Science, Faculty of
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UBCV
Ye, Bin
Doctor of Philosophy - PhD
University of British Columbia
Genome stability is essential for survival in all living organisms. Perturbations of genome stability can lead to cancer and other diseases in human. Dysfunction of genome care-taking genes and exposure to external genotoxins are the two major drivers of genomic instability. Here, we used a model organism, C. elegans to study these factors and how they interact to affect genome stability. We first generated a chemi-genetic interaction map as a preliminary reference for characterizing genes and agents. From there, we elucidated the mechanisms of action for an emerging anticancer therapeutic CX-5461. CX-5461 turned out to be a multi-modal agent that exhibits genotypic sensitivity, mutagenicity and photosensitivity. Next, we identified a few genetic interactions within and between double-strand break repair pathways in C. elegans, which have implications in the response to both endogenous and exogenous DNA damage. We also characterized a less-understood gene SMRC-1, an annealing helicase. Loss of SMRC-1 results in copy number variations, and SMRC-1 is synthetic lethal with a number of DNA damage response genes. Our work concludes that C. elegans is a great platform to untangle the sophisticated complexity in genome maintenance, and provide meaningful insights for prospective anticancer therapeutic development.
C. ELEGANS AS A PLATFORM TO CHARACTERIZE GENES AND THERAPEUTICS AFFECTING GENOME STABILITY by Bin Ye B.Sc., Xiamen University, 2014 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Genome Science and Technology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December 2019 © Bin Ye, 2019 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: C. elegans as a platform to characterize genes and therapeutics affecting genome stability submitted by Bin Ye in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Genome Science and Technology Examining Committee: Philip Hieter, Medical Genetics Supervisor Stefan Taubert, Medical Genetics Supervisory Committee Member Michel Roberge, Biochemistry & Molecular Biology Supervisory Committee Member Peter Lansdorp, Medical Genetics University Examiner Don Moerman, Zoology University Examiner Additional Supervisory Committee Members: Samuel Aparicio, Pathology Supervisory Committee Member Supervisory Committee Member iii ABSTRACT Genome stability is essential for survival in all living organisms. Perturbations of genome stability can lead to cancer and other diseases in human. Dysfunction of genome care-taking genes and exposure to external genotoxins are the two major drivers of genomic instability. Here, we used a model organism, C. elegans to study these factors and how they interact to affect genome stability. We first generated a chemi-genetic interaction map as a preliminary reference for characterizing genes and agents. From there, we elucidated the mechanisms of action for an emerging anticancer therapeutic CX-5461. CX-5461 turned out to be a multi-modal agent that exhibits genotypic sensitivity, mutagenicity and photosensitivity. Next, we identified a few genetic interactions within and between double-strand break repair pathways in C. elegans, which have implications in the response to both endogenous and exogenous DNA damage. We also characterized a less-understood gene SMRC-1, an annealing helicase. Loss of SMRC-1 results in copy number variations, and SMRC-1 is synthetic lethal with a number of DNA damage response genes. Our work concludes that C. elegans is a great platform to untangle the sophisticated complexity in genome maintenance, and provide meaningful insights for prospective anticancer therapeutic development. iv LAY SUMMARY It is critical for all living organisms to faithfully maintain and pass down genetic information encoded in their DNA. This involves a mechanism called genome stability maintenance. Failure to maintain genome stability can lead to dire consequences, such as cancers in human. Here, we are using the nematode worm to study factors that can affect this mechanism. Many genes that function in worms have similar roles in humans, and worms are easier to maneuver, making them a great platform for biomedical research. In this study, we characterized genes and therapeutics that can affect genome stability maintenance and how these two factors interact. This provides insights that can shed light on future development of anticancer therapy. v PREFACE Part of the research described in Chapter 3 was published in "Hong Xu, Marco Di Antonio, Steven McKinney, Veena Mathew, Brandon Ho, Nigel J. O’Neil, Nancy Dos Santos, Jennifer Silvester, Vivien Wei, Jessica Garcia, Farhia Kabeer, Daniel Lai, Priscilla Soriano, Judit Banath, Derek S. Chiu, Damian Yap, Daniel D. Le, Frank B. Ye, Anni Zhang, Kelsie Thu, John Soong, Shu-chuan Lin, Angela Hsin Chin Tsai, Tomo Osako, Teresa Algara, Darren N. Saunders, Jason Wong, Jian Xian, Marcel B. Bally, James D. Brenton, Grant W. Brown, Sohrab P. Shah, David Cescon, Tak W. Mak, Carlos Caldas, Peter C. Stirling, Phil Hieter, Shankar Balasubramanian & Samuel Aparicio (2017). CX-5461 is a DNA G-quadruplex stabilizer with selective lethality in BRCA1/2 deficient tumours. Nature communications 8, 14432”. The paper was designed and written by Dr. Hong Xu, Dr. Marco Di Antonio, Dr. Veena Mathew, Dr. Nigel J. O’Neil, Dr. Sohrab P. Shah, and Dr. Samuel Aparicio. Frank Bin Ye contributed to the C. elegans experiments in the paper, and a small fraction of Figure 2.2 was used to generate figures in the publication. The research described in Chapter 2 and 3 will be submitted as "Frank B. Ye, Akil Hamza, Tejomayee Singh, Stephane Flibotte, Samuel Aparicio, Philip Hieter, Nigel J. O’Neil. C. elegans as a platform to characterize the anti-cancer therapeutic CX-5461". The paper was designed and co-written by Frank Bin Ye, Dr. Nigel J. O’Neil and Dr. Philip Hieter. Frank Bin Ye performed most of the experiments. Dr. Stephane Flibotte performed the bioinformatic analysis on sequencing data. Dr. Akil Hamza performed the yeast experiments, and Tejomayee Singh performed the mammalian cell experiments. vi The research described in Chapter 4 and 5 was designed and performed mostly by Frank Bin Ye, and will be submitted for two future publications. The bioinformatic analysis of sequencing data was performed by Dr. Stephane Flibotte. vii TABLE OF CONTENTS ABSTRACT…………………………………………………………………………. iii LAY SUMMARY……………………..……………..……………………………… iv PREFACE……………..……………..……………………..……………..…………. v TABLE OF CONTENTS……………..……………..………………..……………… vii LIST OF TABLES……………..……………..………………..……………..……… xii LIST OF FIGURES……………..……………..……………..……………..……….. xiii LIST OF SYMBOLS, ABBREVIATIONS & ACRONYMS……………………….. xv ACKNOWLEDGEMENTS……………..……………..……………..……………… xviii DEDICATION……………..……………..……………..………………..…………. xix CHAPTER 1: INTRODUCTION……………..……………..……………..………... 1 1.1 Genome stability and cancer ……………..……………..……………..….. 1 1.2 Genes that affect genome stability…………….………………………….. 2 1.2.1 DNA metabolism genes…………….…………………………..… 2 1.2.2 DNA damage response genes……………………………..…….... 3 1.2.3 DNA repair pathways …………………………………..………… 4 1.2.4 The interplay between DNA repair pathways………..…………… 8 1.3 Carcinogens and therapeutics that affect genome stability………..………. 10 1.3.1 Carcinogens………….……………...……………..……………… 10 1.3.2 Therapeutics...……………..……………..……………..………… 11 1.4 Investigating genome instability..……………..……………..…………… 12 1.4.1 Assays for measuring genome instability: micronuclei test to HPRT assay…………….……………...………..…………..…… 12 1.4.2 Defining the genome stability gene spectrum..…………………… 13 1.4.3 Mutational signatures..……………..………………….……..…... 15 1.5 C. elegans as a platform to study factors affecting genome stability…….. 16 1.6 Research objectives……………..……………..……………..…………... 19 CHAPTER 2: GENERATION OF A CHEMI-GENETIC INTERACTION MAP OF WELL-CHARACTERIZED DNA DAMAGING AGENTS……………..………… 23 viii 2.1 Introduction ……………..……………..……………..………………...... 23 2.2 Methods……………..……………..……………..……………..………... 26 2.3 Results……………..……………..……………..……………..………… 29 2.3.1 Construction of a DNA genome stability mutant panel………… 29 2.3.2 Genotypic sensitivity profile of olaparib ……………..……..…. 29 2.3.3 Genotypic sensitivity profile of camptothecin (CPT)……………. 30 2.3.4 Genotypic sensitivity profile of etoposide (ETOP)……………... 30 2.3.5 Genotypic sensitivity profile of UVA-trimethylpsoralen (UVA- TMP) ………..…………..…………..…………..………………. 31 2.3.6 Genotypic sensitivity profile of ultraviolet C (UVC)…………….. 31 2.4 Discussion …………….…………..……………..……………..………… 32 2.4.1 Each agent has a distinctive genotypic sensitivity profile……….. 32 2.4.2 Some genes are required for many different DNA damage responses………..…………..…………..………………………. 36 2.4.3 The chemi-genetic interaction map can be a reference for characterizing new damaging agents and DNA replication and repair genes………………..…………..……………………….. 37 CHAPTER 3: UNDERSTANDING THE MECHANISMS OF ACTION OF CX-5461………..…………..……………………….………..…………..……………. 41 3.1 Introduction …………….……………..……………..………………….. 41 3.2 Methods …………….……………..……………..…………………..….. 41 3.3 Results……………..……………..……………..……………..………… 45 3.3.1 Genotypic sensitivity profile of CX-5461 ……………..……..… 45 3.3.2 CX-5461 comparison to loss of the G-quadruplex associated helicase dog-1……..…………..……………..…………………. 46 3.3.3 CX-5461 can intercalate into DNA secondary structures………. 47 3.3.4 UVA irradiation enhances CX-5461 cytotoxicity, causing both replication and transcription-associated lesions……..………….. 48 3.3.5 CX-5461 causes phototoxicity by generating reactive oxygen ix species……..…………..……………..……………..…………… 50 3.3.6 CX-5461 is mutagenic, UVA enhances its mutagenicity……….. 51 3.3.7 Mutational profile of CX-5461 (-/+ UVA)……………..……..…. 52 3.4 Discussion: CX-5461 is A multi-modal anticancer drug………………… 55 3.4.1 Multiple DNA repair pathways are required for CX-5461-induced DNA lesions……..…………..……………..…………………… 55 3.4.2 CX-5461 can affect non-G4 DNA………..…………..…………. 56 3.4.3 The double-edged sword of therapeutic photosensitivity………. 57 3.4.4 CX-5461 has a complex mutational profile……………………… 58 3.4.5 CX-5461 could function in a similar manner as ellipticines…….. 59 CHAPTER 4: GENETIC INTERACTIONS BETWEEN AND WITHIN DOUBLE-STRAND BREAK REPAIR PATHWAYS IN C. ELEGANS..……………………. 70 4.1 Introduction ……………..……………..……………..………………….. 70 4.1.1 HDR factors ……………..……..…....…………..……………… 71 4.1.2 NHEJ factors ……………..……..…....…………..…………….. 73 4.1.3 MMEJ factors ……………..……..…....…………..……………. 74 4.1.4 The C. elegans germ line ……………..……..…....…………….. 74 4.1.5 Genetic interactions ……………..……..…....…………………. 75 4.2 Methods ……………..……………..……………..……………..……… 76 4.3 Results……………..……………..……………..……………..………… 78 4.3.1 Loss of NHEJ rescues meiotic defects in helq-1 single mutants but not other HDR mutants. ……..…………..……………..………. 78 4.3.2 Loss of MMEJ lead to transgenerational sterility in HDR mutants 79 4.3.3 helq-1 brd-1 double mutants are synthetic lethal..……………….. 80 4.3.4 brd-1 rfs-1 double mutants are synthetic sick..…………………... 82 4.3.5 Revisiting the synthetic lethality in helq-1 rfs-1 double mutants... 84 4.3.6 Loss of NHEJ rescues the drug sensitivity in some but not all HDR-compromised mutants……..…………..…………………. 86 x 4.3.7 Loss of MMEJ sensitizes HDR mutants to camptothecin (CPT), etoposide (ETOP) and CX-5461……..…………..……………… 86 4.4 Discussion……………..……………..……………..…………….………. 88 4.4.1 Redundancy between HDR factors..…………. ..………………... 88 4.4.2 MMEJ is both a primary and back-up repair pathway dependent on the nature of DNA lesion ……………..……………………… 93 4.4.3 Illegitimate NHEJ activity can be toxic in certain HDR- compromised background……..…………..……………………. 96 CHAPTER 5: CHARACTERIZATION OF THE ANNEALING HELICASE SMRC-1 IN C. ELEGANS……..…………..……………..……………..…………. 110 5.1 Introduction ……………..……………..……………..………………….. 110 5.2 Methods ……………..……………..……………..……………………… 112 5.3 Results……………..……………..……………..……………..………… 115 5.3.1 SMRC-1 is required for maintaining genome stability..………… 115 5.3.2 SMRC-1 is required for survival for DDA treatment..………….. 117 5.3.3 Genetic interactions between loss of SMRC-1 and other helicases 117 5.3.4 Genetic interactions between loss of SMRC-1 and other DDR genes……..…………..……………..……..…………..………… 119 5.4 Discussion……………..……………..……………..…………….………. 120 5.4.1 SMRC-1 is required for genome integrity....………… ..……….. 120 5.4.2 SMRC-1 is required for genome integrity in the absence of RTEL-1……..…………..…………………..…………………… 122 5.4.3 SMRC-1 is required for genome integrity in the absence of MUS- 81……..…………..……………..……..…………..……………. 123 5.4.4 smrc-1 genetically interacts with factors required for DNA metabolism……..…………..………………..…………..……… 124 CHAPTER 6: CONCLUSION……..…………..……………..……………..…….. 133 6.1 Anticancer drugs have complex, multimodal mechanisms of action…… 133 6.2 C. elegans is a good platform with which to study factors affecting xi genome stability……..…………..………………..………..…………..… 135 BIBLIOGRAPHY ……..…………..…………..…………..……………..………… 139 APPENDIX ……..…………..…………..…………..……………..………………. 159 xii LIST OF TABLES Table 2.1 Summary of the DNA damaging agents used to generate the chemi- genetic interaction map.…………..…………………………………… 39 Table 3.1 Phenotypes of indicated mutants (Chapter 3) …………..…………….. 68 Table 3.2 Summary of mutations on CX-5461-treated and CX-5461 + UVA mutants…………..……………………..……………………..………. 69 Table 4.1 Phenotypes of indicated mutants (Chapter 4) …………..…………….. 109 Table 5.1 Phenotypes of indicated mutants (Chapter 5) …………..…………….. 132 Table A1.1 Raw data for generating the sensitivity scores for Olaparib………… 166 Table A1.2 Raw data for generating the sensitivity scores for Camptothecin….. 167 Table A1.3 Raw data for generating the sensitivity scores for Etoposide………. 168 Table A1.4 Raw data for generating the sensitivity scores for UVA- Trimethylpsoralen. …………………………………………………… 169 Table A1.5 Raw data for generating the sensitivity scores for UVC. …………… 170 Table A1.6 Raw data for generating the sensitivity scores for CX-5461………... 171 Table A2. Worm strains generated in this thesis…………..…………………..….. 172 xiii LIST OF FIGURES Figure 1.1 Schematics of different DNA repair pathways……..…….…………..… 21 Figure 2.1 A schematic of quantitative acute assay…………..……………………. 39 Figure 2.2 The chemi-genetic interaction map of different agents…………..…….. 40 Figure 3.1 Genotypic sensitivity to CX-5461..…………..……………………..….. 61 Figure 3.2 Effect of G-quadruplex stabilization on CX-5461-sensitive mutants….. 62 Figure 3.3 CX-5461 stabilizes DNA duplex structures.…………..……………….. 63 Figure 3.4 CX-5461 is a photosensitizer in C. elegans, human cancer cell lines and yeast.…………..…………………..……………….………..………….. 64 Figure 3.5 CX-5461 + UVA can result in photodamage by generating ROS……… 66 Figure 3.6 Exposure to CX-5461 or CX-5461 + UVA results high frequencies of mutations…………..…………………..……………….………..……… 67 Figure 4.1 A schematic of DSB repair pathways in C. elegans..…………………… 98 Figure 4.2 cku-80 genetically interacts helq-1 single mutants but not other HDR mutants or MMEJ mutants..…………………..…………………..…… 98 Figure 4.3 polq-1 genetically interacts with HDR gene/mutants (helq-1, brd-1, and rfs-1).…………..…………………..……………….………..………… 99 Figure 4.4 helq-1 brd-1 double mutants are synthetic lethal…………………. 100 Figure 4.5 brd-1 rfs-1 double mutants are synthetic sick.…………………………. 102 Figure 4.6 helq-1 rfs-1 double mutants are synthetic lethal……………………….. 104 Figure 4.7 Loss of NHEJ rescues the DNA damage agent sensitivity in some but not all HDR-compromised mutants……………………….……………….. 106 Figure 4.8 Loss of MMEJ sensitizes HDR mutants to DNA damage agents…….. 107 Figure 4.9 Model of different DSB repair pathways required for CPT, ETOP and CX-5461…………………………………………………………….…. 108 Figure 5.1 SMRC-1 is required for maintaining genome stability………………... 128 Figure 5.2 SMRC-1 is required for tolerance of genotoxins…………………….… 129 Figure 5.3 Genetic interactions with loss of SMRC-1…………………………….. 130 Figure A3.1: Coverage plot of CX-5461-induced genome rearrangements………. 159 xiv Figure A3.2. Coverage plot of CX-5461 + UVA-induced genome rearrangements 160 Figure A4.1 Representative images of DAPI-stained bodies in diakinesis of cku-80 and polq-1 single mutants…………………………………………… 163 Figure A5.1 Coverage plot of quantile deviations and fractions of variant basis of Million Mutation Project collection strains VC20168 and VC40734.. 164 Figure A5.2 Coverage plot of genome rearrangements of F10 gk176502 and F10 gk784642.…………………………………………………………….. 165 xv LIST OF ABBREVIATIONS & ACRONYMS ATM Ataxia telangiectasia mutated ATR Ataxia telangiectasia and RAD3-related BTR BLM–RMI–TOP3α CDK Cyclin-dependent kinase CNV Copy number variation CPT Camptothecin CRISPR Clustered regularly interspaced short palindromic repeats DAPI 4',6-diamidino-2-phenylindole dihydrochloride DDA DNA damaging agent Del Deletion DMSO Dimethyl sulfoxide DNA Deoxyribonucleic acid DNA-PK DNA-dependent protein kinase DSB Double-strand break dsDNA Double-stranded DNA Dup Duplication EMS Ethyl methanesulfonate ENU N-ethyl-N-nitrosourea ETOP Etoposide FA Fanconi anemia FS Frame shift G4 G quadruplex GG-NER Global genome nucleotide excision repair H2DCFDA 2’,7’-dichlorodihydrofluorescein diacetate HDR Homology-directed repair Him High incidence of males HPRT Hypoxanthine phosphorybosyl transferase IC50 Half maximal inhibitory concentration xvi ICL Interstrand crosslink IDL Insertion/deletion loop Inv Inverstion M-Z- Maternally negative and zygotically negative M+Z- Maternally positive and zygotically negative MBN Mung bean nuclease MGMT O6-methylguanine-DNA methyltransferase MH Microhomology MMEJ Microhomology-mediated end joining MMP Million mutation project MMR Mismatch repair MNM Multi-nucleotide mutation NER Nucleotide excision repair NGI Negative genetic interaction NGM Nematode growth media NHEJ Non-homologous end joining Ola Olaparib PARP Poly (ADP-ribose) polymerase PCR Polymerase chain reaction ROS Reactive oxygen species SDSA Synthesis dependent strand annealing SEM Standard error of the mean shRNA Small hairpin RNA SIOD Schimke immuno-osseous dysplasia SL Synthetic lethality or synthetic lethal SNV Single nucleotide variation SS Synthetic sickness of synthetic sick SSB Single-strand break ssDNA Single-stranded DNA xvii TC-NER Transcription-coupled nucleotide excision repair TFIIH Transcription factor II H TLS Translesion synthesis TMP Trimethylpsoralen Trans Translocation UV Ultra-violet WT Wild-type xviii ACKNOWLEDGEMENTS I thank my supervisor Dr. Philip Hieter for his guidance, support and patience during my graduate studies. I am deeply indebted to Dr. Nigel O’Neil for his generous mentorship, which was crucial to my graduate training. I thank my committee members, Dr. Samuel Aparicio, Dr. Stefan Taubert, and Dr. Michel Roberge. Their scientific input and suggestions were helpful for my thesis. I thank all the members in Hieter Lab, present and past, Dr. Melanie Bailey, Dr. Akil Hamza, Leanne Amitzi, Tejomayee Singh, Rafaela Marodin, Anthony Oppedisano, Walid Omar, Maureen Driessen, Dr. Supipi Duffy, Megan Kofoed, Sidney Ang, and Sivan Reytan. I am grateful for the fun and chats that helped me get through my graduate study. I also thank the friends I have met in Vancouver and Canada that made my graduate study experience a memorable one. I acknowledge scholarship support from the Genome Science and Technology Program, Mitacs Inc., St John’s College, and the University of British Columbia. Thanks to my parents for their enduring support throughout the degree. xix DEDICATION To my parents for their continual support. CHAPTER 1: INTRODUCTION 1.1 Genome stability and cancer Mechanisms that ensure genome stability are critical for the faithful transmission of genetic information to daughter cells upon cell division. To maintain genome stability, organisms have evolved a complex maintenance system to safeguard and protect the genome against both endogenous and exogenous factors that could affect its integrity. Genome integrity is essential for normal cell growth and development, and the perturbation of genome stability can lead to cancer and other diseases (Aguilera and Garcia-Muse, 2013; Jeggo et al., 2016; Negrini et al., 2010). An increasing body of evidence supports genome instability as an enabling characteristic of carcinogenesis (Hanahan and Weinberg, 2011). Genome instability drives genetic diversity by the sequential accumulation of genetic alterations. These genetic alterations range from structural and numerical chromosomal changes, which result in partial gain or loss of chromosomes and chromosomal rearrangements, to single nucleotide variations at the base level. These genetic alterations increase the probability of acquiring a set of mutations that promote tumor initiation and development (Sieber et al., 2003; Stratton et al., 2009). Genome instability enables cancer cell evolution, but there is a limit to the amount of genome instability that cancer cells can withstand (Andor et al., 2017). This is because cancer cells have a reduced set or downregulation of genome maintenance genes, which in effect represents a sub-lethal loss-of-function of processes that in aggregate are essential for cell viability (Helleday et al., 2008; Jeggo et al., 2016). Cancer cells are often sensitized to DNA damaging agents that 1further increase genomic instability (Cheung-Ong et al., 2013). For example, the platinum-based compounds such as cisplatin, can crosslink DNA and cause DNA lesions (Deans and West, 2011). Cancer cells defective in homology-directed repair (HDR) are unable to resolve cisplatin-induced DNA damage which results in cell death (Helleday, 2010). Therefore, genome instability drives tumorigenesis but it can also be exploited for anticancer treatment. The key to providing more informed anticancer therapy is a better understanding of the causes and effects of genome instability. Although it is not always clear how genome instability arises, dysfunction of genome maintenance genes and exposure to external genotoxins are two major causes of genomic instability (Aguilera and Garcia-Muse, 2013). Therefore, characterizing how genome stability genes and genotoxic agents affect genome stability is crucial to the development of anticancer strategies. 1.2 Genes that Affect Genome Stability Cells have a sophisticated system dedicated to maintaining genome stability involving a plethora of interconnected genes. These genes encompass a wide range of biological processes, from maintaining normal DNA metabolism, responding to DNA damage, to mediating the segregation of chromosomes to daughter cells during cell division. This thesis focuses on those genes required for normal DNA metabolism and the DNA damage response. Here, we catalog some of the genes and processes that were investigated as part of this thesis and provide brief phenotypic descriptions of relevant mutations in the model organism Caenorhabditis elegans. 21.2.1 DNA metabolism genes Under normal conditions, many genes are required for DNA metabolism, such as DNA replication and transcription. These genes encode protein products that are directly involved in replication and transcription, such as helicases and polymerases, as well as genes that regulate replication and transcription, such as chromatin modifiers. Disrupting these genes often has a negative effect on DNA metabolism. For example, defects in polymerase can result in a compromised fidelity of replication, while loss of certain helicases can increase the risk of mutations (London et al., 2008). Among DNA metabolism genes, many have been found dysregulated in cancer. For instance, many cancers have upregulated RecQ-family helicases (Futami et al., 2010; Su et al., 2010), making them potential biomarkers and targets for anticancer drugs. 1.2.2 DNA damage response genes DNA damage response genes safeguard the genome by counteracting DNA damage, which if not resolved could lead to genome instability. In most cases, the response to DNA damage is a multi-step process: recognition of damage, cell cycle arrest, repair or bypass of lesions, activation of apoptosis if the damage cannot be repaired, and restart of the cell cycle. Perturbation of any step in the DNA damage response could affect the resolution of DNA lesions and contribute to genome instability. DNA damage checkpoints DNA damage checkpoints initiate a cascade of signalling responses in response to DNA damage. DNA damage checkpoint genes initiate cell cycle arrest and activate and recruit DNA repair proteins to sites of DNA damage. Two major DNA damage checkpoint pathways are ataxia 3telangiectasia mutated (ATM)-dependent signalling and ataxia telangiectasia and RAD3-related (ATR)-dependent signaling. Both ATM and ATR can regulate cyclin-dependent kinases (CDKs) to arrest the cell cycle to allow time for DNA repair proteins to resolve the damage (Blackford and Jackson, 2017; Yang et al., 2003). If DNA damage exceeds the ability to repair, cells activate p53-dependent apoptosis (Nowsheen and Yang, 2012). In C. elegans, ATM-1 and ATR-1 are activated in response to replication stress or double-strand breaks to induce cell cycle arrest (Garcia-Muse and Boulton, 2005). ATM-1 and ATR-1 also play a role in regulating the resolution of meiotic crossovers in C. elegans (Li and Yanowitz, 2019). Loss of atm-1 in C. elegans results in genome instability and telomeric defects (Jones et al., 2012). DNA repair pathways Cells have a number of DNA repair pathways that respond to different types of DNA lesions to maintain genome stability and fidelity (Ciccia and Elledge, 2010). Homology-directed repair (HDR), non-homologous end joining (NHEJ), and microhomology-mediated end joining (MMEJ) respond to double-strand breaks (DSBs) (Ceccaldi et al., 2016a). Nucleotide excision repair (NER) removes bulky lesions such as thymine dimers and photoproducts (Marteijn et al., 2014). The Fanconi Anemia (FA) repair pathway resolves DNA interstrand crosslinks (Deans and West, 2011). Here, we give a brief overview of the different DNA repair pathways. 1.2.3 DNA repair pathways Nucleotide excision repair (NER) NER can repair a wide range of DNA lesions that alter DNA structure, such as bulky chemical adducts, Ultra-violet (UV)-induced thymine dimers and interstrand crosslinks. There are two kinds of NER, global genome NER (GG-NER) and transcription-coupled NER (TC-NER). The 4two kinds of NER differ in how the lesions are first recognized. In GG-NER, the lesion is detected by XPC, a protein that binds to distorted DNA (Sugasawa et al., 2001), whereas in TC-NER, the lesion stalls RNA polymerase II, which then recruits a number of proteins, including CSA, CSB and XPA (Marteijn et al., 2014). Then, XPC or XPA recruits the transcription factor II H (TFIIH) complex, and the two helicases within TFIIH further open up the double helix containing the lesion. The strand containing the lesion is then excised by the endonucleases XPF–ERCC1 and XPG, resulting in a 22-30 nucleotide gap. The gap is filled by a DNA polymerase, which could sometimes be error-prone translesion polymerases such as POLK and POLH (Figure 1.1) (Marteijn et al., 2014; Zheng et al., 2003). In C. elegans, loss of NER results in hypersensitivity to UVC radiation and DNA crosslinks in a developmentally dependent manner (Astin et al., 2008; Wilson et al., 2017). Mismatch repair (MMR) MMR repairs mis-paired bases and insertion/deletion loops (IDLs) (Pena-Diaz and Jiricny, 2012) . The MMR pathway initiates with MutSα (MSH2-MSH6) binding to a mismatch, followed by binding MutSβ (PSM2-MLH1). Then, MutSα is activated by PCNA to incise the newly-synthesized strand, creating a nick that is further processed by exonuclease EXO1 to form a gap up to 150 bp (Kunkel and Erie, 2015; Pena-Diaz and Jiricny, 2012). The gap is filled by a DNA polymerase and a ligase (Figure 1.1). In C. elegans, some mismatch repair mutants exhibit genome instability and mutation accumulation (Degtyareva et al., 2002; Tijsterman et al., 2002). In contrast, the germ line-specific MutS mismatch repair mutant, him-14 is defective in meiotic crossing over (Zalevsky et al., 1999). 5Homology-directed repair (HDR) HDR, NHEJ and MMEJ all repair DSBs. The HDR is an error-free repair pathway as it uses homologous DNA as a template for repair. HDR initiates the DSB repair by resecting the ends carried out by MRE11 and CtIP endonucleases, generating two free 3’ overhangs. The free 3’ overhangs can be further resected by helicases or exonucleases (i.e., CtIP, DNA2, BLM, WRN, and EXO1) (Ceccaldi et al., 2016a). The single-stranded DNA associates with the single-stranded DNA (ssDNA) binding protein RPA; this allows BRCA2-mediated recruitment of RAD51 to displace RPA and form RAD51 filaments. RAD51 filaments catalyze the search for homology and subsequent invasion into the homologous sequences on the sister chromatid or the homologous chromosome, forming a displacement D-loop. DNA synthesis occurs spanning the original break site. Finally, the D-loop is resolved by endonucleases or dissolvases such as MUS81 or the BTR complex (BLM–RMI–TOP3α) (Wright et al., 2018) (Figure 1.1). HDR is a multifaceted repair process with several steps. There are variations on how HDR can resolve the DSBs, which are dictated by the nature of the DSB and the available repair proteins. Variants include: synthesis dependent strand annealing (SDSA), break induced replication repair, and homologous recombination (Verma and Greenberg, 2016). In C. elegans, homologous recombination is essential for meiosis, and therefore, survival (Saito and Colaiacovo, 2017). Loss of function mutations affecting key HDR homolog genes, such as brc-2 (C. elegans homolog of human BRCA2) and rad-51 (C. elegans homolog of human RAD51), cannot be maintained (Alpi et al., 2003; Martin et al., 2005). However, several non-essential HDR genes can be mutated and maintained such as brc-1 (C. elegans homolog of human BRCA1), brd-1 (C. elegans homolog of human BARD1) and rfs-1 (C. elegans homolog of human RAD51C). 6Non-homologous end-joining (NHEJ) In contrast to HDR, the NHEJ pathway does not use a template to repair DSBs and therefore does not need to resect the broken DNA ends. NHEJ mediates the ligation of two non-resected blunt DNA ends. The KU70/80 heterodimer binds to the blunt DSB ends. Next, DNA-dependent protein kinases (DNA-PKs) are recruited along with other end-processing factors to keep the two ends in close proximity. Finally, the DNA ligase LIG4 catalyzes the DNA-end ligation (Panier and Boulton, 2014) (Figure 1.1). In C. elegans, loss of NHEJ results in sensitivity to ionizing radiation in somatic cells (Clejan et al., 2006). Inappropriate NHEJ of meiotic DSBs in the C. elegans germ line is prevented by the COM-1 protein, and loss of COM-1 leads to deleterious NHEJ of meiotic DSBs (Lemmens et al., 2013). This illustrates the importance of interplay between different repair pathways to ensure genome stability. Microhomology-mediated end-joining (MMEJ) MMEJ was discovered in recent years (Deriano and Roth, 2013). As in HDR, the MMEJ pathway requires resection of the DSB ends by MRE11 and CtIP (Truong et al., 2013). In contrast to HDR, MMEJ only requires a microhomology, which can be as small as 1 nucleotide, for DSB repair (Koole et al., 2014). When the resected ends anneal with microhomology via complementary base pairing, the flanking single-stranded regions are filled in by polymerase theta POLQ (Figure 1.1). Finally, DNA ends are ligated by DNA ligases LIG1 or LIG3 (Sfeir and Symington, 2015). Since MMEJ anneals micro-homologous sequences within the resected DSB ends before joining, this often results in deletions surrounding the original DSBs (Sfeir and Symington, 2015). In C. elegans, POLQ-1 is required for the interstrand crosslink (ICL) repair (Muzzini et al., 2008) and the bypass of G-quadruplex (G4) structures that form in the absence of the C. elegans FANCJ homolog, dog-1 (Koole et al., 2014). 