate and 10 mM Tris-HCl, pH 6.3) containing 0.05% Tween-20. Beads were centrifuged (200 g, 15 s) and supernatant was completely removed. His–SUMO conjugates on the beads were eluted by adding 30 µl HU sample buffer (8 M urea, 5% SDS, 200 mM Tris-HCl, pH 6.8, 0.1 mM EDTA, 0.5% Bromophenol Blue and 15 mg/ml dithiothreitol) and heating at 70°C for 10 min. Resultant protein extracts were subjected to standard western blotting and probed for SUMO and other proteins of interest using antibodies. 76 3.3 Results 3.3.1 Perturbation of SUMOylation reduces INQ formation. In order to test the role of SUMOylation in regulating INQ dynamics, I used and created mutants that affect different parts of the SUMOylation cascade, which follows an E1-E2-E3 system (Figure 3.1). Figure 3.1 Schematic for the E1, E2, E3 cascade mediated SUMOylation. I used a ts (temperature sensitive) allele of the only yeast SUMO E2 conjugase (UBC9) under non-permissive temperatures and found that the ubc9-1 allele reduced INQ formation for Rpd3 in MMS treated cells (Figure 3.2a). Interestingly, growing cells at 25℃ itself, lowers INQ levels suggesting that a temperature-based rewiring of the transcriptional landscape might affect chaperone induction and therefore sequestration. The ubc9-1 phenotype indicated an important role for SUMOylation in INQ formation. I explored this further by creating a panel of E3 deletions owing to their redundancy between substrates. While siz1Δ and siz2Δ caused no change, cells harboring the mms21-CH mutation, both on their own or combined with siz1Δ or siz2Δ caused a reduction in INQ foci (Fig. 3b). Strikingly, reduction in INQ formation in these 77 mutants coincided with an increase in cytoplasmic inclusion formation, suggesting that SUMOylation could be an important INQ sorting signal (Figure 3.2b). Figure 3.2 Mutations in the SUMOylation cascade reduce INQ formation. a Rpd3 foci levels in a temperature sensitive allele of the SUMO E2 conjugase Ubc9 at the indicated temperatures in MMS. b Rpd3-GFP foci in nucleus and cytoplasm in strains with the indicated SUMO E3 ligase mutations in MMS treated cells. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. In yeast, mono-SUMOylation is essential while polySUMO chains are not. We can separate their functions by working in an Smt3-3KR strain where lysines responsible for polySUMOylation are mutated to arginine (Horigome et al., 2016). Inhibiting this process led to an increase in INQ formation under MMS treatment suggesting a role for polySUMOylation in INQ clearance rather 78 than formation (Figure 3.3a). DNA damage is known to induce a wave of SUMOylation that aids in DNA repair and stress response (Cremona et al., 2012; Psakhye & Jentsch, 2012). Indeed, we saw that DNA damage also induced SUMOylation of the pellet and this coincided with increased ubiquitination, including protein degradation associated-K48 ubiquitin chains, in MMS treated cells (Figure 3.3b). Since Smt3-3KR had more INQ foci indicating a role in clearance, we retested fractionation in these cells and found a slight reduction in K48-Ub chains and more SUMOylated proteins in the insoluble fraction again supporting the role for polySUMOylation in degradation of proteins at INQ (Figure 3.3b). In conclusion, mutations perturbing global mono-SUMOylation reduced INQ formation for Rpd3 under replication stress. While combining E3 ligase mutations and deletions, especially containing the mms21-CH allele, caused a reduction in INQ formation, they concomitantly caused an increase in CytoQ foci indicating a role for SUMOylation as a sorting signal alongside ubiquitination between sequestration compartments. Furthermore, while SUMOylation might be necessary for INQ formation, polySUMOylation might have a role in INQ clearance hinting towards a complex stepwise role for SUMOylation at INQ. 79 Figure 3.3 PolySUMOylation is involved in INQ turnover. a Rpd3-GFP foci levels when blocking polySUMOylation using the Smt3-3KR allele. b Fractionation in WT and Smt3-3KR cells with and without MMS reveals a wave of SUMOylation and ubiquitination upon DNA damage. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. 3.3.2 DNA damage induces SUMOylation of sequestrase chaperones. My results indicated that SUMOylation controls INQ deposition, suggesting that DNA damage might also induce SUMOylation of proteins that reside at INQ. Previous direct tests of INQ substrates Hsh155 or Mrc1 under MMS treatment did not detect any modification (Colding et al., 2021; Mathew et al., 2020). We sought to expand the set of INQ clients in the hopes of finding 80 SUMO modifications in a more comprehensive manner. I chose five substrates, the splicing factors Hsh155 and Cdc40, Rpd3 and Hos2, and Cmr1, and five chaperones, sHsps Btn2 and Hsp42, Apj1, Hsp104 and Cdc48 (Figure 3.4). To ensure uniformity between studies detailing INQ formation for these proteins, I chose GFP fusions for this mini-screen. Additionally, I used DNA damage-dependent SUMOylation of Rfa1 as a positive control for both DNA damage and a successful pulldown of purified SUMO proteins using the established Smt3-Hisx7 nickel bead denaturing pulldown (Mathew et al., 2020; Ulrich & Davies, 2009). Figure 3.4 Schematic of the SUMOylation mini-screen utilizing GFP-constructs of substrates and chaperones found at INQ. While MMS treatment caused clear mono-, di- and tri-SUMOylation of Rfa1 in cells expressing the Smt3-Hisx7 fusion, I found that the five substrates tested were not SUMOylated (Figure 3.5a, b). This was surprising because I had hypothesized that DNA damage induces 81 SUMOylation of INQ client proteins and sequestrases traffic these SUMOylated proteins through their SUMO-interaction motifs (SIMs) to INQ. However, repeating these denaturing purifications with higher concentrations of lysates, beads and antibody did not reveal any evidence of SUMOylation. While this indicates that I could not establish SUMOylation of client proteins, it could be that my method is not sensitive enough to capture these transient modifications. Repeating the search for common PTMs in conditions where specific proteases like Ulp2 are mutated might yield a higher baseline for SUMOylation of modified proteins thereby increasing sensitivity of the purification system. Figure 3.5 INQ substrates are not SUMOylated upon replicative stress. a, b SUMOylation-specific denaturing nickel bead pulldowns (NiPDs) of INQ substrates as listed in Figure 3.4. 82 On the contrary, DNA damage induced clear SUMOylation of both sHsp sequestrases Btn2 and Hsp42 but none of the other chaperones (Figure 3.6a, b). Btn2 functions as an INQ-specific sequestrase whereas Hsp42 also dictates CytoQ formation (Miller et al., 2015). To try to understand the impact of sequestrase SUMOylation on INQ specifically, I focused our follow up studies on mapping and testing the function of Btn2 SUMO sites rather than Hsp42 which has more pleiotropic effects on the cell. To first confirm the result, I used the ubc9-1 ts allele and found that Btn2 could not be SUMOylated when the E2 conjugase was mutated (Figure 3.6c). To date, genetic data indicates that Btn2 is important to drive INQ formation and I hypothesize that SUMOylation could either support or oppose this role. To test this, I needed to create alleles of Btn2 with reduced SUMOylation following DNA damage. Figure 3.6 DNA damage induces SUMOylation of sHsps Btn2 and Hsp42. a SUMOylation-specific denaturing nickel bead pulldowns (NiPDs) of INQ chaperones respectively, as listed in Figure 3.4. b MMS treatment induces SUMOylation of Btn2 and 83 Hsp42. Shown is a NiPD of His-SUMO conjugates in Hsp42-GFP and GFP-Btn2 backgrounds and western blots for GFP, PGK or RFA1. c Mutating the E2 SUMO conjugase using the ubc9-1 temperature sensitive allele at non-permissive temperatures removes Btn2 SUMOylation. NiPD was performed at 30℃. 3.3.3 Identification of the SUMOylated Btn2 domain Similar to other small-heat shock proteins, Btn2 contains a characteristic alpha-crystallin domain flanked by N-terminal and C-terminal domains. The NTD contains the nuclear localization signal and is also responsible for Sis1 (and therefore Hsp104) recruitment while the CTD is highly disordered and appears to be involved in INQ formation through a mechanism that is unclear (Ho et al., 2019). In order to study these protein regions in the context of the SUMOylated domain, I created three constructs: wt (BTN2 full length), ctdΔ (C-terminal domain deleted), and ctd+cldΔ (only the NTD) (Fig. 4d). The experimental design is represented in Figure 3.7 where a tick indicates SUMOylation. Figure 3.7 Schematic and NiPDs of GFP-Btn2 constructs used to determine the SUMOylated region. 84 To interpret the results, the table should be read column-wise whereby SUMOylation of all three constructs would mean the modified domain is the NTD. Similarly, if only the wt and ctdΔ constructs are modified, the CLD is the SUMOylated domain and lastly, if only the full length wt construct is modified, Btn2 is SUMOylated at its CTD. Using pulldown of Smt3-Hisx7, only the full-length wt construct was SUMOylated upon DNA damage indicating that SUMOylation likely occurred within the CTD (Figure 3.8). Owing to the high background in the replicate shown in Figure 3.8a, this experiment was further repeated, and another replicate shown in Figure 3.8b for confirmation. Figure 3.8 Btn2 is SUMOylated at its C-terminal domain. a, b NiPDs of GFP-Btn2 constructs used to determine the SUMOylated region reveal CTD as the SUMOylated domain. * Represents non-specific banding pattern as noted in all wells. 85 I also fused a GFP at the N-terminus since an N-terminal GFP fusion for Btn2 appeared more functional compared to the C-terminal fusion based on the literature (Dubreuil et al., 2019). Indeed, GFP-Btn2 was also nuclear in the wt construct and formed spontaneous nuclear inclusions that appeared to be INQ (Figure 3.9a). This full-length construct also rescued Rpd3 sequestration to INQ in a btn2Δ background indicating that the construct was functional (Figure 3.9b). Figure 3.9 Rescue experiments in a btn2Δ background using different domain constructs. a, b Rpd3 INQ foci rescue experiments in btn2Δ using GFP-Btn2 domain constructs. Representative images are shown and quantified below. All error bars represent ± SEM, n=3, >100 cells each. ****, ns, p>0.1, Fisher’s test. Scale bars: 5µm. 86 Consistent with literature, the ctdΔ failed to rescue INQ formation for Rpd3 suggesting an functional role in sequestration (Ho et al., 2019). However, this could be due to mislocalization of Btn2 since the GFP-Btn2 ctdΔ also formed CytoQ inclusions (Figure 3.9a). Lastly, similar to ctdΔ, the ctd+cldΔ construct also failed to rescue INQ formation which was consistent with the requirement of an ACD to bind substrates. Since the alpha-crystallin domain is deleted in the ctd+cldΔ construct, Btn2 failed to form any nuclear inclusions (Figure 3.9a, b). These data support key roles for the chaperone domain and CTD of Btn2 and implicate the CTD as a candidate for MMS-induced SUMOylation. 