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Functional characterization of Rtt107 in the DNA damage response in Saccharomyces cerevisiae Leung, Grace Pui Yan 2014

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FUNCTIONAL CHARACTERIZATION OF RTT107 IN THE DNA DAMAGE RESPONSE IN SACCHAROMYCES CEREVISIAE  by Grace Pui Yan Leung  B.Sc., The University of British Columbia, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Genetics)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  July 2014  © Grace Pui Yan Leung, 2014 ii  Abstract  Living cells are constantly exposed to DNA damage that if not properly repaired may lead to mutations or cell death. Due to the crucial importance of genome integrity, cells have complex response mechanisms to sense and repair DNA lesions. In my dissertation, I focus on a protein called Rtt107 in the DNA damage response in Saccharomyces cerevisiae. Yeast lacking Rtt107 are hypersensitive to DNA damaging agents, but the exact role of Rtt107 remains unclear. The DNA damage response occurs in the context of chromatin, and thus is affected by chromatin modifiers. I explored the connection between Rtt107 and the histone methyltransferase Dot1. Deletion of DOT1 or loss of its target H3 K79 trimethylation suppressed the DNA damage sensitivity of rtt107∆ mutants. This suppression was mediated by the translesion synthesis bypass pathway, which improved DNA damage resistance, but increased mutation rates. Rtt107 interacts physically with a number of DNA repair proteins, and I characterized its interaction with the SMC5/6 complex, which has numerous roles in chromosome maintenance. Rtt107 was required for the recruitment of SMC5/6 to double-stranded breaks, but not protein-bound nicks. However, these proteins also contributed independently to other genome maintenance functions. I mapped conditional genetic interaction networks to further understand the functions of Rtt107, its interaction partner Slx4, and a poorly characterized helicase, Hrq1. RTT107 had the greatest number of conditional genetic interactions, particularly after exposure to camptothecin, suggesting that it has an important role in responding to this type of lesion. The genetic interaction profiles and associated gene ontology terms of HRQ1 was the most similar to that of SLX4, suggesting that they may work together in the DNA damage response. Focusing more closely iii  on the protein structure of Rtt107, I discovered that the C-terminal BRCT (BRCA1 C-terminal) domains bound to phosphorylated H2A, mediating the recruitment of Rtt107 to sites of DNA lesions. Rtt107 acted as a scaffold to recruit other proteins to these sites, but its requirement differed depending on the type of DNA insult. In summary, my dissertation contributes to the understanding of Rtt107 function, while highlighting key principles in the DNA damage response.  iv  Preface  Chapter 2 of this dissertation was published in the Journal of Biological Chemistry in November 2010 (Lévesque N, Leung GP, Fok AK, Schmidt TI, and Kobor MS, 2010). As co-first author I was involved in the formation of research questions, research design, data collection, data analysis and writing of the manuscript. I worked together with the other co-first authors, Nancy Lévesque and Alexandra Fok, to perform the experiments and create all the figures. I contributed experimentally to all the figures except Figure 2.5. Chapter 3 was published in the Journal of Biological Chemistry in July 2011 (Leung GP, Lee L, Schmidt TI, Shirahige K, and Kobor MS, 2011). As first author, I provided the primary contribution to research design, data collection, data analysis, and manuscript writing. I performed the experiments and created Figure 3.1, Figure 3.4, and Figure 3.5. Linda Lee performed the experiments for Figure 3.2, Figure 3.3, Figure 3.6, and Figure 3.7.  Chapter 4 was published in G3: Genes, Genomes, Genetics in June 2014 (Leung GP, Aristizabal MJ, Krogan NJ, and Kobor MS, 2014). I created all the yeast strains and Maria Aristizabal performed the cE-MAP experiments and analyzed the data. I performed the experiments for Figure 4.4 and Figure 4.7. I was responsible for writing the manuscript and creating all the figures.  Chapter 5 is based on experiments following up the work of Chapter 3. I performed all the experiments and analyzed the data for all the figures in this chapter. In Chapters 2-5, I consistently use “we” to reflect the co-authors that contributed to the studies. v  Table of Contents  Abstract .................................................................................................................................... ii	  Preface ..................................................................................................................................... iv	  Table of Contents .....................................................................................................................v	  List of Tables ............................................................................................................................ x	  List of Figures ......................................................................................................................... xi	  Acknowledgements .............................................................................................................. xiv	  Dedication ............................................................................................................................... xv	  Chapter 1: Introduction ..........................................................................................................1	  1.1	   The Problem of DNA Damage ................................................................................. 1	  1.2	   Detection of DNA Damage is the Initial Step .......................................................... 1	  1.3	   Activation of Signalling Cascades is Mediated by Checkpoint Kinases .................. 2	  1.4	   Checkpoint Adaptors Transmit Signals via Phosphorylation ................................... 4	  1.5	   Effector Kinases Facilitate Downstream Consequences ........................................... 5	  1.6	   Post-translational Modifications Play a Key Role in the DNA Damage Response .. 7	  1.7	   Multiple Repair Pathways Function in Response to DNA Lesions .......................... 9	  1.8	   DNA Lesions can be Bypassed with Post-replication Repair Pathways ................ 11	  1.9	   Recovery or Adaptation Follows After the DNA Damage Response ..................... 12	  1.10	   Chromatin is Intricately Involved in the DNA Damage Response ......................... 13	  1.11	   Rtt107 Functions in the DNA Damage Response ................................................... 15	  1.12	   Slx4 is an Interaction Partner of Rtt107 .................................................................. 18	  1.13	   Rtt107 Functions Together with the SMC5/6 Complex ......................................... 20	  vi  1.14	   Summary ................................................................................................................. 21	  Chapter 2: Loss of H3 K79 Trimethylation Leads to Suppression of Rtt107-Dependent DNA Damage Sensitivity Through the Translesion Synthesis Pathway ...........................24	  2.1	   Introduction ............................................................................................................. 24	  2.2	   Materials and Methods ............................................................................................ 28	  2.2.1	   Yeast strains ........................................................................................................ 28	  2.2.2	   Sensitivity measurements .................................................................................... 31	  2.2.3	   Protein extracts and protein blot analysis ........................................................... 31	  2.2.4	   Flow cytometric analysis and BrdU incorporation experiments ......................... 32	  2.2.5	   Quantitative bimater assay .................................................................................. 33	  2.2.6	   Microscopy ......................................................................................................... 33	  2.2.7	   Measurement of mutation rates ........................................................................... 34	  2.2.8	   Pulsed-field gel electrophoresis .......................................................................... 34	  2.3	   Results ..................................................................................................................... 35	  2.3.1	   Elimination of H3 K79 methylation suppressed the sensitivity of rtt107∆ and slx4Δ mutants to the DNA-damaging agent MMS ......................................................... 35	  2.3.2	   Deletion of DOT1 suppressed DNA damage sensitivity in the absence of MMS-induced phosphorylation of Rtt107 or Slx4 .................................................................... 41	  2.3.3	   The requirement of Rtt107 for resumption of cell cycle after S phase damage was partially suppressed by lack of Dot1 ....................................................................... 43	  2.3.4	   The TLS pathway was required for the suppression of the MMS sensitivity of rtt107∆ mutants by deletion of DOT1 ............................................................................ 48	  vii  2.3.5	   Rtt107 had functions in maintaining genomic integrity that were independent of Dot1 activity .................................................................................................................... 50	  2.4	   Discussion ............................................................................................................... 52	  Chapter 3: Rtt107 is Required for Recruitment of the Smc5/6 Complex to DNA Double-Strand Breaks .........................................................................................................................57	  3.1	   Introduction ............................................................................................................. 57	  3.2	   Materials and Methods ............................................................................................ 60	  3.2.1	   Yeast strains and plasmids .................................................................................. 60	  3.2.2	   Growth and DNA damage sensitivity assays ...................................................... 63	  3.2.3	   Analytical-scale interaction assays, immunoprecipitation, and phosphatase treatment ......................................................................................................................... 64	  3.2.4	   Yeast two-hybrid assays ..................................................................................... 66	  3.2.5	   Chromatin immunoprecipitation (ChIP) ............................................................. 66	  3.2.6	   Fluorescence microscopy .................................................................................... 67	  3.3	   Results ..................................................................................................................... 68	  3.3.1	   Rtt107 physically interacted with the SMC5/6 complex .................................... 68	  3.3.2	   Rtt107 was required for recruitment of the SMC5/6 complex to a DNA double-stranded break ................................................................................................................. 72	  3.3.3	   Mutations in the SMC5/6 complex resulted in phosphorylation of Rtt107 in the absence of DNA damage agents ..................................................................................... 75	  3.3.4	   Rtt107 and the SMC5/6 Complex had independent functions in the DNA damage response ............................................................................................................. 77	  3.4	   Discussion ............................................................................................................... 79	  viii  Chapter 4: Conditional Genetic Interactions of RTT107, SLX4, and HRQ1 Reveal Dynamic Networks Upon DNA Damage in S. Cerevisiae ...................................................84	  4.1	   Introduction ............................................................................................................. 84	  4.2	   Materials and Methods ............................................................................................ 87	  4.2.1	   Yeast strains ........................................................................................................ 87	  4.2.2	   Conditional epistatic miniarray profiling (cE-MAP) .......................................... 89	  4.2.3	   Growth & DNA damage sensitivity assays ........................................................ 89	  4.2.4	   Gene ontology analysis ....................................................................................... 90	  4.3	   Results ..................................................................................................................... 90	  4.3.1	   Genetic interaction profiles were considerably altered when exposed to DNA damaging agents .............................................................................................................. 90	  4.3.2	   RTT107 exhibited more conditional genetic interactions than SLX4 or HRQ1 .. 96	  4.3.3	   Genes with condition-specific interactions were enriched for functions in the DNA damage response ................................................................................................. 102	  4.3.4	   Deletion of HST3 and MRC1 suppressed the DNA damage sensitivity of rtt107∆ but not slx4∆ mutants .................................................................................................... 104	  4.4	   Discussion ............................................................................................................. 107	  Chapter 5: Rtt107 Acts as a Scaffold at DNA Lesions by Anchoring to Phosphorylated H2A........................................................................................................................................112	  5.1	   Introduction ........................................................................................................... 112	  5.2	   Materials and Methods .......................................................................................... 115	  5.2.1	   Yeast strains & plasmids ................................................................................... 115	  5.2.2	   Growth & DNA damage sensitivity assays ...................................................... 116	  ix  5.2.3	   Protein extracts and immunoblot analysis ........................................................ 116	  5.2.4	   Chromatin immunoprecipitation (ChIP) ........................................................... 117	  5.3	   Results ................................................................................................................... 118	  5.3.1	   Rtt107 BRCT domains were important for proper protein levels ..................... 118	  5.3.2	   Rtt107 recruitment to sites of DNA damage was mediated via binding of its fifth and sixth BRCT domains to γH2A ............................................................................... 120	  5.3.3	   Rtt107 was completely required for recruitment of Slx4, but not Dpb11, to DNA lesions 124	  5.4	   Discussion ............................................................................................................. 127	  Chapter 6: Conclusion .........................................................................................................130	  References .............................................................................................................................141	   x  List of Tables   Table 2.1 Yeast strains used in this study. .............................................................................. 28	  Table 3.1 Yeast strains used in this study. .............................................................................. 61	  Table 3.2 Primers used in this study for qPCR ....................................................................... 67	  Table 4.1 Yeast strains used in this study. .............................................................................. 88	  Table 4.2 Genes that showed condition-specific interactions with query genes that were statistically significant after multiple test correction (q<0.05). .............................................. 96	  Table 4.3 GO terms that were significantly enriched in the list of genetically interacting genes under all conditions using DAVID. ............................................................................ 103	  Table 5.1 Yeast strains used in this study. ............................................................................ 115	  Table 5.2 Primers used in this study for qPCR ..................................................................... 118	   xi  List of Figures   Figure 1.1 DNA damage signalling pathways consists of sensor, adaptor, and effector proteins. ..................................................................................................................................... 2	  Figure 1.2 The DNA damage signalling pathway is mediated by cross-talk within a network of protein complexes. ................................................................................................................ 5	  Figure 1.3 Rtt107 interacts with multiple protein complexes. ................................................ 17	  Figure 2.1 Abrogation of H3 K79 trimethylation suppressed the MMS sensitivity of strains lacking Rtt107 or Slx4. ........................................................................................................... 37	  Figure 2.2 Abrogation of H3 K79 trimethylation suppressed the MMS sensitivity of strains lacking Slx4. ........................................................................................................................... 39	  Figure 2.3 Suppression of rtt107∆ MMS sensitivity by deletion of DOT1 was not dependent on the phosphorylation of Slx4 and vice versa. ...................................................................... 42	  Figure 2.4 Requirement of Rtt107 for resumption of DNA replication and cell cycle after DNA damage was partially suppressed by deletion of DOT1. ............................................... 44	  Figure 2.5 Requirement of Rtt107 for resumption of replication after DNA damage was partially suppressed by deletion of DOT1. ............................................................................. 46	  Figure 2.6 Nuclear division delay of rtt107∆ mutants was not suppressed by deletion of DOT1 in the absence and presence of MMS. ......................................................................... 47	  Figure 2.7 Suppression of the rtt107Δ MMS sensitivity by deletion of DOT1 was dependent on the TLS pathway. ............................................................................................................... 49	  xii  Figure 2.8 Genomic instability of the rtt107∆ mutants was not suppressed by deletion of DOT1. ..................................................................................................................................... 51	  Figure 2.9 Model for repressive effect of chromatin modifications on DNA damage survival in rtt107∆ mutants. ................................................................................................................. 54	  Figure 3.1 Rtt107 physically interacted with the SMC5/6 complex. ...................................... 69	  Figure 3.2 Rtt107 interacted with multiple subunits of the SMC5/6 complex. ...................... 70	  Figure 3.3 Rtt107 interacted with the SMC5/6 complex via the Nse6 subunit in a yeast two-hybrid analysis. ....................................................................................................................... 71	  Figure 3.4 Rtt107 was required for recruitment of the SMC5/6 complex to a double-stranded break but not to a protein-bound nick. .................................................................................... 73	  Figure 3.5 In the absence of MMS, Rtt107 was phosphorylated in mutants expressing a compromised SMC5/6 complex. ............................................................................................ 76	  Figure 3.6 Rtt107 and the SMC5/6 complex had independent functions. .............................. 78	  Figure 3.7 Rtt107 and Slx4 had functions that were independent of the SMC5/6 complex. .. 79	  Figure 4.1 Schematic diagram of the cE-MAP workflow. ..................................................... 91	  Figure 4.2 Genetic interaction profiles changed in response to DNA damaging agents. ....... 93	  Figure 4.3 Distribution of S-scores differed between conditions or between query genes, as shown by cumulative probability plots. .................................................................................. 95	  Figure 4.4 cE-MAP data recapitulated the drug-specific genetic interactions between RTT107 and DOT1 or BRE1. ................................................................................................................ 98	  Figure 4.5 RTT107 had more significant genetic interactions than SLX4 or HRQ1. ............ 100	  Figure 4.6 Different patterns of genetic interactions were observed for genes that significantly changed their interactions in response to DNA damaging conditions. ............ 102	  xiii  Figure 4.7 Positive S-scores from the cE-MAP data were based on suppression of DNA damage sensitivity of rtt107∆ mutants. ................................................................................ 106	  Figure 5.1 Mutants of Rtt107 lacking N-terminal BRCT domains had lower protein levels................................................................................................................................................ 119	  Figure 5.2 Rtt107 mutants with additional BRCT domains had lower protein levels. ......... 120	  Figure 5.3 Rtt107 had physical and genetic links to γH2A. ................................................. 121	  Figure 5.4 Rtt107 recruitment to sites of DNA damage was mediated by its BRCT5/6 binding to γH2A. ................................................................................................................................ 122	  Figure 5.5 The protein levels of the Rtt107 K887M mutant was comparable to wild-type. 123	  Figure 5.6 Yeast containing Rtt107 mutants that were no longer recruited to DNA lesions were sensitive to DNA damaging agents. ............................................................................. 124	  Figure 5.7 Slx4 but not Dpb11 recruitment to DNA lesions was completely dependent on Rtt107. ................................................................................................................................... 126	  Figure 6.1 H2A S129 phosphorylation and H3 K79 methylation lead to inhibition of the TLS pathway via Rad53 activation. .............................................................................................. 132	    xiv  Acknowledgements   Firstly, I would like to thank my supervisor Dr. Michael Kobor for his mentorship and guidance throughout my scientific training. He took the risk to train me when I was only a naïve second year undergraduate, and it is his passion and enthusiasm for science that inspired me to pursue a career in scientific research. I consider it a special privilege to be one of the founding members of the lab and watch as the Kobor Lab grew and expanded over the years to encompass an incredible variety of research projects.  I would also like to thank my supervisory committee members Dr. Philip Hieter, Dr. Elizabeth Conibear, and Dr. Matthew Lorincz for their guidance and advice throughout my doctoral studies, as well as Dr. Hugh Brock for his support in the Genetics Graduate Program. I am thankful for the funding that the Natural Sciences and Engineering Research Council of Canada granted me to support my studies.  I am incredibly grateful for all the members of the Kobor Lab, past and present, who have made this lab a wonderful place to work in. I will always have many fond memories of the celebrations, disasters, hilarious moments, and everyday helpful discussions in the lab. I am especially thankful for Maria and Phoebe; we were an inseparable trio that journeyed together through the ups and downs of life as a graduate student. It was immeasurably easier and more enjoyable to struggle together and celebrate together, and I am truly grateful for their friendship and support.  