7Translesion synthesis (TLS) Bypass Translesion synthesis does not repair DNA lesions. It incorporates nucleotides opposite the site of DNA damage to allow replicative bypass at the expense of low fidelity replication. The bypass often involves several polymerase-switching processes, including the dissociation of the replicative polymerase from the replication fork, the recruitment of the TLS polymerase to the fork, and subsequent displacement of the TLS polymerase with a replicative polymerase. The switch is regulated by posttranslational modifications, such as monoubiquitylation of PCNA in response to DNA damage that displaces replicative polymerase with the TLS polymerase, and ubiqtuitylation of TLS polymerases for removal (Figure 1.1) (Friedberg et al., 2005; Zhao and Washington, 2017). TLS polymerases Rev1, Pol ζ and Pol η are conserved from yeast to human (Sale, 2013). TLS polymerases in C. elegans protect against a wide range of DNA damaging agents (Roerink et al., 2012). Fanconi anemia (FA) Pathway The FA pathway repairs DNA interstrand crosslinks by coordinating a number of different repair pathways. The canonical FA pathway starts with damage recognition by FANCM during DNA replication. FANCM mediates the regression of both replication forks and recruits the FA core complex, including FANCA, FANCB, FANCC, FANCE, FANCF, FANCG and FANCL, which then monoubiquitylates FANCD2-FANCI. Monoubiquitylated FANCD2 recruits the nucleases FAN1, MUS81 and XPF that incise both ends of the lesion on one strand, unhooking the crosslink. Following that, the nascent strand opposite to the unhooked ICL is extended by a translesion polymerase, whereas the DSB induced by the incision is repaired by HDR (Figure 1.1) (Ceccaldi et al., 2016b; Deans and West, 2011). C. elegans appears to lack the core complex 8components but does have homologs of FANCM, FANCJ and FANCD2-FANCI, which when mutated result in sensitivity to interstrand crosslinking agents (Youds et al., 2009). 1.2.4 The interplay between DNA repair pathways Different repair pathways have lesion specificity, but some pathways are functionally compensatory. For example, HDR, NHEJ and MMEJ all repair DSBs, and pathway usage depends on the nature of the DSB, the cell cycle stage, and the availability of repair templates. Much of this repair pathway choice is mediated by the antagonistic relationship between the proteins 53BP1 and BRCA1 that affects end resection (Hustedt and Durocher, 2016; Panier and Boulton, 2014). It has recently been shown that the shieldin complex acts as an effector of 53BP1 in DSB-repair pathway choice by preventing DSB end resection (Gupta et al., 2018; Noordermeer et al., 2018). The interplay between DNA repair pathways is particularly important in cancer because repair mechanisms are often dysregulated, and the reduced replication and repair fidelity can generate greater genetic diversity. For example, upregulation of an error-prone repair mechanism like MMEJ has been found to be common in breast cancer and is associated with poor clinical outcomes (Higgins et al., 2010; Lemee et al., 2010). Dysregulated DNA repair mechanisms could result in a compromised capability to deal with intrinsic DNA lesions as well external chemical insults, which can be exploited for anticancer therapy (Nickoloff et al., 2017). Much work has focused on exploiting the DNA damage response to treat cancer (Nickoloff et al., 2017). An emerging approach is to exploit the concept of synthetic lethality (SL). DNA repair deficiencies in cancer can be made intolerable by inhibiting parallel functional DNA repair 9pathways (Chan and Giaccia, 2011; O'Neil et al., 2017). One successful example is PARP (Poly (ADP-ribose) polymerase) inhibitors, which have been approved to treat BRCA1/2-deficient breast and ovarian cancer patients (Lord et al., 2015; Tutt et al., 2010). The knowledge of mechanism underlying the SL between BRCA1/2 and PARP is still incomplete. Early work showed that co-inhibition of two parallel repair pathways (HDR and single-strand break repair) resulted in cell death (Fong et al., 2009). More recent work argued that PARP inhibitors induced trapping of PARP on the DNA that required functional HDR to bypass (Hopkins et al., 2015). A major challenge for targeting genome instability in tumors is that the genome instability causes tumor evolution that can rewire DNA repair pathways and acquire resistance to therapy. It has been shown that loss of factors that regulate the NHEJ repair mechanism can restore HDR and confer resistance to PARP inhibitors in Brca1-/- cells (Bouwman et al., 2010; Bunting et al., 2012; Bunting et al., 2010). Therefore, some genetic alterations provide an opportunity for anticancer therapeutic targets, while others can result in resistance. Identifying the genetic vulnerabilities that positively and negatively affect genome stability is useful for developing new therapeutic approaches. 1.3 Carcinogens and therapeutics that affect genome stability 1.3.1 Carcinogens Cancer can also be driven by exogenous factors. These factors, also known as carcinogens, promote tumorigenesis, often by causing genetic changes or mutations though interaction with DNA. For instance, individuals that have used aristocholic acid as part of traditional medicine are at high risk of liver and urothelial cancers, as aristocholic acid can form covalent bonds with 10DNA resulting in mutagenic lesions and genome instability (Arlt et al., 2002; Rosenquist and Grollman, 2016). Other nonchemical carcinogens, such as X-rays, or ultraviolet light, can also produce DNA damage, which if not appropriately resolved could result in mutations. 1.3.2 Therapeutics Counterintuitively, many anticancer therapeutics used to treat cancer patients negatively affect genome stability. The mutagenic activity of an anticancer therapeutic can be either its primary mode-of-action or a side-effect (Allan and Travis, 2005). Cancer cells often have DNA damage response or cell cycle regulatory defects that make them more sensitive to DNA damaging agents. This can result in a greater therapeutic index for DNA damaging agents in cancer cells compared to non-cancer cells. Consideration of the mutagenic effects of therapeutics is critical in developing anticancer strategies because these mutagenic therapeutics increase the risk of developing a secondary cancer in patients. One example is Hodgkin lymphoma patients, who have an increased chance of developing lung cancer after they receive ionizing radiation for treatment (Gilbert et al., 2003). In particular, ionizing radiation generates DSBs, which are toxic to cells, making it an effective anticancer treatment; however, error-prone repair of ionizing radiation-induced DSBs can create new sources of genome instability. Furthermore, the landscape of DNA repair competence can vary greatly post-treatment. Cells that escape a treatment may have developed new mutations that confer therapeutic resistance. Although anticancer therapeutics can cause genome instability that promotes the development of secondary tumors or confers resistance, the overall benefit of DNA damaging therapeutics is evident in the wide-range of common DNA damaging anti-cancer therapeutics. Hence, the key to 11improving DNA damaging therapeutics is a better understanding of how different cancer-associated genotypes respond to specific therapeutic agents to develop genotype-specific anticancer therapies that improve efficacy and eliminate or limit ineffective or harmful treatments. 1.4 Investigating genome instability As mentioned above, genome stability is affected by intrinsic and extrinsic factors. Decades of effort have been devoted into developing platforms and methods to study the causes and effects of genome instability. In general, the causes and effects of the genome instability are conserved across species. DSBs, single nucleotide mutations, and chromosome rearrangements are common outcomes observed in bacteria to yeast to humans. In general, the processes that maintain genome stability are highly conserved. In some instances, the specific players may be different but the overall maintenance and repair processes are remarkably similar. 1.4.1 Assays for measuring genome instability: micronuclei test to HPRT assay Assays for genome instability can be physical assays that measure chromosomal aberrations, such as the micronuclei assay which detects micronuclei formation, or the comet assay which detects DNA breaks. These assays allow direct visualization of genomic alterations events but are labor-intensive and thus challenging to conduct in a high-throughput manner. Other assays have been developed to look for phenotypic changes in colonies or cells. The most commonly used genotoxicity assay is the bacteria-based Ames test, which can assess the mutagenicity of chemicals and chemotherapeutics (Ames et al., 1973; Mortelmans and Zeiger, 2000; Wang et al., 2009). However, bacteria have different cellular conditions as compared to human cells. Despite 12some conserved DNA repair pathways between bacteria and eukaryotes such as MMR, bacteria lack many genome stability maintenance mechanisms that eukaryotes use in response to a mutagen, which can result in false negatives when assessing mutagenicity. Yeast-based genotoxicity tests have been developed by taking advantage of auxotrophic selection to screen mutants that acquire genotoxin-induced mutations. These yeast-based tests can assay genotoxins that induce various genome alterations, including gene deletion, gene conversion and mitotic recombination events (Schafer et al., 2008). Furthermore, hypoxanthine phosphorybosyl transferase (HPRT) mutagenesis assay offers a genotoxicity test in mammalian cell lines (Johnson, 2012). In these cell-based tests, the clonal population that have acquired the genotoxin-induced mutations can be sequenced to elucidate the genotypes or genetic changes. However, all these assays rely largely on the generation of mutations in single reporter genes that result in detectable phenotypic changes. This severely limits the spectrum of mutations that can be detected by these methods. 1.4.2 Defining the genome stability gene spectrum Large-scale screens have attempted to identify all the genes required for genome maintenance. Yeast has provided an excellent screening platform to identify genome maintenance genes, because of its low cost and easy manipulation as well available knockout mutant collections. A set of assays have been developed in yeast to conduct comprehensive screens for genome stability genes. In particular, these assays are able to detect increased rates of chromosome mis-segregation, gross chromosomal rearrangement, loss of heterozygosity, and gene conversion (Stirling et al., 2011; Yuen et al., 2007). Many genome maintenance genes identified in yeast screens are conserved in humans (Stirling et al., 2011; Yuen et al., 2007). Later work has 13complemented the gene lists and uncovered genes that caused genome instability when overexpressed (Duffy et al., 2016). These findings have implications for developing therapeutic strategies when combined with synthetic lethal or synthetic dosage lethal screens (Duffy et al., 2016; McLellan et al., 2012). Advances in cell culture, and RNAi-based gene knock-down approaches have made large-scale screening possible in human cancer cell lines. Large scale RNAi screens using mammalian cell lines have delineated genes and cellular processes required for genome integrity. Several have used DNA damage response markers (e.g. phospho-H2AX antibody) as a readout to identify key pathways mediating genome maintenance (Kavanaugh et al., 2015; Paulsen et al., 2009). Some screens further identified the therapeutic-sensitive and -resistant genotypes in multiple cancer cell lines (Barretina et al., 2012; Basu et al., 2013; Garnett et al., 2012). This is meaningful for better understanding the interaction between genotypes and therapeutics, and providing insights of the mechanisms of anti-cancer therapeutics and biology of genome care-taking genes. However, the complex heterogeneous genetic background of cancer cells and the inherent genome instability of these lines can confound the analysis. Recently, the emergence of the genome-editing technology CRISPR (clustered regularly interspaced short palindromic repeats) has made it possible to conduct more powerful and complete loss-of-function genetic screens. Moreover, further advances in sequencing technology enables an economic approach to sequence the genome. CRISPR and advanced sequencing technology has accelerated the era of high-throughput genome-wide screens. For instance, a CRISPR knockout screen for resistance to the DNA Topoisomerase II inhibitor etoposide has 14successfully identified TOP2A and CDK6 (Wang et al., 2014), while another CRISPR screen identified new factors that mediate resistance to the PARP inhibitor olaparib (Zimmermann et al., 2018). 1.4.3 Mutational signatures Advances in next-generation sequencing, have also made it possible to directly characterize the mutational footprints associated with genome stability, including genomic instability caused by loss of function of genome maintenance genes or by treatment with genotoxic agents. Mutational footprints are informative, because they are the visible by-products of the interactions between DNA damage and DNA repair mechanisms. Unique mutational footprints are also known as mutational signatures. Whole-genome sequencing analysis of cells or strains lacking specific repair mechanism can provide mutational signatures associated with repair deficiencies (Meier et al., 2014; Segovia et al., 2017; Serero et al., 2014). Several studies have revealed deep mutational signatures in various cancers and their association with genetic deficiencies and carcinogen exposure (Alexandrov et al., 2013; Roberts et al., 2013). These mutational signature catalogs can be informative for assigning genetic deficiencies or mutagen exposures to newly sequenced tumors. For instance, a tumor with a mutational signature containing a large number of substitutions and small insertions or deletions is likely to be defective in DNA mismatch repair, while another tumor could have a signature consistent with ultraviolet-light-induced mutations. Recently, Kucab and colleagues elucidated comprehensive mutational signatures of a number of carcinogens and anticancer therapeutics (Kucab et al., 2019). This lays the foundation for therapy-induced mutational signatures in human stem cells that provides key insights into the mechanisms of mutagenesis. 15Mutational signatures are, however, limited to those that do not cause severe effects on viability. Mutational signatures involving large chromosomal alterations that result in copy number variations (CNVs) are underrepresented because large CNVs, unlike point mutations, tend to be infrequent in cell populations because of the negative selection against such deleterious genomic changes (Gordon et al., 2012). In addition, there are limitations of short read sequencing in finding large structural chromosomal alterations (Pollard et al., 2018). Nonetheless, mutational signatures provide valuable information for the understanding of DNA damaging factors in cancer etiology with potential implications for prevention and treatment. 1.5 C. elegans as a platform to study factors affecting genome stability Studies in yeast and mammalian cells provide useful insights into how genes and therapeutics affect genome stability. However, there are limitations associated with yeast and mammalian cell line experiments. For instance, yeast lack many DNA damage response and repair genes such as PARP or BRCA1, and genetic heterogeneity in mammalian cell lines can confound the analysis of interactions between genotypes and therapeutics. Here, we are using a model organism, Caenorhabditis elegans to address these limitations. C. elegans is a multicellular invertebrate animal that combines the technical advantages of a microorganism (such as yeast) with greater biological complexity and a gene complement more akin to human. The nematode model was first proposed by Nobel Laureate Sydney Brenner in the early 1960s to be employed as a model for studying developmental biology and neurobiology (Goldstein, 2016). C. elegans feeds on bacteria and is easy and inexpensive to culture in the laboratory. C. elegans adults are only about 1mm long and 0.1mm in width, and it only takes three days for C. elegans to develop from a fertilized oocyte to a multicellular adult nematode worm. Availability of both hermaphrodites 16and males makes genetic manipulations quick and simple. Over the past decades, its simplicity, small size, relatively small genome, evolutionarily conserved mechanisms, and suite of experimental tools have made C. elegans an instrumental platform for elucidating biological processes implicated in human diseases, including cancer (Corsi et al., 2015). C. elegans can be used to study genes required for genome stability to gain insights into human cancer. Genome maintenance mechanisms are well conserved between C. elegans and human, and the majority of human genome stability-associated genes have homologs in C. elegans. For example, C. elegans homologs of the cancer-associated BRCA1 and BRCA2 genes have been identified and shown to be required for DNA repair, whereas no homologs are present in the yeast S. cerevisiae (Boulton et al., 2004; Martin et al., 2005). There are near isogenic C. elegans mutant strains available for most genes, including many well-characterized DNA replication and repair mutants. This provides a powerful resource for studying the biology of genome stability genes. C. elegans is multicellular and therefore has many more sophisticated phenotypes that can be associated with genome instability, including brood size, embryo survival, frequency of spontaneous males, and animal morphologies. There are many tools available for C. elegans such as genetic balancers that suppress meiotic recombination and allow for sterile or lethal mutations to be maintained. Genetic balancers can be used to capture and maintain lethal mutations (such as complex chromosomal rearrangements and CNVs) that tend to be lost in mutagenesis screens or mutational signature analysis in mammalian cells. C. elegans has a relatively small genome (~100Mb), and thus it is more economical to conduct deep whole-genome sequencing than mammalian cells. 17C. elegans can also be used to study the effect of chemicals or environmental factors on genome stability (Craig et al., 2012; Shin et al., 2019). These assays rely on the phenotypic changes such as embryo viability or spontaneous male frequency to identify whether an agent increases genome instability. Combining chemical sensitivity studies with different genome stability mutants allows the identification of anticancer chemi-genetic interactions. This can add to the knowledge of genotypic dependencies, mechanisms of therapeutics, and insights into anticancer treatment. C. elegans has also proven useful for determining mutational frequencies (Denver et al., 2005; Rosenbluth et al., 1983; Rosenbluth et al., 1985). There is a large body of research in C. elegans that has revealed distinct mutational signatures related to different carcinogens and DNA repair deficiencies (Meier et al., 2014; Meier et al., 2018; van Schendel et al., 2016). For instance, self-propagating C. elegans MMR mutants (pms-2 and mlh-1) exhibited extremely high base substitutions and 1-bp insertions or deletions, recapitulating the mutational signatures observed in MMR-deficient cancer samples (Meier et al., 2018). Examples such as these demonstrate the evolutionary conservation between C. elegans and human, highlighting the potential of C. elegans research to gain an understanding of human diseases associated with genome stability. A good example of the utility of C. elegans in exploring genome instability is the study of the C. elegans FANCJ homolog, dog-1. The dog-1 mutants display occasional specific morphological defects, such as notched-head, molting defects, and the Dumpy phenotype (Cheung et al., 2002). These specific phenotypes were mapped, and PCR-based sequencing analysis of these dog-1-induced mutations revealed that all were caused by genetic alterations in poly-guanine tracts, 18suggesting that DOG-1 was required to maintain poly-guanine tracts in the genome (Cheung et al., 2002). The dog-1 mutants are sensitive to DNA damaging agents that create interstrand crosslinks (Youds et al., 2008), which is consistent with the role of the human dog-1 homolog FANCJ, a member of the Fanconi Anemia repair pathway (Bridge et al., 2005). Using the genetic balancer eT1, a reciprocal translocation of half of chromosome III and half of chromosome V, a dog-1-induced mutation screen identified complex chromosomal rearrangement events and CNVs in the absence of dog-1 (Zhao et al., 2008). These chromosomal rearrangements often resulted in lethality in mammalian cells, but were able to be maintained using eT1 balancer in C. elegans. Further genetic interaction and whole-genome sequencing studies revealed that these dog-1-induced poly-guanine deletions were mediated by polymerase theta (Koole et al., 2014). 1.6 Research objectives Cancer is driven by genetic and environmental factors. These genetic and environmental factors can also enhance or suppress the efficacy of anti-cancer therapeutics. Key to the adoption of targeted anticancer therapies is a better understanding of the interactions between therapeutic agents and the tumor genotype and environment. Here, we used C. elegans to explore how genes, therapeutics and their interactions affect genome stability. The first aim of this thesis is to generate a chemi-genetic interaction map for well-characterized therapeutics or genotoxic agents in C. elegans (Chapter 2), which can be used as a reference for investigating new anticancer therapeutics. We then applied the chemi-genetic interaction map to further investigate the mechanism of action for an emerging anticancer therapeutic agent, CX-5461 (Chapter 3). The second aim is to use C. elegans to characterize genetic interactions within and between DSB repair pathways under different conditions, since DSB repair is a process often dysregulated in 19cancer (Chapter 4). The third aim is to study the biology of a less understood genome stability gene, the annealing helicase, SMRC-1 (Chapter 5). We characterized the mutational signature of loss of SMRC-1 and identified novel genetic interactions with SMRC-1. 20UVCUVCGG-NER TC-NERXPCRNAPIICSA, CSBXPATFIIHXPGXPF-ERCC1POLHLIG1NERMSH2 MSH6PMS2 MLH1EXO1POLDMMRLIG1KU70/80POLQEnd-resectionNHEJHDRMMEJDSBMUS81/BTR complexCtIP, MRE11LIG4 BRCA2, RAD51Homology search & strand invasionRecognitionGap-fillingLigation RecognitionNicking Error-removalRe-synthesisLigationExcisionUnwinding21BlockPOLHPCNA-UbPCNA POLDPOLDTLSPolymerase-switchPolymerase-switchBlockFANCM,FA core complexFANCD2, FANCIFAN1, MUS81, XPFHDRNERTLSFAFigure 1.1 Schematics of different DNA repair pathways. See main texts for details. 22CHAPTER 2: GENERATION OF A CHEMI-GENETIC INTERACTION MAP OF WELL-CHARACTERIZED DNA DAMAGING AGENTS 2.1 Introduction Advances in genome sequencing technology are transforming cancer treatment (Hyman et al., 2017). It is now routine to identify genetic and epigenetic changes that differentiate a patient’s tumor cells from non-tumor cells. Tumor-specific genetic alterations not only reveal the biological changes driving tumor progression but also uncover genetic vulnerabilities that can be exploited to selectively target the tumor with therapeutics. This means that therapeutic selection can be informed by the genetic variations in a tumor rather than tumor location. This tumor agnostic cancer treatment has the potential to offer individualized, highly specific therapies with fewer side-effects and reduce the overtreatment of tumors. Personalized oncogenomic approaches have had success treating tumors that have failed to respond to standard therapies (Laskin et al., 2015; Stockley et al., 2016; Swanton et al., 2016). The key to successful tumor agnostic therapeutic selection is understanding how tumor-specific mutations affect the response and resistance to specific chemotherapeutic agents. Each drug or chemotherapeutic has a unique genotypic sensitivity profile and each genetic mutation exhibits a distinctive drug sensitivity profile. Most chemotherapeutic agents target the uncontrolled proliferation of cancer cells, which is often associated with loss of function of DNA replication or repair genes. Many cancer-specific alterations of DNA replication and repair genes have been identified, including BRCA1, BRCA2, and TP53 (King et al., 2003; Olivier et al., 2010). Cells harbouring these mutations have been shown to be more responsive to certain chemotherapies (Helleday et al., 2008). While some genotype-chemotherapy connections have been identified, it 23is likely that many interactions have not yet been discovered. There is a pressing need to characterize chemi-genetic interactions between chemotherapeutics and tumor-specific DNA replication and repair gene mutations. One approach to this challenge is to generate a chemi-genetic interaction map, in which each gene mutation is exposed to each chemotherapeutic and the effect on viability is quantified. A chemi-genetic interaction map connects specific gene mutations to drug sensitivity and resistance. A chemi-genetic interaction map can link chemotherapeutics to patients based on the mutations in their tumor, and can be used to characterize the mechanisms of action for chemotherapeutics. The map could also be used as a reference, so that one can compare a genotypic sensitivity profile of a less-understood agent to well characterized agents and speculate on its mechanisms of action. Similarly, one can predict the biological function of a DNA damage response gene based on the drug sensitivity profiles of the mutant. Many efforts have been devoted to generating a genotypic sensitivity profile for DNA damaging agents in yeasts using a comprehensive gene knock-out library (Chang et al., 2002; Hanway et al., 2002). Genome-wide small hairpin RNA (shRNA) screens in cancer cell lines have been developed to uncover genes that are required for drug resistance (O'Connell et al., 2010). More recently, CRISPR-Cas9 genome-wide screens have also identified a number of genetic vulnerabilities for different drugs (Hart et al., 2015; Zimmermann et al., 2018). Although there are discrepancies between screens due to differences in the genetic backgrounds of the cell lines, it is still a useful resource to identify genetic changes that give rise to the drug sensitivity or resistance and provide guidance for potential personalised cancer treatment. 24In this chapter, we used C. elegans to generate a chemi-genetic interaction map, comprised of DNA replication and repair genes and agents that could induce DNA damage. Since many DNA replication and repair processes are well conserved between C. elegans and human, it is likely that these chemi-genetic interactions can be extrapolated to human cancer cells. We included five different well-characterized DNA damaging agents (DDAs): olaparib, camptothecin, etoposide, UVA + trimethylpsoralen, and ultraviolet (UV) C radiation. The agents were chosen to cover of a broad spectrum of DNA damage types. Each agent generates a different type of DNA lesion that should require a specific set of repair mechanism(s) to resolve (summarized in Table 2.1). Specifically, olaparib inhibits PARylation activity, and traps PARP proteins on DNA, which can block DNA replication (Pommier et al., 2016). Olaparib has been approved by FDA to treat breast and ovarian cancer patients with germline BRCA1/2 mutations (Lord et al., 2015; Tutt et al., 2010). Camptothecin is a Topoisomerase I inhibitor, and it binds to the DNA-Topoisomerase I complex, which prevents normal DNA-religation, resulting in single-strand breaks (SSBs) that can be converted to double-strand breaks (DSBs) upon collision with the replication machinery (Pommier, 2013). Two camptothecin derivative compounds, topotecan and irinotecan, have been approved by FDA to treat different types of cancers (Pommier, 2013). Etoposide is a Topoisomerase II inhibitor, and it binds to DNA-Topoisomerase II complex, which blocks strand re-ligation resulting in DSBs (Nitiss, 2009). Etoposide has been used in combination with other chemotherapeutics as the first-line treatment for small cell lung cancers (Pommier, 2013). UVA + trimethylpsoralen induces interstrand DNA crosslinks that impede DNA replication and transcription (Deans and West, 2011). Crosslinking agents such as the platinum-based cisplatin, carboplatin, and oxaliplatin are commonly used to treat cancer (Huang and Li, 2013). UVC 25radiation causes DNA bulky lesions, such as thymine dimers (Rastogi et al., 2010). UVC is not used as an anti-cancer therapeutic. 2.2 Methods Nematode strains were maintained as described previously (Brenner, 1974). The alleles used in this study were: atm-1(tm5027), brd-1(dw1), rfs-1(ok1372), cku-80(ok861), lig-4(ok716), hsr-9(ok759), polq-1(tm2026), polh-1(lf31), polk-1(lf29), fcd-2 (tm1298), fan-1(tm423), fncm-1(tm3148), msh-2(ok2410), ercc-1(tm1981), xpa-1(ok698), mus-81(tm1937), rcq-5(tm424), rtel-1(tm1866), helq-1 (tm2134), him-6(ok412), dog-1(gk10), wrn-1(gk99), let-418(n3536), him-1(e879), hda-3(ok1991), gld-1(op236), cep-1(gk138), dvc-1(ok260), and rev-3 (gk919715). Bristol N2 was used as wild type in all experiments. rev-3(gk919715) was Million Mutation Project allele (Thompson et al., 2013) provided by the Moerman lab and outcrossed to N2 six times. Some strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440), and some knockout alleles were provided by the Shohei Mitani laboratory. Some strains were generated by the International C. elegans Gene Knockout Consortium (Consortium, 2012) and by the National Bioresource Project of Japan. UVA irradiation UVA Source: predominantly 365 nm, Black-Ray®UV Bench Lamp (Model: XX-15L). Before each UVA exposure, the light source output was determined by a longwave ultraviolet measuring meter (Model: J-221). Different UVA exposures were achieved by varying the exposure times. 26Quantitative Acute Assay for olaparib, camptothecin, and etoposide Synchronized one-day-old adults were incubated in agents (500 µM olaparib, 100 nM camptothecin, 100 µM etoposide) or Dimethyl sulfoxide (DMSO) solvent diluted in M9 buffer containing OP50, carbenicillin (50 µg/ml) and 1X nystatin for ~18 hours. Following treatment, the animals were allowed to recover for 30 to 60 minutes on OP50-containing Nematode growth media (NGM) plates. Next, at least ten worms were transferred in triplicate to NGM plates for a 4-hour interval (18 to 22 hours post-treatment), and then removed. The numbers of arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment (Figure 2.1). For animals that produced few progeny, more treated adult worms were picked to lay embryos in the 4-hour interval. Quantitative Acute Assay for UVA-Trimethylpsoralen Synchronized one-day-old adults were washed off plates and resuspended in 10 µg/mL TMP diluted in 500 µL M9 buffer containing 0.1% Triton for one hour in the dark. Following treatment, the animals were washed in M9 and then transferred to two OP50-containing NGM plates. Next, one plate was exposed to 100 J/m2 UVA irradiation. Both plates were kept in the dark to avoid UVA from light in the laboratory. At least ten worms were transferred in triplicate to NGM plates for a 3-hour interval and then removed. The numbers of arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment. For animals that produced few progeny, more UVA-TMP treated adult worms were picked to lay embryos in the 3-hour interval. Quantitative Acute Assay for UVC Synchronized one-day-old adults were exposed to 50 J UVC irradiation (or mock treatment). UVC source was provided by UV-cross-linker (Ultraviolet Translinker TL-2000, ULTRA-27VIOLET PRODUCTS) with 254 nm bulbs (crosslinker). Animals were kept in the dark overnight. At least ten worms were transferred in triplicate to NGM plates for a 2.5-hour interval (18-21 hours post-irradiation) and then removed. The numbers of arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment. For animals that produced few progeny, more irradiated adult worms were picked to lay embryos in the 2.5-hour interval. Chemi-genetic interaction map generation The chemi-genetic interaction map is generated by using sensitivity scores, and then color-coded (Figure 2.2). The sensitivity score is the double-normalized embryo survival. The detailed calculation is as followed. For each mutant, Embryo survival % = !"