A Genome-Scale RNA Interference Screen Identifies Novel Regulators of DNA Double-Strand Break Repair

Jordan Young

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Molecular Genetics University of Toronto

Jordan Young

Doctor of Philosophy

Department of Molecular Genetics, University of Toronto, 2016 Abstract

All living organisms are continuously challenged by agents in their normal cellular environment that can inflict damage to their genetic material. DNA damage can have negative implications on cellular fitness by disrupting genomic processes such as DNA replication and gene expression. In addition, DNA lesions can lead to gene mutation and gross chromosomal rearrangements, events that are implicated in the development of cancer. DNA double-strand breaks (DSBs) are considered one of the most severe types of DNA damage. To protect genome integrity, organisms have evolved several DSB repair mechanisms. Homologous recombination (HR) is an essential DSB repair pathway that is of critical importance during the S- and G2-phases of the cell cycle. HR plays a central role in promoting genome stability and preventing tumourigenesis. For example, mutations in the genes coding for the HR factors BRCA1 and BRCA2 are responsible for familial breast and ovarian cancer. The limiting step in the HR pathway is the generation of single-stranded DNA (ssDNA) by DNA end resection. In this thesis, I describe the establishment of an immunofluorescence-based assay that monitors end resection by quantitative image-based cytometry (QIBC). Employing this assay, I conducted a plate-based RNA interference (RNAi) screen using a library that targets 18,452 genes. As expected, the top hits in my screen were known regulators of end resection including CtIP and all three subunits of the MRE11-RAD50-NBS1 (MRN) complex. I also outline the validation of one the strongest candidate end resection activators identified in the screen, the zinc finger protein, ZNF335. I demonstrate that ZNF335 promotes DSB repair by HR through the enhancement of DNA end resection. In the final chapter of this thesis, I describe the establishment of a cell microarray- based platform for conducting RNAi screens. The platform circumvents the long liquid-handling robotic procedures and large quantities of reagents that are common for plate-based screens.

Acknowledgements

The completion of the work presented in this thesis has been the most challenging but rewarding endeavor of my young scientific career. I would like to thank, first and foremost, my supervisor Daniel Durocher for giving me the opportunity to work in his lab. My doctoral research had its ups and downs, but Dan always displayed an infectious enthusiasm that kept me moving forward with a positive attitude. I could not have asked for a more brilliant and creative scientific mentor to guide me through this work and I feel so privileged to have been trained in Dan’s lab. The knowledge and skills I have acquired in the Durocher lab are invaluable and I know they will endow me with a successful career in science.

I would also like to thank all of my committee members past and present: Jason Moffat, Bret Pearson, and Corey Nislow. Their unwavering support of my scientific goals, even when the project did not go as planned, was commendable. Furthermore, their thoughtful suggestions were essential to the completion of this thesis.

Thinking back at what I have learned during my time in the Durocher lab is astonishing. I could not have acquired these skills without the help of numerous Durocher lab members both past and present. Throughout my time in the Durocher lab, I have met so many talented and brilliant scientists from all over the globe. I would like to thank two post-doctoral fellows that have especially mentored me: Cristina Escribano-Diaz and Lara O’Donnell.

I would also like to extend acknowledgement to my colleagues at the Lunenfeld- Tanenbaum Research Institute that were essential to this research: Gagan Gupta (Pelletier lab), Thomas Sun and Alessandro Datti (Robotics facility), Mikhail Bashkurov and Cyrus Handy (High-content imaging facility), and Zhen-Yuan Lin (Gingras lab). Their technical expertise, guidance, and patience were remarkable.

Last (but not least) I would like to thank my family and friends for their support and encouragement. I would especially like to acknowledge those that eventually stopped asking the following questions: “when are you finishing school?” and “how long have you been in school for?” I would like to thank Nicole and Chazz for putting up with my absence on many evenings and weekends. Love you guys.

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Table of Contents

Acknowledgements ...... iii

Table of Contents ...... iv

List of Tables ...... ix

List of Figures ...... x

List of Appendices ...... xii

List of Abbreviations ...... 1

Chapter I: Introduction

1.1 Statement of contributions, rights, and permissions ...... 4

1.2 Damage to the DNA double helix ...... 5

1.3 DNA double-strand breaks and how they arise ...... 5

1.3.1 Endogenous sources of DNA double-strand breaks ...... 7

1.3.2 Exogenous sources of DNA double-strand breaks ...... 8

1.3.3 Developmentally programmed DNA double-strand breaks ...... 9

1.4 The cellular response to DNA double-strand breaks ...... 10

1.4.1 DNA damage-induced signal transduction by ATM, DNA-PKcs, and ATR ...... 11

1.4.2 DNA double-strand break-induced cell cycle checkpoints ...... 13

1.4.3 DNA double-strand break-induced senescence and apoptosis ...... 17

1.5 Repair of DNA double-strand breaks ...... 18

1.5.1 Non-homologous end joining ...... 18

1.5.2 Mitotic homologous recombination ...... 21

1.5.3 DNA end resection is the rate-limiting step of homologous recombination ...... 27

1.5.4 Cell cycle-regulated suppression of homologous recombination in G1 cells downstream of end resection ...... 29

1.6 Post-translational modifications and the regulation of DNA double-strand break repair 31

1.6.1 Ubiquitin-dependent signaling at DNA double-strand breaks ...... 31

1.6.2 SUMOylation at DNA double-strand breaks ...... 32

1.6.3 Protein acetylation at DNA double-strand breaks ...... 32

1.6.4 PARylation is one of the earliest protein modifications detected at DNA double-strand breaks ...... 33

1.7 Relevance of DSB repair to human physiology ...... 33

1.7.1 DSB repair promotes tumour suppression ...... 34

1.7.2 DSB repair is important during cerebral cortical development ...... 34

1.8 Rationale and research objective ...... 37

1.9 High-throughput functional discovery utilizing RNA interference screens ...... 37

Chapter II: Genome-scale siRNA screen for regulators of DNA end resection

2.1 Statement of contributions, rights, and permissions ...... 41

2.2 Summary ...... 42

2.3 Introduction ...... 43

2.4 Results ...... 44

2.4.1 Establishment of immunofluorescence-based assays to monitor DNA end resection ...... 44

2.4.2 Quantitative image-based cytometry to monitor DNA end resection ...... 48

2.4.3 Automating DNA end resection assays using liquid-handling robotics ...... 49

2.4.4 Genome-scale siRNA screen utilizing a pooled siRNA library ...... 52

2.4.5 Secondary confirmation screen utilizing cherry-picked siRNA pools ...... 55

2.4.6 Re-screening deconvolved siRNAs for the top resection activator candidates .... 57

2.5 Discussion ...... 59

Chapter III: The zinc finger protein, ZNF335, promotes DNA end resection

3.1 Statement of contributions, rights, and permissions ...... 65

3.2 Summary ...... 66

3.3 Introduction ...... 67 v

3.4 Results ...... 71

3.4.1 Analysis of four siRNA duplexes targeting the ZNF335 messenger RNA ...... 71

3.4.2 ZNF335 depleted cells have defective pRPA32 (S4/S8), RPA32, and BrdU focus formation ...... 73

3.4.3 U2OS cells depleted of ZNF335 are sensitive to DSB-inducing agents ...... 73

3.4.4 Depletion of ZNF335 decreases the efficiency of DSB repair by HR ...... 75

3.4.5 ZNF335 promotes the phosphorylation of CHK1 at a characterized ATR consensus site ...... 78

3.4.6 Defective end resection in ZNF335 depleted cells can be rescued by expressing an RNAi-resistant open reading frame ...... 80

3.4.7 The function of ZNF335 in DNA end resection is dependent on its four C- terminal C2H2 zinc finger domains ...... 80

3.4.8 ZNF335 localizes to sites of DNA damage generated by laser microirradiation . 83

3.4.9 ZNF335 depletion does not affect the expression of the core end resection activators ...... 87

3.4.10 Immunoprecipitation coupled to mass spectrometry (IP-MS) identifies ZNF335 candidate interaction partners ...... 90

3.5 Discussion ...... 93

Chapter IV: Solid-phase transfections in arrayed drops (SPIDR): a cell microarray- based platform for high-content screening

4.1 Statement of contributions, rights, and permissions ...... 98

4.2 Summary ...... 99

4.3 Introduction ...... 100

4.4 Results ...... 101

4.4.1 Design and fabrication of cell microarrays ...... 101

4.4.2 Establishment of solid-phase siRNA transfections in human cultured cells ...... 101

4.4.3 Testing for cell and siRNA cross contamination between samples ...... 104

4.4.4 Solid-Phase transfections In arrayed DRops (SPIDR): a novel method to prevent sample cross contamination on cell microarrays ...... 109

4.4.5 Pilot RNAi screen utilizing the SPIDR platform ...... 109 vi

4.5 Discussion ...... 111

Chapter V: Future directions

5.1 Validation of candidate end resection activators ...... 116

5.2 Validation of candidate end resection inhibitors ...... 117

5.3 Elucidating the mechanism by which ZNF335 promotes end resection ...... 118

5.3.1 Generation of a ZNF335 knockout cell line by genome editing ...... 118

5.3.2 ZNF335 recruitment to DSB sites ...... 118

5.3.3 Investigation of end resection factor recruitment to DSB sites ...... 119

5.3.4 Candidate ZNF335 protein interaction partners ...... 119

5.4 ZNF335 and microcephaly ...... 120

Chapter VI: Materials and methods

6.1 Tissue culture ...... 122

6.1.1 Cell lines ...... 122

6.1.2 RNA interference ...... 122

6.1.3 Generating stable cell lines by lentiviral transduction ...... 122

6.2 Fluorescence microscopy ...... 124

6.2.1 Immunofluorescence ...... 124

6.2.2 Quantitative image-based cytometry ...... 125

6.2.3 Laser microirradiation ...... 125

6.3 Automated genome-scale RNAi screen ...... 127

6.4 Reverse transcription and quantitative PCR ...... 127

6.5 Western blot analysis ...... 128

6.5.1 Whole cell extract preparation ...... 128

6.5.2 SDS-PAGE and immunoblotting ...... 128

6.6 Cell cycle analysis by flow cytometry ...... 128

6.7 Clonogenic survival assays ...... 129 vii

6.8 Homologous recombination assay utilizing the DR-GFP reporter system ...... 129

6.9 Plasmids ...... 130

6.10 Immunoprecipitation coupled to mass spectrometry ...... 130

6.11 Solid-phase reverse siRNA transfections on 1536-well cell microarrays ...... 131

6.12 Solid-phase transfections in arrayed drops (SPIDR) ...... 132

Chapter VII: References……………………………………………………………………..132

Appendix I: Genome-scale RNAi screen data……………………………………………….165

Appendix II: Secondary confirmation screen data………………………………………….167

Appendix III: 53BP1 focus formation SPIDR pilot screen data……………………………169

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List of Tables

Table 1.1. Clinical features of some genome instability syndromes associated with defective DSB repair……………………………………………………………………………………….34 Table 2.1. Number of candidate resection activator deconvolved siRNAs that decreased DNA end resection in S and G2 phase U2OS cells……………………………………………….……60 Table 3.1. DSB repair proteins harbouring zinc finger domains………………………………..68 Table 3.2. Candidate protein interaction partners for full-length ZNF335………………...…...90 Table 3.3. Candidate protein interaction partners for ZNF335 Δ1-1014………………………..91 Table 6.1. List of siRNA duplexes used in this study………………………………………….122 Table 6.2. List of primary antibodies used in this study……………………………………….125

List of Figures

Figure 1.1. Mechanisms of DNA double-strand break formation………………………………..5 Figure 1.2. Activation of DNA damage responsive kinases at DNA double-strand breaks…….11 Figure 1.3. Cell cycle checkpoints activated in response to DNA double-strand breaks……….14 Figure 1.4. DNA double-strand break repair by non-homologous end joining…………………18 Figure 1.5. DNA double-strand break repair by mitotic homologous recombination…………..22 Figure 1.6. The regulation of DNA double-strand break repair pathway choice……………….29 Figure 2.1. DNA end resection assay monitoring RPA32 focus formation…………………….44 Figure 2.2. DNA end resection assay monitoring pRPA32 (S4/S8) focus formation…………..45 Figure 2.3. DNA end resection assay monitoring BrdU focus formation………………………46 Figure 2.4. Measuring end resection by quantitative image-based cytometry………………….49 Figure 2.5. Application of the Kolmogorov-Smirnov test to analyze end resection……………50 Figure 2.6. Automating DNA end resection assays using liquid-handling robotics……………52 Figure 2.7. Genome-scale siRNA screen for regulators of DNA end resection………………...53 Figure 2.8. Pathway enrichment analysis for candidate resection activators…………………...55 Figure 2.9. Secondary confirmation screen utilizing the fluorescent ubiquitylation-based cell cycle indicator (FUCCI) system………………………………………………………………....57 Figure 2.10. Re-screening deconvolved siRNA pools for top candidate resection activators….59 Figure 3.1. Knockdown efficiency, growth, and cell cycle position analysis for siRNA duplexes targeting ZNF335 messenger RNA……………………………………………………………...71 Figure 3.2. ZNF335 is an activator of DNA end resection……………………………………...73 Figure 3.3. ZNF335 deficient cells are sensitive to DSB-inducing agents……………………...75 Figure 3.4. ZNF335 promotes DSB repair by HR………………………………………………76 Figure 3.5. ZNF335 promotes CHK1 serine 345 phosphorylation……………………………...78 Figure 3.6. Expression of siRNA-resistant ZNF335 rescues the observed end resection defect in ZNF335 depleted cells…………………………………………………………………………...80 Figure 3.7. The four C-terminal C2H2 zinc finger domains of ZNF335 are sufficient for its function in end resection…………………………………………………………………………81 Figure 3.8. ZNF335 accumulates at sites of laser microirradiation……………………………..84 Figure 3.9. The accumulation of ZNF335 at laser stripes is PARP-dependent………………...85

Figure 3.10. The PARP-dependent accumulation of ZNF335 at sites of microirradiation is not required for its function in DNA end resection………………………………………………....87 Figure 3.11. ZNF335 depletion does not affect the expression or protein stability of the core end resection activators……………………………………………………………………………...88 Figure 4.1. Formats for high-content screening………………………………………………..101 Figure 4.2. Solid-phase reverse siRNA transfections in HeLa cells…………………………...102 Figure 4.3. Cell migration between samples is minimal on 1536- and 3456-well cell microarrays……………………………………………………………………………………..104 Figure 4.4. Cross contamination of siRNA complexes is evident on cell microarrays………..105 Figure 4.5. Pre-soaking cell microarrays before cell flooding decreases knockdown efficiency……………………………………………………………………………………….107 Figure 4.6. Solid-phase transfections in arrayed drops (SPIDR)……………………………..109 Figure 4.7. High-content siRNA screen utilizing the SPIDR platform………………………..111

List of Appendices

All appendices are presented in an electronic format and are written on the included DVD.

Appendix I: Genome-scale RNAi screen data……………………………………………….165

Appendix II: Secondary confirmation screen data………………………………………….167

Appendix III: 53BP1 focus formation SPIDR pilot screen data……………………………169

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List of Abbreviations

DNA deoxyribonucleic acid RNA ribonucleic acid mRNA messenger RNA DSB double-strand break SSB single-strand break ssDNA single-stranded DNA ROS reactive oxygen species HR homologous recombination CPT camptothecin IR ionizing radiation NCS neocarzinostatin ETOP etoposide HU hydroxyurea AID activation-induced deaminase NHEJ non-homologous end joining PIKK phosphatidylinositol 3-kinase-related kinase MMEJ microhomology-mediated end joining D-loop displacement loop SDSA synthesis-dependent strand annealing dHJ double Holliday junction SSA single-strand annealing LOH loss of heterozygosity PARP poly(ADP-ribose) polymerase PAR poly(ADP-ribose) PARG poly (ADP-ribose) glycohydrolase A-T ataxia-telangiectasia NBS Nijmegen breakage syndrome RNAi RNA interference siRNA small interfering RNA esiRNA endoribonuclease-prepared siRNA shRNA short hairpin RNA BrdU bromodeoxyuridine SEM standard error of the mean QIBC quantitative image-based cytometry KS Kolmogorov–Smirnov test IPA Ingenuity pathway analysis FUCCI fluorescence ubiquitination cell cycle indicator GESS genome-wide enrichment of seed sequences lncRNA long non-coding RNA

IP immunoprecipitation MS mass spectrometry PI propidium iodide PE plating efficiency SF surviving fraction GFP green fluorescent protein RFP red fluorescent protein DR-GFP direct repeat GFP ORF open reading frame PCR polymerase chain reaction NLS nuclear localization signal 4-OHT 4-hydroxytamoxifen DMSO dimethyl sulfoxide DOX doxycycline SPIDR solid-phase transfections in arrayed drops SBS Society for Biomolecular Screening Gy gray DAPI 4',6-diamidino-2-phenylindole CV coefficient of variation FDR false discovery rate ChIP chromatin immunoprecipitation IB immunoblotting IF immunofluorescence

Chapter I

Introduction

1.1 Statement of contributions, rights, and permissions

There are no statements to report regarding contributions, rights, and permissions.

1.2 Damage to the DNA double helix

A cells genetic material is continually altered through reacting with molecules present in the normal cellular environment. DNA damage can take many forms and it is estimated that replicating cells encounter approximately 106 DNA lesions per cell cycle (Lindahl and Barnes, 2000). Many types of DNA damage can be encountered, including base and sugar damage as well as breaks on one or both strands of the sugar-phosphate backbone. These DNA lesions can have severe effects on cellular fitness by disrupting genomic processes like transcription and replication. Furthermore, DNA damage that is not properly repaired can result in mutation and even genome rearrangements, leading to cell death or neoplastic transformation. Genome integrity is maintained by recognition, signaling, and subsequent repair of DNA damage, which reverse the deleterious consequences of DNA lesions and inhibit their transmission to daughter cells (Hoeijmakers, 2001). These signaling-based DNA damage responses have the ability to activate cell cycle checkpoints, coordinate DNA repair, regulate gene expression, and induce apoptosis if the damage load is too high (Jackson and Bartek, 2009).

1.3 DNA double-strand breaks and how they arise

The research presented in this thesis will center on the cellular response to one particular type of DNA lesion, the DNA double-strand break (DSB). DSBs are generated when the two complementary strands of DNA are severed in close proximity (within 10 base pairs) such that the remaining base-pairing and chromatin structure are no longer able to keep the broken strands together (Fig. 1.1A). Indeed, DSBs are often a consequence of two single-strand breaks (SSBs) that are on opposite strands and in close proximity to each other. Dividing cells encounter 10-50 DSBs during each cell cycle, which is substantially less than the frequency of other DNA lesions (Lieber, 2010). However, DSBs are the most deleterious type of DNA lesion because they do not leave an intact complementary strand to be used as a template to restore lost or damaged nucleotides. Studies in budding yeast have demonstrated that a single unrepaired DSB can lead to permanent cell cycle arrest and subsequent programmed cell death (Bennett et al., 1993). DSBs are also required for the formation of gross chromosomal rearrangements that can lead to amplifications, deletions, and gene fusions. These genomic events have been demonstrated to cause malignant transformation in several cancer types (van Gent et al., 2001).

Figure 1.1. Mechanisms of DNA double-strand break formation.

(A) “Two-ended” DSB that has formed as a consequence of close proximity SSBs on opposite DNA strands. Therefore, two-ended DSBs can have small overhang structures at both DNA ends. (B) A leading strand encounters a bulky DNA lesion (i.e. alkylated base) during replication. The DNA lesion will stall the replication fork which can be converted by a nuclease into a “one-ended” DSB. (C) If the leading strand collides with a SSB a “one-ended” DSB will form without the requirement of a nuclease. (D) Topoisomerase and topoisomerase-like (i.e. SPO11- TOPOVIB) enzymes can generate DSBs where the enzyme is covalently linked to the DSB end. (E) Chromosome ends (telomeres) can be recognized as DSBs if not properly protected by the Shelterin complex. Therefore, Shelterin dysfunction can be considered an endogenous source of DSB formation.

1.3.1 Endogenous sources of DNA double-strand breaks

1.3.1.1 Reactive oxygen species

DSBs can arise as a consequence of reactive oxygen species (ROS) that are generated during normal aerobic metabolism. Oxygen is an essential element for energy production but is also dangerous because of the high susceptibility of DNA to attack by ROS, an observation referred to as the “oxygen paradox” (Davies, 1995). A common by-product of several metabolic reactions (including mitochondrial respiration) is the highly reactive hydroxyl free radical. The two main modes of attack to the DNA molecule by a hydroxyl radical is the addition to a double bond or the abstraction of a hydrogen atom from either a DNA base or a deoxyribose sugar (Davies, 1995). Oxidation of the deoxyribose sugar can lead to strand breaks in the sugar-phosphate backbone of DNA and if two of these breaks occur on opposite strands and in close proximity a DSB can form.

1.3.1.2 DNA replication

Another predominant endogenous source of DSBs occurs during S-phase when replication forks encounter DNA lesions. Replicative DNA polymerases, which carry out the bulk of DNA synthesis, are unable to use damaged DNA strands as a template and, consequently, are stopped at most DNA lesions. A stalled replication fork can be re-established by nuclease-mediated conversion into a one-ended DSB which is also called a collapsed fork (Fig. 1.1B) (Fricke and Brill, 2003; Kaliraman et al., 2001). In addition, collapsed replication forks can form when a SSB on the leading strand is encountered during DNA synthesis (Fig. 1.1C). One-ended DSBs are repaired by a DSB repair pathway called homologous recombination (HR) which will be the focus of section 1.6.3.

1.3.1.3 Topoisomerase action

Topoisomerase enzymes create strand breaks to alleviate supercoiled DNA topologies and facilitate genomic processes including replication, transcription, and DNA repair (Vos et al., 2011; Wang, 2002). Topoisomerases can be assigned as either type I or II, depending on their cleavage of one or two strands of DNA, respectively (Liu et al., 1979, 1980; Poccia et al., 1978). DNA cleavage is linked to the formation of a transient but covalent enzyme-DNA adduct at the break terminus (Fig. 1.1D) (Lynn and Wang, 1989). Topoisomerase-DNA linkage prevents the

release of nicked or broken DNA strands before the enzyme can ligate the ends. However, topoisomerase I cleavage in the vicinity of a SSB can induce DSB formation (Jaxel et al., 1988). Moreover, abortive topoisomerase I activity can lead to one-ended DSB formation if encountered by a replication fork. Abortive topoisomerase II activity after cleavage of both DNA strands and before ligation can directly result in a DSB (Fig. 1.1D) (Brown et al., 1979). Remarkably, a class of chemotherapeutic drugs exploits abortive topoisomerase activity by binding to the enzyme, stabilizing it on cleaved DNA ends, and inhibiting ligation. For example, the topoisomerase type I poison, camptothecin (CPT), binds to the enzyme and after cleavage it intercalates within DNA ends to inhibit ligation (Hsiang et al., 1985; Staker et al., 2002).

1.3.1.4 Telomere dysfunction

Telomeres are repetitive DNA sequences present at the ends of eukaryotic chromosomes and are bound by components of the Shelterin complex (de Lange, 2005). Shelterin protects telomeres from being identified as DSB ends and therefore dysfunction of this complex can lead to aberrant DSB recognition by the cell (Fig. 1.1E) (Karlseder et al., 1999). Furthermore, telomeres shorten every cell division cycle as a consequence of normal replication at the end of chromosomes (Harley et al., 1990; Makarov et al., 1997). Telomere shortening can eventually eliminate telomeric repeats, ablating Shelterin binding and enabling the cell to recognize the chromosome end as a DSB (Harley et al., 1990; Karlseder et al., 2002). Therefore, as a population of cells continues to divide, there will be a greater risk of telomeres being recognized as DSBs.

1.3.2 Exogenous sources of DNA double-strand breaks

DSBs can also arise due to the action of exogenous sources including ionizing radiation (IR), heavy metals, and radiomimetic chemical clastogens such as bleomycin and neocarzinostatin (NCS) (Povirk, 1996; Ward, 1988). These agents can react with water within cells to form hydroxyl free radicals. DSBs occur as a consequence of oxidative attack on the sugar-phosphate backbone of DNA. Chemical inhibitors of topoisomerase enzymes including CPT and etoposide (ETOP) are also potent inducers of DSBs (Hsiang et al., 1985; Minocha and Long, 1984). Finally, exogenous chemicals that cause replication stress can induce the formation of one-ended DSBs. For example, the ribonuclease reductase inhibitor, hydroxyurea (HU), depletes the free pool of deoxyribonucleotides used for DNA synthesis, resulting in replication fork stalling

(Krakoff et al., 1968). Stalled replication forks can eventually be processed by nucleases into DSBs (Fricke and Brill, 2003; Kaliraman et al., 2001).

1.3.3 Developmentally programmed DNA double-strand breaks

1.3.3.1 Formation of DNA double-strand breaks in meiosis

Paradoxically, DSBs are not always a deleterious genomic event, and can be formed in a programmed manner to promote several developmental processes required for normal organism physiology. For instance, DSBs are generated in cells of reproductive organs during meiosis I and increases genome diversity in offspring and is required for proper gametogenesis (Kolodkin et al., 1986; Romanienko and Camerini-Otero, 2000). The second meiotic division is similar to mitosis in that it separates the centromeres of sister chromatids, whereas the first meiotic division separates homologous maternal and paternal chromosomes. Meiosis I poses a challenge because homologous chromosomes, unlike sister chromatids, are not necessarily in close proximity. Homologs must locate each other and ‘pair up’ before segregation. Direct association between homologous chromosomes promotes each spindle pole body to attach to one homolog, so that each daughter cell receives only one copy of every chromosome. Therefore, defects in homologous chromosome pairing are associated with aneuploidy and aberrant gametogenesis (Sherman et al., 1994). Homologous chromosome pairing is initiated by developmentally programmed DSBs that are generated at ‘hot spots’ across the length of each chromosome in early meiosis I (Kolodkin et al., 1986). DSBs are generated by the SPO11-TOPOVIBL complex that functions as a heterotetramer and introduces coordinated single nicks on opposite strands leading to a covalent protein-DNA intermediate (Fig. 1.1D) (Keeney et al., 1997; Liu et al., 1995; Robert et al., 2016; Vrielynck et al., 2016). DSBs trigger recombination between homologous chromosomes, which not only keep them physically connected but also promote genetic diversity by permitting the exchange of genetic information between maternal and paternal alleles. Not surprisingly, SPO11-null mice are defective in meiotic recombination and have a severe deficiency in gametogenesis that results in infertility (Romanienko and Camerini- Otero, 2000). The repair of meiotic DSBs must be completed before chromosome segregation and this process is the focus section 1.6.3.5.

1.3.3.2 Formation of DNA double-strand breaks in developing lymphocytes

Programmed DSBs also occur in developing B- and T-lymphocytes to help promote immunoglobulin and T-cell receptor diversity (Bassing and Alt, 2004). Sequence variability at immunoglobulin and T-cell receptor loci is critical for the recognition of diverse pathogens by the adaptive immune response. Immunoglobulin and T-cell receptor proteins contain variable regions that specify antigen binding (Edelman et al., 1969). Through a process termed V(D)J recombination, exons that encode variable regions contain variable (V), diversity (D), and joining (J) segments that can be combined in different ways to generate mature immunoglobulin and T-cell receptor genes (Hesse et al., 1987; Titani et al., 1965). Each segment is flanked by signal sequences that are recognized by the RAG1-RAG2 nuclease, which generates DSBs (Oettinger et al., 1990; Schatz et al., 1989). RAG1-RAG2-generated DSBs are recognized and repaired by a specific DSB repair pathway called non-homologous end joining (NHEJ; introduced in section 1.5.1) (Taccioli et al., 1994).

DSBs are also generated in developing B-lymphocytes to trigger a process called class- switch recombination. During B-cell differentiation, class-switch recombination can fuse different immunoglobulin constant regions to a specific variable region (Kataoka et al., 1980; Nossal et al., 1971). For example, after recognition of an epitope, B-cells mature by changing their immunoglobulin constant domain from membrane-bound to soluble. Differential constant domains also allow the immunoglobulin to interact with a variety of effector molecules and promote an effective adaptive immune response. During class-switch recombination, DNA strand breaks are generated by the concerted action of activation-induced deaminase (AID) and transcription in the immunoglobulin switch regions (Muramatsu et al., 2000). AID triggers the deamination of cytosine to uracil resulting in U-G mismatches that are processed by nucleases to yield DSBs. NHEJ or alternative end-joining (the focus of section 1.5.1 and 1.5.2, respectively) can ligate variable exons to specific constant exons during class-switch recombination (Casellas et al., 1998; Manis et al., 2002; Manis et al., 2004; Ward et al., 2004; Yan et al., 2007).

1.4 The cellular response to DNA double-strand breaks

The response to DSBs is a classical signal transduction pathway in which a signal, in this case a DSB, is first detected by sensor proteins and then transduced to downstream effectors. The focus

of this section will be on the mechanisms of DSB end detection and how this initiates signal transduction. I will outline the targets of DSB-induced signal transduction activity and how these effector proteins modulate cell cycle checkpoints, senescence, and apoptosis.

1.4.1 DNA damage-induced signal transduction by ATM, DNA-PKcs, and ATR

The critical signal transducers in the DSB response are three phosphatidylinositol-3 kinase-like kinases (PIKKs): ATM, DNA-PKcs, and ATR. Upon activation, they phosphorylate a vast set of proteins on serine or threonine residues present within serine/threonine-glutamine (S/T-Q) consensus motifs (Kim et al., 1999). Phosphoproteomic studies have identified hundreds of putative PIKK targets that are phosphorylated in response to DNA damage (Matsuoka et al., 2007; Mu et al., 2007; Roitinger et al., 2015; Smolka et al., 2007; Stokes and Comb, 2008). Although all three kinases have some overlapping substrates, each PIKK also has distinct functions in the response to DNA damage.