Finally I want to thank my parents for their constant love and spoken and unspoken support as I progressed through my graduate education. xv  Dedication        “What I see in Nature is a grand design that we can comprehend only imperfectly,  and that must fill a thinking person with a feeling of humility…” -Albert Einstein 1  Chapter 1: Introduction   1.1 The Problem of DNA Damage DNA, the carrier of genetic information, is vulnerable to damage and modifications due to the chemical instability of its structure, errors in the replication machinery, and exposure to external genotoxic agents (Lindahl. 1993). If these aberrations in DNA structure are not properly repaired, they could compromise genomic integrity or ultimately lead to cell death.  Due to the vital importance of preserving the DNA structure, cells have a complex network of mechanisms to counter genomic insults called the DNA damage response (DDR). The DDR occurs in several closely related stages, starting with detection of DNA damage, and followed by cell cycle arrest and the appropriate repair mechanisms. If the repair is successful, the cell must restart the cell cycle, but if unsuccessful the cell is directed towards bypass pathways or apoptosis. The major proteins involved in the DDR are conserved from unicellular eukaryotes to humans, thus much of the molecular details have been elucidated in model organisms such as the budding yeast, Saccharomyces cerevisiae (Finn et al. 2012; Putnam et al. 2009). In this dissertation, the genes and proteins from S. cerevisiae will be discussed unless otherwise indicated.    1.2 Detection of DNA Damage is the Initial Step An enormous variety of lesions can be induced in DNA, ranging from alkylation to double-stranded breaks. Some lesions are immediately detected and repaired (described in more detail below), whereas others trigger activation of signalling cascades. The proteins involved 2  in these pathways are broadly categorized into sensors, adaptors, and effectors (Figure 1.1). When cells sustain DNA damage, they detect them using sensor proteins that recognize specific DNA structures. The exposed DNA ends in double stranded breaks (DSBs), arguably the most severe DNA lesion, are recognized by the MRX complex, composed of Mre11, Rad50, and Xrs2 (Ogawa et al. 1995). Alternatively, ssDNA exposed by replication stress or processing of other DNA lesions are recognized and bound by replication protein A (RPA) (Zou and Elledge. 2003). Although there are many types of DNA damage, they can all lead to these two types of DNA structures, thus facilitating efficient detection (Finn et al. 2012).   Figure 1.1 DNA damage signalling pathways consists of sensor, adaptor, and effector proteins.  Figure adapted from (van Attikum and Gasser. 2005). Proteins from S. cerevisiae are grouped by their primary role in the DNA damage response. Replication stress activates proteins specific to this type of DNA lesion, although some factors respond to other types of DNA damage as well.       1.3 Activation of Signalling Cascades is Mediated by Checkpoint Kinases Upon detection of DNA damage, signalling cascades are triggered to activate the appropriate downstream responses. These signals are primarily initiated by the central checkpoint 3  kinases, which belong to the phosphoinositol-3-kinase-related kinases (PIKK) family (Finn et al. 2012). Mec1 and Tel1 are the yeast homologues of mammalian ATR (ATM and Rad 3-related) and ATM (ataxia-telangiectasia mutated), respectively. The two kinases are partially redundant, since yeast lacking both kinases are synergistically sensitive to DNA damaging agents (Morrow et al. 1995). Although Mec1 is generally thought to be the primary kinase since mec1Δ mutants are more sensitive to DNA damaging agents than tel1Δ mutants, each kinase responds to specific types of signals (Putnam et al. 2009). Tel1 is activated by the MRX complex binding to DSB ends, whereas Mec1 is activated by RPA bound to ssDNA through its partner Ddc2 (Bandhu et al. 2014; Nakada et al. 2003; Rouse and Jackson. 2002; Zou and Elledge. 2003). However, both kinases interact with similar downstream machinery, providing a possible explanation for their partial redundancy.  Mec1 is also activated by additional pathways, which are differentially favoured depending on the cell cycle phase. One of these pathways is mediated by the checkpoint clamp 9-1-1 complex, which is an alternate version of the proliferating cell nuclear antigen (PCNA) replication clamp (Majka and Burgers. 2007). Loading of the 9-1-1 complex onto ss/ds junction DNA activates Mec1 through the 9-1-1 subunit Ddc1 (Bonilla et al. 2008; Majka et al. 2006). The replication protein Dpb11 is also able to activate Mec1, and Dpb11 itself is recruited to stalled replication forks by the 9-1-1 complex or DNA polymerase epsilon (Mordes et al. 2008; Navadgi-Patil and Burgers. 2008; Navadgi-Patil et al. 2011; Puddu et al. 2011). Another DNA replication factor, Dna2, is able to activate Mec1 using a mechanism of action similar to that of Ddc1 and Dpb11 (Kumar and Burgers. 2013).  4  1.4 Checkpoint Adaptors Transmit Signals via Phosphorylation After Mec1 and Tel1 are activated, they trigger a signalling cascade via the phosphorylation of the checkpoint adaptor Rad9 (Emili. 1998; Vialard et al. 1998) (Figure 1.2). Rad9 is recruited to sites of DNA damage via its Tudor and BRCT domains, which bind to H3 K79 methylation and H2A S129 phosphorylation, respectively (Grenon et al. 2007; Hammet et al. 2007). Rad9 is then phosphorylated by the cyclin-dependent kinase Cdc28 (Soulier and Lowndes. 1999), which forms a binding site for Dpb11 (Pfander and Diffley. 2011; Wang et al. 2012). Binding of Dpb11 brings Mec1 into proximity and promotes Mec1 phosphorylation of Rad9, which induces dimerization of Rad9 (Emili. 1998; Soulier and Lowndes. 1999; Vialard et al. 1998). Rad53 subsequently binds the phospho-site of Rad9 via the Rad53 FHA domains (Durocher et al. 2000; Sun et al. 1998). Recruitment of Rad53 facilitates Mec1 phosphorylation, as well as Rad53 auto-phosphorylation, which triggers full activation of Rad53 (Gilbert et al. 2001; Sweeney et al. 2005). Aside from Rad53, Rad9 is also required for Mec1 phosphorylation and activation of the additional checkpoint effector kinase Chk1 (Blankley and Lydall. 2004; Sanchez et al. 1999).  5   Figure 1.2 The DNA damage signalling pathway is mediated by cross-talk within a network of protein complexes.  Figure adapted from (Pfander and Diffley. 2011). Mec1 and its partner Ddc2 is recruited and activated by the parallel pathways involving the ssDNA binding proteins RPA and the 9-1-1 checkpoint clamp. Rad9 is recruited by binding to H3 K79 methylation and H2A S129 phosphorylation via its Tudor and BRCT domains, respectively. Dpb11 acts as a bridge to bring together Rad9 and Mec1, facilitating phosphorylation of Rad9 by Mec1, and subsequent phosphorylation of Rad53. Green arrows indicate phosphorylation. B1-B4, BRCT domains. AAD, ATR activation domain.  Mrc1 is the checkpoint adaptor specific for replication stress (Alcasabas et al. 2001). Mrc1 is a component of the replication machinery, and thus does not need to be recruited in the event of a stalled replication fork (Katou et al. 2003; Osborn and Elledge. 2003). When Mec1 is recruited to stalled replication forks via RPA binding to exposed ssDNA, it phosphorylates Mrc1, which in turn activates Rad53 (Katou et al. 2003; Osborn and Elledge. 2003).  1.5 Effector Kinases Facilitate Downstream Consequences In all eukaryotes there are two key effector kinases, Rad53 and Chk1, but in budding yeast Rad53 is the principal kinase while Chk1 has a minor role (Finn et al. 2012). Chk1 is much less studied and is believed to function only in G2/M (Liu et al. 2000; Sanchez et al. 1999). 6  These effector kinases continue on to phosphorylate a host of targets, which mediate a wide range of responses including cell cycle arrest, transcription changes, or histone modifications (described in more detail below).   Cell cycle arrest induced by checkpoint activation is presumably for the purpose of ensuring there is enough time for complete repair of the DNA lesion. If DNA damage occurs in G1 phase, Rad53 phosphorylates the G1 transcription factor Swi6, which inhibits expression of CLN1 and CLN2, which encode the G1 cyclins required for G1/S transition (Sidorova and Breeden. 1997). During replication stress, Rad53 is important for stabilization of stalled replication forks and inhibition of late origin firing (Lopes et al. 2001; Shirahige et al. 1998; Sogo et al. 2002; Tercero and Diffley. 2001). However, the largest contribution of Rad53 to cell viability during replication stress is prevention of irreversible replication fork collapse, although the exact mechanism for fork stabilization is still unclear (Segurado and Tercero. 2009; Tercero et al. 2003). Rad53 inhibits late origin firing by phosphorylating and inhibiting the replication initiation protein Sld3 and its regulator Dbf4 (Lopez-Mosqueda et al. 2010; Zegerman and Diffley. 2010).  There are multiple mechanisms that cooperate to induce G2/M arrest. Rad53 phosphorylates the polo-like kinase Cdc5, which subsequently inhibits the mitotic exit network through inhibition of Bub2/Bfa1 (Sanchez et al. 1999; Smits et al. 2000). Rad53 also works together with Chk1 to inhibit the metaphase to anaphase transition by preventing the degradation of yeast securin, Pds1 (Agarwal et al. 2003; Sanchez et al. 1999). Furthermore, Rad53 inhibits 7  the transcription of the G2/M genes by phosphorylation of the transcriptional activator Ndd1 (Edenberg et al. 2014; Yelamanchi et al. 2014).  One of the best characterized transcriptional effects of the DDR is the induction of the ribonucleotide reductase (RNR) genes mediated by the phosphorylation of the Dun1 kinase by Rad53 (Chen et al. 2007; Zhou and Elledge. 1993). Activated Dun1 mediates upregulation of Rnr2, the rate limiting step in deoxyribonucleotide (dNTP) synthesis, and degradation of Sml1, an inhibitor of ribonucleotide reductase (Huang and Elledge. 1997; Zhao and Rothstein. 2002). Survival of DNA damage in yeast is dependent on this increase in levels of dNTPs, which is why the lethality of mec1∆ or rad53∆ mutants is suppressed by deletion of SML1 (Chabes et al. 2003; Zhao et al. 1998). Aside from induction of the RNR genes, the DDR also influences the expression of many other genes, which were identified by a number of genome-wide studies. However, the majority of these gene changes are not unique to the DDR, but overlap with those induced by the environmental stress response (Gasch et al. 2001; Jaehnig et al. 2013; Workman et al. 2006).   1.6 Post-translational Modifications Play a Key Role in the DNA Damage Response A key theme in the DDR is the use of post-translational modifications (PTMs) to activate appropriate responses to the DNA lesions, thus acting as a “molecular switch”. Phosphorylation represents an important type of PTM, as exemplified by the central role of kinases in the DDR, and has been studied extensively. A number of studies have identified more than 50 targets of the checkpoint kinases by systematically analyzing phosphorylation 8  in the whole proteome in response to DNA damaging agents (Chen et al. 2010; Smolka et al. 2007).   Correspondingly, many proteins in the DDR also contain phospho-binding domains (Mohammad and Yaffe. 2009). One of the first phospho-binding domains identified was the 14-3-3 proteins (Muslin et al. 1996; Yaffe et al. 1997). Later studies discovered the forkhead associated (FHA) domain, which specifically binds to phosphothreonine and not phosphoserine (Durocher et al. 1999; Hofmann and Bucher. 1995). In contrast, the BRCT domains (BRCA1 C terminal) have a strong preference for phosphoserine (Manke et al. 2003; Yu et al. 2003). BRCT domains were originally identified in the protein encoded by the human breast cancer susceptibility gene BRCA1 and have subsequently been found in many DDR proteins (Bork et al. 1997; Koonin et al. 1996). One of these BRCT-domain containing proteins is Rtt107, the focus of my dissertation, and in Chapter 5 I describe the functional characterization of the BRCT domains in Rtt107.      In recent years, other types of PTMs have also been found to have important roles in the DDR, such as ubiquitination and sumoylation (Jackson and Durocher. 2013). Moreover, there is significant crosstalk between these two modifications. A classic example of ubiquitination in the DDR is the ubiquitination of PCNA to regulate the choice of post-replication repair pathways (Hoege et al. 2002; Lee and Myung. 2008). PCNA can also be sumoylated, adding an additional layer of regulation to its functions (Hoege et al. 2002). Recent large-scale studies have identified many proteins that are sumoylated in response to 9  DNA damage, and demonstrate that this PTM is important for mediating protein-protein interactions (Cremona et al. 2012; Psakhye and Jentsch. 2012).  1.7 Multiple Repair Pathways Function in Response to DNA Lesions The cell has a variety of specialized repair pathways which are designed to deal with specific types of DNA lesions. In a few specific cases, the DNA lesion can be directly reversed. Cyclobutane pyrimidine dimers caused by ultraviolet light are repaired by the Phr1 photolyase enzyme which removes the dimer and restores the DNA structure (Sancar. 2008). Another enzyme called Mgt1 is a DNA methyltransferase that removes methyl groups from O6-methylguanine and O4-methylthymine modified bases, thereby preventing base mispairings (Sassanfar and Samson. 1990; Xiao et al. 1991). Aside from these pathways, other repair pathways require more extensive processing of the DNA lesion.    Base excision repair (BER) repairs damage due to oxidative stress, hydrolysis, or deamination (Boiteux and Jinks-Robertson. 2013). First, the modified base is released from the deoxyribose by DNA N-glycosylase enzymes, of which there are a number that each recognize specific types of modified bases. Subsequently, an endonuclease/lyase nicks the DNA backbone at the resulting apyrimidinic/apurinic (AP) site, and a 3’- or 5’-phosphodiesterase removes the remaining deoxyribose phosphate residue. Finally, the gap that is created is filled by a DNA polymerase and sealed by a DNA ligase.   Nucleotide excision repair (NER) can also repair some of the same lesions as BER, but it is additionally able to remove large helix distorting lesions including 6-4 photoproducts, 10  adducts from chemical carcinogens, and intramolecular crosslinking from oxidative damage (Boiteux and Jinks-Robertson. 2013). There are two sub-pathways of NER, primarily differentiated by the lesion recognition step – global genome NER (GG-NER) and transcription coupled NER (TC-NER). In GG-NER, dedicated proteins recognize the lesion, whereas in TC-NER, the lesion is detected when RNA polymerase II is blocked. After the DNA lesion is detected, a pre-incision complex is formed to further unwind the DNA helix, and then endonucleases nick the DNA on either side of the complex to release a 25-30 nt long oligonucleotide containing the DNA lesion. As with BER, the final steps involve repair synthesis and ligation.   Mismatch repair (MMR) is responsible for correcting replication errors, thus limiting mutagenesis (Boiteux and Jinks-Robertson. 2013; Hsieh and Yamane. 2008). Because of its key role in replication, MMR is coordinated with PCNA, the replication sliding clamp. The pathway works by introducing a nick into the leading strand of replication, or using the existing nick in the lagging strand. The strand containing the mismatched base is removed by degradation or strand displacement, and the gap is filled by repair synthesis and ligation.   In general, these repair pathways do not activate the checkpoint signaling pathways if the repair is carried out quickly. However, the checkpoint can be activated if there is a build-up of ssDNA, which is recognized by RPA and subsequently activates Mec1 as described above. For example, the Exo1 nuclease can process the NER intermediates to create extended ssDNA regions, thus competing with repair synthesis to activate the Mec1 checkpoint kinase (Giannattasio et al. 2010).  11   The two major repair pathways for DSBs are homologous recombination (HR) and non-homologous end joining (NHEJ), since the previously described pathways are unable to repair broken DNA ends. HR is carried out by the RAD52 epistasis group and involves processing of the DSB by nucleases to generate ssDNA 3’ overhangs (Heyer et al. 2010). These single-stranded tails are coated by proteins to form a recombinase filament which invades into a homologous template to form a D-loop intermediate. DNA polymerase extends the 3’ end of the invading strand, and the gap is filled by a variety of mechanisms, only some of which results in crossover products.      NHEJ is primarily carried out by the yeast Ku proteins, which bind to the DNA ends and align the overhangs that are subsequently ligated together by a DNA ligase (Daley et al. 2005; Lieber. 2010). In a small proportion of cases, the ends are aligned by microhomologies, which may lead to small insertions, deletions, or substitutions.    The MRX complex that binds to DSB ends is involved in both the HR and NHEJ pathways, and is important for regulation of pathway choice (Huertas. 2010). HR occurs primarily in S and G2 phase and uses the sister chromatid as the preferred template, whereas NHEJ is used in G1 phase.  1.8 DNA Lesions can be Bypassed with Post-replication Repair Pathways There are also post-replication repair (PRR) pathways which allow bypass of DNA lesions, and are not true repair pathways (Boiteux and Jinks-Robertson. 2013). PRR consists of error-12  prone and error-free subpathways, which are regulated by ubiquitination of PCNA (Hoege et al. 2002; Lee and Myung. 2008). The first is the translesion synthesis (TLS) pathway which uses specialized low-fidelity DNA polymerases to allow for replication through DNA lesions (Waters et al. 2009). Consequently, this pathway is error-prone and causes increased frequency of mutations. I describe in Chapter 2 the unexpected connection between Rtt107 and the TLS pathway. The second pathway is error-free post-replication repair, which uses template switching of sister chromatids and appears to be similar to the HR pathway (Boiteux and Jinks-Robertson. 2013).   1.9 Recovery or Adaptation Follows After the DNA Damage Response In contrast to the well-studied DNA damage detection and repair pathways, much less is known about the recovery stage. After repair is complete, the checkpoint must be deactivated to allow resumption of the cell cycle. This process involves many players including phosphatases, proteases and helicases (Clemenson and Marsolier-Kergoat. 2009; Harrison and Haber. 2006). Consistent with the concept of molecular switches, phosphatases play an important role in the recovery stage to remove phosphorylation signals and thus deactivate the checkpoint. For example, yeast lacking the Rad53 phosphatases fail to resume DNA synthesis after removal of DNA damage (Leroy et al. 2003; O'Neill et al. 2007).   If the damage is not repaired, cells may be directed toward apoptosis or attempt to restart the cell cycle despite the potential for mutations. Under certain conditions, cells can escape arrest even in the continued presence of DNA lesions through adaptation (Bartek and Lukas. 2007). Very little is known about the mechanisms or regulation of adaptation, and only a few 13  proteins involved in this process have been identified (Lee et al. 1998; Sandell and Zakian. 1993; Toczyski et al. 1997).  1.10 Chromatin is Intricately Involved in the DNA Damage Response DNA in its native context is not a naked double helix, but packaged into chromatin structures, therefore the DDR by necessity involves chromatin. Initially the proposed model of chromatin involvement was “access, repair, restore”, which described the role of chromatin simply as a matter of regulating accessibility to the DNA (Green and Almouzni. 2002). However with further studies it has become clear that chromatin modifiers play a much more complex role – not only to regulate access to DNA, but also to signal in the DDR (Papamichos-Chronakis and Peterson. 2013).   The most well characterized and earliest chromatin response in the DDR is the phosphorylation of H2A on Ser 129 in yeast, or the histone variant H2A.X on Ser 139 in humans (Downs et al. 2000; Mannironi et al. 1989; Rogakou et al. 1998). In S. cerevisiae, H2A is primarily phosphorylated by Mec1 very rapidly after the damage occurs, and the modification is spread in both directions away from the site of the lesion to cover approximately 50 kb of chromatin (Cobb et al. 2005; Downs et al. 2000; Shroff et al. 2004). Phosphorylated H2A creates a binding platform for a variety of players in the DDR, including the checkpoint adaptor Rad9, as well as the chromatin modifiers NuA4 histone acetyltransferase complex, and the Ino80 and Swr1 chromatin remodeling complexes (Downs et al. 2004; Hammet et al. 2007). Consistent with its central role in the DDR, lack of H2A.X in mice results in DNA damage sensitivity and increased genomic instability 14  (Bassing et al. 2002; Celeste et al. 2002). Similarly, Schizosaccharomyces pombe cells containing a non-phosphorylatable mutant of H2A are hypersensitive to DNA damaging agents and are defective in checkpoint arrest (Nakamura et al. 2004). Surprisingly, the S. cerevisiae H2A S129A mutant is only mildly sensitive to DNA damaging agents and can still activate checkpoint arrest (Downs et al. 2000; Keogh et al. 2006; Redon et al. 2003).  Although H2A phosphorylation is the only histone modification triggered by the DDR, there are other histone marks that also influence the DDR. This includes methylation of H3 on Lys 79 by Dot1, regulated by ubiquitination of H2B on Lys 123 by the Rad6-Bre1 complex, which together are required for checkpoint activation in specific cell cycle phases (Dover et al. 2002; Giannattasio et al. 2005; Shilatifard. 2006; Wood et al. 2003; Wysocki et al. 2005). In part, this role in checkpoint activation is mediated through a functional linkage to the checkpoint adaptor Rad9 (Grenon et al. 2007; Wysocki et al. 2005). Furthermore, the Bre1- and Dot1-mediated histone modifications are also involved in regulation of various repair pathways, such as nucleotide excision repair, sister chromatid recombination, global genome nucleotide excision repair, and repair of ionizing radiation damage (Chaudhuri et al. 2009; Conde et al. 2009; Game et al. 2005; Grenon et al. 2007; Lazzaro et al. 2008; Tatum and Li. 2011). Interestingly, Dot1 appears to negatively regulate the translesion synthesis bypass pathway (Conde and San-Segundo. 2008), and characterization of this function in relation to Rtt107 activity is described in Chapter 2. Further examples of histone modifications in the DDR are the replication-associated histone marks, acetylation of H3 on Lys 56 by Rtt109 and phosphorylation by H3 on Thr 45 by Cdc7, which function specifically in the response to replication stress (Baker et al. 2010; Downs. 2008).    15   In addition to histone modifications, chromatin remodelers and histone chaperones also play a role in the DDR. As previously mentioned, H2A phosphorylation recruits the remodeling complexes Ino80 and Swr1 (Downs et al. 2004). These remodelers were thought to reorganize chromatin into a more open conformation (van Attikum and Gasser. 2009), but further work has revealed a more intimate connection to the DDR (Morrison and Shen. 2009). Ino80 evicts nucleosomes near DSBs, thus allowing efficient homologous repair, but it is also important for checkpoint adaptation (Papamichos-Chronakis et al. 2006; Tsukuda et al. 2009). Both Fun30 and Swr1 remodelers promote DNA end processing (Chen et al. 2012; Costelloe et al. 2012; Eapen et al. 2012; Kalocsay et al. 2009; van Attikum et al. 2007). The histone chaperones Asf1 and CAF-1 also impinge on the DDR, primarily through their well characterized role in nucleosome reassembly (Avvakumov et al. 2011).  1.11 Rtt107 Functions in the DNA Damage Response RTT107 (Regulator of Ty1 Transposition) was first identified in a screen for increased Ty1 transposition and later was independently identified in a targeted silencing screen and named ESC4 (Establish Silent Chromatin) (Andrulis et al. 2004; Scholes et al. 2001). Evidence from several genome-wide screens pointed to a role in the DDR (Chang et al. 2002; Hanway et al. 2002), and subsequently this function of Rtt107 was more extensively characterized by several groups.   Yeast lacking Rtt107 are hypersensitive to the DNA alkylating agent methyl methanesulfonate (MMS), the replication inhibitor hydroxyurea (HU), and the topoisomerase 16  I inhibitor camptothecin (CPT), but not to ultraviolet or ionizing radiation (Chang et al. 2002; Hanway et al. 2002; Rouse. 2004). Consistent with sensitivity of rtt107Δ cells to MMS, HU, and CPT, which all cause DNA damage specifically in S phase, Rtt107 is required for the resumption of DNA replication after DNA damage and is recruited to stalled replication forks (Roberts et al. 2008; Rouse. 2004). The acetyltransferase Rtt109 and the cullin Rtt101 is required for recruitment of Rtt107 to chromatin upon exposure to MMS or CPT (Roberts et al. 2008).  