The Recognition and Mobility of DNA Double-Strand Breaks

Jonathan Strecker

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

The Recognition and Mobility of DNA Double-Strand Breaks Jonathan Strecker

Doctor of Philosophy

Graduate Department of Molecular Genetics University of Toronto

2016 Abstract

DNA double-strand breaks (DSBs) pose a threat to cell survival and genomic integrity, and remarkable mechanisms exist to deal with these breaks. A single DSB activates a signalling response that profoundly impacts cell physiology, not least through the engagement of DSB repair pathways and the arrest of cell division. Here I study these processes in the budding yeast

Saccharomyces cerevisiae and investigate two central themes to this response. First, I examine how the natural ends of chromosomes, telomeres, are differentiated from DSBs by generating

DNA ends with increasing telomeric character. I discover a striking transition in the activity of the telomerase inhibitor Pif1 at these ends and propose that this is the dividing line between DSBs and telomeres. Second, I investigate a phenomenon whereby a DSB increases the mobility of chromosomes within the nucleus. This increase in mobility is dependent on the Mec1 kinase and is proposed to promote repair by homologous recombination. I identify that the Mec1-dependent phosphorylation of Cep3, a kinetochore component, is required to stimulate chromatin mobility following DNA breakage and provide a new model for how a DSB affects the constraints on chromosomes. Unexpectedly, I find that increased mobility is not required for DSB repair and instead propose that Cep3 helps arrest the cell cycle in response to a DSB. Finally, my investigation into Cep3 phosphorylation provides new insight into the role of chromatin remodelers and the histone variant H2A.Z in DSB-induced chromatin mobility.

Acknowledgments

I would like to thank my supervisor Daniel Durocher for his mentorship and support, it has been an absolute privilege to begin my journey in such a thriving and successful environment. Many of your lessons and philosophies will undoubtedly remain with me during my career.

I thank my committee members Brenda Andrews and Karim Mekhail for their thoughtful comments and suggestions over the years.

Finally, I am grateful to members of the Durocher lab past and present, with particular mention to Wei Zhang who taught me the basics of budding yeast and whose results and ideas led to the projects described here.

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Table of Contents

Acknowledgments...... iii Table of Contents ...... iv List of Figures ...... viii List of Abbreviations ...... x Chapter 1 Introduction ...... 1 1.1 DNA double-strand breaks threaten genome integrity ...... 1 1.1.1 The causes and consequences of DSBs ...... 1 1.1.2 Genome instability and cancer development ...... 2 1.2 The DNA double-strand break response ...... 3 1.2.1 The sensing and signalling of DSBs ...... 3 1.2.2 Repair of DSBs ...... 6 1.2.3 DSBs arrest cell division ...... 12 1.3 DNA ends in the cell ...... 13 1.3.1 The structure and function of telomeres ...... 13 1.3.2 Telomerase activity and regulation...... 15 1.3.3 Interplay between DSBs and telomeres ...... 18 1.3.4 The telomerase inhibitor Pif1 ...... 18 1.4 DSB repair in the context of the nucleus ...... 20 1.4.1 Spatial organization of the nucleus ...... 20 1.4.2 Nuclear position and genomic integrity...... 22 1.4.3 Chromatin is not static ...... 24 1.4.4 Chromatin mobility following a DSB ...... 26 1.5 Summary and rationale ...... 27 Chapter 2 Materials and Methods ...... 29 2.1 General yeast strains and growth ...... 29 2.2 Telomere addition strains ...... 29 2.3 Chromatin mobility strains ...... 30 2.4 Telomere addition analysis ...... 31 2.5 Genomic DNA extraction ...... 31 2.6 Southern blots for telomere addition and length ...... 31

2.7 PCR mutagenesis screens ...... 32 2.8 Chromatin mobility analysis ...... 33 2.9 Antibodies and immunoblotting ...... 33 2.10 Cell cycle arrest ...... 34 2.11 Recombinant protein production ...... 34 2.12 Rad53 kinase reactions ...... 35 2.13 Visualization of kinetochores ...... 35 2.14 Analysis of SPB-CEN dynamics ...... 36 2.15 DNA damage sensitivity ...... 36 2.16 HR repair analysis ...... 36 2.17 Checkpoint analysis ...... 37 2.18 A-like faker (ALF) assay ...... 37 2.19 Chromosome transmission fidelity (CTF) assay ...... 37 2.20 Chromatin immunoprecipitation (ChIP) ...... 38 2.21 Peptide pulldown assays ...... 38 2.22 Break-induced replication (BIR) assay ...... 38 2.23 Break-induced replication PCR assay ...... 39 2.24 Cell senescence assays ...... 39 2.25 Yeast two-hybrid ...... 39 2.26 Statistics ...... 40 Chapter 3 A Pif1-dependent threshold separates DSBs and telomeres ...... 41 3.1 Introduction ...... 42 3.2 Results ...... 42 3.2.1 Identification of a threshold for Pif1 sensitivity ...... 42 3.2.2 Pif1 is not inhibited by DNA damage kinases ...... 46 3.2.3 Artificial telomerase recruitment does not outcompete Pif1 ...... 50 3.2.4 The DSB-telomere transition recapitulates the differential regulation of Pif1 ...... 51 3.2.5 Investigating the molecular trigger of the DSB-telomere transition ...... 53 3.2.6 Cdc13 and the fate of DNA ends ...... 54 3.2.7 Pif1 does not limit elongation of longer telomeric seeds ...... 60 3.3 Discussion ...... 64 Chapter 4 DNA damage signalling targets the kinetochore to promote chromatin mobility ...... 67

4.1 Introduction ...... 68 4.2 Results ...... 68 4.2.1 A system for measuring chromatin mobility ...... 68 4.2.2 Relieving constraints on chromosomes mimics DSB-induced mobility ...... 69 4.2.3 Identification of a kinetochore mutant that affects DSB-induced mobility ...... 69 4.2.4 Cep3 is phosphorylated in response to DNA damage ...... 75 4.2.5 DSB signalling modulates centromere attachment to the spindle pole body ...... 77 4.2.6 The unbroken chromosome arm retains telomeric constraint ...... 83 4.2.7 Cep3 regulates the global chromatin response to a DSB ...... 85 4.2.8 DSB-induced chromatin mobility is dependent on the centromeric constraint ...... 90 4.2.9 DSB-induced chromatin mobility can be dispensable for repair ...... 91 4.2.10 Cep3 phosphorylation promotes checkpoint arrest ...... 94 4.3 Discussion ...... 96 Chapter 5 Pursuing the molecular events downstream of Cep3 phosphorylation ...... 99 5.1 Introduction ...... 100 5.2 Results ...... 100 5.2.1 The Rpd3L complex is required for DSB-induced chromatin mobility ...... 100 5.2.2 Mimicking Htz1 acetylation abolishes DSB-induced chromatin mobility ...... 103 5.2.3 Increasing chromatin mobility in the absence of DNA damage ...... 107 5.2.4 A centromeric function for chromatin remodelers in DSB-induced mobility ...... 109 5.2.5 Aurora B is required for DSB-induced mobility ...... 111 5.3 Discussion ...... 113 Chapter 6 Summary and future directions ...... 117 Appendix A: The kinesin-14 motor complex promotes general chromatin mobility and atypical DSB repair ...... 124 Introduction ...... 125 Results ...... 125 Kinesin-14 promotes chromatin mobility independent of DNA damage ...... 125 Cik1-Kar3 are important for DNA repair by break-induced replication ...... 128 Discussion ...... 131 Appendix B: Summary of DNA ends ...... 133 Appendix C: Summary of chromatin mobility parameters ...... 135

Appendix D: List of yeast strains...... 140 References ...... 153

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List of Figures Figure 1.1. DSB repair pathways ...... 8 Figure 1.2. Repair of DSBs by homologous recombination ...... 11 Figure 1.3. Analysis of chromatin mobility ...... 25

Figure 3.1. Characterizing a threshold of Pif1 activity at DNA ends ...... 44 Figure 3.2. Pif1 activity at DNA ends reveals a DSB-telomere transition ...... 47 Figure 3.3. Pif1 is not inactivated by Tel1 at short telomeres ...... 48 Figure 3.4. Loss of MEC1 or RAD53 does not affect Pif1 at short telomeres ...... 49

Figure 3.5. A genetic screen to identify Pif1 mutants that inhibit telomerase at the TG82 end ..... 50 Figure 3.6. Artificial telomerase recruitment does not overcome Pif1 activity ...... 51 Figure 3.7. The DSB-telomere transition recapitulates the differential regulation of Pif1 ...... 52 Figure 3.8. The DSB-telomere transition does not require Rap1 ...... 54 Figure 3.9. The N-terminal dimerization mutant Cdc13-L91A impairs telomere addition ...... 57

Figure 3.10. A screen to identify Cdc13 mutants that prevent telomere addition at the TG34 end 59 Figure 3.11. Mutations in cdc13-sp alleles ...... 61

Figure 3.12. Cdc13 mutations which sensitize the TG34 end to Pif1 activity ...... 62 Figure 3.13. The preferential extension of short telomeres is independent of Pif1 ...... 63 Figure 3.14. A model for the length-dependent regulation of telomerase at DNA ends ...... 64

Figure 4.1. A system for measuring chromatin mobility ...... 70 Figure 4.2. Relieving constraints on chromosomes mimics DSB-induced chromatin mobility ... 72 Figure 4.3. A kinetochore component is required for DSB-induced chromatin mobility ...... 73 Figure 4.4. Cep3-S575 is required for DSB-induced mobility in multiple contexts ...... 74 Figure 4.5. Cep3 is phosphorylated after DNA damage ...... 75 Figure 4.6. Cep3 phosphorylation is dependent on Mec1 signalling ...... 78 Figure 4.7. Rad53 and Dun1 are required for Cep3 phosphorylation and DSB-induced chromatin mobility ...... 79 Figure 4.8. Rad53 phosphorylates Cep3 in vitro ...... 81 Figure 4.9. Cep3 modulates centromere attachment to the SPB ...... 82 Figure 4.10. Disrupting kinetochore-microtubule attachment in cep3-S575A mutants ...... 83 Figure 4.11. The broken chromosome arm is more mobile than the unbroken arm ...... 84

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Figure 4.12. Telomeres are not detached following a DSB ...... 86 Figure 4.13. The global chromatin response requires a centromere-proximal DSB ...... 87 Figure 4.14. Global mobility requires Cep3-S575 but retains telomeric based constraint ...... 88 Figure 4.15. Loss of telomeric tethering does not increase mobility of a broken chromosome ... 89 Figure 4.16. The mobility of an episome is increased after DNA damage ...... 90 Figure 4.17. Centromeric constraint dictates the increase in mobility after a DSB ...... 92 Figure 4.18. Increased chromatin mobility is dispensable for DSB repair ...... 93 Figure 4.19. Cep3 phosphorylation promotes cell cycle arrest and genome stability ...... 95 Figure 4.20. Proposed model of how a DSB increases chromatin mobility ...... 96

Figure 5.1. Cep3-S575A does not affect chromatin enrichment ...... 101 Figure 5.2. Valproic acid is a potent inhibitor of DSB-induced chromatin mobility ...... 102 Figure 5.3. Rpd3 is required for DSB-induced chromatin mobility ...... 104 Figure 5.4. The Rpd3L complex is a putative Cep3-pS575 interactor ...... 105 Figure 5.5. Mimicking Htz1 acetylation impairs DSB-induced mobility and checkpoint arrest 108 Figure 5.6. htz1-4KR promotes chromatin mobility in the absence of a DNA break ...... 110 Figure 5.7. The arp8Δ mutant fails to modulate SPB-CEN attachment after a DSB ...... 112 Figure 5.8. Ipl1 promotes DSB-induced chromatin mobility ...... 114

Figure 6.1. In vitro reconstitution of the DSB-telomere transition...... 118 Figure 6.2. Does increased chromosome mobility function in the proofreading of donor sites? 122

Appendix A Figure 1. Cik1-Kar3 promote general chromatin mobility ...... 126 Appendix A Figure 2. Kar3 ATPase activity is required for chromatin mobility ...... 127 Appendix A Figure 3. Cik1-Kar3 promotes break-induced replication...... 129 Appendix A Figure 4. Cik1-Kar3 promotes chromatin mobility during BIR ...... 131

List of Abbreviations αAA...... alpha-aminoadipic acid Ala, A...... alanine APC/C...... anaphase promoting complex/cyclosome ATP ...... adenosine triphosphate ATM ...... ataxia telangiectasia mutated ATR...... ataxia telangiectasia related BIR...... break-induced replication bp ...... base pairs CDK...... cyclin dependent kinase CEN...... centromere ChIP...... chromatin immunoprecipitation C-NHEJ...... classical non-homologous end-joining DTT ...... dithiothreitol DNA...... deoxyribonucleic acid dHJ...... double Holliday junction dsDNA...... double strand DNA DSB...... DNA double-strand break E. coli ...... Escherichia coli GFP...... green fluorescent protein HO...... homothallic HR ...... homologous recombination h ...... hour HU...... hydroxyurea kb ...... kilobases kDa ...... kilodalton MMEJ...... microhomology-mediated end-joining MAT ...... mating type locus min ...... minute MMS...... methyl methane sulphonate NHEJ ...... non-homologous end-joining OB ...... oligonucleotide/oligosaccharide binding PCR...... polymerase chain reaction PI3K...... phosphatidyl inositol 3-kinase rDNA...... ribosomal DNA RNA...... ribonucleic acid RPA...... replication protein A SAC...... spindle assembly checkpoint SDS...... sodium dodecyl sulfate Ser, S...... serine SSA ...... single strand annealing S. cerevisiae...... Saccharomyces cerevisiae ssDNA ...... single-strand DNA Thr, T...... threonine WT ...... wild type YFP...... yellow fluorescent protein

Chapter 1 Introduction 1.1 DNA double-strand breaks threaten genome integrity

1.1.1 The causes and consequences of DSBs

It is imperative that cells safeguard their genomes against alterations and properly segregate genetic material to their daughters. This task is challenged by the constant DNA damage that occurs to cells which threatens genomic stability (Lindahl, 1993). A variety of DNA lesions can arise in the order of thousands per cell division including pyrimidine dimers, base deamination, oxidation damage, single-stranded DNA nicks, and interstrand crosslinks, and numerous enzymatic pathways exist to counteract these alterations (Lindahl and Wood, 1999; Sancar and Tang, 1993). The most dangerous lesion, however, is thought to be the DNA double-strand break (DSB). While DSBs occur at the lowest frequency of all DNA lesions, estimated at 10-50 DSBs per human cell per cell division (Haber, 1999; Vilenchik and Knudson, 2003), work in the budding yeast Saccharomyces cerevisiae has shown that a single DSB in a dispensable episome is capable of inducing cell death (Bennett et al., 1993).

DSBs can arise from a variety of endogenous and exogenous sources including reactive oxygen species and ionizing radiation. The greatest source of physiological DSBs, however, is from DNA replication and DSBs can form if the replication machinery dissociates at a stalled replication fork, termed fork collapse. Replication fork stalling can occur due to single-strand DNA nicks and at common fragile sites within the genome, some of which begin replication early in S-phase (Barlow et al., 2013). Fragile sites are found across species and sites of replication fork stalling largely correlate with sites of DNA damage (Szilard et al., 2010). Although DSBs are cytotoxic to cells, paradoxically DSBs are required for several physiological processes including V(D)J recombination during antibody diversification where segments of the immunoglobulin gene are randomly shuffled to allow for specific antibodies against a countless number of novel antigens (Schatz and Ji, 2011). The recombination of homologous chromosomes in meiosis also requires DSBs mediated by the topoisomerase Spo11 for chromosome crossovers and proper segregation (Keeney, 2001). Finally, the recent advent of CRISPR-Cas9 systems typically relies on

2 programmed DSBs to initiate genome editing (Cong et al., 2013) and understanding how cells deal with these breaks remains a key issue in harnessing their potential.

1.1.2 Genome instability and cancer development

DSBs threaten genomic integrity and the inability to repair DSBs is therefore a dangerous situation for cells; the consequences of which are most clearly observed in cancer. The first connection between DNA mutations and cancer was made over a century ago by the observation of visible chromosome abnormalities in cancer cells (Boveri, 1914; von Hansemann, 1890). We now appreciate that human cancers can exhibit striking genome instability, both in changes at the nucleotide level, and larger structural chromosomal rearrangements or changes in copy number (Lengauer et al., 1998). DNA sequencing analysis reveals unprecedented insight into the molecular steps of cancer progression including hundreds to thousands of individual mutations across diverse sub-clones within a single tumour (Nik-Zainal et al., 2016; Nik-Zainal et al., 2012), and the positive selection of oncogenic driver mutations in normal skin (Martincorena et al., 2015).

The development of cancer has two key connections with DNA damage and DSBs. First, mutations in the molecular pathways that safeguard our genomes can lead to the accumulation of additional mutations and cell transformation (Khanna and Jackson, 2001). The increased rate of acquired mutations, called the mutator phenotype, has been described in both yeast and human cells, and can fuel further genomic instability (Bielas et al., 2006; Loeb et al., 1974). It is therefore unsurprising that the mutation of many DSB repair factors causes an increased risk of cancer. Notable clinical examples include breast and ovarian cancer linked to mutations in the BRCA1 and BRCA2 tumour suppressor genes (Ford et al., 1998), Bloom syndrome caused by mutations in the BLM helicase (Ellis et al., 1995), and ataxia-telangiectasia caused by defects in the ATM gene (ataxia-telangiectasia mutated) (Savitsky et al., 1995; Taylor et al., 1975). In addition to cancer predisposition, mutations in DSB repair pathways can also include symptoms of developmental, immunological, and neurodegenerative defects, highlighting the importance of these repair pathways in the overall biology of organisms.

The second connection between cancer and DNA damage is how the latter can be used for cancer therapy. Chemicals that damage DNA can lead to the development of cancer (Loeb and Harris, 2008), and it is therefore counterintuitive that many modern chemotherapies also exploit the cytotoxic properties of DSBs to kill cancer cells (Lord and Ashworth, 2012). The sensitivity of

3 cancer cells to DNA damage may result from the increased background levels of damage, or through the loss of particular DNA repair pathway which increases the dependence on other pathways (Jackson and Bartek, 2009). A classic example is the use of platinum salts which cause inter- and intra-strand DNA crosslinks and pose a particular problem to cells defective in the pathways required for their repair (Lord and Ashworth, 2012). A recent case exemplifying the benefit of understanding DNA damage repair pathways is the use of PARP1 inhibitors which are particularly toxic to BRCA1 and BRCA2 deficient cells (Farmer et al., 2005).

1.2 The DNA double-strand break response

1.2.1 The sensing and signalling of DSBs

DNA damage elicits a profound cellular response across species (Witkin, 1976; Zhou and Elledge, 2000) and budding yeast has been a particularly instrumental model system to unravel the response to a DSB. The strengths of this organism have included the ease of genetic manipulations for making gene knockouts, the design of powerful genetic screens, and the ability to create experimental systems monitoring specific aspects of DSB signalling and repair (Aylon and Kupiec, 2004). Many such systems harness the HO endonuclease, physiologically used for mating type switching, to induce a single DSB at a specific location in the genome (Haber, 2012). Despite divergence from a common ancestor approximately one billion years ago (Douzery et al., 2004), budding yeast and humans share a large number of orthologous genes and over one third of the yeast genome has an identifiable human counterpart (O'Brien et al., 2005). Highlighting this conservation is the observation that nearly half of essential yeast genes with a direct human ortholog can be replaced with their human counterpart (Kachroo et al., 2015).

The DSB response is a classic signal transduction pathway consisting of sensors which detect broken DNA ends and activate DNA damage kinases which propagate the signalling response to downstream effector proteins. This signalling response communicates the presence of DNA damage throughout the cell, coordinates repair of the break, and delays cell cycle progression to allow time for repair. While this introduction will focus on pathways and components characterized in budding yeast, key components in mammalian cells will also be highlighted.

Two key complexes bind and recognize DSBs including the MRX complex, comprised of Mre11, Rad50 and Xrs2, and the Ku complex, consisting of the Yku70 and Yku80 subunits. MRX is one of the first protein complexes at DNA ends (Lisby et al., 2004; Trujillo et al., 2003) and can directly bind DNA and link opposing DSB ends (Chen et al., 2001a; de Jager et al., 2001). All three subunits of the MRX complex are important for its function in DSB signalling (D'Amours and Jackson, 2001; Grenon et al., 2001; Ivanov et al., 1994) and Mre11 exhibits both endonuclease and exonuclease activity (Paull and Gellert, 1998; Trujillo and Sung, 2001). These enzymatic activities are key in the processing of DSB ends and Mre11 nicks and resects DNA in a 3’ to 5’ direction towards the DSB to facilitate the loading of additional resection machinery (Cannavo and Cejka, 2014). The MRX complex can also tether DSB ends together mediated by the long coiled coil domains and zinc hooks of Rad50 (Hopfner et al., 2002). The second complex, the Ku heterodimer, forms a ring shape and loads onto DSB ends (Walker et al., 2001). Ku is proposed to adopt a “two-face” conformation, with the Yku70 subunit facing outward towards the DSB end, and promotes repair by NHEJ (Ribes-Zamora et al., 2007). Ku binds preferentially to double- stranded DNA (dsDNA) versus single-stranded DNA (ssDNA) (Blier et al., 1993; Falzon et al., 1993) and DNA end processing therefore antagonizes Ku binding.

The sensing of DSB ends activates the phosphatidyl inositol 3-kinase (PI3K)-related family of kinases which propagate the DSB signal and coordinate the cellular response. These kinases include Mec1 and Tel1 in budding yeast (ATM, ATR, and DNA-PK in mammals), and are central to the DNA damage response. The consensus phosphorylation site of Mec1 and Tel1 is serine or threonine residues followed by glutamine (S/T-Q) and both kinases can compensate for each other to some degree. Tel1 is recruited to DSB sites through the Xrs2 component of the MRX complex (Nakada et al., 2003), while Mec1 is recruited to ssDNA through its cofactor Ddc2 (Paciotti et al., 2000; Rouse and Jackson, 2000; Wakayama et al., 2001; Zou and Elledge, 2003). Mec1 plays a more prominent role than Tel1 in response to DNA damage and the exposure of ssDNA from DNA resection is therefore a key requirement in activation of the DNA damage response (Garvik et al., 1995). DNA damage signalling also requires the 9-1-1 clamp complex comprised of Ddc1-Rad17- Mec3 (Majka and Burgers, 2003) which has similarity to the PCNA clamp used in DNA replication (Venclovas and Thelen, 2000). The 9-1-1 complex is loaded by the Rad24-containing replication factor C complex (Ellison and Stillman, 2003; Kim and Brill, 2001) and is required for Mec1

5 signalling, but does not require Mec1 for localization to DSBs (Kondo et al., 2001; Melo et al., 2001), suggesting that it recognizes DNA damage independently.

Mec1 activation requires ssDNA and therefore plays a more prominent role during the S and G2 phases of the cell cycle due to the stimulation of DNA end resection by the cyclin-dependent kinase Cdk1 (Ira et al., 2004). One critical target of Mec1 and Tel1 is the phosphorylation of histone H2A at serine 129 (called γ-H2A upon phosphorylation) which can serve as a marker of DSB sites (Downs et al., 2000; Rogakou et al., 1999). Similarly, phosphorylation of the histone variant H2A.X at serine 139 marks DSBs in human cells (Rogakou et al., 1998). γ-H2A plays a critical role in the recruitment of chromatin remodelers to DSB sites including SWR1 and INO80 (Downs et al., 2004; Morrison et al., 2004; van Attikum et al., 2004) and cohesion which physically tethers sister chromatids (Strom et al., 2004). Chromatin remodelers promote DSB repair by facilitating access to DNA through the movement and eviction of nucleosomes (Price and D'Andrea, 2013; Soria et al., 2012).

Mec1 and Tel1 lie at the top of the signalling cascade and activate additional downstream kinases including Rad53 and Chk1 (CHK2 and CHK1 in humans). Rad53 is phosphorylated and activated by Mec1 through the Rad9 adaptor protein (Schwartz et al., 2002; Sweeney et al., 2005) or via Mrc1 in S phase (Alcasabas et al., 2001; Osborn and Elledge, 2003). Chk1 activation is similarly dependent on Rad9 (Sanchez et al., 1999). Rad53 binds and activates an additional kinase, its paralog Dun1 (Bashkirov et al., 2003; Chen et al., 2007), and the phosphorylation and resulting gel mobility shift of both Rad53 and Dun1 after DNA damage serves as an indicator of DNA damage signalling.

Activation of DNA damage kinases results in profound changes to the phosphoproteomic landscape following a DSB. Notable targets in budding yeast include the ribonucleotide reductase inhibitor Sml1, whose phosphorylation and degradation after DNA damage increases nucleotide levels (Zhao et al., 1998; Zhao and Rothstein, 2002), and DNA repair proteins such as Rad55 (Herzberg et al., 2006) and Nej1 (Ahnesorg and Jackson, 2007). The identification of in vivo DNA damage phosphorylation targets is a key objective in the field and has been largely driven by quantitative phosphoproteomic mass spectrometry following DNA damage induction. Comparison of phosphopeptides from wild-type and kinase null mutants has revealed over eighty substrates which are dependent on DNA damage kinases (Chen et al., 2010; Smolka et al., 2007). Work in

6 mammalian cells has revealed a similarly extensive network with hundreds of targets dependent on the ATM and ATR kinases (Bennetzen et al., 2010; Bensimon et al., 2010; Matsuoka et al., 2007). The remarkable scale of the DNA damage response suggests that many cellular pathways are manipulated following DNA damage and an important objective is to understand the role of these modifications in maintaining genomic integrity. While these mass spectrometry studies are a powerful resource, they do not guarantee that identified sites will have a biological function and careful in-depth studies are therefore required to confirm individual sites and to test for functional consequences, typically through non-phosphorylatable alanine and phosphomimetic glutamic acid substitutions.

In addition to widespread phosphorylation, DNA damage also induces a transcriptional response involving hundreds of up- and down-regulated genes (Gasch et al., 2001) and the extensive modification of transcription factor binding sites (Workman et al., 2006). The DSB response in mammalian cells is initially controlled through phosphorylation based signals, but also makes extensive use of regulatory ubiquitylation events to coordinate the hierarchical recruitment of repair factors to sites of damage (Panier and Durocher, 2009; Panier et al., 2012).

Altogether, the DSB response has a profound impact on numerous aspects of cell physiology, but the two most fundamental goals of this response are to coordinate the faithful repair of the break, and to stop cell cycle progression. A third outcome, especially important in multicellular organisms, is to activate cell death by apoptosis (Roos and Kaina, 2006).

1.2.2 Repair of DSBs

DSBs are repaired by two primary mechanisms: homologous recombination (HR), and nonhomologous end-joining (NHEJ). Both pathways are mutually exclusive and commitment to either process therefore requires a careful decision by the cell (Figure 1.1). As the name implies, NHEJ consists of a ligation reaction to rejoin DNA ends and does not depend on the DNA sequence at either end. Importantly, NHEJ can occur throughout the cell cycle and an essential requirement is that DNA ends are minimally processed and remain double-stranded. Core components of the classical NHEJ (C-NHEJ) pathway involve the Ku complex which binds double-stranded DNA ends and facilitates the recruitment of downstream components: DNA ligase IV (Dnl4) which catalyzes end-joining and is stabilized by Lif1 (Herrmann et al., 1998), and Nej1 which stabilizes

Ku and Dnl4-Lif1 (Chen and Tomkinson, 2011; Frank-Vaillant and Marcand, 2001; Valencia et al., 2001). The MRX complex is also required for NHEJ in budding yeast (Boulton and Jackson, 1998; Moore and Haber, 1996), but not human cells (Harfst et al., 2000) and is thought to physically juxtapose DSB ends through its tethering function (Hopfner et al., 2002). Endonuclease induced DSBs retain 5’ phosphates and 3’ hydroxyl groups and are therefore suitable for direct rejoining, however, additional processing of DNA is required in situations where these modifications are not present. Although several factors have been implicated in this process, it remains poorly understood, particularly in budding yeast. Additional factors linked to NHEJ include the DNA polymerase Pol4 which performs gap filling (Daley et al., 2005), the Rad27 flap endonuclease (Wu et al., 1999), and Tdp1 which acts specifically at ends with 5’ overhangs and promotes high-fidelity NHEJ (Bahmed et al., 2010).

NHEJ joins DNA ends in a sequence-independent manner thus making it a highly versatile repair pathway. Although the tethering of DSB ends promotes their re-ligation (Clerici et al., 2005; Kaye et al., 2004; Lee et al., 2008), NHEJ does not discriminate between ends and has the potential to generate chromosomal translocations and telomere fusions that can drive genomic instability in human cancers (Bunting and Nussenzweig, 2013). NHEJ is often referred to as an error-prone pathway of DSB repair, and is now associated with gene disrupting indels during CRISPR-Cas9 genome editing; however, this is not a completely fair assessment of its fidelity as systems involving site-specific nucleases will undergo cycles of cutting and repair until the cleavage site is mutated. NHEJ can therefore repair clean endonuclease-induced breaks precisely and it is proposed that the condition and processing of DSBs ends is a large factor in repair fidelity and can shift repair to alternative end-joining pathways (Betermier et al., 2014). Although NHEJ plays an important role in DSB repair in humans, it does less so in budding yeast which primarily relies on HR (Boulton and Jackson, 1996a).

Repair by HR was first identified in budding yeast following the transformation of plasmids which were observed to undergo integration into the genome at homologous sites (Orr-Weaver and Szostak, 1983; Orr-Weaver et al., 1981; Szostak et al., 1983). As opposed to NHEJ, HR utilizes intact homologous sequence as a template for repair and this is most frequently provided by the sister chromatid (Johnson and Jasin, 2000). As expected, HR is therefore limited to the S and G2 phases of the cell cycle when the sister chromatid is present following DNA replication (Ira et al., 2004).

Figure 1.1. DSB repair pathways

Figure 1.1. DSB repair pathways. Two primary pathways repair DSBs including nonhomologous end-joining (NHEJ) and homologous recombination (HR). NHEJ requires Ku (green rings) and Dnl4 (blue hexagon) for end-joining. HR involves initial processing by MRN (green circles), DNA resection to reveal ssDNA which is coated by RPA (blue circles), loading of the Rad51 recombinase (red circles), and strand invasion and subsequent DNA synthesis. A third alternative pathway, called microhomology-mediated end-joining (MMEJ) lies between HR and NHEJ.

The key determinant in the choice between NHEJ and HR lies in the initiation of DNA end resection. Unprocessed DNA ends can be channeled into either pathway and attempt repair by NHEJ (Frank-Vaillant and Marcand, 2002), but the generation of ssDNA from 5’ to 3’ DNA resection commits an end to repair by HR. Resection is blocked by the Ku complex (Clerici et al., 2008; Lee et al., 1998; Maringele and Lydall, 2002), and Rad9 (Lazzaro et al., 2008), analogous to the role of 53BP1 which prevents end resection in mammalian cells (Bunting et al., 2010). These blocks to end resection are critical during G1 phase when repair by NHEJ is preferred and entry into S phase shifts the balance towards end resection and HR (Chapman et al., 2012). This cell cycle regulation is largely controlled by the phosphorylation of Sae2 by Cdk1 (Huertas et al., 2008), CtIP in humans (Huertas and Jackson, 2009), which promotes DNA end resection and

9 elegantly ties this process to cell cycle progression. Evidence in human cells suggests that additional factors downstream of end resection are also cell cycle regulated to control HR including the BRCA1-PALB2 interaction (Orthwein et al., 2015).

The phosphorylation of Sae2 stimulates Mre11 activity and triggers short-range end resection towards the DSB, which is dependent on the exonuclease activity of Mre11 (Cannavo and Cejka, 2014). This end processing facilitates long-range 5’ to 3’ DNA end resection by the Exo1 exonuclease, and the Dna2 nuclease in concert with the BLM helicase homolog Sgs1 (Zhu et al., 2008). Genetic evidence reveals that the MRX complex is not required for resection in the absence of Ku suggesting these factors antagonize each other at the DSB end to promote and inhibit end resection respectively (Mimitou and Symington, 2010). DNA end resection occurs at 4 kb per hour (Zhu et al., 2008) and exposed ssDNA is rapidly bound and protected by the heterotrimeric replication protein A complex (RPA). RPA is encoded by the Rfa1, Rfa2, and Rfa3 subunits in budding yeast and RPA binds preferentially to ssDNA in a sequence-independent manner and is critical for DNA replication (Brill and Stillman, 1989; Wold et al., 1989). RPA contains a combined four oligonucleotide/oligosaccharide-binding (OB) folds which bind to ssDNA in 8 and 30 nucleotide modes depending on the number of engaged OB-folds (Bochkareva et al., 2002). RPA was identified in yeast to promote DNA strand exchange (Heyer et al., 1990) and is essential for DNA recombination (Longhese et al., 1994). One proposed function of RPA in this process is to remove secondary structures from ssDNA and to limit self-annealing (Wold, 1997). Finally, RPA also recruits Ddc2-Mec1 and is important for the initiation of DNA damage signalling (Zou and Elledge, 2003).

Repair by HR requires a mechanism to anneal DNA strands and assess their sequence homology and a fundamental question is how this process occurs. The identification of homologous donor sequence is carried out by the Rad51 recombinase, a homolog of the bacterial strand exchange protein RecA (Shinohara et al., 1992). Rad51 replaces RPA on ssDNA to form a nucleoprotein filament, called the presynaptic complex, and the exchange of RPA for Rad51 on resected DNA requires Rad52 (New et al., 1998; Shinohara and Ogawa, 1998; Sung, 1997a) and is promoted by Rad55 and Rad57 (Johnson and Symington, 1995; Sung, 1997b). The Rad51 nucleoprotein filament is critical for strand invasion into potential donor sequences, no small feat given the stability of dsDNA. This invasion step allows pairing of the presynaptic filament to a complementary DNA strand and generates a displaced strand (or D-loop). D-loop formation is

10 supported by additional mediators including Rad54 (Petukhova et al., 1998). The structure of RecA bound to both ssDNA and dsDNA reveals that contacts between the complementary strand and the presynaptic filament occur through DNA base-pairing alone, establishing a mechanism to determine homology (Chen et al., 2008). This structure also revealed non-uniform stretching of Rad51-bound DNA to establish distinct three-nucleotide interaction sites (Chen et al., 2008); an observation confirmed by recent single-molecule analysis of Rad51 binding (Lee et al., 2015).

Once complementary DNA sequence is identified, DNA synthesis can extend 3’ ends to copy genetic material lost at the original DSB site. DNA Polζ was initially implicated in this synthesis (Holbeck and Strathern, 1997), but it is now thought that DNA Polδ (Maloisel et al., 2008) and Polε (Hicks et al., 2010) are the primary enzymes which synthesize new DNA during repair. Several outcomes can result during repair by HR (Figure 1.2) including synthesis-dependent strand annealing in which the invading strand is simply displaced by Srs2 following DNA synthesis (Ira et al., 2003). Alternatively, capture of the second DNA end followed by gap filling and ligation can result in the classic double Holliday junction (Holliday, 1964). These structures can be disassembled through several means to generate either cross-over or non-crossover products. The preferred outcome of double Holliday junctions is dissolution, a process involving convergent branch migration by Sgs1-Top3 (Ira et al., 2003) and decatenation of DNA by Top3, a process stimulated by Rmi1 (Cejka et al., 2010b; Cejka et al., 2012). Dissolution is therefore a key pathway to suppress sister chromatid exchange and mutation of BLM, the Sgs1 ortholog in humans, is characterized by greatly elevated rates of sister chromatid exchange. A second mechanism for solving Holliday junctions is called resolution and involves DNA strand cleavage (Parsons and West, 1988), typically by the Mus81-Mms4 nuclease (Boddy et al., 2001), or by the Yen1 nuclease as a last resort before mitosis (Blanco et al., 2014). Resolution can result in either crossover or non-crossover products depending on which strands are nicked. Finally, if a DSB only contains homology on one end, break-induced replication can occur which involves strand invasion and DNA replication to the end of the chromosome resulting in loss of heterozygosity (Lydeard et al., 2007; Smith et al., 2007).

Rad52 plays an essential role in all recombination-based processes and can be visualized at sites of DNA damage when fused to fluorescent protein markers (Lisby et al., 2003; Lisby et al., 2001). Rad52 can also promote Rad51-independent recombination events (Malkova et al., 1996; Rattray and Symington, 1994), most notably during intrachromosomal exchanges (Bartsch et al., 2000).

The annealing of direct repeats exposed during end resection can also stimulate repair by single- strand annealing (SSA) which requires Rad52, but not Rad51 (Ivanov et al., 1996).

Figure 1.2. Repair of DSBs by homologous recombination

Figure 1.2. Repair of DSBs by homologous recombination. Synthesis-dependent strand annealing (SDSA), the classical double-strand break repair model (DSBR) featuring a double Holliday junction (dHJ), and break-induced replication (BIR) in which only one end has homology. Double Holliday junctions can undergo dissolution (D) or resolution (R), the latter which can result in crossover events.

Although DSB repair is often presented as a dichotomy between NHEJ and HR, a third pathway is gaining appreciation which shares features of both. Termed microhomology-mediated end joining (MMEJ), this pathway facilitates the joining of DNA ends in the absence of Ku and exhibits small regions of homology at the junction site (Figure 1.1). This pathway was first described in Ku-deficient budding yeast as an error-prone repair pathway which requires at least 6 bp of annealed homology (Boulton and Jackson, 1996b; Ma et al., 2003; Yu and Gabriel, 2003) and fill- in DNA synthesis by DNA Polβ and Polδ (Lee and Lee, 2007). MMEJ also occurs in mammalian cells with as little as 1 bp of homology at junction sites (Simsek and Jasin, 2010) and was first identified by the surprising persistence of telomere fusions in the absence of DNA ligase IV (Maser et al., 2007) and class switch recombination in C-NHEJ deficient cells (Yan et al., 2007). MMEJ

12 requires Mre11 and resection initiation (Truong et al., 2013), but is inhibited by RPA in yeast (Deng et al., 2014), suggesting that the end processing requirement for MMEJ lies somewhere between NHEJ and HR. The translesion synthesis polymerase Polθ has been implicated in MMEJ in mammalian cells (Ceccaldi et al., 2015; Mateos-Gomez et al., 2015) and the ability of MMEJ to generate deletions and chromosome translocations suggests it may be an underappreciated contributor to genomic instability in cancer cells (Sfeir and Symington, 2015).

1.2.3 DSBs arrest cell division

Besides promoting DSB repair, a second critical function of DSB signalling is to block cell division and allow time for repair. Seminal work in budding yeast first identified that cell cycle arrest after DNA damage requires RAD9, and although rad9Δ mutants bypass G2/M arrest and continue to divide, this has disastrous consequences for their long-term survival (Weinert and Hartwell, 1988). Remarkably, the arrest of rad9Δ mutants by a microtubule poison restores viability (Weinert and Hartwell, 1988), indicating that Rad9 is not required for DSB repair per se but promotes a cellular checkpoint that monitors for damage. A single HO-induced DSB in budding yeast causes a 12 to 14 h G2/M arrest (Lee et al., 1998) and two parallel pathways support this block to cell division: one coordinated through the Rad53 and Dun1 kinases, and the second through the Chk1 kinase (Gardner et al., 1999; Sanchez et al., 1999). A critical event in mitosis is the cleavage of sister cohesion by the separase protease, Esp1 in budding yeast (Uhlmann et al., 1999). Esp1 is inactivated by securin, Pds1, thereby ensuring that sister chromatids remain together in metaphase until anaphase when Pds1 is degraded (Ciosk et al., 1998). Pds1 is ubiquitylated and targeted for degradation by the anaphase promoting complex/cyclosome (APC/C) in concert with the activator Cdc20 (Visintin et al., 1997) and DNA damage intervenes in this pathway through the Chk1-mediated phosphorylation of Pds1 which blocks ubiquitylation and stabilizes both Pds1 and cohesion (Cohen-Fix and Koshland, 1997; Yamamoto et al., 1996).

Although the phosphorylation of Pds1 can account for the role of Chk1 in G2/M arrest (Wang et al., 2001), how Rad53 and Dun1 promote cell cycle arrest following DNA damage is less understood. One Rad53-dependent target is Cdc20 and phosphorylation blocks the interaction with Pds1 (Agarwal et al., 2003), however, the observation that rad53Δ mutants exhibit a greater checkpoint defect than pds1Δ mutants argues for the existence of yet-to-be identified substrates

(Gardner et al., 1999; Sanchez et al., 1999). One such Rad53 substrate appears to be Bfa1 which may inhibit mitotic exit (Hu et al., 2001).

It should be noted that additional checkpoint mechanisms exist in budding yeast following DNA damage including a Rad9-dependent delay to the G1/S transition (Siede et al., 1993), the DNA replication checkpoint (Santocanale and Diffley, 1998), and an intra-S checkpoint which slows S phase progression (Paulovich et al., 1997). The mechanism of cell cycle arrest in budding yeast also differs slightly from mammalian cells in which both CHK1 and CHK2 kinases phosphorylate the APC/C activator CDC25 after DNA damage leading to its destruction and a block to cell cycle progression (Falck et al., 2001; Furnari et al., 1997; Matsuoka et al., 1998; Peng et al., 1997; Sanchez et al., 1997). Unlike budding yeast, mammalian cells also possess a prominent G1 arrest in response to DNA damage which is dependent on CHK1 and p53 (Chehab et al., 2000).

Upon completion of DSB repair cells must turn off the DNA damage checkpoint, a process termed recovery. Key components include the Srs2 helicase which removes Rad51 (Vaze et al., 2002; Yeung and Durocher, 2011), and the PP4C phosphatase responsible for dephosphorylating γ-H2A (Keogh et al., 2006a; Nakada et al., 2008). Rad53 is also deactivated through dephosphorylation by the PP2C and Pph3-Psy2 phosphatase complexes (Leroy et al., 2003; O'Neill et al., 2007). Cells can also resume cycling in the presence of DNA damage, termed adaptation, and the polo-like kinase Cdc5 (Toczyski et al., 1997), the Ku complex (Lee et al., 1998), and the Rad54 homolog Rdh54 (Lee et al., 2001) have been implicated in this process. While adaptation allows cells to escape immediate arrest, it predictably can lead to additional genomic instability (Galgoczy and Toczyski, 2001).

1.3 DNA ends in the cell

1.3.1 The structure and function of telomeres

The sensitive and robust response to DSBs poses an interesting problem to the cell as each chromosome also contains two linear ends. This problem was first identified by Herman Müller and Barbara McClintock over 75 years ago who recognized that the natural ends of chromosomes, called telomeres, are different from those created by a chromosome break and are somehow hidden

14 from the cell (McClintock, 1941; Muller, 1938). Strikingly, the loss of a single telomere in budding yeast activates the DNA damage checkpoint and arrests cell growth (Sandell and Zakian, 1993). But what delineates a telomere from an internal break? The simple answer lies in the unique repetitive DNA sequence at the ends of all eukaryotic chromosomes, first identified as repetitive TTGGGG sequence in Tetrahymena (Blackburn and Gall, 1978). In budding yeast, telomeres consist of 300 ± 75 bp of heterogeneous TG1-3 repeats (Zakian, 1996), in contrast to the perfect TTAGGG repeats found in humans (Moyzis et al., 1988).

Budding yeast telomeres are bound by two protein complexes which specifically recognize telomeric sequence. First, Cdc13 binds to ssDNA at the distal end of telomeres and is critical for the capping of DNA ends. This discovery was made using temperature sensitive cdc13-1 cells which undergo telomere degradation (specifically of the C-rich strand) and a Rad9-depenendent checkpoint arrest at non-permissive temperature (Garvik et al., 1995). This result highlights two essential properties of telomeres and of Cdc13 in particular, namely to prevent degradation of DNA ends, and to block the DNA damage checkpoint. Cdc13 performs this role in complex with Stn1 and Ten1 which together form the heterotrimeric CST complex (Gao et al., 2007; Grandin et al., 2001; Pennock et al., 2001; Petreaca et al., 2007). The CST complex is proposed to be a telomeric specific version of RPA and also makes extensive use of OB-folds for DNA binding (Gao et al., 2007). Cdc13 binds to a minimum 11 bp sequence of single-stranded telomeric DNA through its canonical OB-fold containing DNA binding domain (Hughes et al., 2000; Lin and Zakian, 1996), and an N-terminal OB-fold has also been shown to promote protein dimerization and bind DNA of 37 bp and longer (Mitchell et al., 2010; Sun et al., 2011).

The second telomeric binding protein in budding yeast is Rap1 which binds to double-stranded DNA. Rap1 is estimated to bind every 18 bp of telomere sequence (Gilson et al., 1993; Ray and Runge, 1999) and uses a double Myb domain to interact with DNA (Graham et al., 1999). Rap1 has an important role in preventing NHEJ and telomere fusions (Pardo and Marcand, 2005), but also contributes to end capping and prevents DNA degradation (Anbalagan et al., 2011; Bonetti et al., 2010). Rap1 can inhibit NHEJ directly (Marcand et al., 2008) and also coordinates the recruitment of additional partners through its C-terminal domain including the Rap1 interacting factors Rif1 and Rif2, and the silent information regulator proteins Sir3 and Sir4 (Palladino et al., 1993; Wotton and Shore, 1997). Two of these downstream factors, Rif2 and Sir4, also inhibit NHEJ (Marcand et al., 2008).

Rap1 in combination with Rif1 and Rif2 negatively regulates telomere length and forms an elegant counting mechanism. As telomeres lengthen the number of Rap1 binding sites and bound Rif1 and Rif2 increases which inhibits further telomere extension, (Levy and Blackburn, 2004; Marcand et al., 1997). Conversely, short telomeres are marked by low levels of Rif2 which allows for the preferential binding of Tel1 (McGee et al., 2010). Although previously introduced as a core DNA damage kinase, Tel1 was first identified as a mutant with short telomeres and is critical for telomere elongation (Lustig and Petes, 1986). Tel1 is therefore negatively regulated by the telomere counting mechanism and one identified mechanism is that Tel1 competes with Rif2 for binding to Xrs2 (Hirano et al., 2009).

Telomeres locally repress gene transcription, a phenomenon called the telomere position effect, and mediated by Rap1 and its interaction with Sir3 and Sir4 (Gottschling et al., 1990; Moretti et al., 1994; Rine and Herskowitz, 1987). Transcriptional silencing requires the histone deacetylase Sir2, the only enzymatic component of the Sir proteins, and Sir2 is recruited through an interaction with Sir4 (Moazed et al., 1997). Deacetylation of histone tails by Sir2 promotes the binding of additional Sir3 and Sir4 molecules and allows transcription silencing to locally spread (Rusche et al., 2003).

Although budding yeast has been a key model in understanding fundamental telomere biology, the specific mechanism of end-protection differs in mammalian cells which utilize the shelterin complex to inhibit DSB signalling and repair through a variety of mechanisms. Although shelterin also contains a Rap1 subunit, the function of budding yeast Rap1 is more comparable to the TRF1 and TRF2 subunits of shelterin which bind to double-stranded TTAGGG repeats and inhibit ATM signalling and C-NHEJ (Karlseder et al., 1999). Analogous to Cdc13, the POT1 subunit binds single-stranded telomeric DNA to inhibit ATR signalling (Denchi and de Lange, 2007), and additional protection against alt-NHEJ and DNA resection is provided by the Ku complex and 53BP1 respectively (Sfeir and de Lange, 2012). A CST complex comprised of Ctc1-Stn1-Ten1 is also found in mammals and is linked to end protection, synthesis of the C-rich strand, and the negative regulation of telomerase (Chen et al., 2012; Miyake et al., 2009; Wang et al., 2012).