1.4.1.1 ATM activation

In response to DSBs, ATM is recruited to and activated by the DSB sensor complex MRE11- RAD50-NBS1 (MRN), a multifunctional complex that is critical for the early stages of the DSB response (Fig. 1.2A) (Carson et al., 2003; Nakada et al., 2003; Uziel et al., 2003). ATM autophosphorylation results in its dissociation from inactive dimers into active monomers that can bind to damaged chromatin (Bakkenist and Kastan, 2003). Furthermore, ATM is acetylated by TIP60 which further stimulates its activation (Kaidi and Jackson, 2013; Sun et al., 2005). The histone H2A variant H2AX is a particularly important substrate for ATM which phosphorylates it on the conserved C-terminal serine 139 residue (Burma et al., 2001; Downs et al., 2000; Rogakou et al., 1998). Phosphorylated H2AX (also called γH2AX) marks a chromatin domain that is recognized by the checkpoint mediator protein MDC1 (Stucki et al., 2005). In a positive- feedback loop, MDC1 promotes further ATM activation through interactions with NBS1 which enhances the accumulation of the MRN complex and of activated ATM on damaged chromatin (Chapman and Jackson, 2008; Melander et al., 2008; Spycher et al., 2008; Wu et al., 2008a).

1.4.1.2 DNA-PKcs activation

The KU70/80 heterodimer is a DSB end sensor complex that recruits and actives DNA-PKcs

Figure 1.2. Activation of DNA damage responsive kinases at DNA double-strand breaks.

(A) ATM activation. The DSB is first detected by the MRN complex which can directly bind to DNA ends. ATM is recruited to DSB sites by the MRN complex which results in its dissociation from inactive dimers to active monomers. ATM acetylation by TIP60 is also important for its activation. (B) DNA-PKcs activation. The DSB ends are first detected by KU70/80. DNA-PKcs is recruited to DSB sites through a direct association with KU70/80. The presence of DNA ends and KU70/80 stimulates DNA-PKcs activation. Two DNA-PKcs molecules can dimerize across a DSB to tether the broken ends together in a so-called ‘synaptic complex’ which holds the broken ends in close proximity and enhances re-joining. DNA-PKcs dimerization also stimulates its kinase activity. (C) ATR responds to the accumulation of ssDNA in the genome which can arise as a consequence of replication stress or DSB end resection during HR. First, ATR is recruited to RPA bound ssDNA through an interaction with ATRIP. The checkpoint clamp 9-1-1 and TOPBP1 are also required for ATR activation.

(Gottlieb and Jackson, 1993; Hartley et al., 1995). The complex formed at the break site consisting of DNA, KU70/80, and DNA-PKcs is referred to as simply “DNA-PK” (Fig. 1.2B). Once bound to KU70/80, the catalytic activity of DNA-PKcs is activated. DNA-PKcs kinase activity is essential for DSB repair by NHEJ but the exact mechanism for why this is the case is currently unknown. DNA-PK has been shown to phosphorylate core NHEJ factors such as KU70/80, XRCC4, and XLF, but surprisingly these phosphorylation events are not required for NHEJ (Douglas et al., 2005; Yu et al., 2008; Yu et al., 2003). The most important NHEJ DNA- PK target appears to be DNA-PKcs itself. In response to DSBs, upwards of 40 autophosphorylation sites have been documented for DNA-PKcs (Davis et al., 2014). One critical autophosphorylation site appears to be serine 2056 as ablating this site resulted in less efficient NHEJ (Chen et al., 2005).

1.4.1.3 ATR activation

ATR is activated in response to replication stress, when stalled forks result in the accumulation of ssDNA bound by the ssDNA binding heterotrimeric replication protein A (RPA14-RPA32- RPA70) complex (Zou and Elledge, 2003). ATR recognition of RPA-ssDNA is carried out by an ATR-associated factor called ATRIP (Cortez et al., 2001; Zou and Elledge, 2003). However, ATR/ATRIP localization to RPA-ssDNA is not sufficient for kinase activation. ATR activation also requires the RAD9-RAD1-HUS1 (or 9-1-1) complex and TOPBP1 (Delacroix et al., 2007; Lee et al., 2007; Majka et al., 2006; Roos-Mattjus et al., 2002). DSBs formed in the S- and G2- phases of the cell cycle can be repaired by HR. The first step of HR involves the generation of ssDNA overhangs at DSB ends through a process called DNA end resection. RPA binds ssDNA formed as a consequence of end resection providing another platform for ATR activation (Fig. 1.2C). Therefore, ATR is activated as a consequence of both replication stress and DSBs in S and G2 cells.

1.4.2 DNA double-strand break-induced cell cycle checkpoints

Activated ATM and ATR kinases are critical for transducing signals to effector proteins that halt cell cycle progression (Liu et al., 2000; Matsuoka et al., 1998). In contrast, DNA-PKcs appears to play a less pivotal role in the activation of DSB-induced cell cycle checkpoints (Burma et al., 1999; Jhappan et al., 2000). During the response to DSBs, it is paramount for the cell to arrest the cell cycle and provide time for repair machineries to re-join the breaks before the start of

processes like replication or mitosis. The replication or segregation of broken chromosomes can have deleterious outcomes for the cell, including the generation of daughter cells with aneuploidy and genome rearrangements (Cahill et al., 1998; Chan et al., 1999). DNA damage- induced checkpoints occur at entry into S-phase (the G1/S checkpoint), within S-phase (the S- phase checkpoint), and entry into mitosis (the G2/M checkpoint).

1.4.2.1 G1/S checkpoint

The G1/S checkpoint is triggered by ATM-dependent signaling and subsequent enrichment of MDC1 on damaged chromatin in G1 cells (Goldberg et al., 2003; Lou et al., 2003; Stewart et al., 2003). Although the precise mechanism of how MDC1 promotes checkpoint activation remains unclear, it appears to be at the level of enhancing ATM activity (Lou et al., 2006; Mochan et al., 2003). ATM targets that are required for G1/S checkpoint activation include the transcription factor p53 and the checkpoint kinases CHK1 and CHK2 (Fig. 1.3A) (Canman et al., 1998; Chen et al., 1999; Gatei et al., 2003; Matsuoka et al., 1998). P53 is stabilized upon phosphorylation on serine 15 by ATM, as it can no longer be inhibited by MDM2 (Canman et al., 1998; Dulic et al., 1994; Shieh et al., 1997). P53 then activates the transcription of p21 which is a potent inhibitor of the G1/S-promoting cyclin E/cyclin-dependent kinase 2 (CDK2) complex and therefore, induces a G1 arrest (Harper et al., 1993; Xiong et al., 1993). ATM also phosphorylates and activates CHK1 and CHK2 in G1 cells which in turn can phosphorylate the phosphatase CDC25A (Falck et al., 2001; Mailand et al., 2000). Phosphorylation of CDC25A induces its ubiquitylation by SCFβ-TRCP and subsequent degradation in the proteasome, potentiating the phosphorylation of CDK2 at threonine 14 and 15 (Busino et al., 2003; Costanzo et al., 2000; Jin et al., 2003; Mailand et al., 2000). Phosphorylated CDK2 is unable to promote DNA synthesis and entry into S-phase, thereby causing a G1 arrest. In addition, activated CHK2 can also phosphorylate p53 to further enhance p21 expression and the G1/S checkpoint (Hirao et al., 2000).

1.4.2.2 S-phase checkpoint

Eukaryotes replicate their genomes from multiple origins that are distributed across each chromosome. Origins of replication are activated throughout S-phase of the cell cycle such that some origins fire early and others fire late. In response to DNA damage during S-phase, cells

Figure 1.3. Cell cycle checkpoints activated in response to DNA double-strand breaks.

ATM and ATR kinases are activated in response to DSBs and transduce signals to effector proteins that halt cell cycle progression. In response to DNA damage, the cell cycle can be arrested at various positions including at the G1/S transition (G1/S checkpoint), within S-phase (the S-phase checkpoint), and at the G2/M transition (the G/M checkpoint). Interestingly, the mechanisms that control cell cycle arrest are different for the three checkpoints. (A) The G1/S checkpoint relies on the inhibition of the S-phase-promoting cyclin E/CDK2 kinase by either p21 or through the inactivation of the CDC25A phosphatase. (B) The S-phase checkpoint relies on the inhibition of Treslin by CHK1 and CHK2. Treslin promotes origin firing during S-phase by directly interacting with TOPBP1 and CDC45. In response to DNA damage, CHK1/2-mediated phosphorylation inhibits the function of Treslin in promoting DNA replication. (C) The G2/M checkpoint relies on the inhibition of the mitosis-promoting cyclin B/CDK1 kinase by the inactivation of the CDC25C phosphatase. Various mechanisms have been described for inhibiting CDC25C in G2 cells exposed to agents that induce DSBs (see main text). Initiation of the G2/M checkpoint does not dependent on p53 but a slower transcriptional program controlled by p53 is required to maintain the checkpoint.

activate a checkpoint that can inhibit later firing origins of replication that have not yet been initiated. The S-phase checkpoint is best understood in the budding yeast Saccharomyces cerevisiae and is controlled by Mec1 (ATR in humans) and the checkpoint kinase Rad53 (CHK2 in humans) (Santocanale and Diffley, 1998; Shirahige et al., 1998). In response to DNA damage, Sld3 (Treslin/TICRR in humans) is phosphorylated by Rad53 which inhibits its function in promoting the firing of late replication origins (Fig. 1.3B) (Boos et al., 2013; Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010). Phosphorylation of Sld3 (Treslin) abrogates its CDK- dependent interaction with the origin firing factors Dpb11 (TOPBP1 in humans) and Cdc45 (Boos et al., 2011; Zegerman and Diffley, 2010).

1.4.2.3 G2/M checkpoint

The G2/M checkpoint prevents G2 cells from entering mitosis when DSBs are present. Similar to cells in S-phase, G2 cells can activate both the ATM and ATR kinases in response to DSBs (Fig. 1.3C). The critical target of the G2/M checkpoint is the mitosis-promoting activity of the cyclin B/CDK1 kinase (Lundgren et al., 1991). The ATM/ATR-dependent activation of CHK1 and CHK2 in G2 cells results in the phosphorylation of the phosphatase CDC25C on serine 216, creating a 14-3-3 binding site (Peng et al., 1997; Sanchez et al., 1997). Binding of 14-3-3 proteins to phosphorylated CDC25C sequesters it in the cytoplasm and inhibits its phosphatase activity towards nuclear cyclin/CDK substrates (Dalal et al., 1999; Yang et al., 1999). Inhibition of the CDC25C phosphatase enables the nuclear accumulation CDK1 that is phosphorylated on tyrosine 15 by WEE1 (Krek and Nigg, 1991; Lundgren et al., 1991; Norbury et al., 1991). WEE1-dependent phosphorylation of CDK1 inhibits its kinase activity and arrests cells in G2. The checkpoint mediator proteins 53BP1, RNF8, BRCA1 are also critical for establishing a robust G2/M checkpoint, likely through the promotion of ATM/ATR and CHK1/CHK2 activity (Fernandez-Capetillo et al., 2002; Huen et al., 2007; Kolas et al., 2007; Mochan et al., 2004; Wang et al., 2002; Yarden et al., 2002). One particular 14-3-3 family member, 14-3-3σ, is expressed in a p53-dependent manner and is required for the maintenance of the G2/M checkpoint (Hermeking et al., 1997). Interestingly, 14-3-3σ, unlike other family members, cannot bind to phosphorylated CDC25C and inhibits entry into mitosis by an alternative mechanism (Chan et al., 1999). In response to DNA damage, 14-3-3σ inhibits mitotic entry by directly sequestering cyclin B/CDK1 in the cytoplasm. The G2/M checkpoint can also be activated in a CHK1/2-independent manner by the p38 and MAPKAP kinase-2 (MK2) kinases (Bulavin et al.,

2001; Manke et al., 2005). In response to DNA damage, p38 and MK2 are activated by ATM/ATR and, like CHK1/2, can control the checkpoint response through the phosphorylation- dependent inhibition of CDC25C. MK2 can directly phosphorylate CDC25C on serine 216 which creates a binding site for 14-3-3 proteins. MK2 can also promote the G2/M checkpoint in the cytoplasm where it controls the posttranscriptional modulation of gene expression (Reinhardt et al., 2010). MK2 can phosphorylate the RNA binding proteins hnRNPA0, TIAR, and PARN, which then bind to and stabilize the messenger RNA of the cell cycle inhibitor gene GADD45α. The maintenance of the G2/M checkpoint relies on a transcriptional program regulated by p53, leading to the upregulation of cell cycle inhibitors including p21, GADD45α, and 14-3-3 proteins (Agarwal et al., 1995; Hermeking et al., 1997; Wang et al., 1999). However, unlike the G1/S checkpoint, cells can initiate a robust G2/M checkpoint in the absence of p53 (Kastan et al., 1991; Levedakou et al., 1995). This observation spurred efforts to therapeutically interfere with the G2/M checkpoint as a potential strategy to sensitize p53-deficient cancer cells to radiation- or chemotherapy-induced DNA damage (McNeely et al., 2014; Russell et al., 1995; Wang et al., 1996; Wang et al., 2001). The most promising therapeutic strategies for inhibiting the G2/M checkpoint in p53-deficient tumours have involved small molecule inhibitors of the ATR, CHK1, and WEE1 kinases.

1.4.3 DNA double-strand break-induced senescence and apoptosis

Cells that experience a degree of DNA damage that is beyond repair can undergo permanent cell cycle arrest (senescence) or programmed cell death. DSB-induced activation of ATM and ATR leads to the phosphorylation and activation of p53 (Canman et al., 1998; Siliciano et al., 1997; Tibbetts et al., 1999). The p53 transcription factor not only mediates the G1/S transient checkpoint but also initiates senescence or apoptotic programs if too many DSBs are present or if their repair is delayed or defective (Di Leonardo et al., 1994; Lowe et al., 1993). The molecular mechanisms that control these p53-dependent cell fate decisions in response to genotoxic stress are largely unknown. It has been suggested that cell fate decisions may be controlled by the level of p53 protein (Batchelor et al., 2011; Batchelor et al., 2008; Loewer et al., 2010). For example, in response to DSBs by ionizing radiation, the levels of p53 exhibit a series of pulses with fixed amplitude and frequency. Higher doses of ionizing radiation increase the number of pulses without affecting their amplitude. Remarkably, precisely timed drug additions that produce a sustained p53 protein pulse can push cells towards senescence and apoptosis rather than a

transient cell cycle arrest (Purvis et al., 2012). Controlling the p53 pulse can have critical therapeutic implications when treating tumours that overexpress the oncogenic p53 inhibitor MDMX (Chen et al., 2016). Tumour cells treated with a MDMX inhibitor initiate a rapid pulse in p53 protein levels followed by low-amplitude oscillations. Remarkably, the exposure of cells with ionizing radiation during the p53 pulse coincides with apoptosis. In stark contrast, programmed cell death is inhibited when cells are treated with ionizing radiation after the pulse, when p53 levels demonstrate low-amplitude oscillations.

1.5 Repair of DNA double-strand breaks

In addition to the modulation of cell cycle progression and apoptosis, the PIKK-mediated cellular response to DSBs also involves the coordination of repair enzymes to promote the re-joining of DSB ends. Eukaryotic cells have evolved two major pathways to re-ligate DSBs: nonhomologous end joining (NHEJ) and homologous recombination (HR).

1.5.1 Non-homologous end joining

The fastest way to re-join a DSB is to ligate it back together utilizing NHEJ (Fig. 1.4). First, DSB ends are detected and bound by the KU70/80 heterodimer (Mimori and Hardin, 1986). Structural studies have shown that KU70/80 forms a ring with a hole that fits double-stranded DNA ends (Walker et al., 2001). It is thought that KU70/80 is the first factor to bind DSBs due to its high nuclear abundance and strong affinity for DNA ends (Mimori and Hardin, 1986). KU70/80 acts as a molecular scaffold for the recruitment of NHEJ effectors including nucleases, polymerases, and at least one ligase. As discussed previously, the extreme C-terminus of KU80 is required for the recruitment of DNA-PKcs (Gell and Jackson, 1999). An interaction between two DNA-PKcs molecules across a DSB can tether the broken ends together in a so-called ‘synaptic complex’ which holds the broken ends in close proximity and inhibits attack by nucleases (DeFazio et al., 2002). DNA-PKcs dimerization across a DSB also stimulates its kinase activity which is required for DSB repair by NHEJ (Kurimasa et al., 1999; Meek et al., 2007). In the final step of NHEJ, the DSB ends are re-joined by the action of DNA ligase IV in complex with XRCC4, XLF, and PAXX (Ahnesorg et al., 2006; Grawunder et al., 1997; Ochi et al., 2015; Wilson et al., 1997). For ligation to occur there must be undamaged nucleotides at both break ends. DSBs generated by ROS often have complex chemical structures and the multi-

Figure 1.4. DNA double-strand break repair by non-homologous end joining.

Repair of DSBs by NHEJ relies on the detection of DSB ends by the KU70/80 heterodimer. The DNA-PKcs kinase is recruited to DSB sites by KU70/80 where it dimerizes across a DSB to form a ‘synaptic complex’. DNA-PKcs dimerization and binding to KU70/80 stimulates its kinase activity. Next, the DSB ends can be processed by enzymes to ensure they are compatible for ligation. The DNA ligase IV complex is responsible for the final re- joining step.

functional nuclease Artemis is responsible for removing damaged nucleotides and secondary structures that may inhibit DSB re-joining (Ma et al., 2002; Moshous et al., 2001). Polynucleotide kinase/phosphatase (PNKP) also plays an important role in promoting DNA ligase IV-dependent joining by ensuring that 5’ DNA termini are phosphorylated and 3’ termini are not (Chappell et al., 2002; Koch et al., 2004). Moreover, ligation is inhibited when adenylate groups are covalently linked to the 5’ termini of breaks. Aprataxin (APTX) catalyzes the nucleophilic release of adenylate groups resulting in termini that can again serve as a substrate for DNA ligase IV (Rass et al., 2007). Several DNA polymerases have important roles in NHEJ. Polymerase µ is a highly versatile enzyme that has both template-dependent and template- independent synthesis capabilities and can add nucleotides to DSB ends (Mahajan et al., 2002a). When the resulting short 3’ overhangs share even one nucleotide of complementarity, ligation by the DNA ligase IV complex is enhanced. Remarkably, polymerase µ, together with KU70/80 and the DNA ligase IV complex, can polymerize across a discontinuous template strand, thereby crossing from one DNA end to another (Nick McElhinny et al., 2005). In addition, polymerase λ only has template-dependent synthesis activity and fills in short gaps that form after annealing of 1-4 base pairs of homology at the DSB end (Garcia-Diaz et al., 2009; Lee et al., 2004). Through a largely unknown mechanism, 53BP1 and its effector proteins RIF1 and MAD2L2 have also been shown to bind near to the sites of DSB ends and promote long-range NHEJ reactions, particularly during V(D)J and class-switch recombination (Boersma et al., 2015; Chapman et al., 2013; Di Virgilio et al., 2013; Difilippantonio et al., 2008; Dimitrova et al., 2008; Escribano- Diaz et al., 2013; Ward et al., 2004; Xu et al., 2015; Zimmermann et al., 2013).

1.5.1.1 Alternative end-joining

DSBs can also be repaired by an alternative end-joining mechanism termed microhomology- mediated end joining (MMEJ). Annealing of 5-25 base pair microhomologous sequences at each of the broken DSB ends is a requirement for MMEJ (Roth and Wilson, 1986). MMEJ can be KU70/80-indepedent but requires the activity of poly(ADP-ribose) polymerase 1 (PARP-1) (Audebert et al., 2004). In order for microhomologies to be exposed for annealing, limited DNA end resection is required to generate short 3’ ssDNA overhangs (Moore and Haber, 1996). Furthermore, small segments of microhomology can be introduced to the DSB ends by the template-independent activity of polymerase θ (Ceccaldi et al., 2015; Mateos-Gomez et al., 2015). Next, the exposed ssDNA homologies at each DSB end can anneal together and the

template-dependent activity of polymerase θ fills in the gaps before ligation. In contrast to canonical NHEJ which employs DNA ligase IV, ligation during MMEJ is conducted by DNA ligase III (Audebert et al., 2004). If microhomologies are present internally at the DSB ends 3’ flaps will be generated and will need to be resolved by nucleases which can result in deletions (Ma et al., 2003). Therefore, MMEJ is a mutagenic DSB repair pathway and the cellular relevance of such an error-prone mechanism still remains unclear. However, it is conceivable that MMEJ may contribute to the stability of small repetitive elements in genomes such as centromeres and telomeres (Capper et al., 2007). In addition, MMEJ is particularly relevant in developing B-cells where it plays an important role in class-switch recombination (Boboila et al., 2012).

1.5.2 Mitotic homologous recombination

NHEJ can re-join DSBs in the G1-, S-, and G2-phases of the cell cycle. In contrast, HR occurs mainly in S and G2 cells, after replication has generated an identical sister chromatid. Indeed, the two sister chromatids, by virtue of being identical copies of each other and for being in close proximity, are the preferred template for HR. As a consequence of this cell cycle-phase preference, HR occurs primarily in dividing cells, whereas terminally differentiated cells rely on NHEJ (Gao et al., 1998). As most mammalian cell types are non-dividing, the majority of DSBs are likely repaired by NHEJ. However, HR is a critical DNA repair process during proliferative stages of development and somatic cell renewal in mammals. HR can re-join DSBs generated in both mitotically and meiotically dividing cells. Although the mechanistic details of mitotic and meiotic HR are similar, some differences do exist and will be highlighted in section 1.6.3.5. In contrast to mammalian cells, HR is the primary DSB repair pathway in the budding yeast Saccharomyces cerevisiae. The mechanistic details of HR were first uncovered in budding yeast but are highly conserved throughout evolution. For this section, I will focus mainly on primary research that was conducted in budding yeast. A description of yeast HR genes will be given but all corresponding human homologs with different names will be supplied in parentheses.

1.5.2.1 DNA end resection initiates homologous recombination

The first step in HR is the detection of DSB ends by the Mre11-Rad50-Xrs2 (MRE11-RAD50- NBS1) or MRX (MRN) complex (Fig. 1.5) (Raymond and Kleckner, 1993). In addition to activating Tel1 (ATM), MRX promotes a process termed DNA end resection that generates 3’

ssDNA tails at each end of the DSB (Moreau et al., 2001; Nairz and Klein, 1997; Paull and Gellert, 1998). End resection is activated during the S- and G2-phases of the cell cycle by CDK- mediated phosphorylation of Sae2 (CtIP) which physically associates with the MRX complex and promotes the nuclease activity of Mre11 (Huertas et al., 2008; Huertas and Jackson, 2009; Sartori et al., 2007). Therefore, in addition to the proximal availability of the sister chromatid, the CDK-dependent initiation of end resection by Sae2 (CtIP) also restricts HR to S and G2 cells. The predicted nuclease activity required for processing DSB ends into 3’ ssDNA overhangs is a 5’-3’ exonuclease. However, Mre11 lacks this activity and is instead a bifunctional nuclease, containing both endonuclease and 3’-5’ exonuclease activity (Paull and Gellert, 1998). It was later determined that Mre11 utilizes both its endonuclease and exonuclease activity to promote end resection (Garcia et al., 2011). First, using its endonuclease activity, Mre11 nicks the strand to be resected up to 300 base pairs from the 5’ terminus of the DSB end. The nick enables end resection in a bidirectional manner, whereby Mre11 exonuclease activity promotes 3’-5’ resection towards the DSB end and the exonucleases Exo1 and Dna2 carry out resection in the 5’-3’ direction away from the DSB end (Garcia et al., 2011; Zhu et al., 2008).

End resection can be separated into two functional steps: short-range (or initiation) and long-range. As Mre11/Sae2 (CtIP) initiate end resection proximal to the DSB end and in the direction towards the break, only short tracts of 3’ ssDNA tails are generated. Therefore, Mre11/Sae2 (CtIP)-dependent 3’ to 5’ processing is termed short-range end resection. The downstream events for HR require long stretches of ssDNA and this is achieved by the exonuclease activity of Exo1 and Dna2, in collaboration with the RecQ helicase Sgs1 (BLM) (Zhu et al., 2008). These nucleases resect DNA in the 5’ to 3’ direction (away from the DSB end) in a process called long-range end resection. In human cells, an additional nuclease, the 3’- 5’ exonuclease EXD2, has been implicated in end resection and can process DSB ends in parallel with MRE11 (Broderick et al., 2016). End resection generates long stretches of ssDNA that are rapidly coated by the RPA complex (Longhese et al., 1994). Through a positive feedback mechanism, the RPA complex stimulates further end resection by Exo1 and Dna2 while also inhibiting the degradation of ssDNA tails and the formation of hairpin structures (Chen et al., 2013).

Figure 1.5. DNA double-strand break repair by mitotic homologous recombination.

Repair of DSBs by HR in mitotically dividing cells primarily occurs in the presence of a sister chromatid (in the S- and G2-phases of the cell cycle). The rate-limiting step for HR is the generation of 3’ ssDNA overhangs at each end of a DSB by DNA end resection. Bi-directional end resection is conducted by nucleases including MRE11, EXO1, and DNA2. The defining step of HR is the loading of the RAD51 recombinase onto ssDNA which facilitates strand invasion and homology search in the sister chromatid.

1.5.2.2 Rad51 assembly and the search for homology

After end resection, RPA bound to ssDNA is exchanged for the Rad51 recombinase (Sugiyama et al., 1997). Rad51 catalyzes the defining step of HR, strand exchange, during which ssDNA invades the sister chromatid, displacing the complementary strand of the duplex to form a displacement loop (D-loop) (Petukhova et al., 2000; Sugawara et al., 1995). However, RPA impedes Rad51 loading to ssDNA and the recombinase needs accessory factors, called recombination mediators. The prototypical recombination mediator is Rad52 which is recruited to resected DSBs through an interaction with RPA (New et al., 1998). Rad52 also interacts with Rad51 to stimulate RPA removal from ssDNA and replacement with Rad51 (Song and Sung, 2000; Stasiak et al., 2000). Rad51 forms a right-handed nucleoprotein filament when loaded onto ssDNA (Ogawa et al., 1993). Formation of filaments is also stimulated by Rad52 (McIlwraith et al., 2000; Sung and Robberson, 1995). In contrast to budding yeast, mammalian cells lacking RAD52 do not display DNA damage sensitivity and only have minor defects in HR (Rijkers et al., 1998). In mammalian cells, the BRCA1-PALB2-BRCA2 complex targets RAD51 to ssDNA and thereby promotes RAD51 to replace RPA and form a nucleoprotein filament (Buisson et al., 2010; Carreira et al., 2009; Dray et al., 2010; Jensen et al., 2010; Moynahan et al., 1999; Sy et al., 2009; Yang et al., 2005). HR is also stimulated by a group of mediator proteins called the RAD51 paralogs. Five RAD51 paralogs have been identified in mammalian species and they interact with one another to form two distinct complexes: RAD51B-RAD51C-RAD51D-XRCC2 (BCDX2) and RAD51C-XRCC3 (CX3) (Masson et al., 2001). Loss of the RAD51 paralogs leads to severe HR defects, DNA damage sensitivity, chromosome abnormalities, and aberrant RAD51 focus formation (French et al., 2002; Godthelp et al., 2002; Johnson et al., 1999; Pierce et al., 1999). The Caenorhabditis elegans homologs of the BCDX2 and CX3 complexes, RFS-1/RIP-1, can bind to RAD51 nucleoprotein filaments to promote their stability and flexible confirmation, which facilitates strand exchange with the sister chromatid (Taylor et al., 2015). Rad51 nucleoprotein filaments also stimulate searching for homologous sequence after strand invasion (Sugawara et al., 1995). If homology is found, repair synthesis by polymerases will replace missing nucleotides at the break site by using the invading sister chromatid as a primer (Holmes and Haber, 1999).

1.5.2.3 Mechanisms for disengaging from the sister chromatid and completing repair

At this step in the pathway, two sub-pathways of HR emerge: synthesis-dependent strand annealing (SDSA) and double Holliday junction (dHJ) formation (Fig. 1.5). The predominant sub-pathway appears to be SDSA, in which the D-loop is ablated, leading to annealing of the newly synthesized strand with the resected strand of the other DSB end (Ira et al., 2006; Kurkulos et al., 1994; Nassif et al., 1994). Gaps are filled in by polymerases and ends are ligated to re-establish the integrity of the chromatid. Another potential outcome when a sister chromatid is invaded by a nucleoprotein filament is that the newly synthesized strand can be captured by the other resected DSB end forming a dHJ (Parsons and West, 1988; West et al., 1983). Two mechanisms exist for processing dHJs: resolution and dissolution. Resolution involves specialized nucleases that cleave dHJs, allowing the strands to pass over each other and anneal to their respective chromatids (Ip et al., 2008). Crossover products, where a strand destined for one chromatid is annealed to the other, may arise as a consequence of resolution. Crossovers have the potential to cause chromosome rearrangements when recombination occurs between non-allelic repetitive sites (i.e. gene paralogs) (Montgomery et al., 1991). In the second mechanism, dissolution, the dHJs are migrated toward each other by the RecQ helicase Sgs1 (BLM) before cleavage (Karow et al., 2000). Junction migration results in a hemicatenane structure that is eliminated by the Sgs1-Top3-Rmi1 (BLM-TOPIIIα-RMI1-RMI2) complex resulting in non- crossover products (Cejka et al., 2010; Chang et al., 2005; Ira et al., 2003; Singh et al., 2008; Wu et al., 2006; Wu and Hickson, 2003; Xu et al., 2008; Yin et al., 2005). HR is rarely associated with crossovers so the preferred HR sub-pathways are likely SDSA and dHJ dissolution as both promote the formation of non-crossover products. An in vitro biochemical study demonstrated that the formation of dHJs is actively blocked by RAD51 (Wu et al., 2008b). RAD51-dependent inhibition of dHJ formation could act as a simple mechanism for promoting SDSA and preventing crossovers during mitotic HR.