Rtt107 is phosphorylated at several S/T-Q motifs by the checkpoint kinase Mec1 in response to DNA damage, and this modification is important for Rtt107 function (Roberts et al. 2006; Rouse. 2004; Ullal et al. 2011). Rtt107 was also identified in a large screen as a sumoylation target after exposure to DNA damaging agents, but the functional significance of this PTM has not been investigated (Cremona et al. 2012).   Aside from the DDR, Rtt107 also has functions in genomic stability. Yeast lacking Rtt107 have increased foci of the DDR proteins Ddc2 and Rad52, indicating higher levels of spontaneous DNA damage (Alvaro et al. 2007; Roberts et al. 2006). Moreover, Rtt107 itself forms nuclear foci during S-phase, and the numbers of foci are increased upon exposure to MMS (Chin et al. 2006). rtt107Δ mutants also have increased chromosomal instability, and elevated rates of spontaneous gross chromosomal rearrangements and sister chromatid recombination (Rouse. 2004; Ullal et al. 2011; Yuen et al. 2007).   Rtt107 contains six BRCT domains, which bind to phosphorylated peptides (Rouse. 2004). The crystal structure of the two C-terminal BRCT domains of Rtt107 has been solved and shown to bind to the phospho-peptide of H2A (Li et al. 2012). This interaction mediates the 17  recruitment of Rtt107 to sites of DNA damage, as described in Chapter 5. The four N-terminal BRCT domains of Rtt107 are thought to mediate the interaction with other DDR proteins. Rtt107 physically interacts with a number of DNA repair proteins, including the Slx4/Slx1 nuclease complex, the SMC5/6 complex, and the DNA replication protein Dpb11 (Chin et al. 2006; Leung et al. 2011; Ohouo et al. 2010; Roberts et al. 2006) (Figure 1.3).    Figure 1.3 Rtt107 interacts with multiple protein complexes.  Rtt107 interacts with the SMC5/6 complex via its N-terminal portion binding to the Nse6 subunit of the SMC5/6 complex. Rtt107 also interacts with Slx4 via the N-terminal portion of Rtt107. After exposure to DNA damage (indicated by a red shadow), Dpb11 interacts with Slx4 and is stabilized by Rtt107. Multiple post-translational modifications occur after DNA damage, including phosphorylation of Rtt107 and Slx4, and sumoylation of Rtt107 and Smc5.  The exact function of Rtt107 in the DDR is still unclear, as well as its roles in maintaining genome stability. Evidence thus far seems to indicate a role for Rtt107 in responding to damage occurring in S phase.  Moreover, the presence of BRCT domains in its structure points towards a scaffolding role. In Chapter 4, I describe the use of genetic interaction profiling in DNA damaging conditions to further characterize Rtt107 function. The conditional genetic network revealed an important role for Rtt107 in the response to CPT-18  induced lesions. Additional investigation is needed to elucidate the specific DDR pathways that Rtt107 is involved in.   The homologue of Rtt107 in Schizosaccharomyces pombe is Brc1 (BRCT containing protein 1), which is named for its six BRCT domains. Like Rtt107, Brc1 is important for recovery after replication stress and for maintenance of genome stability (Bass et al. 2012; Verkade et al. 1999). Brc1 also has several functional connections to the SMC5/6 complex and the Slx4/Slx1 endonuclease (Sheedy et al. 2005; Verkade et al. 1999). In humans, the putative homologue of Rtt107 is Pax2 transactivation domain-interacting protein (PTIP) which also has roles in regulation of DDR signaling pathways and interacts with many key DDR players, although PTIP also has additional functions in embryonic development and transcription (Callen et al. 2013; Gohler et al. 2008; Gong et al. 2009; Jowsey et al. 2004; Munoz and Rouse. 2009; Wang et al. 2010; Wu et al. 2009; Yan et al. 2011). Notably, crystal structures of both Brc1 and PTIP show that they bind phosphorylated H2A via the fifth and sixth BRCT domains (Williams et al. 2010; Yan et al. 2011).  1.12 Slx4 is an Interaction Partner of Rtt107 SLX4, named for “synthetic lethal of unknown function”, was first identified in a screen for genes that were synthetic lethal with SGS1, which encodes for the RecQ helicase (Mullen et al. 2001). This synthetic lethality is due to the functional redundancy of Sgs1-Top3 endonuclease activity with Slx4-Slx1 (Fricke and Brill. 2003). The Slx4-Slx1 structure-specific endonuclease is responsible for maintaining ribosomal DNA structure during replication (Kaliraman and Brill. 2002). In addition to interacting with Slx1, Slx4 also forms 19  a mutually exclusive complex with the endonuclease Rad1/Rad10 (Flott et al. 2007). Slx4 also has potential functional links to Hrq1, a novel RecQ helicase that was recently identified (Barea et al. 2008; Kwon et al. 2012). A close relationship between SLX4 and HRQ1 was suggested by the correlation of their genetic interaction profiles under DNA damaging conditions, which is described in Chapter 4.    Consistent with the multiple interaction partners, the mammalian homologue SLX4 interacts with a number of endonucleases and forms a “molecular toolkit” for Holliday junction resolution and telomere maintenance (Castor et al. 2013; Garner et al. 2013; Sengerova et al. 2011; Wan et al. 2013; Wilson et al. 2013; Wyatt et al. 2013). Human SLX4 is a member of the Fanconi anemia pathway that repairs interstrand crosslinks (Crossan et al. 2011; Kim et al. 2011; Stoepker et al. 2011; Yamamoto et al. 2011). Mutations in SLX4 cause a subtype of Fanconi anemia (Kim et al. 2011; Schuster et al. 2013; Stoepker et al. 2011), but efforts to discover mutations in breast cancer cases in multiple cohorts revealed only a minor contribution of SLX4 mutations (Bakker et al. 2013; Catucci et al. 2012; de Garibay et al. 2013; Fernandez-Rodriguez et al. 2012; Landwehr et al. 2011; Shah et al. 2013). Interestingly, a recent study reported that SLX4 is targeted by the HIV auxiliary protein Vpr during viral infection to cause G2/M arrest (Laguette et al. 2014).      Deletion of SLX4 confers many of the same DNA repair defects observed in cells lacking RTT107 and, like Rtt107, facilitates resumption of DNA replication after DNA damage (Roberts et al. 2006). This function of Slx4 in the DDR appears to be independent of Slx1, as deletion of SLX1 does not confer sensitivity to DNA damaging agents and Slx1 is 20  dispensable for the interaction between Rtt107 and Slx4 (Roberts et al. 2006). Further supporting a close relationship between Slx4 and Rtt107, they are mutually required for their respective Mec1-dependent phosphorylation (Levesque et al. 2010; Roberts et al. 2006). Upon exposure to DNA damaging agents, Slx4 and Rtt107 bind to Dpb11, a protein that has roles in DNA replication, checkpoint control, and DDR (Garcia et al. 2005; Ohouo et al. 2010). Rtt107/Slx4 competes with the checkpoint adaptor Rad9 to bind to Dpb11, thus regulating the activation of the downstream checkpoint kinase Rad53 (Ohouo et al. 2013).  1.13 Rtt107 Functions Together with the SMC5/6 Complex Another interaction partner of Rtt107 is the SMC5/6 complex, which is a complex involved in DNA repair and chromosome segregation (Strom and Sjogren. 2007). The core subunits of this complex, Smc5 and Smc6, are members of the structural maintenance of chromosomes (SMC) proteins, which also include cohesin (SMC1/3) and condensin (SMC2/4). In addition to Smc5 and Smc6, the complex includes six non-Smc elements (Nse1–6), and all the subunits are encoded by essential genes in S. cerevisiae. The SMC5/6 complex functions in the DDR, affecting the homologous recombination pathway (De Piccoli et al. 2009). It also has additional non-repair roles such as ensuring proper replication of the ribosomal DNA and relieving topological stress in replicating chromosomes (Kegel and Sjogren. 2010). Furthermore, similar to the other SMC complexes, SMC5/6 is involved in facilitating proper chromosome segregation during meiosis.   The function of the SMC5/6 is conserved in humans, as the human complex is also involved in DNA repair and recombination (Potts et al. 2006; Stephan et al. 2011; Wu et al. 2012). 21  Moreover, human SMC5/6 is also important for proper DNA replication under unperturbed conditions (Gallego-Paez et al. 2014). The SMC family of proteins is of fundamental importance for biology, as mutations in genes encoding human SMC proteins have been associated with the Cornelia de Lange and Roberts phocomelia syndromes (Ball et al. 2014).   Rtt107 and SMC5/6 are mutually required for their recruitment to DSBs, revealing a close functional connection between the two that may explain the significance of their physical interaction (Leung et al. 2011; Ullal et al. 2011). Interestingly, the interaction between Rtt107 and SMC5/6 is independent of Slx4, which shares many of Rtt107’s other functions (Leung et al. 2011). The function of the SMC5/6 complex impinges on Rtt107 phosphorylation in a complex manner (Ullal et al. 2011), suggesting a multifaceted regulatory network between them that has yet to be fully elucidated. The work characterizing this interaction between Rtt107 and the SMC5/6 complex is described in Chapter 3.  1.14 Summary The DDR is necessarily complex to properly respond to the multitude of DNA lesions that may occur so that each challenge can be met in their specific contexts. The cell utilizes a plethora of protein machineries that fulfill these roles, and although we have identified many of them, there still remains many more whose functions are unclear. In this dissertation, I characterize the functions of one such protein, Rtt107, and its interaction partners, using a combination of biochemical and genetic approaches.   22  In Chapter 2 I describe the relationship between Rtt107 and the histone methyltransferase Dot1. Loss of trimethylation of H3 Lys 79 suppressed the DNA damage-associated phenotypes of rtt107∆ mutants, but not other phenotypes involved with spontaneous genomic instability. The suppression was mediated by activation of the translesion synthesis pathway, which allows bypass of DNA lesions.   In Chapter 3 I demonstrate that Rtt107 physically interacted with the SMC5/6 complex, and that Rtt107 was required for recruitment of SMC5/6 to double-stranded breaks but not protein-bound nicks. Rtt107 became phosphorylated even in the absence of DNA damaging agents when subunits of SMC5/6 were mutated, suggesting a complex relationship beyond the DDR. Moreover, Rtt107 and SMC5/6 contributed independently to the maintenance of genome stability.   In Chapter 4 I describe the genetic interaction profiles of RTT107, SLX4, and the novel helicase HRQ1 under three different DNA damaging agents. These conditional genetic interactions revealed functions specific to the DDR, and exhibited some interesting drug-specific patterns.  RTT107 had the most genetic interactions under campothecin, indicating an important role for Rtt107 in response to this type of lesion. The genetic interaction profiles for HRQ1 and SLX4 and the associated gene ontology terms suggested that these two genes had similar functions in the DDR.   23  In Chapter 5 I show that Rtt107 was recruited to sites of DNA damage via binding of its fifth and sixth BRCT domains to phosphorylated Ser 129 of H2A. Rtt107 in turn was required for Slx4 recruitment, but Dpb11 was only partially dependent on Rtt107 for recruitment.  In Chapter 6 I discuss my results from Chapters 2-5 and highlight several themes that emerged, which illustrate some common principles in the DDR. I also discuss the areas in which there are outstanding questions that require further exploration.      24  Chapter 2: Loss of H3 K79 Trimethylation Leads to Suppression of Rtt107-Dependent DNA Damage Sensitivity Through the Translesion Synthesis Pathway1   2.1  Introduction Multiple mechanisms cooperate to maintain genome integrity, thus ensuring proper transmission of genetic information from one generation to the next.  DNA damage is detected by sensors that activate the DNA damage checkpoint, which in turn elicits various cellular responses including cell cycle arrest, DNA repair, apoptosis, and/or DNA damage-induced transcriptional programs (Harper and Elledge. 2007; Putnam et al. 2009).  In Saccharomyces cerevisiae, the kinase proteins Mec1 and Tel1, the yeast homologues of mammalian ATR (ATM and Rad3-related) and ATM (ataxia-telangiectasia mutated), are crucial for transducing signals in the S phase checkpoint response (Abraham. 2001; Shiloh. 2001).  The downstream signaling cascade leads to cell cycle arrest, replication fork stabilization, and DNA damage repair (Segurado and Tercero. 2009).  Following successful DNA repair, the checkpoint must be deactivated to allow resumption of cell cycle and restart of stalled replication forks.  While one of the main steps in this process is dephosphorylation of the effector kinase Rad53, checkpoint deactivation is further coordinated by many                                                 1 This chapter is published in the Journal of Biological Chemistry. Levesque, N., G. P. Leung, A. K. Fok, T. I. Schmidt and M. S. Kobor, 2010. Loss of H3 K79 trimethylation leads to suppression of Rtt107-dependent DNA damage sensitivity through the translesion synthesis pathway. J. Biol. Chem. 285: 35113-35122. See Preface on page iv for details of my contributions.  25  different proteins including phosphatases, proteases, and helicases (Conde and San-Segundo. 2008; Harrison and Haber. 2006; O'Neill et al. 2007; Szyjka et al. 2008).  In the event of irreparable DNA damage, tolerance mechanisms allow bypass of DNA lesions therefore enabling cells to survive (Lee and Myung. 2008).  One of these pathways is the translesion synthesis (TLS) pathway which uses special error-prone polymerases to allow replication past DNA lesions, resulting in an increased mutation frequency (Waters et al. 2009).   One of the downstream phosphorylation targets of Mec1 is Rtt107/Esc4, which is required for the reinitiating replication after repair of alkylating DNA damage (Roberts et al. 2006; Rouse. 2004).  Accordingly, yeast lacking the non-essential RTT107 gene or carrying an allele encoding for a non-phosphorylatable Rtt107 protein are hypersensitive to different DNA damaging agents (Rouse. 2004). These include the DNA alkylating agent methyl methane-sulfonate (MMS), the nucleotide reductase inhibitor hydroxyurea (HU), and the topoisomerase I poison camptothecin (CPT) (Chang et al. 2002; Roberts et al. 2006; Rouse. 2004).  Moreover, rtt107∆ mutants have a chromosome instability phenotype and an increased incidence of Rad52 foci, indicative of homologous recombination occurring due to stalled DNA replication forks (Alvaro et al. 2007; Yuen et al. 2007).  Aside from these roles in genome integrity, Rtt107 functions to repress the mobility of Ty1 transposons and to establish silent chromatin (Andrulis et al. 2004; Scholes et al. 2001).  Rtt107 contains several BRCA1 C-terminal (BRCT) homology domains, which often serve as phospho-binding modules to recruit signaling complexes and repair factors to DNA damage-induced lesions (Mohammad and Yaffe. 2009; Rouse. 2004).  Consistent with a role 26  as a scaffold for protein-protein interactions during the DNA damage response, Rtt107 interacts with a number of DNA repair and recombination proteins (Chin et al. 2006; Roberts et al. 2006; Roberts et al. 2008).  Of these, Rtt107’s interaction with the structure-specific endonuclease Slx4 is best characterized and indicative of a close functional relationship between the two proteins.  Slx4 is required for Mec1-dependent phosphorylation of four SQ/TQ motifs in the C-terminal half of Rtt107 and, like Rtt107, facilitates resumption of DNA replication after DNA damage (Roberts et al. 2006; Rouse. 2004).   In addition to the complex regulation of the DNA damage response by signaling cascades, chromatin structures in the cell also play many roles in regulating access to DNA during the repair process. One example of the emerging interface between chromatin and DNA damage response pathways is the DNA damage-induced recruitment of Rtt107 to chromatin by the H3 K56 acetyltransferase Rtt109 and the cullin Rtt101 (Roberts et al. 2008). There are many other chromatin modifications involved in the DNA damage response, such as the well-studied H2A phosphorylation and H3K79 methylation pathways (van Attikum and Gasser. 2009). H2A S129 is phosphorylated by Mec1 in response to DNA damage, triggering the assembly of many repair proteins and chromatin modifiers acting at subsequent steps (Chambers and Downs. 2007; Downs et al. 2000; Redon et al. 2003; van Attikum and Gasser. 2009). To allow resumption of cell cycle and DNA replication after successful completion of DNA repair, H2A S129 needs to be dephosphorylated by either Pph3 or Glc7, depending on the exact nature of the initial damage (Bazzi et al. 2010; Keogh et al. 2006). Dot1-mediated H3 K79 methylation, which is regulated by Bre1-mediated H2B K123 ubiquitination, is required for G1 and S phase checkpoints (Dover et al. 2002; Giannattasio et 27  al. 2005; Shilatifard. 2006; Wood et al. 2003; Wysocki et al. 2005). In part, this requirement is mediated through a functional linkage to the Rad9 adaptor protein (Grenon et al. 2007; Wysocki et al. 2005). Several lines of evidence suggest that Dot1 plays an additional role in DNA repair pathways, such as nucleotide excision repair, sister chromatid recombination and repair of ionizing radiation damage (Chaudhuri et al. 2009; Conde et al. 2009; Game et al. 2005). In contrast, Dot1 negatively regulates the error-prone TLS pathway through an unknown mechanism, thereby allowing bypass of DNA replication blocks (Conde and San-Segundo. 2008). Aside from Dot1’s function in DNA damage, it is also involved in gene silencing as well as differential H3 K79 methylation during the cell cycle (Schulze et al. 2009; Singer et al. 1998).  This study established a close connection between Rtt107 and the pathway resulting in a specific chromatin modification, H3 K79 trimethylation.  Specifically, loss of H3 K79 trimethylation suppressed the DNA damage sensitivity of rtt107Δ and slx4Δ mutants.  This suppression was not linked to restoration of Rtt107 or Slx4 phosphorylation, but instead was dependent on the presence of a functional TLS pathway. Moreover, deletion of DOT1 partially suppressed the cell cycle delay and the defect in resuming DNA replication of rtt107Δ mutants during the recovery from MMS-induced DNA damage. In contrast, deletion of DOT1 rescued neither the chromosome instability phenotype nor the increased incidence of spontaneous Rad52 foci caused by loss of Rtt107.    28  2.2 Materials and Methods  2.2.1 Yeast strains All yeast strains used in this study are listed in Table 2.1 and created using standard yeast genetic techniques (Amberg et al. 2005). Complete gene deletions and integration of a triple FLAG tag at the 3’ end of genes (Gelbart et al. 2001) were achieved using one-step gene integration of PCR-amplified modules (Longtine et al. 1998). Plasmid shuffling experiments were performed using 5-FOA as a counterselecting agent for the URA3 plasmid (pRS316, HHT2-HHF2), and shuffling in plasmids containing histone H3 K79 mutations (pRS314, hht2-HHF2) (Yang et al. 2008). Catalytically inactive Dot1 mutants were expressed from pRS315 plasmids (van Leeuwen et al. 2002), and a non-phosphorylatable mutant of Rtt107 (four SQ motifs substituted by AQ) was expressed from a plasmid (pRS315, rtt107-4AQ) which was a generous gift from Grant Brown and Tania Roberts (University of Toronto). BrdU-Incorporating (BrdU-Inc) wild-type and mutant strains containing constitutively expressed Herpes simplex virus thymidine kinase (HSV-TK) and human equilibrative nucleoside transporter (hENT1) were generated by genetic crosses with a previously published parental strain (Viggiani and Aparicio. 2006).  Table 2.1 Yeast strains used in this study.   Strain Relevant Genotype Source* MKY5  MATα ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1   MKY6  MATa can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY7 MATa ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1  MKY399 MATα can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY10  MATα his4  MKY11 MATa his4   29  Strain Relevant Genotype Source* MKY921 MKY5, dot1:: HIS3    MKY922 MKY5,  rtt107::KANMX6 lys2Δ  MKY923 MKY7, rtt107::KANMX6 dot1:: HIS   MKY924 MKY7, slx4:: KANMX6   MKY925 MKY5,  slx4::KANMX6 dot1:: HIS3   MKY927 MKY5,  bre1::HYGMX  lys2Δ  MKY928 MKY5, rtt107::KANMX6   MKY929 MKY5,  rtt107::KANMX6 bre1::HYGMX lys2Δ  MKY930 MKY5,  slx4::KANMX6 lys2Δ  MKY931 MKY5,  slx4::KANMX6 bre1::HYGMX lys2Δ  MKY932 MKY5, [pRS315]  MKY933 MKY5, [pRS315, DOT1]  MKY934 MKY5, [pRS315, dot1G401R]  MKY935 MKY5, [pRS315, dot1G401A]  MKY936 MKY7, rtt107::KANMX6 dot1::HIS3 [pRS315]  MKY937 MKY7, rtt107::KANMX6 dot1::HIS3 [pRS315, DOT1]  MKY938 MKY7, rtt107::KANMX6 dot1::HIS3  [pRS315, dot1G401R]  MKY939 MKY7, rtt107::KANMX6 dot1::HIS3 [pRS315, dot1G401A]  MKY940 MATa/α ADE2/ade2-1 can1-100/can1-100 his3-11/his3-11 leu2-3,112/leu2-3,112 LYS2/lys2Δ   MKY941 MKY940, rtt107::KANMX6/rtt107::NATMX6   MKY942 MKY940, dot1::HIS3/dot1::HIS3   MKY943 MKY940, rtt107::KANMX6/rtt107::KANMX6 dot1::HIS3/ dot1::HIS3   MKY944 MKY7, Rad 52-GFP::NATMX6    MKY945 MKY7, rtt107::KANMX6 Rad 52-GFP::NATMX6   MKY946 MKY7, dot1::HIS3 Rad 52-GFP::NATMX6   MKY947 MKY7, rtt107::KANMX6 dot1::HIS3 Rad 52-GFP   MKY949 MKY7, dot1::HIS3   MKY950 MKY7, rtt107::KANMX6   MKY951 MKY399, rtt107::KANMX6   MKY952 MKY6, dot1::HIS3   MKY953 MKY399, rev1::HYGMX   MKY954 MKY6, rtt107::KANMX6 dot1::HIS3   MKY955 MKY399, dot1::HIS3 rev1::HYGMX   MKY956 MKY399, rtt107::KANMX6 rev1::HYGMX   MKY957 MKY6, rtt107::KANMX6 dot1::HIS3 rev1::HYGMX   MKY958 MKY5, rev3::HYGMX   MKY959 MKY5, rtt107::KANMX6 dot1::HIS3   MKY960 MKY5, dot1::HIS3 rev3::HYGMX   MKY961 MKY5, rtt107::KANMX6 rev3::HYGMX   MKY962 MKY5, rtt107::KANMX6 dot1::HIS3 rev3::HYGMX   MKY963 MKY399, hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS316, HHT2-HHF2] (Yang et al. 2008) 30  Strain Relevant Genotype Source* MKY964 MKY6, rtt107::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS316, HHT2-HHF2]  MKY965 MKY399, hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79A-HHF2]  MKY966 MKY6, rtt107::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79A-HHF2]  MKY992 MKY5, SLX4-3XFLAG::NATMX6   MKY993 MKY5, rtt107::KANMX6 SLX4-3XFLAG::NATMX6   MKY994 MKY5, dot1::HIS3 SLX4-3XFLAG::NATMX6   MKY995 MKY5, rtt107::KANMX6 dot1::HIS3 SLX4-3XFLAG::NATMX6   MKY996 MKY5, RTT107-3XFLAG::NATMX6 lys2Δ  MKY997 MKY5, slx4::KANMX6 RTT107-3XFLAG::NATMX6   MKY998 MKY7, dot1::HIS3 RTT107-3XFLAG::NATMX6 lys2Δ  MKY999 MKY7, slx4::KANMX6 dot1::HIS3 RTT107-3XFLAG::NATMX6 lys2Δ  MKY1000 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79R-HHF2]  MKY1001 MKY399, rtt107::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79R-HHF2]  MKY1040 MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0  MKY1041 MKY1040, rtt107:: HIS3   MKY1042 MKY1040, dot1:: HYGMX   MKY1043 MKY1040, rtt107::HIS3 dot1::HYGMX    MKY1055 MKY5, rtt107::KANMX6 [pRS315, RTT107]  MKY1056 MKY5, rtt107::KANMX6 [pRS315, rtt107-4AQ]  MKY1057 MKY5, rtt107::KANMX6 [ pRS315]  MKY1058 MKY5, rtt107::KANMX6 dot1::HIS3 [pRS315, RTT107]  MKY1059 MKY5, rtt107::KANMX6 dot1::HIS3 [pRS315, rtt107-4AQ]  MKY1060 MKY5, rtt107::KANMX6 dot1::HIS3 [pRS315]  MKY1082 MKY5, slx4::KANMX6 dot1::HIS3 [pRS315]  MKY1083 MKY5, slx4::KANMX6 dot1::HIS3 [pRS315, DOT1]  MKY1084 MKY5, slx4::KANMX6 dot1::HIS3 [pRS315, dot1 G401A]  MKY1085 MKY5, slx4::KANMX6 dot1::HIS3 [pRS315, dot1 G401R]  MKY1086 MKY6, slx4::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, HHT2-HHF2]  MKY1087 MKY6, slx4::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79A-HHF2]  MKY1088 MKY6, slx4::KANMX6 hht1-hhf1::LEU2 hht2-hhf2::HIS3 [pRS314, hht2K79R-HHF2]  MKY1094 MKY6, ars603.5::URA3 ars608::HIS3 ars609::TRP1 LEU2::BrdU-Inc (Viggiani and Aparicio. 2006) MKY1095 MKY6, ars603.5::URA3 ars608::HIS3 ars609::TRP1 LEU2::BrdU-Inc rtt107::KANMX6  MKY1096 MKY6, ars603.5::URA3 ars608::HIS3 ars609::TRP1 LEU2::BrdU-Inc dot1::HYGMX  MKY1097 MKY6, ars603.5::URA3 ars608::HIS3 ars609::TRP1 LEU2::BrdU-Inc rtt107::KANMX6 dot1::HYGMX   * unless otherwise indicated, all strains were constructed for this work (details available upon request) or were from the laboratory collections. All strains are in the W303 background, except strains MKY1040-MKY1043, which are in the S288C background. 31  2.2.2 Sensitivity measurements Overnight cultures grown in YPD or SC-Leu at 30°C were diluted to 0.3 O.D600.  The cells were tenfold serially diluted and spotted onto solid YPD plates or SC-Leu plates with MMS (Sigma) at various concentrations.  The plates were then incubated at 30°C for 2-3 days.  2.2.3 Protein extracts and protein blot analysis Overnight cultures were diluted to 0.3 O.D600 and grown in YPD to 0.8 O.D600, and then treated with 0.03% MMS for 1 h.  The FLAG-tagged Slx4 protein was extracted by an alkaline method using 0.2 M NaOH (Kushnirov. 