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Coordinated activities of chromatin modifying complexes in DNA regulation in Saccharomyces cerevisiae Lu, Ya Ting Phoebe 2015

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 COORDINATED ACTIVITIES OF CHROMATIN MODIFYING COMPLEXES IN DNA REGULATION IN SACCHAROMYCES CEREVISIAE by  Ya Ting Phoebe Lu  B.SC., The University of British Columbia, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Genetics)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   March 2015  © Ya Ting Phoebe Lu, 2015   ii  Abstract   As the sole carrier of genetic information, DNA does not exist as a naked template in the eukaryotic genome; instead DNA exists as a nucleoprotein complex. Wrapped around specialized histone proteins, DNA and histones are assembled into the organized chromatin structure. The chromatin structure is regulated by a wide assortment of factors to facilitate access to the genetic material while at the same time compacting the genome. Perhaps it is not surprising that the coordinated activities of multiple chromatin-modifying complexes are necessary to regulate the many biological process occurring on DNA. We are beginning to understand how chromatin-modifying mechanisms cooperate to regulate gene promoters and define unique chromatin neighbourhoods. In this dissertation, I explore the functional connections between chromatin remodeling complexes, histone chaperones and histone variants in various aspects of chromatin biology. Focusing on the synthetic genetic interaction between ASF1 and YAF9, I uncovered the cooperative activities of histone chaperone and the SWR1-C H2A.Z exchange complex in maintaining heterochromatin boundaries. Furthermore, I also identified that H2A.Z occupancy at gene promoters is partially dependent on Asf1-mediated H3K56 acetylation. I specifically studied the functional relationship between H3K56 acetylation and H2A.Z occupancy. I determined that acetylation of H3K56 was required for maintaining H2A.Z levels at gene promoters. Furthermore, I discovered that H3K56 acetylation was also important in positioning H2A.Z containing nucleosomes at promoters. Lastly, I explored the specific features required for NuA4 structure and histone acetylation function. I uncovered a novel regulatory relationship between the Eaf1 scaffold protein and the Epl1 C-terminus that   iii  anchors the catalytic module to the NuA4 complex. In addition, I demonstrated that the Epl1 C-terminus and the Eaf1 HSA domain are the two key domains regulating the cellular equilibrium of NuA4 and picNuA4. Collectively, the work presented in the dissertation adds to our understanding of the interface between chromatin remodeling complexes, histone chaperones and histone variants in the regulation of chromatin biology and highlights the important role chromatin structure plays in basic cellular processes.     iv  Preface   Chapter 1 of this dissertation was published in part in the journal of Biochemistry and Cell Biology as a mini-review in 2009 (Lu PYT, Levesque N, and Kobor MS, 2009). As co-first author, I analyzed the large-scale genetic interaction and together with Nancy Levesque, we designed the outline of the review and wrote the manuscript.   Chapter 2 of this dissertation was published in whole in Genetics (Lu PYT and Kobor MS 2014). As first author, I generated the strains, designed and conducted all the experiments in this manuscript. I was responsible for making the figures and writing the paper.    Chapter 3 is based on experiments following up the work published in Chapter 2. I constructed the strains and conducted the biochemical and ChIP-on-chip experiments for this chapter. I analyzed the microarray data and wrote the algorithms used to examine the localization of promoter-associated H2A.Z peaks.    I currently lead the project described in Chapter 4, which includes contributions from Maria Aristizabal and Nancy Levesque. I provided primary contributions to the formation of research questions, research design, data collection and data analysis. Maria Aristizabal performed the E-MAP screen in collaboration with N. Krogan (UCSF) and F. Holstege laboratory (U of Utrecht) carried out the microarray expression experiments. I constructed the strains and performed the biochemical characterization of NuA4 for Figures 4.1, 4.4, 4.5, 4.6 and contributed to all figures.    v   In Chapter 2-4, I consistently use “we” to reflect the co-authors that contributed to the studies.    vi  Table of Contents   Abstract .......................................................................................................................................... ii	  Preface ........................................................................................................................................... iv	  Table of Contents ......................................................................................................................... vi	  List of Tables .................................................................................................................................. x	  List of Figures ............................................................................................................................... xi	  List of Abbreviations ................................................................................................................. xiii	  Acknowledgements ...................................................................................................................... xv	  Dedication .................................................................................................................................. xvii	  Chapter 1: Introduction ................................................................................................................1	  1.1	   Chromatin Structure ........................................................................................................... 2	  1.2	   Histone Chaperones in Nucleosome Assembly and Disassembly ..................................... 4	  1.2.1	   Asf1 Histone Chaperone ............................................................................................. 6	  1.3	   Nucleosome Positioning .................................................................................................... 7	  1.4	   Modifications of the Chromatin Structure ......................................................................... 9	  1.4.1	   Post-Translational Modification of Histones ............................................................ 10	  1.4.2	   ATP-Dependent Chromatin-Remodeling Complexes .............................................. 12	  1.4.3	   Histone Variants ........................................................................................................ 13	  1.5	   Coordinated Activities of Chromatin-Modifying Factors ................................................ 15	  1.5.1	   SWR1-C Structure and Function .............................................................................. 17	  1.5.2	   NuA4 Structure and Function ................................................................................... 18	    vii  1.5.3	   H2A.Z at the Intersection between SWR1-C and NuA4 .......................................... 20	  1.6	   Conservation of NuA4 and SWR1-C in Mammals .......................................................... 23	  1.7	   Summary .......................................................................................................................... 24	  Chapter 2: Maintenance of Heterochromatin Boundary and Nucleosome Composition at Promoters by the Asf1 Histone Chaperone and SWR1-C Chromatin Remodeler in Saccharomyces cerevisiae .............................................................................................................26	  2.1	   Introduction ...................................................................................................................... 26	  2.2	   Materials and Methods ..................................................................................................... 30	  2.2.1	   Yeast Strains and Plasmids ....................................................................................... 30	  2.2.2	   Growth and Genotoxic Sensitivity Assays ............................................................... 34	  2.2.3	   RT-qPCR .................................................................................................................. 34	  2.2.4	   ChIP-qPCR and ChIP-on-chip .................................................................................. 37	  2.3	   Results .............................................................................................................................. 38	  2.3.1	   ASF1 Genetically Interacted with Genes Encoding for Subunits of SWR1-C ......... 38	  2.3.2	   ASF1 and YAF9 were Required for Maintaining Heterochromatin-Proximal Gene Expression ................................................................................................................. 39	  2.3.3	   Cooperative Regulation of SIR Occupancy by Asf1 and Yaf9 was Telomere-Specific ..................................................................................................................... 42	  2.3.4	   Asf1 and SWR1-C Affected Heterochromatin-Proximal Gene Expression through Different Mechanisms ............................................................................................... 45	  2.3.5	   Dysregulation of Heterochromatin Boundaries in asf1Δ yaf9Δ was not the Underlying Cause of the Severe Growth Defects ..................................................... 47	    viii  2.3.6	   The Synergistic Genetic Interaction between ASF1 and YAF9 was Mediated via Rtt109-Dependent H3K56 Acetylation ..................................................................... 49	  2.3.7	   Asf1 and H3K56 Acetylation Promoted H2A.Z Occupancy at Subtelomeric Gene Promoters .................................................................................................................. 51	  2.4	   Discussion ........................................................................................................................ 53	  Chapter 3: H3K56 Acetylation is Required for Positioning H2A.Z and for Maintaining H2A.Z Occupancy at Gene Promoters .......................................................................................58	  3.1	   Introduction ...................................................................................................................... 58	  3.2	   Materials and Methods ..................................................................................................... 61	  3.2.1	   Yeast Strains and Plasmids ....................................................................................... 61	  3.2.2	   Growth and Genotoxic Sensitivity Assays ............................................................... 62	  3.2.3	   ChIP-qPCR and ChIP-on-chip .................................................................................. 63	  3.2.4	   Chromatin Association Assay ................................................................................... 65	  3.3	   Results .............................................................................................................................. 65	  3.3.1	   Asf1-Dependent H2A.Z Promoter Occupancy was not Mediated by the SAS Histone Acetyltransferase ....................................................................................................... 65	  3.3.2	   Loss of Asf1 did not Alter Global H2A.Z Levels in Chromatin .............................. 70	  3.3.3	   Genome-Wide H2A.Z Occupancy was Altered by Loss of H3K56 Acetylation ..... 72	  3.3.4	   Defects in H3K56 Acetylation Resulted in Reduced H2A.Z Promoter Occupancy . 76	  3.3.5	   H3K56 Acetylation Positioned H2A.Z at Gene Promoters ...................................... 78	  3.4	   Discussion ........................................................................................................................ 82	  Chapter 4: Eaf1 HSA Domain and Epl1 C-terminus Coordinate NuA4 Stability and Function ........................................................................................................................................88	    ix  4.1	   Introduction ...................................................................................................................... 88	  4.2	   Materials and Methods ..................................................................................................... 91	  4.2.1	   Yeast Strains and Plasmids ....................................................................................... 91	  4.2.2	   Growth and Genotoxic Sensitivity Assays ............................................................... 93	  4.2.3	   Large-Scale Affinity Purification ............................................................................. 93	  4.2.4	   RT-qPCR .................................................................................................................. 94	  4.2.5	   ChIP-qPCR and ChIP-on-chip .................................................................................. 95	  4.2.6	   mRNA Expression Profile ........................................................................................ 95	  4.2.7	   E-MAP ...................................................................................................................... 96	  4.2.8	   Chromatin Association Assay ................................................................................... 96	  4.3	   Results .............................................................................................................................. 97	  4.3.1	   Epl1 C-terminus Truncation Suppressed the Effects of EAF1 Deletion by Stabilizing picNuA4 Complex Formation ................................................................................... 97	  4.3.2	   Loss of Eaf1 and Epl1 C-terminus led to NuA4-Specific Defects at Ribosomal Genes ....................................................................................................................... 101	  4.3.3	   Distinct and Overlapping Cellular Roles for Eaf1 and the Epl1 C-terminus .......... 103	  4.3.4	   The Eaf1 HSA Domain was Required for NuA4 Stability and Function ............... 107	  4.3.5	   Epl1 C-terminus was Required for NuA4 Association with Chromatin ................. 111	  4.4	   Discussion ...................................................................................................................... 114	  Chapter 5: Conclusion ...............................................................................................................121	  References ...................................................................................................................................132	     x  List of Tables   Table 2.1 Yeast strains used in this study ..................................................................................... 31	  Table 2.2 Primer sequences used in this study ............................................................................. 35	  Table 3.1 Yeast strains used in this study ..................................................................................... 62	  Table 3.2 Primer sequences used in this study ............................................................................. 64	  Table 4.1 Yeast strains used in this study ..................................................................................... 92	  Table 4.2 Primers used in this study ............................................................................................. 94	     xi  List of Figures   Figure 1.1 DNA wrapped around a core of eight histones ............................................................. 3	  Figure 1.2 Sequential activity of NuA4 and SWR1-C on chromatin ........................................... 16	  Figure 1.3 Shared subunits of SWR1-C, NuA4, and INO80 ........................................................ 16	  Figure 1.4 SWR1-C Structure ....................................................................................................... 18	  Figure 1.5 NuA4 Structure ............................................................................................................ 19	  Figure 2.1 ASF1 genetically interacted with genes encoding for non-essential subunits of SWR1-C subunits ..................................................................................................................................... 39	  Figure 2.2 ASF1 and YAF9 were required for maintaining heterochromatin-proximal gene expression ..................................................................................................................................... 41	  Figure 2.3 Regulation of Sir2 occupancy by Asf1 and Yaf9 was telomere-specific .................... 45	  Figure 2.4 Asf1 regulated HMR-proximal gene expression in a Sir2-independent manner ......... 47	  Figure 2.5 Silencing defects were not causal for asf1Δ yaf9Δ synthetic growth defect ............... 49	  Figure 2.6 Synergistic genetic interaction between ASF1 and YAF9 was mediated by Rtt109-dependent H3K56 acetylation ....................................................................................................... 50	  Figure 2.7 Asf1 and H3K56 acetylation promoted H2A.Z occupancy at subtelomeric and euchromatic gene promoters ......................................................................................................... 52	  Figure 3.1 Asf1-dependent H2A.Z promoter occupancy did not require the SAS histone acetyltransferase ............................................................................................................................ 67	  Figure 3.2 Asf1-dependent H2A.Z deposition at ChrIII R was not mediated by SAS ................. 69	  Figure 3.3 Loss of Asf1 does not alter global H2A.Z levels in chromatin ................................... 71	    xii  Figure 3.4 Genome-wide H2A.Z occupancy was altered by loss of H3K56 acetylation ............. 73	  Figure 3.5 Loss of H3K56 acetylation led to an overall decrease in H2A.Z levels ...................... 75	  Figure 3.6 Defects in H3K56 acetylation resulted in reduced H2A.Z promoter occupancy ........ 77	  Figure 3.7 Loss of H3K56 acetylation resulted in a shift of H2A.Z position at gene promoters . 79	  Figure 3.8 H3K56 acetylation defects led to a shift in H2A.Z position ....................................... 81	  Figure 4.1 Epl1 C-terminus truncation suppressed the effects of EAF1 deletion by stabilizing picNuA4 complex formation ...................................................................................................... 101	  Figure 4.2 Loss of Eaf1 and Epl1 C-terminus led to NuA4-specific defects at ribosomal genes..................................................................................................................................................... 102	  Figure 4.3 Genetic and gene expression analysis revealed similarities and distinctions between eaf1 and epl1 mutants ................................................................................................................. 106	  Figure 4.4 The Eaf1 HSA domain was required for NuA4 stability and function ..................... 110	  Figure 4.5 epl1-CΔ truncation was epistatic to eaf1 internal truncation mutants ....................... 110	  Figure 4.6 Epl1 C-terminus was required for NuA4 association to chromatin .......................... 113	  Figure 4.7 Model for the regulatory mechanisms of Eaf1 and Epl1 C-terminus in picNuA4 stability ........................................................................................................................................ 119	  Figure 5.1 Model for Asf1 and SWR1-C interactions in Saccharomyces cerevisiae ................. 122	     xiii  List of Abbreviations   ATP  Adenosine triphosphate ChIP  Chromatin immunoprecipitation DDR   DNA damage response DNA   Deoxyribose nucleic acid DSB  Double stranded break E-MAP Epistatic miniarray profiling HAT  Histone acetyltransferase HDAC  Histone deacetylase HSA  Helicase SANT associated HU  Hydroxyurea INO80  Inositol 80 MMS  Methyl methane sulfonate MYST  MOZ, Ybf2/Sas3, Sas2, and Tip60 NFR  Nucleosome free region NuA4  Nucleosome acetyltransferase of H4 ORF  Open reading frame PCR  Polymerase chain reaction picNuA4 piccolo NuA4 PTM  Post-translational modification RNA  Ribonucleic acid   xiv  RSC  Remodel the structure of chromatin SAGA  Spt-Ada-Gcn5-acetyltransferase SANT  Swi3, Ada2, N-Cor and TFIIIB SRCAP Snf2-related CBP activator protein SWR1-C Swi2/Snf2-related ATPase 1 – complex TAP  Tandem affinity purification TIP60  Tat interacting protein (60kDa) TSS  Transcriptional start site    xv  Acknowledgements   I am grateful for the many people who have been a part of my life through this long and challenging journey.    Firstly, I would like to thank my supervisor, Dr. Michael Kobor for giving me the freedom to explore novel and interesting research questions as a mere undergraduate student more than eight years ago. For it was this first taste of scientific curiosity, the ability to question and test an idea that inspired me to pursue a career in scientific research. Thank you for your unwavering mentorship, guidance, and support.   I would also like to thank my supervisory committee, Dr. Carolyn Brown, Dr. Leann Howe and Dr. Stefan Taubert, for their time and guidance over the years. Thank you for always keeping me on track and for continuously challenging me. Thank you to Dr. Hugh Brock for teaching me the basics of genetics and for your continuous support in the Genetics Graduate Program. Furthermore, I am thankful for the funding that the Canadian Institute of Health Research has provided me to support my studies and for the incredible research conferences the travel grant has allowed me to attend.   Thank you to the many Kobor lab members (past, present and honorary) for keeping me sane through the long and seemingly never ending list of experiments, through the trials and tribulations of failed experiments, and the celebration of getting the perfect silver stain! Words   xvi  cannot express how grateful I am to have had the privilege of working alongside each and every one of you.  I am truly thankful of all the lasting friendship I have made over the years and I look forward to celebrating all of your successes in the future.  Finally, I would like to thank my family for their constant love and support. My mom and dad for always making sure I had everything I need and their heartfelt efforts to understand what I was doing. Thank you to my amazing sisters, Jane and Daphne, for their, sometimes, unconventional ways of being supportive and loving. Lastly, thank you to Jaques, my personal editor, my cheerleading squad and my shoulder to lean on. Thank you for always being here with me.    xvii  Dedication          This thesis is dedicated to my parents for their love, endless support and encouragement.    1 Chapter 1: Introduction   DNA is traditionally believed to be the sole carrier of genetic information that is passed down to the next generation. Confirmed more than half a century ago by groundbreaking experiments by Avery-MacLeod-McCarty and Hershey-Chase, the identification of DNA as the transforming principle was cemented. From that moment onward, an explosion of research on DNA commenced. Within ten years, the structure of the DNA double helix was solved, the genetic code was deciphered and the central dogma of molecular biology was conceived. In the eukaryotic nucleus, DNA does not exist as a naked template; instead, it is compacted into a dynamic structure known as chromatin. Specialized histones proteins facilitate the condensation and packaging of DNA into the chromatin structure, allowing vast amounts of DNA to fit within a cell nucleus only a few microns in diameter. Intense research in molecular biology over the last two decades has revealed that chromatin structure plays an immense role in controlling the accessibility of the genetic information stored within the DNA nucleotides. Furthermore, crosstalk between multiple chromatin-modifying complexes and their targets results in a dynamic chromatin landscape, which has profound influences on gene expression, DNA replication, DNA repair, and DNA recombination. Pioneering work in the field of chromatin biology recognized that the information contained in chromatin structure could also be inherited, much like DNA (Eissenberg and Reuter 2009; Girton and Johansen 2008; Ekwall et al. 1997). Thus, the field of research in epigenetics was initiated;     2 which is defined in this dissertation as the study of “the structural adaptation of chromosomal regions so as to register, signal, or perpetuate altered activity states” (Bird 2007).   1.1 Chromatin Structure Chromatin, in its very initial conception by Walther Flemming in 1880, referred to the stainable thread-like material found in the cell nucleus (Flemming 1882). Today, we recognize chromatin as an organizational structure that packages billions of nucleotides of DNA into the eukaryotic nucleus. The dynamic capacity of chromatin to “open” up its structure is vital in accessing much of the information contained within the condensed structure, allowing cells to carry out basal cellular functions and respond to the changing environment.  The fundamental unit of chromatin is the nucleosome, consisting of 147 bp of DNA wrapped 1.7 superhelical turns around a histone octamer comprising two molecules each of H2A, H2B, H3, and H4 (Luger et al. 1997). The eight canonical histones in the nucleosome core particle are small and highly basic. Each core histone is composed of a globular protein domain and a flexible tail that protrudes from the core of the nucleosome (Figure 1.1). A combination of hydrogen bonds, salt linkages and non-polar interactions between the histone core and the negatively charged phosphate backbone of DNA contribute to the overall stability of the nucleosome (Luger et al. 1997; Li and Reinberg 2011). The histone tails, which make up 20% of the octamer mass, are largely unstructured and serve as a platform for a large number of post-translational covalent modifications in addition to mediating interactions with nearby nucleosomes. For example, the basic patch in the H4 N-terminal tail     3 interacts with an acidic patch on the H2A/H2B heterodimer of a neighbouring nucleosome for compaction of a nucleosome array (Dorigo et al. 2003; Fan et al. 2004; Gordon et al. 2005; Luger et al. 1997).      Figure 1.1 DNA wrapped around a core of eight histones Schematic representation of DNA wrapped around two each of H2A, H2B, H3, and H4. The replacement of canonical histone H2A for the histone variant H2A.Z and K56acetylation on histone H3 are two major chromatin modifying mechanisms examined in this dissertation.   