1.5.2.4 Single-strand annealing is an alternative homology-based DNA double-strand break repair mechanism

An alternative HR-based DSB repair pathway is called single-strand annealing (SSA). Akin to the alternative end joining pathway MMEJ, SSA also involves the hybridization of homologous regions at each end of a resected DSB (Fishman-Lobell et al., 1992). In contrast to MMEJ, SSA

involves the base pairing of longer homologous regions. Therefore, SSA requires end resection and is mainly active in the S- and G2-phases of the cell cycle (Clerici et al., 2005; Huertas et al., 2008). Moreover, a key difference between MMEJ and SSA is that SSA requires the help of Rad52 for annealing longer regions of ssDNA (Fishman-Lobell et al., 1992; Ivanov et al., 1996). SSA may be important for the repair of larger repetitive regions in the genome such as ribosomal DNA loci (Sfeir and Symington, 2015). Like MMEJ, SSA is also mutagenic because annealing of repetitive sequences that are internal to DSB ends will generate 3’ flap structures that are cleaved by nucleases, resulting in potentially harmful deletions (Fishman-Lobell et al., 1992).

1.5.2.5 Meiotic homologous recombination

SPO11/TOPOVIBL-generated DSBs during meiosis promote the exchange of genetic material between homologous chromosomes and are essential for proper meiotic chromosome segregation (Kolodkin et al., 1986). DSBs formed by SPO11/TOPOVIBL are repaired by HR. The steps of meiotic HR are similar to that of mitotically dividing cells but with several key differences. First, Spo11 forms a covalent protein-DNA linkage at DSB ends which needs to be removed for efficient strand invasion (Keeney et al., 1997). Utilizing their nuclease activities, the short-range end resection machinery composed of the MRX complex and Sae2 (CtIP) can remove Spo11 from DSB ends (Neale et al., 2005). Secondly, two recombinase proteins, Rad51 and its meiosis- specific paralog Dmc1, are required for meiotic HR (Bishop et al., 1992; Shinohara et al., 1992). Both Rad51 and Dmc1 can form nucleoprotein filaments on ssDNA (Gupta et al., 2001; Ogawa et al., 1993). Interestingly, Rad51 mutants proficient at filament formation but defective in strand invasion can effectively complete meiotic HR (Cloud et al., 2012). In contrast, Dmc1 mutants with defective strand invasion activity cannot repair Spo11-generated DSBs. Therefore, Dmc1 appears to be important for the strand invasion step of meiotic HR whereas Rad51 is required for nucleoprotein filament formation. In mammalian cells, loading of RAD51 and DMC1 onto resected DSB ends in meiosis is BRCA2-dependent (Siaud et al., 2004; Thorslund et al., 2007). However, the requirement of BRCA1 and PALB2 in recombinase loading at SPO11/TOPOVIBL-generated DSBs has not been established.

1.5.3 DNA end resection is the rate-limiting step of homologous recombination

1.5.3.1 Control of DNA end resection length dictates recombination efficiency

DNA end resection is the main focus of my doctoral research and I will further describe the current knowledge regarding its importance and regulation in eukaryotic cells. The first essential step for all homology-based repair pathways is the generation of ssDNA by end resection. Long ssDNA tracts at DSB ends are required to form Rad51 nucleoprotein filaments of the appropriate length. When the length of the filament is increased, homology is found faster (Forget and Kowalczykowski, 2012). In contrast, short filaments are ineffective at sister chromatid strand invasion and homology searching. Therefore, resection length is correlated to HR efficiency. In budding yeast, resection length away from a DSB end can vary from 2000 to 4000 nucleotides (Chung et al., 2010). These long ssDNA tracts are dependent on the processive 5’-3’ nucleases responsible for long-range end resection, Exo1 and Dna2 (Zhu et al., 2008). In cells lacking both Exo1 and Dna2, resection lengths are substantially shorter which inhibits the completion of HR (Chung et al., 2010).

However, hyper-resection or the over-production of ssDNA in the genome can also have adverse cellular consequences. Hyper-resection can increase the proportion of ectopic HR reactions where inappropriate genomic regions with similar nucleotide sequences (i.e. gene paralogs) are used as a template resulting in genome rearrangements (Montgomery et al., 1991). Hyper-resection can also increase the likelihood of exposing repetitive regions that can engage in mutagenic SSA reactions (Fishman-Lobell et al., 1992). Not surprisingly, mechanisms have evolved to curtail end resection, keeping HR in check. Compared to proteins that promote end resection, less is known about inhibitory factors. One such factor, mammalian DNA helicase B (HELB), is recruited to ssDNA by interacting with RPA and it uses its 5’-3’ ssDNA translocase activity to curtail long-range end resection mediated by EXO1 and DNA2 (Tkac et al., 2016). HELB is recruited in an RPA-dependent manner illustrating an elegant feedback mechanism to inhibit on-going end resection. End resection can also be inhibited by controlling CtIP protein levels through the action the prolyl isomerase PIN1 (Steger et al., 2013). PIN1 inhibits CtIP by modulating the isomerization of prolines within several of its CDK sites. PIN1 activity promotes CtIP ubiquitylation and subsequent proteasomal degradation. Therefore, PIN1 deficient cells

have high CtIP levels and hyper-resect DSB ends which leads to more mutagenic repair by SSA. Another impediment to end resection is the presence of nucleosomes (Adkins et al., 2013). Not surprisingly, nucleosomes antagonize long-range resection by Exo1 and Dna2 more than short- range resection by MRX and Sae2 (CtIP). Two members of the SNF2 ATPase family of chromatin remodeling enzymes, Fun30 (SMARCAD1) and SRCAP, appear to be the major remodeling enzymes that are recruited to DSB sites to facilitate long-range end resection (Costelloe et al., 2012; Dong et al., 2014).

1.5.3.2 DNA end resection versus end protection: the cross-roads of DSB repair pathway choice

NHEJ and HR are the principal pathways for DSB repair and the choice between them depends on the species, cell type, cell cycle stage, and type of DNA damage. The greatest determinant of DSB repair pathway choice is position within the cell cycle. NHEJ is active in the G1-, S-, and G2-phases of the cell cycle, whereas HR is active after replication in S and G2 cells, where a suitable homologous template is available. Cell cycle-regulated choice between DSB repair pathways is critical for the promotion of genome integrity. For instance, HR is required in S- phase to re-establish collapsed replication forks by repairing one-ended DSBs. Inappropriate NHEJ of these breaks is associated with genome rearrangements and cell death (Bouwman et al., 2010; Bunting et al., 2010; Saberi et al., 2007). In the same vein, inappropriate HR reactions in G1-phase can result in strand invasion and homology-driven copying of a homologous chromosome rather than a sister chromatid, potentially resulting in loss of heterozygosity (LOH) (Little and Benjamin, 1991).

The major control step in the choice between NHEJ and HR is the generation of ssDNA by end resection; once a DSB end is resected NHEJ cannot be performed. However, Ku70/80 binds DSB ends rapidly, before end resection has initiated and can protect DSB ends from the long-range end resection machinery (Fig. 1.6A) (Shao et al., 2012; Sun et al., 2012). In budding yeast, the inhibitory action of Ku70/80 on end resection in S and G2 cells is alleviated by the CDK-dependent activity of the MRX complex and Sae2 (CtIP) (Fig. 1.6A) (Mimitou and Symington, 2010; Shim et al., 2010). One model is that the MRX complex and Sae2 (CtIP) directly removes Ku70/80 from DSB ends. After end resection, Ku70/80 is unable to bind DSB ends and the break is now committed to being repaired by HR, SSA, or MMEJ (Mimori and Hardin, 1986). Moreover, end resection is suppressed in G1 cells through CtIP ubiquitylation and

degradation by the proteasome (Lafranchi et al., 2014). CtIP ubiquitylation is conducted by the late mitosis/G1-specific E3 ubiquitin ligase APC/C (CDH1). In addition to KU70/80, 53BP1 and its effector proteins RIF1, PTIP, and MAD2L2 play an important role in protecting DSB ends from end resection in mammalian cells (Fig. 1.6B) (Boersma et al., 2015; Bunting et al., 2010; Chapman et al., 2013; Di Virgilio et al., 2013; Escribano-Diaz et al., 2013; Xu et al., 2015; Zimmermann et al., 2013). RIF1 accumulation at DSBs is strongly inhibited by BRCA1-CtIP in the S- and G2-phases of the cell cycle, promoting end resection and HR (Escribano-Diaz et al., 2013). In contrast, 53BP1 and RIF1 antagonize the accumulation of BRCA1 at DSB sites during the G1-phase, promoting NHEJ.

1.5.4 Cell cycle-regulated suppression of homologous recombination in G1 cells downstream of end resection

In addition to the antagonism of end resection in G1 by KU70/80, 53BP1, RIF1, PTIP, and MAD2L2, HR is also inhibited in G1 cells downstream of end resection through the ubiquitin- dependent regulation of the BRCA1-PALB2-BRCA2 mediator complex (Orthwein et al., 2015). In S and G2 cells, BRCA1 directly interacts with PALB2 and promotes the recruitment of the HR factors BRCA2 and RAD51 to DSB sites. In G1 cells, the BRCA1-PALB2 interaction is ablated by PALB2 ubiquitylation which is carried out by an E3 ubiquitin ligase composed of KEAP1, CUL3, and RBX1. Cells deficient in KEAP1 can assemble a stable BRCA1-PALB2- BRCA2 complex in all cell cycle phases. In S and G2 cells, a deubiquitylase, USP11, is responsible for keeping PALB2 in a hypo-ubiquitylated state, promoting the assembly of the BRCA1-PALB2-BRCA2 complex and subsequent loading of RAD51 (Orthwein et al., 2015; Schoenfeld et al., 2004; Wiltshire et al., 2010). Remarkably, it has been demonstrated that HR can be activated in G1 cells by simply overriding the triple block to resection by expressing a hyper-active phospho-mimetic CtIP mutant (T847E) in a 53BP1 and KEAP1 deficient background (Huertas and Jackson, 2009; Orthwein et al., 2015). In addition, HR is also inhibited in G1 cells by the action of the microRNAs miR-1255b, miR-148b*, and miR-193b* (Choi et al., 2014). These microRNAs target the transcripts of several HR factors including BRCA1, BRCA2, and RAD51, which decreases their expression in G1 cells.

Figure 1.6. The regulation of DNA double-strand break repair pathway choice.

(A) DSB ends can be simultaneously recognized by the KU70/80 and MRN complexes. In G1 cells, CtIP is in an inactive state and cannot promote MRE11-dependent end resection. Therefore, KU70/80 can function in stimulating NHEJ. In S and G2 cells, CtIP is activated by CDK-dependent phosphorylation which stimulates the function of MRE11 in initiating end resection. MRE11/CtIP-dependent resection antagonizes the function of KU70/80 possibly by directly removing it from DSB ends. Resected DSBs are committed to repair by HR. (B) The ubiquitin-dependent recruitment of 53BP1 to DSB sites is also important for regulating the choice between NHEJ and HR. In G1 cells, 53BP1 recruits its effector proteins RIF1 and MAD2L2 which inhibit end resection and promote NHEJ. In S and G2 cells, BRCA1 and CtIP inhibit recruitment of RIF1/MAD2L2 to DSB sites, stimulating end resection and HR.

1.6 Post-translational modifications and the regulation of DNA double-strand break repair

1.6.1 Ubiquitin-dependent signaling at DNA double-strand breaks

PIKK activation also sets into motion an ubiquitylation-based signaling cascade on chromatin flanking DSB sites that is required for recruiting checkpoint and repair factors. In response to DSBs, MDC1 can bind to γH2AX where it is also phosphorylated by ATM (Stucki et al., 2005). Phosphorylated MDC1 in turn is recognized by the FHA domain of the RING-type E3 ubiquitin ligase RNF8 which catalyzes the ubiquitylation of proteins at DSB sites (Huen et al., 2007; Kolas et al., 2007; Mailand et al., 2007). RNF8-dependent ubiquitylation events are recognized by the ubiquitin binding domains of RNF168, another E3 ubiquitin ligase (Doil et al., 2009; Stewart et al., 2009). RNF8/RNF168 promote the conjugation of lysine 63 (K63)-linked ubiquitin to their substrates which include the histones H1 and H2A (Doil et al., 2009; Mailand et al., 2007; Mattiroli et al., 2012; Stewart et al., 2009; Thorslund et al., 2015). The critical outcome of RNF8/RNF168-dependent ubiquitylation is the recruitment of DSB repair and signaling proteins to chromatin surrounding the break site. Proteins that localize to DSBs in a RNF8-dependent manner include 53BP1, RAD18, BRCA1, RAP80, and HERC2 (Bekker-Jensen et al., 2010; Huang et al., 2009; Huen et al., 2007; Kolas et al., 2007; Mailand et al., 2007; Silverman et al., 2004; Wang and Elledge, 2007). The RNF8 pathway can promote NHEJ, especially in developing lymphocytes during class-switch recombination (Ramachandran et al., 2010). Whether or not the RNF8 pathway has a functional role in directly promoting HR is less clear. Importantly, by promoting the recruitment of 53BP1, RIF1, PTIP, and MAD2L2, the RNF8 pathway plays a critical function in regulating the choice between NHEJ and HR.

In addition to RNF8/RNF168 many other E3 ubiquitin ligases have been implicated in controlled the cellular response to DSBs. The ubiquitin ligase RNF138, in collaboration with its E2 conjugating enzyme UBE2D, is also recruited to DSB sites and promotes end resection and HR (Ismail et al., 2015; Schmidt et al., 2015). Following DSB formation, KU70/80 and the MRN complex rapidly and independently bind to DSB sites (Britton et al., 2013). The zinc finger domains of RNF138 are thought to recognize short tracks of ssDNA generated by MRN/CtIP, whereas its ubiquitin-binding domains likely stabilize it at DSB sites (Ismail et al., 2015; Schmidt et al., 2015). RNF138 can ubiquitylate KU70/80 which removes it from DSB ends and

promotes end resection (Ismail et al., 2015). RNF138 can also mediate ubiquitylation of CtIP to enhance its recruitment to DSB sites and stimulate end resection (Schmidt et al., 2015).

1.6.2 SUMOylation at DNA double-strand breaks

Many components of the SUMOylation pathway are recruited to DSB sites and promote both NHEJ and HR (Galanty et al., 2009; Morris et al., 2009). The SUMO-specific E3 ligase PIAS1 promotes the recruitment of BRCA1 and RAP80 to DSB sites whereas PIAS4 enhances the localization of RNF168 and subsequent conjugation of K63-linked ubiquitin chains. Only a handful of SUMOylated substrates at DSB sites have been identified including MDC1, RNF168, HERC2, 53BP1, BRCA1, RPA70, and BLM (Danielsen et al., 2012; Dou et al., 2010; Galanty et al., 2009; Morris et al., 2009; Ouyang et al., 2009). SUMOylation at DSB sites has been directly implicated in the regulation of HR. First, RPA70 is able to physically interact with the SUMO protease SENP6 in S-phase which keeps RPA70 in a hypo-SUMOylated state (Dou et al., 2010). In response to DNA damage, SENP6 dissociates from RPA70 allowing for the conjugation of SUMO2/3. SUMOylated RPA70 promotes the recruitment of RAD51 and thus stimulates HR. Next, the RecQ helicase BLM, implicated in both end resection and dHJ dissolution, is also SUMOylated (Ouyang et al., 2009). The expression of BLM mutants that cannot be SUMOylated results in DNA damage sensitivity and impaired RAD51 recruitment. Interplay between ubiquitylation and SUMOylation has also be demonstrated to promote HR through the action of the SUMO-targeted ubiquitin ligase RNF4 (Galanty et al., 2012; Yin et al., 2012). RNF4 is recruited to DSB sites through its N-terminal SUMO-interacting motifs which bind to SUMOylated proteins such as 53BP1, MDC1, and RPA. RNF4-mediated ubiquitylation regulates the rate of turnover of its substrates by targeting them for proteasomal degradation. Remarkably, RNF4 also promotes the recruitment of the proteasome to DSB sites (Galanty et al., 2012). Importantly, RNF4-mediated ubiquitylation and subsequent proteasomal degradation of RPA70 at DSB sites promotes HR by enhancing the exchange of RPA for RAD51 (Galanty et al., 2012; Yin et al., 2012).

1.6.3 Protein acetylation at DNA double-strand breaks

Several studies have outlined the importance of acetylation for regulating protein localization and function at DSBs. For example, both NBS1 and RAD51 are kept in a hypo-acetylated state by SIRT1 which promotes their recruitment to DSBs (Oberdoerffer et al., 2008). Furthermore,

CtIP deacetylation by SIRT6 is important for stimulating its activity in end resection (Kaidi et al., 2010). Protein deacetylation is also important for DSB repair by NHEJ (Miller et al., 2010). Deacetylation of H3K56 by HDAC1 and HDAC2 is important for the recruitment of KU70/80 and Artemis to DSBs. Furthermore, TIP60 forms a stable complex with ATM and promotes its acetylation in response to DNA damage (Sun et al., 2005). Suppression of TIP60 blocks ATM activation and prevents the ATM-dependent phosphorylation of both p53 and CHK2. In response to DNA damage, TIP60 is activated by c-Abl-dependent tyrosine phosphorylation (Kaidi and Jackson, 2013). The mechanism for c-Abl activation is less clear but may involve DNA damage- induced chromatin reorganization.

1.6.4 PARylation is one of the earliest protein modifications detected at DNA double-strand breaks

PARPs are enzymes that catalyze the addition of poly(ADP-ribose) (PAR) chains on protein substrates (Stone and Shall, 1973). PARP enzymes have been implicated in PARylating proteins present at both SSBs and DSBs (Durkacz et al., 1980; Ikejima et al., 1990; Rulten et al., 2011). Importantly, PAR chains are transient protein modifications that are removed quickly at DNA breaks by the action of PAR glycohydrolase (PARG) (Schreiber et al., 2006). PARP-1 physically associates with XRCC1 and is required for sensing SSBs and promoting the recruitment of downstream repair enzymes to re-join the broken ends (Caldecott et al., 1996). PARP-1 also enhances DSB repair by promoting the recruitment of several repair enzymes to DNA damage sites including MRE11, NBS1, APLF, and TIMELESS (Ahel et al., 2008; Bryant et al., 2009; Haince et al., 2008). PARP-1 is required for the repair of DSBs by the alternative KU70/80- indepentent MMEJ pathway (Audebert et al., 2004). Moreover, PARylation is important for the localization of chromatin remodeling factors to DSB sites including ALC1, CHD2, CHD4, EZH2, PCGF2, and BMI (Luijsterburg et al., 2016; Seeber et al., 2013). PARP-dependent chromatin decompaction at DNA damage sites likely promotes accessibly to DNA repair enzymes.

1.7 Relevance of DSB repair to human physiology

Proper repair of DSBs is critical for the maintenance of genome integrity and cellular fitness. It is not surprising that germline mutations in DSB repair genes give rise to a set of human maladies, termed genome instability syndromes. One of the first characterized genome instability

syndromes is Ataxia-Telangiectasia (A-T), which is caused by autosomal-recessive mutations in the ATM gene (Savitsky et al., 1995). Cells from A-T patients are defective in checkpoint activation and DSB signaling and repair. As a consequence, clinical features include immunodeficiency (due to impaired V(D)J and class-switch recombination), neurodegeneration, and a predisposition to the development of cancer. A list of other genome instability disorders is summarized in Table 1.1.

1.7.1 DSB repair promotes tumour suppression

A fundamental hallmark of cancer is genome instability (Hanahan and Weinberg, 2000). DSB repair is critical for maintaining genome stability and thus for inhibiting tumourigenesis. This is illustrated by the cancer predisposition observed in some genome instability syndromes. Defective DSB repair has the unique ability to elicit chromosome rearrangements that can drive cancer development. For example, aberrant DSB repair in V(D)J or class-switch recombination can fuse proto-oncogenes to antigen receptor loci resulting in the development of lymphoid tumours (Schlissel et al., 2006).

1.7.1.1 Defective HR is associated with a predisposition to breast and ovarian cancers

Mutations in genes that promote DSB repair by HR have been associated with the development of breast and ovarian cancers (Kato et al., 2000; Miki et al., 1994; Narod et al., 1993; Soria- Bretones et al., 2013; Wooster et al., 1994). The greatest risk factor for breast and ovarian cancer are germline mutations in one of the breast cancer susceptibility genes, BRCA1, BRCA2, or PALB2, which are required for HR. Furthermore, mutations in the RAD51 paralogs RAD51C and RAD51D can predispose individuals to ovarian cancer, whereas mutations in RAD51B can lead to breast cancer susceptibility (Golmard et al., 2013; Loveday et al., 2011; Meindl et al., 2010). Therefore, many HR genes can also be considered tumour suppressor genes.

1.7.2 DSB repair is important during cerebral cortical development

The analysis of genome instability syndromes has allowed researchers to gain perspectives on the relative importance of DSB repair for the development of different tissues. One common feature of these syndromes is neonatal microcephaly, a clinical term used to describe a reduced head circumference in newborns that is greater than three standard deviations below the mean

Syndrome Gene Radio- Immuno- Neonatal Cancer Reference sensitivity deficiency Microcephaly Predisposition HR: Ataxia ATM + + ND + (Savitsky et telangiectasia al., 1995)

Nijmegen NBS1 + + + + (Varon et breakage al., 1998)

Ataxia MRE11 + ND ND ND (Stewart et telangiectasia- al., 1999) like

Nijmegen RAD50 + ND + ND (Waltes et breakage-like al., 2009)

Seckel ATR + ND + + (O'Driscoll ATRIP ND ND + ND et al., 2003; Ogi et al., CtIP + ND + ND 2012; Qvist DNA2 ND ND + ND et al., 2011; Shaheen et al., 2014)

Jawad CtIP ND ND + ND (Qvist et al., 2011)

Bloom BLM + + ND + (Ellis et al., 1995)

NHEJ: LIG4 LIG4 + + + + (O'Driscoll deficiency et al., 2001)

ART-SCID Artemis + + ND + (Moshous et al., 2001)

XLF-SCID XLF + + + ND (Buck et al., 2006)

Table 1.1. Clinical features of some genome instability syndromes associated with defective DSB repair.

List of genome instability syndromes (not exhaustive) outlined the causative gene and common clinical features. ‘+’ indicates that the clinical feature has been described in the literature for the disorder. ‘ND’ indicates that the feature has not been described for the syndrome.

(Alcantara and O'Driscoll, 2014). Microcephaly arises as a consequence of defective proliferation of neuroprogenitor cells within the developing cerebral cortex (Bond et al., 2002). Neuroprogenitor proliferation defects have been attributed to both increased apoptosis and cell cycle arrest (Chen et al., 2009; Li et al., 2012). The fact that microcephaly is a common symptom of genome instability disorders suggests that the normal re-joining of DSBs is critical to the developing cerebral cortex. It currently remains unclear why DSB repair appears to be more important in the developing brain compared to other tissues. One hypothesis is that neuroprogenitor cells may have a lower threshold for apoptosis making it easier to activate programmed cell death in the presence of unrepaired DSBs. Indeed, apoptosis is a fundamental component of nervous system development and is required for regulating neural cell numbers, tissue remodeling, and eliminating mis-specified cells (Yamaguchi and Miura, 2015). In addition, it is also possible that neuroprogenitor cells have a lower threshold for activating cell cycle checkpoints in response to DSBs. It is interesting to note that patients with A-T do not display microcephaly (Savitsky et al., 1995). One explanation is that apoptosis or cell cycle checkpoint activation could be ATM-dependent in neuroprogenitor cells. Therefore, unrepaired DSBs would be allowed to persist and the damaged cells could accumulate in the developing brain, potentially contributing to the observed neurodegenerative phenotypes described in A-T patients.

1.7.2.1 Seckel syndrome and DNA end resection

Seckel syndrome is a genome instability disorder with clinical features that include intrauterine growth retardation, dwarfism, a ‘bird-like’ facial appearance, and cognitive delay (Shanske et al., 1997). The defining feature of Seckel syndrome is severe microcephaly. A mutation in the ATR gene was first determined to be a cause for Seckel syndrome (O'Driscoll et al., 2003). Later, another Seckel syndrome patient was found to have a mutation in the ATR interacting protein ATRIP, further suggesting that ATR signaling is important for the development of the cerebral cortex and for the prevention of microcephaly (Ogi et al., 2012). As previously discussed, ATR is activated by genomic regions containing ssDNA bound by RPA. Regions of ssDNA-RPA can arise as a consequence of replication fork stalling and DNA end resection during HR. Remarkably, the core end resection factors CtIP and DNA2 were also found to be mutated in two separate cases of Seckel syndrome, suggesting that ATR activation at resected DSBs may be critical for cerebral cortical development (Qvist et al., 2011; Shaheen et al., 2014). It cannot be

excluded that the role of CtIP and DNA2 in HR, rather than ATR activation, contributes to the promotion of neuroprogenitor cell proliferation in the developing cerebral cortex. However, the observation that genes promoting HR downstream of end resection have not been implicated in microcephaly suggests that the function of CtIP and DNA2 in ATR activation, rather than HR, is important for cerebral cortical development. Interestingly, Nijmegen breakage syndrome (NBS) and Nijmegen breakage syndrome-like (NBS-like), where components of the resection- promoting MRN complex are mutated, also display neonatal microcephaly (Varon et al., 1998; Waltes et al., 2009).

1.8 Rationale and research objective

DSB repair by HR is a critical pathway for maintaining genome stability and preventing tumourigenesis. Therefore, at the onset of my doctoral research I sought out to uncover new factors that regulate HR using a functional genomics approach. I decided to focus my attention on the rate-limiting step of HR, DNA end resection. Several core resection factors are known in human cells, including the MRN complex, CtIP, EXO1, DNA2, and BLM. However, these factors were first identified in budding yeast which begs the question if additional factors are required for end resection in human cells. Furthermore, the mechanistic detail regarding the regulation of the known core resection activators is not clear. It is also an open question how resection length is regulated in human cells. Lastly, an important outstanding question is how resection is regulated in the context of chromatin structure. My research objective is to conduct an RNAi screen in human cells to mine for new regulators of end resection in an attempt to characterize new factors and to potentially shed light on these unresolved questions.

1.9 High-throughput functional discovery utilizing RNA interference screens

First discovered in Caenorhabditis elegans and conserved in most eukaryotic species, RNAi is a post-transcriptional gene silencing process that is mediated by double-stranded RNA (Fire et al., 1998). Long precursor RNAs are processed by the ribonuclease DICER into the effectors of RNAi called small interfering RNAs (siRNAs). To elicit target messenger RNA depletion in human cells, siRNAs must not be larger than 30 base pairs as this leads to the activation of the anti-viral interferon response (Stark et al., 1998). RNAi can be used to perturb the expression of specific genes of interest. Genetic perturbation is accomplished by cellular introduction of RNAi

reagents such as synthetic siRNAs, endoribonuclease-prepared siRNAs (esiRNAs), or siRNA precursors such as short hairpin RNAs (shRNAs) that are designed to target a specific transcript (Moffat and Sabatini, 2006).

RNAi reagents can be employed for systematic high-throughput screens that assay specific cellular phenotypes. To facilitate large-scale screens, a number of genome-scale RNAi libraries have been developed by academic and commercial entities and can be delivered to cultured cells in one of two screening formats: arrayed or pooled (Moffat and Sabatini, 2006). For arrayed screening, each well of a multi-well plate can contain an RNAi reagent targeting a specific gene. To ensure gene silencing, some libraries employ several RNAi reagents that hybridize to different locations on a particular transcript. Moreover, reagents that target the same transcript can be pooled into one well to minimize the number of samples in a screen. One advantage of the arrayed format is that the gene target for each well can be easily identified during downstream analysis. In addition, more complex cellular phenotypes can be quantified for each sample using a method termed high-content screening where automated microscopes take images of RNAi-treated cells in each well. Sophisticated image analysis software programs are then employed to measure a wide range of subtle cellular phenotypes. For example, our laboratory has conducted high-content siRNA arrayed screens that monitor the localization of proteins to DSB sites which can be visualized cytologically as subnuclear foci (Kolas et al., 2007; O'Donnell et al., 2010; Stewart et al., 2009). A plethora of information about these foci can be measured by high-content analysis including number per nuclei, size, shape, texture, and intensity. RNAi libraries can also be screened in a pooled format. Genome-wide shRNA plasmid libraries can be packaged into viral particles and pooled. The pool of virus is infected into a population of cells and after the duration of the screen the unique shRNA sequences that were integrated into the genome can be PCR amplified using vector-derived primers. The representation of each shRNA sequence (or molecular barcode) in the population can be identified by next-generation sequencing or hybridization to a library-specific microarray (Berns et al., 2004; Paddison et al., 2004; Vizeacoumar et al., 2013). Pooled RNAi screens are powerful for determining the effect of gene perturbation between two cell populations. For instance, populations could have different genetic backgrounds or could have been treated with different stressors. If the depletion of a specific gene sensitizes cells in a particular condition, the shRNA sequence will drop out of the population. In contrast, if depletion of a gene promotes survival the

shRNA sequence will be enriched in the population. One disadvantage of pooled screening is that it relies on the quantification of cellular fitness and cannot be used for the analysis of more subtle subcellular phenotypes. The following chapter will describe an arrayed genome-scale RNAi screen that I performed utilizing an siRNA library that targets 18,452 human genes. The quantification of DNA end resection was conducted in cells treated with each library reagent using high-content analysis.