2000). Slx4-FLAG proteins were visualized using anti-FLAG M2 antibodies (Sigma) and SuperSignal enhanced chemiluminescence (Pierce Chemical). The procedure for analytical-scale immunoprecipitation of the FLAG-tagged Rtt107 protein was adapted from a previous report (Kobor et al. 2004). Briefly, yeast cells were harvested, and lysed in TAP-IP Buffer (50mMTris [pH 7.8], 150 mM NaCl, 1.5 mM MgAc, 0.15% Nonidet P-40, 1 mM DTT, 10 mM NaPPi, 5 mM EGTA, 5 mM EDTA, 0.1mM Na3VO4, 5 mM NaF, CompleteTM Protease inhibitor mixture) using acid-washed glass beads and mechanically disrupting using a bead beater (BioSpec Products). Rtt107-FLAG fusion proteins were captured using anti FLAG M2 agarose beads (Sigma), and subsequently washed in TAP-IP buffer. Captured material was analyzed by protein blotting with anti-FLAG M2 (Sigma) and visualized using the Odyssey Infrared Imaging System (Licor).  32  2.2.4 Flow cytometric analysis and BrdU incorporation experiments Cells were prepared under the same conditions for flow cytometric analysis and BrdU incorporation experiment. For the latter, we used wild-type and mutant strains containing the BrdU-Inc cassette (Viggiani and Aparicio. 2006) to allow for BrdU uptake in yeast. Briefly, cells were arrested in G1 by addition of 2 µg/ml of α-factor for 2 h at 30°C in YPD, then washed with sterile 1X PBS and resuspended in YPD containing 0.03% MMS for 1 h.  MMS was removed by treating with 2% sodium thiosulfate and washing with sterile 1X PBS.  The cells were resuspended in YPD and incubated at 30°C during the recovery phase. For the BrdU incorporation experiment, 400 µg/ml of BrdU was added to the cultures during the recovery phase. Aliquots were removed at the indicated times and processed further for flow cytometric analysis or measurement of BrdU incorporation.   For flow cytometric analysis, cells were collected in 70% ethanol with 0.2M Tris-HCl pH 7.5 and prepared as previously described (Haase and Lew. 1997). Samples were analyzed using the BD FACS Calibur instrument and the Flow Jo software (Tree Star Inc. OR). For the BrdU incorporation experiments, cells were collected in buffer containing 100 mM EDTA (pH 8.0), 10 mM Tris-HCl (pH 8.0), and 0.1% NaH3. Total genomic DNA was extracted by bead-beating and use of the DNeasy kit (QIAGEN) and sonicated using Bioruptor (Diagenode).  The DNA concentration was adjusted to 20 ng/µl, then heat-denatured and snap-cooled.  1 µg of DNA was spotted onto a nitrocellulose membrane (Bio-Rad) pre-soaked with 2X SSC using the Convertible Filtration Manifold System (GIBCO BRL) and subjected to ultraviolet crosslinking in a Stratalinker (Stratagene).  Subsequently, the 33  membrane was blocked with 5% milk in TTBS, probed with an anti-BrdU antibody (GE Healthcare), and visualized using the Odyssey Infrared Imaging System (Licor).  2.2.5 Quantitative bimater assay The procedure for the bimater assay was modified from a previous method to allow quantification (Yuen et al. 2007). Briefly, 12 independent colonies from each homozygous diploid strain were grown in YPD overnight at 30°C and diluted to 2.0 O.D600. Cells were plated on to solid YPD at appropriate dilutions to determine the total number of cells.  Equal volumes of MATa mating tester cultures (10.0 O.D600) and the homozygous diploid strain cultures (2.0 O.D600) were plated onto solid media containing no amino acids and incubated at 30°C for 3-4 days. Mating rates and 95% confidence intervals were calculated with the Ma-Sandri-Sarkar Maximum Likelihood Estimator (MSS-MLE) method using the online Fluctuation AnaLysis CalculatOR (FALCOR) (Hall et al. 2009).  2.2.6 Microscopy Nuclear morphology was determined by treating cells as for flow cytometric analysis, except α-factor incubation was omitted and SC-complete medium was used to minimize auto-fluorescence.  Aliquots were removed at the indicated times and treated with 4% formaldehyde solution (Sigma) for 10 min. Cells were immobilized on a glass slide with a solution of 1 mg/ml polylysine (Sigma) and then stained with 4, 6-diamidino-2-phenylindole (DAPI; Sigma). Cells with medium to large buds were counted as being in G2/M phase.  Ambiguous cases where cells with separate nuclei were insufficiently spread were considered to be in G1 phase.  34   To visualize Rad52-GFP foci, cells were grown at 30°C in SC-complete medium to logarithmic phase, and then immobilized on a glass slide with a solution of 1.0% agarose in ddH2O. Multiple images were obtained at 0.3 µm intervals along the z axis and Rad52-GFP foci were counted by inspection of all focal planes. At least 300 cells were counted for each time point. All imaging was done with the Zeiss Axioplan 2 fluorescence microscope using the Metamorph software.  Statistical significance was assessed using Student’s t test.  2.2.7 Measurement of mutation rates Forward mutation rates were measured by mutations at the CAN1 locus, which when mutated renders cells sensitive to canavanine (Grenson et al. 1966). Cells from 12 independent colonies for each strain were grown in YPD to logarithmic phase, 0.005% MMS was added to half of each culture, and cells were further incubated at 30°C for 20 hours. Cells plated on SC-Arg were diluted 1 in 200000 while cells plated on SC-Arg containing 50 µg/ml canavanine (Sigma) were diluted by a factor of 2. Plates were incubated at 30°C for 2 days and colonies were counted. The mutation rates and 95% confidence intervals were calculated with the Ma-Sandri-Sarkar Maximum Likelihood Estimator (MSS-MLE) method using the online Fluctuation AnaLysis CalculatOR (FALCOR) (Hall et al. 2009).  2.2.8 Pulsed-field gel electrophoresis  Briefly, cells were arrested in G1 by addition of 2 µg/ml of α-factor for 2 h at 30°C in YPD, then washed with sterile 1X PBS and resuspended in YPD containing 0.03% MMS for 1 h.  MMS was removed by treating with 2% sodium thiosulfate and washing with sterile 1X PBS.  35  The cells were resuspended in YPD and incubated at 30°C during the recovery phase. Aliquots were removed at the indicated times and processed further.   Cells were treated using the CHEF Yeast Genomic DNA Plug Kit (Bio-Rad, CA). As described by the manufacturer, cells were washed with cold 50 mM EDTA and resuspended in Cell Suspension Buffer containing 0.75% agarose and Lyticase, and incubated for 2 h at 37oC. The agarose plugs were incubated with Proteinase K overnight at 50oC, then washed and loaded onto a 1.0% agarose gel in 0.5X Tris-borate EDTA. Pulsed-field gel electrophoresis was carried out as described by the manufacturer (GE Healthcare, UK). Briefly, the gel was subjected to electrophoresis at 100 V with a switch time of 60–120 s for 24 h with constant recirculation of the buffer at 8.5°C. Following electrophoresis, the gels were stained with ethidium bromide (0.5 µg/ml) and photographed.  2.3 Results  2.3.1 Elimination of H3 K79 methylation suppressed the sensitivity of rtt107∆ and slx4Δ mutants to the DNA-damaging agent MMS Rtt107 and its interaction partner Slx4 are required for yeast cells to survive exposure to DNA damage conditions, such as those caused by the alkylating agent MMS (Chang et al. 2002; Roberts et al. 2006). Given the importance of the natural chromatin template during the DNA damage response, and the existing link between Rtt107 and the histone acetyltransferase Rtt109 (Bassal and El-Osta. 2005; Giannattasio et al. 2005; Roberts et al. 2008), we hypothesized that chromatin modifications might affect the requirement for Rtt107 36  and Slx4 during DNA damage repair. For this purpose, we created strains that, in addition to deletion of either RTT107 or SLX4, lacked genes encoding several chromatin modifiers with roles in the DNA damage response to test whether their absence enhanced or suppressed the sensitivity of rtt107∆ or slx4∆ mutants to MMS. While the majority of double mutants grew equally well as rtt107∆ or slx4∆ mutants, we found that deletion of DOT1, a non-essential gene encoding a histone methyltransferase catalyzing mono-, di- and trimethylation of histone H3 K79, almost completely rescued the MMS sensitivity of rtt107∆ and slx4∆ mutants (Figure 2.1A).   37   Figure 2.1 Abrogation of H3 K79 trimethylation suppressed the MMS sensitivity of strains lacking Rtt107 or Slx4.  Tenfold serial dilutions of the indicated strains were plated onto media containing 0.0075% or 0.01% MMS. (A) Loss of Dot1 suppressed MMS sensitivity of rtt107∆ and slx4∆ mutants. (B) Loss of Dot1’s catalytic activity, (C) H3 K79A, K79R, or (D) loss of Bre1 suppressed MMS sensitivity of rtt107∆ mutants. (E) Bre1 affected mainly H3 K79 trimethylation, not di- or monomethylation.  Whole cell extracts of indicated strains were analyzed by protein blotting with anti-H3 K79 tri-, di- or monomethyl antibodies. Antibodies against H3 were used as a loading control.  38  To determine whether this effect was dependent on the catalytic activity of Dot1, alleles encoding catalytically inactive Dot1 proteins were compared to the complete loss of Dot1 and to the presence of wild-type Dot1. The strains carrying dot1G401R and dot1G401A alleles, encoding for catalytically inactive forms of Dot1, suppressed the MMS sensitivity phenotype similar to the complete deletion (Figure 2.1B). As expected, re-introducing wild-type DOT1 in rtt107∆dot1∆ double mutants restored MMS sensitivity to levels similar to that of rtt107∆ single mutants. These data suggested that eliminating the catalytic activity of Dot1 enabled cells lacking Rtt107 to survive otherwise detrimental conditions during exposure to MMS. The same results were obtained for slx4∆ mutants, except that slx4∆ mutants were less sensitive to MMS than rtt107∆ mutants (Figure 2.2A).  39   Figure 2.2 Abrogation of H3 K79 trimethylation suppressed the MMS sensitivity of strains lacking Slx4. Tenfold serial dilutions of the indicated strains were plated onto media containing 0.0075%, 0.01%, or 0.0125% MMS. (A) Loss of Dot1’s catalytic activity, (B) H3 K79A, K79R, or (C) loss of Bre1 suppressed MMS sensitivity of slx4∆ mutants.  The only known target of Dot1 methyltransferase activity to date is the K79 residue located in the core of histone H3, but formally it is possible that Dot1, similar to other chromatin modifiers, has other enzymatic targets not yet identified. To examine whether the suppression of the MMS sensitivity of rtt107∆ mutants by loss of Dot1 was due to lack of H3 K79 methylation, strains with mutated forms of H3 K79 which cannot be methylated were tested for their ability to survive chronic MMS exposure in the absence of Rtt107. Changing lysine 79 to either alanine or arginine rescued the DNA damage sensitivity of the rtt107Δ mutants, 40  analogous to the DOT1 deletion (Figure 2.1C). Therefore, we concluded that the reversal of the MMS sensitivity of rtt107∆ mutants was due to the loss of Dot1-mediated H3 K79 methylation. Similarly, H3 K79A and H3 K79R mutants also suppressed the MMS sensitivity of the slx4∆ strain (Figure 2.2B).   Methylation of H3 K79 by Dot1 is regulated through crosstalk with another histone modification, mono-ubiquitination of H2B K123, which is catalyzed by the Bre1/Rad6 enzyme complex (Dover et al. 2002; Foster and Downs. 2009; Ng et al. 2002; Shahbazian et al. 2005; Shilatifard. 2006; Wood et al. 2003). Thus we wanted to test whether upstream regulators of H3 K79 methylation would have a similar effect on the MMS sensitivity of rtt107∆ and slx4∆ mutants.  Indeed, deletion of BRE1 also rescued the MMS sensitivity of the strains lacking Rtt107 or Slx4 (Figure 2.1D and Figure 2.2C).   To learn more about the biochemical nature underlying the observed effects, we assessed the total levels of mono-, di- or trimethylated H3 K79 in whole cell extracts. Interestingly, whereas Dot1 broadly catalyzes mono-, di- and trimethylation of H3 K79 (Lacoste et al. 2002; Ng et al. 2002; van Leeuwen et al. 2002), Bre1 primarily affected K79 trimethylation (Figure 2.1E). These results suggested that specifically a lack of H3 K79 trimethylation caused the suppression of the MMS sensitivity of rtt107∆ and slx4∆ mutants.  41  2.3.2 Deletion of DOT1 suppressed DNA damage sensitivity in the absence of MMS-induced phosphorylation of Rtt107 or Slx4 In response to DNA damage induced by various agents, Rtt107 and Slx4 are phosphorylated on several S/T residues by the checkpoint kinase Mec1 (Roberts et al. 2006; Rouse. 2004). Phosphorylation of Rtt107 is essential for its function in the DNA damage response and depends on Slx4 (Roberts et al. 2006; Rouse. 2004). It was in principle possible that, in slx4∆dot1∆ double mutants, an alternate pathway directed Rtt107 phosphorylation in the absence of Slx4, thereby enabling cells to survive the otherwise detrimental MMS-induced DNA damage. To test this possibility, Rtt107 phosphorylation was measured in strains lacking Slx4, Dot1, or both simultaneously. As expected, exposure to MMS induced phosphorylation of Rtt107 in wild-type strains but not in strains lacking Slx4 (Figure 2.3A) (Roberts et al. 2006). Although deletion of DOT1 suppressed the MMS sensitivity of slx4Δ mutants, it did not overcome the requirement of Slx4 for Rtt107 phosphorylation (Figure 2.3A). Loss of Dot1 had no effect on Rtt107 phosphorylation in response to MMS when Slx4 was present (Figure 2.3A). Therefore, the suppression by dot1Δ did not involve a restoration of Rtt107 phosphorylation in the absence of Slx4, arguing against an alternative pathway for Rtt107 phosphorylation. Consistent with the physical interaction and close functional relationship between Rtt107 and Slx4, we found that MMS-induced phosphorylation of Slx4 was dependent on Rtt107, but not on Dot1 (Figure 2.3B). Analogous to the results obtained for Rtt107, eliminating DOT1 did not restore Slx4 phosphorylation in the absence of Rtt107 (Figure 2.3B).   42   Figure 2.3 Suppression of rtt107∆ MMS sensitivity by deletion of DOT1 was not dependent on the phosphorylation of Slx4 and vice versa.  (A) Cells expressing Rtt107-Flag were untreated or treated with 0.03% MMS for 1 h. Analytical-scale immunoprecipitations of Rtt107-Flag were performed and analyzed by protein blotting with anti-Flag antibodies. The reduced mobility of Rtt107-Flag indicated phosphorylation of the protein. Background bands (*) were used as a loading control. (B) Cells expressing Slx4-Flag were treated as described in (A). Whole cell extracts were analyzed by protein blotting with anti-Flag antibodies, with reduced mobility of Slx4-Flag being indicative of phosphorylation.  Antibodies against tubulin were used as a loading control. (C) Deletion of DOT1 suppressed the MMS sensitivity of the mutants expressing the non-phosphorylatable Rtt107-4AQ. Tenfold serial dilutions were plated onto SC-Leu containing 0.005% MMS.  To test whether the suppression by deletion of DOT1 was linked to the phosphorylation of Rtt107 at specific SQ sites, we utilized mutants expressing the non-phosphorylatable form of Rtt107. Using a plasmid expressing rtt107-4AQ in cells lacking Dot1, we tested the MMS sensitivity of the double mutants. Consistent with the importance of Rtt107 phosphorylation, deletion of DOT1 suppressed the MMS sensitivity of the rtt107 4AQ mutants (Figure 2.3C). Taken together, these results indicated that MMS sensitivity of mutants lacking Rtt107 phosphorylation was suppressed by deletion of DOT1. 43   2.3.3 The requirement of Rtt107 for resumption of cell cycle after S phase damage was partially suppressed by lack of Dot1 To further understand the molecular mechanism leading to the suppression of rtt107∆ MMS sensitivity, we tested whether loss of Dot1 could compensate for the requirement of Rtt107 during the restart of DNA replication (Roberts et al. 2006; Rouse. 2004). Cells arrested in G1 were released into S phase in the presence of MMS for 1 h, and restart of DNA replication was directly measured by 5-bromo-2’-deoxyuridine (BrdU) incorporation into nascent genomic DNA (Figure 2.4A). Since BrdU was added after MMS treatment, it serves as a quantitative indicator of newly replicated DNA during the recovery process. As expected, BrdU levels increased in wild-type cells during the course of the experiment, indicating successful resumption of DNA replication (Figure 2.4B). In contrast, BrdU levels in rtt107∆ mutants were consistently lower than the wild-type at each time point. Whereas the levels of BrdU incorporation in dot1∆ mutants increased similar to the wild-type cells, rtt107∆dot1∆ mutants incorporated BrdU at intermediate levels between wild-type and rtt107∆ mutants (Figure 2.4C). This result suggested that loss of Dot1 could partly suppress the defect of rtt107∆ mutants in resuming DNA replication.  44   Figure 2.4 Requirement of Rtt107 for resumption of DNA replication and cell cycle after DNA damage was partially suppressed by deletion of DOT1.  (A) Diagram of experimental strategy used for BrdU incorporation experiment. (B) BrdU incorporation into nascent DNA indicated rtt107∆dot1∆ mutants more efficiently resumed DNA replication after DNA damage than rtt107∆ mutants. (C) Quantification of newly replicated DNA as measured by BrdU signals. All values are relative to wild-type at 30 min. Error bars represent standard deviations of values from 3 independent experiments. (D) Diagram of experimental strategy used for FACS analysis. (E) FACS analysis showed rtt107∆dot1∆ mutants recovered from DNA damage earlier than rtt107∆ mutants. ASY, asynchronous cells.  45  Next, we used FACS analysis to test whether loss of Dot1 would have a similar effect on the resumption of cell cycle progression after DNA damage (Figure 2.4D). At the end of the MMS treatment (0 h), wild-type and rtt107∆ mutants were initially arrested in S phase due to activation of the DNA damage checkpoint (Roberts et al. 2006; Rouse. 2004), whereas dot1∆ mutants had proceeded through S phase as judged by the shift of the signal to 2N, consistent with a requirement for Dot1 in the DNA damage checkpoint as described previously (Giannattasio et al. 2005; Wysocki et al. 2005) (Fig. 3E). 2 h after removal of MMS, rtt107∆ mutants were still in S phase whereas all other strains had progressed to G2/M. Further differences between the mutants were observed as strains continued to recover from DNA damage. For example, at 5 h a substantial fraction of cells in the wild-type and dot1∆ mutant had undergone cell division as judged by the appearance of a G1 peak and S phase fraction while rtt107∆ mutants had predominantly a 2N peak, suggesting that they were still residing in G2/M. These differences persisted until 8 h after recovery when the G1 peak and S phase fraction first appeared in the rtt107∆ mutants. The rtt107∆dot1∆ double mutants had an intermediate phenotype, as the G1 peak and S phase fraction became visible at 6 h, which was earlier than rtt107∆ mutants but later than wild-type or dot1∆ mutants (Figure 2.4E). Consistent with this, the delayed appearance of intact chromosomes during recovery in the rtt107∆ mutants was partially rescued by concurrent loss of Dot1 as visualized by pulsed-field gel electrophoresis (PFGE) (Figure 2.5).   46   Figure 2.5 Requirement of Rtt107 for resumption of replication after DNA damage was partially suppressed by deletion of DOT1.  PFGE analysis indicated rtt107∆dot1∆ mutants had more efficient DNA replication after DNA damage than rtt107∆ mutants. * : unresolved chromosomes.    The defect of rtt107∆ mutants in completing the G2/M phase of the cell cycle during recovery from transient DNA damage can also be observed by examining nuclear morphology (Roberts et al. 2006). In wild-type and dot1∆ mutants, the percentage of cells in G2/M increased after exposure to MMS, reached a peak at 3 h of recovery and started to decrease as the cells completed mitosis (Figure 2.6A,B). As expected, the percentage of rtt107∆ mutants in G2/M also reached a peak at 3 h, but the increased level lasted up to 5 h after MMS treatment.  As judged by the percentage of G2/M cells, the kinetics of recovery from DNA damage in rtt107∆dot1∆ double mutants was slower than wild-type and dot1∆ mutants, but faster than rtt107∆ mutants, although this did not reach statistical significance (Figure 2.6A). Another known phenotype of rtt107∆ mutants is the delay of nuclear division, as judged by the higher percentage of large budded cells with elongated nuclei spanning the bud neck. Consistent with previous reports, nuclear division was delayed in rtt107∆ mutants when compared to wild-type at 5 h after exposure to MMS (Figure 2.6C). In contrast, dot1∆ 47  mutants did not show any delay of nuclear division. A similar phenotype was also observed in asynchronous cultures not exposed to MMS. In both conditions, deletion of DOT1 did not rescue the defect caused by loss of Rtt107.   Figure 2.6 Nuclear division delay of rtt107∆ mutants was not suppressed by deletion of DOT1 in the absence and presence of MMS.  Cells were treated with 0.03% MMS for 1h, then washed and resuspended in complete media for DNA damage recovery, and stained with DAPI at the indicated timepoints. (A) The increased percentage of cells in G2/M of rtt107∆ mutants after DNA damage was not suppressed by deletion of DOT1. The percentage of cells with medium to large buds (% G2/M) was calculated by dividing the number of cells with medium to large buds by the total number of cells. (B) Representative differential interference contrast images are shown on the left and the corresponding DAPI images on right. (C) The increased percentage of cells exhibiting nuclear division delay in rtt107∆ mutants was not suppressed by loss of Dot1. The percentage of cells with delayed nuclear division was calculated by dividing the number of large-budded cells with unsegregated nuclei by the total number of cells. In both (A) and (C), at least 200 cells were counted in 3 independent experiments. Error bars represent standard deviations of the values.  * : p<0.05; ** : p<0.005 when compared to the wild-type strain in the same timepoint.  48  2.3.4 The TLS pathway was required for the suppression of the MMS sensitivity of rtt107∆ mutants by deletion of DOT1 Next we sought to determine the pathway by which deletion of DOT1 suppressed the requirement for RTT107 during MMS exposure. In addition to the suppression of the DNA damage sensitivity of rtt107∆ and slx4∆ mutants reported here, lack of Dot1 suppresses the MMS sensitivity of strains lacking a variety of repair proteins, and this effect is dependent on the TLS polymerases Polζ and Rev1 (Conde and San-Segundo. 2008). To address whether the suppression of rtt107Δ DNA damage sensitivity by loss of Dot1 was similarly dependent on the TLS pathway, we constructed triple mutants lacking two main components of the TLS pathway: Rev3 (catalytic subunit of Polζ) or Rev1 (dC-transferase) (Kunz et al. 2000). Lack of Dot1 did not suppress the MMS sensitivity of rtt107Δ mutants in the absence of Rev3 (Figure 2.7A). Similarly, Rev1 was necessary for the reversal of rtt107∆ sensitivity by deletion of DOT1, suggesting that the dot1Δ suppression was dependent on the TLS pathway in general and not specifically on Rev3 (Figure 2.7B). Very low concentrations of MMS were used in this assay due to the extreme MMS sensitivity of the triple mutants. It is interesting to note that both dot1Δrev3Δ and rtt107Δrev3Δ double mutants were very sensitive to MMS, and a similar phenotype was also observed for dot1Δrev1Δ and rtt107Δrev1Δ mutants. This indicated that a functional TLS pathway became more important for DNA damage resistance in the absence of Dot1 or Rtt107.   49   Figure 2.7 Suppression of the rtt107Δ MMS sensitivity by deletion of DOT1 was dependent on the TLS pathway.  Deletion of REV3 or REV1 in rtt107∆dot1∆ mutants resulted in loss of the suppression. (A) rev3∆ or (B) rev1∆ mutants in combination with the indicated deletions of RTT107 and/or DOT1 were plated in tenfold serial dilutions onto YPD containing 0.0025% or 0.005% MMS. (C) Loss of Dot1 resulted in increased mutation rates in presence of MMS. Mutation rates of indicated strains with and without 0.005% MMS were determined using 12 independent colonies. Mutation rates and 95% confidence intervals were calculated using the Ma-Sandri-Sarkar Maximum Likelihood Estimator (MSS-MLE) Method.  The TLS pathway is error-prone and its activation would therefore be expected to cause an increased mutation rate. Using the CAN1 forward mutagenesis assay, we observed a 4 to 5-50  fold increased mutation rate in the presence of 0.005% MMS in both dot1Δ and dot1Δrtt107Δ mutants (Figure 2.7C). Together, these results suggested that deletion of DOT1 led to activation of the TLS pathway, thereby allowing the survival of rtt107∆ mutants.  2.3.5 Rtt107 had functions in maintaining genomic integrity that were independent of Dot1 activity Rtt107’s role in maintenance of genome stability is not restricted to its specific function of restarting cell cycle during S phase after DNA damage.  