1.3.2 Telomerase activity and regulation

The second problem addressed by telomeres, termed the end-replication problem, arises due to the nature of lagging strand replication in eukaryotic cells and the progressive shortening of chromosomes due to the removal of the distal RNA primer. Telomeric DNA is maintained and elongated by telomerase, an enzyme first identified in Tetrahymena extracts which can add new repeats to chromosome ends (Greider and Blackburn, 1985). Additional characterization of telomerase identified that the minimal components include a reverse transcriptase subunit and an RNA component that serves as a primer for extension (Greider and Blackburn, 1987, 1989). In agreement with this idea, telomerase in budding yeast is minimally composed of the Est2 (ever shorter telomeres) reverse transcriptase subunit and TLC1 RNA, although additional subunits including Est1 and Est3 are required for in vivo activity (Lendvay et al., 1996; Lingner et al., 1997; Lundblad and Szostak, 1989). The TLC1 RNA component acts as a critical scaffold and coordinates the binding of the other subunits and regulatory factors (Dandjinou et al., 2004; Zappulla and Cech, 2004), for example Est1 which binds to a bulge stem RNA structure (Seto et al., 2002), and the Pop1, Pop6, and Pop7 proteins which stabilize the holoenzyme (Lemieux et al., 2016). Cells defective for telomerase activity are capable of dividing for 50 to 100 generations until their telomeres reach a critical length and they undergo senescence.

Cdc13 is also required for telomerase activity in vivo and was initially identified as Est4 due to the inability of Cdc13 mutants to lengthen telomeres (Lendvay et al., 1996). Cdc13 recruits telomerase to DNA ends through an interaction with Est1 (Evans and Lundblad, 1999, 2002; Wu and Zakian, 2011), and this is most conclusively revealed by the cdc13-2 and est1-60 charge swap alleles which are individually telomerase deficient but can be combined to restore telomerase activity (Bianchi et al., 2004). Est1 is dispensable in cells expressing a fusion of Cdc13 and Est2 arguing that telomerase recruitment is the essential function of Est1 (Evans and Lundblad, 1999); however, Est1 also has a role in activating telomerase (Evans and Lundblad, 2002; Taggart et al., 2002) and is required for Est3 binding (Sabourin et al., 2007). Est1 interacts with a region of Cdc13 called the recruitment domain and residues S249 and S255 within this region are important for telomerase activity (Tseng et al., 2006). These residues are proposed to be phosphorylated by Mec1 and Tel1 to promote telomerase recruitment (Tseng et al., 2006), but this model is challenged (Gao et al., 2010). In addition to an interaction with Est1, Cdc13 also binds the DNA Polα complex and coordinates extension of the C-rich strand following telomerase extension (Lue et al., 2014; Qi and Zakian, 2000). In contrast to the positive regulation of telomerase, Cdc13 also negatively

17 regulates telomerase which requires the interaction between Cdc13 and Stn1 (Chandra et al., 2001; Puglisi et al., 2008). The involvement of Stn1 in the negative regulation of telomerase is supported by the observation that the overelongation of telomeres in cdc13-5 mutants can be suppressed by Stn1 overexpression (Chandra et al., 2001).

Telomerase purified from synchronized cellular extracts is active throughout the cell cycle; however, telomere elongation in vivo is observed in late S and G2 phase, but not in G1 (Diede and Gottschling, 1999; Marcand et al., 2000). This observation hints at a regulatory model whereby telomere accessibility or the recruitment of telomerase is cell cycle controlled, but not the catalytic activity of telomerase itself. Although Cdc13 and Est2 can be found at telomeres throughout the cell cycle, the binding of Est1 and Est3 is strictly limited to late S and G2, coinciding with telomerase activity (Taggart et al., 2002; Tuzon et al., 2011). The abundance of Est1 also peaks during this time (Osterhage et al., 2006) together supporting a model in which the assembly of the telomerase holoenzyme is tightly controlled to regulate telomerase activity. A second mechanism for the cell cycle regulation of telomerase is through the phosphorylation of Cdc13-T308 by Cdk1 which is proposed to favour the positive regulation of telomerase (Li et al., 2009). Phosphorylation at this site promotes the Cdc13-Est1 interaction at the expense of the interaction with Stn1 and the shortened telomere phenotype observed in cdc13-T308A mutants can be rescued by the stn1- ΔC199 allele which fails to interact with Cdc13 (Li et al., 2009).

Finally, telomerase is not the only mechanism to extend telomeres and a second pathway, called the alternative-lengthening of telomeres (ALT), relies on DNA recombination to add telomere sequence. This process was first discovered in telomerase null est1Δ yeast cells from the rare appearance of spontaneously arising colonies, or survivors (Lundblad and Blackburn, 1993). Survivors can maintain telomeres through the expansion of subtelomeric repeats (Type I), or distal

TG1-3 repeats (Type II) and the genetic requirements for both types are slightly varied although they both largely depend on Rad52 and Pol32 (Chen et al., 2001b; Lebel et al., 2009; Lydeard et al., 2007; Teng and Zakian, 1999). Similarly, human somatic cells do not express telomerase and eventually arrest after reaching a critical limit of telomere shortening. Cancer cells must therefore overcome this block and while many find a way to express telomerase, others use ALT pathways which resemble the Type II survivors of budding yeast (Shay and Bacchetti, 1997; Shay and Wright, 2011).

1.3.3 Interplay between DSBs and telomeres

The fate of DSBs and telomeres requires opposite outcomes and it is therefore surprising that key proteins are involved in the regulation of both ends. Understanding how cells differentiate their ends is therefore a fundamental question and has important consequences for genomic stability. As previously noted, the DNA damage kinase Tel1 plays a pivotal role in telomere maintenance, but mec1Δ mutants also display shortened telomeres and tel1Δ mec1Δ double mutants are completely unable to elongate telomeres and undergo senescence (Ritchie et al., 1999). The Ku complex, involved in NHEJ at DSBs, is also found at telomeres and the Yku80 subunit interacts with TLC1 RNA to promote telomere elongation (Stellwagen et al., 2003). As opposed to the outward facing Yku70 subunit, Yku80 is faces towards the centromere to mediate the telomeric functions of Ku (Ribes-Zamora et al., 2007).

One of the ways to stabilize a broken chromosome is to simply add a new telomere and this process is called de novo telomere addition or telomere healing. This pathway plays prominently in the generation of nanochromosomes in the ciliates Paramecium and Tetrahymena (Baroin et al., 1987; Forney and Blackburn, 1988; Spangler et al., 1988), but also occurs at spontaneous DSBs in budding yeast, Drosophila, and in human cancer cells (Biessmann et al., 1990; Chen et al., 1998; Fouladi et al., 2000; Kramer and Haber, 1993; Sabatier et al., 2005). Telomere addition has relevance to human disorders and new telomeres have been detected at terminal deletions on chromosomes 16 and 22 (Wilkie et al., 1990; Wong et al., 1997). The sequence added during telomere addition in budding yeast is remarkably similar to native telomeres and arises from multiple cycles of TLC1 annealing and telomerase-mediated extension of the 3’ DNA end (Pennaneach et al., 2006). Consistent with this idea, analysis of telomere addition events indicates that they preferentially occur at sites containing TG1-3 sequence with homology to TLC1 (Kramer and Haber, 1993; Mangahas et al., 2001; Putnam et al., 2004). The commitment to telomere addition appears to be a one-time decision by the cell and the analysis of hundreds of DSBs repaired by NHEJ or HR has failed to identify any intervening telomeric repeats (Putnam et al., 2004, 2005). This has lead to the proposal that DSBs which undergo telomere addition are sequestered into nuclear regions that prevent conventional repair (Pennaneach et al., 2006).

1.3.4 The telomerase inhibitor Pif1

Although telomere addition can stabilize the broken DNA end, it has profound consequences for the cell as the acentric chromosome fragment will be prone to loss during cell division. The activity of telomerase must therefore be tightly regulated at DSBs and this is accomplished in budding yeast by the telomerase inhibitor Pif1 (Schulz and Zakian, 1994; Zhou et al., 2000). Pif1 was first discovered in a screen for factors involved in mitochondrial DNA maintenance (Foury and Kolodynski, 1983) and was later identified to exhibit helicase activity (Lahaye et al., 1991). Pif1 was rediscovered in a genetic screen investigating the stability of subtelomeric genes which identified that mutation of Pif1 results in telomere addition at internal sites (Schulz and Zakian, 1994). Pif1 has both mitochondrial and nuclear isoforms encoded from separate translational start sites and the mutation of pif1-m2 abolishes the nuclear isoform while leaving mitochondrial Pif1 intact. The loss of nuclear Pif1 in pif1-m2 cells results in longer telomeres and a 240-fold increase in telomere addition at DSBs which is dependent on telomerase (Bochman et al., 2010; Myung et al., 2001; Schulz and Zakian, 1994; Zhou et al., 2000). The telomere defects in pif1-m2 mutants are not as severe as is pif1Δ cells suggesting that some protein still manages to get into the nucleus (Schulz and Zakian, 1994).

Pif1 is a superfamily 1 (SF1) helicase with 5’ to 3’ activity and a conserved Walker A box motif (Bochman et al., 2010). Budding yeast also contain a related helicase, Rrm3, implicated in the replication of ribosomal DNA and replication fork progression (Ivessa et al., 2000; Keil and McWilliams, 1993), and both Pif1 and Rrm3 have some similarity to the bacterial helicase RecD (Zhang et al., 2006). In addition to the inhibition of telomerase, Pif1 also promotes bubble migration during break-induced replication (Saini et al., 2013; Wilson et al., 2013) and unwinds G-quadruplex structures (Paeschke et al., 2013; Ribeyre et al., 2009; Sanders, 2010).

The helicase activity of Pif1 preferentially unwinds RNA-DNA hybrids in vitro (Boule et al., 2005) and inhibits telomerase by removing the TLC1 RNA template from telomeres (Li et al., 2014; Phillips et al., 2015). Consistent with this model, Pif1 is detected at telomeres by chromatin immunoprecipitation (ChIP) experiments (Zhou et al., 2000). The direct inhibition of telomerase by Pif1 is also supported by the identification of a telomerase mutant, est2-up34, which is insensitive to Pif1 inhibition (Eugster et al., 2006). Despite these observations, it is unknown if Pif1 performs the same function at DSBs to prevent telomere addition. Such a mechanism predicts that the absence of Pif1 should result in the preferential addition of telomeres at TLC1 binding sites however this has not been observed in the analysis of hundreds of telomere addition events

(Putnam et al., 2004). Intriguingly, Pif1 appears to distinguish between DSBs and telomeres as the pif1-4A mutation is unable to inhibit telomere addition, but does not cause a change in telomere length (Makovets and Blackburn, 2009). Although Pif1 normally inhibits telomere addition, DSBs flanked with longer telomeric sequence are elongated in the presence of Pif1 (Zhang and Durocher, 2010). Pif1 is therefore an important factor in the fate of DNA ends and understanding its regulation in this process will shed light on how cells differentiate DSBs and telomeres.

It should be noted that Pif1 is not the only mechanism to limit telomere addition at DSBs and the phosphorylation of Cdc13-S306 by Mec1 also limits telomerase activity (Zhang and Durocher, 2010). Phosphorylation of this residue inhibits Cdc13 accumulation at DSB ends, particularly those with less than 11 bp of telomeric sequence, and is opposed by the Pph3 phosphatase (Zhang and Durocher, 2010).

1.4 DSB repair in the context of the nucleus

1.4.1 Spatial organization of the nucleus

The signalling and repair of DSBs occurs in the context of chromatin and in the spatial confines of the nucleus. DNA is highly organized and as a first step is wrapped around core histone proteins to form nucleosomes and the chromatin fibre. Additional layers of compaction order the chromatin fibre into chromosomes which themselves are not randomly positioned in the nucleus. The notion that chromosomes adopt specific conformations was first made by Carl Rabl from the observation of salamander chromosomes which cluster at one nuclear pole with extended chromosomes arms. Rabl proposed that this conformation would be held throughout interphase and thus form distinct nuclear domains (Rabl, 1885).

The prediction of nuclear domains is supported by modern labelling and imaging techniques including fluorescent in-situ hybridization (FISH) across a variety of species (Marshall et al., 1996; Nagele et al., 1995). Additional methods such as chromosome conformation capture also reveal spatial organization of the genome (Dekker et al., 2002), although sometimes exhibiting discordant results with FISH data (Williamson et al., 2014). Nevertheless, in budding yeast, both approaches provide similar pictures of a highly organized nucleus and which mimics the chromosomes observed by Rabl in salamanders (Berger et al., 2008; Duan et al., 2010). This arrangement of

21 chromosomes, called the Rabl conformation, is characterized by the clustering of centromeres at the centrosome, called the spindle pole body (SPB) in yeast, with chromosome arms radiating towards the nuclear periphery (Bystricky et al., 2004; Jin et al., 2000).

Telomeres and heterochromatin domains are found at the nuclear envelope in numerous organisms, a location which is thought to promote their transcriptional silencing (Towbin et al., 2009). Budding yeast is no exception and their telomeres are found clustered into four to eight foci at the nuclear periphery (Gotta et al., 1996). The perinuclear tethering of telomeres is mediated through two pathways, one involving Sir4, and the other which requires Ku (Hediger et al., 2002; Taddei et al., 2004). As previously mentioned, Sir4 is recruited to telomeres by Rap1 and can bind to the nuclear periphery through the anchoring proteins Esc1 or Mps3, the latter being specific to S phase (Andrulis et al., 2002; Bupp et al., 2007; Luo et al., 2002). The second tethering pathway involving the Ku complex can interact with Sir4, but also facilitates tethering via binding of Yku80 to TLC1 in S phase (Schober et al., 2009; Taddei et al., 2004). The Yku70 subunit also appears involved in tethering in specifically during G1 phase (Taddei et al., 2004). Although perinuclear tethering was initially thought to be required for transcriptional silencing, these two events can be separated (Taddei et al., 2004; Tham et al., 2001). Highly expressed inducible genes can also be relocated to the nuclear periphery (Taddei et al., 2006), highlighting a complex relationship between the nuclear envelope and transcription.

Budding yeast undergo a closed mitosis and the SPB coordinates microtubule organization and function from its position embedded in the nuclear envelope. Centromeres are tethered by a single microtubule linking each kinetochore to the SPB and this attachment persists throughout the cell cycle, except for a brief period in S phase (Kitamura et al., 2007). In contrast to the epigenetically determined centromeres of most eukaryotes, budding yeast has “point” centromeres consisting of approximately 125 bp of DNA which includes three centromere-determining elements (CDE) (Bloom and Carbon, 1982; Fitzgerald-Hayes et al., 1982). The addition of centromeric sequence to an episome is therefore sufficient for microtubule attachment and the ability to undergo mitotic segregation like ordinary chromosomes (Clarke and Carbon, 1980). Kinetochores are composed of distinct protein subcomplexes that assemble upon centromeric DNA although the order and requirements for assembly is not fully understood (Biggins, 2013). Direct centromeric DNA binding proteins include Cbf1, which binds to CDEI sequence (Cai and Davis, 1990; Cai and Davis, 1989), and the CBF3 complex, comprised of Ndc10, Cep3, Ctf13, and Skp1, which binds

22 to CDEIII sequence (Connelly and Hieter, 1996; Goh and Kilmartin, 1993; Lechner and Carbon, 1991). An additional feature of centromeres is the protein Cse4, a histone H3 variant. Cse4 is incorporated into a single unique nucleosome wrapped by CDEII DNA and is required for proper chromosome segregation (Furuyama and Biggins, 2007; Stoler et al., 1995).

Over fifty individual proteins localize to the kinetochore and notable components include Mif2 (Meluh and Koshland, 1995), the Mtw1/MIND complex (Goshima and Yanagida, 2000), the Ndc80 complex (Janke et al., 2001; Wigge and Kilmartin, 2001), the Ctf19/COMA (Ortiz et al., 1999), and the Dam1 ring complex which directly binds microtubules (Cheeseman et al., 2001; Jones et al., 1999). Kinetochores are required for the faithful segregation of chromosomes and sister kinetochores must attach to microtubules from opposing poles before cell division. This process is called biorientation and requires the Mps1 kinase (Maure et al., 2007; Shimogawa et al., 2006). Conversely, improperly attached kinetochores that lack tension are released from microtubules by the Aurora B kinase, thus providing another attempt at biorientation (Biggins et al., 1999; Tanaka et al., 2002). Similar to the DNA damage checkpoint, kinetochore-microtubule attachment is monitored by the spindle assembly checkpoint (SAC), a surveillance mechanism that blocks cell division if problems are detected (Musacchio and Salmon, 2007). This response requires several checkpoint proteins including Mps1, Bub1, Bub3, and Mad1 (Hoyt et al., 1991; Li and Murray, 1991) leading to the inactivation of the APC/C activator Cdc20 though binding by the SAC proteins Mad2 and Mad3 (Chao et al., 2012; Lau and Murray, 2012).

1.4.2 Nuclear position and genomic integrity

In addition to gene expression, the nuclear periphery has important links to genomic stability and is thought to shield certain loci from inappropriate repair events (Mekhail and Moazed, 2010). A striking example is the segregation of repetitive ribosomal DNA (rDNA) to the nucleolus (Duan et al., 2010), a region that excludes Rad52 and which is thought to be a protective mechanism against recombination and the amplification of rDNA (Park et al., 1999; Torres-Rosell et al., 2007). The association of rDNA with the nuclear envelope is critical for its stability and is mediated through the Cohibin complex (Mekhail et al., 2008). Remarkably, breaks formed in rDNA require relocalization to the nuclear interior for repair (Torres-Rosell et al., 2007), consistent with the observation that recombination at the MAT locus also occurs in the nuclear interior (Bystricky et al., 2009). Multiple DSBs have also been observed to cluster into single foci called repair centres

23 within the nuclear interior suggesting that this location is recombination proficient (Lisby et al., 2003; Lisby et al., 2001).

In contrast to DSB repair in the nuclear interior, several studies have revealed that DSBs without homologous donor sequence or slowly repaired DSBs relocate to the nuclear periphery (Kalocsay et al., 2009; Oza et al., 2009). The nuclear envelope protein Mps3 has a critical role in this relocalization and promotes telomere addition at broken DNA ends (Oza et al., 2009; Schober et al., 2009). Mps3 is not the only component at the nuclear periphery linked to genomic stability and the Nup84 nuclear pore complex is also important for repair of subtelomeric DSBs and the relocalization of collapsed replication forks to the periphery (Nagai et al., 2008; Therizols et al., 2006). Together these results support a model in which difficult to repair DSBs are sequestered to the periphery for repair by atypical pathways although the exact mechanism for DSB relocation and repair pathway choice remains unclear.

The non-random organization of chromatin has consequences for all DNA transactions, but perhaps none more intriguing than the process of HR which requires a physical interaction between the broken chromosome and a donor. The genomic integration of transformed plasmid DNA suggested that a search for homology is an essential step in HR (Orr-Weaver and Szostak, 1983; Orr-Weaver et al., 1981; Szostak et al., 1983) and early in vitro experiments hinted that homologous pairing is a rapid and non-rate limiting step (Gonda and Radding, 1983). The efficient recombination between chromosomes in budding yeast also raises the possibility that the whole genome might be scanned, however, a genome-wide search has been argued to be physically impossible (Weiner et al., 2009). Although strand invasion and the base pairing of complementary DNA strands is mediated by the Rad51 recombinase, how homologous sequences find each within the three-dimensional nucleus is a key unresolved scientific question. Possible scenarios range from directed models using localized sliding of the Rad51 filament to more random probing mechanisms promoted by the ability of the filament to bridge two non-contiguous DNA sequences (Renkawitz et al., 2014). The detection of Rad51 ChIP signals, argued to indicate homology search, reveals a predominantly intrachromosomal search which is decreased at further distances from the DSB, although signals can be detected on other chromosomes (Renkawitz et al., 2013). The spatial proximity between loci is also an important factor in their ability to recombine, consistent with a more localized search model (Agmon et al., 2013).

HR is not the only repair pathway with spatial considerations and similar questions surround the nature of chromosomal translocations joined by NHEJ. The principle contention is whether points of chromosomal exchange are prepositioned before breakage, or whether they make contact after breakage (Aten et al., 2004; Soutoglou et al., 2007).

1.4.3 Chromatin is not static

The non-random organization of chromosomes within the nucleus implies and necessitates the movement of genetic material. Initial observations using fluorescent recovery after photobleaching postulated that chromatin it is relatively immobile in the nucleus with occasional movements (Abney et al., 1997; Cremer et al., 1982); however, the study of chromatin mobility was greatly aided by the development of new tools to track individual loci through the integration of LacO arrays capable of binding the Lac repressor (LacR) fused to green fluorescent protein (GFP) (Robinett et al., 1996). The study of single loci also required new quantitative methods for analysis and borrowed a method from particle physics called mean square displacement (MSD) analysis, calculated as the average squared distance a particle has travelled over increasing time intervals (Berg, 1993). This robust analysis averages large numbers of data points to generate quantifiable mobility parameters and plots which reveal the nature of the tracked movement. While randomly moving particles maintain a linear slope over time (proportional to the diffusion coefficient), loci contained within a physical volume reach a plateau as the MSD becomes independent of time (Figure 1.3ab). This plateau is proportional to the radius of confinement, or the area the particle resides in during the course of the experiment.

Seminal work using tagged loci in budding yeast revealed that chromatin is in fact dynamic and moves within the nucleus in a constrained random walk (Marshall et al., 1997). While chromosomes are expected to be minimally contained within the nucleus, these experiments revealed that individual loci are more locally constrained and explore only a small subregion of the nucleus (Marshall et al., 1997). The fundamental diffusion of chromatin is remarkably well conserved in budding yeast, Drosophila, and in human cells, where it undergoes rapid locally constrained movement within a radius of 0.5-1 µm (Chubb et al., 2002; Heun et al., 2001; Marshall et al., 1997; Vazquez et al., 2001). Longer range movements of chromatin can also be detected but are relatively rare events (Heun et al., 2001).

Figure 1.3. Analysis of chromatin mobility

Figure 1.3. Analysis of chromatin mobility. a, Time-lapse imaging of fluorescent particles allows for MSD analysis b, Hypothetical MSD plots of particles moving randomly versus those who are contained within a confined space, i.e. the nucleus. The diffusion coefficient (D) is proportional to the initial MSD slope and number of physical dimensions (d), while the radius of confinement can be determined from the plateau of the MSD curve.

The mobility of chromatin is dependent on ATP but the reason for this is not fully understood (Weber et al., 2010, 2012). Several ATP-requiring factors have been implicated including DNA polymerases, the transcription machinery, and chromatin remodelers (Dion and Gasser, 2013), and future work will be required to identify additional factors that promote and regulate chromatin mobility. Gene transcription has strong links to chromatin movement and key experiments have revealed that silenced plasmids are constrained at the nuclear periphery while loss of silencing promotes movement within the nucleus (Gartenberg et al., 2004). However, the simple correlation between transcription and mobility breaks down as inducible genes can also become tethered to nuclear pores and less mobile upon activation (Cabal et al., 2006; Taddei et al., 2006).

Several factors are known to constrain mobility including the contents of the crowded nucleus and the physical anchoring of chromosomes to nuclear compartments like the nucleolus or nuclear pores (Chubb et al., 2002; Dion and Gasser, 2013). The nature of the DNA fibre also constrains the mobility of individual loci and this constraint is most evident from the observation that an excised extrachromosomal fragment can move freely within the whole nucleus (Gartenberg et al.,

2004). Work in budding yeast has implicated the tethering of centromeres to the SPB as a key constraint to mobility (Verdaasdonk et al., 2013) while the attachment of telomeres to the nuclear periphery through Ku and Sir4 is also known to limit chromatin motion (Hediger et al., 2002).

1.4.4 Chromatin mobility following a DSB

Remarkably, the presence of a DSB in budding yeast increases the mobility of chromosomes and this response is dependent on the Mec1 kinase (Dion et al., 2012; Mine-Hattab and Rothstein, 2012). Several HR proteins have also been implicated in this phenomenon including Rad51 and Sae2 (Mine-Hattab and Rothstein, 2012), as well as the DNA damage checkpoint protein Rad9 (Dion et al., 2012). Loss of SAE2 delays the activation of DNA damage signalling, and increased chromatin mobility can be observed in sae2Δ mutants at later time points, consistent with the idea that Mec1 signalling drives chromosome motion (Mine-Hattab and Rothstein, 2012). Furthermore, activation of the DNA damage checkpoint through the colocalization of Ddc1 and Ddc2 in the absence of a DSB (Bonilla et al., 2008) is able to modestly increase chromatin movement (Seeber et al., 2013). Although it is clear that activation of the DNA damage checkpoint stimulates increased chromatin mobility, the target of Mec1-dependent signalling is unknown and remains a key unaddressed question in this phenomenon.

Surprisingly, unbroken chromosomes also exhibit increased mobility following a DSB elsewhere in the nucleus, albeit to a lesser extent than the broken chromosome (Mine-Hattab and Rothstein, 2012). This phenomenon is termed the global mobility response and is dependent on Mec1 and Rad53, but not Rad51, and may require increased levels of DNA damage (Seeber et al., 2013). Several other factors have also been implicated in DSB-induced mobility including the INO80 and SWR1 chromatin remodelers, and the histone variant H2A.Z, leading to a model in which chromatin remodeling activity increases flexibility of the DNA fibre (Horigome et al., 2014; Seeber et al., 2013). Evidence for this mechanism is provided by the tethering of Ino80-LexA to chromatin which can stimulate chromatin mobility, and which depends on the catalytic activity of Ino80 (Neumann et al., 2012).

Increased chromatin mobility is proposed to facilitate homology search during HR (Dion et al., 2012; Mine-Hattab and Rothstein, 2012) and provides an attractive mechanism to facilitate spatial exploration of the nucleus. The global movement of unbroken chromosomes would also increase

27 the movement of donor sequences and computational modeling suggests a two-fold decrease in search time when the second locus is mobile as opposed to fixed in the nucleus (Gehlen et al., 2011). Despite the conceptual appeal of this model, the link between increased chromatin mobility and homology search has not been conclusively tested as DSB-mobility defective mutations, for example INO80 mutants, have additional roles in DSB repair (Jeggo and Downs, 2014). The identification of specific mobility mutations or separation-of-function mutations is therefore required to directly test this hypothesis.

Whether DSBs exhibit increased mobility in mammalian cells is less clear and seems to depend on both the type of DNA damage used and the experimental method. Early experiments using fixed cells revealed that DSBs occupy stable positions in the nucleus following ultra-soft X-rays (Nelms et al., 1998), but that DSBs can also cluster (Aten et al., 2004), suggesting that some DSBs can move within the nucleus. DSBs marked by the repair factor 53BP1 also exhibit increased mobility in response to ionizing radiation (Krawczyk et al., 2012; Lottersberger et al., 2015) and 53BP1 promotes the mobility of uncapped telomeres to promote NHEJ (Dimitrova et al., 2008; Lottersberger et al., 2015). Telomeres seem particularly prone to movement and have also been observed to undergo long-range directional movement to facilitate lengthening by ALT (Cho et al., 2014). Nevertheless, not all DNA ends display increased mobility and experiments following GFP-tagged histone H2B revealed that most DSBs are immobile within the nucleus (Kruhlak et al., 2006). Experiments monitoring the mobility of chromatin bound LacI-GFP before and after an I-SceI endonuclease induced DSB have also failed to detect changes in mobility (Roukos et al., 2013; Soutoglou et al., 2007) and one active pathway that limits mobility involves the Ku complex which tethers broken ends and is proposed to protect against chromosome translocations (Soutoglou et al., 2007).

1.5 Summary and rationale

The formation of a single DSB activates a highly conserved signalling response which communicates the presence of damage to the cell. This response is driven by the Mec1 and Tel1 kinases in budding yeast which phosphorylate numerous targets to coordinate DSB repair and cell cycle arrest. The identification and characterization of these DNA damage checkpoint targets remains a key goal in understanding the cellular response to DSBs. In particular, a DSB triggers

28 increased chromosome movement which is dependent on Mec1 activity but the target responsible for this phenomenon is unknown.

Repair by HR requires strand invasion of a homologous donor sequence which can be located on the nearby sister chromatid or on a separate chromosome. How the homology search occurs within the nucleus is a key question and is proposed to be aided by increased chromosome movement. This model can be directly tested with specific mutations once the mechanism of DSB-induced mobility is more fully understood.

In contrast to the response at DSBs, telomeres protect chromosomes ends from recognition by the DSB response and telomerase must also be carefully regulated to suppress telomere addition. This is primarily accomplished by the Pif1 helicase; however, short telomeres undergo preferential elongation even in the presence of Pif1 suggesting the presence of a regulatory mechanism which connects telomere length and Pif1 activity. The investigation of this mechanism will further reveal how cells deal with their chromosome ends and what is the dividing line between DSBs and telomeres.

Chapter 2 Materials and Methods 2.1 General yeast strains and growth

Strains were constructed by standard allele replacement, PCR-mediated gene deletion or epitope- tagging methods, or via transformations of the indicated plasmids. The desired mutations were selected by prototrophy or drug selection and verified by PCR or sequencing. Standard yeast media and growth conditions were used. Cells were grown in supplemented minimal medium (SD: 2 g/L amino acids dropout mix, 5 g/L ammonium sulfate, 1.7 g/L yeast nitrogen base) or in rich medium (XY: 20 g/L bactopeptone, 10 g/L yeast extract, 0.1 g/L adenine, 0.2 g/L tryptophan), containing 2% glucose, 2% raffinose or 3% galactose as indicated.

2.2 Telomere addition strains

Telomeric repeats were cloned into the pVII-L plasmid which features an HO endonuclease cut site, a URA3 selection marker, and homology arms for integration at the ADH4 locus (Gottschling et al., 1990). Longer telomeric repeats were assembled using commercial gene synthesis (Mr. Gene) while Quikchange mutagenesis (Stratagene) was performed for further manipulation of repeat sequences. Insertions and deletions of up to 30 bp of TG repeats were robustly obtained in a single round of mutagenesis. Quikchange mediated shortening of a large TG250 sequence also yielded a wide range of shorter repeats. All repeats were verified by DNA sequencing before integration.

The TG82-HO cassette on Chr VII was replaced by integrating SalI-EcoRI digested pVII-L plasmids and selecting for colonies on SD-URA. Single integration of the plasmid and HO cleavage at the locus was confirmed by Southern blot. Telomere addition strains were constructed in a rad52Δ background with a covering pRS414-Rad52 plasmid to facilitate genome manipulation through homologous recombination. Strains were cured of the plasmid by random loss and colonies were screened by replica-plating to SD-trp.Pif1 mutations were generated by Quikchange mutagenesis (Stratagene) on a pAUR101-pif1-m1 nuclear specific construct and integrated at the AUR1 locus in pif1-m2 cells. The est2-up34 mutation was generated by pop-in/pop-out gene replacement.

2.3 Chromatin mobility strains

Candidate kinetochore phosphorylation targets were cloned into pAUR101 (Takara Clontech) and serine-to-alanine mutations introduced by Quikchange (Stratagene). Wild-type or mutant alleles were integrated at the AUR1 locus and the endogenous allele deleted. Htz1 and Ipl1 mutant strains were constructed using a similar approach. Endogenous CEP3 was mutated using a pop-in/pop- out allele replacement method by cloning cep3-S575A into pRS306 and integrating the HpaI digested plasmid. The CEP3 locus was PCR-amplified from 5’fluoroorotic acid resistant pop-out colonies and sequenced to confirm the mutation. Endogenous CEP3 was also mutated by transforming a PCR overlap extension product fusing cep3-S575A and a NATMX cassette placed 100 bp downstream of the stop codon. Colonies were screened by PCR and the PCR products sequenced to confirm the mutation.

Strains for centromere inactivation experiments were generated by integrating the GAL1/10 promoter adjacent to CEN3 or CEN5. Centromere inactivation was confirmed by plating cells onto XY galactose which led to decreased viability.

A 117 bp HO cut site was integrated on Chr IV at a distance of 27, 101, 510 and 998 kb from CEN4 in a strain expressing LacI-GFP and Nup49-mCherry. The LacOx256 arrays from pSR13 were integrated at various genomic positions using a two-step targeting strategy (Rohner et al., 2008). pRS415-LacOx256 was cloned by inserting the BamHI-SalI fragment containing the LacO array from pSR2 (Rohner et al., 2008) into pRS415. The 117 bp HO cut site was also inserted at the AMD2 locus on Chr IV, and the ADH4 locus on Chr VII in cells expressing Nup49-mCherry and plasmid-borne pRS415-Rad52-YFP.

The JKM179 Nup49-GFP microscopy strain (DDY2784) was a gift from Sue Jaspersen. Cells expressing Mtw1-3xGFP were kindly provided by Sue Biggins. The Nup49-mCherry construct was kindly provided by Marc Meneghini. pRS415-Rad52-YFP was a gift from Rodney Rothstein. pSR2 and pSR13 were kindly provided by Susan Gasser. The HR reporter strains were kind gifts from Martin Kupiec. The ALF and CTF reporter strains were kindly provided by Phil Hieter and Grant Brown respectively.

2.4 Telomere addition analysis

Telomere addition assays were performed as previously described(Zhang and Durocher, 2010). Briefly, yeast cultures were grown overnight in XY + glucose to log phase and subcultured into XY + raffinose (2%) for overnight growth to a density of 2.5-7.5x106. Nocodazole (Sigma Aldrich) was added at 15 µg/mL for 2h to synchronize cells in G2/M before addition of galactose to induce HO endonuclease expression. Cells were plated on XY glucose plates before addition of galactose, and 4 h after galactose, and grown for 2 days. The total number of colonies were counted and colonies were replica-plated to media containing α-aminoadipic acid (αAA) to identify cells which had lost the distal LYS2 gene on Chr VII. The protein encoded by LYS2 converts αAA into a toxic semi-aldehyde and renders LYS2 positive cells sensitive to excess αAA. Frequency of telomere addition was calculated as the percent of surviving post-galactose colonies that were αAA resistant. An alternative calculation, (αAA resistant colonies/ (pre-galactose colonies - αAA sensitive colonies), revealed the same threshold of Pif1 activity between the TG18 and TG34 ends, but with increased variability between experiments.

2.5 Genomic DNA extraction

Genomic DNA was isolated using a phenol-chloroform extraction protocol. Briefly, overnight cultures of cells were grown to saturation, pelleted, and resuspended with 200 µL ‘Smash & Grab’ lysis buffer (10 mM Tris, pH 8.0, 1 mM EDTA, 100 mM NaCl, 1% SDS, 2% Triton X-100). 200 µL of glass beads (Sigma Aldrich, 400-600 µm diameter) were added along with 200 µL phenol- chloroform (1:1). Cells were lysed by vortexing for 5 min before addition of 200 µL TE buffer (10 mM Tris-HCl pH 8, 1 mM EDTA). Samples were centrifuged at 4°C and DNA from the upper layer precipitated with the addition of 1 mL ice-cold 100% ethanol and centrifuged at 4°C. The DNA pellets were resuspended in 200 µL TE with 300 µg RNAse A (Sigma) and incubated at 37°C for 30 min. DNA was again precipitated with the addition of 1 mL ice-cold 100% ethanol and 10 µL of 4M ammonium acetate, dried, and resuspended in TE.

2.6 Southern blots for telomere addition and length

Fifteen micrograms quantities of genomic DNA were digested overnight with SpeI (for TG82 strains) or EcoRV (for all other TG repeat lengths). Digested DNA was run on a 1% agarose gel

32 in 0.5X TBE buffer (45 mM Tris-borate, 1 mM EDTA) at 100V for 6 hours, denatured in the gel for 30 min with 0.5 M NaOH and 1.5 mM NaCl, and neutralized for 30 min with 1.5 M NaCl and 0.5 mM Tris-Cl pH 7.5. DNA was transferred to Hybond N+ membrane (GE Healthcare Life Sciences) using overnight capillary flow and 10X SSC buffer (1.5 M NaCl, 150 mM sodium citrate). Membranes were UV crosslinked (Stratagene) and blocked at 65°C with Church hybridization buffer (250 mM NaPO4 pH 7.2, 1 mM EDTA, 7% SDS). Radiolabelled probes complementary to the ADE2 (for TG82 strains) or URA3 gene (for all other TG repeat lengths) were generated from purified PCR products using the Prime-It Random labelling kit (Stratagene) and α32-dCTP. Membranes were probed overnight, washed three times with 65°C Church hybridization buffer and exposed overnight with a phosphor screen (GE Healthcare Life Science) before imaging on a Storm or Typhoon FLA 9000 imager (GE Healthcare Life Sciences). Quantification of the added telomere signal (above CUT band) was performed in ImageQuant GE Healthcare Life Sciences) by subtracting the background signal before HO induction followed by normalization to the internal loading control (INT). Telomere length analysis was performed by digesting demonic DNA with XhoI and probing with a Y’-TG probe generated from the pYT14 plasmid (Shampay et al., 1984).

2.7 PCR mutagenesis screens

Mutant alleles were generated by error-prone PCR using Taq polymerase (New England Biolabs) and 0.25 mM MnCl, and purified using spin columns (Qiagen). The Pif1 mutagenesis screen was performed in TG82 pif1-m2 cells co-transformed with gapped pRS416-pif1-m1 and purified inserts and repaired plasmids were selected on SD-ura. The Cdc13 mutagenesis screen was performed in

TG34 cdc13Δ cells containing a covering YEp-CDC13-URA3 plasmid and co-transformed with gapped pRS425-CDC13 plasmid and PCR inserts and repaired plasmids selected on SD-ura before replica-plating to 5-Fluoroorotic acid (5-FOA) to remove the covering plasmid. Mutant cdc13 alleles which are defective in capping should be inviable in this step. Colonies from both screens were patched onto raffinose plates and grown for 2 days before replica plating to galactose plates for 4 hours, and finally to αAA plates after diluting cells with an agar plate. Plasmids were rescued using a phenol-chloroform extraction and transformed into Escherichia coli. Plasmids were sequenced to identify mutations and retransformed into the parental yeast strain to confirm the phenotype results from the plasmid mutation.

2.8 Chromatin mobility analysis

Yeast cultures were grown overnight to a density of 2.5-5x106 cells/mL in XY + raffinose, or SD + raffinose for plasmid selection as required. Galactose was normally added for 3 hours to induce a DSB as required, DSBs in diploid cells were induced for 2 h. Cells were mounted in concanavalin A- (Sigma Aldrich) coated 8-well Lab-Tek II chambers (Thermo Scientific Nunc) and maintained at 30°C during imaging in SD + raffinose. Live-cell time-lapse microscopy was performed using a DeltaVision widefield microscope (GE Healthcare Life Sciences) with a 100x/1.40 NA plan Apochromat oil immersion objective (Olympus) and CoolSNAP HQ2 CCD Camera (512x512, 128.8 nm/pixel, 2x2 bin) and limited to a 30 min window following DSB induction. Single plane images were acquired with GFP (475 nm excitation, 200 ms exp) or YFP (513 nm excitation, 200 ms exp) every 1.5 s for a total of 180 s while mCherry images were captured every 5th frame (575 nm excitation, 200 ms exp). Cells were excluded from analysis if they displayed cellular movement or misshapen nuclei, or if the tracked particle was lost during time-lapse acquisition. Nuclear alignment was performed using Nup49-mCherry images with a custom MATLAB (Mathworks) script while particles were tracked using the SpotTracker plugin in ImageJ to yield X,Y coordinates (Sage et al., 2005). MSD analysis was performed as previously described (Neumann et al., 2012) using a custom MATLAB script. I used the slope (m) of the first five time intervals (1.5 s-7.5 s) to determine the 2D diffusion coefficient (D) given that D = m/2d, where d is the number of dimensions. The radius of confinement was calculated using the formula Rc =5/4√MSD calculated from the average MSD value of the final 20 time points (121.5 s to 150 s). Data presented in Appendix C represent mean ± s.e.m.

2.9 Antibodies and immunoblotting

The following antibodies were used in this study: mouse anti-Myc 9E10 (sc-40, Santa Cruz Biotech, 1:5000), mouse anti-Pgk1 22C5D8 (459250, Life Technologies, 1:20000), rabbit anti- Clb2 y-180 (sc-9071, Santa Cruz Biotech, 1:500), rabbit anti-Htz1-K14ac (Upstate 07-719, 1:1000), and an affinity purified Rad53 antibody (1:5000). A phosphospecific Cep3-S575 antibody was generated by immunizing rabbits with a peptide containing phosphorylated Cep3-S575

(CKLRQE-pSLLEEE-NH2, Bio Basic) conjugated to keyhole limpet hemocyanin (Covance). Crude serum was purified with positive selection against Cep3-pS575 peptides immobilized to

NHS-Activated Sepharose (GE Healthcare Life Sciences). The base elution was concentrated and used for immunoblots at 1 µg/mL, and non-phosphorylated Cep3-S575 competitor peptide

(CKLRQESLLEEE-NH2, Bio Basic) was added at 10 µg/mL to increase specificity. The following secondary antibodies were used in this study: sheep anti-mouse conjugated to horseradish peroxidase (HRP; NA931, GE Healthcare Life Sciences, 1:5000), and a goat anti-rabbit conjugated to HRP (111-035-144, Jackson ImmunoResearch, 1:5000).

Trichloroacetic acid extracts were prepared from log phase yeast cells lysed with glass beads. Cells were treated with 250 µg/mL Zeocin for 1 h or with galactose for 3 h to induce an HO-DSB where appropriate. Lysates were separated by SDS-PAGE, transferred to nitrocellulose membrane (GE Healthcare Life Sciences), and analyzed by immunoblotting using Tris-buffered saline with 2% bovine serum albumin as a blocking agent. Key Western blots (Figure 3bc) were repeated three times to confirm the observation, synchronized cell experiments were performed twice.

2.10 Cell cycle arrest

Cells were arrested in the G1 phase of the cell cycle by adding 200 nM α-factor for 3 h, while G2/M arrest was achieved by a 2 h treatment with 15 µg/mL nocodazole (Sigma Aldrich). In both cases, arrest was confirmed by visual analysis of cellular morphology.

2.11 Recombinant protein production

Rad53-6xHis proteins were purified from Escherichia coli cells as previously described(Wybenga- Groot et al., 2014). Catalytically active Rad53 contains the A235S mutation, referred to here as wild-type (WT), to facilitate expression in E. coli. Briefly, BL-21 cells transformed with pET-15b

WT- or D339A- Rad53-6xHis were induced overnight at 18°C (A600 = 0.6, 0.2 mM IPTG). Cell pellets were resuspended in 20 mM HEPES pH 7.5, 200 mM NaCl, 10 mM imidazole, 1 mg/mL lysozyme, and cOmplete Protease Inhibitor Cocktail (Roche), and lysed by sonication. The crude lysate was applied 3 times to Ni-NTA agarose columns by gravity flow (Qiagen) at 4°C and washed with 20 mM HEPES pH 7.5, 200 mM NaCl buffer containing 10 mM then 30 mM imidazole. Bound proteins were eluted with buffer containing 350 mM imidazole, dialyzed into 20 mM HEPES pH 7.5, 100 mM NaCl, and flash frozen in liquid nitrogen.

A C-terminal fragment of Cep3 encompassing residues 563-608 was cloned into pMAL-c2 to generate wild-type or S575A Cep3-MBP fusion proteins for use as kinase substrates. Cep3-MBP was purified from BL-21 E. coli cells after a 2 h induction at 37°C (A600 = 0.8, 1 mM IPTG). Cell pellets were resuspended in HCB buffer (20 mM HEPES pH 7.5, 200 mM NaCl, 1 mM EDTA, 1 mM DTT) with 1 mg/mL lysozyme, and cOmplete Protease Inhibitor Cocktail (Roche), and lysed by sonication. Crude lysate was incubated with Amylose resin at 4°C overnight (New England BioLabs) and washed with HCB buffer. Bound proteins were eluted with HCB containing 50 mM maltose, dialyzed into HCB, and flash frozen in liquid nitrogen.

2.12 Rad53 kinase reactions

Two microgram quantities of MBP-Cep3 substrates were incubated with 500 ng of WT- or

D339A-Rad53-6xHis in kinase buffer (50 mM HEPES, pH 7.5, 250 mM NaCl, 20 mM MgCl2, 20 32 mM MnCl2, 2 mM DTT, 20 mM ATP) spiked with 1 µL of γ -ATP for 30 min at 30°C. 10 µL of Ni-NTA agarose (Qiagen) was added to deplete Rad53-6xHis and the reactions were stopped by boiling in Laemmli SDS-PAGE Sample Buffer. Samples were separated by SDS-PAGE, transferred to nitrocellulose membrane, and analyzed by phosphorimaging.

2.13 Visualization of kinetochores

Yeast expressing Mtw1-3xGFP were grown to mid-log phase in XY + glucose media and Zeocin was added at 50 or 250 µg/mL for 90 min. As a control, ndc80-1 mutants were shifted to 37°C (non-permissive) for 90 min. Cells were fixed with 4% (w/v) paraformaldehyde for 20 minutes and washed with KPO4/sorbitol buffer (1.2 M sorbitol, 100 mM potassium phosphate pH 7.5). Fixed cells were imaged on a DeltaVision Elite widefield microscope (GE Healthcare Life Science) with a 100x/1.40 NA plan Apochromat oil immersion objective (Olympus) and a sCMOS camera (1024 x 1024, 64.4 nm/pixel, 1x1 bin). Z-stacks containing 32 planes with a step size of 0.2 µm were acquired (475 nm excitation, 200 ms exposure) and deconvolved with SoftWoRx (Applied Precision). Maximum intensity projections were scored for detached kinetochore phenotypes as determined by the proportion of G2/M cells with more than two discernable Mtw1- 3xGFP foci. 100-200 G2/M cells were scored per condition in a minimum of 3 independent experiments. Representative images for figures were scaled 2-fold with bilinear interpolation in ImageJ.

2.14 Analysis of SPB-CEN dynamics

SPBs were labelled with endogenously expressed Spc42-tdimer2 (a DsRed variant) in cells containing a LacOx256 array at CEN5 and an HO cut site 27 kb from CEN4. Cells were prepared and immobilized for live-cell microscopy as outlined in our chromatin mobility analysis. Cells were imaged at 30°C on a DeltaVision Elite widefield microscope (GE Healthcare Life Science) with a 100x/1.40 NA plan Apochromat oil immersion objective (Olympus) and a sCMOS camera (1024 x 1024, 128.8 nm/pixel, 2x2 bin). Single plane images were acquired with GFP (475 nm excitation, 200 ms exp) and mCherry (513 nm excitation, 200 ms exp) channels every 1.5 s for a total of 180 s. As a DSB results in G2/M arrest, I selected G/2M cells with separated SPBs in asynchronous cycling populations as a control. Cells were excluded for analysis if they displayed cellular movement or if either fluorescent signal was lost during time-lapse acquisition. The positions of the SPB and CEN5 were tracked using the SpotTracker plugin in ImageJ to yield X,Y coordinates for both particles(Sage et al., 2005). For consistency, I tracked the mother SPB in all cells which was distinguished by its higher fluorescent signal intensity. The radial SPB-CEN 2 2 distance was calculated using d =√((XSPB-XCEN) +(YSPB-YCEN) ) for each frame, yielding 121 distances per cell. SPB-CEN dynamics were analyzed using mean square SPB-CEN distance changes as previously described(Dorn et al., 2005) using a custom MATLAB script. SPB-CEN distances changes between each time lapse frame were divided into antipoleward (increase in SPB- CEN distance) and poleward motion (decrease in SPC-CEN distance) and the average speed of the directional movement calculated.