# %&'(#) *+ ",!-"#. /,)0,(!"# %&'(#) *+ ",!-"#. /,)0, 2 !"# %&'(#) *+ ,))#3!#. #'()4*3) x100%. Then, the embryo survival percentage of each mutant treated with DDA is normalized to that with mock treatment. For each mutant, Normalized embryo survival = 6'()4* 3&)070,/ % ),!# (!)#,!#. 97!" ::;)6'()4* 3&)070,/ % '*-< . Next, the sensitivity score is generated by normalizing the normalized embryo survival rate of the mutant to that of wild-type N2. For each mutant, Sensitivity score = =*)',/7>#. #'()4* 3&)070,/ ('&!,%!)=*)',/7>#. #'()4* 3&)070,/ (=?) . The higher the score, the less sensitive the mutant is to the DDA, and the lower the score, the more sensitive the mutant is to the DDA. Lastly, the map is color-coded with light blue representing the least sensitive and red representing the most sensitive. The sensitivity score takes the effect of both the DDAs and the genotypes into account. 282.3 Results 2.3.1 Construction of a genome stability mutant panel As mentioned in Chapter 1, many genes can affect genome stability, ranging from DNA metabolic genes to DNA damage response genes. Here, we constructed a C. elegans genome stability mutant panel that covers a wide range of functions involved in safeguarding the genome integrity, especially DNA replication and repair. The mutant panel consisted of 30 different mutants in total in addition to wild-type N2, which served as a negative control (Figure 2.2). We included a cohesin mutant (him-1), chromatin modifier mutants (let-418 and hda-3), an RNA-binding mutant (gld-1), checkpoint mutants (atm-1 and cep-1), an endonuclease mutant (mus-81), a number of helicase mutants (dog-1, helq-1, him-6, rcq-5, wrn-1 and rtel-1), a kinase mutant (cdk-5), FA repair pathway mutants (fncm-1, fan-1 and fcd-2), NER mutants (xpa-1 and ercc-1), a MMR mutant (msh-2), HDR mutants (brd-1 and rfs-1), NHEJ mutants (cku-80, lig-4 and hsr-9), a MMEJ mutant (polq-1), translesion synthesis mutants (polh-1, rev-3 and polk-1) and a protease mutant (dvc-1) (Figure 2.2). 2.3.2 Genotypic sensitivity profile of olaparib (Ola) We first generated the genotypic sensitivity profile of a PARP inhibitor, olaparib. We treated the mutant panel with 500 µM olaparib and scored progeny viability of treated animals. Three out of thirty mutants showed strong sensitivity to olaparib. The DNA damage response checkpoint mutant, atm-1, and the endonuclease mutant, mus-81, exhibited the greatest sensitivity – about 56% and 67% embryo viability after olaparib treatment (Figure 2.2). To our surprise, the brd-1 mutant was not sensitive to olaparib. This is in contrast to human cells, where tumors deficient in 29BRCA1 or BARD1 exhibited severe proliferation defects when treated with olaparib (Tutt et al., 2010). Another HDR mutant, helq-1 had mild olaparib-sensitivity, which produced around 80% viable progeny after olaparib treatment (Figure 2.2). 2.3.3 Genotypic sensitivity profile of camptothecin (CPT) Next, we generated the genotypic sensitivity profile of the Topoisomerase I inhibitor, camptothecin with the mutant panel. About a third of the mutants we tested exhibited CPT sensitivity. Similar to olaparib, the endonuclease mutant, mus-81, and the checkpoint mutant, atm-1, were among the most sensitive strains (Figure 2.2). Notably, the cohesin mutant, him-1, was also sensitive to CPT, as over 60% progeny were arrested as dead embryos upon CPT treatment. The NHEJ mutants (cku-80, lig-4 and hsr-9) and the MMEJ mutant (polq-1) were not sensitive to CPT treatment, whereas HDR mutants (including brd-1, rfs-1 and helq-1) were hypersensitive, all of which have reduced embryo viability (>80% arrested embryos) after CPT treatment (Figure 2.2). The mismatch repair mutant, msh-2, was also sensitive to CPT (Figure 2.2). 2.3.4 Genotypic sensitivity profile of etoposide (ETOP) Next, we generated genotypic sensitivity profile of etoposide, an inhibitor of Topoisomerase II. The mus-81 mutant exhibited the greatest sensitivity to ETOP, and the cohesin mutant him-1, was hypersensitive to ETOP (Figure 2.2). Although ETOP and CPT shared many sensitive genotypes, there were clear differences in the sensitivity levels. For instance, HDR mutants (brd-1, rfs-1, helq-1) were less sensitive to ETOP than to CPT. The MMEJ mutant, polq-1, which did 30not exhibit CPT-sensitivity, was sensitive to ETOP treatment (Figure 2.2). In addition, two helicase mutants, rtel-1 and dog-1, were very sensitive to ETOP treatment (Figure 2.2). 2.3.5 Genotypic sensitivity profile of UVA-trimethylpsoralen (TMP) Next, we generated the sensitivity profile for UVA-trimethylpsoralen, a DNA interstrand crosslinking (ICL) agent (Deans and West, 2011). TMP exhibited a very different genotypic sensitivity profile compared to other DNA damaging agents. Notably, C. elegans animals deficient in Fanconi Anemia (FA) pathways, which repairs ICLs, were sensitive to TMP. The fan-1 and fncm-1 mutants produced more than 70% arrested embryos upon UVA-TMP treatment (Figure 2.2). Nucleotide excision repair (NER) mutants, xpa-1 and ercc-1, showed high embryo lethality - ~ 70% and 80%, respectively (Figure 2.2). Mutants that lack translesion synthesis capacity, polh-1 and rev-3, were also extremely sensitive to UVA-TMP, producing more than 85% arrested progeny (Figure 2.2). Among the three major double-strand break (DSB) repair pathways, both MMEJ and HDR were required for TMP-resistance, as polq-1, brd-1 and rfs-1 single mutants were all sensitive to UVA-TMP (Figure 2.2). 2.3.6 Genotypic sensitivity of ultraviolet C (UVC) Next, we generated the genotypic sensitivity profile for ultraviolet C exposure. As was expected for UVC-induced DNA damage, the TLS mutants, polh-1 and rev-3, were extremely sensitive to UVC (Figure 2.2). The polh-1 mutant produced almost no viable progeny (>99% arrested embryos) after UVC exposure. The NER mutants xpa-1 and ercc-1 produced more than 60% dead embryos upon UVC exposure (Figure 2.2). Mutants deficient in resolving DSBs, such as HDR, NHEJ and MMEJ mutants, were not sensitive to UVC (Figure 2.2). 312.4 Discussion 2.4.1 Each agent has a distinctive genotypic sensitivity profile Taking into account the magnitude of sensitivity, each DNA damaging agent has a unique sensitivity profile. DNA damage induced by an agent can be complex consisting of different types of lesions requiring more than one repair mechanism. For example, UVC radiation can cause a range of DNA lesions, such as thymine dimers and bulky (6-4) photoproducts (Cadet et al., 2005). In other cases, the DNA damage requires the coordination of multiple repair pathways to repair the damage, as is the case for ICLs (Youds et al., 2009). These complexities result in sensitivity profiles that touch on multiple repair pathways. In most cases, DNA damage is primarily repaired or resolved by some mechanisms over others. For example, NER pathway mutants (xpa-1 and ercc-1) showed much greater sensitivity to UVA-TMP than HDR mutants (brd-1 and rfs-1) (Figure 2.2), suggesting that repair of TMP-induced lesions is more dependent on NER than HDR in C. elegans. Another way to look at the chemi-genetic interaction map is by comparing the profiles between agents. One can discern the primary repair mechanisms for each lesion induced by different agents. For example, CPT and ETOP are both topoisomerase poisons. Despite the fact that both CPT and ETOP can result in DSBs, the primary mechanism to resolve lesions induced by each agent varies. Three major pathways could resolve DSBs: HDR, MMEJ, and NHEJ. In both CPT and ETOP-induced lesions, NHEJ is dispensable for the repair, as evidenced by the fact that the cku-80 mutant showed no embryonic death upon either treatment (Figure 2.2). The data also suggest that HDR repair pathway is the primary repair pathway used for CPT-induced lesions because HDR mutants (brd-1, and rfs-1) were much more sensitive to CPT than the MMEJ 32mutant polq-1 (Figure 2.2), which did not exhibit CPT-sensitivity. By contrast, MMEJ mutant polq-1 had much stronger ETOP-sensitivity than HDR mutants (Figure 2.2), indicating that MMEJ is primary repair mechanism for resolving ETOP-induced DNA damage. This raises the possibility that the type of DSBs or lesions arising from CPT and ETOP can be different. The interplay between repair pathways is explored further in Chapter 4. We also identified features for each agent. Compared to human cell line studies, a high concentration of olaparib was needed (500 µM) to generate the sensitivity profile in C. elegans and only three mutants were found to be olaparib-sensitive. It is possible that either poor uptake into the worm or detoxification of olaparib required a high initial concentration to achieve an effective concentration in the worm (Johnstone, 1994). Alternatively, it is probable that 18 hours of exposure of adult worms to olaparib was not sufficient to induce DNA damage in the germ lines. In contrast to human counterparts (Min et al., 2013; Walsh, 2015), the HDR mutants (brd-1 and rfs-1) showed no sensitivity to olaparib using quantitative acute assays (Figure 2.2). This could be due to the properties of the assay. The readout for sensitivity is the embryonic survival of progeny (embryo) from the treated animals. The rapid early embryonic cell divisions in C. elegans do not utilize HDR pathways and instead rely heavily on TLS (Holway et al., 2006), which could bypass lesions caused by inhibited PARP. C. elegans had a genotypic sensitivity profile for CPT that was similar to the human cell lines (O'Connell et al., 2010). HDR-deficient human cells are hypersensitive to CPT, and C. elegans brd-1 and rfs-1 mutants were extremely sensitive to CPT (Figure 2.2). Furthermore, the mismatch repair gene MSH-2 was required for CPT-resistance likely because of its role in 33recombinational repair, similar to its human counterparts (Jacob et al., 2001; Pichierri et al., 2001). The NHEJ pathway has been proposed to antagonize the HDR pathway in response to CPT, and loss of NHEJ factors renders resistance to CPT (Adachi et al., 2004). Consistent with this, none of the C. elegans NHEJ mutants (cku-80, lig-4, and hsr-9) were sensitive to CPT (Figure 2.2). Our finding that the cohesin mutant, him-1, exhibited CPT-sensitivity was also consistent with a previous report that human glioblastoma H4 cells with a null mutation in the cohesin gene STAG2 were more sensitive to CPT than H4 cells that had the STAG2 mutation repaired (Bailey et al., 2014). Although some studies demonstrated that POLQ-deficient cells were sensitive to CPT (Wang et al., 2019b; Yousefzadeh et al., 2014), our results did not find the polq-1 mutant sensitive, indicating that MMEJ is not the primary repair pathway for CPT-induced lesions in C. elegans. This could be due to worm-specific differences in repair pathway choice or the result of different genetic backgrounds in the human cell lines. The genotypic sensitivity profile of ETOP overlapped with that of CPT (Figure 2.2), as many ETOP-sensitive mutants were also sensitive to CPT, albeit with varying degrees. The repair mechanism for ETOP-induced lesions appears to be well conserved between C. elegans and humans, as we found C. elegans homologs of many human ETOP-sensitive genes were also required for resistance to ETOP. Similar to human counterparts (Treszezamsky et al., 2007; Wang et al., 2019b), both HDR genes and MMEJ genes are required for ETOP-induced DNA damage. Moreover, our data suggested that MMEJ is the primary repair mechanism for ETOP-induced lesions since the MMEJ mutant, polq-1, exhibited much stronger sensitivity than HDR mutants (brd-1 and rfs-1) (Figure 2.2). It is as yet uncertain why C. elegans favors MMEJ over HDR to counteract ETOP-induced DNA lesions. One possibility is that ETOP did not directly 34cause DSBs but instead results in lesions that POLQ-1 could process via its TLS activity (Yoon et al., 2019). This hypothesis is supported by the observation that two C. elegans FANCJ helicase family mutants, dog-1 and rtel-1 were both sensitive to ETOP but not to CPT. DOG-1 is required to unwind G4 structures during replication (London et al., 2008; Youds et al., 2008), and in the absence of DOG-1, POLQ-1 is needed to bypass these structures (Koole et al., 2014). It is possible that ETOP is generating analogous structures that can be resolved either by DOG-1 helicase activity or POLQ-1-mediated bypass, in a similar manner to G4 structures. Alternatively, loss of FANCJ could result in chromatin re-organization that further impedes the repair of TOP2-induced lesions (Schwab et al., 2013). However, it is unknown whether TOP2 has high affinity towards DNA sequences that tend to form secondary structures, or that trapping of TOP2 by ETOP induces formation of secondary structures in the vicinity. The genotypic sensitivity profile of UVA-TMP illustrates the requirement for multiple repair processes in the resolution of interstrand crosslinks in C. elegans. The canonical interstrand crosslink repair involves cooperation of multiple pathways (Deans and West, 2011). XPA-RPA and MutSβ complexes have been previously shown to function together to recognize ICL, which likely induced structural distortion at the site of damage (Zhao et al., 2009). In support of this, our data showed that both xpa-1 and msh-2 mutants were sensitive to UVA-TMP (Figure 2.2). The sensitivity of MMEJ mutant polq-1 to UVA-TMP is consistent with a recent report showing that MMEJ plays a role in counteracting interstrand crosslink lesions in C. elegans (Figure 2.2) (van Schendel et al., 2016). That core FA mutants (fan-1, fcd-2 and fncm-1) were only sensitive to UVA-TMP, but not to other agents we tested (Figure 2.2), suggests that the FA pathway is very specific to the resolution of ICLs (Deans and West, 2011). TLS is required for the final step 35of ICL-repair to ligate the gap generated by endonucleases. In agreement with a previous report that HDR pathway is required for ICL-repair, such as after treatment with nitrogen mustard (Muzzini et al., 2008), both brd-1 and rfs-1 exhibited UVA-TMP-sensitivity (Figure 2.2). There are structural differences between ICLs created by different agents. UVA-TMP generates a specific type of abasic site different from nitrogen mustard and cisplatin after initial incision (Semlow et al., 2016). It has been shown that UVA-TMP-induced ICL-repair did not result in formation of DSBs due to activity of NEIL3 glycosylase (Semlow et al., 2016); however, it does not appear that C. elegans has a NEIL3 homolog that could prevent the formation of DSBs in response to TMP. The genotypic sensitivity profile of UVC elucidates how bulky lesions are repaired in C. elegans. Similar to their human counterparts, mutants deficient in NER and TLS are extremely sensitive to UVC (Figure 2.2). The repair mechanism is likely very similar in both worms and humans. When UVC generates bulky lesions such as thymine dimers, XPA-RPA initiates the recognition process along with RNA polymerase during transcription, XPF-ERCC1 and XPG cleave the lesions, and TLS fills the gap. 2.4.2 Some genes are required for many different DNA damage responses From our chemi-genetic interaction map, some mutants exhibited sensitivity to all agents tested. This could be that the gene is required for a general response to all DNA damage or that it is a key factor indispensable for many types of DNA repair. For instance, the endonuclease mutant, mus-81, showed strong sensitivity to all agents tested (Figure 2.2). MUS81 has multiple roles in the DNA damage response. It has been shown to cleave stalled replication forks to generate a 36substrate for HDR (Hanada et al., 2007) and act as a late cell cycle stage fail-safe for resolving unrepaired DNA lesions that block replication to ensure proper cell division (Naim et al., 2013; Ying et al., 2013). Therefore, it would be expected that mus-81 mutants would be sensitive to many different types of DNA damage. This property makes the mus-81 mutant a good positive control for testing agents that induce DNA damage. The DNA damage response checkpoint mutant, atm-1, was also sensitive to all agents tested (Figure 2.2). Human ATM-deficient cells were also shown to be sensitive to olaparib, CPT, ETOP, UVC and other DNA damaging agents (Bailey et al., 2014; Fedier et al., 2003; Hannan et al., 2002). The ATM checkpoint is activated in DNA damage response and can phosphorylate a number of substrates depending on the type of DNA damage (Marechal and Zou, 2013). These substrates are proteins that are required for removing or resolving the damage. Given the role that ATM-1 plays in the DNA damage checkpoint, it is not surprising that it is sensitive to many different types of DNA lesions. 2.4.3 The chemi-genetic interaction map can be a reference for characterizing new damaging agents and DNA replication and repair genes The chemi-genetic interaction map can be useful as a reference for understanding how less-understood DNA damaging agents work. New agents can be tested with the same panel of C. elegans DNA replication and repair mutants, and new genotypic sensitivity profiles can be generated. Comparing the genotypic sensitivity profile of a less-understood agent with those of well-characterized ones provides a preliminary hypothesis about what type of DNA lesions are induced by the agent under investigation. Agents that induce similar types of lesions require 37similar repair mechanism to resolve the lesions, so there will be overlap in the mutants that show sensitivity to the agents. Conversely, the chemi-genetic interaction map can be used to investigate the biological function of less-understood genes involved in DNA replication and repair. Testing the animals deficient in the gene of interest with all the agents in the chemi-genetic interaction map creates the drug-sensitivity profile for the gene of interest. Drug-sensitivity profile allows us to determine whether the gene of interest is involved in general DNA damage response that is indispensable for repair (such as mus-81 and atm-1), or is dedicated to repair only certain types of DNA lesions (such as genes involved in Fanconi Anemia pathway). Although there are differences between how cells repair certain kinds of DNA damage between C. elegans and human, the chemi-genetic interaction map here is solely generated using worm data. Even if there are differences between worm and human repair pathway choices for a given lesion, the C. elegans panel is still informative because similar types of lesions should require the same pathways for repair or tolerance. The chemi-genetic interaction map is a growing resource that will become even more useful as more mutants and DNA damaging agents are tested and catalogued. 38Recover 50 Young adults Lay embryos(3 biological replicates)Remove adults~4 hours~18 hours Figure 2.1 A schematic of quantitative acute assay. About 50 young adults were treated with agents for ~ 18 hours (alternatively one hour in trimethylpsoralen followed by UVA irradiation, or transient UVC irradiation). Then treated adults were often transferred to a plate to recover for 30 to 60 minutes, before they were picked to new plates in 3 biological replicates (at least 10 worms on each plate). Adults were given a four-hour interval to lay embryos before they were removed. The numbers of unhatched embryos and new larvae were counted on the following day. Leave overnightScore~30-60 minsDNA damaging agent Type(s) of lesions Known DNA repair pathway(s) to resolve the lesionsClinical implicationOlaparib Protein trapping and replication block HDROlaparib is used to treat breast and ovarian cancer patients with germline BRCA mutations.CamptothecinReplication block, single-strand breaks, and double-stranded breaksHDRCamptothecin derivatives (topotecan and irinotecan) are used to treat various types of cancer, including ovarian, cervical and lung cancer.Etoposide Replication block, double-strand breaks HDREtoposide is used in combination with other chemotherapy to treat small cell lung cancer and testicular cancer.UVA + trimethylpsoralen (UVA+TMP)DNA interstrand crosslinks, replication blockFA, HDR, MMEJ, TLS and NERUVA+TMP is not used in the clinics. Many platinum-based anticancer drugs (cisplatin, carboplatin, and oxaliplatin), which cause DNA interstrand crosslinks, are used to treat various types of cancer, including ovarian cancer, lung cancer and head and neck cancer. UVCDNA bulky lesions, including cyclobutane pyrimidine dimers and [6-4]-photoproductsNER and TLS UVC is not used in the clinic. Table 2.1 Summary of the DNA damaging agents used to generate the chemi-genetic interaction map. 39Figure 2.2 The chemi-genetic interaction map of different agents. Indicated mutant animals were treated with corresponding agents, and their embryo survival percentage was calculated. The color range (from blue to red) represent differential sensitivity of mutants as indicated in the scale bar. The calculation of the sensitivity score is detailed in Chapter 2 Method, and raw data are in Table A1. ND, not determined. 0.85 0.75 0.5 0.25 0Not sensitive Most sensitive Function Human homolog Strain Ola CPT ETOP UVA + TMP UVC CX-5461/// wild-type N2 1.00 1.00 1.00 1.00 1.00 1.00Cohesin SMC1A him-1 1.01 0.33 0.08 0.82 0.74 0.39Chromatin remodeling CHD4 let-418 1.02 1.00 1.00 0.50 1.04 0.76HDAC1 hda-3 0.87 1.15 0.83 0.70 0.93 0.96RNA binding QKI gld-1 0.95 0.80 0.95 1.04 0.88 0.87DNA Damage CheckpointATM atm-1 0.56 0.36 0.73 0.67 0.63 0.01TP53 cep-1 1.03 0.60 0.94 0.96 0.71 0.98Endonuclease MUS81 mus-81 0.67 0.07 0.06 0.13 0.36 0.03HelicaseFANCJ dog-1 0.92 0.91 0.50 0.83 0.91 0.95HELQ helq-1 0.80 0.12 0.77 0.48 0.79 0.11BLM him-6 1.15 0.97 ND 0.86 0.82 NDRECQL5 rcq-5 1.02 1.09 0.98 1.03 0.94 0.99RTEL rtel-1 0.96 0.99 0.49 0.63 1.09 0.79WRN wrn-1 0.98 0.92 1.15 0.97 1.14 0.98SMARCAL smrc-1 ND 0.12 0.43 0.75 0.70 0.24Kinase CDK5 cdk-5 1.03 0.84 ND 0.87 0.89 NDFanconi Anemia PathwayFANCM fncm-1 1.00 1.01 1.05 0.28 0.82 0.91FAN1 fan-1 0.90 1.01 1.07 0.11 0.98 0.95FANCD2 fcd-2 1.00 1.00 1.13 0.55 0.99 0.87HDR repairBARD brd-1 1.04 0.17 0.82 0.52 0.93 0.59RAD51C rfs-1 0.96 0.19 0.55 0.49 1.00 0.55Nucleotide Excision RepairERCC1 ercc-1 1.07 1.00 1.02 0.19 0.17 0.08XPA xpa-1 0.95 0.99 0.92 0.31 0.36 0.27MMR MSH2 msh-2 1.02 0.42 0.79 0.25 1.12 1.03NHEJTP53BP1 hsr-9 0.90 0.92 ND 0.99 0.90 0.98KU80 cku-80 1.02 1.00 1.05 0.89 1.02 0.99LIG4 lig-4 0.97 0.93 1.08 1.03 0.98 1.00MMEJ POLQ polq-1 1.01 0.95 0.32 0.55 1.06 0.44Translesion Synthesis (TLS)POLH polh-1 0.99 0.62 0.69 0.00 0.02 0.09REV3 rev-3 0.98 0.96 1.02 0.16 0.51 0.01POLK polk-1 1.04 1.08 ND 1.01 1.04 NDProtease SPRTN dvc-1 ND 1.03 0.91 0.70 0.95 0.8540CHAPTER 3: UNDERSTANDING THE MECHANISMS OF ACTION OF CX-5461 3.1 Introduction CX-5461 is a quinolone-containing small molecule with potential anti-cancer properties. It was originally described as an RNA Pol I inhibitor that demonstrated in vivo anticancer activity in murine xenograft models (Drygin et al., 2011; Haddach et al., 2012). CX-5461 can trigger ATM/ATR checkpoint activation (Negi and Brown, 2015) and rapamycin-associated signaling (Li et al., 2016). More recently, we contributed to a study that showed that CX-5461 binds and stabilises G-quadruplexes (G4s) and causes selective killing of BRCA1/2-deficient tumors (Xu et al., 2017). This has led to a CX-5461 clinical trial in breast cancer patients with homologous recombination-deficient tumors (Hilton et al., 2018). Clinical trials have identified side-effects of CX-5461, including photosensitivity and palmar-plantar erythrodysesthesia (Hilton et al., 2018; Khot et al., 2019). It is imperative to further investigate the mechanism of action of CX-5461 so as to have a better understanding of genotypic sensitivities and resistance, and potential deleterious effects. 3.2 Methods The alleles used in this study were: BC 2200 dpy-18(e364)/eT1(III); unc-46(e177)/eT1(V), atm-1(tm5027), brd-1(dw1), rfs-1(ok1372), cku-80(ok861), lig-4(ok716), hsr-9(ok759), polq-1(tm2026), polh-1(lf31), polk-1(lf29), fcd-2 (tm1298), fan-1(tm423), fncm-1(tm3148), msh-2(ok2410), ercc-1(tm1981), xpa-1(ok698), mus-81(tm1937), rcq-5(tm424), rtel-1(tm1866), helq-1 (tm2134), him-6(ok412), dog-1(gk10), wrn-1(gk99), let-418(n3536), him-1(e879), hda-3(ok1991), gld-1(op236), cep-1(gk138), dvc-1(ok260), smrc-1(gk176502), and rev-3(gk919715). Bristol N2 was used as wild type in all experiments. smrc-1(gk176502), and rev-3(gk919715) 41were Million Mutation Project (Thompson et al., 2013) alleles provided by the Moerman lab and outcrossed to N2 six times. Some strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440), and some knockout alleles were provided by the Shohei Mitani laboratory. Some strains were generated by the International C. elegans Gene Knockout Consortium (Consortium, 2012) and by the National Bioresource Project of Japan. UVA irradiation UVA Source: predominantly 365 nm, Black-Ray®UV Bench Lamp (Model: XX-15L). Before each UVA exposure, the light source output was determined by a longwave ultraviolet measuring meter (Model: J-221). Different UVA exposures were achieved by varying the exposure times. Quantitative acute assay Synchronized one-day-old adults were incubated in CX-5461 (in NaH2PO4) diluted in M9 buffer containing OP50, carbenicillin (50 µg/ml) and 10 ng/ml nystatin for ~18 hours. Following treatment, the animals were allowed to recover for 30 to 60 minutes on OP50-containing NGM plates before UVA irradiation (if applicable) and then plated ten worms per plate in triplicate on NGM plates for a 4-hour interval (18 to 22 hours post-treatment), and then removed. The numbers of both arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment. For animals that produced few progeny, more treated adult worms were picked to lay embryos in the 4-hour interval. All results were from at least 30 treated animals (3 plates with at least 10 animals per plate). The calculation of the sensitivity score was detailed in Chapter 2 Method. 42Mutagenesis screen for CX-5461 Strain BC2200 dpy-18/eT1(III);unc-46/eT1(V) was used in the mutagenesis screen. dpy-18/eT1(III);unc-46/eT1(V) hermaphrodites were treated with or without CX-5461 for 18 hours before 150 J/m2 UVA irradiation, and 200 single dpy-18/eT1(III);unc-46/eT1(V) F1s were picked in each condition. A sterile phenotype at F1 is considered to indicate the acquisition of a dominant lethal mutation, and lines in which F2 or later generations that do not have Dpy Unc animals are considered to indicate the acquisition of a recessive lethal mutation in the balanced regions of chromosome III or V. Genome Sequencing The lines that acquired recessive lethal mutations were maintained for at least three generations. Worms were rinsed off with deionized water and concentrated by centrifugation. Genomic DNA was purified using Puregene® Core Kit A (Qiagen). Whole-genome sequencing was performed at the Novogene Bioinformatics Institute (Beijing, China). Sequence reads were mapped to the C. elegans reference genome version WS230 (http://www.wormbase.org) using the short-read aligner BWA, which gave an average sequencing depth for each sample ranging from 22x to 57x with a median of 34x. Single-nucleotide variants and small insertions/deletions were identified and filtered with the help of the SAMtools toolbox (Li et al., 2009). Candidate variants at genomic locations for which the parental N2 strain had an agreement rate with the reference genome lower than 95% were eliminated from further consideration. Each variant was annotated with a custom Perl script and gene information downloaded from WormBase version WS230. Copy numbers were estimated from the alignments with a procedure analogous to that of Itani et al. using 5 kb wide overlapping sliding windows with the alignments from the parental strain used as the reference (Itani et al., 2015). 43L1 exposure assay Gravid animals were bleached to get synchronized L1 larvae by hypochlorite treatment (0.5 M NaOH, 2% hypochlorite). After overnight starvation, about 100 L1 larvae of each mutant strain were incubated in 50 µl of M9 buffer containing OP50, carbenicillin (50 µg/ml), 10 ng/ml nystatin, 500 µM NaH2PO4 with and without 100 µM CX-5461 for ~18 hours. Following treatment, worms were allowed to recover for 0.5 hour on OP50-containing NGM plates before they were irradiated with the indicated amount of UVA exposure. Live animals were imaged after 4 days following UVA exposure to assay developmental defects using EVOS® XL Cell Imaging System. Reactive oxygen species (ROS) measurement with H2DCFDA Adult worms treated in CX-5461 for 18 hours were added with 25 µM 2’,7’-dichlorodihydrofluorescein diacetate (H2DCFDA), and incubated for another hour in the dark before initial fluorescence measurement by a microplate reader. After initial measurement, worms were irradiated with the indicated amount of UVA exposure, and then immediately measured (Yoon et al., 2017). CX-5461-DNA shifting experiment PCR products (~40 ng/µL) or sheared 6 kb genomic DNA (~5 ng/µL) were mixed with CX-5461 at room temperature. The CX-5461-DNA mixtures were incubated at 94°C for 25 minutes to denature the dsDNA, and then temperature was monitored and decreased stepwise until the target temperature was reached (if applicable). For the nuclease cleavage experiment, mung bean nuclease (MBN) (BioLabs® M0250S, 10,000 units/ml) was added to the CX-5461-DNA mixtures at room temperature (1.75 µL MBN per 100 µL reaction). Then, the mixture was incubated for one hour at either 25°C or 40°C. The final CX-5461-DNA products were loaded on 441% agarose gel containing SYBR-safe DNA stain for visualization. To note, PCR products were generated using standard PCR protocol, and worm genomic DNA was fragmented with g-tube (Covaris®), which allows genomic DNA to be sheared to a certain size (Kong et al., 2017). Generational survival assay Animals were plated individually and maintained at room temperature. Starting with 20 separate lines at P0 generation, a single L4 stage animal was transferred to a fresh plate for each generation. A line was scored as unsustainable when the parent worm was either sterile or laid only dead embryos. 3.3 Results 3.3.1 The genotypic-sensitivity profile of CX-5461 We used the quantitative acute assay and the mutant panel to generate a genotypic sensitivity profile for CX-5461, as described in previous chapter. Fourteen genotypes demonstrated sensitivity to CX-5461 (Figure 2.2, sensitivity score below 0.85). The translesion synthesis (TLS) mutants rev-3 and polh-1, the checkpoint mutant atm-1, and the structure-specific endonuclease mutant mus-81 were the most sensitive strains to CX-5461. Deficiency in homology-directed repair (HDR) and microhomology-mediated end joining (MMEJ) repair pathways, both of which repair DSBs moderately sensitized worms to CX-5461. Simultaneous loss of MMEJ and HDR repair pathways (helq-1 polq-1, polq-1 brd-1, and polq-1 rfs-1) further sensitized worms to CX-5461 (Figure 3.1B). The smrc-1 anti-helicase mutants were exquisitely sensitive to CX-5461. NER mutants (xpa-1, and ercc-1) also showed CX-5461 sensitivity. Taken together, it appears that multiple DNA repair pathways are required for tolerance of CX-5461. 