Nucleosome core particles are joined by sections of linker DNA, which form the iconic “beads on a string” structure (Figure 1.1) (Kornberg 1974; Olins and Olins 1974). Together, the nucleosome core particle and the linker DNA form repeating units of the chromatin polymer. The 11 nm “beads on a string” structure of chromatin is largely unfolded, and represents a transcriptionally active and accessible form of chromatin commonly referred to as euchromatin. Incorporation of the linker histone, H1, promotes condensation of the chromatin polymer into the 30 nm structure (Li and Reinberg 2011; Robinson et al. 2006; Robinson and Rhodes 2006). The precise mechanism of how the 30 nm structure is further compacted into a series of higher order structures resulting in the mitotic chromosome Figure'1'K56ace'    4 remains an active area of research. It is clear, however, that inter-fiber interactions play a large role (Li and Reinberg 2011).  Compaction of DNA into these higher structures leads to formation of the transcriptionally silent heterochromatin structure (Li and Reinberg 2011). In an interphase nucleus, the chromatin polymer exists as a dynamic equilibrium between euchromatin and heterochromatin, simultaneously allowing access to genetic information and compaction of the eukaryotic genome. We are beginning to understand the vital contributions and the dynamic regulation of this higher order chromatin organization to everyday cellular functions.   1.2 Histone Chaperones in Nucleosome Assembly and Disassembly   In eukaryotes, all biological processes involving genomic DNA occur on a chromatin template. Although the euchromatic structure promotes access, unwinding of DNA from the histone octamer is still necessary. Hence, the chromatin structure is constantly undergoing rounds of assembly and disassembly, allowing processes such as DNA replication, transcription, and DNA damage repair to occur without obstruction from the nucleosome.   Histone chaperones were initially perceived as passive histone binding proteins that facilitated histone transport into the nucleus and nucleosome assembly (Park and Luger 2008). Histone chaperones are now being recognized as key proteins that play an active role     5 in the formation of nucleosomes and contribute to the regulation of the dynamic structure of chromatin and its function. In fact, mixing DNA and free histones in the absence of their chaperones produces insoluble aggregates rather than nucleosomes. Histone chaperones can be broadly characterized as chaperones for H2A/H2B heterodimers or H3/H4 heterodimers (De Koning et al. 2007; Eitoku et al. 2008). In addition, some histone chaperones bind to variants of the canonical histones, such as the Chz1 histone chaperone that bind H2A.Z/H2B heterodimers preferentially over H2A/H2B heterodimers (Luk et al. 2007).   One important role for histones chaperones is guiding interactions between histones and DNA in the stepwise process of nucleosome assembly. In the current model, two H3/H4 heterodimers are first deposited onto DNA to form a tetrasome. Next, two pairs of H2A/H2B heterodimers are added and the DNA is wrapped around the resultant histone octamer, thus completing the nucleosome (Luger et al. 1997). During the nucleosome building process in the cell, histone chaperones are required for providing free histones for assembly, stabilizing charged histones until they are deposited onto the DNA and for promoting the correct points of contact between histones and DNA (Eitoku et al. 2008; Das et al. 2010; Ransom et al. 2010). In a similar manner, histone chaperones also coordinate every step of the nucleosome disassembly process. Additionally, histone chaperones are required for shuttling newly synthesized histones into the nucleus, promoting post-translational modification of histones, and maintaining a pool of free histone in the nucleus (Eitoku et al. 2008).       6 1.2.1 Asf1 Histone Chaperone Not surprisingly, many classes of histone chaperones exist to perform the wide range of functions they are required for in the cell. Of particular interest in this dissertation is Asf1 (Anti-Silencing Function 1), a highly conserved histone chaperone for the H3/H4 heterodimer. First discovered in 1997 in a screen looking for disruptors of telomere silencing in Saccharomyces cerevisiae, Asf1 is now one of the best characterized histone chaperones (Le et al. 1997; Singer et al. 1998). Fundamentally, Asf1 is involved in the assembly of the H3/H4 heterodimer into chromatin during replication-dependent and replication-independent nucleosome assembly, as well as nucleosome assembly and disassembly during transcription (Tyler et al. 1999; Adkins et al. 2004; 2007; Adkins and Tyler 2004; Schwabish and Struhl 2006; Mousson et al. 2007; Gkikopoulos et al. 2009; Takahata et al. 2009). In the majority of these processes, Asf1 hands off the H3/H4 heterodimer to other histone chaperones and chromatin-remodellers for nucleosome assembly. Through its cooperation with multiple chromatin-modifying complexes, Asf1 is implicated in DNA replication, DNA damage response, telomere silencing and transcription (Mousson et al. 2007; Adkins et al. 2004; Raisner and Madhani 2008; Emili et al. 2001; Schwabish and Struhl 2006; Miller et al. 2010). In addition to nucleosome assembly, Asf1 also presents histone H3 for post-translational modification. Binding of Asf1 to the H3/H4 heterodimer is required for H3K56 acetylation by the histone acetyltransferase, Rtt109 (Recht et al. 2006; Tsubota et al. 2007; Schneider et al. 2006; Driscoll et al. 2007). Ultimately, the broad functions of histone chaperones can be attributed to their intimate relationship with a large range of chromatin-modifying complexes, which combined, direct nucleosome assembly and disassembly.        7 1.3 Nucleosome Positioning   As hinted at by the regulatory function of histone chaperones, nucleosomes are not randomly scattered in the genome but are often found in discreet increments and at fixed positions. It is now clear that the positioning of nucleosomes is a tightly regulated process of critical importance to various biological processes.  High-resolution mapping of nucleosomes has revealed that nucleosomes are often depleted at enhancer, promoter and terminator regions, but enriched over genes and intergenic regions (Struhl and Segal 2013; Jiang and Pugh 2009; Lee et al. 2007; Mavrich et al. 2008; Shivaswamy et al. 2008; Whitehouse et al. 2007; Lam et al. 2008). It is apparent from these genome-wide experiments that nucleosomes are positioned in a very precise manner and that this is particularly true for genes. In the yeast genome, the majority of genes have a region just upstream of the transcriptional start site (TSS) that lacks nucleosomes, termed the nucleosome free region (NFR). The 5’ NFR is approximately 150 bp in length and is flanked by two highly positioned nucleosomes referred to as the -1 and +1 nucleosomes (Jansen and Verstrepen 2011). However, while the nucleosomes near the 5’ NFR adopt fixed positions, the position of nucleosomes becomes more random with increasing distance from the promoter. Interestingly, consistent with eviction of promoter nucleosomes by Pol II during transcription, highly expressed genes have less defined NFRs (Lee et al. 2004; Weiner et al. 2010; Cairns 2009).   Though there have been much debate regarding the underlying determinants of nucleosome positioning, collectively we now know that the combined effects of DNA sequence and chromatin-remodeling complexes are the two main factors influencing nucleosome     8 positioning in Saccharomyces cerevisiae (Struhl and Segal 2013). DNA sequence influences on nucleosome positioning are mainly attributed to the structural flexibility of the 147 bp that is wrapped around the histone octamer (Segal et al. 2006). G-C interactions are more rigid, whereas A-T interactions connected by two hydrogen bonds instead of three, offer more flexibility and facilitate DNA bending around the nucleosome. In addition, poly(dA:dT) tracts often found at gene promoters are important for nucleosome depletion and promoter accessibility (Segal and Widom 2009; Raveh-Sadka et al. 2012; Iyer and Struhl 1995).   Evidence that DNA sequence alone does not dictate nucleosome positioning comes from in vitro experiments demonstrating that chromatin-modifying complexes are required for nucleosomes to adopt their in vivo positioning patterns (Wippo et al. 2011). Furthermore, the Isw2 remodeler is required for positioning the +1 nucleosome onto unfavaourable DNA at select gene promoters in vivo (Whitehouse and Tsukiyama 2006; Whitehouse et al. 2007). The RSC remodeling complex is also important for sliding nucleosomes onto DNA sequences that are otherwise unfavorable for nucleosome assembly (Hartley and Madhani 2009). Thus, it appears that multiple chromatin remodelers in addition to the underlying DNA sequence are needed to confer specificity in establishing the position of nucleosomes. Although the precise contribution of each of these factors remains a key question in the field, there is no question that they all contribute to the positioning of nucleosomes.   The positions of promoter nucleosomes are highly regulated in part because of their ability to control the activation or repression of genes. Experiments with classical inducible genes such as PHO5 and GAL1 and genome-wide correlation experiments have demonstrated that     9 nucleosomes can regulate transcription by changing their positions (Almer and Hörz 1986; Fedor and Kornberg 1989; Lee et al. 2004; 2007). By physically blocking a transcription factor binding site or preventing docking of the PolII transcription machinery, the position of nucleosomes can indirectly control the activation of genes. Nevertheless, it is important to bear in mind that nucleosome position alone does not account for all transcriptional regulation but rather contributes to and plays an important role in the cellular dynamics of transcriptional control.   1.4 Modifications of the Chromatin Structure  In an effort to balance the need to compact DNA and the need to access genetic information contained within DNA, the chromatin structure must maintain a dynamic equilibrium of compaction and accessibility. The coordinated activities of various chromatin-modifying factors are crucial in maintaining distinct chromatin neighbourhoods, which ensure appropriate gene expression, as well as DNA replication, repair and recombination. Of the four main mechanisms regulating chromatin structures in eukaryotic cells, DNA methylation is the only alteration that does not exist in yeast. Methylation of cytosines in the context of CpG dinucleotides in gene promoter regions is generally associated with gene repression and heterochromatic regions. Another mechanism is the addition of post-translational modifications on histone proteins (Peterson and Laniel 2004; Turner 2005). Furthermore, nucleosomes can be moved in cis or in trans by ATP-dependent chromatin-remodeling complexes (Eberharter and Becker 2004; Lusser and Kadonaga 2003). Lastly, canonical histones can be replaced by histone variants to mark defined chromatin regions (Sarma and     10 Reinberg 2005; Zlatanova and Thakar 2008). The latter three mechanisms are of particular interest in this dissertation and their roles in chromatin regulation are expanded upon below.   1.4.1 Post-Translational Modification of Histones A wealth of work has established that the addition of chemical groups or short protein peptides on histones provides an additional layer of regulatory control on the chromatin structure. Largely achieved by specialized “writer” enzymes, the types of post-translational modification (PTMs) on histones include acetylation, methylation, phosphorylation, ubiquitination, sumoylation, and ADP-ribosylation (Kouzarides 2007). The majority of PTMs occur on the N-terminal tails of histones that protrude from the nucleosome core particle. These tail modifications often serve as a docking platform or a signaling beacon for downstream factors. Nonetheless, some PTMs do occur on the globular domains of histones such as acetylation of H3K56 and these generally alter the overall stability of nucleosomes or regulate inter-nucleosome stability.   The first PTM reported, acetylation of lysine residues, remains one of the best understood (Allfrey et al. 1964). Lysine acetylation of histones is a reversible modification that is achieved by the combined actions of histone acetyltransferases (HATs) and histone deacetylases (HDACs). Characterized by a conserved acetyl-coA-factor binding domain, HATs catalyze the transfer of an acetyl group from acetyl-coA to a histone lysine residue (Li and Reinberg 2011; Shahbazian and Grunstein 2007). Acetylation of lysine neutralizes its positive charge, thereby weakening the interactions with the negatively charged DNA and resulting in a more open chromatin structure (Grunstein 1997). HATs can be classified into     11 two categories based on their substrate specificity. The first category of HATs acetylates histones within chromatin. For example, the essential NuA4 HAT of the MYST (MOZ, Ybf2, Sas2, and Tip60) family acetylates the lysine residues of nucleosomal H4, H2A, and H2A.Z (Allard et al. 1999; Eberharter et al. 1998; Babiarz et al. 2006; Doyon and Côté 2004; Keogh et al. 2006; Millar et al. 2006). These PTMs generally occur at specific chromatin locations and are critical in controlling the chromatin state at that region. For example, NuA4-targeted H4 acetylation at ribosomal gene promoters is required for the transcriptional activation of these genes (Reid et al. 2000). The second category of HATs acetylates newly synthesized histones that are associated with histone chaperones, and not nucleosomes. A fairly novel class of HATs, the activity and functions of these HATs are starting to be understood. Rtt109 is one particular example that has been extensively studied since its discovery (Masumoto et al. 2005; Han and Zhang 2007; Driscoll et al. 2007). In a mutually exclusive manner, Rtt109 requires the assistance of two histone chaperones, Vps75 and Asf1, to acetylate H3K9 and H3K56, respectively (Tsubota et al. 2007; Fillingham et al. 2008; Driscoll et al. 2007).  Furthermore, PTMs sometimes play pivotal roles in the recognition of histone chaperones for their respective substrates. For instance, H3K36me reduces the binding of H3/H4 dimer to the histone chaperone Asf1, thereby suppressing histone exchange (Venkatesh et al. 2012).  Upon the discovery of the PTMs, scientists started looking for “readers” that would recognize and make use of the modified histone peptides. The first reading modules discovered were the chromodomains that recognized methylated lysines and the bromodomains that recognized acetylated lysines residues (Dhalluin et al. 1999; Bannister et     12 al. 2001). We now know that these “readers”, together with PTMs, play key roles in mediating crosstalk between chromatin-modifying complexes and transcriptional regulators (Bannister and Kouzarides 2011). Rather than directing transcriptional activation or repression by altering chromatin structure, PTMs often serve as a signal platform to recruit downstream effectors (readers) responsible for underlying transcriptional changes. Combinatorial studies undertaken in recent years led to the proposal of a “histone code”, where akin to the genetic code, the combination of PTMs on a single nucleosome would result in a unique downstream response (Strahl and Allis 2000). While it is still up for debate whether the histone code can be generalized, emerging evidence suggests that the combinatorial reading capacity of multi-subunit chromatin-modifying complexes increases binding strength and specificity (Huh et al. 2012; Li et al. 2009). In particular, recent structural analysis of NuA4 demonstrates that the HAT complex makes multiple contacts with the entire nucleosome, suggesting that it could recognize multiple PTMs on a single nucleosome (Chittuluru et al. 2011).   1.4.2 ATP-Dependent Chromatin-Remodeling Complexes ATP-dependent chromatin-remodeling complexes are a specialized group of complexes that use the energy from ATP hydrolysis to disrupt DNA-histone contacts in order to carry out their respective functions (Cairns 2007). To facilitate crosstalk, chromatin remodelers have “reading” domain(s) that interact with other chromatin factors and recognize post-translational modifications of histones. Chromatin remodelers can be categorized into three main classes with specialized functions in either assembly of nucleosomes, restructuring the composition of nucleosomes or promoting genome access. Often working with histone     13 chaperones, the assembly class of chromatin remodelers facilitates the formation and positioning of nucleosomes. The second class of chromatin remodelers edits the compositions of nucleosomes, often replacing canonical histones with histone variants. The last class of chromatin remodeler enables access to chromatin, achieved either by sliding or ejecting the histone octamer. A major focus of this dissertation is the second class of chromatin remodelers, namely the SWR1-C ATP-dependent chromatin-remodeling complex. Part of the INO80 family of remodelers, SWR1-C alters the composition of a standard nucleosome by removal of H2A/H2B dimers and incorporation of histone variant H2A.Z/H2B dimers, resulting in the formation of unique chromatin neighbourhoods characterized by H2A.Z occupancy (Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004).   1.4.3 Histone Variants Histone variants are non-allelic variants of canonical histones encoded by unique genes. Often possessing similar primary amino acid sequences and core structures, the incorporation of histone variants in place of canonical histones creates distinct chromosome neighbourhoods that confer specialized functions. In contrast to canonical histones whose expression is cell cycle regulated and peaks at S-phase during DNA replication, expression of histone variants is not cell cycle-dependent (Talbert and Henikoff 2010; Szenker et al. 2011). Instead, their expression is tightly regulated throughout the cell cycle in order to promote their availability and deposition. Histone variants exist for all canonical histones except for histone H4 (Talbert and Henikoff 2010). The H3 family of variants tends to be less diverse whereas the H2A and H2B families of variants have substantial sequence variations.      14 Similar to canonical histones, histone variants also associate with histone chaperones to regulate histone supply and chromatin incorporation (De Koning et al. 2007). For instance, the mammalian Asf1 histone chaperone must recognize the five amino acid differences between H3.1 and H3.3; handing off H3.1/H4 dimer to the CAF-1 complex for replication-dependent nucleosome assembly and delivering H3.3/H4 dimer to the HIRA complex for replication-independent assembly (Tagami et al. 2004). Furthermore, the sequence difference within histone variants can alter the molecular interactions between the histone octamer and DNA. In particular, many lines of evidence suggest that the incorporation of the H2A.Z histone variant reduces nucleosome stability (Ishibashi et al. 2009; Abbott et al. 2001; Watanabe et al. 2013; Bönisch and Hake 2012; Jin et al. 2009). Additionally, the acidic patch that is extended in H2A.Z compared to H2A regulates chromatin compaction by promoting chromatin fiber folding (Suto et al. 2000; Fan et al. 2002; 2004; Subramanian et al. 2013). Furthermore, histone variants often have different PTMs compared to their canonical counterparts, adding another level of complexity. For example, while H2A is acetylated at lysine 4 and 7, the H2A.Z histone variant is acetylated at lysine 3, 8, 10, and 14 (Babiarz et al. 2006; Millar et al. 2006; Suka et al. 2001; Vogelauer et al. 2000; Altaf et al. 2010). The unique features within histone variants provide specific interactions with binding partners, distinct biochemical properties to nucleosomes, and influence higher-level chromatin organization. Thus, histone variants are an exciting avenue of research to understand the creation and functions of distinct chromatin neighbourhoods.        15 1.5 Coordinated Activities of Chromatin-Modifying Factors   It is clear from recent observations that chromatin-modifying factors are not exclusive, but function in parallel with other remodelers to regulate chromatin organization and function. As a major focus in this dissertation, I decided to direct my attention on three specific chromatin-modifying processes that act on nucleosomes. In Saccharomyces cerevisiae, SWR1-C and NuA4 complexes converge on H2A.Z to generate distinct chromatin neighbourhoods (Figure 1.2). NuA4 first acetylates H4, which facilitates the recruitment of SWR1-C through its bromodomain-containing subunit, Bdf1 (Ranjan et al. 2013; Altaf et al. 2010). SWR1-C uses the energy from ATP hydrolysis to exchange canonical histone H2A for the histone variant H2A.Z, hence altering nucleosome composition (Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004). Once deposited, the N-terminus of H2A.Z is acetylated by NuA4 (Babiarz et al. 2006; Keogh et al. 2006; Millar et al. 2006). Intriguingly, yeast NuA4 and SWR1-C have four subunits in common, an observation that potentially accounts for their cooperative functions (Figure 1.3) (Lu et al. 2009).                        16                                                                      Figure 1.2 Sequential activity of NuA4 and SWR1-C on chromatin Model of the sequential relationship between NuA4 and SWR1-C. 1: NuA4 is anchored to chromatin through the shared module and acetylates histone H4. 2: SWR1-C is recruited to chromatin with the help of the shared module and recognition of acetylated H4 by Bdf1. SWR1-C then deposits histone variant H2A.Z into chromatin. 3: NuA4 subsequently acetylates the newly deposited H2A.Z.                                      Figure 1.3 Shared subunits of SWR1-C, NuA4, and INO80 Venn diagram of the overlap between SWR1-C, NuA4, and INO80 subunits.   Figure'2'Ac-H4 Ac-H4 Ac-H4 Ac-H4 Ac-H4 Ac NuA4 SWR1-C H2A.Z Ac-H2A.Z SWR1-C Arp4 Swc4 Act1 Yaf9 123Arp4 Swc4 Act1 Yaf9 NuA4 NuA4 Arp4 Swc4 Act1 Yaf9 Ac Ac-H4 Ac-H4 Figure'3'Esa1'Eaf1'Epl1'Swr1'Swc2'Swc3'Swc5'Swc6'Yaf9''Swc4''Arp4'Act1'''Rvb1'Rvb2'Yng2'Eaf6'Eaf3'Eaf5'Eaf7'Tra1'Swc7'Bdf1'Arp6'Ino80'SWR1HC'H2A.Z'deposiMon'complex'NuA4'Histone'acetyltransferase'INO80'H2A.Z'removal'complex'    17 1.5.1 SWR1-C Structure and Function SWR1-C was the first complex identified as solely dedicated to the deposition of histone variants in S. cerevisiae (Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004). To date, the only cellular function of this fourteen-subunit complex appears to be deposition of histone variant H2A.Z into chromatin at distinct regions such as the promoters of genes, double stranded breaks (DSB), and heterochromatin boundaries. It was recently elucidated that SWR1-C is recruited to gene promoters by recognition of NFRs flanked by nucleosomes by its Swr1 and Swc2 subunits (Ranjan et al. 2013; Yen et al. 2013). Beyond its central role in SWR1-C catalytic activity, Swr1 also acts as a scaffold for the assembly of numerous SWR1-C components (Nguyen et al. 2013; Wu et al. 2005; 2009). Swr1 is a large protein with several distinct regions such as the HSA (helicase-SANT-associated) and ATPase domains that are involved in diverse aspects of SWR1-C function (Figure 1.4). The HSA domain is often associated with helicases and mediates protein-protein interactions (Szerlong et al. 2008; Trotter et al. 2008). The ATPase domain of Swr1 is distinguished from canonical ATPase domains by the presence of an insertion region (Mizuguchi et al. 2004). These two domains are important for interactions with various members of SWR1-C. For example, the ATPase domain of Swr1 is required for binding to Swc2, Arp6, Swc6, Rvb1, and Rvb2, whereas the N-terminal portion is essential for association with Yaf9, Swc4, Act1, Arp4, Swc7, Bdf1, and H2A.Z (Wu et al. 2009; 2005; Nguyen et al. 2013). The N-terminal region of Swr1 can be further divided into two sections, N1 and N2, whereby N2 contains the HSA domain that physically associates with the shared module, composed of Act1, Arp4, Yaf9 and Swc4 (Szerlong et al. 2008). The majority of SWR1-C subunits have been functionally characterized and most of them have been found to be required for H2A.Z exchange in vitro.     18 Interestingly, SWR1-C can perform histone exchange in the absence of Bdf1, Swc7, and Swc3, suggesting that these subunits have auxiliary functions (Wu et al. 2009; 2005).                          Figure 1.4 SWR1-C Structure Schematic representation of the complex association map of SWR1-C showing Swr1 as the assembly platform.   1.5.2 NuA4 Structure and Function NuA4 is a large thirteen-subunit complex that acetylates histones H2A, H2A.Z and H4 and is the only essential HAT in S. cerevisiae (Allard et al. 1999; Eberharter et al. 1998; Smith et al. 1998; Millar et al. 2006; Babiarz et al. 2006; Keogh et al. 2006). The essential function of NuA4 lies within its catalytic subunit, Esa1. In recent years, we are beginning to appreciate NuA4’s role as a general lysine acetyltransferase for many non-histone targets (Mitchell et al. 2008; 2013; Lin et al. 2008; 2009). As a multifunctional HAT complex, NuA4 is implicated in transcriptional regulation, the DNA damage response (DDR), chromosome segregation, heterochromatin boundary establishment, life span, and cell autophagy (Boudreault et al. 2003; Bird et al. 2002; Downs et al. 2004; Lin et al. 2008; Krogan et al. 2004; Zhang et al. 2004; Babiarz et al. 2006; Zhou et al. 2010; Lin et al. 2009; Yi et al. Ac-H4Ac-H4Ac-H4Ac-H4Ac-H4AcNuA4SWR1-CH2A.ZAc-H2A.ZSWR1-CArp4Swc4Act1Yaf9123Arp4Swc4Act1Yaf9NuA4NuA4Arp4Swc4Act1Yaf9A BNuA4SWR1-CFigure 1AcAc-H4Ac-H4Swr1Swc5Arp6Swc7Swc6 Swc2Swc3Act1Arp4Swc4Bdf1ATPasedomainHSARvb1Rvb2N1N2H2A.ZH2BH2A.ZH2BYaf9Act1Yaf9Arp4Swc4 Eaf1Tra1Eaf7Eaf5Eaf3Yng2Epl1Esa1HSASANTeaf6N-term.    19 2012; Lu et al. 2011; Mitchell et al. 2011). The main scaffold component of NuA4 is Eaf1, which is required for binding of various NuA4 sub-modules via its distinct domains (Figure 1.5). For instance, the SANT region of Eaf1 is required for binding to Tra1 and the N-terminal region of Eaf1 associates with the Eaf3/5/7 sub-module (Auger et al. 2008). Furthermore, the HSA domain of Eaf1 is required for binding of the shared module composed of Act1, Arp4, Yaf9 and Swc4. In S. cerevisiae, NuA4 exists in two different forms within the cell. Namely, as the large thirteen-subunit NuA4 complex and as a smaller piccolo NuA4 complex consisting of only Esa1, Yng2, Eaf6, and Epl1 subunits (Auger et al. 2008; Boudreault et al. 2003; Chittuluru et al. 2011). Whereas NuA4 acetylates histones in a gene-specific manner, piccolo NuA4 is involved in global acetylation of H2A and H4 (Boudreault et al. 2003). In Chapter 4 of this dissertation, I expand upon our existing knowledge of the functional and structural relationship between picNuA4 and NuA4.                          Figure 1.5 NuA4 Structure Schematic representation of the complex association map of NuA4 showing Eaf1 as the scaffolding subunit for the modular complex.   