3.3.4 Mapping the modified lysines in the Btn2 C-terminal domain. To reaffirm CTD SUMOylation, I mutated all CTD lysines to arginine to abolish SUMOylation (CTD-KallR). Consistent with the hypothesis, the wt construct was SUMOylated under DNA damage while the CTD-KallR was not (Figure 3.10). Since the CTD-KallR was highly mutated (26 lysines mutated to arginine), I sought to create a minimally mutated Btn2 to test the function of SUMOylation. To this extent, I used the SUMO-prediction software JASSA to create four different constructs of the CTD where nearby lysines were mutated to arginine to avoid jumping of the modification (Beauclair et al., 2015; Klug et al., 2013). This approach created the following constructs: 2KR (predicted site - K234), 6KR (predicted site - K249), 6KR-II (predicted site - K290), and a 12KR construct (no high confidence predictions in this region (Figure 3.11a). 87 Figure 3.10 Mutating all CTD lysines to arginine abolishes Btn2 SUMOylation. Denaturing NiPD of CTD-KallR, where all lysines are mutated to arginine, reveals loss of Btn2 SUMOylation when compared to wild type. I repeated the nickel bead pulldowns under MMS to reveal that out of the four constructs created, only Btn2-6KR showed significant loss of Smt3 signal (Figure 3.11b). The SUMOylation status of the remaining constructs, Btn2-2KR, -6KR-II, and 12KR was either unaffected or milder than wt. For example, Btn2-2KR appeared to have lower levels of SUMOylation compared to wt, but since the input protein levels for each construct were unaffected, the reduction in SUMOylation could be attributed to efficiency of the pulldown itself. I also noticed a slower migrating band above Btn2 SUMOylation which persisted even in Btn2-6KR. However, since this does not appear in previous NiPDs, it could be a feature of GFP-Btn2 plasmids themselves. 88 Figure 3.11 Btn2 SUMOylation is lost in a Btn2-6KR construct. a Schematic of Btn2 CTD showing the location of each group of lysine mutations. Neighboring lysines were mutated to arginine together to avoid jumping of the modification. b Testing Btn2 CTD lysine-to-arginine constructs reveals Btn2-6KR SUMOylation is significantly reduced upon DNA damage. 89 Having confirmed loss of SUMOylation in Btn2-6KR, I sought to compare its induction and clearance under DNA damage with Btn2-WT. To this extent, I treated both strains with an MMS for a short period of 30 minutes, washed the cells into MMS-free media and collected cells at different time points. Western blots for GFP-Btn2 revealed that the induction of Btn2, and clearance over time in a Btn2-6KR background appeared similar to Btn2-WT. Furthermore, I treated both Btn2-WT and Btn2-6KR with the translation inhibitor, cycloheximide (CHX), to study changes in protein stability. Pulse chase CHX experiments also showed no significant difference between the two. At this point, all experiments were performed in a plasmid-based system. However, imaging these plasmids produced lots of variation and did not yield any clear results. Figure 3.12 Induction, clearance, and stability of Btn2-6KR appears similar to Btn2-WT. a Cells were treated with MMS for 30 minutes followed by washout into MMS-free media and collected at various time points. b Pulse chase experiments were performed after treating cells with cycloheximide (CHX) to halt translation and collected at various time points. Cells from both a and b lysed and probed for GFP and PGK1 through western blots. 90 To avoid potential pitfalls with a plasmid system for genetic studies, I introduced these mutations to the genome to create either Btn2-WT or Btn2-6KR with a URA3 selection marker upstream of Btn2. Loss of SUMOylation was reassessed and confirmed after integration (Figure 3.13). In summary, I found DNA damage induced SUMOylation of sHsps Btn2 and Hsp42. Using a domain-specific approach, I found that Btn2 was SUMOylated at its CTD in the K249 region, a domain that appears to be important for INQ formation. Figure 3.13 Btn2 SUMOylation is still lost in Btn2-6KR when integrated into the genome. Shown is a NiPD of Btn2-WT and Btn2-6KR constructs that were integrated into the endogenous Btn2 locus. 91 3.4 Discussion The potential role of SUMO modification at INQ was first identified in 2015 with the discovery of Smt3 localization to INQ during stress (Gallina et al., 2015). SUMO modifications are known to alter the solubility of target proteins and are increasingly appreciated as regulators of membraneless phase-separated compartments (Reviewed in (Keiten-Schmitz et al., 2021)). Surprisingly, I found that it is small heat shock proteins, not INQ client proteins, that were SUMO modified in our screen. Our initial hypothesis was that DNA damage induces SUMOylation of INQ clients thereby giving one explanation for the unique set of proteins sequestered to INQ. However, the discovery that none of the INQ substrates tested are SUMOylated disproves it. While I did not test all proteins found at INQ, we assumed that 10 proteins out of the 37 found at INQ is a good representation of the INQ proteome. Moreover, DNA damage induced SUMOylation of only the two sHsps Btn2 and Hsp42 and not the other chaperones suggesting functional importance in INQ regulation. Interestingly, perturbation of SUMOylation using E3 ligase mutations reduced INQ formation but consequently increased CytoQ foci. This raises the possibility of SUMOylation as a sorting signal for proteins being sequestered to these compartments, similar to ubiquitin as a sorting factor between JUNQ and IPOD for cytoplasmic misfolded proteins (Kaganovich et al., 2008). However, since I did not find any substrates to be modified, sHsp modification by SUMO could dictate the deposition site instead. While future experiments should focus on this aspect of deposition choice, one must note that mutating enzymes in the SUMO cascade is much like brute force as it affects SUMOylation throughout the cell, affecting a multitude of proteins equally. It 92 is therefore ideal to create minimally mutated constructs of the modified proteins to test SUMOylation-specific protein roles. On the other hand, blocking polySUMOylation caused an increase in Rpd3 INQ foci. PolySUMO chains have well documented roles in protein degradation by recruiting chaperones like Cdc48/VCP/p97 or StUbLs such as Slx5/8 and Uls1 that target them to the proteasome (H. G. Lee et al., 2023; Uzunova et al., 2007). Indeed, the increase in INQ levels suggests impaired clearance and introduces the dual nature of SUMO at INQ wherein reduction of SUMOylation reduces INQ formation but polySUMOylation aids in INQ clearance. Finally, I map the modified domain and lysines on Btn2 and find that Btn2 is modified in its CTD at the 6KR region. Creation of a Btn2-6KR construct wherein the modified lysines are mutated to arginine significantly reduced SUMOylation and can be used in future experiments to test the role of Btn2 SUMOylation in regulating INQ dynamics be it formation or clearance. 93 Chapter 4: Btn2 SUMOylation regulates INQ clearance. This chapter is based on an article currently under review: Kumar, A, Mathew, V., Stirling, P.C. “DNA damage induced SUMOylation of Btn2 regulates nuclear protein quality control”. (2023) This was a collaborative effort with Dr. Veena Mathew who aided in method development under the supervision of Dr. Peter Stirling. 4.1 Background Post-translational modifications (PTMs) have ubiquitous roles in regulating cellular processes. Generally reversible, these covalent modifications can significantly affect the structure and therefore activity of proteins. Moreover, PTMs can change protein interactions, affect protein stability, solubility, and even localization thereby affecting the entire lifespan of a protein. To expand on the knowledge of PTMs and their effect on INQ biology, I explored the role of the PTM SUMO in Chapter 3. I found that DNA damage induces SUMOylation of the sHsp Btn2 and created a SUMO-deficient Btn2-6KR construct to test the role of Btn2 SUMOylation at INQ. Since deleting BTN2 abolished INQ formation, and DNA damage caused INQ formation, I reasoned that DNA damage induced SUMOylation of Btn2 might play an essential role in controlling INQ formation. To this extent, in Chapter 4, I characterize the SUMO-deficient Btn2-6KR construct and find that loss of SUMOylation does not affect Btn2 induction, clearance or stability. Additionally, Btn2-94 6KR performed similar to Btn2-WT under DNA damage and among the properties tested, loss of SUMOylation only affected Btn2 solubility. Surprisingly, INQ levels were significantly higher for multiple substrates tested in a Btn2-6KR background indicating that while SUMOylation might be important for INQ formation, SUMOylation of Btn2 specifically is dispensable. To explain the gain-of-function phenotype in Btn2-6KR cells, I perform epistasis experiments and show that Btn2 SUMOylation is parallel to Apj1-mediated degradation but epistatic with Hsp104-dependent refolding at INQ. Btn2 SUMOylation opposes K48-ubiquitin chain formation deposited by the StUbL Slx5/8 when the Apj1 pathway is blocked, irrespective of Hsp104 status. This work reveals a previously unknown Btn2 PTM and its role in regulating the fate of INQ inclusions. This places Btn2 at the nexus of protein refolding and degradation at INQ, delineating the complexity of PQC at sequestration sites. 4.2 Material and Methods 4.2.1 Experimental model and Subject details Yeast (Saccharomyces cerevisiae) strains used in this study (s288c), database IDs, genotypes, primers, and plasmids used in this study are listed in Appendix 1. All strains were grown under nutrient rich YPD medium or synthetic medium lacking amino acids for auxotrophic selection unless otherwise indicated. For all MMS treatment conditions, log cells were exposed to 0.05% MMS (∼99%; Sigma) in synthetic complete (SC) medium for 2 h unless otherwise noted. For growth curves and washouts, MMS was then washed out and replaced with fresh MMS-free medium. Plasmid constructs were created using the pRS316 plasmid backbone and the NotI and Kpn1 restriction sites. Btn2 CTD lysine-to-arginine mutants were created using gene blocks that were purchased from Integrated DNA Technologies. Constructs included necessary fragments to 95 stitch a full length Btn2 into pRS316 that were used for further experiments. Primers are listed in Appendix 1. 4.2.2 Live-cell imaging, image acquisition, analysis, and statistical methods Imaging and subsequent analysis were performed as described previously (Mathew et al., 2017). Log-phase yeast were mounted on slides pre-treated with concanavalin A, in SC growth medium. Immobilized cells were imaged using an Objective HCX PL APO 1.40 NA oil immersion 100× objective on an inverted Leica DMi8 microscope with a motorized DIC (differential interference contrast) turret (for DIC imaging) and a filter cube set for FITC/TRITC (for GFP and mCherry fluorescence imaging). The images were captured at room temperature using a scientific complementary metal oxide semiconductor (sCMOS) camera (ORCA Flash 4.0 V2; Hamamatsu), collected using MetaMorph Premier acquisition software and post processed [including gamma adjustments, counting of cells with/without foci, Z-stacking, projection of maximum intensity and foci intensity measurements] using ImageJ (National Institutes of Health). For all microscopy experiments, the significance of the differences was determined using Prism version 8 or higher (Graphpad). For foci and nuclear intensity measurements, samples were compared with t-tests or ANOVA; Prism performs F-tests for variance as part of this analysis. For comparisons of proportions, Fisher tests were used, and P-values were Holm–Bonferroni corrected in the event of multiple comparisons. Sample sizes and specific statistical details for each image analysis are listed in the figure legends. 