During normal cell cycle progression, cells lacking Rtt107 have increased number of Rad52 and Ddc2 foci, indicative of spontaneous DNA damage and /or replication fork stalling (Alvaro et al. 2007; Roberts et al. 2006). Consistent with this, we observed that the percentage of cells with Rad52-GFP foci in S/G2/M phase was approximately 7-fold higher in rtt107∆ mutants than in wild-type (Figure 2.8A, B). Deletion of DOT1 in the rtt107∆ background did not significantly alter the number of cells containing Rad52-GFP foci. However, consistent with published data, we observed a 2-fold increase in the number of cells with Rad52-GFP foci in dot1∆ mutants, suggesting that RTT107 was epistatic to DOT1 in suppressing spontaneous DNA damage and/or replication fork stalling (Conde and San-Segundo. 2008).  51   Figure 2.8 Genomic instability of the rtt107∆ mutants was not suppressed by deletion of DOT1.  (A) The increased number of Rad52-GFP foci in rtt107∆ mutants was not suppressed in absence of Dot1. The percentage of cells in G1 or S/G2/M phase containing Rad52-GFP foci was calculated by dividing the number of cells in G1 or S/G2/M phase containing Rad52-GFP foci by the total number of cells in G1 or S/G2/M, respectively. At least 200 cells were counted in 3 independent experiments. Error bars represent standard deviations of the values. * : p<0.05; ** : p<0.005 when compared to the wild-type strain for the same cell cycle phase. (B) Representative differential interference contrast images are shown on the left and the corresponding GFP images on the right. (C) Deletion of DOT1 did not suppress the increased loss of heterozygosity in rtt107∆ mutants.  Mating rates of homozygous diploids of indicated strains were determined by using 12 independent colonies. Mating rates and 95% confidence intervals were calculated using the Ma-Sandri-Sarkar Maximum Likelihood Estimator (MSS-MLE) Method.  Further indicative of a broader role of Rtt107 in genome stability is the chromosome instability (CIN) phenotype of rtt107∆ mutants (Yuen et al. 2007). Compared to wild-type cells, rtt107∆ homozygous diploid mutants have higher loss of heterozygosity (LOH) at the MATa and MATα loci, which is due to either enhanced mitotic recombination between homologous chromosomes, chromosome loss, rearrangement or gene conversion (Yuen et al. 2007). Using a quantitative version of the original Bimater Screen (BiM) used to define 52  RTT107 as a CIN gene (Gerring et al. 1990), we tested whether this phenotype was suppressed by loss of Dot1. Consistent with increased LOH, strains lacking Rtt107 had a 4-fold increase in mating rate when compared to wild-type or dot1∆ mutants (Figure 2.8C). Deletion of DOT1 did not rescue the CIN phenotype of rtt107∆ mutants, but rather resulted in a further increase in the mating rate to 6-fold as compared to wild-type. Hence it appeared that rather than rescuing the requirement for Rtt107 in preventing LOH, Dot1 cooperated with Rtt107 in this process.  2.4 Discussion In this study, we uncover a close functional relationship between chromatin and the cellular processes regulated by the BRCT domain-containing protein Rtt107 and its interaction partner Slx4. Loss of Dot1, likely mediated by loss of histone H3 K79 trimethylation, suppressed the DNA damage sensitivity of rtt107∆ mutants through a mechanism that was dependent on the presence of a functional TLS pathway. The DNA damage-induced phosphorylation of Rtt107 and Slx4, which was mutually dependent, was not restored in the absence of Dot1. Furthermore, deletion of DOT1 partially reversed the cell cycle progression and replication fork restart defect caused by the lack of Rtt107. In contrast, other genomic instability defects of rtt107∆ mutants were worsened or unaffected by loss of DOT1. Together, these data point to a complex functional relationship between Rtt107 and Dot1 in both the DNA damage response and preservation of genome integrity.  We propose a model to explain the inhibitory effect of H3 K79 trimethylation on growth during DNA damage conditions in yeast cells lacking Rtt107 or Slx4 (Figure 2.9). Bre1-53  mediated H2B K123 ubiquitination is required for Dot1 to catalyze H3 K79 trimethylation, which in turn prevents rtt107∆ and slx4∆ mutants from surviving DNA damage conditions. This effect likely is mediated through inhibition of the TLS pathway by H3 K79 trimethylation, either directly or indirectly through a nexus to the Dot1-mediated DNA damage checkpoint. We favour a direct mechanism that could involve binding to the H3 K79 trimethylation mark by a protein that inhibits TLS. Alternatively, H3 K79 trimethylation might directly create a chromatin conformation that in some way is refractory to TLS. An indirect enhancement of TLS might be caused by the compromised DNA damage checkpoint due to loss of Dot1, allowing rtt107∆ mutants to survive DNA damage conditions. However, currently there is no evidence linking Dot1’s roles in TLS and DNA damage checkpoint. Moreover, UV exposure of DNA damage checkpoint deficient mutants does not result in an increased mutation rate, thereby disfavouring a link between the DNA damage checkpoint and TLS (Pages et al. 2009). Lastly, it is formally possible that the suppression is an indirect effect of an altered transcriptional response caused by lack of Dot1, Bre1 or H3 K79 methylation, which could involve reduced expression of an unknown inhibitor of the TLS. In any case, given our finding that the suppression of rtt107∆ phenotypes was linked to loss of trimethylation of H3 K79, it is tempting to speculate that specific genomic regions might be more prone to mediate this effect than others. This is supported by a genome-wide analysis showing that regions containing H3 K79 trimethylation are distinct from those containing H3 K79 dimethylation (Schulze et al. 2009). Further studies are required to elucidate the precise mechanism whereby Dot1-mediated H3 K79 trimethylation inhibits the TLS pathway.    54   Figure 2.9 Model for repressive effect of chromatin modifications on DNA damage survival in rtt107∆ mutants.  During DNA damage response, Bre1-mediated H2B K123Ub and by extension, Dot1-mediated H3 K79Me3, are required for checkpoint function. In rtt107∆ mutants, the presence of H3K79Me3 is inhibitory to the yeast survival in DNA damage conditions. Loss of Dot1 increases the activity of the TLS pathway which bypasses the requirement of Rtt107 for cell survival.  Our data showed that Mec1-mediated phosphorylation of Rtt107 was dependent on Slx4, consistent with earlier reports (Roberts et al. 2006). We also showed that MMS-induced phosphorylation of Slx4 was dependent on Rtt107, suggesting a mutual requirement of these two proteins for their respective phosphorylation. Interestingly, neither Rtt107 nor Slx4 was phosphorylated in slx4∆ or rtt107∆ mutants, respectively, when DOT1 was also deleted. Furthermore, the MMS sensitivity of a strain containing a non-phosphorylatable form of Rtt107 was rescued by deletion of DOT1. Together, this biochemical and genetic evidence suggested that DNA damage dependent phosphorylation of Rtt107 is essential for resistance to MMS only when Dot1 is present to methylate H3 K79. Although it is not clear what mechanistic change is triggered by Rtt107 phosphorylation, it is likely to involve a DNA 55  damage-induced protein-protein interaction. Whatever the mechanism might be, it is clear that Rtt107 phosphorylation in the DNA damage response becomes dispensable when H3 K79 trimethylation is inhibited.    The suppression of rtt107∆ by deletion of DOT1 was restricted to situations of induced DNA damage, suggesting that the functional interaction between Rtt107 and Dot1 was context-dependent. Confirming published data from a high-throughput screen for regulators of Rad52 foci formation, we found that loss of Rtt107 caused a significant increase in the number of Rad52 foci positive cells (Alvaro et al. 2007). In contrast to the suppression of MMS sensitivity of rtt107∆ mutants, this phenotype was not rescued by loss of Dot1. This suggested that Rtt107 had a role in preventing spontaneous DNA damage – likely caused by stalled DNA replication forks – which was not negatively regulated by H3 K79 methylation. Furthermore, our work uncovered additional evidence for a complex relationship between Dot1 and Rtt107 in the maintenance of genomic integrity. Rtt107 was required for chromosome stability, as determined by a genetic assay (Yuen et al. 2007). While loss of Dot1 alone did not affect chromosome stability, it enhanced the defect caused by loss of Rtt107. This suggested that in the absence of Rtt107, Dot1 plays a minor role in maintenance of chromosome stability. Taken together, these data point to multiple activities of Rtt107, where only those induced by external DNA damaging agents were suppressed by deletion of DOT1. Presumably, both the increased number of Rad52 foci and the chromosome instability in rtt107∆ mutants were not suppressed by deletion of DOT1 because the TLS pathway is unlikely to be activated in these conditions.  56  The suppression of DNA damage sensitivity by loss of Dot1 reported here is interesting in light of other findings suggesting certain chromatin modifications act as negative regulators of DNA replication, recombination and repair. For example, the H3 K36 histone methyltransferase Set2 and the ATP-dependent chromatin remodeler Chd1 exert an inhibitory effect on DNA replication, as deletions of these genes suppress the HU sensitivity of mutations in several genes involved in DNA replication (Biswas et al. 2008). In addition, deletion of CHD1 can suppress the lethality normally caused by disruption of the gene encoding either Mec1 or Rad53 DNA damage checkpoint kinases (Biswas et al. 2008). The UV sensitivity and G2/M checkpoint defects of rad9∆ and mec1-21 mutants can be suppressed by loss of genes encoding components of the Rpd3/Sin3 histone deacetylase (Scott and Plon. 2003). Rpd3 and the aforementioned Set2 also repress meiotic recombination at the HIS4 meiotic recombination hotspot (Merker et al. 2008). However, not all chromatin modifications involved in DNA metabolism exert a negative effect, as mutations in the genes encoding members of the histone acetyltransferase complex NuA4, the ATP-dependent chromatin remodelers RSC, Swi/Snf, SWR1-C, and INO80 cause sensitivity to MMS, as does loss of the histone variant H2A.Z, or the Mec1-dependent phosphorylation targets in H2A (Karagiannis and El-Osta. 2007; Morrison and Shen. 2009). Together, all these data suggest a complex and differentiated role for the chromatin template in DNA repair, recombination and replication. Our work revealed that Dot1’s role in the DNA damage response is multifaceted and extends to regulation of the TLS pathway and maintenance of chromosome stability, although the mechanisms are still unclear. The challenge of future research will be to uncover the intricate network between chromatin modifiers and DNA damage response effectors. 57  Chapter 3: Rtt107 is Required for Recruitment of the Smc5/6 Complex to DNA Double-Strand Breaks2    3.1 Introduction Eukaryotic cells have evolved complex mechanisms to maintain genome integrity which is essential for genetic inheritance and cell viability. One of the central causes of genome instability is the failure to repair damaged DNA resulting from the constant assault of chemicals, radiation or biological processes. Many different types of DNA lesions can occur, the most severe being double-strand breaks (DSBs). In Saccharomyces cerevisiae, DSB ends are immediately sensed and bound by the MRX (Mre11–Rad50–Xrs2) complex (Ogawa et al. 1995). A signalling cascade is triggered, leading to the activation of the kinases Mec1 and Tel1, the yeast homologues of mammalian ATR (ATM and Rad 3-related) and ATM (ataxia-telangiectasia mutated), and recruitment of a whole host of DNA damage response proteins to the DSB (Harper and Elledge. 2007; Lisby and Rothstein. 2009).   One of the downstream phosphorylation targets of Mec1 is Rtt107/Esc4, which is required for reinitiating replication after repair of alkylating DNA damage (Roberts et al. 2006; Rouse. 2004). Deletion of the RTT107 gene results in hypersensitivity to DNA damaging agents such as the DNA alkylating agent methyl methane-sulfonate (MMS), the nucleotide                                                 2 This chapter is published in the Journal of Biological Chemistry. Leung, G. P., L. Lee, T. I. Schmidt, K. Shirahige and M. S. Kobor, 2011. Rtt107 is required for recruitment of the SMC5/6 complex to DNA double strand breaks. J. Biol. Chem. 286: 26250-26257. See Preface on page iv for details of my contributions. 58  reductase inhibitor hydroxyurea (HU), and the topoisomerase I poison camptothecin (CPT) (Chang et al. 2002; Roberts et al. 2006; Rouse. 2004). However, the requirement of Rtt107 for resistance to DNA damaging agents is alleviated when the chromatin regulatory pathway leading to H3 K79 trimethylation is inhibited (Levesque et al. 2010). Nevertheless, even in the absence of DNA damaging agents, rtt107Δ mutants exhibit chromosome instability and an increased incidence of Rad52 foci, indicative of a failure to properly process DNA damage or stalled DNA replication forks (Alvaro et al. 2007; Yuen et al. 2007).   Rtt107 contains several BRCT (BRCA1 C-terminal) homology domains, which often serve as phospho-binding modules to recruit signaling complexes and repair factors to DNA damage-induced lesions (Mohammad and Yaffe. 2009; Rouse. 2004). Consistent with a role as a scaffold for protein-protein interactions during the DNA damage response, Rtt107 interacts with a number of DNA repair and recombination proteins (Chin et al. 2006; Ohouo et al. 2010; Roberts et al. 2006; Roberts et al. 2008).  The best characterized Rtt107-interacting partner is the replication-specific endonuclease Slx4, which interacts with the N-terminal BRCT domains of Rtt107 (Roberts et al. 2006). Slx4 is required for Mec1-dependent phosphorylation of Rtt107 and, like Rtt107, facilitates resumption of DNA replication after DNA damage (Roberts et al. 2006). However, it has become clear over the last few years that Rtt107 also has Slx4-independent functions, and vice versa. Consistent with this, the defects in DNA damage response are generally more severe in rtt107∆ mutants than those of slx4∆ mutants, and the rtt107∆ slx4∆ double mutants are more sensitive to MMS than either of the single mutants (Roberts et al. 2006).   59  While the Rtt107-Slx4 interaction is well characterized, the Slx4-independent functions of Rtt107 and the proteins associated with these have yet to be elucidated. As an example, Rtt107 was recently identified to associate with the evolutionary conserved SMC5/6 complex (Ohouo et al. 2010). The latter is a large multi-subunit complex comprised of the Smc5-Smc6 heterodimer and six non-Smc elements (Nse1-6), which are all encoded by essential genes in budding yeast (De Piccoli et al. 2009; Fujioka et al. 2002; Zhao and Blobel. 2005). Smc5 and Smc6 are members of the structural maintenance of chromosome (SMC) proteins, a group that includes condensin (Smc1-3) and cohesin (Smc2-4) (De Piccoli et al. 2009).  The SMC5/6 complex is important for numerous chromosome maintenance activities, including DNA repair, chromosome segregation and telomere function (De Piccoli et al. 2009). Genome-wide mapping of the budding yeast SMC5/6 complex revealed that it localizes to centromeric regions, long chromosome arms, and rDNA arrays in unchallenged cells, as well as stalled DNA replication forks and DSBs (Lindroos et al. 2006). Interestingly, the recruitment of the SMC5/6 complex to these diverse chromosomal regions is differentially regulated, suggesting the involvement of multiple mechanisms and protein interaction partners (Lindroos et al. 2006).   In this study we characterized the interaction between Rtt107 and the SMC5/6 complex, and found that this association was mediated via the N-terminal BRCT domain of Rtt107 and the Nse6 subunit of the SMC5/6 complex. Here, we determine that the function underlying this interaction was a requirement of Rtt107 for the recruitment of the SMC5/6 complex to DNA DSBs, although not to certain DNA lesions such as protein-bound nicks. Moreover, this interaction was independent of Slx4 and DNA damage, consistent with this being a Slx4-60  independent role for Rtt107. Our results suggest that Rtt107 and the SMC5/6 complex cooperate together at DSBs, but they also have distinct functions in the DNA damage response.  3.2 Materials and Methods  3.2.1 Yeast strains and plasmids All yeast strains used in this study are listed in Table 3.1 and created using standard yeast genetic techniques (Ausubel. 1987).  Complete gene deletions and integration of TAP, FLAG, HA, or VSV tags at the 3’ end of genes were achieved using one-step gene integration of PCR-amplified modules (Funakoshi and Hochstrasser. 2009; Gelbart et al. 2001; Longtine et al. 1998; Rigaut et al. 1999). The mec1Δ mutants were created by complete deletion of MEC1 in sml1Δ strains. The nse5-R103G allele was created in K. Shirahige’s laboratory using standard procedures. The yeast strains containing the smc6-9 or nse3-SB1 allele were generous gifts from Luis Aragon (Imperial College London) and Philip Hieter (University of British Columbia), respectively. The GFP::NATMX6 plasmid and the pJ69-4a/α yeast strains were obtained from Elizabeth Conibear (University of British Columbia). The yeast two-hybrid plasmids were constructed by first cloning the genes into the pCR8GW-Topo Gateway Entry vector (Invitrogen) and subsequent cloning into either the pGal4 DBD-Dest or pGal4 AD-Dest gateway destination vectors obtained from Stefan Taubert (University of British Columbia).   61   Table 3.1 Yeast strains used in this study.   Strain Relevant Genotype Source* MKY6  MATa can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY7 MATa ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1  MKY399 MATα can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY102 MKY7, RTT107-TAP::TRP1  MKY103 MKY7, SLX4-TAP::TRP1  MKY107 MKY7, NSE5-3XFLAG::KANMX6  MKY108 MKY7, RTT107-TAP::TRP1 NSE5-3XFLAG::KANMX6  MKY109 MKY7, SLX4-TAP::TRP1 NSE5-3XFLAG::KANMX6  MKY123 MKY7, RTT107-TAP::TRP1 NSE5-3XFLAG::KANMX6 slx4::HIS3  MKY124 MKY7, NSE5-3XFLAG::KANMX6 rtt107::HIS3 [pRS314, 4XTAP]  MKY125 MKY7, NSE5-3XFLAG::KANMX6 rtt107::HIS3 [pRS314, 4XTAP-RTT107aa1-1070]  MKY126 MKY7, NSE5-3XFLAG::KANMX6 rtt107::HIS3 [pRS314, 4XTAP-RTT107aa1-512]  MKY127 MKY7, NSE5-3XFLAG::KANMX6 rtt107::HIS3 [pRS314, 4XTAP-RTT107aa512-1070]  MKY139 MKY7,  rtt107::HIS3 [pRS314, 4XTAP]  MKY140 MKY7, rtt107::HIS3 [pRS314, 4XTAP-RTT107aa1-1070]  MKY141 MKY7, rtt107::HIS3 [pRS314, 4XTAP-RTT107aa1-512]  MKY142 MKY7, rtt107::HIS3 [pRS314, 4XTAP-RTT107aa512-1070]  MKY1237 PJ69-4a/α (MATa/α trpl-901/trpl-901 leu2-3,112/leu2-3,112 ura3-52/ura3-52 his3-200/his3-200 ga14A/ga14A ga18OA/ga18OA LYSZ::GALl-HIS3/LYSZ::GALl-HIS3 GAL2-ADE2/GAL2-ADE2  metZ::GAL7-lacZ/metZ::GAL7-lacZ) (James et al. 1996) MKY1238 MKY1237, [pGAL4 DBD-Dest] [pGAL4 AD-Dest]    MKY1239 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-Dest]    MKY1240 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-RTT107aa1-1070]    MKY1241 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-SMC5]     MKY1242 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-SMC6]    MKY1243 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-NSE1]    MKY1244 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-MMS21]    MKY1245 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-NSE3]    MKY1246 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-NSE4]    MKY1247 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-NSE5]    MKY1248 MKY1237, [pGAL4 DBD-RTT107aa1-1070] [pGAL4 AD-NSE6]    MKY1249 MKY1237, [pGAL4 DBD-RTT107aa1-512] [pGAL4 AD-Dest]    MKY1250 MKY1237, [pGAL4 DBD-RTT107aa1-512] [pGAL4 AD-NSE6]    MKY1251 MKY1237, [pGAL4 DBD-RTT107aa512-1070] [pGAL4 AD-Dest]    MKY1252 MKY1237, [pGAL4 DBD-RTT107aa512-1070] [pGAL4 AD-NSE6]    62  Strain Relevant Genotype Source* MKY1253 MKY1237, [pGAL4 DBD-Dest] [pGAL4 AD-NSE6]    MKY1254 MKY1237, [pGAL4 DBD-NSE6] [pGAL4 AD-Dest]    MKY1255 MKY1237, [pGAL4 DBD-NSE6] [pGAL4 AD-SMC5]    MKY1256 MKY1237, [pGAL4 DBD-NSE6] [pGAL4 AD-NSE5]    MKY1101 JKM139 (MATa hoΔ hmlΔ::ADE1 hmrΔ::ADE1 ade1-100 leu2-3,112 lys5 trp1::hisG ura3-52 ade3::GAL::HO) (Haber. 2002) MKY1102 JKM179 (MATα hoΔ hmlΔ::ADE1 hmrΔ::ADE1 ade1-100 leu2-3,112 lys5 trp1::hisG ura3-52 ade3::GAL::HO) (Haber. 2002) MKY1257 MKY1102, RTT107-3XFLAG::NATMX6  MKY1258 MKY1101, SMC5-3XFLAG::NATMX6  MKY1259 MKY1101, SMC5-3XFLAG::NATMX6 rtt107::KANMX6  MKY1260 ade2-1 trp1-1 his3-11 his3-15 ura3-1 leu2-3 leu2-112 Cir0 LEU2::GAL10-Flp(H305L)::leu2Δ1 fob1::HIS Parent from (Nielsen et al. 2009) MKY1261 MKY1260, RTT107-3XFLAG::NATMX6  MKY1262 MKY1260, 3XRFB-G418-FRT RTT107-3XFLAG::NATMX6  MKY1263 MKY1260, SMC5-3XFLAG::NATMX6  MKY1264 MKY1260, 3XRFB-G418-FRT SMC5-3XFLAG::NATMX6  MKY1265 MKY1260, SMC5-3XFLAG::NATMX6 rtt107::KANMX6  MKY1266 MKY1260, 3XRFB-G418-FRT SMC5-3XFLAG::NATMX6 rtt107::KANMX6  MKY1267 MKY6, RTT107-3XFLAG::KANMX6  MKY1268 MKY6, RTT107-3XFLAG::KANMX6 nse3-SB1::URA3 Parent from (Ben-Aroya et al. 2008) MKY1269 MKY6, RTT107-3XFLAG::KANMX6 nse5-R103G::NATMX6  MKY1270 MKY6, RTT107-3XFLAG::KANMX6 smc6-9-3XHA::HIS3 Parent from (Torres-Rosell et al. 2005) MKY419 MKY6, RAD52-GFP::NATMX6  MKY1271 MKY6, RAD52-GFP::NATMX6 rtt107::KANMX6  MKY1272 MKY6, RAD52-GFP::NATMX6 smc6-9-3XHA::HIS3 Parent from (Torres-Rosell et al. 2005) MKY1273 MKY6, RAD52-GFP::NATMX6 smc6-9-3XHA::HIS3 rtt107::KANMX6  MKY1274 MKY6, RAD52-GFP::NATMX6 nse3-SB1::URA3 Parent from (Ben-Aroya et al. 2008) MKY1275 MKY6, RAD52-GFP::NATMX6 nse3-SB1::URA3 rtt107::KANMX6   MKY1276 MKY6, RAD52-GFP::NATMX6 nse5-R103G::HYGMX  MKY1277 MKY6, RAD52-GFP::NATMX6 nse5-R103G::HYGMX rtt107::KANMX6  MKY317 MKY6, rtt107::KANMX6  MKY426 MKY6, rtt107::KANMX6 nse5-R103G::NATMX6  MKY541 MKY6, smc6-9-3XHA::HIS3 Parent from (Torres-Rosell et al. 2005) MKY542 MKY6, smc6-9-3XHA::HIS3 rtt107::KANMX6  MKY543 MKY399, SMC6-3XHA::HIS3  MKY544 MKY399, smc6-9-3XHA::HIS3 slx4::KANMX6  MKY547 MKY6, nse5-R103G::NATMX6 slx4::KANMX6  63  Strain Relevant Genotype Source* MKY1278 MKY6, nse5-R103G::NATMX6   MKY1279 MKY6, slx4::KANMX6  MKY1280 MKY6, nse3-SB1::URA3  Parent from (Ben-Aroya et al. 2008) MKY1281 MKY6, nse3-SB1::URA3 rtt107::KANMX6  MKY1282 MKY6, nse3-SB1::URA3 slx4::KANMX6  MKY414 MKY6, SMC6-3XHA::HIS3  MKY1283 MKY6, MMS21-3XHA::HIS3  MKY1284 MKY6, NSE1-3XFLAG::NATMX6  MKY1285 MKY6, RTT107-3XVSV::KANMX6  MKY1286 MKY6, NSE1-3XFLAG::NATMX6 RTT107-3XVSV::KANMX6  MKY1287 MKY6, SMC6-3XHA::HIS3 RTT107-3XVSV::KANMX6  MKY1288 MKY399, MMS21-3XHA::HIS3 RTT107-3XVSV::KANMX6  MKY1320 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX  MKY1321 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX mec1::HIS3  MKY1322 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX nse3-SB1::URA3  MKY1323 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX nse3-SB1::URA3 mec1::HIS3  MKY1324 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX nse5-R103G::NATMX6  MKY1325 MKY399, RTT107-3XFLAG::KANMX6 sml1::HYGMX nse5-R103G::NATMX6 mec1::HIS3  MKY1326 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX smc6-9-3XHA::HIS3  MKY1327 MKY6, RTT107-3XFLAG::KANMX6 sml1::HYGMX smc6-9-3XHA::HIS3 mec1::NATMX6  MKY1334 MKY6, SLX4-3XFLAG::NATMX6 SMC5-3XHA::HIS3  MKY1335 MKY6, RTT107-TAP::TRP1 SLX4-3XFLAG::NATMX6 SMC5-3XHA::HIS3   * unless otherwise indicated, all strains were constructed for this work (details available upon request) or were from the laboratory collections. All strains are in the W303 background, except for strains MKY1237-MKY1256.  3.2.2 Growth and DNA damage sensitivity assays Overnight cultures grown in YPD were diluted to 0.3 A600 and grown in YPD to ~1-2 A600.  The cells were tenfold serially diluted and spotted onto solid YPD plates or plates with MMS, CPT, or HU (Sigma) at various concentrations. The plates were then incubated at the indicated temperature for 2-3 days.  64  3.2.3 Analytical-scale interaction assays, immunoprecipitation, and phosphatase treatment Overnight cultures were diluted to 0.3 A600 and grown in YPD to 0.8 A600, and cells were collected for immunoprecipitation. The procedure for analytical-scale immunoprecipitation of the epitope tagged proteins was adapted from a previous report (Kobor et al. 2004). Briefly, yeast cells were harvested, and lysed in TAP-IP Buffer (50mM Tris [pH 7.8], 150 mM NaCl, 1.5 mM MgAc, 0.15% Nonidet P-40, 1 mM DTT, 10 mM NaPPi, 5 mM EGTA, 5 mM EDTA, 0.1mM Na3VO4, 5 mM NaF, CompleteTM Protease inhibitor mixture (Roche)) using acid-washed glass beads and mechanically disrupting using a bead beater (BioSpec Products). Epitope tagged fusion proteins were captured using IgG Sepharose beads (Amersham Biosciences), anti-FLAG M2 agarose beads (Sigma), anti-VSV agarose beads (Abcam), or anti-HA agarose beads (Sigma) and subsequently washed in TAP-IP buffer.  Captured and copurifying proteins were detected by immunoblotting with anti-FLAG M2 (Sigma), anti-HA (Applied Biological Materials) or anti-VSV (Bethyl) antibodies and visualized using the Odyssey Infrared Imaging System (Licor).   