Ac-H4Ac-H4Ac-H4Ac-H4Ac-H4AcNuA4SWR1-CH2A.ZAc-H2A.ZSWR1-CArp4Swc4Act1Yaf9123Arp4Swc4Act1Yaf9NuA4NuA4Arp4Swc4Act1Yaf9A BNuA4SWR1-CFigure 1AcAc-H4Ac-H4Swr1Swc5Arp6Swc7Swc6 Swc2Swc3Act1Arp4Swc4Bdf1ATPasedomainHSARvb1Rvb2N1N2H2A.ZH2BH2A.ZH2BYaf9Act1Yaf9Arp4Swc4 Eaf1Tra1Eaf7Eaf5Eaf3Yng2Epl1Esa1HSASANTeaf6N-term.    20 1.5.3 H2A.Z at the Intersection between SWR1-C and NuA4  The histone variant H2A.Z is the most notable nexus between NuA4 and SWR1-C. Elucidation of the cellular function of H2A.Z has been greatly facilitated by its non-essential role in S. cerevisiae. Reassuringly, the majority of these functions were subsequently confirmed in higher eukaryotes in which H2A.Z is essential. In yeast, H2A.Z is involved in heterochromatin boundary formation, transcriptional activation and repression, chromosome stability, and chromosome segregation (Adam et al. 2001; Larochelle and Gaudreau 2003; Meneghini et al. 2003; Zhang et al. 2005; Santisteban et al. 2000; Sharma et al. 2013). Consistent with a role in transcriptional regulation, H2A.Z is primarily found in the promoter regions of genes where it often flanks the NFR proximal to the TSS (Guillemette et al. 2005; Li et al. 2005; Raisner et al. 2005). However, the exact role of H2A.Z in transcriptional regulation remains a puzzle, as genome-wide studies in yeast and metazoans do not reveal a simple regulatory relationship between loss of H2A.Z and gene activity (Abbott et al. 2001; Li et al. 2005; Meneghini et al. 2003; Suto et al. 2000; Zhang et al. 2005; Albert et al. 2007; Gévry et al. 2007; Buchanan et al. 2009; Hardy et al. 2009).   The role of H2A.Z at the boundary of heterochromatin and euchromatin is one example that nicely illustrates the functional interplay between NuA4 and SWR1-C. Global mRNA profiling indicated that H2A.Z is necessary for proper expression of many genes located in sub-telomeric regions (Meneghini et al. 2003). In the majority of cases, aberrant spreading of the heterochromatin SIR proteins into euchromatin regions caused reductions in gene expression in the absence of H2A.Z, suggesting that H2A.Z forms a barrier that limits SIR spread (Meneghini et al. 2003). Interestingly, the ability of H2A.Z to maintain proper     21 heterochromatin boundaries is dependent on its acetylation by NuA4 (Babiarz et al. 2006). The targeted deposition of H2A.Z into telomeres is also dependent on the acetylation of H4K16 by the SAS HAT complex, further demonstrating the interplay between H2A.Z deposition and H4 acetylation (Shia et al. 2006). As I will illustrate in Chapter 3, however, it appears that the SAS-mediated H2A.Z deposition activity is a telomere-specific mechanism.    SWR1-C is the principal complex responsible for the incorporation of H2A.Z into chromatin whereas NuA4 has two regulatory roles in H2A.Z biology. First, NuA4 is important for the association of H2A.Z and SWR1-C with chromatin. NuA4-dependent H2A and H4 acetylation stimulates and targets SWR1-C to specific gene promoters for H2A.Z exchange (Babiarz et al. 2006; Durant and Pugh 2007; Altaf et al. 2010). In addition to the acetylation of histones by NuA4, it appears that acetylation of the H3 N-terminal by another HAT, SAGA, also contributes to H2A.Z deposition (Raisner et al. 2005). Second, given NuA4’s role in H2A acetylation and the high degree of homology between H2A and H2A.Z, it is perhaps not surprising that NuA4 also acetylates H2A.Z (Babiarz et al. 2006; Keogh et al. 2006; Millar et al. 2006). H2A.Z is first deposited into chromatin by SWR1-C, and then subsequently acetylated by NuA4 (Babiarz et al. 2006; Keogh et al. 2006). Interestingly, acetylation of H2A.Z becomes essential in the absence of H4 acetylation as seen in the synthetic lethality of a strain containing both the non-acetylable forms of H2A.Z (htz1-K3, 8, 10, 14R) and H4 (hhf2-K5, 8, 12R) (Babiarz et al. 2006). Lastly, it was recently shown that INO80 catalyzes the opposite reaction as SWR1-C; INO80 replaces nucleosomal H2A.Z for H2A (Papamichos-Chronakis et al. 2011).        22 Initial biochemical purifications of SWR1-C provided hints to its connection to NuA4 and INO80, as all three complexes share the Arp4 and Act1 subunits (Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004). During the DDR, there are intriguing physical and functional overlaps between SWR1-C, NuA4, and INO80 (Morrison and Shen 2009; van Attikum and Gasser 2009; Papamichos-Chronakis and Peterson 2013; Sinha and Peterson 2009; Papamichos-Chronakis et al. 2006). Upon DNA damage, one of the first PTMs that initiate the DDR cascade is the phosphorylation of H2A.X by the checkpoint kinases, Mec1 and Tel1 (Shroff et al. 2004). This modification covers a large region flanking the DSB that ranges from 50-100 kb (Rogakou et al. 1999). Arp4, a shared subunit of SWR1-C, NuA4, and INO80, is required for recruitment of these complexes to sites of DNA damage via direct physical interaction with phosphorylated H2A.X (Downs et al. 2004). NuA4 is recruited to DSBs in order to acetylate histone H4. This process results in a more accessible chromatin structure and allows for recruitment of DNA repair machinery and various chromatin-modifying complexes, including SWR1-C and INO80 (Bird et al. 2002; Downs et al. 2004; Tamburini and Tyler 2005; Bennett et al. 2013). In addition to acetylated H4, phosphorylated H2A.X also directly recruits SWR1-C and INO80 to DSBs (Morrison et al. 2004; Papamichos-Chronakis et al. 2006; van Attikum et al. 2007). H2A.Z is transiently and rapidly deposited into chromatin at DSBs immediately following DNA damage (Kalocsay et al. 2009). A strain lacking H2A.Z displays DNA resection defects and damage checkpoint defects, suggesting that H2A.Z has an early role in the DSB repair pathway (Kalocsay et al. 2009; Van et al. 2015). Following DDR, H2A.Z and H2A.X are subsequently removed from the DSB in an INO80-dependent manner (van Attikum et al. 2007; Papamichos-Chronakis et al. 2011).      23  1.6 Conservation of NuA4 and SWR1-C in Mammals  Many components of SWR1-C and NuA4 are structurally and functionally conserved throughout eukaryotic evolution, highlighting the importance of these complexes in various cellular processes of eukaryotic development. The SRCAP complex functions as the human counterpart of the yeast SWR1-C to deposit H2A.Z into chromatin (Wong et al. 2007). Similar to NuA4, the primary function of human TIP60 complex is to acetylate histones H2A and H4, but it can also exchange H2A.Z/H2B for H2A/H2B dimers (Ikura et al. 2000; Cai et al. 2003; Doyon et al. 2004). H2A.Z exchange is mediated by two distinct subunits in mammals, p400 (in the TIP60 complex) and the SRCAP subunit (in the SRCAP complex) (Gévry et al. 2007; Svotelis et al. 2009; Martinato et al. 2008). TIP60 and SRCAP complexes share several subunits in addition to the unique subunits that differentiate the two complexes. Biochemical and structural evidence suggests that NuA4 and SWR1-C exist as a single, massive complex in higher eukaryotes (Doyon et al. 2004). It is speculated that the two major scaffold proteins for NuA4 and SWR1-C, Eaf1 and Swr1, fused to form the p400 protein in higher eukaryotes, since p400 contains SANT, HSA, and ATPase domains (Auger et al. 2008; Doyon et al. 2004; Lu et al. 2009). Evidently, a fusion protein containing distinguishable domains of Swr1 and Eaf1 in S. cerevisiae can partially replace the function of both subunits in H2A.Z deposition and H4 acetylation (Auger et al. 2008). Furthermore, a TIP60-like complex is reconstituted in yeast in the presence of the Eaf1-Swr1 fusion protein, suggesting a physical merger of SWR1-C and NuA4.        24 1.7 Summary  The chromatin structure is essential for the compaction of the eukaryotic genome into the cell nucleus. A wide variety of chromatin-modifying complexes regulate this organizational structure in order to maintain an equilibrium of accessibility and compaction. While we are beginning to appreciate the complex mechanisms underlying chromatin biology, there still remain many more questions on how these factors work together. In this dissertation, I examined the cooperative activities of the Asf1 histone chaperone and SWR1-C chromatin-remodeling complex at heterochromatic regions and gene promoters. Furthermore, I dissected the complex dynamics of the NuA4 histone acetyltransferase and its catalytic module using a variety of genome-wide and biochemical approaches.   In Chapter 2, I describe the genetic and functional connections between Asf1 and SWR1-C in the maintenance of native heterochromatin structures and nucleosome composition at promoters. Asf1 and a subunit of SWR1-C, Yaf9, are both required for maintaining expression of heterochromatin-proximal genes and they worked cooperatively to prevent repression of telomere-proximal genes by limiting the spread of SIR complexes to nearby regions. Furthermore, genetic and molecular analysis revealed that H3K56 acetylation was required for efficient deposition of H2A.Z at subtelomeric and euchromatic gene promoters, pointing to a role for Asf1-dependent H3K56 acetylation in SWR1-C biology.   In Chapter 3, I investigate the functional connection between Asf1 and SWR1-C on H2A.Z containing nucleosomes. Loss of H2A.Z at heterochromatin boundaries was not due to loss     25 of SAS recruitment by Asf1. By examining H2A.Z occupancy genome-wide, I found that the acetylation of lysine 56 on histone H3 was required for H2A.Z occupancy at a subset of promoters. Furthermore, loss of H3K56 acetylation resulted in alterations of H2A.Z position at gene promoters normally enriched for this histone modification.   In Chapter 4, I examined the consequences on NuA4 structure and function in the presence of two genetic perturbations, loss of the entire Eaf1 protein and loss of the Epl1 C-terminus. Eaf1 is the main scaffold protein of NuA4 and the C-terminus of Epl1 anchors the catalytic module to Eaf1. Large-scale genetic and gene expression analysis and comprehensive biochemical assays identified the Eaf1 HSA domain and the Epl1 C-terminus as two key domains in the NuA4 complex that regulate the cellular equilibrium of NuA4 and picNuA4.   In Chapter 5, I discuss my results in Chapter 2-4 and highlight how my findings fit within and contribute to our current understanding of chromatin biology. I explore the implications of my findings and discuss future experiments that would elucidate the molecular mechanisms governing the coordinated activities of chromatin-modifiers on the chromatin structure.       26 Chapter 2: Maintenance of Heterochromatin Boundary and Nucleosome Composition at Promoters by the Asf1 Histone Chaperone and SWR1-C Chromatin Remodeler in Saccharomyces cerevisiae1   2.1  Introduction  The fundamental building block of chromatin is a nucleosome composed of 146 bp of DNA wrapped around a histone octamer. Protein complexes involved in post-translational modification of histones, nucleosome movement or replacement alter chromatin dynamics to regulate various chromosomal processes. Often, these chromatin-modifying processes intersect and interact cooperatively to regulate chromatin structure.   Transcriptionally silent heterochromatin structures are a prime example of the multilayered activities of chromatin modifying complexes. In budding yeast Saccharomyces cerevisiae, there are three well-defined regions of silent chromatin: the mating loci (HMR and HML), the rDNA locus and telomeres. These regions are characterized by a distinct set of histone modifications and associated factors that distinguish them from adjacent transcriptionally active euchromatin (Rusche et al. 2003). Chief among these is the Silent Information Regulator (SIR) complex, which not only constitutes the main structural component during                                                 1 This chapter is published in Genetics. Lu, P.Y.T. and Kobor, M.S., 2014. Maintenance of heterochromatin boundary and nucleosome composition at promoters by the Asf1 histone chaperone and SWR1-C chromatin remodeler in Saccharoymyces cerevisiae.Genetics. 197(1): 133-145      27 establishment and maintenance of heterochromatin but also harbors an enzymatic function (Rusche et al. 2003). Specifically, the Sir2 subunit is a NAD+ dependent histone deacetylase (HDAC) that deacetylates H4K16, a process critical for promoting initial SIR complex formation at heterochromatin loci as well as for formation of heterochromatin boundaries. The establishment of heterochromatin boundaries is biologically important to prevent antagonistic silencing of neighbouring euchromatic genes by encroaching heterochromatic structures (Rusche et al. 2003; Imai et al. 2000).   Several additional histone modifying enzymes contribute to demarcating the boundary between heterochromatin and euchromatin, including the Something About Silencing 2 (SAS2) complex, the Dot1 histone methyltransferase, and the highly conserved histone variant H2A.Z (Takahashi et al. 2011; Verzijlbergen et al. 2009; Shia et al. 2006; Meijsing and Ehrenhofer-Murray 2001; Osborne et al. 2009). Similar to the other factors, loss of H2A.Z from the heterochromatin-euchromatic boundary results in the spread of SIR complexes to nearby subtelomeric genes and subsequent repression of these genes (Meneghini et al. 2003). In part, the role of H2A.Z at boundaries is mediated by acetylation of its N-terminal lysine residues, although the magnitude of this effect varies among different studies (Babiarz et al. 2006; Millar et al. 2006; Keogh et al. 2006).   Like many of the factors sculpting heterochromatin boundaries, H2A.Z also has roles in euchromatic regions. H2A.Z, for instance, is enriched at the majority of gene promoters and often resides in the two nucleosomes flanking the nucleosome free region (NFR) (Raisner et al. 2005). H2A.Z is more commonly localized to lowly expressed gene promoters and is     28 largely absent from highly expressed genes (Guillemette et al. 2005; Li et al. 2005; Zhang et al. 2005). Despite these strong correlations, the causal role of H2A.Z for gene expression is more nebulous as genome-wide expression studies found that loss of H2A.Z affects only a minority genes in yeast (Meneghini et al. 2003). Rather than affecting steady-state gene expression, H2A.Z may facilitate the induction of genes in response to changing environments (Adam et al. 2001; Larochelle and Gaudreau 2003; Lemieux et al. 2008; Halley et al. 2010).   H2A.Z is deposited into chromatin by SWR1-C, an ATP-dependent chromatin remodeling complex that recognizes the NFR and exchanges H2A-H2B dimers with H2A.Z-H2B dimers at the two flanking nucleosomes (Ranjan et al. 2013; Yen et al. 2013; Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004). Illustrative of the crosstalk between chromatin remodelers, SWR1-C and the NuA4 histone acetyltransferase (HAT) have interconnected activities converging on H2A.Z chromatin neighbourhoods. Acetylation of H4 lysine residues by NuA4 promotes the recruitment of SWR1-C for H2A.Z deposition, and subsequent acetylation of newly incorporated H2A.Z by NuA4 is required for gene activation and heterochromatin formation (Altaf et al. 2010; Ranjan et al. 2013; Durant and Pugh 2007; Babiarz et al. 2006). The interplay between SWR1-C and NuA4 likely involves a module of four subunits that is shared between the two complexes (Lu et al. 2009). For instance, at a subset of telomeres, the Yaf9 shared subunit is vital to H2A.Z incorporation, H4 acetylation, and proper telomere-specific gene expression changes (Zhang et al. 2004; Wang et al. 2009).      29 Yaf9 contains an evolutionary conserved YEATS domain, whose closest structural relative in budding yeast is Asf1(Wang et al. 2009). These two proteins share a common essential function, although the nature thereof has yet to be defined.  Asf1 plays a number of roles in heterochromatin function, transcription regulation, and cellular response to DNA damage (Mousson et al. 2007; Lin and Schultz 2011; Chen et al. 2008; Williams et al. 2008; Sharp et al. 2001). Asf1 functions as an anti-silencing factor as both its deletion and overexpression have an antagonistic effect on silencing of yeast mating type loci (Le et al. 1997; Singer et al. 1998). Regulation of silencing by Asf1 occurs through a redundant pathway with chromatin assembly factor 1 (CAF-1) (Sharp et al. 2001; Sutton et al. 2001). Mechanistically, Asf1 functions as a highly conserved histone chaperone for the H3/H4 heterodimer to mediate nucleosome assembly and disassembly, and is required to facilitate the acetylation of lysine 56 on H3 by the HAT, Rtt109 (Kolonko et al. 2010; Recht et al. 2006; Tsubota et al. 2007; Fillingham et al. 2008). Similar to Asf1, the acetylation state of H3K56 is important for the maintenance of silent chromatin structures at yeast mating loci and telomeres (Miller et al. 2008; Värv et al. 2010; Xu et al. 2007). Remarkably, Asf1 and SWR1-C are functionally linked as H3K56 acetylated nucleosomes alter the substrate specificity of SWR1-C to cause promiscuous exchange of H2A.Z (Watanabe et al. 2013).     The functional and genetic linkages between Asf1 and SWR1-C suggest that their cellular roles are closely related. Our works revealed that while Asf1 and SWR1-C had distinct functions for the expression of heterochromatin-proximal genes, they had overlapping roles in H2A.Z deposition. Consistent with locus–specific layers of chromatin modifications at heterochromatin, Asf1 was required for normal expression of HMR-proximal but not     30 telomere-proximal genes, whereas Yaf9 regulated silencing of both HMR- and telomere-proximal genes. Furthermore, we found Asf1 worked cooperatively with Yaf9 at the telomeric heterochromatin boundary to restrict SIR complexes at a subset of telomeres. The growth defect of asf1Δyaf9Δ cells could not be attributed to defects in maintaining heterochromatin structure. Instead, we established that Asf1-mediated H3K56 acetylation was required to maintain normal levels of H2A.Z at promoters of subtelomeric genes, hinting that Asf1 may regulate subtelomeric gene expression by influencing H2A.Z occupancy at promoters. We also elucidated that acetylation of H3K56 by Asf1 was also required for H2A.Z occupancy at euchromatic promoters, which suggests a broader role for this histone modification in H2A.Z biology.   2.2 Materials and Methods  2.2.1 Yeast Strains and Plasmids All strains used in this study are listed in Table 2.1. Yeast strains were generated using standard genetic techniques including homologous recombination and genetic crosses followed by tetrad dissection (Ausubel 1987). Complete deletion of genes and 3’ end integration of an in frame 3xFLAG tag were achieved using one-step gene integration PCR-amplified modules (Gelbart et al. 2001). Mating of the sir2Δ strain was achieved with the aid of the URA3 plasmid pRS316[SIR2], 5-FOA was used as a counter-selection agent to evict plasmid following tetrad dissection. Plasmid shuffle experiments were performed using 5-FOA to evict URA3 plasmid pRS316[H3 H4], and select for pRS316[H3K56R H4], pRS316[H3K56Q H4], pRS316[H3K9R H4], pRS316[H3K9Q H4], and     31 pRS316[H3K9RK56R H4]. These plasmids were a generous gift from Ann Kirchmaier (Purdue University). The pASF1, pasf1 H36A/D37A and pasf1 H39A/K41A plasmids were generous gifts from Paul Kaufman (University of Massachusetts Medical School). The pasf1 V94R mutant was obtained from Carl Mann (CEA).   Table 2.1 Yeast strains used in this study Strain number Relevant Genotype  MKY5  W303, MATα ade2-1 can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 LYS2  MKY6  MKY7 MKY399 W303, MATA, ADE2, can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ W303, MATA, ade2-1, can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 LYS2 W303, MATα, ADE2, can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY352 MKY6, asf1Δ::NATMX  MKY1596 MKY6, yaf9Δ::KANMX  MKY1597 MKY6, asf1Δ::NATMX yaf9Δ::KANMX  MKY1598 MKY6, htz1Δ::HYGMX6  MKY1599 MKY6, asf1Δ::NATMX6  htz1Δ::HYG  MKY646 MKY6, swr1Δ::HIS  MKY754 MKY6, asf1Δ::NATMX swr1Δ::HIS  MKY672 MKY6, swc2Δ::HIS  MKY385 MKY6, asf1Δ::NATMX swc2Δ::HIS  MKY394 MKY6, swc3Δ::HIS  MKY396 MKY6, asf1Δ::NATMX swc3Δ::HIS  MKY903 MKY6, swc5Δ::HIS  MKY467 MKY6, asf1Δ::NATMX swc5Δ::HIS  MKY463 MKY6, swc6Δ::HIS  MKY458 MKY6, asf1Δ::NATMX swc6Δ::HIS  MKY1600  MKY7, SIR2-FLAG::KANMX  MKY1601 MKY7, SIR2-FLAG::KANMX asf1Δ::HIS       32 Strain number Relevant Genotype  MKY1602 MKY7, SIR2-FLAG::KANMX yaf9Δ::NATMX  MKY1603 MKY7, SIR2-FLAG::KANMX asf1Δ::HIS yaf9Δ::NATMX  MKY1605 MKY6, asf1Δ::NATMX yaf9Δ::HIS sir2Δ::KANMX  MKY1606 MKY6, asf1Δ::NATMX swr1Δ::HIS sir2Δ::KANMX  MKY1607 MKY6, asf1Δ::NATMX sir2Δ::KANMX  MKY1608 MKY6, yaf9Δ::HIS sir2Δ::KANMX  MKY1609 MKY6, swr1Δ::HIS sir2Δ::KANMX  MKY1610 MKY399, asf1Δ::NATMX [pRS414, ASF1]  MKY1611 MKY399, asf1Δ::NATMX [pRS414, asf1 H36A/D37A]  MKY1612 MKY399, asf1Δ::NATMX cac2Δ::HYGMX [pRS414, ASF1]  MKY1613 MKY399, asf1Δ::NATMX yaf9Δ::KANMX [pRS414, ASF1]  MKY1614 MKY399, asf1Δ::NATMX cac2Δ::HYGMX [pRS414, asf1 H36A/D37A]  MKY1615 MKY399, asf1Δ::NATMX yaf9Δ::KANMX [pRS414, asf1 H36A/D37A]  MKY1616 MKY399, asf1Δ::NATMX cac2Δ::HYGMX yaf9Δ::KANMX [pRS414, ASF1]  MKY1617 MKY399, asf1Δ::NATMX cac2Δ::HYGMX yaf9Δ::KANMX [pRS414, asf1 H36A/D37A]  MKY1618 MKY399, asf1Δ::NATMX [pRS414, asf1 H39A/K41A]  MKY1619 MKY399, asf1Δ::NATMX cac2Δ::HYGMX [pRS414, asf1 H39A/K41A]  MKY1620 MKY399, asf1Δ::NATMX yaf9Δ::KANMX [pRS414, asf1 H39A/K41A]  MKY1621 MKY399, asf1Δ::NATMX cac2Δ::HYGMX yaf9Δ::KANMX [pRS414, asf1 H39A/K41A]  MKY1622  MKY6 [pRS314]  MKY1623 MKY6, asf1Δ::NATMX [pRS314]  MKY1624 MKY6, yaf9Δ::KANMX [pRS314]  MKY1625 MKY6, asf1Δ::NATMX yaf9Δ::KANMX [pRS314]  MKY1626 MKY6, asf1Δ::NATMX [pRS314, ASF1-MYC]  MKY1627 MKY6, asf1Δ::NATMX [pRS314, asf1 V94R-MYC]  MKY1628 MKY6, asf1Δ::NATMX yaf9Δ::KANMX [pRS314, ASF1-MYC]  MKY1629 MKY6, asf1Δ::NATMX yaf9Δ::KANMX [pRS314, asf1 V94R-MYC]  MKY1630 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2-HHF2]  MKY1631 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2-HHF2]      33 Strain number Relevant Genotype  MKY1632 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K56Q HHF2]  MKY1633 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K56R HHF2]  MKY1634 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2 K56Q HHF2]  MKY1635 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2 K56R HHF2]  MKY555 MKY6, rtt109Δ::KANMX  MKY1636 MKY6, rtt109Δ::KANMX yaf9Δ::HIS  MKY677 MKY6, rtt109Δ::KANMX swr1Δ::HIS  MKY674 MKY6, rtt109Δ::KANMX swc2Δ::HIS  MKY696 MKY6, rtt109Δ::KANMX swc3Δ::HIS  MKY763 MKY6, rtt109Δ::KANMX swc5Δ::HIS  MKY690 MKY6, rtt109Δ::KANMX swc6Δ::HIS  MKY1637 MKY6, rtt109Δ::KANMX, htz1Δ::HYGMX  MKY1638 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K9R HHF2]  MKY1639 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K9Q HHF2]  MKY1640 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2 K9Q HHF2]  MKY1641 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2 K9R HHF2]  MKY1642 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K9R K56R HHF2]  MKY1643 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::KANMX [pRS314, HHT2 K9R K56R HHF2]  MKY353 MKY5, HTZ1-FLAG::KANMX  MKY355 MKY5, HTZ1-FLAG::KANMX asf1Δ::NATMX  MKY354 MKY5, HTZ1-FLAG::KANMX yaf9Δ::HIS  MKY356 MKY5, HTZ1-FLAG::KANMX asf1Δ::NATMX  yaf9Δ::HIS  MKY1644 MKY7, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2-HHF2]  MKY1645 MKY7, HTZ1-FLAG::KANMX  hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2-HHF2]  MKY1646 MKY7, HTZ1-FLAG::KANMX  hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::NATMX [pRS314, HHT2-HHF2]  MKY1647 MKY7, HTZ1-FLAG::KANMX  hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K56R HHF2]  MKY1648 MKY7, HTZ1-FLAG::KANMX  hht1-hhf1::LEU2 hht2-hhf2::HIS yaf9Δ::NATMX [pRS314, HHT2 K56R HHF2]      34 2.2.2 Growth and Genotoxic Sensitivity Assays Overnight cultures grown in YP-dextrose were diluted to OD600 0.5. Cells were 10-fold serially diluted and spotted onto solid YPD plates or plates with 10mM hydroxyurea. For strains containing TRP1 plasmids, the cultures were grown in SC –TRP media and serially diluted cells were spotted onto SC –TRP plates or plates containing 10mM hydroxyurea. The plates were then incubated at the indicated temperature for 36 hours. The 16°C plates were incubated for 96 hours.  2.2.3 RT-qPCR Overnight cultures were diluted to OD600  of 0.15 and grown in YP-dextrose to an OD600 of 0.5. Ten OD600 units were harvested for RNA extraction and purification using a Qiagen RNeasy minikit as per manufacturer protocol. cDNA was synthesized using QuantiTect Reverse Transcription Kit (Qiagen). cDNA was analyzed using a Rotor-Gene 6000 (Qiagen) and PerfeCTa SYBR green FastMix (Quanta Biosciences). mRNA levels were normalized to TUB1 mRNA levels. Samples were analyzed in triplicates for three independent RNA preparations. Statistical significance was assessed using Student’s t test.  Primer sequences are listed in Table 2.2.            35 Table 2.2 Primer sequences used in this study Name Method Primer sequence (5’-3’) YCR094W ORF F YCR094W ORF R RT-qPCR RT-qPCR TTTCACTCTTTGGTGGCACA TCGCTCTTGGTTGGAAGATT YCR095C ORF F  RT-qPCR CTTTGCAGAGAGCCAGAAGTG YCR095C ORF R RT-qPCR TCAAATCGTCTCCTAGAACTCCAC GIT1 ORF F RT-qPCR ATCGGTTCTGTAGTAGGCG GIT1 ORF R YCR099C ORF F  YCR099C ORF R YCR100C ORF F YCR100C ORF R RDS1 ORF F RDS1 ORF R TUB1 ORF F TUB1 ORF R ChrIII 286.5kbp F ChrIII 286.5kbp R ChrIII 288kbp F ChrIII 288kbp R ChrIII 289.5kbp F ChrIII 289.5kbp R ChrIII 291kbp F ChrIII 291kbp R HMR ORF F HMR ORF R ChrIII 296.5kbp F ChrIII 296.5kbp R ChrIII 298kbp F  ChrIII 298kbp R  RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP TTACCAGTCCAGCCATTGG TCAGATTGCCCTTTCCGATA GGAATGTTCATGGCCTCAAT CCAGATGGATCAGGCTCAAA TCGATCGCATACAGGACACT AAGCCGTGAGATTGAAATGG  CTCCATCTGGCACAACAGAA  TCTTGGTGGTGGTACTGGTT TGGATTTCTTACCGTATTCAGCG CGCACTGCGAACAAAGAATA CGCTGTGTTAGGAGGCTTTT TGAGGATGTACACGCAGAGAA AGCAAACCCTGGGAGTTCTT ATTGGGCTTCTGGATGACAG AGGAAGGCATGCATTGAACT TGATTTAAGCGTGCGTGAAG GGTCAACATAAAGTGGCGAGA TGGCGGAAAACATAAACAGA TGCTTGGGGTGATATTGATG TCAACATGGTGTTCCAAAGC TGATGCAGTGGCGATATCAT GAACACACTCAATGGCCAGA TGGTCCATTTGCTACGATCA     36 Name Method Primer sequence (5’-3’) ChrIII 299.5kbp F ChrIII 299.5kbp R ChrIII 301kbp F ChrIII 301kbp R ChrIII 309kbp F ChrIII 309kbp R ChrIII 310.5kbp F ChrIII 310.5kbp R ChrIII 312kbp F ChrIII 312kbp R ChrIII 313.5kbp F ChrIII 313.5kbp R YCR094W Prom F YCR094W Prom R ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP TTTCGACGGCTGATAACCTT AAAGAAGGGGAGCGAAAAAG GGAATGTTCATGGCCTCAAT TCAGATTGCCCTTTCCGATA GGTATTTCCAACGCAAAGGA CGGCCAATTATTTCATGTCC CGAGCATGTCTGATGAGGTC CCGAATGCAAAAATGTGAAG TGAGCCCCACTAACTTGCTT TACCAGCATCACCTTCAGCA TTTGATGCCTTCTGGGATTT TTACATTCCGCCTGCCTAAC GCAAACCCCTCTACAATCCA CAAAAGTGAAAGCGACCCATA YCR095C Prom F  ChIP TACCGTATGCGGTATAATGA YCR095C Prom R ChIP GTCTCCACTTTAGAACATCT GIT1 Prom F ChIP TTCATGAATTTCCTTACTGGAC GIT1 Prom R YCR099C Prom F  YCR099C Prom R YCR100C Prom F YCR100C Prom R RDS1 Prom F ChIP ChIP ChIP ChIP ChIP ChIP GTTGACTAGTCACAAGAAACAG TGCTACTGGTGATCTGGGAAA CTGATCCATCTGGCGTTGTA GCAAGGATTCTGACTTTACTGG  CTCGTTATGCCCGTCATCTT TGTGCTATCTAAGAGGATGGTTCA  RDS1 Prom R PRP8 ORF F PRP8 ORF R GID7 Prom F GID7 Prom R STE50 Prom F ChIP ChIP ChIP ChIP ChIP ChIP CAGCAGCCAATTTCATGTTC  GGATGTATCCAGAGGCCAAT AACCCGCGTATTAAGCCATA CTTGTGTGCGCTTGTAGCAT CCCGTTTCCGATGATATTGT AATGGTAGACACCGGACTCG     37 Name Method Primer sequence (5’-3’) STE50 Prom R BUD3 Prom F BUD3 Prom R MRK1 Prom F MRK1 Prom R BZZ1 Prom F BZZ1 Prom R YDC1 Prom F YDC1 Prom R ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP AGTGGAGCAGGCTGTACGTT GAGATTTCGATCACCATCGT  CTTCCTCATTGAAGATCAAAGC  AGGGGCAGCATTATCCGTTA CTTCCCCGCAGTATGGTAGA CGCTTAGCAAAGGAATAGACAGA TGCTCAATGTCCTTCAACCA CGGAGAGTGGCATAGCATG TGCAATAGAAAGAAAGCAAGCG   2.2.4 ChIP-qPCR and ChIP-on-chip Chromatin immunoprecipitation (ChIP) experiments were performed as described previously (Schulze et al. 2009). In brief, 250 ml of cells were grown in YP-dextrose to an OD600 of 0.5-0.6 from OD600 of 0.15 and were crosslinked with 1% formaldehyde for 20 minutes before chromatin was extracted. The chromatin was sonicated (Bioruptor, Diagenode, Spart, NJ: 10 cycles, 30s on/off, high setting) to yield an average DNA fragment of 500bp. Anti-FLAG antibody (4.2µl, Sigma) was coupled to 60µl of protein A magnetic beads (Invitrogen). After reversal of the crosslinking and DNA purification, the immunoprecipitated and input DNA were analyzed by quantitative real-time PCR (qPCR). Samples were analyzed in triplicate for three independent ChIP experiments. Statistical significance was assessed using Student’s t test.  