96 4.2.3 Western Blotting and protein stability time course For western blotting, whole-cell extracts of logarithmic phase cells were prepared using trichloroacetic acid (TCA) extraction and blotted with experiment-specific antibodies. For the stability time course, overnight cultures of the indicated strains were diluted to below an OD600 nm of 0.2 and allowed to progress into the logarithmic phase before collection. The cells were treated with or without cycloheximide (CHX) at a concentration of 200 μg/ml for the indicated times (0–120 min). Final samples of 2×107 cells were collected by centrifugation, and whole-cell extracts were prepared by TCA extraction and used for immunoblotting. 4.2.4 Fractionation of the insoluble pellet Cellular fractionation was performed as described in (den Brave et al., 2020). In brief, logarithmic cells were collected after or before MMS treatment. Frozen cells were resuspended in a fractionation buffer (100mM HEPES, 1% Triton X-100, 300mM NaCl, protease inhibitor cocktail) and lysed by bead beating. Lysates were precleared at 100g, 4℃ for 5 minutes. Supernatant (whole cell lysate) was recovered and fractionated at 16000g, at 4℃ for 10 minutes to separate into soluble (S) and insoluble (P) fractions. Equal amounts were loaded onto gels followed by immunoblotting. 4.3 Results 4.3.1 Non-SUMOylatable Btn2 is a functional sequestrase. Having engineered Btn2-6KR as an allele with minimal residual SUMOylation I sought to characterize its effects on INQ biology. Since SUMOylation occurred upon DNA damage, I first tested growth under varying MMS concentrations and temperatures but observed no major 97 difference between Btn2-WT and Btn2-6KR (Figure 4.1a). I noticed that a combination of growth at 25℃ and 0.01% MMS together slowed down growth for all the strains tested. Since the images for all plates were taken on Day 2 after pinning, the slow growth was also included. However, since I noticed no differences between BY4741, Btn2-WT and Btn2-6KR, I concluded that a strain carrying a non-SUMOylatable Btn2 does not gain MMS or temperature sensitivity. I also tested growth rates for these strains after recovery from a 2hr exposure to MMS. Similar to the spot assays, Btn2-WT and Btn2-6KR had similar recovery growth rates (Figure 4.1b). 98 Figure 4.1 Loss of SUMOylation does not induce MMS sensitivity or growth defects. a Spot dilution assay comparing Btn2-WT and Btn2-6KR growth at 25°C, 30°C, 34°C, and 37°C combined with varying MMS concentrations of 0.001%, 0.005% and 0.01%. Images were taken after two days. b Growth curves analysis of Btn2-WT and Btn2-6KR cells after 2 h of MMS treatment followed by washout into MMS-free media compared with untreated cells. 4.3.2 Btn2 localization is unaffected by its SUMOylation status. Next, I focused on Btn2 localization. SUMOylation has been shown to govern localization for various proteins such that mutation of the modified lysine causes mislocalization to a different subcellular compartment (D. Wen et al., 2017). Using DAPI as a marker for DNA, I found that both Btn2-WT and Btn2-6KR localize to the nucleus and therefore SUMOylation status does not affect Btn2 localization (Figure 4.2a). Interestingly, Btn2-WT formed spontaneous inclusions in the nucleus which have been noted as INQ in another study (Miller et al., 2015). As explained in Section 1.2.3.1.2, sHsps form substructures containing an outer shell and an inner stable core (Figure 1.5b) indicating that a focus of Btn2 probably contains hundreds to thousands of Btn2 molecules in these foci, as opposed to substrates like Rpd3 that do not form INQ foci until stressed. Since these cells are unstressed, it introduces the possibility of Btn2 marking the INQ deposition site, acting as a landmark for trafficking and deposition of misfolded proteins. Surprisingly, GFPBtn2-6KR formed more spontaneous inclusions compared to Btn2-WT indicating a possible role for SUMOylation in regulating the presence of Btn2 at INQ (Figure 4.2b). 99 Figure 4.2 Btn2 localizes to the nucleus irrespective of its SUMOylation status. a Representative images for ethanol-fixed DAPI-stained Btn2-WT and Btn2-6KR cells Scale bars: 5µm. b Quantification of spontaneous GFPBtn2 foci in the nucleus in untreated cells. Error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, Fisher’s test. 4.3.3 Probing the role of Btn2 SUMOylation in its degradation. SUMOylation is also directly linked to protein degradation as it can act as a signal for ATPases such as Cdc48 that target SUMOylated proteins for degradation (H. G. Lee et al., 2023). Moreover, SUMOylation can serve as a platform for StUbLs that can ubiquitinate SUMOylated proteins for degradation (Psakhye et al., 2019; Talhaoui et al., 2018). To test whether the SUMOylation status of Btn2 affects its stability, I repeated the cycloheximide (CHX) chase experiment performed in Figure 3.12b, but with the genomically integrated Btn2-WT and Btn2-6KR strains. Additionally, I combined the CHX chase with fractionation to test whether SUMOylation affects protein stability in the pellet versus the soluble fraction. 100 Figure 4.3 Btn2 SUMOylation might regulate its degradation. a GFP-Btn2 stability and solubility with or without the 6KR mutation following cycloheximide chase. Pulse chase experiments were performed after treating cells with cycloheximide to halt translation and collected at various time points. b, c NiPD of GFP-Btn2 cells in WT vs proteasome defective cim3-1 ts allele at two non-permissive temperatures of 30°C and 37°C respectively. All cells were treated with MMS. SUMO-defective Btn2-6KR appeared to have a similar stability to Btn2-WT (Figure 4.3a). This was recapitulated in the soluble and pellet fraction with Btn2 degrading at the same rate. To my surprise, CHX treatment removed Btn2 from the pellet within 30 minutes of treatment which might explain why our previous study reported loss of INQ formation in cells treated with both MMS and CHX at the same time (Mathew et al., 2017). This makes it difficult to study INQ protein half-lives and stability using CHX. One explanation for the rapid dissolution of INQ and removal of Btn2 from the insoluble fraction (both of which could be related) due to CHX 101 treatment could be that halting translation frees up chaperones like Hsp70s that can together act on INQ and other inclusions for refolding or degradation. However, to further interrogate Btn2 stability, I performed isolation of SUMO-modified Btn2 in a cim3-1 proteasome-defective mutant at non-permissive temperatures of 30℃ and 37℃. This analysis showed a mild change in SUMO-modified Btn2 at 37℃ while having no change in Btn2 input levels. While this change was modest compared to stabilization of SUMO modified forms of other proteins found in the literature (Psakhye et al., 2019), it suggested that the small fraction of Btn2 that is SUMOylated could be associated with proteasomal degradation (Figure 4.3b,c). 4.3.4 Loss of SUMOylation affects Btn2 solubility. While fractionating cells in the CHX chase experiments above, I noticed that Btn2-6KR was more abundant in the insoluble fraction than Btn2-WT (Figure 4.4). Loss of SUMOylation appeared to make Btn2 more insoluble, consistent with the role of SUMO as a solubility tag (Butt et al., 2005; Liebelt et al., 2019; Sarangi et al., 2015) and could explain the increased spontaneous foci for Btn2 in Figure 4.2b. 102 Figure 4.4 SUMOylation regulates Btn2 solubility. Fractionation of untreated Btn2-WT and Btn2-6KR cells followed by GFP western blots or an anti-Pgk1 loading control. W = whole cell extract; S = supernatant; P = pellet. Fold change was quantified with respect to the insoluble pellet in a Btn2-WT background from three independent experiments. To attribute the increase in insolubility to loss of SUMOylation and not the mutations, I used alphafold2 via the colabfold notebook to predict structures for Btn2-WT and Btn2-6KR (Mirdita et al., 2022). These were fed to two softwares CamSol and Aggrescan 3D which predict protein solubility and aggregation potential per residue (Kuriata et al., 2019; Sormanni et al., 2015). When comparing Btn2-WT and Btn2-6KR in the region specific to the mutations (represented in the bin) the mutations to arginine were actually predicted to make the binned region more soluble (Figure 4.5) 103 Figure 4.5 Computationally predicted soluble regions of Btn2. CamSol and Aggrescan3D plots for Btn2-WT and Btn2-6KR. Graphs were plotted using scores provided by the respective softwares. The rectangular bin represents the Btn2-6KR mutation region. Furthermore, using FoldX, we calculated the predicted energy difference (ΔΔG) between Btn2-WT and Btn2-6KR to be -0.2kJ/mol which is far lower than ΔΔG values for destabilizing mutations of >1kJ/mol (Zambrano et al., 2015). These analyses support my contention that the mutations in Btn2-6KR did not destabilize or affect protein solubility (Figure 4.5). Rather the change in solubility could be attributed to loss of SUMOylation. Overall, non-SUMOylatable Btn2-6KR appears to be as stable as Btn2-WT to localize normally and to permit cellular fitness 104 after DNA damage and enable INQ formation. However, Btn2-6KR does appear more insoluble following DNA damage suggesting potential differences in the state of INQ or its lifetime. 4.3.5 Btn2-SUMO restrains INQ accumulation. Next, I sought to understand the role of Btn2 SUMOylation at INQ. Since Btn2-6KR was more insoluble and higher levels of spontaneous Btn2 foci, I wondered whether INQ as a compartment would also be more insoluble. To test this, I chose four substrates (Rpd3, Hos2, Hsh155, Cmr1) and four PQC components (Cdc48, Slx8, Apj1, Hsp104). I included Slx8 here as a proxy for the Slx5/8 complex that acts as a StUbL to degrade SUMOylated proteins (Mullen & Brill, 2008; Uzunova et al., 2007). In order to confirm INQ localization, all proteins were colocalized with Hos2-mCherry under MMS treatment. This analysis showed that loss of Btn2 SUMOylation in Btn2-6KR resulted in elevated sequestration of INQ substrates Rpd3, Hos2, Hsh155 and Cmr1, and PQC factors Cdc48 and Slx8 (Figure 4.6a). Fractionation for Rpd3 further supported this data, as Rpd3 was more insoluble in Btn2-6KR compared to Btn2-WT after MMS treatment (Figure 4.6b). These data suggest that SUMOylation of Btn2 is not the signal required for sequestration or INQ formation since it appears to be unaffected in Btn2-6KR. Interestingly, I found a small yet significant decrease for localization of proteins Apj1 and Hsp104, both of which have well established roles in INQ clearance (Figure 4.6a). This indicated that Btn2 SUMOylation and therefore INQ solubility might play a role in INQ clearance and recovery by affecting one or both pathways. 105 Figure 4.6 Elevated sequestration in Btn2-6KR cells. a Loss of Btn2 SUMOylation causes elevated sequestration for multiple INQ proteins. Representative images are shown below. b Fractionation of Rpd3 in both Btn2-WT/Btn2-6KR strains in untreated and MMS treated cells. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, **, p<0.002, *, p<0.03, Fisher’s test. Scale bars: 5µm. Studying sequestration compartments in yeast has widely increased our understanding of PQC pathways, however the physiological impact on fitness has remained an open question. The importance of sequestration was finally shown in 2019 where researchers showed that INQ and JUNQ formation is essential for cell growth when heavily burdened with misfolded proteins (Ho et al., 2019). They showed that cells lacking both the disaggregase HSP104 and the NEF FES1 are dependent on Btn2 and Hsp42 such that their fitness defects are further exacerbated if these sHsps are deleted. Assuming sequestration as a scale wherein blocking sequestration in 106 fes1Δhsp104Δ cells results in cell death, I hypothesized that elevated sequestration should be on the opposite side of the spectrum, and therefore rescue the growth defects. Remarkably, and consistent with a pro-survival role for INQ, the elevated levels of INQ in Btn2-6KR actually correlated with partial rescue of temperature sensitivity in fes1Δhsp104Δ (Figure 4.7). Thus, a hyper-sequestration phenotype wherein loss of Btn2 SUMOylation results in elevated sequestration for INQ substrates possibly due to inefficient INQ dissolution appears to be beneficial in chaperone-deficient genetic backgrounds. Figure 4.7 Temperature sensitivity of fes1Δhsp104Δ cells is rescued by Btn2-6KR. Btn2-6KR growth effects in a fes1Δhsp104Δ background as seen from a spot dilution assay. Equal ODs of the indicated strains were serially diluted and spotted on YPD at 25°C, 30°C, 34°C and 37°C. 4.3.6 Btn2 SUMOylation mediates INQ clearance. Elevated sequestration in a Btn2-6KR background was not associated with an increase in protein levels for proteins tested such as Rpd3 and Btn2 (Figure 4.4, 4.6b). This suggested that Btn2 SUMOylation could either be involved in INQ clearance or could restrict sequestrase activity, such that the loss of the modification could improve sequestration. The latter would require purification of Btn2, in vitro SUMOylation by purified SUMOylation E1, E2 and E3 enzymes 107 and addition of model misfolding substrates to test binding and holdase activity. The former hypothesis, however, could be tested using washout experiments after MMS stress in different genetic backgrounds. Two established and independent pathways that govern INQ clearance are the Hsp104-mediated protein refolding pathway and the Apj1-mediated degradation pathway (den Brave et al., 2020). I sought to determine whether Btn2-6KR-induced insolubility is epistatic with either of the two or is a third pathway mediating INQ clearance. I treated cells harboring Btn2-WT or Btn2-6KR combined with apj1Δ, hsp104Δ, or apj1Δhsp104Δ with MMS for two hours followed by a washout into MMS-free media and measured the rate of clearance for Rpd3 INQ foci. In WT cells, I noted that despite Btn2-6KR having a higher percentage of Rpd3 INQ foci than Btn2-WT, both strains successfully cleared INQ such that after two hours only 20% of cells harbored INQ (Figure 4.8). In contrast, while apj1Δ Btn2-WT cells could clear INQ, apj1Δ Btn2-6KR cells were completely defective in clearance during the time course of this experiment and instead continued to buildup Rpd3 foci (Figure 4.8). Therefore, preventing Apj1-mediated degradation and concomitantly blocking Btn2 SUMOylation resulted into a complete inability to clear INQ in the time course of this experiment (Figure 4.8). 108 Figure 4.8 Btn2-6KR is independent of Apj1-degradation and epistatic with Hsp104-refolding. a WT, apj1Δ, hsp104Δ, and apj1Δhsp104Δ cells were treated with MMS for two hours followed by a washout into MMS-free media. Shown are Rpd3-GFP foci percentages in these backgrounds after two hours of treatment and after one and two hours of washout. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. Since Apj1 and Hsp104 execute independent pathways, I hypothesized that Btn2-6KR and Hsp104 should be epistatic. Indeed, deletion of Hsp104 significantly impaired resolution of INQ foci after MMS washout but no additional defect was seen in cells expressing Btn2-6KR (Figure 4.8). As expected, deleting both Apj1 and Hsp104 in Btn2-WT cells blocked INQ clearance and further removing Btn2-SUMOylation did not exaggerate this phenotype (Figure 4.8). 109 Figure 4.9 Rpd3 INQ measurements reflect Btn2 SUMO-Hsp104 epistasis. INQ intensities were measured for strains in (A), and both sets of data reveal epistasis between Btn2 SUMOylation and Hsp104-mediated refolding while indicating that these two pathways may operate in parallel to Apj1-mediated degradation. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, ANOVA. I noted the same results when I compared Rpd3 fluorescence intensities at INQ across different genotypes (Figure 4.9). In MMS-treated Btn2-WT cells, depleting APJ1 or HSP104 individually did not affect INQ intensity suggesting that both pathways can act in parallel, accommodating for each other. Only when both pathways were blocked in apj1Δhsp104Δ Btn2-WT cells, Rpd3 intensity at INQ was significantly increased compared to WT cells. In contrast, combining Btn2-6KR with apj1Δ showed a remarkable increase in Rpd3 INQ intensity suggesting they could indeed be parallel pathways regulating Rpd3 INQ clearance. In line with that, depleting only 110 HSP104 in Btn2-6KR cells did not yield any change indicating epistasis. Similarly, apj1Δhsp104Δ Btn2-6KR cells did show an increase for Rpd3 INQ intensity, however, it was similar to apj1Δ Btn2-6KR cells. Monitoring cytoplasmic inclusion clearance revealed that while apj1Δ Btn2-6KR cells cleared these inclusions slower compared to apj1Δ Btn2-WT cells, these cytoplasmic sites were eventually cleared. Furthermore, cytoplasmic foci were also cleared in WT, hsp104Δ, and apj1Δ hsp104Δ cells regardless of Btn2 SUMOylation status indicating that the phenotypes shown in Figure 4.8 are an INQ-specific defect (Figure 4.10). Figure 4.10 Clearance of cytoplasmic inclusions is unaffected by Btn2 SUMOylation. Quantification of peripheral CytoQ foci in INQ clearance-deficient washout experiments (related to Figure 4.8). All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. 111 In addition, I also noted an increase in cells containing ≥2 cytoplasmic foci in Btn2-6KR cells regardless of chaperone status (Figure 4.11) similar to Hsp42 mutants as noted in another study (Grousl et al., 2018). Reports have independently noted the existence of both JUNQ, and smaller dispersed cytoplasmic foci termed CytoQ foci which have been suggested to coalesce to form JUNQ (Escusa-Toret et al., 2013). Since these foci are membrane-less, it suggests a process of coalescence or fusion of small inclusions into larger phase separated organelles. For multiple smaller inclusions to fuse, they need to have similar solubility, which could be mediated by both ubiquitin and SUMO. Therefore, the >2 CytoQ foci phenotype could be a result of loss of SUMOylation since it affects solubility of these inclusions that could in turn block fusion. The physiological relevance of this phenotype however remains to be investigated. Figure 4.11 Btn2-6KR cells have an increased frequency of cells with >2 CytoQ foci. Quantification of cells with greater than 1 peripheral aggregate in the strains from Figure 4.10. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. 112 In order to probe a possible role for Btn2 SUMOylation in either refolding or degradation of proteins at INQ, I utilized the GFP-tagged firefly luciferase protein construct (FFL-GFP). Reactivation of heat-aggregated FFL-GFP has been used to differentiate between protein refolding and degradation. Dissolution of FFL-GFP aggregates and therefore increase in in vivo luciferase activity after heat shock is dependent on Hsp104 while deleting degradation pathways accelerates the rate of recovery. I compared recovery of luciferase activity between Btn2-WT and Btn2-6KR and found no significant change between the two constructs (Figure 4.12). Figure 4.12 Btn2-SUMO does not affect luciferase refolding. Indicated strains were transformed with a plasmid expressing firefly luciferase (FFL) fused with GFP. Cells were subjected to heat shock at 42℃ for 20minutes to aggregate FFL-GFP. Reactivation was determined by measuring luciferase activity at different time points following heat shock. Data points plotted are relative to luciferase activity measured before heat shock. Quantification shows SEM from two independent experiments. 113 While this suggests that SUMOylation of Btn2 is not involved in either protein refolding or degradation, an obvious caveat to this experiment is the localization of FFL-GFP itself. Since it does not contain a strong NLS, FFL-GFP is diffused throughout the cell and therefore any changes could not have been solely attributed to Btn2. Additionally, I repeated the same experiment in apj1Δ, hsp104Δ, and apj1Δ hsp104Δ cells but could not obtain a consistent result. Therefore, while I could not derive concrete observations from this experiment regarding the role of Btn2-SUMO in protein refolding or degradation, I concluded that epistasis of Btn2-SUMO and Hsp104 could suggest a role in refolding running in parallel with the degradative function of Apj1. Switching focus to the ubiquitination status of the insoluble pellet in these genetic backgrounds could provide valuable information on the role of Btn2 SUMOylation in INQ clearance. 4.3.7 Btn2-SUMO regulates ubiquitin levels in sequestration compartments. Since apj1Δhsp104Δ cells had defective INQ clearance that could not be exacerbated with a non-SUMOylatable Btn2, we wondered whether there were differences in the SUMO and ubiquitin status of these inclusions. To test this, we performed cellular fractionation and probed for total SUMO, ubiquitin, the degradation specific K48-ubiquitin chains and signalling K63-ubiquitin chains. As expected, apj1Δ cells had an accumulation of total and K48-linked ubiquitinated and SUMOylated proteins and further deleting Hsp104 did not exacerbate this phenotype (Figure 4.13) reaffirming their roles in degradation and refolding respectively. 114 Figure 4.13 Btn2 SUMOylation impacts ubiquitin and SUMOylation status of sequestration inclusions. Fractionation in WT, apj1Δ, hsp104Δ, and apj1Δhsp104Δ cells treated with MMS and western blot for levels of total Ubiquitin (Ub), K48-, K63-linked Ubiquitin chains, and SUMO/Smt3 chains. Normalized quantifications were acquired from the insoluble fractions shown in the western blots, performed in duplicates. 