2.15 DNA damage sensitivity

Overnight yeast cultures were serially diluted 5-fold and spotted onto XY plates containing methyl methanesulfonate (MMS, Sigma Aldrich), Zeocin (Life Technologies), hydroxyurea (HU, Sigma Aldrich), or camptothecin (CPT, Sigma Aldrich). Plates were grown at 30°C for 2-3 days.

2.16 HR repair analysis

DSB repair by HR was measured using strains designed to test various spatial configurations of repair (Agmon et al., 2013). Overnight yeast cultures from individual colonies were grown in XY + raffinose to mid- log phase and plated onto XY plates containing glucose as a control and

37 galactose to induce a DSB. Repair efficiency was calculated by dividing the number of surviving colonies on the galactose plate by the number of colonies on the glucose control plate. 4-8 independent replicates were performed for each genotype. The kinetics of repair was monitored in NA60 by inducing a DSB in XY + raffinose cultures and taking samples in a time course. Genomic DNA was isolated by phenol-chloroform extraction and the cut locus was amplified by PCR and purified using PCR purification columns (Qiagen). PCR products were digested with ClaI, separated on an agarose gel, and stained with ethidium bromide. Genomic DNA from a colony that survived on XY + galactose was used as a positive control.

2.17 Checkpoint analysis

Overnight yeast cultures grown in XY + raffinose were plated onto XY + galactose plates. G1 cells were isolated using an Axioskop Tetrad microscope (Zeiss). Cell cycle progression was monitored every 30 min and checkpoint arrest length was measured from the time of a large-budded dumbbell morphology to the appearance of the first new bud. 100-200 cells were measured for each genotype in a minimum of two independent experiments.

2.18 A-like faker (ALF) assay

ALF assays were performed as previously described(Warren et al., 2004; Yuen et al., 2007). MATα his3 cells that lose the MAT locus, designated “a-like fakers”, are capable of mating to a MATα his1 tester strain and are identified by prototrophic selection. Individual colonies of each 2 7 genotype were patched in 1.2-cm squares on XY + glucose and grown overnight. 5x10 cells from overnight cultures of the YPH316 MATα tester strain were plated on XY + glucose and allowed to dry before patches were transferred to the tester mating lawn by replica plating. Mating plates were grown for an additional 24 h before replica plating to SD-his. Surviving HIS+ colonies in each patch were counted after 4 days growth and 24-60 patches were scored for each genotype.

2.19 Chromosome transmission fidelity (CTF) assay

CTF assays were performed as previously described(Yuen et al., 2007). Briefly, strains were grown on SD-ura + glucose plates to maintain selection of the chromosome fragment. Individual colonies were grown overnight in XY + glucose cultures to mid-log phase and plated on SD +

38 glucose media with low adenine (5 mg/L) to allow for red pigmentation to develop. Plates were grown for 6-8 days and the number of half-sectored colonies were counted (indicating loss of the chromosome fragment in the first cell division), and divided by the total number of colonies. 3-8 independent replicates were performed for each genotype.

2.20 Chromatin immunoprecipitation (ChIP)

Strains expressing 13xMyc tagged CEP3 or cep3-S575A alleles were grown in XY + raffinose and a DSB induced by addition of galactose for 3 hours. Cells were fixed in 1% formaldehyde for 20 min and the sheared chromatin fraction immunoprecipitated overnight using an anti-Myc 9E10 antibody (Santa Cruz Biotech) and Pan-Mouse IgG Dynabeads (Life Technologies). DNA was purified using PCR purification columns (Qiagen) and fold-enrichment determined by quantitative real-time PCR using Power SYBR Green Mix (Life Technologies). Enrichment of a probe adjacent to CEN3 was compared between samples and normalized to the TSC11 locus 193 kb from CEN3. Five independent replicates were performed for each condition.

2.21 Peptide pulldown assays

Extracts were generated from 50 mL overnight cultures of a strain expressing Rxt3-13xMyc. Cell pellets were resuspended in buffer containing 50 mM HEPES, 150 mM NaCl, 1 mM DTT, 0.1% Triton X-100, 1 mM EDTA, and cOmplete Protease Inhibitor Cocktail (Roche), and lysed by vortexing with glass beads. Extracts were spun to remove insoluble debris and protein concentration measured by BCA assay (Pierce). 2 µg quantities of biotinylated Cep3 peptides (Bio Basic) encompassing S575 in unphosphorylated (LNKLRQESLLEEEDE), phosphorylated (LNKLRQEpSLLEEEDE), and a scrambled phosphorylated form (NLLEKERpSEQELLED) were incubated with 4 mg of cellular extracts for 2 h at 4°C. 20 µL Streptavidin Dynabeads (ThermoFisher Scientific) were added and reactions were incubated for 1 h. Beads were washed three times with buffer containing 150 mM NaCl and 0.1% Triton-X100 and the beads boiled in Laemmli SDS-PAGE Sample Buffer. Samples were separated by SDS-PAGE and analyzed by immunoblot.

2.22 Break-induced replication (BIR) assay

BIR assays were performed as previously described (Lydeard et al., 2007). Briefly, JRL 346 derived strains were grown overnight in XY + raffinose to log phase and plated onto XY containing either glucose or galactose. Cells surviving the DSB were replica-plated to plates containing hygromycin B to confirm the outcome of repair. DSB repair by BIR was calculated by dividing the number of hygromycin B sensitive colonies by the initial glucose colonies. Genetic assays were performed with two separate clones of each mutant for a minimum combined n=6.

2.23 Break-induced replication PCR assay

JRL 346 derived strains were grown in XY + raffinose and 3% galactose added in log phase to induce the DSB. Time points were taken at 0, 3, 6, 9, 12, 19, and 24 hours post induction and genomic DNA extracted. Semi-quantitative PCR was used to measure the unique BIR repair product using primers P1 and P2 as previously described (Lydeard et al., 2007). PCR was performed using NEB Taq polymerase under linear conditions and products separated in 1% agarose stained with ethidium bromide. Gels were imaged using a Bio-Rad Gel Doc XR+ with constant exposure parameters and the resulting images analyzed in ImageQuant using volume quantitation. Percent repair by BIR was calculated by normalizing the BIR amplicon signal to the loading control HTA2 amplicon, subtracting the t=0 signal, and normalizing to genomic DNA from a clone that had undergone BIR. Error bars represent standard deviation between three biological replicates.

2.24 Cell senescence assays

Cells were grown to saturation in 10 mL cultures of XY with 2% glucose and diluted to 5x105 cells mL every 24 h. Cell density was measured using a hemocytometer. Samples were taken at days 0, 4 and 8 for telomere length analysis by Southern blotting. Genomic DNA was digested with XhoI. The approximate number of generations was calculated on the basis of the initial growth on plates after sporulation (3 days) and the cell density during liquid culturing.

2.25 Yeast two-hybrid

Cells were transformed with pGBK and pGAD-T7 plasmids (Clonetech) and selected on SD-trp- leu. Overnight yeast cultures were serially diluted 5-fold and spotted onto SD-trp-leu-his plates

40 containing Zeocin (Life Technologies) and 3-Amino-1,2,4-triazole (3-AT; Sigma Aldrich), a competitive inhibitor of the HIS3 reporter. Plates were grown at 30°C for 2-3 days.

2.26 Statistics

All statistical analysis was performed with GraphPad Prism v5.02 (GraphPad Software) using Student’s t-test or one-way analysis of variance (ANOVA) followed by Bonferroni post-hoc analysis.

Chapter 3 A Pif1-dependent threshold separates DSBs and telomeres

Statement of contributions: I performed all of the experiments described in this chapter.

A manuscript related to this work is currently in preparation for publication.

3.1 Introduction

A fundamental question in chromosome biology is how cells differentiate between spontaneous DSB ends and telomeres, the natural ends of chromosomes. The failure to properly deal with each end has severe consequences for the cell as the inappropriate repair of telomeres can lead to chromosome fusions and mitotic breakage. Similarly, the activity of telomerase at DSBs can generate new telomeres, at a cost of the genetic information distal to the break. While DSBs and telomeres reflect extreme positions on the spectrum, a continuum of DNA ends exist between them including critically short telomeres and DSBs occurring in telomeric-like sequence: all of which require a decision, should the end be repaired or elongated by telomerase?

The activity of telomerase must be tightly regulated at DSB sites and this is accomplished in S. cerevisiae by the telomerase inhibitor Pif1 (Schulz and Zakian, 1994; Zhou et al., 2000). Pif1 preferentially unwinds RNA-DNA hybrids in vitro (Boule et al., 2005) and is thought to remove the TLC1 RNA template from telomeres (Li et al., 2014; Phillips et al., 2015), but it is not clear whether Pif1 performs the same function at DSBs. Arguing against this model is the observation that telomere addition events do not preferentially occur at TLC1 binding sites in the absence of Pif1 (Putnam et al., 2004).

Previous work in our lab revealed that Pif1 is active at DNA ends containing 18 bp of telomeric repeats (referred to as TG18), but has no effect at the TG82 end (Zhang and Durocher, 2010). This result suggests that the TG82 substrate is interpreted as a short telomere by the cell and allowed to elongate in a manner unchecked by Pif1. I sought to investigate the molecular basis of this DNA end-fate decision using the activity of Pif1 as a cellular sensor.

3.2 Results

3.2.1 Identification of a threshold for Pif1 sensitivity

To characterize the dividing line between DSBs and telomeres I used a genetic system in which galactose-inducible HO endonuclease can be expressed to create a single DSB at the ADH4 locus on Chr VII-L (Figure 3.1ab) (Diede and Gottschling, 1999; Gottschling et al., 1990). By placing different lengths of telomeric (TG1-3)n sequence adjacent to the HO cut site I can study how these

DNA ends are dealt with using two readouts: a genetic assay for telomere addition based on the loss of the distal LYS2 marker, and by Southern blot to monitor the length of the DNA end. The HO endonuclease cut site in this system contributes one additional thymine to the inserted telomeric seed accounting for a one base pair discrepancy from past reports. Previous work indicated that Pif1 is active at TG18, but not TG82 (Zhang and Durocher, 2010) so I first constructed strains containing 34, 45, 56, and 67 bp of telomeric repeats in both wild-type and nuclear Pif1 deficient pif1-m2 cells (see Appendix B for all TG repeat sequences). I observed similar rates of telomere addition at all DNA ends in both backgrounds indicating that 34 bp of telomeric repeat is sufficient to render a DNA end insensitive to Pif1 (Figure 3.1c). To account for variations between different DNA ends in HO cutting efficiency and the ability to recruit telomerase, normalization of telomere addition frequency to pif1-m2 cells provides a clear readout of Pif1 activity at each DNA end yielding insight into the cellular end-fate decision (Figure 3.1d).

Analysis of the DNA ends by Southern blot also revealed robust telomere addition at the TG34 substrate in PIF1 cells mirroring the genetic assay (Figure 3.1e).

The standard genetic assay for telomere addition includes a nocodazole arrest before DSB induction as telomerase is active in S and G2 (Diede and Gottschling, 1999), however, asynchronously dividing cells also exhibited the same phenotype at the TG18 and TG34 ends (Figure 3.1f). To study telomere addition by telomerase and not by recombination based processes, telomere addition strains are constructed in a rad52Δ background. The presence of

RAD52 in this assay also does not affect the observed behavior of Pif1 at the TG18 and TG34 ends (Figure 3.1f).

To further refine the threshold, I added 4 bp increments of TG repeat sequence to the centromeric side of the TG18 substrate yielding strains with 22, 26, 30, 34, and 38 bp of telomeric repeats. Importantly, with the exception of length, these strains contain the same DNA sequence and share a common distal end. Analysis of telomere addition revealed that Pif1 is active at DNA ends up to

TG26 while the frequency of telomere addition increased at the TG30 end and beyond (Figure 3.2a). As telomeric repeats are heterogeneous in nature, I next determined if this phenotype is dependent on the particular sequence used. I selected three random sequences from telomeric DNA and constructed strains with DNA ends containing either 26 or 36 bp of each sequence. Consistent with my initial observations, telomere addition was blocked by Pif1 at all TG26 ends, whereas the corresponding TG36 ends resulted in telomere addition in the presence of Pif1 (Figure 3.2bc).

Figure 3.1. Characterizing a threshold of Pif1 activity at DNA ends

Figure 3.1. Characterizing a threshold of Pif1 activity at DNA ends. a, Schematic of a system to study the fate of DNA ends. Telomeric repeats are placed adjacent to an HO cut site (HOcs) on at the ADH4 locus on Chr VII. Telomere addition can be measured using a genetic assay based on the loss of the distal LYS2 gene as measured by resistance to α-aminoadipic acid. Southern blotting with a probe complementary to URA3 (black bar) allows for visualization of DNA end stability. b, Sequence of the TG18 substrate and the overhang produced by the HO endonuclease. The C-rich strand runs 5’ to 3’ towards the centromere and is resected following DSB induction to expose a 3’ G-rich overhang. c, d, Telomere addition frequency at DNA ends containing 18-82 bp of TG sequence (c), and normalized to pif1-m2 cells for each DNA end (d). Data represents the mean ± s.d. from a minimum of n=3 independent experiments. See Appendix B for the sequences of all DNA ends. e, Southern blot of DNA ends containing TG18 and TG34 ends in wild-type and pif1- m2 cells following HO induction. A URA3 probe was used to label the ura3-52 internal control (INT) and the URA3 gene adjacent to the TGn-HO insert (PRE) which is cleaved by HO endonuclease (CUT). Newly added telomeres products are visualized as the heterogeneous smear above the CUT band. f, Telomere addition frequency in rad52Δ cells synchronized with 15 µg/mL nocodazole for 2 h (+ noc), asynchronous cells (- noc), and in cells containing a pRS415-RAD52 plasmid. Data represents the mean ± s.d. from n=3 independent experiments.

Visualization of the combined TG repeat substrates revealed a striking transition with regards to Pif1 function (Figure 3.2d). By using Pif1 as a cellular sensor I propose that the 26 to 34 bp window of telomeric sequence is the dividing line between what the cell interprets as a DSB, and what is a critically short telomere. These data suggest that DNA ends containing 34 bp or more of telomeric DNA are allowed to elongate in a manner unimpeded by Pif1 and I herein refer to this phenomenon as the DSB-telomere transition.

3.2.2 Pif1 is not inhibited by DNA damage kinases

One attractive mechanism for the observed DSB-telomere transition is that Pif1 might be inactivated at DNA ends containing longer telomeric repeats. Prime candidates for this regulation include the central DNA damage kinases and previous work has identified that Tel1 promotes telomerase elongation of the TG82 end (Frank et al., 2006), and targets short telomeres for elongation (Sabourin et al., 2007). These results raised the possibility that Tel1 antagonizes Pif1 at short telomeres so I deleted TEL1 in both wild-type and pif1-m2 backgrounds and followed the

TG82 DNA end by Southern blot. Although telomere addition was reduced in tel1Δ cells, I observed a similar decrease in pif1-m2 cells indicating that TEL1 and PIF1 function in separate pathways (Figure 3.3ab). Consistent with this observation, the loss of TEL1 did not affect the observed DSB-telomere transition at the TG18 and TG34 DNA ends (Figure 3.3c). Pif1 contains five consensus S/T-Q Mec1 and Tel1 phosphorylation sites in Pif1; however, their mutation in the pif1-5AQ allele (S148A, S180A, T206A, S707A, T811A) also did not decrease telomere addition at the TG34 end (Figure 3.3d). I next tested for the involvement of Mec1 and Rad53 in regulating Pif1, but the deletion of these kinases also failed to inhibit telomerase in a Pif1-specific manner at the TG82 end (Figure 3.4a-d), together suggesting that Pif1 is unlikely to be regulated by DNA damage kinases on the telomeric side of the transition.

I reasoned that Pif1 might be regulated through unanticipated post-translational modifications or protein interactions and next performed a PIF1 mutagenesis screen to identify gain-of-function mutations that inhibit telomere addition at the TG82 end. Briefly, a gapped plasmid containing the pif1-m1 nuclear isoform was transformed with error-prone PCR amplified pif1 products (error rate of 0.008 nucleotides/position) and telomere addition assessed using a modified plate-based genetic assay (Figure 3.5ab).

Figure 3.2. Pif1 activity at DNA ends reveals a DSB-telomere transition

Figure 3.2. Pif1 activity at DNA ends reveals a DSB-telomere transition. a, b, Telomere addition frequency normalized to pif1-m2 cells at DNA ends containing 18-38 bp of TG sequence (a), and ends containing 26 bp or 36 bp versions of natural telomeric (TG1-3)n sequence (b). Data represents the mean ± s.d. from n=3 independent experiments. c, Southern blot of DNA ends containing the TG26b and TG36b ends in wild-type (WT) and pif1-m2 cells following HO induction. A URA3 probe was used to label the ura3-52 internal control (INT) and the URA3 gene adjacent to the TGn-HO insert (PRE) which is cleaved by HO endonuclease (CUT). d, Summary of telomere addition frequency normalized to pif1-m2 across the spectrum of TG repeat substrates.

Screening of approximately 2 500 Pif1 mutants resulted in the identification of five suppressors of telomere addition; however, rescued plasmids from these strains failed to reproduce the result when re-transformed indicating that the phenotype was due to a secondary mutation. Together these data challenge the hypothesis that Pif1 is inactivated at the TG34 and TG82 DNA ends and I next considered alternative explanations for the observed DSB-telomere transition.

Figure 3.3. Pif1 is not inactivated by Tel1 at short telomeres

Figure 3.3. Pif1 is not inactivated by Tel1 at short telomeres. a, b, Southern blot of the TG82 DNA end in wild-type (WT) and pif1-m2 cells combined with a TEL1 deletion following HO induction. An ADE2 probe was used to label the ade2Δ1 internal control (INT) and the ADE2 gene adjacent to the TGn-HO insert (PRE) which is cleaved by HO endonuclease (CUT). Quantification of the newly added telomere signal (b) calculated by subtracting the pre-cut background and normalizing to the INT control. Data represents the mean ± s.d. from n=2 independent experiments. c, Telomere addition frequency at the TG18 and TG34 DNA ends in tel1Δ mutants. Data represents the mean ± s.d. from n=3 independent experiments. d, Telomere addition frequency at the TG34 DNA end in pif1-m2 cells (-) and cells expressing a wild-type (WT) or pif1- 5AQ (S148A, S180A, T206A, S707A, T811A) nuclear specific pif1-m1 allele. Data represents the mean ± s.d. from n=3 independent experiments.

Figure 3.4. Loss of MEC1 or RAD53 does not affect Pif1 at short telomeres

Figure 3.4. Loss of MEC1 or RAD53 does not affect Pif1 at short telomeres. a-d, Southern blot of the TG82 DNA end in sml1Δ and sml1Δ pif1-m2 cells combined with the deletion of MEC1 (a) or RAD53 (c) following HO induction. SML1 was deleted to suppress the lethality of mec1Δ and rad53Δ. An ADE2 probe was used to label the ade2Δ1 internal control (INT) and the ADE2 gene adjacent to the TGn-HO insert (PRE) which is cleaved by HO endonuclease (CUT). Quantification of the newly added telomere signal in mec1Δ cells (b), from n=1 experiment, and rad53Δ cells (d), from n=2 independent experiments. Data represents the mean ± s.d.

Figure 3.5. A genetic screen to identify Pif1 mutants that inhibit telomerase at the TG82 end

Figure 3.5. A genetic screen to identify Pif1 mutants that inhibit telomerase at the TG82 end. a, Schematic of a screen in TG82 pif1-m2 cells using a plate-based genetic assay for telomere addition. Repaired mutant pif1-m1 plasmids were selected on SD-ura before DSB induction. Plates were grown for 2-3 days with the exception of galactose, which was grown for 4 hours. An agar plate was used to reduce cell number before final selection. The protein encoded by LYS2 converts αAA into a toxic semialdehyde and renders LYS2 positive cells sensitive to excess αAA. b, Example plate from the screen. Pif1 mutants which prevent telomere addition are identified by the inability to grow on αAA media (green box). The TG18-HO strain was used as a control as telomere addition is normally inhibited by Pif1 at this substrate (white box).

3.2.3 Artificial telomerase recruitment does not outcompete Pif1

A simple explanation for the observed DSB-telomere transition is that longer telomeric repeats might have an increased ability to recruit telomerase. If correct, this model predicts that artificially increasing telomerase recruitment to the TG18 end might be sufficient to overcome Pif1 inhibition. The primary mechanism of telomerase recruitment involves an interaction between the DNA binding protein Cdc13 and the Est1 telomerase subunit (Nugent et al., 1996; Pennock et al., 2001) and to test this possibility I expressed the Cdc13-Est1 and Cdc13-Est2 fusion proteins (Evans and Lundblad, 1999). In agreement with previous work, expression of both fusions resulted in greatly elongated telomeres (Figure 3.6a); however, it did not increase telomere addition at the TG18 DNA end in the presence of Pif1 (Figure 3.6b). One caveat to this result is whether the fusion proteins are in fact capable of interacting with the TG18 end. To test this, I deleted the endogenous copy of EST1 and mutated the Cdc13-Est1 fusion to include the est1-60 (K444E) mutation thereby disrupting the interaction with endogenous Cdc13 (Pennock et al., 2001). As a result, telomerase activity in these cells should only arise from the provided fusion protein and my observation of

51 telomere addition in pif1-m2 cells suggests that the Cdc13-Est1 fusion can elongate the TG18 substrate in vivo (Figure 3.6b). Together these data indicate that Pif1 is able to effectively suppress telomere addition even in the presence of enhanced telomerase recruitment and suggests that telomerase recruitment does not underlie the observed DSB-telomere transition.

Figure 3.6. Artificial telomerase recruitment does not overcome Pif1 activity

Figure 3.6. Artificial telomerase recruitment does not overcome Pif1 activity. a, Southern blot for telomere length in TG18-HO wild-type (WT) and pif1-m2 cells with an empty plasmid (-) and those expressing plasmid-based Cdc13-Est1 or Cdc13-Est2 fusions. Cells were passaged for approximately 75 generations before genomic DNA extraction and a Y’TG probe was used to label telomere sequences. b, Telomere addition frequency of the cells described in panel a, and strains expressing a cdc13-est1-60 (K444E) fusion. Data represents the mean ± s.d. from n=3 independent experiments.

3.2.4 The DSB-telomere transition recapitulates the differential regulation of Pif1

While I previously hypothesized that Pif1 might be inhibited at DNA ends which resemble telomeres, an alternative regulatory model is that Pif1 is only activated at DNA ends with minimal telomeric sequence. Consistent with this model, Pif1 is reported to be phosphorylated after DNA damage in a Mec1-Rad53-Dun1-dependent manner and further characterization of this activity led to the identification of the pif1-4A mutant (T763A/S765A/S766A/S769A) which is unable to inhibit telomere addition at DSBs (Makovets and Blackburn, 2009). Importantly, mimicking

52 phosphorylation with the phosphomimetic pif1-4D allele can restore Pif1 activity (Makovets and

Blackburn, 2009). I first confirmed the function of these mutants at the TG18 DNA end by integrating nuclear specific pif1-m1 alleles at the AUR1 locus in pif1-m2 cells (Figure 3.7a). If

Pif1 phosphorylation only occurs at DNA ends with minimal telomeric repeats, such as TG18, then mimicking phosphorylation may be sufficient to inhibit telomere addition at DNA ends with longer telomeric repeats. Contrary to this prediction, the pif1-4D mutant did not restore Pif1 activity at the TG34 DNA end (Figure 3.7a) indicating that phosphorylation of these sites does not regulate the DSB-telomere transition.

Figure 3.7. The DSB-telomere transition recapitulates the differential regulation of Pif1

Figure 3.7. The DSB-telomere transition recapitulates the differential regulation of Pif1. a, Telomere addition frequency at the TG18 and TG34 DNA ends in pif1-m2 cells (-) and cells expressing a wild-type (WT), pif1-4A (T763A/S765A/S766A/S769A), or pif1-4D (T763A/S765A/S766A/S769A) nuclear specific pif1-m1 allele. Data represents the mean ± s.d. from n=3 independent experiments. b, Southern blot for telomere length in wild-type and pif1-m2 cells combined with the est2-up34 mutation. Cells were passaged for approximately 75 generations before genomic DNA extraction and a Y’TG probe was used to label telomere sequences. c, Telomere addition frequency at the TG18 and TG34 DNA ends in est2up-34 mutants. Data represents the mean ± s.d. from n=3 independent experiments.

Several lines of evidence indicate that Pif1 functions differently at both DSBs and telomeres. First, the pif1-4A mutation affects telomere addition at DSBs, but has no effect on telomere length

(Makovets and Blackburn, 2009). The inability of the pif1-4D allele to inhibit telomerase at TG34 therefore provides indirect evidence that this DNA end is interpreted by the cell as a minimal telomere. A second mutation that affects Pif1 activity has also been identified: the est2-up34

53 mutation, located in the finger domain of the telomerase reverse transcriptase subunit (Eugster et al., 2006). Interestingly, the est2-up34 mutant results in over-elongated telomeres in wild-type but not pif1-m2 cells indicating that the est2-up34 allele can at least partially bypass Pif1 inhibition (Eugster et al., 2006). To test if this holds true at DSBs I generated the est2-up34 mutation in strains with a TG18 DNA end. Although I observed increased telomere length in PIF1 est2-up34 cells (Figure 3.7b), telomere addition was not increased (Figure 3.7c), indicating that the est2- up34 mutation can bypasse Pif1 at telomeres but not at DSBs. Together these data support the idea that Pif1 possesses distinct functions at DSBs and telomeres, and that these differences are recapitulated in the TG18 and TG34 DNA ends on either side of the DSB-telomere transition.

3.2.5 Investigating the molecular trigger of the DSB-telomere transition

My results thus far have failed to identify a modification of Pif1 that can explain the DSB-telomere transition so I next focused on whether a unique property of the TG34 DNA end renders it insensitive to Pif1 activity. Two of the first factors recruited to DNA ends are the MRX and Ku complexes which function in both DSB repair and telomere maintenance, however, the loss of either complex had no effect on either side of the transition (Figure 3.8a). Indeed, the binary nature of the DSB-telomere transition strongly suggests that the discrete binding of a telomeric protein triggers insensitivity to Pif1. As Cdc13 binds to single-stranded DNA at the distal end, an attractive prediction is that Rap1 bound to the double-stranded telomeric DNA of longer repeats might inhibit Pif1. This model nicely correlates with the observed length of the DSB-telomere transition, as Cdc13 and Rap1 bind DNA sequences of 11 bp (Hughes et al., 2000) and 18 bp respectively (Gilson et al., 1993; Ray and Runge, 1999). Rap1 has also been previously identified to stimulate telomere addition (Grossi et al., 2001; Lustig et al., 1990; Ray and Runge, 1998).

Rap1 is an essential protein and binds the consensus DNA sequence of 5’- ACACCCATACACC

-3’ containing an invariable CCC core (Graham and Chambers, 1994; Grossi et al., 2001; Wahlin and Cohn, 2000). Importantly, substitution of the middle cytosine to guanine in this motif abolishes Rap1 binding (Graham and Chambers, 1994; Grossi et al., 2001). To test whether Rap1 is required to bypass Pif1 activity at DNA ends I first generated synthetic telomeric sequences with strict

(TGTGG)n or (TG)n repeats in both 26 bp and 36 bp lengths. Unlike natural telomeric sequences, both sequences lack a CCC motif on the opposing strand. Despite these alternations, I still observed

54 increased telomere addition at TG36 ends in wild-type cells (Figure 3.8b), suggesting that Rap1 binding is not required for this phenomenon.

In a second approach, I constructed an array of four consecutive Rap1 binding sites with a spacer adjacent to the HO cut site containing 14 bp of random DNA or a TG14 sequence (Figure 3.8c). Disrupting individual Rap1 binding sites with a cytosine to guanine substitution yielded arrays with zero to four functional binding sites (Figure 3.8c). Consistent with previous work, telomere addition was stimulated by a functional Rap1 binding site, but surprisingly all the tested arrays were insensitive to Pif1, including the array with zero functional binding sites (Figure 3.8d).

Arrays with an adjacent TG14 spacer sequence displayed similar results with overall decreased rates of telomere addition possibly due to decreased HO cutting efficiency (Figure 3.8e). These result suggest that Rap1 binding is not required to bypass Pif1 activity and that additional features of the arrays are instead responsible. Finally, as telomere length regulation by Rap1 is coordinated through two downstream negative regulators of telomerase: Rif1 and Rif2, I asked whether these proteins are important for the DSB-telomere transition. Consistent with a Rap1-independent mechanism, telomere addition at the TG34 end was unaltered in rif1Δ rif2Δ mutants (Figure 3.8f).

Together, these results suggest that the resistance of the TG34 end to Pif1 is independent of Rap1. This possibility is perhaps not surprising as there are hundreds of Rap1 sites throughout the genome involved in transcriptional regulation (Lieb et al., 2001; Shore and Nasmyth, 1987).

3.2.6 Cdc13 and the fate of DNA ends

Cdc13 binds a minimum 11 bp TG sequence through its canonical OB-fold DNA binding domain (Hughes et al., 2000) (Figure 3.9a) suggesting that three molecules of Cdc13 might render DNA ends insensitive to Pif1. The distinct nature of the DSB-telomere transition argues that this process might involve the assembly of a higher order protein complex conferring unique properties to the

TG34 DNA end.

Figure 3.8. The DSB-telomere transition does not require Rap1

Figure 3.8. The DSB-telomere transition does not require Rap1. a, Telomere addition frequency at the TG18 and TG34 DNA ends in mre11Δ and yku70Δ mutants. Data represents the mean ± s.d. from n=3 independent experiments. b, Telomere addition frequency normalized to pif1-m2 cells at DNA ends containing 26 bp and 36 bp (TGTGG)n and (TG)n repeats. Data represents the mean ± s.d. from n=3 independent experiments. c, Schematic of an array of Rap1 binding sites adjacent to the HO cut site on Chr VII with a random 14 bp spacer or a TG14 spacer. Rap1 binding is disrupted by a cytosine to guanine mutation in the C-rich strand (position highlighted in red) to yield arrays of zero to four functional Rap1 sites. d, e, Telomere addition frequency at the Rap1 arrays described in panel c with a non-telomeric spacer (d) or TG14 spacer (e) between the HO cut site and the first Rap1 binding site. Data represents the mean ± s.d. from n=3 independent experiments. f, Telomere addition frequency at the TG18 and TG34 DNA ends in rif1Δ, rif2Δ, and rif1Δ rif2Δ double mutants. Data represents the mean ± s.d. from n=3 independent experiments.

Alternatively, the Cdc13 N-terminal OB-fold domain (OB1) forms dimers (Mitchell et al., 2010; Sun et al., 2011) and can also bind telomeric ssDNA repeats of 37 and 43 bp in vitro, but not 18 and 27 bp (Mitchell et al., 2010). I hypothesized that Cdc13 dimerization and the unique N- terminal binding mode might allow longer DNA ends to bypass Pif1 and sought to test this idea by disrupting dimerization with the cdc13-L91A mutation (Mitchell et al., 2010). Consistent with this model, telomere addition at the TG34 end was inhibited by Pif1 in cdc13-L91A cells (Figure 3.9b); however, further investigation revealed a growth defect in these mutants which was suppressed by pif1-m2 (Figure 3.9c).

Figure 3.9. The N-terminal dimerization mutant Cdc13-L91A impairs telomere addition

Figure 3.9. The N-terminal dimerization mutant Cdc13-L91A impairs telomere addition. a, Schematic of Cdc13 domain architecture consisting of four OB-fold domains (OB1-4) and a telomerase recruitment domain (RD). b, Telomere addition frequency at the TG18 and TG34 DNA ends in cdc13Δ cells expressing wild-type (WT) or cdc13-L91A from a low copy (pRS415) or high copy plasmid (pRS425). Data represents the mean ± s.d. from n=3 independent experiments. c, Spot assays to determine cell viability in cdc13Δ cells with a covering YEp-CDC13-URA3 plasmid and pRS415 or pRS425 plasmids expressing wild-type Cdc13 (WT) or Cdc13-L91A. 5-fold serial dilutions of yeast cultures were grown on SD-leu as a control, and on SD-leu+5-FOA to determine viability in the absence of the covering plasmid. Plates were grown at 30°C for 2-3 days.

This result was reminiscent of the defective cdc13-1 allele which is also suppressed by PIF1 loss (Addinall et al., 2008; Downey et al., 2006). High copy plasmid expression of cdc13-L91A was able to rescue the growth defect, but also increased telomere addition at the TG34 substrate (Figure 3.9b) arguing that the initially observed defect in cdc13-L91A mutants was not solely due to impaired N-terminal dimerization.

I sought to more precisely dissect out a unique function of Cdc13 that occurs at the TG34 end and performed a mutagenesis screen to identify CDC13 alleles which have become sensitive to Pif1 activity (Figure 3.10a). The screening of approximately 6 000 mutants led to the identification of fifteen hits which strongly reduced growth on αAA indicating that telomere addition was impaired. As this screen was performed in wild-type cells, I next determined if the mutations could support telomere addition in the absence of PIF1. Recovered plasmids were re-transformed into wild-type and pif1-m2 cells, and telomere addition analysis of the hits revealed two clones with minor phenotypes, five clones with greatly reduced telomere addition in both wild-type and pif1-m2 cells, and eight clones in which telomere addition was impaired in wild-type cells but relatively unaffected in pif1-m2 cells (Figure 3.10b). This observation suggested that this third group of mutations specifically sensitize the TG34 end to the activity of Pif1 and are herein referred to as cdc13-sp alleles (sensitive to Pif1).

DNA sequencing revealed an excessively high mutation average of eleven amino acid substitutions per cdc13-sp allele and methodical mapping experiments led to the identification of causative amino acid substitutions in six of the eight cdc13-sp mutants (Figure 3.11, highlighted in red). Three alleles had contributions from multiple substitutions: I87N and Y758N in cdc13-sp1, H12R and F728I in cdc13-sp72, and E566V, N567, and Q583K in cdc13-sp3 (Figure 3.12a). Cdc13-I87, like L91A, is also implicated in OB1 dimerization (Mitchell et al., 2010), again hinting that blocking this function may facilitate Pif1 activity. The moderate phenotype of Cdc13-I87N was likely only identified in the screen due to further exacerbation of the defect by Y758N (Figure 3.12a). The most important mutation in cdc13-sp3 was identified to be Q583K with a minor contribution from E556V/N567D; interestingly, all three residues are found in the DNA binding domain suggesting that weakening the association with telomeric DNA can also sensitize the TG34 end to Pif1.

Figure 3.10. A screen to identify Cdc13 mutants that prevent telomere addition at the TG34 end

Figure 3.10. A genetic screen to identify Cdc13 mutants that prevent telomere addition at the TG34 end. a, Schematic of a screen in TG34 cdc13Δ cells using a plate-based genetic assay for telomere addition. Repaired mutant cdc13 plasmids were selected on SD-leu and the covering YEp-CDC13-URA3 removed by plating on 5-FOA before DSB induction. This step also eliminates all inviable cdc13 mutations. Plates were grown for 2-3 days with the exception of galactose, which was grown for 4 hours. An agar plate was used to reduce cell number before final selection. b, Example re-testing plate from the screen. Cdc13 mutants which prevent telomere addition are identified by the inability to grow on αAA media (green box), compared to positive control wild- type cells which add telomeres (red box) and TG18 cells which do not (white box). c, Telomere addition frequency at the TG34 DNA end in PIF1 and pif1-m2 cells in a cdc13Δ background expressing recovered pRS425-Cdc13 mutants from the screen. Data represents the mean ± s.d. from n=1 experiment for hits #7-81, and n=2 independent experiments for all cdc13-sp alleles.

Conversely, three powerful single mutations could completely recapitulate the phenotype of the remaining three alleles and included S255P, F236S, and Q256H (Figure 3.12a). These three residues all map to the Cdc13 recruitment domain suggesting that weakening the association with

telomerase is another means to facilitates Pif1 activity at TG34. While mutations of Cdc13-F236 and Q256 have not previously been reported, the surrounding residues P235, F237, and S255 have been identified to impair telomerase function (Gao et al., 2010; Tseng et al., 2006). Telomere length in cdc13-sp alleles was reduced in both wild-type and pif1-m2 backgrounds, and the severity of the defect generally correlated the telomere addition phenotype (Figure 3.12b).

The diversity of Cdc13 mutations that can sensitize the TG34 substrate to Pif1 suggests that generally disrupting Cdc13 function facilitates Pif1 activity and shifts the balance away from telomere addition. In agreement with this idea, the classic telomerase null cdc13-2 allele (Lendvay et al., 1996; Nugent et al., 1996) was also sensitive to Pif1, a phenotype mirrored in cdc13-1 mutants grown at permissive temperature (Figure 3.12c), a mutant now known to disrupt OB2 dimerization and Stn1 binding (Mason et al., 2013). Analysis of hits from my screen which decreased telomere addition in both wild-type and pif1-m2 cells revealed powerful double mutations of critical residues including S255L/Q256R in clone 37, I87T/F236Y in clone 40, and F236S/E252K in clone 48, suggesting that further disruption of Cdc13 eventually impairs telomere addition even in the absence of PIF1. In line with this idea, the F235S/E252K/Q583K triple mutant severely impaired telomere addition even in pif1-m2 cells (Figure 3.12c).

3.2.7 Pif1 does not limit elongation of longer telomeric seeds

A final observation regarding the DSB-telomere transition involves the extension of longer

telomeric repeats. While the TG34 and TG82 substrates are rapidly elongated by telomerase

following DNA cleavage, repeats containing TG250 are inert when observed by Southern blot (Negrini et al., 2007). This observation is consistent with the known ability of cells to preferentially elongate short telomeres (Marcand et al., 1999; Teixeira et al., 2004). To further characterize this phenomenon, I generated strains containing 96, 119, 142, and 162 bp of telomeric sequence and

followed the DNA ends by Southern blot. While minor elongation was detectable at the TG119

DNA end, the TG96 and TG142 ends were inert within the timescale of the experiment (Figure

3.13ab). The increased extension of TG119 compared to the TG96 end suggests that specific features

of the TG96 substrate make it refractory to telomerase and not the length per se. A likely candidate

61 for this regulation is the number and affinity of Rap1 proteins that can bind to the array (Negrini et al., 2007). The limited extension of the TG162 substrate raised the possibility that Pif1 might counteract telomerase at longer repeats thereby promoting the preferential elongation of short telomeres. This possibility is supported by the recent observation that Pif1 association is increased at long telomeres (Phillips et al., 2015). Contrary to this idea, I found that the loss of Pif1 did not increase elongation of the TG162 DNA end (Figure 3.13cd) suggesting that additional regulatory mechanisms are at play. Together these results suggest that DNA ends containing 34 bp to approximately 120 bp of telomeric repeat are recognized as critically short telomeres and are subject to immediate elongation.

Figure 3.11. Mutations in cdc13-sp alleles

Figure 3.11. Mutations in cdc13-sp alleles. Mutations highlighted in red were identified by mapping experiments to contribute to the mutant phenotype. Mutations highlighted in blue target important Cdc13 residues with untested substitutions and are predicted to contribute to the defect.

Figure 3.12. Cdc13 mutations which sensitize the TG34 end to Pif1 activity

Figure 3.12. Cdc13 mutations which sensitize the TG34 end to Pif1 activity. a, Telomere addition frequency at the TG34 DNA end in PIF1 and pif1-m2 cells in a cdc13Δ background expressing wild-type or mutated pRS425-Cdc13. Data represents the mean ± s.d. from n=3 independent experiments. b, Southern blot for telomere length of the strains in panel a. Cells were passaged for approximately 75 generations before genomic DNA extraction and a Y’TG probe was used to label telomere sequences. c, Telomere addition frequency at the TG34 DNA end in PIF1 and pif1-m2 cells in a cdc13Δ background expressing wild-type or mutated pRS425-Cdc13. The cdc13-1 mutant was grown at a permissive temperature of 23ºC. Data represents the mean ± s.d. from n=3 independent experiments.

Figure 3.13. The preferential extension of short telomeres is independent of Pif1

Figure 3.13. The preferential extension of short telomeres is independent of Pif1. a, b, Southern blot of DNA ends containing 82-162 bp of (TG1-3)n sequence following HO induction. A URA3 probe was used to label the ura3-52 internal control (INT) and the URA3 gene adjacent to the TGn-HO insert (PRE) which is cleaved by HO endonuclease (CUT). Quantification of the newly added telomere signal (b) calculated by subtracting the pre-cut background and normalizing to the INT control. Data represents n=1 experiment. c, d, Southern blot of the TG162 DNA end in wild-type (WT) and pif1-m2 cells following HO induction. Probed and quantified (d) as described for panel a and b. Data represents n=1 experiment.

3.3 Discussion

The work presented here sheds light on how cells distinguish between DSBs and short telomeres and reveals a striking transition in the fate of DNA ends with regards to the activity of the telomerase inhibitor Pif1. My findings agree with previous reports which identified that linear plasmid substrates containing 41 bp of telomeric repeats are efficiently converted into telomeres

(Lustig, 1992) and that a TG22 sequence can promote telomere addition (Hirano and Sugimoto, 2007). The discovery here that increased telomere addition is due to the apparent inactivity of Pif1 is a novel insight and future work will be required to fully characterize the molecular switch that occurs at the DSB-telomere transition.

The observed behavior of Pif1 complements several known mechanisms which tightly integrate telomeric sequence length and the regulation of telomerase (Figure 3.14). The identified activity of Pif1 at telomeric repeats under 34 bp joins a Mec1-dependent mechanism which inhibits Cdc13 binding at repeats under 11 bp (Zhang and Durocher, 2010), highlighting the importance of inhibiting telomerase at DSBs. Conversely, I propose that DNA ends containing telomeric sequences of 34 bp to approximately 120 bp are recognized as critically short telomeres by the cell and are preferentially elongated in manner dependent on Tel1 (Arneric and Lingner, 2007; Chang et al., 2007). A final mechanism, the canonical counting mechanism of telomeres, limits telomere extension through the negative regulators Rif1 and Rif2 (Hirano et al., 2009; Levy and Blackburn, 2004; Marcand et al., 1997; McGee et al., 2010).

Figure 3.14. A model for the length-dependent regulation of telomerase at DNA ends

Figure 3.14. A model for the length-dependent regulation of telomerase at DNA ends. See the discussion for more detail.

Although my data indicates that Pif1 is not responsible for the relative stability of the TG162 repeat sequence, other work has proposed that Pif1 does promote the preferential elongation of short telomeres by specifically interacting with longer telomeres (Phillips et al., 2015).

In order to maintain genome stability, one might imagine that the length of telomeric repeat sequence necessary to overcome Pif1 should be greater than any natural sequence occurring within in the genome. Moreover, any longer sequences should therefore be prone to conversion into new telomeres and might be under negative selection during evolution due to the loss of genetic material. Consistent with this idea, the two longest (TG1-3)n repeats in the correct orientation outside of telomeric regions in budding yeast include a 35 bp sequence on Chr VII, and a 31 bp sequence on Chr VI (Mangahas et al., 2001).

My investigation into the molecular trigger of the DSB-telomere transition points to a key role for the DNA binding protein Cdc13. This conclusion is supported by work revealing that microsatellite repeats which contain Cdc13 binding sites stimulate telomere addition (Piazza et al., 2012), and recently that a hotspot on Chr V promotes Cdc13 binding and telomere addition (Obodo et al., 2016). Furthermore, the tethering of Cdc13, but not Rap1, to this site was shown to stimulate telomere addition (Obodo et al., 2016).

The ability of the Cdc13 OB1 domain to dimerize and bind DNA provides an attractive solution to the DSB-telomere transition; however, my results clearly indicate that sensitivity to Pif1 is not a unique property and can result from a variety of mutations throughout Cdc13, most notably in the recruitment domain. Although my genetic assay is based on the proportion of telomere addition events from viable colonies, this investigation is further confounded by the fact that the growth of defective CDC13 mutants is suppressed by the loss of PIF1. Weakening the ability of Cdc13 to recruit telomerase provides a satisfying answer as to why Pif1 regains activity at the TG34 end, but is unable to explain why the TG34 end is resistant to Pif1 in the first place, especially in light of the observation that fusing telomerase to Cdc13 is not able to overwhelm Pif1 at the TG18 substrate. Interestingly, the mammalian CST complex can bind single-stranded telomeric DNA 32 bp and longer (Miyake et al., 2009) suggesting that Cdc13 in combination with Stn1 and Ten1 may also possess unique binding properties.

One key unresolved issue is the mechanism by which Pif1 inhibits telomerase on either side of the DSB-telomere transition and my results with the pif1-4A and -4D alleles suggest that these

66 activities may be distinct. It is clear that Pif1 can remove telomerase RNA from telomeres (Boule et al., 2005), but genetic data suggests that Pif1 also has telomerase-independent activity as PIF1 loss also increases growth in cdc13-1 tlc1Δ cells (Dewar and Lydall, 2010). One potential mechanism for Pif1 at DSBs is that it promotes DNA end resection. This possibility is based on the observation that Pif1 facilitates end resection in cdc13-1 mutants (Dewar and Lydall, 2010) and that resection is known to impair telomere addition (Chung et al., 2010). Consistent with this model, telomere addition events occur closer to DSB sites in pif1-m2 cells (Chung et al., 2010). Pif1 is also proposed to function with the nuclease/helicase Dna2 in end resection and Okazaki fragment processing as the lethality of DNA2 loss is suppressed by removal of PIF1 and hints that Pif1 unwinds DNA flaps that are processed by Dna2 (Budd et al., 2006; Ribeyre et al., 2009).

This model therefore predicts that the TG18 end may be resected with the help of Pif1, and that resection is blocked by the TG34 end, both of which are directly testable using quantitative amplification of single-stranded DNA assays (Holstein and Lydall, 2012). Consistent with this prediction, a TG22 end was previously observed to partially suppress DNA end resection compared to a TG11 substrate (Hirano and Sugimoto, 2007). The model that Pif1 may function at DSBs by promoting end resections also provides a satisfying explanation as to why tethering telomerase to the TG18 end did not increase telomere addition.

In conclusion, using Pif1 as a cellular indicator for the DNA-end fate decision reveals a novel threshold that recapitulates several properties of DSBs and telomeres. I propose that the TG34 DNA end, approximately ten percent of a healthy budding yeast telomere, is interpreted by the cell as a minimal telomere and future work will be required to characterize its unique properties.

Chapter 4 DNA damage signalling targets the kinetochore to promote chromatin mobility

Statement of contributions: I performed all of the experiments described in this chapter.

The text and figures presented in this chapter were modified with permission from the following publication:

Strecker, J., Gupta, G.D., Zhang, W., Bashkurov, M., Landry, M.C., Pelletier, L., and Durocher, D. (2016). DNA damage signalling targets the kinetochore to promote chromatin mobility. Nat Cell Biol 18, 281-290. doi:10.1038/ncb3308

4.1 Introduction

In budding yeast, Mec1 signalling following a DSB results has been reported to increase the mobility of chromatin within the nucleus and is proposed to promote repair by HR (Dion et al., 2012; Mine-Hattab and Rothstein, 2012). While sister chromatids are physically tethered to damaged chromatin, HR can also occur between distant interchromosomal sequences thereby requiring a genome-wide search for homology (Renkawitz et al., 2014). The increased mobility of chromatin following chromosome breakage therefore provides an attractive mechanism to facilitate spatial exploration of the nucleus.

Although HR factors and chromatin remodelers have been implicated in DSB-induced mobility (Horigome et al., 2014; Mine-Hattab and Rothstein, 2012; Seeber et al., 2013), the target of Mec1- dependent signalling that triggers increased mobility is unknown. I sought to identify the Mec1 target as a means to uncover the mechanism and function of DSB-induced chromatin mobility.

