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UBC Theses and Dissertations

Defects in messenger RNA processing and biogenesis of RNA polymerases contribute to eukaryotic genome… Minaker, Sean Wilson 2013

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   Genome instability has been observed in mutants involved in various aspects of transcription and RNA processing. The prevalence of this mechanism among essential chromosome instability (CIN) genes remains unclear. In this thesis, it is shown that RNA biogenesis mutants exhibit elevated sensitivity to DNA damaging agents. A secondary screen for increased Rad52 foci in CIN mutants, representing ~25% of essential genes, identified seven essential subunits of the mRNA cleavage and polyadenylation (mCP) machinery. Genome-wide analysis of fragile sites by ChIP-chip of phosphorylated-H2A in these mutants supported a transcription-dependent mechanism of DNA damage characteristic of RNA:DNA hybrids known as R-loops, which were subsequently observed in mCP mutants. Among the CIN mutants with elevated Rad52 foci levels were the GPN proteins, a poorly-characterized and deeply evolutionarily conserved family of three paralogous small GTPases, Gpn1, 2 and 3. The founding member, GPN1/NPA3/XAB1, is proposed to function in nuclear import of RNA polymerase II along with a recently described protein called Iwr1. Here, it is shown that the previously uncharacterized protein Gpn2 binds both Gpn3 and Npa3/Gpn1, and that temperature-sensitive alleles of Saccharomyces cerevisiae GPN2 and GPN3 exhibit genetic interactions with RNA polymerase II mutants, hypersensitivity to transcription inhibition and defects in RNA polymerase II nuclear localization. Importantly, previously unrecognized RNA polymerase III localization defects were observed in GPN2, GPN3 and IWR1 mutant  	
    backgrounds but no localization defects of unrelated nuclear proteins or of RNA polymerase I were found. In this study, it was shown that the nuclear import defect of iwr1Δ, but not the GPN2 or GPN3 mutant defects, is partially suppressed by fusion of a nuclear localization signal to the RNA polymerase II subunit Rpb3. These data, combined with strong genetic interactions between GPN2 and IWR1 suggest that the GPN proteins function upstream of Iwr1 in RNA polymerase II and III biogenesis. We propose that the three GPN proteins execute a common function in RNA polymerase assembly and subsequent transport. These findings demonstrate how mRNA cleavage and polyadenylation and proper RNA polymerase assembly contribute to maintenance of genome 	
    integrity and may be relevant to certain human cancers.	
   A version of Chapter 2 has been published. Stirling PC*, Chan YA*, Minaker SW*, Aristizabal MJ, Barrett I, Sipahimalani P, Kobor MS, Hieter P (2012).  R-loop-mediated  genome  instability  in  mRNA  cleavage  and  polyadenylation mutants. Genes and Development; Jan 15;26(2):163-75. Copyright © 2012, Genes and Development by Cold Spring Harbor Laboratory Press. *These authors contributed equally to this work. The figures and data represent work I directly contributed to with the exception of the R-loop staining experiments performed by Y. Chan. P. Sipahimalani and P. Stirling contributed to microscopy of yeast strains and data analysis of chIP-chip data was performed by P. Stirling. M. Aristizabal provided guidance for microarray experiments that I performed in the laboratory of M. Kobor. T. P. Stirling, Y. Chan, S. Minaker and P. Hieter wrote the manuscript upon which this section is based.  The chemical genomics screen presented in Chapter 2 as figure 2.1-2.3 and Table 2.1 and 2.2 represents my contribution to a published work. van Pel, DM, Stirling, PC, Minaker, SW, Sipahimalani, P, Hieter, P. Saccharomyces cerevisiae Genetics Predicts Candidate Therapeutic Genetic Interactions at the Mammalian Replication Fork. G3 (Bethesda). 2013 Feb;3(2):273-82. Copyright © 2013 van Pel et al. I contributed technical work in preparing the screen while P. Stirling conducted data analysis. D. van Pel, P. Stirling and P. Hieter prepared the manuscript from which this data was extracted.  	
    A version of Chapter 3 has been published. Minaker, SW, Filiatrault, MC, Ben-Aroya, S, Hieter, P, Stirling PC. Biogenesis of RNA Polymerases II and III Requires the Conserved GPN small GTPases in Saccharomyces cerevisiae. Genetics. 2013 Mar;193(3):853-64. Copyright © 2013 The Genetics Society of America. I contributed the majority of the experimental work included here. P. Stirling contributed to cell imaging and data analysis and strain construction. M. Filiatrault performed some of the imaging and plating assays during the revision process. S. Minaker, P. Stirling and P. Hieter wrote the manuscript upon which this section was based.  	
    Table of Contents Abstract................................................................................................................ ii	
   Preface ................................................................................................................ iv	
   Table of Contents ............................................................................................... vi	
   List of Tables ...................................................................................................... xi	
   List of Figures ................................................................................................... xii	
   List of Abbreviations ........................................................................................ xv	
   Acknowledgements ........................................................................................ xvii	
   Dedication ....................................................................................................... xviii	
   Chapter 1. Introduction....................................................................................... 1	
   1.1 Genome instability is a major contributing factor in human cancers ........... 1	
   1.2 CIN as a potential therapeutic target in cancer therapy .............................. 3	
   1.3 Budding yeast is an excellent model organism for discovering and characterizing CIN genes .................................................................................. 5	
   1.4 The fidelity of mRNA processing is essential for maintaining genomic integrity .............................................................................................................. 8	
   1.5 R-loop formation compromises DNA integrity through spontaneous DNA damage ........................................................................................................... 13	
   1.6 Mutation of eukaryotic transcription machinery results in CIN and DNA damage sensitivity ........................................................................................... 15	
   1.7 Research scope of this thesis ................................................................... 17	
    Chapter 2. Defects in mRNA processing promote a loss of genomic integrity in Saccharomyces cerevisiae ........................................................... 19	
   2.1 Background information ............................................................................ 19	
   2.2 Materials and methods .............................................................................. 21	
   2.2.1 Yeast strains and plasmids ................................................................. 21	
   2.2.2 Chemical genomic screening .............................................................. 21	
   2.2.3 Rad52-YFP foci screening and CIN assays ....................................... 22	
   2.2.4 ChIP-chip ............................................................................................ 22	
   2.2.5 ChIP-chip statistical analysis .............................................................. 23	
   2.2.6 Yeast chromosome spreads ............................................................... 23	
   2.2.7 Growth curve analysis ........................................................................ 23	
   2.3 Results....................................................................................................... 24	
   2.3.1 Selective killing of CIN mutants by genome-destabilizing chemicals . 24	
   2.3.2 Rad52 foci screening of essential CIN genes ..................................... 29	
   2.3.3 Fragile sites in mCP mutants differ in transcribed ORFs and near replication origins ......................................................................................... 33	
   2.3.4 CIN in mRNA cleavage and polyadenylation mutants occurs via Rloops ............................................................................................................ 39	
   2.3.5 Comparative analysis of mCP mutants with known R-loop forming mutants ........................................................................................................ 42	
   2.3.6 CIN mutants for GPN family members result in moderate accumulation of DNA double strand breaks ....................................................................... 46	
   2.4 Discussion ................................................................................................. 49	
    2.4.1 Chemical genomic and Rad52 foci screening of essential CIN genes 49	
   2.4.2 Profiling fragile site changes in Rad52 foci forming mRNA processing mutants ........................................................................................................ 51	
   2.4.3 Mechanism of genome instability in mCP mutants ............................. 52	
   Chapter 3. Biogenesis of RNA polymerase II and III requires the GPN small GTPases in Saccharomyces cerevisiae .......................................................... 56	
   3.1 Background information ............................................................................ 56	
   3.2 Materials and methods .............................................................................. 58	
   3.2.1 Yeast growth, strains and plasmids .................................................... 58	
   3.2.2 GPN mutant allele sequencing ........................................................... 59	
   3.2.3 Microscopy .......................................................................................... 59	
   3.2.4 Localization scoring and statistics ...................................................... 60	
   3.2.5 TAP pull-downs and western blotting ................................................. 61	
   3.2.6. Nuclear proteome scale GFP microscopy screen ............................. 62	
   3.3 Results....................................................................................................... 63	
   3.3.1 Mutational analysis of the GPNs: an evolutionarily conserved protein family............................................................................................................ 63	
   3.3.2 GPN2 exhibits genetic and physical interactions with GPN3 and NPA3/GPN1 and IWR1 ................................................................................ 68	
   3.3.3 GPN2 and GPN3 mutants are defective in localization and stability of RNAPII subunits........................................................................................... 71	
   3.3.4 GPN2, GPN3 and IWR1 mutants are defective in localization of RNAPIII ........................................................................................................ 79	
    3.3.5 Fusing a nuclear localization signal to Rpb3p partially bypasses IWR1 but not GPN mutants ................................................................................... 82	
   3.3.6 A nuclear proteome wide GFP localization screen reveals that GPN2 mutation affects localization of transcription associated proteins ................ 85	
   3.4 Discussion ................................................................................................. 88	
   3.4.1 Gpn2 and Gpn3 are part of an evolutionarily conserved family required for RNA polymerase II biogenesis genomic integrity ................................... 88	
   3.4.2. Gpn2 and Gpn3 function is required for nuclear localization of RNA polymerase III but not RNA polymerase I .................................................... 90	
   3.4.3. The GPN proteins appear to function upstream of Iwr1 in RNA polymerase II biogenesis ............................................................................. 91	
   Chapter 4. Concluding remarks and future directions ................................. 95	
   4.1 Expanding our understanding of contributing factors to DNA damage ..... 95	
   4.2 ChIP-chip identifies enrichment of genes adjacent to replication origins .. 96	
   4.3. R-loop formation as a mechanism for DNA damage in mRNA processing mutants. ........................................................................................................... 97	
   4.4 RNA processing mutations and human disease ....................................... 98	
   4.5 Mutation of the GPN family of GTPases and genomic instability ............ 101	
   4.6 The functional relationship between GPN2 and GPN3 proteins ............. 102	
   4.7 RNA polymerase III assembly and nuclear import requires both the GPN proteins and Iwr1 ........................................................................................... 103	
   4.8 Extending the function of the GPN proteins beyond RNA polymerase assembly ....................................................................................................... 105	
    4.9 Concluding remarks ................................................................................ 106	
   References ....................................................................................................... 108	
   Appendix .......................................................................................................... 126	
    List of Tables Table 2.1 Rationale for chemicals used in genome-wide analysis ..................... 25 Table 2.2 RNA biogenesis genes with mutant alleles that exhibit enhanced drug sensitivity to hydroyurea, MMS and benomyl. ............................................. 26 Table 3.1. Mutations in GPN2 and GPN3 alleles ................................................ 65 Table 3.2 Validated hits from screen for mislocalization of GFP tagged nuclear proteins in a gpn2-2 mutant background shows enrichment for transcription and splicing factors. ..................................................................................... 88 Table 4.1 Frequency of mutation and copy number changes in select human tumors using data from the cBio Cancer Genomics Portal for 18 human splicing gene orthologs to yeast CIN genes. ............................................. 100 Table A1 Yeast strains used in Chapter 2. ....................................................... 129 Table A2 Yeast strains used in Chapter 3. ....................................................... 133 Table A3 Plasmids used in this thesis.. ............................................................ 136  	
    List of Figures 	
   Figure 2.1 Gene ontology (GO) derived cellular functional groups uncovered by chemical screening of essential gene mutants.. .......................................... 27 Figure 2.2 The spectrum of genotoxic drug sensitivities of essential genes is enriched for CIN genes ................................................................................ 29 Figure 2.3 A screen for DNA damage foci in essential CIN genes. .................... 31 Figure 2.4 Genetic interactions between foci generating mutants and rad52Δ..	
   Figure 2.5 High-resolution mapping of yeast γ-sites	
   Figure 2.6 mCP mutants are enriched for γ-sites adjacent to origins of replication. 	
   Figure 2.7 clp1-ts and pcf11-2 mutant γ-sites link DNA damage to transcription. 	
   Figure 2.8 DNA damage in mCP mutants is biased towards genes with higher transcriptional frequency.	
   Figure 2.9 Transcription coupled R-loops are the likely cause of CIN in mCP mutants.	
   Figure 2.10 Genetic interactions of mCP mutants and known R-loop forming mutants.	
   Figure 2.11 mCP mutants exhibit recombination rates similar to THO mutants..	
   Figure 2.12 Spontaneous DNA damage and genetic interactions with splicing machinery in GPN mutants..	
   Figure 3.1 Phylogenetic tree of GPN orthologues.	
   Figure 3.2	
  Generation and characterization of GPN2 and GPN3 ts-alleles.	
    Figure 3.3 Chromosomal instability and DNA damage sensitivities of GPN mutants..	
   Figure 3.4 Gpn2 physically interacts with both Gpn1 and Gpn3 but does not appear to homodimerize.	
   Figure 3.5 Genetic interactions between GPN2 mutants and NPA3/GPN1, GPN3 and IWR1 mutants.	
   Figure 3.6 RNA polymerase II mislocalization in GPN mutants.	
   Figure 3.7 Overexpression of GPN family members and IWR1 does not strongly complement growth and Rpb1 localization defects in GPN mutants.	
   Figure 3.8 GTP binding is required for function of the GPN proteins.	
   Figure 3.9 Functional impact of GPN mutations on RNAPII.	
   Figure 3.10 Effect of the GPN-IWR1 system on RNA polymerase III localization. 	
   Figure 3.11 RNA polymerase III is mislocalized in iwr1Δ mutants.	
   Figure 3.12 Cellular effects of Rpb3-NLS fusion on RNAPII localization.	
   Figure 3.13 NLS fusion constructs partially rescues the RNA polymerase mislocalization of iwr1Δ mutants.	
   Figure 3.14 Examples of GFP microscopy displaying effects of gpn2-2 mutation on localization of nuclear proteins.	
   Figure 3.15 Model for GPN1, 2 and 3 function in biogenesis of RNA polymerases II and III.	
   Figure A1.Transcriptional frequency of ORFs associated with mCP mutantinduced g-H2A signal in their 3’-UTRs.	
    Figure A2. Rpa135-GFP is not mislocalized in GPN2, GPN3 or IWR1 mutants. 	
   Figure A3. Effect of IWR1-NLS mutation on RNA Polymerase II and III subunits. 	
    List of Abbreviations 	
   6-AU  6-azauracil  ALF  A-like faker  ANOVA  Analysis of variance  ARS  Autonomously replicating sequence  BiM  Bimater  CCT  Chaperonin-containing tailless  ChIP-chip  Chromatin immunoprecipitation and microarray	
    CIN  Chromosomal instability  CLL  Chronic lymphocytic leukemia  Ctf  Chromosome transmission fidelity  DAmP  Decreased Abundance by mRNA Perturbation  DAPI  4',6-diamidino-2-phenylindole  GCR  Gross chromosomal rearrangement  GFP  Green fluorescent protein  GPN  Glycine-proline-asparagine  HR  Homologous recombination  HRP  Horseradish peroxidase  HU  Hydroxyurea  IP  Immunoprecipitation  LOH  Loss of heterozygosity  mCP  mRNA cleavage and polyadenylation  MEF  mouse embryonic fibroblasts  	
    MMS  Methyl methanesulfonate  NHEJ  Non-homologous end joining  NLS  Nuclear localization signal  ORF  Open reading frame  RNAP  RNA polymerase  SC  Synthetic complete (medium)  SGA  Synthetic genetic array  SL  Synthetic lethal  Ts  Temperature-sensitive  TAP  Tandem affinity purification  UV  Ultraviolet  WT  wild type  YPD  Yeast peptone dextrose (medium)  	
   I thank my supervisor Dr. Philip Hieter and my colleagues and mentors Dr. Peter Stirling and Dr. Shay Ben-Aroya for their guidance during my graduate studies. I also thank my committee members Dr. LeAnn Howe and Dr. Leonard Foster for their scientific input and helpful suggestions during my thesis work.  I would like to thank my peers in the laboratory, particularly Megan Filliatrault and Payal Sipahamlani for their contributions to this research, friendship and regular coffee and cookie runs. Thanks also to my labmates Derek van Pel, Nigel O’Neil and Lee Tang for providing a welcome distraction from the trials and tribulations of benchwork as friends and fermentation science product enthusiasts.  I acknowledge personal financial support from the Canadian Institutes of Health Research and the University of British Columbia and support for the Hieter Lab provided by the National Institutes of Health and the Howard Hughes Medical Institute. Thanks also to the staff of the Michael Smith Laboratories, particularly David Thomson who was most helpful in dealing with administrative issues.  I would like to thank my parents for introducing me to biology as a child and allowing me to choose my own path in life while supporting me constantly. Most of all, I would like to thank my wife Peggy for her support and tolerance throughout my degree and for pushing me to carry on through the ups and downs. Meeting her was my greatest discovery during graduate school.  	
    Dedication  To my wife Peggy, for her love and support. To my grandparents, who never got the chance for the education they deserved.  	
    Chapter 1. Introduction 1.1 Genome instability is a major contributing factor in human cancers All eukaryotes must ensure faithful replication and stable transmission of their genomes during cell division. Genome instability can have dire consequences for regulation, cell growth and viability, and can contribute to a variety of complex phenotypes in multicellular organisms, including cancer.  In colon cancer, for  example, genome instability commonly manifests itself as either a mutator phenotype, characterized by an increased mutation rate (Leach et al., 1993), or as chromosomal instability (CIN) characterized by an increased rate of gain, loss or rearrangement of whole chromosomes (Lengauer et al., 1997).  A consequence of  CIN is aneuploidy where cells no longer have a normally balanced number of chromosomes.  Increases in gene copy number in aneuploid cells have been  correlated with higher levels of transcription (Lyle et al., 2004, Torres et al, 2008), potentially leading to loss of control of cell proliferation due to overproduction of growth factors. Conversely, a CIN phenotype may lead to loss of heterozygosity (LOH), and possible loss of cell cycle checkpoint genes or genes that prevent uncontrolled cell division, including tumor suppressors. Therefore, the genetic basis and molecular mechanisms of CIN are of great interest in understanding the nature of cancer development and progression. It has been over a century since Boveri postulated that the acquisition of abnormal chromosome numbers might be a contributing factor in tumor development (Boveri, 1902, Satzinger, 2008). In fact, the majority of human tumors exhibit an abnormal karyotype resulting from chromosomal instability (Rajagopalan 	
    and Lengauer, 2004, Weaver and Cleveland, 2006). CIN is thought to arise through inactivating mutation in or aberrant expression of genes required for accurate chromosome transmission, such as the mitotic spindle assembly checkpoint that ensures that all chromosomes are properly attached to the mitotic spindle before anaphase proceeds.  For example, mutations in the mitotic spindle checkpoint  kinase BUB1B are associated with aneuploidy and tumor predisposition in patients with a recessive disease called mosaic variegated aneuploidy (Hanks et al., 2004). BUB1B has also been found to be overexpressed in breast cancer cell lines with chromosomal instability (Yuan et al., 2006).  Another spindle checkpoint gene,  MAD2, is found mutated in breast cancers (Hernando et al., 2001). MAD2 haploinsufficiency leads to CIN, aneuploidy and tumor formation in a mouse model (Michel et al., 2001). In addition to proper spindle assembly, sister chromatid cohesion (SCC) must be regulated to ensure that sister chromatids do not prematurely separate before anaphase.  PTTG1 encodes securin, an inhibitory protein that binds the  protease separase, and prevents premature cleavage of cohesin, the protein complex that mediates SCC (Zou et al., 1999). PTTG1 was originally found to be overexpressed in rat pituitary tumors; overexpression was subsequently shown to cause tumor formation in nude mice (Pei, et al., 1997). Altered PTTG1 expression is found in a variety of tumor types (Reviewed in Schvartzman et al, 2010). A recent study showed that mutation of PTTG1 results in increased chromosomal instability in vitro, suggesting that PTTG1 is a CIN gene that could also be mutated in human cancers (Mora-Santos et al., 2013).  	
    The cohesins themselves have been cited as  2	
    possible cancer CIN genes including STAG2, which was found mutated or deleted in a variety of tumors, with aneuploidy observed due to STAG2 inactivation in vitro (Solomon et al., 2011, Kim et al., 2012). Moreover, sequencing of a panel of CIN candidate genes in colon tumors has revealed enrichment for mutation in genes encoding cohesin proteins (Barber et al., 2009), providing additional evidence that CIN derived from defective maintenance of sister chromatid cohesion could contribute to tumor development. Despite a multitude of studies implying that CIN, caused by mutation or overexpression of CIN candidate genes, can contribute to cancer development and progression, the complete spectrum of cancer-associated mutations in CIN genes is unknown.  The application of next generation sequencing technologies to tumor  samples will uncover many other CIN candidate genes, but ascribing functions to these variants remains a daunting task.  Fortunately, model organisms provide  powerful experimental tool sets for both discovering and understanding the mechanisms of CIN genes and pathways, as discussed further in Section 1.3.  1.2 CIN as a potential therapeutic target in cancer therapy 	
   Mutations in CIN genes represent a unique genetic state that may sensitize tumors to treatment with certain classes of chemotherapeutic agents. An approach that has gained considerable attention in discovering novel therapeutic targets for selective killing of cancer cells harboring known CIN mutations is synthetic lethality (SL) based therapeutics (Hartwell et al., 1997, Kaelin, 2005, Soncini et al., 2012, Porcelli et al., 2012). Synthetic lethality is a genetic concept where the simultaneous mutation of two genes in the same cell results in death, while mutation of either gene  	
    alone does not affect viability (Lucchesi, 1968, Berlin et al., 1990). In tumors with known CIN mutations, targeting the protein products of genes that are SL with CIN genes mutated in cancer could provide a strategy for selective killing of tumors. One drawback to many of the range of DNA damaging agents and microtubule destabilizing drugs currently used in chemotherapy is toxicity to non-cancerous tissues such as bone marrow (Bakhoum and Compton, 2012). Large scale siRNA screens for genes that are SL with commonly mutated CIN genes may prove fruitful in identifying novel drug targets for SL-based chemotherapy which have fewer detrimental effects on normal cells and tissues. One example of an SL-based chemotherapeutic approach is the application of poly (ADP-ribose) polymerase (PARP) inhibitors to specific killing of cancer cells carrying mutations in the tumor suppressor gene BRCA1, which itself is a CIN gene (Stolz et al., 2010).  siRNA depletion or chemical inhibition of PARP in BRCA1  deficient cells results in lethality (Bryant et al., 2005, Farmer et al., 2005). Clinical trials are currently underway to assess the efficacy of PARP inhibition for treating breast cancer, with mixed results to date (Audeh et al., 2010, Tutt et al., 2010). PARP inhibitors may also prove effective in treatment of cancers harboring mutations in other CIN genes, for example, cohesin (O’Neil et al., 2012).  The  development of next-generation sequencing technology that could be applied in a clinical setting, combined with expanded knowledge of the spectrum of SL genetic interactions for CIN genes, holds promise for developing tumor-specific treatments.  	