115 Importantly, the ability to SUMOylate Btn2 impacted the ubiquitin status of the fractionated inclusions. While Btn2-6KR itself did not cause any change, apj1Δ Btn2-6KR cells had higher total, K48- and K63-linked ubiquitin compared to apj1Δ Btn2-WT cells (Figure 4.13). This was particularly interesting because it suggested a role for Btn2 SUMOylation, independent on Apj1, in protein degradation. Moreover, the increase in degradation-specific K48-linked ubiquitin chains could explain why INQ clearance is blocked in apj1Δ Btn2-6KR cells. I also noted a similar increase for K48-Ub chains in apj1Δhsp104Δ Btn2-6KR cells compared to apj1Δhsp104Δ Btn2-WT cells, strongly suggesting a role for Btn2 SUMOylation in negatively regulating the build-up of K48-Ub chains in apj1Δhsp104Δ inclusions (Figure 4.13). Contrary to K48-Ub chains which could be linked to the microscopy data studying INQ clearance in Figure 4.8, K63-Ub chains in the insoluble pellet posed a more complicated picture. While apj1Δ cells had an accumulation of K63-Ub chains compared to WT and hsp104Δ cells, combining apj1Δ and hsp104Δ brought back K63-Ub to WT levels, indicating a possible role for Hsp104 in K63-Ub buildup by refolding ubiquitinated substrates destined for degradation resulting in production of unanchored K63-Ub chains (Figure 4.13). Another explanation for the loss of K63-Ub buildup could be that when both Apj1-mediated degradation and Hsp104-dependent refolding are blocked, a third pathway could be prioritized, for example nucleophagy, that could help turnover K63-Ub chains. Surprisingly, loss of Btn2 SUMOylation in apj1Δhsp104Δ cells restored K63-Ub accumulation further supporting its role in negatively regulating ubiquitination in apj1Δhsp104Δ inclusions (Figure 4.13). 116 In support of a SUMO-dependent degradation role for Apj1, apj1Δ cells had higher Smt3 staining in the pellet. While this seemed to slightly increase in apj1Δ Btn2-6KR cells, Smt3 levels in all genetic backgrounds tested remained consistent (Figure 4.13). 4.3.8 Slx5/8 activity on the MMS-induced insoluble proteome. While the degradation-specific role of Apj1 has been established (den Brave et al., 2020), the mechanism through which it targets proteins for degradation is yet to be discovered. A negative genetic interaction with the StUbL Slx5 suggested that Apj1 might be involved in the degradation of SUMOylated substrates (Sahi et al., 2013). To this extent, the authors deleted both APJ1 and SLX5 and noted a strong increase in whole cell SUMOylation indicating parallel roles in SUMO-mediated degradation. Since it is an Hsp40, we can assume it works in concert with an Hsp70 for protein degradation. Additionally, increased ubiquitination of the MMS-induced insoluble proteome in apj1Δ cells which was further exacerbated in apj1Δ Btn2-6KR cells suggests the action of a StUbL K48-Ub ligase that targets SUMOylated proteins for degradation. Slx5/8 is a good candidate for this activity since it is a StUbL that localizes to INQ but its effect on INQ SUMOylation and ubiquitination is unknown (Fig. 5d). To investigate this, I first tested Btn2 SUMOylation in slx5Δ cells. Deleting SLX5 increased Btn2 SUMOylation, introducing visible polySUMO chains that were not observed in WT conditions, suggesting a constant action of either StUbLs or SUMO proteases at INQ maintaining polySUMOylation Btn2 (Figure 4.14a). Measuring Btn2 stability with cycloheximide chase experiments showed a slower rate of 117 degradation in slx5Δ cells indicating that Slx5/8 could serve to turnover Btn2 under normal conditions (Figure 4.14b). Figure 4.14 Btn2 SUMOylation and turnover is controlled by Slx5/8. a Deleting Slx5 causes an increase in Btn2 mono-SUMOylation and polySUMOylation. NiPD was performed in WT and slx5Δ cells treated with MMS. b Btn2 stability after a cycloheximide chase shows prolonged lifetime in cells lacking SLX5. Since both Apj1 and Slx5 can act on SUMOylated substrates, I also tested the effect of apj1Δ on Btn2 SUMOylation. Interestingly, cells lacking APJ1 also had higher Btn2 SUMOylation compared to WT cells but also had higher expression of Btn2 protein itself (Figure 4.15). The increased basal expression of Btn2 protein levels in apj1Δ cells makes it harder to interpret the increase in SUMOylation since a stronger Btn2-SUMO band could be attributed to increased protein levels that can be modified. Additionally, apj1Δ could stabilize Btn2 by preventing its degradation, thereby explaining the increase in Btn2 SUMOylation. I also used the synthetically sick apj1Δslx5Δ mutant for this experiment and surprisingly, Btn2 protein levels are restored to 118 WT levels but with an additive effect on the modification. The double mutant had elevated Btn2 SUMOylation compared to other backgrounds tested (Figure 4.15). First, this reiterated the notion that Apj1 and Slx5 are two independent SUMO-degradation pathways and second, showed that both pathways act on SUMOylated Btn2 to possibly regulate its degradation. Figure 4.15 Slx5 and Apj1 regulate Btn2 SUMOylation levels. Denaturing NiPD in WT, slx5Δ, apj1Δ, and slx5Δapj1Δ cells treated with MMS and western blots for GFP, PGK1 and RFA1. Aside from its turnover, stabilization of Btn2 SUMO forms in apj1Δ cells supported the idea that Btn2 might be involved in the recruitment of a StUbL such as Slx5/8 to INQ through SUMOylation. To test this, I monitored the localization of Slx8-GFP alongside Hos2-mCherry in WT, apj1Δ, hsp104Δ, and apj1Δhsp104Δ cells. In agreement with my previous results (Figure 4.6), recruitment of Slx8-GFP to INQ was not dependent on Btn2 SUMOylation status since I 119 saw Slx8 foci at INQ in both Btn2-WT and Btn2-6KR cells although I did note higher levels of foci in the backgrounds tested (Figure 4.16). Interestingly, deletion of APJ1 significantly increased Slx8 foci suggesting a functional significance such that the increase in pellet ubiquitination in these cells might be due to the activity of the Slx5/8 complex. Figure 4.16 Slx5/8 recruitment to INQ is not dependent on Btn2 SUMOylation. Slx8-GFP INQ foci levels in WT, apj1Δ, hsp104Δ, and apj1Δhsp104Δ cells with Btn2-WT/Btn2-6KR following MMS treatment. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. To this end, I tested ubiquitination and SUMOylation status of the pellet proteome in apj1Δ, hsp104Δ, and apj1Δhsp104Δ cells by combining it with an SLX5 deletion. Consistently, I found that accumulation of K48-Ub chains in apj1Δ and apj1Δhsp104Δ cells was dependent on Slx5 (Figure 4.17). This did not coincide with an increase in the SUMOylated fraction of the insoluble pellet indicating that contrary to previous results stating Apj1 and Slx5 are parallel 120 pathways for degradation of SUMOylated substrates, these results place Slx5/8 in the Apj1-mediated degradation pathway at INQ as the E3 ligase responsible for ubiquitinating proteins with K48-polyUb chains. Figure 4.17 Slx5 is the E3 ligase responsible for depositing K48-Ub chains in the MMS-induced insoluble proteome. 121 Fractionation in WT, apj1Δ, apj1Δslx5Δ, hsp104Δ, hsp104Δslx5Δ, apj1Δhsp104Δ, and apj1Δhsp104Δslx5Δ cells treated with MMS and western blot for levels of total Ubiquitin (Ub), K48-, K63-linked Ubiquitin chains, and SUMO/Smt3 chains. . Normalized quantifications were acquired from the insoluble fractions shown in the western blots, performed in duplicates. On the other hand, increased K63-Ub in apj1Δ cells was not lost in an apj1Δslx5Δ background supporting the idea that another E3 ligase deposits K63 chains (Figure 4.17). Furthermore, hsp104Δ and apj1Δhsp104Δ cells did not have increased K63-Ub of the MMS-induced insoluble fraction, consistent with the results for the same perturbations in a Btn2-WT background in Figure 4.13. However, further deleting SLX5 increased K63-Ub chain formation in these backgrounds indicating that Slx5-mediated K48-Ub chain deposition could oppose K63-Ub deposition by another E3 ligase at INQ. Owing to its roles in signalling pathways, K63-Ub chain formation upon loss of K48-Ub for degradation could represent a backup pathway such as nucleophagy when both degradation and refolding are blocked at INQ. To further support the hypothesis that Slx5 and Apj1 together mediate protein degradation at INQ, I performed MMS washouts in slx5Δ Btn2-WT and slx5Δ Btn2-6KR cells. Coupling slx5Δ with Btn2-6KR resulted in totally defective INQ clearance while not affecting CytoQ clearance, similar to apj1Δ Btn2-6KR cells (Figure 4.18). 122 Figure 4.18 INQ clearance is blocked in slx5Δ Btn2-6KR cells. a, b slx5Δ Btn2-WT and slx5Δ Btn2-6KR cells were treated with MMS for two hours followed by a washout into MMS-free media. Shown are quantified percentages of Rpd3-GFP INQ foci in a and peripheral CytoQ foci in b in these backgrounds after two hours of treatment and after one and two hours of washout. All error bars represent ± SEM, n=3, >100 cells each. ****, p<0.0001, ***, p<0.0002, **, p<0.002, *, p<0.03, ns, p>0.1, Fisher’s test. 4.3.9 Discussion In conclusion, I place Btn2 SUMOylation at the core of the triage decision at INQ. I show that while Btn2 SUMOylation is epistatic with Hsp104-mediated refolding, it appears to be parallel to Apj1-mediated degradation. Additionally, I utilize fractionation to show that Btn2 SUMOylation negatively regulates K48-Ubiquitin chain buildup in the MMS-induced insoluble proteome when Apj1 is absent. Furthermore, contrary to previous studies showcasing Apj1 and Slx5 to be 123 parallel pathways for degradation of SUMOylated proteins, I highlight the role of the Slx5/8 complex as the primary E3 ligase in the MMS-induced insoluble proteome in apj1Δ cells. Surprisingly, localization of Slx8-GFP in apj1Δ is unaffected of Btn2 SUMOylation status. This is intriguing as I still note increased ubiquitination in apj1Δ Btn2-6KR cells. One explanation could be that Btn2 SUMOylation acts as a primary substrate for Slx5/8, thus shielding other proteins at INQ from ubiquitination. However, upon loss of Btn2 SUMOylation, Slx5/8 can ubiquitinate substrates more freely. This could also explain why I do not notice increased ubiquitination upon MMS treatment in WT cells since Slx5/8-ubiquitinated proteins are triaged by Apj1 for degradation. While this could explain one such phenotype, additional work will be required to delineate the intricate crosstalk between the PQC pathways at INQ. 124 Chapter 5: Conclusion 5.1 Summary of data The genome, chromatin states, and transcriptome respond dynamically to stressful conditions. It is now clear that the nuclear proteome is no exception and must be protected and remodeled during times of stress. Yeast sequester endogenous nuclear proteins into the INQ compartment, especially during times of genotoxic stress. The INQ then serves as an organizing center for triage decisions leading to protein reactivation and refolding or degradation. Since only a subset of nuclear proteins seems to relocalize to the INQ following MMS treatment, I speculate based on the literature and our observations that both biophysical properties of these proteins and the regulated functional needs of the cell determine sequestration in the INQ. Interestingly, sequence and functional characteristics of the INQ proteome analyzed through bioinformatics revealed no unique features suggesting a role for a common post-translational modification (PTM) dictating sequestration to the INQ upon replication stress. The aim of my thesis was to elucidate the role of DNA damage induced SUMOylation in regulating INQ dynamics. To this end, I established Rpd3 as a novel INQ marker under DNA damage and used it to highlight the importance of SUMOylation in controlling INQ biology. Furthermore, I identified the DNA damage induced SUMOylation of sHsps Btn2 and Hsp42 and studied the role of Btn2 SUMOylation in INQ clearance. In Chapter 2, I used C-terminal GFP fusions to