Primer sequences are listed in Table 2.2. For microarray analysis, after reversal of crosslinking and DNA purification, the DNA was amplified with two rounds of T7 RNA polymerase amplification and hybridized to Affymetrix 1.0R S. cerevisiae tiling microarray.     38 A modified version of the model-based analysis of tiling arrays (MAT) algorithm was used to reliable detect Sir2 occupancy across the genome. The data was normalized using both input DNA and a mock IP control.   2.3 Results  2.3.1 ASF1 Genetically Interacted with Genes Encoding for Subunits of SWR1-C Yeast lacking both ASF1 and YAF9 have an exacerbated growth defect compared to cells with a single deletion of either gene, suggesting these genes share a redundant function (Figure 2.1) (Wang et al. 2009). Given that YAF9 is required for H2A.Z deposition by SWR1-C, we asked if the synthetically sick phenotype of asf1Δyaf9Δ double mutants broadly reflected defects in SWR1-C function. We characterized genetic interactions between ASF1 and genes that encode all unique subunits of SWR1-C. Under unperturbed growth conditions, loss of ASF1 resulted in synthetic sick genetic interactions with genes encoding eight SWR1-C subunits (Figure 2.1). The growth fitness of the double mutants was further reduced when cells were grown at 16°C or in conditions that induce DNA replication stress (HU). The decrease in fitness of all double mutants under stressed conditions was generally comparable to asf1Δyaf9Δ double mutants, although nuanced differences were present (Figure 2.1). Although ASF1 showed strong synthetic interaction with genes encoding for the entire SWR1-C complex, a similar pattern and severity of genetic interactions was not observed with members of the NuA4 complex (data not shown). Taken together, the genetic data suggested that the synthetic lethality between ASF1 and YAF9 primarily reflected Yaf9’s     39 function within SWR1-C, and that SWR1-C in its entirety had an important functional relationship with the Asf1 histone chaperone.    Figure 2.1 ASF1 genetically interacted with genes encoding for non-essential subunits of SWR1-C subunits Double mutants containing asf1Δ and a subunit of SWR1-C grew significantly more slowly than the respective single mutants. Ten-fold serial dilutions of the indicated strains were plated on YPD media, with or without 10mM HU and incubated at the indicated temperatures.    2.3.2 ASF1 and YAF9 were Required for Maintaining Heterochromatin-Proximal Gene Expression To further examine the interplay between Asf1 and Yaf9, we dissected their roles at heterochromatin boundaries, focusing on the well-characterized subtelomeric region encompassing the right telomere of chromosome III and the HMR locus. NuA4 does not YPD 30CWildTypeasf1∆yaf9∆asf1∆,  yaf9∆htz1∆asf1∆.  htz1∆swr1∆asf1∆,  swr1∆swc2∆asf1∆,  swc2∆swc3∆asf1∆,  swc3∆YPD 16C10mM HUWildTypeasf1∆yaf9∆asf1∆,  yaf9∆swc5∆asf1∆,  swc5∆swc6∆asf1∆,  swc6∆Figure 1.      40 acetylate H4 at Chr III R, which allowed us to assay SWR1-C-specific activities of Yaf9 (Zhang et al. 2004). A unique aspect of this 35kb region of DNA is that it encompasses three distinct heterochromatin boundaries: two located on either side of the HMR and one at the telomere heterochromatin boundary (Figure 2.2A). By RT-qPCR, we quantified expression levels of endogenous genes to interrogate native silencing defects, circumventing issues associated with embedded reporter genes such as URA3. Consistent with previous work, gene expression at all loci tested was significantly reduced in yaf9Δ strains (Figure 2.2B) (Zhang et al. 2004). Surprisingly, the effect of ASF1 in this 35kb region differed depending on the gene’s proximity to either the HMR or the telomere. Loss of ASF1 led to reduced expression of genes flanking the HMR, while ASF1 was dispensable for proper expression of subtelomeric genes (Figure 2.2B). Based both on proximity to the HMR/telomere and differential expression patterns in asf1Δ cells, we classified YCR094, YCR095 and GIT1 as HMR-influenced genes and YCR099, YCR100 and RDS1 as telomere-influenced genes. Interestingly, changes in transcript levels in the asf1Δ yaf9Δ double mutant did not reflect the genetic interaction phenotype. mRNA levels of the three genes flanking the HMR in the asf1Δ yaf9Δ double mutant were comparable to either of the single mutants alone, whereas mRNA levels of YCR099C, YCR100C, and RDS1 in the double mutant were reduced to the same level as the yaf9Δ mutant (Figure 2.2B).       41  Figure 2.2 ASF1 and YAF9 were required for maintaining heterochromatin-proximal gene expression (A) Schematic representation of ORFs at the boundaries of the HMR locus and the right telomere of Chromosome III. Arrows represent ORFs and provide transcriptional direction information. Locations of PCR primers for Sir2-FLAG ChIP analysis are indicated with arrows 1 to 12 spaced in 1.5kb increments. (B) As measured by RT-qPCR, mRNA levels of the indicated genes in wildtype, asf1Δ, yaf9Δ and asf1Δ yaf9Δ strains are represented by black, light gray, dark gray and white bars, respectively. asf1Δ strains exhibited reduced mRNA levels of HMR-flanking genes. Loss of YAF9 led to the repression of all heterochromatin-proximal genes tested. mRNA levels for all genes were normalized to levels of TUB1 mRNA. (C) Sir2 spread beyond the telomere and HMR in yaf9Δ and asf1Δ yaf9Δ strains. yaf9Δ exhibited a small but significant increase in Sir2 levels at regions previously devoid of Sir2. asf1Δ yaf9Δ showed a synergistic increase in Sir2 levels immediately adjacent to the telomere that spread up to 15kb away. Sir2 levels measured by ChIP-qPCR were normalized to Sir2 level at the PRP8 ORF. (B), (C) (D) Error bars represent standard error of values of three biological replicates.  *, p-value < 0.10; **, p-value < 0.05 when compared with the Wildtype strain using a two-tailed student t-test. Figure 2.   ABHMRYCR095C GIT1 YCR100C  RDS1Chromosome IIIYCR094WYCR099C1    2     3    4                       5    6    7    8                      9   10  11   12WildtypeSIR2-FLAGasf1 SIR2-FLAGyaf9 SIR2-FLAGasf1 yaf9 SIR2-FLAGC DHMR-flanking Subtelomeric35kbp02468101626360246810121416286500bp 288000bp 289500bp 291000bp HMR orf 296500bp 298000bp 299500bp 301000bpSIR2 levels normalized over PRP8309000bp 310500bp 312000bp 313500bpSIR2 levels normalized over PRP81               2               3              4                    5                6              7              8                                    9             10           11            12   00.20.40.60.811.21.4YCR094 YCR095 GIT1 YCR099 YCR100 RDS1mRNA abundance normalize over TUB1 Wildtypeasf1yaf9asf1 yaf9* ****** ** **** **************************** ******* ******* ****    42 We then asked whether repression of HMR-influenced genes in asf1Δ cells was due to the spread of SIR complexes beyond the HMR locus. We examined Sir2 levels via ChIP-qPCR in 1.5kb increments on either side of the HMR locus and outside of the Chr III R telomeric region (Figure 2.2A). Sir2 levels were enriched at the HMR locus and depleted at all other sites tested in wildtype cells as expected (Figure 2.2C). In agreement with previous studies, loss of YAF9 led to Sir2 spreading into regions previously devoid of SIR complexes, with the highest levels of Sir2 occupancy occurring on both sides flanking the HMR (Figure 2.2C) (Meneghini et al. 2003). Even though cells lacking ASF1 exhibited reduced transcriptional activity of HMR-adjacent genes, significant enrichment of Sir2 was not observed more than 1.5kb away from the HMR nor at the subtelomeric regions (Figure 2.2C and Figure 2.2D). In particular, the promoters of YCR094W, YCR095C and GIT1 were not enriched for Sir2 in asf1Δ cells (Figure 2.2C). Strikingly, while yaf9Δ cells showed significant increases in Sir2 levels at the subtelomeric regions, asf1Δ yaf9Δ cells exhibited dramatic Sir2 spreading immediately adjacent to the telomere (Figure 2.2C and Figure 2.2D).  These data suggested that Asf1 works cooperatively with SWR1-C to regulate SIR occupancy adjacent to the telomere. The increase in local concentrations of Sir2 was not reflected in gene expression levels since mRNA levels of telomere-associated genes in asf1Δ yaf9Δ double mutants were similar to yaf9Δ single mutants.   2.3.3 Cooperative Regulation of SIR Occupancy by Asf1 and Yaf9 was Telomere-Specific To ascertain how Asf1 and Yaf9 regulate Sir2 positioning, we characterized genome-wide Sir2 occupancy by ChIP-on-chip in the mutant strains. We focused our analysis on telomere-    43 proximal regions 30kb away from the chromosomal ends. In accordance with our ChIP-qPCR analysis of Chr III R, Sir2 occupancy was restricted to the HMR and to the region immediately adjacent to the telomere in wildtype and asf1Δ cells whereas in both the yaf9Δ strain and the double mutant, Sir2 spread beyond the normal heterochromatic regions (Figure 2.3A). We also demonstrated that over the 17kb region between the telomere and HMR, the genome-wide ChIP analysis recapitulated the dramatic increase in Sir2 levels when ASF1 was deleted in a yaf9Δ background. Across all chromosomal ends, the loss of ASF1 did not significantly alter Sir2 occupancy although the loss of YAF9 led to an increase in Sir2 occupancy at subtelomeres (data not shown). Interestingly, the principles derived from Chr III R appeared to not be limited to this unique region. We found similar patterns on three more chromosomal ends where both Asf1 and Yaf9 were required to restrict the spread of Sir2 (Figure 2.3B). Conversely, the patterns of Sir2 occupancy on the remaining 28 telomere ends were highly comparable between the yaf9Δ and the asf1Δ yaf9Δ mutants (data not shown). We also noted that Asf1 was important for regulating Sir2 occupancy over the highly conserved Y’ element that is present in the majority of yeast telomeres. Loss of YAF9 led to minimal changes to Sir2 occupancy within the Y’ element but the combined deletion of ASF1 and YAF9 led to a synergistic increase in Sir2 occupancy over the telomere element (Figure 2.3C). The highly repetitive nature of telomeric DNA makes it difficult to verify whether our observed trend holds true for all Y’ elements.       44  138 P. Y. T. Lu and M. S. Kobor    45 Figure 2.3 Regulation of Sir2 occupancy by Asf1 and Yaf9 was telomere-specific ChIP-on-chip profiles of Sir2. Thirty kilo-basepairs of chromosome ends were plotted along the x-axis against the relative occupancy (R/O) of Sir2 in wildtype, asf1Δ, yaf9Δ and asf1Δ yaf9Δ cells. ORFs are indicated as light gray rectangles above the x-axis for Watson genes and below the x-axis for Crick genes. (A) ChIP-on-chip profile of Sir2 along the right telomere of Chr III. Arrows 1 through 12 represent locations assayed by ChIP-qPCR in Figure 2. (B) Telomere-proximal profiles of Chr VIII L, VIII R, and IR showed increased Sir2 occupancy in asf1Δ yaf9Δ cells similar to Chr III R. (C) Genome-wide profiling of Sir2 highlighted differences in Asf1 and Yaf9 function over heterochromatin-specific Y’ elements. Chr XVI L, XIV L, and VII R are shown.     2.3.4 Asf1 and SWR1-C Affected Heterochromatin-Proximal Gene Expression through Different Mechanisms We next tested whether Sir2 accumulation caused gene expression changes across the right arm of chromosome III by measuring mRNA levels in cells lacking SIR2. To test whether Yaf9’s role at the Chr III R telomere was indeed linked to its function in SWR1-C as our previous experiments suggested, we included strains lacking SWR1, the catalytic subunit of SWR1-C. Not surprisingly, strains lacking YAF9 or SWR1 had very similar expression profiles at the Chr III R region suggesting that the decreased mRNA levels of these genes resulted from reduced SWR1-C activity (Figure 2.4A and Figure 2.4B). The combined analysis of gene expression in double and triple mutants highlighted distinct functions and causal interdependencies in this region. Firstly, loss of SIR2 restored expression levels of all six genes in yaf9Δ and swr1Δ cells (Figure 2.4A and Figure 2.4B).  In agreement with our Sir2 ChIP results, loss of SIR2 did not restore mRNA levels of HMR-flanking genes in asf1Δ cells, indicating that repression of these genes was not mediated by the spread of Sir2 into adjacent euchromatin (Figure 2.4C). Secondly, deletion of SIR2 in asf1Δ yaf9Δ and asf1Δ swr1Δ strains restored the mRNA levels of the genes between the HMR and telomere, but not those flanking the left side of the HMR (Figure 2.4D and Figure 2.4E). Both YCR094W and YCR095C that depended on Asf1 for expression remained repressed even when SIR2 was     46 deleted (Figure 2.4D and Figure 2.4E). Unlike the other two HMR-flanking genes, the expression of GIT1 was restored to at least wildtype levels in both asf1Δ yaf9Δsir2Δ and asf1Δ swr1Δsir2Δ (Figure 2.4D and Figure 2.4E). YCR099C and YCR100C exhibited higher than wildtype levels of mRNA in the asf1Δ swr1Δ sir2Δ triple mutant (Figure 2.4E). Together, these data suggested that in the absence of a functional SWR1-C, SIR complexes spread into nearby euchromatic regions, whereas asf1Δ-dependent repression of HMR-flanking genes was not due to SIR activity.             47  Figure 2.4 Asf1 regulated HMR-proximal gene expression in a Sir2-independent manner mRNA levels of heterochromatic genes in the indicated strains were carried out in parallel. mRNA levels were determined by qRT-PCR and normalized to TUB1. Data shown are the average of three independent experiments. Error bars represent standard error of values of the three biological replicates. *, p-value < 0.10; **, p-value < 0.05 when compared with the Wildtype strain using a two-tailed student t-test. (A) Deletion of SIR2 rescued all expression defects observed in yaf9Δ strains. mRNA levels of wildtype (black), yaf9Δ (light gray), sir2Δ (dark gray) and yaf9Δ sir2Δ (white) strains. (B) Expression profile of tested genes in cells lacking SWR1 resembled those lacking YAF9. Loss of SIR2 in swr1Δ background also rescued the expression of all tested genes to wildtype levels. mRNA levels of wildtype (black), swr1Δ (light gray), sir2Δ (dark gray) and swr1Δ sir2Δ (white) strains. (C) Loss of SIR2 in strains lacking ASF1 was unable to rescue the repression of HMR-flanking genes. mRNA levels of wildtype (black), swr1Δ (light gray), asf1Δ (dark gray) and asf1Δ sir2Δ (white) strains. (D) Loss of SIR2 in an asf1Δ yaf9Δ background was able to rescue all Yaf9-specific transcriptional defects but not Asf1-specific defects around the HMR. mRNA levels of wildtype (black), asf1Δ (light gray), yaf9Δ (dark gray), asf1Δ yaf9Δ (white) and asf1Δ yaf9Δ sir2Δ (medium gray) strains. (E) Similarly, deletion of SIR2 rescued gene expression of subtelomeric genes to wildtype levels in asf1Δ swr1Δ strains, but not the HMR-flanking genes. mRNA levels of wildtype (black), asf1Δ (light gray), swr1Δ (dark gray), asf1Δ swr1Δ (white) and asf1Δ swr1Δ sir2Δ (medium gray) strains.  2.3.5 Dysregulation of Heterochromatin Boundaries in asf1Δ yaf9Δ was not the Underlying Cause of the Severe Growth Defects The complex relationship between Sir2 localization and gene expression described above raised the possibility that the growth phenotype of asf1Δ yaf9Δ was a reflection of additional AFigure 4B CD EHMR-flanking Subtelomeric HMR-flanking Subtelomeric HMR-flanking SubtelomericHMR-flanking Subtelomeric HMR-flanking Subtelomeric00.511.52YCR094 YCR095 GIT1 YCR099 YCR100 RDS1mRNA abundance normalized over TUB1Wildtypeyaf9sir2yaf9 sir200.511.52YCR094 YCR095 GIT1 YCR099 YCR100 RDS1mRNA abundance normalized over TUB1Wildtypeswr1sir2swr1 sir200.511.522.5YCR094 YCR095 GIT1 YCR099 YCR100 RDS1Wildtypeasf1sir2asf1 sir2mRNA abundance normalized over TUB100.511.52YCR094 YCR095 GIT1 YCR099 YCR100 RDS1mRNA abundance normalized over TUB1Wildtypeasf1yaf9asf1 yaf9asf1 yaf9 sir200.511.52YCR094 YCR095 GIT1 YCR099 YCR100 RDS1mRNA abundance normalized over TUB1Wildtypeasf1swr1asf1 swr1asf1 swr1 sir2****** ** ****** * ** ****** **** ** ******** ****** ************ ** **yaf9 sir2 swr1 sir2 asf1 sir2asf1yaf9 sir2 asf1swr1 sir2    48 cellular processes. We therefore expected that aberrant SIR chromatin structures, caused by the simultaneous loss of ASF1 and YAF9, did not contribute to the pronounced growth defects. Indeed, loss of SIR2 did not suppress the synergistic genetic interaction between ASF1 and YAF9, indicating that the growth defect was due to other functions of Asf1 and SWR1-C (Figure 2.5A).    To gain a better mechanistic understanding of the genetic interaction between ASF1 and YAF9, we focused on Asf1’s role in nucleosome assembly at heterochromatin by using two asf1 alleles (H36A/D37A and H39A/K41A) that exhibit reduced silencing at telomeres when CAF-1 is mutated (Sharp et al. 2001; Daganzo et al. 2003). Plasmids containing the asf1 alleles were transformed into strains lacking both ASF1 and a gene encoding for the CAC2 subunit of CAF-1. As expected, asf1H36A/D37A and asf1H39A/K41A mutants alone did not exhibit a significant growth defect (Figure 2.5B) (Daganzo et al. 2003). Strains lacking CAC2 with an asf1 allele showed slight growth defects compared to wildtype strains (Figure 2.5B). Consistent with previous work, the asf1H39A/K41A allele exhibited a stronger phenotype than asf1H36A/D37A (Figure 2.5B). cac2Δ yaf9Δ asf1H36A/D37A and cac2Δ yaf9Δ asf1H39A/K41A strains, however, did not demonstrate a strong synergistic interaction (Figure 2.5B). Our genetic data therefore revealed that the synergistic genetic interaction between ASF1 and YAF9 was not due to Asf1’s functions in various aspects of heterochromatin biology.        49   Figure 2.5 Silencing defects were not causal for asf1Δ yaf9Δ synthetic growth defect   (A) Deletion of SIR2 did not suppress the growth defects observed in asf1Δ yaf9Δ and asf1Δ swr1Δ strains. Ten-fold serial dilution of strains were plated and incubated at the indicated condition. (B) Heterochromatin-specific nucleosome assembly mutants of ASF1 did not genetically interact with YAF9. Ten-fold serial dilutions of indicated strains carrying pASF1, pasf1 H36A/D37A, or pasf1H39A/K41A were plated on SC-TRP media and incubated for two to three days at the indicated conditions.    2.3.6 The Synergistic Genetic Interaction between ASF1 and YAF9 was Mediated via Rtt109-Dependent H3K56 Acetylation To test if Asf1’s ability to act as a global chaperone in nucleosome assembly was important for its genetic interaction with YAF9, we used the well-characterized asf1V94R mutant to assay whether a complete loss of histone H3/H4 binding can replicate the synergistic interaction between ASF1 and YAF9 (Mousson et al. 2005). We found that similar to asf1Δyaf9Δ, asf1V94R yaf9Δ grew much slower at 16°C and showed increased sensitivity to a low concentration of hydroxyurea, implicating Asf1 and H3 interaction in this pathway (Figure 2.6A). Since the physical association between Asf1 and H3 is important to mediate the acetylation of lysine 56 on H3 by Rtt109, we subsequently tested whether RTT109 also displayed a genetic interaction with YAF9 and genes encoding for other subunits of SWR1-C. YPD 30CYPD 16C10mM HUWildTypeasf1∆yaf9∆sir2∆asf1∆,  yaf9∆asf1∆.  sir2∆yaf9∆,  sir2∆asf1∆,  yaf9∆,  sir2∆WildTypeasf1∆swr1∆sir2∆asf1∆,  swr1∆asf1∆.  sir2∆swr1∆,  sir2∆asf1∆,  swr1∆,  sir2∆Figure 5ABSc -trp 30CSc -trp16C10mM HUpASF1pasf1H39A/K41Acac2∆  +  pASF1yaf9∆  +pASF1cac2∆∆  +  pasf1H39A/K41Ayaf9∆  +  pasf1H39A/K41Acac∆,  yaf9∆  +  pASF1cac2∆,  yaf9∆  +  pasf1H39A/K41ApASF1pasf1H36A/D37Acac2∆  +  pASF1yaf9∆  +pASF1cac2∆  +  pasf1H36A/D37A  yaf9∆  +  pasf1H36A/D37Acac∆,  yaf9∆  +  pASF1cac2∆,  yaf9∆  +  pasf1H36A/D37Aasf1∆ asf1∆YPD 30CYPD 16C10mM HUWildTypeasf1∆yaf9∆sir2∆asf1∆,  yaf9∆asf1∆.  sir2∆yaf9∆,  sir2∆asf1∆,  yaf9∆,  sir2∆WildTypeasf1∆swr1∆sir2∆asf1∆,  swr1∆asf1∆.  sir2∆swr1∆,  sir2∆asf1∆,  swr1∆,  sir2∆Figure 5ABSc -trp 30CSc -trp16C10mM HUpASF1pasf1H39A/K41Acac2∆  +  pASF1yaf9∆  +pASF1cac2∆∆  +  pasf1H39A/K41Ayaf9∆  +  pasf1H39A/K41Acac∆,  yaf9∆  +  pASF1cac2∆,  yaf9∆  +  pasf1H39A/K41ApASF1pasf1H36A/D37Acac2∆  +  pASF1yaf9∆  +pASF1cac2∆  +  pasf1H36A/D37A  yaf9∆  +  pasf1H36A/D37Acac∆,  yaf9∆  +  pASF1cac2∆,  yaf9∆  +  pasf1H36A/D37Aasf1∆ asf1∆    50 As previously seen in the asf1Δyaf9Δ double mutants, RTT109 displayed a synthetic genetic interaction with YAF9. Furthermore, the growth fitness of all double mutants was significantly compromised compared to their respective single mutants under stress-induced conditions (Figure 2.6B).     Figure 2.6 Synergistic genetic interaction between ASF1 and YAF9 was mediated by Rtt109-dependent H3K56 acetylation (A) YAF9 genetically interacted with asf1V94R. (B) RTT109 genetically interacted with all non-essential genes encoding for SWR1-C subunits.  (C) H3K56R genetically interacted with YAF9. Ten fold-serial dilutions of strains were plated and incubated at the labeled conditions.    To confirm if the genetic interaction we observed was mediated via acetylation of H3K56, we took advantage of existing H3K56 alleles containing a point mutation at lysine 56 to either glutamine or arginine, mimicking a constitutively hyperacetylated lysine or constitutively hypoacetylated lysine, respectively (Miller et al. 2008). In agreement with published data, the yaf9ΔH3K56R strain had significantly reduced growth fitness compared to either of the single mutants alone, while the yaf9ΔH3K56Q strain did not display a strong Figure 6WildTypeasf1∆yaf9∆asf1∆,  yaf9∆asf1∆  +  pASF1  asf1∆  +  pasf1V94Rasf1∆,  yaf9∆  +  pASF1asf1∆,  yaf9∆  +  pasf1V94RWildTypertt109∆yaf9∆rtt109∆,  yaf9∆htz1∆rtt109∆.  htz1∆swr1∆rtt109∆,  swr1∆swc2∆rtt109∆,  swc2∆swc3∆rtt109∆,  swc3∆WildTypertt109∆yaf9∆rtt109∆,  yaf9∆swc5∆rtt109∆,  swc5∆swc6∆rtt109∆,  swc6∆H3/H4yaf9∆  H3/H4H3K56QH3K56Ryaf9∆  H3K56Q  yaf9∆  H3K56RSc -TRP 30C Sc -TRP 30CSc -TRP 16C Sc -TRP 16C10mM HU 10mM HUYPD 30CYPD 16C10mM HUA B C    51 growth defect (Figure 2.6C) (Wang et al. 2009). Furthermore, similar experiments testing for H3K9 acetylation, which is also mediated by Rtt109, revealed that the genetic interaction was specific to Asf1’s role in H3K56 acetylation and not Rtt109-mediated H3K9 acetylation (data not shown). These results indicated that the genetic interaction between ASF1 and YAF9 was a result of failure to acetylate H3K56 in an Asf1/Rtt109-dependent manner.  2.3.7 Asf1 and H3K56 Acetylation Promoted H2A.Z Occupancy at Subtelomeric Gene Promoters We again utilized our six genes of interest to explore the genetic connection between H3K56 acetylation and SWR1-C in more detail, since all but YCR099 have robust H2A.Z promoter occupancy. Because we previously demonstrated that H3K56ace-containing promoters preferentially lose H2A.Z in a yaf9 hypomorph, we next addressed whether H3K56 acetylation and Asf1 are involved in H2A.Z occupancy at gene promoters (Wang et al. 2009). We used ChIP-qPCR to assay H2A.Z localization at the promoters of heterochromatin-proximal genes in strains that contain either asf1Δ or the H3K56R non-acetylable mutation. H2A.Z was enriched to varying degrees at the promoters of all genes we examined and as expected, a loss of YAF9 led to a total depletion of H2A.Z at each promoter (Figure 2.7A). Remarkably, deletion of ASF1 resulted in a significant decrease in H2A.Z levels across all promoters tested; this result demonstrated that Asf1 enhanced H2A.Z deposition but was not required (Figure 2.7A). Next, we asked if the reduction of H2A.Z at subtelomeric gene promoters seen in asf1Δ cells was due to a loss of H3K56ace. We found that H2A.Z enrichment in the H3K56R mutant was reduced to levels similar to those of     52 asf1Δ, suggesting that Asf1-mediated H3K56 acetylation promoted H2A.Z localization at the promoters of heterochromatin-proximal genes (Figure 2.7B).    Figure 2.7 Asf1 and H3K56 acetylation promoted H2A.Z occupancy at subtelomeric and euchromatic gene promoters H2A.Z occupancy at promoters of indicated genes was measured by ChIP-qPCR using the anti-FLAG antibody (Sigma). H2A.Z enrichment was normalized to the reference gene PRP8. Error bars represent standard error of three biological replicates. *, p-value < 0.10; **, p-value < 0.05 when compared with the Wildtype strain using a two-tailed student t-test. (A) Occupancy of H2A.Z at heterochromatin-proximal gene promoters was reduced significantly in cells lacking ASF1. (B) Similarly, promoter H2A.Z levels of subtelomeric genes were significantly reduced in a H3K56R mutant. (C) (D) Asf1 and H3K56 acetylation are both required for normal H2A.Z occupancy at euchromatic promoters. Loss of YAF9 led to a complete loss of H2A.Z at gene promoters.     A BHMR-flanking Subtelomeric02468101214no tagHTZ1-FLAGH3K56R HTZ1-FLAGyaf9 HTZ1-FLAGyaf9 H3K56R HTZ1-FLAGHMR-flanking Subtelomeric02468101214YCR094 YCR095 GIT1 YCR099 YCR100 RDS1H2A.Z levels normalized to PRP8no tagHTZ1-FLAGasf1 HTZ1-FLAGyaf9 HTZ1-FLAGasf1 yaf9 HTZ1-FLAGH2A.Z levels normalized to PRP8Figure 7YCR094 YCR095 GIT1 YCR099 YCR100 RDS1asf1  H3K56R05101520GID7 STE50 BUD3 MRK1 BZZ1 YDC1no tagHtz1-FLAGasf1 Htz1-FLAGyaf9 Htz1-FLAGasf1 yaf9 Htz1-FLAG024681012GID7 STE50 BUD3 MRK1 BZZ1 YDC1no tagHtz1-FLAGH3K56R Htz1-FLAGyaf9 Htz1-FLAGH3K56R yaf9 Htz1-FLAGH2A.Z levels normalized to PRP8H2A.Z levels normalized to PRP8DCasf1  H3K56REuchromatic genes Euchromatic genes****** ************** ************************************************************* ********* **************** **** ****************    53 Since our genetic data suggested that the relationship between Asf1 and H2A.Z was not exclusive to the heterochromatic region, we extended the analysis to include euchromatic promoters that are H2A.Z-enriched. We selected six genes from either previously published primer sets or genome-wide H2A.Z ChIP data sets. As expected, H2A.Z was highly enriched at the promoter of these genes in wildtype cells (Figure 2.7C and Figure 2.7D). Analogous to the effects at heterochromatic gene promoters, loss of ASF1 also led to a reduction in H2A.Z levels at all six genes we assayed (Figure 2.7C). Furthermore, H2A.Z levels were also significantly reduced in the H3K56R mutant, suggesting that the acetylation of H3K56 by Asf1 is important for H2A.Z occupancy at euchromatic gene promoters (Figure 2.7D).   2.4 Discussion  We have teased apart the region-specific and global interactions between Asf1 and SWR1-C, particularly its Yaf9 subunit, and expanded upon the established linkages between them. First, we identified region-specific changes in gene expression in the absence of either Asf1 or Yaf9 within a 35kb region on the right telomere of chromosome III. Whereas loss of SWR1-C function resulted in repression of heterochromatin-proximal genes as a result of Sir2 spread, loss of Asf1 led to repression of only HMR-proximal genes in a Sir2-independent manner. Nevertheless, we found that Asf1 cooperated with Yaf9 to modulate the telomere-heterochromatin boundary of Chr III R to prevent the spread of SIR complexes into the subtelomeric region. This interaction was recapitulated at three additional telomere ends upon genome-wide analysis of Sir2 occupancy.  