The protocol for phosphatase treatment was adapted from a previous report (Flott and Rouse. 2005). Briefly, analytical-scale immunoprecipitation was performed as described above, and washed with TAP-IP buffer lacking NaPPi, EGTA, EDTA, Na3VO4, and NaF. Beads were resuspended in 40 µl of 1X NEBuffer for PMP (50 mM HEPES, 100 mM NaCl, 2 mM DTT, 0.01% Brij 35, 1 mM MnCl2) with or without the addition of 200 units of λ-phosphatase (New England Biolabs) in the presence or absence of EDTA (100 mM), and incubated for 30 65  min at 30°C. After the incubation, beads were washed twice with the modified TAP-IP buffer, and analyzed as above.  Rtt107-TAP complexes were purified from extracts obtained from 2 l cultures that were harvested in late logarithmic phase. Our protocol was modified from published protocols to allow immunoprecipitation of the TAP tag using Calmodulin beads (Kobor et al. 2004). Briefly, cells were disrupted with a coffee grinder in the presence of dry ice pellets and resuspended in 0.8 volumes/weight of lysis buffer (40 mM Hepes-KOH [pH 7.3], 200 mM NaCl, 0.1% Tween-20, 10% glycerol, 5 mM β-mercaptoethanol, Complete Protease inhibitor cocktail [Roche]). Crude extracts were prepared by centrifugation in a SS34 rotor for 20 min at 14,000 rpm. These were then incubated with 250 µl of Calmodulin beads (Stratagene) for 120 min at 4 °C. Beads were washed with 2 x 200 µl of lysis buffer with 2 mM CaCl2 followed by 200 µl of lysis buffer with 0.5mM CaCl2. Finally, the proteins were eluted by adding 2 x 300 µl of elution buffer (50 mM Tris-Cl [pH 8.0], 100 mM NaCl, 5 mM EGTA, 10% glycerol, 1 mM DTT) to the beads and incubating for 30 min at 4 °C. A fraction of the protein eluates were then mixed with SDS sample buffer, and the rest was incubated with either anti-FLAG M2 agarose beads or anti-HA agarose beads (Sigma), and washed with wash buffer (10 mM HEPES-KOH [pH 7.3], 200 mM NaCl, 0.1% Tween-20, 10% glycerol, 1 mM EDTA, 1 mM DTT). The beads were resuspended in SDS sample buffer and analyzed as described above.  66  3.2.4 Yeast two-hybrid assays Two-hybrid assays were performed as described previously (James et al. 1996). In brief, two-hybrid strain pJ69-4a was transformed with GBD vector or GBD fusion plasmids; transformants were selected and grown on SC-Leu medium. Similarly, two-hybrid strain pJ69-4alpha was transformed with GAD vector or GAD fusion plasmids, selected and grown on SC-Trp medium. The resulting transformants were then mated, and the diploid cells containing both the GAD and GBD constructs were selected on SC-Leu-Trp plates. Overnight cultures of the dipoid strains grown in SC-Leu-Trp were diluted to ~0.3 A600 and grown in SC-Leu-Trp to ~1-2 A600.  The cells were tenfold serially diluted and spotted onto solid SC-Leu-Trp plates, SC-Leu-Trp-His plates, SC-Leu-Trp-Ade plates or SC-Leu-Trp-His plates containing 1 mM 3-amino-1,2,4-triazole (3-AT) (Fisher). The plates were then incubated at 30°C for 5 days.  Positive interactions were indicated by cell growth on SC-Leu-Trp-Ade and/ or SC-Leu-Trp-His (+ 1 mM 3-AT). Each construct was tested with vector alone as a control.  3.2.5 Chromatin immunoprecipitation (ChIP) ChIP experiments were performed as described previously (Schulze et al. 2009). In brief, yeast cells (250 ml) were grown in a rich medium to an A600 of 0.5-0.6 and were crosslinked with 1% formaldehyde for 20 min before chromatin was extracted. The chromatin was sonicated (Bioruptor, Diagenode; Sparta, NJ: 10 cycles, 30 s on/off, high setting) to yield an average DNA fragment of 500 bp. Anti-FLAG antibody (Sigma, 4 µl) were coupled to 60 µl of protein A magnetic beads (Invitrogen). After reversal of the crosslinking and DNA purification, the immunoprecipitated and input DNA were analyzed by quantitative real-time 67  PCR (qPCR) using Rotor-Gene 6000 (QIAGEN). Samples were analyzed in triplicate for three independent ChIP experiments. Primer sequences are listed in Table 3.2.  Table 3.2 Primers used in this study for qPCR   Forward Reverse Location MAT CCCATCGTCTTGCTCTTGTT ATCCGTCCCGTATAGCCAAT 0.2 kb from HO site TAF2 CAAGGATGCCCTTGTTTTGT TTTTGACGGCCAATCTTTTC 1 kb from HO site YCR043C* CCAAGGAACTAATGATCTAAGCACA CATGTTGGTACTCTAAATCACCTCC 5 kb from HO site IMG1* TGGATCATGGACAAGGTCCTAC GGCGAAAACAATGGCACTCT 10 kb from HO site PWP2* GACACACTTTACTTTGGCTTGGTT GACTTCCAAAGACTTAAGCGCA 20 kb from HO site FRT† AAGTTCGACATGGGCTTCAG TGATCTGCATGGGTTCGTTA 0.1 kb from FRT site ROG3† ATCTTTGCCAAATTGCTTCG TTTGTCGGGCCATGAGTTAT 0.85 kb from FRT site ATG18† GAAACTTCCCGTTGAAACCA TATCCATCCGAGGAAACGAC 2.7 kb from FRT site LSB3† AAGAACGAGCCCTTGACTGA TCGACGATGATGATGACGAT 5.8 kb from FRT site CDC14 TATTCGCCGTAGAAGGTTGG TTGGCGGCTTATATCCCTTA 12 kb from FRT site PRP8 GGATGTATCCAGAGGCCAAT AACCCGCGTATTAAGCCATA  Reference locus * Primer sequences from (Kim and Haber. 2009) † Primer sequences from (Nielsen et al. 2009)   3.2.6 Fluorescence microscopy To visualize Rad52-GFP foci, cells were grown at 25°C in SC-complete medium to logarithmic phase, briefly sonicated to loosen cell aggregates, and then immobilized on a glass slide with a solution of 1.0% agarose in ddH2O.  Multiple images were obtained at 0.3 µm intervals along the z axis and Rad52-GFP foci were counted by inspection of all focal planes. The cells were categorized as G1 (unbudded), S (small bud), or G2/M (medium to large bud). At least 400 cells were counted for each strain.  All imaging was done with the Zeiss Axioplan 2 fluorescence microscope using the Metamorph software.  Statistical significance was assessed using Student’s t test.  68  3.3 Results  3.3.1 Rtt107 physically interacted with the SMC5/6 complex Having previously characterized the interaction of Rtt107 with Slx4 (Roberts et al. 2006), we wished to expand our understanding of Rtt107 function by studying its interaction with the SMC5/6 complex, recently identified by a mass spectrometry approach (Ohouo et al. 2010). We first confirmed the physical interaction between Rtt107 and the SMC5/6 complex found by Ohouo et al by analytical scale interaction assays, testing immunoprecipitated epitope-tagged Rtt107 for co-purifying SMC5/6 subunits (Figure 3.1, Figure 3.2, and data not shown). Since Rtt107 interacted with all the subunits tested, we chose Nse5 as a representative subunit to further characterize the interaction. The interaction of Nse5-FLAG and Rtt107-TAP was independent of DNA damage induced by MMS (Figure 3.1A). Interestingly, Slx4 did not associate with SMC5/6, suggesting that only a fraction of cellular Rtt107 associated with SMC5/6 (Figure 3.1A). Furthermore, Slx4 was not required for the interaction between Rtt107 and SMC5/6, nor did its absence affect the relative level of Nse5 interacting with Rtt107, supporting the model that separate pools of Rtt107 interacted with either Slx4 or SMC5/6 (Figure 3.1B). To measure this more directly, Slx4-FLAG or Smc5-HA was immunoprecipitated from purified Rtt107-TAP complexes. In the FLAG immunoprecipitation fraction, Smc5-HA was depleted, while the opposite trend was observed in the HA immunoprecipitation fraction (Figure 3.1C). This suggested that the majority of Rtt107 interacted with Slx4 or SMC5/6 separately, although there may be a minor population that interacts with both.    69   Figure 3.1 Rtt107 physically interacted with the SMC5/6 complex.  (A) Rtt107-TAP but not Slx4-TAP co-immunoprecipitated with Nse5-FLAG independently of exposure to 0.025% MMS for 2 h. Analytical-scale TAP purifications were performed on whole-cell extracts of the indicated strains. Immunoblotting was performed using anti-rabbit IgG or anti-FLAG antibodies.  (B) Rtt107-TAP co-immunoprecipitated with Nse5-FLAG independently of Slx4. (C) Separate pools of Rtt107 interacted with Slx4 or SMC5/6. Eluates from large-scale TAP purifications were subsequently immunoprecipitated with anti-FLAG or anti-HA agarose beads. Immunoblotting was performed using anti-rabbit IgG, anti-HA, or anti-FLAG antibodies. (D) The N-terminal portion of Rtt107 was responsible for the interaction with Nse5-FLAG. The truncation mutants of Rtt107 were from a previous study (Roberts et al. 2006).  70   Figure 3.2 Rtt107 interacted with multiple subunits of the SMC5/6 complex.  Analytical-scale immunoprecipitations were performed on whole-cell extracts of the indicated strains. Immunoblotting was performed using the indicated antibodies. Pgk1 in the input fraction was used as a loading control. (A) Rtt107-VSV co-purified with Smc6-HA. (B) Nse1-FLAG co-purified with Rtt107-VSV. (C) Mms21-HA co-purified with Rtt107-VSV.  BRCT domains are known to mediate protein-protein interactions among a variety of DNA damage repair proteins including the Rtt107-Slx4 interaction (Roberts et al. 2006).  Rtt107 contains four BRCT domains in the N-terminus and two BRCT domains in the C-terminus (Figure 3.1D). We expressed and purified TAP-tagged truncation mutants comprising the N- or C-terminal portion of Rtt107 from a low-copy plasmid (Roberts et al. 2006) in an rtt107∆ NSE5-FLAG strain and tested their ability to co-purify with Nse5-FLAG. In contrast to the C-terminal fragment, the N-terminal fragment of Rtt107 co-purified with Nse5-FLAG, suggesting that Nse5 interacted with the N-terminal BRCT domains of Rtt107 (Figure 3.1D).    To decipher which subunit within the SMC5/6 complex was mediating the contact with Rtt107, we utilized a pair-wise yeast two-hybrid interaction matrix between Rtt107 fused to the Gal4 DNA binding domain (GBD) and the individual SMC5/6 subunits fused to the Gal4 activation domain (GAD).  As expected, Nse6 displayed a weak two-hybrid interaction with Smc5 and a strong interaction with Nse5, consistent with previous reports (Figure 3.3) (Duan et al. 2009). Cells expressing both full-length GBD-Rtt107 (FL) and GAD-Nse6 were able to 71  grow better on SC–His medium than the GAD-vector control, but not on SC–Ade medium, indicating that the full-length GBD-Rtt107 (FL) exhibited a weak two-hybrid interaction with Nse6, but not with the other subunits of the SMC5/6 complex. In addition, GAD-Nse6 displayed a strong yeast two-hybrid interaction with the N-terminal fragment of Rtt107 and very little interaction with the C-terminal fragment of Rtt107, consistent with our co-immunoprecipitation data. Together, these results suggested that Rtt107 interacted with the SMC5/6 complex via its N-terminal BRCT domains, and that this interaction was likely mediated by the Nse6 subunit.   Figure 3.3 Rtt107 interacted with the SMC5/6 complex via the Nse6 subunit in a yeast two-hybrid analysis.  Rtt107 was fused to the Gal4 DNA binding domain (GBD) and the individual SMC5/6 subunits were fused to the Gal4 activation domain (GAD). Physical interaction was indicated by growth of the yeast on SC–His and/ or SC–Ade selection media. The N-terminal and C-terminal fragments of Rtt107 used here were equivalent to the constructs in Figure 3.1C.   72  3.3.2 Rtt107 was required for recruitment of the SMC5/6 complex to a DNA double-stranded break To identify a function for the interaction between Rtt107 and the SMC5/6 complex, we first examined the DNA damage response, since both partners are involved in this process (De Piccoli et al. 2009; Roberts et al. 2006; Rouse. 2004). Given that the SMC5/6 complex is recruited to a DSB (De Piccoli et al. 2006; Lindroos et al. 2006), we tested whether Rtt107 physically associated with this genomic lesion as well. To this end, we FLAG-tagged Rtt107 in a strain that contained the HO cut site in the mating locus and the HO endonuclease under control of the galactose promoter, allowing for creation of a single DSB at a specific locus (Haber. 2002). After 2 hours of galactose induction, we performed chromatin immunoprecipitation (ChIP) and measured Rtt107-FLAG enrichment at various distances from the HO cut site by quantative real-time PCR (qPCR). Interestingly, Rtt107 was significantly enriched at regions near the DSB created by the HO endonuclease after galactose induction (Figure 3.4A). While Rtt107 was recruited at low levels at the DSB itself, its enrichment slowly increased up to a maximum at 5 kb from the DSB, and then slowly decreased with increasing distance from the DSB, up to 20 kb. Since Rtt107 physically interacted with the SMC5/6 complex, we next tested whether Rtt107 was required for recruitment of the SMC5/6 complex to the DSB. As expected, Smc5-FLAG was recruited to regions near the HO cut site after galactose induction, and this enrichment reached a maximum at 5 kb from the DSB, thus mirroring the Rtt107 binding pattern (Figure 3.4B). However, Smc5-FLAG enrichment was dramatically reduced in the absence of Rtt107, particularly in the region 5-20 kb away from the DSB where the majority of Smc5-FLAG was observed (Figure 3.4B).  73    Figure 3.4 Rtt107 was required for recruitment of the SMC5/6 complex to a double-stranded break but not to a protein-bound nick.  Fold enrichment of Rtt107-FLAG or Smc5-FLAG was determined by ChIP-qPCR. Enrichment at the target loci were normalized to the PRP8 reference locus and the corresponding input DNA. Graphs show averages from 3 independent experiments and error bars represent standard deviations. (A) Rtt107 was recruited to regions near the DSB after 2 hours of galactose induction in asychronous cells. (B) Smc5 recruitment to the DSB was dependent on Rtt107. * p < 0.05 comparing WT to rtt107Δ after galactose induction. (C) Rtt107 was recruited to regions near the protein-bound nick in the presence of the FRT target site 2 hours after release into S phase. (D) Smc5 was similarly recruited to regions near the protein-bound nick independently of Rtt107.  Since the above experiment represented only one type of DNA lesion that can be encountered by cells, we turned to another system that allows interrogation of a protein-bound nick in order to further expand our understanding of the roles of Rtt107 and the SMC5/6 complex in 74  the DNA damage response. In this system, a protein-bound nick is introduced at a specific locus in the genome by expressing a ligation-defective Flp recombinase that remains covalently bound to the DNA after forming a nick at its recognition target site (termed the FRT site) (Nielsen et al. 2009). During S phase, DNA replication forks run into the protein-bound nick and become stalled or collapsed, thus mimicking DNA damage produced by camptothecin. We expressed either Rtt107-FLAG or Smc5-FLAG in this strain and measured its enrichment near the site of DNA damage by ChIP-qPCR. Cells were arrested in G1 with alpha factor, treated with galactose to induce expression of the mutant Flp recombinase, released into S phase, and collected after 2 hours. Rtt107-FLAG was recruited to this type of DNA lesion as well, albeit at lower levels than at a DSB, and was enriched up to 12 kb from the protein-bound nick (Figure 3.4C). Similarly, Smc5-FLAG was recruited up to 6 kb away from the protein-bound nick (Figure 3.4D). Although the relative Smc5-FLAG enrichment was lower at a protein-bound nick compared to a DSB, the levels were comparable to the fold enrichment measured by qPCR at other chromsomal loci enriched for the SMC5/6 complex identified in ChIP-on-chip studies (data not shown; (Lindroos et al. 2006)). Intriguingly, the absence of Rtt107 had no significant effect on Smc5-FLAG recruitment to a protein-bound nick, thus starkly contrasting with the situation at the HO-induced DSB (Figure 3.4D). Taken together, this data suggested that Rtt107 and the SMC5/6 complex cooperate only in response to specific DNA lesions, although they both may have broad roles in the DNA damage response.  75  3.3.3 Mutations in the SMC5/6 complex resulted in phosphorylation of Rtt107 in the absence of DNA damage agents Rtt107 is phosphorylated upon exposure to DNA damage agents and this modification is important for its role in the DNA damage response (Roberts et al. 2006; Rouse. 2004).  To explore a potential regulatory relationship between Rtt107 and the SMC5/6 complex, we tested whether Rtt107 phosphorylation was dependent on the SMC5/6 complex. We FLAG-tagged Rtt107 in strains containing the nse3-SB1, nse5-R103G, or the smc6-9 hypomorph alleles (Ben-Aroya et al. 2008; Torres-Rosell et al. 2005), and exposed them to 0.03% MMS for 1 hour. Surprisingly, in the cultures not exposed to MMS, Rtt107-FLAG from the strains with an altered SMC5/6 complex exhibited partially retarded migration through the gel when compared to the wild-type strain (Figure 3.5A, compare lanes 2-4 to lane 1). This result suggested that Rtt107 was modified in the absence of MMS, most likely by phosphorylation, although not to the same extent as the phosphorylation induced by DNA damaging agents. After exposure to MMS, Rtt107 was phosphorylated regardless of whether the genes encoding for subunits of the SMC5/6 complex were mutated (Figure 3.5A). To confirm that the retarded migration of Rtt107 was caused by phosphorylation, we treated the protein extracts with λ-phosphatase alone or in the presence of the phosphatase inhibitor EDTA. The retarded migration was completely abolished in the presence of λ-phosphatase, an effect which was reversed by the addition of EDTA (Figure 3.5B). Moreover, this phosphorylation was dependent on the checkpoint kinase Mec1, as the retarded migration was eliminated in strains lacking MEC1 (Figure 3.5C).   76   Figure 3.5 In the absence of MMS, Rtt107 was phosphorylated in mutants expressing a compromised SMC5/6 complex.  (A) Cells expressing Rtt107-FLAG were untreated or treated with 0.03% MMS for 1 h. Analytical-scale immunoprecipitations of Rtt107-FLAG were performed and analyzed by immunoblotting with anti-FLAG antibodies. The reduced mobility of Rtt107-FLAG indicated phosphorylation of the protein. Cross-reaction bands were used as a loading control. (B) As in (A), except immunoprecipitates were left untreated or incubated with λ-phosphatase (λ-PP; 200 units) for 30 min at 30°C in the presence or absence of EDTA (100 mM). (C) Cells expressing Rtt107-FLAG with and without MEC1 were treated as in (A). All strains contained sml1Δ to suppress the lethality of mec1Δ mutants. 77   3.3.4 Rtt107 and the SMC5/6 Complex had independent functions in the DNA damage response To further understand the functional relationship between Rtt107 and the SMC5/6 complex in the DNA damage response, we examined the effect of deleting RTT107 on cell growth during continuous exposure to DNA damaging agents when SMC5/6 function was compromised. The nse5-R103G mutant alone was sensitive to high temperature and to DNA damaging agents. In comparison, the rtt107∆ nse5-R103G double mutant grew much slower at the semi-permissive temperature (34oC) and on low concentrations of the DNA damaging agents MMS, HU and CPT (Figure 3.6A). We then tested the effect of deleting SLX4 to compare its involvement in SMC5/6 functions to that of RTT107. The slx4∆ nse5-R103G double mutant also exhibited slower growth than either single mutant in all conditions tested (Figure 3.6A). A similar pattern of growth phenotypes was observed when rtt107∆ or slx4Δ was combined with hypomorph alleles encoding for other SMC5/6 subunits (Figure 3.7 and data not shown). The overall poorer growth of all double mutants suggested that both RTT107 and SLX4 contributed to other DNA damage response pathways that were partially redundant with the functions of the SMC5/6 complex.   As an additional indicator of involvement in the DNA damage response, we measured Rad52-GFP foci to determine whether Rtt107 and the SMC5/6 complex cooperate in limiting spontaneous DNA damage. In the absence of exogenous DNA damaging agents, rtt107∆ mutants exhibited an increased fraction of cells with Rad52-GFP foci, as previously reported (Figure 3.6B) (Alvaro et al. 2007; Levesque et al. 2010). The strains containing a 78  compromised SMC5/6 complex exhibited a range of phenotypes, from wild-type levels to increased levels of spontaneous Rad52-GFP foci, due to the hypomorphic nature of the alleles (Figure 3.6B). The double mutants displayed a tendency for higher levels of Rad52-GFP foci than the respective single mutants, suggesting that Rtt107 and SMC5/6 contribute independently to controlling spontaneous DNA damage (Figure 3.6B). Taken together, these data indicated that Rtt107 and SMC5/6 functioned separately in the DNA damage response in addition to their shared roles as one complex.   Figure 3.6 Rtt107 and the SMC5/6 complex had independent functions.  (A) Double mutants containing nse5-R103G and rtt107Δ or slx4Δ grew significantly more slowly than the respective single mutants. Tenfold serial dilutions of the indicated strains were plated onto media containing 79  various DNA damaging agents and incubated at the indicated temperatures. (B) rtt107Δ mutants exhibited an increased level of Rad52-GFP foci whereas mutants expressing hypomorph alleles of genes encoding SMC5/6 exhibited a range of phenotypes. The percentage of cells in G1, S, or G2/M phase containing Rad52-GFP foci was calculated by dividing the number of cells in G1, S, or G2/M phase containing Rad52-GFP foci by the total number of cells in that cell cycle phase. At least 150 cells were counted in a minimum of 3 independent experiments. Error bars represent standard deviations of the values.   Figure 3.7 Rtt107 and Slx4 had functions that were independent of the SMC5/6 complex.  Tenfold serial dilutions of the indicated strains were plated onto media containing various DNA damaging agents and incubated at the indicated temperatures. Double mutants containing (A) smc6-9 or (B) nse3-SB1 and rtt107Δ or slx4Δ grew significantly more slowly than the respective single mutants.   3.4 Discussion In this study, we characterized the interaction between Rtt107 and the SMC5/6 complex, and documented that this interaction was independent of DNA damage and of Slx4, the best characterized Rtt107 interaction partner. We revealed that Rtt107 was required for recruitment of the SMC5/6 complex to DNA DSBs but not at a protein-bound nick. The 80  relationship of these proteins extended to the phosphorylation status of Rtt107, as mutating genes encoding for SMC5/6 resulted in Rtt107 phosphorylation in the absence of DNA damaging agents. Although Rtt107 and the SMC5/6 complex clearly worked together in some aspects of the DNA damage response, we demonstrated that they also had independent functions.   We have confirmed the physical interaction between Rtt107 and the SMC5/6 complex first identified by mass spectrometry (Ohouo et al. 2010), and found that it was mediated by the N-terminal BRCT domains of Rtt107. A unique feature of the interaction between Rtt107 and the SMC5/6 complex was its independence from Slx4, in contrast to the Slx4-dependent interaction between Rtt107 and the DNA replication and repair protein Dpb11  (Ohouo et al. 2010). Rtt107 forms a complex with Slx4 to provide resistance to DNA alkylating agents and promote recovery from DNA damage, and loss of their function can be suppressed by inhibiting the pathway leading to H3 K79 trimethylation (Levesque et al. 2010; Roberts et al. 2006). Whereas many of the established functions of Rtt107 involve Slx4, the interaction between Rtt107 and the SMC5/6 complex did not, suggesting that this interaction represented a distinct role for Rtt107. Our data were consistent with a model in which separate pools of Rtt107 in the cell interact with either Slx4 and Dpb11 or the SMC5/6 complex. Interestingly, both Slx4 and the SMC5/6 complex bound the N-terminus of Rtt107 (Roberts et al. 2006). The functional relationship of Rtt107 and SMC5/6 was further supported by the finding that Rtt107 was phosphorylated by Mec1 when the SMC5/6 complex was compromised, even in the absence of DNA damage induced by exogenous agents. This suggested that Rtt107 acted directly or indirectly as an indicator of malfunction of the SMC5/6 complex.    81   At least one of the functions of the Rtt107-SMC5/6 complex appears to be at the site of DSBs, since both partners were recruited there upon induction of a DSB and Rtt107 was required for recruitment of the SMC5/6 complex. However this does not represent a sequential recruitment to the DSB, since Rtt107 interacted with the SMC5/6 complex constitutively, in the absence of DNA damaging agents. Therefore, they must be recruited to the DSB together as one complex upon receiving a signal from the DNA damage site. Since Rtt107 was required for the recruitment of the SMC5/6 complex, it is likely that the triggering signal occurs via Rtt107. As such, we propose a model whereby Rtt107 is recruited to the DSB by binding to phosphorylated H2A S129, a histone modification that occurs immediately after DSB formation (Downs et al. 2000). In support of this model, Brc1, the Schizosaccharomyces pombe homologue of Rtt107, binds specifically to a phosphorylated H2A peptide in vitro via the phospho-binding BRCT domains of Brc1 (Williams et al. 2010). In addition, given that Mre11, one of the early sensors of DSBs, is also required for the recruitment of the SMC5/6 complex to DSBs, it is tempting to speculate that Mre11 is responsible for recruiting Rtt107, and consequently the SMC5/6 complex, to the site of DNA damage (Lindroos et al. 2006; Lisby et al. 2004). Although we have determined that Rtt107 was required for the recruitment of the SMC5/6 complex to DSBs, the question remains whether Rtt107 has subsequent functions in regulating the function of the SMC5/6 complex at DSBs, since both rtt107Δ and smc5/6 mutants have defects in sister chromatid recombination (De Piccoli et al. 2006; Rouse. 