    1.3 Budding yeast is an excellent model organism for discovering and characterizing CIN genes The model eukaryote Saccharomyces cerevisiae is an excellent experimental system for discovering and characterizing mutations that lead to chromosomal instability. Over the past four decades, genetic screening of yeast has proved fruitful in identifying genes that are involved in cell cycle regulation (Hartwell et al, 1973) and maintenance of minichromosomes (Maine et al., 1984), data sets that were subsequently found to include CIN genes conserved in humans. An early assay for chromosomal instability in yeast exploited the endogenous mating type MAT locus on chromosome III to monitor chromosome loss (Haber, 1974, Liras et al., 1978).	
   Diploid MATa/MATα	
   yeast do not mate unless either the MATa	
   or MATα	
   locus on chromosome III is lost or homozygosed in a rare mitotic segregant, which uncovers expression of the haploid specific mating pathway genes. Therefore, the rate of loss of heterozygosity (LOH) at the MAT locus in diploid mutants strains can be monitored by determining the frequency of cells able to mate with haploid strains, in what is known as the bimater (BiM) assay. In another assay, CIN in haploid MATα	
   strains can be monitored through detection of rare mitotic segregants able to mate with other MATα	
  strains (an a-like faker phenotype), an event that only occurs when the MATα	
   locus is lost (via chromosome loss or terminal deletion) and the haploid yeast revert to the default MATa mating type and are rescued by mating (Strathern et al., 1981, Yuen et al., 2007).	
    A third assay for chromosome loss is the chromosome transmission fidelity  (ctf) assay (Spencer et al., 1990), which is a visual assay for chromosome loss.  	
    Yeasts harboring an ade2-101 mutation accumulate a red pigment, giving red colonies.  The ctf assay monitors chromosome loss by introducing an artificial  chromosome fragment containing a suppressor tRNA that restores the normal white colony color; red colony sectoring in cells grown on media containing minimal adenine is a quantitative indicator of chromosome loss in the ctf assay.  In the  original ctf screen, Spencer et al. used ethyl methanesulfonate mutagenesis of yeast and isolated 136 ctf mutants in ~50 genes, which were eventually identified as genes involved in sister chromatid cohesion, DNA replication and kinetochore function. In addition to testing mutants for whole chromosome loss and LOH, yeast can also be used to screen for CIN mutants with increased rates of gross chromosomal rearrangements (GCR). Yeasts with a functional CAN1 gene, which codes for an arginine permease, are sensitive to the toxic arginine analog, canavanine (Walker, 1955, Whelan et al., 1979). Wild type yeast are also sensitive to 5-fluoroorotic acid which is converted to the toxic 5-fluorouracil by the URA3 gene product (Boeke et al., 1987). The Myung laboratory generated an assay for GCR by inserting a URA3 gene adjacent to the CAN1 gene approximately 22 kb from the end of the left arm of chromosome V, in a ura3Δ genetic background (Smith et al., 2004, Motegi and Myung, 2007).  By scoring for simultaneous loss of both markers, one can  determine a rate of GCR in mutants of interest by quantifying yeast growth on media containing 5-FOA (loss of URA3) and canavanine (loss of CAN1), indicators of GCR at chromosome V.  	
    The construction of the non-essential yeast gene deletion collection (Winzeler et al., 1999) was a great advance for expanding the known yeast CIN genes. Yuen et al. screened ~4700 yeast deletion strains for CIN using the BiM, ALF and ctf assays, identifying 293 null mutants with a CIN phenotype (Yuen et al., 2007). Not unexpectedly, the genes identified in this screen were enriched for functions in DNA replication and repair, and chromosome segregation, including yeast orthologs of cancer-related genes such as MAD1, MAD2, and BUB1. BLAST analysis of these 293 CIN genes revealed that 103 of them were highly conserved in higher eukaryotes and could be factors in CIN development in cancer. The non-essential deletion collection has also been screened for GCR, revealing 10 novel CIN genes and validating previously known mutants with increased rates of GCR (Smith et al., 2004). In order to gain a complete picture of the yeast CIN mutational spectrum, screens for genome instability had to be extended to the essential genes. Temperature sensitive (ts) mutants are commonly used for studying essential gene function, as protein function can effectively be shut off through a shift to a higher temperature. However, ts mutants can also be used to determine the effects of partial loss of function, by growing strains at a semi-permissive temperature. By using a mutagenic PCR technique, our laboratory systematically generated ts or hypomorphic alleles for ~500 genes for which no ts mutant previously existed and screened these novel mutants for a CIN phenotype (Ben-Aroya et al., 2008, BenAroya et al., 2010b, Stirling et al., 2011). The screen for essential CIN genes by Stirling et al., included these new ts mutants and a set of ‘decreased abundance  	
    through mRNA perturbation’ (DAmP) alleles for essential genes (Yan et al., 2008, Breslow et al., 2008). The latter have reduced protein levels through disruption of the 3’ UTR. A recently assembled collection of ts mutants from the yeast research community, representing ~500 genes (Li et al., 2011) was also screened for CIN, bringing the total essential gene mutant alleles assayed to ~2000, covering 95% of essential yeast genes. In total, 257 essential genes were identified as mutable to a CIN phenotype and when screens for LOH in yeast (Anderson et al., 2008) and gross chromosomal rearrangement (Smith et al., 2004, Kanellis et al., 2007) are considered, a total of 692 yeast ORFs have been identified as mutable to CIN. Overall, 485 of these yeast CIN genes were found to display sequence homology with human genes (Stirling et al., 2011), and represent potential cancer CIN genes. Gene ontology (GO) term analysis of these essential CIN genes revealed enrichment for some unexpected biological pathways such as lipid synthesis, secretion, nuclear transport, transcription and RNA processing.  Notable among  these highly conserved CIN genes are several members of the mRNA processing machinery that are discussed further in Chapter 2.  1.4 The fidelity of mRNA processing is essential for maintaining genomic integrity Screening for essential genes mutable to a CIN phenotype in yeast revealed multiple genes involved in splicing and mRNA processing (Stirling et al., 2011). The biogenesis of mRNAs involves several post-transcriptional processing steps including 5’ capping, splicing, cleavage and polyadenylation, and finally export to the cytoplasm for translation. Despite the fact that only 283 yeast genes contain introns  	
    and the majority of introns are deleted without adverse effects on cell growth or viability (Parenteau et al., 2008), mRNA processing appears to be crucial for maintaining genomic integrity. Since essentially every mammalian gene contains introns, splicing is likely to be crucial for genomic stability in humans as well. Obviously, defects in splicing may result in indirect effects on genome stability through incorrectly spliced protein products but there is a growing body of work suggesting that malfunction of splicing machinery can lead directly to DNA damage. Pioneering studies by Li and Manley demonstrate that a mammalian splicing factor, ASF/SF2 which is part of the serine and arginine rich (SR) protein family of RNA binding proteins (Valcárcel and Green, 1996), directly affects genome stability. Knockdown of ASF/SF2 in DT40 chicken cells results in loss of genomic integrity, cell cycle arrest and eventually cell death (Li and Manley, 2005). Furthermore, Li and Manley demonstrated that depletion of ASF/SF2 results in the accumulation of R-loops, RNA:DNA hybrids that result from stalled transcription machinery. The authors proposed that unresolved R-loops promote DNA damage by leaving singlestranded DNA exposed to chemical insult and oxidative damage (Li and Manley, 2005b). Further work on the ASF/SF2 splicing factor revealed that overexpression of the RNA binding protein RNPS1 suppresses many of the phenotypes of ASF/SF2 depletion (Li et al., 2007) leading to a proposed mechanism that ASF/SF2 functions to  bind  mRNA  precursors,  thus  preventing  R-loop  formation,  and  that  overexpression of RNPS1 provides an RNA binding protein to compensate for loss of ASF/SF2. A further example of an SR protein with a role in cancer development is SC35, which is overexpressed in HPV-positive cervical cancers (Mole et al.,  	
    2009). Depletion of SC35 in mouse embryonic fibroblasts (MEFs) results in the accumulation of double strand breaks and a p53-dependent G2/M arrest (Xiao et al., 2007), raising the possibility that CIN is the cancer-relevant phenotype in tumors overexpressing SC35. SFPQ/PSF is a pre-mRNA splicing factor (Patton et al., 1993) which may contribute directly to genome integrity through dual roles in mRNA processing and DNA repair. SFPQ/PSF physically interacts with the RAD51D protein, a mammalian homologue of the yeast RAD51 protein that is involved in homologous recombination (HR) mediated DNA repair (Morozumi et al., 2009, Rajesh et al., 2011). In a recent study, it was shown that knockdown of SFPQ/PSF in MEFs results in increased sensitivity to DNA damaging agents and reduced HR-mediated DNA repair (Changanamkandath et al., 2010).  Moreover, SFPQ depleted cells  showed defects in sister chromatid cohesion and a 5-fold increase in spontaneous chromosomal aberrations. SFPQ-depleted cells also exhibit synthetic lethality with RAD51D; that is, cells lacking both SPQF and RAD51D are inviable. Since SFPQ also physically interacts with NONO, a protein involved in non-homologous end joining (NHEJ) DNA repair (Yang et al., 1993), the authors suggested that the SFPQ-RAD51D synthetic lethality stems from a potential role for SFPQ in both HR and NHEJ mediated DNA repair pathways (Changanamkandath et al., 2010). A splicing factor commonly mutated in human cancers is SF3B1, which is a core subunit of the mammalian RNA splicing machinery in the U2-snRNP component of the spliceosome (Klimek, 2011).  Genome sequencing of chronic  lymphocytic leukemia (CLL) patients has shown that SF3B1 is mutated in up to 15%  	
    of cases (Wang, 2011). Mutated SF3B1 has been reported in breast carcinomas, melanoma and pancreatic cancer through data in the Catalogue of Somatic Mutations in Cancer (COSMIC) database (Forbes et al., 2012).  In an effort to  explain why mutation of SF3B1 contributes to CLL development, Quesada et al. screened for truncated transcripts of possible SF3B1 targets in SF3B1 mutants and identified increased levels of C-terminally truncated, FOXP1, a transcription factor with altered expression in B- cell lymphoma (Brown et al., 2008), possibly linking defective FOX1P1 splicing to CLL. Since FOXP1 is overexpressed or deleted in a variety of tumor types, some research groups have suggested that it may function as either a tumor suppressor or an oncogene (Koon et al., 2007). Aberrant splicing by SF3B1 could result in defective FOXP1 function. While SF3B1 mutation has not been correlated with a CIN phenotype, this provides an example of how the indirect effects of splicing defects may still contribute to oncogenesis. A potential contributing factor to chromosome instability is aberrant splicing of factors required for accurate chromosome segregation. Shugosin (Sgo) is a conserved protein involved in maintenance of centromeric cohesion during mitosis. Overexpression of truncated Sgo variants that could result from incorrect splicing has been shown to result in CIN in HEK293 cells (Suzuki et al., 2006) raising the possibility that aberrant regulation of Sgo splicing and expression may be a contributing factor in some human cancers.  Another possible example of CIN  indirectly resulting from splicing defects is MAD1β, an	
   isoform of the MAD1 protein that functions in the spindle assembly checkpoint to delay the onset of anaphase until all kinetochores have stable bipolar connections (Nezi and Musacchio., 2009).  	
    Expression of MAD1β,	
   which lacks exon 4, results in mitotic checkpoint impairment with chromosome bridge formation and aberrant chromosome numbers (Sze et al., 2008). Nascent mRNA cleavage and polyadenylation is also of interest in human disease and it has been shown that cancer cell lines display altered patterns in the 3’-UTR of ~840 mRNAs (Singh et al., 2009). Cleavage and polyadenylation of premRNAs requires recognition and binding of the cleavage site in the mRNA by a conserved complex of proteins known as the CPSF in humans and CPF in yeast, followed by mRNA cleavage and recruitment of the poly-A polymerase which adds poly-adenosine monophosphate tails to the mRNAs in preparation for nuclear export (Chan et al., 2011).  Studies on global changes in mRNA 3’-UTR processing  revealed over 800 genes with alternative mRNA processing in B-cell lymphoma cells lines. These changes affected expression levels of some proteins including UBE2A, a ubiquitin conjugating enzyme involved in UV induced DNA repair (Singh et al., 2009). In the same study, the authors note that cell lines with this aberrant mRNA processing also exhibited significant changes in expression of mRNA cleavage factors CSTF3 and PCF11, which the authors suggest may contribute to the observed global alteration of polyadenylation in this cancer type (Singh et al., 2009). In addition to mRNA cleavage, overexpression of poly-A polymerase has also been linked to human cancers including breast and colon cancers (Danckwardt et al., 2008, Pendurthi et al., 1994) and the polyadenylation factor CstF-50 has been found to interact with a BRCA1-associated protein, BARD1, that inhibits polyadenylation, possibly blocking mRNA processing at sites of DNA damage (Kleiman et al., 1999).  	
    There is strong evidence to suggest that splicing factors and the mRNA cleavage and polyadenylation machinery physically interact to co-ordinate proper mRNA processing (Lutz et al., 1996, Awasthi et al., 2003, Kyburz et al., 2006). Since genomic integrity and regulation of cell growth are compromised in the absence of normal mRNA processing in cancer cell lines, understanding the mechanism of CIN in yeast mRNA processing mutants may contribute to our knowledge of the role that mRNA processing plays in cancer cell biology.  1.5 R-loop formation compromises DNA integrity through spontaneous DNA damage The formation and retention of RNA:DNA hybrids known as R-loops can also have disastrous consequences for genomic integrity. The association of the RNA splicing machinery with the elongating RNA polymerase is critical for mRNA biogenesis since splicing is required for export of mRNAs (Dreyfuss et al., 1997). In their 2002 paper, Jimeno et al. described the characterization of the yeast THO complex, which includes Tho2, Hpr, Mft1 and Thp2 (Jimeno et al., 2002). They demonstrated that deletion of any member of the THO complex results in transcriptional  defects  and  hyper-recombination  between  direct  repeats.  Overexpression of the Sub2 RNA helicase suppressed the transcriptional defects of THO mutants, possibly due to the fact that Sub2 is also an mRNA export factor which promotes exit of the mRNA from the transcription complex.  The THO  complex was also linked to increased UV sensitivity when the so-called “global genome repair” genes RAD7 and RAD16 were mutated (Gonzalez-Barrera et al., 2002). A proposed mechanism for the observed UV sensitivity is that mRNA  	
    elongation is compromised in tho mutants and the stalled RNA polymerase recruits the transcription-coupled nucleotide excision repair machinery (Hanawalt and Spivek, 2008) erroneously, thus stalling repair of actual UV induced lesions. Additionally, in 2003, the group of Andres Aguilera suggested that the THO complex was required to prevent the formation of extended RNA:DNA hybrids now known as R-loops. (Huertas et al., 2003), since RNase H overexpression suppressed the hyper-recombination phenotype associated with tho mutants. In addition to R-loop formation, one possible consequence of stalled transcription is the collision between RNA polymerase II and the replication fork. The short cell cycle of Escherichia coli in rich media means that the cells are continuously replicating DNA and therefore require a mechanism to prevent stalling of replication forks at actively transcribed genes. DinG is an E. coli helicase that removes R-loop structures in conjunction with an RNase (Moser et al., 1997) to prevent the blockage of DNA replication forks (Voloshin et al., 2007). Overexpression of RNase H can suppress dinG mutant phenotypes and R-loop formation while deletion of the Rnase H encoding rnh gene in dinG mutants severely affects cell growth (Boubakri et al., 2010, Voloshin et al., 2007). DinG appears to function with Mfd which acts to remove stalled RNA polymerases, by moving behind the RNA polymerase and causing RNAPII dissociation upon contact (Park et al., 2002). DinG is conserved as Rad3 in budding yeast and XPD in humans but these homologues have been shown to function in nucleotide excision repair (Boubakri et al., 2010). In their 2012 paper, Alzu et al. showed that the yeast Sentaxin RNA/DNA helicase Sen1 associates with the replication fork and enables DNA replication  	
    through transcribed regions. Mutation of SEN1 results in accumulation of RNA:DNA hybrids and replication fork instability at transcribed regions in response to treatment with the DNA damaging agent hydroxyurea (Alzu et al., 2012).  Therefore, the  authors suggested that Sentaxin is an important factor in preventing genome instability due to collision between transcription and DNA replication in eukaryotic cells.  This result has clinical relevance as sentaxin is found mutated in the  neurodegenerative diseases ataxia with oculomotor apraxia type 2 and amyotrophic lateral sclerosis 4 (Alzu et al., 2012).  1.6 Mutation of eukaryotic transcription machinery results in CIN and DNA damage sensitivity Eukaryotic RNA polymerase II (RNAPII) contains 12 subunits with 10 found in the structures solved using X-ray crystallography (Cramer et al., 2001, Westover et al., 2004). Of these 10 subunits, genes encoding for Rpo21, Rpb3, Rpb4, Rpb5 and Rpb7 were identified as mutable to a CIN phenotype in yeast (Stirling et al., 2011). One possibility is that the CIN phenotype is due to reduced transcription of other genes involved in maintenance of genomic stability. Alternatively, RNA polymerase may have a direct role in protecting the genome. For example, RNA polymerase turnover is also known to be important in response to DNA damage by UV radiation (Chen et al., 2007). Normally, the largest RNAPII subunit Rpb1 is ubiquitinated and degraded by the 26S proteasome in response to UV radiation but deletion of the non-essential RNAPII subunit Rpb9 results in retention of Rpb1 (Chen et al., 2007). This links an RNA polymerase II subunit to transcription-coupled DNA repair as it is  	
    thought that this turnover eliminates stalled RNAPII complexes at UV induced DNA lesions. Transcription and elongation factors also play roles in maintaining genomic integrity. Deletion of the elongation factor Spt2 results in an approximately 7-fold increase in recombination rates in a Rad51-dependent fashion (Nourani et al., 2006). Spt2 is related to mammalian HMG proteins involved in chromatin dynamics and is also mutable to a CIN phenotype. In addition many chromatin remodeling factors were found to be mutated in various CIN screens (Stirling et al., 2011). While it is clear that proper function and regulation of RNA polymerase is essential for both global transcription and UV-induced NER, one aspect of RNAPII biology that is poorly understood is how RNA polymerase is assembled and transported into the nucleus. RNA polymerase II is assembled in the cytoplasm from three protein subcomplexes and subsequently imported into the nucleus (Cramer et al., 2012). A useful tool for studying RNA polymerase II biogenesis is alpha-amanitin, a toxic cyclic peptide from Amanita species of mushrooms that has been shown to trigger degradation of Rpb1 (Nguyen et al., 1996), and is therefore a potent inhibitor of transcription. Alpha-amanitin treatment of mammalian cells has proven useful for understanding the intermediates of RNAPII assembly as it interferes with normal RNA polymerase biogenesis (Boulon et al., 2010). Studies on alpha-amanitin-treated cells have revealed that a network of chaperones including HSP90, RPAP3 and the R2TP/prefoldin complex regulate RNA polymerase II assembly (Boulon et al., 2010).  	
    Quantitative mass spectrometry using stable isotope labeling of amino acids in cell culture (Ong et al., 2002) revealed interactions between RNAPII assembly intermediates and a family of conserved small GTPases, known as GPN1, GPN2 and GPN3. These GPN family proteins were found to be required for proper nuclear localization of RNAPII subunits (Boulon et al. 2010, Carre et al. 2011, Calera et al. 2011) but their mechanism of action remains unclear. Notably, the yeast orthologs of the GPN family are also mutable to a CIN phenotype (Ben-Aroya et al., 2008, Alonso et al., 2011). The cellular role of the yeast orthologs of the GPN family is discussed in detail in Chapter 3.  1.7 Research scope of this thesis 	
   The work in this thesis aims to understand the mechanisms of chromosomal instability for a group of essential yeast genes involved in RNA processing and RNA polymerase assembly, many of which are poorly characterized. This work not only contributes to our understanding of genomic instability but also highlights some novel and conserved functions of a previously unrecognized class of CIN genes, including genes involved in RNA polymerase II assembly. In Chapter 2, we tested the hypothesis that a proportion of essential CIN mutants spontaneously acquire DNA double strand breaks and identified several mRNA processing mutants with high incidence of Rad52 foci (Lisby et al., 2001). This effort identified an R-loop-dependent mechanism of CIN and DNA damage related to collisions between the transcription machinery and the DNA replication fork.  	
    Chapter 3 explores the conserved function of the GPN family of small GTPases in the biogenesis and nuclear import of RNA polymerase II and also identifies a novel role for these proteins in the biogenesis of RNA polymerase III. The functional relationship between the individual GPN proteins and the genomic instability phenotype of GPN mutants is also explored.  As eukaryotic RNA  polymerase biogenesis is poorly understood, this research represents a novel contribution to this conserved and essential process.  	