In agreement with these effects likely being region-specific, genetic analysis revealed that global loss of SIR2 did not rescue the growth     54 defect in asf1Δ yaf9Δ double mutants. Next, we demonstrated that a strong synergistic genetic interaction existed between genes encoding for SWR1-C and factors in the H3K56 acetylation pathway. Loss of H3K56 acetylation, either by the deletion of the Asf1 histone chaperone or mutation of the K56 residue, diminished H2A.Z levels at the promoters of subtelomeric and euchromatic genes.   Maintenance of euchromatin-heterochromatin boundaries requires the concerted activities of chromatin remodeling complexes. RT-qPCR analysis of genes adjacent to the right telomere of Chr III showed that SWR1-C, but not Asf1, regulated this heterochromatin boundary. After careful dissection of how Asf1 and Yaf9 function in restricting Sir2 spread around the HMR and the Chr III R subtelomeric region, however, we found that the connection between Asf1 and SWR1-C was more complex. Repression of telomere-proximal genes in yaf9Δ cells was due to the loss of a functional heterochromatin boundary, which resulted in a spread of SIR complexes over nearby genes. On the other hand, loss of ASF1 alone did not affect the function of the heterochromatin boundary, as evidenced by the lack of Sir2 spread in asf1Δ cells and normal expression levels of subtelomeric genes. We demonstrated by ChIP-qPCR and by ChIP-on-chip that combined loss of ASF1 and YAF9 led to increased spread of Sir2 and elevated levels of Sir2, suggesting that Yaf9 and Asf1 cooperated to restrict the spread of SIR proteins into nearby subtelomeres. Despite the dramatic enhancement of Sir2 occupancy over the subtelomeric region, mRNA levels of the three telomere-proximal genes we examined were reduced to the same level in the asf1Δ yaf9Δ double mutant as compared to the yaf9Δ single mutant. It has been suggested that SIR complexes function in a dosage-dependent manner in a reporter gene assay when SIR3 and SIR4 are overexpressed (Strahl-    55 Bolsinger et al. 1997). However, our combined occupancy and expression approach at Chr III R supported a threshold effect for SIR-dependent silencing. Specifically, the presence of Sir2 over the locus, regardless of whether it constitutes a two-fold increase of Sir2 occupancy in yaf9Δ or up to five-fold increase of Sir2 occupancy in asf1Δ yaf9Δ, led to the same level of gene repression of subtelomeric genes. Our genome-wide analysis of Sir2 occupancy patterns also demonstrated that the functional connection between Asf1 and SWR1-C was telomere-specific. Altogether, our findings showed that SWR1-C and Asf1 act cooperatively at a subset of telomere-proximal ends to limit the spread of SIR complexes onto nearby subtelomeric regions and that the SIR complex mediated silencing through a threshold mechanism.   The HMR is unique in that the silent cassette is flanked by silencers that initiate the formation of silenced chromatin (Guillemette et al. 2005). H2A.Z protects genes on either sides of the HMR from SIR-mediated silencing and loss of the histone variant leads to repression of the nearby genes (Meneghini et al. 2003; Babiarz et al. 2006; Li et al. 2005). Consistent with this, we found that deletion of YAF9 and SWR1 led to decreased transcript levels of HMR-proximal genes. Furthermore, asf1Δ and asf1Δ yaf9Δ cells also exhibited the same level of transcriptional defects, suggesting that SWR1-C and Asf1 regulate gene expression in the same pathway. Further analysis, however, revealed that repression of YCR094, YCR095 and GIT1 in SWR1-C mutants resulted from a defective heterochromatin boundary and Asf1-dependent expression defects were not a result of Sir2 spreading. The lack of correlation between SIR occupancy and changes in gene expression indicated that the silencing of HMR-proximal genes in asf1Δ cells was not due to SIR spread. Moreover, the persistence of a     56 transcriptional defect in asf1Δ yaf9Δ sir2Δ and asf1Δ swr1Δ sir2Δ triple mutants further supported that decreased mRNA levels of YCR094 and YCR095 resulted from an Asf1-specific transcriptional defect that was not linked to either Sir2 or H2A.Z-dependent boundary activity. Therefore, even though loss of ASF1 and YAF9 both led to gene repression around the HMR, Asf1 and SWR1-C mediated these effects through distinct mechanisms. Perhaps, by promoting H3K56 acetylation, Asf1 is directly involved in transcriptional activation of these subtelomeric genes by promoting nucleosome clearance at promoters (Adkins et al. 2004; Korber et al. 2006; Williams et al. 2008; Tolkunov et al. 2011).   In addition to region-specific effects, we also systematically characterized the genetic interaction profiles of ASF1 and RTT109 with all non-essential genes encoding for subunits of the SWR1-C complex.  We found that all SWR1-C-encoding genes displayed a synthetically sick genetic interaction with ASF1 and RTT109 in an H3K56 acetylation-dependent manner. Given the severity of the genetic interaction, our data suggest that the functional relationship between H3K56 acetylation and SWR1-C goes beyond their interaction at heterochromatin. To add to our current understanding of SWR1-C-mediated H2A.Z deposition, we demonstrated that the level of chromatin-associated H2A.Z was also dependent on the histone chaperone Asf1. Deletion of ASF1 or introduction of an unacetylable allele of H3K56 (H3K56R) resulted in an intermediate reduction in H2A.Z levels at promoters of heterochromatin-proximal and euchromatic genes. Based on previous genome-wide expression data in htz1Δ, it was not surprising that changes in H2A.Z levels in cells lacking ASF1 were not correlated to changes in the expression of these lowly expressed genes.  We speculate based on our data that Asf1 might directly influence the deposition of     57 H2A.Z through an H3K56ace-dependent pathway. Reduced H2A.Z occupancy in the H3K56R mutant suggests that acetylation promotes H2A.Z deposition. Alternatively, it is possible that H2A.Z-containing nucleosomes are more stable when H3K56 is also acetylated within the same nucleosome. While our study focused on specific heterochromatin-proximal and euchromatic loci, a recent publication demonstrated that the H3K56Q mutant promotes removal of the H2A.Z/H2B heterodimer by SWR1-C across the yeast genome (Watanabe et al. 2013). Hence, it remains to be seen whether the locus-specific effects of H3K56 acetylation-dependent H2A.Z deposition extends into all genomic regions similar to H3K56Q. Moreover, further studies are required to elucidate how both the absence and the presence of this histone modification can lead to loss of H2A.Z at promoters.      58 Chapter 3: H3K56 Acetylation is Required for Positioning H2A.Z and for Maintaining H2A.Z Occupancy at Gene Promoters   3.1  Introduction  The eukaryotic genome is packaged into the nucleus as a compact structure called chromatin. The fundamental repeating unit of chromatin is the nucleosome, a 146 bp DNA fragment wrapped around a histone core containing two copies each of H2A, H2B, H3 and H4. Over the last few years, it has become clear that nucleosomes are not assembled randomly across the genome (Brogaard et al. 2012; Jiang and Pugh 2009; Lee et al. 2007; Mavrich et al. 2008; Shivaswamy et al. 2008; Whitehouse and Tsukiyama 2006). Instead, they are highly organized and tightly regulated by a number of factors. The underlying DNA sequence, dedicated chromatin remodeling complexes and various transcription factors all contribute to and influence the position of nucleosomes. This is particularly true at yeast gene promoters where two highly positioned nucleosomes flank the nucleosome free region (NFR) just upstream of the transcriptional start site (TSS) (Mavrich et al. 2008; Shivaswamy et al. 2008).  In addition to the four canonical histones that constitute the major protein component of the nucleosome, nucleosomes may also contain histone variants that mark unique chromatin domains. H2A.Z is one of the most highly conserved histone variants throughout eukaryotic     59 evolution and its three-dimensional structure is very similar to canonical cousin, H2A (Suto et al. 2000). Studies in yeast have shown that H2A.Z is required for a wide range of genome functions including gene expression, maintenance of silencing boundaries, DNA repair, cell cycle progression and chromosome stability (Malik and Henikoff 2003; Meneghini et al. 2003; Kobor et al. 2004; Mizuguchi et al. 2004; Kalocsay et al. 2009; Dhillon et al. 2006; Krogan et al. 2004; Rangasamy et al. 2004). As evidence of its central role in a variety of cellular functions, H2A.Z is essential for viability in higher eukaryotes such as flies, frogs and mice (Faast et al. 2001; Iouzalen et al. 1996; van Daal and Elgin 1992).   H2A.Z is considered a genome-wide signature of eukaryotic promoters and enhancer elements. Despite being found in only 10% of all nucleosomes, the majority of chromatin-bound H2A.Z variants are deposited at the +1 and -1 nucleosomes flanking the NFR (Albert et al. 2007; Barski et al. 2007; Mavrich et al. 2008). Deposition of H2A.Z at specific genomic regions is solely catalyzed by the multi-subunit SWR1-C ATPase-dependent chromatin-remodeling complex (Kobor et al. 2004; Krogan et al. 2004; Mizuguchi et al. 2004). Using the energy from ATP hydrolysis, SWR1-C evicts two nucleosomal H2A/H2B units for two H2A.Z/H2B heterodimers, resulting in a variant nucleosome containing two copies of H2A.Z (Luk et al. 2010). While the mechanism of SWR1-C mediated H2A.Z exchange is well understood from in vitro experiments, the mechanisms of SWR1-C recruitment to sites of H2A.Z exchange in vivo remains an area of significant interest. Early studies demonstrated that NFR formation by the Reb1 general transcription regulatory protein and the RSC chromatin remodeler is sufficient to induce H2A.Z deposition (Raisner et al. 2005; Hartley and Madhani 2009). In addition, it was recently elucidated that the NFR     60 and the adjoining nucleosome plays a direct role in targeting SWR1-C to sites of H2A.Z exchange at gene promoters (Ranjan et al. 2013; Yen et al. 2013).   Consistent with H2A.Z’s role in maintaining telomeric boundaries, SWR1-C also catalyzes H2A.Z exchange at the boundaries of euchromatin and heterochromatin (Meneghini et al. 2003; Zhang et al. 2004). Loss of H2A.Z and SWR1-C activity at heterochromatin boundaries result in repression of subtelomeric genes as a consequence of SIR complex spread (Meneghini et al. 2003; Lu and Kobor 2014). Intriguingly, two post-translational modifications have been shown to be critical for H2A.Z occupancy at telomeres. Firstly, H4K16 acetylation by the SAS histone acetyltransferase is required targeting H2A.Z deposition at the right telomere of ChrVI (Shia et al. 2006). Secondly, in Chapter 2 of this dissertation, we identified that Asf1-dependent H3K56 acetylation is required for maintaining H2A.Z levels at subtelomeric regions of ChrIII R (Lu and Kobor 2014). Physical interaction between Asf1 and the SAS complex suggests the two complexes might act in the same pathway to regulate H2A.Z deposition (Osada et al. 2005; 2001).   The primary role of Asf1 is a histone chaperone for the H3/H4 heterodimer. However, research in the last decade has revealed a key role of Asf1 in many auxiliary functions. For example, Asf1 is part of a class of histone chaperones that promote post-translational modifications (PTM) of histones. Binding of the H3/H4 heterodimer by Asf1 exposes the globular domain to Rtt109 thus allowing for acetylation of H3K56 (Recht et al. 2006; Tsubota et al. 2007; Schneider et al. 2006; Driscoll et al. 2007; Adkins et al. 2007). Furthermore, Asf1 is connected to a number of biological functions from DNA replication,     61 DNA repair, and transcription-coupled nucleosome assembly and disassembly (Tyler et al. 1999; Adkins et al. 2004; Adkins and Tyler 2004; Schwabish and Struhl 2006; Adkins et al. 2007; Mousson et al. 2007; Gkikopoulos et al. 2009; Takahata et al. 2009). By promoting physical interactions with various chromatin-remodeling complexes, Asf1 plays a key role in establishing unique chromatin neighbourhoods.   Here, we examined the functional connection between H3K56 acetylation and H2A.Z occupancy with respect to known regulatory mechanisms for H2A.Z deposition. Firstly, we studied whether Asf1-mediated H2A.Z occupancy at heterochromatin boundaries is dependent on H4K16 acetylation by the SAS complex, and secondly, whether H3K56 acetylation is required for H2A.Z occupancy genome-wide. We determined that H3K56 acetylation did not regulate H2A.Z occupancy in a SAS-dependent manner. Instead, H3K56 acetylation was required for maintaining promoter H2A.Z levels genome-wide. Our data suggested a key role for H3K56 acetylation in positioning H2A.Z nucleosomes flanking the NFR at a subset of gene promoters.   3.2 Materials and Methods  3.2.1 Yeast Strains and Plasmids All strains used in this study are listed in Table 3.1. Yeast strains were generated using standard genetic techniques including homologous recombination and genetic crosses followed by tetrad dissection (Ausubel 1987). Complete deletion of genes and 3’ end integration of an in frame 3xFLAG tag (Gelbart et al. 2001) were achieved using one-step     62 gene integration PCR-amplified modules. Plasmid shuffle experiments were performed using 5-FOA to evict URA3 plasmid pRS316[H3 H4], and select for pRS316[H3K56R H4], pRS316[H3K56Q H4], pRS316[H3K9R H4], pRS316[H3K9Q H4], and pRS316[H3K9RK56R H4]. These plasmids were a generous gift from Ann Kirchmaier (Purdue University).  Table 3.1 Yeast strains used in this study Strain number Relevant Genotype  MKY6 W303, MATA, ADE2, can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ  MKY1754  MKY1755 MKY1756 MKY6, HTZ1-FLAG::KAN MKY6, HTZ1-FLAG::KAN, asf1::NAT MKY6, HTZ1-FLAG::KAN, sas2::HIS  MKY1757 MKY6, HTZ1-FLAG::KAN, asf1::NAT, sas2::HIS  MKY1758 MKY6, SWC2-VSV::KAN  MKY1759 MKY6, SWC2-VSV::KAN, asf1::HIS  MKY1760 MKY6, SWC3-VSV::KAN  MKY1761 MKY6, SWC3-VSV::KAN, asf1::HIS  MKY1630 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2-HHF2]  MKY1633 MKY6, hht1-hhf1::LEU2 hht2-hhf2::HIS [pRS314, HHT2 K56R HHF2]   3.2.2 Growth and Genotoxic Sensitivity Assays Overnight cultures grown in YP-dextrose were diluted to OD600 0.5. Cells were 10-fold serially diluted and spotted onto solid YPD plates or plates with 10mM hydroxyurea. The plates were then incubated at the indicated temperature for 36 hours. The 16°C plates were incubated for 96 hours.      63 3.2.3 ChIP-qPCR and ChIP-on-chip Chromatin immunoprecipitation (ChIP) experiments were performed as described previously (Schulze et al. 2009). In brief, 250 ml of cells were grown in YP-dextrose to an OD600 of 0.5-0.6 from OD600 of 0.15 and were crosslinked with 1% formaldehyde for 20 minutes before chromatin was extracted. The chromatin was sonicated (Bioruptor, Diagenode, Spart, NJ: 10 cycles, 30s on/off, high setting) to yield an average DNA fragment of 500bp. Anti-FLAG antibody (4.2µl, Sigma) was coupled to 60µl of protein A magnetic beads (Invitrogen). After reversal of the crosslinking and DNA purification, the immunoprecipitated and input DNA were analyzed by quantitative real-time PCR (qPCR). Samples were analyzed in triplicate for three independent ChIP experiments. Statistical significance was assessed using Student’s t test.  Primer sequences are listed in Table 3.2.   For microarray analysis, after reversal of crosslinking and DNA purification, the DNA was amplified with two rounds of T7 RNA polymerase amplification and hybridized to Affymetrix 1.0R S. cerevisiae tiling microarray. Experiments were carried out in duplicates. A modified version of the model-based analysis of tiling arrays (MAT) algorithm was used to normalize and average the duplicates. The data was normalized using both input DNA and a mock IP control. Relative occupancy scores were calculated for each probe using a 300bp sliding window. Enriched features had at least 50% of the probes contained in the feature above the 1.5 threshold. Promoters were defined as the 500bp upstream of the ORF.         64 Table 3.2 Primer sequences used in this study Name Method Primer sequence (5’-3’) HMR left F HMR left R HMR right F HMR right R ChrIII tel R F ChrIII tel R R YCR094W Prom F YCR094W Prom R YCR095C Prom F  ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP CTACCTTGTTGTATATAGGC AACAGCGATAGTGCGAGGAT TCTACAATGCAACCCCACAA TTGAACCGATGATCTCCACA AAGGCGGTGTTATTTGTTGC TTATGCGGCCGAATGATACT GCAAACCCCTCTACAATCCA CAAAAGTGAAAGCGACCCATA TACCGTATGCGGTATAATGA YCR095C Prom R ChIP GTCTCCACTTTAGAACATCT GIT1 Prom F ChIP TTCATGAATTTCCTTACTGGAC GIT1 Prom R YCR099C Prom F  YCR099C Prom R YCR100C Prom F YCR100C Prom R RDS1 Prom F ChIP ChIP ChIP ChIP ChIP ChIP GTTGACTAGTCACAAGAAACAG TGCTACTGGTGATCTGGGAAA CTGATCCATCTGGCGTTGTA GCAAGGATTCTGACTTTACTGG  CTCGTTATGCCCGTCATCTT TGTGCTATCTAAGAGGATGGTTCA  RDS1 Prom R PRP8 ORF F PRP8 ORF R T6R-H F T6R-H R T6R-E F T6R-E R T6R-C F T6R-C R ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP ChIP GAATCCATCAGAGCATTTCA GGATGTATCCAGAGGCCAAT AACCCGCGTATTAAGCCATA GAAAGTTTGGATGCTAGCAAGGGC GCATAGCCTTTGAAAACGGCG GTCTCGTAGGTAGCTTTCAC CGGTGTTCCTTTACAAACCC CGTTCTTCTTGGCCCTTATC CATCATCGGTGGTTTTGTCGTG      65 3.2.4 Chromatin Association Assay Chromatin association assays were performed as previously described (Wang et al. 2009). In brief, cells were diluted to OD600 0.15 and collected at logarithmic phase. Following collection, cells were incubated in pre-spheroblast buffer (100 mM PIPES/KOH [pH 9.4], 10 mM DTT, 0.1% sodium azide) for 10 minutes at room temperature rotating, then spheroblasted for 20 minutes at 37C with 20mg/ml Zymoylase-100T (Seikagaku Corporation). Spheroblasts were washed with wash buffer (50 mM HEPES/KOH [pH 7.5], 100mM KCl, 2.5 mM MgCl2, 0.4 M sorbitol) and resuspended in equal volume of EB (50 mM HEPES/KOH [pH 7.5], 100mM KCl, 2.5 mM MgCl2, 1 mM DTT, 1 mM PMSF, and Complete Protease Inhibitor cocktail [Roche]). Cells were lysed with 1% Triton X-100. Whole cell extracts were saved and the remaining lysate were separated into the chromatin pellet and supernatant fraction by centrifugation through EBSX (EB + 0.25% Triton X-100 and 30% sucrose). The three fractions were subsequently analyzed by SDS-PAGE and immunoblotted.    3.3 Results  3.3.1 Asf1-Dependent H2A.Z Promoter Occupancy was not Mediated by the SAS Histone Acetyltransferase In chapter 2, we found that H2A.Z occupancy at subtelomeric promoters is dependent partly on the Asf1 histone chaperone. Loss of H3K56 acetylation, either by deleting ASF1 or by introducing an unacetylable mutation (H3K56R), resulted in a decrease in H2A.Z levels at     66 the promoters of genes proximal to the telomere of Chr III. Based on previous work, we tested whether Asf1 recruits the SAS complex to promote H2A.Z deposition at ChrIII R (Osada et al. 2005; Shia et al. 2006). We first created strains that lacked genes encoding for ASF1 and the catalytic subunit of the SAS complex, SAS2. While strains lacking SAS2 grew better than the asf1Δ strain, we found that the asf1Δ sas2Δ double mutant more closely resembled the asf1Δ strain (Figure 3.1A).   The acetylation of H4K16 by the SAS histone acetyltransferase complex is a key chromatin modification that restricts the spread of SIR silencing complexes at heterochromatin boundaries (Kimura et al. 2002; Suka et al. 2002; Oki and Kamakaka 2004). To examine if Asf1 was required for H4K16 acetylation at the well-characterized heterochromatin boundaries of ChrIII R, we first compared levels of H4K16 acetylation in strains lacking ASF1 or SAS2 (Figure 3.1B). Normalized for nucleosome content, we found that upon ASF1 deletion, there was a significant reduction of H4K16 acetylation at all three boundaries (Figure 3.1C). As expected, sas2Δ cells had very low levels of acetylated H4K16 at the heterochromatin boundaries of ChrIII R. To test whether the SAS complex is required for H2A.Z occupancy in a similar manner to Asf1, we assessed the levels of H2A.Z at the ChrIII R heterochromatin boundaries in sas2Δ strains. Unlike asf1Δ, a strain lacking SAS2 had H2A.Z levels similar to that of a wildtype strain, indicating that neither the SAS complex nor H4K16 acetylation was required for H2A.Z deposition at the boundaries of ChrIII R (Figure 3.1D).       67  Figure 3.1 Asf1-dependent H2A.Z promoter occupancy did not require the SAS histone acetyltransferase (A) Ten-fold serial dilution assay of indicated strains plated on YPD media with or without HU and incubated at the labeled temperatures. (B) Schematic representation of the boundaries of the HMR locus and the ChrIII R telomere. (C) H4K16 acetylation partially depended on Asf1. ChIP-qPCR of H4K16 acetylation at heterochromatin boundaries of ChrIII R in wildtype, asf1Δ, and sas2Δ strains. Enrichment of H4K16 acetylation was normalized to nucleosome density as measured by H3. (D) H2A.Z deposition at heterochromatin boundaries of ChrIII R does not depend on SAS2. ChIP-qPCR of H2A.Z-FLAG in wildtype, asf1Δ, and sas2Δ strains. H2A.Z enrichment was normalized to enrichment level at the PRP8 ORF. (C)(D) Error bars represent standard errors of the means for three independent experiments.     A"D"C"B"Figure"1"Chr$III$R$YCR094W( RDS1(GIT1((YCR099C((YCR100C(HMR(YCR095C(le)$of$HMR$ right$of$HMR$ telomere$right$0"0.5"1"1.5"2"2.5"3"3.5"4"4.5"HMR"Le." HMR"right" ChrIII"telomere"right"H2A.Z"enrichment"normalized"over"PRP8$noAtag"HTZ1"asf1"HTZ1"sas2"HTZ1"H2A.Z"ChIP"0"0.2"0.4"0.6"0.8"1"1.2"1.4"le+"of"HMR" right"of"HMR" ChrIII"telomere"right"H4K16ace"level"normalized"to"H3" Wildtype"asf1Δ"sas2Δ"H4K16ace"ChIP"    68  In Chapter 2, we found that Asf1 was also required for maintaining H2A.Z levels at the promoters of subtelomeric genes, therefore, we also examined whether the SAS complex was involved in this aspect of H2A.Z occupancy (Figure 3.2A). Unlike Asf1, Sas2 was also not required for maintaining H2A.Z levels at subtelomeric gene promoters, with the exception of the RDS1 gene that lies closest to the ChrIII R telomere (Figure 3.2B). Our observations at the right telomere of ChrIII suggested that the SAS complex might have distinct H2A.Z regulatory functions at different telomeres. We next examined the role of Asf1 and its relationship with the SAS complex at ChrVI since the SAS complex is required for H2A.Z deposition at this heterochromatin domain (Figure 3.2C) (Shia et al. 2006). We verified that H2A.Z was lost in the subtelomeric region of ChrVI upon SAS2 deletion in a distance-dependent manner (Figure 3.2D). However, the profile of H2A.Z occupancy differed between asf1Δ and sas2Δ cells. When the histone chaperone was deleted, there was a consistent reduction in overall H2A.Z levels across all sites assayed, irrespective of their distance away from the heterochromatin boundary (Figure 3.2D). In accordance with our findings at the subtelomeric genes of ChrIII R, the levels of H2A.Z were reduced by half (Lu and Kobor 2014). Furthermore, H2A.Z level in the asf1sas2 double mutant was reduced to the same level as the asf1Δ mutant at both ChrIII R and ChrVI R telomeric ends. Taken together, this set of experiments suggested that Asf1 does not promote H2A.Z occupancy through a SAS complex-mediated mechanism. Furthermore, Asf1 seems to be important for maintaining H2A.Z levels across telomeres.       69  Figure 3.2 Asf1-dependent H2A.Z deposition at ChrIII R was not mediated by SAS (A) Schematic representation of ORFs at the right telomere of ChrIII. (B) H2A.Z promoter enrichment at ChrIII R subtelomeric region did not depend on SAS2. (C) Schematic representation of ORFs at the right telomere of ChrVI. (D) H2A.Z promoter enrichment at ChrVI R subtelomeric regions depended on ASF1 and SAS2. (B) (D) ChIP-qPCR of H2A.Z was performed in the indicated strains and normalized to H2A.Z enrichment over PRP8. Error bars represent standard errors of the means for three independent experiments.   Figure"2"D"B"A"C"Chr$III$R$YCR094W(YCR095C(RDS1(GIT1((YCR099C((YCR100C(HMR(35$kb$Chr$VI$R$RPN12(YFR054C(20$kb$YFR055W(HXK1(RET2(T6R9E$ T6R9C$T6R9H$    70 3.3.2 Loss of Asf1 did not Alter Global H2A.Z Levels in Chromatin The consistent reduction in H2A.Z levels in asf1Δ cells at subtelomeric promoters prompted us to ask if bulk H2A.Z deposition and SWR1-C recruitment was dependent on Asf1. In wildtype cells, the majority of cellular H2A.Z is chromatin bound (Wang et al. 2009). Using a centrifugation-based assay to separate chromatin-associated proteins from the rest of the whole cell lysate (referred to as the supernatant fraction), we found that cells lacking the Asf1 histone chaperone had similar levels of H2A.Z in the supernatant and chromatin fractions as compared to wildtype cells (Figure 3.3A). Similarly, Asf1 did not seem to play a role in the recruitment of the SWR1-C complex to chromatin, as chromatin association of key subunits of the SWR1-C complex (Swc2 and Swc3) was not significantly altered upon deletion of ASF1 (Figure 3.3B). Unlike H2A.Z, the majority of Swc2 and Swc3 proteins were found in the chromatin fraction and there were no detectable amounts of these SWR1-C subunits in the supernatant (Figure 3.3B).   As H3K56 acetylation is a hallmark of newly deposited histones during DNA replication, we were prompted to investigate the relationship between Asf1 and H2A.Z during S-phase (Kaplan et al. 2008; Masumoto et al. 2005; Recht et al. 2006; Han and Zhang 2007). Cells lacking ASF1 are particularly sensitive to hydroxyurea (HU), a genotoxic agent commonly used to arrest cells during S-phase (Tyler et al. 1999; Franco and Kaufman 2005; Ramey et al. 2004). Therefore, cells were synchronized in G1-phase by alpha-factor arrest and collected at S-phase as determined by fluorescence assisted cell sorting (FACS). In wildtype cells, the bulk of cellular H2A.Z was in chromatin in both asynchronous and S-phase cells (Figure 3.3C). Despite the prominence of H3K56 acetylation in S-phase, in asf1Δ cells, the     71 majority of H2A.Z was associated with chromatin in both asynchronous and S-phase (Figure 3.3C). Furthermore, the levels of H2A.Z in the supernatant were comparable between the cell cycle phases in asf1Δ cells (Figure 3.3C). Taken together, our data suggested that Asf1 was not required for global SWR1-C recruitment or bulk H2A.Z deposition and that the relationship between the histone chaperone and H2A.Z was not cell cycle-dependent.                Figure 3.3 Loss of Asf1 does not alter global H2A.Z levels in chromatin  Chromatin association assay was performed on the indicated strains. WCE, whole cell extract; SUP, supernatant; CHR, chromatin pellet. (A) Loss of ASF1 did not alter H2A.Z association to chromatin. (B) Loss of ASF1 did not alter Swc2 and Swc3 recruitment to chromatin. (C) Loss of ASF1 did not alter H2A.Z association to chromatin during S-phase. The relative amounts of H2A.Z-FLAG, Swc2-VSV, and Swc3-VSV were determined by immunoblotting with anti-FLAG and anti-VSV antibodies. Antibodies against histone H4 and Pgk1 were used as loading controls for the chromatin pellet and supernatant, respectively.  