Chapter II

Genome-scale siRNA screen for regulators of DNA end resection

2.1 Statement of contributions, rights, and permissions

Thomas Sun and Alessandro Datti (LTRI robotics facility) developed liquid-handling robotic procedures to preform automated siRNA transfections and immunofluorescence. In addition, they helped in all robotic troubleshooting.

Mikhail Bashkurov (LTRI high-content screening facility) helped in all aspects of automated confocal microscopy and downstream image analysis.

2.2 Summary

DNA end resection is the rate-limiting step of HR. Efficient end resection promotes the assembly of a long RAD51 nucleoprotein filament which will enhance the process of sister chromatid strand invasion and homology search. End resection is also at the cross-roads of DSB repair pathway choice. Once end resection has commenced, DSB ends are no longer compatible for KU70/80 binding and re-joining by NHEJ. Therefore, the regulation of end resection initiation at DSB sites is an important mechanism to control the choice between HR and NHEJ throughout the cell cycle. The molecular details of end resection have been mostly dissected in budding yeast, leading one to speculate if additional factors are necessary in human cells. Here, I report the establishment of a cell-based assay that measures end resection in an automated and quantitative manner in human cells. I employ this assay to screen a genome-scale siRNA library that targets 18,452 genes and identify candidate end resection activators and inhibitors. The success of the screen is demonstrated by the identification of known end resection factors including all three subunits of the MRN complex and CtIP. I show that the completion of several confirmation screens allows the categorization of real end resection regulators from false- positives.

2.3 Introduction

My goal when I started my doctoral research was to uncover novel genes that function in DSB repair by HR, specifically at the step of DNA end resection. To achieve this goal, I devised an RNAi screen in human cells. Three cell-based assays are routinely used in the DSB repair field to measure end resection and each relies on the quantification of nuclear immunofluorescence intensity which makes them strong candidates for an arrayed high-content RNAi screen. Two of the three assays exploit the function of the RPA complex which rapidly coats ssDNA formed as a consequence of end resection at DSB sites. The first assay monitors the formation of subnuclear RPA foci by conducting RPA32 immunofluorescence (Fig. 2.1A). The second assay monitors RPA focus formation by utilizing an antibody that detects phosphorylated RPA32 (Fig. 2.2A). In response to DSBs, RPA32 becomes hyper-phosphorylated on multiple serine and threonine residues by the DNA damage responsive kinases (Brush et al., 1994; Liu and Weaver, 1993; Shao et al., 1999). RPA32 hyper-phosphorylation prevents the complex from associating with ssDNA at DNA replication forks, which may increase the free pool of RPA that is available for binding resected DSB ends (Vassin et al., 2004). The mechanistic detail of how the recruitment of hyper-phosphorylated RPA32 is inhibited at replication centers in unknown. The kinetics of RPA32 phosphorylation and dephosphorylation are also important for HR. The PP4 phosphatase complex interacts with RPA32 and can dephosphorylate it (Lee et al., 2010). PP4-depleted cells have an increase in hyper-phosphorylated RPA32 that is not bound to chromatin. The free pool of phosphorylated RPA32 sequesters RAD51 away from DSB sites, thereby inhibiting HR. The last assay measures the relative amount of ssDNA generated by end resection through native immunostaining of the thymidine analog, bromodeoxyuridine (BrdU). BrdU is added to the medium and is incorporated into the genome during DNA replication. After long term incubation of cells with BrdU, cells are treated with a DSB-inducing agent and processed for BrdU immunofluorescence. The antibody only detects BrdU in the context of ssDNA and can be used as a more direct readout of the amount of ssDNA formed by end resection (Fig. 2.3A). My first course of action was to establish these cell-based assays so that I could select the most appropriate method for a high-content RNAi screen.

2.4 Results

2.4.1 Establishment of immunofluorescence-based assays to monitor DNA end resection In order to conduct an RNAi screen for regulators of end resection, I first needed to establish a cell-based assay to monitor resection in human cultured cells. As discussed above, three immunofluorescence-based assays are routinely utilized in the DSB repair field to monitor resection by microscopy, including the quantification of RPA32, phospho-RPA32 (S4/S8), and BrdU focus formation. Several DSB-inducing drugs are frequently used to induce detectable amounts of end resection in human cells, including the topoisomerase I poison CPT and the radiomimetic drug NCS. CPT induces DSBs only in the S phase of the cell cycle and I surmised that it would be important in a screen to monitor end resection occurring in other cell cycle phases. In contrast, NCS indiscriminately induces DSBs in all cell cycle phases and was selected for the rest of the study. Next, to test immunostaining and NCS treatment conditions I manually seeded and reverse-transfected human bone osteosarcoma U-2 OS (U2OS) cells with control scrambled or CtIP siRNA (Fig. 2.1B, 2.2B, and 2.3B). U2OS cells were used in this study because they possess a flat morphology and are highly adherent to glass, making them an ideal model cell line for conducting automated high-throughput microscopy-based screens (Ponten and Saksela, 1967). In addition, U2OS cells are readily transfected with siRNAs and they are routinely utilized as a model cell line to study DSB repair. After 48 hours-post siRNA transfection (to allow time for target mRNA depletion), cells were treated with various concentrations of NCS for different time periods and were processed for either RPA32, phospho- RPA32 (S4/S8), or BrdU immunostaining. Next, I visualized cells on a LSM780 Zeiss confocal microscope utilizing a 60X oil immersion objective to observe NCS-induced sub-nuclear focus formation. To circumvent high levels of pan nuclear background staining with the RPA32 and BrdU antibodies, cells were pre-extracted with nuclear extraction buffer before fixation to remove any proteins that were not bound to chromatin. Decreased background staining allowed for the visualization of chromatin-associated RPA32 and BrdU foci. I then manually quantified the percentage of cells in each treatment condition that contained more than 15 RPA32, pRPA32 (S4/S8), or BrdU foci (Figure 2.1C, 2.2C, 2.3C, respectively). I selected a threshold of 15 because some treatments contained upwards of 100 foci per nucleus making it difficult to count individual foci. Furthermore, approximately 5-10 foci were present in cells not treated with NCS and in cells depleted of the core resection regulator, CtIP. Therefore, background foci are present

Figure 2.1. DNA end resection assay monitoring RPA32 focus formation. (A) The RPA heterotrimeric complex rapidly binds to ssDNA generated by resection at DSB ends. (B) The accumulation of RPA at DSB sites can be observed cytologically as subnuclear foci after RPA32 immunostaining. U2OS cells transfected with scrambled control or CtIP siRNAs were either mock treated (-NCS) or treated with 50 ng/ml NCS for 3 hours. Incubation with NCS was followed by pre-extraction, fixation, and RPA32 immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. Scale bar represents 5 µm. (C) Manual quantification of the percentage of cells with greater than 15 RPA32 nuclear foci after incubation with various concentrations of NCS and at different time points (Mean ± Standard error of the mean (SEM); N ≥ 3).

Figure 2.2. DNA end resection assay monitoring pRPA32 (S4/S8) focus formation. (A) The RPA heterotrimeric complex rapidly binds to ssDNA generated by resection at DSB ends. RPA32 become hyper-phosphorylated on multiple serine and threonine residues by PIKKs. Hyper-phosphorylated RPA32 can be detected by employing a phospho-specific antibody that recognizes phosphorylated serines 4 and 8. (B) U2OS cells transfected with scrambled control or CtIP siRNAs were either mock treated (-NCS) or treated with 100 ng/ml NCS for 3 hours. Incubation with NCS was followed by fixation and pRPA32 (S4/S8) immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. Scale bar represents 5 µm. (C) Manual quantification of the percentage of cells with greater than 15 pRPA32 (S4/S8) nuclear foci after incubation with various concentrations of NCS and at different time points (Mean ± SEM; N ≥ 3).

Figure 2.3. DNA end resection assay monitoring BrdU focus formation. (A) The generation of ssDNA at DSB sites can be directly measured by incubating cells with the thymidine analog BrdU and conducting BrdU immunostaining. The antibody can only detect BrdU in the context of ssDNA and thus is a valuable tool for monitoring end resection. (B) U2OS cells incubated with BrdU and transfected with scrambled control or CtIP siRNAs were either mock treated (-NCS) or treated with 50 ng/ml NCS for 3 hours. Incubation with NCS was followed by pre-extraction, fixation, and BrdU immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. Scale bar represents 5 µm. (C) Manual quantification of the percentage of cells with greater than 15 BrdU nuclear foci after incubation with various concentrations of NCS and at different time points (Mean ± SEM; N ≥ 3).

48 in U2OS cells that are not a result of DNA end resection at DSB sites. RPA-bound ssDNA can also occur at replication forks and active sites of transcription and these background foci may be a result of these genomic processes. As expected, I observed an increase in the number of RPA32, pRPA32 (S4/S8), and BrdU foci in U2OS cells treated with NCS. These foci were largely dependent on the core resection activator CtIP and increased in number with increasing concentrations of NCS. Furthermore, the foci also increased in number with time, with maximal levels observed from 3 to 5 hours after the addition of NCS. A large population of cells in each condition were devoid of foci and this observation is consistent with the fact that end resection is actively inhibited during the G1 phase of the cell cycle (Bunting et al., 2010; Huertas and Jackson, 2009). Representative micrographs shown in Figure 2.1, 2.2, and 2.3 are of U2OS cells 3 hours post-NCS addition.

2.4.2 Quantitative image-based cytometry to monitor DNA end resection

Genome-scale RNAi screens that quantified the number and intensity of nuclear DNA damage foci required high magnification micrographs (40-60X) and many fields needed to be imaged per sample in order to acquire data on a sufficient number of cells (Kolas et al., 2007; O'Donnell et al., 2010; Stewart et al., 2009). The requirement of more fields resulted in long imaging times per plate, creating a bottle-neck in the screening pipeline. Quantitative image-based cytometry (QIBC) is a plate-based method to rapidly scan multiwell plates using low magnification (4-10X) objective lens (Toledo et al., 2013). QIBC enables users to image the entire well and thus acquire data on thousands of cells for every sample (akin to flow cytometry). Importantly, the analysis of more cells can increase the statistical quality of an RNAi screen (Birmingham et al., 2009). First, I tested whether an increase in the total nuclear RPA32, pRPA32 (S4/S8), or BrdU intensity could be detected in response to NCS using the Celigo plate cytometer (Brooks Automation), which rapidly scans plates with a 4X objective (Fig. 2.6C). U2OS cells were seeded and reverse- transfected with control or CtIP siRNAs in a 96-well plate. After 48 hours, cells were treated with NCS and processed for RPA32, pRPA32 (S4/S8), or BrdU immunofluorescence. Using pRPA32 (S4/S8) stained cells as an example, representative cytometry images are depicted in Figure 2.4A with the corresponding cell-by-cell analysis outlined in Figure 2.4B. Image analysis was conducted using software packaged with the Celigo plate cytometer. In brief, DAPI stained nuclei were segmented and the intensity of either RPA32, pRPA32 (S4/S8), or BrdU immunostaining was measured under a nuclear mask. Cell-by-cell intensities were plotted in

frequency distributions and in response to NCS bi-modal distributions were observed. Bi-modal distributions were expected as end resection is a cell cycle-regulated process occurring primarily in the S and G2 phases. Next, an arbitrary intensity threshold was established to determine the percentage of cells that were positive (Fig. 2.4C). The percentage of NCS-induced RPA-, pRPA32 (S4/S8)-, or BrdU-positive cells decreased substantially in cells depleted of CtIP. Furthermore, the dynamic range between the control and CtIP siRNAs was similar to that observed when manually quantifying focus formation using a 60X objective (as outlined in Fig. 2.1, 2.2, and 2.3). Therefore, QIBC is a robust method that can be utilized to more rapidly quantify end resection in U2OS cells.

As end resection assays display non-Gaussian bi-modal distributions, I decided to employ a non-parametric statistical analysis called the two-sample Kolmogorov-Smirnov (KS) test. For this test, cell-by-cell intensity data was plotted in a cumulative frequency distribution for a reference sample (i.e. siCTRL) and a test sample (i.e. siCtIP). As an example, cumulative frequency distributions of pRPA32 (S4/S8) nuclear intensities are shown in Figure 2.5A. The KS score is the maximum vertical distance between the reference and test distributions. The KS test was conducted on RPA32, pRPA32 (S4/S8), and BrdU nuclear intensity distributions from cells depleted of several known resection factors including CtIP, MRE11, RAD50, and NBS1 (Fig. 2.5B). Cumulative frequency distributions and KS tests were conducted using MATLAB (The MathWorks Inc.) in collaboration with Mikhail Bashkurov in the Lunenfeld-Tanenbaum Research Institute’s (LTRI) high-content screening facility.

2.4.3 Automating DNA end resection assays using liquid-handling robotics

With the establishment of cell-based end resection assays, the next step towards the commencement of an RNAi screen was to test whether siRNA transfections and immunostaining procedures could be carried out in an automated manner using liquid-handling robotics. For this I initiated a collaboration with Alessandro Datti and Thomas Sun in the LTRI’s robotics facility. We developed a protocol on the 96 tip Biomek FX (Beckman-Coulter; Fig. 2.6A) liquid handler to seed and transfect U2OS cells with siRNAs in 384-well plates. In addition, we programmed a procedure to conduct immunostaining on the Dimension 4 robotic platform (Fig. 2.6B) utilizing various peripheral instruments including a Biomek FX liquid handler (Beckman-Coulter) and an Embla plate washer (Molecular Devices). To test our protocol, we seeded and transfected U2OS

Figure 2.4. Measuring end resection by quantitative image-based cytometry. (A) U2OS cells seeded in a 96-well plate and incubated with scrambled control or CtIP siRNA were treated with 100 ng/ml NCS for 3 hours before pRPA32 (S4/S8) immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. QIBC using the Celigo plate cytometer and image analysis software was conducted to segment DAPI-stained nuclei and measure the mean pRPA32 (S4/S8) nuclear intensity. Scale bar represents 200 µm. (B) Cell-by-cell pRPA32 (S4/S8) nuclear intensities were plotted in frequency distributions. (C) The percentage of NCS-induced RPA32-, pRPA32 (S4/S8)-, and BrdU-positive nuclei was determined by setting an arbitrary intensity threshold (Mean ± SEM; N ≥ 3).

Figure 2.5. Application of the Kolmogorov-Smirnov test to analyze end resection. (A) Cell-by-cell pRPA32 (S4/S8) mean nuclear intensity were plotted in a cumulative frequency distribution. The Kolmogorov-Smirnov (KS) score was determined by measuring the maximal vertical distance between the reference (siCTRL) and test distributions (siCtIP). (B) KS scores were calculated for RPA32, pRPA32 (S4/S8), and BrdU immunostaining and multiple control siRNAs (Mean ± SEM; N ≥ 3).

cells with control, CtIP, MRE11, or NBS1 siRNA in 384-well plates using the Biomek FX. After 48 hours to allow for target depletion, the plates were loaded onto the Dimension 4 platform for the addition of NCS and subsequent RPA32, pRPA32 (S4/S8), or BrdU immunostaining. The cells were then imaged on the Celigo for QIBC (Fig. 2.6C). The RPA32 and BrdU immunostaining procedures required a pre-extraction with mild detergent before fixation. Unfortunately, after testing various washing pressures with the Embla plate washer, we were unable to find a condition that did not remove pre-extracted cells from the 384-well plates. In contrast, pRPA32 (S4/S8) immunofluorescence did not require a pre-extraction and these cells adhered very well to the plates. A heat map of pRPA32 (S4/S8) nuclear intensity KS scores for the test plate are depicted in Figure 2.6D. KS scores were similar to those observed when I conducted the transfection and immunostaining procedures manually, demonstrating that the assay can be effectively automated using liquid handling robotics. Furthermore, I also observed an excellent dynamic range between the negative and positive control siRNAs. At this stage of the project, I decided to commence a genome-scale siRNA screen that monitors pRPA32 (S4/S8) nuclear intensity in response to NCS treatment.

2.4.4 Genome-scale siRNA screen utilizing a pooled siRNA library

Using the automated end resection assay outlined above, I screened the genome-scale SMARTpool siRNA library (Dharmacon/GE Healthcare) that targets 18,452 genes (Fig. 2.7A). For each gene, the library contains a pool of four distinct siRNAs that hybridize to different regions of the target transcript. Each 384-well screening plate contained both negative (scrambled CTRL siRNA) and positive (CtIP siRNA) controls within the outside columns. After imaging and raw cell-by-cell nuclear pRPA32 (S4/S8) intensities were determined, Mikhail Bashkurov calculated the KS score for each library siRNA by comparing it to a scrambled control siRNA sample on the same plate. I set an arbitrary cut-off for hit identification at a KS score of -19 for resection activators because it included many of the known activators as hits and limited the list to a manageable ~2.5% of the data set (451/18,452). Known end resection activators, including all three subunits of the MRN complex and CtIP were identified as hits in the screen (Fig. 2.7B). In addition, several kinases responsible for RPA32 hyper- phosphorylation, DNA-PKcs and ATR, were also identified as hits. EXO1 and DNA2 are both 5’ to 3’ exonucleases known to carry out long-range end resection but were not identified as hits. I hypothesized that these two nucleases may be able to functionally compensate in long-range

Figure 2.6. Automating DNA end resection assays using liquid-handling robotics. (A) BioMek FX (Beckman-Coulter) is a 96 tip liquid dispensing robot and was used to deliver siRNA complexes, NCS, and immunostaining reagents to 384-well screening plates. (B) The Dimension 4 robotic platform facilitated the systematic use of multiple peripheral instruments including a temperature- and carbon dioxide-controlled incubator, a plate washer, and a BioMek FX liquid handler. The platform enabled immunostaining procedures to run on multiple 384-well screening plates at the same time. (C) Celigo plate cytometer (Brooks Automation) which rapidly scans multiwell plates using a 4X objective lens. (D) Heat-map of a 384-well plate illustrating the pRPA32 (S4/S8) nuclear intensity KS score for U2OS cells incubated with various control siRNAs. U2OS cells were seeded and transfected with siRNAs using the BioMek FX. Cells were treated with NCS followed by pRPA32 (S4/S8) immunofluorescence using the Dimension 4 platform. QIBC was conducted using the Celigo plate cytometer and image analysis software. KS scores were calculated using a script generated in MATLAB (Mean ± SEM; N = 2).

Figure 2.7. Genome-scale siRNA screen for regulators of DNA end resection. (A) Scatter plot depicting the NCS-induced pRPA32 (S4/S8) nuclear intensity KS scores for 18,452 siRNA pools. (B) KS scores from the RNAi screen for known end resection activators. (C) U2OS cells seeded in a 96-well plate and incubated with various combinations of scrambled control, EXO1, or DNA2 siRNA. Cells were treated with 100 ng/ml NCS for 3 hours before pRPA32 (S4/S8) immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. QIBC using the Celigo plate cytometer and image analysis software was conducted to segment DAPI-stained nuclei and measure the pRPA32 (S4/S8) nuclear intensity. KS scores were calculated using a script generated in MATLAB. (N = 3) (D) KS scores from the RNAi screen for known end resection inhibitors.

resection when the other is lost. To test this, I manually co-depleted both EXO1 and DNA2 with siRNAs and observed a dramatic decrease in end resection compared to cells treated with either EXO1 or DNA2 siRNA alone (Fig. 2.7C). Next, I set the cut-off for putative resection inhibitors at a KS score of +13 which also corresponded to ~2.5% of the data set (494/18,452). In contrast to resection activators, the screen was not as successful at identifying known resection inhibitors (Fig. 2.7D). The helicase HELQ was identified as an end resection inhibitor. However, HELQ is known to promote repair of replication-associated DSBs by HR (Adelman et al., 2013; Takata et al., 2013). HELQ functions at collapsed replication forks to promote HR downstream of RAD51 loading onto resected ssDNA. Cells deficient in HELQ have an increased accumulation of γH2AX, RPA32, and RAD51 foci in response to agents that induce replication stress. These foci persist for longer time periods in cells lacking HELQ, demonstrating a defect in the repair of replication-associated DSBs. The increase in NCS-induced hyper-phosphorylation of RPA32 observed in the RNAi screen for HELQ depleted U2OS cells is consistent with these data.

Next, I conducted pathway enrichment analysis for the identified candidate resection activators utilizing software available through Qiagen called Ingenuity Pathway Analysis (IPA). IPA is a rigorously updated database (called the Ingenuity Knowledge Base) that catalogs human genes into functional cellular processes. The list of 451 candidate resection activators was imported to IPA and pathway enrichment P values were determined using the Fisher’s exact test (Fig. 2.8). As expected, genes functionally implicated in DSB repair by HR were enriched in the data set. As end resection is a cell cycle-regulated process, it was not surprising that genes involved in cell cycle control were also enriched.

2.4.5 Secondary confirmation screen utilizing cherry-picked siRNA pools

RNAi screens are known to elicit a high number of false-positive hits (Echeverri et al., 2006). One potential source of false-positives in my screen were siRNAs that could alter the normal cell cycle profile of U2OS cells. Changes in cell cycle progression would impact the quantification of end resection, as it is a cell cycle-regulated process, occurring in only S and G2 cells. Therefore, to normalize any differences in cell cycle progression, I established a secondary confirmation assay employing the fluorescent ubiquitylation-based cell cycle indicator (FUCCI) system to measure end resection in only S and G2 cells (Sakaue-Sawano et al., 2008). Stable FUCCI cell lines express fragments of the cell cycle-regulated proteins Geminin (S/G2/M phases) and CDT1

Figure 2.8. Pathway enrichment analysis for candidate resection activators. Pathway enrichment analysis was conducted for the identified candidate resection activators using Ingenuity Pathway Analysis (IPA). IPA is a rigorously updated database that categorizes human genes into functional cellular processes. The list of 451 candidate resection activators was imported to IPA and pathway enrichment P values were determined using the Fisher’s exact test. The percentage of candidate resection activator genes that overlap with the list of genes for each functional process is depicted by the bar graph.

(G1 phase) that are fused to a green fluorescent protein (mAG) and a red fluorescent protein (mKO), respectively. Cells that express both reporters (and are yellow) are considered to be at the G1/S transition (Fig. 2.9A).

As a first line of attack, I focused my attention on confirming candidate resection activators. We cherry-picked 360 of the top activators for a confirmation screen but excluded annotated ribosomal, proteasomal, or solute carrier proteins (-40 candidate activators). These genes were excluded because I wanted to focus my attention on candidates that may have a more direct function in promoting end resection. Briefly, U2OS FUCCI cells were seeded into 384- well plates and transfected with the 360 cherry-picked siRNAs from the SMARTpool library (Dharmacon/GE Healthcare). Two days post-transfection the cells were treated with NCS for 3 hours and then immunostained for pRPA32 (S4/S8) using a secondary antibody conjugated to the far red fluorophore, Alexa647. DNA was counterstained with DAPI followed by 4-channel QIBC using the 10X objective of the InCell 6000 automated confocal microscope (GE Healthcare). Images were analyzed by segmenting DAPI-stained nuclei and identifying S and G2 phase cells (Geminin-mAG-positive cells) using an intensity threshold. As expected, there was an enrichment (141/360) of candidate resection activator siRNA pools that decreased the proportion of cells in S and G2 phases of the cell cycle (Fig. 2.9A). Of these 141 siRNA pools, 82 did not elicit any detectable end resection defect. Therefore, cell cycle position was a large source of false-positive hits in the primary screen. In addition, the percentage of pRPA32 (S4/S8)-positive S and G2 cells was determined by setting an intensity threshold. Importantly, there were still many candidate resection activators, that when depleted, decreased pRPA32 (S4/S8) nuclear intensity in only S and G2 phase cells. This list of activators included CtIP and all three subunits of the MRN complex. A cut-off of 0.8 for the relative percentage of pRPA32- positive cells was set to identify hits. I chose this cut-off because it included many of the known resection activators and resulted in a manageable list of 154 confirmed hits.

2.4.6 Re-screening deconvolved siRNAs for the top resection activator candidates

It has been well documented that siRNAs have the propensity to bind off target mRNAs and therefore elicit the down-regulation of multiple genes (Sigoillot et al., 2012). It is likely that some of the 154 candidate resection activators are false positives due to off target mRNA down- regulation. Therefore, I ordered deconvolved siRNAs for the top 70 pools (of 154) identified in

Figure 2.9. Secondary confirmation screen utilizing the fluorescent ubiquitylation-based cell cycle indicator (FUCCI) system. (A) Model of the FUCCI system and images of a QIBC end resection assay in U2OS FUCCI cells treated with 100 ng/ml NCS for 3 hours. Scale bar represents 200 µm. (B) Secondary confirmation screen for cherry-picked siRNA pools targeting the top 360 candidate resection activators identified in the primary screen.

the secondary screen and re-screened these duplexes individually. I ranked the 70 candidate resection activators based on the number of duplexes that caused a decrease in the percentage of pRPA32 (S4/S8)-positive S and G2 cells by more than 20% relative to the control siRNA (Fig.2.10A). Candidate activators where four out of four duplexes resulted in a pRPA32 (S4/S8) reduction were ranked highest. Within the list of highest ranking candidates was CtIP where all four duplexes decreased pRPA32 (S4/S8) nuclear intensity in S and G2 cells. All three subunits of the MRN complex had at least three of the four duplexes that decreased end resection. The incidence of siRNA off target effects in the screen was likely high as 34.3% of the top 70 candidate activators only had one siRNA duplex that decreased end resection (Fig. 2.10B). It is also possible, albeit unlikely, that only one of the four siRNA duplexes resulted in the appropriate level of knockdown to elicit a detectable resection defect. The 70 candidate resection activators and how many of the four siRNAs caused a decrease in end resection below the 0.8 cut-off are listed in Table 2.1.

2.5 Discussion

In this chapter, I described the establishment of an automated DNA end resection assay in human cells and the utilization of this assay to screen a genome-scale siRNA library. Overall, the screen was successful as evidenced by the identification of known resection activators including CtIP, MRE11, RAD50, NBS1, RPA32, RPA70, WRN, POLE3 (CHRAC17), and SRCAP (Dolganov et al., 1996; Dong et al., 2014; Featherstone and Jackson, 1998; Lan et al., 2010; Petrini et al., 1995; Sartori et al., 2007; Sturzenegger et al., 2014). In contrast, the identification of known end resection inhibitors including 53BP1, PTIP, RIF1, MAD2L2, HELB, and PIN1 was not successful (Boersma et al., 2015; Bunting et al., 2010; Chapman et al., 2013; Di Virgilio et al., 2013; Escribano-Diaz et al., 2013; Steger et al., 2013; Tkac et al., 2016; Xu et al., 2015; Zimmermann et al., 2013). One possible explanation for this discrepancy was that the dose of NCS used in the screen was too high and resulted in a level of end resection that was at the maximum detectable limit. Being at or close to the detection limit would make it difficult to observe increases in pRPA32 (S4/S8) nuclear intensity after siRNA-mediated knockdown. Therefore, I decided to focus my attention on the identified candidate resection activators.

Several known end resection activators were not identified in the primary screen including EXO1, DNA2, BLM, EXD2, SMARCAD1, SIRT6, RNF4, and RNF138 (Costelloe et

Figure 2.10. Re-screening deconvolved siRNA pools for top candidate resection activators. (A) The siRNA pools for the top 70 candidate resection activators identified in the secondary screen were deconvolved and the four distinct siRNA duplexes were transfected individually. U2OS FUCCI cells incubated with the deconvolved siRNAs were treated with 100 ng/ml NCS for 3 hours followed by pRPA32 (S4/S8) immunofluorescence and QIBC. The results are displayed as a heat-map where the rows represent each candidate resection activator and the columns represent the four individual siRNA duplexes. Candidate resection activators that had a greater number of siRNA duplexes that decreased the percentage of pRPA32 (S4/S8)-positive S/G2 cells were ranked highest. (B) Breakdown of how many siRNA duplexes for each candidate had at least a 20% decrease in the percentage of pRPA32 (S4/S8)-positive S/G2 cells compared to the control siRNA.

4 of 4 3 of 4 2 of 4 1 of 4 0 of 4 IK USPL1 SESN2 ACSM1 CAPN15 ZNF335 SF3B3 CCNC CCNI MX2 SLU7 INSM1 CREB3L4 TRIM64C RPAP2 SF3B2 DNA-PKcs POM121L2 C2ORF69 LEO1 CtIP TP53I13 TNKS1BP1 MEF2D NAA10 RAD50 MS4A7 IL27RA CDC40 WRN C11ORF35 AKNAD1 NHP2L1 MRE11 GOSR1 HIVEP1 ZNF771 DDX25 C9ORF152 NME6 ATRIP KLF17 LRRC61 ZNF32 EIF4A3 NBS1 RNF169 FAM216A C9ORF106 PHF5A POLE3 NDN RPA32 ZCCHC10 BCAS2 SRCAP SMARCE1 PSMD14 MAP1S ZNF821 C9ORF156 TRIM11 RBBP9 ASCC1 ANKRD16 FKBP10 FAM107B LEPRE1 RAB6B ZNF512B

Table 2.1. Number of candidate resection activator deconvolved siRNAs that decreased DNA end resection in S and G2 phase U2OS cells. Genes in green font are known regulators of DSB repair. Genes in orange font are characterized messenger RNA processing factors.

al., 2012; Galanty et al., 2012; Gravel et al., 2008; Ismail et al., 2015; Kaidi et al., 2010; Nimonkar et al., 2011; Nimonkar et al., 2008; Schmidt et al., 2015; Broderick et al., 2016). My results suggest that EXO1 and DNA2 were not identified as hits because the nucleases act redundantly and each can functionally compensate for the loss of the other. It is possible that the other known activators were not identified because of poor knockdown efficiency or they elicited moderate resection defects that did not make the hit cut-off. To confirm the resection activators identified in the screen, I cherry-picked siRNA pools for the top hits and re-screened them in a cell cycle phase-specific resection assay utilizing the FUCCI system. A large proportion of the cherry-picked siRNA pools elicited an accumulation of cells in G1 and did not cause a detectable defect in end resection. The high occurrence of false-positive hits due to the cell cycle may have led to real resection activators not being identified. For example, known end resection activators like SMARCAD1 and RNF138 had KS scores of -15.9 and -8.2, respectively, and may have been identified as hits if a lower false-positive rate was achieved. In the future, more candidate activators outside of the -19 KS score cut-off could be re-screened utilizing the FUCCI system.