    Chapter 2. Defects in mRNA processing promote a loss of genomic integrity in Saccharomyces cerevisiae 2.1 Background information 	
   Yeast genome-wide screens for CIN phenotypes have culminated in an extensive compilation of nearly 700 CIN genes across an array of cellular processes (Stirling et al., 2011). This list provides a useful resource for the study of candidate CIN genes in humans and also reveals many cellular components for which the mechanism of genome instability is obscure (Andersen et al., 2008, Kanellis et al., 2007, Smith et al., 2004, Stirling et al., 2011, Yuen et al., 2007). A widely recognized mechanism leading to CIN is excessive DNA damage or defective DNA repair (Jackson and Bartek 2009). DNA damage occurs primarily due to the perils of normal DNA replication, where it tends to occur at fragile sites marked by H2A phosphorylation in yeast or H2AX phosphorylation in mammalian cells (i.e. mapped recently in Szilard et al., 2010). However, DNA damage can also occur as a result of a host of environmental factors, mutations and intracellular stresses. This damage is repaired effectively by extensive DNA repair machinery (Jackson and Bartek 2009). Many diverse pathways seem to converge on DNA damage and it may be that a significant number of CIN mutations act through inappropriate DNA damage or repair (Alvaro et al., 2007, Jackson and Bartek 2009). One more recently appreciated group of CIN genes, associated with increases in DNA damage, mutation and hyper-recombination, is a subset of genes involved in transcription and RNA processing. Mutations in topoisomerase I, Sen1/SENATAXIN, THO/TREX, and the SR protein splicing factor ASF/SF2 have each been linked to genome instability via a common mechanism (El Hage et al., 	
    2010, Gomez-Gonzalez et al., 2009, Li and Manley 2005, Mischo et al., 2011). These mutants induce the formation of persistent, transcription-associated RNA:DNA hybrids which form R-loops. The R-loop structures expose damage-prone single-stranded DNA (ssDNA) on the nonsense strand and may act as a block for replication fork progression, consistent with observations that mammalian transcription-associated recombination requires DNA replication and that R-loop mediated genome instability in E. coli is caused by replication fork collisions (Gan et al., 2011, Gomez-Gonzalez et al., 2009, Gottipati et al., 2008, Prado and Aguilera 2005). At present, the extent of cellular processes that contribute to R-loop-based genome instability is unclear. In an effort to extend previous studies on drug sensitivities of yeast mutants for non-essential genes (Giaever et al., 2002), a chemical genomic screen of temperature-sensitive and DAmP alleles of essential genes was conducted. Screening known genome destabilizing chemicals enriched for CIN mutants as well as mutants with roles in mRNA processing and transcription. To identify CIN processes that increase cellular demands on the DNA repair/recombination machinery, we performed a visual screen for Rad52-marked recombination centers in mutants of 305 essential CIN genes. This represents direct tests of mutants in more than 25% of essential genes. Remarkably, of 44 strains with increased Rad52 foci, we identified seven subunits of the mRNA cleavage and polyadenylation (mCP) machinery. These mCP proteins have been implicated in transcription elongation, termination and mRNA export due to their role in RNA processing (Brodsky and Silver 2000, Luna et al., 2005, Tous et al., 2011). Chromatin immunoprecipitation  	
    and microarray (ChIP-chip) using phosphorylation of H2A as a marker of DNA damage revealed fragile site differences between mCP mutants and wildtype (WT) that map to a set of transcribed ORFs linked to replication origins, supporting a transcription-dependent mechanism for DNA damage.  2.2 Materials and methods 2.2.1 Yeast strains and plasmids 	
   Yeast strains were grown in YPD or synthetic complete media lacking the appropriate amino acid where nutritional selection was required. Serial dilution assays and growth curve analysis were performed as described (McLellan et al., 2009, Stirling et al., 2011). See Table A1 in the appendix for a list of yeast strains and plasmids. 2.2.2 Chemical genomic screening 	
   For chemical screening, arrays of yeast mutants were pinned in triplicate onto YPD containing either 0.01% MMS, 50 mM HU, 10 µg/mL Benomyl or 1 ng/mL rapamycin. After 24 hours growth, each plate was again pinned onto chemicalcontaining media in triplicate leading to nine total replicates passaged on YPD + chemical. Plate images were collected after another 24 hours growth on a flatbed scanner. Image analysis and scoring for both SGA and chemical sensitivities was performed as described (McLellan et al., 2012; Stirling et al., 2011). Chemicals were selected based on diverse mechanisms representing genotoxic or non-genotoxic anti-cancer strategies (Table 2.1).  	
    2.2.3 Rad52-YFP foci screening and CIN assays 	
   Rad52-YFP fusions were made by direct transformation and introduced into the relevant strains by SGA (Tong et al., 2004). Log-phase cultures expressing Rad52-YFP were shifted from 25°C to 37°C for 3.5 hours and then prepared for microscopy as described (Carroll et al., 2009). Both DIC and fluorescence images were collected with Metamorph (Molecular Devices) and analyzed using ImageJ (http://rsbweb.nih.gov/ij/index.html). In Fig. 2.8, cells growing at 25°C were pretreated with 100 µg/mL of 6AU for 20 minutes before shifting to 37°C for 3.5 hours and imaged as for untreated cells. For each putative hit, triplicate experiments were performed and ≥100 cells were counted per replicate. Budding indices were derived from these images. Chromosome instability assays were performed as described (Spencer et al., 1990, Yuen et al., 2007). 2.2.4 ChIP-chip 	
   ChIP-chip experiments were performed as described (Schulze et al., 2009) using a ChIP-grade anti-phospho-Ser129 H2A (ab15083; Abcam), except ts-alleles were inactivated by shifting to 37°C for 3 hours before cross-linking. Data analysis and plotting of enriched features was performed using custom R, MatLab and python scripts essentially as described (Schulze et al., 2009, Takahashi et al., 2011). All profiles were generated in duplicate except the h2a-S129A control. The enrichment threshold was 2.5 fold for all analyses. Features were considered enriched if 25% of the feature length (bp) met the enrichment threshold when normalized to h2a-s129A and if 10% of the feature met the enrichment criteria for mutants normalized to WT.  	
    2.2.5 ChIP-chip statistical analysis 	
   To determine whether the data recovered more of a particular feature than would be predicted at random a Monte Carlo simulation on the relevant dataset was run using five randomly generated start positions for a particular set of features (e.g. ARSs), while maintaining the total number of features within a chromosome (Schulze et al., 2009). 500 simulations were run for each feature to generate a mean and standard deviation. These values were compared to the observed score using the cumulative normal distribution, which calculates the probability of seeing a lower score if a value was selected at random (www.stattrek.com). Transcriptional frequency categories and Rad52 foci counts were compared to controls using a Student’s t-test (http://www.graphpad.com). Overall transcriptional frequencies for smaller samples were compared using the Mann-Whitney test. Significance levels are indicated in the main text or figure legends. 2.2.6 Yeast chromosome spreads 	
   Chromosome spreads were done essentially as described except that cells were shifted to 37°C for 4-6 hours before lysis (Hartman et al. 2000). The S9.6 antibody (Hu et al. 2006) was used at 1µg/mL in blocking buffer (1x PBS with 5% BSA and 0.2% skim milk powder) and detected with Cy3-conjugated goat antimouse antibody (Jackson Labs, #115-165-003). 2.2.7 Growth curve analysis Growth curve data was collected using a TECAN M200 pro plate reader as described (McLellan et al. 2009). The areas under three replicate growth curves were calculated and converted to a fitness score expressed as a percentage of WT 	
    fitness. Predicted fitness of double mutants was calculated by multiplying the two single mutant fitness scores. Double mutant fitness scores that were observed as >10% less than predicted were scored as synthetic sick (SS).  2.3 Results 2.3.1 Selective killing of CIN mutants by genome-destabilizing chemicals 	
   Recent work identifying and evaluating gene-drug interactions in cancer has revealed that compounds targeting a specific genotype typically yield better selective killing than drugs having a more pleiotropic cytotoxic effect (Barretina et al., 2012; Garnett et al., 2012). In this study we screened the essential gene mutants for drug sensitivity in order to understand the suite of genotypes that is targeted by lessselective therapeutic strategies whose effects challenge entire CIN pathways, in this case DNA replication, repair and mitosis. We generated a comprehensive view of gene-drug interactions for four distinct classes of cytotoxic chemical, MMS, HU, benomyl and, as a non-genotoxic control, rapamycin (Table 2.1). Since these data are available already for all non-essential gene deletions, we tested only DAmP and ts alleles in essential genes; 1945 alleles were tested representing ~90% of essential yeast genes (Ben-Aroya et al., 2008; Breslow et al., 2008; Li et al., 2011). Adding these data for essential genes is particularly relevant to CIN because nearly half of reported CIN genes are essential (Stirling et al., 2011).  	
    Table 2.1 Rationale for chemicals used in genome-wide analysis. Chemical Methyl methanesulfonate  Mode of Action Alkylating agent; directly damages DNA bases  Analogues in chemotherapy Nitrogen-mustard based (e.g. ifosfamide, chlorambucil); Other (e.g. temozolomide)  Hydroxyurea  Inhibitor of ribonucleotide reductase; causes stalled replication forks due to reduced nucleotide pool  Hydroxyurea (i.e. marketed as Droxia, Hydrea)  Benomyl  Binds tubulin heterodimers preventing microtubule assembly  Vinblastine, Vincristine, Vinorelbine, Vindesine, Paclitaxel  Rapamycin  Binds FK506-binding protein to inhibit TORC1 signaling  Sirolimus and derivatives (e.g. Everolimus (Afinitor); Temsirolimus (Torisel))  Growth of the essential gene mutant strains in the presence of chemicals was measured in high-throughput array format (see Materials and Methods). 123, 122, 47 and 33 genes met our stringent cut-off for a negative chemical-genetic interaction with HU, benomyl, MMS and rapamycin (van Pel et al., 2013) Notably, multiple RNA biogenesis factors were found to exhibit drug sensitivities, including multiple splicing factors and transcription associated genes (Table 2.2).  	
    Table 2.2 RNA biogenesis genes with mutant alleles that exhibit enhanced drug sensitivity to hydroyurea, MMS and benomyl.  	
    Gene  Function  Drug  CLF1  Splicing  Hydroxyurea, benomyl  DBP9  rRNA processing  Hydroxyurea  FAF1  rRNA processing  Hydroxyurea  LSM8  mRNA processing  Hydroxyurea  MED4  transcription initiation  Hydroxyurea  NOP14  rRNA processing  Hydroxyurea  POP6  tRNA, rRNA, mRNA processing  Hydroxyurea  POP7  tRNA, rRNA, mRNA processing  Hydroxyurea  PRP2  Splicing  Hydroxyurea  PRP39  Splicing  Hydroxyurea  PRP40  Splicing  Hydroxyurea  RBA50  mRNA transcription  Hydroxyurea  RCL1  rRNA processing  Hydroxyurea  RMP1  RNA processing  Hydroxyurea  RPB4  RNA polymerase  Hydroxyurea  RPB9  RNA polymerase  Hydroxyurea  RRN11  rRNA transcription  Hydroxyurea  RRN6  rRNA trnscription  Hydroxyurea  SEN1  mRNA processing  Hydroxyurea  SNM1  mRNA and rRNA processing  Hydroxyurea  PRP11  Splicing  Benomyl  PRP16  Splicing  Benomyl  PRP22  Splicing  Benomyl  PRP43  Splicing  Benomyl  RPC37  RNA pol III subunit  Benomyl  SAD1  Splicing  Benomyl  SAS10  rRNA processing  Benomyl  SNU114  Splicing  Benomyl, MMS  YSH1  mRNA processing  Benomyl  YHC1  Splicing  Benomyl  IFH1  rRNA transcription  MMS  PRP40  Splicing  MMS  ROX3  mRNA transcription  MMS  SPT15  transcription factor  MMS  TAF6  transcription initiation  MMS  26	
    Grouping the essential genes sensitized to each chemical by gene ontology allowed us to build a network of pathways affected by the four chemicals (Figure 2.1). Many expected interactions emerged: for example, MMS and HU impacted DNA replication and repair, whereas benomyl was strongly associated with mitotic spindle defects (Figure 2.1). However, analysis of the essential genes also provided several new insights: for example, splicing mutants were highly sensitized to benomyl. This is potentially due to the presence of an intron in the TUB1 gene which, if improperly spliced, would lead to a toxic excess of β tubulin relative to α tubulin (Biggins et al., 2001; Burns et al., 2002).  Figure 2.1 Gene ontology (GO) derived cellular functional groups uncovered by chemical screening of essential gene mutants. The network represents the raw data found in van Pel et al 2013. Chemicals are indicated with blue nodes. Red nodes represent functional groups where node size indicates the number of genes in that group and edge thickness indicates the number of connections to a particular chemical.  When we enumerated genes that are sensitive to one of the four test chemicals and also display a CIN phenotype, we saw clear enrichment of CIN genes sensitive to benomyl, MMS and HU, but no enrichment of CIN genes impacted by 	
    the non-genotoxic control rapamycin (Figure 2.2A). Indeed, benomyl-sensitive mutants exhibit a greater than 7-fold enrichment of CIN genes. This is perhaps explained by the fact that benomyl would seem most likely to cause whole chromosome loss, as it functions to disrupt microtubules, and chromosome loss is a phenotype commonly measured in detecting CIN genes (Stirling et al., 2011; Yuen et al., 2007). The majority of all reported CIN genes (i.e. 445/692, 64%) were sensitized to at least one of the genome destabilizing drugs (Figure 2.2B). Specifically focusing on the orthologs of cancer gene census genes – that is, genes believed to play a causative role in tumorigenesis (Futreal et al., 2004) – also showed that the majority of CIN genes in this subset were sensitive to one of the genome destabilizing chemicals (Figure 2.2C). Though these findings are not unexpected given the known modes of action of MMS, HU and benomyl, they do expand our knowledge base of CIN processes and/or genes that may be targeted through therapeutics and highlight the importance of transcription and mRNA processing in maintaining genomic integrity in response to chemical insult.  	
    Figure 2.2 The spectrum of genotoxic drug sensitivities of essential genes is enriched for CIN genes. (A) Compilation of new chemical sensitive genes with the literature highlights the enrichment of CIN genes within genotoxic drug sensitivity profiles. Enrichment indicates the quotient of the percentage of CIN genes in each chemical sensitivity list (van Pel et al, 2013 in Press) and the percentage of CIN genes in the entire genome. (B) Overlap of MMS, HU or Benomyl sensitive mutants with known CIN C) Number of CIN genes with orthologs on the cancer gene census that are sensitive to one of the chemicals tested.  2.3.2 Rad52 foci screening of essential CIN genes 	
   Rad52p is essential for homologous recombination and organizes into repair centers in response to double-strand breaks and other recombination events (Lisby et al., 2001, Mortensen et al. 2009). Increases in Rad52 foci can therefore indicate a number of genome destabilizing conditions including increased double-strand breaks, inefficient resolution of recombination intermediates or hyper-recombination (Alvaro et al., 2007). To determine which CIN mutants cause increased or prolonged engagement of the homologous recombination machinery, we introduced a RAD52YFP fusion under control of its native promoter by synthetic genetic array (SGA)  	
    (Tong et al., 2004). The resultant strains expressing a mutant CIN gene and RAD52YFP were screened visually for increased levels of Rad52 foci (see Materials and Methods). We screened 360 alleles of 305 essential CIN genes, including 306 tsand 54 DAmP alleles (Stirling et al., 2011). Similar to previous studies we retested mutants where ≥15% of cells had Rad52 foci in the primary screen (Alvaro et al., 2007). Triplicate retesting produced a list of 46 alleles in 44 unique genes whose mutation elicits an increased level of Rad52 foci (Fig. 2.3). Eighteen additional mutants had increased but variable levels of Rad52 foci that did not meet our threshold across replicates (Stirling et al., 2012). The largest functional group of mutants with increased Rad52 foci affected DNA replication consistent with the Sphase function of Rad52 and its role in repairing damage caused by collapsed replication forks (Lisby et al., 2001). Interestingly, mutations in multiple genes involved in the proteasome, Smc5/6 complex, early secretion, transcription and mRNA processing all caused increased rates of Rad52 foci (Fig. 2.3). These data stress the value of screening essential gene mutant collections since a systematic screen of non-essential gene deletions uncovered a non-overlapping set of biological processes (Alvaro et al., 2007).  	
    Figure 2.3 A screen for DNA damage foci in essential CIN genes. Percentage of cells with Rad52 foci. Bars are color-coded to denote cell cycle arrest as large budded cell (black), foci formation only in budded cells (dark grey) or foci formation at all stages (light grey). Multi-member biological groups are labeled above.  Published genetic interactions exist between rad52Δ and 12 of 44 genes identified in our screen (i.e. BioGrid/DRYGIN databases). The presence of Rad52 foci in a mutant may indicate the dependence of that mutant on RAD52 function (Alvaro et al., 2007). To further examine this hypothesis, we directly tested selected mutants from different biological pathways for genetic interactions with rad52Δ. For pol31-ts, sld7-ts, rpn5-1, orc6-ts and rna15-58 there were clear growth defects in the absence of RAD52 (Fig. 2.4). For ssl1T242I and pcf11-2 a growth defect was apparent when a low level of HU was present, whereas nse3-ts and sec12-4 did not show obvious growth defects in this assay (Fig. 2.4).  	
    Figure 2.4 Genetic interactions between foci generating mutants and rad52Δ. Equal ODs of the indicated strains were serially diluted and spotted on YPD +/- 5mM HU at 30°C.  Overall our screen indicates that a large proportion of essential CIN genes (i.e. 14%, 44 of 306 genes) exhibit significant levels of mutant-induced Rad52 foci and thus that disruption of unexpected cellular pathways creates a requirement for homology-directed DNA repair. Of particular interest was the identification of seven mRNA cleavage and polyadenylation genes in the Rad52 foci screen (Fig. 2.3). The mCP machinery is essential for processing nascent RNAs to free polyadenylated species (Gross and Moore 2001). Interestingly, unlike the majority of mutants identified in the screen, which arrested as large-budded G2/M cells, most mCP mutants exhibited Rad52 foci at all cell cycle stages, including G1. In comparison to those Rad52 foci-forming mutants involved in DNA replication, the role of mCP genes in genome integrity is not well characterized and we decided to investigate it further.  	
    2.3.3 Fragile sites in mCP mutants differ in transcribed ORFs and near replication origins Phosphorylation of H2A-Ser129 is a mark of DNA damage analogous to human γ-H2AX modification in response to DNA damage and is critical for DSB repair (Downs et al., 2000). Recent analysis of sites of H2A-S129 phosphorylation in the yeast genome revealed a set of fragile loci or “γ-sites” using chromatin immunoprecipitation on microarray (ChIP-chip) (Szilard et al., 2010). It is known that h2a-S129A mutants accumulate Rad52 foci and that both Rad52 and H2A phosphorylation contribute to efficient DNA repair (Downs et al., 2000, Mortensen et al., 2009, Szilard et al., 2010). We reasoned that mapping differences in fragile sites between WT and mCP mutants could suggest a mechanism for CIN in the mutants. We performed γ-H2A ChIP-chip on the two mCP alleles with the highest levels of Rad52 foci, pcf11-2 and clp1-ts. We applied double-T7 amplified anti-phosphoSer129-H2A chromatin immunoprecipitates from log-phase WT, mCP mutant or h2a-S129A control cells to a high-density tiling microarray containing 3.2 million probes with an average 5bp resolution and 20bp overlap between probes (Schulze et al., 2009). Normalizing our data to the h2a-S129A control produced a profile of yeast γ-sites (Fig. 2.5; see also Stirling et al., 2012).  	
    Figure 2.5 High-resolution mapping of yeast γ-sites. Overview of γ-sites for Chromosome VI indicating chromosomal features (purple box = ARS, grey box = ORF). The colored traces represent duplicates of WT (red and black), and one replicate each of clp1-ts (green) and pcf11-2 (blue). Large sub-telomeric γ−site regions are noted with black bars, seven enriched ARS are noted with vertical dotted lines and three examples of repressed genes (<1 mRNA/hr) are noted with dotted lines connecting to red-boxes to define the gene boundaries.  Overall our WT ChIP-chip profiles confirmed the findings of Szilard et al., (2010), for example, identifying γ-sites enriched in subtelomeric regions, replication origins, long terminal repeats, repressed ORFs, MAT and near rDNA and centromeres (Fig. 2.5, Stirling et al., 2012 Supplemental information). Initial examination of the mutant data revealed similar γ-site peak profiles in replicates of clp1-ts, pcf11-2 and WT strains (Fig. 2.5). For example, Fig. 2.6A shows similar mutant and WT γ-sites associated with a repressed gene, HXT10, and a replication origin in the indicated 50kb segment of chromosome VI. To highlight regions of  	
    mutant-specific γ-H2A signal, which could be linked to mutant-induced DNA damage, we normalized the mutant profiles to that of the WT (Fig. 2.6B). This analysis revealed reproducible and widespread differences in γ-sites adjacent to replication origins (e.g. ARS603.5 in Fig. 2.6B).  Figure 2.6 mCP mutants are enriched for γ-sites adjacent to origins of replication (A) Representative pcf11-2 and clp1-ts γ-site profiles for a 50kB segment of ChrVI from the third row in Figure 2.5. Enriched γ-sites at HXT10, which is repressed in glucose, and ARS605 are indicated. (B) Replicates of pcf11-2 (black, green) and clp1-ts (blue, red) γ-site profiles normalized to WT to identify regions of enhanced signal common to both mutants. The same 50kB region of ChrVI from Fig. 2.5 is shown after normalization to the WT γ-site profile. Common differences are marked with a black bar.  To  quantify  the  observed  connection  between  mCP  mutant  γ-site  enhancement and ARS-linked sites we scanned a 2kb window on either side of all ARSs for enriched regions ≥500bp in length. This analysis found significantly more sites than would be expected at random for both clp1-ts and pcf11-2 mutants (i.e. P=0.99997; Fig. 2.7 compares the observed values to the predicted number of peaks if ARS start coordinates were randomized). 64 ARSs met these stringent criteria for 3 of 4 mutant replicates (i.e. the sites are found at least once in both clp1ts and pcf11-2 and found in two replicates of either clp1-ts and pcf11-2; (Fig. 2.7 and Stirling et al., 2012 supplemental Table S4).  	
    Very few ARSs were themselves  35	
    enriched in the mCP-specific γ-sites; instead the peaks appeared in ORFs in close proximity to the ARS. The peaks of 60 of the 64 ARS-linked sites could be assigned to single genes. 65% of these genes were oriented towards the ARS, similar to the genome-wide frequency of ORF-ARS collision (60%). Notably, ARS-linked γ-H2A enriched ORFs oriented in such a colliding fashion had a higher average transcriptional frequency than the complete set of colliding ORF-ARS pairs, while this trend was not seen for non-colliding ORF-ARS pairs (Fig. 2.7, right panel and Stirling et al., 2012). These observations are consistent with a mechanism contingent on R-loop formation since they implicate the collision of transcription and DNA replication as a source of DNA damage in mCP mutants.  	