Figure"3"A"B"C"    72 3.3.3 Genome-Wide H2A.Z Occupancy was Altered by Loss of H3K56 Acetylation To reconcile the differences between our observations of H2A.Z deposition at bulk levels versus promoter-specific differences, we explored the relationship between Asf1 and H2A.Z more thoroughly by analyzing H2A.Z occupancy genome-wide. ChIP-on-chip (Chromatin Immunoprecipitation on microarray) was used to precisely map changes in H2A.Z occupancy in cells lacking ASF1 and in cells containing an unacetylable substitute for lysine 56, H3K56R. Consistent with previous studies, H2A.Z was predominantly found at gene promoters using our enrichment criteria, with 3508 promoters enriched for H2A.Z in wildtype cells (Figure 3.4A). In both asf1Δ and H3K56R cells, the majority of H2A.Z peaks were found to be associated with gene promoters (Figure 3.4B). However, the H2A.Z profiles of both mutants had notable distinctions from the wildtype H2A.Z profile. At a subset of gene promoters, H2A.Z was completely lost, whereas at some promoters there was a reduction in the level of promoter-associated H2A.Z (Figure 3.4B). Overall, H2A.Z was completely lost in roughly 15% of promoters that were enriched for H2A.Z in the wildtype, with 3032 and 2940 enriched promoters enriched in asf1Δ and H3K56R cells, respectively (Figure 3.4A).        73  Figure 3.4 Genome-wide H2A.Z occupancy was altered by loss of H3K56 acetylation Averaged H2A.Z profiles over the entire yeast genome were obtained from two independent experiments. (A) Venn diagram of gene promoters enriched for H2A.Z in wildtype, asf1Δ, and H3K56R. (B) ChIP-chip profile of H2A.Z-FLAG-enriched regions in wildtype, asf1Δ, and H3K56R strains. Sample genomic regions were plotted for ChrIII and ChrIV along the x-axis against the relative occupancy of H2A.Z. ORFs are indicated as light gray rectangles above the x-axis for Watson genes and below the axis for Crick genes.      Figure"4"A"B"    74 Correspondingly, there was a slight overall decrease in H2A.Z occupancy in cells lacking ASF1 or H3K56 acetylation when all genes were visualized by CHROMATRA (Figure 3.5A, Figure 3.5B, and Figure 3.5C) (Hentrich et al. 2012). As expected, the majority of H2A.Z in wildtype cells was found in close proximity to the TSS of genes and was in low abundance along the ORF (Figure 3.5A). Very little H2A.Z was observed at the promoters of highly transcribed genes in all strains assayed (Figure 3.5A). Unlike the robust H2A.Z 500bp signal on either side of the TSS in wildtype cells, there appeared to be an overall decrease in H2A.Z enrichment at the 5’ end of all genes in cells lacking ASF1 or H3K56 acetylation (Figure 3.5B and Figure 3.5C).  Furthermore, changes in H2A.Z levels were independent of transcription rate and transcript length (Figure 3.5B and Figure 3.5C). The striking similarity of the H2A.Z profiles of asf1∆ and H3K56R cells suggested a common mechanism in H2A.Z regulation. The H2A.Z profiles of the two mutants had a Spearman correlation of 0.946, suggesting that the acetylation of H3K56 was the key regulatory mechanism mediating their effects on H2A.Z occupancy (Figure 3.5D).              75  Figure 3.5 Loss of H3K56 acetylation led to an overall decrease in H2A.Z levels (A) (B) (C) Comparison of H2A.Z enrichment of all genes in wildtype, asf1Δ, and H3K56R. CHROMATRA plots of relative H2A.Z occupancy across all transcripts sorted by length and transcriptional frequency. All genes are aligned by their TSS and include a 500bp region upstream of the TSS. Enrichment scores are binned into 150bp segments. Transcripts were grouped into five classes according to their transcriptional frequency as per Holstege et al. (1998). (D) Spearman correlation of H2A.Z enrichment profile of WT, asf1Δ, and H3K56R.    Figure"5"A" B"C"Transcrip)onal,Frequency,(mRNA/h), >50,16?50,4?16,1?4,<1,TSS,0,,,,,450,,,,900,bp,H2A.Z,WT,Transcrip)onal,Frequency,(mRNA/h), >50,16?50,4?16,1?4,<1,TSS,0,,,,,450,,,,900,bp,H2A.Z,asf1Δ,Transcrip)onal,Frequency,(mRNA/h), >50,16?50,4?16,1?4,<1,TSS,0,,,,,450,,,,900,bp,H2A.Z,H3K56R,D"1.00$1.00$1.00$0.85$0.85$ 0.86$0.86$ 0.95$0.95$H2A.Z&WT&&H2A.Z&asf1Δ&&H2A.Z&H3K56R&&H2A.Z&WT&& H2A.Z&asf1Δ&& H2A.Z&H3K56R&&    76 3.3.4 Defects in H3K56 Acetylation Resulted in Reduced H2A.Z Promoter Occupancy Next, differences in H2A.Z relative occupancy were considered, with a specific focus on promoters that were significantly enriched for the H2A.Z variant in wildtype cells. As expected, an average gene profile of all genes whose promoters were enriched for H2A.Z revealed that H2A.Z was highly enriched around the TSS, depleted over the ORF, and enriched to a small degree at the 3’ end of genes (Figure 3.6A). The average gene profile of H2A.Z enrichment in cells lacking ASF1 clearly demonstrated an overall reduction in H2A.Z levels at both the 5’ IGR (intergenic region) and the 3’IGR (Figure 3.6A). In addition, cells containing an unacetylable arginine in place of lysine 56 exhibited a similar H2A.Z average gene profile, with a strong decrease in H2A.Z levels at the 5’ end of genes and a similar decrease at the 3’ end of genes (Figure 3.6A). Comparing the relative H2A.Z occupancy, in a gene-by-gene manner, of all 3509 H2A.Z-containing promoters revealed an overall decrease in H2A.Z occupancy in asf1Δ and H3K56R cells (t-test p-value = 4.54e-117 in asf1Δ mutant and p-value = 8.41e-172 in H3K56R mutant) (Figure 3.6B and Figure 3.6C). Demonstrating that while the majority of genes were still considered enriched under our selection criteria, there was a significant decrease in the relative occupancy of H2A.Z at these promoters.       77  Figure 3.6 Defects in H3K56 acetylation resulted in reduced H2A.Z promoter occupancy (A) Average gene profile of H2A.Z enrichment of all genes enriched at its promoters in wildtype cells.  Each ORF was divided into 40 bins (independent of gene length), and average enrichment values were calculated for each bin. 1500bp upstream of the TSS and 1500bp downstream of the 3’ UTR were assigned to 20 bins. The average enrichment value for each bin was plotted. (B)(C) H2A.Z promoter occupancy scores were significantly decreased in asf1Δ (p-value = 4.54e-117) and H3K56R (p-value = 8.41e-172) cells. Promoter occupancy scores for H2A.Z were plotted for all promoters enriched for H2A.Z in wildtype cells. Promoters are ordered from highest to lowest based on the H2A.Z enrichment score of wildtype.    −3−2−1012345678-505101520H2A.Z promoter enrichmentRelative H2A.Z occupancy Wild type  asf1 -505101520H2A.Z promoter enrichmentRelative H2A.Z occupancy Wild type  H3K56R Figure"6"A"B" C"Rela=ve$occupancy$Enriched$ORF$5’$IGR$ 3’$IGR$9$Wildtype$9(asf1Δ(9(H3K56R(    78 3.3.5 H3K56 Acetylation Positioned H2A.Z at Gene Promoters  While a subset of H2A.Z peaks were lost or reduced when H3K56 acetylation was abolished, the majority of gene promoters remained enriched for H2A.Z in asf1Δ and H3K56R mutants. Closer examination of the H2A.Z genome-wide profile revealed that there was a subtle yet clear shift in the genomic location of H2A.Z peaks in asf1Δ and H3K56R mutants compared to wildtype.  Representative regions in ChrIII and ChrIX highlighted a shift in the H2A.Z enrichment profile towards the left arm of both chromosomes (Figure 3.7A). Analogous to all earlier analyses of the H2A.Z profiles of asf1Δ and H3K56R in this chapter, the shift patterns observed in the two mutants were also strikingly similar. Additionally, the magnitude of the shifted peaks remained consistent with wildtype cells despite the change in their location (Figure 3.7A).   To determine the significance of the observed shift between wildtype and the H3K56 acetylation mutant strains, we developed a new method to define the change in location of the H2A.Z peak relative to the TSS of its corresponding gene (for the analysis, this is defined as the closest gene). Using the TSS of genes as a reference point, we calculated the distance from TSS to the midpoint of H2A.Z peaks in wildtype and mutants, and compared the differences in H2A.Z position. Only genes whose promoters were enriched for H2A.Z in wildtype, asf1Δ, and H3K56R cells were used. Furthermore, a criterion was set so that the midpoint of the associated H2A.Z peak must be within 500bp upstream or downstream of the TSS. Following this set of selection criteria, 2399 genes were left whose promoter had an associated H2A.Z peak in all three strains.       79             Figure 3.7 Loss of H3K56 acetylation resulted in a shift of H2A.Z position at gene promoters  (A) Representative genomic regions were plotted for ChrIII and ChrIX along the x-axis against the relative occupancy of H2A.Z normalized to input in wildtype, asf1Δ, and H3K56R strains. ORFs are indicated as light gray rectangles above the x-axis for Watson genes and below the axis for Crick genes. (B) (C) Distribution of H2A.Z positions did not change at H2A.Z-enriched promoters (D) (E) Position of H2A.Z peaks were significantly shifted at H3K56 acetylated promoters in H3K56R cells (p-value = 0.021). (B) (C) (D) (E) Violin plot representing the distribution of H2A.Z peak position relative to TSS for each strain.  -500-2500250500WT asf1base pairH2A.Z promoters-500-2500250500WT asf1base pairH3K56ace promoters-500-2500250500WT H3K56Rbase pairH2A.Z promoters-500-2500250500WT H3K56Rbase pairH3K56ace promotersFigure"7"B"D"WT" asf1Δ& WT" H3K56R&WT" H3K56R&WT" asf1Δ&Distance$away$from$TSS$(bp)$Distance$away$from$TSS$(bp)$Distance$away$from$TSS$(bp)$Distance$away$from$TSS$(bp)$A"C"E"    80 First, we examined H2A.Z position in these 2399 genes enriched for the histone variant in all three mutant strains. As expected, for the majority of genes in wildtype cells, H2A.Z was located just upstream of the TSS, with a mean position of -103bp relative to the TSS (Figure 3.7B). Thus, it is likely that the single large peak from our microarray analysis represents an average profile of the -1 and +1 H2A.Z containing nucleosomes. Comparable to the wildtype distribution, the majority of H2A.Z was found approximately 100bp upstream of the TSS in asf1Δ and H3K56R (Figure 3.7B and Figure 3.7C). Taken together, the distribution of promoter H2A.Z position in asf1Δ and H3K56R resembled the wildtype distribution suggesting that H2A.Z position did not change.   Next, since a shift in H2A.Z upon loss of H3K56 acetylation was expected only at promoters that are normally acetylated at H3K56, we examined genes whose promoter was normally enriched for H3K56 acetylation (Rufiange et al. 2007). Using the genome-wide H3K56 acetylation profile of asynchronous wildtype cells, we identified 1369 promoters enriched for H3K56 acetylation, of which 452 also had an associated H2A.Z peak in WT, asf1Δ and H3K56R cells (Rufiange et al. 2007). Strikingly, there was a notable shift in the positions of H2A.Z peaks in both asf1Δ and H3K56R mutants at H3K56 acetylated promoters (Figure 3.7 D, 7E). The distribution of H2A.Z peak position was significantly shifted towards the TSS in the H3K56R mutant with a p-value of 0.021, whereas the change in position of the asf1Δ mutant had a p-value of 0.086  (Wilcox test for paired, non-normal distribution) (Figure 3.7E). On average, H2A.Z position in H3K56R shifted closer to the TSS than in the asf1Δ mutant.        81 Similar to before, a gene-by-gene approach was taken to examine the 452 promoters where H2A.Z and H3K56ace colocalized. To get a sense of the magnitude and direction of the change in H2A.Z position upon ASF1 deletion and H3K56 mutation, we plotted the distance to TSS for this subset of H2A.Z peaks and compared the wildtype profile against the mutant (Figure 3.8). As a whole, there was a large range in the change of H2A.Z peak positions in the mutants when compared to the wildtype. In both asf1Δ and H3K56R, there was a proportion of H2A.Z peaks that remained relatively unchanged compared to wildtype H2A.Z position as well as a subset of H2A.Z peaks that shifted drastically away from the wildtype position (Figures 3.8A and Figure 3.8B). Importantly, the H2A.Z peaks shifted both upstream and downstream of its wildtype position with loss of ASF1 or H3K56 acetylation (Figure 3.8A and Figure 3.8B). Taken together, our data demonstrated that H3K56 acetylation was required for H2A.Z positioning at a subset of gene promoters.       Figure 3.8 H3K56 acetylation defects led to a shift in H2A.Z position Scatterplot of H2A.Z position relative to the TSS of the corresponding gene in wildtype, asf1Δ, and H3K56R cells. (A) Loss of ASF1 resulted in a shift of H2A.Z peaks both upstream and downstream of the wildtype position. (B) A change in H2A.Z position was observed in H3K56R compared to wildtype.  -400-2000200400Distance away from TSS in WT versus H3K56R strainsDistance away from TSS Wild type  H3K56R -400-2000200400Distance away from TSSDistance from TSS Wild type  asf1 Figure"8"A"B"-400-2000200400Distance away from TSSDistance from TSS Wild type  asf1 -400-2000200400Distance away from TSS in WT versus H3K56R strainsDistance away from TSS Wild type  H3K56R -400-2000200400Distance away from TSSDistance from TSS Wild type  asf1 Figure"8"A"B"-400-2000200400Distance away from TSSDistance from TSS Wild type  asf1     82 3.4 Discussion  In this chapter, we provide evidence that Asf1 regulated H2A.Z occupancy and positioning at gene promoters by promoting H3K56 acetylation. Despite indications from previous studies, Asf1-dependent H2A.Z occupancy at ChrIII was not mediated through H4K16 acetylation by the SAS HAT complex. Instead, H2A.Z deposition at telomeres was dependent on H3K56 acetylation and the Asf1 histone chaperone. We mapped the genome-wide occupancy profile of H2A.Z in asf1Δ and H3K56R cells by ChIP-chip to gain a comprehensive and detailed understanding of how H3K56 acetylation could be regulating H2A.Z occupancy. Loss of H3K56 acetylation resulted in a slight, albeit significant, reduction in H2A.Z levels and alterations in H2A.Z position at gene promoters. At a subset of genes, H2A.Z was depleted at promoters that were occupied by the histone variant in the wildtype. At the majority of affected promoters, there was significant decrease in the level of H2A.Z in strains that lack H3K56 acetylation. Interestingly, we also observed a significant shift in the position of promoter H2A.Z upon the loss of H3K56 acetylation, suggesting a key role for the histone modification in positioning H2A.Z. Taken together, the results in this chapter present a model in which H3K56 acetylation at gene promoters played a key role in governing H2A.Z biology, both in regulating the levels and position of this histone variant.   In Chapter 2 of this dissertation, we uncovered that Asf1 was required to maintain normal H2A.Z levels at the promoters of both subtelomeric and euchromatic genes. In this chapter, we explored this relationship further to tease apart the mechanistic and functional relationship between the Asf1 H3/H4 histone chaperone and the H2A.Z histone variant.     83 Previous reports proposed a recruitment-based model in which Asf1 recruited the SAS complex, a H4K16 histone acetyltransferase, to the heterochromatin boundary (Osada et al. 2005; 2001). H4K16ace by the SAS complex has been shown to be required for SWR1-C-dependent H2A.Z deposition at one heterochromatin boundary (Shia et al. 2006). We found that while H2A.Z occupancy at the ChrVI heterochromatin boundary was indeed dependent on the SAS complex, this did not hold true at the heterochromatin boundaries of ChrIII R. At ChrIII R, even though loss of ASF1 resulted in reduced H4K16 acetylation levels, loss of H4K16ace through sas2Δ did not alter H2A.Z levels. While the reduction in H4K16 acetylation was likely due to a role for Asf1 in recruiting the SAS complex to heterochromatic boundaries, our findings suggested that the relationship between the SAS complex and H2A.Z was telomere-specific. Furthermore, H2A.Z occupancy level at subtelomeric gene promoters in asf1Δsas2Δ was similar to asf1Δ at ChrVI R and ChrIII, supporting the model that Asf1 acts upstream of the SAS complex. While the focus of this chapter was in H2A.Z at heterochromatin, the functional relationship between Asf1 and the SAS complex likely extends beyond their roles in H2A.Z regulation. A number of earlier publications highlight this relationship as cells lacking Asf1 or SAS function exhibit similar growth deficiencies, defects in establishing silent chromatin structure, and genetic interaction partners (Miller et al. 2010; Raisner and Madhani 2008; Osada et al. 2005; Meijsing and Ehrenhofer-Murray 2001).   Analysis of the relationship between Asf1 and the bulk levels of chromatin-bound H2A.Z suggested that bulk H2A.Z occupancy and SWR1-C recruitment did not depend on Asf1. This differed from our region-specific examination of the subtelomeric gene promoters, as     84 we saw a consistent decrease in H2A.Z levels at multiple telomeres as well as euchromatic genes (Lu and Kobor 2014). One possible explanation for this may be due to the differences between the two assays. A centrifugation-based separation of chromatin from whole cell lysate may not be able to distinguish SWR1-C bound H2A.Z that are associated to chromatin, but not deposited, from H2A.Z incorporated into the nucleosome. Similar observations in cells lacking YAF9 support this idea and indicate that a chromatin association assay cannot pick up the differences between H2A.Z that are associated with chromatin versus H2A.Z that are incorporated in the nucleosome (Wang et al. 2009).   Using ChIP-on-chip, we generated a detailed global H2A.Z occupancy profile in the absence of H3K56 acetylation. The genome-wide H2A.Z profile highlighted three distinct effects that the loss of H3K56 acetylation had on H2A.Z biology in chromatin. The loss of H3K56 acetylation resulted in either the absolute loss of H2A.Z at gene promoters, a reduction in the levels of H2A.Z at gene promoters, or a shift in H2A.Z position at gene promoters; highlighting a key role for this histone modification in the regulation of chromatin-bound H2A.Z. The most exciting observation that we uncovered as a result of our genome-wide H2A.Z profiling was the observed shift in H2A.Z occupancy peaks in H3K56 acetylation mutants. In wildtype cells, almost 50% of H3K56 acetylated promoters are also enriched with H2A.Z, supporting a functional overlap of these two chromatin marks. While nucleosomes with or without acetylated H3K56 have the same stability in vitro, (H3/H4)2 tetrasomes are more stable than (H3K56ace/H4)2 tetrasomes, suggesting that the modified tetrasome has the capacity to promote nucleosome movement (Andrews et al. 2010; Muthurajan et al. 2004). It is tempting to speculate the heightened chromatin fluidity of     85 H3K56-acetylated tetrasomes allow for fine-tuning of nucleosome position during H2A.Z/H2B exchange by SWR1-C, whereas this capability is lost in cells lacking H3K56 acetylation. Indeed, in vitro FRET experiments have shown that the constitutive acetyl mimic, K56Q, have heightened nucleosome thermal mobility compared to unmodified nucleosomes (Ferreira et al. 2007). This proposed mechanism is distinct from a recent report demonstrating that H3K56 acetylation promotes the removal of H2A.Z by SWR1-C by disrupting the locking mechanism of Swc2 (Watanabe et al. 2013). It is interesting that H3K56 acetylation has such a dynamic role in the regulation of H2A.Z. This could be the underlying reason for why H3K56 acetylation is such a transient and highly regulated histone modification (Maas et al. 2006; Kaplan et al. 2008; Ozdemir et al. 2006).   Interestingly a recent paper demonstrated that the H3K56Q, designed to mimic H3K56 acetylation, also lead to reduced H2A.Z occupancy (Watanabe et al. 2013). Furthermore, our own ChIP-on-chip data also revealed a similar reduction in H2A.Z level in a H3K56Q mutant (Lu, data not shown). Taken together with the data presented in this chapter, it seems as if both the constitutive presence and absence of H3K56 acetylation regulate H2A.Z occupancy. It is tempting to speculate that perhaps, it is the tight regulation of this transient histone modification that is the key in its role in H2A.Z position and occupancy. Indeed, while H3K56 acetylation is required for chromatin reassembly after DNA damage repair, removal of the histone modification is also required for release from cell cycle checkpoint after repair (Hyland et al. 2005; Tjeertes et al. 2009; Chen et al. 2008; Masumoto et al. 2005; Wurtele et al. 2012; Celic et al. 2006; Maas et al. 2006). Further research is needed to determine if a     86 similar requirement for regulating the status of H3K56 acetylation exist in regulating H2A.Z occupancy.   In this chapter, we utilized two distinct genetic perturbations that lead to loss of H3K56 acetylation and their effect on H2A.Z biology. Surprisingly, deletion of ASF1 and the unacetylable H3 allele, H3K56R, both exhibited very similar defects in H2A.Z occupancy. That being said, loss of H3K56 acetylation through H3K56R consistently resulted in a stronger decrease in H2A.Z levels and a bigger shift in H2A.Z nucleosomes compared to the asf1Δ mutant. Considering that H3K56R mutant very specifically compromises Asf1’s role as a chaperone in promoting H3K56 acetylation, it is possible that a complete deletion of the histone chaperone would lead to compensation by other chromatin remodeling factors. Nevertheless, the overall effect on H2A.Z position and occupancy between the two mutations were quite similar. With a Spearman correlation of 0.946 and over 90% overlap of promoters enriched for H2A.Z between asf1Δ and H3K56R, our data emphasizes the importance H3K56 acetylation plays in regulating the position and occupancy level of H2A.Z at a subset of gene promoters.    It is exciting to think that perhaps a more precise method of measuring nucleosome position would yield a better understanding of the precise mechanism of H2A.Z positioning by acetylated H3K56. With our current data set from the ChIP-chip analysis, we developed a method to analyze the change in location of each H2A.Z peak found at gene promoters. As it currently stands, the peak shift analysis also includes all H2A.Z peaks even if they were not shifted. This was due to the inability to set objective cutoffs deeming significantly shifted     87 peaks. By mapping H2A.Z containing nucleosomes through MNase digestion followed by ChIP-seq, we would better understand and appreciate the significance of H3K56 acetylation in H2A.Z nucleosome positioning.      88 Chapter 4: Eaf1 HSA Domain and Epl1 C-terminus Coordinate NuA4 Stability and Function   4.1  Introduction Chromatin structure facilitates DNA compaction inside the cell nucleus and contributes to the regulation of nuclear processes such as transcription, DNA repair and DNA replication (Ehrenhofer-Murray 2004; Rando and Chang 2009). In eukaryotes, the structure of chromatin can be modified by several fundamental mechanisms, often involving specialized and dedicated protein complexes. For instance, ATP-dependent remodeling enzymes use energy derived from ATP hydrolysis to slide or exchange nucleosomes on the DNA template (Eberharter and Becker 2004; Clapier and Cairns 2009). Additionally, the composition of nucleosomes can be altered by incorporation of the less abundant histone variants, who differ from their canonical counterparts in their primary amino acid sequence and function (Sarma and Reinberg 2005; Zlatanova and Thakar 2008). Finally, both canonical histones and their variants are subjected to numerous post-translational chemical modifications, including acetylation, methylation, phosphorylation, ubiquitination and ADP-ribosylation (Bannister and Kouzarides 2011).   Budding yeast, Saccharomyces cerevisiae, contain several histone acetyltransferase (HAT) complexes, including NuA4 (Nucleosome acetyltransferase of H4). NuA4, the only essential HAT in yeast, is composed of 13 subunits and not only acetylates the N-terminal tails of     89 histones H4, H2A, and H2A.Z but also additional non-histone substrates (Babiarz et al. 2006; Doyon et al. 2004; Jeong et al. 2011; Keogh et al. 2006; Lin et al. 2008; 2009; Mitchell et al. 2011). NuA4 is required for a diverse set of cellular functions, including cell cycle progression, DNA double-stranded break repair, establishment of heterochromatin-euchromatin boundaries at subtelomeric regions, and transcription of ribosomal genes (Bird et al. 2002; Boudreault et al. 2003; Clarke et al. 1999; Reid et al. 2000; Smith et al. 1998; Zhang et al. 2004). Its catalytic activity resides within the Esa1 subunit, while other subunits coordinate specific functions of NuA4 and contribute to its structural stability. NuA4 is a modular complex that assembles on the Eaf1 subunit and as such, different regions of the Eaf1 protein mediate the association of distinct components of NuA4. For example, the N-terminal region of Eaf1 is responsible for the association of the Eaf5/7/3 sub-module, while its SANT (SWI3-ADA2-N-CoR-TFIIIB) domain binds Tra1, a subunit involved in NuA4 recruitment to specific promoters via Tra1’s interaction with acidic transcription activators such as Gcn4, Hap4, and Gal4 (Auger et al. 2008; Brown et al. 2001). In addition, the HSA (helicase-SANT-associated) domain of Eaf1 interacts with a module containing Swc4, Arp4, Yaf9, and Act1 (Auger et al. 2008; Szerlong et al. 2008). The Swc4/Arp4/Act1/Yaf9 module provides an intriguing link to the H2A.Z histone variant as these four proteins are also components of SWR1-C, an ATP-dependent chromatin-remodelling complex that deposits H2A.Z into chromatin (Kobor et al. 2004; Krogan et al. 2003; Mizuguchi et al. 2004; Zhang et al. 2004). Consistent with a broad requirement of Eaf1 for NuA4 assembly and function, cells lacking EAF1 have strongly reduced global levels of H4 tetra-acetylation and sensitivity for a wide range of genotoxic agents (Kobor et al. 2004; Krogan et al. 2004).       90 Previous work has shown that the catalytic module of NuA4 exists as the functional Piccolo NuA4 (picNuA4) subcomplex composed of Esa1, Yng2, Epl1, and Eaf6 subunits (Auger et al. 2008; Boudreault et al. 2003; Chittuluru et al. 2011). Binding of Esa1 and Yng2 to the EPcA domain of Epl1 is important for the association and function of picNuA4 (Selleck et al. 2005). A number of studies have suggested that picNuA4 is anchored to NuA4 through a physical interaction between the Epl1 C-terminus and the Eaf1 scaffolding subunit, as removal of the C-terminal region of Epl1 results in the dissociation of the picNuA4 submodule from NuA4. In contrast to the locus-specific recruitment, acetylation, and activation of NuA4, picNuA4 is thought to catalyze non-targeted global histone acetylation (Boudreault et al. 2003). In vitro, picNuA4 has a preference in acetylating nucleosomal histones over free histones unlike its larger cousin (Nourani et al. 2004; Reid et al. 2000; Boudreault et al. 2003; Friis et al. 2009; Selleck et al. 2005). Recent structural analysis identified the Yng2 subunit and the Epl1 N-terminus as the key binding surfaces for picNuA4 association to the nucleosome core particle (Chittuluru et al. 2011). Although rooted in the discovery of picNuA4, untargeted histone acetylation is emerging as an important, yet poorly understood, property of HAT complexes.   