demonstrate that Rpd3 is sequestered to nuclear inclusions under DNA damage. Using nuclear and INQ markers, I show that these inclusions are in fact INQ sites, and that sequestration is mediated by sHsps Btn2 and Hsp42. Furthermore, I 125 establish that INQ formation for Rpd3 is not due to an artifact of the genotoxins used, but rather a general response to DNA damage. In addition, I study the role of various INQ clearance mechanisms and show that multiple chaperones govern the dynamics of proteins present at INQ. In Chapter 3, using Rpd3 as a reporter for INQ, I identified the dual nature of SUMOylation in regulating INQ biology. I find that while perturbation of global SUMOylation reduces INQ formation, blocking polySUMOylation affects INQ turnover. Importantly, I performed direct tests for SUMOylation on one-third of the currently known INQ proteome and identified the novel DNA damage induced SUMOylation of sHsps Btn2 and Hsp42. Furthermore, I mapped the modified region of Btn2 and engineered a SUMO-deficient Btn2 mutant to understand the role of Btn2 SUMOylation at INQ. In Chapter 4, I compared Btn2-WT and the SUMO-deficient Btn2-6KR to understand the role of Btn2 SUMOylation. For the biochemical characteristics tested, I show that SUMOylation controls Btn2 solubility which in turn resulted in elevated sequestration for proteins tested. When combined with INQ clearance deficient mutants, Btn2-6KR was epistatic with Hsp104-mediated refolding while being parallel to Apj1-controlled degradation. Additionally, I identified the role of Btn2 SUMOylation in opposing the buildup of K48- and K63-ubiquitin chains in these clearance deficient mutants, placing it at the crossroads of both protein degradation and refolding at INQ. Overall, the data presented in this thesis identifies targets for SUMO modification at the INQ for the first time. My data suggests that Btn2 SUMOylation is dispensable for INQ formation and 126 instead impacts solubility and recovery processes. sHsps bound to non-native client proteins and are thought to exist in a substructured system - a stable inner core with few sHsps bound to substrates and a dynamically exchanging sHsp shell that enables access of refolding or degradation machinery to substrates (Żwirowski et al., 2017). My data suggests that SUMOylation might modify the solubility or accessibility of substrate proteins sequestered by Btn2 to other PQC machinery that enables dissolution of INQ upon stress removal. Cells unable to SUMOylate Btn2 promote a less-soluble INQ state that is incapable of dissolving Rpd3 at INQ in the absence of the Apj1 co-chaperone, which has been linked to INQ protein degradation (den Brave et al., 2020). Finally, I observed that cells unable to SUMO modify Btn2 accumulate K48-linked ubiquitin modified proteins in the insoluble fraction when Apj1 was absent. K48 chains on INQ proteins only accumulate in cells with functional Slx5, a SUMO targeted E3 ubiquitin ligase with known roles in genome maintenance (Horigome et al., 2016; Thu et al., 2016). When Btn2 cannot be SUMOylated the Slx5/8 complex seems to localize in excess to INQ, loss of Apj1 reveals K48-Ub chain accumulation, and delayed recovery. This data supports a model in which either Slx5/8 is hyperactive without Btn2-SUMO, or that Slx5/8 substrate proteins at INQ cannot be extracted efficiently, slowing K48-modified substrate removal and INQ recovery (Figure 5.1). In the future it will be important to determine the function and contribution of Hsp42 SUMOylation that we discovered to this process. Since Hsp42 plays important roles at cytoplasmic quality control compartments, it is possible that SUMOylation will have more complex roles in regulating triage to these various sites under different conditions. 127 Figure 5.1 Btn2 SUMOylation regulates INQ dynamics. Briefly, in WT cells (top), Btn2 SUMOylation likely occurs after INQ formation and helps to regulate normal ubiquitination and turnover of substrates during stress recovery. In cells that cannot SUMOylate Btn2, INQ formation is unaffected (bottom left) rather Btn2 itself and substrates like Rpd3 shift more strongly to the insoluble fraction suggesting a potential change in state. Cells that further lack the degradation pathway mediated by Apj1 (bottom right) completely fail to clear INQ and exhibit hyper-accumulation of ubiquitinated proteins in the insoluble fraction. The figure was created using BioRender. 128 5.2 DNA damage causes INQ formation. Sequestration in yeast was first described by Frydman and colleagues by introducing artificially misfolding proteins such as VHL-GFP and Ubc9-ts GFP and tracking their movement to inclusions that they named JUNQ and the IPOD (Kaganovich et al., 2008). Additional work from the Bukau and Mogk group described the presence of an intranuclear sequestration site that was labelled INQ (Miller et al., 2015). While there initially were some opinions that the JUNQ and the INQ could be the same structures, recent work has confirmed that JUNQ and INQ are different entities separated by the nuclear membrane (Sontag et al., 2023). While most of what we understand about sequestration comes from these seminal papers utilizing misfolding reporters coupled with heat shock and proteasomal impairment, our lab and others have discovered the relocalization of endogenous proteins to INQ under DNA damage (Gallina et al., 2015; Mathew et al., 2017). This raises an important question. Does DNA damage cause misfolding of endogenous proteins? This is a possibility since the chemical agent MMS, that is the accepted standard drug for inducing INQ has been found to cause RNA modifications that could result in misfolding (Thomas et al., 2020). To answer this fundamental question, I took a two-step approach. First, I used multiple DNA damaging agents, each with a unique mechanism of action and found that regardless of the type of stressor, DNA damage results in INQ formation for Rpd3. Specifically, exposure to hydroxyurea (HU), a small molecule inhibitor of ribonucleotide reductase, caused INQ formation (Koç et al., 2004). Second, I used mutations in the DNA repair cascade to recapitulate the DNA damage phenotype without any external stresses. In line with our hypothesis, Rpd3 was sequestered to INQ in these conditions as well. It appears that INQ 129 formation could be a new facet of the DNA damage response, one that might be essential for cell survival when other PQC circuits are burdened. 5.3 The significance of the INQ proteome. Our understanding of the INQ resident proteome is derived from high-throughput cytological GFP screens that have revealed the sequestration of 37 proteins to INQ (Table 1.1). Considering the presence of ~6000 protein coding genes in yeast, it is remarkable that only a minor subset of proteins are transported to INQ under DNA damage. This specificity could be due to several reasons. First and foremost, it might be a caveat of the screening system itself. GFP fusions have been shown to affect protein function, could misfold the protein and be the reason for sequestration. While this might add false positives to the screens performed, fractionation probed with antibodies specific to protein and not a tag indicate that endogenous proteins do in fact become insoluble and form a part of these inclusions. Additionally, retesting localization in brighter and smaller fluorophores might reveal new INQ residents. Second, are all proteins at INQ misfolded? This fundamental question remains unanswered. Based on the function and experimental validation of how sHsps work, we know that these sequestrases bind to exposed hydrophobic regions in misfolded proteins and sequester them to site-specific compartments. However, while INQ might contain misfolded proteins, it also appears to contain inactive forms of properly folded proteins, such that their sequestration controls or rather curbs their activity. The hypothesis of INQ sequestration to regulate protein activity is an exciting idea since the cell does not have to spend extra energy to remake a subset of proteins. That said, if we think of INQ as a “storage locker” for proteins involved or necessary 130 in stress response, the INQ proteome might change with the stress used to induce its formation. For example, the 37 proteins listed in Table 1.1 are from studies using MMS as an INQ inducer. If we were to use HU or heat shock instead, the INQ proteome could be different and stress specific. Additionally, proteins at MMS-induced INQ appear to the involved in the stability of the replisome and their sequestration could be beneficial for the stress response to MMS. While this does not explain how sHsps might bind to functional proteins, it adds a tightly regulated layer to stress response and recovery in a cell. 5.4 Importance of sequestration in humans, SUMO, and PML bodies What can INQ tell us about nuclear PQC in response to DNA damage across species? SUMOylation of human proteins in the DNA damage response has been recognized for many years, for example in the regulation of BLM helicase localization to PML bodies (Eladad et al., 2005). Indeed, PML has been shown to function in nuclear PQC in a SUMO dependent manner along with the SUMO targeted ubiquitin ligase RNF4 (Guo et al., 2014). Bringing these studies together provides dramatic parallels with the yeast system I describe here. Moreover, just like the relationship between INQ and CytoQ frequency I observed, the nuclear SUMO dependent network described in humans also cross-talks with cytoplasmic stress granules (Keiten-Schmitz et al., 2020). While PML protein is not conserved, and PML bodies seem to serve multiple functions in humans, organizing principles related to PQC functions seen at INQ are clearly evident. Additionally, sequestration as a PQC response in humans has been discovered and elucidated at the aggresome (reviewed in (Chin et al., 2008)). 131 While cytoplasmic sequestration appears to be conserved, nuclear sequestration and compartmentalization of human misfolded proteins in a specific site remains to be discovered. The human nucleolus acts as a PQC hub safeguarding misfolded proteins and epigenetic regulators that functionally recover upon removal of heat shock in an HSP70-dependent manner (Azkanaz et al., 2019; Frottin et al., 2019). This is in direct relation to INQ and the sequestration of functionally active proteins upon stress. It is possible that during evolution, the capacity to form INQ and sequester proteins was acquired by the nucleolus, however this remains to be proven. Studies of humanized yeast suggest that at least human small heat shock proteins HspB2 and HspB3 have the potential to serve as sequestrases (Shrivastava et al., 2022). Moreover, HspB2 and B3 localize to the nucleus and can form phase separated droplets in human cells, supporting the potential for a conserved pathway of nuclear PQC by small heat shock proteins with sequestrase functions (Joosten et al., 2023). Future studies should help to further define key conserved features of nuclear protein quality control in response to DNA damaging stresses and the contributions of small heat shock proteins, and SUMO to this process. 