    Figure 2.7 clp1-ts and pcf11-2 mutant γ-sites link DNA damage to transcription. (A) Linkage of mutant specific peaks to replication origins. Peaks within 2kb of replication origins were identified in the (mutant-WT) difference profiles and compared (left panel) to predicted values generated by Monte Carlo simulation of randomized ARS positions. * indicates p<0.0001. The right panel shows the relative orientation, with respect to ARS, of ORFs encompassed by ARS-linked γ-site differences across 3 of 4 mutant replicates (pointed end indicates the direction of transcription). The average transcriptional frequency of ARScolliding ORFs in γ-sites was significantly higher than for all ARS-colliding ORFs across the entire genome (p<0.05).  In addition to the ARS-linked γ-sites, there was mutant-specific γ-H2A enrichment at a set of 918 ORFs throughout the genome. When we examined the transcriptional frequency of these 918 ORFs we found significantly fewer of the lowest transcription category (i.e. <1 mRNA/hr) compared to all ORFs (Fig. 2.8A; Holstege et al., 1998). This represents a dramatic shift from the WT γ-sites which strongly enriched repressed or weakly transcribed genes and underrepresented higher transcriptional categories (Fig. 2.8A; Szilard et al., 2010). If we map the relative position of γ-site differences within the 918 ORFs there is a slight bias toward the 3’-end of the gene for both pcf11 and clp1-ts mutants compared to enriched ORFs in the global γ-site profile of WT cells (Appendix Figure A1). Moreover, examining the 55 genes whose 3’-UTR contains a γ-site in both clp1-ts  	
    and pcf11-2 profiles (i.e. 3 of 4 replicates covered by >50% of a 500bp window downstream), reveals that the associated genes tend to have higher transcriptional frequencies (Stirling et al., 2012). Together these data suggest that at least some transcription occurs at ORFs specifically enriched for γ-H2A in mCP mutants and that damage may be biased to the site of mCP function at 3’-ends. One potential reason that the bias is not more pronounced is that various 3’-end processing factors, including the mCP, also affect transcription elongation and could be causing damage within genes as well as downstream (Tous et al., 2011). If DNA damage in pcf11-2 and clp1-ts mutants occurs at transcribed ORFs it is plausible that reducing transcription could mitigate the increase in Rad52 foci. To test this hypothesis we treated cells with 6-azauracil (6AU), an inhibitor of transcription elongation, and scored the presence of Rad52 foci. WT cells showed an increase in Rad52 foci when treated with 6AU (Fig. 2.8B). Remarkably, both clp1ts and pcf11-2 strains showed a significant decrease in Rad52 foci when treated with 6AU (Fig. 2.8B). This effect was not seen in the strong hits srm1-ts or cdc24-11 which both retained high levels of Rad52 foci in 6AU (Fig. 2.8B). We conclude from this experiment that at least some of the DNA damage in mCP mutants is occurring within transcribed regions.  	
    Figure 2.8 DNA damage in mCP mutants is biased towards genes with higher transcriptional frequency. (A) Transcriptional frequency of ORFs with enhanced γ-H2A signal in WT cells and mCP mutant normalized to WT samples. The distribution of transcription frequencies (Holstege et al., 1998) is shown for all ORFs (grey bars), ORFs covered at least 25% by a WT γ-site (black bars) and ORFs covered at least 10% by a γ-site difference in clp1-ts and pcf11-2 ChIP-chip profiles. (B) Effect of the transcription inhibitor 6azauracil on Rad52 foci formation in WT and mutant strains. Asterisks indicate for (A) significant deviations from all ORFs within a transcription frequency category (hypergeometric test) and (B) significant differences in Rad52 foci levels (students t-test; * p<0.05, **p<0.005).  2.3.4 CIN in mRNA cleavage and polyadenylation mutants occurs via R-loops 	
   Our data show that mutants in the mCP pathway accumulate Rad52 foci and our ChIP-chip data suggest a connection between DNA replication and the role of mCP in transcription. It is known that transcription can act as a replication fork barrier (Deshpande and Newlon 1996, Takeuchi et al., 2003), and that mutations in mCP components can cause transcription defects (Birse et al., 1998, Prado and Aguilera 2005). To investigate the connection of these phenotypes to CIN, we first confirmed reported mCP phenotypes in three standard CIN assays, investigating primarily chromosome loss (Chromosome Transmission Fidelity (Ctf)), loss of heterozygosity (BiMater (BiM)) and rearrangement/gene conversion (a-like faker (ALF)). Mutants in CFT2, CLP1, FIP1, PCF11 and RNA15 had a robust Ctf phenotype and a weak ALF phenotype; RNA15, CLP1 and PCF11 mutants also had  	
    a detectable BiM phenotype (Stirling et al., 2012). Each mCP mutant was hypersensitive to the genotoxic agents hydroxyurea (HU), cisplatin, bleomycin and UV (Fig. 2.9A). These agents may synergize with the DNA damage that occurs in mCP mutants; for example, failed transcription termination in mCP mutants could act as a block to replication forks thereby sensitizing cells to replication inhibitors like HU or cisplatin (Birse et al., 1998, Prado and Aguilera 2005). Indeed, HU and certain mCP mutations both increase recombination rates; thus the effects may be additive (Luna et al., 2005). Our data and the literature support the notion that transcriptional defects underlie genome instability in mCP mutants. One common way by which this occurs is through formation of transcription-coupled RNA:DNA hybrids called R-loops. The identification  of  pcf11-2  and  clp1  ts γ–site  differences  proximal genes, combined with the suppression of Rad52 foci by 6AU  in  ARS-  (Fig.  2.8)  make this an especially attractive model . Consistent with this explanation of our CIN data, we find that known R-loop forming mutations in the helicase SEN1 and the THO components MFT1 and THP2 lead to strong CTF and ALF phenotypes (Stirling et al, 2012) To confirm that the mCP mutants accumulate R-loops, we transiently grew cells at the non-permissive temperature of 37°C and harvested them for chromosome spreads. We performed immunofluorescent detection of RNA:DNA	
   hybrids using the S9.6 monoclonal antibody in chromosome spreads of WT, mCP mutants, known or predicted R-loop forming mutants including rnh1Δrnh201Δ	
  double mutants, and a panel of Rad52 foci forming strains from other biological pathways  	
    (Figure 2.9). For each allele of the mCP machinery tested we observed an increased number of chromosome spreads with RNA:DNA hybrid staining.	
  The rnh1Δrnh201Δ	
   double mutant also showed high levels of RNA:DNA hybrids confirming the role of endogenous RNaseH is actively suppressing R-loop formation (Figure 2.9).	
   Indeed the rnh1Δrnh201Δ	
  strain exhibits a strong chromosome instability phenotype (Stirling et al., 2012).  As predicted, mutants in the THO subunits MFT1 and THP2 and in  SEN1 caused RNA:DNA hybrids (Figure 2.9; Gomez-Gonzalez et al., 2009, Mischo et al., 2011). Deletion of the transcription termination factor RTT103 also increased RNA:DNA hybrids (Figure 2.9). Among the panel of Rad52 foci-forming control strains, only srm1-ts showed an increase in RNA:DNA hybrids (Figure 2.9). SRM1 plays a role in nucleocytoplasmic transport of mRNAs which could account for accumulation of RNA:DNA hybrids in srm1-ts mutants. As an antibody specificity control we show that pre-treatment of chromosome spreads with recombinant RNaseH in vitro significantly reduces R-loop signal (Stirling et al., 2012). Together these data suggest that mCP mutants are similar to THO and SEN1 mutants in that the mechanism of genome instability requires the accumulation of RNA:DNA hybrids.  	
    Figure 2.9 Transcription coupled R-loops are the likely cause of CIN in mCP mutants. (A) 10-fold serial dilution spot assays of mCP mutant strains on indicated media. ctf4Δ is included as a sensitive control strain. An additional YPD control (right) is included for cisplatin sensitivity because these plates have a different pH. (B) Immunofluorescence of RNA:DNA hybrids1 in chromosome spreads. Representative spreads from WT and pcf11ts10 cells (blue, DNA; red, RNA:DNA hybrid; left panel) and quantification of the immunofluorescence data for the indicated mutants (right panel) are shown. “R-loop positive controls” indicate strains known or predicted to form R-loops and “Rad52 foci positive controls” indicate strains which form Rad52 foci (Fig. 1) but are not predicted to form Rloops. Significant differences from WT (p<0.05) are indicated by *. 1  R-loop staining performed by YA Chan  2.3.5 Comparative analysis of mCP mutants with known R-loop forming mutants Known R-loop forming mutants (e.g. THO complex, SEN1) work in parallel with mCP in the continuum of transcription termination, and mRNA processing and export. Since mCP mutants have defects in transcription elongation, termination, mRNA processing and export, we wanted to address their similarity to known R-loop  	
    forming mutants to probe the mechanism by which mCP-mutant-induced R-loops cause genome instability (Brodsky and Silver 2000, Luna et al., 2005, Tous et al., 2011). Mutants in THO, and the SEN1 helicase exhibit CIN phenotypes and RNA:DNA hybrid formation in our cytological assay (Stirling et al., 2012). Moreover, representative alleles in mCP (i.e. fip1-ts, clp1-ts, rna15-58 and alleles of pcf11) exhibit synthetic growth defects or lethality with deletion of the THO subunit THP2, the termination factor RTT103, or with the sen1-1 ts-allele, as assessed by tetrads and growth curve analysis (Figure 2.10). This analysis confirms that the cellular defects caused by our mCP mutants are buffered by the action of THO and SEN1 and vice versa.  	
    Figure 2.10 Genetic interactions of mCP mutants and known R-loop forming mutants. A summary of genetic interactions between mCP mutants and components of THO, SEN1 and the transcription termination machinery. Double mutants were isolated by tetrad analysis and scored as viable or synthetic lethal. Growth defects were quantified using growth curve analysis over a 24 hour period. A summary table is shown below (SS = synthetic sickness, SL= synthetic lethality). * indicates that pcf11-2 was SS with thp2Δ and pcf11-ts10 was SS with sen1-1, both were synthetic lethal with rtt103Δ. Error bars indicated three standard deviations from the observed score.  Another phenotype common to R-loop forming mutations is hyperrecombination, and a weak phenotype has been reported for certain components of mCP (Chavez et al., 2000, Luna et al., 2005, Mischo et al., 2011). To clarify these observations we subjected the mCP mutants to a well characterized direct-repeat recombination assay (Prado et al., 1997). At 30°C the mCP mutants clp1-ts, rna15-  	
    58, and pcf11-ts10 exhibited significant hyper-recombination (Figure 2.11). In this assay mutations in SEN1 or the THO complex lead to strong hyper-recombination phenotypes; however, the introduction of a transcription terminator after the first homology segment blocks hyper-recombination in THO mutants but stimulates recombination in the sen1-1 allele (Figure 2.11; (Mischo et al., 2011). Thus, while THO and SEN1 mutants both induce R-loops, this reporter system mechanistically differentiates RNA packaging defects (i.e. THO mutants) from SEN1 helicase defects. Unlike sen1-1, hyper-recombination in mCP mutants is not increased by the transcription terminator but rather it is generally reduced (Stirling et al., 2012). Thus, mCP mutants behave like THO mutants, consistent with the known interdependence of THO and mCP function in mRNA processing and export (Luna et al., 2005).  	
    Figure 2.11 mCP mutants exhibit recombination rates similar to THO mutants. Recombination rates of indicated mutants among direct repeats in the absence (plasmid LNA) or presence (plasmid LNAT) of a transcription terminator (Prado et al., 1997). Recombination substrates are schematized above the graph where the shaded region indicates homology segments between the two moieties of LEU2. Recombination rates are reported with 95% confidence intervals. The fold change from the WT control is indicated above each bar.  2.3.6 CIN mutants for GPN family members result in moderate accumulation of DNA double strand breaks Despite not meeting the stringent cutoffs for Rad52 foci counts included in the published data from Figure 2.3, temperature-sensitive and hypomorphic mutant alleles  for  two  poorly-characterized  paralogous  essential  CIN  genes  YOR262W/GPN1 and YLR243W/GPN3 were also subjected to Rad52 foci screening, because of the previously described CIN phenotype of YOR262W (Ben-  	
    Aroya et al, 2008). As shown in Figure 2.12, several alleles display elevated levels of Rad52 foci when incubated at 37oC for 3 hours. The human orthologues of the GPN proteins exhibit defects in nuclear localization of RNA polymerase II (Boulon et al, 2010, Calera et al 2011), suggesting that they play an indirect role in eukaryotic transcription. GPN2 mutants were tested for genetic interactions with mRNA processing mutants, since the literature suggested a conserved role in RNA polymerase II assembly/import. As shown in figure 2.12B, gpn2-1 mutants exhibit genetic interactions with mutants for genes involved in mRNA cleavage and polyadenylation, which provides support for a conserved role in mRNA biogenesis. These observations provide the rationale for the studies on GPN2 and GPN3 function described in Chapter 3.  	
   Figure 2.12 Spontaneous DNA damage and genetic interactions with splicing machinery in GPN mutants. A) Some mutant strains of GPN family members display significantly elevated Rad52 foci levels. Log phase cell cultures were shifted to 37oC before imaging and scored (see Methods and Materials). B) Conditional synthetic genetic interactions of gpn2-1 with mCP mutants. As shown, the gpn2-1 cft2-ts double mutant has strongly reduced fitness at 25oC while the gpn2-1 rna15-58 double mutant exhibits synthetic lethality at 30oC.  	
    2.4 Discussion 2.4.1 Chemical genomic and Rad52 foci screening of essential CIN genes 	
   Focused secondary screens are a powerful way to define specific functions within primary functional genomic data (e.g. Carroll et al., 2009, Doheny et al., 1993, Michaelis et al., 1997, Stirling et al., 2011). The chemical genomics screening presented here provides data for mutants of essential genes that expand the chemical sensitivity dataset to encompass the majority the genome.  It also  highlights the key processes involved in resistance to chemical insult, including transcription and RNA processing (Table 2.1, Figure 2.1). While it is possible that many of the drug response phenotypes observed in these mutants are due to general defects in transcription and translation, cancer therapies are often developed to target DNA repair and the mitotic spindle in tumors (Barakat et al., 2012, Capasso, 2012). Therefore, the drug sensitivities observed for these mutants may be of clinical relevance as tumors with mutations in the human orthologs of these  genes  could  theoretically  be  sensitized  to  the  specific  types  of  chemotherapeutics assessed in these screens. By direct screening of more than one-quarter of essential genes we link 44 essential CIN mutants to increased Rad52 foci formation including seven mRNA cleavage and polyadenylation factors with a previously unappreciated role in genome integrity (Fig. 2.3). Rad52 foci indicate organization of the homologous recombination machinery into a repair center in response to DNA damage (Lisby et al., 2001). Our genetic data support the idea that increased Rad52 foci often indicate a requirement for Rad52 function (Fig. 2.4; Alvaro et al., 2007).  	
    Aside from the mCP mutants that were the focus of follow-up studies, the Rad52 foci screen identified mutants with diverse functions. The highest levels of foci were seen in srm1-ts and cdc24-11 alleles (Fig. 2.4). SRM1 strongly interacts with nucleosomes and mutants cause gross chromosomal defects, although we also found evidence that srm1-ts may cause RNA:DNA hybrids potentially due to its role in RNA export (Fig. 2.9; Koerber et al.,, 2009). CDC24 encodes a guaninenucleotide exchange factor for the small GTPase Cdc42 and, while the mechanism of cdc24-11 induced Rad52 foci is unknown, a recent high-content screen identified foci of the DNA damage marker Ddc2-GFP in alleles of CDC24 (Li et al., 2011). Mutations affecting DNA replication were the class that most frequently exhibited enhanced Rad52 foci and usually arrested as large budded, G2/M cells (Fig. 2.3). DNA replication mutants were not significantly identified in the Rad52 foci screen of non-essential genes because so many of the components are essential. The same is true of essential transcription/mRNA processing, and proteasome subunits that were not identified in the literature (Alvaro et al., 2007). Conversely, Alvaro et al (2007) identified DNA repair, mitotic checkpoint and other genes not found in our survey of essential CIN mutants. Screening of both essential and non-essential gene mutants therefore provided complementary, non-overlapping datasets. Given the relatively large number of mutants screened for foci, the cutoff at 15% of nuclei with Rad52 foci removed multiple mutant strains as candidates for further investigation while providing a manageable data set. However, a suite of mutant strains for an uncharacterized pair of yeast genes known as GPN family members were screened for Rad52 foci because of the CIN phenotype of GPN2  	
    mutants (Ben-Aroya et al, 2008). Three GPN mutant strains accumulated elevated levels of Rad52 foci. Since mammalian experiments suggest that the GPN proteins function in RNA polymerase II nuclear import (Boulon et al, 2010) the genomic instability observed in GPN2 mutants could be due to adverse effects on transcription.  Further analyses of the GPN mutants are discussed in detail in  Chapter 3. 2.4.2 Profiling fragile site changes in Rad52 foci forming mRNA processing mutants By comparing fragile sites in WT and mCP mutant yeast we formulated a putative mechanism involving transcription coupled R-loops which we confirmed by direct tests (Figs. 2.3-2.9). The robustness of the WT γ−site profiles to perturbations by different mutants was somewhat surprising but similar to findings in strains lacking RRM3, a helicase which facilitates replisome processivity (Szilard et al., 2010). Despite the similarities in WT and mutant γ-sites, we were able to extract meaningful differences within transcribed ORFs adjacent to ARSs and with a slight bias toward the 3’end of the ORF (Fig. 2.6, 2.7 and 2.8). These observations are consistent with the function of mCP in transcription termination and 3’-end processing of pre-mRNAs and suggest that DNA damage may occur near the site of mCP function. Given the role of mCP in transcription we were surprised that we did not recover the most highly transcribed genes; indeed, this category was underrepresented (Fig. 2.8A). It is possible that highly transcribed genes may have mechanisms in place to protect the genome from the deleterious effects of high transcription. This idea has been supported by work in mammalian cells showing  	
    that highly expressed genes are oriented co-directionally with replication origins (Huvet et al., 2007, Tuduri et al., 2009). Based on the success of our approach, we predict that mapping changes in fragile sites (e.g. with ChIP-chip of γ-H2A or another DNA repair protein) in other CIN mutant backgrounds could be an informative method to suggest basic mechanisms. 2.4.3 Mechanism of genome instability in mCP mutants 	
   R-loops underlie genome instability in a growing number of transcription and RNA processing mutants, including SPT2, TOP1, SEN1, THO complex and ASF/SF2 (El Hage et al., 2010, Gomez-Gonzalez et al., 2009, Li and Manley 2005, Mischo et al., 2011, Sikdar et al., 2008, Tuduri et al., 2009). R-loops have been linked to various genome instability phenotypes including hyper-recombination, replication defects, DNA damage and chromosome loss. Our Rad52 screening and ChIP-chip studies suggested a mechanism for genome instability in mCP mutants that involves DNA damage induced by the formation of transcription-coupled Rloops (Fig. 2.3-2.8). The fact that Rad52 foci are suppressed by reducing transcription with 6AU (Fig. 2.8B) shows that recombination is linked to transcription in the mCP mutants. This study also provides evidence to suggest an R-loop dependent mechanism, first via immunofluorescent detection of RNA:DNA hybrids and second, by comparison of the R-loop staining with Rad52 foci formation (Figure 2.9). Further support comes from experiments demonstrating that the CIN phenotype of mCP mutants can be suppressed by overexpression of RNase H1 which prevents R-loop formation (Stirling et al, 2012)  	
    mCP mutants exhibit reported defects in transcription termination, elongation and mRNA export (Birse et al., 1998, Brodsky and Silver 2000, Tous et al., 2011). More recently, mRNA 3’-end processing has been shown to directly influence transcription re-initiation (Mapendano et al., 2010). Therefore, several potential molecular mechanisms could explain the R-loop formation underlying genome instability in mCP mutants. Importantly, by comparing hyper-recombination phenotypes to mutants in THO and SEN1 we find that mCP mutants behave similarly to THO mutants (Figure 2.11). This is consistent with the established strong interdependence of THO and mRNA 3’-end processing factors (Luna et al., 2005). It is known that THO occupancy at gene 3’-ends is decreased by ~50% when mCP genes are mutated and mCP occupancy at gene 3’-ends is significantly increased in THO mutants (Luna et al., 2005, Rougemaille et al., 2008). Thus, mCP acts upstream of THO in the biogenesis of mRNA export complexes, but THO is required to disassemble mCP machinery at 3’-ends. Therefore, our data best support a mechanism where mCP mutants facilitate RNA:DNA hybrid formation by reducing THO recruitment and/or increasing the time transcription machinery is present at gene 3’-ends due to inefficient termination. While the molecular mechanism of R-loop formation and propagation is complex even for better-characterized mutants in THO and SEN1, we can also make a statement globally about the impact of R-loop-induced DNA damage in mCP mutants. Our ChIP-chip data shows that mCP mutant-induced DNA damage occurs within transcribed genes and that the damaged 3’-UTRs tend to be associated with higher transcriptional frequencies (Stirling et al., 2012). More importantly,  	
    transcribed genes colliding with adjacent replication origins are enriched in DNA damage in mCP mutants, supporting a model where collision with DNA replication is a major source of DNA damage in mCP mutants (Figure 2.8 and Stirling et al., 2012). This is consistent with literature showing that collision of R-loops with the replisome is commonly associated with genome instability (Gan et al., 2011, Prado and Aguilera 2005, Sikdar et al., 2008, Tuduri et al., 2009). In contrast, the fact that mCP mutants can increase Rad52 foci in G1 cells suggests that DNA replication is not strictly required to elicit foci, although it may play a major role (Fig. 2.3). It may be that mCP mutants cause lesions requiring Rad52 in more than one way and it will be interesting in future studies to determine the relative contributions of transcription and replication to genome stability in these mutants. One confounding factor for the interpretation of these mechanisms is that we are studying a set of partial loss-offunction mutations in essential genes whose protein products form a multi-functional and dynamic protein complex. Therefore, despite their phenotypic overlap, there are certain to be gene and allele-specific effects within the mCP mutants described here that modulate the phenotypes observed. A genome-wide compilation of yeast CIN genes identified dozens of transcription and RNA processing related CIN mutants whose R-loop status is unknown (Stirling et al., 2011). We predict that other CIN mutants in these pathways also induce R-loop formation. Interestingly, a high-throughput study of γ-H2AX DNA damage foci in mammalian cells identified many RNA processing genes, suggesting that this pathway is generally important for genome integrity  	
    (Paulsen et al., 2009). Moreover, these authors found that overexpression of RNaseH could suppress γ-H2AX foci in some instances, supporting the R-loop model for diverse RNA processing pathways (Paulsen et al., 2009). The prevalence of R-loop-mediated CIN in cancer is unclear although it is tempting to speculate that it could occur in cancers where the orthologs of known R-loop forming mutants are disrupted (e.g. TOP1 or FIP1; (Cools et al., 2003, Iwase et al., 2003).  	