Here we study the two scenarios represented by loss of the entire Eaf1 protein and the loss of the Epl1 C-terminus to better understand the precise nature of the relationship between NuA4 and picNuA4. Remarkably, removal of the Epl1 C-terminus suppressed the bulk histone acetylation defects and growth defects associated with loss of EAF1 through the stabilization of the picNuA4 complex. In contrast, NuA4-specific defects, such as decreased promoter H4 acetylation and expression of ribosomal genes, were not rescued by removal of the Epl1 C-    91 terminus. Large-scale genetic and gene expression analyses revealed commonalities and distinct properties between eaf1Δ and epl1-CΔ mutants, providing further evidence that these two genetic alterations resulted in distinct perturbations in NuA4 HAT function. Structure-function studies identified the HSA domain as being the key domain for picNuA4 formation, likely by being the docking platform for the Epl1 C-terminus. Lastly, artificially releasing picNuA4 via the removal of the Epl1 C-terminus revealed that picNuA4 was poorly associated to chromatin despite being sufficient to acetylate H4 and H2A.Z. Taken together, the results presented in this chapter suggested that the interface between these two key subunits likely played an important role in maintaining the equilibrium of active NuA4 and picNuA4 in the cell.   4.2 Materials and Methods  4.2.1 Yeast Strains and Plasmids All strains used in this study are listed in Table 4.1. Yeast strains were generated using standard genetic techniques including homologous recombination and genetic crosses followed by tetrad dissection (Ausubel 1987). Complete deletion of genes, EPL1 truncations, and 3’ end integration of an in frame epitope tag (TAP or 3xFLAG) were achieved using one-step gene integration PCR-amplified modules (Gelbart et al. 2001). The EAF1 gene was PCR-amplified from genomic DNA and cloned into pRS316 (URA3) centromeric vector containing a 3x-HA tag. Internal deletions of EAF1 were performed by adapting the Quick     92 Change site-directed mutagenesis method (Stratagene) following manufacturer’s protocol. All mutations were confirmed by DNA sequencing. Table 4.1 Yeast strains used in this study Strain number Relevant Genotype MKY6 W303, MATA, ADE2, can1-100 his3-11 leu2-3,112 trp1-1 ura3-1 lys2Δ MKY1762  MKY1763 MKY1764 MKY6, EPL1-TAP::TRP MKY6, EPL1-TAP::TRP, eaf1::HIS MKY6, epl1-485-TAP::TRP MKY1765 MKY6, epl1-485-TAP::TRP, eaf1::HIS MKY1766 MKY6, epl1-380-TAP::TRP MKY1767 MKY6, epl1-380-TAP::TRP, eaf1::HIS MKY583 BY4741, MATα, his3Δ1 leu2Δ0 LYS2 met15Δ0 ura3Δ0 Δcan1::MATαPr-HIS3 Δlyp1::MATαPr-LEU2 MKY1419 MKY1428 MKY1429 MKY1430 MKY1431 MKY 1768 MKY1769 MKY1770 MKY1771 MKY1772 MKY583, eaf1 1-982-3XHA::NAT MKY583, eaf1Δ-3XHA::NAT MKY583, EPL1-3xFLAG::KAN MKY583, epl1-485-3xFLAG::KAN MKY583, epl1-380-3xFLAG::KAN MKY6, pRS316 MKY6, EPL1-TAP::TRP, pRS316 MKY6, EPL1-TAP::TRP, eaf1::HIS [pRS316] MKY6, EPL1-TAP::TRP, eaf1::HIS [pRS316, EAF1] MKY6, EPL1-TAP:::TRP, eaf1::HIS [pRS316, eaf1-HSAΔ] MKY1773 MKY6, EPL1-TAP::TRP, eaf1::HIS [pRS316, eaf1-SANTΔ] MKY1774 MKY1775 MKY1776 MKY1777 MKY1778 MKY1779 MKY6, EPL1-TAP::TRP, eaf1::HIS [pRS316, eaf1-HSAΔSANTΔ] MKY6, epl1-485-TAP::TRP, eaf1::HIS [pRS316] MKY6, epl1-485-TAP::TRP, eaf1::HIS [pRS316, EAF1] MKY6, epl1-485-TAP::TRP, eaf1::HIS [pRS316, eaf1-HSAΔ] MKY6, epl1-485-TAP::TRP, eaf1::HIS [pRS316, eaf1-SANTΔ] MKY6, epl1-485-TAP ::TRP, eaf1::HIS [pRS316, eaf1-HSAΔSANTΔ]     93  4.2.2 Growth and Genotoxic Sensitivity Assays Overnight cultures grown in YP-dextrose were diluted to OD600 0.5. Cells were 10-fold serially diluted and spotted onto solid YPD plates or plates with 0.005% MMS (Sigma), 50mM hydroxyurea (HU), or 1% formamide (Sigma). For strains containing URA3 plasmids, the cultures were grown in SC –URA media and serially diluted cells were spotted onto SC –URA plates or plates containing genotoxic agents. The plates were then incubated at the indicated temperature for 36 hours.   4.2.3 Large-Scale Affinity Purification Purification of native protein complexes were performed using extracts from strains encoding the TAP tag fused in-frame to the 3’ end of genes. Purifications were performed from 1L cultures that were harvested at an OD600 of 1.00. Large-scale purification was adapted from protocols previous described with minor modifications (Mitchell et al. 2008). Briefly, cells were lysed with a coffee grinder with dry ice pellets and resuspended in TAP buffer (20 mM Hepes [pH 8], 350 mM NaCl, 10% glycerol, 0.1% Tween-20, 1x phosphatase inhibitor mix, Complete Protease Inhibitor cocktail [Roche]). NP-40 was added to a final concentration of 1% prior to centrifugation at 3000g for 10 minutes at 4C. Crude extracts were incubated with 200µl of IgG (Millipore) crosslinked M-270 Epoxy beads (Invitrogen) as per manufacturer’s protocol for 3 hours at 4C rotating. The beads were then washed with 4ml of TAP buffer three times. The protein complex was eluted in 30µl of 0.1M citrate [pH 3.1] and loaded into 4-20% gradient gel (Bio-rad) for silver stain or western analysis.       94 4.2.4 RT-qPCR Overnight cultures were diluted to OD600  of 0.15 and grown in YP-dextrose to an OD600 of 0.5. Ten OD600 units were harvested for RNA extraction and purification using a Qiagen RNeasy minikit as per manufacturer protocol. cDNA was synthesized using QuantiTect Reverse Transcription Kit (Qiagen). cDNA was analyzed using a Rotor-Gene 6000 (Qiagen) and PerfeCTa SYBR green FastMix (Quanta Biosciences). mRNA levels were normalized to ACT1 mRNA levels. Samples were analyzed in triplicates for three independent RNA preparations. Statistical significance was assessed using Student’s t test.  Primer sequences are listed in Table 4.2.  Table 4.2 Primers used in this study Name Method Primer sequence (5’-3’) EPL1 F EPL1 R RPL19B F RPL19B R RPS11B F RPS11B R RPS3 F RPS3 R RPL19B F RPL19B R RPS11B F RPS11B R RPS3 F RPS3 R RT RT RT RT RT RT RT RT ChIP ChIP ChIP ChIP ChIP ChIP GGACGATCCCTTTAATCCTG CTCGAACGGCCATTTGAATC TGCAAGCGTTAATTCGATTG ACAACAGAAGCGGCAAGTCT GAAAAGCAGGACCTCGAATG TGGCATTAGTCGGGTGAAAT CACTCCAACCAAGACCGAAG TCTGACGACACCGTAAGCAG CTCATCGCTATGGGAATTGG TGGTGACTTTTTATTCGATTTGG CATGTTTCCCGCTTTGTTTT GATTTTCAACAGACGCAGCA TTTGTTTCCGTAACATCCATACC AAGAGCTGCATTGATTTGGAA      95 4.2.5 ChIP-qPCR and ChIP-on-chip Chromatin immunoprecipitation (ChIP) experiments were performed as described previously (Schulze et al. 2009). In brief, 250 ml of cells were grown in YP-dextrose to an OD600 of 0.5-0.6 from OD600 of 0.15 and were crosslinked with 1% formaldehyde for 20 minutes before chromatin was extracted. The chromatin was sonicated (Bioruptor, Diagenode, Spart, NJ: 10 cycles, 30s on/off, high setting) to yield an average DNA fragment of 500bp. Anti-FLAG antibody (4.2µl, Sigma) was coupled to 60µl of protein A magnetic beads (Invitrogen). After reversal of the crosslinking and DNA purification, the immunoprecipitated and input DNA were analyzed by quantitative real-time PCR (qPCR). Samples were analyzed in triplicate for three independent ChIP experiments. Statistical significance was assessed using Student’s t test.  Primer sequences are listed in Table 4.2. For microarray analysis, after reversal of crosslinking and DNA purification, the DNA was amplified with two rounds of T7 RNA polymerase amplification and hybridized to Affymetrix 1.0R S. cerevisiae tiling microarray. A modified version of the model-based analysis of tiling arrays (MAT) algorithm was used to reliably detect Epl1 occupancy across the genome. The data was normalized using both input DNA and a mock IP control.   4.2.6 mRNA Expression Profile Expression profiling was performed as described previously (van de Peppel et al. 2003). Briefly, EAF1 and EPL1 alleles were processed four times from two independently inoculated cultures. Dual-channel 70-mer oligonucleotide arrays were used with a common reference wild-type RNA. After RNA isolation, all steps were operated using robotic liquid handlers and scores were calculated as previously described (van Bakel and Holstege 2004;     96 van de Peppel et al. 2003). Differentially expressed genes were determined by p-value of 0.01 and minimum fold change of 1.7 compared to wildtype. Database for Annotation, Visualization, and Integrated Discovery (DAVID) was used for Gene Ontology enrichment analysis (Dennis et al. 2003). Multiple testing correction was done using the Benjamini method.    4.2.7 E-MAP E-MAP assay was performed as described previously (Schuldiner et al. 2006). Briefly, using a Singer robot, EAF1 and EPL1 alleles were crossed to a library of 1,536 mutants representing a number of processes including transcription, RNA processing and chromatin biology. All strains were screened three to four times and scores were calculated as previously described (Collins et al. 2006; Schuldiner et al. 2006).   4.2.8 Chromatin Association Assay Chromatin association assay were performed as previously described (Wang et al. 2009). In brief, cells were diluted to OD600 0.15 and collected at logarithmic phase. Following collection, cells were incubated in pre-spheroblast buffer (100 mM PIPES/KOH [pH 9.4], 10 mM DTT, 0.1% sodium azide) for 10 minutes at room temperature rotating, then spheroblasted for 20 minutes at 37C with 20mg/ml Zymoylase-100T (Seikagaku Corporation). Spheroblasts were washed with wash buffer (50 mM HEPES/KOH [pH 7.5], 100mM KCl, 2.5 mM MgCl2, 0.4 M sorbitol) and resuspended in equal volume of EB (50 mM HEPES/KOH [pH 7.5], 100mM KCl, 2.5 mM MgCl2, 1 mM DTT, 1 mM PMSF, and Complete Protease Inhibitor cocktail [Roche]). Cells were lysed with 1% Triton X-100.     97 Whole cell extracts were saved and the remaining lysate were separated into the chromatin pellet and the supernatant fraction by centrifugation through EBXS (EB + 0.25% Triton X-100 and 30% sucrose). The three fractions were subsequently analyzed by SDS-PAGE and immunoblotted. Immunoblots were scanned with the Odyssey Infrared Imaging System (Licor).     4.3 Results  4.3.1 Epl1 C-terminus Truncation Suppressed the Effects of EAF1 Deletion by Stabilizing picNuA4 Complex Formation Many lines of evidence suggested that the physical interaction between the Epl1 C-terminus and Eaf1 anchors the catalytic module to the rest of the NuA4 complex (Auger et al. 2008; Boudreault et al. 2003). To dissect the functional and structural relationship between Epl1 and Eaf1 more closely, we first examined the genetic interaction between EAF1 and EPL1 by combining a complete deletion of the non-essential EAF1 gene with one of two EPL1 C-terminal truncation mutants (Boudreault et al. 2003). Two variants of epl1-CΔ exist, allowing us to comprehensively interrogate its C-terminus. In addition to the N-terminal EPcA domain, the epl1-485 allele encoded protein contains central stretches of alanine and glutamine repeats whereas the epl1-380 allele does not (Figure 4.1A).  Loss of Eaf1 and the Epl1 C-terminus result in a variety of growth and functional defects typically associated with loss of NuA4 function, such as sensitivity to genotoxic agents such as methyl-methanesulfonate (MMS), hydroxyurea (HU), and formamide, and reduced acetylation of H4     98 (Figure 4.1B and Figure 4.1C) (Auger et al. 2008; Babiarz et al. 2006; Boudreault et al. 2003; Kobor et al. 2004). Surprisingly, upon combining eaf1Δ in parallel with epl1 1-380 or epl1 1-485 alleles, sensitivity to genotoxic agents of these eaf1 epl1-CΔ double mutants was not as severe as the single eaf1 mutant (Figure 4.1B). The partial suppression was particularly noticeable in the healthier epl1-485 strain, whereby truncation of its C- terminus in an eaf1Δ background was sufficient to partially rescue the growth phenotype to that of an epl1-485 single mutant (Figure 4.1B). Given that loss of EAF1 has been shown to reduce the levels of bulk H4 acetylation, we next evaluated the effect of further removing the C-terminus of Epl1 on these core NuA4 activities(Kobor et al. 2004; Krogan et al. 2004). In contrast to the reduction observed in the eaf1Δ mutant, removal of the Epl1 C-terminus had no effect on the level of H4 tetra-acetylation (Figure 4.1C). Interestingly, the H4 acetylation levels in the eaf1Δ epl1-CΔ double mutants were similar to that of the epl1-CΔ single mutants and wildtype, strongly implying that removal of the Epl1 C-terminus can rescue bulk acetylation defects caused by loss of Eaf1.   Based on the ability of the EPL1 C-terminal truncation mutants to rescue the defects in H4 acetylation caused by loss of Eaf1, we tested whether this was rooted in changes to NuA4 complex integrity. Proteins associated with a TAP-tagged Epl1 in cells lacking Eaf1 and/or the C-terminus of Epl1 were identified by a small-scale TAP purification followed by silver stain. In agreement with previous reports, loss of EAF1 resulted in a striking decrease in the levels of NuA4 subunits associated with Epl1-TAP, suggesting that the NuA4 complex was largely disassociated in the absence of Eaf1 (Figure 4.1D) (Mitchell et al. 2008). As expected, pull down of epl1-485-TAP and epl1-380-TAP led to purification of Yng2 and     99 Esa1, demonstrating release of the picNuA4 complex upon truncation of the Epl1 C-terminus (Figure 4.1D) (Auger et al. 2008). Most importantly, by combining the EPL1 C-terminal truncation mutants with eaf1Δ, we found that both Epl1-CΔ proteins were sufficient to reconstitute the picNuA4 complex in the absence of Eaf1 (Figure 4.1D). Consistent with the phenotypic data, the restoration picNuA4 complex integrity in an eaf1Δ background suggests that the suppression of eaf1Δ-specific defects such as growth and H4 acetylation was mediated through the stabilization of picNuA4 upon Epl1 C-terminus truncation.  The small-scale TAP purification experiments also revealed that the levels of Epl1 varied among the different strains. Further examination on whole cell extracts demonstrated that Epl1 protein levels were dependent on its own C-terminus and Eaf1 (Figure 4.1E). Specifically, loss of EAF1 led to reduced levels of Epl1 protein while truncation of the Epl1 C-terminus increased Epl1 protein levels. In addition, RT-qPCR data showed that differences in Epl1 protein levels did not occur at the transcriptional level but were rather likely due to protein instability (Figure 4.1F). Taken together, these results suggested that interactions between the Epl1 C-terminus and Eaf1 lie at the core of NuA4 complex stability and function.       100    A"B"C"E" F"Figure"1"D"!Epl1!TAP)full)length)!Epl1!485!TAP)!Epl1!380!TAP)!cross)reac;on)Wildtype)Epl1%TAP)eaf1Δ)Epl1%TAP)epl1%485%TAP)epl1%485%TAP)eaf1Δ)Epl1%380%TAP)eaf1Δ)epl1%380%TAP)    101 Figure 4.1 Epl1 C-terminus truncation suppressed the effects of EAF1 deletion by stabilizing picNuA4 complex formation  (A) Schematic of the Epl1-CΔ mutations.  (B) Ten-fold serial dilutions of the indicated strains were plated on YPD media, with or without genotoxic agents and incubated for 3 days at the indicated temperature. (C) Whole cell extracts of indicated strains were analyzed by protein blotting with anti-H4 tetra acetylation antibodies. Antibodies against H4 were used as loading control. (D) TAP purification of NuA4 using Epl1-TAP demonstrated dissociation of NuA4 subunits in eaf1Δ mutant and stabilization of picNuA4 in epl1-CΔ mutants. Purified fractions from indicated strains were loaded onto a 4-20% gradient SDS-PAGE gel and visualized by silver staining. Bands corresponding to NuA4 subunits are indicated on the left. Untagged Epl1 was used as a negative control. Pull down of Epl1-TAP in EPL1-TAP eaf1Δ strain was verified by western (data not shown). (E) Whole cell extract were analyzed for total Epl1-TAP level by protein blotting with IgG antibody. IgG unspecific band was used as loading control. (F) mRNA level for EPL1 were comparable across all strains examined. mRNA level was measured by RT-qPCR and normalized to levels of TUB1 mRNA.   4.3.2 Loss of Eaf1 and Epl1 C-terminus led to NuA4-Specific Defects at Ribosomal Genes The loss of NuA4 complex in eaf1Δ and epl1-CΔ cells suggested abnormalities in NuA4-specific functions in the cell. In yeast, NuA4 is recruited to promoters of ribosomal protein (RP) genes in an activator-dependent manner and H4 acetylation at these specific genomic regions is required for RP gene expression (Reid et al. 2000). Utilizing chromatin immunoprecipitation (ChIP) followed by qPCR, we determined that epl1 1-380 and eaf1Δ single mutants both exhibited decreased H4 tetra-acetylation at promoters of RPL19B, RPS11B, and RPS3 genes (Figure 4.2A). While both single mutants have substantial H4 acetylation defect, the eaf1Δ epl1 1-380 double mutant also showed a similar decrease in H4 tetra-acetylation at these loci suggesting that they act in the same pathway (Figure 4.2A). This result is in agreement with previous models that indicate picNuA4 does not target H4 acetylation at specific loci such as the RP genes. Consistently, the H4 acetylation pattern closely mirrored the mRNA expression patterns of these genes. As assayed by RT-qPCR, expression of RPL19B, RPS11B, and RPS3 was reduced in both single mutants and in the     102 double mutant (Figure 4.2B). Thus, these results revealed that targeted acetylation by NuA4 at specific loci were lost under both epl1-CΔ and eaf1Δ conditions. Furthermore, while loss of Epl1 C-terminus was sufficient to suppress global H4 acetylation defects in an eaf1Δ background, this inhibition did not extend to loci targeted by NuA4.    Figure 4.2 Loss of Eaf1 and Epl1 C-terminus led to NuA4-specific defects at ribosomal genes  (A)ChIP of H4 tetra-acetylation at promoters of RPL19B, RPS11B, and RPS3 in wildtype, epl1 1-380, eaf1Δ, and epl1 1-380 eaf1Δ. Enrichment of H4 tetra-acetylation was normalized to nucleosome density as measured by H3. (B) mRNA levels were measured by RT-qPCR for the indicated transcripts. mRNA levels for all genes were normalized to levels of ACT1 mRNA. Error bars represent standard errors of the means for three independent experiments. A"B"Figure"2"H4#tetra(acetyl#ChIP#*"*"*" *" *"Expression#level#*"*"*" *"*"*" *"*"*"    103  4.3.3 Distinct and Overlapping Cellular Roles for Eaf1 and the Epl1 C-terminus Built on our findings that a complete deletion of Eaf1 and truncation of the Epl1 C-terminus led to vastly different consequences on NuA4 complex structure and integrity, we decided to perform genome-wide assays to examine the biological consequences of these two mutations in the cell. A combination of whole genome expression profile and Epistatic Mini-Array Profiling (E-MAP) profiling were used to thoroughly examine eaf1Δ, epl1-485 and epl1-380.  As a whole, the yeast transcriptome changed dramatically upon deletion of EAF1 and truncation of Epl1, with an average Spearman correlation of 0.20 against wildtype (Figure 4.3A). However, the gene expression profiles of the three mutants are quite similar with the lowest correlation of 0.799 between eaf1Δ and epl1-485, and the highest correlation of 0.987 between the two epl1-CΔ mutants (Figure 4.3A). This can be attributed to the relatively small number of genes that are differentially expressed in eaf1Δ, epl1-485 and epl1-380 compared to wildtype (Figure 4.3B). There were roughly 150 genes whose expression was significantly altered upon either deletion of EAF1 or truncation of EPL1 (Figure 4.3B). Altogether, only 18 down-regulated genes and 61 up-regulated were shared between eaf1Δ, epl1-485 and epl1-380 (Figure 4.3B). Although NuA4 has been primarily linked to gene activation, our analysis revealed that genes were both up-regulated and down-regulated in the three mutant strains (Figure 4.3B). While epl1-485 and epl1-380 share a significant number of differentially expressed genes, it is clear that there are distinct subsets of genes that are differentially expressed between eaf1Δ and epl1-CΔ (Figure 4.3B and Figure 4.3C).       104 Functional characterization suggested that genes that required either Eaf1 or Epl1 for proper expression were involved in various metabolic processes such as “carbohydrate catabolic process” (p-value = 7.30E-05), “glycoside metabolic process” (p-value = 1.60E-04) and “protein catabolic process” (p-value = 2.7E-04). Furthermore, both Eaf1 and Epl1 were also required for expression of genes in “response to temperature stimulus” (p-value = 3.00E-19). While expression of ribosomal genes was altered in the expression profile of all three strains, they did not pass our stringent significant threshold (data not shown).       105       C"A"Figure"3"F"D"EAF1%HA'eaf1Δ'EPL1%flag'epl1%380%flag'epl1%485%flag'Genes%with%increased%mRNA%levels%eaf1Δ&epl1)380&epl1)485&(7)$ (20)$(42)$(46)$(1)$ (61)$(3)$Genes%with%decreased%mRNA%levels%eaf1Δ&epl1)380&epl1)485&(8)$ (21)$(28)$(36)$(1)$ (18)$ (6)$B"EAF1%HA'eaf1Δ'epl1%380%flag'epl1%485%flag'EPL1%flag'EAF1%HA'eaf1Δ'epl1%380%flag'epl1%485%flag'EPL1%flag'EAF1%HA'eaf1Δ'epl1%380%flag'epl1%485%flag'EPL1%flag'EAF1%HA'eaf1Δ'epl1%380%flag'epl1%485%flag'EPL1%flag' !!!Nega%ve'Gene%c'Interac%ons'eaf1Δ&epl1)380&epl1)485&(63)! (42)!(146)!(27)!(24)!(29)!(9)!!!!eaf1Δ&epl1)380&epl1)485&(89)!(111)!(113)!(45)!(23)!(193)!(28)!Posi%ve(Gene%c(Interac%ons(E"CCR4$NOT(complex(CCR4(POP2(CAF40(MOT2(CAF130(CDC39(–(DAmP(NOT3(NOT5(EAF1$HA(eaf1Δ(epl1$380$flag(epl1$485$flag(EPL1$flag((RPS4B(RPS22A(RPS23A(RPS4A(RPS28B(RPS0A(RPS11A(RPS19B(RPS8A(RPS16A(RPS6B(RPS6A(RPS27B(RPS21B(RPS14A(RPS30A(RPS29A(RPS1B(ASC1(RPS19B(RIBOSOME(BIOGENESIS(COMPASS'SWD1'SWD3'BRE2'SPP1' "HOS2'HDA1'HOS1'UBP3'BRE5'HOG1'' SAS2'SAS5'SAS4'' HAT2'HAT1'SAS'Acetyla<on'Ubiqui<n''protease' SAGA'TAF12'–'DAmP'TAF10'–'DAmP'SGF73'SPT8'SPT7'SGF29'CHD1'    106 Figure 4.3 Genetic and gene expression analysis revealed similarities and distinctions between eaf1 and epl1 mutants  (A) Spearman correlation of gene expression profile of wildtypes and mutant strains. (B) Venn diagram of overlap in up-regulated and down-regulated genes in eaf1Δ, epl1-485, and epl1-380. (C) Unsupervised hierarchical cluster matrix of eaf1Δ and epl1 C-Δ mutants with their respective up- and down- regulated genes. Yellow indicates upregulated genes, and blue represents downregulated genes. (D) Spearman correlation of genetic interaction profiles of wildtypes and mutant strains.  Unsupervised hierarchical cluster matrix of eaf1Δ and epl1 C-Δ mutants with the respective positive and negative genetic interactions. (E) Venn diagram of shared negative genetic interactions and shared positive genetic interactions between eaf1Δ, epl1-485, and epl1-380. (F) Genetic interaction profiles of eaf1Δ and epl1 C-Δ with indicated complexes. Blue indicates aggravating interactions, yellow represents alleviating interactions, and gray denotes missing data.  Next, the mutant strains carrying eaf1Δ or epl1-CΔ derivatives were analyzed by Epistatic Mini Array Profiling (E-MAP), a technique that enables quantitative measurements of aggravating and alleviating genetic interactions against a library of 1,536 mutant genes involved in transcription, RNA processing, and chromatin biology (Schuldiner et al. 2006). Similar to the expression data, correlation analysis of the genetic profiles revealed that the two epl1-CΔ mutants clustered closer to each other than to the eaf1Δ mutant (Figure 4.3D). In line with growth differences between epl1-485 and epl1-380, the two alleles genetically interact with distinct subsets of genes (Negative genetic interactions: p-value = 1.36E-92, Positive genetic interactions: p-value = 2.03E-207) (Figure 4.3E).   E-MAP analysis also revealed similarities and differences between eaf1Δ and epl1-CΔ mutants. For instance, negative genetic interactions existed for both sets of mutants with genes involved in the SAS, HAT2 and SAGA histone acetyltransferase (Figure 4.3F) (Krogan et al. 2004; Mitchell et al. 2008). In contrast, epl1-CΔ and eaf1 mutants differed in their interactions with genes encoding for COMPASS, CCR4-NOT complex and subunits of various histone deacetylases (Figure 4.3F). Despite having mostly alleviating interactions with ribosomal subunit encoding genes, both negative and positive interaction existed for     107 eaf1Δ and epl1-CΔ (Figure 4.3F). Interestingly, all three mutants display strong alleviating interaction with BRE5 and UBP3, genes encoding for a ubiquitin protease implicated in degradation of ribosomal proteins and DNA damage response (Figure 4.3F) (Kraft et al. 2008; Bilsland et al. 2007). Collectively, our large-scale genetic and gene expression analyses demonstrated that EAF1 and EPL1 have overlapping yet divergent functions, which is not easily reconciled with these proteins acting similarly in regulating picNuA4 levels and activity.   4.3.4 The Eaf1 HSA Domain was Required for NuA4 Stability and Function Given the drastic decrease in NuA4 levels in eaf1Δ cells, we asked whether specific domains of EAF1 were important for the stability of Epl1 and the NuA4 complex. To this end, internal truncation alleles of EAF1 lacking the HSA and/or SANT domains were created (Figure 4.4A). While cells lacking the Eaf1 HSA domain had phenotypes similar to deletion of EAF1, including defects in both H4 and H2A.Z acetylation and sensitivity to genotoxic stress, cells lacking the Eaf1 SANT domain had histone acetylation levels and growth fitness comparable to those of wildtype cells (Figure 4.4B and Figure 4.4C). Given that dissociation of the NuA4 complex likely was responsible for the drastic phenotype in eaf1Δ, it prompted us to examine NuA4 stability in the EAF1 internal deletion mutants. Consistent with the phenotypic similarity between eaf1Δ and eaf1 HSAΔ, loss of the HSA domain alone led to the dissociation of the NuA4 complex, indicating that the HSA domain of Eaf1 is important for NuA4 stability (Figure 4.4D). On the other hand, we were able to purify the complete NuA4 complex in the absence of the Eaf1 SANT domain (Figure 4.4D). Samples from the small-scale TAP purification were further analyzed by western blotting to confirm the presence of     108 Epl1-TAP in all strains. Similar to before, we observed a decrease in Epl1 level in the absence of EAF1. Furthermore, strains with the HSA internal deletion also exhibited a similar decrease in Epl1 levels, suggesting that the HSA domain is important for full-length Epl1 stability (Figure 4.4E). In agreement with the results from the silver stain, which demonstrated that the HSA domain was required for NuA4 and picNuA4 stability, we could not detect any Epl1-associated Esa1 in cells lacking the Eaf1 HSA domain (Figure 4.4E). Similar to before, experiments on whole cell extracts confirmed that the stability of the NuA4 catalytic module was dependent on the HSA domain and to a lesser extent on the SANT domain (Figure 4.4F).   Our results placed the Eaf1 HSA domain and the Epl1 C-terminus at the center of NuA4 complex formation and function. As many lines of evidence, including our own, suggest that the Epl1 protein is anchored to Eaf1 through its C-terminus, we would predict that truncation the Epl1 would suppress the eaf1 HSAΔ growth (Auger et al. 2008; Szerlong et al. 2008). Loss of the Eaf1 HSA domain led to a similar growth defect compared to a complete loss of Eaf1 (Figure 4.5A). Truncating the C-terminus of Epl1 in either an eaf1Δ background or an eaf1 HSAΔ background rescued the growth impediment to levels similar to the epl1-CΔ single mutant (Figure 4.5A). Taken together, our observations suggested that the truncation of Epl1 C-terminus partially suppressed eaf1-HSAΔ defects.      