4.2 Results

4.2.1 A system for measuring chromatin mobility

To study chromatin mobility I performed live-cell time-lapse microscopy to follow the Lac repressor (LacI) fused to green fluorescent protein (GFP; yielding LacI-GFP) bound to an array of the lacO operator sequence (LacOx256) located 6 kb from the HO endonuclease cut site at the MAT locus (Figure 4.1a). Images were collected on a widefield fluorescent microscope every 1.5 s for 180 s, before and 3 h after DSB induction. The nuclear periphery was labelled with Nup49- mCherry to facilitate nuclear alignment and LacI-GFP foci were tracked and analyzed using mean- square displacement (MSD) analysis, a robust measure of mobility (Dion and Gasser, 2013). To reduce light exposure, I utilized 2x2 binning which did not affect the sensitivity of chromatin tracking (Figure 4.1b), and cells survived imaging with minimal growth delay (Figure 4.1c). Despite several technical differences, this experimental system robustly detects increased chromatin mobility after DSB induction that is consistent with previous results (Dion et al., 2012; Mine-Hattab and Rothstein, 2012) (Figure 4.1d; refer to Appendix C for calculated mobility

69 parameters). Increased mobility was also dependent on the INO80 chromatin remodeler (Figure 4.1e) as previously described (Seeber et al., 2013).

4.2.2 Relieving constraints on chromosomes mimics DSB- induced mobility Budding yeast chromosomes are attached minimally at two sites: at telomeres, which are clustered at the nuclear periphery, and at kinetochores, which are tethered to the spindle pole body (SPB) via a microtubule throughout most of the cell cycle (Figure 4.2a) (Gasser, 2002). Both sources of attachment constrain chromatin mobility (Hediger et al., 2002; Verdaasdonk et al., 2013), and I thus asked whether relieving these constraints could recapitulate the motion observed after chromosome breakage. I disrupted telomeric attachment by deleting SIR4 (Gartenberg et al., 2004; Hediger et al., 2002), whereas kinetochore attachment was impaired by forcing centromeric transcription with the galactose inducible GAL1/10 promoter (Hill and Bloom, 1987). Manipulation of either constraint resulted in modest increases in mobility (Figure 4.2b), but their combined disruption was sufficient to mimic chromatin mobility observed after a DSB (Figure 4.2c). Neither SIR4 deletion nor centromere (CEN) inactivation further increased chromatin mobility following a DSB (Figure 4.2d), suggesting that relief of the constraints imposed by telomere attachment to the periphery and kinetochore attachment to the SPB (referred to hereafter as the centromeric constraint) are included in the DSB response. I first sought to identify how DSBs relieve the centromeric constraint and hypothesized that DNA damage signalling may target the kinetochore or SPB.

4.2.3 Identification of a kinetochore mutant that affects DSB-mobility

As DSB-induced chromatin mobility is dependent on Mec1 (Dion et al., 2012; Mine-Hattab and Rothstein, 2012), I mined phosphoproteomic data (Chen et al., 2010; Smolka et al., 2007) to identify kinetochore or SPB proteins that are phosphorylated in a Mec1-dependent manner. This search revealed six candidates: Bir1-S751, Cbf1-S45/48, Cep3-S575, Dad1-S89, Dad3-S43, and Glc7-S3. I generated serine-to-alanine mutations and observed wild-type mobility in the bir1, cbf1, dad1, dad3, and glc7 mutants before and after a DSB (Figure 4.3a).

Strikingly, the cep3-S575A mutant exhibited normal mobility in the absence of DNA damage, but was unable to increase mobility after chromosome breakage (Figure 4.3abc). As these strains expressed either wild-type or mutant Cep3 at the AUR1 locus with endogenous CEP3 deleted, I confirmed this phenotype using a S575A mutation at the CEP3 locus in both haploid and diploid cells (Figure 4.4ab).

Figure 4.1. A system for measuring chromatin mobility

Figure 4.1. A system for measuring chromatin mobility. a, Schematic of the primary system used for chromatin mobility analysis and representative images for chromatin mobility analysis. HOcs represents the HO endonuclease cut site. Chromatin is tracked by integrating LacOx256 arrays in cells expressing LacI-GFP and Nup49-mCherry. Scale bar represents 2 µm. b, X-axis displacement of a single tracked MAT locus over time in 1x1 or 2x2 binned images. c, Growth comparison of unexposed and imaged cells. Cells were held in a DeltaVision microscope at 30°C in SD media supplemented with raffinose and imaged at the indicated time points. Scale bar represents 4 µm. d, e, Mean square displacement (MSD) analysis of the MAT locus in cells before a DSB and 3 h after HO induction (d) and in INO80 mutant arp8Δ cells (e). All MSD data represents the mean ± s.e.m. from 3 independent experiments. Mobility parameters and the combined number of tracked cells for each genotype are presented in Appendix C.

Figure 4.2. Relieving constraints on chromosomes mimics DSB-induced chromatin mobility

Figure 4.2. Relieving constraints on chromosomes mimics DSB-induced chromatin mobility. a, Diagram of physical constraints imposed on budding yeast chromosomes. SPB, spindle pole body. b, MSD analysis of the MAT locus in wild-type (WT) cells and in strains where telomeric attachment (TEL off) or centromeric attachment (CEN off) is artificially disrupted in the absence of a DSB. c, MSD analysis of the MAT locus with the combined disruption of both constraints (CEN off + TEL off) in the absence of a DSB. d, MSD analysis of the MAT locus where telomeric (TEL off) or centromeric attachment (CEN off) is artificially relieved in the presence of a DSB.

Chromatin mobility can also be measured by tracking the DSB repair protein Rad52 fused to yellow fluorescent protein (Rad52-YFP), which accumulates at DSB sites (Lisby et al., 2003; Lisby et al., 2001). I observed decreased Rad52-YFP mobility in cep3-S575A cells compared to wild-type after DSB induction at the AMD2 (Chr IV) or ADH4 (Chr VII) locus (Figure 4.4cd). Together, these data indicate that Cep3-S575 is required for the DSB-induced mobility of multiple loci, including chromatin engaged in DSB repair.

Figure 4.3. A kinetochore component is required for DSB-induced chromatin mobility

Figure 4.3. A kinetochore component is required for DSB-induced chromatin mobility. a, MSD analysis of the MAT locus in strains with wild-type or serine-to-alanine (SxA) mutations of candidate Mec1 phosphorylation targets. b, Radius of confinement (a) and diffusion coefficients (b) of wild-type and cep3-S575A cells following a DSB. Data represents the mean ± s.e.m. pooled from 3 independent experiments (WT control n = 131 cells, WT DSB n = 81 cells, cep3-S575A control n = 189 cells, cep3-S575A DSB n = 111 cells). *** p<0.001; One-way ANOVA. c, Representative mobility traces of wild-type and cep3-S575A cells.

Figure 4.4. Cep3-S575 is required for DSB-induced mobility in multiple contexts

Figure 4.4. Cep3-S575 is required for DSB-induced mobility in multiple contexts. a, MSD analysis of the MAT locus in wild-type cells and a cep3-S575A mutant generated at the endogenous CEP3 locus by the pop-in/pop-out method. b, MSD analysis of the MAT locus in wild-type or cep3-S575A/cep3-S575A diploid cells containing an HO cut site on one homolog. c, d, MSD analysis of Rad52-YFP in response to a DSB at the AMD2 locus on Chr IV (c) or the ADH4 locus on Chr VII (d).

4.2.4 Cep3 is phosphorylated in response to DNA damage

Cep3 is a zinc-finger DNA binding protein that recognizes the centromeric CDEIII sequence as part of the CBF3 complex (Lechner, 1994; Lechner and Carbon, 1991). To confirm that Cep3 is a bona fide target of DNA damage signalling I generated an antibody against phosphorylated Cep3- S575, which is located in an unstructured C-terminal acidic activation domain (Figure 4.5a) (Purvis and Singleton, 2008). This antibody detected phosphorylation of both untagged and Myc- tagged Cep3 in response to the DNA-damaging agent Zeocin in a manner dependent on the S575 residue (Figure 4.5b). A single HO-induced DSB, as used in mobility experiments, was also sufficient for Cep3-S575 phosphorylation (Figure 4.5c). Cep3 phosphorylation is not simply a consequence of cell cycle arrest as it was observed only following Zeocin treatment in synchronized G1 or G2/M cells (Figure 4.5d).

Figure 4.5. Cep3 is phosphorylated after DNA damage

Figure 4.5. Cep3 is phosphorylated after DNA damage. a, Schematic of Cep3 domain architecture, the location of S575 (red), and the basic organization of a S. cerevisiae kinetochore. b, Immunoblot analysis using a phosphospecific Cep3-pS575 antibody on whole cell extracts from cells expressing endogenous Cep3 (WT or S575A) or Myc-tagged Cep3 (WT-13xMyc or S575A-13xMyc) treated with the DNA damaging agent Zeocin. Cells were treated with Zeocin (250 µg/mL) for 1 h (+), or left untreated (-). Myc and Pgk1 antibodies were used to control for loading. Checkpoint activation was confirmed by probing for Rad53, which undergoes a phosphorylation-dependent mobility shift in response to DNA damage. *, the Myc tag is cleaved from a fraction of Cep3-13xMyc to yield untagged Cep3. c, Immunoblot analysis using a phosphospecific Cep3-S575 antibody on whole cell extracts from cells expressing Myc-tagged Cep3 (WT or S575A) in response to a HO-induced DSB. Rad53 was probed to confirm checkpoint activation, Myc was used to control for loading. d, Immunoblot analysis of asynchronous (ASN), α-factor arrested (G1, or nocodazole arrested (G2/M) cells treated with Zeocin (250 µg/mL) for 1 h. Levels of the mitotic cyclin Clb2 were ascertained to confirm cell cycle synchronization. Pgk1 and Rad53 were probed to control for loading and to confirm checkpoint activation respectively.

I next analyzed the genetic requirements for Cep3 phosphorylation. As anticipated, deletion of MEC1 greatly reduced Cep3-S575 phosphorylation (Figure 4.6a), consistent with the impaired DSB-induced mobility of mec1Δ mutants (Figure 4.6b). Loss of TEL1 had minimal effect on Cep3 phosphorylation or DSB-induced mobility (Figure 4ac), but a mec1Δ tel1Δ double mutant further reduced mobility after a DSB (Figure 4.6d). Cep3 is unlikely to be a direct target of Mec1, as S575 does not conform to the consensus S/T-Q motif. Cep3-S575 phosphorylation detected by mass spectrometry was also dependent on Rad53 (Chen et al., 2010; Smolka et al., 2007), which I confirmed using my Cep3-pS575 antibody (Figure 4.6a). This result highlights the value and reliability of phosphoproteomic studies in identifying in vivo kinase-specific substrates. Cep3- S575 has similarity to the Rad53 consensus target site (Figure 4.7a) and I observed Cep3 phosphorylation as early as 15 min after DNA damage induction, concomitant with Rad53 activation (Figure 4.7b). In my experimental system, DSB-induced mobility is abolished in rad53Δ cells (Figure 4.7c) and rad53Δ cep3-S575A double mutants (Figure 4.7d). Cep3 is also likely phosphorylated by the Rad53 paralog Dun1 (Figure 4.7a) as I observed a partial reduction of DSB-induced mobility and Cep3 phosphorylation in dun1Δ cells (Figure 4.7ef).

To test whether Rad53 can phosphorylate Cep3-S575 in vitro I purified recombinant MBP-Cep3 and His6-Rad53 from E. coli. Using purified proteins, I observed in vitro phosphorylation of MBP- Cep3 which was dependent on Cep3-S575 and the catalytic activity of Rad53 (Figure 4.8ab). I conclude that the DNA damage-induced phosphorylation of Cep3 on S575 is due to the combined action of the Rad53 and Dun1 kinases downstream of Mec1 and Tel1.

4.2.5 DSB signalling modulates centromere attachment to the spindle pole body

To test whether DSB signalling directly modulates the attachment of centromeres to the SPB, I adapted a system that monitors the integrity of kinetochore clusters by detecting the GFP-tagged kinetochore protein Mtw1 (Mtw1-3xGFP) (Pinsky et al., 2006). As a control I used ndc80-1 cells, which exhibit severe kinetochore detachment phenotypes at the non-permissive temperature (Figure 4.9a). I observed that treatment with Zeocin resulted in a dose-dependent increase in detached kinetochores, as measured by the percentage of cells with more than two Mtw1-3xGFP

78 foci (Figure 4.9ab). This increase in kinetochore detachment was largely abrogated in cep3-S575A cells (Figure 4.9b) suggesting that Cep3 phosphorylation modulates attachment to the SPB.

Figure 4.6. Cep3 phosphorylation is dependent on Mec1 signalling

Figure 4.6. Cep3 phosphorylation is dependent on Mec1 signalling. a, Immunoblot analysis using a phosphospecific Cep3-pS575 antibody on whole cell extracts from cells of the indicated genotype. Cells were treated with Zeocin (250 µg/mL) for 1 h (+), or left untreated (-). Pgk1 antibodies were used to control for loading. Checkpoint activation was confirmed by probing for Rad53. b, c, d, MSD analysis of the MAT locus in mec1Δ (b), tel1Δ (c), and tel1Δ mec1Δ (d) mutants. SML1 was deleted to suppress mec1Δ and rad53Δ lethality.

Figure 4.7. Rad53 and Dun1 are required for Cep3 phosphorylation and DSB-induced chromatin mobility

Figure 4.7. Rad53 and Dun1 are required for Cep3 phosphorylation and DSB-induced chromatin mobility. a, Amino acid sequence surrounding Cep3-S575 and the consensus phosphorylation target sites of the Rad53 and Dun1 kinases. Matching residues are highlighted in red, Ψ represents aliphatic amino acids. b, Time-course of Cep3-13xMyc phosphorylation in response to Zeocin (250 µg/mL) determined by immunoblot analysis on whole cell extracts with a phosphospecific Cep3-pS575 antibody. Myc antibodies were used to control for loading, checkpoint activation was confirmed by probing for Rad53. c, d, e, MSD analysis of the MAT locus in rad53Δ (c), cep3-S575A rad53Δ (d), and dun1Δ (e) mutants. SML1 was deleted to suppress rad53Δ lethality. f, Immunoblot analysis of Cep3-S575 phosphorylation in dun1Δ cells following 1 h of Zeocin treatment (250 µg/mL). Pgk1 antibodies were used to control for loading, Rad53 was probed to confirm checkpoint activation.

Figure 4.8. Rad53 phosphorylates Cep3 in vitro

Figure 4.8. Rad53 phosphorylates Cep3 in vitro. a, In vitro kinase reaction components separated by SDS-PAGE and stained with GelCode Blue. SA denotes Cep3-S575A. *, WT His6- Rad53 contains two degradation products at 75 and 48 kDa. g, Autoradiograph of incorporated 32P from the in vitro reactions shown in f.

As an orthogonal approach, I investigated SPB-CEN dynamics using time-lapse microscopy. I labelled SPBs with Spc42-tdimer2 and placed a LacOx256 array at CEN5 in cells containing an HO cut site on Chr IV. I tracked the positions of CEN5 and the mother SPB in G2/M cells before and after a DSB and calculated the SPB-CEN distance for each frame. While I observed no change in mean SPB-CEN distance (Figure 4.9c), analysis by mean squared distance change (Dorn et al., 2005) revealed a robust increase in SPB-CEN dynamics following DSB induction in wild-type cells that was abolished in cep3-S575A mutants (Figure 4.9d). Further characterization of these dynamics revealed a Cep3-S575-dependent increase in the speed of both antipoleward (AP) and poleward (P) CEN5 movements after a DSB (Figure 4.9e).

These results suggest that Cep3 phosphorylation relieves centromeric constraint by modulating SPB-CEN attachment. In support of this possibility, inactivation of CEN3 by forced transcription increased DSB-induced chromatin mobility of the MAT locus in cep3-S575A cells (Figure 4.10ab). This increase was specific to centromere inactivation of the tracked chromosome as CEN5 inactivation had no effect on mobility (Figure 4.10ac).

Figure 4.9. Cep3 modulates centromere attachment to the SPB

Figure 4.9. Cep3 modulates centromere attachment to the SPB. a, Representative images of Mtw1-3xGFP-labelled kinetochore clusters in wild-type cells that were untreated (control) or treated with Zeocin (250 µg/mL for 90 min), and in ndc80-1 kinetochore mutants. Scale bar represents 2 µm. b, Quantitation of detached kinetochores after Zeocin treatment in wild-type and cep3-S575A mutants as measured by the frequency of G2/M cells with greater than two Mtw1- 3xGFP foci. Data represents the mean ± s.d., n = 3-9 independent experiments. c, Box-and-whisker plot of SPB-CEN distance in G2/M cells before and after a DSB. Whiskers represent the 10-90 percentiles in all panels. d, Mean square SPB-CEN distance change analysis in response to a DSB in wild-type and cep3-S575A cells. e, Box-and-whisker plot of the speed of CEN antipoleward (AP) and poleward (P) movement following a DSB in wild-type and cep3-S575A cells. For all panels: *** p<0.001; one-way ANOVA.

Figure 4.10. Disrupting kinetochore-microtubule attachment in cep3-S575A mutants

Figure 4.10. Disrupting kinetochore-microtubule attachment increase mobility in cep3- S575A mutants. a, Schematic of mobility strains allowing for the conditional inactivation of CEN3 or CEN5. b, c, MSD analysis of the MAT locus on Chr III in cep3-S575A cells with CEN3 (b) or CEN5 (c) inactivation.

4.2.6 The unbroken chromosome arm retains telomeric constraint

I next turned my focus to how a DSB can relieve telomeric constraint and envisioned two possible scenarios: DNA damage signalling could modify telomeric tethering proteins, or physically breaking the DNA fibre could relieve contact to the nuclear periphery. In agreement with the first possibility, DNA damage signalling is known promote the relocalization of Sir3 from telomeres (Martin et al., 1999; McAinsh et al., 1999; Mills et al., 1999). To differentiate between these scenarios, I generated strains with an HO cut site on Chr IV-R and placed a LacOx256 array on either Chr IV-R or the uncut Chr IV-L (Figure 4.11a). As expected, both arrays exhibited similar mobility in the absence of a DSB (Figure 4.11b); however, the mobility of the uncut Chr IV-L arm following a DSB was less than the cut Chr IV-R arm induction indicating the unbroken arm retains constraint (Figure 4.11b). The increased mobility of the unbroken Chr IV-L arm was completely dependent on Cep3-S575 (Figure 4.11c) indicating that both chromosomes arms

84 respond to the modulation of centromeric constraint. Moreover, mobility of the unbroken arm increased upon loss of SIR4 and mirrored the mobility of the cut Chr IV-R arm (Figure 4.11d). These data provide strong evidence that telomeric constraint is only relieved on the broken chromosome arm and suggests that breaking the DNA relieves attachment to the nuclear periphery.

Figure 4.11. The broken chromosome arm is more mobile than the unbroken arm

Figure 4.11. The broken chromosome arm is more mobile than the unbroken arm. a, Schematic of strains to follow the mobility of both chromosome arms following an HO-induced DSB on Chr IV (red triangle). b, MSD analysis of the Chr IV-R and Chr IV-L arms before and after a DSB. c, MSD analysis of the uncut Chr IV-L arm in wild-type and cep3-S575A cells. d, MSD analysis of the cut Chr IV-R arm and the uncut Chr IV-L arm after a DSB in sir4Δ mutants.

To directly test whether DNA damage signalling results in telomeric detachment from the nuclear periphery, I followed a LacOx256 array on the telomeric side of a Chr IV-R HO cut site (Figure 4.12a). I hypothesized that the physical release of telomeres from the periphery should result in a large increase in chromatin mobility, and be epistatic to the additional loss of tethering factors such as Sir4. In contrast to this prediction, I observed only a slight increase in mobility on the telomeric side of the break (Figure 4.12b). Additionally, mobility of the telomeric side was further increased after SIR4 loss (Figure 4.12c), suggesting that Sir4-mediated tethering to the nuclear periphery is not disrupted after a DSB. The mild increase in mobility after a DSB on the telomeric side was not dependent on Mec1 (Figure 4.12b), or Cep3-S575 (Figure 4.12c), indicating that it does not result from DSB signalling or increased SPB-CEN dynamics. Rather, I propose that breaking the DNA relieves centromeric constraint from the acentric fragment, just as a DSB relieves the telomeric constraint from the centric chromosome fragment.

4.2.7 Cep3 regulates the global chromatin response to a DSB

One surprising discovery is that the presence of a DSB in budding yeast can also increase the mobility of undamaged chromosomes (Mine-Hattab and Rothstein, 2012; Seeber et al., 2013). As each chromosome is tethered to the SPB by a microtubule, the modulation of centromeric constraint provides an attractive mechanism to regulate the global (or in trans) mobility response (Figure 4.13a). Intriguingly, initial reports using endonuclease-induced DSBs were conflicted over whether unbroken chromosomes are in fact more mobile (Dion et al., 2012; Mine-Hattab and Rothstein, 2012). As the DSB-to-CEN distance differed in these systems, I considered the possibility that the position of the DSB relative to the centromere might impact mobility. I monitored a LacOx256 array on Chr V in strains containing an HO cut site in trans located 27, 101, 510, or 998 kb from CEN4 (Figure 4.13b). Notably, only the DSB induced 27 kb from CEN4 increased mobility of the unbroken chromosome (Figure 4.13c). Similar to the mobility of the unbroken chromosome arm, the increase in mobility of the unbroken chromosome Chr V was less than that of the MAT locus on broken Chr III (Figure 4.13d). Mobility of the unbroken chromosome was completely dependent on Cep3-S575 (Figure 4.14a) indicating that this mobility response is due to the modulation of kinetochore-microtubule attachment.

Figure 4.12. Telomeres are not detached following a DSB

Figure 4.12. Telomeres are not detached following a DSB. a, Schematic of a strain to follow the mobility the telomeric side of the broken Chr IV. b, c, d, e, MSD analysis of the Chr IV-RTEL array before and after a DSB in wild-type (b), mec1Δ (c), cep3-S575A (d), or sir4Δ (e) cells.

As unbroken chromosomes should retain their telomeric attachment to the nuclear periphery (Figure 4.13a), I asked whether removal of Sir4 could further increase mobility and account for the difference in mobility between broken and unbroken chromosomes. I found that SIR4 deletion increased the DSB-induced mobility of an unbroken chromosome (Figure 4.14b) to levels that

87 mirrored its broken counterpart (Figure 4.14c). This behaviour is in contrast to the mobility of broken chromosomes, which is not increased in sir4Δ mutants following a DSB (Figure 4.15a). The insensitivity of broken chromosomes to loss of telomeric tethering is not specific to sir4Δ as both yku70Δ and yku70Δ sir4Δ mutants exhibited minimally increased chromatin mobility after a DSB despite having pronounced effects on mobility in the absence of damage (Figure 4.15bc).

Figure 4.13. The global chromatin response requires a centromere-proximal DSB

Figure 4.13. The global chromatin response requires a centromere-proximal DSB. a, A proposed model for how a DSB could affect the mobility of all chromosomes. b, Schematic of strains used to follow the mobility of an unbroken chromosome Chr V at the MAK10 locus (Chr V:53601) in trans to a DSB on Chr IV. HO cut sites on Chr IV (red triangles) are located at various distances relative to the centromere. c, MSD analysis of the MAK10 locus on unbroken Chr V in the strains outlined in b. d, MSD analysis of the MAT locus on Chr III (in cis) and the MAK10 locus on unbroken Chr V in trans to a DSB 27kb from CEN4.

Figure 4.14. Global mobility requires Cep3-S575 but retains telomeric based constraint

Figure 4.14 The global mobility response requires Cep3-S575 but retains telomeric based constraint. a, b, MSD analysis of the MAK10 locus on unbroken Chr V in trans to a DSB 27 kb from CEN4 in cep3-S575A (a) and sir4Δ (b) cells. c, MSD analysis of the MAT locus, in cis to a DSB on Chr III, and the MAK10 locus on unbroken Chr V in trans to a DSB 27 kb from CEN4 in a sir4Δ background.

In parallel experiments I assessed whether a centromere-proximal DSB could promote the mobility of a CEN/ARS-LacOx256 plasmid (Figure 4.16a). While the mobility of the episome was highly constrained as previously observed (Marshall et al., 1997), its mobility also increased after a DSB (Figure 4.16b), and in a manner that was dependent on Cep3-S575 (Figure 4.16c). The increased

89 mobility of the episome was also dependent on the DSB-to-CEN distance as a DSB 998 kb from CEN4 had no effect (Figure 4.16d).

Figure 4.15. Loss of telomeric tethering does not increase mobility of a broken chromosome

Figure 4.15. Loss of telomeric tethering factors does not increase mobility of a broken chromosome. a, b, c, MSD analysis of the MAT locus, in cis to a DSB, in sir4Δ (a), yku70Δ (b), and yku70Δ sir4Δ (c) cells.

From these results I conclude that Cep3 phosphorylation after DNA damage promotes chromatin mobility by relieving constraint from all centromere containing DNA providing a compelling mechanism for the global chromatin mobility response.

Figure 4.16. The mobility of an episome is increased after DNA damage

Figure 4.16. The mobility of an episome is increased after DNA damage. a, Schematic of strains used to follow the mobility of a CEN/ARS-LacOx256 in trans to a DSB on Chr IV at 27 kb or 998 kb from CEN4. b, c, MSD analysis of a CEN/ARS-LacOx256 episome in wild-type (b) and cep3-S575A (c) cells in response to a DSB 27 kb from CEN4. d, MSD analysis of a CEN/ARS- LacOx256 episome in wild-type cells response to a DSB 998 kb from CEN4.

4.2.8 The increase in mobility following a DSB is dependent on centromeric constraint

Centromeric constraint varies along the length of the chromosome and is highest at centromere proximal loci (Verdaasdonk et al., 2013). My model therefore predicts that loci close to the centromere should have a larger increase in DSB-induced mobility compared to loci far from the centromere. To test this prediction, I generated strains with LacOx256 arrays 89, 497, and 985 kb from CEN4 and assessed chromatin mobility before and after a DSB induced 12 kb from each array (Figure 4.17a). As expected, proximity to the centromere dictated mobility in the absence of a DSB (Figure 4.17b). Strikingly, all three positions responded differently to a DSB (Figure 4.17c), but ended up with similar mobility (Figure 4.17d): the array 89 kb from CEN4 strongly increased its mobility, similar to the MAT locus (79 kb from CEN3), the array 497 kb from CEN4 exhibited a limited response, and the array 985 kb from CEN4 had no detectable increase in mobility. These data are consistent with a model in which the primary mechanism of DSB-induced chromatin mobility is through the relief of centromeric constraint.

4.2.9 DSB-induced chromatin mobility can be dispensable for repair

An attractive role for increased chromatin mobility after DNA breakage is to stimulate homology search during HR; however, factors know to affect DSB-induced mobility, such as Rad9 or INO80, have additional roles in DSB repair (Jeggo and Downs, 2014). Therefore, whether increased chromatin mobility promotes DSB repair remains an open question. As Cep3 is a kinetochore component not directly associated with DNA repair, I used the cep3-S575A mutation as a tool to probe the function of chromatin mobility in genome maintenance. I first treated cep3-S575A cells with a variety of genotoxic agents; unexpectedly, I failed to detect any sensitivity (Figure 4.18a). As a control, I used an arp8Δ mutant, which is also defective in DSB-induced chromatin mobility (Seeber et al., 2013) and exhibits a similar mobility phenotype to cep3-575A mutants (Figure 4.1e, Figure 4.3a). I next assessed the efficiency of DSB repair by HR using strains in which two homologous cassettes are located at different positions in the genome to explore various spatial configurations for repair (Figure 4.18b) (Agmon et al., 2013). An HO cut site at one cassette can be repaired using the second cassette, the donor, as template for HR repair.

Figure 4.17. Centromeric constraint dictates the increase in mobility after a DSB

Figure 4.17. Centromeric constraint dictates the increase in mobility after a DSB. a, Schematic of strains to observe DSB-induced mobility at three loci on Chr IV at different positions relative to the centromere. Each strain features an HO cut site (red triangle) 12 kb from the LacOx256 array. b, MSD analysis of the indicated loci in the absence of a DSB. c, MSD analysis of the Chr IV loci before and after a DSB. d, MSD analysis of the Chr IV loci following DSB induction.

Figure 4.18. Increased chromatin mobility is dispensable for DSB repair

Figure 4.18. Increased chromatin mobility is dispensable for DSB repair. a, Spot assays to assess DNA damage sensitivity. 5-fold serial dilutions of yeast cultures were grown on XY (control) plates or plates containing methyl methanesulfonate (MMS), hydroxyurea (HU), Zeocin, or camptothecin (CPT). b, Schematic of HR repair strains indicating the relative location of each homologous cassette and the HO cut site (red). c, Repair efficiency of HR strains in the indicated genetic backgrounds. Data represents the mean ± s.e.m., n = 4-10 independent experiments. d, Schematic of the HR repair strain NA60 and primers P1 and P2 used to amplify the cut site on Chr IX. Repair of the DSB by HR is accompanied by the appearance of a ClaI site provided by the donor recombination cassette on Chr XIII. e, Agarose gel of ClaI-digested PCR products in response to a DSB in wild-type and cep3-S575A cells. DNA was extracted from cells harvested at the indicated time points, the cut locus was amplified by PCR and digested with ClaI. Gene conversion (GC) products are cleaved by ClaI and form two lower molecular weight bands. A colony that survived plating on galactose was used as a positive control (+).

I found that in contrast to arp8Δ cells, which displayed reduced HR efficiency in nearly all tested strains, the cep3-S575A mutation did not significantly affect repair in the eight tested spatial configurations (Figure 4.18c). Following repair kinetics by monitoring the transfer of a unique ClaI endonuclease site from the donor to the cut locus also failed to detect a delay in cep3-S575A cells (Figure 4.18de). Together, these data challenge the notion that increased mobility following chromosome breakage promotes homology search and I therefore sought an alternative explanation for this phenomenon.

4.2.10 Cep3 phosphorylation promotes checkpoint arrest

A key insight into the potential role of Cep3 phosphorylation came by taking a step back from chromatin mobility and I considered the cellular consequences of relieving centromeric constraint. As unattached kinetochores trigger the spindle assembly checkpoint (SAC) leading to mitotic arrest (Rieder et al., 1995), I hypothesized that Cep3 phosphorylation might engage the SAC. DSBs in budding yeast induce G2/M arrest through parallel Chk1 and Rad53 pathways (Sanchez et al., 1999; Wang et al., 2001); however, the SAC has also been implicated, acting in part through an unidentified centromeric target (Dotiwala et al., 2010). I measured the duration of arrest following an irreparable DSB and observed a decrease in checkpoint duration in cep3-S575A cells compared to wild-type (Figure 4.19a), which was further reduced in cep3-S575A chk1Δ mutants indicating that Cep3 functions in parallel to Chk1. The loss of CHK1 had minimal effect on DSB- induced chromatin mobility (Figure 4.19b, left panel), indicating that defective checkpoint arrest does not impair chromatin motion.

I next asked if Cep3 phosphorylation promotes cell cycle arrest through engagement of the SAC, which is dependent on Mad2 (Musacchio and Salmon, 2007). The effect of MAD2 deletion on checkpoint duration was epistatic to cep3-S575A (Figure 4.19a), indicating that Cep3 and Mad2 function in the same pathway with respect to the DNA damage checkpoint. Furthermore, the severe checkpoint arrest defect of chk1Δ mad2Δ cells was also insensitive to cep3-S575A mutation (Figure 4.19a). Chromatin mobility was unaffected by MAD2 loss (Figure 4.19b, right panel), suggesting that engagement of the SAC is not required for DSB-induced mobility, and rather that it might be a consequence of relieving the centromeric constraint.

Figure 4.19. Cep3 phosphorylation promotes cell cycle arrest and genome stability

Figure 4.19. Cep3 phosphorylation promotes cell cycle arrest and genome stability. a, Length of checkpoint arrest in response to an irreparable DSB at the MAT locus. Each dot represents a single cell from 3 independent experiments (116-199 total cells per genotype); red bars represent the median arrest length. b, MSD analysis of the MAT locus in chk1Δ and mad2Δ cells. c, a-like faker (ALF) assay as measured by the number of surviving HIS+ colonies per patch. Data represents the mean ± s.e.m. from 24-60 patches in 3 independent experiments. d, Chromosome transmission fidelity (CTF) assay as measured by the frequency of half-sectored colonies. Data represents the mean ± s.e.m., n = 3-8 independent experiments. For all panels: *** p<0.001, ** p<0.01, *p<0.05; one-way ANOVA.

Finally, as defects in checkpoint arrest can lead to precocious chromosome segregation and genomic instability, I tested whether cep3-S575A cells showed evidence of such events in two independent assays: the a-like faker (ALF) assay, which detects spontaneous loss of the MAT locus in MATα cells by mating test (Warren et al., 2004; Yuen et al., 2007), and the chromosome transmission fidelity (CTF) assay (Yuen et al., 2007) that monitors loss of an artificial chromosome fragment. Both assays revealed that the cep3-S575A mutation leads to increased chromosome

96 instability and that this increase is exacerbated in cep3-S575A chk1Δ double mutants (Figure 4.19cd).

4.3 Discussion

Here I report that the Mec1- and Rad53-dependent phosphorylation of Cep3 is required for increased chromatin mobility after DNA breaks. My results suggest that DSB-induced mobility results from the modulation of two physical constraints: Cep3 phosphorylation modulates SPB- kinetochore attachment, while breaking the DNA relieves telomeric constraint to the nuclear periphery (Figure 4.20).

20. Proposed model of how a DSB increases chromatin mobility

Figure 4.20. Proposed model of how a DSB increases chromatin mobility. Cep3 is phosphorylated after a DSB in a Mec1/Rad53/Dun1-dependent manner and increases chromatin mobility by relieving the centromeric constraint imposed on chromosomes. A DSB also increases the mobility of the cut chromosome arm by relieving a telomeric-based constraint. Cep3 phosphorylation promotes cell cycle arrest through a Mad2-dependent mechanism.

Antagonizing centromeric attachment by Cep3 phosphorylation provides an attractive mechanism for the mobility of unbroken chromosomes as all centromeres are tethered by a microtubule and physically clustered by the spindle pole body. An intriguing observation concerns the DSB-to- CEN distance and the ability to stimulate chromosome motion and my discovery that in trans

97 mobility requires a centromere proximal DSB explains a previous discrepancy in the literature (Dion et al., 2012; Mine-Hattab and Rothstein, 2012), and the reported dose-dependent effect of Zeocin (Seeber et al., 2013), as increasing the number of DSBs will increase the probability of a single DSB occurring near the centromere.

The requirement for a centromere-proximal break may reflect the ability of Mec1/Rad53 to reach CEN-bound Cep3 and is reminiscent of Rad51 and phosphorylated histone H2A (γH2A) ChIP signals that spread along broken chromosomes and can jump to centromeres of unbroken chromosomes when centromere-proximal DSBs are induced (Renkawitz et al., 2013). As the spreading of Rad51 and γH2A to other chromosomes is indicative of homology search, the phosphorylation of CEN-bound Cep3 may require homology search, rather than promote it. In support of this possibility, Rad51 is necessary for DSB-induced mobility (Mine-Hattab and Rothstein, 2012). ChIP experiments with my Cep3-S575 phosphospecific antibody would help elucidate the kinetics of Cep3 phosphorylation at the centromere.

The ability for cep3-S575A cells to undergo efficient repair by HR suggests that increased chromatin mobility may be dispensable for homology search. Importantly, I do not conclude that mobility per se is not required for interchromosomal HR; rather I propose that increased mobility after DNA breakage is not necessary. Although this conclusion opposes a popular model in the field (Dion and Gasser, 2013), this possibility is regardless consistent with data indicating that homology search is concentrated locally on broken chromosomes (Renkawitz et al., 2013) and that HR is greatly influenced by the pre-existing physical proximity of loci (Agmon et al., 2013; Lee et al., 2016). Furthermore, the recent identification that the DSB-mobility defective sae2Δ and htz1Δ mutants are also fully competent for repair HR provides support for my argument against the homology search model (Lee et al., 2016).

Finally, my results indicate that a function for Cep3 phosphorylation is to stimulate checkpoint arrest via the Mad2-dependent SAC. This discovery builds on previous genetic results which suggested that DNA damage can engage the SAC (Dotiwala et al., 2010; Garber and Rine, 2002; Maringele and Lydall, 2002) and provides a key substrate in this response. Interestingly, the SAC response can be modulated depending on the number of unattached chromosomes (Dick and Gerlich, 2013) and the amount of Mad2 (Collin et al., 2013); cells may therefore be able to fine- tune engagement of the SAC as a part of the DNA damage checkpoint. Rad53 phosphorylation has

98 also been implicated in the SAC (Clemenson and Marsolier-Kergoat, 2006) suggesting that crosstalk between pathways functions in both directions. My identification of Cep3 as a candidate link between the spindle assembly and DNA damage checkpoints further suggests that cooperation between these checkpoint systems is a powerful means to promote genome stability in eukaryotes.

Chapter 5 Pursuing the molecular events downstream of Cep3 phosphorylation

Statement of contributions: I performed all of the experiments described in this chapter.

Some of the text and figures presented in this chapter were modified with permission from the following publication:

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5.1 Introduction

The identification of Cep3-S575 provides new insight into how a DSB modifies the constraints on chromosomes and the function of this phenomenon; however, several key questions remain. First, the mechanism by which Cep3 phosphorylation antagonizes microtubule constraint is yet to be determined. Although the CBF3 complex has been implicated with spindle stability (Bouck and Bloom, 2005) and septin dynamics (Gillis et al., 2005), my observation that centromere inactivation can restore mobility in cep3-S575A mutants argues for a centromeric function.

Second, while the phosphorylation of Cep-S575 can explain the requirement of Mec1 and Rad53 in DSB-induced mobility, additional factors have also been implicated in this process; namely the INO80 and SWR1 chromatin remodelers, and the histone variant Htz1. It is currently unclear whether these mutants are connected to Cep3 or if they are involved in separate processes. INO80 is recruited to DSB sites (Downs et al., 2004; Morrison et al., 2004; van Attikum et al., 2004) and is proposed to promote chromatin mobility by increasing the flexibility of the DNA fibre (Neumann et al., 2012; Seeber et al., 2013). Although my results do not exclude a role for INO80 at DSBs, the ability to mimic DSB-induced mobility through the disruption of centromeric and telomeric constraints makes these results difficult to reconcile. Further insight into this biological phenomenon is therefore required to understand the contributions of these factor and I next sought to elucidate the molecular mechanism downstream of Cep3 phosphorylation.

5.2 Results

5.2.1 The Rpd3L complex is required for DSB-induced chromatin mobility

The crystal structure of Cep3 places S575 in an unstructured region adjacent to the zinc cluster DNA binding domain (Figure 5.1a), hinting that phosphorylation might modulate DNA binding (Purvis and Singleton, 2008); however, my ChIP experiments revealed no difference in centromeric enrichment following a DSB (Figure 5.1b). Cep3 shares homology with Gal4 transcription factors and the S575 residues is located in an acidic activation domain (Figure 5.2a) which often recruit transcriptional co-regulators in Gal4 family proteins including histone

101 acetyltransferase and deacetyltransferase (HDAC) enzymes. I serendipitously discovered that the HDAC inhibitor valproic acid (Figure 5.2b), but not nicotinamide (Figure 5.2c), strongly affected DSB-induced chromatin mobility and confirmed that valproic acid treatment at this concentration was not toxic to cells (Figure 5.2d). HDAC inhibition has been reported to inhibit Mec1- dependent DNA damage signalling (Robert et al., 2011); however, I observed minimal impact on Rad53 and Cep3-S575 phosphorylation following valproic acid treatment at five times the concentration used in mobility experiments (Figure 5.2e). Together these results suggest that an HDAC inhibited by valproic acid might function downstream of Cep3.

Figure 5.1. Cep3-S575A does not affect chromatin enrichment

Figure 5.1. Cep3-S575A does not affect chromatin enrichment. a, Crystal structure of Cep3 (Purvis and Singleton, 2008) with added unstructured acidic activation domain residues (blue) and the zinc cluster (ZC) (PDB ID 2VEQ). The predicted position of S575 is highlighted in red. b, Chromatin immunoprecipitation (ChIP) of Cep3-13xMyc and Cep3-S575A-13xMyc in response to a DSB at the MAT locus. Fold enrichment at CEN3 is normalized to the non-centromeric control locus TSC11. Data represents the mean ± s.d., n = 5 independent experiments

Valproic acid inhibits Class I and II HDACs which includes Hda1, Hos1, Hos2, Hos3, and Rpd3 in budding yeast. I generated deletion strains for each factor and identified RPD3 as the sole component required for DSB-induced mobility (Figure 5.3ab).

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Figure 5.2. Valproic acid is a potent inhibitor of DSB-induced chromatin mobility

Figure 5.2. Valproic acid is a potent inhibitor of DSB-induced chromatin mobility. a, Schematic of Cep3 and Gal4 domain architecture revealing conserved N-terminal DNA binding domains and C-terminal acidic activation domains. The location of Cep3-S575 is highlighted in red. b, MSD analysis of the MAT locus in cells treated with 5 mM nicotinamide (NAM) (b), or 2 mM valproic acid (VPA) (c). d, Spot assays to assess sensitivity to VPA. 5-fold serial dilutions of yeast cultures were grown on XY (control) plates or plates containing 2 mM VPA. e, Immunoblot analysis using a phosphospecific Cep3-pS575 antibody on whole cell extracts. Cells were treated with a high 10 mM dose of VPA for 2 h before Zeocin was added for 1 h (250 µg/mL). Pgk1 antibodies were used to control for loading, checkpoint activation was confirmed by probing for Rad53.

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Deletion of the Class III HDACs SIR2 and HST4 also had no affect on chromatin mobility further indicating the specificity of this phenotype (Figure 5.3a). Consistent with my results, loss of the Rpd3 cofactor SIN3 mirrored the mobility defect of rpd3Δ cells (Figure 5.4c). Importantly, I still observed Cep3-S575 phosphorylation in rpd3Δ and sin3Δ mutants following an HO-induced DSB (Figure 5.4d), suggesting that the loss of Rpd3 is unlikely to impair DSB-induced mobility simply by inhibiting DNA damage signalling.

Rdp3 functions in two separate multisubunit complexes, the small (Rpd3S) and the large (Rpd3L) complex, both of which require Sin3 (Yang and Seto, 2008). I identified Rpd3L as the responsible complex as increased mobility requires SDS3, but not RCO1 (Figure 5.4ab) (Carrozza et al., 2005a; Carrozza et al., 2005b). Intriguingly, high throughput yeast two-hybrid screens identified an interaction between Cep3 and the Rxt3 subunit of the Rpd3L complex (Wong et al., 2007); however, in my hands this interaction does not require Cep3-S575 or DNA damage (Figure 5.4c). In an second approach, I tagged endogenous Rxt3 with a 13xMyc tag and made cellular extracts for in vitro peptide pulldowns. Precipitation of biotinylated Cep3 peptides encompassing S575 revealed an interaction with Rxt3-13xMyc and which required phosphorylated Cep3-S575 and the correct sequence of surrounding amino acids as a scrambled peptide containing phosphoserine in the same position did not pulldown Rxt3-13xMyc (Figure 5.4d). Consistent with the possibility that Rxt3 might interact with phosphorylated Cep3 in vivo, deletion of RXT3 also impaired increased chromatin mobility after a DSB (Figure 5.4e).

5.2.2 Mimicking Htz1 acetylation abolishes DSB-induced chromatin mobility

The identification of Rpd3L as a putative downstream effector of Cep3-pS575 raises the question of what Rpd3L deacetylates to promote increased chromatin mobility. Several acetylation marks have been implicated in centromere function in S. cerevisiae including histone H3-K56 (Millar and Grunstein, 2006) and histone H4-K16 (Choy et al., 2011), however, my search was aided by the identification of two additional mobility mutants: SWR1 and Htz1 (Horigome et al., 2014). The SWR1 chromatin remodeler promotes the exchange of histone H2A for the histone variant H2A.Z (Krogan et al., 2003), encoded by HTZ1 in budding yeast, and the loss of either SWR1 or Htz1 impairs DSB-induced chromatin mobility (Horigome et al., 2014).

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Figure 5.3. Rpd3 is required for DSB-induced chromatin mobility

Figure 5.3. Rpd3 is required for DSB-induced chromatin mobility. a, b, c, MSD analysis of the MAT locus in cells deleted for the indicated HDACs. d, Immunoblot analysis using a phosphospecific Cep3-pS575 antibody on whole cell extracts in HDAC mutants following an HO- induced DSB. Pgk1 was probed to control for loading, checkpoint activation was confirmed by probing for Rad53.

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Figure 5.4. The Rpd3L complex is a putative Cep3-pS575 interactor

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Figure 5.4. The Rpd3L complex is a putative Cep3-pS575 interactor. a, b, MSD analysis of the MAT locus in rco1Δ (a), and sds3Δ (b) mutants. c, Yeast two-hybrid assay in cells expressing pGBK-Cep3 wild-type or S575A bait and control plasmids or pGAD-Rxt3 as prey. 5-fold serial dilutions of yeast cultures were grown on SD-leu-trp plates or SD-leu-trp-his plates to test for expression of the HIS3 reporter. Zeocin (ZEO) was added at 25 µg/mL to induce DNA damage where indicated, the HIS3 competitive inhibitor 3-Amino-1,2,4-triazole (3-AT) was added at 30 mM to increase interaction specificity of the system. d, Peptide pulldowns with biotinylated Cep3 peptides containing unphosphorylated S575 (non-phospho), phosphorylated S575 (phospho), or a peptide containing phosphorylated S575 at the same position with the remaining residues randomly shuffled (scrambled-phospho). Peptides were incubated with cellular extract from cells expressing Rxt3-13xMyc and precipitated with Streptavidin beads. e, MSD analysis of the MAT locus in rxt3Δ cells.

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Htz1 is found throughout the genome but is also enriched in pericentric chromatin (Millar et al., 2006). Furthermore, Htz1 is acetylated on four N-terminal lysines (K3, K8, K10, and K14) (Krogan et al., 2004), and I hypothesized that these residues might be targeted for deacetylation by Rpd3L. My model predicts that Htz1 deacetylation by Rpd3L should promote increased chromatin mobility; therefore, mimicking constitutive acetylation via glutamine substitution should impair this response. I first mutated all four N-terminal lysines to generate the htz1-4KQ (K3Q/K8Q/K10Q/K14Q) mutant and assessed chromatin mobility before and after a DSB. Similar to cep3-S575A and rpd3Δ cells, mobility in htz1-4KQ mutants was unaffected in the absence of a DSB but did not increase following chromosome breakage (Figure 5.5a). This phenotype was recapitulated in htz1-K14Q cells (Figure 5.5b) suggesting that K14 plays a particularly important role in this process.

Consistent with the possibility that Htz1 is involved in this process, Htz1 is known to be deacetylated after DNA damage in a Mec1-dependent manner, and that deacetylation is reduced, but not abolished, in hda1Δ mutants suggesting a second HDAC is also involved (Bandyopadhyay et al., 2010). I observed that Htz1-K14 deacetylation after DNA damage was completely abolished in hda1Δ rpd3Δ double mutants indicating a role for Rdp3 in this process (Figure 5.5c). The requirement for Cep3-S575 in Htz1 deacetylation is less clear as measured by immunoblot (Figure 5.5c). As the involvement of Cep3 is likely limited to centromeric chromatin comprising only a small percent of the global Htz1 pool (Millar et al., 2006), future ChIP experiments will be required to test the genetic dependence of this phenotype. If Htz1 deacetylation lies downstream of Cep3 phosphorylation, then a critical prediction is that htz1-K14Q cells should mirror other phenotypes observed in cep3-S575A cells. In agreement with this idea, the length of checkpoint arrest in response to an irreparable DSB was also decreased in htz1-K14Q cells, mirroring the phenotype of cep3-S575A mutants (Figure 5.5d).