    Chapter 3. Biogenesis of RNA polymerase II and III requires the GPN small GTPases in Saccharomyces cerevisiae 3.1 Background information 	
   Previous efforts in our laboratory to systematically catalogue CIN phenotypes among essential yeast genes have uncovered the cellular pathways required to maintain genome stability (Stirling et al., 2011); both predictable and less-predictable pathways emerged as highly enriched for CIN genes. In addition, the CIN gene catalogue identified a suite of conserved CIN genes that are poorly characterized (Ben-Aroya et al., 2008; Stirling et al., 2011). In principle, any perturbation of a conserved process or CIN gene could be responsible for modulating genome stability in human cancer. YOR262W (hereafter referred to as GPN2) was identified as a conserved and essential CIN gene in one of our recent efforts (Ben-Aroya et al., 2008). GPN2 belongs to a highly conserved family of small GTPases and exists in yeast and humans with two paralogues, GPN1 (yeast NPA3) and GPN3 (YLR243W, hereafter referred to as GPN3).  In archaea, a single GPN gene  encodes a protein with a conserved glycine-proline-asparagine motif in the G domain that gives the family its name.  The 3-D structure of the archaeal GPN  protein (PAB9855) reveals a homodimeric molecule with a canonical GTPase domain and the purified protein exhibits GTPase activity in vitro (Gras et al., 2007) The first characterized human GPN orthologue (GPN1/XAB1/NPA3) was shown to bind the nucleotide excision repair protein XPA and was named XAB1 (XPA binding protein 1). The XPA-XAB1 interaction was presciently, although indirectly, suggested to play a role in nuclear localization of XPA because deletion of 	
    amino acids required for nuclear localization disrupted XAB1 binding (Nitta et al., 2000). Subsequent mass spectrometry studies of partially assembled RNA polymerase II (RNAPII) complexes identified the Gpn1, Gpn2 and Gpn3 proteins and revisited the possibility of a role for GPNs in nuclear transport (Boulon et al., 2010; Forget et al., 2010). These observations initiated directed studies of GPN1 function in human tissue culture and yeast. In yeast, reduction of function of NPA3/GPN1 leads to chromatid cohesion defects, cell cycle defects and cytoplasmic mislocalization of the RNAPII subunits Rpb1 and Rpb3p (Alonso et al., 2011; Staresincic et al., 2011). Mutations of NPA3/GPN1 designed to abrogate its GTP binding or hydrolysis activities also cause defects in RNAPII nuclear transport. Work in human cells shows that both GPN1 and GPN3 are required for nuclear import of RNAPII, leaving the function of GPN2 unknown (Carre and Shiekhattar. 2011). Interestingly, another poorly-characterized protein, Iwr1, was recently shown to be important in the localization of RNA polymerase II, presumably in co-operation with the GPNs (Czeko et al., 2011). Unlike the GPNs and indeed unlike RNA polymerases themselves, Iwr1 contains a bipartite nuclear localization signal (NLS) which may serve to direct the nascent RNA polymerase to a karyopherin-mediated nuclear import pathway (Czeko et al., 2011). IWR1 function has also been linked to the transcriptional activity of all three nuclear RNA polymerases (Esberg et al., 2011). Importantly, IWR1 is dispensable for cell viability unlike the three GPN genes; therefore, its role in nuclear import of RNA polymerases must be buffered by some other cellular activity.  	
    This chapter includes characterization of mutant alleles of the previously uncharacterized yeast GPN2 and GPN3 orthologues. Mutations in these genes cause a chromosome loss phenotype and sensitivity to UV and genome destabilizing chemicals. The GPNs also exhibit physical and genetic interactions with one another supporting a common function for this protein family. Remarkably, rather than having independent functions in the nuclear transport of different substrates as might have been predicted, mutants in either GPN2 or GPN3 cause defects in the localization of protein subunits of both RNAPII and III. Finally, we show that fusion of a NLS to the RNAPII subunit Rpb3p does not restore nuclear localization of Rpb1 in GPN mutants, while partially rescuing the defects in iwr1Δ. These findings, combined with information from the literature, support a model in which all three GPNs serve independently essential functions in the biogenesis of RNA polymerase II and III that are upstream of the NLS activity contained in Iwr1.  3.2 Materials and methods 3.2.1 Yeast growth, strains and plasmids 	
   Yeast strains and plasmids used in this study are listed in the appendix Table A2. Ts-alleles were generated as described (Ben-Aroya et al., 2008). Other strains were constructed by PCR-mediated one step gene replacement using a published set of tagging cassettes and standard PEG/lithium acetate transformation (Longtine et al., 1998). For the RPB3- and RPC40-NLS fusions, the SV40 NLS sequence (PKKKRKV) was incorporated into mutagenic primers designed to fuse the NLS to the C-terminus of the protein followed by the KanMX marker from pFA6-KanMX6 (Longtine et al., 1998). The TAP and GFP-tagged strains were obtained from the  	
    proteome-wide TAP (Open Biosystems) and GFP collections (Laboratory of Brenda Andrews).  Site-directed mutagenesis was performed using the Quickchange  Lightning Kit (Agilent Technologies) and GPN2 and GPN3 clones from the MoBYORF collection (Ho et al., 2009). Yeast were grown in rich media (YPD) or synthetic media as indicated and at the temperatures indicated. Hydroxyurea or 6-azauracil (Sigma) was added at the indicated concentrations to YPD or SC-uracil media respectively. Genetic interactions were assessed by tetrad analysis. Viable double mutants were subjected to spot dilution or growth curve assays to confirm enhanced sickness of double mutants. For spot plating assays, overnight cultures of the indicated strains were normalized to the same OD600 and subsequently spotted in 10-fold serial dilutions onto the indicated media and grown for 24 to 48 hours before imaging. 3.2.2 GPN mutant allele sequencing 	
   Phenol-chloroform preparations of mutant strain genomic DNA were made and the GPN genes were amplified using flanking primers. PCR products were purified using the ChargeSwitch kit (Invitrogen) and cycle sequencing amplification was conducted using the BigDye kit (Applied Biosystems).  Sequencing was  performed by the NAPS unit in the Michael Smith Laboratories at the University of British Columbia. 3.2.3 Microscopy 	
   Overnight cultures were diluted and allowed to grow for several hours to reenter log phase and subsequently shifted to the indicated temperatures for 3 hours. For live cell imaging, log phase cells were mounted on concanavalin A coated 	
    slides, washed and imaged in SC media essential as described (Carroll et al., 2009; Stirling et al., 2012b). In some cases 1 mg/ml DAPI was added to live cells immediately prior to imaging to mark the nucleus. Live and fixed cells were imaged with the appropriate filter sets on a Zeiss Axioscope using Metamorph software (Molecular Devices). 3.2.4 Localization scoring and statistics 	
   GFP localization was assessed qualitatively by counting the proportion of cells with strong nuclear signal (i.e. the predominant localization in wild-type) and comparing the wild-type control to GPN mutants using a Student’s t-test. We also extracted a quantitative measure of mislocalization by defining a pixel area within the DAPI-stained nucleus and another, equally-sized area immediately adjacent in the unstained cytosol. The nuclear localization score expresses the total fluorescence  pixel  intensity  within  these  two  areas  as  a  ratio  (i.e.  nucleus/cytoplasm). Pooling intensity ratio data for three replicates in which at least 30 cells were scored permitted computation of the mean and standard deviation of scores. Where two samples were compared, a Student’s t-test was used to determine significance (p-values are reported in the figures). In situations where greater than two samples were compared, a one-way ANOVA followed by Tukey’s post-hoc test was used to determine whether the samples were significantly different (α is reported in the figures).  	
    3.2.5 TAP pull-downs and western blotting 	
   TAP pull-downs were performed essentially as described (Kobor et al., 2004). Briefly, 100 of mL TAP-tagged and control cultures were grown to an OD of 1.0, collected via centrifugation and lysed in TAP-IP buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1.5 mM magnesium acetate, 10 mM sodium pyrophosphate, 5 mM EDTA, 5 mM EGTA, protease inhibitors) using glass beads and a Precellys 24 tissue homogenizer. Lysates were cleared via centrifugation at 14,000 rpm for 15 minutes and then incubated with IgG beads (GE Healthcare) for 90 minutes with rotation. Beads containing bound proteins were washed four times with TAP-IP buffer followed by elution with 2x SDS-PAGE buffer. TAP pull-downs involving Gpn3-TAP required growth in yeast media containing 2% galactose as the N-terminally tagged Gpn3 was only functional when over-expressed from a GAL1 promoter. The pulldown fractions and lysates were loaded onto Mini-Protean TGX acrylamide gels (BioRad) and transferred to nitrocellulose membranes and blocked using PBS containing 0.02% Tween-20 and 5% skim milk powder.  Blots were  probed with mouse monoclonal anti-myc antibodies (Roche Cat #11667203001, 1:2500) or rabbit anti-TAP (Thermo Scientific Prod. # CAB1001, 1:2000) followed by anti-mouse horseradish peroxidase (HRP) or anti-rabbit HRP conjugated antibodies (1:15,000). Blots were imaged using Biorad Chemi-Doc system. For western blots to assay protein levels, 5 mL log phase cultures were collected by centrifugation, resuspended in TAP-IP lysis buffer and lysed in the Precellys with acid-washed glass beads. Lysates were cleared by centrifugation and quantitated using the Bradford protein assay (Bio-Rad). 10 µg total protein was  	
    subjected to electrophoresis on an SDS-PAGE gel, and transferred to a nitrocellulose membrane for blocking. Membranes were washed and incubated with anti-Rpb3 antibody (Neoclone Cat. #: W0012, 1:1000 dilution) followed by goat antimouse horse-radish peroxidase (HRP) secondary antibody (1:3000). Blots were imaged, stripped (200 mM glycine, 0.1%SDS, 1% Tween 20, pH2.2) and reprobed with anti-PGK1 antibody (Invitrogen Cat. # 6457, 1:10,000) followed by goat antimouse HRP (1:15,000) followed by imaging. 3.2.6. Nuclear proteome scale GFP microscopy screen 	
   A mini-array of GFP tagged strains was generated for proteins annotated as localized to the nucleus or nuclear periphery. This array was mated with a gpn2-2 mutant strain containing an mCherry-tagged histone H2A and haploid selection was done using SGA technology (Tong et al, 2004). Wild type and mutant haploids were grown in 96 well plates and manually screened for localization of the GFP tagged proteins using 12-well microscope slides. Images were scored for differences using Metamorph Imaging Software. Putative hits and controls were validated by tetrad dissection and additional microscopy.  	
    3.3 Results 3.3.1 Mutational analysis of the GPNs: an evolutionarily conserved protein family Three distinct GPN GTPases have been identified in humans and yeast. Each GPN is more closely related to its inter-species orthologue than its intraspecies paralogues suggesting that three functionally conserved sub-families may exist. To understand the evolutionary roots of the GPNs, we identified GPN1, 2 and 3-like sequences using BLAST-P, from a set of diverse eukaryotes and archaea and built a phylogenetic tree (Figure 3.1). Our phylogenetic analysis suggests that while GPN-like proteins are also found in diverse archaeal species, they are not universally conserved in archaea and are never found in bacteria (Figure 3.1). This analysis identified likely members of each GPN1, 2 and 3 group in all the diverse eukaryotes examined, including even one of the most minimal eukaryotic genomes known from the microsporidian Encephalitzoon cuniculi (Figure 3.1;(Katinka et al., 2001). The remarkable conservation of three distinct GPN proteins across eukaryotes suggests that each GPN serves an important function not served by the paralogous GPN proteins. The importance of each GPN is supported by the fact that NPA3/GPN1, GPN2 and GPN3 are all essential genes in S. cerevisiae (Giaever et al., 2002).  	
    Figure 3.1 Phylogenetic tree of GPN orthologues. A ClustalW multiple alignment of the BLAST-P top hit GPN1, 2 or 3 orthologues from the indicated species was plotted using Treeview. Colored bubbles highlight the four largegroups identified in the tree which correspond to GPN1, GPN2, GPN3 or archaeal GPN. Genus and species information for eukaryotic abbreviations and archaeal genera is shown adjacent.  	
    NPA3 (GPN1) is the best characterized yeast GPN, whereas GPN2 and GPN3 are considerably less studied. To generate tools to specifically interrogate the functions of GPN2 and GPN3 we created mutant alleles of each gene using a mutagenic PCR approach described previously (Ben-Aroya et al 2008). Four gpn2 mutants and three gpn3 mutants were isolated that exhibit hypomorphic and, in some cases, temperature-sensitive (ts) growth (Figure 3.2). The mutant alleles were sequenced and variants mapped onto the crystal structure of archaeal GPN PAB0955 to assess the potential functional impact. Each mutant had between 1 and 5 amino acid changes resulting from missense mutations (Table 3.2). In some cases the mutations appeared in the GTPase domain, while in others the putative dimerinterface was more directly affected (summarized in Figure 3.2). While the specific functional consequence of each mutation remains to be assessed, each allele was a partial loss-of-function mutation that decreased cellular fitness. We selected alleles that showed hypomorphic growth with temperature-dependent phenotypes and low incidence of suppressors, gpn2-2 and gpn3-1 (Figure 3.2) for further study.  Table 3.1 Mutations in GPN2 and GPN3 alleles. Allele  Amino acid changes  Potential impact based on archaeal GPN structure  gpn2-2  C19 predicted in an alpha helix adjacent to GTP  gpn2-4  C19S E97D, I153T, K176E, K181R, E331G* E98A, K125R, R161K, E203G, E338G* L228P  gpn3-1  I58T, H123R, S148L, F150S  Potentially dimer interface Predicted dimer interface or protein core  gpn3-9  M27T, C134R, N205D  Predicted dimer interface or protein core  gpn3-10  G153S, I169T, S241G  Predicted dimer interface or protein core  gpn2-1 gpn2-3  K176 predicted to contact GTP K125 predicted directly in dimer interface  Predicted locations based on alignment to PDBID: 1YR9 * Residue not found in crystal structure  	
    Figure 3.2	
   Generation and characterization of GPN2 and GPN3 ts-alleles. (A) Schematic of the 347 amino acid Gpn2 and 272 amino acid Gpn3 proteins with amino acid substitutions in each mutant allele used in this study overlaid. B) Growth curves of GPN2 and GPN3 mutant alleles. Equal cell numbers were inoculated and grown at the semi-permissive temperature of 30°C for 24 hours. The identity of each allele is indicated on the graph. The graph represents the mean of three replicates. Each allele grew significantly worse than wild-type as measured by the area under the curve (McLellan et al., 2012).  GPN2 was originally described as a chromosomal instability mutant displaying sister chromatid cohesion defects, a phenotype which was also  	
    subsequently observed for GPN1 mutants (Alonso et al., 2011; Ben-Aroya et al., 2008). Some hypomorphic alleles of GPN3 created here also show a visible Chromosome Transmission Fidelity (CTF) phenotype (Figure 3.3; (Spencer et al., 1990; Stirling et al., 2012a). Thus each member of the GPN family appears important for maintaining genome integrity. Some of the GPN2 and GPN3 mutants also exhibited increased sensitivity to hydroxyurea, a nucleotide pool poison that increases stalling of replication forks (Figure 3.3). Since human GPN1 has been reported to bind the UV-damage repair protein XPA/Rad14 we also tested UV sensitivity in GPN mutants and found that they were hypersensitive to a UV dose of 25 J/m2 (Figure 3.3; (Nitta et al., 2000). Importantly mutants in RNAPII show a CTF phenotype and other genome integrity defects (Stirling et al., 2011; Stirling et al., 2012b). Together, this suggests that the CIN phenotype of GPN mutants could be a result of reduced function of RNA polymerase.  	
    Figure 3.3 Chromosomal instability and DNA damage sensitivities of GPN mutants. A) GPN3 mutations cause chromosome integrity defects. Red sectoring is indicative of loss of an artificial chromosome containing a suppressor tRNA that prevents the accumulation of red pigment due to the ade2-101 mutation (Spencer et al., 1990). (B) GPN2 and GPN3 mutants exhibit heightened sensitivity to the DNA damaging agent hydroxyurea at 30oC. (C) GPN mutants exhibit sensitivity to exposure to 25 J/m2 ultraviolet radiation exposure followed by growth at the semi-permissive temperature of 34oC for 48 hours.  3.3.2 GPN2 exhibits genetic and physical interactions with GPN3, NPA3/GPN1 and IWR1 The mutants we describe for GPN2 and GPN3 share genome integrity phenotypes consistent with the possibility that the GPNs cooperate to execute some common cellular function. Indeed, physical interactions between GPN family members in both human and yeast cells have previously been shown (Boulon et al., 2010; Staresincic et al., 2011). In yeast, two-hybrid data suggest that Npa3/Gpn1  	
    binds to both Gpn2 and Gpn3 (Uetz et al., 2000). We found that Gpn2 also binds to both Npa3/Gpn1 and Gpn3 by affinity purification and western blotting of epitopetagged protein (Figure 3.4A). It should be noted that while C-terminal Gpn2 and Npa3 epitope fusions (e.g. 13-myc or TAP) supported robust growth as the sole source of the protein, Gpn3 fusions had to be N-terminal and were not completely functional, only supporting growth when over-expressed from a GAL promoter (unpublished observation). Gpn3 is the smallest GPN protein (~9kDa smaller than Gpn2) and is missing a fungal-specific C-terminal domain found in Gpn2; whether this accounts for its sensitivity to epitope tagging is unknown. Together with our data, interactions between all yeast GPNs have now been observed and the structural and in vitro data suggest that these interactions may involve oligomerization (Figure 3.4; (Gras et al., 2007; Staresincic et al., 2011)). Crystallographic studies of the archaeal GPN protein PAB0955 suggest that homodimerization occurs between PAB0955 proteins (Gras et al., 2007). In humans, both homo- and heterodimerization have been observed with purified Gpn1 and Gpn3 (Carre and Shiekhattar. 2011). Thus, eukaryotic GPN interactions could occur through homodimerization, heterodimerization or simply in the context of a large protein complex. To begin to clarify this issue, we determined whether Gpn2 could homodimerize in vivo using a diploid strain containing one 13-myc tagged copy and one TAP-tagged copy of Gpn2. Pulldown of the TAP-tagged Gpn2 did not result in co-precipitation of 13-myc tagged Gpn2 beyond background, suggesting that S. cerevisiae Gpn2 may not homodimerize and indeed that the stoichiometry of Gpn2 in stable cellular protein complexes may be one (Figure 3.4).  	
    Figure 3.4 Gpn2 physically interacts with both Gpn1 and Gpn3 but does not appear to homodimerize. Co-precipitation tests of Gpn2-13-Myc by TAP fusions of Npa3 or Gpn3 (A) or Gpn2-TAP (B) as measured by western blotting with anti-TAP or anti-Myc antibodies. WCE, Whole Cell Extract.  To assess a common or partially overlapping cellular function for GPNs we generated double mutants among NPA3, GPN2, GPN3 and IWR1 mutant alleles. Tetrad analysis of double heterozygous mutants of gpn2-2 NPA3::DAmP (Decreased Abundance by mRNA Perturbation; (Breslow et al., 2008) showed synthetic lethality between these two genes (Figure 3.5A). The gpn2-2 gpn3-1 double mutants could be isolated at 25°C but growth curve analysis revealed a strong genetic interaction between GPN2 and GPN3 at higher temperatures (Figure 3.5B). IWR1 plays a role in nuclear localization of RNAPII and would therefore be predicted to synergize with other GPN mutants if they play a role similar to NPA3 in the process (Czeko et al., 2011; Staresincic et al., 2011). Consistently, iwr1Δ gpn2-2 double mutants show a severe synergistic temperature-sensitive growth defect, supporting a connection between GPN2 and nuclear transport of RNAPII (Figure 3.5C). Interestingly, similar to otherwise non-essential core RNAPII mutants (i.e. RPB4 and RPB9; (Woychik and Young. 1989; Woychik et al., 1991), the function of  	
    Iwr1 becomes essential at 37°C, as IWR1 mutant cells failed to grow at this temperature (Figure 2E).  Figure 3.5 Genetic interactions between GPN2 mutants and NPA3/GPN1, GPN3 and IWR1 mutants. (A) Tetrad dissection of gpn2-2 NPA3::DAMP double mutants. Triangles indicate where double mutant colonies should grow. (B) Growth curve assay of gpn2-2, gpn3-1 and double mutants at the indicated temperature. (C) Spot dilution assays of WT, gpn2-2, iwr1Δ and double mutant cells. Cells were grown for 2 days at the indicated temperature.  3.3.3 GPN2 and GPN3 mutants are defective in localization and stability of RNAPII subunits Core subunits of RNAPII were shown to mislocalize from the nucleus to the cytoplasm when GPN1 or GPN3 was disrupted by siRNA in human cells or by mutation of NPA3/GPN1 in yeast (Carre and Shiekhattar. 2011; Staresincic et al., 2011). To examine the impact of our GPN2 and GPN3 mutants we expressed a functional GFP-tagged version of Rpb1 in each mutant. We found that both gpn2-2 and gpn3-1 mutants show defects in Rpb1-GFP localization, as there is a strong  	
    cytoplasmic accumulation of Rpb1-GFP compared to the wild-type genetic background (Figure 3.6A). This is not specific to Rpb1 as both Rpb2-GFP and Rbp3-GFP also mislocalized in gpn2-2 mutants (data not shown). To assess the levels of endogenous RNA polymerase subunits we took advantage of a specific antibody to yeast Rpb3. Western blots showed a notable reduction in the levels of Rpb3 in gpn2-2 and gpn3-1 mutants at both 25°C and 37°C, when compared to wild type (Figure 3.6B). Together these data demonstrate that each GPN family member plays a role in the localization of RNAPII subunits and that soluble subunit levels are also altered either because of instability or reduced expression. To phenotypically link reduced Rpb3 protein levels to some of the GPN2/3 mutant allele phenotypes, we exploited an RPB3-DAmP allele that expresses reduced levels of Rpb3 (data not shown). Similar to GPN2 and GPN3 mutants, the RPB3-DAmP allele was hypersensitive to both HU and UV treatments, suggesting that reduced RNA Pol II subunit levels could account for at least some of the GPN mutant phenotypes (Figure 3.6C).  	