45CX-5461 genotypic sensitivity profile resembled those of UVA-TMP, etoposide and camptothecin (Figure 2.2 & 3.1A). 3.3.2 CX-5461 comparison to loss of the G-quadruplex associated helicase dog-1 CX-5461 can bind and stabilize G-quadruplexes, which in turn can affect DNA replication (Xu et al., 2017). Loss of the C. elegans FANCJ homolog, dog-1, was found to result in deletions at replication blocking G-quadruplexes (Cheung et al., 2002; Zhao et al., 2008). These deletions were shown to be the by-product of MMEJ-mediated bypass of dog-1-stabilized G-quadruplexes (Koole et al., 2014). The MMEJ POLQ-1 polymerase was also needed for the bypass or repair CX-5461-mediated lesions (Figure 2.2 & 3.1B). To test whether the effect of CX-5461 treatment phenocopies the loss of dog-1 activity, we crossed the dog-1 single mutant to CX-5461-sensitive mutants and generated dog-1-containing double mutants, and compared the multigenerational fitness between single and double mutants. As expected, many CX-5461-sensitive mutants exhibited sensitivity to loss of dog-1. The polq-1 and mus-81 mutant animals were the most sensitive to loss of dog-1, as fewer than 50% of the lines survived to the fourth generation and the brood sizes of both dog-1;polq-1 and dog-1 mus-81 were significantly reduced at the F1 generation (Figure 3.2A,B, Table 3.1). HDR mutants, brd-1 and rfs-1, were both mildly sensitive to loss of dog-1. In addition, the anti-helicase mutants smrc-1, also showed moderate sensitivity to loss of dog-1, as no more than 20% of the lines survive at F8 (Figure 3.2B). Nevertheless, not all CX-5461-sensitive mutants were sensitive to loss of dog-1. For instance, the TLS mutant rev-3 was not sensitive to dog-1-induced G-quadruplex stabilization, although it was hypersensitive to CX-5461 (Figure 2.2 & 3.2A,B, Table 463.1). This suggests that CX-5461 could cause more than G-quadruplex stabilization. 3.3.3 CX-5461 can intercalate into DNA secondary structures. The chemi-genetic signature of CX-5461 suggested that CX-5461 could affect DNA replication. In silico analysis predicts that the pharmacophore of CX-5461 can intercalate into a crystal structure of DNA (PDB code 1Z3F) (Canals et al., 2005) in a manner similar to the antineoplastic agent ellipticine (Andrews et al., 2013). Therefore, we set out to test whether CX-5461 can intercalate into DNA. First, we incubated 25 µM CX-5461 with a PCR product (719 bp) and observed a mobility shift on 1% agarose gel. The band shift suggested that CX-5461 could impede double-strand DNA (dsDNA) migration (Figure 3.3A). When dsDNA was denatured and re-annealed in the presence of a high concentration of CX-5461 (>100 µM), the effect was more prominent, with DNA remaining in the well and not migrating into the gel. This suggests that CX-5461 interacts more readily with ssDNA. In addition, we also incubated CX-5461 with genomic DNA that had been sheared to 6 kbp fragments. CX-5461 impeded the migration of genomic DNA with the DNA remaining in the well at high concentrations of CX-5461 (>200 µM). When the genomic DNA was denatured, the single strands could not re-anneal with their complementary partners. We saw no dsDNA bands in the lane without CX-5461 (Figure 3.3B). However, addition of CX-5461 was able to promote the re-annealing of the denatured genomic DNA or prevented the denaturation of the DNA at the denaturation temperature. Second, we tested whether CX-5461 distorted the DNA structure. Intercalation of ellipticine can 47distort the DNA helix (Canals et al., 2005). To test whether CX-5461 intercalation could distort DNA, perhaps by stabilizing G-quadruplexes, we tested whether Mung bean nuclease (MBN), which cleaves ssDNA, could cleave CX-5461-distorted DNA. We incubated 30 µM CX-5461 with a PCR product (890 bp) predicted to form a G-quadruplex and a PCR product (719 bp) without a predicted G-quadruplex, and measured the nuclease activity on the agarose gel. Contrary to our expectation that CX-5461 would expose ssDNA for MBN cleavage, we found that CX-5461 protected DNA from MBN cleavage at both 25°C and 40°C (Figure 3.3C), indicating that CX-5461 reduced the amount of ssDNA available for cleavage, thereby enhancing the thermal stability of DNA. 3.3.4 UVA irradiation enhances CX-5461 cytotoxicity, causing both replication and transcription-associated lesions. In the course of investigating the properties of CX-5461, we noticed occasional CX-5461-sensitivity in wild-type animals. The varying WT sensitivity was found to be due to light exposure from the light source of the dissection microscope, which emitted UVA radiation. This observation was similar to reports of photosensitivity in the CX-5461 clinical trial (Hilton et al., 2018; Khot et al., 2019). We speculated that CX-5461 might exhibit photosensitivity in a manner similar to other therapeutics. To re-create the CX-5461-induced photosensitivity in C. elegans, young adults were treated with CX-5461 for ~16 hours and then irradiated with UVA. We found that wild-type N2 were not sensitive to CX-5461 or UVA alone, but were sensitive to CX-5461 followed by UVA irradiation (Figure 3.4A). The toxicity is dose-dependent with both increased UVA exposure or 48CX-5461 concentration. Importantly, the effect of UVA-induced CX-5461 toxicity is only observed in worms that have been treated with CX-5461 and then exposed to UVA irradiation, but not in worms treated with in vitro UVA-irradiated-CX-5461. This demonstrates that UVA is not changing the properties of CX-5461 to create a more toxic compound. Next, we tested whether the photosensitivity is limited to the germ line by testing the sensitivity of L1 larvae. We synchronized L1 larvae by starvation, exposed the arrested L1 larvae to CX-5461 for 16 hours followed by UVA irradiation and then released the L1 larvae from arrest by feeding. The effect of the exposure was assessed by measuring development 96 hours later. We found that L1 wild-type larvae were not sensitive to CX-5461 or UVA alone, as these animals developed normally and produced F1 progeny. In contrast, CX-5461 + UVA exposed animals failed to develop to the adult stage (Figure 3.4B). This demonstrated that the CX-5461 photosensitivity also affects somatic cells in C. elegans. Similar results were seen in both mammalian cell culture and yeast (Figure 3.4 C,D). We found that most mutants do not exhibit greater sensitivity to CX-5461 + UVA relative to wild-type animals if one takes into account the CX-5461 sensitivity associated with the mutant. The exceptions were the translesion synthesis (TLS) mutant rev-3 and the nucleotide excision repair (NER) mutant xpa-1, which were much more sensitive to CX-5461 + UVA than predicted based on their CX-5461 sensitivity (Figure 3.4E). Since NER mutants were sensitive to CX-5461 and most NER activity is transcription-coupled, we speculated that CX-5461 could cause transcription-blocking lesions. Replication-arrested NER defective L1 mutant larva fail to re-initiate development after exposure to UVC, which 49causes transcription blocking lesions (Astin et al., 2008). To test whether CX-5461 causes transcription blocking lesions, replication-arrested xpa-1 L1 mutant larvae were treated with CX-5461 followed by UVA irradiation. NER mutant xpa-1 L1 larvae failed to develop to later larval stages after CX-5461 + UVA treatment (Figure 3.5A), similar to replication-arrested xpa-1 L1 when exposed to UVC. This suggests that CX-5461 can also cause transcription-blocking lesions upon UVA irradiation. In contrast, the replication-associated CX-5461 hypersensitive mutant, mus-81, reinitiated development after L1 CX-5461 exposure and develop into sterile adults, a phenotype associated with defects in post-embryonic cell proliferation. Consistent with this, the TLS mutant rev-3 had a very similar phenotype as mus-81 upon CX-5461 treatment (Figure 3.5B). These strong effects of CX-5461 + UVA suggested that replication and transcription blocking lesions were formed. To further investigate the nature of these CX-5461 + UVA -induced lesions, we tested the CX-5461-sensitivity of the dvc-1 mutant. DVC-1 has been shown to remove DNA-protein adducts (Stingele et al., 2016). The dvc-1 mutant was completely sterile and exhibited a protruding vulva phenotype when exposed to CX-5461 + UVA treatment at L1, consistent with the possibility that CX-5461 + UVA could cause lesions that block DNA replication and transcription, like by generating DNA-protein adducts. 3.3.5 CX-5461 causes phototoxicity by generating reactive oxygen species Many phototoxicities of quinolone compounds are due to generation of reactive oxygen species (ROS) after light exposure (de Guidi et al., 2011). To investigate whether CX-5461 generates ROS after UVA exposure, we used an intracellular fluorescent probe, 2’,7’-dichlorodihydrofluorescein diacetate (H2DCFDA) to assay ROS in CX-5461 + UVA exposed C. elegans (Yoon et al., 2017). We found that increasing either UVA irradiation or CX-5461 50concentration increased the fluorescent signal, indicating the production of more ROS (Figure 3.5C,D). The increased ROS could explain the increased sensitivity to CX-5461 + UVA of xpa-1 and rev-3 mutants, because NER and TLS are needed to repair ROS-generated DNA damage induced by UVA irradiation (van Bostelen and Tijsterman, 2017). Taken together, these data suggest that CX-5461 causes photo-toxicity by generating ROS. 3.3.6 CX-5461 is mutagenic, UVA enhances its mutagenicity Since CX-5461 could stabilize G-quadruplexes (Xu et al., 2017), it could lead to replication-associated mutagenic events (Cheung et al., 2002; London et al., 2008). This fact taken together with the observation that several DNA repair pathways were required for CX-5461 tolerance, it was probable that CX-5461 was mutagenic. To assay the mutagenicity of CX-5461, we used the genetic balancer strain BC2200 dpy-18/eT1(III);unc-46/eT1(V) to measure the frequency of mutagenic events caused by CX-5461 in the absence and presence of UVA irradiation. The eT1 balancer is a reciprocal translocation of approximately half of chromosome III and half of chromosome V with dpy-18 and unc-46 balanced by eT1(III) and eT1(V), respectively. It can capture both single nucleotide variants (SNVs) and copy number variants (CNVs) in balanced regions, including terminal deletion events and translocations (Rosenbluth et al., 1985). Two hundred phenotypically wild-type (genotype: dpy-18/eT1(III);unc-46/eT1(V)) F1s from CX-5461-treated BC2200 animals were picked for each condition and scored for phenotypes. Sterility at F1 is considered to indicate the acquisition of a dominant lethal mutation, and lines 51which produce no Dpy Unc animals in F2 or later generations are considered to indicate the acquisition of a recessive lethal mutation in the balanced regions on chromosome III or V. UVA alone or buffer did not cause lethal mutations (less than 0.5% based on the screen). CX-5461 alone resulted in 4 recessive lethals out of 200 (2%). UVA exposure increased the mutagenicity of CX-5461 by more than four times, giving rise to 19 recessive lethals out of 200 (9.5%). CX-5461 and CX-5461 + UVA led to 14 and 19 dominant lethals, respectively (Figure 3.6A). 3.3.7 Mutational profile of CX-5461 (-/+ UVA) Taking advantage of the genetic balancer eT1, we were able to maintain and sequence the 23 lines with recessive lethal mutations. The CX-5461-treated lines contained a range of different mutation types, including large copy number variations (CNVs) and single nucleotide variations (SNVs) (Figure 3.6B,C). By analyzing the mutations on the balanced regions of chromosome III and V, we were able to identify the probable cause of recessive lethality in the 23 lines. Many of the lethal mutations were large CNVs on the balanced regions (2/4 and 7/19 for CX-5461 and CX-5461 + UVA, respectively), whereas others were likely due to SNVs on essential genes on the balanced regions (Table 3.2). CX-5461-induced CNVs Both MMEJ and HDR pathways are required for tolerance of CX-5461 (Figure 2.2 & 3.1B), and these two pathways respond to double-strand breaks (DSBs). Likewise, the high frequency of CNVs confirmed the necessity of pathways needed for repair of DSBs. The breakpoints of some 52of the CNVs were flanked by regions of microhomology, consistent with the MMEJ signature (Figure 3.6B). Many simple, tandem, and inverted repeats were also found surrounding the breakpoints of CNVs. However, we did not find enrichment of any motifs such as G-quadruplex forming sequences around the breakpoints. CX-5461-induced SNVs All lines exposed to CX-5461 contained a high frequency of SNVs. Specifically, we focused on homozygous mutations outside balanced regions and heterozygous mutations on balanced regions. CX-5461 + UVA induced more homozygous (mean 84.11+/-8.5 SEM (standard error of the mean)) and balanced heterozygous SNVs (mean 44.21 +/- 5.1 SEM) than CX-5461 alone (homozygous mean 27.25 +/- 6.6 SEM, balanced heterozygous mean 18.25 +/- 7.1 SEM) (Figure 3.6C). This was consistent with a higher frequency of balanced lethals seen in animals exposed to CX-5461 + UVA. There was no evident difference in the profiles of heterozygous or homozygous SNVs caused by CX-5461 with or without UVA exposure. Taking all the SNVs together, we found that SNVs were distributed throughout the genome with no trend towards coding regions or non-coding regions (Figure 3.6D). Interestingly, we noticed that around 12% SNVs (517 out of 4284) were in multi-nucleotide mutation (MNM) clusters, in which 2 to 13 mutations occur within a 1000 bp region (Figure 3.6E). Most SNVs (>80%) in MNMs were less than 15 bases from the neighboring mutations. MNMs are a common feature in the genomes of many organisms. About 2% of human genetic polymorphisms are MNM events, and clustered mutations are often present in tumors (Roberts et al., 2012). De novo MNMs have been observed in model organisms and human (Lynch et al., 2008; Ossowski et al., 2010; Schrider et al., 2013; Schrider et al., 2011). An 53analysis of MNMs in human genomes suggested that MNMs can result from error-prone translesion polymerase activity (Harris and Nielsen, 2014), which is consistent with our observation that TLS mutants are very sensitive to CX-5461. We hypothesized that the SNVs could be the product of repair or bypass of CX-5461-stabilized G-quadruplexes. To test this, we searched for G-quadruplex forming sequences around each SNV using the QuadBase2 webserver (Dhapola and Chowdhury, 2016). By assessing 100 bp 5’ and 3’ of the SNVs, we found that SNVs were not strongly correlated with G-quadruplex forming sequences. By comparison, only 0.75% of mutation-flanking regions contained predicted G-quadruplexes in CX-5461-treated genomes compared to 0.45% of the mutation-flanking regions in the ethyl methanesulfonate (EMS)-mutated genomes of the Million Mutation Project (Thompson et al., 2013). CX-5461-induced SNVs exhibited a unique mutational signature. More than 80% of SNVs were A to X mutations, with nearly half of all SNVs being A to T transversions (Figure 3.6F). Examination of the DNA context surrounding the A to X mutations revealed preferences for bases neighboring the mutated adenine. About 70% of the bases immediately 3’ (+1) of the mutated adenine were thymine, whereas cytosine and thymine were overrepresented at the 5’ (-1) position (Figure 3.6G). Despite the higher frequency in CX-5461 + UVA-treated genome, there is no difference between the types of SNVs, suggesting that the UVA increases the mutagenicity of CX-5461 but does not affect mutational mechanism. 54We also looked whether there was any motif that was prone to CX-5461 mutagenesis. Analyzing the flanking regions (-/+ 100 bp) of sites that were mutated in more than one line (47 sites in total), we observed that more than 50% of them contained inverted or tandem repeats (25/47), and that only 3 out of 47 were predicted to form G-quadruplexes. In a control set of EMS-induced mutations from the Million Mutation Project strains, only 20.2% (753/3719) contained inverted or tandem repeats flanking EMS-induced mutations. Based on these data, it appears that CX-5461 is prone to inducing lesions in regions that contain tandem or inverted repeats. 3.4 Discussion: CX-5461 is a multi-modal anticancer drug Originally identified as RNA polymerase I inhibitor and then proposed to be a more general G-quadruplex stabilizing agent, CX-5461 appears to have other characteristics associated with antineoplastic activity. Our work sheds light on the photosensitivity, chemi-genetic profile, and mutational signature of CX-5461, and suggests that CX-5461 is a complex multimodal anticancer drug that may be more broadly applicable to anti-cancer therapy than previously expected. 3.4.1 Multiple DNA repair pathways are required for CX-5461-induced DNA lesions. CX-5461 binds or interacts with DNA to generate lesions that block replication and transcription. Our observation that CX-5461 affects dsDNA migration on agarose gel and inhibits nuclease degradation of dsDNA supports the hypothesis that CX-5461 can bind DNA and alter DNA structure. The nature of CX-5461-DNA interactions can also be inferred through the comparison of chemi-genetic signatures between CX-5461 and other better-characterized agents. For instance, many genotypes are also sensitive to UVA-TMP, an inter-crosslinking (ICL) agent, but 55CX-5461 is unlikely to result in ICLs, because Fanconi Anemia (FA) genes are not required for CX-5461 tolerance. Most genes required for CPT-tolerance and ETOP-tolerance are also needed for resolution of CX-5461-induced damage, indicating that CX-5461 could result in DNA replication blocks in a manner similar to CPT and ETOP. The mechanism of CX-5461 can be differentiated from those of CPT and ETOP by the differential requirements for DSB repair pathways in C. elegans. HDR rather than MMEJ contributes to the repair of CPT under normal conditions, since HDR mutants (brd-1 and rfs-1) but not MMEJ mutants (polq-1) are sensitive to CPT; whereas MMEJ but not HDR is the primary pathway for ETOP-induced lesions (Figure 2.2). In contrast, both HDR and MMEJ contributed to the repair of CX-5461-induced lesions, as HDR single mutants (brd-1 and rfs-1) and MMEJ mutants (polq-1) are all sensitive to CX-5461. Moreover, simultaneous loss of HDR and MMEJ leads to greater CX-5461-sensitivity than expected, as seen in polq-1 brd-1 and polq-1 rfs-1 double mutants (Figure 3.1B). 3.4.2 CX-5461 can affect non-G4 DNA CX-5461 can stabilize G-quadruplexes (Xu et al., 2017). MMEJ is needed for the bypass of G-quadruplexes (Koole et al., 2014) and CX-5461 tolerance in C. elegans (Figure 2.2 & 3.1B). It is possible that CX-5461 stabilizes G-quadruplexes and other secondary structures that form barriers to replication and transcription. Aspects of the CX-5461 genotypic sensitivity profile are similar to that of the genetic interactions with dog-1, the C. elegans homolog of FANCJ that unwinds G-quadruplexes (Youds et al., 2008). However, if CX-5461 exclusively affects G-quadruplexes, we would expect that loss of dog-1 should fully phenocopy CX-5461 treatment. 56Although many CX-5461-sensitive genotypes are also sensitive to loss of dog-1, including polq-1, brd-1, rfs-1 and smrc-1, all of which have been implicated in resolution of G-tract in the absence of dog-1 (Koole et al., 2014; Ward et al., 2007; Yang et al., 2019; Youds et al., 2006), the CX-5461-hypersensitive mutant rev-3 did not exhibit a genetic interaction with the dog-1 mutant. This suggests that CX-5461 could cause lesions that are not G-quadruplex-associated. 3.4.3 The double-edged sword of therapeutic photosensitivity We also investigated the phototoxicity of CX-5461. CX-5461 caused photosensitivity in some patients (Hilton et al., 2018; Khot et al., 2019). This is similar to many other therapeutics that exhibit phototoxicity, such as vemurafenib, tamoxifen, and docetaxel (Zuba et al., 2016). We phenocopied CX-5461 photosensitivity in C. elegans, and demonstrated that an increase in either UVA irradiation or the concentration of CX-5461 enhanced the cytotoxicity. We also determined that the toxicity enhancement was due to generation of reactive oxygen species (ROS). Despite that, we did not see a ROS-induced mutational signature from genomes treated with CX-5461 + UVA. It is worth noting that either a high amount of UVA irradiation or a high concentration of CX-5461 alone is required to affect the growth and viability of wild-type animals. Worm DNA repair pathways could be sufficient to repair ROS-induced damage from CX-5461 + UVA. This is supported by the observation that the only mutants that were more sensitive to CX-5461 + UVA were TLS and NER mutants, both of which have previously been shown to repair UVA-mediated ROS damage (van Bostelen and Tijsterman, 2017) . Photosensitivity is a double-edged sword. While phototoxicity is a deleterious side-effect, the ability to generate ROS in response to UVA light could be leveraged for photo-dynamic therapy, 57in which targeted light is used to activate a photosensitizer within cancer cells leading to cell death (Banerjee et al., 2017; Yanovsky et al., 2019). Given the strong ROS-mediated phototoxicity and positive drug properties of CX-5461, CX-5461 may be an option for photo-dynamic therapy with fiber-optic light guides to deliver UVA to the tumor site (Mallidi et al., 2016). Since all genotypes of worm animals exhibit photosensitivity to CX-5461, it is likely that many different types of cancer would benefit from CX-5461. 3.4.4 CX-5461 has a complex mutational profile We showed that CX-5461 is mutagenic and that the mutagenicity increases with UVA irradiation. The mutation frequencies for CX-5461 and CX-5461 + UVA were comparable to exposure to 5 mM and 25 mM Ethyl Methane Sulfonate (EMS), a common alkylating mutagen, or 1000 and 2500 rads gamma radiation (Rosenbluth et al., 1985). Consistent with the complex genotypic sensitivity profile, CX-5461-treated genomes have a complex mutational signature that includes both CNVs and SNVs. Since mutational signatures are characteristic, and dictated by the type of DNA damage and DNA repair process, we can use these signatures to better understand the mechanism of action of CX-5461. Most CX-5461-treated genomes had CNVs, which are usually associated with formation and resolution of DSBs. This was also supported by the requirement of HDR and MMEJ for CX-5461-tolerance. In addition to CNVs, a high frequency of SNVs was also found in CX-5461-treated genomes. The CX-5461-induced A to X mutations were similar to the mutational signature observed in human urothelial cancers that have been exposed to aristocholic acids (Hoang et al., 2013; Poon et al., 2013). However, the extended sequence context differed 58between CX-5461 (CATG) and aristolochic acid (T/CAG). In addition, aristolochic acid results almost exclusively in A-T changes, whereas CX-5461 caused A-X changes. The mutational signature of aristocholic acid shows a strong transcriptional strand bias (Poon et al., 2013), and our data does not support a strand bias for CX-5461 mutagenesis. Despite this, we showed evidence that the NER pathway (which is often transcription-associated) is required for the tolerance of CX-5461, and that CX-5461 + UVA could lead to lesions that arrested L1 development, which has been used as an indirect readout of transcriptional competence (Astin et al., 2008). Given the high frequency of SNVs, the hypersensitivity of TLS mutants, and the presence of MNMs, it is likely that error-prone TLS activity is responsible for the A-X changes in CX-5461-treated genomes in a similar manner as A-T change in tumor cells exposed to aristolochic acids (Hashimoto et al., 2016). 3.4.5 CX-5461 could function in a similar manner as ellipticines The exact mechanism of how CX-5461 might interact with DNA to form blocking lesions is still unknown. Interestingly, CX-5461 shares a number of characteristics with another antineoplastic compound, ellipticine. Ellipticines have been shown to bind and stabilize G-quadruplexes, and selectively inhibit RNA polymerase I (Wei et al., 2018). A number of studies have also reported that ellipticines intercalate into DNA (Andrews et al., 2013) and lead to inhibition of topoisomerase II upon oxidation by cytochromes P450 (Stiborova et al., 2001). Ellipticines also induce reactive oxygen species (Kim et al., 2011). These characteristics lead us to speculate that CX-5461 acts in a manner similar to ellipticine in causing DNA lesions and activating cellular responses. An in silico analysis demonstrated that the pharmacophore of CX-5461 can intercalate into the DNA sequence CGATCG in a similar fashion to ellipticine, resulting in a slight 59unwinding of ApT and lengthening of DNA (Andrews et al., 2013). This could account for the propensity of adenine as a substrate for CX-5461 mutagenesis mediated by TLS. The gel shift experiments showed that 25 µM CX-5461 could result in DNA band shift on an agarose gel, suggesting that CX-5461 binds to and distorts DNA migration. This in vitro gel shift assay, while not a perfect simulation of the interactions that occur within a cell, can be used to shed light on the interaction between CX-5461 and DNA. Similarly, although the half maximal inhibitory concentration (IC50) for killing BRCA2-deficient HCT-116 cells is 4.8 nM, and the IC50 for RNA polymerase I inhibition in HCT-116 cells, is 142 nM (Drygin et al., 2011; Xu et al., 2017), CX-5461-induced G4-stabilization was confirmed by an in vitro melting assay at the micro-molar level (Xu et al., 2017). It could be that higher concentrations of CX-5461 are required to see measurable effects in vitro, and that the effect could be more localized to specific DNA structures at lower concentrations of CX-5461. Additionally, we do not yet know whether CX-5461 forms covalent bonds with DNA, as some ellipticine metabolites have been shown to, or whether metabolic processing of CX-5461 is required for formation of covalent bonds between CX-5461 and DNA. In summary, CX-5461 is a multimodal DNA damaging agent that shares properties with the antineoplastic agent ellipticine. Ellipticines have failed in stage 1 and 2 clinical trials due to strong adverse side effects (Andrews et al., 2013). It is possible that CX-5461 could provide the anti-cancer properties of ellipticine but with fewer adverse effects. 60Figure 3.1 Genotypic sensitivity to CX-5461. (A) Genotypic sensitivity profile of CX-5461. Venn diagram shows that the CX-5461-sensitive mutants also exhibited sensitivity to other DNA-damaging agents, including Etoposide (ETOP), Camptothecin (CPT), and UVA-irradiated trimethylpsoralen (UVA-TMP). (B) Loss of polq-1 sensitizes HDR-associated mutants (brd-1, rfs-1, and helq-1) to CX-5461. Embryo survival rate of indicated mutants treated with CX-5461. Error bars represent the standard error of the mean. Student t tests were used to compare the embryo survival rates between indicated mutants. *, P <0.05; **, P <0.01; ***, P<0.001. A CX-5461-sensitive strainsETOPUVA-TMPCPTatm-1mus-81him-1smrc-1polh-1brd-1rfs-1helq-1polq-1rtel-1dog-1xpa-1ercc-1rev-3B61Figure 3.2 Effect of G-quadruplex stabilization on CX-5461-sensitive mutants. (A) Brood size, embryonic lethality and male percentage in dog-1 and dog-1-containing double mutants. Error bars represent the standard error of the mean. Student t tests were used to compare brood sizes between indicated animals. * denotes p value less than 0.05, ** <0.01, ***<0.001. (B) Percentage of sustainable lines by generational fitness assay. M+Z-: maternally positive and zygotically negative. M-Z-: maternally negative and zygotically negative.00.511.52050100150200250300350N2dog-1mus-81dog-1 mus-81 (M+Z-)dog-1 mus-81 (M-Z-)polq-1dog-1;polq-1 (M+Z-)dog-1;polq-1 (M-Z-)atm-1atm-1 dog-1 (M+Z-)atm-1 dog-1 (M-Z-)rev-3dog-1;rev-3(M+Z-)dog-1;rev-3(M-Z-)Male percentage %Embryo lethality/ Brood size Embryo Lethality Brood Size Male %0204060801000 1 2 3 4 5 6 7Percentage of LinesGeneration% sustainable linesN2dog-1brd-1rfs-1smrc-1dog-1;brd-1dog-1;rfs-1dog-1;smrc-10204060801000 1 2 3 4 5 6 7Percentage of LinesGeneration% sustainable lines N2dog-1polq-1mus-81rev-3dog-1;polq-1mus-81 dog-1dog-1;rev-3atm-1 dog-1ABns ** ns nsns nsns ***62Figure 3.3 CX-5461 stabilizes DNA duplex structures. (A and B) CX-5461 binds to and shifts DNA. CX-5461 binding is enhanced by DNA denaturation and re-annealing. Non G4-containing PCR products (719 bp) were used in A, and sheared worm genomic DNA (about 6 kbp) was used in B. (C) CX-5461 stabilizes PCR products and the complex were more resistant to Mung Bean Nuclease (MBN) cleavage. Two different PCR products were used in the study, G4-containing PCR products (890 bp) and non-G4-containing PCR products (719 bp).G4-containing PCR product Non-G4-containing PCR productCX-5461 (µM) 0 4 40 2004000 4 40 200400denatured “re-annealed”BCdenatured “re-annealed”ACX-5461 (30 µM)MBN0 25 40 1252500 40 250CX-5461(µM)63ABCD64EFigure 3.4 CX-5461 is a photosensitizer in C. elegans, human cancer cell lines and yeast. (A) Viability of WT C. elegans embryos from adult animals exposed to CX-5461 and irradiated with UVA. Left-fixed [CX-5461]; Right-fixed UVA dose. Error bars represent standard error of the mean. (B) Representative images of WT C. elegans populations 96 hours after CX-5461 +UVA exposure of synchronized WT L1 larvae. The large animals are the treated P0 individuals. (C) HCT116 and HT29 colorectal cancer cell lines were treated with increasing concentrations of CX-5461 and exposed to UVA irradiation in 96-well format and cell nuclei counted after 96 hours. Student tests were used to compare the fraction of cells under different conditions. ****, P< 0.0005; ******, P<0.000005 . (D) Growth curve analysis of the relative fitness of wild-type and rad52 yeast exposed to CX-5461 + UVA radiation. Left-fixed [CX-5461]; Right-fixed UVA dose. (E) Differential sensitivity of worm mutants upon CX-5461 + UVA. No UVA65Figure 3.5 CX-5461 + UVA can result in photodamage by generating ROS. (A) UVA enhances the toxicity of CX-5461 that affect replication and transcription. The image shows the growth and development of worms four days after L1 larva-treatment. Upon CX-5461 treatment and UVA irradiation, mus-81 mutants became completely sterile, whereas xpa-1 mutants were arrested in L1. (B) CX-5461 could lead to DNA-protein adduct or bulky lesions. Upon CX-5461 treatment and UVA irradiation, dvc-1 and rev-3 mutants became mostly sterile and exhibited Pvl (protruding vulva) phenotypes (red arrows). (C) Intracellular ROS levels were measured in CX-5461 + UVA treated wild-type C. elegans with constant [CX-5461] and with (D) a constant UVA dose. Student t tests were used to compare the fluorescent intensity of different conditions. *, P < 0.05; **, P <0.01; *****, P<0.00001. The black bars represent fluorescent intensity before UVA irradiation, the grey bars represent the fluorescent intensity after UVA irradiation. NaH2PO4 100 µM CX-5461100 µM CX-5461 +100 J/m2 UVAdvc-1rev-3DBAC DFluorescence intensity Fluorescence intensity NaH2PO4 100 J/m2 UVA100 µM CX-5461 + 100 J/m2 UVA100 µM CX-5461after UVAbefore UVA66E FGFBA C D67Genotypes Brood size Embryo Survival Rate % Male % nN2 256.71±12.00 99.58±0.22 0.03±0.02 14dog-1 274.80±12.47 98.44±1.78 0.15±0.07 20mus-81 140.40±17.78 94.80±3.47 0.24±0.09 10mus-81 dog-1 (M+Z-) 187.90±13.25 90.42±6.72 0.18±0.12 10mus-81 dog-1 (M-Z-) 67.75±16.18 51.29±12.76 0.53±0.20 8polq-1 228.30±36.64 98.51±9.87 0.00±0.00 10dog-1;polq-1 (M+Z-) 233.10±19.71 93.39±5.16 0.05±0.03 20dog-1;polq-1 (M-Z-) 143.29±23.85 84.45±8.12 0.19±0.51 14atm-1 148.60±22.58 98.52±0.61 0.00±0.00 10atm-1 dog-1 (M+Z-) 159.30±10.83 97.87±0.35 0.00±0.00 10atm-1 dog-1 (M-Z-) 190.70±18.04 96.64±2.07 0.06±0.04 10rev-3 225.70±18.74 99.11±0.39 0.05±0.03 10dog-1;rev-3(M+Z-) 254.90±24.10 96.47±2.20 0.00±0.00 10dog-1;rev-3(M-Z-) 200.30±19.32 98.80±0.27 0.00±0.00 10Table 3.1 Phenotypes of indicated mutants. n denotes the number of parent worms whose progeny were scored. All values are ± standard error of the mean. M+Z-: maternally positive and zygotically negative. M-Z-: maternally negative and zygotically negative. Figure 3.6 Exposure to CX-5461 or CX-5461 + UVA results high frequencies of mutations. (A) Number of balanced recessive lethal mutations and dominant sterile mutations. n=200 for each condition. (B) Coverage plot of CX-5461 + UVA-induced genome rearrangements in sample CXU12. Whole genome (Left). Detailed coverage plot of chromosome II (Top right) and chromosome III (bottom right). Sequence at the fusion shown on right. Microhomology in bold. (C) Number of homozygous and heterozygous balanced SNVs/genome. Welch’s t-test was used to compare the number of SNV per genome under different conditions. *** p<0.0005, * p<0.05. (D) Distribution of SNVs in coding and non-coding elements. (E) Distance between SNVs in multi-nucleotide mutations. (F) SNV mutational signature of CX-5461. (G) Extended sequence context of CX-5461-induced SNVs. 68Table 3.2 Summary of mutations on CX-5461-treated and CX-5461 + UVA mutants. (SNV, single nucleotide variation. CNV, copy number variation. Del, deletion. MH, microhomology. FS, frameshift. Dp, duplication. Inv, inversion. Trans, translocation.)Line SNVsBalanced heterozygousSNVsHomozygousSNVsBalanced CNVsPutative lethal mutationCX-5461 1 68 14 52 III Del2 58 5 51 V Del3 348 47 11 chc-1 stop4 46 13 19 F54C8.4 stopCX-5461+UVA1 159 38 62 plrg-1 FS2 283 52 117 III Del MH3 190 34 107 mrpl-1 4 241 60 130 III Del strd-1/mlc-75 144 37 80 V Del MH6 258 57 143 III Dp multiple7 178 45 87 hpo-268 121 35 68 III Del/Inv 9 179 52 95 V Del III Inv multiple10 201 33 107 T05H4.1011 151 35 74 V Dp npp-1612 138 23 56 III Del13 54 11 15 V Trans14 485 89 157 V Del let-41315 243 43 84 klp-716 154 22 39 pri-117 222 39 79 ncx-218 168 29 43 III Del19 195 10 31 ?69CHAPTER 4: GENETIC INTERACTIONS BETWEEN AND WITHIN DOUBLE-STRAND BREAK REPAIR PATHWAYS IN C. ELEGANS 4.1 Introduction Cells are exposed to endogenous and exogenous damage that results in DNA lesions, including double-strand breaks (DSBs), which if not properly resolved can lead to genome instability. Cells have evolved various DNA damage repair pathways to counteract the deleterious effects of DNA damage. As mentioned in Chapter 1, cells can directly ligate DSBs using the non-homologous end joining (NHEJ) pathway, or resect the DSB ends to allow homology-based repair, including the homology-directed repair (HDR) and the microhomology-mediated end joining (MMEJ) (Ceccaldi et al., 2016a) (Figure 1.1 and 4.1). Each pathway has substrate specificity and a set of specific proteins involved. While the mechanisms of DNA repair pathways are well characterized, how these repair pathways interact to ensure error-free repair is less understood. Here, we studied the interplay within and between the different DSB repair pathways in response to endogenous meiotic DSBs and DNA damaging agent-induced DSBs. Components of the DSB repair process are well conserved across metazoans. Therefore, we used C. elegans as a model to investigate genetic interactions among the three DSB repair pathways. The key factors involved in these three DSB repair pathways are fairly well characterized. By assaying and comparing the phenotypes of single, double, triple or quadruple mutants, we are able to shed light on how different DSB repair pathways interact and are regulated in response to intrinsic meiotic DSBs and extrinsic chemical-induced DSBs. 704.1.1 HDR factors BRC-2 and RAD-51 HDR involves homology-templated repair. RAD51 filaments are essential for homology searching. The loading and stabilization of RAD51 filaments at ssDNA regions, after DSBs have been extensively resected, requires BRCA2 (Prakash et al., 2015) (Figure 4.1). BRCA2 is the tumor suppressor gene commonly mutated in breast and ovarian cancers (King et al., 2003). The C. elegans homologs of BRCA2 and RAD51, brc-2 and rad-51 are both emb (embryonic lethal), and therefore cannot be maintained as a viable strain (Alpi et al., 2003; Martin et al., 2005). For this reason, we focused on non-essential HDR factors and their genetic interactions. BRD-1 BARD1, the interacting partner of the tumor suppressor gene BRCA1, has been extensively studied. BRCA1 is one of the best characterized tumor suppressor genes and is frequently mutated in breast and ovarian cancer (Futreal et al., 1994; King et al., 2003). The BRCA1/BARD1 heterodimer is critical for homologous recombination-mediated repair of DSBs, by promoting DNA break resection and preventing non-homologous end joining (Densham et al., 2016). Loss of BRCA1/BARD1 is associated with sensitivity to a number of chemotherapeutic agents, including cisplatin and etoposide (Bunting et al., 2012; Treszezamsky et al., 2007). Both BRCA1- and BARD1-null mutants are embryonic lethal in mammals (Hakem et al., 1996; McCarthy et al., 2003). The C. elegans BRCA1 and BARD1 homologs, BRC-1 and BRD-1, are not essential for embryogenesis but are required for repairing DNA lesions caused by camptothecin or ionizing radiation (Boulton et al., 2004; Park et al., 2016). In addition, it was reported that the BRC-1/BRD-1 heterodimer is also needed for non-crossover resolution of meiotic DSBs via an inter-sister pathway (Adamo et al., 2008) (Figure 4.1). More recent studies 71showed that BRC-1/BRD-1 promotes stage-specific RAD-51 recruitment in recombination- and synapsis-impaired mutants (Janisiw et al., 2018; Li et al., 2018). RFS-1 RAD51 paralogs are mediators of HDR repair pathways that act in concert with BRCA2, and RAD51 to facilitate HDR events. RFS-1/RIP-1/SWS-1 is the heterodimeric RAD51 paralog complex in C. elegans, and has been shown to physically interact with RAD-51 and BRC-2 (Martin et al., 2005). Unlike BRC-2 and RAD-51 in C. elegans, RFS-1 is dispensable for repairing SPO-11-generated and ionizing radiation-induced DSBs. However, RFS-1 is required for RAD-51 recruitment at replication blocking lesions, including those caused by camptothecin or DNA crosslinking agents (Ward et al., 2007). RFS-1/RIP-1/SWS-1 binds and remodels RAD-51 filaments to a flexible conformation that is prone to strand exchange (Taylor et al., 2015; Taylor et al., 2016). It appears that RFS-1/RIP-1/SWS-1 plays multiple roles in regulating RAD-51 filaments, from recruiting to remodeling, from stabilizing to unloading (Figure 4.1). It will be useful to study how RFS-1 interacts with other HDR factors to better understand the biological role of RFS-1/RIP-1/SWS-1. HELQ-1 HELQ belongs to the HEL308 gene family, and can unwind DNA with a 3' to 5' polarity in an ATP-dependent manner (Han et al., 2016). HELQ mutations are observed in ovarian cancers, however, whether or how HELQ might contribute to tumorigenesis and tumor development is unclear (Pelttari et al., 2016). Mammalian HELQ is recruited to the stalled replication forks and unwind parental strands to recruit repair factors (Tafel et al., 2011). The worm homolog, helq-1 is required for tolerance of interstrand crosslinking agents (ICLs) (Muzzini et al., 2008), as is rfs-1. helq-1 and rfs-1 operate redundantly to disassemble RAD-51 filaments from double-strand 72breaks in meiosis in C. elegans germ line (Ward et al., 2010) (Figure 4.1). Consistent with this observation, rfs-1 is synthetic lethal with helq-1, most likely due to persistent meiotic RAD-51 foci (Ward et al., 2010). More recently, human HELQ has been shown to interact with RAD51 paralogs and function in the ICL repair pathway (Takata et al., 2013). It remains largely unknown how HELQ interacts with other DNA repair factors and how it is regulated. 4.1.2 NHEJ factors CKU-80 NHEJ directly ligates two broken DNA ends with no need for sequence homology, which can result in the loss of bases or translocations of DNA. KU70 and KU80 are NHEJ factors that form heterodimer that binds to the broken end of a DSB, preventing the broken end from nuclease processing. The KU70/80 heterodimer recruits ligase IV to seal the breaks (Figure 4.1). NHEJ is the predominant DSB repair pathway in non-cycling somatic cells (Ceccaldi et al., 2016a). In C. elegans, loss of NHEJ components sensitizes late stage embryos to ionizing radiation, resulting in somatic developmental defects (Clejan et al., 2006). CKU-70/80 activity is normally suppressed in wild type germ cells, but improper CKU-70/80 activity can occur in the absence of the nuclease MRE-11 or COM-1/CtIP (Lemmens et al., 2013; Yin and Smolikove, 2013), which function to resect the broken ends at DSBs (Figure 4.1). It has been shown that the resection of DSBs dictates repair choice between HDR and NHEJ (Ceccaldi et al., 2016a; Panier and Boulton, 2014). To ensure proper repair pathway choice, DSB end resection is a highly regulated process that can be affected by cell stage, the post-translational modifications of the chromatin surrounding DSBs, and a number of accessory factors including 53BP1 and shieldin (Ceccaldi et al., 2016a; Noordermeer et al., 2018). Studies in mammalian cells have shown that knock down 73of KU70/80 alleviates sensitivity of BRCA-deficient mutants to PARP inhibitors (Bunting et al., 2012). It is not known whether a similar antagonism exists between cku-70/80 and other HDR factors in C. elegans. 4.1.3 MMEJ factors POLQ-1 POLQ encodes the polymerase theta (Pol θ), which similar to HELQ, belongs to the HEL308 family. Pol θ has a family A DNA polymerase motif at the C-terminus and DNA helicase-like motif at the N-terminus. Pol θ is involved in MMEJ, an end joining DSB repair mechanism that does not involve KU proteins. MMEJ requires DNA end resection and mediates the joining of resected DSBs that harbor microhomology (Figure 4.1). The C. elegans homolog, polq-1, was first described to be required for tolerance of interstrand crosslinking (ICLs) in a pathway distinct from helq-1, another HEL308 family member (Muzzini et al., 2008). POLQ-1 plays an important role in preventing catastrophic chromosomal events at endogenous replication blocking sites (Koole et al., 2014; Roerink et al., 2014). POLQ-1 generates small indels initiated from G4 loci in the absence of dog-1, the C. elegans FANCJ homolog. POLQ-1 also generates small deletions in EMS and UVA+TMP-treated TLS mutant animals (van Schendel et al., 2016). In mammalian cell lines, loss of BRCA1/2 and POLQ is synthetic lethal, which suggests Pol θ as a potential therapeutic target for tumors carrying mutations in HDR genes (Mateos-Gomez et al., 2015). We used the C. elegans polq-1 mutant to study the interplay of MMEJ with other DSB repair pathways with respect to different types of DNA lesions. 744.1.4 The C. elegans germ line The C. elegans germ line is extremely useful for studying genes involved in DNA replication and repair, particularly DSB repair, because all stage of meiosis can be observed in a spatial-temporal gradient. The germ line starts at the distal tip, where stem germ nuclei undergo mitosis in the mitotic zone. Adjacent to the mitotic zone, nuclei progress to the transition zone, where meiosis begins and at this stage meiotic DSBs are introduced by the SPO-11 endonuclease (Dernburg et al., 1998). After the transition zone, nuclei progress into pachytene which is followed by diplotene and diakinesis, during which DSBs have to be properly resolved. In diakinesis, crossover recombination occurs and condensed chromosomes can be seen as six dumbbell-shaped bivalents. The C. elegans germ line allows direct visualization of the effect on chromosomes when DSB repair factors are disrupted. 4.1.5 Genetic interactions A genetic interaction occurs when the phenotypic effects of one gene are modified by one or several other genes. Negative genetic interactions (NGIs) occurs when mutation of two genes in a double mutant leads to a fitness defect greater than that predicted for the additive effect of the two single gene mutations alone. NGIs can manifest as synthetic lethality (SL) or synthetic sickness (SS). SL is an extreme situation where perturbations of two genes result in loss of viability, while SS is a sub-lethal fitness defect. Positive genetic interactions occur when a double mutant has a less severe fitness defect than expected based on the additive effects of the two single mutants, which could be epistasis or phenotypic suppression. 75Complete loss of the HDR pathway leads to inviability in C. elegans, because homologous paring and recombination are required for meiosis (Hillers et al., 2017). Loss of NHEJ factors can partially alleviate severe HDR phenotypes, but cannot fully rescue fertility (Lemmens et al., 2013; Macaisne et al., 2018; Yin and Smolikove, 2013). It remains largely unknown how different HDR factors genetically interact to maintain HDR functions in meiosis, and how other non-HDR repair mechanisms (including NHEJ and MMEJ) affect HDR to counteract both endogenous and exogenous DNA lesions. Here in this chapter, we focus on the genetic interactions between and within DSB repair pathways in C. elegans. 4.2 Methods Construction of mutant strains Alleles used in this chapter include: atm-1(tm5027), brd-1(dw1), rfs-1(ok1372), cku-80(ok861), polq-1(tm2026), and helq-1(tm2134). Some strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440), and some knockout alleles were provided by the Shohei Mitani laboratory. Some strains were generated by the International C. elegans Gene Knockout Consortium (Consortium, 2012) and by the National Bioresource Project of Japan. Double mutants, triple mutants and quadruple mutants were made by genetic crossing using strains containing the mutant gene(s) of interest. In all cases, mutant genotypes were confirmed by PCR. To maintain mutants with fitness defects, the hT2 balancer was used to keep mutations as heterozygotes. The hT2 balancer balances all genes aforementioned except cku-80 and helq-1. The cku-80 gene is located extremely close to the hT2 (III) translocation site but is not covered by hT2 (III). However, since cku-80 is close to the balanced region where there was very little crossing over between cku-80 76and hT2, cku-80 was maintained as a heterozygote. Most homozygous double-, triple- and quadruple mutants used in this chapter were newly isolated segregants from balanced heterozygotes. The helq-1 single mutant exhibits genomic instability, so it was frequently outcrossed to wild-type N2 and re-segregated to prevent accumulation of background mutations. Brood Scoring Single P0 L4 larvae animals of each genotype were picked to fresh NGM plates and transferred daily. F1 progeny were scored on the next day for arrested embryos and hatched larvae, and re-scored two days later for adult males and hermaphrodites. The total brood was measured as the sum of arrested embryos and hatched larvae. The embryonic lethality was defined as the percentage of arrested embryos in the total brood. The male percentage was defined as the percentage of adult males in total adults. DAPI staining diakinesis bodies Staged 24 hour-old adult worms were picked into 10µl of M9 buffer, and then transferred to 100µl of 150nM 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) in 70% ethanol. The animals were stained in the dark at 4°C overnight. Then they were destained twice by soaking in M9 buffer for one hour each time. DAPI-stained animals were mounted on slides with 3% agarose pads and visualized at 63x magnification on a Zeiss microscope. Stacked images were taken by Digital CMOS camera C11440-42U30 and processed by Zeiss Zen software. DAPI-stained diakinesis bodies were counted in the three oocytes most proximal to the spermatheca in the germ line. A DAPI-stained diakinesis body was defined as a continuous DAPI-stained chromatin structure, regardless of its size. Generational survival assay Ten P0 animals were plated individually and maintained at room temperature. Each generation, a 77single L4 stage animal was transferred to a fresh plate. A line was scored as unsustainable when the parent worm was either sterile or produced only arrested embryos. Quantitative acute assay Synchronized one-day-old adults were incubated in the DNA damaging agent (CX-5461 in 50mM NaH2PO4, etoposide (ETOP) in DMSO, or camptothecin (CPT) in DMSO) diluted in M9 buffer containing OP50, carbenicillin (50 µg/ml) and 10 ng/ml nystatin for ~18 hours. Following treatment, the animals were plated to allow recovery for 30 to 60 minutes on OP50-containing NGM plates, and then plated ten worms per plate in triplicate on NGM plates for a 4-hour interval (18 to 22 hours post-treatment), and then removed. The numbers of both arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment. All results were from at least 30 treated animals. For animals that produced few progeny, more treated adult worms were picked to lay embryos in the 4-hour interval. 4.3 Results 4.3.1 Loss of NHEJ rescues meiotic defects in helq-1 single mutants but not other HDR mutants. Previously, brd-1, helq-1 and rfs-1 were shown to be involved in HDR repair (Boulton et al., 2004; Ward et al., 2007; Ward et al., 2010), and appeared to function at different steps of HDR in response to different types of DNA lesions requiring HDR. All three HDR genes are not essential. All three deletion mutants brd-1(dw1), helq-1(tm2314) and rfs-1(ok1372) exhibit a Him (High Incidence of Males) phenotype, which is associated with mis-segregation of the X chromosome. NHEJ and MMEJ mutants, cku-80(ok681) and polq-1(tm2025), respectively, have 78no apparent Him phenotype. First, we set out to test for genetic interactions between these three distinct repair pathways, since others have observed that simultaneous loss of two DSB repair pathways result in compromised viability (Mateos-Gomez et al., 2015; Schimmel et al., 2017). Genetic crosses were carried out between single mutants each deficient in one DSB repair pathway: polq-1(tm2025) and cku-80(ok861) were representatives for MMEJ and NHEJ pathways, and brd-1(dw1), helq-1(tm2314) and rfs-1(ok1372) were representatives for different aspects of HDR. Surprisingly, no apparent negative genetic interactions were observed in M+Z- (maternally positive and zygotically negative) cku-80 brd-1 or cku-80 rfs-1 double mutants (Figure 4.2, Table 4.1). In contrast, loss of cku-80 significantly reduced the percentage of males in the helq-1 mutant animals. In order to eliminate the possibility that maternal contribution might be masking genetic interactions, M-Z- double mutants were also scored and no obvious phenotypic difference was observed. 4.3.2 Loss of MMEJ lead to transgenerational sterility in HDR mutants Although POLQ deficiency has been reported to cause extensive chromosomal aberrations and lethality in BRCA1/2 mutant cells (Mateos-Gomez et al., 2015), we did not observe a strong phenotype in the P0 generation (M+Z-) of polq-1 brd-1, and polq-1 rfs-1 animals (Figure 4.3A). We did observe a small but statistically significant reduction in brood size and male percentage in helq-1 polq-1 P0 animals. We speculate that these M+Z- double mutants had maternal POLQ-1, BRD-1 or RFS-1 proteins that contributed to their development, and that the effect of loss of polq-1 was ameliorated. Therefore, we tested the phenotypes of double mutants at F1, which 79were M-Z-. In all three cases, the brood sizes of F1 double mutants were significantly smaller than those of double mutants at P0 (Figure 4.3A,B, Table 4.1), indicating negative genetic interactions between HDR and MMEJ. To further assess the NGIs, we performed a multigenerational survival assay. The multigenerational survival assay determines whether the mutant can be maintained. The assay can infer whether mutants are accumulating germline DNA damage (Ahmed and Hodgkin, 2000). The generational survival assay found that the helq-1 polq-1 double mutants exhibited the strongest negative genetic interaction as all ten lines were inviable by the F6 generation. The polq-1 brd-1 and polq-1 rfs-1 double mutants exhibited a weaker but still striking, negative genetic interaction with > 50% of the lines becoming inviable by the F10 generation, compared to less than 20 % of the single mutants. 4.3.3 helq-1 brd-1 double mutants are synthetic lethal brd-1, helq-1 and rfs-1 are all involved in HDR. Although loss of any one of these genes results in meiotic defects and sensitivity to DNA damaging agents, these genes are not essential for HDR. This emphasizes the fact that the HDR pathway is complex with many different mechanisms to achieve DSB repair. It is possible that brd-1, helq-1, and rfs-1 act in the same processes or they could represent parallel processes within HDR. The loss of two non-essential HDR genes may be epistatic, or they may be synergistic. A previous study showed that helq-1 is SL with rfs-1 due to an overlapping function between HELQ-1 and RFS-1 in unloading of RAD-51 filaments in meiotic DSB repair (Ward et al., 2010). We asked whether simultaneous loss of helq-1 and brd-1, or brd-1 and rfs-1 would also lead to exacerbated defective meiotic phenotypes. 80Both helq-1 and brd-1 single mutants exhibited Him phenotypes (0.79% ± 0.18% and 1.14% ± 0.30%, respectively) but very little embryonic lethality. The homozygous helq-1 brd-1 double mutants displayed embryonic lethality in 98% of the progeny from P0, and the 2% that survived likely did so because of maternal contribution (Figure 4.4A), as most hatched F1 failed to reach adulthood. No chromatin bridges were observed in helq-1 brd-1 double mutants, suggesting that there were no mitotic defects. Next, we asked whether the synthetic lethality was caused by meiotic defects. We first examined whether there were abnormalities in wild type N2, helq-1, and brd-1 single mutants and helq-1 brd-1 double mutants at diakinesis. Over 75% of N2, helq-1 and brd-1 single mutants displayed normal six DAPI-stained dumbbell-shaped bivalents in diakinesis (Figure 4.4B). By contrast, helq-1 brd-1 double mutants exhibited chromosomal defects with misshapen chromosomes in diakinesis (Figure 4.4B), reminiscent of the phenotype seen in rad-51 mutants (Alpi et al., 2003; Colaiácovo et al., 2003; Macaisne et al., 2018). Although 64% of helq-1 brd-1 double mutants contained six DAPI-stained bodies in diakinesis, the DNA in diakinesis that was not organized into discrete bivalents as in rad-51 mutants. This indicated that synthetic lethality in helq-1 brd-1 double mutants is likely due to meiotic defects. Previous studies showed that loss of the NHEJ component cku-80 partially alleviates the meiotic defects in rad-51 mutants (Macaisne et al., 2018). Our data demonstrated that cku-80 loss reduced the Him phenotype in helq-1 single mutants. We speculated that loss of cku-80 could alleviate the phenotypes in helq-1 brd-1 double mutants. To test this, we generated helq-1 cku-80 brd-1 triple mutants and assayed phenotypes. cku-80 loss led to a partial suppression of fitness 81defects observed in helq-1 brd-1 double mutants (Figure 4.4A, Table 4.1), as helq-1 cku-80 brd-1 triple mutants have a survival rate of 30%, significantly higher than helq-1 brd-1 double mutants. Close examination of diakinesis found a slight improvement in the DAPI-stained body phenotype, as we observed more six-dumbbell shaped diakinesis bodies in helq-1 cku-80 brd-1 triple mutants (about 80%) (Figure 4.4B,C). Loss of MMEJ can also alleviate the meiotic defects in rad-51 single mutants (Macaisne et al., 2018). We tested whether loss of POLQ-1 could alleviate the meiotic defects in helq-1 brd-1 double mutants. In contrast to loss of cku-80, loss of polq-1 did not improve the embryonic survival. Instead, the brood size of helq-1 brd-1 reduced by almost 30% upon loss of POLQ-1 (Figure 4.4A, Table 4.1) and the DAPI-stained bodies in helq-1 polq-1 brd-1 triple mutants were indistinguishable from those of helq-1 brd-1 double mutants (Figure 4.4B,C), indicating that MMEJ has little contribution to the chromosomal abnormalities in the meiotic DSB repair in helq-1 brd-1 double mutants. Surprisingly, simultaneous loss of both MMEJ and NHEJ had little effect on helq-1 brd-1 double mutants (Figure 4.4A,B,C) suggesting that the partial rescue observed in helq-1 cku-80 brd-1 mutants is dependent on POLQ-1 activity. 4.3.4 brd-1 rfs-1 double mutants are synthetic sick brd-1 and rfs-1 single mutants share many phenotypes, including Him, camptothecin-sensitivity, and UVA-TMP-sensitivity (Figure 2.2). Based on their similar characteristics and chemical sensitivity profiles, we might expect that brd-1 rfs-1 would be epistatic and thus the brd-1 rfs-1 double mutants should exhibit similar phenotypes as the single mutants. However, brd-1 rfs-1 double mutants were synthetic sick. The homozygous double mutants exhibited severe fitness 82defects: more than 50% of progeny of P0 animals were arrested embryos, compared to less than 6% embryonic lethality in the brd-1 or rfs-1 single mutants (Figure 4.5A, Table 4.1). Moreover, the brd-1 rfs-1 double mutants had an extremely high percentage of males (about 14%), whereas the single mutants had less than 2% of males. We then set out to examine the DAPI-stained bodies in diakinesis. The brd-1 and rfs-1 single mutants all exhibited six wild-type bivalents, while the brd-1 rfs-1 double mutants often had seven or eight DAPI-stained diakinesis bodies (Figure 4.5B). We found that approximately 40% of brd-1 rfs-1 double mutant oocytes had more than six DAPI-stained diakinesis bodies, which could account for the Him phenotype (Figure 4.5C). In those brd-1 rfs-1 double mutants that had more than six DAPI-stained bodies at diakinesis, the extra DAPI-stained bodies did not resemble chromosome fragments but rather dissociated univalent(s). The high frequency of more than six DAPI-stained bodies in diakinesis likely reflects a repair defect in the brd-1 rfs-1 double mutants. To address whether this abnormality is due to defective meiotic DSB repair, we generated spo-11; brd-1 rfs-1 triple mutants. SPO-11 is an essential enzyme in the evolutionarily conserved mechanism to induce meiotic DSBs in C. elegans germ lines. The spo-11 mutant lacks meiotic DSBs, and therefore there are no meiotic crossovers or formation of chiasma. The spo-11 mutant has 12 separate univalents in diakinesis (Dernburg et al., 1998). In contrast to the brd-1 rfs-1 double mutants, the spo-11; brd-1 rfs-1 triple mutants displayed the same 12 DAPI-stained bodies as spo-11 mutants (Figure 4.5B), indicating that the NGI between BRD-1 and RFS-1 is due to defective repair of meiotic DSBs induced by SPO-11. 83Next, we asked whether loss of cku-80 would affect the viability of brd-1 rfs-1 double mutants. We found that cku-80 brd-1 rfs-1 triple mutants exhibited very similar phenotypes as brd-1 rfs-1 double mutants: ~50% arrested embryos, and a high percentage of males (Figure 4.5A, Table 4.1). Cytological analysis also found no difference in DAPI-stained diakinesis bodies between the double and triple mutants (Figure 4.5B,C). This suggested that the meiotic DSB repair defects in brd-1 rfs-1 were not the result of inappropriate utilization of the NHEJ pathway. 4.3.5 Revisiting the synthetic lethality in helq-1 rfs-1 double mutants It appeared that loss of NHEJ factors (e.g. cku-80) could partially suppress the chromosomal instability in helq-1 mutants. We decided to investigate whether NHEJ played a role in causing the meiotic defects in helq-1 rfs-1. The previous model for synthetic lethality observed in the helq-1 rfs-1 double mutants was that simultaneous loss of HELQ-1 and RFS-1 results in failure to unload RAD-51 filaments (Ward et al., 2010). This model proposed that SPO-11-induced meiotic DSBs were committed to HDR repair, and that the inability to resolve the HDR intermediates led to chromosomal aberrations in helq-1 rfs-1 double mutants. However, the chromosomal aggregates observed in diakinesis in helq-1 rfs-1 double mutants resembled the phenotype of com-1 and mre-11S mutants (Lemmens et al., 2013; Yin and Smolikove, 2013), both of which are partially rescued by loss of cku-80. It is possible that there exists an alternative mechanism for the synthetic lethality in the helq-1 rfs-1 double mutants. Therefore, we hypothesized that improper end joining events contributed to the chromosomal aggregates and lethality in the helq-1 rfs-1 double mutants. 84To test this hypothesis, we generated helq-1 cku-80 rfs-1 and helq-1 polq-1 rfs-1 triple mutants. Surprisingly, both triple mutants have much larger brood sizes (119 ± 10 and 205 ± 13 for helq-1 cku-80 rfs-1 and helq-1 polq-1 rfs-1, respectively), compared to helq-1 rfs-1 double mutants (23 ± 5) (Figure 4.6A, Table 4.1). This is strong evidence that both NHEJ and MMEJ activities negatively contribute to the brood size in the helq-1 rfs-1 double mutants. Notably, loss of cku-80 also partially rescued the embryonic survival rate to 14%, whereas loss of polq-1 did not significantly reduced embryonic lethality (Figure 4.6A, Table 4.1). We then examined the DAPI-stained bodies in diakinesis in both triple mutants. We found that loss of either NHEJ or MMEJ rescued the aggregated chromosome phenotypes, resulting in distinct bivalents; however, the chromatin morphology in both triple mutants was still different from that of wild-type animals (Figure 4.6B,C). Of note, flocculent chromosomes with long tails were often observed in the helq-1 polq-1 rfs-1 and helq-1 cku-80 rfs-1 triple mutants. Taken together, these data suggest that both end joining pathways contribute to the reduction in brood size and aggregated chromosome phenotypes. We also tested whether simultaneous loss of both NHEJ and MMEJ would further mitigate the defects in the helq-1 rfs-1 double mutants. Interestingly, elimination of both POLQ-1 and CKU-80 did not result in further increase in either brood size or embryonic viability as compared to single loss of POLQ-1 (Figure 4.6A, Table 4.1). Examination of DAPI-stained diakinesis bodies found no discernable difference between helq-1 cku-80 polq-1 rfs-1 quadruple mutants and the two triple mutants (Figure 4.6B,C). This implied that there still existed another DSB repair pathway that resolved meiotic DSBs in the quadruple mutants (Macaisne et al., 2018). 854.3.6 Loss of NHEJ rescues the sensitivity to DNA damaging agents in some but not all HDR-compromised mutants Our results have shown that cku-80 deficiency was able to partially rescue the meiotic defects in the helq-1 single mutants, but not the brd-1 or rfs-1 single mutants. We set out to determine whether loss of cku-80 could suppress sensitivity of mutants to DNA damaging agents (DDAs). We tested the Topoisomerase I inhibitor camptothecin (CPT), Topoisomerase II inhibitor etoposide (ETOP) and the G-quadruplex-stabilizing agent CX-5461. Using quantitative acute assays, we determined that NHEJ loss could partially rescue both CPT- and CX-5461-sensitivity in the helq-1 mutants (Figure 4.7A,B), as the helq-1 cku-80 double mutants had much higher embryo survival after treatment. By contrast, loss of cku-80 had no effect on the rfs-1 single mutants, as the cku-80 rfs-1 double mutants had similar embryo survival upon CPT or CX-5461 treatment to the rfs-1 mutants (Figure 4.7A,B). The cku-80 brd-1 double mutants showed a mild suppression of CPT-sensitivity compared to brd-1 single mutants (Figure 4.7C). We also observed a partial suppression of CPT-sensitivity in the atm-1 mutants by loss of cku-80 (Figure 4.7D). ATM-1 has been shown to activate and regulate HDR pathways (Balmus et al., 2019). 