I also screened deconvolved siRNAs for the top candidate resection activators confirmed in the secondary screen. There was likely a high degree of siRNA off-target effects in the primary screen as ~35% of the candidate resection activators demonstrated end resection defects with only one out of the four deconvolved siRNAs. Several groups have documented the high degree of off-target transcript binding that can occur in RNAi screens (Paulsen et al., 2009; Sigoillot et al., 2012; Sudbery et al., 2010). A bioinformatics study by Sigoillot et al. (2012) described a method to analyze the seven base pair seed sequences of the top scoring siRNAs in a screen to determine if there is enrichment in the binding to any specific transcript. The method was called genome-wide enrichment of seed sequence matches (GESS) and MAD2L1 was identified as a prominent off-target transcript in an RNAi screen for genes required for the spindle assembly checkpoint (Sigoillot et al., 2012). GESS analysis was also conducted for an RNAi screen searching for new regulators of DSB repair and identified an enrichment of siRNA seed sequences that corresponded to the RAD51 transcript (Adamson et al., 2012). Future work should focus on conducting the GESS analysis on the resection activators identified in my RNAi primary screen. It will be interesting to determine if there is an enrichment of siRNA seed sequences that can target known resection activators.

The deconvolved screen identified 42 high-confidence candidate resection activators where at least two of the four siRNA duplexes resulted in an end resection defect (Table 2.1). Remarkably, 11 of these resection activators are already known to function in DSB repair. Other interesting functional categories in this list included RNA splicing proteins and long non-coding RNAs (lncRNAs). Proteins containing zinc finger domains were also evident among the candidate resection activators and included ZNF335, ZNF771, INSM1, KLF17, ZCCHC10, and ZNF821. Zinc finger domains can bind to DNA and numerous characterized DSB repair proteins are known to harbour this domain (Table 3.1). ZNF335 immediately caught my attention as it was found to be mutated in a rare genetic syndrome that displays neonatal microcephaly (Yang et al., 2012). Microcephaly is a common clinical feature of patients harbouring mutations in DSB repair genes. The next chapter will focus on the functional characterization of ZNF335 in the promotion of end resection and HR.

Chapter III

The zinc finger protein, ZNF335, promotes DNA end resection

3.1 Statement of contributions, rights, and permissions

Zhen-Yuan Lin in Anne-Claude Gingras’ laboratory (LTRI) conducted all immunoprecipitation coupled to mass spectrometry experiments.

3.2 Summary

Numerous genome instability syndromes have been documented to display neonatal microcephaly as a clinical feature. The ZNF335 gene was recently found to be mutated in a syndrome that causes some of the worst cases of microcephaly ever documented. In addition, ZNF335 was consistently one of the highest scoring candidate resection activators identified in the RNAi screen outlined in Chapter II. Here, I validate the results of the screen and demonstrate that ZNF335 promotes DNA end resection. I uncover that ZNF335-deficient cells are sensitive to agents that induce DSBs and that ZNF335 can promote DSB repair by HR. I show that the four C-terminal zinc finger domains of ZNF335 are required for its function in end resection. In addition, I show that ZNF335 is recruited to sites of DNA damage generated by laser microirradiation and this accumulation is short-lived and dependent on the activity of PARP. However, I provide evidence that PARP activity is not required for end resection. This data suggests that ZNF335 recruitment to sites of laser microirradiation is not necessary for its function in promoting end resection. Lastly, I show that ZNF335 does not regulate the expression or protein stability of the core end resection factors. I conclude that ZNF335 is a new factor that can promote end resection and HR.

3.3 Introduction

Zinc finger protein domains are found throughout evolution, from bacteria to humans. Approximately 3% of the human genome encodes proteins that contain zinc finger domains and most of these proteins are completely uncharacterized. Zinc fingers are small self-contained domains that are stabilized by one or more zinc ions and have the capacity to bind both nucleic acids (RNA and DNA) and proteins (Laity et al., 2001). For nucleic acid binding, zinc fingers are usually present as repetitive nucleotide-binding modules giving them the unique ability to specifically bind longer stretches of nucleotides. Numerous zinc finger domain types have been discovered but here I will focus on the classical 22-25 amino acid C2H2 finger which is stabilized by a single zinc ion bound to a pair of cysteines and a pair of histidines. The two cysteines and two histidines are fundamental for zinc binding and proper folding of the zinc finger (Lee et al., 1989). In addition, C2H2 zinc fingers contain three other conserved amino acids in the region between the last cysteine and first histidine. These three amino acids (tyrosine, phenylalanine, and leucine) form a hydrophobic structural core which is also crucial for the folding of the zinc finger module (Lee et al., 1989). The structure of a single zinc finger consists of an antiparallel β-sheet with a loop formed by the two cysteines and an α-helix with a loop formed by the two histidines (Pavletich and Pabo, 1991). These two structural units are held together by the zinc ion. The zinc finger binds DNA through its α-helix where it forms hydrogen bonds at helical positions -1, 3, and 6 to a triplet sequence of nucleotides on one strand of DNA (Pavletich and Pabo, 1991). Later it was found that helical position 2 can also interact with a nucleotide on the opposite strand of DNA (Fairall et al., 1993). Collections of zinc finger mutants have been generated and aided in the formulation of rules that related particular amino acids (at helical positions -1, 2, 3, and 6) to four corresponding nucleotides (Klug, 2010). However, the rules relied on DNA being in the canonical B form and strict adherence to the rules did not always correlate with successful binding to other DNA sequences. A more powerful method to engineer zinc finger domains that can bind specific DNA sequences is to utilize affinity selection from libraries of zinc finger mutants by phage display (Choo and Klug, 1994).

Zinc fingers are the most common domain in the entire human proteome and thus have been implicated in a diverse set of cellular processes including transcription, mRNA processing, translation, protein-protein interactions, and post-translational modifications (Laity et al., 2001). It is not surprising that numerous human developmental disorders and disease states have been

linked to zinc finger protein dysfunction. Importantly, many DSB repair factors employ zinc finger domains to carry out their function at sites of DNA damage. These domains facilitate several functional properties to promote DNA repair including DNA binding, poly-ADP ribose (PAR) binding, protein-protein interactions, and substrate SUMOylation and ubiquitylation. An outline of DSB repair factors that possess zinc finger domains can be found in Table 3.1.

Several candidate resection activators identified in the RNAi screen (outlined in Chapter II) harbour zinc finger domains, including ZNF335. The human ZNF335 gene is located on chromosome 20 and contains 28 exons that encode a 1,342 amino acid protein. ZNF335 appears to be vertebrate-specific and contains 13 C2H2-type zinc finger domains that are dispersed throughout the protein (see protein schematic in Figure 3.7A). ZNF335 was first identified as a coactivator of nuclear hormone receptor signaling through an interaction with nuclear receptor coregulatory (Mahajan et al., 2002b). Nuclear hormone receptors are ligand-dependent transcription factors that control gene expression programs for numerous physiological, developmental, and metabolic processes (Aranda and Pascual, 2001). More recently, the ZNF335 gene was found to be mutated in a rare genetic syndrome that causes one of the most severe cases of microcephaly ever documented (Yang et al., 2012). The syndrome was reported in a large consanguineous Arab Israeli pedigree where seven individuals displayed neonatal microcephaly with head circumferences as small as nine standard deviations below the mean. Mapping using single-nucleotide polymorphism arrays identified a single 2 megabase region that was homozygous in all affected pedigree members. Sequencing of this region determined the presence of 40 genes but only one homozygous nonsynonymous change – a G to A transition at nucleotide position 3332 in the coding sequence of the ZNF335 gene. The c.3332g>a mutation resulted in an amino acid change from an arginine to a histidine at position 1111 which is located in the final (thirteenth) zinc finger domain. The mutation is also located at the final position of the splice donor site for exon 21 and resulted in the accumulation of a larger transcript due to intron retention. RNA-sequencing experiments determined that the large transcript retains the two introns flanking exon 21. Patient cells also expressed messenger RNA of the expected size which suggested that some normal splicing did occur. Immunoblotting of whole cell extracts from patient lymphoblast cells showed severely reduced ZNF335 protein at the expected size which likely resulted from translation of normally spliced messenger RNA. The antibody used by Yang et al. (2012) was produced by injecting rabbits with a peptide

DSB repair factor Zinc finger DSB Function Reference type repair pathway PARP-1 PARP-type HR, NHEJ Early recruitment of several DSB repair (Beck et al., factors (i.e. APLF) 2014) APLF PBZ-type NHEJ Nuclease component of DNA ligase IV (Iles et al., complex 2007) KAT5 (TIP60) C2HC-type HR, NHEJ Acetylates ATM after DNA damage to (Ikura et al., promote repair and checkpoint 2000) activation TOP3A GRF-type HR Component of BLM complex that (Wu and promotes Holliday junction dissolution Hickson, 2003) INO80B HIT-type HR, NHEJ Component of chromatin remodeling (Morrison et al., complex that relaxes chromatin 2004) structure at DSB sites ZNHIT1 HIT-type HR Component of SRCAP chromatin (Dong et al., remodeling complex that promotes end 2014) resection PIAS1 MIZ-type HR, NHEJ E3 SUMO ligase that promotes (Galanty et al., RNF8/RNF168-dependent 2009) ubiquitylation at DSB sites PIAS4 MIZ-type HR, NHEJ E3 SUMO ligase that promotes (Galanty et al., RNF8/RNF168-dependent 2009) ubiquitylation at DSB sites HERC2 ZZ-type HR, NHEJ HECT E3 ubiquitin ligase that (Bekker-Jensen promotes RNF8/RNF168-dependent et al., 2010) ubiquitylation at DSBs TRIM28 (KAP-1) PHD-type HR, NHEJ Important for ATM-dependent DSB (Ziv et al., repair in heterochromatin 2006) BRCA1 RING-type HR Critical factor involved in resection and (Huen et al., RAD51 loading 2010) RNF4 RING-type HR (SUMO)-targeted E3 ubiquitin ligase (Galanty et al., that regulates RPA turnover at resected 2012; Yin et al., DSBs 2012) RNF8 RING-type HR, NHEJ E3 ubiquitin ligase that promotes the (Huen et al., recruitment of critical repair factors to 2007; Kolas et DSB sites al., 2007; Mailand et al., 2007) RNF168 RING-type HR, NHEJ E3 ubiquitin ligase that promotes the (Doil et al., recruitment of critical repair factors to 2009; Stewart et DSB sites al., 2009) RNF138 RING-type HR E3 ligase that ubiquitylates CtIP and (Ismail et al., promotes its recruitment to DSB sites 2015; Schmidt et al., 2015)

Table 3.1. DSB repair proteins harbouring zinc finger domains.

corresponding to the final 42 C-terminal amino acids of ZNF335. Intron retention before and after exon 21 would result in a premature stop codon and a protein product that does not contain the C-terminal region that the antibody was raised against. Therefore, it is possible that this truncated ZNF335 protein accumulated in patient cells but was simply not detected by the authors.

After characterizing the mutation in human ZNF335, Yang et al. (2012) engineered null Znf335 mutations in mice and determined that homozygous loss of Znf335 led to early embryonic lethality at day 7.5 (E7.5). The essential requirement of Znf335 in mice pointed to the hypothesis that the human c.3332g>a mutation may be hypomorphic as affected patients were born to term, albeit with severe clinical symptoms including microcephaly and small birth weight and length. The potential hypomorphic nature of this mutation could be explained by the presence of a small amount of normally sized R1111H-mutated ZNF335 protein that was detected in patient cells. Conditional knockdown of Znf335 in the developing mouse brain caused defects in neural progenitor proliferation (Yang et al., 2012). Furthermore, the low number of Znf335-depleted neurons that did form in the cerebral cortex had abnormal neuronal morphology including small cell bodies and a lack of vertical apical dendritic processes. Importantly, these neuronal phenotypes were also observed post-mortem in affected human patients. Immunoprecipitation coupled to mass spectrometry (IP-MS) uncovered that human ZNF335 interacted with components of the trithorax group of proteins including MLL, SETD1A, ASH2L, RBBP5, and WDR5. The trithorax complex functions in histone methylation and transcriptional activation (Schuettengruber et al., 2011). Chromatin immunoprecipitation followed by next-generation sequencing in early mouse embryos determined that ZNF335 bound to many gene promoters and promoted H3K4 methylation (Yang et al., 2012). ZNF335 was associated with the promoter of the repressor element 1 (RE1)-silencing transcription factor (REST) gene which is a critical epigenetic regulator of neurogenesis. REST acts as a transcriptional repressor by recruiting histone deacetylases (HDACs) and is expressed in neural stem cells to maintain progenitor cell fate and inhibit differentiation into neurons (Ballas et al., 2005). The expansion and maintenance of neural progenitor cells is critical for later neurogenesis. Yang et al. (2012) surmised that ZNF335 could function upstream of REST by promoting its expression. Importantly, it was not ruled out that neural progenitor cells lacking ZNF335 could have cell proliferation defects due to other factors including enhanced apoptosis

71 or activation of cell cycle checkpoints. Proliferation defects due to aberrant mitosis or excessive DNA damage are of particular interest because most genetic syndromes that display neonatal microcephaly are caused by mutations in genes regulating centrosome and DNA repair biology (Thornton and Woods, 2009). In the following study, I demonstrate that ZNF335 promotes DNA end resection and DSB repair by HR. I argue that the function of ZNF335 in DSB repair may contribute to the severe microcephaly observed in patients with the c.3332g>a mutation.

3.4 Results

3.4.1 Analysis of four siRNA duplexes targeting the ZNF335 messenger RNA

The four ZNF335 siRNAs used in this study have identical sequences to those used in the deconvolved confirmation screen. I first determined the efficiency of each siRNA duplex in depleting the ZNF335 transcript and protein in U2OS cells. I conducted reverse transcription and subsequent quantitative PCR using RNA extracted from U2OS cells incubated with ZNF335 siRNAs. Quantitative PCR was conducted with a primer and probe set specific for ZNF335 (Solaris/GE Healthcare). The relative abundance of ZNF335 transcript in each sample was determined by normalizing to the abundance of the reference gene GAPDH. Compared to cells treated with the scrambled control siRNA, the abundance of ZNF335 messenger RNA was reduced by at least 2-fold with 3 out of the 4 siRNAs (Fig. 3.1A). Knockdown efficiency was confirmed at the protein level by western blot analysis utilizing a commercial ZNF335 antibody that targets the C-terminus (Fig. 3.1B). I then conducted a growth rate analysis for U2OS cells treated with the four ZNF335 siRNA duplexes. U2OS cells were reverse-transfected in 6-well plates and the number of cells was manually quantified at various time points (Fig. 3.1C). The growth of U2OS cells incubated with ZNF335 siRNAs correlated with the observed knockdown efficiency. Next, I determined the cell cycle profile of U2OS cells treated with the siRNAs utilizing propidium iodide (PI) staining and flow cytometry (Fig. 3.1D). The cell cycle profile of cells treated with the control siRNA and those treated with ZNF335 siRNA #1 and 3 were similar. Cells incubated with siRNA #2 had a small decrease in the proportion of cells in G1 whereas siRNA #4 had a small increase. Overall, these small differences in cell cycle position cannot account for the end resection defect observed for ZNF335 knockdown in the RNAi screen.

Figure 3.1. Knockdown efficiency, growth, and cell cycle position analysis for siRNA duplexes targeting ZNF335 messenger RNA. (A) U2OS cells were transfected with the indicated siRNAs and incubated for 48 hours. Total RNA was extracted from cells and cDNA was synthesized by reverse transcription. Quantitative real-time PCR was conducted using Solaris primers and probe (GE Healthcare) specific to ZNF335 and the reference gene GAPDH (Mean ± SEM; N ≥ 3). (B) U2OS cells incubated with siRNAs targeting ZNF335 were processed for immunoblotting using a ZNF335 specific antibody. (C) U2OS cells incubated with ZNF335 siRNAs were counted at various time points post- transfection to assess their growth rates. (D) Cell cycle profiles for U2OS cells incubated with each ZNF335 siRNA duplex. Cell cycle position was determined by measuring DNA content by staining nuclei with propidium iodide before flow cytometry analysis.

3.4.2 ZNF335 depleted cells have defective pRPA32 (S4/S8), RPA32, and BrdU focus formation

I confirmed the results obtained from the screen by monitoring pRPA32 (S4/S8) focus formation in U2OS FUCCI cells depleted of ZNF335 (Fig. 3.2A). U2OS FUCCI cells were seeded and reverse-transfected with control or ZNF335 siRNA. After 48 hours to allow time for knockdown, cells were treated with 100 ng/ml NCS for 3 hours and then processed for pRPA32 (S4/S8) immunofluorescence. Compared to the control, knockdown of ZNF335 resulted in less pRPA32 (S4/S8) foci per nucleus in S and G2 phase cells. The defect in RPA32 phosphorylation could also be detected by western blot analysis (Fig. 3.2D). To demonstrate that ZNF335 knockdown specifically decreased end resection and not just RPA32 phosphorylation I also processed cells for RPA32 and BrdU immunofluorescence (Fig. 3.2B,C). The depletion of ZNF335 decreased both RPA32 and BrdU focus formation compared to the control siRNA.

3.4.3 U2OS cells depleted of ZNF335 are sensitive to DSB-inducing agents

The observation that ZNF335 is important for end resection (and possibly DSB repair) led me to hypothesize that cells lacking ZNF335 may be more sensitive to incubation with agents that induce DSBs. To test this, I conducted clonogenic survival assays (Franken et al., 2006) on U2OS cells treated with control, CtIP, or ZNF335 siRNAs. After 48 hours of siRNA incubation, I sparsely seeded multiple densities of the siRNA-treated cells into 6-well plates and then exposed the cells (for 1 hour) to various drugs that cause DSBs, including NCS, ETOP, and CPT. I allowed individual colonies to grow for two weeks and then fixed and stained them with methanol and crystal violet, respectively. First, I calculated the plating efficiency of cells treated with each of the siRNAs (and no DSB-inducing drugs) using the following formula:

100

Next, I counted the number of colonies on each plate exposed to the various concentrations of the DSB-inducing drugs and calculated the surviving fraction for each treatment with the following formula:

Figure 3.2. ZNF335 is an activator of DNA end resection. (A) U2OS FUCCI cells incubated with scrambled control, CtIP, or ZNF335 siRNA were treated with 100 ng/ml NCS for 3 hours followed by pRPA32 (S4/S8) immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. The percentage of S and G2 U2OS FUCCI cells with more than 15 pRPA32 (S4/S8) foci was quantified (Mean ± SEM; N ≥ 3). Scale bars represent 5 µm. (B) U2OS cells transfected with control, CtIP, or ZNF335 siRNA were treated with 50 ng/ml NCS for 3 hours followed by pre-extraction and RPA32 immunostaining. The percentage of cells with more than 15 RPA32 foci was quantified (Mean ± SEM; N ≥ 3). (C) U2OS cells incubated with BrdU and control, CtIP, or ZNF335 siRNA were treated with 50 ng/ml NCS for 3 hours followed by pre-extraction and BrdU immunofluorescence. The percentage of cells with more than 15 BrdU foci was quantified (Mean ± SEM; N ≥ 3). (D) U2OS cells incubated with control, CtIP, or ZNF335 siRNA were treated with 100 ng/ml NCS for 3 hours and then processed for pRPA32 (S4/S8), RPA32, CtIP, ZNF335, and GAPDH immunoblotting.

There was only a small decrease in the plating efficiency for U2OS cells depleted of CtIP and ZNF335 compared to the control siRNA (Fig. 3.3A). As reported previously (Sartori et al., 2007), cells depleted of CtIP were substantially more sensitive (decreased surviving fraction) to DSB-inducing drugs compared to cells incubated with the control siRNA. In addition, ZNF335 depleted cells were also sensitive to DSB-inducing drugs but to a lesser degree than cells treated with CtIP siRNA (Fig. 3.3B). Therefore, these data further support a role for ZNF335 in promoting DSB repair.

3.4.4 Depletion of ZNF335 decreases the efficiency of DSB repair by HR

To assess HR efficiency, I established an automated assay to quantify the intensity of the recombinase RAD51 at DSB sites by immunofluorescence. The detection of RAD51 loading at resected DSBs can act as a surrogate for measuring the relative number of DSBs that are undergoing HR. Before being processed for RAD51 and γH2AX immunostaining, cells were treated with 5 Gy of ionizing radiation (IR) and were allowed to recover for three hours. Cells were imaged on an automated confocal microscope and DSBs (γH2AX foci) were detected using image analysis software. The intensity of RAD51 immunostaining was quantified at each γH2AX focus. To test if the depletion of ZNF335 affected RAD51 focus formation I seeded and reverse-transfected U2OS cells with control, BRCA2, CtIP, or ZNF335 siRNA in a 96-well plate. After 48 hours of siRNA incubation, cells were exposed to IR followed by a three hour recovery period to allow time for end resection and subsequent RAD51 loading. Cells were then processed for RAD51 and γH2AX immunofluorescence (Fig. 3.4A). In line with CtIP knockdown, U2OS cells transfected with siRNAs targeting ZNF335 had a reduction in the intensity of RAD51 at DSB sites compared to the control siRNA (Fig. 3.4B,C).

I also utilized the well-established Direct Repeat (DR)-GFP reporter system to measure HR in ZNF335 depleted cells. Briefly, DR-GFP U2OS cells were previously engineered to have a single I-SceI endonuclease consensus site surrounded by two non-functional GFP alleles integrated into their genome (Pierce et al., 1999). Once expression of I-SceI is introduced, a DSB

Figure 3.3. ZNF335 deficient cells are sensitive to DSB-inducing agents. U2OS cells incubated with scrambled control, CtIP, or ZNF335 siRNA #3 were seeded into 6-well plates at various densities: 250, 500, 1000, 2000, 4000, and 8000 cells per well. The day after seeding, cells were treated with various concentrations of NCS, ETOP, or CPT for 1 hour and then allowed to grow until colonies were visible by eye (~2 weeks). Colonies were fixed, stained with crystal violet, and then counted. Only seeding densities that resulted in a manageable number of colonies (1-100) were counted. (A) The plating efficiency (PE) was calculated for each siRNA (PE = number of colonies formed / number of cells seeded) (Mean ± SEM; N ≥ 3). (B) The surviving fraction (SF) was then calculated for each drug treatment (SF = number of colonies formed after treatment / number of cells seeded x PE). The surviving fraction for each treatment was plotted using a log scale (Mean ± SEM; N ≥ 3).

Figure 3.4. ZNF335 promotes DSB repair by HR. (A) U2OS cells incubated with scrambled control, BRCA2, CtIP, or ZNF335 siRNA were treated with 5 grays ionizing radiation and 3 hours later were immunostained for RAD51 and γH2AX. DNA was counterstained with DAPI to visualize nuclei. Scale bar represents 5 µm. (B) Cells were imaged on an automated confocal microscope and using image analysis software γH2AX foci were segmented and the intensity of RAD51 at each focus was measured. (C) A RAD51 focus intensity threshold of 1500 was applied to calculate the percentage of RAD51- positive γH2AX foci or DSBs (Mean ± SEM; N ≥ 3). (D) U2OS DR-GFP cells were incubated with siRNAs and then transfected with an I-SceI expressing plasmid in 96-well plates. After 48 hours of I-SceI expression, cells were fixed and DNA was stained with DAPI. Cells were imaged on an automated confocal microscope and the percentage of GFP-positive cells was calculated using image analysis software and a GFP intensity threshold (Mean ± SEM; N ≥ 3).

will be created between the two GFP sequences. If properly repaired by HR, the two sequences will be joined and the U2OS cells will express a functional GFP (Fig. 3.4D). I established an automated method in 96-well plates to determine the number of GFP-positive U2OS cells by QIBC. To examine if ZNF335 promotes the efficiency of DSB repair by HR, I seeded and reverse-transfected U2OS DR-GFP cells with control, BRCA2, CtIP, or ZNF335 siRNA in a 96- well plate. After 24 hours, I forward-transfected pCBASceI (Addgene #26477) to introduce the expression of I-SceI. At 48 hours post-I-SceI transfection, I fixed the cells and stained the DNA with DAPI. Cells were imaged on an automated confocal microscope and the percentage of GFP- positive nuclei was determined using image analysis software. Compared to control siRNA, U2OS DR-GFP cells depleted of ZNF335 had approximately a two-fold reduction in the percentage of cells that were GFP-positive, suggesting a defect in the completion of HR (Fig. 3.4D).

3.4.5 ZNF335 promotes the phosphorylation of CHK1 at a characterized ATR consensus site

As discussed in Chapter I, the DNA damage responsive kinase, ATR, is activated by the presence of RPA-coated ssDNA (Zou and Elledge, 2003). An enrichment of this genomic structure can arise through various mechanisms, namely during times of replication stress and as a consequence of end resection at DSB sites during HR. It has been demonstrated that defects in end resection can lead to decreased ATR activation (Jazayeri et al., 2006). Therefore, I posited that cells depleted of ZNF335, in addition to having defects in end resection, may demonstrate a decreased ability to activate ATR. The DNA damage checkpoint kinase, CHK1, is phosphorylated by ATR on two well characterized sites, serine 317 and 345 (Zhao and Piwnica- Worms, 2001). One common method to assay ATR activation is to detect these phosphorylation events in cellular extracts utilizing phospho-specific antibodies. To determine if cells depleted of ZNF335 have a defect in CHK1 phosphorylation by ATR, I seeded and reverse-transfected U2OS cells with control, CtIP, or ZNF335 siRNA. At 48 hours-post transfection, I treated cells with NCS for three hours followed by cellular lysis and western blot analysis with an antibody targeting phospho-CHK1 (S345). Remarkably, cells depleted of ZNF335 had a defect in the phosphorylation of serine 345 on CHK1, similar to that observed upon CtIP knockdown (Fig. 3.5A). In ZNF335 depleted cells, defective ATR activation was due to decreased end resection and not aberrant replication-associated ATR activation as treatment with the replication stressor,

Figure 3.5. ZNF335 promotes CHK1 serine 345 phosphorylation. (A) Whole cell extracts from U2OS cells incubated with scrambled control, CtIP, or ZNF335 siRNA and treated with 50 ng/ml NCS for 3 hours were separated by SDS-PAGE and examined by pCHK1 (S345), CtIP, ZNF335, or GAPDH immunoblotting. (B) U2OS cells incubated with scrambled control, CtIP, or ZNF335 siRNA and treated with 10 mM HU for 30 minutes were processed for pCHK1 (S345), CtIP, ZNF335, or GAPDH immunoblotting.

HU, elicited normal levels of CHK1 phosphorylation (Fig. 3.5B).

3.4.6 Defective end resection in ZNF335 depleted cells can be rescued by expressing an RNAi-resistant open reading frame

To exclude the possibility that the observed defect in end resection was due to an off-target effect of the siRNA, I cloned the ZNF335 open reading frame (ORF) with a N-terminal GFP tag into the lentiviral inducible expression vector, pCW57.1 (Root lab; Addgene #41393). I then generated a siRNA-resistant ORF by creating silent mutations in the binding site for siRNA #3 by PCR mutagenesis. Next, I packaged this construct into lentivirus, infected U2OS cells, and selected transduced cells with puromycin to generate a stable cell line. In response to induction by doxycycline, GFP-ZNF335 accumulated entirely in the nucleus and was devoid in the nucleolus (Fig. 3.6A). Furthermore, exogenous GFP-tagged ZNF335 protein ran at the correct size by western blot (~250 kDa) and was completely insensitive to siRNA #3 (Fig. 3.6B). In addition, expression of siRNA-resistant GFP-ZNF335 completely rescued the end resection defect observed upon ZNF335 depletion with siRNA #3 using the pRPA32 (S4/S8) QIBC resection assay described in Chapter II (Fig. 3.6C). This data further validates that ZNF335 promotes end resection and that the observed phenotype is not a result of an off-target effect of the siRNA.