    Figure 3.6 RNA polymerase II mislocalization in GPN mutants. (A) Representative GFP micrographs showing the localization of Rpb1-GFP in WT, gpn2-2 and gpn3-1 mutants. Scale bar indicates 4 mm. (B) Mutation of GPN2 or GPN3 leads to reduced levels of an RNA polymerase II subunit. Rpb3 protein levels in whole cell lysates of the indicated strains were assessed by western blot with anti-Rpb3 specific antibodies. Anti-PGK1 blots are shown to indicate equal loading. C) RPB3-DAmP allele sensitivity to HU and UV treatment comparable to a rad52Δ control. Log phase cultures were normalized to the same density and serially diluted 10 -fold before plating and, if applicable, UV exposure, followed by growth at 30°C for 48 hours.  Based on observed genetic interactions between GPN mutants, one question was whether the RNAPII mislocalization or temperature-sensitivity could be suppressed through overexpression of the GPN proteins or IWR1.  High copy  number plasmids from the MOBY yeast ORF collection (Ho et al., 2009) were transformed into the gpn2-1 and gpn2-2 Rpb1-GFP mutant strains to look for suppression of phenotypes. In all cases, only the wild-type GPN2 gene could fully suppress the mutant phenotype although mild suppression of the gpn2-1 ts  	
    phenotype by GPN1 was observed at 34oC (Figure 3.7A) and overexpression of GPN3 appeared detrimental to gpn2-1 mutant growth at lower temperatures. A similar result was observed in the Rpb1-GFP gpn2-2 mutant strain as only the GPN2 plasmid could rescue the nuclear localization defect of RNAPII in the mutant background (Figure 3.7B). These results support a non-overlapping function for the GPN family members.  	
    Figure 3.7 Overexpression of GPN family members and IWR1 does not strongly complement growth and Rpb1 localization defects in GPN mutants. A) gpn2-1 mutant strains containing MOBY 2µ plasmids grown on SC-leu at a range of temperatures for 48 hours B) gpn2-2 Rpb1-GFP strains containing overexpression plasmids grown in SC-leu at 30oC and imaged using by fluorescence microscopy and. All images represent gpn2-2 mutants with the exception of the wild-type (WT) control on in the leftmost panel.  Previous work has implicated conserved aspartic acid (D) and glutamine (Q) residues involved in GTP binding and hydrolysis respectively, as important for the 	
    function of NPA3/GPN1 in localization of RNA polymerase II (Staresincic et al., 2011). To assess the importance of these same residues for cell viability in Gpn2 and Gpn3, we engineered point mutations at the orthologous sites via site-directed mutagenesis.  Plasmids containing GPN2 with D106A or Q110L mutations and  GPN3 with D104A or Q108L mutations were transformed into heterozygous deletion mutant strains followed by sporulation and tetrad dissection. As shown in Figure 3.8A, the gpn2D106A mutation results in a non-functional protein that cannot complement the deletion mutant while the gpn2Q110L mutant protein is functional. A similar result was observed for analogous point mutants in GPN3 (Figure 3.8B). These data suggest that GTP binding is essential for Gpn2 and Gpn3 function but GTP hydrolysis may not be, although at this point it is unclear to what degree the Q110 and Q108 mutations actually impair GTP hydrolysis. Additionally, the previous work on NPA3/GPN1 involved complementing ts-degron alleles with the point mutants, rather than knockouts, which may account for the differences observed here (Staresincic et al., 2011).  	
    Figure 3.8 GTP binding is required for function of the GPN proteins. A)Tetrad analysis of a heterozygous deletion mutant for GPN2 transformed with wild-type and mutant GPN2 plasmids reveals the requirement for the GTP binding D106 residue for Gpn2 function but not Q110. (D) Mutation of the conserved D104 and Q108 residues in Gpn3 concurs with the observed Gpn2 results.  Given the disrupted localization and levels of some RNAPII subunits it would be expected that GPN mutants would be sensitized to additional perturbation of transcription. Consistently, the GPN mutants showed increased sensitivity to the transcription elongation inhibitor 6-azauracil (Figure 3.9A). Moreover, gpn2-2 showed synthetic lethality or synthetic growth defects when combined with mutants in the core RNAPII subunits RPB1 (rpo21-1), RPB2 (rpb2-6) or the transcriptional regulator ESS1 (ess1-H164R) (Figure 3.9B). gpn3-1 also showed synthetic lethality 	
    with rpb2-6 (Figure 3.9C). These data, together with published data (Staresincic et al.,, 2011), show that loss-of-function mutations in any GPN family member lead to incomplete RNAPII localization and function (Staresincic et al., 2011).  Figure 3.9 Functional impact of GPN mutations on RNAPII. (A) Spot-dilution assay for 6azauracil (6-AU) sensitivity in the indicated GPN2 and 3 mutants. Genetic interactions were tested by (B) tetrad analysis of gpn2-2 crossed with RNA polymerase II transcription mutants and (C) tetrad analysis of gpn3-1 crossed with rpb2-6.  	
    3.3.4 GPN2, GPN3 and IWR1 mutants are defective in localization of RNAPIII Nuclear localization of model nuclear proteins was not obviously affected by GPN mutation (Minaker et al., 2013). However, it remains possible that transport of an unknown subset of the yeast nuclear proteome is affected by GPNs. Among the best candidates for additional substrates of the GPNs are the RNA polymerase I and III complexes. While comparatively little is known about the assembly of RNAPI and III, they have paralogous subunit architectures to RNAPII and share a number of subunits. Therefore, GFP fusions to representative RNA polymerase I and III subunits were crossed to GPN2 and GPN3 mutant strains and visualized. Fluorescence microscopy revealed that the RNAPIII subunit Rpc53-GFP is mislocalized in both gpn2-4 and gpn3-1 mutant backgrounds, whereas the RNAPI subunit Rpa135-GFP appeared unaffected by either GPN mutation (Figure 3.10A and Appendix Figure A2). It should be noted that introduction of the GFP-tagged versions of several RNA polymerase I and III subunits into the GPN mutant background resulted in lethality suggesting genetic interactions of GPN mutants with both RNAPI and III, indeed this is why the gpn2-4 allele was used instead of gpn2-2 (data not shown). While the localization of Rpc53-GFP in GPN mutants was qualitatively different than wildtype, a clear nuclear signal usually remained (Figure 3.10A). While this might be expected given the hypomorphic nature of the GPN alleles, it prompted us to develop a quantitative measure of nuclear localization to which we could assign statistical significance. Figure 3.10B reports the ratio of GFP signal in the nucleus versus the cytoplasm for the data in Figure 3.10A. Importantly, we confirmed our observations using an independent RNA polymerase III subunit,  	
    Rpo31-GFP, in gpn2-2 and gpn3-1 mutants using the same quantification scheme (Figure 3.10B, lower panel). This method recapitulates the qualitative observations and shows a significant reduction in nuclear Rpc53-GFP levels in gpn2-4 and gpn31 cells and Rpo31-GFP levels in gpn2-2 and gpn3-1 (a=0.01). When we directly tested for genetic interactions with RNAPI and III subunit tsalleles, we identified synthetic slow growth interactions between gpn2-2 and alleles in all three RNA polymerases (RPA190 , RPO31 and RPC34), a RNA polymerase I transcription factor (RRN3), and between gpn3-1 and RPC34 and potentially RPA190 (Figure 3.10C). Since transcription by all three RNA polymerases is required to coordinate cellular activities (for example, ribosome assembly), it is not surprising that the GPN mutants could exhibit genetic interactions with all three RNAPs while only affecting localization of RNAPII and III. Alternatively, it is possible that in different GPN mutant backgrounds or with different GFP fusions, RNAPI mislocalization would become visible.  	
    Figure 3.10 Effect of the GPN-IWR1 system on RNA polymerase III localization. (A) Nuclear localization of Rpc53-GFP in gpn2-4 and gpn3-1 mutants. (B) Schematic of nuclear localization scoring system and quantification of Rpc53-GFP localization from panel A. a indicates the results of Tukey’s posthoc analysis of a one-way ANOVA for the three datasets. (C) Spot dilution assays of GPN2 (left) and GPN3 (right) mutants in combination with RNAPI and III subunit mutant alleles.  Since the GPNs may cooperate with IWR1 in RNAPII localization we also sought to explore the effects of IWR1 mutations on RNAPI and III localization. For this experiment we introduced the IWR1 deletion into four GFP-fusion strains: Rpo31-GFP and Rpc37-GFP, subunits of only RNAPIII; Rpc40-GFP, a subunit of both RNAPI and RNAPIII; and Rpa135, a subunit of only RNAPI. We observed clear mislocalization of Rpc37, Rpo31 and Rpc40 in the iwr1Δ strains but did not see any mislocalization of Rpa135-GFP, indicating that Iwr1 and the GPNs are affecting  	
    RNAPIII but not obviously affecting RNAPI (Figure 3.11A and Appendix Figure A2). Quantification of the nuclear/cytoplasmic GFP ratios from Figure 3.11A showed that the mislocalization of RNAPIII subunits in iwr1Δ was statistically significant (Figure 3.11B; p<0.0001).  Figure 3.11 RNA polymerase III is mislocalized in iwr1Δ mutants. A) Localization of Rpc37GFP and Rpc40-GFP in iwr1Δ mutants. B) Quantification of Rpc37- Rpc40- and Rpo31GFP fusions using the scoring system from Figure 3.10. p-values indicate the results of a Student’s t-test.  3.3.5 Fusing a nuclear localization signal to Rpb3p partially bypasses IWR1 but not GPN mutants In light of the data presented in Figures 3.5, 3.10 and 3.11 and in recent publications on the GPN proteins and the NLS-containing protein Iwr1 (Di Croce, 2011), we propose a co-operative role for these proteins in RNAPII biogenesis and nuclear import. However, it is unclear whether the GPN proteins primarily function in assembly or nuclear import. To address this, and begin to dissect the functional  	
    contributions of GPNs and IWR1, we hypothesized that fusion of a strong NLS directly to Rpb3p could bypass the nuclear localization defect in iwr1Δ and potentially GPN mutants. To test this hypothesis in vivo, we generated a fusion of the SV40 NLS (PKKKRKV) to RPB3. As RPB3 is an essential gene, and the fusion strain grew normally, we inferred that the fusion protein was functional. Remarkably, the Rpb3-NLS fusion protein exacerbated the growth defect of gpn2-2 mutants at 37°C while partially rescuing the growth defect of iwr1Δ mutants at 34°C (Figure 3.12). While the RPB3-NLS fusion was competent to support robust growth, it did slightly but significantly reduce the nuclear/cytoplasmic GFP ratio in Rpb1- and Rpb2-GFP bearing strains (Figure 3.12B). This could indicate that aberrant nucleartargeting of RNAPII subunits may not permit efficient RNAP assembly.  Figure 3.12 Cellular effects of Rpb3-NLS fusion on RNAPII localization. (A) Fitness of gpn22, gpn3-1 and iwr1Δ mutants with or without Rpb3-NLS fusion. Spot dilution assays of the indicated strains were performed as in Figure 3.5. (B) Effect of NLS-fusion on nuclear localization of Rpb1-GFP and Rpb2-GFP in wild-type cells.  	
    When we examined the localization of Rpb1-GFP in the GPN and IWR1 mutants containing the Rpb3-NLS fusion, we observed qualitatively similar mislocalization in the gpn2-2 and gpn3-1 mutants but observed partial rescue in the iwr1Δ strain (Figure 3.13A). Quantification of these data confirmed a significantly higher nuclear/cytoplasmic GFP ratio in the iwr1Δ strains bearing the NLS-fusion than the strain without the NLS-fusion (Figure 3.13B). To prove that this function was also important for RNA polymerase III, we fused the same NLS sequence to RPC40 and found that this could partially restore localization of Rpo31-GFP in iwr1Δ strains (Figure 3.13C). Together these data show that addition of an alternative nuclear import signal can partially rescue IWR1 but not GPN2 or GPN3 mutants. In support of the literature on Gpn1 and Iwr1, these data imply that the Gpn2 and Gpn3 proteins are working upstream of import to assemble functional RNAPII complexes while at least part of the function of Iwr1 is specifically tied to nuclear import. Moreover, these data suggest that the NLS contained within IWR1 is critical for its role in RNA polymerase III localization. Indeed, we found that, while expression of full-length IWR1 from a plasmid rescues RNAPII and III localization in iwr1Δ, expression of partial or complete deletions of the NLS sequence failed to improve localization of Rpb1-, Rpo31- or Rpc37-GFP fusion (Appendix Figure A3).  	
    Figure 3.13 NLS fusion constructs partially rescue the RNA polymerase mislocalization of iwr1Δ mutants. A) Qualitative localization defects of Rpb1-GFP in the indicated mutant background with (below) and without (above) the RPB3-NLS fusion construct. (B) Quantification of Rpb1-GFP localization in the strains from Panel A as in Figure 3.10. (C) Quantification of Rpc37-GFP localization in iwr1Δ cells with or without the Rpc40-NLS fusion. p-values indicate the results of Student’s t-tests.  3.3.6 A nuclear proteome wide GFP localization screen reveals that GPN2 mutation affects localization of transcription associated proteins Our data and the literature demonstrate a role for GPNs in assembly and subsequent nuclear import of RNA polymerase II; however, it is unclear whether the GPNs have a broader role in the assembly and import of other proteins. To address this we tested for mislocalization of other nuclear proteins using the GFP-tagged yeast collection. The majority of the ~1350 nuclear proteins retained normal  	
    localization in the gpn2-2 mutants that we screened. This result indicates that the canonical nuclear import pathways are likely functioning normally in the GPN2 mutant background. Over 25% of the proteins with obvious mislocalization were found to be RNA polymerase II and III subunits as expected. However, it was noted that several transcription and splicing factors appeared to be mislocalized in the gpn2-2 mutant background (Figure 3.14, Table 3.2). Six RNA polymerase II or III subunits and 17 other nuclear proteins have been validated from tetrad crosses as mislocalized in the gpn2-2 mutant background (Table 3.2). The mislocalization of entire nuclear protein complexes besides the RNA polymerases was not observed, suggesting that the primary role of Gpn2 is in RNA polymerase II and III assembly. However, since many of the nuclear proteins observed showed equal staining of the nucleus and cytoplasm, more subtle effects of the GPN proteins cannot be formally excluded without quantification. Notably Rad6, an E2 ubiquitin-conjugating enzyme associated with DNA repair (Jentsch et al, 1987, Sung et al, 1991), is known to function in H2B ubiquitination during transcription elongation and has been found to physically associate with RNAPII (Xiao et al, 2005). Thus, the DNA damage sensitivity of the GPN mutants could be due to aberrant nuclear localization of transcription associated repair proteins such as Rad6. A possible explanation for the mislocalization of other proteins is binding to mislocalized RNA polymerase II subunits, thus sequestering them inappropriately in the cytoplasm.  	
    Figure 3.14 Examples of GFP microscopy displaying effects of gpn2-2 mutation on localization nuclear proteins. These included multiple RNA polymerase II and III subunits in addition to the splicing and elongation factors Msl1 and Elf1. 	
    Table 3.2 Validated hits from a screen for mislocalization of GFP tagged nuclear proteins in a gpn2-2 mutant background shows enrichment for transcription and splicing factors. GENE	
    3.4 Discussion 3.4.1 Gpn2 and Gpn3 are part of an evolutionarily conserved family required for RNA polymerase II biogenesis genomic integrity The GPNs represent a highly conserved family of small, apparently active GTPases that evolved in the common ancestor of eukaryotes and archaea (Figure 3.1).  Moreover, experiments with Npa3/Gpn1 suggest the potential for the  nucleotide switch-like behavior seen in better characterized GTPases like Ras or the nuclear import regulator Ran (Staresincic et al., 2011). The precise role of the GTPase activity of the GPNs is not clear, although there is evidence that the affinity  	
    of Npa3/Gpn1 for RNAPII is regulated by its GTP binding status (Staresincic et al., 2011). Our mutational analysis suggests that at least GTP-binding by Gpn2 and Gpn3 is an absolute requirement of cell viability; thus, all three GPN proteins require GTP. Based on this study (Figures 3.6) and the literature it appears that GPN1, GPN2 and GPN3 activities are all required for normal RNA polymerase II nuclear localization (Carre and Shiekhattar. 2011; Staresincic et al., 2011). While GPNs are also conserved in many archaea, a conserved function cannot be related to nuclear transport but could feasibly relate to RNAP assembly. Consistent with a distinct function for Iwr1, no significant BLAST-P hits for Iwr1 are observed in archaea (data not shown), supporting the notion that Iwr1 is most important for nuclear import and is accordingly not present in archaea. Prior to this study GPN2 and GPN3 were virtually uncharacterized in yeast and thus, our focus was accordingly on understanding their function, particularly that of GPN2. The genome instability phenotypes of the GPN mutants (Figure 3.3) could be ascribed to their role in RNA polymerase biogenesis, as defects in RNAPII subunits are known to elicit many of the same phenotypes (e.g. (Stirling et al., 2011; Stirling et al., 2012b). The causes of transcription-associated CIN are potentially diverse including RNA:DNA hybrid formation, loss of specific transcripts or defects in DNA repair (Aguilera and Garcia-Muse. 2012; Herrero and Moreno. 2011; Lagerwerf et al., 2011). In this study, reducing the levels of RNAPII subunits by using a DAmP allele was sufficient to recapitulate some of the phenotypes of the GPN2 and GPN3 mutants, suggesting that one important role for GPNs is maintaining sufficient levels of RNA polymerase subunits (Figure 3.6). The reported interaction between  	
    GPN1/XAB1 and XPA/Rad14p has not so far been observed in high-throughput yeast interaction studies or explored thoroughly in mammalian cells, although this could be another possible connection to genome integrity (Nitta et al., 2000). 3.4.2. Gpn2 and Gpn3 function is required for nuclear localization of RNA polymerase III but not RNA polymerase I The mechanism by which the GPN proteins might co-operate is not clear. Mass spectrometry studies have identified all three proteins in a complex, and direct tests have validated Gpn1-Gpn2 and Gpn2-Gpn3 interactions (Figure 3.4; (Boulon et al., 2010; Carre and Shiekhattar. 2011; Forget et al., 2010; Staresincic et al., 2011)). In this study we observed genetic interactions between gpn2-2, gpn3-1 and NPA3::DAmP alleles, also supporting a common function. Archaeal GPNs homodimerize in vitro and, while in vitro studies have suggested the capability of human Gpn1 and Gpn3 to homodimerize, we could not find evidence of a Gpn2 homodimer by co-precipitation from cell lysates (Figure 3.4; Carre and Shiekhattar, 2011; Gras et al., 2007). Indeed, the question of why virtually all eukaryotes have retained three GPN genes remains unclear. While an appealing model would propose that each GPN is responsible for a different RNA polymerase (i.e. three essential GPNs for three essential RNAPs), our results do not support this idea. GPN2 and GPN3 mutants displayed defects in both RNAPII and III localization but did not mislocalize RNAPI subunits (Figure 3.10-3.14 and Appendix Figure A1). While this could suggest that GPNs truly have no role in RNAPI assembly, it is possible that our hypomorphic alleles simply did not perturb some specific aspect of GPN function required for RNAPI assembly. Moreover, the role of NPA3/GPN1 in  	
    localization of RNAPI and III has not been thoroughly assessed and could affect RNAPI. Finally, the assembly of RNAPI appears to be fundamentally different from RNAPII as individual subunits are brought to the rDNA prior to assembly (Dundr et al., 2002). Therefore, GPNs may play a role in assembly within the nucleolus itself or be responsible for the localization of single subunits or subcomplexes not assessed in the present study. The case may be similar for IWR1 mutants, as we showed defects in RNAPIII localization but not RNAPI in the absence of Iwr1 (Figure 3.11). In the literature single GFP markers of RNAPI and III used previously did not mislocalize in IWR1 mutants (Czeko et al., 2011), whereas other data suggest that Iwr1 functions in initiation of transcription for all three nuclear RNA polymerases (Esberg et al., 2011). Some of these discrepancies could be due to the choice of specific assay, which may be important for detecting defects in RNA polymerases with different assembly pathways. 3.4.3. The GPN proteins appear to function upstream of Iwr1 in RNA polymerase II biogenesis These data together with the literature support a model in which all three GPNs act upstream of Iwr1 to assemble RNA polymerase II and III (Figure 3.17). The inability of Rpb3-NLS fusions to rescue GPN mutants, while partially rescuing the fitness and localization defects in iwr1Δ suggests that the role of GPNs is predominantly upstream of import at the level of RNA polymerase assembly, which supports and extends the previous model for RNA polymerase II biogenesis (Wild and Cramer. 2012). We consistently found that Rpb3 was unstable suggesting it was not undergoing normal biogenesis in GPN mutants; however, based on the data  	
    collected we cannot rule out that this is due to reduced transcription (Figure 3). To perform an assembly and stability function, the GPNs are likely acting as part of a network of molecular chaperones known to interact with RNA polymerases. In particular, Hsp90 and the R2TP complex have been shown to play a role, in complex with the GPNs, in assembling RNA polymerases (Boulon et al., 2010). Interestingly, there are also potential physical connections to the CCT (ChaperoninContaining Tailless complex polypeptide 1) complex which is primarily involved in folding actin and tubulin subunits. However, many other less abundant proteins have been implicated as CCT interactors including RNAPII and RNAPIII subunits (Dekker et al., 2008; Yam et al., 2008). Notably, tubulin seems to be important for RNA polymerase assembly and the CCT complex is known to co-operate with the hexameric Prefoldin chaperone, subunits of which are found in R2TP, which has been proposed to play a role in RNAP assembly (Forget et al., 2010; Vainberg et al., 1998). The potentially elaborate chaperone pathway for RNA polymerase assembly is not unprecedented; for example, tubulin heterodimer assembly requires at least eight chaperones and cofactors (reviewed in Lundin et al., 2010). The unbiased cytological screen of the nuclear proteome for additional substrates of Gpn2 action identified primarily RNA polymerase subunits and factors associated with transcription and RNA processing. This indicates that Gpn function is probably very focused on RNA polymerase assembly, with secondary mislocalizations due to physical interactions with mislocalized RNA polymerase subunits. As shown in Figure 3.14, the mislocalization phenotype is often subtle compared to the RNA polymerase subunits themselves, such as that seen for the  	
    elongation factor Elf1. However, this sequestration of these protein factors in the cytoplasm may have consequences for transcription and mRNA processing.  Figure 3.15 Model for GPN1, 2 and 3 function in biogenesis of RNA polymerases II and III. After synthesis RNA polymerase subcomplexes begin to assemble in the cytoplasm. The three GPN proteins bind with the assembly intermediates and aid the formation of the RNA polymerase complex, possibly cooperating with the Hsp90/CCT chaperone machinery. Iwr1 association with the RNA polymerases provides the NLS required for import through the nuclear pore complex (NPC). Once in the nucleus, the RNA polymerase is able to associate with transcription factors (TF) and carry out transcription activity while the GPN proteins and Iwr1 are recycled from the nucleus via a nuclear export pathway, potentially driven by a nuclear export sequence in Gpn1 (Reyes-Pardo et al., 2012).  There are several outstanding questions regarding the cellular role of the GPN proteins: Given their common phenotypes, what specialized roles do each GPN play that makes all three genes essential? Is the substrate repertoire of the GPNs for protein complex assembly limited to nuclear RNA polymerases and  	
    transcription-associated proteins? More broadly, the existence of the GPN and Iwr1 system suggests that regulated assembly and transport of RNAPs is important for proper functioning or regulation of transcription since RNAPs do not encode NLSs in their primary sequence.  	