5.5 Limitations and caveats While I use fractionation to support the microscopy data, the current protocol separates the whole cell lysate into a soluble and an insoluble fraction. This means that for all fractionation experiments listed in this thesis, I cannot differentiate between cytoplasmic and nuclear proteins and therefore cannot use it to hypothesize about specific sites such as INQ or JUNQ. I was unable to set up a protocol that combines both fractionation and isolation of the nucleus. However, going forward a new protocol focusing on the insoluble fraction of the nucleus coupled with mass spectrometry will help deliver insights into the INQ proteome. 132 SUMOylation as a PTM is notorious for its transient nature. Modified proteins are constantly under the action of StUbLs or SUMO proteases that cleave the SUMO off a substrate, de-SUMOylating it in the process (Sriramachandran & Dohmen, 2014). To this extent, purification of SUMO proteins is carried out in a denaturing environment where proteases are made inactive (Ulrich & Davies, 2009). While I can sufficiently pull down SUMOylated RPA in my pulldown system, mild SUMOylation of Btn2 and Hsp42 might indicate that a small fraction of the sHsps get SUMOylated but also speak to the sensitivity of the system. To improve sensitivity and efficiency of the pulldowns, these experiments can be repeated in the presence of the SUMO protease inhibitor N-ethylmaleimide (Xiao et al., 2015). Additionally, deleting Slx5, or proteases like Ulp2 might increase the SUMOylated fraction of a protein, thus increasing yield of a SUMO pulldown (Psakhye et al., 2019). 5.6 Future directions 5.6.1 Hsp42 SUMOylation Hsp42 is essential for sequestering proteins to JUNQ, and the IPOD but also to INQ under DNA damage as shown in Figure 2.1. Furthermore, Hsp42 functions similar to Btn2, in that it also contains a crystallin-like domain to bind substrates, but also contains a prion-like domain and an intrinsically disordered domain that aid in chaperone activity and stability (Grousl et al., 2018). While I explored the role of Btn2 SUMOylation at INQ, the importance of Hsp42 SUMOylation remains unanswered. It is important to note that only a small fraction of Hsp42 appears to get modified compared to the total protein level. While this could suggest that SUMOylation of Hsp42 is not imperative for function, it could also be explained by a constant action of SUMO 133 proteases. Since Hsp42 and Btn2 have similar functions and domain architecture, Hsp42 could also be modified at its CTD. Surprisingly, the Hsp42-CTD has only one lysine which could be SUMOylated. Future directions will include mutating this lysine residue to ultimately create a non-SUMOylatable Hsp42 and understanding the role of Hsp42 SUMOylation in regulating sequestration. 5.6.2 Identification of the structural component of INQ deposition As discussed earlier in this chapter, the type of stress might control the proteome that localizes to INQ. To that extent, a common structural component for the substrates found at an MMS-induced INQ remains elusive. Interestingly, WD40 repeats in the protein Cmr1 have been shown to be essential for its relocalization to INQ and these repeats are found in four other MMS-induced INQ proteins (Table 1.1; Apc4, Cdc20, Cdc40, Dug2) (Gallina et al., 2015). Moreover, the chaperonin CCT has been found to interact with WD40 repeats in the protein WDR68 to promote its folding, and importantly interact with WDR76 (the human homolog of Cmr1) via its WD40 domain, suggesting the first structural basis for INQ sequestration (Dayebgadoh et al., 2019; Miyata et al., 2014). Additionally, WD40 repeats also bind ubiquitin, which is interesting from an INQ quality control standpoint (Pashkova et al., 2010). Since SUMOylation does not appear to be the common PTM shared between INQ substrates, ubiquitin could be driving proteins near the replisome to INQ for safety. While ubiquitin has been shown to be dispensable for heat shock-induced INQ, it appears to play an important role in MMS-induced INQ sequestration, controlling relocalization of Mrc1 and Mus81 (Colding et al., 2021; Miller et al., 2015; Saugar et al., 2017). Going forward, dissecting the structural component required for INQ deposition will provide novel insights into the specificity of proteins that are sequestered under 134 different types of stress. Exploiting paralogs where one protein localizes to INQ, while the other does not might provide a simple system to test this hypothesis. 5.6.3 Why sequester Rpd3? If INQ sequestration for endogenous proteins is targeted, then sequestration of Rpd3 must be beneficial for the stress response. For example, the presence of Mus81 at INQ is determined by the phosphorylation state of Mms4 such that prematurely phosphorylating Mms4 released Mus81 from INQ (Saugar et al., 2017). Accordingly, increased recombination intermediates that need to be resolved by Mus81-Mms4 released Mus81 from INQ, connecting its sequestration to its function. Similarly, sequestration of Hsh155 is associated with lowered splicing, and release of Mrc1 from INQ is involved in replisome progression upon removal of MMS (Colding et al., 2021; Mathew et al., 2017). It is therefore conceivable that sequestration of Rpd3 might play an important role in stress response, by altering gene expression upon MMS treatment or by affecting DNA replication. Rpd3, through its HDAC activity, has been implicated in delaying the firing of late replicating origins (Aparicio et al., 2004). It is possible that Rpd3 sequestration to INQ could affect the chromatin state at late origins, allowing early firing, promoting recovery from MMS. My data on spontaneous Rpd3 INQ formation in sml1Δ further supports this data as they have faster replication velocity, and early firing of late origins (Poli et al., 2012). Additionally, no studies have characterized the role of acetylation in INQ biology. Rpd3 could be sequestered to INQ to mediate deacetylation of modified proteins, however this remains to be explored. 135 5.6.4 The role of phase separation at INQ. Liquid-liquid phase separation (LLPS) has emerged as a ubiquitous phenomenon for sequestration of DNA, RNA, and proteins to form membrane-less organelles (Banani et al., 2017). Phase separated entities can range from DNA repair foci to P-bodies and could also explain the physical nature of the sequestration compartments mentioned in this thesis (Luo et al., 2018; Oshidari et al., 2020). While I do not talk about this process nor its physiological relevance, the INQ, JUNQ and IPOD are membrane-less and specifically for JUNQ, might explain how smaller CytoQ foci could coalesce to form a larger JUNQ focus (Escusa-Toret et al., 2013). Similarly, both INQ and the IPOD which only appear as one focus, could behave like G-bodies that have gel-like characteristics (Fuller et al., 2020). While the presence of an intrinsically disordered region does not necessarily indicate phase separation (Borcherds et al., 2021), the IDRs in both Btn2 and Hsp42 could be involved in this process. Indeed, SUMOylation of these IDRs (experimentally proven for Btn2, hypothesized for Hsp42) further supports this hypothesis as SUMO itself has key roles in phase separation (Keiten-Schmitz et al., 2021). SUMOylation of purified Btn2 and Hsp42 in vitro will help interrogate the possible roles of phase separation at INQ. Ultimately, the work presented in this thesis delves into DNA damage induced sequestration and the role of the PTM SUMOylation in regulating this process. Future work focusing on the questions laid out here will advance our knowledge on protein sequestration, its physiological relevance under DNA damage and the pathways regulating the formation and dissolution of these compartments. 136 Bibliography Ajit Tamadaddi, C., & Sahi, C. (2016). 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Boeke, New York University, New York, NY Rpd3-GFP PSYL984 MATa ura3∆0 leu2∆0 met15∆0 or MET15 RPD3-GFP::HIS3 Huh et al. 2003 Rpd3-GFP, Hta2-mCherry PSYL1013 MATa ura3∆0 leu2∆0 met15∆0 or MET15 RPD3-GFP::HIS3 HTA2-mCherry::URA3 This study Rpd3-GFP, Hos2-mCherry PSYL1010 MATa ura3∆0 leu2∆0 met15∆0 or MET15 RPD3-GFP::HIS3 HOS2-mCherry::URA3 This study btn2Δ, Rpd3-GFP PSYL1255 MATa ura3∆0 leu2∆0 met15∆0 or MET15 btn2Δ::KanMX RPD3-GFP::HIS3 This study hsp42Δ, Rpd3-GFP PSYL1022 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp42Δ::KanMX RPD3-GFP::HIS3 This study Sin3-GFP PSYL1198 MATa ura3∆0 leu2∆0 met15∆0 or MET15 SIN3-GFP::HIS3 Huh et al. 2003 Ume1-GFP PSYL1199 MATa ura3∆0 leu2∆0 met15∆0 or MET15 UME1-GFP::HIS3 Huh et al. 2003 Ume6-GFP PSYL1200 MATa ura3∆0 leu2∆0 met15∆0 or MET15 UME6-GFP::HIS3 Huh et al. 2003 Sap30-GFP PSYL1201 MATa ura3∆0 leu2∆0 met15∆0 or MET15 SAP30-GFP::HIS3 Huh et al. 2003 Pho23-GFP PSYL1202 MATa ura3∆0 leu2∆0 met15∆0 or MET15 PHO23-GFP::HIS3 Huh et al. 2003 Eaf3-GFP PSYL1203 MATa ura3∆0 leu2∆0 met15∆0 or MET15 EAF3-GFP::HIS3 Huh et al. 2003 Rco1-GFP PSYL1204 MATa ura3∆0 leu2∆0 met15∆0 or MET15 RCO1-GFP::HIS3 Huh et al. 2003 Snt2-GFP PSYL1282 MATa ura3∆0 leu2∆0 met15∆0 or Huh et al. 2003 169 MET15 SNT2-GFP::HIS3 Ecm5-GFP PSYL1283 MATa ura3∆0 leu2∆0 met15∆0 or MET15 ECM5-GFP::HIS3 Huh et al. 2003 sml1Δ, Rpd3-GFP PSYL1473 MATa ura3∆0 leu2∆0 met15∆0 or MET15 sml1Δ::KanMX RPD3-GFP::HIS3 This study sml1Δ, mec1Δ, Rpd3-GFP PSYL1264 MATa ura3∆0 leu2∆0 met15∆0 or MET15 sml1Δ::KanMX mec1Δ::HygMX RPD3-GFP::HIS3 This study tel1Δ, Rpd3-GFP PSYL1475 MATa ura3∆0 leu2∆0 met15∆0 or MET15 tel1Δ::KanMX RPD3-GFP::HIS3 This study sml1Δ, mec1Δ, tel1Δ Rpd3-GFP PSYL1477 MATa ura3∆0 leu2∆0 met15∆0 or MET15 sml1Δ::KanMX mec1Δ::HygMX tel1Δ::NatMX RPD3-GFP::HIS3 This study mrc1Δ, Rpd3-GFP PSYL1394 MATalpha ura3∆0 leu2∆0 met15∆0 or MET15 mrc1Δ::KanMX RPD3-GFP::HIS3 This study tof1Δ, Rpd3-GFP PSYL1395 MATalpha ura3∆0 leu2∆0 met15∆0 or MET15 tof1Δ::KanMX RPD3-GFP::HIS3 This study csm3Δ, Rpd3-GFP PSYL1396 MATa ura3∆0 leu2∆0 met15∆0 or MET15 csm3Δ::KanMX RPD3-GFP::HIS3 This study chk1Δ, Rpd3-GFP PSYL1397 MATa ura3∆0 leu2∆0 met15∆0 or MET15 chk1Δ::KanMX RPD3-GFP::HIS3 This study rad9Δ, Rpd3-GFP PSYL1479 MATa ura3∆0 leu2∆0 met15∆0 or MET15 rad9Δ::KanMX RPD3-GFP::HIS3 This study apj1Δ, Rpd3-GFP PSYL1290 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX RPD3-GFP::HIS3 This study Rpd3-GFP trp- PSYL1387 MATa ura3∆0 leu2∆0 met15∆0 or MET15 trp1Δ RPD3-GFP::HIS3 This study apj1Δ, Rpd3-GFP trp- PSYL1389 MATalpha ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX trp1Δ RPD3-GFP::HIS3 This study cdc48-4601, Rpd3-GFP PSYL1318 MATa ura3∆0 leu2∆0 met15∆0 or MET15 cdc48-4601::KanMX RPD3-GFP::HIS3 This study nvj1Δ, Rpd3-GFP PSYL1392 MATa ura3∆0 leu2∆0 met15∆0 or MET15 nvj1Δ::KanMX RPD3- This study 170 GFP::HIS3 vac8Δ, Rpd3-GFP PSYL1342 MATa ura3∆0 leu2∆0 met15∆0 or MET15 vac8Δ::KanMX RPD3-GFP::HIS3 This study atg39Δ, Rpd3-GFP MATa ura3∆0 leu2∆0 met15∆0 or MET15 atg39Δ::KanMX RPD3-GFP::HIS3 This study ubc9-1 ts, Rpd3-GFP PSYL1616 MATa ura3∆0 leu2∆0 met15∆0 or MET15 ubc9-1::URA3 RPD3-GFP::HIS3 This study siz1Δ, Rpd3-GFP PSYL1401 MATa ura3∆0 leu2∆0 met15∆0 or MET15 siz1Δ::KanMX RPD3-GFP::HIS3 This study siz2Δ, Rpd3-GFP PSYL1404 MATa ura3∆0 leu2∆0 met15∆0 or MET15 siz2Δ::KanMX RPD3-GFP::HIS3 This study mms21-CH, Rpd3-GFP PSYL1609 MATa ura3∆0 leu2∆0 met15∆0 or MET15 mms21-CH::KanMX RPD3-GFP::HIS3 This study siz1Δ, siz2Δ, Rpd3-GFP PSYL1603 MATa ura3∆0 leu2∆0 met15∆0 or MET15 siz1Δ::KanMX