4.3.7 Loss of MMEJ sensitizes HDR mutants to camptothecin (CPT), etoposide (ETOP) and CX-5461. MMEJ has been considered as a back-up pathway to resolve DSBs in the absence of HDR and NHEJ. Our data showed that MMEJ was required for transgenerational viability of HDR-compromised mutant animals. Next, we determined whether MMEJ was a primary or a backup mechanism for the DSB repair of exogenous DNA damage when other DSB repair pathways were absent. 86 We first tested the sensitivity of HDR and MMEJ mutants to a Topoisomerase I inhibitor, camptothecin (CPT). HDR mutants all exhibited sensitivity to CPT, whereas MMEJ mutants were not sensitive to CPT (Figure 2.2). HDR single mutants were hypersensitive to CPT (Figure 2.2), so we used a low concentration of CPT. Surprisingly, the loss of POLQ-1 further sensitized all three HDR mutants to CPT, as the helq-1 polq-1, polq-1 rfs-1 and polq-1 brd-1 mutants all had greater embryonic lethality upon CPT treatment than their respective single mutants (Figure 4.8A,B). Because the polq-1 single mutants were not sensitive to CPT and deficiency in HDR resulted in a greater sensitivity, it suggested that MMEJ was not a primary repair mechanism for CPT-induced DNA lesions but could repair some lesions in the absence of HDR. Next, we tested mutants with a Topoisomerase II inhibitor, etoposide (ETOP). Notably, we found that the polq-1 mutants were much more sensitive to ETOP than HDR-compromised animals (Figure 2.2). At a low dose of ETOP, HDR mutant animals had little embryonic death, whereas the MMEJ mutant polq-1 was more sensitive to ETOP than helq-1, rfs-1 and brd-1 mutants (Figure 4.8C). Combined loss of polq-1 and any one of the three HDR genes resulted in significant reduction in embryo viability upon ETOP treatment (Figure 4.8C). Therefore, HDR deficiency further sensitized MMEJ mutants to ETOP. Taken together, the data suggest there exists a negative genetic interaction between HDR and MMEJ in response to ETOP-induced lesions, and that HDR could repair some ETOP damage in the absence of MMEJ. Lastly, we also tested the sensitivity of HDR and MMEJ mutants to CX-5461. In contrast to CPT or ETOP, both HDR single mutants and MMEJ single mutants were sensitive to CX-5461, 87indicating that both HDR and MMEJ were required for CX-5461 tolerance. Again, loss of POLQ-1 led to greater CX-5461-sensitivity in helq-1, brd-1, and rfs-1 mutants (Figure 4.8D,E), suggesting that MMEJ and HDR respond independently to CX-5461-induced DNA damage. 4.4 Discussion 4.4.1 Redundancy between HDR factors We found NGIs between three factors involved in the HDR pathway. First, HELQ-1 is required for survival in the absence of BRD-1/BRC-1. Second, BRD-1 and RFS-1 work synergistically to maintain genome stability. Third, we analyzed the synthetic lethality observed in the helq-1 rfs-1 double mutants and suggest a mechanism that may be contributing to SL. All three factors (BRD-1/BRC-1, HELQ-1, and RFS-1) have been shown to play key roles in homologous recombination, including recruiting, influencing or interacting with RAD-51 filaments at the sites of DNA damage (Boulton et al., 2004; Janisiw et al., 2018; Li et al., 2018; Ward et al., 2010; Yousefzadeh et al., 2014). Although they are all involved in HDR of DNA lesions, none is essential for viability in C. elegans. Therefore, the NGI (SL and SS) between any two genes uncover the compensating functions of each gene pair. A. HELQ-1 & BRD-1 Since helq-1 brd-1 double mutants were synthetic lethal, it is possible that BARD-1 and HELQ-1 share a partially redundant role in meiotic DSB repair. Our data suggest that HELQ-1 is involved in an early step in RAD-51 filament dynamics or resection, since the loss of HELQ-1 resulted in improper NHEJ activity, similar to loss of DNA repair factors that drive DNA resection. This is evidenced by the observation that the helq-1 cku-80 double mutants had mildly yet significantly 88reduced embryonic lethality and male percentage (Figure 4.2), and that loss of CKU-80 suppressed some sensitivity to CPT and CX-5461 in the helq-1 single mutants (Figure 4.7A,B). In mammalian cells, BRCA1/BARD1 promotes HDR and counteracts NHEJ activity by presenting end-resected DSBs through its interaction with CtIP and MRN (Chen et al., 2008), and loss of CKU-80 partially alleviated phenotypes in C. elegans mutants deficient in CtIP and MRN complex (Girard et al., 2018; Lemmens et al., 2013; Yin and Smolikove, 2013). Taken altogether, one possibility is that there exists a collaborative synergy between BRD-1 and HELQ-1 in promoting HDR in early stage of meiotic DSB repair. Another possible explanation for SL in helq-1 brd-1 is co-involvement of HELQ-1 and BRD-1 in late stage of HDR. HELQ-1 affects meiotic recombination by evicting the RAD-51 filaments (Ward et al., 2010). BRC-1/BRD-1, on the other hand, promotes inter-sister repair of SPO-11-induced DSBs (Adamo et al., 2008). In the brc-1 and brd-1 single mutants, there is an increase in RAD-51 foci in late pachytene regions compared to wild-type animals (Adamo et al., 2008), whereas no increase of RAD-51 filament is observed in late pachytene in the helq-1 single mutants despite a proposed role in the disassembly of RAD-51 filaments (Ward et al., 2010). Loss of HELQ-1 further accumulates the RAD-51 filaments in rtel-1 mutants (Ward et al., 2010), and here it is likely that simultaneous loss of HELQ-1 and BRD-1 also results in excess unloaded RAD-51 filaments in late pachytene, indicative of elevated unrepaired DNA damage that potentially gives rise to lethality. 89Close examination of the diakinesis in the helq-1 brd-1 double mutants revealed a near wild-type number of DAPI-stained diakinesis bodies. However, all DAPI-stained diakinesis bodies in the helq-1 brd-1 double mutants were misshapen and reminiscent of those observed in jmjd-5 and nmad-1 single mutants (Amendola et al., 2017; Wang et al., 2019a). JMJD-5 is a histone demethylase that has been shown to act in the late stage of HDR to promote RAD-51 filament removal in a similar manner as HELQ-1 (Amendola et al., 2017), whereas NMAD-1 is a DNA demethylase that removes methyl groups from DNA; loss of NMAD-1 also resulted in increased RAD-51 foci in the germ line (Wang et al., 2019a). Notably, lack of either JMJD-5 or NMAD-1 could alter the global chromatin methylation level, which could affect DSB repair choice at an early stage. It would be interesting to test whether methylation might be affecting the recruitment of HELQ-1 and BRD-1. B. BRD-1 & RFS-1 The synthetic sickness in brd-1 rfs-1 double mutants illustrates the synergy between BRD-1 and RFS-1. The brd-1 rfs-1 double mutants exhibit greater genome instability phenotypes (Him, and embryonic lethality) than either single mutant, suggesting that co-loss of BRD-1 and RFS-1 compromises meiotic DSB repair likely through affecting HDR. The brd-1 rfs-1 double mutants displayed a high frequency of more than six DAPI-stained diakinesis bodies, whereas the spo-11; brd-1 rfs-1 triple mutants displayed a phenotype similar to spo-11 mutants that have a twelve-univalent phenotype (Figure 4.5B), indicating that the aberrant diakinesis in brd-1 rfs-1 is dependent on the repair of SPO-11-induced DSBs. From this data, we conclude that that the NGI between BRD-1 and RFS-1 is a consequence of defective meiotic DSB repair. 90The brd-1 rfs-1 mutants frequently contained seven DAPI-stained diakinesis bodies, which is consistent with the extremely high male percentage observed in brd-1 rfs-1 double mutants. In contrast to the helq-1 brd-1 double mutants, the brd-1 rfs-1 double mutants did not have misshapen DAPI-stained bodies, suggesting a different mechanism leading to meiotic DSB repair defects. Both BRD-1 and RFS-1 promote HDR at different types of lesions (Boulton et al., 2004; Ward et al., 2007). It is possible that loss of both gives rise to insufficient RAD-51 loading and thus compromised HDR activity. However, this is unlikely the case, because CKU-80 loss did not rescue the fitness defects in the brd-1 rfs-1 double mutants (Figure 4.5B,C), and failure to initiate HDR could result in utilization of NHEJ. BRC-1/BRD-1 stabilizes RAD-51 filaments on new DSBs in mid- to late pachytene (Li et al., 2018). RFS-1/RIP-1/SWS-1 have been shown to remodel pre-synaptic RAD-51 filaments to a stabilized and flexible confirmation that promote homologous recombination (Taylor et al., 2015). Therefore, this raised another hypothesis that the combined loss of BRD-1 and RFS-1 results in destabilized RAD-51 filaments that weaken HDR competence. This could explain why deficiency in NHEJ did not improve the phenotype of the brd-1 rfs-1 double mutants, because RAD-51 filaments have already committed repair to HDR, and therefore the DSBs are likely to be resected and not substrates for CKU-80 binding. Altogether, simultaneous loss of BRD-1 and RFS-1 greatly compromised meiotic DSB repair, and hence caused genome instability phenotypes, such as Him and high embryonic lethality. Future work could assay RAD-51 foci and the state of the synaptonemal complex across 91different stages of meiosis in brd-1 rfs-1 double mutants to further elucidate the basis of NGI between BRD-1 and RFS-1. C. RFS-1 & HELQ-1 A previous study has characterized the SL in helq-1 rfs-1 double mutants as excessive unloading of RAD-51 from RAD-51 filaments in late meiotic DSB repair (Ward et al., 2010). Here, we present mechanistic insight into the repair pathways that are used in the absence of both HELQ-1 and RFS-1. Our data suggest that in the helq-1 rfs-1 double mutants, DSB repair mechanisms other than HDR could take place, such as NHEJ and MMEJ, resulting in chromosomal fusions. Loss of either CKU-80 or POLQ-1 or both, remarkably increased the brood size of helq-1 rfs-1 double mutants. Moreover, there are no chromosomal fusion phenotypes observed any of the triple mutants helq-1 cku-80 rfs-1 and helq-1 polq-1 rfs-1, or the quadruple mutants helq-1 cku-80 polq-1 rfs-1. This suggest that illegitimate end joining may be ligating broken meiotic DSB ends when HDR is defective. Interestingly, CKU-80 loss slightly but significantly increased the embryo viability of the helq-1 rfs-1 double mutants, whereas loss of POLQ-1 or both POLQ-1 and CKU-80, while rescuing brood size, led to complete embryo lethality, suggesting that MMEJ was needed for viability (discussed more in 4.4.3). These results are reminiscent of the phenotypes observed in polq-1;rad-51, cku-70;rad-51, and polq-1 cku70; rad-51 (Yang et al., 2019). It is still unknown how homologous recombination intermediates in the helq-1 rfs-1 double mutants would activate or trigger end joining repair. One possibility is that RFS-1 deficiency results in destabilized RAD-51 filaments that are susceptible to nuclease degradation (Taylor et al., 2015). Despite presence of a large quantity of RAD-51 filaments, homologous recombination 92is not stimulated. It is also observed in the brc-1 single mutants that there is high retention of RAD-51 in late pachytene, although the meiotic DSB repair is largely HDR-dependent (Adamo et al., 2008). 4.4.2 MMEJ is both a primary and back-up repair pathway dependent on the nature of the DNA lesion It has long been thought that MMEJ is a back-up pathway to repair DSB lesions, but recent work has suggested that MMEJ can independently resolve DNA lesions in the presence of both HDR and NHEJ (Koole et al., 2014; Roerink et al., 2014). The activity of MMEJ has been frequently found in the human genome, as many repetitive elements that constitute a large proportion in the genome, have signatures consistent with MMEJ activity (Sfeir and Symington, 2015). Another study demonstrated that POLQ-1 is involved in generating the small indels that are abundantly present in wild isolates of C. elegans (Koole et al., 2014). Here, our data also elucidated the versatile role that MMEJ plays in response to different types of DNA lesions in C. elegans. Under normal conditions, when there is no exogenous DNA damage, most DSBs are either SPO-11-induced meiotic DSBs or DSBs formed at hard to replicate regions such as G-quadruplexes or tandem repeats. MMEJ has been shown to initiate the formation of small deletions at endogenous G4 sites when DOG-1 is absent (Koole et al., 2014). It is likely that MMEJ is the primary repair mechanism to resolve DNA lesions formed at G4, compared to other repair pathways such as HDR and TLS, since the dog-1; polq-1 mutants exhibited a much more severe fitness defect than the dog-1;brd-1 and dog-1;rev-3 mutants (Figure 3.2B). It has also been previously reported that POLQ-1 is involved in the repair of the transposon-induced breaks and CRISPR/Cas9-induced 93breaks in meiosis (van Schendel et al., 2015). However, POLQ-1 is not required for meiosis under normal conditions, as polq-1 mutants have a near wild-type phenotype by cytological analysis of the germ line (Figure A4.1). Our data suggest that POLQ-1 is required for genome integrity when HDR is compromised. Co-depletion of POLQ-1 and HDR factors result in a progressive decline in viability. This is corroborated by the evidence that the brood sizes of the helq-1 polq-1, polq-1 rfs-1 and polq-1 brd-1 mutants at F1 (M-Z-) are all significantly smaller than those of their M+Z- counterparts. The reduced brood size could be due to under-proliferated germ lines, or defects in sperm meiosis. Besides, generational survival assay suggests that the double mutants exhibit greater genome instability by accumulating mutations that result in sterility. Additionally, POLQ-1 played a crucial role in chromosomal abnormalities when both HELQ-1 and RFS-1 were absent, because loss of POLQ-1 rescued the chromosomal aggregates and increased the brood size in the helq-1 rfs-1 double mutants (Figure 4.6A, Table 4.1). This suggested that POLQ-1 could ligate broken DSB ends. To counteract exogenous DNA damage, we found that MMEJ could be both a back-up repair pathway and a dedicated repair mechanism depending on the type of DNA lesions cells encounter (Figure 4.9). The topoisomerase I inhibitor CPT has been shown to cause replication blocks, which if not repaired could lead to fork collapse and DSB. Our data showed that the polq-1 mutants were not sensitive to CPT, although some POLQ-/- human cell lines exhibited mild sensitivity upon CPT treatment (Wang et al., 2019b; Yousefzadeh et al., 2014). It is likely that MMEJ is not the primary repair mechanism for CPT-induced lesions in C. elegans. We 94found that HDR mutants exhibited strong CPT-sensitivity, indicating HDR as the primary repair mechanism for resolving CPT-induced lesions. We also observed increased sensitivity to CPT when both HDR and MMEJ pathways were defective (Figure 4.8A,B), suggesting that POLQ-1 could be used as a back-up pathway for CPT-lesions when HDR pathway is compromised (Figure 4.9). This is possible because both HDR and MMEJ requires end resection, and MMEJ could use resected ends to resolve CPT lesions at a cost of causing small deletions. The topoisomerase II inhibitor ETOP, however, induces a different type of lesion than CPT. Consistent with that, ETOP utilizes a different repair mechanism than CPT. Our data showed that the polq-1 mutants exhibited markedly stronger sensitivity even at a low dose of ETOP, compared to HDR mutants (rfs-1, helq-1 and brd-1) (Figure 4.8C). This suggests that MMEJ rather than HDR is the primary repair pathway in response to ETOP-induced lesions (as discussed in Chapter 2). Loss of HDR factors further sensitized the polq-1 mutants to ETOP (Figure 4.8C), demonstrating that HDR could be a back-up repair mechanism to resolve ETOP-induced lesions when the primary MMEJ pathway is not available. This is the converse of CPT, in which HDR is the primary pathway and MMEJ is the backup pathway (Figure 4.9). CX-5461 induces lesions distinct from CPT or ETOP. It has been shown that CX-5461 stabilizes G-quadruplex and blocks DNA replication (Xu et al., 2017). The G-quadruplex-induced lesions have been shown to be repaired by polymerase theta-mediated MMEJ (Koole et al., 2014), and MMEJ also repaired CX-5461-induced lesions. Based on our data, it appears that HDR is also required for some lesions induced by CX-5461. If HDR and MMEJ are truly redundant pathways for responding to CX-5461-induced lesions, mutants deficient in only one single pathway should 95not be sensitive. Simultaneous loss of MMEJ and HDR significantly reduced the embryonic survival in worms exposed to CX-5461 (Figure 4.8D,E), indicating that two pathways work independently to repair CX-5461-induced DNA damage (Figure 4.9). Similarly, a study in A549/DR cancer cells has shown that HDR and MMEJ synergize for cell survival in response to cisplatin (Dai et al., 2016). 4.4.3 Illegitimate NHEJ activity can be toxic in certain HDR-compromised background Many studies have shown that improper NHEJ activity could lead to deleterious cellular consequences. This is often because NHEJ is an error-prone DSB repair pathway that directly ligates the broken DNA ends, and this can result in deletions or translocations. During meiosis in C. elegans, SPO-11-induced meiotic DSBs are processed by the MRN complex and COM-1, which antagonizes CKU-70/80. The meiotic defects of mre-11S, com-1, and nbs-1 mutant animals are partially restored by loss of cku-80 (Girard et al., 2018; Lemmens et al., 2013; Yin and Smolikove, 2013). We showed that HELQ-1 might also be involved in the early processing of DSB ends to prohibit NHEJ activity. Loss of cku-80 suppressed the HIM phenotype in helq-1 single mutants (Figure 4.2). Additionally, the role of HELQ-1 counteracting NHEJ activity appeared not only limited to meiotic DSBs, but also DNA lesions induced by DNA damaging agents, such as CPT and CX-5461 (Figure 4.7A,B). We also observed NHEJ deficiency mildly suppressed the CPT-sensitivity in brd-1 mutants, similar to human counterparts (Figure 4.7C) (Bunting et al., 2012). The effect of the loss of NHEJ on the helq-1 mutants was also visible in genetic interactions. Loss of cku-80 partially suppressed the lethality in both helq-1-containing double mutants, helq-961 brd-1 and helq-1 rfs-1 (Figure 4.4A, 4.6A). But the cku-80 suppression of the double mutants was completely ameliorated by additional loss of MMEJ, as both (M+Z-) quadruple mutants helq-1 cku-80 polq-1 brd-1 and helq-1 cku-80 polq-1 rfs-1 produced no viable progeny (Figure 4.4A, 4.6A). Taken together, these data indicate a hierarchy for repair mechanisms of meiotic DSBs. Under normal conditions, DSBs are resolved by HDR mechanisms, and end processing factors like MRN and CtIP and other helicases such as HELQ-1 prevent inappropriate NHEJ mechanism. When HDR is defective, NHEJ can act, leading to deleterious effects. When both HDR and NHEJ are defective, MMEJ can join DSBs in a less deleterious manner than NHEJ. Animals lacking all three major DSB repair pathways were unable to resolve meiotic DSBs properly. 97Figure 4.1 A schematic of DSB repair pathways in C. elegans. See main texts for details.CKU-70/80POLQ-1End-resectionNHEJHDRMMEJDSBBRC-1/BRD-1COM-1, MRE-11LIG-4 BRC-2, RAD-51Homology search & strand invasionSPO-11-induced DSBs or DNA damaging agentsFigure 4.2 cku-80 genetically interacts helq-1 single mutants but not other HDR mutants or MMEJ mutants.Brood size, embryonic lethality, and % males in single mutants and cku-80-containing double mutants. At leasteight hermaphrodites were scored for their progeny. Error bars represent the standard error of the mean.Student t tests were used to compare brood sizes between indicated animals (upper) and male percentagebetween indicated animals (bottom). Male percentage of cku-80 was no different from that of M+Z- and M-Z-helq-1 cku-80, but significantly different from M-Z- cku-80 brd-1(***) and M-Z- cku-80 rfs-1(***). * denotes pvalue less than 0.05, ** <0.01, ***<0.001.012345050100150200250300350N2cku-80helq-1helq-1 cku-80 (M+Z-)helq-1 cku-80 (M-Z-)brd-1cku-80 brd-1(M+Z-)cku-80 brd-1(M-Z-)rfs-1cku-80 rfs-1 (M+Z-)cku-80 rfs-1 (M-Z-)Male percentage %Embryo lethality/ Brood sizeEmbryo Lethality Brood Size Male %nsnsns nsnsns*** ns*nsnsns980204060801000 1 2 3 4 5 6 7 8 9 10 11 12Percentage of LinesGeneration% of sustainable linesN2helq-1rfs-1brd-1polq-1helq-1 polq-1polq-1 rfs-1polq-1 brd-1012345050100150200250300350N2 polq-1 helq-1 helq-1polq-1(M+Z-)helq-1polq-1(M-Z-)brd-1 polq-1brd-1(M+Z-)polq-1brd-1(M-Z-)rfs-1 polq-1rfs-1(M+Z-)polq-1rfs-1(M-Z-)Male percentage %Embryo lethality/ Brood sizeEmbryo Lethality Brood Size Male %Figure 4.3 polq-1 genetically interacts with HDR gene/mutants (helq-1, brd-1, and rfs-1). (A) Brood size, embryonic lethality, and % males in HDR single mutants and polq-1-containing double mutants of maternally positive and zygotically negative (M+Z-) P0 and maternally positive and zygotically negative (M-Z-) F1. At least eight hermaphrodites were scored for their progeny. Student t tests were used to compare brood sizes between indicated animals (upper) and male percentage between indicated animals (bottom). To note, the brood size of polq-1 mutant was significantly different from that of M-Z- polq-1 brd-1(*), and M-Z- polq-1 rfs-1(*). * denotes p value less than 0.05, ** <0.01, ***<0.001. Error bars represent the standard error of the mean. (B) Viability of independent lines of N2, HDR single mutants, and polq-1-containing double mutants at 23℃. All starting P0 were M+Z-, and n = 10. The helq-1, rfs-1, brd-1, and polq-1 lines were largely overlapped between F7 and F11. **** ns*ns*nsns**nsnsnsAB99050100150200250300350N2brd-1helq-1helq-1 brd-1helq-1 cku-80 brd-1helq-1 polq-1 brd-1helq-1 cku-80 polq-1 brd-1Embryo lethality/ Brood size Embryo Lethality Brood SizeN2 helq-1 brd-1helq-1 brd-1 67 6 7 6helq-1 cku-80 brd-1 6 6 7 666B6 666A*ns ns*100Figure 4.4 helq-1 brd-1 double mutants are synthetic lethal. (A) Brood size and embryonic lethality in helq-1 and brd-1 single mutants, helq-1 brd-1 double mutants and other associated mutants. The double, triple and quadruple mutants were scored as M+Z- worms. Error bars represent the standard error of the mean. Student t tests were used to compare embryo survival rate between indicated animals. * denotes p value less than 0.05, ** <0.01, ***<0.001. (B) Representative images of DAPI-stained bodies in diakinesis of respective mutants. Numbers below in the brackets indicate the number of DAPI-stained bodies per nuclei. Scale bar denotes 5 μm. (C) Quantification of DAPI-stained bodies at diakinesis from animals of the indicated mutants. ws denotes weird shaped. helq-1 polq-1 brd-1 6 6 6 66 6 7 6 helq-1 cku-80 polq-1 brd-1 020406080100<6 ws 6 7 8 9 10 11 12Number of DAPI-stained bodies% of DAPI-stained diakinesis bodies N2brd-1 (n=28)helq-1 (n=40)helq-1 brd-1 (n=25)helq-1 cku-80 brd-1 (n=31)helq-1 polq-1 brd-1 (n=37)helq-1 cku-80 polq-1 brd-1 (n=22)C101048121620050100150200250300350N2 brd-1 rfs-1 brd-1rfs-1cku-80brd-1rfs-1Male percentage %Embryo lethality/ Brood sizeEmbryo Lethality Brood SizeMale %brd-1 rfs-110 67 8rfs-16spo-11 12 12 126BAns102Figure 4.5 brd-1 rfs-1 double mutants are synthetic sick. (A) Brood size and embryonic lethality in brd-1 and rfs-1 single mutants, brd-1 rfs-1 double mutants and cku-80 brd-1 rfs-1 triple mutants. The double and triple mutants were scored as M+Z- worms. Student t tests were used to compare the brood size, embryo survival rate and HIM between indicated animals. No statistical significance was found. (B) Representative images of DAPI-stained bodies at diakinesis of respective mutants. Numbers below indicate the number of DAPI-stained bodies per nuclei. Scale bar denotes 5 μm. (C) Quantification of DAPI-stained bodies at diakinesis from animals of the indicated mutants. 020406080100<6 ws 6 7 8 9 10 11 12Number of DAPI-stained bodies% of DAPI-stained diakinesis bodies brd-1 rfs-1 (n=31)cku-80 brd-1 rfs-1 (n=20)N2 (n=18)brd-1 (n=28)rfs-1 (n-=26)cku-80 brd-1 rfs-112 12 12spo-11; brd-1 rfs-18 7 67C103050100150200250300350N2 helq-1 rfs-1 helq-1rfs-1helq-1cku-80rfs-1helq-1polq-1rfs-1helq-1cku-80polq-1rfs-1Embryo lethality/ Brood size Embryo Lethality Brood Sizehelq-1 rfs-1 helq-1 cku-80 rfs-1 helq-1 polq-1 rfs-1 6 6 6 68 7 6 62 4 5BA*ns***ns104Figure 4.6 helq-1 rfs-1 double mutants are synthetic lethal. (A) Brood size and embryonic lethality in helq-1 and rfs-1 single mutants, helq-1 rfs-1 double mutants and associated triple and quadruple mutants. The double, triple and quadruple mutants were scored as M+Z- worms. Error bars represent the standard error of the mean. Student t tests were used to compare embryo survival rate between indicated animals. * denotes p value less than 0.05, ** <0.01, ***<0.001. In addition, Student t tests were used to compare brood size between helq-1 rfs-1 and helq-1 cku-80 rfs-1, and helq-1 rfs-1 and helq-1 polq-1 rfs-1. p-value for both was smaller than 0.001. (B) Representative images of DAPI-stained bodies in diakinesis of respective mutants. Numbers below indicate the number of DAPI-stained bodies per nuclei. Scale bar denotes 5 μm. (C) Quantification of DAPI-stained bodies at diakinesis from animals of the indicated mutants. helq-1 cku-80 polq-1 rfs-1 6 6 6 5020406080100<6 ws 6 7 8 9 10 11 12Number of DAPI-stained bodies% of DAPI-stained diakinesis bodies N2helq-1rfs-1helq-1 rfs-1 (n=13)helq-1 cku-80 rfs-1 (n=25)helq-1 polq-1 rfs-1 (n=25)helq-1 cku-80 polq-1 rfs-1 (n=14)C105Figure 4.7 Loss of NHEJ rescues the DNA damage agent sensitivity in some but not all HDR-compromised mutants. (A - D) Embryo survival rate of indicated mutants treated with camptothecin (CPT) or CX-5461. CPT was dissolved in DMSO, CX-5461 was dissolved in 50mM Na2HPO4. mus-81 is used as positive control in above experiments. All experiments were conducted in three replicates. Error bars represent the standard error of the mean. P values were calculated using two-tailed student t test. * denotes p value less than 0.05, ** <0.01, ***<0.001. **020406080100N2 mus-81 cku-80 brd-1 cku-80brd-1Embryo survival rate %DMSO 100 nM CPT*020406080100N2 cku-80 helq-1 helq-1cku-80rfs-1 cku-80rfs-1Embryo Survival Rate %DMSO 100nM CPT*** ns ** ns020406080100N2 cku-80 helq-1 helq-1cku-80rfs-1 cku-80rfs-1Embryo survival rate %Buffer 100 µM CX-5461DCA BD020406080100N2mus-81cku-80atm-1atm-1;cku-80Embryo survival rate %DMSO 100 nM CPT106020406080100N2 mus-81 polq-1 brd-1 polq-1brd-1Embryo survival rate % Buffer 100 µM CX-5461020406080100N2 polq-1 helq-1 helq-1polq-1rfs-1 rfs-1polq-1Embryo Survival Rate %DMSO 50nM CPT* **020406080100N2 mus-81 polq-1 brd-1 polq-1brd-1Embryo survival rate %DMSO 50 nM CPT**020406080100N2 mus-81 polq-1 helq-1 helq-1polq-1brd-1 polq-1brd-1rfs-1 polq-1rfs-1Embryo survival rate %DMSO 50 µM ETOP******** ***polq-1 rfs-1 polq-1rfs-1Buffer 100 µM CX-5461*020406080100helq-1 polq-1 helq-1polq-1Embryo Survival Rate %Buffer 20 µM CX-5461** *******ECA BDFigure 4.8 Loss of MMEJ sensitizes HDR mutants to DNA damage agents. (A - E) Embryo survival rate of indicated mutants treated with camptothecin (CPT), etoposide (ETOP) or CX-5461. CPT and ETOP were dissolved in DMSO, CX-5461 was dissolved in 50mM Na2HPO4. mus-81 is used as positive control in above experiments. All experiments were conducted in three replicates. Error bars represent the standard error of the mean. P values were calculated using two-tailed student t test. * denotes p value less than 0.05, ** <0.01, ***<0.001. 107Figure4.9ModelofdifferentDSBrepairpathwaysrequiredforCPT(A),ETOP(B),andCX-5461(C).(A)CPT-inducedlesionsisrepairedprimarilyviaHDRpathway(left).IntheabsenceofHDR,MMEJcanbeabackuppathway.(B)ETOP-inducedlesionsisrepairedprimarilyviaMMEJpathway(left).WhenMMEJiscompromised,HDRcanbeabackuppathway.(C)CX-5461-inducedlesionsrequirebothHDRandMMEJpathwaystorepair.Thickarrowsdenotetheprimarypathway,thinarrowsdenotethebackuppathwaywhentheprimaryoneisoverwhelmedorabsent,andintermediatearrowsin(C)indicatethatbothpathwaysarerequired.CPT-induced DNA lesionsHDRMMEJpolq-1helq-1, brd-1, rfs-1 AETOP-induced DNA lesionsBCX-5461-induced DNA lesionsCHDRMMEJpolq-1helq-1, brd-1, rfs-1 HDRMMEJpolq-1helq-1, brd-1, rfs-1 108Genotype Brood Size Embryo Survival Rate % Male % nN2 256.71±12.00 99.58±0.22 0.03±0.02 14helq-1 258.40±20.16 98.30±0.37 0.79±0.18 10brd-1 261.20±24.31 97.55±0.46 1.14±0.30 10rfs-1 223.50±23.85 95.53±2.48 1.85±0.49 10cku-80 260.30±11.35 99.77±0.10 0.08±0.07 10polq-1 228.30±36.64 98.51±9.87 0.00±0.00 10helq-1 cku-80 (M+Z-) 213.56±7.65 99.27±0.19 0.10±0.07 9helq-1 cku-80 (M-Z-) 244.80±17.29 99.14±0.31 0.04±0.04 10cku-80 brd-1 (M+Z-) 232.80±23.26 97.59±0.24 0.66±0.23 10cku-80 brd-1 (M-Z-) 268.40±14.47 98.92±0.55 1.37±0.15 10cku-80 rfs-1 (M+Z-) 267.60±15.38 91.52±1.09 2.35±0.32 10cku-80 rfs-1 (M-Z-) 220.80±17.19 95.10±0.44 2.59±0.28 10cku-80 polq-1 266.70±11.42 98.65±0.36 0.11±0.06 10helq-1 polq-1 (M+Z-) 189.20±16.69 87.21±6.81 0.00±0.00 10helq-1 polq-1 (M-Z-) 129.30±19.09 91.11±4.05 0.00±0.00 10polq-1 brd-1 (M+Z-) 217.70±11.63 97.29±0.47 1.21±0.23 10polq-1 brd-1 (M-Z-) 98.30±20.06 88.71±10.62 2.08±0.98 10polq-1 rfs-1 (M+Z-) 233.30±32.02 86.71±3.84 1.54±0.37 10polq-1 rfs-1 (M-Z-) 131.30±23.83 96.73±0.83 0.88±0.21 10helq-1 brd-1 (M+Z-) 158.80±10.36 1.89±0.46 ND 10helq-1 rfs-1 (M+Z-) 22.56±5.07 7.20±1.28 ND 16brd-1 rfs-1 (M+Z-) 169.73±19.62 49.96±2.89 13.66±1.27 15helq-1 brd-1 rfs-1 (M+Z-) 90.80±6.50 0.11±0.20 ND 10helq-1 cku-80 polq-1 (M+Z-) 228.30±19.51 91.90±5.11 0.00±0.00 10cku-80 polq-1 rfs-1 (M+Z-) 247.90±29.22 95.93±0.60 0.76±0.15 10cku-80 polq-1 brd-1 (M+Z-) 193.40±27.80 50.83±10.65 1.36±1.01 10helq-1 cku-80 brd-1 (M+Z-) 171.40±18.20 30.46±7.87 3.48±5.45 10helq-1 polq-1 brd-1 (M+Z-) 97.00±10.41 10.62±3.51 0.00±0.00 10helq-1 cku-80 rfs-1 (M+Z-) 118.93±10.25 10.37±1.17 8.70±2.72 15helq-1 polq-1 rfs-1 (M+Z-) 204.60±13.09 0.10±0.06 ND 10cku-80 brd-1 rfs-1 (M+Z-) 167.60±10.42 45.76±2.11 13.70±1.28 10helq-1 cku-80 polq-1 rfs-1 (M+Z-) 210.25±25.03 0.00±0.00 ND 16helq-1 cku-80 polq-1 brd-1 (M+Z-) 136.90±8.99 3.80±0.73 ND 10Table 4.1 Phenotypes of indicated mutants. n denotes the number of parent worms whose progeny were scored. All values are ± standard error of the mean. M+Z-: maternally positive and zygotically negative. M-Z-: maternally negative and zygotically negative. ND: not determined. 