    Chapter 4. Concluding remarks and future directions 4.1 Expanding our understanding of contributing factors to DNA damage 	
   In attempting to understand CIN mechanisms in yeast mutants, the work described in this thesis extends the previous screening of Rad52 foci done using the non-essential yeast deletion collection (Alvaro et al., 2007). This previous work primarily identified genes directly involved in DNA replication and repair, mitochondrial function, chromosome segregation and chromatin remodeling. However, since retention of genomic integrity is essential for cell viability, screening for DNA damage only in non-essential genes has the potential to exclude important contributing factors.  Through screening of the essential CIN mutants, the work  described in Chapter 2 adds to the known yeast genes that can be mutated to promote spontaneous DNA damage in S. cerevisiae, and by extension, may identify conserved genes with similar roles in higher eukaryotes. The detailed examination of Rad52-foci formation as a secondary screen of the set of CIN mutants is a rational approach, since DNA repair and CIN are relevant to cancer. It will be important to extend the Rad52-foci screen to all the remaining essential genes in the future. Indeed, Li et al. screened the yeast community ts collection for DNA damage marked by Ddc2-GFP, another marker of DNA damage (Li et al., 2011), primarily identifying damage in DNA replication and repair mutants.  A complete  documentation of all mutants carrying mutations in essential genes that cause acquisition  of  spontaneous  damage  could  be  particularly  informative  for  uncharacterized essential genes with as yet unknown roles in genome stability.  	
    It is not surprising that a significant proportion of the top hits from this Rad52 foci screen are directly involved in genome maintenance, such as DNA replication, mitosis, or proteasome function (Fig 2.3). However, it was striking that a group of mutants involved in a protein complex that mediates 3’-end processing of mRNAs should all form high levels of foci, and that additional transcription mutants were identified as well. As shown in Figure 2.1 and Table 2.2, transcription and mRNA processing mutants are sensitized to DNA damage and, although these phenotypes may be due to indirect effects such as reduced expression of DNA repair genes, the data here suggest a direct mechanism of DNA damage that includes a requirement for homologous recombination-mediated DNA repair in these mutants (Figure 2.4).  4.2 ChIP-chip identifies enrichment of genes adjacent to replication origins One issue that all cells must deal with is the potential for collision between the replication fork and the transcription machinery, an event which may result in defective DNA replication and possibly DNA damage (Boubakri et al., 2010). As shown in Chapter 2, ChIP-chip analysis is a useful tool for identifying DNA damage sites in mutants enriched for Rad52-foci (Szilard et al., 2010, Stirling et al., 2012). By using an antibody against phosphorylated histone H2A as a marker of DNA damage in ChIP-chip analysis, we discovered that mCP mutants acquire DNA damage at sites of transcribed genes adjacent to origins of DNA replication (Figure 2.5- 2.7). The immediate hypothesis was that defects in the mRNA processing machinery were promoting stalling of the RNA polymerase and consequent fork collision.  	
    Previous studies had indicated that R-loop formation contributes to  96	
    spontaneous DNA damage in cells with defective mRNA processing (Li et al., 2007; Xiao et al., 2007), therefore R-loop formation seemed a possible contributing factor in these yeast mutants.  4.3. R-loop formation as a mechanism for DNA damage in mRNA processing mutants. Subsequent analysis of R-loop formation in chromosome spreads of mRNA processing mutants showed that the pcf11 and clp1 mutants, which were analyzed using phospho-H2A ChIP-chip, also acquired elevated levels of RNA:DNA hybrids (Figure 2.9). As discussed in Chapter 1, mutations in the mRNA export factor THO cause both an elevated level of R-loop formation and a hyper-recombination phenotype.  The mCP mutants in this study also exhibit hyper-recombination in  addition to negative-genetic interactions with THO mutants (Figure 2.10 and 2.11), providing additional evidence for R-loop formation contributing to genomic instability in mCP mutants. A recent study by Wahba et al. supports the results observed in this thesis (Wahba et al., 2011) as yeast RNA biogenesis mutants displayed increased rates of gross chromosomal rearrangements which could be suppressed through overexpression of RNaseH. Further investigation into the genome instability phenotype of a transcriptional regulator, SIN3, provided evidence that sin3Δ mutants form R-loops and elevated levels of Rad52 foci, both of which could be suppressed through overexpression of RNase H (Wahba et al., 2011). Based on these data, a likely mechanism for the DNA damage phenotype observed for the mCP mutants is R-loop formation, which stalls RNA polymerases leading to collisions with the replication fork, and the subsequent requirement for DNA repair.  	
    Clearly R-loop formation is a deleterious state that can be caused by numerous players in the production and processing of mRNA. A possible future line of investigation would be to use the RNA:DNA hybrid immunofluorescence method to extend screening for R-loops in other yeast CIN mutants, particularly uncharacterized mutants that are also sensitive to DNA damaging agents. Alternatively, a screen for ts or drug sensitive mutants that respond positively to RNase H overexpression could be a high-throughput method for identifying R-loop forming mutants. These approaches have the potential to uncover new mRNA processing genes and mechanisms underlying CIN and DNA damage in yeast mutants. A project currently underway in our laboratory is the application of the anti RNA:DNA hybrid antibody in a ChIP-chip based approach to determine the sites of R-loops in the genome.  Assuming that R-loops are preserved during the  crosslinking and chromatin fractionation steps, it would be informative to determine if R-loop formation correlates with DNA damage and transcription, and to assess changes in the distribution of R-loops in a variety of mutant backgrounds.  4.4 RNA processing mutations and human disease 	
   Mis-regulation of mRNA biogenesis is a potential contributing factor to CIN. One of the mCP mutants described in Chapter 2 is fip1-ts, a mutant variant of the Fip1 protein that normally interacts with poly-A polymerase during mRNA cleavage and polyadenylation.  The human orthologue of FIP1, FIP1L1 is found as a  truncated fusion protein in 10-20% of of hyper-eosinophilic syndrome/ eosinophilic leukemias (Cools et al., 2003, Gotlib and Cools, 2008). In these cases, the Nterminus of FIP1L1 is fused to the tyrosine kinase domain of PDGFRα, deregulating  	
    PDGFRα activity, which affects cell proliferation. Indeed truncation of FIP1 results in a CIN phenotype and R-loop formation in yeast, suggesting that the truncated form of FIP1L1 in human leukemias may also affect genome integrity (Stirling et al., 2012). It is worth noting that the expression of aberrantly spliced isoforms of proteins required for genome stability or growth regulation are observed in human cancer cell lines (Li et al., 2007, Wang et al., 2011, Sze et al., 2008). Genetic alteration of splicing and other mRNA processing factors is observed in human cancers as shown in Table 4.1, which was generated using cancer genome data from the cBio Cancer Genomics Portal (http://cbioportal.org, Cerami et al., 2012). Whether these somatic mutations cause CIN, and if so, whether they exert their effects via direct or indirect mechanisms on chromosomal DNA is of great interest.  	
    Table 4.1 Frequency of mutation and copy number changes in select human tumors using data from the cBio Cancer Genomics Portal for 18 human splicing gene orthologs to yeast CIN genes. The percentage of cancers with at least one alteration is in the gene set listed is presented below. 	
   Gene  Breast invasive carcinoma  HNRPNC SRSF1  Colon and Rectum Adenocarcinoma  Lung squamous cell carcinoma  Ovarian Serous Cystadenocarcinoma  <1%  0%  2%  5%  5%  0%  4%  1%  SF1  <1%  3%  1%  1%  SNORD84  <1%  <1%  <1%  2%  TXNL4A  1%  <1%  3%  2%  SNRPC  <1%  1%  <1%  2%  SART1  1%  <1%  3%  <1%  AAR2  2%  15%  2%  2%  SNRPNP70  <1%  2%  2%  <1%  ZMAT2  <1%  <1%  3%  <1%  PRPF31  2%  1%  <1%  3%  PRPF4  1%  2%  <1%  1%  PRPF6  4%  9%  3%  9%  SF3B1  2%  4%  4%  3%  NHP2L1  <1%  0%  0%  <1%  XAB2  <1%  <1%  2%  3%  EFTUD2  <1%  4%  2%  <1%  <1% 21% of 484 cases  2%  3% 31% of 179 cases  2%  TFIP11 Frequency  29% of 212 cases  31% of 316 cases  Splicing is one type of RNA processing that is already a promising target for therapeutic use. Spliceostatins, drugs based on bacterially derived compounds that target the human spliceosome (Reviewed in Bonnal et al., 2012), are being developed with much focus on targeting cancers with SF3B1 mutations. As the genetic interaction networks for more essential RNA processing mutants are determined using SGA technology (Li et al., 2011), this may reveal more potential  	
    synthetic lethal interactions that could be targeted in cancer treatment, particularly as next generation sequencing technology develops to the point where whole tumor genome re-sequencing becomes a diagnostic tool.  4.5 Mutation of the GPN family of GTPases and genomic instability 	
   The yeast CIN mutational spectrum includes RNA polymerase I, II and III subunits (Stirling et al., 2011) and mutation of the large RNAPII subunit RPB1/RPO21 results in significantly increased levels of Rad52-foci (Figure 2.3). Since transcription, mRNA processing, and RNA polymerase II turnover in response to DNA damage are all required for normal genomic integrity, it is logical that disruption of RNA polymerase assembly could compromise the genome. All three GPN GTPases have been identified as CIN mutants with GPN1 and GPN2 mutants displaying defective sister chromatid cohesion (Ben-Aroya et al. 2008, Alonso et al. 2011, Minaker et al., 2012). The CIN phenotype, together with the increased Rad52 foci levels (Figure 2.12B) and DNA damage sensitivity of the mutant alleles (Figure 3.3), suggested a possible role in the DNA replication and repair or chromosome segregation machinery. A survey of the literature for information on the human orthologs of the GPNs implied a role in RNA polymerase II nuclear import (Boulon et al., 2010). Nonetheless, an SGA screen for gpn2-2 genetic interactions did not cluster with genes in any specific pathway, despite hundreds of significant interactions (S. Minaker, unpublished observations). This represents a challenge of working with uncharacterized genes with pleiotropic function. In the context of recent studies in mammalian cells, we tested and were able to show that the GPN mutants exhibit the  	
    same RNA polymerase II mislocalization in yeast as was observed when siRNA was used to knockdown the human orthologs (Boulon et al., 2010, Carre et al., 2011). In Chapter 3 it was shown that mutation of GPN2 and GPN3 leads to depletion of Rpb3 and the RPB3:DAmP allele exhibits DNA damage sensitivities similar to the GPN mutants.  The GPN mutants studied here do not form R-loops (YA Chan,  unpublished observation) so DNA damage resulting from defective transcription of DNA repair genes seems a plausible explanation. GST pull-down experiments suggest Rpb3 interacts with multiple subunits of the mammalian RNA polymerase (Acker et al., 1997); therefore, depletion of Rpb3 likely results in fewer functional RNA polymerase complexes.  Moreover, transcript level analysis of GPN1 mutants  suggest that they exert global effects on transcription (Staresincic et al., 2011).  4.6 The functional relationship between GPN2 and GPN3 proteins 	
   GPN2 was the first CIN GPN family member to be identified and much of the early work focused in deciphering its molecular function. As shown in Figure 3.4, Gpn2 physically interacts with both Gpn1 and Gpn3 but does not appear to homodimerize, unlike the Archaeal GPN protein that exists as homodimers, at least in vitro (Gras et al., 2007). It remains unclear whether all three GPN proteins form a heterotrimer or if Gpn2 binds either or both of the paralogues as heterodimers. Future work on the GPN family could also determine the stoichiometry of the GPN complex. Since the GPN proteins are conserved as three members throughout Eukarya and as a single member in Archaea, they likely evolved through gene duplication events. Notably, the coiled-coil domain in Gpn2 in Figure 3.2 is only found in fungal species, implying a fungal specific role in Gpn2 function.  	
    It appears that GTP binding is essential for Gpn2 and Gpn3 function as point mutants at a key aspartic acid residue required for GTP binding were non-functional, leading to spore death (Figure 3.8). However, mutants in a conserved glutamine residue in either Gpn2 or Gpn3 (reportedly required for GTP hydrolysis) were able to support cell viability. One possibility is that GTP binding by all GPN proteins is required but, GTP hydrolysis is not required by all three family members as long as GTPase activity was retained by some of the family members. Alternatively, there could be structural differences between the Gpn2 and Gpn3 proteins and the Ras GTPase used as a model for mutagenesis (Staresincic et al., 2011) such that the mutated glutamine does not play an essential role in GTP hydrolysis in the GPN proteins.  There does not appear to be overlapping function between the GPN  proteins (Figure 3.6) as overexpression of any one GPN could not provide strong suppression of the growth or RNA polymerase localization defects in gpn2 mutants. Perhaps the GPNs have essential secondary functions unrelated to RNA polymerase assembly, acting independently of the other family members.  In  Chapter 3, observed synthetic lethality and severe growth defects in GPN double mutants was shown, but future investigations using double and possibly triple mutants for GTP hydrolysis could help elucidate the requirement for GTPase activity in these proteins.  4.7 RNA polymerase III assembly and nuclear import requires both the GPN proteins and Iwr1 An early hypothesis was that each GPN was specific to a different RNA polymerase, thus explaining why three GPN proteins had evolved. This does not  	
    appear to be the case, but the effect of GPN mutations on RNA polymerase III assembly is quite apparent. Since the iwr1Δ mutant also accumulated GFP tagged RNAPIII subunits in the cytoplasm, and the localization of RNAPIII is dependent on the presence of the NLS, it is possible that the GPNs and Iwr1 mediate RNAPIII assembly and subsequent nuclear import in a similar fashion to RNAPII. Notably, no RNAPI mislocalization phenotype was observed in any of the GPN or iwr1Δ mutants examined here, possibly resulting from a different assembly mechanism (Dundr et al., 2002). Since no conditional mutants for GPN1 were available for this study, perhaps future investigations will reveal a function for GPN1 in RNAPI biogenesis. Alternatively, the GPN proteins may still play a functional role in RNAPI assembly, but this may occur within the nucleus and would not be obvious using the GFP microscopy techniques applied here. A physical interaction between Gpn2 and RNAPIII was not observed in this study, despite attempts at pull-downs using several tagged RNAPIII subunits (data not shown).  This was not unexpected as the first observation of a physical  interaction between RNAPII assembly intermediates and the mammalian GPN proteins required the specific condition of α-amanitin treatment (Boulon et al., 2010). GPN binding to RNAPIII could be transient and not retained under the conditions used for the pull-downs in these experiments. Moreover, protein interaction studies have identified few validated physical interactions for the GPN proteins (Krogan et al., 2006). Future improvements in protein interaction screening that allow for better detection of weak or transient interactions would be most informative in determining the spectrum of GPN physical interactions. Alternatively, in vivo methods such as  	
    the protein complementation assay (Remy and Michnick, 2004, Ear and Michnick, 2009) could be used to screen for physical interactions between the GPN proteins and RNAPIII, assuming that the GPN proteins bind directly and not through an intermediate protein.  4.8 Extending the function of the GPN proteins beyond RNA polymerase assembly Once the RNAP mislocalization phenotype was verified in the GPN mutants, a major question was whether they affected the assembly/nuclear localization of other proteins. In order to answer this question, we took advantage of the GFP collection to screen all proteins with annotated nuclear localization for changes in localization in the gpn2-2 mutant background. The results of this screen imply that the primary role of Gpn2 is RNAPII and RNAPIII assembly but it was notable that the localizations of some other mRNA biogenesis proteins were affected (Figure 3.15, Table 3.2).  This could be due to the accumulation of RNAPII subunits in the  cytoplasm resulting in inappropriate sequestration of factors that would normally associate with the RNA polymerase in the nucleus. It cannot be ruled out that the transcriptional effects of depleted RNAPII also affect the expression and localization of other proteins identified in this mislocalization screen. Notably, there was no enrichment for mislocalized DNA replication or repair complexes, observations that support that a DNA damage sensitivity phenotype resulting from defects in mRNA biogenesis. RNA polymerase II turnover in response to DNA damage is an important feature of the UV induced DNA repair pathway (discussed in 1.7). The nuclear  	
    GFP-tagged protein screen suggests that Rad6, an E2 ligase associated with DNA repair, was mislocalized in the gpn2-2 mutant background, and furthermore, rad6Δ mutants are hypersensitive to UV exposure (Game and Kaufman, 1999). Since human Gpn1 interacts with the UV repair protein XPA1 (Nitta et al., 2000), there may be a link between GPN function and repair of UV induced damage. Perhaps Gpn2 mediates shuttling of Rad6 into the nucleus with newly assembled RNA polymerase  II  subunits  in  order  to  mediate  transcription  coupled  NER.  Mislocalization of the yeast XPA1 ortholog Rad14 was not observed in gpn2-2 mutants, but this may only occur in GPN1 mutants if the physical interaction between the orthologs is conserved. Future work on assembly and import of RNA polymerases should focus on identifying other factors required for nuclear import as it is clear that in both GPN and iwr1Δ mutants, RNA polymerase still enters the nucleus.  This would include  identifying a karyopherin that mediates nuclear import of the assembled complexes. Additionally, there is no clear homologue of IWR1 in mammals, but perhaps a yet undiscovered protein performs an analogous function in RNA polymerase import.  4.9 Concluding remarks 	
   The work presented in this thesis increases our understanding of the mechanisms of CIN using yeast as a model system and describes previously unobserved  phenotypes  associated  with  new  temperature-sensitive  mRNA  processing mutants. In addition, newly described mutants of the GPN protein family were shown to be essential for biogenesis of RNA polymerases II and III, with genomic instability possibly resulting from faulty RNA polymerase assembly. The  	
    majority of the genes described here are conserved in higher eukaryotes; therefore, the research presented here could have implications for understanding mechanisms of human diseases associated with genome instability and/or faulty regulation of transcription and mRNA biogenesis.  	
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    Figure A1 Transcriptional frequency of ORFs associated with mCP mutant-induced g-H2A signal in their 3’-UTRs (i.e. g-sites covering >50% of 500bp window 3’ of genes in 3 of 4 mCP mutant replicates). A significant (*p<0.01) decrease in repressed genes (<1mRNA/hr) and concomitant increase in genes with high transcriptional frequencies (16-50 mRNA/hr) is observed in those genes with mCP mutant-induced γ-H2A enrichment in their 3’-UTRs.  	
   Figure A2 Rpa135-GFP is not mislocalized in GPN2, GPN3 or IWR1 mutants. Representative GFP and DIC micrographs of Rpa135-GFP in the indicated background. Below, quantification of nuclear/cytoplasmic GFP ratio indicates normal or increased levels (i.e. in the case of gpn2-2) of Rpa135-GFP in the nucleus. The asterisk indicates the results of Tukey’s posthoc analysis of a one-way ANOVA are α=0.01.  	
    Figure A3 Effect of IWR1-NLS mutation on RNA Polymerase II and III subunits. IWR1 deletion strains bearing the indicated GFP fusion proteins (colored bars) and transformed with the indicated plasmids (along the X-axis) were imaged and quantified as in Figure 3.10. α indicates a significant difference by ANOVA between the plasmid expressing WT IWR1 and all other groups.  	
    Table A1 Yeast strains used in Chapter 2.  	