5.2.3 Increasing chromatin mobility in the absence of DNA damage

The requirement of Cep3-S575 phosphorylation in DSB-induced mobility raised the possibility that mimicking phosphorylation might increase mobility in the absence of DNA damage. Phosphomimetic cep3-S575E cells were viable but unable to increase mobility in the absence of DNA damage (Figure 5.6a). Additionally, cep3-S575E mutants had a mild mobility defect

108 following chromosome breakage suggesting that the S575E mutation is not fully functional as a phosphomimetic residue. Figure 5.5. Mimicking Htz1 acetylation impairs DSB-induced mobility and checkpoint arrest

Figure 5.5. Mimicking Htz1 acetylation impairs DSB-induced mobility and checkpoint arrest. a, b, MSD analysis of the MAT locus in htz1-4KQ (a) and htz1-K14Q (b) mutants. c, Immunoblot analysis using an Htz1-K14 acetylation specific antibody and a phosphospecific Cep3-S575 antibody on whole cell extracts from cells of the indicated genotype treated with 250 µg/mL Zeocin. Pgk1 was probed to control for loading, checkpoint activation was confirmed by probing for Rad53. d, Length of checkpoint arrest in response to an irreparable DSB at the MAT locus. Each dot represents a single cell (72-199 total cells per genotype), red bars represent the median arrest length.

Htz1 provides a second avenue to control chromatin mobility and I reasoned that mimicking Htz1 deacetylation with arginine substitutions might increase mobility. The htz1-K14R mutation had a

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mild phenotype (Figure 5.6b), but mutation of all four N-terminal lysines in the htz1-4KR (K3R/K8R/K10R/K14R) mutant was sufficient to increase chromatin mobility in the absence of a DSB (Figure 5.6c), similar to centromere inactivation (Figure 4.2b). Moreover, the htz1-4KR mutation did not increase mobility after a DSB (Figure 5.6c), consistent with the idea that Htz1 deacetylation is included in the DSB-induced mobility response.

My previous work identified that the simultaneous disruption of centromeric and telomeric tethering could mimic the behavior of broken chromosomes. In an attempt to recapitulate this result, I deleted SIR4 in htz1-4KR cells and strikingly observed mobility mirroring the wild-type response to a DSB (Figure 5.6d). A prevailing idea in the field is that the recruitment of chromatin remodelers drives mobility after a DSB which I reasoned should still impact htz1-4KR sir4Δ mutants. If, however, DSB-induced mobility results solely from the modulation of chromosomal constraints then a DSB should have no effect. In agreement with the second outcome, a DSB did not increase chromatin mobility in htz1-4KR sir4Δ cells (Figure 5.6d). Finally, I asked whether mimicking Htz1 deacetylation could bypass the requirement for Cep3 phosphorylation in DSB- induced mobility. I observed that the htz1-4KR mutation could suppress the cep3-575A mobility defect indicating that these components function in the same genetic pathway (Figure 5.6e).

5.2.4 A centromeric function for chromatin remodelers in DSB- induced mobility

The SWR1 and INO80 remodelers are thought to promote chromatin mobility by increasing flexibility of the DNA fibre; however, my data argues that the manipulation of physical constraints on chromosomes is sufficient to explain their behavior. While these observations do not strictly exclude a role for remodelling at DSBs, they leave little room for the functional significance of such activities and I therefore sought alternative explanations for their involvement in chromosome movement. The pivotal role of SWR1 for the incorporation of Htz1 provides a simple explanation for its requirement in DSB-induced mobility; however, the role of INO80 in this process is equivocal.

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Figure 5.6. htz1-4KR promotes chromatin mobility in the absence of a DNA break

Figure 5.6. Mimicking Htz1 deacetylation promotes chromatin mobility in the absence of a DNA break. a-e, MSD analysis of the MAT locus in cep3-S575E (a), htz1-K14R (b), htz1-4KR (K3R, K8R, K10R, K14R) (c), htz1-4KR sir4Δ (d), and cep3-S575A htz1-4KR (e) mutants.

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The INO80 complex can bind and activate Rad53 (Kapoor et al., 2015) raising the possibility that INO80 mutants might have DSB-induced mobility defects due to compromised DNA damage signalling. This, however, seems unlikely to fully explain the phenotype as I observed only a minor effect on Cep3 phosphorylation in the arp5Δ and arp8Δ INO80 mutants compared to mec1Δ cells (Figure 5.7a, Figure, 4.6a), but a stronger mobility defect (Figure 4.1e, Figure 4.6b).

Interestingly, INO80 is required for proper centromere structure (Chambers et al., 2012) and subunits of the complex, including Arp4 and Ino80, bind directly to centromeres (Ogiwara et al., 2007). These observations raised the possibility that INO80 mutants might impair SPB-CEN dynamics downstream of Cep3 phosphorylation. Consistent with this idea, ARP8 loss also decreased checkpoint duration in response to an irreparable DSB, and in a manner that was epistatic to cep3-S575A (Figure 5.7b), suggesting that INO80 and Cep3 act in a common pathway. To determine if INO80 mutants are unable to modulate SPB-CEN attachment, I generated strains containing conditional centromeres in arp8Δ cells. I observed that CEN3 inactivation was sufficient to increase DSB-induced mobility in arp8Δ cells (Figure 5.7cd) indicating that arp8Δ mutants fail to relieve centromeric constraint following a DSB.

5.2.5 Aurora B is required for DSB-induced mobility

Despite the identification of Rpd3L and Htz1, it remains unclear how these events might modulate kinetochore attachment, and likely involves additional effectors. An attractive candidate is the Aurora B kinase which generates unattached kinetochores in response to erroneous microtubule attachment (Carmena et al., 2012; Pinsky et al., 2006). To approach the problem from another angle, I asked whether Aurora B is required for the increased chromatin mobility observed after a DSB.

Aurora B functions in the conserved chromosome passenger complex (CPC) and is encoded by the essential IPL1 gene in budding yeast. To control Ipl1 levels I integrated the F-box protein TIR1 from Oryza sativa (OsTIR1) and a C-terminal auxin-inducible degron tag (3xAID) into the IPL1 gene to allow targeted protein degradation (Kubota et al., 2013; Nishimura et al., 2009). Addition of the auxin indole-3-acetic acid (IAA) inhibited cell growth in these cells (Figure 5.8a) suggesting that the system can functionally deplete Ipl1-3xAID. While the expression of OsTIR1 alone had no effect on chromatin mobility (Figure 5.8b), cells expressing Ipl1-3xAID were unable

112 to increase chromatin mobility after a DSB following auxin treatment (Figure 5.8c) indicating that the Ipl1 protein is required for this phenomenon. The Ipl1-3xAID fusion is likely hypomorphic as I observed a slight decrease in DSB-induced mobility with drug solvent alone (Figure 5.8c).

Figure 5.7. The arp8Δ mutant fails to modulate SPB-CEN attachment after a DSB

Figure 5.7. The arp8Δ mutant fails to modulate SPB-CEN attachment after a DSB. a, Immunoblot analysis of Cep3-S575 phosphorylation in two separate clones each of the INO80 mutants arp5Δ and arp8Δ. Cells were treated with Zeocin (250 µg/mL) for 1 h (+), or left untreated (-). Pgk1 was probed to control for loading, checkpoint activation was confirmed by probing for Rad53. b, Length of checkpoint arrest in response to an irreparable DSB at the MAT locus. Each dot represents a single cell from at least two independent experiments (171-199 total cells per genotype); red bars represent the median arrest length. c, d, MSD analysis of the MAT locus in INO80 mutant arp8Δ cells with CEN3 (c) or CEN5 (d) inactivation.

In a second approach to regulate Ipl1 activity, I introduced the M181G and T244G mutations to generate ATP analogue sensitive ipl1-as6 cells (Kung et al., 2005) and treatment with the ATP

113 analogue PP1 Analog II (1NMPP1) was able to effectively suppress cell growth (Figure 5.8d). In agreement with my auxin-inducible degron results, short-term 1NMPP1 treatment decreased chromatin mobility following a DSB compared to the DMSO control (Figure 5.8e). Notably, this phenotype was observed by adding 1NMPP1 only thirty minutes prior to imaging, two and a half hours after DSB induction, suggesting that continual Ipl1 activity is required for DSB-induced mobility. This result is consistent with the notion that the modulation of SPB-CEN attachment following a DSB is transient in nature and requires constant signalling to be maintained.

5.3 Discussion

The experiments presented here hint at molecular steps which promote increased chromatin mobility after DNA breakage; however, diligent work will be required to substantiate these findings. While my genetic data suggest that the deacetylation of Htz1 by the Rpd3L complex lies downstream of Cep3 phosphorylation, additional work will be required to firmly place them in this pathway. First, the hypothesis that Rpd3L interacts with phosphorylated Cep3 at the centromere can be directly tested by ChIP experiments. As Rpd3 does not bind chromatin directly, this approach would likely require careful optimization of multiple crosslinking agents (Kurdistani et al., 2002). My ability to generate phosphorylated MBP-Cep3 also provides a tool to investigate phosphorylation-dependent protein interactions and the purification of recombinant Rxt3 could test for a direct interaction in vitro. As Rxt3 contains no known domains, phosphoprotein binding or otherwise, the interaction identified by peptide pulldown from cellular extracts may be indirect.

Although I observed Cep3 phosphorylation in rpd3Δ mutants, known connections between HDACs and DNA damage signalling demand cautious interpretation for a direct role of Rpd3 in DSB-induced mobility. Of note, valproic acid has been reported to inhibit Mec1 signalling (Robert et al., 2011), Rpd3 can deacetylate Rad53 to prevent overactivation (Tao et al., 2013), and loss of Rpd3 can restore the DNA damage checkpoint in RAD9 and MEC1 mutants in response to UV damage (Scott and Plon, 2003). In this regard, the identification of Htz1 mutations affecting DSB- induced mobility was a key discovery and provides specific tools to investigate this pathway. Importantly, mimicking constitutive Htz1 acetylation and deacetylation can both impair and promote chromatin mobility respectively, strongly arguing for a direct role of Htz1 modification in this process. The ability of htz1-4KR cells to suppress the cep3-S575A mobility defect provides

114 strong evidence that Htz1 and Cep3 function in a common pathway; however, several aspects of this model remain to be tested.

Figure 5.8. Ipl1 promotes DSB-induced chromatin mobility

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Figure 5.8. Ipl1 promotes DSB-induced chromatin mobility. a, Spot assays to test an auxin degron system of the essential IPL1 gene. 5-fold serial dilutions of yeast cultures were grown on XY (control) plates or plates containing 0.5 mM indole-3-acetic acid (IAA) to induce Ipl1-3xAID degradation. b, c, MSD analysis of the MAT locus in cells expressing OsTIR1 alone (b), or in combination with Ipl1-3xAID (c). IAA or ethanol as a control was added 1 hour prior to imaging, 2 hours after DSB induction. d, Spot assays to test the ATP analogue sensitive ipl1-as6 allele. 5- fold serial dilutions of cultures were grown on XY plates containing DMSO or 10 µM ATP analogue PP1 Analog II (1NMPP1). e, MSD analysis of the MAT locus in ipl1-as6 cells. 1NMPP1 or DMSO as a control was added 30 min prior to imaging, 2.5 hours after DSB induction.

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A key prediction is that deacetylation of Htz1 at the centromere occurs after DNA damage, and that this deacetylation requires Cep3-S575 and Rdp3L. This would be best addressed by ChIP experiments using acetylation-specific Htz1 antibodies. The importance of Htz1 acetylation for chromosome segregation has been previously concluded from segregation defects in htz1-K14R cells (Keogh et al., 2006b; Krogan et al., 2004), although the mechanism is unclear. My work provides a new interpretation for these data and suggests that mimicking constitutive deacetylation perturbs SPB-CEN dynamics.

My results suggest that the chromatin mobility defects observed in SWR1 and INO80 mutants are linked to Cep3 and likely result from misincorporation of Htz1 in pericentric chromatin. The best evidence that INO80 might promote mobility at DSB sites is that tethering of LexA-Ino80 to chromatin can increase mobility (Neumann et al., 2012); however, this result cannot exclude perturbations to centromeric constraint from ectopic Ino80 expression. Expression of Ino80 without a LexA tag in this system would be required to solidify this result. My observation that DSB-induced mobility includes a signal traveling to the centromere suggests that this process may be highly sensitive to chromatin remodelling activity and might require INO80.

The high baseline chromatin mobility, and even viability, of htz1-4KR sir4Δ mutants is an intriguing observation that warrants further investigation. Although these cells appear normal during routine growth, it is likely that several cellular defects exist and these mutants may provide an unique tool to probe the consequences of increased chromosome dynamics. Prime candidates include chromosome segregation errors, cell cycle abnormalities, and changes in gene expression.

While this works raises more loose ends than desired, the potential involvement of Ipl1 provides an attractive mechanism to regulate SPB-CEN attachment. Key experiments to help address this would be to determine if the detachment of kinetochores following DNA damage, as visualized by Mtw1-3xGFP, requires Ipl1 activity. Similar experiments have revealed that Ipl1 generates the unattached kinetochores in ndc80-1 mutants (Pinsky et al., 2006), and that Ipl1 can bind and phosphorylate the CBF3 complex (Biggins et al., 1999). How deacetylated Htz1 might be involved in Aurora B recruitment or activation remains to be determined; however, this notion is not completely unfounded as mammalian H2A.Z is known to interact with the INCENP subunit of Aurora B (Rangasamy et al., 2003).

Chapter 6 Summary and future directions

How cells differentiate their DNA ends is an important question in chromosome biology. Here I identify that the inhibition of telomerase at DSBs by Pif1 is tightly integrated with telomeric length and uncover an additional regulatory mechanism that promotes genome stability. Future work will be required to identify the exact property of the TG34 DNA end that confers insensitivity to Pif1; however, my results clearly implicate Cdc13 in this process. This investigation has been troubled by my inability to identify a specific molecular function of Cdc13 that triggers the DSB-telomere transition, and the suppression of defective cdc13 alleles by loss of PIF1 which confounds genetic analysis. Both problems suggest that a biochemical approach may be more tractable and is inspired by the recent reconstitution of the multi-protein machineries involved in DNA end resection (Cejka et al., 2010a; Niu et al., 2010) and DNA replication (Yeeles et al., 2015).

The goal of this approach is to modify existing in vitro telomerase extension assays with purified Pif1 and Cdc13 to identify how these proteins regulate end protection and telomere extension. Purified Pif1 has previously been shown to inhibit telomerase at oligonucleotide substrates (Boule et al., 2005) and I will first investigate whether telomerase can extend TG18 and TG34 containing oligonucleotide substrates in the presence of Pif1 (Figure 6.1a). Additional components may also be required for the recapitulation of the DSB-telomere transition observed in vivo; prime candidates include the CST subunits Stn1 and Ten1, and Polα. As Pif1 has also been shown to promote resection at uncapped telomeres (Dewar and Lydall, 2010), a second readout for this assay will be to look at resistance of substrates to Exo1 and I hypothesize that the Cdc13-bound TG34 end may form a barrier to resection (Figure 6.1b). This idea is based on the observations that telomere addition events in pif1-m2 cells are not targeted to TLC1 binding sites (Pennaneach et al., 2006), but are located further from DSB sites (Chung et al., 2010). This can be accomplished by blocking the double-stranded DNA end of the substrate with biotin-Streptavidin.

Pif1 also has the ability to patrol DNA where it binds to a dsDNA-ssDNA junction and reels in the 3’ ssDNA tail (Zhou et al., 2014). This activity can be measured using a Förster resonance energy transfer (FRET) assay as Pif1 brings the FRET donor and acceptor into close proximity. Although Pif1 in this system is able to reel in 70 bp lengths of ssDNA, these experiments were not performed in the presence of Cdc13 (Zhou et al., 2014). I will therefore assess whether Pif1 can also patrol

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DNA substrates containing TG18 or TG34 sequence in the presence of Cdc13 or CST complex (Figure 6.1c).

Figure 6.1. In vitro reconstitution of the DSB-telomere transition.

Figure 6.1. In vitro reconstitution of the DSB-telomere transition. a, Schematic of a 200 bp telomere addition substrate featuring a distal 3’ end containing the TG18 or TG34 substrate. Pif1 exhibits 5’ to 3’ helicase activity, pictured here as translocating towards the end of the ssDNA, but Pif1 could also move in the opposite direction on the complementary strand. The ability of telomerase to extend substrates in this system will be measured gel electrophoresis. b, Schematic of a similar system to measure resistance to Exo1-mediated DNA end resection. The opposite end of the DNA substrate is biotinylated and blocked with Streptavidin to prevent Exo1 activity. c, Schematic of a FRET assay to monitor the Pif1 patrolling function. A 3’ FRET donor (red circle) is brought into proximity of the FRET acceptor at the DNA junction (green circle) as Pif1 reels ssDNA in.

A key observation is that the activity of Pif1 at the TG18 DNA end can be inhibited and promoted by the pif1-4A and pif1-4D mutants respectively, but not at the TG34 DNA end. Understanding the molecular function of these phosphorylation sites is therefore likely to provide clues into events that occur at the DSB-telomere transition. A simple first experiment would be to measure DNA end resection at uncapped telomeres in pif1-4A and pif1-4D cells. Immunoprecipitation of Myc- tagged Pif1 mutants followed by mass spectrometry would also be an attractive method to identify potential protein interactions regulated by these phosphorylation sites. Finally, purified Pif1-4A and Pif1-4D could be tested using in vitro helicase assays (Boule et al., 2005) to determine how these mutations alter Pif1 function.

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The main conclusion from my work concerns how a DSB increases chromosome movement and I have identified a sought-after Mec1-dependent target that is required for this phenomenon. Moreover, my work provides a mechanistic framework for the dynamics of chromosomes after breakage and highlights chromosomal constraints as key factors in this process. By identifying a specific mutant, not otherwise connected to DSB repair, my work suggests that the enhanced mobility of chromatin is not important for homology search or repair. But how can these results be interpreted in light of other work and the known spatial organization of the nucleus? Particularly puzzling is the observation that irreparable DSBs relocate to the nuclear periphery, implying a need for chromatin mobility. Recent work however has also challenged this association as the DSB-induced mobility defects of arp8Δ and rad51Δ mutants do not impair the ability of DSBs to move to the nuclear periphery (Horigome et al., 2014). While artificially tethering the SWR1 subunit Arp6 to chromatin using a LexA fusion is able to promote relocalization in the absence of a DSB, it does not affect chromatin mobility, revealing that increased mobility is neither necessary nor sufficient for the relocalization of DSBs to the periphery (Horigome et al., 2014). One shared conclusion from this work and my own results is that the increase in chromatin mobility is not required and suggests that the normal baseline mobility of chromosomes is adequate for functional movement within the nucleus. A key consideration related to this is the time-scale of various experiments; while mobility is studied on the order of minutes, DSBs are processed for hours before repair. The ability of individual loci to explore, even when impaired, may simply not be a limiting factor in cell processes, especially in the small nuclear volume of budding yeast.

The ability to disconnect chromatin mobility from functional repair processes and DSB relocalization suggests that the random local diffusion of chromatin, while a robust phenotype, is not always informative. For example, the current way of observing chromosome behavior has no relevance on whether or not it will relocate to the periphery, yet we know mobility must somehow be involved. Additional methods will therefore be required in the future to study mobility in more specific ways. This could include longer time-lapse experiments, and to incorporate nuclear position into the analysis, both of which address current limitations in my analysis.

The kinesin-14 motor complex is a good example where new approaches will be required to investigate chromatin mobility (Appendix A). Cik1-Kar3 promotes chromatin mobility and atypical DSB repair; however, the direct link between these phenotypes is questionable in light of other results suggesting that mobility does not impact repair. For example, Cik1-Kar3 is only

120 required for DSB repair by telomere addition and BIR, processes thought to involve relocalization to and from the nuclear periphery, but as previously mentioned the relationship between chromatin mobility and relocalization to the nuclear periphery is not direct. Mobility-defective cep3-S575A mutants are also proficient in both telomere addition and BIR, questioning the general role of chromatin mobility in these processes. One possibility is that Cik1-Kar3 promotes a specific function or movement at the nuclear periphery that is not easily detected or appreciated in my analysis. For example, Cik1-Kar3 might facilitate the movement or hand-off of DNA ends between nuclear pores or complexes in the nuclear periphery like Mps3 and Nup84. These movements might reflect very small distances, but could underlie the repair defects of Cik1-Kar3 mutants.

A fundamental problem is that our study of mobility is currently based on extrapolation. We observe the behavior of a chromosome for a few minutes and hypothesize what this behavior might translate to over longer periods of time. This has led to numerous correlative observations, including the link between chromatin mobility and DNA transcription until later studies revealed that these processes can be separated (Dion and Gasser, 2013). Powerful new approaches combining long term live-cell imaging and single-cell sequencing could provide clear insight into the role of chromatin mobility and nuclear positioning in DNA repair outcomes. This idea is inspired by the ‘Look-Seq’ technique first applied to the isolation and shattering of chromosomes in the micronuclei of mammalian cells (Zhang et al., 2015) and the goal would be to follow a DSB from its origin and to take snapshots of DSB repair after different movements in the nucleus. This approach would require careful optimization to balancing spatial resolution and photobleaching, and the imaging of individual cells in 384-well plates followed by cell rapid fixation throughout the steps of DSB repair. The study of individual cells in this manner could provide strong links between specific mobility events and DSB repair, particularly in situations where two cells of the same genotype exhibit different chromosomal motion.

The ability to image chromatin in real-time is likely a limiting factor in these experiments and new approaches to study DSB mobility retrospectively could be useful in the field. Such systems could tether LacI to a methyltransferase enzyme, for example to the GpC-specific methyltransferase M.CviP from Chlorella virus (Xu et al., 1998) which has previously been used to map nucleosome positions in single yeast cells (Small et al., 2014). DNA extraction and sodium bisulfite treatment would then convert unmethylated cytosines to uracil allowing for the detection of methylation marks and indirectly the chromatin interactions that were made by the DSB end. Similar

121 experiments could fuse LacI and the promiscuous biotin ligase BirA (Roux et al., 2013; Roux et al., 2012) to identify protein interactions observed during the course of nuclear exploration. These systems would require tightly controlled expression of the LacI fusion proteins to limit background interactions that occur before DSB induction.

While I have identified that one function of Cep3 phosphorylation is to promote cell cycle arrest, it is currently unclear if increased chromatin mobility itself has a function that is yet to be determined, or if it simply results as a by-product of physics. A potential insight into this question comes from the observation that the mutation of cep3-S575A actually increases DSB repair in several assays including in a BIR system (Figure 6.2ac), and in an reporter strain containing two interstitial loci which undergo very efficient repair by HR (Figure 6.2bd). These results suggest that increased chromatin mobility may actually inhibit repair in certain contexts and one possible explanation is that increased mobility may promote the proofreading of homologous donor sequence by limiting Rad51 binding to energetically less favourable DNA sequences. In this model, Rad51 filament binding to an imperfect complementary sequence may be displaced by the added forces imposed by chromosome movement thereby promoting reannealing to other sites. As BIR can involve multiple rounds of strand invasion during extension (Donnianni and Symington, 2013; Smith et al., 2007), excess mobility may be particularly detrimental to this process.

This hypothesis can be directly tested by measuring the mobility and spatial retention of Rad52- YFP labelled DSBs in the presence of donor sites containing varying degrees of sequence homology (Figure 6.2e). I hypothesize that DSBs will exhibit slower movement and a persistent interaction upon co-localization with perfect homologous donors, but not with donor sequences featuring less sequence complementarity. Furthermore, the use of cep3-S575A mutants in this system will determine if this preference is due to increased chromosome movement. Genetic assays in strains which feature multiple donors simultaneously can also be used to determine if cep3-S575A cells undergo DSB repair using sequences with less homology (Figure 6.2f). As chromatin movement increases with temperature (Neumann et al., 2012), performing these experiments at both 20°C and 30°C may also reveal changes in donor preference due to differences in kinetic energy. As the bulk of my experiments have previously followed the behavior of an irreparable DSB, it will also be important in the future to study mobility during active HR repair.

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A final consideration involves the relationship of my work to the movement of mammalian DSBs and due to the unique nuclear architecture of budding yeast, directly analogous mechanisms are unlikely. While mammalian chromosomes have physical interactions with nuclear structures including the nucleolus, nuclear periphery, nuclear speckles, and PML bodies (Chubb et al., 2002; Molenaar et al., 2003; Wiesmeijer et al., 2008), future work will be required to characterize if these attachments are modified following DNA damage.

The recent connection between microtubules and the mobility of mammalian DSBs is intriguing but also indicates an opposite regulatory role in which microtubules “poke” the nucleus to stimulate motion (Lottersberger et al., 2015). Given the similar diffusion coefficient of chromatin between organisms, the 200- to 400-fold increase of nuclear volume in mammalian cells also poses a significant barrier for the ability of chromosome movement to impact cellular functions. One potential conserved mechanism from my work might rather involve crosstalk between the DNA damage and spindle assembly checkpoints as several ATR kinase targets have been identified in SAC components as well as the centromeric protein CENPF (Matsuoka et al., 2007).

Figure 6.2. Does increased chromosome mobility function in the proofreading of donor sites?

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Figure 6.2. Does increased chromosome mobility function in the proofreading of donor sites? a, Schematic of the tGI 354 HR reporter featuring a HO endonuclease cut site on Chr V which can be repaired by a donor template on Chr III (Ira et al., 2003). b, Schematic of the JRL346 BIR reporter, see Appendix A Figure 3 for more information. c, Repair efficiency of tGI 354 strains in the indicated genetic backgrounds. Data represents the mean ± s.d. from n=3 independent experiments. d, Repair efficiency by BIR in strains of the indicated genotype as measured by the proportion of surviving canavanine sensitive colonies. Data represents the mean ± s.d. from n=3 independent experiments. e, Schematic of yeast strains to study the mobility of DSBs in the presence of perfect (ex. CATAGG) or imperfect template donors (ex. TATAGG). Cells express Nup49-mCherry, Rad52-YFP, and LacO-CFP to facilitate nuclear alignment and visualization of both the donor and template DNA. f, Schematic of a genetic assay to test if increased chromosome mobility favours repair by a perfectly complementary donor sequence. To control for the physical location of the donor, variant strains with shuffled donors will be required. Template choice can be determined by unique restriction sites at each donor and confirmed by PCR amplification and DNA sequencing of the repaired DSB site. For all panels: *** p<0.001, ** p<0.01; Student’s t- test.

Appendix A: The kinesin-14 motor complex promotes general chromatin mobility and atypical DSB repair

Statement of contributions: While I performed all of the experiments presented in this appendix, this project was initiated and led by Wei Zhang. Notably, Wei discovered that Cik1-Kar3 is important for telomere addition and first postulated that kinesin-14 might play a role in chromatin mobility. Wei also discovered that Cik1-Kar3 promote telomerase-independent telomere maintenance and my data is simply confirmation of this observation (Appendix A Figure 3h).

Some of the text and figures presented in this chapter were modified with permission from the following publication:

Chung, D.K., Chan, J.N., Strecker, J., Zhang, W., Ebrahimi-Ardebili, S., Lu, T., Abraham, K.J., Durocher, D., and Mekhail, K. (2015). Perinuclear tethers license telomeric DSBs for a broad kinesin- and NPC-dependent DNA repair process. Nat Commun 6, 7742. doi:10.1038/ncomms8742

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Introduction

Previous work in our lab identified CIK1 in a screen for genes required for telomere addition (Zhang and Durocher, 2010). Cik1 forms a complex with the Kar3 ATPase, together forming the kinesin-14 motor in S. cerevisiae. Kar3 is targeted to the plus-end of microtubules by Cik1 (Page et al., 1994) and can translocate to the minus-end and promote microtubule destabilization (Sproul et al., 2005). Kar3 has been implicated in several microtubule functions including karyogamy (Meluh and Rose, 1990), mitotic spindle stability (Gardner et al., 2008), and kinetochore attachment (Tanaka et al., 2005), making the involvement of Cik1-Kar3 in telomere addition surprising to say the least. As this type of repair is thought to involve relocalization to the nuclear periphery and association with the anchoring protein Mps3 (Oza et al., 2009), one possibility is that kinesin-14 might be involved in the transport of DNA ends. My main objective was to characterize chromatin mobility in kinesin-14 mutants and to investigate if Cik1-Kar3 are required for other types of DSB repair.

Results

Kinesin-14 promotes chromatin mobility independent of DNA damage

I first measured chromatin mobility at the MAT locus in cik1Δ and kar3Δ cells and observed greatly reduced mobility in both mutants (Appendix A Figure 1ab). Importantly, chromatin mobility still increased following a DSB suggesting that Cik1-Kar3 promotes the general mobility of chromatin independent of DNA damage. Kar3 forms a separate complex with a Cik1 paralog, Vik1 (Manning et al., 1999); however, loss of VIK1 had no impact on chromosome movement (Appendix A Figure 1c) indicating that this function is specific to the Cik1-containing complex. To test whether other kinesins exhibit similar activity I next deleted CIN8, the kinesin-5 motor. Loss of CIN8 revealed only a mild defect following DSB induction (Appendix A Figure 1d) indicating that kinesin-14 may have a particularly important role in chromosome movement.

To determine if the ATPase activity of Kar3 is required for this function I used the kar3-898 mutant which specifically fails to hydrolyze ATP upon microtubule binding (Song and Endow, 1998). The

126 reintroduction of wild-type KAR3, but not kar3-898, led to increased chromatin mobility in kar3Δ cells (Appendix A Figure 2a-c) indicating that Kar3 ATPase activity is necessary for this process.

Appendix A Figure 1. Cik1-Kar3 promote general chromatin mobility

Appendix A Figure 1. Cik1-Kar3 promote general chromatin mobility. a-d, MSD analysis of the MAT locus in cik1Δ (a), kar3Δ (b), vik1Δ (c), and cin8Δ cells (d).

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Appendix A Figure 2. Kar3 ATPase activity is required for chromatin mobility

Appendix A Figure 2. Kar3 ATPase activity is required for chromatin mobility. a-c, MSD analysis of the MAT locus in kar3Δ cells containing pRS414-KAR3 (a), pRS414-kar3-898 (b), or an empty pRS414 plasmid (c).

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Cik1-Kar3 are important for DNA repair by break-induced replication

Although chromatin mobility in cik1Δ mutants increases after a DSB, the resulting mobility is still less than that of wild-type cells in the absence of DNA damage (Appendix A Figure 1a). Despite this severe mobility defect, cik1Δ cells were proficient at DSB repair by HR in the four tested strains (Appendix A Figure 3ab) reinforcing the argument that chromosome movement is not required for interchromosomal recombination.

In addition to its role in telomere healing, Wei Zhang also identified that Cik1-Kar3 is important for DSB repair in the YMV2 strain (Zhang, 2012) in which a DSB can be repaired by single-strand annealing or break-induced replication (BIR), the later which utilizes homology with one DNA end to facilitate DNA replication to the end of the chromosome (Vaze et al., 2002) (Jain et al., 2009). Similar to telomere addition, BIR promotes the repair of subtelomeric DSBs and is thought to require localization to the nuclear periphery and interactions with nuclear pore complexes (Therizols et al., 2006). To test whether Cik1-Kar3 is specifically required for repair by BIR I used a system in which a DSB adjacent to a partial CAN1 sequence on Chr V can be repaired by strand invasion and replication of a homologous sequence on Chr XI (Appendix A Figure 3c) (Lydeard et al., 2007). Repair by BIR in this system results in canavanine sensitive colonies and is dependent on the Polδ subunit Pol32 as previously described (Appendix A Figure 3d) (Lydeard et al., 2007). Deletion of CIK1 and KAR3 decreased repair by BIR in this assay and revealed delayed kinetics of repair intermediates (Appendix A Figure 3de). Furthermore, the reintroduction of the kar3-1 and kar3-898 ATPase mutants was unable to rescue the kar3Δ defect (Appendix A Figure 3f) indicating that Kar3 ATPase activity is required for BIR. Cells harbouring the cep3-S575A mutation were proficient in both BIR and telomere addition (Appendix A Figure 3dg) suggesting that increased chromatin movement following a DSB is not required for these repair outcomes.

BIR is involved in the telomerase-independent alternative lengthening of telomeres (Lydeard et al., 2007) suggesting that Cik1-Kar might also promote telomere maintenance in this manner. Serial propagation of telomerase null cik1Δ and kar3Δ mutants revealed accelerated cellular senescence concomitant with shortened telomeres (Appendix A Figure 3hi) indicating that Cik1- Kar3 is important for telomerase-independent telomere maintenance.

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Appendix A Figure 3. Cik1-Kar3 promotes break-induced replication.

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Appendix A Figure 3. Cik1-Kar3 promotes break-induced replication. a, Schematic of HR repair strains indicating the relative location of each homologous cassette and the HO cut site (red). b, Repair efficiency of HR strains in the indicated genetic backgrounds. Data represents the mean ± s.d. from n=3 independent experiments. c, Schematic of the JRL 346 strain which measures DSB repair by BIR. An HO cut site on Chr V can be repaired using a priming template on Chr XI leading to CAN1+ HPH- colonies. d, e, Repair efficiency by BIR in strains of the indicated genotype as measured by the proportion of surviving canavanine sensitive colonies (d), and by semi- quantitative PCR using primers P1 and P2 which specifically amplify the BIR product (e). Data represents the mean ± s.d. from n=6 (d) and n=3 (e) independent experiments f, Repair efficiency of JRL 346 kar3Δ cells transformed with empty pRS414 or pRS414 expressing KAR3, kar3-1, or kar3-898. Data represents the mean ± s.d. from n=6 independent experiments. g, Telomere addition frequency in TG5-HO pif1-m2 cells of the indicated genotypes. Data represents the mean ± s.d. from n=3 independent experiments. h, Serial culturing of the indicated genotypes following tetrad dissection. i, Southern blot for telomere length of the cultures shown in panel h. The calculated number of generations for each time point is indicated. For all panels: *** p<0.001, ** p<0.01; One-way ANOVA.

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The lack of correlation between the general movement of DNA ends within the nucleus and DSB repair questions whether the repair defects in cik1Δ and kar3Δ mutants are in fact caused by defective chromosome movement. While this is a difficult issue to address directly, I next asked whether Cik1-Kar3 also promotes chromatin mobility in the context of BIR and to do so added Nup49-mCherry and Rad52-YFP to the JRL 346 strain. Analysis of Rad52-YFP mobility revealed lower movement compared to the MAT locus, possibly due to associations with the nuclear periphery; however, mobility was again decreased in cik1Δ and kar3Δ mutants (Appendix A Figure 3a). This defect could be rescued with wild-type KAR3 expression, but not the kar3-1 mutant (Appendix A Figure 3b), suggesting that the ATPase function of Kar3 is also required for mobility in this context.

Appendix A Figure 4. Cik1-Kar3 promotes chromatin mobility during BIR

Appendix A Figure 4. Cik1-Kar3 promotes chromatin mobility during BIR. a, MSD analysis of Rad52-YFP in response to a DSB induced in the JRL 346 BIR reporter. b, MSD analysis of Rad52-YFP in JRL 346 kar3Δ cells transformed with empty pRS414 or pRS414 expressing KAR3 or kar3-1.

Discussion

The movement of DNA within the nucleus is dependent on ATP, yet the reason for this dependence is not fully understood. Nucleosome remodelers are likely candidates; however, in the case of INO80 and SWR1, their deletion only affects the increase in chromatin mobility following a DSB. Here I identify that Cik1-Kar3 is an ATPase that promotes the general movement of chromosomes.

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As kinetochore attachment represents a major constraint on chromosomes, the observation that all four nuclear kinesins in S. cerevisiae localize to the kinetochore (Tytell and Sorger, 2006) suggests that other motors might also promote mobility in certain contexts, although Cin8 appears to have a limited role in this process.

The involvement of Cik1-Kar3 in DSB repair is intriguing and appears to be limited to telomere addition and BIR. Both pathways use mutagenic repair, are thought to occur slowly, and require contacts with the nuclear periphery. The known relocalization of slowly repaired DSBs to the nuclear periphery (Oza et al., 2009) hints that this process might be promoted by Cik1-Kar3; however, recombination and telomerase elongation is thought to occur in the nuclear interior (Bystricky et al., 2009; Ferreira et al., 2011; Nagai et al., 2008), suggesting that Cik1-Kar3 may also be important for moving DSBs away from the nuclear periphery or to specific nuclear pore complexes. This second option is attractive as it is now known that increased mobility is not sufficient, or required, to relocalize DSBs to the nuclear periphery (Horigome et al., 2014).

These results are correlative in nature and future work will be required to identify more directly whether Cik1-Kar3 in fact promotes atypical repair by promoting DSB mobility. The observation that cep3-S575A cells are proficient at both BIR and telomere addition indicates that increased mobility is not strictly required for these repair pathways, consistent with the idea that Cik1-Kar3 might play a specific role in the transport of DNA ends.

Appendix B: Summary of DNA ends TG repeat Sequence (5’ to 3’) TG6 ACCACA TG12 ACACACCCACAC TG18 ACACCACACCCACACACA TG22 ACACCACACCCACACACACACC TG26 ACACCACACCCACACACACACCCACA TG30 ACACCACACCCACACACACACCCACACCCA TG34 ACACACACACCACACCCACACCCACACACCACAC TG34v2 ACACCACACCCACACACACACCCACACCCACACA TG38 ACACCACACCCACACACACACCCACACCCACACACCAC TG45 ACCCACACACCCACACCCACACACCACACCCACACACACCACACC ACACCCACCACACCCACACACCCACACCCACACACCACACCCACACACA TG 56 CCACACC ACACACACACCACACCCACCACACCCACACACCCACACCCACACACCAC TG 67 ACCCACACACACCACACC ACACACACACCACACCCACACCCACACACCACACCACACACACCACACC TG 82 CACCACACCCACACACCCACACCCACACACCAC TG26-A ACCACACACCCACACACCACACCCAC TG36-A ACCACACACCCACACACCACACCCACACACACCACA TG26-B ACCACACACACCACACCCACACCACA TG36-B ACCACACACACCACACCCACACCACACCCACACACC TG26-C ACCACACCACACCCACACACCACACC TG36-C ACCACACCACACCCACACACCACACCCACACACACC TG26-(TGTGG) ACCACACCACACCACACCACACCACA TG36-(TGTGG) ACCACACCACACCACACCACACCACACCACACCACA TG26-(TG) ACACACACACACACACACACACACAC TG36-(TG) ACACACACACACACACACACACACACACACACACAC ACCACACCACACACCACACACCCACACCCACACACCCACACCCACACCA TG 96 CACACCCACACCACACACCCACACCCACACACCCACACCCACACCAC ACCACACCACACACCACACACCCACACCCACACCACACCACACCACACA TG119 CCACACACCCACACCCACACACCCACACCCACACCACACACCCACACCA CACACCACACACCCACACCCA ACCACACCACACACCACACACCCACACCCACACCACACACCCACACCCA TG142 CACCACACCCACACCCACACCACACCACACCACACACCACACACCCACA CCCACACACCCACACCCACACCACACACCCACACCACACACACC ACCACACCACACACCACACACCCACACCCACACCACACACCCACACCCA CACCACACCCACACCCACACCACACCACACCACACACCACACACCCACA TG 162 CCCACACACCCACACCCACACCACACACCCACACCACACACACCCACAC CACACCCACACCACA ACACCACACACCCACACCCCACACCCACACCCACACCACACACCCACAC CCACACCACACCCACACCACACCACACACACCCACACCACACACACACC CACACCACACACCCACACCCACACACACCCACACCACACACACCCACAC TG 251 CACACACCCACACCCACACACCCACACCACACCCACACCACACCACACC CACACACCCACACCACACACACCACACACCCACACCACACCCACACCAC ACACCC

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GTAGATCCTAAGCGGTACACGCATACACCTTGATGTACACGCATACACC Rap1 x0 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT GTAGATCCTAAGCGGTACACCCATACACCTTGATGTACACGCATACACC Rap1 x1 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT GTAGATCCTAAGCGGTACACCCATACACCTTGATGTACACCCATACACC Rap1 x2 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT GTAGATCCTAAGCGGTACACCCATACACCTTGATGTACACCCATACACC Rap1 x3 TTGATGTACACCCATACACCTTGATGTACACGCATACACCTTGAT GTAGATCCTAAGCGGTACACCCATACACCTTGATGTACACCCATACACC Rap1 x4 TTGATGTACACCCATACACCTTGATGTACACCCATACACCTTGAT ACACACACCCACACGTACACGCATACACCTTGATGTACACGCATACACC Rap1 +TG x0 14 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT ACACACACCCACACGTACACCCATACACCTTGATGTACACGCATACACC Rap1 +TG x1 14 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT ACACACACCCACACGTACACCCATACACCTTGATGTACACCCATACACC Rap1 +TG x2 14 TTGATGTACACGCATACACCTTGATGTACACGCATACACCTTGAT ACACACACCCACACGTACACCCATACACCTTGATGTACACCCATACACC Rap1 +TG x3 14 TTGATGTACACCCATACACCTTGATGTACACGCATACACCTTGAT ACACACACCCACACGTACACCCATACACCTTGATGTACACCCATACACC Rap1 +TG x4 14 TTGATGTACACCCATACACCTTGATGTACACCCATACACCTTGAT

Appendix C: Summary of chromatin mobility parameters Genotype Particle tracked Locus Condition Rc (µm) D (10-3 µm2/s) # cells Wild-type GFP-LacI MAT control 0.480±0.010 1.458±0.034 264 DSB 0.584±0.012 1.949±0.051 331 sml1Δ GFP-LacI MAT control 0.502±0.011 1.443±0.035 192 DSB 0.605±0.016 2.195±0.109 170 sml1Δ mec1Δ GFP-LacI MAT control 0.492±0.013 1.523±0.044 159 DSB 0.567±0.017 1.906±0.072 120 sml1Δ rad53Δ GFP-LacI MAT control 0.457±0.012 1.416±0.041 194 DSB 0.491±0.016 1.533±0.062 123 sml1Δ tel1Δ GFP-LacI MAT control 0.490±0.012 1.576±0.042 184 DSB 0.600±0.014 2.226±0.078 187 sml1Δ mec1Δ tel1Δ GFP-LacI MAT control 0.507±0.012 1.553±0.039 165 DSB 0.514±0.012 1.622±0.055 161 dun1Δ GFP-LacI MAT control 0.488±0.012 1.503±0.046 168 DSB 0.540±0.014 1.836±0.058 182 chk1Δ GFP-LacI MAT control 0.494±0.017 1.325±0.046 105 DSB 0.559±0.017 1.764±0.063 151 mad2Δ GFP-LacI MAT control 0.484±0.012 1.415±0.035 187 DSB 0.570±0.021 2.002±0.088 116 arp8Δ GFP-LacI MAT control 0.516±0.011 1.606±0.044 192 DSB 0.539±0.015 1.807±0.056 165 CEP3 GFP-LacI MAT control 0.506±0.010 1.596±0.042 262 DSB 0.653±0.016 2.208±0.075 228 cep3-S575A GFP-LacI MAT control 0.514±0.009 1.533±0.034 336 DSB 0.534±0.012 1.763±0.052 236 cep3-S575A sml1Δ GFP-LacI MAT control 0.484±0.014 1.431±0.045 136 rad531Δ DSB 0.518±0.019 1.670±0.087 83 BIR1 GFP-LacI MAT control 0.504±0.018 1.427±0.049 97 DSB 0.622±0.031 2.059±0.127 57 bir1-S751A GFP-LacI MAT control 0.531±0.020 1.352±0.054 81 DSB 0.584±0.020 2.095±0.087 114 CBF1 GFP-LacI MAT control 0.522±0.018 1.559±0.075 84 DSB 0.621±0.026 2.023±0.132 65 cbf1-S45/48A GFP-LacI MAT control 0.486±0.018 1.494±0.054 113 DSB 0.615±0.025 2.180±0.097 102 DAD1 GFP-LacI MAT control 0.463±0.017 1.454±0.065 81 DSB 0.629±0.026 1.986±0.108 62 dad1-S89A GFP-LacI MAT control 0.474±0.018 1.435±0.071 86 DSB 0.639±0.028 2.161±0.108 61

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DAD3 GFP-LacI MAT control 0.487±0.018 1.457±0.052 98 DSB 0.636±0.026 2.332±0.147 57 dad3-S43A GFP-LacI MAT control 0.486±0.017 1.391±0.056 102 DSB 0.637±0.026 2.012±0.091 74 GLC7 GFP-LacI MAT control 0.519±0.021 1.628±0.078 67 DSB 0.596±0.025 2.132±0.115 67 glc7-S3A GFP-LacI MAT control 0.541±0.022 1.531±0.062 88 DSB 0.613±0.020 2.198±0.112 108 cep3-S575A GFP-LacI MAT control 0.532±0.016 1.611±0.054 139 (endogenous) DSB 0.560±0.016 1.936±0.072 113 Wild-type GFP-LacI MAT control 0.464±0.012 1.235±0.037 180 (diploid) DSB 0.587±0.018 1.993±0.093 134 cep3-S575A GFP-LacI MAT control 0.517±0.01 1.474±0.031 306 (diploid) DSB 0.524±0.012 1.579±0.040 208 HOcsΔ GAL-CEN3 GFP-LacI MAT control 0.463±0.009 1.297±0.027 287 GAL 0.515±0.011 1.527±0.036 294 sir4Δ GFP-LacI MAT control 0.516±0.018 1.440±0.056 92 DSB 0.560±0.026 1.656±0.080 81 yku70Δ GFP-LacI MAT control 0.514±0.012 1.622±0.021 161 DSB 0.609±0.021 2.23±0.101 102 yku70Δ sir4Δ GFP-LacI MAT control 0.530±0.020 1.674±0.062 116 DSB 0.580±0.021 2.124±0.105 125 HOcsΔ GAL-CEN3 GFP-LacI MAT control 0.529±0.015 1.421±0.037 133 sir4Δ GAL 0.558±0.016 1.578±0.046 150 GAL-CEN3 GFP-LacI MAT control 0.527±0.017 1.565±0.053 107 DSB/GAL 0.623±0.018 2.097±0.074 160 Wild-type Rad52-YFP ADH4 DSB 0.544±0.014 1.197±0.072 304 cep3-S575A Rad52-YFP ADH4 DSB 0.456±0.011 0.959±0.024 345 Wild-type Rad52-YFP AMD2 DSB 0.580±0.020 1.575±0.070 153 cep3-S575A Rad52-YFP AMD2 DSB 0.502±0.017 1.130±0.043 132 cep3-S575A GAL- GFP-LacI MAT control 0.489±0.015 1.514±0.054 117 CEN3 DSB/GAL 0.660±0.023 2.586±0.172 84 cep3-S575A GAL- GFP-LacI MAT control 0.520±0.018 1.528±0.055 129 CEN5 DSB/GAL 0.531±0.021 1.827±0.091 81 arp8Δ GAL-CEN3 GFP-LacI MAT control 0.509±0.014 1.458±0.044 154 DSB/GAL 0.603±0.021 2.358±0.124 88 arp8Δ GAL-CEN5 GFP-LacI MAT control 0.524±0.014 1.693±0.048 151 DSB/GAL 0.553±0.017 1.956±0.082 105 Wild-type GFP-LacI ChrV:53601 control 0.488±0.013 1.226±0.033 210 (27kb DSB-CEN) DSB in trans 0.577±0.011 1.468±0.032 357