RPD3-GFP::HIS3 This study siz1Δ, mms21-CH, Rpd3-GFP PSYL1605 MATa ura3∆0 leu2∆0 met15∆0 or MET15 siz1Δ::KanMX mms21-CH::kanMX RPD3-GFP::HIS3 This study siz2Δ, mms21-CH, Rpd3-GFP PSYL1607 MATa ura3∆0 leu2∆0 met15∆0 or MET15 siz2Δ::KanMX mms21-CH::kanMX RPD3-GFP::HIS3 This study smt3-3KR PSYL1774 MATa ura3∆0 leu2∆0 met15∆0 or MET15 smt3-3KR::KanMX This study smt3-3KR, Rpd3-GFP PSYL1615 MATa ura3∆0 leu2∆0 met15∆0 or MET15 smt3-3KR::KanMX RPD3-GFP::HIS3 This study Hos2-GFP PSYL687 MATa ura3∆0 leu2∆0 met15∆0 or MET15 HOS2-GFP::HIS3 Huh et al. 2003 Cdc40-GFP PSYL985 MATa ura3∆0 leu2∆0 met15∆0 or MET15 CDC40-GFP::HIS3 Huh et al. 2003 Hsh155-GFP PSY1502 MATa ura3∆0 leu2∆0 met15∆0 or MET15 HSH155-GFP::HIS3 Huh et al. 2003 Cmr1-GFP PSYL1451 MATa ura3∆0 leu2∆0 met15∆0 or MET15 CMR1-GFP::HIS3 Huh et al. 2003 GFP-Btn2 PSYL1529 MATalpha his3∆1 leu2∆0 met15∆0 ura3∆0 lys+ GFP-BTN2 E. Conibear, University of British Columbia, Vancouver, British 171 Columbia, Canada Hsp42-GFP PSYL1454 MATa ura3∆0 leu2∆0 met15∆0 or MET15 HSP42-GFP::HIS3 Huh et al. 2003 Apj1-GFP PSYL1450 MATa ura3∆0 leu2∆0 met15∆0 or MET15 APJ1-GFP::HIS3 Huh et al. 2003 Hsp104-GFP PSYL1455 MATa ura3∆0 leu2∆0 met15∆0 or MET15 HSP104-GFP::HIS3 Huh et al. 2003 Cdc48-GFP PSYL560 MATa ura3∆0 leu2∆0 met15∆0 or MET15 CDC48-GFP::HIS3 Huh et al. 2003 ubc9-1 ts, GFP-Btn2 PSYL1778 Matalpha his3∆1 leu2∆0 met15∆0 ura3∆0 lys+ ubc9-1::URA3 GFP-BTN2 This study GFP-Btn2 WT PSYL1784 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::GFP-BTN2 This study GFP-Btn2 6KR PSYL1785 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::GFP-btn2-6KR This study Btn2-WT PSYL1788 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 This study Btn2-6KR PSYL1790 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR This study btn2-WT, Rpd3-GFP PSYL1792 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 RPD3-GFP::HIS3 This study btn2-6KR, Rpd3-GFP PSYL1793 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR RPD3-GFP::HIS3 This study btn2-WT, Hos2-GFP PSYL1806 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 HOS2-GFP::HIS3 This study btn2-6KR, Hos2-GFP PSYL1808 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR HOS2-GFP::HIS3 This study btn2-WT, Hsh155-GFP, Hos2-mCherry PSYL1828 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 HSH155-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Hsh155-GFP, Hos2-mCherry PSYL1829 MATa ura3∆0 leu2∆0 met15∆0 or MET15URA3::btn2-6KR HSH155-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-WT, Cmr1-GFP, Hos2-mCherry PSYL1822 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 CMR1-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Cmr1-GFP, Hos2-mCherry PSYL1823 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR CMR1-GFP::HIS3 HOS2-mCherry::URA3 This study 172 btn2-WT, Cdc48-GFP, Hos2-mCherry PSYL1824 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 CDC48-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Cdc48-GFP, Hos2-mCherry PSYL1825 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR CDC48-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-WT, Slx8-GFP, Hos2-mCherry PSYL1832 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 SLX8-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Slx8-GFP, Hos2-mCherry PSYL1833 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR SLX8-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-WT, Apj1-GFP, Hos2-mCherry PSYL1826 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 APJ1-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Apj1-GFP, Hos2-mCherry PSYL1827 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR APJ1-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-WT, Hsp104-GFP, Hos2-mCherry PSYL1830 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::BTN2 HSP104-GFP::HIS3 HOS2-mCherry::URA3 This study btn2-6KR, Hsp104-GFP, Hos2-mCherry PSYL1831 MATa ura3∆0 leu2∆0 met15∆0 or MET15 URA3::btn2-6KR HSP104-GFP::HIS3 HOS2-mCherry::URA3 This study btn2Δ PSYL1009 MATa ura3∆0 leu2∆0 met15∆0 or MET15 btn2Δ::KanMX This study fes1Δ, hsp104Δ PSYL1853 MATa ura3∆0 leu2∆0 met15∆0 or MET15 fes1Δ::HphMX hsp104Δ::NatMX This study fes1Δ, hsp104Δ, btn2-WT PSYL1873 MATa ura3∆0 leu2∆0 met15∆0 or MET15 fes1Δ::HphMX hsp104Δ::NatMX URA3::BTN2 This study fes1Δ, hsp104Δ, btn2-6KR PSYL1874 MATa ura3∆0 leu2∆0 met15∆0 or MET15 fes1Δ::HphMX hsp104Δ::NatMX URA3::btn2-6KR This study apj1Δ, btn2-WT, Rpd3-GFP PSYL1855 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::BTN2 RPD3-GFP::HIS3 This study apj1Δ, btn2-6KR, Rpd3-GFP PSYL1857 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::btn2-6KR RPD3-GFP::HIS3 This study hsp104Δ, btn2-WT, Rpd3-GFP PSYL1875 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX URA3::BTN2 RPD3-GFP::HIS3 This study 173 hsp104Δ, btn2-6KR, Rpd3-GFP PSYL1876 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX URA3::btn2-6KR RPD3-GFP::HIS3 This study apj1Δ, hsp104Δ, btn2-WT, Rpd3-GFP PSYL1877 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::BTN2 RPD3-GFP::HIS3 This study apj1Δ, hsp104Δ, btn2-6KR, Rpd3-GFP PSYL1878 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::btn2-6KR RPD3-GFP::HIS3 This study slx5Δ, GFP-Btn2 PSYL1618 Matalpha his3∆1 leu2∆0 met15∆0 ura3∆0 lys+ slx5Δ::KanMX GFP-BTN2 This study apj1Δ, btn2-WT, Slx8-GFP PSYL1912 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::BTN2 SLX8-GFP::HIS3 This study apj1Δ, btn2-6KR, Slx8-GFP PSYL1913 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::btn2-6KR SLX8-GFP::HIS3 This study hsp104Δ, btn2-WT, Slx8-GFP PSYL1914 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX URA3::BTN2 SLX8-GFP::HIS3 This study hsp104Δ, btn2-6KR, Slx8-GFP PSYL1915 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX URA3::btn2-6KR SLX8-GFP::HIS3 This study apj1Δ, hsp104Δ, btn2-WT, Slx8-GFP PSYL1916 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::BTN2 SLX8-GFP::HIS3 This study apj1Δ, hsp104Δ, btn2-6KR, Slx8-GFP PSYL1917 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::btn2-6KR SLX8-GFP::HIS3 This study apj1Δ, btn2-WT PSYL1896 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::BTN2 This study apj1Δ, btn2-6KR PSYL1897 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX URA3::btn2-6KR This study hsp104Δ, btn2-WT PSYL1898 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX URA3::BTN2 This study hsp104Δ, btn2-6KR PSYL1900 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::KanMX This study 174 URA3::btn2-6KR apj1Δ, hsp104Δ, btn2-WT PSYL1902 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::BTN2 This study apj1Δ, hsp104Δ, btn2-6KR PSYL1903 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX URA3::btn2-6KR This study apj1Δ PSYL1888 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX This study hsp104Δ PSYL1890 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::NatMX This study apj1Δ, hsp104Δ PSYL1892 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX This study btn2Δ, hsp104Δ PSYL1894 MATa ura3∆0 leu2∆0 met15∆0 or MET15 btn2Δ::KanMX hsp104Δ::NatMX This study slx5Δ, btn2-WT PSYL1859 MATa ura3∆0 leu2∆0 met15∆0 or MET15 slx5Δ::KanMX URA3::BTN2 This study slx5Δ, btn2-6KR PSYL1860 MATa ura3∆0 leu2∆0 met15∆0 or MET15 slx5Δ::KanMX URA3::btn2-6KR This study apj1Δ, slx5Δ PSYL1918 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX slx5Δ::KanMX This study hsp104Δ, slx5Δ PSYL1922 MATa ura3∆0 leu2∆0 met15∆0 or MET15 hsp104Δ::NatMX slx5Δ::KanMX This study apj1Δ, hsp104Δ, slx5Δ PSYL1920 MATa ura3∆0 leu2∆0 met15∆0 or MET15 apj1Δ::KanMX hsp104Δ::NatMX slx5Δ::kanMX This study 175 A.2 Primers used in this study. Primer name Primer sequence (5'-3') Primer description Btn2-Full length-F AGGTGCGGCCGCTTTCATTATGGACTACTTTAC Forward primer binding to Btn2 promoter region, contains NotI site Btn2-Full length-R CCCGGTACCATAATTAGTTATTTAATGCCG Reverse primer binding to Btn2 terminator region, contains KpnI site. Also used as the reverse primer for the second product of Btn2-CTD mutant stitch PCRs Btn2-CTDΔ-R CCCGGTACCTTAAATTTCTTCCTCATGCTTAAC Reverse primer binding to the end of Btn2 CLD region, contains a stop codon and KpnI site. Also used as the reverse primer for the first product of Btn2-CTD mutant stitch PCRs Btn2-CTD+CLDΔ-R CCCGGTACCTTAATGGTTCTCCTTTAGCTCACCT Reverse primer binding to the end of Btn2 NTD region, contains a stop codon and KpnI site 176 Btn2 CTD stitch-F ATTGGTTTAGTTAAGCATGAGGAAGAAATT Forward primer for first half of the Btn2 stitch for CTD mutant PCRs Btn26KR integration-F CAACAACCAAAAGAAAATAACTAATAGACCCCATTACAATATAGAAGAGTGCACCACGCTTTTCAAT Forward primer for integrating Btn2-WT and Btn2-6KR from a pRS316 background to the genome Btn26KR integration-R ATAATTAGTTATTTAATGCCG Reverse primer for integrating Btn2-WT and Btn2-6KR from a pRS316 background to the genome A.3 Plasmids used in this study. Plasmid description Relevant Features Source pRS316 CEN6/ARSH4, ampR, URA3 P. Hieter, University of British Columbia, Vancouver, British Columbia, Canada YEplac181 ampR CUP1 YEplac181-HIS-Smt3::LEU2 H. Ulrich et al. 2009 (obtained from Addgene) pRS316 - btn2wt Btn2 ORF along with promoter and terminator This study 177 region, ampR, URA3 pRS316 - btn2ctdΔ Btn2 ORF without the C-terminal domain, along with promoter and terminator region, ampR, URA3 This study pRS316 - btn2ctd+cldΔ Btn2 ORF, only the N-terminal domain, along with promoter and terminator region, ampR, URA3 This study pRS316 - btn2ctdKallR Btn2 ORF, with all C-terminal domain lysines mutated to arginine, ampR, URA3 This study pRS316 - btn2ctd2KR Btn2 ORF, with lysines K227 and K234 mutated to arginine, ampR, URA3 This study pRS316 - btn2ctd6KR Btn2 ORF, with lysines K249,K255,257,262,267,268 mutated to arginine, ampR, URA3 This study pRS316 - btn2ctd6KR the II Btn2 ORF, with lysines K282,287,290,297,298,300 mutated to arginine, ampR, URA3 This study pRS316 - btn2ctd12KR Btn2 ORF, with lysines K305,308,314,316,318,323,324,329,341,375,376,402 mutated to arginine, ampR, URA3 This study p426MET25-FFL(firefly luciferase)-GFP ampR, LEU2, Met repressible promoter, heat inactivated at 42 degrees, expressed throughout the cell (no localization markers) K. Morano, Abrams and Morano 2013 pRS315 CEN6/ARSH4, ampR, TRP1 P. Hieter, University of British Columbia, 178 Vancouver, British Columbia, Canada pRS316 - APJ1 Apj1 ORF along with its promoter and terminator region, ampR, TRP1 Sahi et al. 2013 pRS316 - APJ1 H34Q Apj1 ORF with a mutated J domain, ampR, TRP1 Sahi et al. 2013 pRS316 - APJ1 ΔZn Apj1 ORF without the Zinc finger domain, ampR, TRP1 Sahi et al. 2013 pRS316 - APJ1 ΔGFS Apj1 ORF without the G/F domain, ampR, TRP1 Sahi et al. 2013 pRS316 - APJ1 I79S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013 pRS316 - APJ1 L198S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013 pRS316 - APJ1 L200S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013 pRS316 - APJ1 L277S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013 pRS316 - APJ1 V310S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013 pRS316 - APJ1 I318S Apj1 ORF hydrophobic cleft mutant, TRP1 Sahi et al. 2013
Thesis/Dissertation
eng
Medical Genetics
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Stirling, Peter C.
DNA damage induced SUMOylation regulates intranuclear protein quality control in Saccharomyces cerevisiae
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