2004).   82  Using the recently established Flp-nick system (Nielsen et al. 2009), we demonstrated that both Rtt107 and the SMC5/6 complex were recruited to a protein-bound nick. This was consistent with previous studies showing that both Rtt107 and the SMC5/6 complex are recruited to stalled replication forks (Lindroos et al. 2006; Roberts et al. 2008), which may occur at the protein-bound nick introduced by the mutant Flp recombinase. Interestingly, Smc5-FLAG recruitment was not affected by the absence of Rtt107, in contrast to the situation at DSBs. This result supported the model that SMC5/6 is involved in multiple DNA repair pathways that are differentially regulated, consistent with previous studies (Lindroos et al. 2006). Taken together, this data suggested that Rtt107 and the SMC5/6 complex cooperate only in response to specific DNA lesions, although they both may have broad roles in the DNA damage response.  While our data strongly suggested a shared function of Rtt107 and the SMC5/6 complex at DNA DSBs, their distinct biochemical associations and the phenotypes of mutants in either RTT107 or genes encoding SMC5/6 subunits point towards differing additional functions. From a genetic perspective, all the subunits of the SMC5/6 complex are encoded by essential genes, whereas RTT107 is non-essential, indicating the crucial function(s) carried out by SMC5/6 does not require Rtt107.  The genetic interactions between RTT107 and the genes encoding for SMC5/6 supports independent functions, as the double mutants were much sicker than the respective single mutants. Furthermore, they exhibit contrasting genetic interactions with other genes in the DNA damage response. Whereas deletion of DOT1 suppresses the DNA damage sensitivity of rtt107Δ mutants, it had no effect on mutants with a compromised SMC5/6 complex ((Levesque et al. 2010) and data not shown). Conversely, 83  deletion of RAD52 partially suppresses the temperature sensitivity of mutants expressing the hypomorphic alleles encoding for SMC5/6 subunits (Torres-Rosell et al. 2005), but rad52Δ rtt107Δ double mutants have increased sensitivity to DNA damaging agents (Chin et al. 2006; Rouse. 2004).   Our work revealed that a subset of Rtt107 and SMC5/6 functions are shared, most likely at the site of DSBs. However, the details of the regulatory network surrounding this relationship are yet to be discovered. The challenge of future research will be to fully elucidate the functional connection between these two important players in genome integrity. 84  Chapter 4: Conditional Genetic Interactions of RTT107, SLX4, and HRQ1 Reveal Dynamic Networks Upon DNA Damage in S. Cerevisiae3   4.1  Introduction Mapping of genetic interactions has been a valuable and powerful approach to reveal connections within complex biological systems (Baryshnikova et al. 2013). Much of these studies have been done in Saccharomyces cerevisiae because of the tools in place to create double mutants and the availability of vast arrays of mutant libraries. Although providing great biological insights, most screens to date have been conducted under unperturbed growth conditions, whereas many networks in cells respond to environmental stimuli.   A significant type of environmental stimuli is DNA damage, which can be caused by external factors such as exposure to genotoxins or UV, or internal factors such as replication fork stalling or DNA polymerase error (Lindahl. 1993). Cells are constantly exposed to these insults that if not properly repaired may compromise genomic integrity or ultimately lead to cell death.  Due to the vital importance of genomic integrity, cells have complex mechanisms to regulate the DNA damage response. DNA damage is detected by sensors, which trigger a signalling cascade, leading to the activation of the kinases Mec1 and Tel1, the yeast homologues of mammalian ATR (ATM and Rad 3-related) and ATM (ataxia-telangiectasia                                                 3 This chapter is published in G3: Genes, Genomes, Genetics. Leung, G. P., M. J. Aristizabal, N. J. Krogan and M. S. Kobor, 2014. Conditional Genetic Interactions of RTT107, SLX4, and HRQ1 Reveal Dynamic Networks Upon DNA Damage in S. Cerevisiae. G3. 4: 1059-1069. See Preface on page iv for details of my contributions. 85  mutated). These kinases in turn elicit various cellular responses including cell cycle arrest, DNA repair, apoptosis, and/or DNA damage-induced transcriptional program (Finn et al. 2012; Putnam et al. 2009).   One of the downstream phosphorylation targets of Mec1 is Rtt107/Esc4, which is required for reinitiating replication after repair of alkylating DNA damage (Roberts et al. 2006; Rouse. 2004). Deletion of the RTT107 gene results in hypersensitivity to DNA damaging agents such as the DNA alkylating agent methyl methane-sulfonate (MMS), the nucleotide reductase inhibitor hydroxyurea (HU), and the topoisomerase I poison camptothecin (CPT) (Chang et al. 2002; Parsons et al. 2006; Roberts et al. 2006; Rouse. 2004). Rtt107 contains several BRCT (BRCA1 C-terminal) homology domains, which often serve as phospho-binding modules to recruit signaling complexes and repair factors to DNA damage-induced lesions (Mohammad and Yaffe. 2009; Rouse. 2004). Consistent with a role as a scaffold for protein-protein interactions during the DNA damage response, Rtt107 interacts with a number of DNA repair and recombination proteins and is recruited to sites of DNA lesions (Chin et al. 2006; Leung et al. 2011; Ohouo et al. 2010; Roberts et al. 2006; Roberts et al. 2008; Ullal et al. 2011).    The best characterized Rtt107-interacting partner is the replication-specific endonuclease Slx4, which interacts with the N-terminal BRCT domains of Rtt107 (Roberts et al. 2006). Slx4 is required for Mec1-dependent phosphorylation of Rtt107 and, like Rtt107, facilitates resumption of DNA replication after DNA damage (Roberts et al. 2006). However, it has become clear over the last few years that Rtt107 also has Slx4-independent functions, and 86  vice versa. Consistent with this, the defects in DNA damage response are generally more severe in rtt107∆ mutants than in slx4∆ mutants, and rtt107∆ slx4∆ double mutants are more sensitive to MMS than either of the single mutants (Roberts et al. 2006).    Although Slx4 has been studied in the context of its interaction with Rtt107, the SLX4 gene was first identified in a synthetic lethal screen with SGS1, which encodes for a RecQ helicase (Mullen et al. 2001). DNA helicases represent an important class of enzymes involved in the DNA damage response, and have roles in recognition of DNA damage, DNA recombination, and stabilization of stalled replication forks (Brosh. 2013). In S. cerevisiae, Sgs1 was thought to be the only RecQ helicase family member until recently when Hrq1 was identified as a novel RecQ helicase (Barea et al. 2008; Kwon et al. 2012). The functions of Hrq1 have only been preliminarily characterized, but based on the relationship between Slx4 and Sgs1, Hrq1 may also have linkages to Slx4 that have yet to be uncovered.       Initial genome-wide studies to characterize genetic function in response to DNA damage measured the fitness of deletion mutants exposed to a variety of genotoxic insults (Giaever et al. 2004; Hillenmeyer et al. 2008; Parsons et al. 2006). However, these studies only evaluated the requirement of single genes for resistance to DNA damaging agents, while the effects on genetic networks was only studied in a small directed screen (St Onge et al. 2007). Two recent studies utilized genetic interaction mapping to gain new insights into the DNA damage response (Bandyopadhyay et al. 2010; Guenole et al. 2013). In the initial study, all possible double mutants of 418 genes were created and exposed to MMS to evaluate changes in the genetic interaction network (Bandyopadhyay et al. 2010). Using this approach, the 87  authors demonstrated that differential genetic interactions are better able to reveal functions in the DNA damage response, and identified new roles for several genes. A follow-up study expanded on this work and interrogated 55 query genes crossed to a library of more than 2000 genes in MMS, CPT, and zeocin conditions (Guenole et al. 2013). Analysis of the differential genetic networks revealed several genes that were hubs of genetic interactions, and additional experiments demonstrated that these genes had novel roles in the DNA damage response.   Here we use a similar approach of measuring conditional genetic interactions to study further the functions of Rtt107, Slx4, and Hrq1. We analyzed the significantly interacting gene pairs to identify those that emerged or changed in response to DNA damage. Overall, RTT107 exhibited more genetic interactions than SLX4 or HRQ1 in CPT conditions, indicating an important role for Rtt107 in responding to CPT. Furthermore, SLX4 and RTT107 showed distinct, and sometimes even opposing, genetic interactions, even though the protein products exist at least in part as a complex in the cell. Interestingly, the interaction profile and enriched gene ontology terms for HRQ1 most closely resembled that of SLX4, suggesting that they have overlapping functions in the DNA damage response.  4.2 Materials and Methods  4.2.1  Yeast strains All yeast strains used in this study are listed in Table 4.1 and created using standard yeast genetic techniques (Ausubel. 1987).  Complete gene deletions were achieved using one-step 88  gene integration of PCR-amplified modules (Longtine et al. 1998). Mutants for cE-MAP screens were constructed in the BY4742 background, whereas all other strains were constructed in the W303-1A background.   Table 4.1 Yeast strains used in this study. Strain Relevant Genotype Background MKY1649  Matα his3Δ1 leu2Δ0 LYS2+ met15Δ0 ura3Δ0 Δcan1::MATaPr-HIS3 Δlyp1::MATαPr-LEU2 rtt107::NATMX6 BY4742 MKY1650 Matα his3Δ1 leu2Δ0 LYS2+ met15Δ0 ura3Δ0 Δcan1::MATaPr-HIS3 Δlyp1::MATαPr-LEU2 slx4::NATMX6 BY4742 MKY1651 Matα his3Δ1 leu2Δ0 LYS2+ met15Δ0 ura3Δ0 Δcan1::MATaPr-HIS3 Δlyp1::MATαPr-LEU2 hrq1::NATMX6 BY4742 MKY5 MATα ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 W303-1A MKY7 MATa ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 W303-1A MKY1652 MKY5, rtt107::KANMX6 W303-1A MKY1653 MKY5, dot1::HIS3 W303-1A MKY959 MKY5, rtt107::KANMX6 dot1::HIS3 W303-1A MKY1654 MKY5, bre1::HYGMX W303-1A MKY1655 MKY5, rtt107::KANMX6 bre1::HYGMX W303-1A MKY1656 MKY5, slx4::HYGMX W303-1A MKY1657 MKY5, hrq1::NATMX6 W303-1A MKY1658 MKY7, slx4::NATMX6 W303-1A MKY1659 MKY5, mrc1::HIS3 W303-1A MKY1660 MKY5, mrc1::HIS3 rtt107::KANMX6 W303-1A MKY1661 MKY5, mrc1::HIS3 slx4::KANMX6 W303-1A MKY1662 MKY5, hst3::HIS3 W303-1A MKY1663 MKY5, hst4::HYGMX W303-1A MKY1664 MKY5, hst3::HIS3 slx4::KANMX6 W303-1A MKY1665 MKY5, hst3::HIS3 rtt107::KANMX6 W303-1A MKY1666 MKY5, hst4::HYGMX rtt107::KANMX6 W303-1A MKY1667 MKY5, hst4::HYGMX slx4::KANMX6 W303-1A MKY1668 MKY5, hst3::HIS3 hst4::HYGMX W303-1A MKY1669 MKY5, hst3::HIS3 hst4::HYGMX rtt107::KANMX6 W303-1A MKY1670 MKY5, lys2∆ hst3::HIS3 hst4::HYGMX slx4::KANMX6 W303-1A  89  4.2.2 Conditional epistatic miniarray profiling (cE-MAP) Conditional E-MAP screens were performed and normalized as described previously (Collins et al. 2010), with the exception that we expanded the number of plates per query strain to accommodate all the drug conditions tested (see Figure 1). Briefly, deletion mutants of the query genes were crossed, using a Singer robot, to a library of 1,536 mutants covering a number of categories, including kinases/phosphatases and chromatin biology. We used 10 and 15 µg/ml of CPT, 50 and 100 mM of HU, and 0.0075% and 0.0125% of MMS (Sigma), along with two no drug controls. All strains and conditions were screened in triplicate.  Differential S-scores were calculated by subtracting the S-score in YPD from the S-score in each drug condition for each gene pair, and converted to Z-scores. Corresponding p-values were corrected for multiple testing using the fdrtool R package (Strimmer. 2008) and conditional genetic interactions were considered significant for q-values < 0.05. cE-MAP networks were visualized using Cytoscape (Cline et al. 2007).   4.2.3 Growth & DNA damage sensitivity assays Overnight cultures grown in YPD were diluted to 0.5 OD600. The cells were tenfold serially diluted and spotted onto solid YPD plates or plates with MMS, CPT, or HU (Sigma) at the indicated concentrations. The plates were then incubated at 30°C for 2 days and subsequently photographed.  90  4.2.4 Gene ontology analysis  Database for Annotation, Visualization, and Integrated Discovery (DAVID) was used for GO term enrichment analysis (Dennis et al. 2003). For each query gene, a list of significantly interacting genes was compiled including all drug conditions and both positively and negatively interacting genes. Multiple testing correction was done using the Benjamini method and enriched GO terms were considered significant for q-values<0.05.  4.3 Results  4.3.1 Genetic interaction profiles were considerably altered when exposed to DNA damaging agents To gain a global understanding of the functions of Rtt107, Slx4, and Hrq1, we measured genetic interactions with representative genes across the genome under three DNA damaging conditions (MMS, HU, and CPT). To achieve this, we utilized a version of the synthetic genetic array technology, the epistatic miniarray profile (E-MAP), to map genetic interactions for RTT107, SLX4, and HRQ1 under different DNA damaging conditions, which we termed conditional E-MAP (cE-MAP) (Baryshnikova et al. 2010; Collins et al. 2010; Tong et al. 2001). During the mutant selection process, the plates were expanded to accommodate the four different conditions tested, and to measure each condition with three technical replicates (see Figure 4.1 for workflow).    91   Figure 4.1 Schematic diagram of the cE-MAP workflow.  The number of plates was expanded at various points in the process to accommodate for the multiple conditions tested. All screens were performed three times.  The analysis pipeline for E-MAP calculates S-scores which reflect both the strength of the genetic interaction and the statistical confidence (Collins et al. 2010). For a broad assessment of how DNA damage affected the genetic networks, we first calculated the Pearson’s correlations of the query genes’ S-score profiles under the three drug conditions (Figure 4.2A). Supporting the idea that genetic networks respond significantly to external stimuli, the genetic interaction profiles generated under unperturbed growth conditions clustered away from the profiles generated under DNA damaging conditions. Strikingly, the profiles for 92  RTT107 in DNA damaging conditions were closely correlated, regardless of the type of DNA insult. In contrast, the profiles for SLX4 and HRQ1 clustered together by the type of DNA damaging agent. In addition to these major patterns, the genetic interaction profiles of all query genes under the same type of DNA damaging agent were also positively correlated to one another (for example, see the positive correlations between the RTT107, SLX4, and HRQ1 profiles in MMS). Finally, we observed that the two concentrations of drug used produced very similar profiles, thus we averaged the scores of the two concentrations. This improved the confidence of our S-scores since they were represented by six replicates instead of three; these sets of average S-scores were used for all further analyses.  93   Figure 4.2 Genetic interaction profiles changed in response to DNA damaging agents.  (A) Pearson’s correlation revealed that the genetic interaction profiles of RTT107 were more similar to one another regardless of the drug condition, whereas SLX4 and HRQ1 were more similar to one another. (B) Density plots of the S-scores for each query gene showed a broader distribution of S-scores for RTT107 than SLX4 or HRQ1. (C) rtt107∆ mutants were more sensitive to DNA damaging agents than slx4∆ or hrq1∆. 10-fold serial dilutions of the indicated strains were plated onto media containing the indicated drugs.  According to previously published thresholds, S-scores lower than -2.4 or higher than 2.0 are considered significant genetic interactions (Collins et al. 2010). Consistent with the dynamic 94  nature of the DNA damage response, the distribution of the S-scores for each query gene changed significantly between the unperturbed growth conditions and the DNA damaging conditions (Figure 4.2B and Figure 4.3). For example, under HU conditions, the S-score distribution for each query gene became broader, indicating that the number and strength of the genetic interactions increased (Kolmogorov-Smirnov test, two-sided, p-value < 1.5 x 10-7 for all comparisons). Strikingly, rtt107∆ mutants had the broadest distribution of S-scores of the three query mutants, i.e. the most significant genetic interactions, followed by slx4∆ mutants, then hrq1Δ mutants (Figure 4.2B). We note that this was consistent with the relative DNA damage sensitivity of these mutants (Figure 4.2C). The distribution of the S-scores of all conditions for all three query genes were significantly different from one another, although the distributions for SLX4 and HRQ1 looked more similar to one another than to that of RTT107 (Kolmogorov-Smirnov test, two-sided, p-value < 7.8 x 10-16 for all comparisons, Figure 4.3D).  95   Figure 4.3 Distribution of S-scores differed between conditions or between query genes, as shown by cumulative probability plots.  (A) S-scores for RTT107. Kolmogorov-Smirnov test, two-sided, p-values for untreated vs CPT = 6.4e-09, untreated vs HU = 1.4e-07, untreated vs MMS < 2.2e-16. (B) S-scores for SLX4. Kolmogorov-Smirnov test, two-sided, p-values for untreated vs CPT = 5.6e-16, untreated vs HU = 4.9e-10, untreated vs MMS = 0.14. (C) S-scores for HRQ1. Kolmogorov-Smirnov test, two-sided, p-values for untreated vs CPT = 2.5e-03, untreated vs HU < 2.2e-16, untreated vs MMS = 7.0e-05. (D) Combined S-scores for all conditions for each query gene. Kolmogorov-Smirnov test, two-sided, p-values for RTT107 vs HRQ1 < 2.2e-16, RTT107 vs SLX4 < 2.2e-16, HRQ1 vs SLX4 = 7.8e-16.  96  4.3.2 RTT107 exhibited more conditional genetic interactions than SLX4 or HRQ1 To identify genetic interactions under DNA damage conditions that were significantly different from unperturbed growth conditions, we adapted a published method (Bandyopadhyay et al. 2010). Specifically, the differences between the S-score in the DNA damage condition and the S-score in unperturbed growth conditions for each gene pair were subtracted from the average of all the differential scores and divided by the standard error to calculate a Z score. Genes with significant Z scores after correcting for multiple testing are listed in Table 4.2 (q-value < 0.05). In total there were 569 gene pairs found to have significant differential interaction in the drug conditions tested. Of these, 378 were negative interactions (DNA damage-induced sickness or lethality) and 191 were positive interactions (DNA damage-induced epistasis or suppression). To test the reliability of this approach, we looked for known condition-specific genetic interactions. Consistent with our previously published results, deletion of DOT1 and BRE1 suppressed the DNA damage sensitivity of rtt107∆ in MMS (Levesque et al. 2010). Furthermore, the suppression by dot1∆ was additionally observed in HU but not in CPT, whereas the suppression by bre1∆ was limited to MMS, in both the cE-MAP data and the independently constructed mutants (Figure 4.4).   Table 4.2 Genes that showed condition-specific interactions with query genes that were statistically significant after multiple test correction (q<0.05).  Query Drug Interaction Significant Genes HRQ1 CPT Negative ASF1, CHL1, CLB5, CTF4, DCC1, DDC1, MMS1, MMS22, MRE11, PBY1, RAD17, RAD24, RAD52, RAD54, RAD55, RAD57, RAD59, RTT101, RTT109, SAE2 HRQ1 CPT Positive CYC8, GMH1, PDA1 HRQ1 HU Negative ARP4, ASF1, BAS1, BMH1, BRE1, CLB5, ERG5, ERJ5, GCN1, GCN20, GET2, GNP1, HPC2, IRA2, LAT1, LGE1, LST4, MET18, MFT1, MKS1, MRC1, MRE11, NPR1, PBS2, PDB1, PDE2, PMR1, POL32, RAD52, RAD54, RAD55, RAD57, RIM21, RPL34B, RPS21B, RTF1, RTG3, RTT109, SDC1, SEC22, SEC66, SEC72, SGF73, SNX4, SPF1, SUA7, SWD1, SWD3, SWI4, SYC1, UBA3, UBP15, UBP6, URE2, VPS8, YTA7 HRQ1 HU Positive  AIM21, APS3, ARO1, ARP7, BTS1, BUL1, CAP1, COQ2, CUE3, CYC8, CYT1, DCR2, FEN1, HOS2, MSS18, NGL2, PET130, THI6, YMR102C HRQ1 MMS Negative DAL81, FKS1, GFD1, GFD2, GNP1, MMS2, MOG1, MPH1, MRE11, MSH4, PBY1, PET18, PHO5, POL32, RAD10, RAD18, RAD27, RAD59, REV1, REV3, REV7, RVS161, 97  Query Drug Interaction Significant Genes SAK1, SCS7, SRO9, STP1, TRS33, UBC13, VMA21, YGL081W, YSY6 HRQ1 MMS Positive CLB5, HST3, ILM1, KAP122, PMR1, SUR4 RTT107 CPT Negative AGE2, BCK1, BFA1, BUB2, CHS5, CLB2, CLB5, COG5, CRN1, CSG2, DCC1, DDC1, DEP1, ECM33, ELA1, ERV14, FET3, FKS1, FUN30, GAS1, HPC2, IRA2, IRC21, KEX2, LAS21, LEM3, LGE1, LTE1, MAK31, NCL1, OPI3, OST3, PAC1, PBY1, PEP8, PFA4, PMT1, PMT2, PPH21, PPH3, PPM1, PRE9, PSP2, RAD17, RAD24, RAD54, RAD55, RAD57, RAD61, RDH54, RDI1, REV7, RGA1, RRD2, RTF1, RTS1, RXT2, SAP30, SCS7, SEC22, SEC28, SLA1, SMI1, SMY1, SPF1, SRO9, STV1, SUR4, SWI4, TPK3, UBP14, VPS24, VPS27, VPS29, VPS35, VPS5, VPS8, VPS9, YDR061W, YJR088C, YLR426W, YPL150W, ZDS1 RTT107 CPT Positive APQ12, ARP7, BUB1, BUB3, CDC28, CYC8, CYT1, DBF2, DPB4, ELC1, ELG1, ESS1, GAC1, GAL80, HDA1, IKI3, IPT1, JHD2, LIA1, MIP1, MKK2, MMS4, MRC1, MRT4, NAP1, NCS2, NFI1, NOP12, OAF1, PAP2, PEF1, POL32, PSY4, RAD27, RPL11B, RPL8B, RPS1B, RPS4A, RSC4, SAC3, SAM37, SCT1, SFL1, SGF29, SLC1, SMT3, SPT2, SSN2, TED1, TOP1, TRM10, TUB3, TUP1, UBX4, UFO1, YCR050C, YLR287C, YNR004W RTT107 HU Negative AGE2, BAS1, BUB2, CRN1, DUG2, GCN1, IRA2, MAD2, NPR1, OXR1, PBS2, PSP2, SEC22, SER2, SLX9, SLY41, SPF1, SUR4, SWI4, URE2, YDL089W RTT107 HU Positive AEP2, AIM26, ARO1, ARP7, BTS1, BUB3, BUD21, BUD6, BUL1, CDC48, CYC8, CYK3, CYT1, DBP1, ECM5, ERG6, HDA1, JNM1, LAG1, MET22, NAP1, NUP2, PEF1, PET130, PEX17, PMR1, POL30, PPM1, RIM101, RPN13, RPS1B, RPS28B, RPS30A, SCT1, SEF1, SPE3, SPT21, SSE1, THI6, TSA1, UBX4, UFO1 RTT107 MMS Negative BCK1, BFA1, CLB2, CRN1, DDC1, DUG2, FKS1, GFD1, GNP1, HRT1, IRC21, LSM7, LTE1, MBP1, MMS2, PAC1, POL30, PPM1, PTC2, RAD17, RAD18, RAD27, REV7, RTS1, SCS7, SLX9, SRO9, SRS2, STP1, TEL1, TSA1, UBC13, VMA21, YGL081W, YPL041C RTT107 MMS Positive AIM29, APQ12, ARO1, BMH1, BTS1, CSM3, CYT1, DBF2, DOT1, ELC1, FEN1, GAL80, GSF2, HST3, ILM1, LGE1, MRC1, MSS18, NAM7, NPP1, PER1, RAD52, RMD11, ROT2, RPL8B, RPS1B, RPS4A, RSC4, SPT2, TEP1, TMA23, TOP1, TUB3, UFO1 SLX4 CPT Negative ARP4, ASF1, CHL1, CIK1, CLB5, CSM3, CTF4, DCC1, DDC1, DIA2, GAS1, LAS21, LEM3, LGE1, MMS1, MMS22, PBY1, PMR1, PPH3, RAD17, RAD24, RAD52, RAD54, RAD55, RAD57, RAD59, RDI1, RTT101, RTT109, SAE2, SGF73, SRS2, STV1 SLX4 HU Negative AGE2, AIM32, ARL3, ARP4, BAS1, BMH2, BRE1, CLB5, COG5, CWH41, DDC1, ECM30, ERG5, ERJ5, FKH2, GCN1, GCN20, GEF1, GNP1, HCM1, HPC2, INP53, IRA2, LGE1, LST4, MDS3, MRE11, NPR1, OXR1, PBS2, PBY1, PDB1, PDE2, PFA4, PMR1, PRE9, PSP2, QCR10, QNQ1, RAD54, RAD55, RAD57, RPL41B, RPS11A, RTF1, SDC1, SEC22, SEC66, SGF73, SKY1, SNG1, SPF1, STV1, SUA7, SUR4, SWD1, SWD3, SWI4, SYC1, SYF2, UBP15, URE2, VPS27, YDR061W, YER064C, YPR063C, YSY6 SLX4 HU Positive APS3, BTS1, BUL1, CAP1, CDC36, COQ2, CYT1, FEN1, HOS2, IMP2, MDM38, MFT1, MSS18, NGL2, PET123, PET130, RPL43A, RPN13, SCD6, SPT21, TAF9, TOP1, YDL176W, YIP3 SLX4 MMS Negative CKB1, DIA2, ENT4, ERV25, GFD1, GNP1, HST3, IMP2, MMS2, MPH1, PAC1, PET18, POL32, PPH3, PSY3, RAD18, RAD26, RAD27, RAD59, REV3, REV7, RRD1, SAE2, SAP185, SCS7, STP1, TOM7, TSR3, UBC11, UBC13, VMA21, YGL081W SLX4 MMS Positive BUD14, DOT1, MSH4, NAM7, RPS21B  98   Figure 4.4 cE-MAP data recapitulated the drug-specific genetic interactions between RTT107 and DOT1 or BRE1.  (A) Shown are subsets of cE-MAP data. Blue and yellow represent negative and positive genetic interactions, respectively. (B) 10-fold serial dilutions of the indicated strains were plated onto media containing the indicated drugs.  