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Wild-type GFP-LacI ChrV:53601 control 0.508±0.021 1.273±0.051 104 (101kb DSB-CEN) DSB in trans 0.532±0.018 1.387±0.044 138 Wild-type GFP-LacI ChrV:53601 control 0.490±0.017 1.391±0.057 97 (510kb DSB-CEN) DSB in trans 0.542±0.019 1.395±0.043 120 Wild-type GFP-LacI ChrV:53601 control 0.497±0.019 1.419±0.062 84 (998kb DSB-CEN) DSB in trans 0.528±0.018 1.528±0.055 97 cep3-S575A GFP-LacI ChrV:53601 control 0.509±0.015 1.368±0.045 140 (27kb DSB-CEN) DSB in trans 0.523±0.017 1.295±0.039 126 sir4Δ GFP-LacI ChrV:53601 control 0.542±0.018 1.448±0.050 233 (27kb DSB-CEN) DSB in trans 0.604±0.021 1.696±0.066 263 Wild-type GFP-LacI pRS415 control 0.395±0.013 0.886±0.029 144 (27kb DSB-CEN) CEN-ARS DSB 0.449±0.014 1.138±0.074 173 cep3-S575A GFP-LacI pRS415 control 0.398±0.013 0.951±0.061 135 (27kb DSB-CEN) CEN-ARS DSB 0.420±0.014 0.937±0.034 143 Wild-type GFP-LacI pRS415 control 0.414±0.018 0.997±0.074 110 (998kb DSB-CEN) CEN-ARS DSB 0.420±0.014 0.968±0.037 147 Wild-type GFP-LacI ChrIV:351224 control 0.457±0.012 1.331±0.042 173 (uncut Chr IV-L) DSB 0.507±0.013 1.385±0.043 167 cep3-S575A GFP-LacI ChrIV:351224 control 0.473±0.012 1.443±0.042 158 (uncut Chr IV-L) DSB 0.449±0.013 1.244±0.037 153 sir4Δ GFP-LacI ChrIV:351224 control 0.521±0.012 1.487±0.049 172 (uncut Chr IV-L) DSB 0.547±0.015 1.597±0.047 191 Wild-type GFP-LacI ChrIV:539110 control 0.454±0.013 1.345±0.043 147 (89 kb array) DSB 0.555±0.015 1.727±0.063 183 Wild-type GFP-LacI ChrIV: control 0.554±0.014 1.730±0.015 141 (497 kb array) 946605 DSB 0.580±0.015 1.656±0.063 206 Wild-type GFP-LacI ChrIV: control 0.596±0.014 1.921±0.047 187 (985 kb array) 1447374 DSB 0.587±0.019 1.729±0.060 163 Wild-type Spc42/ SPB/CEN5 control 0.339±0.011 0.516±0.044 94

(ΔdSPB-CEN) GFP-LacI DSB in trans 0.404±0.010 0.738±0.037 258 cep3-S575A Spc42/ SPB/CEN5 control 0.356±0.010 0.620±0.036 94

(ΔdSPB-CEN) GFP-LacI DSB in trans 0.354±0.011 0.569±0.045 203 Wild-type GFP-LacI ChrIV: control 0.557±0.013 1.606±0.045 181 (TEL side) 946605 DSB 0.575±0.014 1.884±0.060 193 cep3-S575A GFP-LacI ChrIV: control 0.548±0.014 1.743±0.058 125 (TEL side) 946605 DSB 0.584±0.016 1.994±0.064 128 sml1Δ mec1Δ GFP-LacI ChrIV: control 0.544±0.014 1.736±0.049 164 (TEL side) 946605 DSB 0.583±0.015 1.935±0.058 148 sir4Δ GFP-LacI ChrIV: control 0.582±0.014 1.772±0.049 189 (TEL side) 946605 DSB 0.603±0.020 1.748±0.062 131

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Wild-type GFP-LacI MAT control 0.483±0.015 1.560±0.051 118 No treatment DSB 0.586±0.016 2.011±0.083 141 Wild-type GFP-LacI MAT control 0.480±0.017 1.230±0.044 95 VPA (2 mM) DSB 0.485±0.020 1.345±0.066 83 Wild-type GFP-LacI MAT control 0.468±0.018 1.290±0.052 79 NAM (5 mM) DSB 0.609±0.022 1.874±0.081 102 hda11Δ GFP-LacI MAT control 0.459±0.014 1.295±0.048 114 DSB 0.567±0.017 1.873±0.062 143 hos1Δ GFP-LacI MAT control 0.477±0.017 1.362±0.050 88 DSB 0.556±0.020 1.813±0.063 133 hos2Δ GFP-LacI MAT control 0.504±0.021 1.491±0.071 70 DSB 0.595±0.028 2.094±0.102 74 hos3Δ GFP-LacI MAT control 0.497±0.017 1.445±0.050 111 DSB 0.639±0.022 1.984±0.073 136 sir2Δ GFP-LacI MAT control 0.507±0.015 1.444±0.048 104 DSB 0.605±0.018 2.028±0.097 132 hst4Δ GFP-LacI MAT control 0.485±0.013 1.399±0.046 111 DSB 0.572±0.019 1.840±0.076 115 rpd3Δ GFP-LacI MAT control 0.481±0.015 1.423±0.055 103 DSB 0.488±0.019 1.473±0.095 117 sin3Δ GFP-LacI MAT control 0.496±0.017 1.558±0.079 74 DSB 0.513±0.016 1.506±0.055 163 rxt3Δ GFP-LacI MAT control 0.521±0.017 1.553±0.059 117 DSB 0.497±0.018 1.380±0.061 117 sds3Δ GFP-LacI MAT control 0.509±0.018 1.435±0.069 78 DSB 0.505±0.022 1.331±0.078 71 rco1Δ GFP-LacI MAT control 0.500±0.018 1.517±0.061 81 DSB 0.579±0.016 1.833±0.068 156 cep3-S575E GFP-LacI MAT control 0.519±0.013 1.535±0.038 213 DSB 0.556±0.013 1.854±0.060 213 HTZ1 GFP-LacI MAT control 0.451±0.013 1.407±0.050 137 DSB 0.568±0.016 1.834±0.060 182 htz1-4KQ GFP-LacI MAT control 0.426±0.014 1.151±0.049 111 DSB 0.470±0.015 1.452±0.070 108 htz1-k14Q GFP-LacI MAT control 0.480±0.011 1.449±0.040 180 DSB 0.493±0.011 1.542±0.042 206 htz1-K14R GFP-LacI MAT control 0493.±0.011 1.444±0.037 173 DSB 0.539±0.018 1.661±0.079 142 htz1-4KR GFP-LacI MAT control 0.493±0.016 1.468±0.048 133 DSB 0.560±0.019 1.703±0.069 133

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htz1-4KR sir4Δ GFP-LacI MAT control 0.578±0.014 1.782±0.054 217 DSB 0.572±0.018 1.815±0.070 150 htz1-4KR cep3-S575A GFP-LacI MAT control 0.518±0.014 1.499±0.045 163 DSB 0.599±0.016 2.008±0.068 177 OsTIR1 GFP-LacI MAT control 0.441±0.016 1.231±0.051 87 Ethanol DSB 0.545±0.020 1.590±0.065 143 OsTIR1 GFP-LacI MAT control 0.477±0.015 1.320±0.055 78 IAA (0.5 mM) DSB 0.535±0.018 1.652±0.066 124 OsTIR1 IPL1-3xAID GFP-LacI MAT control 0.438±0.016 1.407±0.072 69 Ethanol DSB 0.496±0.019 1.471±0.072 126 OsTIR1 IPL1-3xAID GFP-LacI MAT control 0.459±0.019 1.258±0.062 79 IAA (0.5 mM) DSB 0.452±0.013 1.155±0.042 180 ipl1-as6 (DMSO) GFP-LacI MAT DSB 0.580±0.018 1.640±0.068 114 ipl1-as6 (NMPP1) GFP-LacI MAT DSB 0.476±0.015 1.337±0.061 127 cik1Δ GFP-LacI MAT control 0.376±0.021 1.053±0.037 28 DSB 0.460±0.022 1.063±0.064 53 kar3Δ GFP-LacI MAT control 0.421±0.018 1.134±0.055 36 DSB 0.459±0.031 1.402±0.131 33 cin8Δ GFP-LacI MAT control 0.514±0.017 1.593±0.066 94 DSB 0.580±0.017 1.831±0.059 135 vik1Δ GFP-LacI MAT control 0.524±0.025 0.621±0.037 51 DSB 1.433±0.070 2.295±0.185 36 kar3Δ GFP-LacI MAT control 0.486±0.029 1.357±0.097 40 pRS414-KAR3 DSB 0.574±0.038 1.713±0.156 29 kar3Δ GFP-LacI MAT control 0.426±0.016 1.146±0.057 56 pRS414-kar3-898 DSB 0.438±0.023 1.349±0.124 40 kar3Δ GFP-LacI MAT control 0.420±0.016 1.083±0.053 67 pRS414 DSB 0.459±0.029 1.171±0.077 36 Wild-type Rad52-YFP BIR DSB 0.467±0.021 0.805±0.032 96 cik1Δ Rad52-YFP BIR DSB 0.383±0.022 0.725±0.038 56 kar3Δ Rad52-YFP BIR DSB 0.371±0.019 0.600±0.038 49 kar3Δ pRS414-KAR3 Rad52-YFP BIR DSB 0.453±0.018 0.896±0.034 112 kar3Δ pRS414-kar3-1 Rad52-YFP BIR DSB 0.398±0.020 0.663±0.034 70 kar3Δ pRS414 Rad52-YFP BIR DSB 0.378±0.020 0.636±0.027 68

Appendix D: List of yeast strains Strain Strain Number Background Genotype Source Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- (Zhang & DDY2458 S288C Δ1::GAL1:HO-LEU2 rad52::HIS pRAD52-TRP VII-L-ADE2-TG82- Durocher, HOcs-LYS2 2010) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- (Zhang & DDY2476 S288C Δ1::GAL1:HO-LEU2 rad52::HIS pRAD52-TRP VII-L-ADE2-TG82- Durocher, HOcs-LYS2 pif1-m2 2010) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3254 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3203 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3376 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2986 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2985 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG45-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2987 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG45-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3129 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG56-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3130 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG56-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2988 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG67-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2990 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG67-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2472 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY2556 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 - This study m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3204 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG22-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3205 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG22-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3206 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3207 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26-HOcs-LYS2 This study pif1-m2

140

141

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3208 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG30-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3209 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG30-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3210 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3211 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3404 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG38-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3406 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG38-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3275 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26a-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3276 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26a-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3277 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36a-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3278 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36a-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3279 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26b-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3280 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26b-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3281 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36b-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3282 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36b-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3283 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26c-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3284 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26c-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3285 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36c-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3286 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36c-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3042 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study tel1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3043 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study pif1-m2 tel1::KANMX

142

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3483 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study tel1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3484 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 tel1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3485 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study tel1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3486 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 tel1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3234 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 AUR1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3236 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3244 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1(5AQ) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3224 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3226 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3230 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1(4A) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3470 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1(4D) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3240 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1(4D) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3141 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study sml1::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3142 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study pif1-m2 sml1::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3144 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study sml1:NATMX: mec1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3039 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study pif1-m2 sml1:NATMX: mec1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3146 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study sml1::NATMX rad53::KANMX

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Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3041 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L-ADE2-TG82-HOcs-LYS2 This study pif1-m2 sml1::NATMX rad53::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3224 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3226 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3470 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 AUR1::pif1-m1(4D) Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3604 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-Cdc13-Est1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3605 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-Cdc13-Est1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3606 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-Cdc13-Est1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3607 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-Cdc13-Est1 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3608 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-Cdc13-Est2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3609 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-Cdc13-Est2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3610 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-Cdc13-Est2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3611 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-Cdc13-Est2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3499 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study est2-up34 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3500 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 est2up-34 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3501 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study est2-up34 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3502 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 est2-up34 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3287 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26d-HOcs-LYS2

144

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3288 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26d-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3289 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36d-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3290 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36d-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3291 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26e-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3292 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG26e-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3293 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36e-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3294 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG36e-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3324 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x0+TG14- This study HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3325 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x0+TG14- This study HOcs-LYS2 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3326 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x1+TG14- This study HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3327 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x1+TG14- This study HOcs-LYS2 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3328 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x2+TG14- This study HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3329 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x2+TG14- This study HOcs-LYS2 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3330 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x3+TG14- This study HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3331 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x3+TG14- This study HOcs-LYS2 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3332 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x4+TG14- This study HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3333 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x4+TG14- This study HOcs-LYS2 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3334 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x0-HOcs-LYS2

145

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3335 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x0-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3336 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x1-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3337 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x1-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3338 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x2-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3339 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x2-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3340 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x3-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3341 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x3-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3342 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x4-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3343 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-Rap1x4-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3475 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study rif1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3476 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 rif1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3477 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study rif1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3478 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 rif1::KANMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3479 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3480 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3481 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3482 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3466 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study rif1::KANMX rif2::NATMX

146

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3467 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 rif1::KANMX rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3468 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study rif1::KANMX rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3469 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG34-HOcs-LYS2 This study pif1-m2 rif1::KANMX rif2::NATMX Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3526 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3527 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3528 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3530 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 This study pif1-m2 ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3589 S288C Δ1::GAL1:HO rad52::HIS VII-L::TG18-HOcs-LYS2 leu2::NAT This study ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 pRAD52-TRP Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- Δ1::GAL1:HO rad52::HIS VII-L::URA3-TG18-HOcs-LYS2 pif1-m2 DDY3591 S288C This study leu2::NAT ura3::HPHMX cdc13::KANMX YEp-URA3-CDC13 pRAD52-TRP Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3534 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3536 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3535 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-cdc13-L91A Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3537 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG18-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-cdc13-L91A Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3538 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3540 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3539 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pRS414-cdc13-L91A

147

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3541 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPHMX cdc13::KANMX pif1-m2 pRS414-cdc13-L91A Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3520 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KANMX YEP24-CDC13 pRAD52 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3522 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KANMX YEP24-CDC13 pRAD52 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3528 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KANMX YEP24-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3530 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KANMX YEP24-CDC13 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3584 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH leu2::NAT cdc13::KANMX YEP24-CDC13 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3586 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH leu2::NAT cdc13::KANMX YEP24-CDC13 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3589 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH leu2::NAT cdc13::KANMX YEP24-CDC13 pRAD52 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 DDY3591 S288C This study ura3::HPH leu2::NAT cdc13::KANMX YEP24-CDC13 pRAD52 pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3248 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG96-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3250 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG119-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3252 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG142-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3133 S288C This study Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG162-HOcs-LYS2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3134 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::URA3-TG162-HOcs-LYS2 This study pif1-m2 Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3529 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KAN YEP24-CDC13 DDY3614 S288C DDY3529 leu2::NATMX This study Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3531 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KAN YEP24-CDC13 pif1-m2 DDY3615 S288C DDY3531 leu2::NATMX This study Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3614 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KAN leu2::NAT YEP24-CDC13

148

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY3615 S288C Δ1::GAL1:HO-LEU2 rad52::HIS VII-L::TG34-HOcs-LYS2 This study ura3::HPH cdc13::KAN leu2::NAT YEP24-CDC13 pif1-m2 DDY3768 S288C DDY3614 pRS425-CDC13 This study DDY3769 S288C DDY3614 pRS425-cdc13-L91A This study DDY3770 S288C DDY3614 pRS425-cdc13-F236S This study DDY3771 S288C DDY3614 pRS425-cdc13-Q256H This study DDY3772 S288C DDY3614 pRS425-cdc13-Q583K This study DDY3773 S288C DDY3614 pRS425-cdc13-I87N This study DDY3774 S288C DDY3614 pRS425-cdc13-Y758N This study DDY3775 S288C DDY3614 pRS425-cdc13-H12R This study DDY3776 S288C DDY3614 pRS425-cdc13-E556V/N567D This study DDY3777 S288C DDY3614 pRS425-cdc13-F728I This study DDY3778 S288C DDY3614 pRS425-cdc13-E252K This study DDY3779 S288C DDY3614 pRS425-cdc13-P235S This study DDY3780 S288C DDY3614 pRS425-cdc13-S255A This study DDY3781 S288C DDY3614 pRS425-cdc13-K50Q This study DDY3782 S288C DDY3614 pRS425-cdc13-F237V This study DDY3783 S288C DDY3615 pRS425-CDC13 This study DDY3784 S288C DDY3615 pRS425-cdc13-L91A This study DDY3785 S288C DDY3615 pRS425-cdc13-F236S This study DDY3786 S288C DDY3615 pRS425-cdc13-Q256H This study DDY3787 S288C DDY3615 pRS425-cdc13-Q583K This study DDY3788 S288C DDY3615 pRS425-cdc13-I87N This study DDY3789 S288C DDY3615 pRS425-cdc13-Y758N This study DDY3790 S288C DDY3615 pRS425-cdc13-H12R This study DDY3791 S288C DDY3615 pRS425-cdc13-E556V/N567D This study DDY3792 S288C DDY3615 pRS425-cdc13-F728I This study DDY3793 S288C DDY3615 pRS425-cdc13-E252K This study DDY3794 S288C DDY3615 pRS425-cdc13-P235S This study DDY3795 S288C DDY3615 pRS425-cdc13-S255A This study DDY3796 S288C DDY3615 pRS425-cdc13-K50Q This study DDY3797 S288C DDY3615 pRS425-cdc13-F237V This study MATalpha Δhml::ADE1 Δhmr::ADE1 ade1-110 leu2-3,112 (Oza et al., DDY2784 JKM179 lys5trp1::hisG ura3::CUP1-GFP-LAC1-URA3 NUP49-GFP- 2009) NATMX ARS313-LACOR-TRP1 ade3::GAL:HO MATalpha Δhml::ADE1 Δhmr::ADE1 ade1-110 leu2-3,112 DDY3165 JKM179 lys5trp1::hisG ura3::CUP1-GFP-LAC1-URA3 NUP49-mCherry- This study HPHMX ARS313-LACOR-TRP1 ade3::GAL:HO DDY3940 JKM179 DDY3165 AUR1::CEP3 cep3::NATMX This study DDY3941 JKM179 DDY3165 AUR1::cep3-S575A cep3::NATMX This study DDY3954 JKM179 DDY3165 AUR1::CBF1 cbf1::NATMX This study DDY3955 JKM179 DDY3165 AUR1::cbf1-A45/48A cbf1::NATMX This study DDY3956 JKM179 DDY3165 AUR1::BIR1 bir1::NATMX This study DDY3957 JKM179 DDY3165 AUR1::bir1-S751A bir1::NATMX This study DDY3958 JKM179 DDY3165 AUR1::DAD1 dad1::NATMX This study DDY3959 JKM179 DDY3165 AUR1::dad1-S89A dad1::NATMX This study DDY3960 JKM179 DDY3165 AUR1::DAD3 dad3::NATMX This study DDY4309 JKM179 DDY3165 AUR1::dad3-S43A dad3::NATMX This study DDY4310 JKM179 DDY3165 AUR1::Glc7 glc7::NATMX This study

149

DDY4311 JKM179 DDY3165 AUR1::glc7-S3A glc7::NATMX This study DDY4117 JKM179 DDY3165 sml1::NATMX This study DDY4119 JKM179 DDY3165 sml1::NATMX mec1::KANMX This study DDY4121 JKM179 DDY3165 sml1::NATMX rad53::KANMX This study DDY4355 JKM179 DDY3165 sml1::NATMX mec1::LEU2 tel1::KANMX This study DDY4356 JKM179 DDY3165 sml1::NATMX tel1::KANMX This study DDY4160 JKM179 DDY3165 dun1::KANMX This study DDY4254 JKM179 DDY3165 CEP3-13xMyc::NATMX This study DDY4256 JKM179 DDY3165 cep3-S575A-13xMyc::NATMX This study DDY4142 JKM179 DDY3165 arp8::NATMX This study DDY4165 JKM179 DDY3165 chk1::KANMX This study DDY4163 JKM179 DDY3165 ura3::LEU2 This study DDY4069 JKM179 DDY3165 ura3::LEU2 cep3-S575A This study DDY4201 JKM179 DDY3165 HOcs::LEU2 pGAL1/10-CEN3::KANMX This study DDY4204 JKM179 DDY3165 HOcs::LEU2 pGAL1/10-CEN3::KANMX sir4::NATMX This study DDY4173 JKM179 DDY3165 sir4::NATMX This study DDY4357 JKM179 DDY3165 yku70::KANMX This study DDY4358 JKM179 DDY3165 sir4::NATMX yku70::KANMX This study DDY4272 JKM179 DDY3165 pGAL1/10-CEN3::KANMX This study DDY4243 JKM179 DDY3165 AUR1::CEP3 cep3::NATMX mad2::KANMX This study DDY4169 JKM179 DDY3165 ura3::LEU2 cep3-S575A chk1::KANMX This study DDY4245 JKM179 DDY3165 AUR1::cep3-S575A cep3::NATMX mad2::KANMX This study DDY4334 JKM179 DDY3165 mad2::NATMX chk1::KANMX This study DDY4335 JKM179 DDY3165 ura3::LEU2 cep3-S575A mad2::NATMX chk1::KANMX This study DDY3165 AUR1::cep3-S575A cep3::NATMX pGAL1/10- DDY4135 JKM179 This study CEN3::KANMX DDY3165 AUR1::cep3-S575A cep3::NATMX pGAL1/10- DDY4137 JKM179 This study CEN5::KANMX DDY4138 JKM179 DDY3165 arp8::NATMX pGAL1/10-CEN3::KANMX This study DDY4140 JKM179 DDY3165 arp8::NATMX pGAL1/10-CEN5::KANMX This study DDY4150 W303 MATa CEP3::KANMX This study DDY4152 W303 MATa cep3-S575A::KANMX This study DDY4231 W303 MATa CEP3::KANMX HOcs::LEU2 This study DDY4233 W303 MATa cep3-S575A::KANMX HOcs::LEU2 This study DDY4064 Diploid DDY4231/DDY3165 This study DDY4065 Diploid DDY4233/DDY4069 This study DDY4336 JKM179 DDY3165 ura3::LEU2 cep3-S575A sml1::KANMX rad53::NATMX This study Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- DDY4018 S288C Δ1::GAL1:HO-LacI-GFP::KANMX Nup49-mCherry::HPHMX This study rad52::HIS pRAD52-TRP DDY4027 S288C DDY4018 ChrIV:476896-HOcs::URA3 ChrV:53601-LacO::LEU2 This study DDY4030 S288C DDY4018 ChrIV:551206-HOcs::URA3 ChrV:53601-LacO::LEU2 This study DDY4037 S288C DDY4018 ChrIV:960174-HOcs::URA3 ChrV:53601-LacO::LEU2 This study DDY4040 S288C DDY4018 ChrIV:1447374-HOcs::URA3 ChrV:53601-LacO::LEU2 This study DDY4028 S288C DDY4027 cep3-S575A::NATMX This study DDY4345 S288C DDY4027 sir4::NATMX This study DDY4346 S288C DDY4018 ChrIV:476896-HOcs::URA3 pRS415-LacOx256 This study DDY4347 S288C DDY4346 cep3-S575A::NATMX This study DDY4348 S288C DDY4018 ChrIV:1447374-HOcs::URA3 pRS415-LacOx256 This study DDY4101 S288C This study

150

Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- Δ1::GAL1:HO::NATMX Nup49-mCherry::HPHMX rad52::HIS pRAD52-TRP DDY4349 S288C DDY4101 AMD2::HOcs pRS415-RAD52-YFP This study DDY4101 AMD2::HOcs cep3-S575A::NATMX pRS415-RAD52- DDY4350 S288C This study YFP DDY4351 S288C DDY4101 ADH4::HOcs pRS415-RAD52-YFP This study DDY4352 S288C DDY4101 ADH4::HOcs cep3-S575A::NATMX pRS415-RAD52-YFP This study MATα ade2-101::NATMX CFIII(CEN3.L)URA3 (Yuen et DDY4323 YKY53 SUPP11mfa1∆::MFA1pr-HIS3 can1∆ his3∆1 leu2∆0 ura3∆0 al., 2007) MET15+ lys2∆0 DDY4325 YKY53 YKY53 AUR1::CEP3 cep3::KANMX This study DDY4328 YKY53 YKY53 AUR1::cep3-S575A cep3::KANMX This study DDY4331 YKY53 YKY53 AUR1::CEP3 cep3::KANMX chk1::HPHMX This study DDY4332 YKY53 YKY53 AUR1::cep3-S575A cep3::KANMX chk1::HPHMX This study DDY4324 YKY53 YKY53 cin8::KANMX This study MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA60 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA68 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA70 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA77 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA87 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA89 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA92 MK225 100 al., 2013) MATa-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1- (Agmon et NA95 MK225 100 al., 2013) DDY4074 MK225 NA60 AUR1::CEP3 cep3::NATMX This study DDY4076 MK225 NA68 AUR1::CEP3 cep3::NATMX This study DDY4078 MK225 NA70 AUR1::CEP3 cep3::NATMX This study DDY4080 MK225 NA77 AUR1::CEP3 cep3::NATMX This study DDY4215 MK225 NA87 AUR1::CEP3 cep3::NATMX This study DDY4219 MK225 NA89 AUR1::CEP3 cep3::NATMX This study DDY4223 MK225 NA92 AUR1::CEP3 cep3::NATMX This study DDY4227 MK225 NA95 AUR1::CEP3 cep3::NATMX This study DDY4082 MK225 NA60 AUR1::cep3-S575A cep3::NATMX This study DDY4084 MK225 NA68 AUR1::cep3-S575A cep3::NATMX This study DDY4086 MK225 NA70 AUR1::cep3-S575A cep3::NATMX This study DDY4088 MK225 NA77 AUR1::cep3-S575A cep3::NATMX This study DDY4217 MK225 NA87 AUR1::cep3-S575A cep3::NATMX This study DDY4221 MK225 NA89 AUR1::cep3-S575A cep3::NATMX This study DDY4225 MK225 NA92 AUR1::cep3-S575A cep3::NATMX This study DDY4229 MK225 NA95 AUR1::cep3-S575A cep3::NATMX This study DDY4337 MK225 NA60 arp8::NATMX This study DDY4338 MK225 NA68 arp8::NATMX This study

151

DDY4339 MK225 NA70 arp8::NATMX This study DDY4340 MK225 NA77 arp8::NATMX This study DDY4341 MK225 NA87 arp8::NATMX This study DDY4342 MK225 NA89 arp8::NATMX This study DDY4343 MK225 NA92 arp8::NATMX This study DDY4344 MK225 NA95 arp8::NATMX This study MATα ade2-1 trp1-1 can1-100 leu2-3,112 his3-11,15 ura3-1 GAL+ DDY4362 W303 This study psi+ ssd1-d2 DDY4363 W303 DDY4362 cep3-S575A This study DDY4364 W303 DDY4362 chk1::KANMX This study DDY4365 W303 DDY4362 cep3-S575A chk1::KANMX This study YPH316 YPH316 MATα his1 (Yuen et DDY4032 S288C DDY4018 ChrIV:551206-HOcs::URA3 ChrIV:539110-LacO::LEU2 al., 2007) DDY4039 S288C DDY4018 ChrIV:960174-HOcs::URA3 ChrIV:946605-LacO::LEU2 This study DDY4018 ChrIV:1447374-HOcs::URA3 ChrIV:1434567- DDY4042 S288C This study LacO::LEU2 DDY4018 ChrIV::551206-HOcs::URA3 ChrIV:351224- DDY4368 S288C This study LacO::LEU2 DDY4369 S288C DDY4368 cep3-S575A::NATMX This study DDY4370 S288C DDY4368 sir4::NATMX This study ura3-1::TUB1-CFP::URA3 leu2-3,112 his3-11 trp1-1 can1-100 (Pinsky et DDY4359 SBY4340 ade2-1 bar1-1 MTW1-3xGFP::HIS3 al., 2006) ura3-1::TUB1-CFP::URA3 leu2-3,112 his3-11 trp1-1 can1-100 (Pinsky et DDY4449 SBY4341 ade2-1 bar1-1 MTW1-3xGFP::HIS3 ndc80-1 al., 2006) DDY4360 W303 SBY4340 AUR1::CEP3 cep3::NATMX This study DDY4361 W303 SBY4340 AUR1::cep3-S575A cep3::NATMX This study Mata-inc ura3-52 lys2-801 ade2-101 ochre trp1-Δ63 his3-Δ200 leu2- Δ1::GAL1:HO-LacI-GFP::KANMX Spc42-tdimer2::HPHMX DDY4371 S288C This study rad52::HIS pRAD52-TRP ChrIV:476896-HOcs::URA3 CEN5::LacO::LEU2 DDY4372 S288C DDY4371 cep3-S575A::NAT This study DDY4034 S288C DDY4018 ChrIV:551206-HOcs::URA3 ChrIV:946605-LacO::LEU2 This study DDY4018 ChrIV:551206-HOcs::URA3 ChrIV:946605-LacO::LEU2 DDY4466 S288C This study cep3-S575A::NATMX DDY4018 ChrIV:551206-HOcs::URA3 ChrIV:946605-LacO::LEU2 DDY4474 S288C This study sml1::NATMX mec1::LYS2 DDY4018 ChrIV:551206-HOcs::URA3 ChrIV:946605-LacO::LEU2 DDY4465 S288C This study sir4::NATMX DDY4112 JKM179 DDY3165 hos1::KATMX This study DDY4113 JKM179 DDY3165 hos2::KATMX This study DDY4114 JKM179 DDY3165 hos3::KATMX This study DDY4111 JKM179 DDY3165 hda1::KATMX This study DDY4070 JKM179 DDY3165 rpd3::KATMX This study DDY4107 JKM179 DDY3165 sin3::KATMX This study DDY4115 JKM179 DDY3165 sir2::NATMX This study DDY4116 JKM179 DDY3165 hst4::NATMX This study DDY4183 JKM179 DDY3165 rxt3::NATMX This study DDY4181 JKM179 DDY3165 sds3::NATMX This study DDY4185 JKM179 DDY3165 rco1::NATMX This study DDY4105 AH109 AH109 CEP3::NAT pGBK-CEP3 pGAD-RXT3 This study

152

DDY4106 AH109 AH109 cep3-S575A::NAT pGBK-cep3-S575A pGAD-RXT3 This study DDY4197 S288C DDY3165 AUR1-CEP3 cep3::NATMX RXT3-13xMyc::KAN This study DDY4056 JKM179 DDY3165 AUR1::htz1-4KQ htz1::NATMX This study DDY4050 JKM179 DDY3165 AUR1::htz1-K14Q htz1::NATMX This study DDY4052 JKM179 DDY3165 AUR1::HTZ1 htz1::NATMX This study DDY4072 JKM179 DDY3165 cep3-S575A hda1::NATMX This study DDY4073 JKM179 DDY3165 hda1::NATMX rpd3::KANMX This study DDY4162 JKM179 DDY3165 cep3-S575E cep3::NATMX This study DDY4054 JKM179 DDY3165 htz1-K14R htz1::NATMX This study DDY4057 JKM179 DDY3165 htz1-4KR htz1::NATMX This study DDY4062 JKM179 DDY3165 htz1-K14R htz1::NATMX sir4::KANMX This study DDY4099 JKM179 DDY3165 cep3-S575A htz1-4KR htz1::NATMX This study DDY4206 JKM179 DDY3165 AUR1::OsTir1 This study DDY4208 JKM179 DDY3165 AUR1::OsTir1 IPL1-3xAID::KANMX This study DDY4212 JKM179 DDY3165 AUR1::IPL1 ipl1::NATMX This study DDY4213 JKM179 DDY3165 AUR1::ipl1-as6 ipl1::NATMX This study DDY3166 JKM179 DDY3165 cik1::NATMX This study DDY3659 JKM179 DDY3165 kar3::NATMX This study DDY3827 JKM179 DDY3165 vik1::NATMX This study DDY3883 JKM179 DDY3165 cin8::KATMX This study DDY3661 JKM179 DDY3165 kar3::NATMX pRS415-KAR3 This study DDY3662 JKM179 DDY3165 kar3::NATMX pRS415-kar3-898 This study DDY3663 JKM179 DDY3165 kar3::NATMX pRS415 control This study DDY4127 MK225 NA60 cik1::NATMX This study DDY4128 MK225 NA68 cik1::NATMX This study DDY4129 MK225 NA70 cik1::NATMX This study DDY4130 MK225 NA77 cik1::NATMX This study Mata::HOcsDEL::hisG ura3 DEL851 trp1DEL63 (Lydeard sup53DEL::leu2DEL::NATMX hmlDEL::hisG hmrDEL::ADE3 DDY3393 JRL346 et al., ade3::GAL10::HO can1,1-1446::HOcs::HPH::DEL AVT2 2007) ykl215c::leu2::hisG::can1DEL1-289 DDY3398 JRL346 DDY3393 pol32::KANMX This study DDY3394 JRL346 DDY3393 cik1::KANMX This study DDY3396 JRL346 DDY3393 kar3::KANMX This study DDY4422 JRL346 DDY3393 cep3-S575A This study DDY4013 JRL346 DDY3393 kar3::KANMX pRS414 This study DDY4014 JRL346 DDY3393 kar3::KANMX pRS414-KAR3 This study DDY4015 JRL346 DDY3393 kar3::KANMX pRS414-kar3-1 This study DDY4016 JRL346 DDY3393 kar3::KANMX pRS414-kar3-898 This study DDY4296 JRL346 DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP This study DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP DDY4297 JRL346 This study cik1::KANMX DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP DDY4298 JRL346 This study kar3::KANMX DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP DDY4299 JRL346 This study kar3::KANMX pRS414-KAR3 DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP DDY4300 JRL346 This study kar3::KANMX pRS414-kar3-1 DDY3393 Nup49-mCherry::URA3 pRS415-Rad52-YFP DDY4301 JRL346 This study kar3::KANMX pRS414

References Abney, J.R., Cutler, B., Fillbach, M.L., Axelrod, D., and Scalettar, B.A. (1997). Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J Cell Biol 137, 1459-1468.

Addinall, S.G., Downey, M., Yu, M., Zubko, M.K., Dewar, J., Leake, A., Hallinan, J., Shaw, O., James, K., Wilkinson, D.J., et al. (2008). A genomewide suppressor and enhancer analysis of cdc13-1 reveals varied cellular processes influencing telomere capping in Saccharomyces cerevisiae. Genetics 180, 2251-2266.

Agarwal, R., Tang, Z., Yu, H., and Cohen-Fix, O. (2003). Two distinct pathways for inhibiting pds1 ubiquitination in response to DNA damage. J Biol Chem 278, 45027-45033.

Agmon, N., Liefshitz, B., Zimmer, C., Fabre, E., and Kupiec, M. (2013). Effect of nuclear architecture on the efficiency of double-strand break repair. Nat Cell Biol 15, 694-699.

Ahnesorg, P., and Jackson, S.P. (2007). The non-homologous end-joining protein Nej1p is a target of the DNA damage checkpoint. DNA Repair (Amst) 6, 190-201.

Alcasabas, A.A., Osborn, A.J., Bachant, J., Hu, F., Werler, P.J., Bousset, K., Furuya, K., Diffley, J.F., Carr, A.M., and Elledge, S.J. (2001). Mrc1 transduces signals of DNA replication stress to activate Rad53. Nat Cell Biol 3, 958-965.

Anbalagan, S., Bonetti, D., Lucchini, G., and Longhese, M.P. (2011). Rif1 supports the function of the CST complex in yeast telomere capping. PLoS Genet 7, e1002024.

Andrulis, E.D., Zappulla, D.C., Ansari, A., Perrod, S., Laiosa, C.V., Gartenberg, M.R., and Sternglanz, R. (2002). Esc1, a nuclear periphery protein required for Sir4-based plasmid anchoring and partitioning. Mol Cell Biol 22, 8292-8301.

Arneric, M., and Lingner, J. (2007). Tel1 kinase and subtelomere-bound Tbf1 mediate preferential elongation of short telomeres by telomerase in yeast. EMBO Rep 8, 1080-1085.

Aten, J.A., Stap, J., Krawczyk, P.M., van Oven, C.H., Hoebe, R.A., Essers, J., and Kanaar, R. (2004). Dynamics of DNA double-strand breaks revealed by clustering of damaged chromosome domains. Science 303, 92-95.

Aylon, Y., and Kupiec, M. (2004). DSB repair: the yeast paradigm. DNA Repair (Amst) 3, 797-815.

Bahmed, K., Nitiss, K.C., and Nitiss, J.L. (2010). Yeast Tdp1 regulates the fidelity of nonhomologous end joining. Proc Natl Acad Sci U S A 107, 4057-4062.

Bandyopadhyay, S., Mehta, M., Kuo, D., Sung, M.K., Chuang, R., Jaehnig, E.J., Bodenmiller, B., Licon, K., Copeland, W., Shales, M., et al. (2010). Rewiring of genetic networks in response to DNA damage. Science 330, 1385-1389.

Barlow, J.H., Faryabi, R.B., Callen, E., Wong, N., Malhowski, A., Chen, H.T., Gutierrez-Cruz, G., Sun, H.W., McKinnon, P., Wright, G., et al. (2013). Identification of early replicating fragile sites that contribute to genome instability. Cell 152, 620-632.

Baroin, A., Prat, A., and Caron, F. (1987). Telomeric site position heterogeneity in macronuclear DNA of Paramecium primaurelia. Nucleic Acids Res 15, 1717-1728.

153

154

Bartsch, S., Kang, L.E., and Symington, L.S. (2000). RAD51 is required for the repair of plasmid double- stranded DNA gaps from either plasmid or chromosomal templates. Mol Cell Biol 20, 1194-1205.

Bashkirov, V.I., Bashkirova, E.V., Haghnazari, E., and Heyer, W.D. (2003). Direct kinase-to-kinase signaling mediated by the FHA phosphoprotein recognition domain of the Dun1 DNA damage checkpoint kinase. Mol Cell Biol 23, 1441-1452.

Bennett, C.B., Lewis, A.L., Baldwin, K.K., and Resnick, M.A. (1993). Lethality induced by a single site- specific double-strand break in a dispensable yeast plasmid. Proc Natl Acad Sci U S A 90, 5613-5617.

Bennetzen, M.V., Larsen, D.H., Bunkenborg, J., Bartek, J., Lukas, J., and Andersen, J.S. (2010). Site- specific phosphorylation dynamics of the nuclear proteome during the DNA damage response. Mol Cell Proteomics 9, 1314-1323.

Bensimon, A., Schmidt, A., Ziv, Y., Elkon, R., Wang, S.Y., Chen, D.J., Aebersold, R., and Shiloh, Y. (2010). ATM-dependent and -independent dynamics of the nuclear phosphoproteome after DNA damage. Sci Signal 3, rs3.

Berg, H.C. (1993). Random Walks in Biology (Princeton, NJ: Princeton University Press).

Berger, A.B., Cabal, G.G., Fabre, E., Duong, T., Buc, H., Nehrbass, U., Olivo-Marin, J.C., Gadal, O., and Zimmer, C. (2008). High-resolution statistical mapping reveals gene territories in live yeast. Nat Methods 5, 1031-1037.

Betermier, M., Bertrand, P., and Lopez, B.S. (2014). Is non-homologous end-joining really an inherently error-prone process? PLoS Genet 10, e1004086.

Bianchi, A., Negrini, S., and Shore, D. (2004). Delivery of yeast telomerase to a DNA break depends on the recruitment functions of Cdc13 and Est1. Mol Cell 16, 139-146.

Bielas, J.H., Loeb, K.R., Rubin, B.P., True, L.D., and Loeb, L.A. (2006). Human cancers express a mutator phenotype. Proc Natl Acad Sci U S A 103, 18238-18242.

Biessmann, H., Mason, J.M., Ferry, K., d'Hulst, M., Valgeirsdottir, K., Traverse, K.L., and Pardue, M.L. (1990). Addition of telomere-associated HeT DNA sequences "heals" broken chromosome ends in Drosophila. Cell 61, 663-673.

Biggins, S. (2013). The composition, functions, and regulation of the budding yeast kinetochore. Genetics 194, 817-846.

Biggins, S., Severin, F.F., Bhalla, N., Sassoon, I., Hyman, A.A., and Murray, A.W. (1999). The conserved protein kinase Ipl1 regulates microtubule binding to kinetochores in budding yeast. Genes Dev 13, 532- 544.

Blackburn, E.H., and Gall, J.G. (1978). A tandemly repeated sequence at the termini of the extrachromosomal ribosomal RNA genes in Tetrahymena. J Mol Biol 120, 33-53.

Blanco, M.G., Matos, J., and West, S.C. (2014). Dual control of Yen1 nuclease activity and cellular localization by Cdk and Cdc14 prevents genome instability. Mol Cell 54, 94-106.

Blier, P.R., Griffith, A.J., Craft, J., and Hardin, J.A. (1993). Binding of Ku protein to DNA. Measurement of affinity for ends and demonstration of binding to nicks. J Biol Chem 268, 7594-7601.

155

Bloom, K.S., and Carbon, J. (1982). Yeast centromere DNA is in a unique and highly ordered structure in chromosomes and small circular minichromosomes. Cell 29, 305-317.

Bochkareva, E., Korolev, S., Lees-Miller, S.P., and Bochkarev, A. (2002). Structure of the RPA trimerization core and its role in the multistep DNA-binding mechanism of RPA. EMBO J 21, 1855-1863.

Bochman, M.L., Sabouri, N., and Zakian, V.A. (2010). Unwinding the functions of the Pif1 family helicases. DNA Repair (Amst) 9, 237-249.

Boddy, M.N., Gaillard, P.H., McDonald, W.H., Shanahan, P., Yates, J.R., 3rd, and Russell, P. (2001). Mus81-Eme1 are essential components of a Holliday junction resolvase. Cell 107, 537-548.

Bonetti, D., Clerici, M., Anbalagan, S., Martina, M., Lucchini, G., and Longhese, M.P. (2010). Shelterin- like proteins and Yku inhibit nucleolytic processing of Saccharomyces cerevisiae telomeres. PLoS Genet 6, e1000966.

Bonilla, C.Y., Melo, J.A., and Toczyski, D.P. (2008). Colocalization of sensors is sufficient to activate the DNA damage checkpoint in the absence of damage. Mol Cell 30, 267-276.

Bouck, D., and Bloom, K. (2005). The role of centromere-binding factor 3 (CBF3) in spindle stability, cytokinesis, and kinetochore attachment. Biochem Cell Biol 83, 696-702.

Boule, J.B., Vega, L.R., and Zakian, V.A. (2005). The yeast Pif1p helicase removes telomerase from telomeric DNA. Nature 438, 57-61.

Boulton, S.J., and Jackson, S.P. (1996a). Identification of a Saccharomyces cerevisiae Ku80 homologue: roles in DNA double strand break rejoining and in telomeric maintenance. Nucleic Acids Res 24, 4639- 4648.

Boulton, S.J., and Jackson, S.P. (1996b). Saccharomyces cerevisiae Ku70 potentiates illegitimate DNA double-strand break repair and serves as a barrier to error-prone DNA repair pathways. EMBO J 15, 5093- 5103.

Boulton, S.J., and Jackson, S.P. (1998). Components of the Ku-dependent non-homologous end-joining pathway are involved in telomeric length maintenance and telomeric silencing. EMBO J 17, 1819-1828.

Boveri, T., ed. (1914). Zur Frage der Entstehung Maligner Tumoren.

Brill, S.J., and Stillman, B. (1989). Yeast replication factor-A functions in the unwinding of the SV40 origin of DNA replication. Nature 342, 92-95.

Budd, M.E., Reis, C.C., Smith, S., Myung, K., and Campbell, J.L. (2006). Evidence suggesting that Pif1 helicase functions in DNA replication with the Dna2 helicase/nuclease and DNA polymerase delta. Mol Cell Biol 26, 2490-2500.

Bunting, S.F., Callen, E., Wong, N., Chen, H.T., Polato, F., Gunn, A., Bothmer, A., Feldhahn, N., Fernandez-Capetillo, O., Cao, L., et al. (2010). 53BP1 inhibits homologous recombination in Brca1- deficient cells by blocking resection of DNA breaks. Cell 141, 243-254.

Bunting, S.F., and Nussenzweig, A. (2013). End-joining, translocations and cancer. Nat Rev Cancer 13, 443-454.

156

Bupp, J.M., Martin, A.E., Stensrud, E.S., and Jaspersen, S.L. (2007). Telomere anchoring at the nuclear periphery requires the budding yeast Sad1-UNC-84 domain protein Mps3. J Cell Biol 179, 845-854.

Bystricky, K., Heun, P., Gehlen, L., Langowski, J., and Gasser, S.M. (2004). Long-range compaction and flexibility of interphase chromatin in budding yeast analyzed by high-resolution imaging techniques. Proc Natl Acad Sci U S A 101, 16495-16500.

Bystricky, K., Van Attikum, H., Montiel, M.D., Dion, V., Gehlen, L., and Gasser, S.M. (2009). Regulation of nuclear positioning and dynamics of the silent mating type loci by the yeast Ku70/Ku80 complex. Mol Cell Biol 29, 835-848.

Cabal, G.G., Genovesio, A., Rodriguez-Navarro, S., Zimmer, C., Gadal, O., Lesne, A., Buc, H., Feuerbach- Fournier, F., Olivo-Marin, J.C., Hurt, E.C., et al. (2006). SAGA interacting factors confine sub-diffusion of transcribed genes to the nuclear envelope. Nature 441, 770-773.

Cai, M., and Davis, R.W. (1990). Yeast centromere binding protein CBF1, of the helix-loop-helix protein family, is required for chromosome stability and methionine prototrophy. Cell 61, 437-446.

Cai, M.J., and Davis, R.W. (1989). Purification of a yeast centromere-binding protein that is able to distinguish single base-pair mutations in its recognition site. Mol Cell Biol 9, 2544-2550.

Cannavo, E., and Cejka, P. (2014). Sae2 promotes dsDNA endonuclease activity within Mre11-Rad50- Xrs2 to resect DNA breaks. Nature 514, 122-125.

Carmena, M., Wheelock, M., Funabiki, H., and Earnshaw, W.C. (2012). The chromosomal passenger complex (CPC): from easy rider to the godfather of mitosis. Nat Rev Mol Cell Biol 13, 789-803.

Carrozza, M.J., Florens, L., Swanson, S.K., Shia, W.J., Anderson, S., Yates, J., Washburn, M.P., and Workman, J.L. (2005a). Stable incorporation of sequence specific repressors Ash1 and Ume6 into the Rpd3L complex. Biochim Biophys Acta 1731, 77-87; discussion 75-76.

Carrozza, M.J., Li, B., Florens, L., Suganuma, T., Swanson, S.K., Lee, K.K., Shia, W.J., Anderson, S., Yates, J., Washburn, M.P., et al. (2005b). Histone H3 methylation by Set2 directs deacetylation of coding regions by Rpd3S to suppress spurious intragenic transcription. Cell 123, 581-592.

Ceccaldi, R., Liu, J.C., Amunugama, R., Hajdu, I., Primack, B., Petalcorin, M.I., O'Connor, K.W., Konstantinopoulos, P.A., Elledge, S.J., Boulton, S.J., et al. (2015). Homologous-recombination-deficient tumours are dependent on Poltheta-mediated repair. Nature 518, 258-262.

Cejka, P., Cannavo, E., Polaczek, P., Masuda-Sasa, T., Pokharel, S., Campbell, J.L., and Kowalczykowski, S.C. (2010a). DNA end resection by Dna2-Sgs1-RPA and its stimulation by Top3-Rmi1 and Mre11-Rad50- Xrs2. Nature 467, 112-116.

Cejka, P., Plank, J.L., Bachrati, C.Z., Hickson, I.D., and Kowalczykowski, S.C. (2010b). Rmi1 stimulates decatenation of double Holliday junctions during dissolution by Sgs1-Top3. Nat Struct Mol Biol 17, 1377- 1382.