   Strain Number Screening genotype for KanMX marked alleles  Relevant Genotype  Screening genotype for URA3 marked alleles  MATa can1Δ::MFA1pr-HIS3::LEU2	
  met15Δ0 or MET15	
   RAD52-YFP::KanMX  BY4741  MATa ura3Δ0	
  met15Δ0 MATα can1Δ mfa1Δ::MFA1pr-HIS3 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2  SB1  MATa can1Δ::STE2pr-SpHis5 lyp1Δ	
  RAD52-YFP::URA3 yfeg::KanMX  Jef Boeke Ben-aroya et al., 2008  YPH1015 Y10540  (w303) MATa hta1-­‐S129A	
  trp1-1 can1100 RAD5+ MATa ura3-52 lys2-801 ade2-101 leu2Δ1 his3Δ200 CFIII(CEN3.L)::HIS3 SUP11 ctf13-30 MATα	
  lys2Δ0 rad52Δ::KanMX  YPH2076  MATa ura3Δ0	
  pcf11-2::KanMX  Li et al., 2011  YPH2077  MATa can1Δ::MFA1pr-HIS3::LEU2	
  met15Δ0 lys2Δ	
  clp1-­‐ts::URA3  Ben-aroya et al., 2008  YPH2078  MATa leu2Δ0	
  pol31-ts::URA3 rad52Δ::KanMX  This Study  YPH2079  MATa leu2Δ0	
  sld7-ts::URA3 rad52Δ::KanMX  This Study  YPH2080  MATα	
  lys2Δ0 rad52Δ::KanMX rpn5-1::KanMX  This Study  YPH2081  MATa leu2Δ0	
  orc6-ts::URA3 rad52Δ::KanMX  This Study  YPH2082  MATα	
  lys2Δ0 rad52Δ::KanMX rna15-58::KanMX  This Study  YPH2083  MATα	
  lys2Δ0 rad52Δ::KanMX ssl1  ::KanMX  This Study  YPH2084  MATa ura3Δ0	
  pcf11-2::KanMX rad52Δ::KanMX  This Study  YPH2085  MATa leu2Δ0	
  nse3-ts::URA3 rad52Δ::KanMX  This Study  YPH2086  MATa ura3Δ0	
  sec12-4::KanMX rad52Δ::KanMX MATa his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 pcf112::KanMX MATa his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 pcf1110::KanMX MATa his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 rna1558::KanMX  This Study  yDD1793  YPH2087 YPH2088 YPH2089  	
    Source  T242I  Daniel Durocher This Study Open Biosystems  This Study This Study This Study  129	
    Strain Number  Source  YPH2093 YPH2094 YPH2095 YPH2096 YPH2097 YPH2098 YPH2099  Relevant Genotype MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 clp1-ts::URA3 MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 fip1-ts::URA3 MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 ade2-101::NatMX CFVII(RAD2.d)::LYS2 cft2-ts::URA3 MATα his3Δ1 ura3Δ0 leu2Δ0 pcf11-2::KanMX MATα his3Δ1 ura3Δ0 leu2Δ0 pcf11-10::KanMX MATα his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 rna15-58::KanMX MATα his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 clp1-ts::URA3 MATα can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 fip1-ts::URA3 MATα can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 cft2-ts::URA3 MATa/α ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 pcf11-2::KanMX/pcf11-2::KanMX  YPH2100  MATa/α ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 pcf11-10::KanMX/pcf11-10::KanMX  This Study  YPH2090 YPH2091 YPH2092  YPH2101 YPH2102 YPH2103 YPH2104 YPH2105 YPH2106 YPH2107 YPH2108 YPH2109 YPH2110 YPH2111 YPH2112 YPH2113 YPH2114 YPH2115  	
    MATa/α ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 lys2Δ0/LYS2 rna1558::KanMX/rna15-58::KanMX MATa/α can1Δ::MFA1pr-HIS3::LEU2 ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 lys2Δ0/lys2Δ0 clp1-ts::URA3/clp1-ts::URA3 MATa/α can1Δ::MFA1pr-HIS3::LEU2 ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 lys2Δ0/lys2Δ0 fip1-ts::URA3/fip1-ts::URA3 MATa/α can1Δ::MFA1pr-HIS3::LEU2 ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 his3Δ1/his3Δ1 lys2Δ0/LYS2 cft2-ts::URA3/cft2-ts::URA3 MATa his3Δ1 ura3Δ0 leu2Δ0 pcf11-2::NatMX MATa his3Δ1 ura3Δ0 leu2Δ0 pcf11-10::NatMX MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 cft2-ts::URA3 MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 clp1-ts::URA3 MATa can1Δ::MFA1pr-HIS3::LEU2 his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0 fip1-ts::URA3 MATa his3Δ1 ura3Δ0 leu2Δ0 rna15-58::NatMX MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX pcf11-2::NatMX MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX pcf11-10::NatMX MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX lys2Δ0 clp1-ts::URA3  This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study  This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study This Study  130	
    Strain Number YPH2116 YPH2117 YPH2118 YPH2119  	
    Relevant Genotype MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX can1Δ::MFA1prHIS3::LEU2 fip1-ts::URA3 MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX lys2Δ0 cft2-ts::URA3 MATα ura3Δ0 leu2Δ0 his3Δ1 rnh1Δ::KanMX rnh201Δ::KanMX rna15-58::NatMX MATa/α	
  trp1Δ63/trp1Δ63 lys2Δ0/LYS2 met15Δ0/MET15 FIP1-GFP::HIS3/FIP1  Source This Study This Study This Study This Study  YPH2120  MATa/α	
  trp1Δ63/trp1Δ63 lys2Δ0/LYS2 met15Δ0/MET15 1-279 fip1 -GFP::HIS3/FIP1  This Study  YPH2121  MATa/α	
  trp1Δ63/trp1Δ63 lys2Δ0/LYS2 met15Δ0/MET15 1-213 fip1 -GFP::HIS3/FIP1  This Study  YPH2122  MATa ura3Δ0	
  lys2Δ0 trp1Δ63 FIP1-GFP::HIS3  This Study  YPH2123  MATa ura3Δ0	
  lys2Δ0 trp1Δ63 fip1  This Study  YPH2124  MATa leu2Δ0	
  lys2Δ0 fip1  YPH2125  MATa leu2Δ0	
  lys2Δ0 FIP1-GFP::HIS3 ade2-101::NatMX CFIII{URA3, SUP11}  This Study  YPH2126  MATa leu2Δ0	
  lys2Δ0 fip1  This Study  YPH2127  MATa leu2Δ0	
  lys2Δ0 FIP1-GFP::HIS3 ade2-101::NatMX CFVII{URA3, SUP11}  This Study  YPH2128  MATa ura3Δ0	
  lys2Δ0 fip1  This Study  YPH2129 YPH2130 YPH2131 SB182  MATa ura3Δ0	
  lys2Δ0 FIP1-GFP::HIS3 RAD52-YFP::KanMX MATa can1Δ::MFA1pr-HIS3::LEU2 fip1-ts::URA3 ade2-101::NATMX CFIII{LYS2, SUP11} MATa can1Δ::MFA1pr-HIS3::LEU2 fip1-ts::URA3 RAD52-YFP::KANMX MATα	
  lys2Δ0 bim1Δ::KanMX  This Study This Study This Study Ben-aroya et al., 2008  SB181  MATa/α	
  lys2Δ0/LYS2 met15Δ0/MET15 chl1Δ::KanMX/chl1Δ::KanMX  Ben-aroya et al., 2008  YPH1726  MATα	
  lys2Δ0 can1ΔMFA1pr-HIS3 ade2-101::NatMX CFIII{URA3, SUP11}  Yuen et al., 2007  YPH1725  MATα	
  lys2Δ0 can1ΔMFA1pr-HIS3 ade2-101::NatMX CFVII{URA3, SUP11}  Yuen et al., 2007  YPH2230  MATα	
  lys2Δ0 rtt103Δ::KanMX  Open Biosystems  YPH2231  MATα	
  lys2Δ0 mft1Δ::KanMX  Open Biosystems  YPH2232  MATα	
  lys2Δ0 thp2Δ::KanMX  Open Biosystems  YPH2233  MATα	
  sen1-1::KanMX  Li et al., 2011  YPH2234  MATα	
  lys2Δ0 trp1Δ63 thp2Δ::KanMX  This Study  1-279  1-279  1-279  -GFP::HIS3  -GFP::HIS3 ade2-101::NatMX CFIII{URA3, SUP11} -GFP::HIS3 ade2-101::NatMX CFVII{URA3, SUP11} 1-279  -GFP::HIS3 RAD52-YFP::KanMX  This Study  131	
    Strain Number  Relevant Genotype  Source  YPH2235  MATα	
  lys2Δ0 trp1Δ63 rna15-58::KanMX  This Study  YPH2236  MATα	
  lys2Δ0 trp1Δ63 sen1-1::KanMX  This Study  YPH2237  MATα	
  his3Δ0 trp1Δ63 clp1-ts::URA3  This Study  YPH2238  MATα	
  his3Δ0 trp1Δ63 fip1-ts::URA3  This Study  YPH2239  MATα	
  lys2Δ0 trp1Δ63 rtt103Δ::KanMX  This Study  YPH2240  MATα	
  his3Δ0 trp1Δ63 pcf11-ts10::KanMX  This Study  YPH2241  MATα	
  lys2Δ0 trp1Δ63 mft1Δ::KanMX  This Study  YPH2242  MATa	
  sen1-1::KanMX rna15-58::KanMX  This Study  YPH2243  MATa	
  thp2Δ::KanMX pcf11-2::KanMX  This Study  YPH2244  MATa	
  lys2Δ0 thp2Δ::KanMX pcf11-ts10::KanMX  This Study  YPH2247  MATa	
  lys2Δ0 thp2Δ::KanMX fip1-ts::URA3  This Study  YPH2249  MATa	
  lys2Δ0 thp2Δ::KanMX rna15-58::KanMX  This Study  YPH2251  MATa	
  sen1-1::KanMX pcf11-2::KanMX  This Study  YPH2252  MATα	
  sen1-1::KanMX pcf11-ts10::KanMX  This Study  YPH2253  MATa	
  sen1-1::KanMX clp1-ts::URA3  This Study  YPH2254  MATa	
  sen1-1::KanMX fip1-ts::URA3  This Study  YPH2255  MATa	
  thp2Δ::KanMX clp1-ts::URA3  This Study  YPH2256 YPH2257 YPH2261 YPH2265  MATa	
  rtt103Δ::KanMX clp1-ts::URA3 MATα leu2Δ0 his3Δ0 ade2-101::NATMX CFIII{URA3, SUP11} sen1-1::KanMX MATα leu2Δ0 his3Δ0 lys2Δ0 ade2-101::NATMX CFIII{URA3, SUP11} thp2Δ::KanMX MATα leu2Δ0 his3Δ0 lys2Δ0 ade2-101::NATMX CFIII{URA3, SUP11} mft1Δ::KanMX  This Study This Study This Study This Study  * YFEG = an essential gene of interest (Your Favorite Essential Gene)  	
    Table A2 Yeast strains used in Chapter 3. Strain Number YPH1344 YPH1345 YPH2511 YPH2512 YPH2513 YPH2514 YPH2515 YPH2516 YPH2517 YPH2518 YPH2519 YPH2520 YPH2521 YPH2522 YPH2523 YPH2524 YPH2525 YPH2526 YPH2527 YPH2528 YPH2529 YPH2530 YPH2531 YPH2532 YPH2533 YPH2534 YPH2535 YPH2536  	
    Relevant Genotype MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 MATalpha his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 MATa/alpha leu2Δ0/leu2Δ0 ura3Δ0/ura3Δ0 GPN2-13myc::KanMX GPN2TAP::HIS3 MATa/alpha his3Δ1/his3Δ1 leu2Δ0/leu2Δ0 lys2Δ0/lys20 GPN2-13myc::KanMX TRP1::GAL1-TAP-GPN3 MATa/alpha his3Δ1/his3Δ1 ura3Δ0/ura3Δ0 leu2Δ0/leu2Δ0 lys2Δ0/lys20 TRP1::GAL1-TAP-GPN3 MATa leu2Δ0 met15Δ0 ura3Δ0 NPA3-TAP::HIS3 MATa leu2Δ0 met15Δ0 ura3Δ0 GPN2-TAP::HIS3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 GPN2-13myc::KanMX MATa his3Δ1 leu2Δ0 ura3Δ0 NPA3-TAP::HIS3 GPN2-13myc::KanMX MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-1::URA3 MATa his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-2::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-2::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-3::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-4::URA3 MATa his3Δ1 leu2Δ0 met15D0 or MET15 gpn3-1::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn3-1::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn3-9::URA3 MATalpha his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn3-10::URA3 MAT leu2Δ0 his3Δ1 met15Δ0 or MET1 ade2-101::NAT his3 lys2 CFVII(RAD2.d)::LYS2 gpn3-9::ura3 MATa leu2Δ0 his3Δ1 met15Δ0 or MET1 ade2-101::NAT his3 lys2 CFVII(RAD2.d)::LYS2 gpn3-10::ura3 MATa ade2-101::NAT met15Δ0 or MET1 his3 ura3 lys2 can1Δ mfa1Δ::MFA1prHIS3 CFVII(RAD2.d)::LYS2 MATa leu2Δ0 lys2Δ0 Smc1-GFP::HIS3GFP gpn2-2::URA3 MATalpha leu2Δ0 lys2Δ0 Ctf4-GFP::HIS3GFP gpn2-2::URA3 MATalpha leu2Δ0 his3Δ1 lys2Δ0 Swc4-GFP::KanMx gpn2-2::URA3 MATa leu2Δ0 met15Δ0 or MET15 gpn2-2::URA3 Rpb1-GFP::HIS3 MATalpha leu2Δ0 lys2Δ0 met15D0 or MET15 gpn3-1::URA3 Rpb1-GFP::HIS3 MATa leu2Δ0 met15Δ0 or MET15 iwr1Δ::KanMX Rpb1-GFP::HIS3 MATalpha leu2Δ0 lys2Δ0 met15Δ0 or MET1 gpn2-2::URA3 Rpb3-NLS::KanMX  Source Jef Boeke Jef Boeke This study This study This study Open Biosystems Open Biosystems This study This study This study This study This study This study This study This study This study This study This study This study This study Ben-Aroya et al. 2008 Open Biosystems Open Biosystems This study This study This study This study This study  133	
    Strain Number YPH2537 YPH2538 YPH2539 YPH2540 YPH2541 YPH2542 YPH2543 YPH2544 YPH2545 YPH2546 YPH2547 YPH2548 YPH2549 YPH2550 YPH2551 YPH2552 YPH2553 YPH2554 YPH2555 YPH2556 YPH2557 YPH2558 YPH2559 YPH2560 YPH2561 YPH2562 YPH2563 YPH2564  	
    Relevant Genotype Rpb1-GFP::HIS3 MATa his3Δ1 leu2Δ0 lys2Δ0 met15D0 or MET15 gpn2-2::URA3 gpn3-1::URA3 MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 gpn2-2::URA3 iwr1Δ::KanMx MATa leu2Δ0 met15Δ0 or MET15 ura3Δ0 iwr1Δ::KanMx MATa leu2Δ0 met15Δ0 or MET15 gpn3-1::URA3 Rpb1-GFP::HIS3 Rpb3NLS::KanMX MATa leu2Δ0 met15Δ0 or MET15 iwr1Δ::KanMx Rpb1-GFP::HIS3 Rpb3NLS::KanMX MATa leu2D0 RPC53-GFP::HIS3 gpn3-1::URA3 MATa leu2Δ0 met15Δ0 or MET15 gpn2-4::URA3 Rpc53-GFP::HIS3 MATa leu2Δ0 met15Δ0 or MET15 gpn3-1::URA3 Rpb3-NLS::KanMX MATa leu2Δ0 met15Δ0 or MET15 gpn2-2::URA3 Rpb3-NLS::KanMX MATa leu2Δ0 met15Δ0 or MET15 iwr1Δ::KanMX Rpb3-NLS::KanMX MATalpha leu2Δ0 ura3Δ0 met15Δ0 or MET15 Rpb1-GFP::HIS3 Rpb3NLS::KanMX MATalpha leu2Δ0 ura3Δ0 met15Δ0 lys2Δ0 or MET15 Rpb2-GFP::HIS3 Rpb3NLS::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET1 ura3Δ0 RPB3-NLS::KanMX MATa leu2Δ0 met15Δ0 or MET15 ura3Δ0 Rpb1-GFP::HIS3 MATa leu2Δ0 ura3Δ0 met15Δ0 or MET15 ura3Δ0 Rpb2-GFP::HIS3 MATa leu2D0 lys2Δ0 ura3Δ0 RPC53-GFP::HIS3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPC40-GFP::HIS3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPC37-GFP::HIS3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPB11-GFP::HIS3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPB2-GFP::HIS3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPC40-GFP::HIS3 iwr1Δ::KanMX MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPC37-GFP::HIS3 iwr1Δ::KanMX MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPB11-GFP::HIS3 iwr1Δ::KanMX MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPB2-GFP::HIS3 iwr1Δ::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ0 gpn2-2::URA3, rpc34-1::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ1 gpn2-2::URA3, rpo31698::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ2 gpn3-1::URA3, rpc34-1::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ3 gpn3-1::URA3, rpa190G728D::KanMX  Source This study This study This study This study This study This study This study This study This study This study This study This study This study Brenda Andrews Brenda Andrews Brenda Andrews Brenda Andrews Brenda Andrews Brenda Andrews Brenda Andrews This study This study This study This study This study This study This study This study  134	
    Strain Number YPH2565 YPH2566 YPH2567 YPH2568 YPH2569 YPH2570 YPH2585 YPH2586 YPH2587 YPH2588 YPH2589 YPH2590 YPH2591 YPH2592 YPH2593 YPH2594 YPH2595 YPH2596 PSY1373 PSY1374 GFP collection  Relevant Genotype MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ4 gpn3-1::URA3, rrn3S213P::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ5 gpn2-2::URA3, rpa190-1::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ6 gpn3-1::URA3, rpa190-1::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ7 gpn2-2::URA3, rrn3S213P::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ8 gpn3-1::URA3, rpo31698::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 lys2Δ9 gpn2-2::URA3, rpa190G728D::KanMX MATa leu2D0 ura3Δ0 met15Δ0 or MET15 RPO31-GFP::HIS3 MATalpha leu2Δ0 ura3Δ0 met15Δ0 lys2Δ0 or MET15 Rpo31-GFP::HIS3 Rpc40NLS::KanMX MATa leu2Δ0 lys2Δ0 met15Δ0 or MET15 gpn3-1::URA3 Rpo31-GFP::HIS3 Rpc40-NLS::KanMX MATa leu2Δ0 met15Δ0 or MET15 gpn2-2::URA3 Rpo31-GFP::HIS3 Rpc40NLS::KanMX MATa leu2Δ0 lys2Δ0 met15Δ0 or MET15 iwr1Δ::KanMX Rpo31-GFP::HIS3 Rpc40-NLS::KanMX MATalpha leu2D0 ura3Δ0 met15D0 or MET15 iwr1Δ::KanMX RPO31-GFP:HIS3 MATalpha leu2Δ0 lys2Δ0 met15Δ0 or MET15 gpn3-1::URA3 Rpo31-GFP::HIS3 MATa leu2Δ0 lys2Δ0 met15Δ0 or MET15 gpn2-2::URA3 Rpo31-GFP::HIS3 MATa/alpha leu2Δ0/leu2Δ0 ura3Δ0/ura3Δ0 gpn2Δ::KanMX/GPN2 MATa/alpha leu2Δ0/leu2Δ0 ura3Δ0/ura3Δ0 gpn3Δ::KanMX/GPN3 MATa leu2Δ0 met15Δ0 ura3Δ0 RPB3-DAmP::KanMX MATa his3Δ1 leu2Δ0 met15Δ0 or MET15 ura3Δ0 Δrad52::KanMX MATalpha leu2Δ0 his3Δ1 ura3Δ0 met15Δ0 lyp1::STE3pr-LEU2 HTA2mCherry::Hph MATalpha leu2Δ0 his3Δ1 met15Δ0 lyp1::STE3pr-LEU2 HTA2-mCherry::Hph gpn2-2::URA3 MATa leu2D0 ura3Δ0 met15Δ0 or MET15 YFNP-GFP::HIS3  Source This study This study This study This study This study This study Brenda Andrews This study This study This study This study This study This study This study Open Biosystems Jef Boeke Breslow et al. Nat. Methods 2008 Open Biosystems This study This study  *YFNP- Your Favorite Nuclear Protein **YFG – Your Favorite Gene  	
    Table A3 Plasmids used in this thesis.  Plasmid	
    LNA  LEU2-5'-LEU2-3', TRP1,	
  CEN  Andres Aguilera  LNAT  LEU2-5'-Cyc1Terminator-LEU2-3', TRP1,	
  CEN  Andres Aguilera  pRS316 (URA3) empty vector  URA3, CEN  Sikorski and Hieter, 1989  pEC069-IWR1  URA3, CEN  Patrick Cramer  pEC098-IWR1 NΔ  URA3, CEN  Patrick Cramer  pEC100-IWR1 NΔCΔ  URA3, CEN  Patrick Cramer  pGPN2-MoBY  GPN2::KanMX, URA3, CEN  Ho et al. Nat. Biotech 2009  pGPN3-MoBY  GPN3::KanMX, URA3, CEN  Ho et al. Nat. Biotech 2009  pGPN2-D106A  GPN2D106A::KanMx, URA3, CEN  This study  pGPN2-Q110L  GPN2Q110L::KanMx, URA3, CEN  This study  pGPN3-D104A  GPN3D104A::KanMx, URA3, CEN  This study  pGPN3-Q108L  GPN3Q108L::KanMx, URA3, CEN  This study  p5586 (MoBY CEN empty vector)  KanMX, URA3, CEN  Ho et al. Nat. Biotech 2009  pGPN1-MoBY  GPN1::KanMx, LEU2, 2µ  Ho et al. Nat. Biotech 2009  pGPN2-MoBY  GPN2::KanMx, LEU2, 2µ  Ho et al. Nat. Biotech 2010  pGPN3-MoBY  GPN3::KanMx, LEU2, 2µ  Ho et al. Nat. Biotech 2011  pIWR1-MoBY  IWR1::KanMx, LEU2, 2µ  Ho et al. Nat. Biotech 2012  p5587 (MoBY 2µ empty vector)  KanMx, LEU2, 2µ  Ho et al. Nat. Biotech 2013  	


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