3.4.7 The function of ZNF335 in DNA end resection is dependent on its four C-terminal C2H2 zinc finger domains

In order to determine what regions of the ZNF335 protein were required for end resection, I generated C- and N-terminal deletions of FLAG-ZNF335 by PCR in the pCW57.1 expression vector (Fig. 3.7A). I packaged these constructs into lentivirus and produced U2OS stable cell lines by transduction and puromycin selection. Remarkably, all constructs that contained the four C-terminal C2H2 zinc fingers were able to rescue the end resection defect observed upon ZNF335 knockdown. In a phospho-proteomics study, ZNF335 was shown to be phosphorylated in response to DNA damage at Serine 416 which is an ATM/ATR consensus site (Matsuoka et al., 2007). In addition, efficient end resection at DSB ends requires the activity of ATM (Jazayeri et al., 2006). Therefore, I hypothesized that ATM-dependent phosphorylation of serine 416 on ZNF335 could be important for its function in promoting end resection. To examine this

Figure 3.6. Expression of siRNA-resistant ZNF335 rescues the observed end resection defect in ZNF335 depleted cells. (A) U2OS cells infected with lentivirus packaged with an empty vector (EV) or a vector with full-length (FL) GFP- tagged ZNF335 containing silent mutations to render it insensitive to RNAi. In transduced cells, the pCW57.1 lentiviral expression vector is inducible by the addition of doxycycline (DOX). Infected U2OS cells were incubated with 5 µg/ml doxycycline (+DOX) or without doxycycline (-DOX) for 24 hours and then fixed. DNA was stained with DAPI to visualize nuclei. (B) U2OS cells infected with lentivirus as in (A) and incubated with control or ZNF335 siRNA before GFP and GAPDH immunoblotting. (C) U2OS cells transduced with siRNA-resistant GFP- ZNF335 were seeded in a 96-well plate and incubated with scrambled control or ZNF335 siRNA. Cells were treated with 100 ng/ml NCS for 3 hours before processing for pRPA32 (S4/S8) and Cyclin A immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. QIBC using an automated confocal microscope and image analysis software was conducted to segment DAPI-stained nuclei and determine the nuclear intensities of Cyclin A and pRPA32 (S4/S8). The bar graph depicts the percentage of S and G2 cells (Cyclin A-positive) with pRPA32 (S4/S8) nuclear intensities over the mean nuclear background intensity (calculated in cells not treated with NCS) (Mean ± SEM; N ≥ 3).

Figure 3.7. The four C-terminal C2H2 zinc finger domains of ZNF335 are sufficient for its function in end resection. (A) U2OS cells stably expressing the Geminin-mAG FUCCI reporter were infected with full-length (FL) FLAG- tagged ZNF335 or the illustrated ZNF335 mutants. Transduced cells were seeded in a 96-well plate and incubated with either scrambled control or ZNF335 siRNA. The next day, cells were treated with either 5 µg/ml doxycycline (+DOX) or without doxycycline (-DOX) for 24 hours. Cells were treated with 100 ng/ml NCS for 3 hours and processed for pRPA32 (S4/S8) immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. QIBC using an automated confocal microscope and image analysis software was conducted to segment DAPI-stained nuclei and determine the nuclear intensities of Geminin-mAG and pRPA32 (S4/S8). The bar graph depicts the percentage of S and G2 cells (Geminin-mAG-positive) with pRPA32 (S4/S8) nuclear intensities over the mean nuclear background intensity (calculated in cells not treated with NCS) (Mean ± SEM; N ≥ 3). (B) Transduced U2OS cells were also processed for FLAG immunostaining to examine the subcellular localization of each ZNF335 mutant.

possibility, I created a serine-to-alanine mutant at amino acid 416 by PCR mutagenesis. The S416A mutant was able to complement the depletion of endogenous ZNF335 suggesting that phosphorylation at this site is not required for the function of ZNF335 in end resection (Fig. 3.7A).

The ZNF335 microcephaly c.3332g>a mutation resulted in an amino acid change from arginine to histidine at position 1111 in the thirteenth C2H2 zinc finger domain (Yang et al., 2012). In addition, the mutation caused aberrant splicing and intron retention which resulted in drastically lower ZNF335 protein levels in patient cells. I created the R1111H amino acid change by PCR mutagenesis in the FLAG-ZNF335 ORF and produced a stable cell line by lentiviral transduction. Expression of the ZNF335 R1111H mutant was able to rescue the end resection defect observed upon siRNA-mediated depletion. The stability of the overexpressed R1111H mutant was similar to that of the wild type protein (Fig. 3.7B). This data suggested that any ZNF335 R1111H protein expressed in patient cells can function in the promotion of end resection. Importantly, these findings did not rule out that patient cells could have defective end resection due to aberrant ZNF335 mRNA splicing and resulting low protein levels. The exogenously expressed ZNF335 R1111H mutant was translated from a messenger RNA that did not contain introns. Therefore, the effect of the c.3332g>a mutation on RNA splicing cannot be examined using this experimental system.

All ZNF335 mutants accumulated entirely in the nucleus except for the Δ271-1342 deletion which localized primarily in the cytoplasm (Fig. 3.7B). I analyzed the ZNF335 ORF for a nuclear localization signal (NLS) and found one canonical monopartite NLS from amino acid 352 to 361 (GRPRKLPRLE). Not surprisingly, the Δ271-1342 mutant does not contain this NLS. However, N-terminal deletions not containing this sequence were able to accumulate in the nucleus, suggesting that there may be another more cryptic NLS in the C-terminus or that ZNF335 can potentially shuttle into the nucleus with an interacting protein.

3.4.8 ZNF335 localizes to sites of DNA damage generated by laser microirradiation

As depletion of ZNF335 resulted in an end resection defect and sensitivity to DNA damaging agents, I posited that ZNF335 may be able to localize to sites of DSBs in order to elicit its function. To test this hypothesis, I treated U2OS cells expressing GFP-ZNF335 with various

DSB-inducing agents and then processed the cells for γH2AX immunofluorescence to mark DSB sites. Using various doses, incubation times, and two DSB-inducing drugs (NCS and ETOP) I was unable to detect any co-localization of GFP-ZNF335 with γH2AX foci (Fig. 3.8A). Next, I utilized a U2OS DSB reporter system in which a mCherry-LacI-FokI nuclease fusion protein creates DSBs within a single genomic locus that is engineered to contain 256 lacO elements (Tang et al., 2013). The FokI fusion protein is equipped with a destabilization domain from a modified estrogen receptor which allows for inducible nuclease expression after the addition of the small molecules Shield-1 and 4-hydroxytamoxifen (4-OHT). Four hours after small molecule addition, I did not detect any GFP-ZNF335 enrichment at the mCherry-LacI-FokI focus (Fig. 3.8B). In contrast, I observed robust localization of GFP-CtIP, suggesting that FokI is in fact generating DSBs at the lacO array.

Laser microirradiation is a technique to increase the local concentration of DSBs in chromatin by using a 355 nm laser to generate a ‘stripe’ of DNA damage across a nucleus (Nelms et al., 1998; Rogakou et al., 1999). In addition to DSBs, microirradiation also results in SSBs and other DNA lesions (Kong et al., 2009). I conducted laser microirradiation on the nuclei of U2OS cells expressing GFP-ZNF335 and observed a robust accumulation to the stripe within 5 minutes (Fig. 3.8C). To test if ZNF335 accumulation at stripes is ATM-dependent, I pre- treated U2OS cells expressing GFP-ZNF335 with 10 µM of the ATM inhibitor KU55933 for 1 hour before laser microirradiation. In all 15 microirradiated nuclei tested, I observed a robust recruitment of GFP-ZNF335 to stripes that was comparable to cells treated with DMSO alone (Fig. 3.9A). In contrast, the accumulation of GFP-ZNF335 at stripes was completely dependent on the activity of PARP, as I did not detect recruitment in U2OS cells pre-treated with 100 nM of the PARP-1/2 inhibitor olaparib (Fig. 3.9A). PARP-1 is rapidly recruited to sites of laser microirradiation where it catalyzes the synthesis of PAR (Mortusewicz et al., 2007). At stripes, the PAR modification is short-lived and is actively removed within minutes by PARG (Mortusewicz et al., 2011). Therefore, factors recruited in a PARP-dependent manner generally associate with sites of DNA damage in a transient manner or are stabilized there by other interactions. To examine the temporal dynamics of GFP-ZNF335 localization to laser stripes I followed 15 nuclei for 30 minutes post-microirradiation, taking images and quantifying GFP- ZNF335 stripe intensity at the 0, 5, and 30 minute time points (Fig. 3.9B). The localization

Figure 3.8. ZNF335 accumulates at sites of laser microirradiation. (A) U2OS cells infected with lentivirus packaged with pCW57.1 containing the full-length (FL) GFP-tagged ZNF335. Cells were incubated with 5 µg/ml doxycycline for 24 hours and then treated with either 50 ng/ml NCS or 1 µM ETOP for 1 hour before γH2AX immunofluorescence. DNA was counterstained with DAPI to visualize nuclei. (B) U2OS mCherry-LacI-FokI cells were infected with lentivirus packaged with pCW57.1 containing either GFP-CtIP or GFP-ZNF335. Cells were incubated with 5 µg/ml doxycycline for 24 hours followed by treatment with Shield-1 and 4-OHT for 4 hours to induce DSBs at the lacO array. Cells were fixed and DNA was stained with DAPI to visualize nuclei. (C) U2OS cells were infected with lentivirus packaged with pCW57.1 containing the full- length (FL) GFP-tagged ZNF335. Cells were incubated with 5 µg/ml doxycycline for 24 hours and a ‘stripe’ of DNA damage was generated across several nuclei using a confocal microscope and a 355 nm laser. After 5 minutes post-microirradiation, cells were processed for γH2AX immunostaining. Scale bars represent 5 µm.

Figure 3.9. The accumulation of ZNF335 at laser stripes is PARP-dependent. (A) U2OS cells expressing GFP-ZNF335 and pre-treated with 1 µM of the ATM inhibitor KU55933 or 100 nM of the PARP inhibitor olaparib for 1 hour before microirradiation. Laser stripes were generated on a confocal microscope with a 355 nm laser. After 5 minutes post-DNA damage, images of live U2OS GFP-ZNF335 cells were captured. GFP-ZNF335 ‘stripe’ intensity was quantified for 15 nuclei using image analysis software and dividing the mean stripe intensity by the mean nuclear background intensity. (B) U2OS cells expressing GFP-ZNF335 were microirradiated and stripe intensities were quantified for 15 nuclei at various time intervals post-DNA damage. Scale bars represent 5 µm.

of GFP-ZNF335 to sites of laser microirradiation was transient as it decreased over the time course and was undetectable in most nuclei at 30 minutes.

Next, I tested the ability of GFP-tagged ZNF335 deletion mutants (first outlined in Fig. 3.7A) to be recruited to sites of laser microirradiation (Fig. 3.10A). All of the N- and C-terminal ZNF335 deletions were able to localize to laser stripes at 5 minutes-post microirradiation. I quantified stripe intensities for 10 microirradiated nuclei for each ZNF335 deletion and determined that recruitment was enhanced in the mutants that contained more C2H2 zinc finger domains (Fig. 3.10B). Mutants that did not possess the four C-terminal zinc finger domains were still able to localize to sites of microirradiation. This is in contrast to functional assays, where the C-terminus of ZNF335 was required for end resection (Fig. 3.7A). I surmised that if ZNF335 recruitment to DSB sites was required for its function in end resection then treatment of cells with olaparib should result in a resection defect. To test this, I pre-treated U2OS cells with olaparib for 30 minutes before being incubated in media containing both NCS and olaparib for three hours. I then processed the cells for pRPA32 (S4/S8) immunofluorescence and conducted QIBC to measure end resection. I did not observe an end resection defect with any of the olaparib concentrations tested (Fig. 3.10C). Moreover, an increase in end resection was detected in U2OS cells incubated with a PARP-1 siRNA. Therefore, these data suggested that the PARP- dependent recruitment of GFP-ZNF335 to sites of DNA damage was not required for its function in resecting DSB ends. It is possible that ZNF335 recruitment to stripes is required for another unknown function or that it is an artefact of the laser microirradiation technique.

3.4.9 ZNF335 depletion does not affect the expression of the core end resection activators

ZNF335 was previously implicated in the regulation of gene expression by activating transcription as a component of the trithorax complex (Garapaty et al., 2009; Yang et al., 2012). Therefore, I posited that ZNF335 could modulate end resection by regulating gene expression or steady-state protein levels of one or more core resection activators. To test this, I treated cells with control or ZNF335 siRNA and monitored protein levels of core resection activators by western blot analysis. I did not detect differences between U2OS cells incubated with control or ZNF335 siRNA in the protein levels of any core activator, including CtIP, MRE11, RAD50, NBS1, EXO1, DNA2, or BLM (Fig. 3.11A-E). Importantly, no trithorax subunits were identified as candidate resection activators in the pRPA32 (S4/S8) genome-scale RNAi primary screen

Figure 3.10. The PARP-dependent accumulation of ZNF335 at sites of microirradiation is not required for its function in DNA end resection. (A) U2OS cells expressing full-length (FL) or deletion mutants of GFP-tagged ZNF335 five minutes post- microirradiation with a 355 nm laser. Scale bar represents 5 µm. (B) GFP-ZNF335 stripe intensity quantification for (A). (C) U2OS FUCCI cells incubated with siRNA before a 3 hour NCS treatment or pre-treated with various concentrations of olaparib for 1 hour before incubation with olaparib and NCS for 3 hours. Cells were then processed for pRPA32 (S4/S8) immunofluorescence and DNA was counterstained with DAPI to visualize nuclei. QIBC using an automated confocal microscope and image analysis software was conducted to segment DAPI- stained nuclei and determine the nuclear intensities of Geminin-mAG and pRPA32 (S4/S8). The bar graph depicts the percentage of S and G2 cells (Geminin-mAG-positive) with pRPA32 (S4/S8) nuclear intensities over the mean nuclear background intensity (calculated in cells not treated with NCS) (Mean ± SEM; N ≥ 3).

Figure 3.11. ZNF335 depletion does not affect the expression or protein stability of the core end resection activators. (A) Whole cell extracts from U2OS cells incubated with scrambled control, CtIP, MRE11, RAD50, NBS1, or ZNF335 siRNA were separated by SDS-PAGE and subjected to MRE11, RAD50, NBS1, ZNF335, and GAPDH immunoblotting. (B) Whole cell extracts from U2OS cells incubated with scrambled control, CtIP, or ZNF335 siRNA were separated by SDS-PAGE and subjected to CtIP, ZNF335, and GAPDH immunoblotting. (C) Whole cell extracts from U2OS cells incubated with scrambled control, EXO1, or ZNF335 siRNA were separated by SDS- PAGE and subjected to EXO1, ZNF335, and GAPDH immunoblotting. (D) Whole cell extracts from U2OS cells incubated with scrambled control, DNA2, or ZNF335 siRNA were separated by SDS-PAGE and subjected to DNA2, ZNF335, and GAPDH immunoblotting. (E) Whole cell extracts from U2OS cells incubated with scrambled control, BLM, or ZNF335 siRNA were separated by SDS-PAGE and subjected to BLM, ZNF335, and GAPDH immunoblotting. (F) Bar graph depicting KS scores from the primary RNAi screen for known components of the trithorax complex.

(Fig. 3.11F), suggesting that ZNF335 may function independently of trithorax to promote end resection.

3.4.10 Immunoprecipitation coupled to mass spectrometry (IP-MS) identifies ZNF335 candidate interaction partners

In an attempt to gain more mechanistic detail regarding the function of ZNF335 in end resection I conducted ZNF335 FLAG IP-MS in collaboration with Zhen-Yuan Lin in Anne-Claude Gingras’ laboratory at the LTRI. I generated HEK293 cell lines that stably express N-terminally FLAG-tagged ZNF335 and a control cell line that expresses FLAG alone. After induction with doxycycline, I treated each cell line with NCS for 1 h before collecting cell pellets. Cell lysis, FLAG immunoprecipitations, mass spectrometry, and analysis were carried out by Zhen-Yuan Lin. SAINT (significance analysis of interactome) was applied to the data and is an algorithm that determines the probably of protein-protein interactions (Choi et al., 2011). High-confidence prey proteins with SAINT scores over 0.9 are listed in Table 3.2. One particular interaction, DCLRE1C (or Artemis), caught our attention because it is a well-characterized nuclease that functions in DSB repair by NHEJ. It will be interesting in the future to determine if Artemis has any role in resecting DSB ends and if this function is facilitated by ZNF335.

Expressing the four C-terminal zinc finger domains of ZNF335 was sufficient to restore end resection in ZNF335 depleted cells. To increase protein interactions that may be relevant to the function of ZNF335 in end resection, I decided to conduct FLAG IP-MS with only the four C-terminal zinc finger domains (ZNF335 Δ1-1014 mutant). Identified prey proteins with SAINT scores over 0.9 are listed in Table 3.3. The highest ranking candidate interactor was the histone chaperone SET which is known to promote chromatin compaction by masking lysine residues from the action of histone acetyltransferases (Seo et al., 2001). Another candidate interactor that sparked my interest was FAM115A. Although the function of this protein is unknown, our laboratory recently identified it as an RPA32 interactor (Tkac et al., 2016). The RecQ helicase, RECQL, was also identified as an interacting partner with the ZNF335 C-terminus. This will be an interesting interaction to validate as two other RecQ helicases, BLM and WRN, function in long-range end resection (Gravel et al., 2008; Sturzenegger et al., 2014). Lastly, PARP-1 was identified as a candidate interacting protein with the ZNF335 C-terminus which was intriguing as I also determined that ZNF335 recruitment to sites of DNA damage generated by microirradiation was PARP-dependent. However, I demonstrated that PARP is not required for

Prey proteins Spectral count SAINT score Function HNRNPA0 51 0.89 mRNA processing DDX21 46 1 mRNA processing HNRNPM 36 1 mRNA processing ENO2 23 1 Metabolic enzyme; enolase STAU1 21 1 mRNA transport and processing AHNAK 17 1 Scaffolding protein with diverse functions DCLRE1C 16 1 Artemis; nuclease implicated in NHEJ IGF2BP1 11 1 mRNA binding protein; regulates translation RBM14 10 0.99 mRNA processing SAFB 7 0.94 Transcription factor SDF4 7 0.9 Exocytosis GTF2F1 6 0.94 Transcription factor MAP7D1 6 0.9 Microtubule organization FUBP3 5 0.9 Transcription factor LARP1 5 0.9 mRNA binding protein; regulates translation UTP14A 5 0.9 rRNA processing; ribosome biogenesis FLII 4 0.9 Actin cytoskeleton organization

Table 3.2. Candidate protein interaction partners for full-length ZNF335. Protein interactions determined by FLAG IP-MS. The spectral count for each prey protein is the average of three biological replicates. SAINT analysis was conducted to determine the probability that the interaction is true and to normalize for protein size. High SAINT scores indicate that the interaction was observed in multiple replicates and was only minimally identified in negative control samples (FLAG only).

Prey proteins Spectral count SAINT score Function SET 121 1 Histone chaperone; inhibits histone acetylation promoting chromatin compaction FAM115A 31 1 Unknown function; Identified RPA interactor PARP1 21 1 Poly (ADP) ribose polymerase; DNA repair PRKAR2A 21 1 Kinase; regulates protein transport to Golgi CPVL 20 1 Metabolic enzyme; carboxypeptidase SELRC1 20 1 Cytochrome c oxidase assembly factor SEC23B 17 0.99 Promotes vesicle budding from ER MGA 12 0.93 Transcription factor SEC24B 12 0.93 Promotes vesicle budding from ER MRPS31 10 0.99 Mitochondrial ribosomal protein IQGAP2 8 1 Cytoskeleton organization KPNA6 8 0.99 Importin; cytoplasm to nucleus import TIMM13 8 0.99 Translocase of inner mitochondrial membrane SLC25A5 7 0.99 Mitochondrial adenine solute carrier EIF4A3 7 0.95 Translation initiation factor PDS5A 6 0.95 Cohesion associated factor; regulates sister chromatid cohesion GNB3 5 0.95 Guanine nucleotide-binding protein MAD2L1 5 0.95 Mitotic spindle assembly checkpoint PRKACA 5 0.95 Protein kinase A subunit; protein kinase A is important to many cellular processes, including differentiation and apoptosis RECQL 5 0.95 DSB repair by HR; Helicase involved in HJ branch migration and replication fork reversal EIF2S3 5 0.92 Translation initiation factor MATR3 5 0.92 mRNA processing MRPS14 5 0.92 Mitochondrial ribosomal protein MRPS21 5 0.91 Mitochondrial ribosomal protein SLFN11 5 0.91 Potential DNA/RNA helicase; unknown function PKN2 4 0.90 Protein kinase N2; numerous cellular processes

Table 3.3. Candidate protein interaction partners for ZNF335 Δ1-1014. Protein interactions determined by FLAG IP-MS. The spectral count for each prey protein is the average of three biological replicates. SAINT analysis was conducted to determine the probability that the interaction is true and to normalize for protein size. High SAINT scores indicate that the interaction was observed in multiple replicates and was only minimally identified in negative control samples (FLAG only).

end resection, suggesting that ZNF335 may interact with PARP-1 to participate in another function.

3.5 Discussion

In this chapter, I demonstrated that ZNF335 promotes DNA end resection and subsequent DSB repair by HR. For these studies, I utilized RNAi-mediated depletion of ZNF335 from human U2OS cells. End resection is cell cycle-regulated, occurring in the S and G2 phases when a sister chromatid is available for template-driven repair. My results suggest that ZNF335 knockdown specifically impairs end resection and is not an artifact of the siRNA causing a G1 cell cycle arrest or delay. I also uncovered that cells depleted of ZNF335 are more sensitive to DSB- inducing drugs which further supports a role for ZNF335 in DSB repair. Importantly, other known end resection factors also cause sensitivity to DSB-inducing agents when their function is perturbed, including the MRN complex and CtIP (Sartori et al., 2007; Stewart et al., 1999; Taalman et al., 1983; Waltes et al., 2009). CtIP depleted U2OS cells were more sensitive to DSB-inducing agents than cells incubated with ZNF335 siRNA, suggesting that CtIP plays a more essential role in the completion of DSB repair and the promotion of cell survival. CtIP is known to be required for the initiation of end resection (Sartori et al., 2007). One hypothesis is that ZNF335 may promote end resection downstream of initiation where it increases the efficiency of resection and HR but is not necessarily required.

A previous phospho-proteomics study uncovered a ZNF335 peptide that was phosphorylated on serine 416, a canonical ATM/ATR consensus site (Matsuoka et al., 2007). My data indicated that phosphorylation at this site was not required for end resection as a serine- to-alanine amino acid change was able to rescue resection upon ZNF335 depletion. Consistently, I determined that the four C-terminal C2H2 zinc finger domains of ZNF335 are required for end resection and serine 416 is not present in this region. Moreover, in response to DNA damage with NCS, I have never observed any mobility shifts when conducting ZNF335 immunoblotting. The importance of the C-terminus enhanced my interest in the microcephaly syndrome as the c.3332g>a mutation is located in this region (Yang et al., 2012). The disease mutation caused an amino acid change from an arginine to histidine in the middle of the thirteenth zinc finger domain. My data demonstrated that the ZNF335 R1111H mutant protein can function normally in promoting end resection. Importantly, the disease mutation also caused aberrant splicing and

94 retention of the two introns flanking exon 21 in some ZNF335 transcripts. Improperly spliced transcript (if not degraded by nonsense mediated mRNA decay), would produce a truncated protein that terminated after exon 20 with the inclusion of 38 mutant amino acids coded for in the subsequent intron before a premature stop codon was reached. The truncated protein would terminate in the middle of the eleventh zinc finger domain meaning the final three C-terminal zinc fingers would be disrupted. If the truncated ZNF335 protein is even produced in patient cells is not known as the authors employed an antibody that only detected the final 42 amino acids of the C-terminus. It will be interesting in the future to overexpress a truncated ZNF335 ORF that terminates in the middle of the eleventh zinc finger domain to determine if it is able to function in promoting end resection and HR.

One important question is whether ZNF335 can promote end resection by directly acting at DSB sites or if it exerts its function indirectly. I was only able to visualize GFP-ZNF335 localization to sites of DNA damage generated by laser microirradiation which is consistent with other end resection regulators including EXO1, SRCAP, and HELB (Bolderson et al., 2010; Dong et al., 2014; Tkac et al., 2016). I observed a robust recruitment of GFP-ZNF335 to laser stripes that was ATM-independent and PARP-dependent. GFP-ZNF335 only localized to sites of microirradiation for a short time period which is in stark contrast to the temporal dynamics of other proteins known to promote end resection. For example, CtIP has been observed to localize to DSB sites for up to four hours after DNA damage (Yuan and Chen, 2009). I determined that the transient accumulation of GFP-ZNF335 at sites of microirradiation was not functionally relevant for end resection as inhibiting recruitment with olaparib treatment had no effect in resection assays. Moreover, I observed that U2OS cells depleted of PARP-1 had an elevated level of end resection in response to NCS treatment. This result was not surprising as NCS is a potent inducer of SSBs and PARP-1 is critical for SSB repair (Molinete et al., 1993). The high density of SSBs in PARP-1-deficient cells treated with NCS will enhance the probability that two close-proximity SSBs will be converted into a DSB. Indeed, PARP-1 depleted cells also form more one-ended DSBs during S phase when replication forks encounter unrepaired SSBs (Bryant et al., 2005; Farmer et al., 2005). In contrast to PARP-1 knockdown, olaparib treatment resulted in a level of NCS-induced end resection that was similar to cells treated with DMSO alone. A possible explanation for this discrepancy is that olaparib results in PARP-1/2 trapping on DNA break ends which could potentially have an inhibitory effect on the end resection

machinery (Murai et al., 2012). The PARP-dependent recruitment of GFP-ZNF335 to stripes could be important for another function or it could be an artefact of the laser microirradiation technique. Indeed, it was recently demonstrated that a plethora of DNA-binding proteins localize transiently to stripes in a PARP-dependent manner (Izhar et al., 2015). The DNA-binding proteins included many transcription factors that have never been functionally implicated in DNA repair. A collection of 576 epitope-tagged proteins were examined for localization to sites of laser microirradiation. Remarkably, of the transcription factors tested, over 70% localized to stripes, and of these, 90% were PARP-dependent. By examining Hoechst staining patterns at laser stripes, evidence of PARP-dependent chromatin decompaction was also observed (Izhar et al., 2015). The authors postulated that PARP-regulated chromatin remodeling at sites of microirradiation may endow accessibility to DNA-binding transcription factors. Indeed, several chromatin remodeling enzymes are known to be recruited to microirradiated chromatin in a PARP-dependent manner, including the NuRD complex and the SNF2-related proteins CHD1 and CHD2 (Ahel et al., 2009; Chou et al., 2010; Luijsterburg et al., 2016; Polo et al., 2010). However, it cannot be ruled out that the DNA-binding domains of some transcription factors may directly bind to PAR. Direct binding would not be a surprising observation as PAR is chemically and structurally similar to the nucleotides present in DNA (Kim et al., 2005). The enrichment of transcription factors at sites of microirradiation begs the question if these proteins have functional roles in DNA repair or if their recruitment is merely an artifact of associating with newly (and transiently) accessible DNA.

DNA end resection is not only important for HR but also for the activation of ATR. RPA coated ssDNA is a required platform for ATR activation and this can occur as a consequence of replication stress or end resection (Zou and Elledge, 2003). Once activated, ATR can phosphorylate hundreds of downstream substrates that regulate DNA repair, cell cycle checkpoints, and apoptosis. As outlined in Chapter I, several DSB repair genes are mutated in genetic syndromes that display neonatal microcephaly (O'Driscoll and Jeggo, 2008). Several microcephaly-causing subtypes of Seckel syndrome are a consequence of mutations in ATR or its interacting protein ATRIP, suggesting that ATR activity is important for normal neurogenesis in the developing cerebral cortex (O'Driscoll et al., 2003; Ogi et al., 2012). Moreover, two factors that regulate end resection, CtIP and DNA2, have been shown to be mutated in Seckel syndrome (Qvist et al., 2011; Shaheen et al., 2014). ZNF335 is also mutated in a rare genetic

96 syndrome that results in severe neonatal microcephaly (Yang et al., 2012). I determined that ATR activity was diminished in cells depleted of ZNF335 by measuring CHK1 phosphorylation. It will be interesting in the future to determine if patient cells harbouring the c.3332g>a mutation in ZNF335 have defective ATR activation and if ZNF335 should be classified as a new causative gene for Seckel syndrome.

Chapter IV

Solid-phase transfections in arrayed drops (SPIDR): a cell microarray-based platform for high-content screening

4.1 Statement of contributions, rights, and permissions

Gagan Gupta in Laurence Pelletier’s laboratory (LTRI) designed the 1536- and 3456-well cell microarrays and provided critical guidance in experimental design and troubleshooting.

Scientific Device Laboratory (Des Plaines, Illinois, USA) fabricated the 1536- and 3456-well cell microarrays using thin cover glass and biologically-compatible black ink to define well positions.

Thomas Sun and Alessandro Datti (LTRI robotics facility) developed all liquid-dispensing procedures on the Caliper Sciclone inL10 and provided critical guidance in experimental design and troubleshooting.

Mikhail Bashkurov (LTRI high-content screening facility) established automated confocal imaging procedures for cell microarrays using the InCell Analyzer 6000 (GE Healthcare) and provided critical guidance for microscopy troubleshooting.

4.2 Summary

High-content screens are typically conducted in multi-well plates which require long liquid handling robotic procedures and large quantities of expensive plates and reagents. These limitations can be circumvented by using cell microarray-based formats where thousands of distinct genetic reagents are printed as individual microscopic “spots” on glass slides. The printed arrays are then flooded with media containing adherent cultured cells which attach and become treated with the reagent present on each spot. Here, I show the establishment of a cell microarray-based platform for conducting high-content RNAi screens. I show the design and fabrication of thin glass cell microarrays that are the size of a standard multi-well plate. I demonstrate that the previously reported conditions for conducting siRNA transfections on cell microarrays are not sufficient in preventing sample cross contamination. I show that the delivery of cells within nanolitre-scale droplets of media to each printed siRNA spot prevents the diffusion of solubilized transfection complexes between samples. This strategy allows gene expression knockdown to occur in a localized fashion and we have named this method Solid- Phase transfections In arrayed DRops (or SPIDR). I demonstrate the utility of SPIDR by screening 1,536 siRNAs for regulators of 53BP1 localization to DSB sites. I conclude that SPIDR is a novel strategy for performing cell microarray-based high-content screens.