109CHAPTER 5: CHARACTERIZATION OF THE ANNEALING HELICASE SMRC-1 IN C. ELEGANS 5.1 Introduction SMARCAL1 belongs to the SNF2 family of ATP-dependent chromatin remodeling enzymes that are conserved across species, including worms and humans. SMARCAL1 has been shown to play important roles in maintaining genome integrity. It has a RPA-interacting motif at its N-teminus, and two tandem HARP domains at the protein centre. At its C-terminus, SMARCAL1 has a helicase domain that contains one DEXDc and one HELICc motif. Bi-allelic loss-of-function mutations in SMARCAL1 cause an autosomal recessive disease called Schimke immune-osseous dysplasia (SIOD) (Boerkoel et al., 2002). The disease is characterized by smaller brain sizes, compromised neuronal development, bone abnormalities, and T-cell immunodeficiency (Deguchi et al., 2008). It has been shown that SMARCAL1 is required for cell viability under treatment with agents that cause replication stress, including UV, mitomycin, and camptothecin (Bansbach et al., 2009; Ciccia et al., 2009). SMARCAL1 is regulated following hydroxyurea treatment or UV radiation (Bansbach et al., 2009). Biochemical experiments demonstrated that SMARCAL1 can act as an anti-helicase by annealing the nascent ssDNA strands to reverse the fork, forming a four-way DNA structure resembling a chicken foot (Betous et al., 2012). The current model for SMARCAL1 is that it prevents genome instability through the bypass of DNA lesions by catalyzing fork reversal and restoration. It appears that SMARCAL1 maintains genome integrity by preventing ssDNA accumulation at stalled replication forks. Loss of SMARCAL1 leads to 110ssDNA that can be a substrate for other repair or resolution mechanisms, which can often be deleterious. There are many factors that play an important role in guarding the genome stability, including the resolution of substrates arising at stalled replication forks. There are very few confirmed anti-helicases. Other anti-helicases have been identified that are able to reverse replication forks, including HLTF and ZRANB3 (Poole and Cortez, 2017). SMRC-1, the C. elegans SMARCAL1 homolog, appears to be the only anti-helicase in C. elegans, and therefore, the worm may provide a simpler model to characterize the role of anti-helicases in genome stability. Recently, smrc-1 mutants have been found to genetically interact with met-2, a histone methyltransferase (Yang et al., 2019). Little is known about how SMARCAL1 interacts with other factors at the replication fork. Loss of SMARCAL1 could have an effect on MUS81, a structure-specific endonuclease, which cleaves the stalled forks to create DSBs for HDR-mediated fork restart (Hanada et al., 2007; Regairaz et al., 2011). SMARCAL1 could also play a role in ssDNA maintenance during HDR. The anti-recombinase RTEL1, prevents recombination by disrupting D-loop formations and template switching (Barber et al., 2008; Leon-Ortiz et al., 2018), this process would generate ssDNA that could be a substrate for SMARCAL1. Other helicases play important roles in genome maintenance, such as FANCJ and HELQ. FANCJ can unwind G-quadruplexes, and loss of FANCJ function induces small deletions in C. elegans (Cheung et al., 2002; London et al., 2008). HELQ is less understood but has been shown to play a role in regulating D-loops (Ward et al., 2010). In addition, the protease SPRTN/DVC1 safeguards the genome by degrading proteins trapped on DNA (Stingele et al., 2016; Vaz et al., 1112016). If DNA-protein adducts are not resolved, they can act as replication blocks, and SMARCAL1 may be required to mediate the replication fork reversal. To address these hypotheses, we identified and characterized the genetic interactions between SMARCAL1 and genes involved in DNA metabolism, with an emphasis on those that resolve stalled forks. 5.2 Methods Construction of mutant strains Alleles used in this chapter include: smrc-1 (gk176502), smrc-1 (gk784642), dog-1 (gk10), rtel-1(tm1866), mus-81(tm1937), helq-1 (tm2134) and dvc-1 (ok260). Some strains were provided by the Caenorhabditis Genetics Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440), and some knockout alleles were provided by the Shohei Mitani laboratory. Some strains were generated by the International C. elegans Gene Knockout Consortium (Consortium, 2012) and by the National Bioresource Project of Japan. Two smrc-1 alleles, gk176502 and gk784642 were obtained by outcrossing the strains VC20168 and VC40734 from the Million Mutation Project (MMP) collection (Thompson et al., 2013) to N2 six times, and then balancing the smrc-1 mutations with the hT2 balancer to maintain smrc-1 as a heterozygote to prevent accumulation of mutations. Double mutants were generated by genetic crossing using strains containing the mutant gene(s) of interest. In all cases, mutants were genotyped by PCR. To maintain mutants with strong fitness defects, the hT2 balancer was used to keep mutations heterozygous. The hT2 balancer is able to balance most genes studied in this 112chapter with the exception of dog-1 and helq-1. Most homozygous double mutants were characterized as fresh segregants from balanced heterozygotes. Brood Scoring L4 larvae of each genotype were picked to fresh NGM plates and transferred each day. Progeny were first scored 24 hours later for dead embryos and hatched larvae, and scored two days later for adult males and hermaphrodites. The total brood is measured as the sum of dead embryos and hatched larvae. The embryonic lethality is defined as the percentage of dead embryos in the total brood. The male percentage is defined as the percentage of males in the total adult population. Generational survival assay Animals were plated individually and maintained at room temperature. Starting with 10 or 20 separate individuals at the P0 generation, a single L4 larva was transferred to a fresh plate at each generation. A line was scored as unsustainable when the parent worm was either sterile or produced only dead embryos. Genome Sequencing One starting P0 and all viable F10 lines of both smrc-1 mutants (gk176502 and gk784642) were prepared for sequencing. Worms were rinsed off with deionized water and pelleted by centrifugation. Genomic DNA was purified using Puregene® Core Kit A (Qiagen). Illumina Hi-Seq DNA sequencing was performed at the Novogene Bioinformatics Institute (Beijing, China). Sequence reads were mapped to the C. elegans reference genome version WS230 (http://www.wormbase.org) using the short-read aligner BWA, which gave each sample a sequencing depth of a least 20x. Copy numbers were estimated from the alignments with a procedure analogous to that of Itani et al. using 5 kb wide overlapping sliding windows with the 113alignments from the parental strain used as the reference (Itani et al., 2015). Copy number alterations in F10 lines were normalized by subtracting the respective P0. DAPI staining Synchronized animals were picked into 10@l of M9 buffer, and then transferred to 100@l of 150nM 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) in 70% ethanol. The animals were stained in the dark at 4 overnight. Then they were destained twice by soaking in M9 buffer for one hour each. DAPI-stained animals were mounted on slides with 3% agarose pads and visualized at 63x magnification on a Zeiss Axioplan microscope using Z-stacking. Stacked images were taken by Hamamatsu ORCA-Flash 4.0 camera and processed by Zeiss Zen software. DAPI-stained diakinesis bodies were counted in the three oocytes most proximal to the spermatheca in the germ line. A DAPI-stained diakinesis body was defined as a continuous DAPI-stained chromatin structure, regardless of its size. Mitotic bridges were defined as unresolved chromatin structures between two distinct nuclei. Quantitative acute assay Synchronized one-day-old adults were incubated in the DNA damaging agent (CX-5461 in 50mM NaH2PO4, etoposide (ETOP) in DMSO, or camptothecin (CPT) in DMSO) diluted in M9 buffer containing OP50, carbenicillin (50 µg/ml) and 10ng/ml nystatin for ~18 hours. The exception was that adults were incubated in Ethyl methanesulfonate (EMS) for only 4 hours. Following treatment, the animals were plated to allow recovery for 30 to 60 minutes on OP50-containing NGM plates, and then plated ten worms per plate in triplicate on NGM plates for a 4-hour interval (18 to 22 hours post-treatment), and then removed. The numbers of both arrested embryos and hatched larvae were counted one day later in order to calculate the percentage of embryo survival after treatment. All results were from at least 30 treated animals. For animals 114that produced few progeny, more treated adult worms were picked to lay embryos in the 4-hour interval. The calculation of the sensitivity score was detailed in Chapter 2 Method. 5.3 Results 5.3.1 SMRC-1 is required for maintaining genome stability In the course of assembling a panel of C. elegans DNA repair mutants, we noticed that two Million Mutation Project (MMP) strains (VC20168 and VC40734) that had a distinct mutational profile. VC20168 and VC40734 both contained many small deletions and duplications (Figure A5.1). Furthermore, both VC20168 and VC40734 exhibited a Him phenotype, which can be indicative of chromosome instability. The MMP strains were generated by EMS (Ethyl methanesulfonate), ENU (N-ethyl-N-nitrosourea), and EMS+ENU mutagenesis followed by 10 generations of self-crossing to homozygose mutations before whole genome sequencing (Thompson et al., 2013). If a mutator homozygosed early in the self-crossing process, this strain would accumulate mutations, which could be distinct from the mutagenesis-induced mutations. The MMP strains contain many homozygous mutations that could be responsible for these phenotypes. Specifically, VC20168 contained 253 homozygous mutations, of which 93 are predicted to result in non-synonymous protein changes, whereas VC40734 contained 373 homozygous mutations, of which 143 are predicted to result in non-synonymous protein changes. Both strains contained potentially deleterious mutations in the smrc-1 gene. VC20168 contains smrc-1 (gk176502), a point mutation in a conserved residue of the DEXDc helicase domain, and VC40734 contains smrc-1 (gk784642), a point mutation that results in a premature stop codon. To test whether the smrc-1 mutations were the cause of genomic instability, smrc-1 single 115mutants were first outcrossed to N2 six times to remove background mutations. After six times of outcrossing, each of the smrc-1 alleles (gk176502 and gk784642) was maintained in separate lines using the hT2 balancer. Then, 10 maternally positive zygotically negative (M+Z-) individual lines (termed as P0) for both smrc-1(gk176502) and smrc-1(gk784642) alleles were bottle-necked by picking one L4 in each generation for 10 generations (F10) to re-create the process used to generate the MMP strains. Notably, not all ten lines were viable at F10. Five out of ten lines containing smrc-1(gk176502) remained fertile at F10, and seven out of ten of lines containing smrc-1(gk784642) had become inviable by F10 (Figure 5.1A) The remaining F10 (M-Z-) smrc-1 and starting P0 (M+Z-) of both alleles were prepared for whole-genome sequencing. In the strains carrying either smrc-1 allele, comparing genomes at F10 to P0 revealed dramatically more small copy number variations in F10 compared to the P0, supporting the hypothesis that the smrc-1 mutations were the cause of the copy number variations (Figure 5.1B, Figure A5.2). Next, we set out to characterize the phenotypes of both M+Z- and M-Z- smrc-1 mutants. Both M+Z- smrc-1 (gk176502) and smrc-1 (gk784642) alleles caused a Him (high incidence of males) phenotype and reduced embryonic viability (Figure 5.1C). M-Z- smrc-1 mutants showed more severe fitness defects than their M+Z- counterparts. In particular, brood size and embryo viability of smrc-1 (gk784642) were significantly reduced in M-Z- animals compared to those in M+Z- animals. This is consistent with a recent study that showed SMRC-1 promotes fertility and embryonic viability in C. elegans (Yang et al., 2019). 1165.3.2 SMRC-1 is required for survival after DDA treatment Next, we asked whether SMRC-1 was required for tolerance of exogenous DNA damage. To address this, we exposed both smrc-1 mutants to DNA damaging agents, including ethyl methanesulfonate (EMS, an alkylating agent), camptothecin (CPT, a Topoisomerase I inhibitor), etoposide (ETOP, a Topoisomerase II inhibitor), and CX-5461 (a G4-stabilizing agent), and then scored the viability of the progeny of treated animals. smrc-1 mutant animals showed reduced viability upon treatment of all these agents (Figure 5.2), suggesting SMRC-1 was required for tolerance to all these DNA damaging agents tested. 5.3.3 Genetic interactions between loss of SMRC-1 and other helicases Due to the role of SMRC-1 in maintaining genome integrity, we expected that smrc-1 would have genetic interactions with other genes that maintain genome stability, such as DNA replication and repair genes. A previous report showed that SMRC-1 was required when the chromatin modifier, MET-2 was lost (Yang et al., 2019). First, we crossed smrc-1(gk176502) males to rtel-1(tm1866) hermaphrodites. RTEL-1 is a helicase that functions as an anti-recombinase. RTEL-1 has been shown to regulate the homeostasis of RAD-51 filaments, and its loss results in increased heterologous and homologous recombination (Barber et al., 2008; Leon-Ortiz et al., 2018). The rtel-1; smrc-1 double mutants were synthetic lethal. The rtel-1; smrc-1 double mutants produced 100% arrested embryos, whereas both the rtel-1 and smrc-1 (gk176502) single mutants produced over 90% viable progeny (Figure 5.3A, Table 5.1). In addition, the brood size was severely reduced in the double mutants compared to the smrc-1 and rtel-1 single mutants. Cytological analysis of rtel-1; smrc-1 117(M+Z-) double mutants revealed that the phenotypes of DAPI-stained diakinesis bodies were near wild-type (Figure 5.3C), suggesting that meiosis was grossly normal in rtel-1; smrc-1 animals. By contrast, many somatic cells in rtel-1; smrc-1 double mutants showed chromatin bridges (Figure 5.3C), which can be indicative of replication stress and incomplete replication. We also observed chromatin bridges in early embryos produced by double mutants. Taken altogether, simultaneous loss of the anti-helicase smrc-1 and the anti-recombinase rtel-1 resulted in profound replication defects in the soma and early embryos. We also generated dog-1; smrc-1 and helq-1 smrc-1 double mutants. The dog-1;smrc-1 (M+Z-) and helq-1 smrc-1 (M+Z-) double mutants did not exhibit significantly more genome instability-associated phenotypes than the single mutants (Figure 5.3A, Table 5.1). We speculated that the double mutants may accumulate DNA lesions that lead to fitness defects over successive generations, since loss of SMRC-1 has been shown to cause more G/C-associated deletions (Yang et al., 2019). To test this, we bottle-necked the double mutants and the single mutants by generational survival assay. Both the helq-1 smrc-1 and dog-1;smrc-1 double mutants became completely sterile by F10 (Figure 5.3B) (Yang et al., 2019), whereas over 40 percent of the helq-1, dog-1, and smrc-1 single mutants remain fertile. This indicated that the double mutants exacerbated the accumulation of DNA lesions that were passed down to later generations and resulted in sterility in a much faster manner than the smrc-1 (gk176502) single mutants. Overall, smrc-1 exhibited synthetic lethality or sickness in combination with rtel-1, helq-1, and dog-1. 1185.3.4 Genetic interactions between loss of SMRC-1 and other DDR genes A previous study proposed a model that cells depend on MUS81-mediated repair when SMARCAL1 is absent (Betous et al., 2012). We hypothesized that loss of smrc-1 would affect the viability of mus-81 mutants in C. elegans. We crossed smrc-1 (gk176502) males to mus-81 hermaphrodites, and mus-81;smrc-1 double mutants were maintained as heterozygotes using the hT2 balancer. Five out of ten M+Z- double mutants scored produced less than 20 progeny, including two that were sterile. On average, the mus-81;smrc-1 (M+Z-) double mutants had a small brood size, 50% less than the brood size of the mus-81 (M+Z-) or smrc-1 (gk176502) single mutants. The double mutant had a decreased embryo survival rate (50.51% ± 7.78%), compared to more than 90% in the mus-81 or smrc-1 (gk176502) single mutants (Figure 5.3A, Table 5.1). The F1 mus-81;smrc-1 double mutants that survived likely did so by maternal rescue, because all F1 were either sterile or produced no viable progeny (Figure 5.3B). Cytological analysis found similar somatic replication defects in mus-81; smrc-1 as observed in rtel-1;smrc-1 double mutants (Figure 5.3C). The DAPI-stained diakinesis bodies in mus-81; smrc-1 double mutants showed little difference to those in mus-81 single mutants, both of which had mild meiotic defects (O'Neil et al., 2013). It has been shown that metalloprotease SPRTN/DVC1 could regulate translesion synthesis as well as degrade DNA-protein crosslink lesions (Ghosal et al., 2012; Mosbech et al., 2012; Stingele et al., 2016; Vaz et al., 2016), and SMARCAL1 mediates fork reversal when the fork encounters DNA lesions. We hypothesized that these two genes could work in concert to counteract DNA lesions. We asked whether simultaneous loss of DVC-1 and SMRC-1 in C. elegans would affect animal survival by generating dvc-1;smrc-1 double mutants. The dvc-1191;smrc-1 double mutants exhibited more severe fitness defects than either the dvc-1 or smrc-1 single mutants (Figure 5.3A, Table 5.1). The double mutants had approximately one third the brood size of either single mutants, and two out of 10 lines of dvc-1;smrc-1 (M+Z-) double mutants were sterile. In addition, the double mutants showed reduced embryo viability (76.61% ± 11.49%) than that of the dvc-1 mutants (98.41 ± 0.68%). 5.4 Discussion SMRC-1 is required for genome integrity. Biallelic mutation of SMARCAL1 in humans is linked to the etiology of Schimke immuno-osseous dysplasia (SIOD). Little is known about how SMARCAL1 interacts with other replication fork stabilizing proteins. Using C. elegans, we investigated the in vivo role of SMRC-1 and studied the genetic interactions between smrc-1 and other genes involved in genome maintenance. 5.4.1 SMRC-1 is required for genome integrity. Close examination of mutational signatures of the two original MMP strains that contain SMRC-1 mutations raised a hypothesis that SMRC-1 loss induced chromosomal abnormalities. Accumulation of these chromosomal alterations over successive generations could eventually lead to sterility or lethality. Recently, Yang and colleagues have showed that CRISPR-edited smrc-1 nonsense and frame-shift alleles could cause reduced fertility and increased embryo lethality (Yang et al., 2019). We found similar genome instability-associated phenotypes using independent nonsense and missense smrc-1 alleles. To further support that SMRC-1 plays a key role in maintaining genome 120integrity, we bottle-necked smrc-1 mutants for 10 generations and sequenced the genome of F10 (M-Z-) and P0 (M+Z-) smrc-1 mutants. Our data indicated that SMRC-1 deficiency resulted in a mutator phenotype characterized by the accumulation of small copy number variations. In addition, generational sterility occurred in later generations confirmed that accumulation of chromosomal alteration events had deleterious effects on viability. It has been previously shown that SMARCAL1 was part of the DNA damage response by its recruitment to mediate the regression of stalled replication forks via interaction with RPA (Bansbach et al., 2009; Ciccia et al., 2009), a protein complex that coats and protects ssDNA. DNA damage is often associated with the presence of ssDNA-RPA complex, which triggers the DNA damage response (Zou et al., 2006). Lack of SMRC-1 could lead to inability to repair certain type of DNA lesions. Therefore, when SMRC-1 is defective, animals could accumulate unresolved endogenous DNA lesions and develop dysfunctional germ lines that produce fewer progeny; or alternatively, if the lesions were tolerable and passed to the next generation, the next generation animals would continue to accumulate unresolved lesions, which could eventually lead to embryonic lethality or sterility. The sensitivity of smrc-1 mutants to various types of exogenous DNA damage demonstrates that SMRC-1 plays a vital role in preserving genome stability. For example, EMS induces methylated guanines that are usually resolved by O6-methylguanine-DNA methyltransferase (MGMT) or the mismatch repair pathway. In the course of repair, single stranded DNA is generated, which is susceptible to further damage in the form of DNA breaks. Our data showed that smrc-1 mutants were extremely sensitive to EMS treatment, indicating that annealing ability of SMRC-1 could 121be critical for resolution of EMS-induced DNA lesions in C. elegans. Likewise, treatment with the topoisomerase poisons camptothecin and etoposide can generate ssDNA intermediates, which might require SMRC-1 to prevent the spreading of ssDNA and DNA breakage. Taken together, SMRC-1 is required for protecting genome integrity from both endogenous and exogenous DNA damage in vivo. 5.4.2 SMRC-1 is required for genome integrity in the absence of RTEL-1 The synthetic lethality between smrc-1 and rtel-1 emphasizes the role of SMRC-1 in maintaining genome integrity. Previous studies showed that loss of SMRC-1 increased meiotic recombination frequency by five-fold (Yang et al., 2019), and that loss of RTEL-1 could elevate meiotic recombination frequency by 2.5-fold (Barber et al., 2008; Yang et al., 2019). It is unknown whether a similar mechanism underlies these increases in meiotic crossover rates. In addition, loss of RTEL1 has been shown to cause increase in heterologous recombination and lead to duplication of genomic regions inserted at some distance from its original copies in mammalian cells (Leon-Ortiz et al., 2018). We have also identified a similar pattern of duplications in smrc-1-deficient mutants (Figure 5.1B, Figure A5.2). It appears that loss of SMRC-1 or RTEL-1 increases the frequency of CNVs. In light of the annealing activity of SMRC-1 and anti-recombination activity of RTEL-1, we propose a speculative model in which the loss of SMRC-1 causes ssDNA accumulation upon replication fork stalling, which becomes a substrate for homology-directed recombination, eventually giving rise to templated-duplications. These duplications are the by-product of an error-prone tolerance mechanism needed for smrc-1 mutant survival. However, additional loss of RTEL-1 further promotes inappropriate recombination and 122template switching, potentially resulting in massive amount of recombination intermediates that could not be resolved. 5.4.3 SMRC-1 is required for genome integrity in the absence of MUS-81 A previous study showed that SMARCAL1-deficient cells rely on MUS81-catalyzed cleavage of damaged forks to prevent DNA lesions (Betous et al., 2012). In agreement with this, we showed that loss of SMRC-1 function is synthetic lethal with the absence of MUS-81. In human cells, MUS81, together with its interacting partners (EME1 or EME2), is able to cleave a variety of DNA structure substrates, and its activity is tightly regulated by WEE1/CDK and other kinases (Duda et al., 2016; Wyatt et al., 2013). Since MUS81 is required for cutting persistent replication intermediates at common fragile sites in mitosis, it is often the backup mechanism to safeguard chromosome segregation (Minocherhomji et al., 2015; Naim et al., 2013; Ying et al., 2013). Similar to mus-81 mutants, smrc-1 mutants exhibited broad sensitivity to many DNA damaging agents tested (Figure 5.2). This suggests that SMRC-1 could be involved in different repair mechanisms to counteract DNA damage, or that SMRC-1 is essential to regulate ssDNA which is a common intermediate for many repair pathways (Zellweger et al., 2015). Mechanistically, Mus81 has been shown to cleave the stalled forks to form DSBs to restart replication (Hanada et al., 2007) as well as to limit Pol32-dependent error prone synthesis in break-induced repair (Mayle et al., 2015). This raises the possibility that MUS-81 and SMRC-1 represent two alternate repair/bypass mechanisms at stalled replication forks. 123It is worth noting that both mus-81 and rtel-1 exhibit synthetic lethality with smrc-1, and that mus-81 rtel-1 double mutants are also synthetic lethal due to accumulated replication intermediates that fail to be resolved in meiosis (Barber et al., 2008). It has also been shown in yeast that Mus81 removes toxic Holliday junctions in the absence of Srs2, the yeast helicase analogous to RTEL1 (Elango et al., 2017). Therefore, it appears that there are distinct mechanisms behind the SL observed mus-81;smrc-1, rtel-1;smrc-1 and mus-81 rtel-1. The next step will be to look for phenotypes that could differentiate the three pairs of synthetic lethal double mutants to elucidate distinctive roles of each gene. 5.4.4 smrc-1 genetically interacts with factors required for DNA metabolism The helq-1 smrc-1, dog-1;smrc-1, and dvc-1;smrc-1 double mutants were all synthetic sick, indicating that smrc-1 negatively interacts with helq-1, dog-1, and dvc-1, all of which are factors involved in very different aspects of DNA metabolism. DOG-1 is the C. elegans FANCJ homolog that has been shown to unwind G-quadruplexes (Youds et al., 2008). Loss of DOG-1 induces a mutator phenotype that primarily causes G-tract deletions that are mediated by MMEJ repair (Cheung et al., 2002; Koole et al., 2014). It has been recently reported that additional loss of SMRC-1 increases the frequency of G-tract deletions in the dog-1 mutant background (Yang et al., 2019). We found that, similar to dog-1 mutants, the loss of SMRC-1 results in a distinctive mutator phenotype characterized by small copy number variations (Figure 5.1B, Figure A5.2). It remains to be determined if the smrc-1 mutator phenotype is catalysed by MMEJ as is the case with dog-1. However, smrc-1-induced rearrangements are frequently small local duplications which are distinct from the small 124deletions flanked by microhomology. It remains to be determined if these small duplications contain microhomology at the breakpoints. One model for dog-1-induced small deletions involves persistence of ssDNA through mitosis (Koole et al., 2014), which could also be a substrate for smrc-1. It is likely that loss of SMRC-1 changes the mutational nature of dog-1 or increases the mutations to an intolerable level. This is consistent with the observation that dog-1;smrc-1 double mutant lines became inviable much faster than dog-1 and smrc-1 single mutant lines (Figure 5.3B). In human cells, a recent study demonstrated that FANCJ and helicase-like transcription factor (HLTF, another fork reversal remodeler) work in opposing roles to keep fork remodeling and replication progression in check (Peng et al., 2018). Combined loss of FANCJ and HLTF results in severe replication stress in cells. Our data suggest there is a synergy between DOG-1 and SMRC-1 in protecting genome integrity. helq-1 is the C. elegans homolog of human HELQ, which is involved in repairing interstrand crosslink (ICL) damage. It has been shown that HELQ interacts with RAD51C to promote efficient homologous recombination (Adelman et al., 2013; Ward et al., 2010). We found that helq-1 mutant animals exhibited sensitivity to a number of DNA damaging agents, including topoisomerase poisons and CX-5461 (See Chapter 2 and 3), and that helq-1 is synthetic lethal with brd-1 (Chapter 4). Little is known about how helq-1 genetically interacts with other helicases. M+Z- helq-1 smrc-1 double mutants did not show strong fitness defects when compared to smrc-1 single mutants. It appears that HELQ-1 deficiency exacerbated the negative effect of the mutator phenotype due to loss of SMRC-1, since helq-1 smrc-1 lost fertility much faster than respective single mutants. The next step would be to look into how HELQ-1 contributes to DNA repair in the absence of SMRC-1. 125 dvc-1 is the C. elegans homolog of human SPRTN metalloprotease. SPRTN regulates the switch between replicative polymerases and translesion synthesis (TLS) polymerases (Ghosal et al., 2012; Mosbech et al., 2012), and removes protein-DNA adducts (Stingele et al., 2016; Vaz et al., 2016). The dvc-1;smrc-1 double mutants were synthetic sick, with a reduced brood size and reduced embryo viability compared to respective single mutants. The negative genetic interaction between dvc-1 and smrc-1 raised two possibilities due to two distinctive roles of DVC-1. One possibility is that TLS and fork remodeling represent two separate repair mechanisms to bypass endogenous DNA lesions without actually resolving the lesions, and that simultaneous loss of TLS regulators and SMRC-1 lead to cell death. The other possibility would involve a role for SMRC-1 in bypassing DNA-protein adducts when they are not degraded by DVC-1. It is noted that both TLS regulation and protease activity can play a role in managing protein adducts trapped on DNA, thus making SMRC-1 a potential target in some genetic backgrounds for anticancer therapeutics that result in protein trapping on DNA. We have identified genetic interactions between SMRC-1 and multiple genes involved in DNA metabolism and confirmed the distinct mutator phenotype that occurs when SMRC-1 is lost. More detailed analysis of the smrc-1-induced rearrangements could shed light on the mechanisms that cause the rearrangements in the absence of SMRC-1. Unfortunately, the complex nature of the rearrangements, which contain multiple small duplications and breakpoints, confound analysis using the standard short reads of Illumina sequencing. To further characterize how SMRC-1 contributes to maintaining genome stability, we could use long-read Nanopore sequencing to identify breakpoints of CNVs in smrc-1 mutant animals to elucidate 126whether there are sequence motifs or structural preferences in the genome where chromosomal abnormality events are likely to occur in the absence of SMRC-1. 1270123050100150200250300350400N2smrc-1 (gk176502) M+Z-smrc-1 (gk784642) M+Z-smrc-1 (gk176502) M-Z-smrc-1 (gk784642) M-Z-Male percentage %Embryo lethality/Brood sizeEmbryo Lethality Brood Size Male %02468100 2 4 6 8 10Number of lines GenerationsNumber of survival lines smrc-1(gk176502)smrc-1(gk784642)N2Figure 5.1 SMRC-1 is required for maintaining genome stability. (A) Viability of independent lines of N2, smrc-1 mutants at 23℃. Both smrc-1 mutants were M+Z-. (B) Coverage plot of genome rearrangements of F10 gk176502 (Left). Coverage plot of genome rearrangements of F10 gk784642 (Right). (C) Brood size and embryonic lethality in smrc-1(gk176502) and smrc-1(gk784642) single mutants, both smrc-1 mutants have been outcrossed to N2 six times, and maintained heterozygous by hT2 balancer before scoring. Error bars represent the standard error of the mean. Student t tests were used to compare brood sizes between indicated animals (upper) and male percentage between indicated animals (bottom). * denotes p value less than 0.05, ** <0.01, ***<0.001.Copy numberCopy numberChromosome I II III IV V XChromosome I II III IV V XA CB********128Figure 5.2 SMRC-1 is required for tolerance of genotoxins. Embryo survival rate of smrc-1 mutants treated with ethyl methanesulfonate (EMS), CX-5461, camptothecin (CPT), and etoposide (ETOP). CPT and ETOP were dissolved in DMSO solution. mus-81 mutant was used as a positive control in all experiments above. All experiments were conducted in at least three replicates. Error bars represent the standard error of the mean. P values were calculated using two-tailed student t test. The embryo survival rates of mutants treated with DDAs were all significantly from that of N2 treated with DDAs. * denotes p value less than 0.05, ** <0.01, ***<0.001. 