We visualized the conditional genetic interactions in a network where nodes represented query or array genes, and edges represented significant conditional genetic interactions. The edges were colored according to the drug condition that the genetic interaction occurred in (Figure 4.5A). As revealed by the network map, there was a subset of genes that interacted with all three query genes, suggesting that they play a more general role in the DNA damage response. These included the homologous recombination genes RAD52, RAD55, and RAD57 (Figure 4.5B). Aside from this group of genes, there were also subsets that interacted with only two out of the three query genes. HRQ1 and SLX4 shared the greatest number of interacting genes, and this represented a significant overlap between these two groups (Fisher’s exact test, greater, p-value = 2.2 x 10-16, Figure 4.5A). Further supporting shared functions of RTT107 and SLX4, there was also a significant overlap between their interacting genes (Fisher’s exact test, greater, p-value = 1.7 x 10-4). Conversely, each query gene had 99  unique genetic interactions, and RTT107 had the greatest number of these (Figure 4.5A). Whereas the majority of the unique genetic interactions with RTT107 occurred under CPT conditions, HRQ1 and SLX4 had minimal numbers of unique interactions in CPT.   100   Figure 4.5 RTT107 had more significant genetic interactions than SLX4 or HRQ1.  (A) Visualization of all the significant genetic interactions. Nodes represent query or array genes, and edges are colored by the drug condition that the interaction occurred in. Blue represents CPT, green represents HU, and red represents MMS. The numbers of unique interactions for each query gene are labelled for each drug 101  condition. Indicated p-values are from Fisher’s exact tests (greater) of the genes that interact only with two out of the three query genes. (B) Enlarged view of a subset of the network indicated by the black box in (A). (C) RTT107 had more positive genetic interactions than SLX4 or HRQ1. RTT107 had more (D) positive and (E) negative genetic interactions in CPT than SLX4 or HRQ1.  When comparing the total number of genetic interactions, RTT107 had many more positively interacting genes than either SLX4 or HRQ1 (Figure 4.5C). After these interactions were separated into each drug condition, it became clear that the biggest difference in interactions occurred during exposure to CPT for both positive and negative interactions (Figure 4.5D, E). Taken together, this data suggested that Rtt107 played an important role in responding to protein-bound nicks induced by CPT. To support this, RTT107 also had a strong positive/epistatic genetic interaction with TOP1, the molecular target of CPT, whereas this interaction was absent for SLX4 and HRQ1 (Figure 4.6A).    102   Figure 4.6 Different patterns of genetic interactions were observed for genes that significantly changed their interactions in response to DNA damaging conditions.  Shown are subsets of cE-MAP data. Blue and yellow represent negative and positive genetic interactions, respectively. (A) Some genetic interactions were specific for the query genes. (B) Other genetic interactions were specific to the drug condition, and were common across all query genes.  4.3.3 Genes with condition-specific interactions were enriched for functions in the DNA damage response To further analyze the functions of RTT107, SLX4, and HRQ1 revealed by the conditional genetic interactions, we looked at the enrichment of gene ontology (GO) terms using the Database for Annotation, Visualization, and Integrated Discovery (DAVID) (Dennis et al. 2003). For each query gene, all the genes that had significant conditional interactions in all 103  three DNA damaging conditions were analyzed for GO term enrichment. Since the library of genes tested with our cE-MAP approach was already enriched for nuclear function, we used this list of genes as the background for the analysis rather than the whole set of genes in the yeast genome. All the significantly enriched GO terms were related to the DNA damage response as expected (Table 4.3), indicating that the treatments induced DNA damage specifically and not a general stress response. However, the specific enriched processes differed between the three query genes. Both SLX4 and HRQ1 were enriched for processes related to DNA metabolism and repair, and post-replication repair was one of the most over-represented terms (5.45 and 6.42 fold enrichment, respectively). Notably, the lists of enriched GO terms for SLX4 and HRQ1 were almost identical, further suggesting that these two genes have similar functions in the DNA damage response. In contrast, RTT107 was significantly enriched only for one GO term, cell cycle checkpoint (2.95 fold enrichment, q<0.03), although many of the GO terms that did not meet the significance cutoff were also related to the cell cycle (data not shown).  Table 4.3 GO terms that were significantly enriched in the list of genetically interacting genes under all conditions using DAVID.  Query GO Term Fold Enrichment q-value (Benjamini) RTT107 cell cycle checkpoint 2.95 2.56E-02 SLX4 DNA metabolic process 2.54 6.22E-07 response to DNA damage stimulus 2.53 1.87E-05 DNA recombination 4.09 1.29E-05 DNA repair 2.59 4.70E-05 recombinational repair 4.91 6.39E-04 DNA replication 3.03 7.59E-04 postreplication repair 5.45 1.94E-02 cellular response to stress 1.78 2.31E-02 HRQ1 DNA metabolic process 2.83 8.29E-08 DNA recombination 4.64 7.05E-06 response to DNA damage stimulus 2.77 5.32E-06 DNA repair 2.90 8.83E-06 DNA replication 3.44 2.28E-04 cellular response to stress 2.08 9.93E-04 postreplication repair 6.52 5.21E-03 recombinational repair 4.89 5.12E-03 104  Query GO Term Fold Enrichment q-value (Benjamini) DNA-dependent DNA replication 3.91 6.35E-03 double-strand break repair 3.67 1.13E-02 double-strand break repair via single-strand annealing 7.34 2.97E-02   4.3.4 Deletion of HST3 and MRC1 suppressed the DNA damage sensitivity of rtt107∆ but not slx4∆ mutants Hierarchical clustering of the significantly interacting genes revealed several patterns of genetic interaction profiles. There were sets of genes that were specific for the query gene, regardless of the DNA damaging agent (Figure 4.6A). Intriguingly, there were certain genes that showed strong positive interactions with RTT107, but negative interactions with SLX4, such as HST3 and MRC1, further supporting the idea that Rtt107 and Slx4 have unique functions.   Conversely, other sets of genes were specific for the DNA damaging agent, and interacted with all three query genes under that condition (Figure 4.6B). For example, the CPT-specific genes included RAD24, RAD17 and DDC1, which encode for components of the 9-1-1 checkpoint clamp and RFC loader complex (Majka and Burgers. 2007). The MMS-specific genes included multiple components of the translesion synthesis (TLS) pathway, such as REV3, REV7, and RAD18 (Sharma et al. 2013). Unexpectedly, the HU-specific genes included several transcription-related genes such as SWD1, SWD3, and SDC1.   Positive S-scores indicate that the double mutant exhibits better fitness than expected (multiplicative product of single mutants’ fitness), but it does not differentiate between 105  suppression and epistasis. To further investigate the nature of the genetic interaction between HST3 and RTT107 and SLX4, we independently constructed deletion mutants, and extended the analysis to include HST4 which was not on the E-MAP library. Hst3 and Hst4 are protein deacetylases that are both responsible for removing histone H3 K56 acetylation, thereby affecting replicative lifespan and response to DNA damage (Miller et al. 2006). Deletion of HST3 clearly suppressed the DNA damage sensitivity of rtt107Δ mutants in all three drugs tested, albeit to a lesser extent in HU (Figure 4.7A). Confirming the striking opposite interactions of RTT107 and SLX4 observed in the E-MAP, HST3 and SLX4 showed a synergistic interaction in CPT and MMS, but not HU. In general, HST4 showed the same genetic interaction profile as HST3. However, the hst3∆hst4∆rtt107∆ triple mutant showed a variable phenotype depending on the DNA damaging agent, and this differed from the hst3∆rtt107∆ or hst4∆rtt107∆ double mutants, portraying a complex relationship between HST3 and HST4 in this genetic interaction.    106   Figure 4.7 Positive S-scores from the cE-MAP data were based on suppression of DNA damage sensitivity of rtt107∆ mutants.  10-fold serial dilutions of the indicated strains were plated onto media containing the indicated drugs. (A) Deletion of HST3 and HST4 suppressed the DNA damage sensitivity of rtt107∆ mutants, but aggravated the sensitivity of slx4∆ mutants. (B) Similarly, deletion of MRC1 suppressed the DNA damage sensitivity of rtt107∆ mutants, but aggravated the sensitivity of slx4∆ mutants.  As a second example of characterizing positive interactions, we focused on MRC1, a gene encoding for an S-phase checkpoint adaptor (Tanaka. 2010), which also showed this pattern of opposite genetic interactions with RTT107 and SLX4. Using independently constructed deletion mutants, we observed that deletion of MRC1 suppressed the DNA damage sensitivity of rtt107Δ mutants to CPT and MMS (Figure 4.7B). Interestingly, the converse was observed in the case of HU, in that deletion of RTT107 mildly suppressed the sensitivity 107  of mrc1∆ mutants. In contrast, the slx4∆mrc1∆ double mutant was clearly more sensitive to CPT and MMS than the slx4∆ single mutant, although the deletion of SLX4 mildly suppressed the HU sensitivity of the mrc1∆ single mutant.  4.4 Discussion In this study we used cE-MAP to generate conditional genetic interaction profiles for RTT107, SLX4, and HRQ1, in order to further investigate their functions in the DNA damage response. We tested CPT, HU, and MMS, which elicited specific genetic interactions with each of the query genes, and the network of interactions observed provides insight into the mechanisms of both the DNA damaging agent and the query genes. Furthermore, we validated two specific examples of genetic interactions emerging from the cE-MAP in direct genetic tests, and identified a novel genetic suppression in each case.   A critical component of the cE-MAP approach was the use of DNA damaging agents. Since we are interested in interrogating the DNA damage response functions of the query genes, it was crucial to evaluate the genetic interactions in conditions when those functions are active. Testing three different DNA damaging agents also provided an opportunity to compare between the responses of the genetic interaction network to each type of DNA insult.   We observed sets of genes that showed significant interactions under each specific drug condition. For each drug condition there was different DNA damage response pathways represented within the sets of genes, supporting the idea that the cell responds specifically to different types of DNA lesions. The CPT-specific genes included components of the 9-1-1 108  checkpoint clamp and RFC loader complex, suggesting that the DNA damage response to protein adducts involves this component of the checkpoint response (Majka and Burgers. 2007). The MMS-specific genes included multiple components of the TLS pathway, which is one of the pathways in post-replication repair that allows cells to replicate past damaged bases or bulky adducts (Sharma et al. 2013). This data suggests that it also plays a role in bypassing alkylated bases. The HU-specific genes included several transcription-related genes, which was unexpected, but could be explained by an indirect effect of the mechanism of HU, which depletes the deoxynucleotide triphosphate (dNTP) pool. Interestingly, one of the HU-specific genes, YER064C, is relatively uncharacterized but was recently shown to change its cellular localization upon exposure to HU, suggesting a role for this gene in response to replication stress (Tkach et al. 2012). Consistent with our study, the previously published E-MAP analysis of the DNA damage response revealed that genes showing significant interactions in CPT are enriched for function in the DNA damage checkpoint, whereas significant genes in MMS are enriched for post-replication repair (Guenole et al. 2013).   In contrast to the drug-specific genes, there were also sets of genes that showed unique interactions with each specific query gene. These interactions suggested that the query genes we interrogated have distinct functions in the DNA damage response. It is of particular interest that RTT107 and SLX4 shared only a subset of genetic interactions, given that the Rtt107 and Slx4 proteins exist as a complex in the cell (Roberts et al. 2006). An attractive model is that there are different pools of Rtt107 and Slx4 protein complexes that contribute to specific functions, since Rtt107’s interaction with SMC5/6 and Slx4’s interaction with 109  Slx1 are independent of each other (Leung et al. 2011; Roberts et al. 2006). The human homologues of Rtt107 and Slx4, PTIP and SLX4 respectively, also have many distinct functions. While PTIP is involved in the DNA damage signaling cascades and DNA repair pathway choice (Callen et al. 2013; Gong et al. 2009; Wu et al. 2009), SLX4 has roles in Holliday junction resolution and telomere length regulation (Castor et al. 2013; Garner et al. 2013; Wan et al. 2013; Wilson et al. 2013; Wyatt et al. 2013). The data from this cE-MAP provides an opportunity to further elucidate the unique functions of Rtt107 and Slx4, which may be further dissected into responses to different DNA lesions.   The cE-MAP data also provided more insight into the function of the helicase Hrq1, which has only been preliminarily characterized (Bochman et al. 2014; Choi et al. 2013; Kwon et al. 2012). Interestingly, the genetic interaction profile of HRQ1 correlated more closely to SLX4 than RTT107, and the sets of genes that interacted with SLX4 or HRQ1 overlapped significantly. In addition, GO analysis of the significantly interacting genes returned almost identical GO terms for HRQ1 and SLX4. The close relationship between HRQ1 and SLX4 revealed by the cE-MAP data is supported by previous studies showing that SLX4 is synthetic lethal with SGS1, the major RecQ helicase in S. cerevisiae (Mullen et al. 2001). Moreover, there is some evidence suggesting that Hrq1 and Slx4 are both involved in interstrand crosslink repair and suppression of telomere addition (Bochman et al. 2014; Ward et al. 2012; Zhang et al. 2006). However, slx4∆ and hrq1∆ mutants display different sensitivities to DNA damaging agents, indicating they function separately as well (Bochman et al. 2014; Choi et al. 2013; Flott and Rouse. 2005). Further experiments are needed to determine the roles of Hrq1 and Slx4 in DNA structure maintenance and the nature of their relationship in 110  these functions. Possible routes of inquiry can be suggested by additional examination of the genetic data.   We followed up on two genes that had particularly striking conditional genetic interactions. Both HST3 and MRC1 showed a strong positive genetic interaction with RTT107 but a negative interaction with SLX4. Although Rtt107 and Slx4 form a complex, these genetic interactions suggest that not only do Rtt107 and Slx4 have independent functions, but they may have opposing functions in these contexts. Using a direct genetic test, we found that deletion of HST3, as well as HST4, suppressed the DNA damage sensitivity of rtt107∆ mutants. The known target of the Hst3 and Hst4 deacetylases is H3 K56 acetylation (H3 K56ac). Whereas deletion of HST3 alone causes an increase in H3 K56ac, deletion of HST4 alone does not change the acetylation levels, and only in the double mutant are all H3 molecules completely acetylated (Celic et al. 2006). Intriguingly, the suppression of the rtt107∆ mutant phenotype in CPT was observed upon deletion of HST3 or both HST3 and HST4, but not HST4 alone. In contrast, deletion of either HST3 or HST4 alone was sufficient to suppress the DNA damage sensitivity of rtt107∆ mutants to MMS and HU. Based on this data, we speculate that the deacetylation of H3 K56ac may be important to the genetic interaction in CPT, whereas it is not relevant in MMS or HU conditions, rather there may be a different function or target of Hst3 and Hst4 involved. Similarly for the genetic interaction with SLX4, deletion of HST3 or HST4 alone exhibited the same phenotype, thus suggesting that deacetylation of H3 K56ac is not involved.    111  We also validated the genetic interaction with MRC1, and found that deletion of MRC1 suppressed the DNA damage sensitivity of rtt107∆ mutants in CPT and MMS, but aggravated the sensitivity of slx4∆ mutants. Interestingly, the situation was different in HU, where deletion of either RTT107 or SLX4 mildly suppressed the sensitivity of mrc1∆ mutants. This is consistent with a model proposed by a previous study suggesting that Rtt107 and Slx4 inhibit the checkpoint adaptor protein Rad9, which is normally not important in replication stress, but becomes crucial in the absence of Mrc1 (Ohouo et al. 2013). However this model does not explain the genetic interactions observed in CPT and MMS, and reflects the distinct responses to various types of DNA lesions, as well as the multiple functions of DNA damage response proteins.   Our study contributes to the growing of body of data that has mapped genetic interactions in response to DNA damage, and further validates it as a fruitful approach that reveals condition-specific functions and pathways in the cell. There remains much ground to be covered as we have only started to characterize the pathways specific for the multitude of environmental conditions that affect all living organisms.   112  Chapter 5: Rtt107 Acts as a Scaffold at DNA Lesions by Anchoring to Phosphorylated H2A   5.1  Introduction A crucial element of the cellular response to DNA damage is the activation and amplification of signalling pathways to ensure proper detection and repair of DNA lesions. The central players in the signalling cascades are the phosphoinositol-3-kinase-related kinases (PIKK) Mec1 and Tel1 (Finn et al. 2012). They are activated via a variety of pathways, which are initiated by the DNA damage sensors including the MRX complex and RPA protein. Mec1 and Tel1 then phosphorylate a host of targets that mediate DNA damage responses including cell cycle arrest, transcription induction and repair pathways.   The cell uses post-translational modifications to respond to external stimuli and exert dynamic control over biological processes. In the DNA damage response, phosphorylation is one of the key modifications, and has been extensively characterized. Correspondingly, many of the proteins in these pathways contain domains that recognize phosphorylation (Mohammad and Yaffe. 2009). One major class of these phospho-recognition domains is the BRCA1 C-terminal (BRCT) domain, which was first identified in the tumour suppressor BRCA1 (Bork et al. 1997; Callebaut and Mornon. 1997; Manke et al. 2003; Yu et al. 2003). BRCT domains preferentially recognize phosphoserine, and a pair of BRCT domains typically works together in tandem to form a binding pocket for the phosphate group (Leung 113  and Glover. 2011). There are multiple examples of BRCT domains mediating phosphorylation-dependent protein-protein interactions that are crucial for protein recruitment to DNA lesions and subsequent pathway activation (Leung and Glover. 2011).   One of the first targets of Mec1 is phosphorylation of H2A S129 (termed γH2A) (Downs et al. 2000). The modification spreads up to 50 kb away from the site of DNA damage, and is believed to form a binding platform for a number of downstream factors (Papamichos-Chronakis and Peterson. 2013; Shroff et al. 2004). These include the chromatin remodelling complexes Swr1 and Ino80, and the histone acetyltransferase complex NuA4 (Downs et al. 2004). The checkpoint adaptor Rad9 is also recruited by γH2A, and this binding is mediated by the pair of BRCT domains in Rad9 (Hammet et al. 2007; Javaheri et al. 2006; Toh et al. 2006). Surprisingly, the H2A S129A mutant is only mildly sensitive to DNA damaging agents and can still activate checkpoint arrest (Downs et al. 2000; Keogh et al. 2006; Redon et al. 2003).   Another target of Mec1 is Rtt107, which is phosphorylated at several S/T-Q motifs (Rouse. 2004). In addition, Rtt107 contains six BRCT domains, suggesting that it acts as a scaffold in the DNA damage response. Rtt107 forms foci upon treatment with methyl methanesulfonate (MMS) or hydroxyurea (HU), and is also recruited locally to sites of induced DNA damage (Chin et al. 2006; Leung et al. 2011; Ullal et al. 2011). Using bulk chromosome spreads, a previous study found that Rtt107 recruitment to chromatin in the presence of stalled replication forks is dependent on the acetyltransferase Rtt109 and the cullin Rtt101 (Roberts 114  et al. 2008). Moreover, a recent study solved the crystal structure of the fifth and sixth BRCT domains of Rtt107 bound to a H2A phospho-peptide (Li et al. 2012).   Rtt107 physically interacts with a number of proteins involved in the DNA damage response. This includes the endonuclease Slx4, which forms a complex with Rtt107 and has a close functional relationship (Roberts et al. 2006). Yeast lacking RTT107 or SLX4 share common phenotypes such as sensitivity to DNA damaging agents and prolonged DNA damage checkpoint activation (Roberts et al. 2006). Moreover, Slx4 is required for Mec1-mediated phosphorylation of Rtt107, and vice versa (Levesque et al. 2010; Roberts et al. 2006). Interestingly, the DNA replication protein Dpb11 interacts with both Rtt107 and Slx4, but only after exposure to DNA damage (Ohouo et al. 2010). This interaction is proposed to counteract hyperactivation of the DNA damage checkpoint (Ohouo et al. 2013).    In this study we sought to examine the roles of the six BRCT domains in Rtt107 and how they contribute to a scaffolding function in the DNA damage response. Unexpectedly, we found that the Rtt107 protein was unstable when the BRCT1/2 or BRCT3/4 pairs of domains were removed. Focusing on the BRCT5/6 pair, we demonstrated that Rtt107 was recruited to DNA lesions via its C-terminal BRCT domains binding to γH2A. Furthermore Slx4 recruitment to DNA lesions was dependent on Rtt107. However Dpb11 recruitment to a double-stranded break (DSB) was only partially dependent on Rtt107, and was completely independent of Rtt107 in the case of a protein-bound nick.         115  5.2 Materials and Methods  5.2.1 Yeast strains & plasmids All yeast strains used in this study are listed in Table 5.1 and created using standard yeast genetic techniques (Ausubel. 1987).  Complete gene deletions and integration of FLAG tags at the 3’ end of genes were achieved using one-step gene integration of PCR-amplified modules (Funakoshi and Hochstrasser. 2009; Longtine et al. 1998). Rtt107 mutants were constructed using overlap extension PCR mutagenesis (Heckman and Pease. 2007). The H2A S129* mutant strain was a generous gift from Susan Gasser (Basel).   Table 5.1 Yeast strains used in this study.  Strain Relevant Genotype Source* MKY5 MATα ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1  MKY6  MATa can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY996 MKY5, RTT107-3XFLAG::NATMX6  MKY1267 MKY6, RTT107-3XFLAG::KANMX6  MKY1694 MKY5, rtt107∆BRCT1/2-3XFLAG::NATMX6  MKY1695 MKY5, rtt107∆BRCT3/4-3XFLAG::NATMX6  MKY1696 MKY6, rtt107∆BRCT5/6-3XFLAG::NATMX6  MKY1057 MKY5, rtt107::KANMX6 [pRS315]  MKY1671 MKY5, rtt107::KANMX6 [pRS315, RTT107-3XFLAG::NATMX6]  MKY1672 MKY5, rtt107::KANMX6 [pRS315, rtt107-K887M-3XFLAG::NATMX6]  MKY1697 MKY5, rtt107::KANMX6 [pRS315, rtt107-820-3XFLAG::NATMX6]  MKY1698 MKY5, rtt107::KANMX6 [pRS315, rtt107-BRCT1-4,1/2-3XFLAG::NATMX6]  MKY1699 MKY5, rtt107::KANMX6 [pRS315, rtt107- BRCT1-4,3/4-3XFLAG::NATMX6]  MKY1101 JKM139 (MATa hoΔ hmlΔ::ADE1 hmrΔ::ADE1 ade1-100 leu2-3,112 lys5 trp1::hisG ura3-52 ade3::GAL::HO) (Haber. 2002) MKY1673 MKY1101, RTT107-3XFLAG::NATMX6  MKY1674 MKY1101, RTT107-3XFLAG::NATMX6 hta1-S129* hta2-S129* Parent from (van Attikum et al. 2004) MKY1675 MKY1101, rtt107-820-3XFLAG::NATMX6  MKY1676 MKY1101, rtt107::KANMX6 [pRS315, RTT107-3XFLAG::NATMX6]  MKY1677 MKY1101, rtt107::KANMX6 [pRS315, rtt107-K887M-3XFLAG::NATMX6]  116  Strain Relevant Genotype Source* MKY1680 MKY1101, SLX4-3XFLAG::NATMX6  MKY1681 MKY1101, SLX4-3XFLAG::NATMX6 rtt107::KANMX6  MKY1682 MKY1101, SLX4-3XFLAG::NATMX6 rtt107-820-3XHA:KANMX6  MKY1687 MKY1101, DPB11-3XFLAG::NATMX6  MKY1688 MKY1101, DPB11-3XFLAG::NATMX6 rtt107::KANMX6  MKY1689 MKY1101, DPB11-3XFLAG::NATMX6 rtt107-820-3XHA:KANMX6  MKY1260 MATa ade2-1 trp1-1 his3-11 his3-15 ura3-1 leu2-3 leu2-112 Cir0 LEU2::GAL10-Flp(H305L)::leu2Δ1 fob1::HIS Parent from (Nielsen et al. 2009) MKY1261 MKY1260, RTT107-3XFLAG::NATMX6  MKY1262 MKY1260, 3XRFB-G418-FRT RTT107-3XFLAG::NATMX6  MKY1678 MKY1260, rtt107-820-3XFLAG::NATMX6  MKY1679 MKY1260, 3XRFB-G418-FRT rtt107-820-3XFLAG::NATMX6  MKY1683 MKY1260, SLX4-3XFLAG::NATMX6  MKY1684 MKY1260, 3XRFB-G418-FRT SLX4-3XFLAG::NATMX6  MKY1685 MKY1260, SLX4-3XFLAG::NATMX6 rtt107::HYGMX  MKY1686 MKY1260, 3XRFB-G418-FRT SLX4-3XFLAG::NATMX6 rtt107::HYGMX  MKY1690 MKY1260, DPB11-3XFLAG::NATMX6  MKY1691 MKY1260, 3XRFB-G418-FRT DPB11-3XFLAG::NATMX6  MKY1692 MKY1260, DPB11-3XFLAG::NATMX6 rtt107::HYGMX   MKY1693 MKY1260, 3XRFB-G418-FRT DPB11-3XFLAG::NATMX6 rtt107::HYGMX  163  Zou, L., and S. J. Elledge, 2003 Sensing DNA damage through ATRIP recognition of RPA-

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