Cejka, P., Plank, J.L., Dombrowski, C.C., and Kowalczykowski, S.C. (2012). Decatenation of DNA by the S. cerevisiae Sgs1-Top3-Rmi1 and RPA complex: a mechanism for disentangling chromosomes. Mol Cell 47, 886-896.

157

Chambers, A.L., Ormerod, G., Durley, S.C., Sing, T.L., Brown, G.W., Kent, N.A., and Downs, J.A. (2012). The INO80 chromatin remodeling complex prevents polyploidy and maintains normal chromatin structure at centromeres. Genes Dev 26, 2590-2603.

Chandra, A., Hughes, T.R., Nugent, C.I., and Lundblad, V. (2001). Cdc13 both positively and negatively regulates telomere replication. Genes Dev 15, 404-414.

Chang, M., Arneric, M., and Lingner, J. (2007). Telomerase repeat addition processivity is increased at critically short telomeres in a Tel1-dependent manner in Saccharomyces cerevisiae. Genes Dev 21, 2485- 2494.

Chao, W.C., Kulkarni, K., Zhang, Z., Kong, E.H., and Barford, D. (2012). Structure of the mitotic checkpoint complex. Nature 484, 208-213.

Chapman, J.R., Taylor, M.R., and Boulton, S.J. (2012). Playing the end game: DNA double-strand break repair pathway choice. Mol Cell 47, 497-510.

Cheeseman, I.M., Enquist-Newman, M., Muller-Reichert, T., Drubin, D.G., and Barnes, G. (2001). Mitotic spindle integrity and kinetochore function linked by the Duo1p/Dam1p complex. J Cell Biol 152, 197-212.

Chehab, N.H., Malikzay, A., Appel, M., and Halazonetis, T.D. (2000). Chk2/hCds1 functions as a DNA damage checkpoint in G(1) by stabilizing p53. Genes Dev 14, 278-288.

Chen, C., Umezu, K., and Kolodner, R.D. (1998). Chromosomal rearrangements occur in S. cerevisiae rfa1 mutator mutants due to mutagenic lesions processed by double-strand-break repair. Mol Cell 2, 9-22.

Chen, L., Trujillo, K., Ramos, W., Sung, P., and Tomkinson, A.E. (2001a). Promotion of Dnl4-catalyzed DNA end-joining by the Rad50/Mre11/Xrs2 and Hdf1/Hdf2 complexes. Mol Cell 8, 1105-1115.

Chen, L.Y., Redon, S., and Lingner, J. (2012). The human CST complex is a terminator of telomerase activity. Nature 488, 540-544.

Chen, Q., Ijpma, A., and Greider, C.W. (2001b). Two survivor pathways that allow growth in the absence of telomerase are generated by distinct telomere recombination events. Mol Cell Biol 21, 1819-1827.

Chen, S.H., Albuquerque, C.P., Liang, J., Suhandynata, R.T., and Zhou, H. (2010). A proteome-wide analysis of kinase-substrate network in the DNA damage response. J Biol Chem 285, 12803-12812.

Chen, S.H., Smolka, M.B., and Zhou, H. (2007). Mechanism of Dun1 activation by Rad53 phosphorylation in Saccharomyces cerevisiae. J Biol Chem 282, 986-995.

Chen, X., and Tomkinson, A.E. (2011). Yeast Nej1 is a key participant in the initial end binding and final ligation steps of nonhomologous end joining. J Biol Chem 286, 4931-4940.

Chen, Z., Yang, H., and Pavletich, N.P. (2008). Mechanism of homologous recombination from the RecA- ssDNA/dsDNA structures. Nature 453, 489-484.

Cho, N.W., Dilley, R.L., Lampson, M.A., and Greenberg, R.A. (2014). Interchromosomal homology searches drive directional ALT telomere movement and synapsis. Cell 159, 108-121.

Choy, J.S., Acuna, R., Au, W.C., and Basrai, M.A. (2011). A role for histone H4K16 hypoacetylation in Saccharomyces cerevisiae kinetochore function. Genetics 189, 11-21.

158

Chubb, J.R., Boyle, S., Perry, P., and Bickmore, W.A. (2002). Chromatin motion is constrained by association with nuclear compartments in human cells. Curr Biol 12, 439-445.

Chung, W.H., Zhu, Z., Papusha, A., Malkova, A., and Ira, G. (2010). Defective resection at DNA double- strand breaks leads to de novo telomere formation and enhances gene targeting. PLoS Genet 6, e1000948.

Ciosk, R., Zachariae, W., Michaelis, C., Shevchenko, A., Mann, M., and Nasmyth, K. (1998). An ESP1/PDS1 complex regulates loss of sister chromatid cohesion at the metaphase to anaphase transition in yeast. Cell 93, 1067-1076.

Clarke, L., and Carbon, J. (1980). Isolation of a yeast centromere and construction of functional small circular chromosomes. Nature 287, 504-509.

Clemenson, C., and Marsolier-Kergoat, M.C. (2006). The spindle assembly checkpoint regulates the phosphorylation state of a subset of DNA checkpoint proteins in Saccharomyces cerevisiae. Mol Cell Biol 26, 9149-9161.

Clerici, M., Mantiero, D., Guerini, I., Lucchini, G., and Longhese, M.P. (2008). The Yku70-Yku80 complex contributes to regulate double-strand break processing and checkpoint activation during the cell cycle. EMBO Rep 9, 810-818.

Clerici, M., Mantiero, D., Lucchini, G., and Longhese, M.P. (2005). The Saccharomyces cerevisiae Sae2 protein promotes resection and bridging of double strand break ends. J Biol Chem 280, 38631-38638.

Cohen-Fix, O., and Koshland, D. (1997). The anaphase inhibitor of Saccharomyces cerevisiae Pds1p is a target of the DNA damage checkpoint pathway. Proc Natl Acad Sci U S A 94, 14361-14366.

Collin, P., Nashchekina, O., Walker, R., and Pines, J. (2013). The spindle assembly checkpoint works like a rheostat rather than a toggle switch. Nat Cell Biol 15, 1378-1385.

Cong, L., Ran, F.A., Cox, D., Lin, S., Barretto, R., Habib, N., Hsu, P.D., Wu, X., Jiang, W., Marraffini, L.A., et al. (2013). Multiplex genome engineering using CRISPR/Cas systems. Science 339, 819-823.

Connelly, C., and Hieter, P. (1996). Budding yeast SKP1 encodes an evolutionarily conserved kinetochore protein required for cell cycle progression. Cell 86, 275-285.

Cremer, T., Cremer, C., Baumann, H., Luedtke, E.K., Sperling, K., Teuber, V., and Zorn, C. (1982). Rabl's model of the interphase chromosome arrangement tested in Chinese hamster cells by premature chromosome condensation and laser-UV-microbeam experiments. Human genetics 60, 46-56.

D'Amours, D., and Jackson, S.P. (2001). The yeast Xrs2 complex functions in S phase checkpoint regulation. Genes Dev 15, 2238-2249.

Daley, J.M., Laan, R.L., Suresh, A., and Wilson, T.E. (2005). DNA joint dependence of pol X family polymerase action in nonhomologous end joining. J Biol Chem 280, 29030-29037.

Dandjinou, A.T., Levesque, N., Larose, S., Lucier, J.F., Abou Elela, S., and Wellinger, R.J. (2004). A phylogenetically based secondary structure for the yeast telomerase RNA. Curr Biol 14, 1148-1158. de Jager, M., van Noort, J., van Gent, D.C., Dekker, C., Kanaar, R., and Wyman, C. (2001). Human Rad50/Mre11 is a flexible complex that can tether DNA ends. Mol Cell 8, 1129-1135.

159

Dekker, J., Rippe, K., Dekker, M., and Kleckner, N. (2002). Capturing chromosome conformation. Science 295, 1306-1311.

Denchi, E.L., and de Lange, T. (2007). Protection of telomeres through independent control of ATM and ATR by TRF2 and POT1. Nature 448, 1068-1071.

Deng, S.K., Gibb, B., de Almeida, M.J., Greene, E.C., and Symington, L.S. (2014). RPA antagonizes microhomology-mediated repair of DNA double-strand breaks. Nat Struct Mol Biol 21, 405-412.

Dewar, J.M., and Lydall, D. (2010). Pif1- and Exo1-dependent nucleases coordinate checkpoint activation following telomere uncapping. EMBO J 29, 4020-4034.

Dick, A.E., and Gerlich, D.W. (2013). Kinetic framework of spindle assembly checkpoint signalling. Nat Cell Biol 15, 1370-1377.

Diede, S.J., and Gottschling, D.E. (1999). Telomerase-mediated telomere addition in vivo requires DNA primase and DNA polymerases alpha and delta. Cell 99, 723-733.

Dimitrova, N., Chen, Y.C., Spector, D.L., and de Lange, T. (2008). 53BP1 promotes non-homologous end joining of telomeres by increasing chromatin mobility. Nature 456, 524-528.

Dion, V., and Gasser, S.M. (2013). Chromatin movement in the maintenance of genome stability. Cell 152, 1355-1364.

Dion, V., Kalck, V., Horigome, C., Towbin, B.D., and Gasser, S.M. (2012). Increased mobility of double- strand breaks requires Mec1, Rad9 and the homologous recombination machinery. Nat Cell Biol 14, 502- 509.

Donnianni, R.A., and Symington, L.S. (2013). Break-induced replication occurs by conservative DNA synthesis. Proc Natl Acad Sci U S A 110, 13475-13480.

Dorn, J.F., Jaqaman, K., Rines, D.R., Jelson, G.S., Sorger, P.K., and Danuser, G. (2005). Yeast kinetochore microtubule dynamics analyzed by high-resolution three-dimensional microscopy. Biophys J 89, 2835- 2854.

Dotiwala, F., Harrison, J.C., Jain, S., Sugawara, N., and Haber, J.E. (2010). Mad2 prolongs DNA damage checkpoint arrest caused by a double-strand break via a centromere-dependent mechanism. Curr Biol 20, 328-332.

Douzery, E.J., Snell, E.A., Bapteste, E., Delsuc, F., and Philippe, H. (2004). The timing of eukaryotic evolution: does a relaxed molecular clock reconcile proteins and fossils? Proc Natl Acad Sci U S A 101, 15386-15391.

Downey, M., Houlsworth, R., Maringele, L., Rollie, A., Brehme, M., Galicia, S., Guillard, S., Partington, M., Zubko, M.K., Krogan, N.J., et al. (2006). A genome-wide screen identifies the evolutionarily conserved KEOPS complex as a telomere regulator. Cell 124, 1155-1168.

Downs, J.A., Allard, S., Jobin-Robitaille, O., Javaheri, A., Auger, A., Bouchard, N., Kron, S.J., Jackson, S.P., and Cote, J. (2004). Binding of chromatin-modifying activities to phosphorylated histone H2A at DNA damage sites. Mol Cell 16, 979-990.

160

Downs, J.A., Lowndes, N.F., and Jackson, S.P. (2000). A role for Saccharomyces cerevisiae histone H2A in DNA repair. Nature 408, 1001-1004.

Duan, Z., Andronescu, M., Schutz, K., McIlwain, S., Kim, Y.J., Lee, C., Shendure, J., Fields, S., Blau, C.A., and Noble, W.S. (2010). A three-dimensional model of the yeast genome. Nature 465, 363-367.

Ellis, N.A., Groden, J., Ye, T.Z., Straughen, J., Lennon, D.J., Ciocci, S., Proytcheva, M., and German, J. (1995). The Bloom's syndrome gene product is homologous to RecQ helicases. Cell 83, 655-666.

Ellison, V., and Stillman, B. (2003). Biochemical characterization of DNA damage checkpoint complexes: clamp loader and clamp complexes with specificity for 5' recessed DNA. PLoS Biol 1, E33.

Eugster, A., Lanzuolo, C., Bonneton, M., Luciano, P., Pollice, A., Pulitzer, J.F., Stegberg, E., Berthiau, A.S., Forstemann, K., Corda, Y., et al. (2006). The finger subdomain of yeast telomerase cooperates with Pif1p to limit telomere elongation. Nat Struct Mol Biol 13, 734-739.

Evans, S.K., and Lundblad, V. (1999). Est1 and Cdc13 as comediators of telomerase access. Science 286, 117-120.

Evans, S.K., and Lundblad, V. (2002). The Est1 subunit of Saccharomyces cerevisiae telomerase makes multiple contributions to telomere length maintenance. Genetics 162, 1101-1115.

Falck, J., Mailand, N., Syljuasen, R.G., Bartek, J., and Lukas, J. (2001). The ATM-Chk2-Cdc25A checkpoint pathway guards against radioresistant DNA synthesis. Nature 410, 842-847.

Falzon, M., Fewell, J.W., and Kuff, E.L. (1993). EBP-80, a transcription factor closely resembling the human autoantigen Ku, recognizes single- to double-strand transitions in DNA. J Biol Chem 268, 10546- 10552.

Farmer, H., McCabe, N., Lord, C.J., Tutt, A.N., Johnson, D.A., Richardson, T.B., Santarosa, M., Dillon, K.J., Hickson, I., Knights, C., et al. (2005). Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 434, 917-921.

Ferreira, H.C., Luke, B., Schober, H., Kalck, V., Lingner, J., and Gasser, S.M. (2011). The PIAS homologue Siz2 regulates perinuclear telomere position and telomerase activity in budding yeast. Nat Cell Biol 13, 867-874.

Fitzgerald-Hayes, M., Clarke, L., and Carbon, J. (1982). Nucleotide sequence comparisons and functional analysis of yeast centromere DNAs. Cell 29, 235-244.

Ford, D., Easton, D.F., Stratton, M., Narod, S., Goldgar, D., Devilee, P., Bishop, D.T., Weber, B., Lenoir, G., Chang-Claude, J., et al. (1998). Genetic heterogeneity and penetrance analysis of the BRCA1 and BRCA2 genes in breast cancer families. The Breast Cancer Linkage Consortium. American journal of human genetics 62, 676-689.

Forney, J.D., and Blackburn, E.H. (1988). Developmentally controlled telomere addition in wild-type and mutant paramecia. Mol Cell Biol 8, 251-258.

Fouladi, B., Sabatier, L., Miller, D., Pottier, G., and Murnane, J.P. (2000). The relationship between spontaneous telomere loss and chromosome instability in a human tumor cell line. Neoplasia (New York, NY) 2, 540-554.

161

Foury, F., and Kolodynski, J. (1983). pif mutation blocks recombination between mitochondrial rho+ and rho- genomes having tandemly arrayed repeat units in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 80, 5345-5349.

Frank-Vaillant, M., and Marcand, S. (2001). NHEJ regulation by mating type is exercised through a novel protein, Lif2p, essential to the ligase IV pathway. Genes Dev 15, 3005-3012.

Frank-Vaillant, M., and Marcand, S. (2002). Transient stability of DNA ends allows nonhomologous end joining to precede homologous recombination. Mol Cell 10, 1189-1199.

Frank, C.J., Hyde, M., and Greider, C.W. (2006). Regulation of telomere elongation by the cyclin- dependent kinase CDK1. Mol Cell 24, 423-432.

Furnari, B., Rhind, N., and Russell, P. (1997). Cdc25 mitotic inducer targeted by chk1 DNA damage checkpoint kinase. Science 277, 1495-1497.

Furuyama, S., and Biggins, S. (2007). Centromere identity is specified by a single centromeric nucleosome in budding yeast. Proc Natl Acad Sci U S A 104, 14706-14711.

Galgoczy, D.J., and Toczyski, D.P. (2001). Checkpoint adaptation precedes spontaneous and damage- induced genomic instability in yeast. Mol Cell Biol 21, 1710-1718.

Gao, H., Cervantes, R.B., Mandell, E.K., Otero, J.H., and Lundblad, V. (2007). RPA-like proteins mediate yeast telomere function. Nat Struct Mol Biol 14, 208-214.

Gao, H., Toro, T.B., Paschini, M., Braunstein-Ballew, B., Cervantes, R.B., and Lundblad, V. (2010). Telomerase recruitment in Saccharomyces cerevisiae is not dependent on Tel1-mediated phosphorylation of Cdc13. Genetics 186, 1147-1159.

Garber, P.M., and Rine, J. (2002). Overlapping roles of the spindle assembly and DNA damage checkpoints in the cell-cycle response to altered chromosomes in Saccharomyces cerevisiae. Genetics 161, 521-534.

Gardner, M.K., Haase, J., Mythreye, K., Molk, J.N., Anderson, M., Joglekar, A.P., O'Toole, E.T., Winey, M., Salmon, E.D., Odde, D.J., et al. (2008). The microtubule-based motor Kar3 and plus end-binding protein Bim1 provide structural support for the anaphase spindle. J Cell Biol 180, 91-100.

Gardner, R., Putnam, C.W., and Weinert, T. (1999). RAD53, DUN1 and PDS1 define two parallel G2/M checkpoint pathways in budding yeast. EMBO J 18, 3173-3185.

Gartenberg, M.R., Neumann, F.R., Laroche, T., Blaszczyk, M., and Gasser, S.M. (2004). Sir-mediated repression can occur independently of chromosomal and subnuclear contexts. Cell 119, 955-967.

Garvik, B., Carson, M., and Hartwell, L. (1995). Single-stranded DNA arising at telomeres in cdc13 mutants may constitute a specific signal for the RAD9 checkpoint. Mol Cell Biol 15, 6128-6138.

Gasch, A.P., Huang, M., Metzner, S., Botstein, D., Elledge, S.J., and Brown, P.O. (2001). Genomic expression responses to DNA-damaging agents and the regulatory role of the yeast ATR homolog Mec1p. Mol Biol Cell 12, 2987-3003.

Gasser, S.M. (2002). Visualizing chromatin dynamics in interphase nuclei. Science 296, 1412-1416.

162

Gehlen, L.R., Gasser, S.M., and Dion, V. (2011). How Broken DNA Finds Its Template for Repair: A Computational Approach. Progress of Theoretical Physics Supplement 191, 20-29.

Gillis, A.N., Thomas, S., Hansen, S.D., and Kaplan, K.B. (2005). A novel role for the CBF3 kinetochore- scaffold complex in regulating septin dynamics and cytokinesis. J Cell Biol 171, 773-784.

Gilson, E., Roberge, M., Giraldo, R., Rhodes, D., and Gasser, S.M. (1993). Distortion of the DNA double helix by RAP1 at silencers and multiple telomeric binding sites. In Journal of molecular biology, pp. 293- 310.

Goh, P.Y., and Kilmartin, J.V. (1993). NDC10: a gene involved in chromosome segregation in Saccharomyces cerevisiae. J Cell Biol 121, 503-512.

Gonda, D.K., and Radding, C.M. (1983). By searching processively RecA protein pairs DNA molecules that share a limited stretch of homology. Cell 34, 647-654.

Goshima, G., and Yanagida, M. (2000). Establishing biorientation occurs with precocious separation of the sister kinetochores, but not the arms, in the early spindle of budding yeast. Cell 100, 619-633.

Gotta, M., Laroche, T., Formenton, A., Maillet, L., Scherthan, H., and Gasser, S.M. (1996). The clustering of telomeres and colocalization with Rap1, Sir3, and Sir4 proteins in wild-type Saccharomyces cerevisiae. J Cell Biol 134, 1349-1363.

Gottschling, D.E., Aparicio, O.M., Billington, B.L., and Zakian, V.A. (1990). Position effect at S. cerevisiae telomeres: reversible repression of Pol II transcription. Cell 63, 751-762.

Graham, I.R., and Chambers, A. (1994). Use of a selection technique to identify the diversity of binding sites for the yeast RAP1 transcription factor. Nucleic Acids Res 22, 124-130.

Graham, I.R., Haw, R.A., Spink, K.G., Halden, K.A., and Chambers, A. (1999). In vivo analysis of functional regions within yeast Rap1p. Mol Cell Biol 19, 7481-7490.

Grandin, N., Damon, C., and Charbonneau, M. (2001). Ten1 functions in telomere end protection and length regulation in association with Stn1 and Cdc13. EMBO J 20, 1173-1183.

Greider, C.W., and Blackburn, E.H. (1985). Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 43, 405-413.

Greider, C.W., and Blackburn, E.H. (1987). The telomere terminal transferase of Tetrahymena is a ribonucleoprotein enzyme with two kinds of primer specificity. Cell 51, 887-898.

Greider, C.W., and Blackburn, E.H. (1989). A telomeric sequence in the RNA of Tetrahymena telomerase required for telomere repeat synthesis. Nature 337, 331-337.

Grenon, M., Gilbert, C., and Lowndes, N.F. (2001). Checkpoint activation in response to double-strand breaks requires the Mre11/Rad50/Xrs2 complex. Nat Cell Biol 3, 844-847.

Grossi, S., Bianchi, A., Damay, P., and Shore, D. (2001). Telomere formation by rap1p binding site arrays reveals end-specific length regulation requirements and active telomeric recombination. Mol Cell Biol 21, 8117-8128.

Haber, J.E. (1999). DNA recombination: the replication connection. Trends Biochem Sci 24, 271-275.

163

Haber, J.E. (2012). Mating-type genes and MAT switching in Saccharomyces cerevisiae. Genetics 191, 33- 64.

Harfst, E., Cooper, S., Neubauer, S., Distel, L., and Grawunder, U. (2000). Normal V(D)J recombination in cells from patients with Nijmegen breakage syndrome. Molecular immunology 37, 915-929.

Hediger, F., Neumann, F.R., Van Houwe, G., Dubrana, K., and Gasser, S.M. (2002). Live imaging of telomeres: yKu and Sir proteins define redundant telomere-anchoring pathways in yeast. Curr Biol 12, 2076-2089.

Herrmann, G., Lindahl, T., and Schar, P. (1998). Saccharomyces cerevisiae LIF1: a function involved in DNA double-strand break repair related to mammalian XRCC4. EMBO J 17, 4188-4198.

Herzberg, K., Bashkirov, V.I., Rolfsmeier, M., Haghnazari, E., McDonald, W.H., Anderson, S., Bashkirova, E.V., Yates, J.R., 3rd, and Heyer, W.D. (2006). Phosphorylation of Rad55 on serines 2, 8, and 14 is required for efficient homologous recombination in the recovery of stalled replication forks. Mol Cell Biol 26, 8396-8409.

Heun, P., Laroche, T., Shimada, K., Furrer, P., and Gasser, S.M. (2001). Chromosome dynamics in the yeast interphase nucleus. Science 294, 2181-2186.

Heyer, W.D., Rao, M.R., Erdile, L.F., Kelly, T.J., and Kolodner, R.D. (1990). An essential Saccharomyces cerevisiae single-stranded DNA binding protein is homologous to the large subunit of human RP-A. EMBO J 9, 2321-2329.

Hicks, W.M., Kim, M., and Haber, J.E. (2010). Increased mutagenesis and unique mutation signature associated with mitotic gene conversion. Science 329, 82-85.

Hill, A., and Bloom, K. (1987). Genetic manipulation of centromere function. Mol Cell Biol 7, 2397-2405.

Hirano, Y., Fukunaga, K., and Sugimoto, K. (2009). Rif1 and rif2 inhibit localization of tel1 to DNA ends. Mol Cell 33, 312-322.

Hirano, Y., and Sugimoto, K. (2007). Cdc13 telomere capping decreases Mec1 association but does not affect Tel1 association with DNA ends. Mol Biol Cell 18, 2026-2036.

Holbeck, S.L., and Strathern, J.N. (1997). A role for REV3 in mutagenesis during double-strand break repair in Saccharomyces cerevisiae. Genetics 147, 1017-1024.

Holliday, R. (1964). A mechanism for gene conversion in fungi. Genet Res 5, 282-304.

Holstein, E.M., and Lydall, D. (2012). Quantitative amplification of single-stranded DNA. Methods Mol Biol 920, 323-339.

Hopfner, K.P., Craig, L., Moncalian, G., Zinkel, R.A., Usui, T., Owen, B.A., Karcher, A., Henderson, B., Bodmer, J.L., McMurray, C.T., et al. (2002). The Rad50 zinc-hook is a structure joining Mre11 complexes in DNA recombination and repair. Nature 418, 562-566.

Horigome, C., Oma, Y., Konishi, T., Schmid, R., Marcomini, I., Hauer, M.H., Dion, V., Harata, M., and Gasser, S.M. (2014). SWR1 and INO80 chromatin remodelers contribute to DNA double-strand break perinuclear anchorage site choice. Mol Cell 55, 626-639.

164

Hoyt, M.A., Totis, L., and Roberts, B.T. (1991). S. cerevisiae genes required for cell cycle arrest in response to loss of microtubule function. Cell 66, 507-517.

Hu, F., Wang, Y., Liu, D., Li, Y., Qin, J., and Elledge, S.J. (2001). Regulation of the Bub2/Bfa1 GAP complex by Cdc5 and cell cycle checkpoints. Cell 107, 655-665.

Huertas, P., Cortes-Ledesma, F., Sartori, A.A., Aguilera, A., and Jackson, S.P. (2008). CDK targets Sae2 to control DNA-end resection and homologous recombination. Nature 455, 689-692.

Huertas, P., and Jackson, S.P. (2009). Human CtIP mediates cell cycle control of DNA end resection and double strand break repair. J Biol Chem 284, 9558-9565.

Hughes, T.R., Weilbaecher, R.G., Walterscheid, M., and Lundblad, V. (2000). Identification of the single- strand telomeric DNA binding domain of the Saccharomyces cerevisiae Cdc13 protein. Proc Natl Acad Sci U S A 97, 6457-6462.

Ira, G., Malkova, A., Liberi, G., Foiani, M., and Haber, J.E. (2003). Srs2 and Sgs1-Top3 suppress crossovers during double-strand break repair in yeast. Cell 115, 401-411.

Ira, G., Pellicioli, A., Balijja, A., Wang, X., Fiorani, S., Carotenuto, W., Liberi, G., Bressan, D., Wan, L., Hollingsworth, N.M., et al. (2004). DNA end resection, homologous recombination and DNA damage checkpoint activation require CDK1. Nature 431, 1011-1017.

Ivanov, E.L., Sugawara, N., Fishman-Lobell, J., and Haber, J.E. (1996). Genetic requirements for the single-strand annealing pathway of double-strand break repair in Saccharomyces cerevisiae. Genetics 142, 693-704.

Ivanov, E.L., Sugawara, N., White, C.I., Fabre, F., and Haber, J.E. (1994). Mutations in XRS2 and RAD50 delay but do not prevent mating-type switching in Saccharomyces cerevisiae. Mol Cell Biol 14, 3414-3425.

Ivessa, A.S., Zhou, J.Q., and Zakian, V.A. (2000). The Saccharomyces Pif1p DNA helicase and the highly related Rrm3p have opposite effects on replication fork progression in ribosomal DNA. Cell 100, 479-489.

Jackson, S.P., and Bartek, J. (2009). The DNA-damage response in human biology and disease. Nature 461, 1071-1078.

Jain, S., Sugawara, N., Lydeard, J., Vaze, M., Tanguy Le Gac, N., and Haber, J.E. (2009). A recombination execution checkpoint regulates the choice of homologous recombination pathway during DNA double- strand break repair. Genes Dev 23, 291-303.

Janke, C., Ortiz, J., Lechner, J., Shevchenko, A., Shevchenko, A., Magiera, M.M., Schramm, C., and Schiebel, E. (2001). The budding yeast proteins Spc24p and Spc25p interact with Ndc80p and Nuf2p at the kinetochore and are important for kinetochore clustering and checkpoint control. EMBO J 20, 777-791.

Jeggo, P.A., and Downs, J.A. (2014). Roles of chromatin remodellers in DNA double strand break repair. Exp Cell Res 329, 69-77.

Jin, Q.W., Fuchs, J., and Loidl, J. (2000). Centromere clustering is a major determinant of yeast interphase nuclear organization. J Cell Sci 113 ( Pt 11), 1903-1912.

Johnson, R.D., and Jasin, M. (2000). Sister chromatid gene conversion is a prominent double-strand break repair pathway in mammalian cells. EMBO J 19, 3398-3407.

165

Johnson, R.D., and Symington, L.S. (1995). Functional differences and interactions among the putative RecA homologs Rad51, Rad55, and Rad57. Mol Cell Biol 15, 4843-4850.

Jones, M.H., Bachant, J.B., Castillo, A.R., Giddings, T.H., Jr., and Winey, M. (1999). Yeast Dam1p is required to maintain spindle integrity during mitosis and interacts with the Mps1p kinase. Mol Biol Cell 10, 2377-2391.

Kachroo, A.H., Laurent, J.M., Yellman, C.M., Meyer, A.G., Wilke, C.O., and Marcotte, E.M. (2015). Evolution. Systematic humanization of yeast genes reveals conserved functions and genetic modularity. Science 348, 921-925.

Kalocsay, M., Hiller, N.J., and Jentsch, S. (2009). Chromosome-wide Rad51 spreading and SUMO-H2A.Z- dependent chromosome fixation in response to a persistent DNA double-strand break. Mol Cell 33, 335- 343.

Kapoor, P., Bao, Y., Xiao, J., Espejo, A., Yang, L., Bedford, M.T., Peng, G., and Shen, X. (2015). Phosphorylation-Dependent Enhancement of Rad53 Kinase Activity through the INO80 Chromatin Remodeling Complex. Mol Cell 58, 863-869.

Karlseder, J., Broccoli, D., Dai, Y., Hardy, S., and de Lange, T. (1999). p53- and ATM-dependent apoptosis induced by telomeres lacking TRF2. Science 283, 1321-1325.

Kaye, J.A., Melo, J.A., Cheung, S.K., Vaze, M.B., Haber, J.E., and Toczyski, D.P. (2004). DNA breaks promote genomic instability by impeding proper chromosome segregation. Curr Biol 14, 2096-2106.

Keeney, S. (2001). Mechanism and control of meiotic recombination initiation. Current topics in developmental biology 52, 1-53.

Keil, R.L., and McWilliams, A.D. (1993). A gene with specific and global effects on recombination of sequences from tandemly repeated genes in Saccharomyces cerevisiae. Genetics 135, 711-718.

Keogh, M.C., Kim, J.A., Downey, M., Fillingham, J., Chowdhury, D., Harrison, J.C., Onishi, M., Datta, N., Galicia, S., Emili, A., et al. (2006a). A phosphatase complex that dephosphorylates gammaH2AX regulates DNA damage checkpoint recovery. Nature 439, 497-501.

Keogh, M.C., Mennella, T.A., Sawa, C., Berthelet, S., Krogan, N.J., Wolek, A., Podolny, V., Carpenter, L.R., Greenblatt, J.F., Baetz, K., et al. (2006b). The Saccharomyces cerevisiae histone H2A variant Htz1 is acetylated by NuA4. Genes Dev 20, 660-665.

Khanna, K.K., and Jackson, S.P. (2001). DNA double-strand breaks: signaling, repair and the cancer connection. Nat Genet 27, 247-254.

Kim, H.S., and Brill, S.J. (2001). Rfc4 interacts with Rpa1 and is required for both DNA replication and DNA damage checkpoints in Saccharomyces cerevisiae. Mol Cell Biol 21, 3725-3737.

Kitamura, E., Tanaka, K., Kitamura, Y., and Tanaka, T.U. (2007). Kinetochore microtubule interaction during S phase in Saccharomyces cerevisiae. Genes Dev 21, 3319-3330.

Kondo, T., Wakayama, T., Naiki, T., Matsumoto, K., and Sugimoto, K. (2001). Recruitment of Mec1 and Ddc1 checkpoint proteins to double-strand breaks through distinct mechanisms. Science 294, 867-870.

166

Kramer, K.M., and Haber, J.E. (1993). New telomeres in yeast are initiated with a highly selected subset of TG1-3 repeats. Genes Dev 7, 2345-2356.

Krawczyk, P.M., Borovski, T., Stap, J., Cijsouw, T., ten Cate, R., Medema, J.P., Kanaar, R., Franken, N.A., and Aten, J.A. (2012). Chromatin mobility is increased at sites of DNA double-strand breaks. J Cell Sci 125, 2127-2133.

Krogan, N.J., Baetz, K., Keogh, M.C., Datta, N., Sawa, C., Kwok, T.C., Thompson, N.J., Davey, M.G., Pootoolal, J., Hughes, T.R., et al. (2004). Regulation of chromosome stability by the histone H2A variant Htz1, the Swr1 chromatin remodeling complex, and the histone acetyltransferase NuA4. Proc Natl Acad Sci U S A 101, 13513-13518.

Krogan, N.J., Keogh, M.C., Datta, N., Sawa, C., Ryan, O.W., Ding, H., Haw, R.A., Pootoolal, J., Tong, A., Canadien, V., et al. (2003). A Snf2 family ATPase complex required for recruitment of the histone H2A variant Htz1. Mol Cell 12, 1565-1576.

Kruhlak, M.J., Celeste, A., Dellaire, G., Fernandez-Capetillo, O., Muller, W.G., McNally, J.G., Bazett- Jones, D.P., and Nussenzweig, A. (2006). Changes in chromatin structure and mobility in living cells at sites of DNA double-strand breaks. J Cell Biol 172, 823-834.

Kubota, T., Nishimura, K., Kanemaki, M.T., and Donaldson, A.D. (2013). The Elg1 replication factor C- like complex functions in PCNA unloading during DNA replication. Mol Cell 50, 273-280.

Kung, C., Kenski, D.M., Dickerson, S.H., Howson, R.W., Kuyper, L.F., Madhani, H.D., and Shokat, K.M. (2005). Chemical genomic profiling to identify intracellular targets of a multiplex kinase inhibitor. Proc Natl Acad Sci U S A 102, 3587-3592.

Kurdistani, S.K., Robyr, D., Tavazoie, S., and Grunstein, M. (2002). Genome-wide binding map of the histone deacetylase Rpd3 in yeast. Nat Genet 31, 248-254.

Lahaye, A., Stahl, H., Thines-Sempoux, D., and Foury, F. (1991). PIF1: a DNA helicase in yeast mitochondria. EMBO J 10, 997-1007.

Lau, D.T., and Murray, A.W. (2012). Mad2 and Mad3 cooperate to arrest budding yeast in mitosis. Curr Biol 22, 180-190.

Lazzaro, F., Sapountzi, V., Granata, M., Pellicioli, A., Vaze, M., Haber, J.E., Plevani, P., Lydall, D., and Muzi-Falconi, M. (2008). Histone methyltransferase Dot1 and Rad9 inhibit single-stranded DNA accumulation at DSBs and uncapped telomeres. EMBO J 27, 1502-1512.

Lebel, C., Rosonina, E., Sealey, D.C., Pryde, F., Lydall, D., Maringele, L., and Harrington, L.A. (2009). Telomere maintenance and survival in saccharomyces cerevisiae in the absence of telomerase and RAD52. Genetics 182, 671-684.

Lechner, J. (1994). A zinc finger protein, essential for chromosome segregation, constitutes a putative DNA binding subunit of the Saccharomyces cerevisiae kinetochore complex, Cbf3. EMBO J 13, 5203-5211.

Lechner, J., and Carbon, J. (1991). A 240 kd multisubunit protein complex, CBF3, is a major component of the budding yeast centromere. Cell 64, 717-725.

Lee, C.S., Wang, R.W., Chang, H.H., Capurso, D., Segal, M.R., and Haber, J.E. (2016). Chromosome position determines the success of double-strand break repair. Proc Natl Acad Sci U S A 113, E146-154.

167

Lee, J.Y., Terakawa, T., Qi, Z., Steinfeld, J.B., Redding, S., Kwon, Y., Gaines, W.A., Zhao, W., Sung, P., and Greene, E.C. (2015). DNA RECOMBINATION. Base triplet stepping by the Rad51/RecA family of recombinases. Science 349, 977-981.

Lee, K., and Lee, S.E. (2007). Saccharomyces cerevisiae Sae2- and Tel1-dependent single-strand DNA formation at DNA break promotes microhomology-mediated end joining. Genetics 176, 2003-2014.

Lee, K., Zhang, Y., and Lee, S.E. (2008). Saccharomyces cerevisiae ATM orthologue suppresses break- induced chromosome translocations. Nature 454, 543-546.

Lee, S.E., Moore, J.K., Holmes, A., Umezu, K., Kolodner, R.D., and Haber, J.E. (1998). Saccharomyces Ku70, mre11/rad50 and RPA proteins regulate adaptation to G2/M arrest after DNA damage. Cell 94, 399- 409.

Lee, S.E., Pellicioli, A., Malkova, A., Foiani, M., and Haber, J.E. (2001). The Saccharomyces recombination protein Tid1p is required for adaptation from G2/M arrest induced by a double-strand break. Curr Biol 11, 1053-1057.

Lemieux, B., Laterreur, N., Perederina, A., Noel, J.F., Dubois, M.L., Krasilnikov, A.S., and Wellinger, R.J. (2016). Active Yeast Telomerase Shares Subunits with Ribonucleoproteins RNase P and RNase MRP. Cell 165, 1171-1181.

Lendvay, T.S., Morris, D.K., Sah, J., Balasubramanian, B., and Lundblad, V. (1996). Senescence mutants of Saccharomyces cerevisiae with a defect in telomere replication identify three additional EST genes. Genetics 144, 1399-1412.

Lengauer, C., Kinzler, K.W., and Vogelstein, B. (1998). Genetic instabilities in human cancers. Nature 396, 643-649.

Leroy, C., Lee, S.E., Vaze, M.B., Ochsenbein, F., Guerois, R., Haber, J.E., and Marsolier-Kergoat, M.C. (2003). PP2C phosphatases Ptc2 and Ptc3 are required for DNA checkpoint inactivation after a double- strand break. Mol Cell 11, 827-835.

Levy, D.L., and Blackburn, E.H. (2004). Counting of Rif1p and Rif2p on Saccharomyces cerevisiae telomeres regulates telomere length. Mol Cell Biol 24, 10857-10867.

Li, J.R., Yu, T.Y., Chien, I.C., Lu, C.Y., Lin, J.J., and Li, H.W. (2014). Pif1 regulates telomere length by preferentially removing telomerase from long telomere ends. Nucleic Acids Res 42, 8527-8536.

Li, R., and Murray, A.W. (1991). Feedback control of mitosis in budding yeast. Cell 66, 519-531.

Li, S., Makovets, S., Matsuguchi, T., Blethrow, J.D., Shokat, K.M., and Blackburn, E.H. (2009). Cdk1- dependent phosphorylation of Cdc13 coordinates telomere elongation during cell-cycle progression. Cell 136, 50-61.

Lieb, J.D., Liu, X., Botstein, D., and Brown, P.O. (2001). Promoter-specific binding of Rap1 revealed by genome-wide maps of protein-DNA association. Nat Genet 28, 327-334.

Lin, J.J., and Zakian, V.A. (1996). The Saccharomyces CDC13 protein is a single-strand TG1-3 telomeric DNA-binding protein in vitro that affects telomere behavior in vivo. Proc Natl Acad Sci U S A 93, 13760- 13765.

168

Lindahl, T. (1993). Instability and decay of the primary structure of DNA. Nature 362, 709-715.

Lindahl, T., and Wood, R.D. (1999). Quality control by DNA repair. Science 286, 1897-1905.

Lingner, J., Cech, T.R., Hughes, T.R., and Lundblad, V. (1997). Three Ever Shorter Telomere (EST) genes are dispensable for in vitro yeast telomerase activity. Proc Natl Acad Sci U S A 94, 11190-11195.

Lisby, M., Barlow, J.H., Burgess, R.C., and Rothstein, R. (2004). Choreography of the DNA damage response: spatiotemporal relationships among checkpoint and repair proteins. Cell 118, 699-713.

Lisby, M., Mortensen, U.H., and Rothstein, R. (2003). Colocalization of multiple DNA double-strand breaks at a single Rad52 repair centre. Nat Cell Biol 5, 572-577.

Lisby, M., Rothstein, R., and Mortensen, U.H. (2001). Rad52 forms DNA repair and recombination centers during S phase. Proc Natl Acad Sci U S A 98, 8276-8282.

Loeb, L.A., and Harris, C.C. (2008). Advances in chemical carcinogenesis: a historical review and prospective. Cancer Res 68, 6863-6872.

Loeb, L.A., Springgate, C.F., and Battula, N. (1974). Errors in DNA replication as a basis of malignant changes. Cancer Res 34, 2311-2321.

Longhese, M.P., Plevani, P., and Lucchini, G. (1994). Replication factor A is required in vivo for DNA replication, repair, and recombination. Mol Cell Biol 14, 7884-7890.

Lord, C.J., and Ashworth, A. (2012). The DNA damage response and cancer therapy. Nature 481, 287-294.

Lottersberger, F., Karssemeijer, R.A., Dimitrova, N., and de Lange, T. (2015). 53BP1 and the LINC Complex Promote Microtubule-Dependent DSB Mobility and DNA Repair. Cell 163, 880-893.

Lue, N.F., Chan, J., Wright, W.E., and Hurwitz, J. (2014). The CDC13-STN1-TEN1 complex stimulates Pol alpha activity by promoting RNA priming and primase-to-polymerase switch. Nat Commun 5, 5762.

Lundblad, V., and Blackburn, E.H. (1993). An alternative pathway for yeast telomere maintenance rescues est1- senescence. Cell 73, 347-360.

Lundblad, V., and Szostak, J.W. (1989). A mutant with a defect in telomere elongation leads to senescence in yeast. Cell 57, 633-643.

Luo, K., Vega-Palas, M.A., and Grunstein, M. (2002). Rap1-Sir4 binding independent of other Sir, yKu, or histone interactions initiates the assembly of telomeric heterochromatin in yeast. Genes Dev 16, 1528-1539.

Lustig, A.J. (1992). Hoogsteen G-G base pairing is dispensable for telomere healing in yeast. Nucleic Acids Res 20, 3021-3028.

Lustig, A.J., Kurtz, S., and Shore, D. (1990). Involvement of the silencer and UAS binding protein RAP1 in regulation of telomere length. Science 250, 549-553.

Lustig, A.J., and Petes, T.D. (1986). Identification of yeast mutants with altered telomere structure. Proc Natl Acad Sci U S A 83, 1398-1402.

169

Lydeard, J.R., Jain, S., Yamaguchi, M., and Haber, J.E. (2007). Break-induced replication and telomerase- independent telomere maintenance require Pol32. Nature 448, 820-823.

Ma, J.L., Kim, E.M., Haber, J.E., and Lee, S.E. (2003). Yeast Mre11 and Rad1 proteins define a Ku- independent mechanism to repair double-strand breaks lacking overlapping end sequences. Mol Cell Biol 23, 8820-8828.

Majka, J., and Burgers, P.M. (2003). Yeast Rad17/Mec3/Ddc1: a sliding clamp for the DNA damage checkpoint. Proc Natl Acad Sci U S A 100, 2249-2254.

Makovets, S., and Blackburn, E.H. (2009). DNA damage signalling prevents deleterious telomere addition at DNA breaks. Nat Cell Biol 11, 1383-1386.

Malkova, A., Ivanov, E.L., and Haber, J.E. (1996). Double-strand break repair in the absence of RAD51 in yeast: a possible role for break-induced DNA replication. Proc Natl Acad Sci U S A 93, 7131-7136.

Maloisel, L., Fabre, F., and Gangloff, S. (2008). DNA polymerase delta is preferentially recruited during homologous recombination to promote heteroduplex DNA extension. Mol Cell Biol 28, 1373-1382.

Mangahas, J.L., Alexander, M.K., Sandell, L.L., and Zakian, V.A. (2001). Repair of chromosome ends after telomere loss in Saccharomyces. Mol Biol Cell 12, 4078-4089.

Manning, B.D., Barrett, J.G., Wallace, J.A., Granok, H., and Snyder, M. (1999). Differential regulation of the Kar3p kinesin-related protein by two associated proteins, Cik1p and Vik1p. J Cell Biol 144, 1219-1233.

Marcand, S., Brevet, V., and Gilson, E. (1999). Progressive cis-inhibition of telomerase upon telomere elongation. EMBO J 18, 3509-3519.

Marcand, S., Brevet, V., Mann, C., and Gilson, E. (2000). Cell cycle restriction of telomere elongation. Curr Biol 10, 487-490.

Marcand, S., Gilson, E., and Shore, D. (1997). A protein-counting mechanism for telomere length regulation in yeast. Science 275, 986-990.

Marcand, S., Pardo, B., Gratias, A., Cahun, S., and Callebaut, I. (2008). Multiple pathways inhibit NHEJ at telomeres. Genes Dev 22, 1153-1158.

Maringele, L., and Lydall, D. (2002). EXO1-dependent single-stranded DNA at telomeres activates subsets of DNA damage and spindle checkpoint pathways in budding yeast yku70Delta mutants. Genes Dev 16, 1919-1933.

Marshall, W.F., Dernburg, A.F., Harmon, B., Agard, D.A., and Sedat, J.W. (1996). Specific interactions of chromatin with the nuclear envelope: positional determination within the nucleus in Drosophila melanogaster. Mol Biol Cell 7, 825-842.

Marshall, W.F., Straight, A., Marko, J.F., Swedlow, J., Dernburg, A., Belmont, A., Murray, A.W., Agard, D.A., and Sedat, J.W. (1997). Interphase chromosomes undergo constrained diffusional motion in living cells. Curr Biol 7, 930-939.

Martin, S.G., Laroche, T., Suka, N., Grunstein, M., and Gasser, S.M. (1999). Relocalization of telomeric Ku and SIR proteins in response to DNA strand breaks in yeast. Cell 97, 621-633.

170

Martincorena, I., Roshan, A., Gerstung, M., Ellis, P., Van Loo, P., McLaren, S., Wedge, D.C., Fullam, A., Alexandrov, L.B., Tubio, J.M., et al. (2015). Tumor evolution. High burden and pervasive positive selection of somatic mutations in normal human skin. Science 348, 880-886.

Maser, R.S., Wong, K.K., Sahin, E., Xia, H., Naylor, M., Hedberg, H.M., Artandi, S.E., and DePinho, R.A. (2007). DNA-dependent protein kinase catalytic subunit is not required for dysfunctional telomere fusion and checkpoint response in the telomerase-deficient mouse. Mol Cell Biol 27, 2253-2265.

Mason, M., Wanat, J.J., Harper, S., Schultz, D.C., Speicher, D.W., Johnson, F.B., and Skordalakes, E. (2013). Cdc13 OB2 dimerization required for productive Stn1 binding and efficient telomere maintenance. Structure 21, 109-120.

Mateos-Gomez, P.A., Gong, F., Nair, N., Miller, K.M., Lazzerini-Denchi, E., and Sfeir, A. (2015). Mammalian polymerase theta promotes alternative NHEJ and suppresses recombination. Nature 518, 254- 257.

Matsuoka, S., Ballif, B.A., Smogorzewska, A., McDonald, E.R., 3rd, Hurov, K.E., Luo, J., Bakalarski, C.E., Zhao, Z., Solimini, N., Lerenthal, Y., et al. (2007). ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science 316, 1160-1166.

Matsuoka, S., Huang, M., and Elledge, S.J. (1998). Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 282, 1893-1897.

Maure, J.F., Kitamura, E., and Tanaka, T.U. (2007). Mps1 kinase promotes sister-kinetochore bi-orientation by a tension-dependent mechanism. Curr Biol 17, 2175-2182.

McAinsh, A.D., Scott-Drew, S., Murray, J.A., and Jackson, S.P. (1999). DNA damage triggers disruption of telomeric silencing and Mec1p-dependent relocation of Sir3p. Curr Biol 9, 963-966.

McClintock, B. (1941). The Stability of Broken Ends of Chromosomes in Zea Mays. Genetics 26, 234-282.