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4.3 Introduction

In parallel to the plate-based high-content screen mining for regulators of DNA end resection (Chapter II), I have also worked to establish an alternative high-content screening platform based on cell microarrays. Plate-based screens require long liquid handling robotic procedures and large quantities of expensive plates and reagents. These limitations can be circumvented by using cell microarray-based formats where hundreds to thousands of distinct genetic reagents are printed as individual microscopic “spots” on glass slides (Fig. 4.1A). The printed and dried arrays are then seeded with adherent cells that attach and become treated with the reagent present on each spot.

Cell microarray technology was first invented in David Sabatini’s laboratory where GFP expression plasmids were printed on glass slides in combination with gelatin and a lipid-based transfection reagent (Ziauddin and Sabatini, 2001). After drying the DNA/gelatin mixture, cells were flooded onto the array and adhered to the semi-aqueous features where they became reverse-transfected in solid-phase. The distance between adjacent spots on cell microarrays is typically less than 0.5 mm which facilitates sample densities almost 200 times that of a 384-well plate. Remarkably, it is theoretically possible to array the entire set of human ORFs or siRNA libraries on a single glass slide the size of a standard multi-well plate (Wu et al., 2002). Furthermore, cell microarrays are highly suitable for multiplexing where hundreds of arrays can be printed in a single day. Cell microarrays also make it possible to screen thousands of reagents under different experimental conditions (i.e. genetic backgrounds or stressors). DNA/gelatin- printed cell microarrays can be stored for long time periods (up to 1 year) and still elicit efficient gene expression knockdown (Ziauddin and Sabatini, 2001). In addition to cDNAs, cell microarrays have now been printed with numerous genetic reagents including siRNAs, shRNA plasmids, and shRNA lentivirus (Bailey et al., 2006; Silva et al., 2004; Wood et al., 2012). However, these technical reports only outlined the printing of small sets of RNAi reagents and no genome-scale high-content screens have been published. Here, I outline the establishment of a cell microarray-based platform for high-content siRNA screens and employ the technology to screen 1,536 siRNAs on a single glass slide.

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4.4 Results

4.4.1 Design and fabrication of cell microarrays

In order to establish a cell microarray-based platform we designed 1536- and 3456-well arrays where each ‘well’ was separated by a thin biologically-compatible cured black ink layer (Fig. 4.1B). The arrays were fabricated by Scientific Device Laboratory (Des Plaines, Illinois, USA) using thin glass and dimensional standards outlined by the Society for Biomolecular Screening (SBS). We envisioned that this thin ink layer would minimize cell migration between samples and allow for visually defined well positions to help in array alignment during downstream imaging routines. Using the 96 tip Caliper Sciclone inL10, small volumes (nano-litre range) can be delivered to either of these arrays with a coefficient of variation (CV) less than 5%.

4.4.2 Establishment of solid-phase siRNA transfections in human cultured cells

I optimized solid-phase reverse siRNA transfections starting with conditions outlined by Rainer Pepperkok’s group (Erfle et al., 2007). To test knockdown efficiency, I utilized an immunofluorescence-based assay that monitors the recruitment of 53BP1 to DSB sites which can be visualized cytologically as subnuclear foci. 53BP1 is a critical DSB response protein that mediates the DNA damage checkpoint and promotes long-range NHEJ reactions (Difilippantonio et al., 2008; Dimitrova et al., 2008; Wang et al., 2002) The recruitment of 53BP1 is ubiquitin- dependent and requires the activity of two E3 ligases, RNF8 and RNF168 (Doil et al., 2009; Huen et al., 2007; Kolas et al., 2007; Mailand et al., 2007; Stewart et al., 2009). I tested various ratios of solid-phase transfection ingredients (gelatin, fibronectin, sucrose, RNAiMAX transfection reagent, and siRNA) by dispensing a small drop (500 nL) of each condition directly to the center of a glass coverslip followed by drying in a vacuum desiccator for 48 hours (Fig. 4.2A). For each condition both scrambled control and RNF168 siRNA was tested. Next, media containing HeLa cells was dispensed on top of the coverslips in 24-well plates. After 48 hours to allow for knockdown, cells were exposed to 10 Grays IR followed by a one hour recovery. HeLa cells were then processed for 53BP1 immunofluorescence. Coverslips were mounted onto glass slides and cells were imaged on an automated confocal microscope. Using image analysis software, 53BP1 focus formation was quantified by measuring the fraction of the nuclear area covered by segmented foci. The solid-phase transfection complex that yielded the most efficient

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Figure 4.1. Formats for high-content screening. (A) high-content screens are typically conducted in 96- or 384-well plates. Plate-based screens require long liquid handling robotic procedures and large quantities of expensive and sometimes limited antibodies. These limitations can be circumvented by using cell microarray-based formats where reagents are arrayed in high density on glass slides and then dried. Cells and immunostaining reagents are flooded onto the array without the need for robotics. (B) In order to establish a cell microarray-based platform we designed and fabricated 1536- and 3456-well arrays where each well is separated by a thin biologically-compatible black ink layer. We envisioned that this layer could help minimize cell migration between samples. Visually defined well positions also helps in array alignment during downstream imaging routines.

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Figure 4.2. Solid-phase reverse siRNA transfections in HeLa cells. (A) Solid-phase transfection mix containing either scrambled control (CTRL) or RNF168 siRNA was manually dispensed to the center of a coverslip. The mix contained 20 ng of siRNA, 5% RNAiMAX, 0.08% gelatin, 32 mM sucrose, and 0.004% fibronectin. The complexes were then dried in a vacuum desiccator for 48 hours. HeLa cells were flooded onto the coverslip and after 48 hours they were exposed to 10 Grays of IR. One hour later cells were immunostained for 53BP1. Scale bar represents 20 µm. (B) Transfection efficiency was comparable to standard liquid-phase transfections as 53BP1 focus formation was abolished to a similar degree upon RNF168 depletion (Mean ± SEM; N ≥ 3). (C) Dried down complexes were stored in a vacuum sealed bag at 4 ºC for various time intervals before being utilized in 53BP1 focus formation assays (Mean ± SEM; N ≥ 3).

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knockdown is depicted in Figure 4.2. Importantly, I observed a similar, if not equal, knockdown efficiency when I conducted a standard liquid-phase siRNA transfection (Fig. 4.2B). For the cell microarray platform to act as a viable method for high-content screens the dried complexes must be stable for long time periods. To test this, the optimized solid-phase transfection mixture was dispensed and dried at the center of multiple glass coverslips and stored in vacuum sealed packages at 4 ºC for either 0, 2, 4, or 6 months before being utilized in 53BP1 focus formation assays. The dried siRNA complexes were stable for up to 6 months as they elicited RNF168 knockdown (Fig. 4.2C).

4.4.3 Testing for cell and siRNA cross contamination between samples

To determine if cultured cells are able to migrate between wells, GFP- and RFP-expressing U2OS cells were delivered in alternate rows on both 1536- and 3456-well arrays using the Sciclone inL10 (Fig. 4.3A). The volume of robotically dispensed cells was small enough not to touch and cross contaminate adjacent wells on the arrays. After eight hours to allow the cells to adhere to the glass, the array was flooded with fresh media. Cells were fixed 48 hours after delivery to the arrays and imaged using an automated confocal microscope. An end point of 48 hours was selected because this would be the duration of a typical siRNA screen. GFP- and RFP- positive cells were quantified using image analysis software. The percent cell cross contamination for each well was determined by dividing the number of inappropriately localized cells by the total number of cells. For example, an inappropriately localized cell would be a GFP- positive cell where RFP-positive cells were dispensed. In this scenario, an average of 0.33% of cells were inappropriately localized on the 1536-well array and only 0.38% on the 3456-well (Fig. 4.3B). Therefore, in the context of a 48 hour experiment, there is minimal cell cross contamination between wells on our fabricated cell microarrays.

Next, I surmised that siRNA complexes may diffuse after flooding arrays and cross contaminate adjacent wells during the course of a screen. To test this, fluorescently conjugated Alexa488 control siRNA was delivered to three clusters (each cluster was made up of four wells) on a 1536 array (Fig. 4.4A). The remaining wells were occupied by control siRNA with no fluorescent dye. Using the Sciclone, 100 nL of the optimized solid-phase transfection mixture was dispensed to each well. After drying, each spot was a diameter of approximately 1 mm and the distance between spots was also 1 mm. The complexes were dried down in a vacuum

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Figure 4.3. Cell migration between samples is minimal on 1536- and 3456-well cell microarrays. (A) GFP- or RFP-expressing U2OS cells were dispensed in alternate rows using the Sciclone InL10. After 8 hours, the array was flooded with media. Cells were fixed 48 hours after delivery and imaged on an automated microscope. GFP- and RFP-positive cells were quantified using image analysis software. (B) The percent cell cross contamination for each well was determined by dividing the number of inappropriately localized cells by the total number of cells.

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Figure 4.4. Cross contamination of siRNA complexes is evident on cell microarrays. (A) Solid-phase transfection mix containing fluorescently conjugated Alexa488 control siRNA was delivered to three clusters (each cluster was made up of four wells) on a 1536 array. The remaining wells contained control siRNA with no fluorescent dye. The complexes were dried down in a vacuum desiccator for 48 hours and then HeLa cells in media were flooded onto the array. After 48 hours to allow for cell seeding and siRNA transfection, the cells were fixed and DNA was stained with DAPI to visualize nuclei. Cells were imaged with an automated microscope and DAPI-stained nuclei were segmented with image analysis software. Next, nuclear masks were expanded to include parts of the cytoplasm. Spot detection was conducted in the cytoplasmic region to identify Alexa488- positive endosomes. (B) Solid-phase transfection mix containing either control (CTRL) or RNF168 siRNA was dispensed using the Sciclone. After drying, HeLa cells were flooded onto the array. Strikingly, RNF168 siRNA was able to diffuse and transfect cells in adjacent wells where control siRNA was delivered. This is demonstrated by the decrease in 53BP1 focus formation in cells located in control siRNA wells that immediately surround RNF168 siRNA wells.

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desiccator for 48 hours and then HeLa cells in media were flooded onto the array. After 48 hours to allow for cell seeding and siRNA transfection, the cells were fixed and DNA was stained with DAPI to visualize nuclei. In mammalian cells, liposome-mediated transfection of nucleic acids occurs through a process called endocytosis (Rizzo et al., 1983). Therefore, successfully transfected cells will contain siRNA-positive endosomes. Using an automated confocal microscope, fluorescent cytoplasmic foci (likely endosomes) were observed in HeLa cells growing where Alexa488-conjugated control siRNA was dispensed (Fig. 4.4A). In contrast, cells growing at distal regions of the 1536-well array did not contain any visible Alexa488-positive endosomes. However, wells adjacent to where Alexa488-conjugated control siRNA was dispensed also contained Alexa488-positive endosomes which suggested that siRNA complexes were able to diffuse and transfect cells in neighbouring wells. I also examined siRNA cross contamination utilizing the established functional assay that monitors 53BP1 focus formation. Solid-phase transfection mix containing either control or RNF168 siRNA was dispensed onto a 1536-well array using the Sciclone. After drying, HeLa cells were flooded onto the array. After 48 hours to allow for knockdown, cells were exposed to IR and then processed for 53BP1 immunostaining. Strikingly, RNF168 siRNA was also able to diffuse and transfect cells in adjacent wells where control siRNA was delivered. Cross contamination was demonstrated by the decrease in 53BP1 focus formation in cells located in control siRNA wells that immediately surrounded wells where RNF168 siRNA was delivered (Fig. 4.4B).

Numerous ratios of the solid-phase transfection ingredients were tested but we were unable to minimize siRNA cross contamination between wells. I attempted to increase the distance between samples on the array by decreasing the total dispensing volume of solid-phase transfection mix. The Sciclone can dispense a minimum of 50 nL and this volume resulted in a spot diameter of approximately 0.5 mm and a distance of 2 mm between spots. However, even with this distance, sample cross contamination was evident. Next, the newly printed arrays were dried for longer time periods before flooding cells and these tests were also unsuccessful at preventing siRNA diffusion. I surmised that pre-soaking or blocking printed arrays with media before flooding the cells could help decrease the surface siRNA concentration for each sample and therefore decrease the observed diffusion. To test this, control and RNF168 siRNA solid- phase transfection mixes were delivered to three 1536-well arrays and, after drying, each was pre-soaked with fresh media for approximately 30, 120, or 300 seconds before flooding HeLa

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Figure 4.5. Pre-soaking cell microarrays before cell flooding decreases knockdown efficiency. (A,B) Solid-phase transfection mix containing either control (CTRL) or RNF168 siRNA was delivered to a 1536- well cell microarray. Three wells received RNF168 siRNA and the remaining wells received CTRL siRNA. After drying, the arrays were pre-soaked in media for various time intervals: 30, 120, and 300 seconds. Next, the arrays were flooded with HeLa cells. After 48 hours to allow cell seeding and knockdown, cells were exposed to 10 grays IR and, one hour later, processed for 53BP1 immunofluorescence. (C) All pre-soaking times resulted in a decreased RNF168 knockdown efficiency, determined by quantifying the fraction of each nucleus with 53BP1 foci (Mean ± SEM; N ≥ 3).

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cells. After 48 hours, the cells were exposed to IR and processed for 53BP1 immunofluorescence (Fig. 4.5A,B). Remarkably, pre-soaking the arrays in media before flooding cells substantially decreased the observed cross contamination of siRNA between wells. Unfortunately, it also markedly decreased knockdown efficiency in cells growing directly on dried down RNF168 siRNA spots, even at the rapid pre-soak time of 30 seconds (Fig. 4.5C).

4.4.4 Solid-Phase transfections In arrayed DRops (SPIDR): a novel method to prevent sample cross contamination on cell microarrays

To circumvent the issue of siRNA cross contamination I tried delivering cells in small droplets of media to each of the arrayed siRNA complexes (Fig. 4.6A, B). I hypothesized that this approach would minimize the diffusion of solubilized transfection complexes between samples allowing gene expression knockdown to occur in a localized fashion. Using the Sciclone, I was able to dispense 500 nL of media containing HeLa cells to each well of a 1536 array and after 48 hours of growth a consistent number of cells per sample was observed (Fig. 4.6C). To examine siRNA cross contamination, RNF168 siRNA was delivered to several rows of a 1536 array. Using the Sciclone, small droplets of media (500 nL) containing HeLa cells were dispensed to each of the arrayed and dried siRNA complexes. After 24 hours of localized transfection in the drops the cell microarray was flooded with fresh media. A total of 48 hours after seeding, cells were exposed to IR and allowed to recover for 1 hour before being processed for 53BP1 immunostaining. After multiple phases of optimization, I determined a concentration of solid- phase transfection ingredients that resulted in minimal cellular toxicity and maximal knockdown. As the transfections are conducted in small nanolitre-scale droplets, the optimal concentration was determined to be 15-fold less than the conditions used for cell flooding. Strikingly, the cells growing in the control siRNA wells adjacent to where RNF168 siRNA was delivered had robust 53BP1 focus formation, demonstrating localized gene expression knockdown (Fig. 4.6D). We have named this method SPIDR for Solid-Phase transfections In arrayed DRops.

4.4.5 Pilot RNAi screen utilizing the SPIDR platform

To test the SPIDR high-content screening platform the ubiquitin conjugation subset of the siRNA SMARTpool library (targeting 606 genes; GE Healthcare/Dharmacon) was dispensed onto a 1536 array. The ubiquitin conjugation subset was selected in a biased manner as 53BP1 localization to DSBs is ubiquitin-dependent and requires the E3 ligases RNF8 and RNF168, both

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Figure 4.6. Solid-phase transfections in arrayed drops (SPIDR). (A,B) To circumvent siRNA cross contamination we have developed a cell microarray platform called SPIDR. Cells are delivered in small droplets of media to each of the arrayed siRNAs to prevent the diffusion of solubilized transfection complexes between samples. This strategy allows gene expression knockdown to occur in a localized fashion. (C) HeLa cells dispensed to a 1536 array using the Sciclone. Small-volume cell delivery worked well with the Sciclone, resulting in a coefficient of variation (CV) less than 10%. (D) HeLa cells were delivered in small droplets to each well of a 1536 array where each well contained dried down solid-phase transfection mix with either control (CTRL) or RNF168 siRNA. After 24 hours, the array is washed with PBS and flooded with fresh media. A total of 48 hours post-seeding, HeLa cells were exposed to IR and one hour later processed for 53BP1 immunostaining. Of the 1536 negative (siCTRL) and positive (siRNF168) controls tested only 4 wells did not demonstrate the expected result, giving a false discovery rate (FDR) of 0.25% (4/1536).

111 of which were present in the subset. The remaining wells on the pilot array were occupied with an siRNA subset that contained controls for assays being tested by our collaborators. Multiple positive control siRNAs that should decrease 53BP1 focus formation were added to the outside columns of the array, including RNF8, RNF168, MDC1, and 53BP1. A total of 32 copies of the array were produced to allow for experimental replicates and to test multiple assays. As a first test, HeLa cells were dispensed to the pilot array and after 24 hours the array was flooded with fresh media. After 48 hours post-seeding, cells were exposed to IR and allowed to recover for 1 hour before being processed for 53BP1 immunofluorescence. Two biological replicates were carried out for this experiment using two copies of the pilot array. After imaging and analysis of each replicate, the mean fraction of the nucleus with 53BP1 foci was measured for each well and data was normalized using the z-score (number of standard deviations from the mean; Fig. 4.7A). Remarkably, not only did most of the added positive controls result in lowered 53BP1 foci levels, but the top three hits in the pilot screen were RNF8, RNF168, and 53BP1. One systematic error observed in the data was that the signal intensity was greater in the center of the arrays compared to the edges. Therefore, in collaboration with Mikhail Bashkurov, another normalization method was employed called B-score analysis which corrected for within-plate row and column effects by an iterative application of the Tukey median polish algorithm (Birmingham et al., 2009). B-score normalization not only corrected the regional signal intensity variations in the data but also was able to pick out weaker hits, including UBC13, the E2 enzyme that functions with RNF8 and RNF168 in substrate ubiquitylation (Doil et al., 2009; Huen et al., 2007; Kolas et al., 2007; Stewart et al., 2009). In conclusion, we have developed a novel method for conducting cell microarray-based screens that circumvents the technical issue of sample cross contamination.

4.5 Discussion

We have established a novel high-content screening platform using transfected cell microarrays. Our fabricated 1536- and 3456-well arrays supported the healthy growth of human cultured cells and were compatible for the delivery of nanolitre volumes using the Caliper Sciclone inL10 small-volume liquid-handling robot. I was also able to successfully dispense to the arrays using another low-volume liquid handler called the Mosquito HTS (TPPlabtech). However, the minimum dispensing volume that we could achieve with these instruments was 50 nL. One potential strategy to minimize cross contamination on cell microarrays is to increase the sample-

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Figure 4.7. High-content siRNA screen utilizing the SPIDR platform. (A) To test the SPIDR screening platform, the ubiquitin conjugation subset of the Dharmacon siRNA library and multiple control siRNAs including RNF8, RNF168, MDC1, and 53BP1 were delivered to a 1536-well array. 53BP1 focus formation was monitored 1 h after 10 Grays of IR. Known modulators of 53BP1 focus formation were strong hits in the screen demonstrating the efficacy of the SPIDR platform. Furthermore, data quality metrics such as Z factor and false discovery rate (FDR) also reflect the utility of this platform. (B) One systematic trend observed in the data was a higher signal of 53BP1 focus formation in the center of the array compared to the edges. To normalize this error, B-score analysis was conducted to correct for within-plate row and column effects by an iterative application of the Tukey median polish algorithm.

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to-sample distance by decreasing the volume of solid-phase transfection mix that is delivered to each well. Recently, new liquid handling robots have been developed that utilize sound waves to transfer extremely low volumes (2.5-10 nL) with high accuracy. These ‘acoustic’ instruments can transfer liquids from multiwell source plates to target plates by delivering sound waves to each well. Another method for delivering small volumes is to use microarray pinning tools which can achieve volumes as low as 1 nL. It will be exciting to utilize these technologies in the future to troubleshoot sample cross contamination on our flooded cell microarrays.

The protocol I employed for conducting solid-phase siRNA transfections on cell microarrays was first established by Rainer Pepperkok’s group (Erfle et al., 2007). The authors observed minimal siRNA cross contamination between samples which was measured by quantifying cell death induced by KIF11 knockdown. KIF11 is a kinesin motor protein that acts on microtubules and is required for chromosome segregation in mitosis (Sawin et al., 1992). KIF11 deficient cells accumulate in mitosis and eventually initiate programmed cell death. Compared to control siRNA spots, Erfle et al. (2007) reported that the number of cells growing on KIF11 siRNA spots was less 48 hours-post transfection. However, KIF11 knockdown efficiency on their fabricated microarrays appeared to be inefficient as they observed only a small dynamic range between control and KIF11 siRNA. The observed cross contamination on our cell microarrays could be a consequence of more efficient RNF168 knockdown. It is also possible that quantification of 53BP1 focus formation may be a more sensitive readout then cell number for assessing cross contamination between samples. For example, KIF11 may have to be completely depleted from cells to observe efficient cell death whereas intermediate knockdown of RNF168 could drastically reduce 53BP1 foci.

To prevent siRNA cross contamination between samples we developed a novel method called SPIDR that involves delivering cells in small droplets of media to each of the arrayed and dried transfection complexes. SPIDR prevents the diffusion of siRNAs between samples, allowing gene expression knockdown to occur in a localized manner. The SPIDR platform allows us to perform high-content siRNA screens using a miniaturized cell microarray format. SPIDR has all the advantages of the original cell microarray concept but with one drawback, cells have to be delivered to the arrays using robotics rather than simple manual flooding. In the future, it will be important to test the SPIDR platform on other biological assays and cell types. We would also like to expand to higher density cell microarrays (i.e. 3456-well) and screen

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larger siRNA sets. Furthermore, with the recent advancement of CRISPR/Cas9 gene editing technology it will be exciting to produce SPIDR cell microarrays with synthetic small guide RNAs (sgRNAs) or sgRNA plasmids to be transfected into Cas9-expressing cells (Wright et al., 2016).

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Chapter V

Future directions

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5.1 Validation of candidate end resection activators

The characterization of other candidate resection activators identified in the RNAi screen should be the focus of upcoming research. C9ORF106 and TP53I13 are candidate resection activator genes of unknown function and have been previously identified by next-generation sequencing experiments to express putative lncRNAs (Bhartiya et al., 2013; Liu et al., 2014). Thousands of lncRNAs are transcribed by the human genome and have important cellular functions that include the regulation of transcription and chromatin structure (Geisler and Coller, 2013). Expression of the TP53I13 gene was previously shown to be transcriptionally upregulated in response to DNA damage in a p53-dependent manner (Hata et al., 2004). Moreover, there has been no experimental evidence that the TP53I13 gene generates a functional protein. The lncRNA, DDSR1, has been recently characterized to function in DSB repair by HR (Sharma et al., 2015). In response to DNA damage, DDSR1 is expressed in an ATM-dependent manner and can physically bind to BRCA1. DDSR1 is important for end resection and HR but the mechanistic detail of how it accomplishes this function is unknown. Future experiments should determine if TP53I13 and C9ORF106 are also lncRNAs that can stimulate end resection and HR.

The N-terminal protein acetyltransferase, NAA10, was also identified as a candidate resection activator. N-terminal protein acetyltransferases are responsible for transferring acetyl moieties to the N-termini of 80-90% of the human proteome (Kalvik and Arnesen, 2013). N- terminal acetylation was thought to mainly promote protein stability but recent work has demonstrated additional roles including the regulation of protein subcellular localization and multiprotein complex integrity (Behnia et al., 2004; Scott et al., 2011). Like ZNF335, NAA10 was also found to be mutated in a disease that displays neonatal microcephaly as a clinical feature (Rope et al., 2011). It will be interesting in the future to determine how N-terminal protein acetylation can impact end resection and if the observed microcephaly is a consequence of defective DSB repair in the developing cerebral cortex.

Pathway enrichment analysis conducted on candidate resection activators identified in the primary screen determined an over-representation of RNA splicing factors. Other genome-scale RNAi screens monitoring DSB repair have also identified RNA splicing as an important functional category (Adamson et al., 2012; Paulsen et al., 2009). Both constitutive and alternative RNA splicing are critical steps in the production of most human proteins (Nilsen and

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Graveley, 2010). RNA sequencing and splicing sensitive microarrays have demonstrated that many messenger RNAs are alternatively splicing in response to DNA damage (Munoz et al., 2009; Paronetto et al., 2011; Savage et al., 2014; Solier et al., 2010; Tresini et al., 2015). Importantly, the end resection factors EXO1, NBS1, and WRN are alternatively spliced in response to DSBs but whether these splicing events are important for promoting end resection is unknown (Savage et al., 2014; Takai et al., 2008). Future experiments should focus on determining if the splicing factors identified in the screen can modulate EXO1, NBS1, or WRN alternative splicing and whether this regulation is required for end resection. Potential alternative splicing of other known resection activators should not be ruled out in these studies. In response to DNA damage, BRCA1 physically interacts with spliceosome components including BCLAF1 and the SF3B complex to promote alternative splicing of several DNA repair proteins including EXO1 (Savage et al., 2014). Remarkably, two SF3B subunits, SF3B2 and SF3B3, were identified as candidate resection activators in my screen. It will be interesting to investigate if any other end resection factors are alternatively spliced in a BRCA1/BCLAF1-dependent manner and if these splicing events are important for protein function in end resection.

5.2 Validation of candidate end resection inhibitors

No known end resection inhibitors were identified in the primary screen. Importantly, it is possible that the primary screen did identify end resection inhibitors with biological relevance to DNA repair as the helicase HELQ was the top siRNA pool that increased RPA32 phosphorylation. HELQ was recently shown to be important for HR repair of replication- associated DSBs (Adelman et al., 2013; Takata et al., 2013). HELQ-deficient cells have an increase in the accumulation of γH2AX, RPA32, and RAD51 foci at long time periods after replication stress. These data likely reflect that HELQ deficient cells are defective at repairing DSBs generated at replication forks and not that HELQ is an active inhibitor of end resection. In support of this hypothesis, Adelman et al. (2013) also determined that HELQ knockdown results in a HR defect using the DR-GFP reporter system. If HELQ was an active inhibitor of end resection, an increase in HR efficiency would be predicted in HELQ-deficient cells. It would be informative in the future to conduct a secondary confirmation screen on the top candidate end resection inhibitors using the FUCCI system.

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5.3 Elucidating the mechanism by which ZNF335 promotes end resection

5.3.1 Generation of a ZNF335 knockout cell line by genome editing

Determining the mechanism by which ZNF335 promotes DNA end resection requires further investigation. First, the generation of a knockout or loss-of-function cell line by CRISPR/Cas9- mediated genome editing should be completed. ZNF335 knockout cells will simplify experiments by eliminating siRNA transfections, allowing for functional studies to be conducted in consistent genetic backgrounds. My preliminary genome editing experiments suggest that ZNF335 may be essential for viability in U2OS cells. The essentiality of ZNF335 is not surprising as its loss resulted in embryonic lethality in mice (Yang et al., 2012). Impending genome editing experiments should focus on deleting the four C-terminal zinc finger domains of ZNF335 which are required for its function in end resection. C-terminally truncated ZNF335 may be able to support other potential functions and thus promote cellular viability. If the C- terminus of ZNF335 is also essential for growth, knockout cells could be generated in the context of exogenously expressed ZNF335. Recombinant ZNF335 could be fused to a destabilizing domain that rapidly results in protein degradation in the absence of a degradation inhibitor. During genome editing experiments, cells would be grown in the presence of the degradation inhibitor which would stabilize the ZNF335 fusion and promote cellular viability. When conducting downstream loss-of-function experiments, the inhibitor is removed from the system resulting in the rapid degradation of exogenous ZNF335.

5.3.2 ZNF335 recruitment to DSB sites

An additional method to determine if ZNF335 is recruited to DSB sites is to conduct chromatin immunoprecipitation (ChIP). ZNF335 ChIP would be carried out after the induction of a single I- SceI generated DSB previously engineered in U2OS cells (Dong et al., 2014; Pierce et al., 1999). Following ZNF335 ChIP, primers designed to the region flanking the DSB are used to conduct quantitative PCR to determine the relative concentration of ZNF335 at the DSB. Another method to examine if ZNF335 can promote end resection by acting directly at DSB sites is to artificially tether exogenous ZNF335 to a genomic region where DSBs have been generated. Tethering could be accomplished by employing the LacO/LacR chromatin-targeting system (Janicki et al., 2004; Shanbhag et al., 2010). A LacO array would be engineered into the genome of U2OS cells

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and after the co-expression of the nuclease LacI-mCherry-FokI and a LacI-GFP-ZNF335 fusion end resection could be measured at the array by conducting pRPA32 (S4/S8), RPA32, or BrdU immunostaining.

ZNF335 could also function in the promotion of end resection by globally binding to chromatin. Global association with chromatin could be enhanced in response to DNA damage. An examination of ZNF335 chromatin binding should be conducted in unperturbed cells and those treated with DSB-inducing agents. Chromatin association could be quantified by isolating the chromatin-bound protein fraction and conducting ZNF335 immunoblotting. Global genome profiling of ZNF335 binding could also be conducted in unperturbed and DNA damaging conditions utilizing ChIP and next-generation sequencing. ZNF335 was previously implicated in the regulation of gene expression by chromatin remodeling through a physical association with the trithorax histone methyltransferase complex (Garapaty et al., 2009; Yang et al., 2012). Importantly, my data suggests that ZNF335 is not important for regulating the expression of several known core resection factors. However, this observation does not rule out that ZNF335 could modulate the expression of other genes that directly or indirectly promote end resection. Therefore, genome-wide expression profiling in ZNF335 wildtype and knockout (or knockdown) cells by RNA-sequencing should be conducted. If any genes are found to be expressed in a ZNF335-dependent manner they should be cross-referenced to candidate resection activators identified in the RNAi screen.