020406080100N2mus-81smrc-1 (gk176502)smrc-1 (gk784642)Embryo Survival Rate %0 EMS25 mM EMS020406080100N2mus-81smrc-1 (gk176502)smrc-1 (gk784642)Embryo survival rate %Buffer100 µM CX-5461020406080100N2mus-81smrc-1 (gk176502)smrc-1 (gk784642)Embryo survival rate %DMSO100 nM CPT020406080100N2mus-81smrc-1 (gk176502)Embryo survival rate %DMSO100 µM ETOP******ns*******ns ***ns********ns12900.511.522.5050100150200250300350N2smrc-1 (gk176502) M+Z-rtel-1rtel-1;smrc-1 (M+Z-)mus-81 M+Z-mus-81; smrc-1 (M+Z-)dog-1dog-1;smrc-1 (M+Z-)helq-1helq-1 smrc-1 (M+Z-)dvc-1dvc-1;smrc-1 (M+Z-)Male percentage %Embryonic lethality/ Brood SizeEmbryo Lethality Brood Size Male %0204060801000 2 4 6 8 10Percentage of LinesGeneration% of Survival linessmrc-1 (gk176502)N2dog-1dog-1;smrc-1mus-81mus-81;smrc-1helq-1helq-1 smrc-1AB* ***** *** **130Figure 5.3 Genetic interactions with loss of SMRC-1. (A) Brood size and embryonic lethality and male percentage single mutants, and smrc-1(gk176502)-containing double mutants. Student t tests were used to compare brood sizes between indicated animals. The brood size of smrc-1(gk176502) was significantly different from rtel-1;smrc-1(*), mus-81;smrc-1(**), and dvc-1;smrc-1(**). * denotes p value less than 0.05, ** <0.01, ***<0.001. Error bars represent the standard error of the mean. (B) Viability of independent lines of N2, smrc-1, rtel-1, helq-1, dog-1 single mutants and smrc-1-containing double mutants at 23℃. (C) Representative images of somatic cells, early embryos and DAPI-stained diakinesis bodies of rtel-1;smrc-1. (D) Representative images of somatic cells and DAPI-stained diakinesis bodies of mus-81;smrc-1 double mutants . Scale bar denotes 5 μm if not indicated. Diakinesis bodies in mus-81;smrc-1Chromatin bridges between intestinal cells in mus-81;smrc-1 Chromatin bridges between intestinal cells in rtel-1;smrc-1 Diakinesis bodies in rtel-1;smrc-1Chromatin bridges in early embryos in rtel-1;smrc-1CD131Table 5.1 Phenotypes of indicated mutants. n denotes the number of parent worms whose progeny were scored. All values are ± standard error of the mean. * indicates the smrc-1 allele gk176502. M+Z-: maternally positive and zygotically negative. M-Z-: maternally negative and zygotically negative. ND: not determined. Genotypes Brood Size Embryo Survival Rate % Male % nN2 256.71±12.00 99.58±0.22 0.03±0.02 14smrc-1 (gk176502) M+Z- 184.40±28.08 91.16±8.80 0.97±0.34 10smrc-1 (gk784642) M+Z- 297.70±15.97 83.67±1.06 0.67±0.20 10smrc-1 (gk176502) M-Z- 159.60±21.59 81.08±7.82 1.08±1.09 10smrc-1 (gk784642) M-Z- 141.30±15.99 73.04±4.28 1.31±0.40 10rtel-1 102.90±20.49 94.17±2.71 0.21±0.32 10mus-81 (M+Z-) 140.40±17.78 94.80±3.47 0.24±0.09 10helq-1 258.40±20.16 98.30±0.37 0.79±0.18 10dog-1 274.80±12.47 98.44±1.78 0.15±0.07 20dvc-1 176.50±27.84 98.41±0.68 1.41±0.57 10rtel-1;smrc-1* (M+Z-) 31.50±8.13 1.27±0.82 ND 10mus-81; smrc-1* (M+Z-) 58.40±20.29 50.51±7.78 0.96±0.38 10helq-1 smrc-1* (M+Z-) 158.80±15.13 82.24±1.44 1.33±0.31 10dog-1;smrc-1* (M+Z-) 134.30±23.31 96.95±0.63 0.48±0.44 10dvc-1;smrc-1* (M+Z-) 49.60±14.19 76.61±11.49 1.24±0.56 10132CHAPTER 6: CONCLUSION 6.1 Anticancer drugs have complex, multimodal mechanisms of action. The promise of genotype targeted anti-cancer therapies raises new challenges. Previously, anti-proliferative drugs targeting the hyper-proliferation of cancer cells were the front line of tumor treatment. Now, genome sequencing technologies are uncovering new actionable biomarkers in tumors, thereby facilitating the development of new highly selective therapeutics. This new wave of therapeutics requires extensive testing not only to assess efficacy but also to confirm their target specificity. It has been recently suggested that off-target toxicity could be the main mechanism of action for many new anticancer drugs undergoing clinical trial (Lin et al., 2019). The efficacy of these drugs was not affected even when their putative targets (genes) were knocked out (Lin et al., 2019). Therefore, understanding the mechanisms of action of new drugs is critical for the effective adoption of new anticancer therapeutics. Using the model metazoan C. elegans, we were able to characterize a new anticancer chemotherapeutic, CX-5461, in a whole live organism. CX-5461 is an example of a therapeutic for which our understanding of the mechanism of action is evolving. CX-5461 was first identified as an RNA polymerase I inhibitor (Drygin et al., 2011). A more recent study showed that the mechanism of action was likely through stabilization of G-quadruplexes, a mechanism that could also account for the RNA polymerase I inhibition (Xu et al., 2017). It is probable that G4-stabilization at sites other than the rDNA-associated G4 was responsible for the selective killing of BRCA-deficient cancer cells, as the concentration of CX-5461 required to kill BRCA-deficient cells was not high enough to inhibit RNA polymerase I (Xu et al., 2017). Interestingly, a wide-range of breast cancer cells were sensitive to CX-5461, some 133of which were BRCA-proficient (Xu et al., 2017), suggesting that a broader range of genotypes may be sensitive to CX-5461 treatment. This led us to test a panel of DNA replication and repair mutants to determine if CX-5461 sensitivity was linked to mutations associated with G4 repair. We also sequenced CX-5461-induced mutations to see if the mutations were at sites that could form G4 DNA structures. In light of our data (as discussed in Chapter 3), it is likely that the mechanism of action of CX-5461 extends beyond G4-stabilization, and may be due, at least in part, to DNA intercalation. CX-5461 could be similar to the anthracycline class of intercalating agents. Anthracyclines, such as daunorubicin and doxorubicin, intercalate into DNA, poison topoisomerases, generate reactive oxygen species, and form covalent DNA adducts (Kizek et al., 2012). Our data strongly supports a model in which CX-5461 intercalates into DNA, resulting in a range of lesions requiring several different repair pathways for tolerance. The therapeutic mechanisms of action and efficacy can be affected by both genetic and environmental factors. Often the effect of environmental factors on therapeutic efficacy is overlooked. For example, the efficacy of some drugs can be affected by the consumption of grapefruit, which contains compounds that can increase the activity and expression of cytochrome P450 CYP1A1 and enhance drug metabolism and detoxification (Hahn-Obercyger et al., 2005). In the case of the antineoplastic ellipticine, P450 metabolism of ellipticine generates metabolites that can covalently bind DNA and result in nucleotide substitution mutations (Stiborova et al., 2004). The metabolism of drugs is primarily performed in the liver (Almazroo et al., 2017). P450-mediated drug metabolism in cell culture is variable depending on the cell line. The importance of P450-mediated metabolism in analyzing DNA damaging agents was demonstrated in a recent study, which showed that some compounds, including ellipticine, were 134only mutagenic after metabolism by rat liver extracts (Kucab et al., 2019). C. elegans is remarkably efficient in P450-mediated metabolism of compounds, and this process is similar to that observed in the human liver (Harlow et al., 2018). It is possible that testing the effects of drugs in C. elegans would also test the effect of drug metabolites, a capacity that would not be available in cultured mammalian cell lines. This could explain differences in CX-5461 genotypic sensitivities observed between human cell lines and C. elegans (Xu et al., 2017). Other environmental factors such as sunlight can also play a role affecting the drug efficacy. The CX-5461 clinical trial resulted in some cases of photosensitivity (Hilton et al., 2018; Khot et al., 2019). We used C. elegans to show that UVA potentiates CX-5461 toxicity by generating ROS (Figure 3.4 and 3.5). To summarize, mechanisms of action for anticancer therapeutics can be complex and dependent on physiological and environmental factors, such as light and diet. Therefore, it is imperative to use in vivo platforms to identify the targeted biological processes and to assess the efficacy and off-target effects. We have demonstrated that C. elegans is a powerful tool with which to investigate the effects of chemotherapeutics in an in vivo model organism. 6.2 C. elegans is a good platform with which to study factors affecting genome stability Maintaining genome integrity is essential for all organisms and the processes that maintain genome stability tend to be evolutionarily conserved. While not all components are conserved at the gene level, the overall repair processes are conserved. For example, the Fanconi anemia (FA) pathway, which repairs interstrand crosslinks (ICLs), is only partially conserved between humans and C. elegans at the protein level. However, mutation of FA pathway genes sensitizes 135C. elegans specifically to ICL agents (Figure 2.2), in a similar fashion as the human counterpart (Deans and West, 2011). Although the C. elegans FA pathway contains fewer components than the human FA pathway, it could represent the minimal requirements for functional ICL repair and simplify the study of the core FA repair processes (Youds et al., 2009). This is also true for other DNA repair pathways in C. elegans. C. elegans has fewer RecQ helicases, PARPs, homologous recombination proteins, and annealing helicases. The smaller number of repair genes in C. elegans makes it easier to investigate the role of specific genes in response to anticancer therapeutics and to dissect the interplay between repair pathways. C. elegans provides a simplified in vivo platform for studying human diseases such as cancer. The simpler DNA replication and repair pathways in C. elegans make it a great model in studying the biology of genes involved in genome maintenance. SMRC-1 is the only annealing helicase known in C. elegans, making it a simple model to investigate the role of annealing helicases. Loss of SMRC-1 has been shown to induce genome instability. Examination of the genome from late generation smrc-1 mutants revealed many small CNVs (Figure 5.1B & Figure A5.2). Despite that, the smrc-1 mutants were still viable at F10, indicating that these chromosomal duplications or deletions may be the byproducts of repair or bypass pathways that are needed in the absence of SMRC-1 for survival. The simpler DNA repair pathways in C. elegans also benefit the dissection of interplay between different repair proteins. This thesis has illustrated the overlapping and competing roles of different repair mechanisms using C. elegans. For instance, MMEJ repair pathway can act as an alternate repair mechanism to repair DNA lesions that are normally repaired by HDR (Figure 1364.9). Even within the HDR repair pathway, there are many proteins that show functional redundancy. In HDR-compromised mutants (such as helq-1), improper NHEJ activity could occur and lead to genome destabilizing effects. Such scenarios are common in a tumor, that the redundancies and alleviation in different repair mechanisms could allow a tumor to survive and evolve through accumulating more mutations that favor survival, making anticancer treatment more complicated (Nickoloff et al., 2017). In addition to the advantages of a simpler repertoire of DNA damage response genes, there is a suite of tools and phenotypic readouts in C. elegans that can be exploited to query the effects of genome stability. For instance, brood size, embryo viability, and the frequency of spontaneous males can be used to assess whether a gene mutation leads to genome instability. The well characterized development of C. elegans allows one to study the effects of genome instability in different tissue types or under different growth conditions. For example, the quantitative acute assay measures the effect of genome instability in embryos, which undergo rapid division and contain no DNA damage checkpoints, whereas the L1 exposure assay measures genome instability affecting post-embryonic somatic cell development and can be used to determine if a DNA damaging agent affects transcription (Astin et al., 2008). In addition, the C. elegans germline nuclei are arranged in a spatio-temporal gradient in which sequential meiotic stages are easily differentiated, making C. elegans a great platform to investigate meiotic events, such as meiotic DSB repair (Lui and Colaiacovo, 2013). Genetic balancers in C. elegans provide tools with which to measure the mutagenicity of the factors of interest (Rosenbluth et al., 1983) and allow for the capture and stable propagation of induced mutations, which is necessary for whole genome sequencing to delineate mutational signatures (Edgley et al., 2006). 137 In this thesis, we have demonstrated how C. elegans can be exploited to investigate the phenotypic effects of internal and external factors that affect genome stability. One could use the same strategies that we used to characterize CX-5461 in C. elegans to study other genome-destabilizing agents, whether they are chemical agents or anticancer therapeutics. We have also generated a chemi-genetic interaction map that is a useful reference for the study of new DNA damaging agents or DNA repair mutants. By generating and comparing the genotypic sensitivity profile of the agent of interest, it is possible to form a hypothesis on the mechanism of action of the agent. 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Acta dermatovenerologica Croatica : ADC 24, 55-64. 1583210321032103210Chromosome Figure A3.1 Coverage plot of CX-5461-induced genome rearrangements (A-D, 4 in total) I II III IV V VI A BC DCopy numberCopy numberCopy numberCopy numberAPPENDIX1594321043210432104321043210432104321043210A BC DE FG HCopy numberCopy numberCopy numberCopy numberCopy numberCopy numberCopy numberCopy number1604321043210432104321043210432104321043210I JK LM NO PCopy numberCopy numberCopy numberCopy numberCopy numberCopy numberCopy numberCopy number161432104321043210Chromosome Figure A3.2 Coverage plot of CX-5461 + UVA-induced genome rearrangements (A-S, 19 in total) Q RSCopy numberCopy numberCopy number162Figure A4.1 Representative images of DAPI-stained bodies in diakinesis of cku-80 and polq-1 single mutants.Both mutants have a near wild-type phenotype that displays six DAPI-stained bodies in diakinesis. Numbersbelow indicate the number of DAPI-stained bodies per nuclei. Scale bar denotes 5 μm.cku-80polq-16 66 6163Figure A5.1 Coverage plot of quantile deviations and fractions of variant basis of Million Mutation Project collection strains VC20168 and VC40734. Pictures were screenshotted from http://genome.sfu.ca/mmp/. (Thompson et al., 2013)164Figure A5.2 Coverage plot of genome rearrangements of F10 gk176502 (A - E) and F10 gk784642 (F - G).Copy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XCopy numberChromosome I II III IV V XADCBDEF G165DMSO 500 µM OlaSensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 99.83±0.29 522 98.25±1.97 411 1.00 0.3392mus-81 90.74±4.62 436 59.80±0.99 345 0.67 0.0046 0.0009atm-1 79.64±6.27 330 44.10±9.61 352 0.56 0.0250 0.0097polq-1 98.52±1.03 479 97.47±1.72 433 1.01 0.5155 0.7388him-1 99.81±0.33 465 99.40±0.59 464 1.01 0.5107 0.4651cku-80 99.07±0.30 533 99.10±0.79 442 1.02 0.9233 0.5217gen-1 99.30±0.64 422 99.78±0.38 414 1.02 0.3478 0.3653fncm-1 77.71±7.04 490 76.38±11.73 499 1.00 0.9097 0.0780fan-1 99.65±0.61 269 88.08±2.74 343 0.90 0.0113 0.0029rfs-1 92.47±4.37 489 87.17±1.91 477 0.96 0.1397 0.0590dog-1 99.73±0.47 318 90.26±8.64 416 0.92 0.2123 0.1588rtel-1 99.77±0.41 398 99.77±0.41 409 1.02 1.0000 0.3618N2 98.62±0.88 501 96.73±1.69 429 1.00 0.3222mus-81 91.77±6.12 306 63.70±7.63 252 0.71 0.0096 0.0207him-6 40.15±2.17 475 45.14±4.79 306 1.15 0.0876 0.0014ercc-1 68.09±2.94 396 71.63±3.98 467 1.07 0.1503 0.0112helq-1 88.22±7.79 399 69.42±2.88 372 0.80 0.0829 0.0044fcd-2 99.61±0.68 485 97.66±0.97 424 1.00 0.1667 0.5232let-418 96.70±2.88 188 96.29±3.40 234 1.02 0.8362 0.8156N2 97.75±2.04 454 94.87±4.75 243 1.00 0.5383mus-81 91.12±3.23 130 35.83±9.27 274 0.41 0.0118 0.0113rcq-5 99.46±0.47 374 98.61±0.86 307 1.02 0.1833 0.2989cdk-5 99.22±0.71 424 99.25±0.66 267 1.03 0.9721 0.2558polk-1 98.47±0.87 403 99.05±0.89 299 1.04 0.3957 0.2015cep-1 98.89±0.40 296 98.90±1.91 288 1.03 0.9969 0.3993brd-1 97.92±3.61 279 99.16±0.87 371 1.04 0.6243 0.2372N2 99.73±0.47 394 98.41±0.36 441 1.00 0.0638mus-81 90.46±8.13 197 33.19±3.59 186 0.37 0.0135 0.0012msh-2 83.85±9.81 364 84.64±2.05 292 1.02 0.9043 0.0093N2 99.43±0.50 356 98.48±0.77 269 1.00 0.1332mus-81 92.96±7.18 117 55.94±5.46 223 0.61 0.0069 0.0046rev-3 90.45±3.85 216 87.82±3.88 200 0.98 0.4479 0.0302polh-1 93.88±2.96 390 91.59±10.68 454 0.99 0.6592 0.3936N2 98.08±0.43 364 99.33±0.58 314 1.00 0.0349mus-81 90.24±3.62 218 38.45±12.26 263 0.42 0.0138 0.0131lig-4 99.65±0.60 289 98.01±1.05 305 0.97 0.0393 0.1837xpa-1 98.72±1.24 287 95.18±2.42 295 0.95 0.1400 0.0808wrn-1 98.77±1.07 332 98.10±1.42 300 0.98 0.3981 0.3982hsr-9 65.94±7.68 304 60.28±12.76 326 0.90 0.6756 0.0365N2 99.71±0.51 341 99.23±0.79 401 1.00 0.4856mus-81 90.62±2.18 247 39.84±13.47 232 0.44 0.0296 0.0175gld-1 94.43±1.95 261 89.30±1.85 247 0.95 0.0606 0.0226hda-3 93.45±0.91 341 80.84±2.31 343 0.87 0.0398 0.0156Table A1.1 Raw data for generating the sensitivity scores for Olaparib. 166DMSO 100 nM CPTSensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 98.55±0.57 547 98.30±1.30 485 1.00 0.6284mus-81 94.27±3.34 433 6.63±1.68 312 0.07 0.0001 0.0001fncm-1 97.49±1.67 485 98.24±0.75 385 1.01 0.6404 0.9284fan-1 99.07±0.80 241 99.59±0.70 267 1.01 0.3492 0.3593rfs-1 93.99±1.52 381 17.42±7.96 308 0.19 0.0048 0.0033dog-1 97.58±1.51 284 88.83±10.94 326 0.91 0.3344 0.2839rtel-1 99.23±0.87 407 98.09±1.07 385 0.99 0.2244 0.8237N2 99.78±0.37 407 97.12±0.76 309 1.00 0.0260mus-81 91.07±2.88 389 0.32±0.56 362 0.00 0.0005 0.0001him-1 80.55±7.22 216 25.80±3.13 218 0.33 0.0047 0.0009smrc-1 84.98±4.64 362 10.10±6.07 331 0.12 0.0054 0.0015N2 98.62±0.88 501 98.43±0.92 397 1.00 0.8329mus-81 91.77±6.12 306 3.21±4.73 266 0.04 0.0025 0.0008him-6 40.15±2.17 475 39.05±8.25 334 0.97 0.8690 0.0059ercc-1 68.09±2.94 396 67.68±1.88 388 1.00 0.5749 0.0026helq-1 88.22±7.79 399 10.43±2.87 362 0.12 0.0060 0.0006atm-1 82.60±2.55 406 29.59±4.23 479 0.36 0.0046 0.0013fcd-2 99.61±0.68 485 99.25±0.75 399 1.00 0.5945 0.4357let-418 96.70±2.88 188 96.53±0.61 346 1.00 0.9114 0.1411polq-1 96.94±2.19 445 92.28±8.14 367 0.95 0.4335 0.3330cku-80 99.32±0.04 444 99.39±0.06 496 1.00 0.3350 0.2295N2 97.75±2.04 454 56.39±1.81 203 1.00 0.0000mus-81 91.12±3.23 130 1.47±1.45 214 0.03 0.0007 0.0001rcq-5 99.46±0.47 374 62.26±1.50 172 1.09 0.0005 0.0620polk-1 98.47±0.87 403 61.34±8.30 203 1.08 0.0186 0.4625N2 99.73±0.47 394 96.15±1.22 298 1.00 0.0232mus-81 90.46±8.13 197 0.95±0.82 242 0.01 0.0032 0.0001msh-2 83.85±9.81 364 33.67±12.08 300 0.42 0.0151 0.0110brd-1 97.85±2.35 370 16.28±3.02 308 0.17 0.0001 0.0009cep-1 96.50±1.49 364 56.05±10.23 218 0.60 0.0263 0.0201N2 99.43±0.50 356 99.33±0.58 278 1.00 0.8655mus-81 92.96±7.18 117 1.66±1.88 176 0.02 0.0002 0.0002rev-3 90.45±3.85 216 86.79±5.68 138 0.96 0.1302 0.0726polh-1 93.88±2.96 390 58.29±9.59 273 0.62 0.0354 0.0161N2 98.08±0.43 364 87.93±7.60 335 1.00 0.1373mus-81 90.24±3.62 218 1.09±1.89 196 0.01 0.0007 0.0038lig-4 99.65±0.60 289 83.22±0.28 262 0.93 0.0004 0.3913wrn-1 98.77±1.07 332 81.84±4.78 246 0.92 0.0158 0.4779cdk-5 99.08±0.81 237 74.92±7.96 210 0.84 0.0396 0.1438N2 99.71±0.51 341 77.70±11.64 288 1.00 0.0819mus-81 90.62±2.18 247 0.34±0.60 254 0.00 0.0002 0.0068gld-1 94.43±1.95 261 58.70±18.38 251 0.80 0.0650 0.0960hda-3 93.45±0.91 341 83.56±2.31 310 1.15 0.2314 0.1622N2 98.10±1.24 368 97.82±2.05 288 1.00 0.6607mus-81 88.92±5.45 131 6.86±6.95 158 0.08 0.0075 0.0014xpa-1 96.07±1.73 321 94.46±1.60 326 0.99 0.2465 0.0109N2 99.28±0.69 393 95.75±3.75 357 1.00 0.2023hsr-9 78.32±9.18 307 69.32±16.32 310 0.92 0.5990 0.1282dvc-1 89.60±7.96 346 88.90±2.88 256 1.03 0.8351 0.0169Table A1.2 Raw data for generating the sensitivity scores for Camptothecin. 167DMSO 100 µM ETOPSensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 90.75±2.88 454 82.93±9.38 316 1.00 0.1980mus-81 75.56±11.80 268 4.24±2.61 273 0.06 0.0126 0.0068polq-1 90.56±2.17 519 26.38±6.32 363 0.32 0.0018 0.0203helq-1 84.31±4.25 519 59.32±5.38 465 0.77 0.0090 0.0677brd-1 86.79±3.26 479 65.12±5.40 491 0.82 0.0490 0.1016rfs-1 65.79±7.82 453 32.89±6.03 511 0.55 0.0496 0.0195N2 91.76±7.59 462 83.98±8.70 386 1.00 0.1954mus-81 89.81±3.39 304 5.53±0.28 326 0.07 0.0005 0.0042gld-1 94.01±3.26 272 81.36±6.01 311 0.95 0.0656 0.3420polh-1 91.88±4.89 252 58.25±3.87 309 0.69 0.0004 0.0265fan-1 98.13±1.14 471 96.34±0.44 466 1.07 0.1848 0.1437fcd-2 91.84±1.40 405 94.78±2.34 424 1.13 0.2582 0.1228fncm-1 94.69±1.79 453 91.33±3.73 418 1.05 0.1977 0.4132N2 97.57±1.83 305 80.10±3.88 331 1.00 0.0230mus-81 88.84±3.41 292 7.76±5.00 268 0.11 0.0003 0.0002wrn-1 97.82±1.33 316 92.09±4.43 405 1.15 0.0912 0.1102lig-4 97.27±1.31 392 86.32±0.37 380 1.08 0.0047 0.1004msh-2 84.72±7.18 228 55.11±5.75 270 0.79 0.0477 0.0374cep-1 92.51±3.05 389 71.52±5.90 374 0.94 0.0068 0.0201N2 98.98±0.36 384 85.26±6.23 347 1.00 0.0681mus-81 81.53±9.15 354 11.60±4.67 323 0.17 0.0022 0.0064him-1 85.33±2.23 293 5.89±2.33 271 0.08 0.0003 0.0022let-418 90.36±2.86 340 78.01±12.55 357 1.00 0.2923 0.2976hda-3 84.84±3.67 338 60.62±9.55 338 0.83 0.0199 0.0342ercc-1 63.96±3.72 299 56.16±4.66 366 1.02 0.1630 0.0055dvc-1 89.25±1.72 357 70.02±12.73 280 0.91 0.1078 0.2374N2 92.26±1.70 580 91.65±2.37 452 1.00 0.7306mus-81 70.07±5.81 447 18.03±4.52 346 0.26 0.0058 0.0026xpa-1 89.20±1.32 484 81.33±8.81 374 0.92 0.2898 0.1761rtel-1 80.70±4.84 240 39.32±1.62 326 0.49 0.0066 0.0008smrc-1 55.54±2.93 365 23.68±5.54 450 0.43 0.0207 0.0041rcq-5 91.60±3.65 551 89.23±5.91 441 0.98 0.3896 0.6569N2 98.66±0.44 371 89.75±6.76 483 1.00 0.1634mus-81 74.27±7.88 292 8.89±2.58 342 0.13 0.0053 0.0021atm-1 92.17±2.70 391 60.95±12.57 344 0.73 0.0443 0.0836rev-3 95.62±1.71 449 89.03±6.15 320 1.02 0.2823 0.9236dog-1 95.10±4.40 368 43.63±8.32 325 0.50 0.0174 0.0336cku-80 95.99±4.62 538 92.01±5.76 422 1.05 0.5307 0.3488Table A1.3 Raw data for generating the sensitivity scores for Etoposide. 168TMP TMP + UVA Sensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 99.82±0.32 506 97.64±1.85 449 1.00 0.1333mus-81 91.42±5.51 247 11.73±10.20 146 0.13 0.0027 0.0032ercc-1 69.87±2.88 516 12.73±2.56 506 0.19 0.0010 0.0007him-6 46.32±6.52 392 39.06±4.62 452 0.86 0.1856 0.0040him-9 77.39±3.17 354 11.45±2.63 359 0.15 0.0000 0.0001cep-1 99.72±0.25 691 93.69±5.28 522 0.96 0.1761 0.2231gld-1 95.24±1.91 192 97.31±2.46 217 1.04 0.4814 0.9037N2 98.11±0.97 791 95.42±0.41 832 1.00 0.0221mus-81 82.86±0.63 375 25.42±13.53 418 0.32 0.0194 0.2547polq-1 98.98±0.98 775 52.93±6.79 648 0.55 0.0077 0.0098brd-1 98.98±0.18 681 49.69±6.63 644 0.52 0.0062 0.0089rfs-1 91.26±2.89 608 43.15±8.42 588 0.49 0.0166 0.0279N2 99.44±0.55 541 98.05±0.77 523 1.00 0.2004mus-81 87.70±5.36 459 14.74±6.45 397 0.17 0.0029 0.0016hda-3 96.61±1.35 441 66.61±3.11 442 0.70 0.0013 0.0027N2 99.22±0.76 369 93.95±2.74 286 1.00 0.0443mus-81 87.44±6.10 250 3.97±2.88 275 0.05 0.0014 0.0000lig-4 98.47±0.52 327 96.02±2.45 308 1.03 0.2444 0.1371wrn-1 99.36±0.66 454 91.56±6.20 608 0.97 0.1839 0.3684xpa-1 97.37±2.47 734 28.76±11.75 726 0.31 0.0132 0.0160N2 99.80±0.35 517 95.39±0.63 574 1.00 0.0162mus-81 96.57±0.09 175 19.07±4.39 173 0.21 0.0010 0.0008hsr-9 95.31±1.81 482 90.22±3.08 452 0.99 0.0507 0.1330N2 98.73±0.36 704 85.17±11.55 786 1.00 0.1707mus-81 91.97±1.10 223 7.14±6.84 171 0.09 0.0026 0.0033him-1 72.16±7.42 482 51.06±2.62 332 0.82 0.0547 0.0438fcd-2 97.45±1.53 760 46.19±16.82 655 0.55 0.0334 0.0175N2 98.88±1.52 570 99.62±0.33 511 1.00 0.4604mus-81 92.88±4.20 351 17.13±10.90 287 0.18 0.0119 0.0058rev-3 93.88±3.02 249 15.42±6.41 239 0.16 0.0031 0.0021cku-80 99.00±0.68 710 88.69±11.14 635 0.89 0.1114 0.1067cdk-5 99.62±0.40 838 87.09±14.76 620 0.87 0.2707 0.2759let-418 92.58±8.29 279 46.85±3.71 242 0.50 0.0208 0.0015N2 99.71±0.25 702 96.35±3.23 679 1.00 0.2331mus-81 89.69±8.15 331 22.82±8.83 314 0.26 0.0039 0.0025dog-1 98.07±1.38 470 78.28±14.62 547 0.83 0.1609 0.1709polk-1 99.86±0.25 794 97.20±2.13 830 1.01 0.1511 0.8029polh-1 73.08±12.52 395 0.00±0.00 390 0.00 0.0096 0.0004rcq-5 99.63±0.34 792 98.76±1.26 582 1.03 0.2822 0.4365msh-2 88.38±3.24 671 21.20±3.27 483 0.25 0.0018 0.0007N2 99.82±0.32 831 94.83±1.78 798 1.00 0.0341mus-81 91.42±5.51 375 5.10±1.72 376 0.13 0.0222 0.0002rtel-1 69.87±2.88 277 50.40±13.74 297 0.63 0.0566 0.0367atm-1 46.32±6.52 691 55.11±9.97 534 0.67 0.0061 0.0212fncm-1 77.39±3.17 589 26.31±2.99 647 0.28 0.0004 0.0004fan-1 99.72±0.25 615 10.67±7.37 621 0.11 0.0019 0.0039helq-1 95.24±1.91 504 40.01±2.09 699 0.48 0.0068 0.0012N2 84.51±1.80 687 50.40±13.74 615 1.00 0.0540mus-81 40.19±12.91 424 1.62±2.81 309 0.01 0.0003 0.0005dvc-1 86.63±8.82 709 55.11±9.97 658 0.70 0.0075 0.0203smrc-1 97.74±0.65 663 26.31±2.99 641 0.75 0.0484 0.0171Table A1.4 Raw data for generating the sensitivity scores for UVA-Trimethylpsoralen. 1690J UVC 50J UVC Sensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 99.19±0.75 582 81.35±5.10 631 1.00 0.0335mus-81 91.73±6.97 256 27.05±4.68 205 0.36 0.0007 0.0062smrc-1 70.73±9.86 382 40.57±3.81 401 0.70 0.0285 0.0007hsr-9 91.46±0.88 452 67.77±1.42 523 0.90 0.0003 0.0370polh-1 92.73±3.79 172 1.46±1.30 163 0.02 0.0007 0.0008N2 99.49±0.89 192 70.51±2.50 606 1.00 0.0041mus-81 87.26±2.39 203 18.56±3.77 338 0.30 0.0022 0.0036xpa-1 91.52±7.83 541 23.53±6.12 328 0.36 0.0062 0.0090gld-1 88.02±2.08 241 54.69±6.38 205 0.88 0.0067 0.0765cep-1 99.44±0.97 336 50.01±4.70 797 0.71 0.0026 0.0089cku-80 100.00±0.00 591 72.33±3.85 528 1.02 0.0064 0.6684wrn-1 99.22±0.37 521 80.34±3.81 600 1.14 0.0116 0.0151N2 99.86±0.25 693 72.71±3.27 719 1.00 0.0051mus-81 94.45±3.34 558 28.51±1.20 378 0.41 0.0014 0.0021let-418 85.42±13.68 350 64.70±2.55 305 1.04 0.0951 0.0383rtel-1 98.69±0.53 390 78.11±2.34 244 1.09 0.0059 0.0364fncm-1 99.48±0.48 574 59.73±3.39 551 0.82 0.0030 0.0277fan-1 99.05±1.65 222 70.90±5.76 616 0.98 0.0080 0.6628fcd-2 99.72±0.49 324 72.24±4.28 511 0.99 0.0066 0.6917polk-1 99.60±0.40 752 75.37±3.33 693 1.04 0.0071 0.4493polq-1 99.33±1.16 476 76.95±3.60 490 1.06 0.0043 0.0545N2 100.00±0.00 222 85.20±6.12 256 1.00 0.0526mus-81 90.58±5.53 346 28.34±6.86 308 0.37 0.0043 0.0072helq-1 91.29±4.36 392 61.62±4.63 448 0.79 0.0117 0.0625rev-3 91.14±3.49 230 39.33±7.01 233 0.51 0.0132 0.0262ercc-1 65.80±2.41 464 9.38±2.97 210 0.17 0.0001 0.0031him-9 83.78±0.97 443 13.03±10.85 117 0.18 0.0085 0.0035cdk-5 99.59±0.35 487 75.22±7.87 457 0.89 0.0311 0.3152rfs-1 94.19±3.59 382 80.24±5.56 355 1.00 0.0206 0.1475N2 100.00±0.00 203 82.30±8.22 169 1.00 0.0650mus-81 89.27±6.37 136 26.69±1.88 232 0.36 0.0055 0.0066dog-1 98.38±1.10 272 73.62±5.16 379 0.91 0.0188 0.3470rcq-5 99.76±0.42 418 77.30±5.07 467 0.94 0.0162 0.1350msh-2 86.27±4.60 454 79.55±2.51 450 1.12 0.1432 0.4963him-6 47.74±4.04 378 32.22±0.97 447 0.82 0.0242 0.0098brd-1 97.94±1.90 484 74.97±1.42 471 0.93 0.0020 0.3092N2 99.08±0.80 419 89.49±1.32 546 1.00 0.0159mus-81 81.10±6.15 413 31.23±6.17 450 0.43 0.0061 0.0029him-1 84.42±6.05 385 56.37±1.41 573 0.74 0.0124 0.0018N2 99.84±0.28 613 77.79±8.06 390 1.00 0.1681mus-81 85.24±10.63 251 23.14±6.64 233 0.35 0.0226 0.0036atm-1 93.68±5.89 263 46.00±4.86 228 0.63 0.0013 0.0107N2 98.98±0.89 405 89.34±3.06 397 1.00 0.0165mus-81 94.60±3.31 236 38.66±9.18 292 0.45 0.0129 0.0162dvc-1 90.40±10.93 436 77.58±2.39 532 0.95 0.1234 0.0636N2 99.81±0.33 542 79.05±4.12 508 1.00 0.0150mus-81 87.82±7.00 351 23.81±9.06 238 0.34 0.0008 0.0047hda-3 99.03±1.01 336 73.17±6.99 413 0.93 0.0174 0.4487N2 98.51±1.37 347 76.77±5.44 651 1.00 0.0286mus-81 94.00±3.97 470 28.74±3.95 293 0.39 0.0047 0.0113lig-4 99.38±0.58 452 75.53±6.87 610 0.98 0.0259 0.8420Table A1.5 Raw data for generating the sensitivity scores for UVC. 170Phosphate Buffer 100 µM CX-5461 Sensitivity scoreP-value (in t-test with mock treatment)P-value (in t-test with N2)Genotype Embryo survival rate %Number of progeny countedEmbryo survival rate %Number of progeny countedN2 99.70±0.52 388 99.08±0.94 322 1.00 0.4325mus-81 93.92±1.01 338 3.24±4.82 270 0.03 0.0006 0.0008cep-1 99.68±0.56 284 96.62±3.12 196 0.98 0.1902 0.3803N2 99.79±0.37 407 99.05±0.94 311 1.00 0.3319mus-81 90.57±2.62 344 2.50±4.33 239 0.03 0.0012 0.0007hda-3 99.83±0.30 528 94.72±3.16 345 0.96 0.0976 0.1973hsr-9 99.75±0.44 435 97.23±3.64 413 0.98 0.3081 0.5068N2 99.81±0.33 449 94.87±1.68 340 1.00 0.0264atm-1 91.57±5.89 154 1.11±1.92 133 0.01 0.0014 0.0001him-1 89.55±3.60 317 32.94±6.17 202 0.39 0.0096 0.0049gld-1 97.52±1.73 202 81.05±2.75 148 0.87 0.0151 0.0437N2 99.22±0.02 384 90.12±8.66 292 1.00 0.2106mus-81 92.50±5.10 265 5.58±4.83 196 0.07 0.0038 0.0064dvc-1 96.09±5.35 178 74.41±5.67 142 0.85 0.0642 0.0885N2 99.17±0.82 351 96.17±2.42 237 1.00 0.0850mus-81 92.23±3.95 298 1.79±3.09 174 0.02 0.0017 0.0000xpa-1 98.69±1.51 368 25.49±5.22 369 0.27 0.0011 0.0006ercc-1 55.28±10.11 232 4.18±1.22 415 0.08 0.0160 0.0001N2 99.32±0.71 465 0.70±300.00 300 1.00 0.0918mus-81 86.31±5.57 363 1.72±290.00 290 0.02 0.0019 0.0001msh-2 96.35±4.51 271 1.80±339.00 339 1.03 0.8917 0.8930N2 98.84±2.01 389 88.45±0.89 390 1.00 0.0089mus-81 87.43±4.64 248 1.45±1.36 206 0.02 0.0016 0.0000let-418 92.71±4.04 248 63.18±10.21 233 0.76 0.0334 0.0434polh-1 98.39±1.76 566 7.80±6.89 171 0.09 0.0011 0.0025N2 99.75±0.43 398 99.67±0.57 312 1.00 0.8932mus-81 86.36±8.55 277 0.44±0.77 269 0.01 0.0034 0.0000lig-4 99.72±0.49 378 99.72±0.49 293 1.00 0.4226 0.9408rcq-5 98.75±0.42 316 98.15±2.04 251 0.99 0.5868 0.3519wrn-1 95.96±7.00 449 94.44±5.30 379 0.98 0.2645 0.2486N2 99.29±0.73 438 99.03±0.96 298 1.00 0.7918mus-81 90.91±4.93 339 1.38±1.20 263 0.02 0.0013 0.0000cku-80 98.79±1.31 349 97.34±0.81 218 0.99 0.2183 0.2307brd-1 97.49±1.46 315 57.68±4.73 343 0.59 0.0058 0.0028polq-1 99.07±1.61 451 43.69±3.93 283 0.44 0.0033 0.0025rfs-1 95.04±0.62 402 55.53±4.41 344 0.59 0.0048 0.0027helq-1 90.51±2.36 406 10.13±5.96 380 0.11 0.0034 0.0011N2 99.22±0.77 375 97.38±2.28 154 1.00 0.3074dog-1 95.33±3.98 277 89.23±4.65 187 0.95 0.3428 0.2383N2 99.72±0.49 302 95.30±2.31 273 1.00 0.0521mus-81 92.70±4.67 316 0.00±0.00 202 0.00 0.0008 0.0002fncm-1 97.94±0.43 430 85.23±2.88 295 0.91 0.0124 0.0406fan-1 98.96±0.96 255 90.27±3.40 266 0.95 0.0593 0.1273fcd-2 98.21±3.10 416 82.10±11.99 295 0.87 0.2058 0.1755smrc-1 88.68±0.93 353 19.92±2.21 207 0.24 0.0001 0.0011rev-3 97.71±10.11 332 1.37±1.22 179 0.01 0.0002 0.0014Table A1.6 Raw data for generating the sensitivity scores for CX-5461. 171Notes for Table A1.1-A1.6. These are the raw data for generating chemi-genetic interaction map of Olaparib (Ola), Camptothecin (CPT), Etoposide (ETOP), UVA-TMP (Trimethylpsoralen), UVC and CX-5461. Different sub-sections in each table indicate different batches of experiments. N2 animals were used as positive control, whereas mus-81 mutants were often used as a negative control. The calculation of sensitivity score is detailed in Chapter 2 Method. Worm Strain GenotypeWPH90 smrc-1(gk176502)/hT2WPH91 smrc-1(gk784642)/hT2WPH92 dvc-1(ok260);smrc-1(gk176502)WPH103 helq-1(tm2134) cku-80(ok861) WPH108 cku-80(ok861) rfs-1(ok1372)WPH114 cku-80(ok861) polq-1(tm2026)WPH124 helq-1(tm2134) rfs-1(ok1372)/helq-1(tm2134) hT2WPH125 helq-1(tm2134) cku-80(ok861) rfs-1(ok1372)/helq-1(tm2134) + hT2WPH132 polq-1(tm2026) rfs-1(ok1372)WPH134 cku-80(ok861) polq-1(tm2026) rfs-1(ok1372) WPH140 helq-1(tm2134) cku-80(ok861) polq-1(tm2026) WPH142 helq-1(tm2134) polq-1(tm2026)WPH145 helq-1(tm2134) polq-1(tm2026)/helq-1(tm2134) hT2WPH147 polq-1(tm2026) rfs-1(ok1372)/hT2WPH148 cku-80(ok861) polq-1(tm2026) rfs-1(ok1372)/+ hT2WPH168 mus-81(tm1937) dog-1(gk10) WPH169 mus-81(tm1937) dog-1(gk10)/hT2 dog-1(gk10)WPH172 atm-1(tm5027) dog-1(gk10) WPH176 dog-1(gk10);polq-1(tm2026)/hT2(III)WPH179 atm-1(tm5027) dog-1(gk10)/hT2 dog-1(gk10)WPH181 dog-1(gk10);rev-3(gk919715)WPH189 helq-1(tm2134) polq-1(tm2026) rfs-1(ok1372)/helq-1(tm2134) hT2WPH191 helq-1(tm2134) cku-80(ok861) polq-1(tm2026) rfs-1(ok1372)/helq-1(tm2134) + hT2WPH201 brd-1(dw1) rfs-1(ok1372)/hT2WPH202 cku-80(ok861) brd-1(dw1) WPH205 polq-1(tm2026) brd-1(dw1)WPH217 polq-1(tm2026) brd-1(dw1)/hT2WPH221 cku-80(ok861) brd-1(dw1) rfs-1(ok1372)/+ hT2WPH222 cku-80(ok861) brd-1(dw1)/+ hT2WPH228 cku-80(ok861) polq-1(tm2026) brd-1(dw1)/+ hT2WPH229 helq-1(tm2134) brd-1(dw1)/hT2WPH229 dog-1(gk10);smrc-1(gk176502)/hT2WPH231 helq-1(tm2134) polq-1(tm2026) brd-1(dw1)/helq-1(tm2134) hT2WPH233 mus-81(tm1937)/hT2; smrc-1(gk176502)/hT2WPH236 rtel-1(tm1866)lhT2;smrc-1(gk176502)/T2WPH243 helq-1(tm2134) smrc-1(gk176502)/helq-1(tm2134) hT2WPH244 helq-1(tm2134) cku-80(ok861) brd-1(dw1)/helq-1(tm2134) + hT2WPH248 cku-80(ok861) polq-1(tm2026)/+ hT2WPH249 smrc-1 (gk176502)WPH250 smrc-1 (gk784642)WPH251 helq-1(tm2134) brd-1(dw1) rfs-1(ok1372)/helq-1(tm2134) hT2WPH279 spo-11(me44)/+;brd-1(dw1) rfs-1(ok1372)WPH281 rev-3(gk919715)WPH282 helq-1(tm2134) cku-80(ok861) polq-1(tm2026) brd-1(dw1)/helq-1(tm2134) + hT2Table A2. Worm strains generated in this thesis. WPH is the Hieter Lab worm collection.172
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C. elegans as a platform to characterize genes and therapeutics affecting genome stability
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