McGee, J.S., Phillips, J.A., Chan, A., Sabourin, M., Paeschke, K., and Zakian, V.A. (2010). Reduced Rif2 and lack of Mec1 target short telomeres for elongation rather than double-strand break repair. Nat Struct Mol Biol 17, 1438-1445.

Mekhail, K., and Moazed, D. (2010). The nuclear envelope in genome organization, expression and stability. Nat Rev Mol Cell Biol 11, 317-328.

Mekhail, K., Seebacher, J., Gygi, S.P., and Moazed, D. (2008). Role for perinuclear chromosome tethering in maintenance of genome stability. Nature 456, 667-670.

Melo, J.A., Cohen, J., and Toczyski, D.P. (2001). Two checkpoint complexes are independently recruited to sites of DNA damage in vivo. Genes Dev 15, 2809-2821.

Meluh, P.B., and Koshland, D. (1995). Evidence that the MIF2 gene of Saccharomyces cerevisiae encodes a centromere protein with homology to the mammalian centromere protein CENP-C. Mol Biol Cell 6, 793- 807.

Meluh, P.B., and Rose, M.D. (1990). KAR3, a kinesin-related gene required for yeast nuclear fusion. Cell 60, 1029-1041.

171

Millar, C.B., and Grunstein, M. (2006). Genome-wide patterns of histone modifications in yeast. Nat Rev Mol Cell Biol 7, 657-666.

Millar, C.B., Xu, F., Zhang, K., and Grunstein, M. (2006). Acetylation of H2AZ Lys 14 is associated with genome-wide gene activity in yeast. Genes Dev 20, 711-722.

Mills, K.D., Sinclair, D.A., and Guarente, L. (1999). MEC1-dependent redistribution of the Sir3 silencing protein from telomeres to DNA double-strand breaks. Cell 97, 609-620.

Mimitou, E.P., and Symington, L.S. (2010). Ku prevents Exo1 and Sgs1-dependent resection of DNA ends in the absence of a functional MRX complex or Sae2. EMBO J 29, 3358-3369.

Mine-Hattab, J., and Rothstein, R. (2012). Increased chromosome mobility facilitates homology search during recombination. Nat Cell Biol 14, 510-517.

Mitchell, M.T., Smith, J.S., Mason, M., Harper, S., Speicher, D.W., Johnson, F.B., and Skordalakes, E. (2010). Cdc13 N-terminal dimerization, DNA binding, and telomere length regulation. Mol Cell Biol 30, 5325-5334.

Miyake, Y., Nakamura, M., Nabetani, A., Shimamura, S., Tamura, M., Yonehara, S., Saito, M., and Ishikawa, F. (2009). RPA-like mammalian Ctc1-Stn1-Ten1 complex binds to single-stranded DNA and protects telomeres independently of the Pot1 pathway. Mol Cell 36, 193-206.

Moazed, D., Kistler, A., Axelrod, A., Rine, J., and Johnson, A.D. (1997). Silent information regulator protein complexes in Saccharomyces cerevisiae: a SIR2/SIR4 complex and evidence for a regulatory domain in SIR4 that inhibits its interaction with SIR3. Proc Natl Acad Sci U S A 94, 2186-2191.

Molenaar, C., Wiesmeijer, K., Verwoerd, N.P., Khazen, S., Eils, R., Tanke, H.J., and Dirks, R.W. (2003). Visualizing telomere dynamics in living mammalian cells using PNA probes. EMBO J 22, 6631-6641.

Moore, J.K., and Haber, J.E. (1996). Cell cycle and genetic requirements of two pathways of nonhomologous end-joining repair of double-strand breaks in Saccharomyces cerevisiae. Mol Cell Biol 16, 2164-2173.

Moretti, P., Freeman, K., Coodly, L., and Shore, D. (1994). Evidence that a complex of SIR proteins interacts with the silencer and telomere-binding protein RAP1. Genes Dev 8, 2257-2269.

Morrison, A.J., Highland, J., Krogan, N.J., Arbel-Eden, A., Greenblatt, J.F., Haber, J.E., and Shen, X. (2004). INO80 and gamma-H2AX interaction links ATP-dependent chromatin remodeling to DNA damage repair. Cell 119, 767-775.

Moyzis, R.K., Buckingham, J.M., Cram, L.S., Dani, M., Deaven, L.L., Jones, M.D., Meyne, J., Ratliff, R.L., and Wu, J.R. (1988). A highly conserved repetitive DNA sequence, (TTAGGG)n, present at the telomeres of human chromosomes. Proc Natl Acad Sci U S A 85, 6622-6626.

Muller, H.J. (1938). The remaking of chromosomes. Collecting Net 13, 181-198.

Musacchio, A., and Salmon, E.D. (2007). The spindle-assembly checkpoint in space and time. Nat Rev Mol Cell Biol 8, 379-393.

Myung, K., Chen, C., and Kolodner, R.D. (2001). Multiple pathways cooperate in the suppression of genome instability in Saccharomyces cerevisiae. Nature 411, 1073-1076.

172

Nagai, S., Dubrana, K., Tsai-Pflugfelder, M., Davidson, M.B., Roberts, T.M., Brown, G.W., Varela, E., Hediger, F., Gasser, S.M., and Krogan, N.J. (2008). Functional targeting of DNA damage to a nuclear pore- associated SUMO-dependent ubiquitin ligase. Science 322, 597-602.

Nagele, R., Freeman, T., McMorrow, L., and Lee, H.Y. (1995). Precise spatial positioning of chromosomes during prometaphase: evidence for chromosomal order. Science 270, 1831-1835.

Nakada, D., Matsumoto, K., and Sugimoto, K. (2003). ATM-related Tel1 associates with double-strand breaks through an Xrs2-dependent mechanism. Genes Dev 17, 1957-1962.

Nakada, S., Chen, G.I., Gingras, A.C., and Durocher, D. (2008). PP4 is a gamma H2AX phosphatase required for recovery from the DNA damage checkpoint. EMBO Rep 9, 1019-1026.

Negrini, S., Ribaud, V., Bianchi, A., and Shore, D. (2007). DNA breaks are masked by multiple Rap1 binding in yeast: implications for telomere capping and telomerase regulation. Genes Dev 21, 292-302.

Nelms, B.E., Maser, R.S., MacKay, J.F., Lagally, M.G., and Petrini, J.H. (1998). In situ visualization of DNA double-strand break repair in human fibroblasts. Science 280, 590-592.

Neumann, F.R., Dion, V., Gehlen, L.R., Tsai-Pflugfelder, M., Schmid, R., Taddei, A., and Gasser, S.M. (2012). Targeted INO80 enhances subnuclear chromatin movement and ectopic homologous recombination. Genes Dev 26, 369-383.

New, J.H., Sugiyama, T., Zaitseva, E., and Kowalczykowski, S.C. (1998). Rad52 protein stimulates DNA strand exchange by Rad51 and replication protein A. Nature 391, 407-410.

Nik-Zainal, S., Davies, H., Staaf, J., Ramakrishna, M., Glodzik, D., Zou, X., Martincorena, I., Alexandrov, L.B., Martin, S., Wedge, D.C., et al. (2016). Landscape of somatic mutations in 560 breast cancer whole- genome sequences. Nature.

Nik-Zainal, S., Van Loo, P., Wedge, D.C., Alexandrov, L.B., Greenman, C.D., Lau, K.W., Raine, K., Jones, D., Marshall, J., Ramakrishna, M., et al. (2012). The life history of 21 breast cancers. Cell 149, 994-1007.

Nishimura, K., Fukagawa, T., Takisawa, H., Kakimoto, T., and Kanemaki, M. (2009). An auxin-based degron system for the rapid depletion of proteins in nonplant cells. Nat Methods 6, 917-922.

Niu, H., Chung, W.H., Zhu, Z., Kwon, Y., Zhao, W., Chi, P., Prakash, R., Seong, C., Liu, D., Lu, L., et al. (2010). Mechanism of the ATP-dependent DNA end-resection machinery from Saccharomyces cerevisiae. Nature 467, 108-111.

Nugent, C.I., Hughes, T.R., Lue, N.F., and Lundblad, V. (1996). Cdc13p: a single-strand telomeric DNA- binding protein with a dual role in yeast telomere maintenance. Science 274, 249-252.

O'Brien, K.P., Remm, M., and Sonnhammer, E.L. (2005). Inparanoid: a comprehensive database of eukaryotic orthologs. Nucleic Acids Res 33, D476-480.

O'Neill, B.M., Szyjka, S.J., Lis, E.T., Bailey, A.O., Yates, J.R., 3rd, Aparicio, O.M., and Romesberg, F.E. (2007). Pph3-Psy2 is a phosphatase complex required for Rad53 dephosphorylation and replication fork restart during recovery from DNA damage. Proc Natl Acad Sci U S A 104, 9290-9295.

173

Obodo, U.C., Epum, E.A., Platts, M.H., Seloff, J., Dahlson, N.A., Velkovsky, S.M., Paul, S.R., and Friedman, K.L. (2016). Endogenous Hot Spots of De Novo Telomere Addition in the Yeast Genome Contain Proximal Enhancers That Bind Cdc13. Mol Cell Biol 36, 1750-1763.

Ogiwara, H., Enomoto, T., and Seki, M. (2007). The INO80 chromatin remodeling complex functions in sister chromatid cohesion. Cell Cycle 6, 1090-1095.

Orr-Weaver, T.L., and Szostak, J.W. (1983). Multiple, tandem plasmid integration in Saccharomyces cerevisiae. Mol Cell Biol 3, 747-749.

Orr-Weaver, T.L., Szostak, J.W., and Rothstein, R.J. (1981). Yeast transformation: a model system for the study of recombination. Proc Natl Acad Sci U S A 78, 6354-6358.

Orthwein, A., Noordermeer, S.M., Wilson, M.D., Landry, S., Enchev, R.I., Sherker, A., Munro, M., Pinder, J., Salsman, J., Dellaire, G., et al. (2015). A mechanism for the suppression of homologous recombination in G1 cells. Nature 528, 422-426.

Ortiz, J., Stemmann, O., Rank, S., and Lechner, J. (1999). A putative protein complex consisting of Ctf19, Mcm21, and Okp1 represents a missing link in the budding yeast kinetochore. Genes Dev 13, 1140-1155.

Osborn, A.J., and Elledge, S.J. (2003). Mrc1 is a replication fork component whose phosphorylation in response to DNA replication stress activates Rad53. Genes Dev 17, 1755-1767.

Osterhage, J.L., Talley, J.M., and Friedman, K.L. (2006). Proteasome-dependent degradation of Est1p regulates the cell cycle-restricted assembly of telomerase in Saccharomyces cerevisiae. Nat Struct Mol Biol 13, 720-728.

Oza, P., Jaspersen, S.L., Miele, A., Dekker, J., and Peterson, C.L. (2009). Mechanisms that regulate localization of a DNA double-strand break to the nuclear periphery. Genes Dev 23, 912-927.

Paciotti, V., Clerici, M., Lucchini, G., and Longhese, M.P. (2000). The checkpoint protein Ddc2, functionally related to S. pombe Rad26, interacts with Mec1 and is regulated by Mec1-dependent phosphorylation in budding yeast. Genes Dev 14, 2046-2059.

Paeschke, K., Bochman, M.L., Garcia, P.D., Cejka, P., Friedman, K.L., Kowalczykowski, S.C., and Zakian, V.A. (2013). Pif1 family helicases suppress genome instability at G-quadruplex motifs. Nature 497, 458- 462.

Page, B.D., Satterwhite, L.L., Rose, M.D., and Snyder, M. (1994). Localization of the Kar3 kinesin heavy chain-related protein requires the Cik1 interacting protein. J Cell Biol 124, 507-519.

Palladino, F., Laroche, T., Gilson, E., Axelrod, A., Pillus, L., and Gasser, S.M. (1993). SIR3 and SIR4 proteins are required for the positioning and integrity of yeast telomeres. Cell 75, 543-555.

Panier, S., and Durocher, D. (2009). Regulatory ubiquitylation in response to DNA double-strand breaks. DNA Repair (Amst) 8, 436-443.

Panier, S., Ichijima, Y., Fradet-Turcotte, A., Leung, C.C., Kaustov, L., Arrowsmith, C.H., and Durocher, D. (2012). Tandem protein interaction modules organize the ubiquitin-dependent response to DNA double- strand breaks. Mol Cell 47, 383-395.

174

Pardo, B., and Marcand, S. (2005). Rap1 prevents telomere fusions by nonhomologous end joining. EMBO J 24, 3117-3127.

Park, P.U., Defossez, P.A., and Guarente, L. (1999). Effects of mutations in DNA repair genes on formation of ribosomal DNA circles and life span in Saccharomyces cerevisiae. Mol Cell Biol 19, 3848-3856.

Parsons, C.A., and West, S.C. (1988). Resolution of model Holliday junctions by yeast endonuclease is dependent upon homologous DNA sequences. Cell 52, 621-629.

Paull, T.T., and Gellert, M. (1998). The 3' to 5' exonuclease activity of Mre 11 facilitates repair of DNA double-strand breaks. Mol Cell 1, 969-979.

Paulovich, A.G., Margulies, R.U., Garvik, B.M., and Hartwell, L.H. (1997). RAD9, RAD17, and RAD24 are required for S phase regulation in Saccharomyces cerevisiae in response to DNA damage. Genetics 145, 45-62.

Peng, C.Y., Graves, P.R., Thoma, R.S., Wu, Z., Shaw, A.S., and Piwnica-Worms, H. (1997). Mitotic and G2 checkpoint control: regulation of 14-3-3 protein binding by phosphorylation of Cdc25C on serine-216. Science 277, 1501-1505.

Pennaneach, V., Putnam, C.D., and Kolodner, R.D. (2006). Chromosome healing by de novo telomere addition in Saccharomyces cerevisiae. Mol Microbiol 59, 1357-1368.

Pennock, E., Buckley, K., and Lundblad, V. (2001). Cdc13 delivers separate complexes to the telomere for end protection and replication. Cell 104, 387-396.

Petreaca, R.C., Chiu, H.C., and Nugent, C.I. (2007). The role of Stn1p in Saccharomyces cerevisiae telomere capping can be separated from its interaction with Cdc13p. Genetics 177, 1459-1474.

Petukhova, G., Stratton, S., and Sung, P. (1998). Catalysis of homologous DNA pairing by yeast Rad51 and Rad54 proteins. Nature 393, 91-94.

Phillips, J.A., Chan, A., Paeschke, K., and Zakian, V.A. (2015). The pif1 helicase, a negative regulator of telomerase, acts preferentially at long telomeres. PLoS Genet 11, e1005186.

Piazza, A., Serero, A., Boule, J.B., Legoix-Ne, P., Lopes, J., and Nicolas, A. (2012). Stimulation of gross chromosomal rearrangements by the human CEB1 and CEB25 minisatellites in Saccharomyces cerevisiae depends on G-quadruplexes or Cdc13. PLoS Genet 8, e1003033.

Pinsky, B.A., Kung, C., Shokat, K.M., and Biggins, S. (2006). The Ipl1-Aurora protein kinase activates the spindle checkpoint by creating unattached kinetochores. Nat Cell Biol 8, 78-83.

Price, B.D., and D'Andrea, A.D. (2013). Chromatin remodeling at DNA double-strand breaks. Cell 152, 1344-1354.

Puglisi, A., Bianchi, A., Lemmens, L., Damay, P., and Shore, D. (2008). Distinct roles for yeast Stn1 in telomere capping and telomerase inhibition. EMBO J 27, 2328-2339.

Purvis, A., and Singleton, M.R. (2008). Insights into kinetochore-DNA interactions from the structure of Cep3Delta. EMBO Rep 9, 56-62.

175

Putnam, C.D., Pennaneach, V., and Kolodner, R.D. (2004). Chromosome healing through terminal deletions generated by de novo telomere additions in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 101, 13262-13267.

Putnam, C.D., Pennaneach, V., and Kolodner, R.D. (2005). Saccharomyces cerevisiae as a model system to define the chromosomal instability phenotype. Mol Cell Biol 25, 7226-7238.

Qi, H., and Zakian, V.A. (2000). The Saccharomyces telomere-binding protein Cdc13p interacts with both the catalytic subunit of DNA polymerase alpha and the telomerase-associated est1 protein. Genes Dev 14, 1777-1788.

Rabl, C. (1885). Über Zelltheilung. Morph Jb 10, 214-330.

Rangasamy, D., Berven, L., Ridgway, P., and Tremethick, D.J. (2003). Pericentric heterochromatin becomes enriched with H2A.Z during early mammalian development. EMBO J 22, 1599-1607.

Rattray, A.J., and Symington, L.S. (1994). Use of a chromosomal inverted repeat to demonstrate that the RAD51 and RAD52 genes of Saccharomyces cerevisiae have different roles in mitotic recombination. Genetics 138, 587-595.

Ray, A., and Runge, K.W. (1998). The C terminus of the major yeast telomere binding protein Rap1p enhances telomere formation. Mol Cell Biol 18, 1284-1295.

Ray, A., and Runge, K.W. (1999). The yeast telomere length counting machinery is sensitive to sequences at the telomere-nontelomere junction. Mol Cell Biol 19, 31-45.

Renkawitz, J., Lademann, C.A., and Jentsch, S. (2014). Mechanisms and principles of homology search during recombination. Nat Rev Mol Cell Biol 15, 369-383.

Renkawitz, J., Lademann, C.A., Kalocsay, M., and Jentsch, S. (2013). Monitoring homology search during DNA double-strand break repair in vivo. Mol Cell 50, 261-272.

Ribes-Zamora, A., Mihalek, I., Lichtarge, O., and Bertuch, A.A. (2007). Distinct faces of the Ku heterodimer mediate DNA repair and telomeric functions. Nat Struct Mol Biol 14, 301-307.

Ribeyre, C., Lopes, J., Boule, J.B., Piazza, A., Guedin, A., Zakian, V.A., Mergny, J.L., and Nicolas, A. (2009). The yeast Pif1 helicase prevents genomic instability caused by G-quadruplex-forming CEB1 sequences in vivo. PLoS Genet 5, e1000475.

Rieder, C.L., Cole, R.W., Khodjakov, A., and Sluder, G. (1995). The checkpoint delaying anaphase in response to chromosome monoorientation is mediated by an inhibitory signal produced by unattached kinetochores. J Cell Biol 130, 941-948.

Rine, J., and Herskowitz, I. (1987). Four genes responsible for a position effect on expression from HML and HMR in Saccharomyces cerevisiae. Genetics 116, 9-22.

Ritchie, K.B., Mallory, J.C., and Petes, T.D. (1999). Interactions of TLC1 (which encodes the RNA subunit of telomerase), TEL1, and MEC1 in regulating telomere length in the yeast Saccharomyces cerevisiae. Mol Cell Biol 19, 6065-6075.

176

Robert, T., Vanoli, F., Chiolo, I., Shubassi, G., Bernstein, K.A., Rothstein, R., Botrugno, O.A., Parazzoli, D., Oldani, A., Minucci, S., et al. (2011). HDACs link the DNA damage response, processing of double- strand breaks and autophagy. Nature 471, 74-79.

Robinett, C.C., Straight, A., Li, G., Willhelm, C., Sudlow, G., Murray, A., and Belmont, A.S. (1996). In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J Cell Biol 135, 1685-1700.

Rogakou, E.P., Boon, C., Redon, C., and Bonner, W.M. (1999). Megabase chromatin domains involved in DNA double-strand breaks in vivo. J Cell Biol 146, 905-916.

Rogakou, E.P., Pilch, D.R., Orr, A.H., Ivanova, V.S., and Bonner, W.M. (1998). DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273, 5858-5868.

Rohner, S., Gasser, S.M., and Meister, P. (2008). Modules for cloning-free chromatin tagging in Saccharomyces cerevisae. Yeast 25, 235-239.

Roos, W.P., and Kaina, B. (2006). DNA damage-induced cell death by apoptosis. Trends in molecular medicine 12, 440-450.

Roukos, V., Voss, T.C., Schmidt, C.K., Lee, S., Wangsa, D., and Misteli, T. (2013). Spatial dynamics of chromosome translocations in living cells. Science 341, 660-664.

Rouse, J., and Jackson, S.P. (2000). LCD1: an essential gene involved in checkpoint control and regulation of the MEC1 signalling pathway in Saccharomyces cerevisiae. EMBO J 19, 5801-5812.

Roux, K.J., Kim, D.I., and Burke, B. (2013). BioID: a screen for protein-protein interactions. Current protocols in protein science / editorial board, John E Coligan [et al] 74, Unit 19 23.

Roux, K.J., Kim, D.I., Raida, M., and Burke, B. (2012). A promiscuous biotin ligase fusion protein identifies proximal and interacting proteins in mammalian cells. J Cell Biol 196, 801-810.

Rusche, L.N., Kirchmaier, A.L., and Rine, J. (2003). The establishment, inheritance, and function of silenced chromatin in Saccharomyces cerevisiae. Annu Rev Biochem 72, 481-516.

Sabatier, L., Ricoul, M., Pottier, G., and Murnane, J.P. (2005). The loss of a single telomere can result in instability of multiple chromosomes in a human tumor cell line. Molecular cancer research : MCR 3, 139- 150.

Sabourin, M., Tuzon, C.T., and Zakian, V.A. (2007). Telomerase and Tel1p preferentially associate with short telomeres in S. cerevisiae. Mol Cell 27, 550-561.

Sage, D., Neumann, F.R., Hediger, F., Gasser, S.M., and Unser, M. (2005). Automatic tracking of individual fluorescence particles: Application to the study of chromosome dynamics. IEEE Transactions on Image Processing 14, 1372-1383.

Saini, N., Ramakrishnan, S., Elango, R., Ayyar, S., Zhang, Y., Deem, A., Ira, G., Haber, J.E., Lobachev, K.S., and Malkova, A. (2013). Migrating bubble during break-induced replication drives conservative DNA synthesis. Nature 502, 389-392.

Sancar, A., and Tang, M.S. (1993). Nucleotide excision repair. Photochemistry and photobiology 57, 905- 921.

177

Sanchez, Y., Bachant, J., Wang, H., Hu, F., Liu, D., Tetzlaff, M., and Elledge, S.J. (1999). Control of the DNA damage checkpoint by chk1 and rad53 protein kinases through distinct mechanisms. Science 286, 1166-1171.

Sanchez, Y., Wong, C., Thoma, R.S., Richman, R., Wu, Z., Piwnica-Worms, H., and Elledge, S.J. (1997). Conservation of the Chk1 checkpoint pathway in mammals: linkage of DNA damage to Cdk regulation through Cdc25. Science 277, 1497-1501.

Sandell, L.L., and Zakian, V.A. (1993). Loss of a yeast telomere: arrest, recovery, and chromosome loss. Cell 75, 729-739.

Sanders, C.M. (2010). Human Pif1 helicase is a G-quadruplex DNA-binding protein with G-quadruplex DNA-unwinding activity. Biochem J 430, 119-128.

Santocanale, C., and Diffley, J.F. (1998). A Mec1- and Rad53-dependent checkpoint controls late-firing origins of DNA replication. Nature 395, 615-618.

Savitsky, K., Bar-Shira, A., Gilad, S., Rotman, G., Ziv, Y., Vanagaite, L., Tagle, D.A., Smith, S., Uziel, T., Sfez, S., et al. (1995). A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science 268, 1749-1753.

Schatz, D.G., and Ji, Y. (2011). Recombination centres and the orchestration of V(D)J recombination. Nature reviews Immunology 11, 251-263.

Schober, H., Ferreira, H., Kalck, V., Gehlen, L.R., and Gasser, S.M. (2009). Yeast telomerase and the SUN domain protein Mps3 anchor telomeres and repress subtelomeric recombination. Genes Dev 23, 928-938.

Schulz, V.P., and Zakian, V.A. (1994). The saccharomyces PIF1 DNA helicase inhibits telomere elongation and de novo telomere formation. Cell 76, 145-155.

Schwartz, M.F., Duong, J.K., Sun, Z., Morrow, J.S., Pradhan, D., and Stern, D.F. (2002). Rad9 phosphorylation sites couple Rad53 to the Saccharomyces cerevisiae DNA damage checkpoint. Mol Cell 9, 1055-1065.

Scott, K.L., and Plon, S.E. (2003). Loss of Sin3/Rpd3 histone deacetylase restores the DNA damage response in checkpoint-deficient strains of Saccharomyces cerevisiae. Mol Cell Biol 23, 4522-4531.

Seeber, A., Dion, V., and Gasser, S.M. (2013). Checkpoint kinases and the INO80 nucleosome remodeling complex enhance global chromatin mobility in response to DNA damage. Genes Dev 27, 1999-2008.

Seto, A.G., Livengood, A.J., Tzfati, Y., Blackburn, E.H., and Cech, T.R. (2002). A bulged stem tethers Est1p to telomerase RNA in budding yeast. Genes Dev 16, 2800-2812.

Sfeir, A., and de Lange, T. (2012). Removal of shelterin reveals the telomere end-protection problem. Science 336, 593-597.

Sfeir, A., and Symington, L.S. (2015). Microhomology-Mediated End Joining: A Back-up Survival Mechanism or Dedicated Pathway? Trends Biochem Sci 40, 701-714.

Shampay, J., Szostak, J.W., and Blackburn, E.H. (1984). DNA sequences of telomeres maintained in yeast. Nature 310, 154-157.

178

Shay, J.W., and Bacchetti, S. (1997). A survey of telomerase activity in human cancer. Eur J Cancer 33, 787-791.

Shay, J.W., and Wright, W.E. (2011). Role of telomeres and telomerase in cancer. Seminars in cancer biology 21, 349-353.

Shimogawa, M.M., Graczyk, B., Gardner, M.K., Francis, S.E., White, E.A., Ess, M., Molk, J.N., Ruse, C., Niessen, S., Yates, J.R., 3rd, et al. (2006). Mps1 phosphorylation of Dam1 couples kinetochores to microtubule plus ends at metaphase. Curr Biol 16, 1489-1501.

Shinohara, A., Ogawa, H., and Ogawa, T. (1992). Rad51 protein involved in repair and recombination in S. cerevisiae is a RecA-like protein. Cell 69, 457-470.

Shinohara, A., and Ogawa, T. (1998). Stimulation by Rad52 of yeast Rad51-mediated recombination. Nature 391, 404-407.

Shore, D., and Nasmyth, K. (1987). Purification and cloning of a DNA binding protein from yeast that binds to both silencer and activator elements. Cell 51, 721-732.

Siede, W., Friedberg, A.S., and Friedberg, E.C. (1993). RAD9-dependent G1 arrest defines a second checkpoint for damaged DNA in the cell cycle of Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 90, 7985-7989.

Simsek, D., and Jasin, M. (2010). Alternative end-joining is suppressed by the canonical NHEJ component Xrcc4-ligase IV during chromosomal translocation formation. Nat Struct Mol Biol 17, 410-416.

Small, E.C., Xi, L., Wang, J.P., Widom, J., and Licht, J.D. (2014). Single-cell nucleosome mapping reveals the molecular basis of gene expression heterogeneity. Proc Natl Acad Sci U S A 111, E2462-2471.

Smith, C.E., Llorente, B., and Symington, L.S. (2007). Template switching during break-induced replication. Nature 447, 102-105.

Smolka, M.B., Albuquerque, C.P., Chen, S.H., and Zhou, H. (2007). Proteome-wide identification of in vivo targets of DNA damage checkpoint kinases. Proc Natl Acad Sci U S A 104, 10364-10369.

Song, H., and Endow, S.A. (1998). Decoupling of nucleotide- and microtubule-binding sites in a kinesin mutant. Nature 396, 587-590.

Soria, G., Polo, S.E., and Almouzni, G. (2012). Prime, repair, restore: the active role of chromatin in the DNA damage response. Mol Cell 46, 722-734.

Soutoglou, E., Dorn, J.F., Sengupta, K., Jasin, M., Nussenzweig, A., Ried, T., Danuser, G., and Misteli, T. (2007). Positional stability of single double-strand breaks in mammalian cells. Nat Cell Biol 9, 675-682.

Spangler, E.A., Ryan, T., and Blackburn, E.H. (1988). Developmentally regulated telomere addition in Tetrahymena thermophila. Nucleic Acids Res 16, 5569-5585.

Sproul, L.R., Anderson, D.J., Mackey, A.T., Saunders, W.S., and Gilbert, S.P. (2005). Cik1 targets the minus-end kinesin depolymerase kar3 to microtubule plus ends. Curr Biol 15, 1420-1427.

179

Stellwagen, A.E., Haimberger, Z.W., Veatch, J.R., and Gottschling, D.E. (2003). Ku interacts with telomerase RNA to promote telomere addition at native and broken chromosome ends. Genes Dev 17, 2384-2395.

Stoler, S., Keith, K.C., Curnick, K.E., and Fitzgerald-Hayes, M. (1995). A mutation in CSE4, an essential gene encoding a novel chromatin-associated protein in yeast, causes chromosome nondisjunction and cell cycle arrest at mitosis. Genes Dev 9, 573-586.

Strom, L., Lindroos, H.B., Shirahige, K., and Sjogren, C. (2004). Postreplicative recruitment of cohesin to double-strand breaks is required for DNA repair. Mol Cell 16, 1003-1015.

Sun, J., Yang, Y., Wan, K., Mao, N., Yu, T.Y., Lin, Y.C., DeZwaan, D.C., Freeman, B.C., Lin, J.J., Lue, N.F., et al. (2011). Structural bases of dimerization of yeast telomere protein Cdc13 and its interaction with the catalytic subunit of DNA polymerase alpha. Cell Res 21, 258-274.

Sung, P. (1997a). Function of yeast Rad52 protein as a mediator between replication protein A and the Rad51 recombinase. J Biol Chem 272, 28194-28197.

Sung, P. (1997b). Yeast Rad55 and Rad57 proteins form a heterodimer that functions with replication protein A to promote DNA strand exchange by Rad51 recombinase. Genes Dev 11, 1111-1121.

Sweeney, F.D., Yang, F., Chi, A., Shabanowitz, J., Hunt, D.F., and Durocher, D. (2005). Saccharomyces cerevisiae Rad9 acts as a Mec1 adaptor to allow Rad53 activation. Curr Biol 15, 1364-1375.

Szilard, R.K., Jacques, P.E., Laramee, L., Cheng, B., Galicia, S., Bataille, A.R., Yeung, M., Mendez, M., Bergeron, M., Robert, F., et al. (2010). Systematic identification of fragile sites via genome-wide location analysis of gamma-H2AX. Nat Struct Mol Biol 17, 299-305.

Szostak, J.W., Orr-Weaver, T.L., Rothstein, R.J., and Stahl, F.W. (1983). The double-strand-break repair model for recombination. Cell 33, 25-35.

Taddei, A., Hediger, F., Neumann, F.R., Bauer, C., and Gasser, S.M. (2004). Separation of silencing from perinuclear anchoring functions in yeast Ku80, Sir4 and Esc1 proteins. EMBO J 23, 1301-1312.

Taddei, A., Van Houwe, G., Hediger, F., Kalck, V., Cubizolles, F., Schober, H., and Gasser, S.M. (2006). Nuclear pore association confers optimal expression levels for an inducible yeast gene. Nature 441, 774- 778.

Taggart, A.K., Teng, S.C., and Zakian, V.A. (2002). Est1p as a cell cycle-regulated activator of telomere- bound telomerase. Science 297, 1023-1026.

Tanaka, K., Mukae, N., Dewar, H., van Breugel, M., James, E.K., Prescott, A.R., Antony, C., and Tanaka, T.U. (2005). Molecular mechanisms of kinetochore capture by spindle microtubules. Nature 434, 987-994.

Tanaka, T.U., Rachidi, N., Janke, C., Pereira, G., Galova, M., Schiebel, E., Stark, M.J., and Nasmyth, K. (2002). Evidence that the Ipl1-Sli15 (Aurora kinase-INCENP) complex promotes chromosome biorientation by altering kinetochore-spindle pole connections. Cell 108, 317-329.

Tao, R., Xue, H., Zhang, J., Liu, J., Deng, H., and Chen, Y.G. (2013). Deacetylase Rpd3 facilitates checkpoint adaptation by preventing Rad53 overactivation. Mol Cell Biol 33, 4212-4224.

180

Taylor, A.M., Harnden, D.G., Arlett, C.F., Harcourt, S.A., Lehmann, A.R., Stevens, S., and Bridges, B.A. (1975). Ataxia telangiectasia: a human mutation with abnormal radiation sensitivity. Nature 258, 427-429.

Teixeira, M.T., Arneric, M., Sperisen, P., and Lingner, J. (2004). Telomere length homeostasis is achieved via a switch between telomerase- extendible and -nonextendible states. Cell 117, 323-335.

Teng, S.C., and Zakian, V.A. (1999). Telomere-telomere recombination is an efficient bypass pathway for telomere maintenance in Saccharomyces cerevisiae. Mol Cell Biol 19, 8083-8093.

Tham, W.H., Wyithe, J.S., Ko Ferrigno, P., Silver, P.A., and Zakian, V.A. (2001). Localization of yeast telomeres to the nuclear periphery is separable from transcriptional repression and telomere stability functions. Mol Cell 8, 189-199.

Therizols, P., Fairhead, C., Cabal, G.G., Genovesio, A., Olivo-Marin, J.C., Dujon, B., and Fabre, E. (2006). Telomere tethering at the nuclear periphery is essential for efficient DNA double strand break repair in subtelomeric region. J Cell Biol 172, 189-199.

Toczyski, D.P., Galgoczy, D.J., and Hartwell, L.H. (1997). CDC5 and CKII control adaptation to the yeast DNA damage checkpoint. Cell 90, 1097-1106.

Torres-Rosell, J., Sunjevaric, I., De Piccoli, G., Sacher, M., Eckert-Boulet, N., Reid, R., Jentsch, S., Rothstein, R., Aragon, L., and Lisby, M. (2007). The Smc5-Smc6 complex and SUMO modification of Rad52 regulates recombinational repair at the ribosomal gene locus. Nat Cell Biol 9, 923-931.

Towbin, B.D., Meister, P., and Gasser, S.M. (2009). The nuclear envelope--a scaffold for silencing? Curr Opin Genet Dev 19, 180-186.

Trujillo, K.M., Roh, D.H., Chen, L., Van Komen, S., Tomkinson, A., and Sung, P. (2003). Yeast xrs2 binds DNA and helps target rad50 and mre11 to DNA ends. J Biol Chem 278, 48957-48964.

Trujillo, K.M., and Sung, P. (2001). DNA structure-specific nuclease activities in the Saccharomyces cerevisiae Rad50*Mre11 complex. J Biol Chem 276, 35458-35464.

Truong, L.N., Li, Y., Shi, L.Z., Hwang, P.Y., He, J., Wang, H., Razavian, N., Berns, M.W., and Wu, X. (2013). Microhomology-mediated End Joining and Homologous Recombination share the initial end resection step to repair DNA double-strand breaks in mammalian cells. Proc Natl Acad Sci U S A 110, 7720-7725.

Tseng, S.F., Lin, J.J., and Teng, S.C. (2006). The telomerase-recruitment domain of the telomere binding protein Cdc13 is regulated by Mec1p/Tel1p-dependent phosphorylation. Nucleic Acids Res 34, 6327-6336.

Tuzon, C.T., Wu, Y., Chan, A., and Zakian, V.A. (2011). The Saccharomyces cerevisiae telomerase subunit Est3 binds telomeres in a cell cycle- and Est1-dependent manner and interacts directly with Est1 in vitro. PLoS Genet 7, e1002060.

Tytell, J.D., and Sorger, P.K. (2006). Analysis of kinesin motor function at budding yeast kinetochores. J Cell Biol 172, 861-874.

Uhlmann, F., Lottspeich, F., and Nasmyth, K. (1999). Sister-chromatid separation at anaphase onset is promoted by cleavage of the cohesin subunit Scc1. Nature 400, 37-42.

181

Valencia, M., Bentele, M., Vaze, M.B., Herrmann, G., Kraus, E., Lee, S.E., Schar, P., and Haber, J.E. (2001). NEJ1 controls non-homologous end joining in Saccharomyces cerevisiae. Nature 414, 666-669. van Attikum, H., Fritsch, O., Hohn, B., and Gasser, S.M. (2004). Recruitment of the INO80 complex by H2A phosphorylation links ATP-dependent chromatin remodeling with DNA double-strand break repair. Cell 119, 777-788.

Vaze, M.B., Pellicioli, A., Lee, S.E., Ira, G., Liberi, G., Arbel-Eden, A., Foiani, M., and Haber, J.E. (2002). Recovery from checkpoint-mediated arrest after repair of a double-strand break requires Srs2 helicase. Mol Cell 10, 373-385.

Vazquez, J., Belmont, A.S., and Sedat, J.W. (2001). Multiple regimes of constrained chromosome motion are regulated in the interphase Drosophila nucleus. Curr Biol 11, 1227-1239.

Venclovas, C., and Thelen, M.P. (2000). Structure-based predictions of Rad1, Rad9, Hus1 and Rad17 participation in sliding clamp and clamp-loading complexes. Nucleic Acids Res 28, 2481-2493.

Verdaasdonk, J.S., Vasquez, P.A., Barry, R.M., Barry, T., Goodwin, S., Forest, M.G., and Bloom, K. (2013). Centromere tethering confines chromosome domains. Mol Cell 52, 819-831.

Vilenchik, M.M., and Knudson, A.G. (2003). Endogenous DNA double-strand breaks: production, fidelity of repair, and induction of cancer. Proc Natl Acad Sci U S A 100, 12871-12876.

Visintin, R., Prinz, S., and Amon, A. (1997). CDC20 and CDH1: a family of substrate-specific activators of APC-dependent proteolysis. Science 278, 460-463. von Hansemann, D. (1890). Ueber asynmetrische Zelltheilung in apithel Krebsen und deren biologische Bedeutung. Virchows Arch Path Anat 119, 299.

Wahlin, J., and Cohn, M. (2000). Saccharomyces cerevisiae RAP1 binds to telomeric sequences with spatial flexibility. Nucleic Acids Res 28, 2292-2301.

Wakayama, T., Kondo, T., Ando, S., Matsumoto, K., and Sugimoto, K. (2001). Pie1, a protein interacting with Mec1, controls cell growth and checkpoint responses in Saccharomyces cerevisiae. Mol Cell Biol 21, 755-764.

Walker, J.R., Corpina, R.A., and Goldberg, J. (2001). Structure of the Ku heterodimer bound to DNA and its implications for double-strand break repair. Nature 412, 607-614.

Wang, F., Stewart, J.A., Kasbek, C., Zhao, Y., Wright, W.E., and Price, C.M. (2012). Human CST has independent functions during telomere duplex replication and C-strand fill-in. Cell Rep 2, 1096-1103.

Wang, H., Liu, D., Wang, Y., Qin, J., and Elledge, S.J. (2001). Pds1 phosphorylation in response to DNA damage is essential for its DNA damage checkpoint function. Genes Dev 15, 1361-1372.

Warren, C.D., Eckley, D.M., Lee, M.S., Hanna, J.S., Hughes, A., Peyser, B., Jie, C., Irizarry, R., and Spencer, F.A. (2004). S-phase checkpoint genes safeguard high-fidelity sister chromatid cohesion. Mol Biol Cell 15, 1724-1735.

Weber, S.C., Spakowitz, A.J., and Theriot, J.A. (2010). Bacterial chromosomal loci move subdiffusively through a viscoelastic cytoplasm. Physical review letters 104, 238102.

182

Weber, S.C., Spakowitz, A.J., and Theriot, J.A. (2012). Nonthermal ATP-dependent fluctuations contribute to the in vivo motion of chromosomal loci. Proc Natl Acad Sci U S A 109, 7338-7343.

Weiner, A., Zauberman, N., and Minsky, A. (2009). Recombinational DNA repair in a cellular context: a search for the homology search. Nat Rev Microbiol 7, 748-755.

Weinert, T.A., and Hartwell, L.H. (1988). The RAD9 gene controls the cell cycle response to DNA damage in Saccharomyces cerevisiae. Science 241, 317-322.

Wiesmeijer, K., Krouwels, I.M., Tanke, H.J., and Dirks, R.W. (2008). Chromatin movement visualized with photoactivable GFP-labeled histone H4. Differentiation; research in biological diversity 76, 83-90.

Wigge, P.A., and Kilmartin, J.V. (2001). The Ndc80p complex from Saccharomyces cerevisiae contains conserved centromere components and has a function in chromosome segregation. J Cell Biol 152, 349- 360.

Wilkie, A.O., Lamb, J., Harris, P.C., Finney, R.D., and Higgs, D.R. (1990). A truncated human chromosome 16 associated with alpha thalassaemia is stabilized by addition of telomeric repeat (TTAGGG)n. Nature 346, 868-871.

Williamson, I., Berlivet, S., Eskeland, R., Boyle, S., Illingworth, R.S., Paquette, D., Dostie, J., and Bickmore, W.A. (2014). Spatial genome organization: contrasting views from chromosome conformation capture and fluorescence in situ hybridization. Genes Dev 28, 2778-2791.

Wilson, M.A., Kwon, Y., Xu, Y., Chung, W.H., Chi, P., Niu, H., Mayle, R., Chen, X., Malkova, A., Sung, P., et al. (2013). Pif1 helicase and Poldelta promote recombination-coupled DNA synthesis via bubble migration. Nature 502, 393-396.

Witkin, E.M. (1976). Ultraviolet mutagenesis and inducible DNA repair in Escherichia coli. Bacteriological reviews 40, 869-907.

Wold, M.S. (1997). Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu Rev Biochem 66, 61-92.

Wold, M.S., Weinberg, D.H., Virshup, D.M., Li, J.J., and Kelly, T.J. (1989). Identification of cellular proteins required for simian virus 40 DNA replication. J Biol Chem 264, 2801-2809.

Wong, A.C., Ning, Y., Flint, J., Clark, K., Dumanski, J.P., Ledbetter, D.H., and McDermid, H.E. (1997). Molecular characterization of a 130-kb terminal microdeletion at 22q in a child with mild mental retardation. American journal of human genetics 60, 113-120.

Wong, J., Nakajima, Y., Westermann, S., Shang, C., Kang, J.S., Goodner, C., Houshmand, P., Fields, S., Chan, C.S., Drubin, D., et al. (2007). A protein interaction map of the mitotic spindle. Mol Biol Cell 18, 3800-3809.

Workman, C.T., Mak, H.C., McCuine, S., Tagne, J.B., Agarwal, M., Ozier, O., Begley, T.J., Samson, L.D., and Ideker, T. (2006). A systems approach to mapping DNA damage response pathways. Science 312, 1054-1059.

Wotton, D., and Shore, D. (1997). A novel Rap1p-interacting factor, Rif2p, cooperates with Rif1p to regulate telomere length in Saccharomyces cerevisiae. Genes Dev 11, 748-760.

183

Wu, X., Wilson, T.E., and Lieber, M.R. (1999). A role for FEN-1 in nonhomologous DNA end joining: the order of strand annealing and nucleolytic processing events. Proc Natl Acad Sci U S A 96, 1303-1308.

Wu, Y., and Zakian, V.A. (2011). The telomeric Cdc13 protein interacts directly with the telomerase subunit Est1 to bring it to telomeric DNA ends in vitro. Proc Natl Acad Sci U S A 108, 20362-20369.

Wybenga-Groot, L.E., Ho, C.S., Sweeney, F.D., Ceccarelli, D.F., McGlade, C.J., Durocher, D., and Sicheri, F. (2014). Structural basis of Rad53 kinase activation by dimerization and activation segment exchange. Cell Signal 26, 1825-1836.

Xu, M., Kladde, M.P., Van Etten, J.L., and Simpson, R.T. (1998). Cloning, characterization and expression of the gene coding for a cytosine-5-DNA methyltransferase recognizing GpC. Nucleic Acids Res 26, 3961- 3966.

Yamamoto, A., Guacci, V., and Koshland, D. (1996). Pds1p, an inhibitor of anaphase in budding yeast, plays a critical role in the APC and checkpoint pathway(s). J Cell Biol 133, 99-110.

Yan, C.T., Boboila, C., Souza, E.K., Franco, S., Hickernell, T.R., Murphy, M., Gumaste, S., Geyer, M., Zarrin, A.A., Manis, J.P., et al. (2007). IgH class switching and translocations use a robust non-classical end-joining pathway. Nature 449, 478-482.

Yang, X.J., and Seto, E. (2008). The Rpd3/Hda1 family of lysine deacetylases: from bacteria and yeast to mice and men. Nat Rev Mol Cell Biol 9, 206-218.

Yeeles, J.T., Deegan, T.D., Janska, A., Early, A., and Diffley, J.F. (2015). Regulated eukaryotic DNA replication origin firing with purified proteins. Nature 519, 431-435.

Yeung, M., and Durocher, D. (2011). Srs2 enables checkpoint recovery by promoting disassembly of DNA damage foci from chromatin. DNA Repair (Amst) 10, 1213-1222.

Yu, X., and Gabriel, A. (2003). Ku-dependent and Ku-independent end-joining pathways lead to chromosomal rearrangements during double-strand break repair in Saccharomyces cerevisiae. Genetics 163, 843-856.

Yuen, K.W., Warren, C.D., Chen, O., Kwok, T., Hieter, P., and Spencer, F.A. (2007). Systematic genome instability screens in yeast and their potential relevance to cancer. Proc Natl Acad Sci U S A 104, 3925- 3930.

Zakian, V.A. (1996). Structure, function, and replication of Saccharomyces cerevisiae telomeres. Annu Rev Genet 30, 141-172.

Zappulla, D.C., and Cech, T.R. (2004). Yeast telomerase RNA: a flexible scaffold for protein subunits. Proc Natl Acad Sci U S A 101, 10024-10029.

Zhang, C.Z., Spektor, A., Cornils, H., Francis, J.M., Jackson, E.K., Liu, S., Meyerson, M., and Pellman, D. (2015). Chromothripsis from DNA damage in micronuclei. Nature 522, 179-184.

Zhang, D.H., Zhou, B., Huang, Y., Xu, L.X., and Zhou, J.Q. (2006). The human Pif1 helicase, a potential Escherichia coli RecD homologue, inhibits telomerase activity. Nucleic Acids Res 34, 1393-1404.

Zhang, W. (2012). Analysis of Telomere Healing of DNA Double-Strand Breaks (Doctoral Dissertation). In Department of Molecular Genetics (University of Toronto).

184

Zhang, W., and Durocher, D. (2010). De novo telomere formation is suppressed by the Mec1-dependent inhibition of Cdc13 accumulation at DNA breaks. Genes Dev 24, 502-515.

Zhao, X., Muller, E.G., and Rothstein, R. (1998). A suppressor of two essential checkpoint genes identifies a novel protein that negatively affects dNTP pools. Mol Cell 2, 329-340.

Zhao, X., and Rothstein, R. (2002). The Dun1 checkpoint kinase phosphorylates and regulates the ribonucleotide reductase inhibitor Sml1. Proc Natl Acad Sci U S A 99, 3746-3751.

Zhou, B.B., and Elledge, S.J. (2000). The DNA damage response: putting checkpoints in perspective. Nature 408, 433-439.

Zhou, J., Monson, E.K., Teng, S.C., Schulz, V.P., and Zakian, V.A. (2000). Pif1p helicase, a catalytic inhibitor of telomerase in yeast. Science 289, 771-774.

Zhou, R., Zhang, J., Bochman, M.L., Zakian, V.A., and Ha, T. (2014). Periodic DNA patrolling underlies diverse functions of Pif1 on R-loops and G-rich DNA. Elife 3, e02190.

Zhu, Z., Chung, W.H., Shim, E.Y., Lee, S.E., and Ira, G. (2008). Sgs1 helicase and two nucleases Dna2 and Exo1 resect DNA double-strand break ends. Cell 134, 981-994.

Zou, L., and Elledge, S.J. (2003). Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300, 1542-1548.