5.3.3 Investigation of end resection factor recruitment to DSB sites

My data suggests that ZNF335 does not promote the expression or protein stability of the core end resection factors (MRN, CtIP, EXO1, DNA2, and BLM). However, an examination of the core end resection factors and their recruitment to DSB sites in ZNF335-deficient cells has not been conducted. An analysis of end resection factor recruitment to DSBs could provide mechanistic insights into how ZNF335 promotes end resection. Furthermore, these data could also pin-point if ZNF335 functions in the promotion of either short- or long-range end resection.

5.3.4 Candidate ZNF335 protein interaction partners

IP-MS experiments revealed several intriguing interactions with the ZNF335 C-terminus including SET, FAM115A, and RECQL. SET is a histone chaperone that promotes chromatin

120 compaction by inhibiting the action of histone acetyltransferases (Seo et al., 2001). SET was shown to inhibit end resection and HR by enhancing chromatin compaction which limited the accessibility of DNA repair factors at DSB sites (Kalousi et al., 2015). One hypothesis is that ZNF335 could inhibit the function of SET to promote end resection. Future experiments should confirm that SET is an inhibitor of resection and that it can indeed physically interact with ZNF335. Next, mutants that disrupt the ZNF335-SET interaction should be uncovered to determine if the physical association is required for SET inhibition by ZNF335. Focus should be given to another ZNF335 interacting partner, FAM115A, which was recently found to physically interact with RPA (Tkac et al., 2016). A ZNF335 interaction with FAM115A (and RPA-coated ssDNA) could facilitate the function of ZNF335 in promoting end resection. The ZNF335- FAM115A interaction should be confirmed and it should be determined if FAM115A has any role in promoting end resection. The ZNF335-RECQL interaction should also be validated as two other RecQ DNA helicases, BLM and WRN, have previously been implicated in the promotion of end resection and HR (Gravel et al., 2008; Nimonkar et al., 2011).

5.4 ZNF335 and microcephaly

Lastly, future work should focus on examining if patient cells harbouring the c.3332g>a microcephaly mutation have defective end resection, HR, and ATR signaling. Patient cells from both homozygous individuals and their heterozygous parents have been previously collected (Yang et al., 2012). Patient cells, along with isogenic wild type cells, should be tested for end resection and HR defects. Furthermore, CRISPR/Cas9 genome editing could be utilized to introduce the microcephaly mutation into cells by simultaneously introducing ZNF335-targeting guide RNAs, Cas9, and a donor template for homology-directed repair of a Cas9 generated DSB. The donor template should contain long homology arms flanking the point mutation to promote HR repair. Yang et al. (2012) generated mice with conditional knockdown of ZNF335 in the cerebral cortex by in utero electroporation of shRNA plasmids in cortical progenitor cells. ZNF335 depletion in cortical progenitor cells resulted in a proliferation defect and a severely reduced brain size at later stages of development. It will be interesting to determine if the proliferation defect observed in cortical progenitor cells depleted of ZNF335 is a consequence of defective DSB repair. The four C-terminal zinc finger domains of ZNF335 are critical for its function in end resection and future work should determine if expression of this region can rescue proliferation in ZNF335 deficient cortical progenitor cells.

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Chapter VI

Materials and methods

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All chemicals were purchased from Sigma-Aldrich unless otherwise indicated.

6.1 Tissue culture

6.1.1 Cell lines

Human cell lines were cultured in an environmental incubator set to 37 °C and 5% CO2. Cells were grown on plastic dishes and flasks and passaged utilizing trypsin (Gibco). Cells were frozen in 10% DMSO/medium using Mr. Frosty freezing containers (Nalgene) according to the manufacturer’s instruction. For long-term storage, cells were kept in a liquid nitrogen tank. All culture media were supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin/streptomycin (Gibco). U-2-OS (U2OS) cells were cultured in McCoy’s medium (Gibco). HEK293 cells and HeLa cells were cultured in DMEM (Gibco).

6.1.2 RNA interference

U2OS and HeLa cells were transfected with siRNA using a reverse transfection mode. Complexes were formed in serum-free media (Opti-MEM; Gibco) by adding siRNA (dissolved in 1X RNA buffer; Thermo Fisher Scientific) and the lipid-based transfection reagent RNAiMAX (Invitrogen) according to the manufacturer’s instruction. For all siRNA transfections, the final concentration of siRNA in media was 10 nM. For reverse transfections, siRNA complexes are added first to the tissue culture vessel followed by the addition of media containing the appropriate number of cells. After 6-8 hours-post transfection, cells were washed once with phosphate buffered saline (PBS; Gibco) and supplemented with fresh media. Table 6.1 outlines the sequences of all siRNA duplexes used in this study.

6.1.3 Generating stable cell lines by lentiviral transduction

U2OS cells stably expressing N-terminally GFP- and FLAG-tagged ZNF335 and CtIP ORFs were generated by lentiviral transduction. Tagged ZNF335 and CtIP ORFs were cloned into the doxycycline-inducible mammalian expression vector pCW57.1 (Root laboratory; Addgene # 41393; see section 6.9). pCW57.1 is compatible for packaging into lentiviral particles. To package lentiviral particles, 4 million HEK293T cells were seeded into a 10 cm dish. The next day, complexes were produced with 10 µg of plasmid DNA and LT1 transfection reagent (Mirus) in serum-free Opti-MEM as instructed by the manufacturer. The lentiviral expression vector pCW57.1 (5 µg) was co-transfected with two lentivirus packaging vectors pPAX2 (2.5

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Target Product number Duplex siRNA sequence (Dharmacon/GE) number Scrambled CTRL D-001210-03 3 Proprietary CtIP D-011376-01 1 GAGCAGACCUUUCUCAGUA CtIP D-011376-02 2 GAAGUGAACAAGAUCAUUA CtIP D-011376-03 3 CAACCAAGAUGUAUCCUUU CtIP D-011376-04 4 GAAUAGGACUGAGUACGGU CtIP Custom siRNA 1 GCUAAAACAGGAACGAAUC ZNF335 D-016173-01 1 GAACAGUGAUGACGAAACA ZNF335 D-016173-02 2 GAACGCCACUUCCGUCCAG ZNF335 D-016173-03 3 GAAAUACCGCAAGUACUAU ZNF335 D-016173-04 4 GUAACGGGCACCUCAAGUU EXO1 D-013120-01 1 GAAGUUUCGUUACAUGUGU EXO1 D-013120-02 2 GUAAAUGGACCUACUAACA EXO1 D-013120-03 3 ACUCGGAUCUCCUAGCUUU EXO1 D-013120-04 4 GUUAGCAGCAUUUGGCAUA DNA2 D-026431-01 1 GCUAAACCGUGAAGCAAGA DNA2 D-026431-02 2 CUACGUCACUUUAAAGAUG DNA2 D-026431-03 3 ACAGUUGCCUGCAUUCUAA DNA2 D-026431-04 4 UGAUAUAGAUACCCCAUUA BRCA2 D-003462-01 1 GAAACGGACUUGCUAUUUA BRCA2 D-003462-02 2 GUAAAGAAAUGCAGAAUUC BRCA2 D-003462-03 3 GGUAUCAGAUGCUUCAUUA BRCA2 D-003462-04 4 GAAGAAUGCAGGUUUAAUA PARP-1 D-006656-02 1 GAAAGUGUGUUCAACUAAU PARP-1 D-006656-03 2 GCAACAAACUGGAACAGAU PARP-1 D-006656-04 3 GAAGUCAUCGAUAUCUUUA PARP-1 D-006656-17 4 GAUAGAGCGUGAAGGCGAA MRE11 D-009271-01 1 GAUGAGAACUCUUGGUUUA MRE11 D-009271-02 2 GAAAGGCUCUAUCGAAUGU MRE11 D-009271-03 3 GCUAAUGACUCUGAUGAUA MRE11 D-009271-04 4 GAGUAUAGAUUUAGCAGAA RAD50 D-005232-01 1 GAAACAAACUGCAGAAUGU RAD50 D-005232-02 2 GAACAAGGAUCUGGAUAUU RAD50 D-005232-03 3 GCUCAGAGAUUGUGAAAUG RAD50 D-005232-05 4 UAACCUCACUGUUGGGAUA NBS1 D-009641-02 1 GGAGGAAGAUGUCAAUGUU NBS1 D-009641-03 2 GAAGAAACGUGAACUCAAG NBS1 D-009641-04 3 GAAAUGGAUUCAGUCAAUA NBS1 D-009641-18 4 ACAUGGGAUUUGAGUGAAA BLM D-007287-01 1 GAGCACAUCUGUAAAUUAA BLM D-007287-03 2 GAGAAACUCACUUCAAUAA BLM D-007287-04 3 CAGGAUGGCUGUCAGGUUA BLM D-007287-05 4 CUAAAUCUGUGGAGGGUUA RNF168 D-007152-01 1 GGAAGUGGCUGAUGACUAU RNF168 D-007152-02 2 GAAAUUCUCUCGUCAACGU RNF168 D-007152-03 3 AGAAGGAGGUGGAUAAAGA RNF168 D-007152-18 4 GAGUAUCACUUACGCGCUA

Table 6.1. List of siRNA duplexes used in this study.

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µg) and pVSV-G (2.5 µg). One day post-transfection, 5 mL of fresh media supplemented with 20 mM HEPES (pH 7.5) was added to the cells. Media containing lentivirus was harvested 48 hours-post transfection. To remove any suspended HEK293T cells the media was centrifuged for 5 min at 1000 rpm and passed through at 0.45 µm filter. Lentivirus was aliquoted and snap frozen in liquid nitrogen before being stored at -80 ºC.

U2OS lentiviral infections were conducted in 6-well plates by seeding 100,000 cells/well. Various ratios of media to lentivirus (2 mL:0 mL; 1.8 mL:0.2 mL; 1.5 mL:0.5 mL; 1 mL: 1 mL; 0.5 mL: 1.5 mL; 0 mL: 2 mL) were used for infecting U2OS cells. Each transduction was carried out in the presence of 10 µg/ml polybrene to increase virus infection efficiency. One day after infection, fresh media containing 2 µg/ml of puromycin was added to the cells. The pCW57.1 expression vector also encodes a puromycin resistance gene. Therefore, only cells successfully transduced with lentivirus will survive puromycin selection. For each transduction, I selected the media to lentivirus ratio that resulted in 10-20% survival after puromycin selection. This survival percentage corresponds to an approximate multiplicity of infection (MOI) of 0.1-0.2. Thus, the majority of U2OS cells stably expressing tagged ZNF335 and CtIP ORFs only contain a few random genome integrations per cell (according to the Poisson statistical distribution). For all experiments involving stable cell lines produced by lentiviral transduction, biological replicates were conducted with cells generated by separate infections.

6.2 Fluorescence microscopy

6.2.1 Immunofluorescence

U2OS, U2OS FUCCI, and HeLa cells were grown on glass coverslips. Cells were fixed with 4% (w/v) paraformaldehyde (PFA) in PBS for 10 min at room temperature and then permeabilized with 0.3% (v/v) Triton X-100 for 30 min at room temperature. Alternatively, cells were pretreated with nuclear extraction buffer (20 mM HEPES pH 7.5, 20 mM NaCl, 5 mM MgCl2, 1mM DTT, 0.5% NP-40, Phosphatase Inhibitor Cocktail, and Protease Inhibitor Cocktail [Roche]) for 5 min on ice before PFA fixation to remove any protein not bound to chromatin. After fixation, cells were washed with PBS three times and then blocked with ADB (Antibody Dilution Buffer; 10% normal goat serum, 0.1% Triton X-100, 0.1% saponin in PBS) for 30 min. Cells were incubated with primary antibody (diluted in ADB) for 1 h at room temperature and washed three times with PBS. A list of primary antibodies utilized in this study is outlined in

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Table 6.2. Cells were the incubated with secondary IgG antibody conjugated to various Alexa Fluor dyes (diluted in ADB; Invitrogen) for 1 h at room temperature. Secondary antibody dilutions also contained 0.5 µg/ml of DAPI to counterstain DNA. Cells were then washed three times with PBS and the coverslips were mounted onto glass slides with Prolong Gold mounting agent (Invitrogen). Micrograph images were taken on an inverted LSM780 confocal microscope (Carl Zeiss) equipped with a 63X oil immersion objective.

6.2.2 Quantitative image-based cytometry

U2OS and U2OS FUCCI cells were grown in 96- and 384-well plates. Cells were processed for immunofluorescence as described in section 6.2.1. Cell seeding, transfections, and immunostaining procedures were either conducted manually using multi-channel pipets or in an automated manner using the 96 tip Biomek liquid-handling robot (Beckman-Coulter). Micrograph images were taken with either the Celigo plate cytometer (Brooks Automation) or the InCell 6000 analyzer (GE Healthcare) using a 4X or 10X objective, respectively. The appropriate number of fields were imaged per well to ensure that all cells were imaged for each sample. Images were uploaded to an image analysis and management platform called Columbus (PerkinElmer). Image analysis software segmented each DAPI-stained nucleus and measured the mean nuclear intensity for all other channels. The percentage of cells with a mean nuclear intensity above a set threshold was calculated. In addition, cell-by-cell mean nuclear intensities for a reference and test sample were plotted in a cumulative frequency distribution to conduct the non-parametric Kolmogorov-Smirnov test. To determine the Kolmogorov-Smirnov score the maximum vertical distance between the two distributions was calculated.

6.2.3 Laser microirradiation

Laser microirradiation was performed with a LSM780 confocal microscope (Carl Zeiss) equipped with an environmental chamber set to 37 ºC and 5% CO2. Cells were treated with 1 µg/ml Hochst for 15 min before microirradiation. Localized DSB tracks were generated with a 355 nm continuous laser (40X oil immersion objective, 2X zoom, 20% output power, 1 iteration, 1 pixel width). Stripe intensities compared to background were measured using ImageJ software.

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Antibody Species Vendor Product IB IF number RPA32 Mouse Abcam Ab2175 1:500 1 h RT 1:500 1 h RT pRPA32 (S4/S8) Rabbit Bethyl A300-245A 1:10,000 1 h RT 1:1,000 1 h RT laboratories

BrdU Mouse GE Healthcare RPN202 - 1:500 1 h RT

ZNF335 Rabbit Bethyl A300-798A 1:500 ON 4 ºC - laboratories

CtIP Rabbit Abcam ab70163 1:1,000 1 h RT -

GAPDH Rabbit Sigma G9545 1:20,000 1 h RT -

Tubulin Mouse Calbiochem DM1A 1:2,000 1 h RT -

RAD51 Rabbit Bioacademia 70-001 1:10,000 1 h RT 1:15,000 1h RT

γH2AX Mouse Millipore JBW301 - 1:2,000 1 h RT pCHK1 (S345) Rabbit Bethyl 2348 1:1,000 1 h RT - laboratories

GFP Goat L. Pelletier (gift) - 1:20,000 1 h RT -

FLAG Mouse Sigma M2 1:500 ON 4 ºC 1:1,000 1 h RT

MRE11 Mouse Abcam 12D7 1:500 1 h RT -

RAD50 Mouse Abcam 13B3 1:1,000 1 h RT -

NBS1 Rabbit Novus NB100-143 1:10,000 1 h RT - Biologicals

EXO1 Rabbit Abgent AP2871a 1:500 ON 4 ºC -

DNA2 Rabbit Abgent AP10182c 1:500 ON 4 ºC -

BLM Rabbit Abcam ab2179 1:500 ON 4 ºC -

53BP1 Mouse BD Biosciences 612523 - 1:5,000 1 h RT

Table 6.2. List of primary antibodies used in this study.

IB: immunoblotting; IF: immunofluorescence; RT: room temperature; ON: overnight

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6.3 Automated genome-scale RNAi screen

The SMARTpool siRNA library (Dharmacon/GE Healthcare) targeting 18,452 genes was screened to uncover new regulators of DNA end resection. The library was housed in round bottom 96-well plates where the first and last columns were kept empty to allow for the addition of negative and positive control siRNA pools to each screening plate. Therefore, 80 library siRNA pools were present on each 96-well library plate making a total of 232 plates to screen. The library siRNA pools were previously dissolved in RNA buffer at a concentration of 2 µM. Each week, 40 96-well library plates were thawed and aliquoted (2.5 µL per well) into sterile round bottom 96-well transfection plates using the Biomek FX liquid handler (Beckman Coulter). One 384-well cell plate can receive siRNA pools from four 96-well transfection plates. Therefore, each week, 10 384-well cell plates (PerkinElmer Cell Carrier) were required. First, 1000 U2OS cells were dispensed to each well of the 10 384-well plates. The following day, the cells were forward transfected with the library siRNA pools. For the transfection, each well of the transfection plates (containing aliquoted siRNA pools) received 57.5 µL of transfection mix (0.3 µL RNAiMAX [Invitrogen] and 57.2 µL Opti-MEM [Gibco]). After 20-30 min to allow complex formation, 10 µl of each library transfection mix was added to the appropriate well of the 10 384-well cell plates. Before addition of the complexes, each well of the cell plates received 40 µL of fresh media. The cells were incubated with the siRNA pools for 48 hours. Next, cells were treated with 100 ng/ml NCS for 3 h followed by immunostaining for pRPA32 (S4/S8) as described in section 6.2.1. NCS addition and immunofluorescence was conducted on the Dimension 4 robotic platform. Plates were imaged and analyzed using the Celigo plate cytometer (Brooks Automation) and packaged image analysis software. The same method was employed for conducting secondary confirmation screens with the exception that U2OS FUCCI cells were used.

6.4 Reverse transcription and quantitative PCR

U2OS cells growing in a 6-well plate were washed twice with PBS followed by lysis and total RNA extraction employing the RNeasy mini kit (Qiagen) and instructions outlined by the manufacturer. Using 2 µg of total RNA, cDNA was generated from polyadenylated messenger RNAs by reverse transcription employing oligo(dT)20 primers (Invitrogen) and SuperScript III reverse transcriptase enzyme (Invitrogen) according to manufacturer’s instructions. Quantitative

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PCR was conducted on 250 ng of the total cDNA preparation by using a primer and FAM-linked probe (Solaris/GE Healthcare) specific to the ZNF335 sequence. The relative quantity of ZNF335 messenger RNA was determined by comparing to the reference gene GAPDH for each sample.

6.5 Western blot analysis

6.5.1 Whole cell extract preparation

Confluent plates of U2OS cells were rinsed with PBS. Cells were then scraped in 1 ml 1X PBS, pelleted by centrifugation at 500 g for 5 min and washed once more with PBS. Cell pellets were frozen in liquid nitrogen and stored at -80 °C. For lysis, cell pellets were thawed on ice and resuspended in lysis buffer (50 mM HEPES-KOH, pH 7.5, 100 mM KCl, 2 mM EDTA, 0.5 % NP-40, 10% glycerol, 0.25 mM sodium orthovanadate, 10 mM NaF, 50 mM 2- glycerolphosphate, pH 7.5, 1mM DTT, 1X protease inhibitors [Roche]) and incubated on ice for 40 min. Cell lysates were clarified by centrifugation at 13,000 g for 20 min at 4 °C. Protein concentration was determined using the bicinchoninic acid (BCA) method (Pierce) according to the manufacturer’s instructions. Lysates were frozen in liquid nitrogen and stored at -80 °C.

6.5.2 SDS-PAGE and immunoblotting

Proteins were separated by SDS-PAGE and transferred onto a nitrocellulose membrane using standard procedures (Sambrook & Russell, 2001). After a wet transfer, the membrane was blocked in 5% skim milk/ TBS-T for 30 min at room temperature and incubated with the indicated primary antibody (diluted in 5% skim milk/ TBS-T) for 1 h at room temperature or overnight at 4 ºC. Primary antibodies and details regarding their use in immunoblotting is outlined in Table 6.2. The membrane was then washed 3 times for 5 min with TBS-T, incubated with a horseradish peroxidase-conjugated secondary antibody for 1 h at room temperature, and washed again 3 times for 5 min with TBS-T. The immunoblot was developed using Supersignal West Dura (Pierce). All incubations were carried out on a horizontal shaker.

6.6 Cell cycle analysis by flow cytometry

U2OS cells growing in a 10 cm dish were trypsinized and counted. One million cells were centrifuged for 5 min at 1000 rpm. The pellet was washed once with PBS and centrifuged again.

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The pellet was re-suspended in propidium iodide staining solution (20 µg/ml propidium iodide, 2 mg DNase-free RNase A, and 0.1% (v/v) Triton X-100 in PBS) and incubated at room temperature for 15 min. Propidium iodide staining intensity was analyzed for 10,000 cells using a FACSCalibur flow cytometer (BD Biosciences) and FlowJo software.

6.7 Clonogenic survival assays

Clonogenic survival assays were conducted as outlined previously (Franken et al., 2006). Briefly, U2OS cells were reverse transfected with the indicated siRNA duplexes in 10 cm dishes. After 48 h, transfected cells were seeded into 6-well plates at various densities per well: 250, 500, 1000, 2000, 4000, and 8000 cells. One day after seeding, each siRNA treatment and density was incubated with various concentrations of NCS, ETOP, and CPT for 30 min. After treatment, cells were washed once with PBS before the addition of fresh media. Colonies were allowed to growth for two weeks before fixation and staining with methanol and 0.5% (w/v) crystal violet, respectively. Only wells containing 5-100 colonies were counted. The plating efficiency of cells treated with each of the siRNAs was calculated using the following formula:

100

Next, the number of colonies on each plate exposed to the various concentrations of DSB- inducing drugs was counted and the surviving fraction was calculated using the following formula:

6.8 Homologous recombination assay utilizing the DR-GFP reporter system

DR-GFP U2OS cells were previously engineered to have a single I-SceI endonuclease consensus site surrounded by two non-functional GFP alleles integrated into their genome (Pierce et al., 1999). U2OS DR-GFP cells were reverse-transfected with siRNAs in a 96-well plate and the following day forward-transfected with pCBASceI (Addgene #26477) using Lipofectamine 2000 (as described by manufacturer; Invitrogen) to introduce the expression of the I-SceI endonuclease. After 48 h post-I-SceI transfection, U2OS DR-GFP cells were fixed with 4%

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(w/v) PFA for 10 min and permeabilized with 0.3% (v/v) Triton X-100 for 10 min. Next, cells were incubated with 0.5 µg/ml DAPI for 15 min to visualize nuclei. Cells were imaged on the automated InCell 6000 Analyzer confocal microscope and the percentage of GFP-positive cells was determined using Columbus image analysis software (PerkinElmer).

6.9 Plasmids

CtIP ORF was obtained from Steve Jackson’s laboratory (University of Cambridge, UK). ZNF335 ORF was obtained from OpenFreezer (LTRI). The CtIP and ZNF335 ORFs were PCR amplified and cloned into the multiple cloning site of pcDNA 5/FRT/TO (Thermo Fisher Scientific) using restriction endonucleases. CtIP and ZNF335 were cloned into two pcDNA 5/FRT/TO vectors, each containing a distinct polylinker to add either an N-terminal GFP or FLAG tag. Silent point mutations were generated by Quikchange PCR (Stratagene) to produce CtIP and ZNF335 ORFs that were resistant to the siRNAs GCUAAAACAGGAACGAAUC and GAAAUACCGCAAGUACUAU, respectively. RNAi-resistant GFP- and FLAG-tagged CtIP and ZNF335 were PCR amplified with 5’ attB1 and 3’ attB2 Gateway-compatible sites (Thermo Fisher Scientific). Using the Gateway BP clonase enzyme, the PCR products were inserted into the Gateway entry vector pDONR221. The LR clonase enzyme was used to transfer GFP- and FLAG-tagged CtIP and ZNF335 into the Gateway destination vector pCW57.1 (Root laboratory; Addgene # 41393). ZNF335 point mutations and deletions were generated in pCW57.1 by Quikchange PCR (Stratagene).

6.10 Immunoprecipitation coupled to mass spectrometry

ZNF335 FLAG affinity purification was conducted using a protocol that allows for simultaneous identification of both soluble and chromatin-bound interaction partners. The protocol described in Lambert et al. (2014) was followed. Briefly, near confluent U2OS cells on five 15 cm plates were collected by scraping and centrifugation for 5 min at 1,000 rpm. Cell pellets were snap frozen in liquid nitrogen and the frozen pellet was lysed with 1 mL of lysis buffer (50 mM HEPES-NaOH pH 8.0, 100 mM KCl, 2 mM EDTA, 0.1% NP40, 10% glycerol, 1 mM PMSF, 1 mM DTT, and protease inhibitor cocktail). Each sample was sonicated for 30 sec (10 sec ON, 2 sec OFF cycles) at amplitude 0.35 using QSONICA 125W sonicator equipped with 1/8” probe to shear DNA. 1 µL of benzonase was added to each sample and incubated at 4 °C for 1 h to further digest chromatin. The lysates were centrifuged at 14,000 rpm for 20 min at 4 °C. Anti-FLAG M2

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magnetic beads were prepared as follow: 25 µL of the 50% slurry was aliquoted for each sample (5 x 15 cm plates), and the beads were washed with 3 x 1 mL of cold lysis buffer. The centrifuged cell lysates were transferred to the appropriate tube containing the anti-FLAG magnetic beads. The mixture was incubated for 2 h at 4 °C with gentle agitation. Beads were pelleted by centrifugation (1,000 rpm for 1 min). The tubes were placed on a cold magnetic rack (placed on ice) to collect the beads on the side of the tubes. The supernatant was removed slowly with a pipette and the beads re-suspended in 1 mL of cold lysis buffer. The beads were then transferred to a fresh tube with 1 mL of 20 mM Tris-HCl (pH 8.0) containing 2mM CaCl2. Following the last wash, the samples were quickly centrifuged and the last drops of liquid removed. The dried beads removed from the magnet were re-suspended in 7.5 µL of 20 mM Tris-HCl (pH 8.0) containing 500 ng of trypsin, and the suspension was incubated at 37 °C with agitation overnight. After this first incubation, the sample was quickly centrifuged, then magnetized and the supernatant transferred to a fresh tube. Another 500 ng of trypsin was added in 5 µL of 20 mM Tris-HCl (pH 8.0) and the resulting sample was incubated at 37 °C for 3-4 hours. Formic acid was added to the sample to a final concentration of 2% and stored at -80 °C.

Samples were analyzed on the AB SCIEX 5600 TripleTOF in Data dependent acquisition (DDA) mode. A quarter of the volume of the digested sample (~3 µL) was analyzed using a homemade column (i.d. 100 µm x 10 cm). The column was coupled to a NanoLC-Ultra 1D plus (Eksigent, Dublin, CA) system with 0.1% formic acid in water as buffer A and 0.1% formic acid in ACN as buffer B. To identify significant interaction partners from the affinity purification data, the data were subjected to SAINT analysis implemented in ProHits. Proteins with average SAINT score (AvgP) greater than 0.9 were considered to be statistically significant. Artifact proteins (such as trypsin and keratin) were manually curated away from the final interaction partner list.

6.11 Solid-phase reverse siRNA transfections on 1536-well cell microarrays

Gelatin-based solid-phase siRNA transfections were conducted according to Erfle et al. (2007). Solid-phase transfection complexes were generated in serum-free Opti-MEM (Gibco) and the final concentration for each ingredient after dispensing 500 nL was 20 ng siRNA, 5% RNAiMAX (Invitrogen), 0.08% gelatin, 32 mM sucrose, and 0.004% fibronectin. 50-100 nL of

132 solid-phase transfection mixes were dispensed to each well on a 1536 array using the Sciclone inL10 (Caliper) small-volume liquid handling robot. Printed arrays were dried in a vacuum desiccator for 48 h before flooding with media containing HeLa cells. After 48 h, arrays were processed for immunostaining (as described in section 6.2.1). After staining, arrays were mounted on a large glass slide in 50 µL of Prolong Gold mounting agent (Invitrogen). Mounted arrays were imaged on the InCell 6000 Analyzer. A 1536 array map generated using the InCell software was aligned with the mounted array before imaging. Image analysis was conducted using Columbus (PerkinElmer).

6.12 Solid-phase transfections in arrayed drops (SPIDR)

Solid-phase transfection complexes for SPIDR were produced in Opti-MEM (Gibco) and the final concentration of each component after dispensing 100 nL using the Sciclone inL10 was 2 ng siRNA, 0.3% RNAiMAX (Invitrogen), 0.005% gelatin, 2 mM sucrose, and 0.0003% fibronectin. After delivering transfection complexes, printed 1536 arrays were dried in a vacuum desiccator for 48 h. Next, the Sciclone inL10 was used to deliver 500 nL of media containing HeLa cells. A HeLa cell suspension was used to allow the delivery of 500 cells per well. The following day, the 1536 array was washed once with PBS and flooded with fresh media. After 48 h post-seeding, cells were processed for immunofluorescence as described in section 6.2.1). After staining, arrays were mounted on a large glass slide in 50 µL of Prolong Gold mounting agent (Invitrogen). Mounted arrays were imaged on the InCell 6000 Analyzer. A 1536 array map generated using the InCell software was aligned with the mounted array before imaging. Image analysis was conducted using Columbus (PerkinElmer).

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Chapter VII

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Appendix I

Genome-scale RNAi screen data

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Appendix I is presented in an electronic format and is written on the included DVD.

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Appendix II

Secondary confirmation screen data

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Appendix IIA: Data from secondary confirmation screen using the FUCCI system and cherry-picked siRNA pools.

Appendix IIB: Data from secondary confirmation screen using the FUCCI system and deconvolved siRNA duplexes.

Appendix IIA and IIB are presented in an electronic format and are written on the included DVD.

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Appendix III

53BP1 focus formation SPIDR pilot screen data

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Appendix III is presented in an electronic format and is written on the included DVD.