109      A"Figure"4"B"C"D" E"F"WT  Epl1-TAP pEAF1 vector peaf1 HSAΔ peaf1 HSAΔ SANTΔ peaf1 SANTΔ eaf1Δ, Epl1-TAP - rIgG against    Epl1-TAP - Esa1 - cross    reaction - Eaf1     110 Figure 4.4 The Eaf1 HSA domain was required for NuA4 stability and function (A) Schematic representation of EAF1 internal region deletions: HSAΔ, SANTΔ, and HSAΔ/SANTΔ. (B) Whole cell extracts of indicated strains were analyzed by protein blotting with anti-H4 tetra- or anti-H2A.Z K14 acetylation antibodies. Antibodies against H4 were used as a loading control. (C) Ten-fold serial dilutions of the indicated strains were plated onto Sc –URA media containing the indicated genotoxic agents. (D) TAP purification of NuA4 using Epl1-TAP demonstrated disassociation of NuA4 in eaf1 HSAΔ. Purified fractions from indicated strains were loaded onto a 4-20% gradient SDS-PAGE gel and visualized by silver staining. Bands corresponding to NuA4 subunits are indicated on the left. Untagged Epl1 was used as a negative control. (E) TAP purifications of indicated strains were loaded on western to verify Epl1-TAP pull down and Esa1 association. IgG and anti-Esa1 antibodies were used to detect Epl1-TAP and Esa1 respectively. (F) Whole cell extract were analyzed for total Epl1-TAP, Eaf1 and Esa1 level by protein blotting. IgG unspecific band was used as loading control.      Figure 4.5 epl1-CΔ truncation was epistatic to eaf1 internal truncation mutants Ten-fold serial dilutions of the indicated strains were plated onto Sc –URA media containing the indicated genotoxic agents.    A"Figure"5"    111 4.3.5 Epl1 C-terminus was Required for NuA4 Association with Chromatin To gain a better understanding of picNuA4 function in the context of the epl1-CΔ strain, genome-wide ChIP-on-chip experiments was performed on Epl1-TAP, epl1-485-TAP, and epl1-380-TAP. We expected that the Epl1-TAP occupancy profile would be representative of binding sites for both NuA4 and picNuA4 whereas the occupancy profile of Epl1 lacking the C-terminus (485 and 380) would be representative of sites enriched for picNuA4. Validating our ChIP-on-chip approach, wildtype Epl1 was highly enriched over ribosomal genes as previous reported (Figure 4.6A). In agreement with our H4 acetylation ChIP-qPCR and RT-qPCR data, Epl1 lacking the C-terminus was significantly depleted from the majority of ribosomal genes previously enriched for Epl1-TAP (Figure 4.6B). Furthermore, we found few sites besides ribosomal protein genes to be enriched with NuA4 under our selection criteria. This is similar to previous efforts at mapping NuA4 occupancy sites through HA-Esa1 as a representative subunit of NuA4 (Reid et al. 2000). Surprisingly, Epl1 occupancy profiles in epl1-485-TAP and epl1-380-TAP strains yielded few significantly enriched sites across the genome. This result suggested that despite an increase in picNuA4 levels and normal H4 acetylation in the C-terminus truncation mutants, picNuA4 was transiently associated with chromatin under these circumstances.   Subsequently, we used bulk fractionation assay to examined picNuA4 occupancy in epl1-CΔ mutants as another measure of picNuA4 association to chromatin. Using centrifugation-based sucrose gradient, chromatin-bound proteins (pellet fraction) were separated from whole cell lysate (supernatant fraction). In the wildtype strain, almost all of the Epl1-TAP in the cell fractioned tightly with the chromatin pellet leaving no detectable amounts of Epl1-TAP in     112 the supernatant fraction (Figure 4.6C). In addition, despite the decreased levels of Epl1-TAP in an eaf1Δ background, Epl1-TAP was enriched in the chromatin fraction with no detectable amounts in the supernatant (Figure 4.6C). Interestingly, upon truncation of the Epl1 C-terminus, a shift in the localization of epl1-485-TAP and epl1-380-TAP was observed. Instead of being highly enriched in the chromatin pellet, the levels of chromatin-bound epl1-CΔ proteins were lower than that of wildtype (Figure 4.6C). Furthermore, there were low levels of epl1-485-TAP and epl1-380-TAP found in the supernatant fraction (Figure 4.6C). Despite a decrease in the levels of chromatin-associated Epl1 in the truncation mutants, the levels of acetylated H4 in chromatin remained the same (Figure 4.6C). Taken together with our genome-wide occupancy profiles, the data suggested that the picNuA4 released in epl1-CΔ strains was still sufficient to acetylate H4 in chromatin despite being loosely associated.       113  Figure 4.6 Epl1 C-terminus was required for NuA4 association to chromatin  (A) ChIP-chip profile of NuA4 and picNuA4 bound regions in EPL1-TAP, epl1-485, and epl1-380. Sample genomic regions were plotted for ChrIII and ChrX along the x-axis against the relative occupancy of Epl1 normalized over input. ORFs are indicated as light gray rectangles above the x-axis for Watson genes and below the x-axis for Crick genes. (B) Epl1 C-terminus was required for NuA4 association to ribosomal genes. Relative occupancy of Epl1 over input signal at promoters of ribosomal genes. (C) Loss of Epl1 C-terminus reduced Epl1 association to chromatin. WCE, whole cell extract; SUP, supernatant; CHR, chromatin pellet. The relative amounts of Epl1-TAP and H4 tetra acetylation was determined by immunoblotting. In each case, the fractionation efficiency was judged by the levels of the control proteins, H2A and Pgk1, found in the chromatin fraction and the supernatant fraction, respectively.   A"Figure"6"ChrII#ChrX#B"Relative occupancy Relative occupancy RPS9B# RPL21A#RPS14B# RPL39#RPS22A#RPL17B#C"!"Untagged"!"Epl1!TAP"!"Epl1!485!TAP"!"Epl1!380!TAP"    114 4.4 Discussion  This study explored the functional and structural connection between two subunits of the NuA4 HAT complex in Saccharomyces cerevisiae. Our experiments focused on dissecting the circuitry between of the Eaf1 protein and the Epl1 C-terminus in regulating NuA4 function and composition. Genome-wide expression and genetic interaction data revealed an expected set of common requirements, but more surprisingly, also remarkably different phenotypic and functional consequences. Detailed biochemical examination revealed that full length Epl1 did not support the formation of a detectable levels picNuA4 complex in the absence of Eaf1. However, deletion of the Epl1 C-terminus restored and stabilized a functional picNuA4 complex in the absence of EAF1, suggesting that interaction between the Epl1 C-terminus and Eaf1 may be necessary for complex stability. Furthermore, Eaf1 was important for maintaining the protein level of individual picNuA4 subunits, including Epl1. We subsequently determined that the HSA domain of Eaf1, the proposed interaction domain of Eaf1 with the Epl1 C-terminus, was required for picNuA4 formation and function. Despite being sufficient to restore global H4 acetylation levels in an eaf1Δ background, the picNuA4 released in epl1-CΔ strains displayed reduced interaction with chromatin. Taken together, our combination of whole-genome investigation and detailed biochemical analysis revealed unexpected differences and similarities of epl1-CΔ and eaf1Δ in NuA4 complex stability and HAT activity.   The present work shed light on two existing and somewhat conflicting models of NuA4 structural integrity. The first model suggests that upon deletion of the scaffolding Eaf1     115 protein, the various submodules of the NuA4 complex remain intact, leaving behind a functional and stoichiometric picNuA4 complex sufficient for un-targeted chromatin acetylation (Auger et al. 2008). The alternative model suggests that Eaf1 is essential for NuA4 complex integrity and that upon deletion of EAF1, there is a significant reduction in the level of NuA4 in the cell (Mitchell et al. 2008). Both models place Eaf1 at the center of NuA4 function through a combination of biochemical and genetic means. However, the defining difference between the two models centers on the importance of Eaf1 in regulating the quantitative balance between the various submodules, and in particular picNuA4. In principle, the data presented in this chapter supports the co-existence of both models. The strong decrease of H4 acetylation in the eaf1Δ mutant suggests that the released picNuA4 had compromised HAT activity (Krogan et al. 2004; Kobor et al. 2004). Supported by the NuA4 purification experiments, we observed a significant reduction in the picNuA4 subunits that co-purified with Epl1-TAP when EAF1 was deleted. However, in contrast to the eaf1Δ mutant, the eaf1Δ epl1-CΔ mutants have a fully active minimal picNuA4 capable of maintaining basal H4 acetylation levels, suggesting that the catalytic module of NuA4 exists as a functional and independent entity. Consistent with published in vitro data, this suggested that the picNuA4 formed under this condition is capable of untargeted chromatin acetylation of the native picNuA4 complex (Boudreault et al. 2003).  The work presented in this dissertation consolidates the two prevailing models in the field: we found that while EAF1 is central for NuA4 stability and function, removal of Epl1 C-terminus was sufficient to restore the growth and H4 acetylation defects of eaf1Δ. Furthermore, while both models suggest that loss of Eaf1 or loss of Epl1 C-terminus equally     116 liberate picNuA4, the data presented in this chapter are difficult to reconcile with such a simple model. Gene expression array analysis identified both overlapping and divergent gene expression clusters between the mutants. Similarly, genetic epistasis analysis uncovered unique genes required to support growth of eaf1Δ and epl1-CΔ. Taken together with our biochemical examination, our data supports that these two genetic manipulations led to distinct consequences in NuA4 structure and function.   Our combinatorial analysis of cells lacking Eaf1 and the Epl1 C-terminus revealed a novel functional relationship between these two key components of the NuA4 HAT complex. In line with previously reports, cells lacking EAF1 displayed decreased fitness under genotoxic stress and reduced global H4 acetylation as a result of the dissociation of the HAT complex (Kobor et al. 2004; Mitchell et al. 2008; Boudreault et al. 2003; Auger et al. 2008; Babiarz et al. 2006). This effect however, was due to the presence of the Epl1 C-terminus, as removal of the C-terminus domain was sufficient to restore the bulk H4 acetylation and growth phenotype of eaf1Δ. Specifically, analytical-scale purification experiments identified the stable assembly of picNuA4 upon removal of the Epl1 C-terminus as the likely mechanism underlying the suppression in eaf1Δ epl1-CΔ double mutants. Importantly, we demonstrated that Eaf1 is only required for picNuA4 function and stability in the presence of full length Epl1 protein, highlighting a potential regulatory role of the Epl1-C terminus in picNuA4 formation. The inverse relationship between the stability of picNuA4 subunits and the presence of Eaf1 in the cell suggests that Eaf1 may also have an important, yet likely indirect, regulatory role in the stability of the NuA4 catalytic module. In line with this potential role of Eaf1, reduced levels of the Eaf3/5/7 RpdS-associated module have been     117 reported in the absence of EAF1 (Rossetto et al. 2014). Detailed domain deletion analysis of Eaf1 identified its HSA domain as a key player in the stability of picNuA4 as deletion of the Eaf1 HSA domain led to growth and global H4 acetylation defects. Similarly, eaf1 HSAΔ cells also exhibited reduced picNuA4 stability akin to the eaf1Δ mutant, while loss of the Eaf1 SANT domain had little effect on cell fitness, NuA4 stability and H4 acetylation. As a whole, the eaf1 HSAΔ mutant phenocopied the eaf1Δ mutant suggesting that the HSA domain was the key region required for picNuA4 function and stability. Taken together, our data suggested that the interface between the Eaf1 HSA domain and the Epl1 C-terminus is required for maintaining the equilibrium of active NuA4 and picNuA4 in the cell.   Taking advantage of the release of a functional picNuA4 upon Epl1 C-terminal truncation, we examined the genome-wide binding profile of NuA4 and picNuA4. Similar to previous efforts to map NuA4, Epl1 was highly enriched over genes encoding for ribosomal proteins (RP) (Reid et al. 2000). In agreement with the current model underlying the functional differences between NuA4 and picNuA4, the peaks associated with RP promoters likely are a result of the targeted recruitment of NuA4 complex by activators such as Rap1 or Abf1 (Reid et al. 2000). Adding to our gene-specific analysis of H4 enrichment and mRNA levels at RPL19b, RPS11B, and RPS3, loss of Epl1 peaks genome-wide at RP promoters in epl1-CΔ confirmed that these peaks corresponded to NuA4 binding and not picNuA4. Distinct from NuA4, picNuA4 is proposed to be diffused over the genome and is responsible for basal acetylation (Boudreault et al. 2003; Auger et al. 2008; Reid et al. 2000). Therefore, the lack of distinct peaks other than the RP peaks in the wildtype Epl1 binding profile suggested that picNuA4 was either lowly enriched or transiently associated with the rest of the genome.     118 Despite the dramatic increase in picNuA4 levels in the cell upon Epl1 C-terminus truncation, there were few novel sites of Epl1-CΔ binding genome-wide.   Interestingly, bulk chromatin association assays provided novel insight into the cellular changes occurring in these mutant strains when the normal equilibrium between NuA4 and picNuA4 levels was disturbed. Upon C-terminal truncation of Epl1, there was a shift from predominantly chromatin-associated Epl1 to equal levels of chromatin-bound and free-floating picNuA4 in these cells. However, the level of acetylated H4 in chromatin remained the same, suggesting that the picNuA4 that remained associated with chromatin is sufficient to maintain basal acetylation. As we observed an increase in picNuA4 levels in epl1-CΔ cells, it is tempting to speculate that non-chromatin bound picNuA4 formed under these circumstances might erroneously acetylate non-chromatin specific targets, thus leading to the dramatic growth defects in these mutants. This would not be unexpected as NuA4 is well documented to have numerous non-chromatin acetylation functions (Mitchell et al. 2011; Downey et al. 2014; Lin et al. 2009).        119  Figure 4.7 Model for the regulatory mechanisms of Eaf1 and Epl1 C-terminus in picNuA4 stability (A) NuA4 and picNuA4 co-exist in wildtype cells. picNuA4 catalytic module associates with NuA4 through the Epl1 C-terminus and Eaf1 scaffold protein. (B). Deletion of Eaf1 and its HSA domain lead to reduced picNuA4 levels, possibly mediated by a degradation pathway. (C) Removal of the Epl1 C-terminus in an eaf1Δ background restores function on chromatin and increases cellular picNuA4 level.  Consolidating this set of multifaceted data, we put forth the following model: adding to previous work suggesting that picNuA4 docks onto Eaf1 through the Epl1 C-terminus, we propose that in addition to this physical interaction, the Eaf1 HSA domain and Epl1 C-terminus are key regulators important for maintaining the cellular balance of NuA4 and picNuA4 (Figure 4.7). We demonstrated that without Eaf1 or its HSA domain, the picNuA4 subunits (Yng2 and Esa1) that dock upon the Epl1 EPcA domain for stability no longer interacts with Epl1. It is tempting to propose that, perhaps, the interaction between Eaf1 (specifically HSA domain) and Epl1 C-terminus is required for Epl1 and picNuA4 stability (Figure 4.7B). It would be interesting to examine whether Eaf1 is indirectly involved in regulating picNuA4 stability through an ubiquitin degradation pathway, especially in light of Figure"7"Eaf1"Eaf7#Eaf3#Eaf5#picNuA4"Tra1#Arp4#Swc4#Act1#Yaf9#Eaf6#Epl1#Esa1#Yng2#NuA4"eaf1"HSAΔ(epl1+CΔ(eaf1(Δ A." B."C."Eaf1"Eaf7#Eaf3#Eaf5#Tra1#Arp4#Swc4#Act1#Yaf9#degradaGon#Eaf7#Eaf3#Eaf5#Tra1#Arp4#Swc4#Act1#Yaf9#Eaf6#Epl1#Esa1#Yng2#(COOH#    120 the genetic connection with the Ubp3/Bre5 ubiquitin protease highlighted by E-MAP. Secondly, our data also demonstrated that upon truncation of the Epl1 C-terminus, picNuA4 function and stability was restored in an eaf1Δ background (Figure 4.7C). Moreover, there was an increase in the cellular levels of picNuA4 in epl1-CΔ cells. Our data underscores the regulatory relationship between Eaf1 and Epl1 C-terminus in picNuA4 formation. Taken together, the evidence presented in this chapter supports a model where the stability and levels of picNuA4 is regulated by the Epl1 C-terminus, and that this regulation occurs in concert with the central scaffold protein, Eaf1.       121 Chapter 5: Conclusion  Since the identification of DNA as the genetic material, immense strides have been made in understanding how DNA stores information and how the information is interpreted. We now recognize that it is not the DNA sequence alone, but also how the DNA is packaged that regulates our cells’ ability to read and utilize the genetic information. Realizing that chromatin structure regulates the accessibility of the information contained within DNA was a key breakthrough in the field of chromatin biology. Decades of intense research highlighted the multiple mechanisms that the cell has evolved to package and regulate access to the DNA sequence. Scientists are now embarking on an exciting journey to tease apart how histone modifications, chromatin-remodeling complexes, histone variants, and histone chaperones work together to coordinate the many biological processes that are dependent on DNA.   My dissertation adds to our current understanding of the coordinated activities of the many chromatin-modifying factors in our cells. Spanning multiple chromatin-modifying mechanisms in S. cerevisiae, this dissertation examines the functional relationships between histone chaperones, histone variants, post-translational modifications, and chromatin-modifying complexes. More specifically, I explored the mechanistic link between the Asf1 histone chaperone and the SWR1-C chromatin-remodeling complex at subtelomeric regions. I determined that Asf1-dependent H3K56 acetylation was an integral component of H2A.Z nucleosome occupancy. Moreover, I characterized key players in NuA4 histone acetyltransferase biology while at the same time identifying crucial domains that regulate the dynamic equilibrium of its structure and HAT function.      122   Figure 5.1 Model for Asf1 and SWR1-C interactions in Saccharomyces cerevisiae (A) Asf1 cooperates with SWR1-C to restrict spread of heterochromatin. (B) Asf1-dependent H3K56 acetylation contributes to H2A.Z nucleosome dynamics at gene promoters.    Our understanding of the broad functions of the Asf1 H3/H4 histone chaperone has come a long way since its identification in an anti-silencing screen almost two decades ago (Le et al. 1997; Singer et al. 1998). In addition to delivering H3/H4 heterodimers to chromatin-remodelers and chaperones for nucleosome assembly, we are now starting to appreciate the functional connections between Asf1 and chromatin-modifying complexes regulating the chromatin landscape. In Chapter 2 of this dissertation, I described one such interaction between Asf1 and the SWR1-C H2A.Z-depositing complex. Using Yaf9 as a representative heterochroma)n+Swr1% Yaf9%H2A.Z+A"B"Swr1% Yaf9%H2A.Z+H3K56ace+ORF+    123 subunit of SWR1-C activity at heterochromatin, I found that Asf1 cooperated with the entire SWR1-C to maintain three heterochromatin boundaries on ChrIII R by preventing the spread of SIR complexes onto nearby subtelomeric regions (Figure 5.1). While we know H2A.Z is required for preventing heterochromatin spread at the boundaries of heterochromatin and euchromatin, the exact mechanism of the cooperative activities between Asf1 and SWR1-C at telomeres remains to be determined (Meneghini et al. 2003). It is likely that Asf1’s role at heterochromatin lies within its key function as a chaperone for H3K56 acetylation, as H3K56 mutants have impaired telomeric silencing in reporter assays and native telomeres (Xu et al. 2007; Yang et al. 2008; Miller et al. 2008). However, while H3K56 mutants exhibit normal telomere occupancy of Sir2, Sir3 and Sir4, I observed increased Sir2 levels at telomeres and spread of Sir2 into subtelomeres in asf1Δ	  yaf9Δ	  cells	  (Xu	  et	  al.	  2007). Interestingly in S. pombe, Asf1/HIRA cooperates with the Clr6 HDAC complex to maintain heterochromatin silencing and associates with SHREC (Snf2/HDAC repressor complex) to promote nucleosome occupancy at heterochromatic loci (Yamane et al. 2011; Kaufman 2011). While these activities in fission yeast are dependent on HP1 and H3K9me3, neither of which exists in S. cerevisiae, it alludes to the existence of a similar heterochromatin regulatory pathway involved in the cooperative activities between Asf1 and SWR1-C in S. cerevisiae.    In addition to Asf1’s role at heterochromatic regions, I found that Asf1 also contributed to maintaining H2A.Z occupancy at euchromatic and subtelomeric gene promoters. This finding is in line with recent publications demonstrating that Asf1 works with chromatin-modifying complexes in a variety of biological contexts. For example, Asf1 cooperates with the SWI/SNF complex to displace nucleosomes at heat shock gene promoters (Erkina and Erkine     124 2014).  Asf1 also recruits chromatin-modifying complexes, such as the SAS complex and the Set2 methyltransferase complex, to promote H4K16 acetylation and H3K36 trimethylation, respectively (Lin et al. 2010; Osada et al. 2005). Given Asf1’s role in the assembly and disassembly of nucleosomes during DNA replication and transcription, it is likely other chromatin-modifying complexes may be active participants in these biological processes.   The genetic relationship between genes encoding for SWR1-C and ASF1 also alludes to a functional connection that extends beyond heterochromatin maintenance. As I demonstrated in Chapter 2, cells lacking Asf1 and a component of SWR1-C are particularly sensitive to cold temperatures and genotoxic agents. The synergistic growth defect in cells lacking functional components of both chromatin-modifying factors suggested that SWR1-C and Asf1 have additional redundant functions. Indeed, both Asf1 and H2A.Z contribute to chromosome positioning at the nuclear periphery. Asf1 and H3K56 acetylation are crucial for the association of yeast telomeres to the nuclear periphery and long-range interactions between heterochromatic loci (Hiraga et al. 2008; Miele et al. 2009). H2A.Z’s role in transcriptional memory appears to be tightly linked to its role in regulating the nuclear-periphery localization of repressed genes such as INO1 and GAL1 (Brickner et al. 2007). However, the connection between nuclear anchoring and transcriptional memory remains controversial as other has found that H2A.Z is not required for transcriptional memory of GAL1 (Halley et al. 2010; Kundu and Peterson 2010). Nevertheless, more experiments are needed to address whether H2A.Z and Asf1 are contributing to the three-dimensional organization of chromosomes independently or redundantly.       125 Taken together, our current understanding of Asf1 would suggest that this histone chaperone has two important functions within the cell. Primarily, Asf1provides free histones for nucleosome assembly and receives H3/H4 heterodimers during nucleosome disassembly. Secondly, it is tempting to speculate that Asf1 is also a key player in establishing distinct chromatin neighbourhoods by recruiting additional chromatin-modifying complexes such as HATs, histone-methyltransferases and chromatin remodelers. It remains to be seen whether the latter function is directly connected to Asf1’s primary role as a histone chaperone.   In Chapter 2 and 3 of this dissertation, I identified that H3K56 acetylation by Rtt109 was required for maintaining H2A.Z levels at discreet regions of subtelomeric and euchromatic gene promoters. A histone modification that requires binding of the H3/H4 heterodimer by Asf1histone chaperone, loss of H3K56 acetylation resulted in global changes in H2A.Z occupancy and position. At a subset of gene promoters, H2A.Z was either completely lost or the levels of the histone variant were reduced. At other gene promoters, the position of H2A.Z peaks shifted with respect to the TSS when H3K56 could no longer be acetylated. Our genome-wide approach in Chapter 3 led us to propose that acetylation of H3K56 was crucial in maintaining both H2A.Z levels and specific positioning of H2A.Z at promoters. Several lines of evidence supported a role for H3K56 acetylation in regulating nucleosome positioning. When H3K56 acetylation was first identified in S. cerevisiae, it was proposed that this PTM promotes nucleosome instability by loosening the DNA-histone interaction based on its location at the entry and exit point of DNA (Xu et al. 2007; Neumann et al. 2009). Subsequent structural analyses showed that binding of DNA to the modified tetramer (H3K56ace/H4)2 is much weaker compared to the unmodified tetramer, suggesting that     126 H3K56 acetylation contributes to nucleosome positioning (Andrews et al. 2010). By promoting DNA breathing in tetrameric form and locking in the nucleosome position once the full nucleosome is assembled, H3K56 acetylation has the potential to facilitate nucleosome positioning during nucleosome assembly (Andrews et al. 2010; Watanabe et al. 2010). In this regard, perhaps it is H3K56’s acetylation status during nucleosome assembly that is important for positioning H2A.Z containing nucleosomes at NFRs.   An interesting avenue of research lies within the precise regulatory mechanism between H3K56 acetylation and H2A.Z deposition. One may hypothesize that the relationship between H3K56 acetylation and H2A.Z is characterized by the PTM’s role in nucleosome positioning; H3K56 acetylation positions the +1 and -1 nucleosomes, thus creating the distinct NFR required for SWR1-C recruitment and H2A.Z deposition (Ferreira et al. 2007; Ranjan et al. 2013). Or perhaps a more intriguing idea is the possibility that H3K56 acetylation is directly involved in positioning H2A.Z-containing nucleosomes during SWR1-C catalyzed histone exchange. The latter hypothesis is in line with recently published data suggesting that the Swc2 subunit of SWR1-C interacts with H3K56 on nucleosomes to “lock” in H2A.Z containing nucleosomes (Watanabe et al. 2013). Furthermore, constitutively acetylated H3K56 also lead to the reduced nucleosomal H2A.Z, suggesting that perhaps the act of acetylating and deacetylation H3K56 was important for regulating H2A.Z position and occupancy (Watanabe et al. 2013)( Phoebe Lu, Data not shown).  At this time it is unclear whether the constitutive presence and absence of H3K56ace are affecting H2A.Z occupancy in the same manner; detailed nucleosome position and biochemical analyses are needed to tease apart these intricate mechanisms.      127  My work in Chapter 4 focused on teasing apart the structural and functional aspects of the NuA4 histone acetyltransferase. By examining the Epl1 and Eaf1 subunits that connect the catalytic sub-module to the rest of the multi-subunit complex, I found that while Eaf1 and its HSA domain were required for NuA4 stability and function, truncation of Epl1 C-terminus in an eaf1Δ background was sufficient to rescue picNuA4 formation and H4 acetylation. Furthermore, epl1-CΔ eaf1Δ double mutants had notably increased Epl1 levels and higher picNuA4 levels compared to wildtype. Taken together, I proposed that Eaf1 and the Epl1 C-terminus play a regulatory role in the formation of picNuA4 in the cell and likely control the cellular equilibrium of NuA4: picNuA4. This model raises the interesting question of how are Eaf1 and the Epl1 C-terminus regulating NuA4 complex stability. One suggestion is that docking of the Epl1 C-terminus onto the Eaf1 HSA domain stabilizes picNuA4. While an attractive model, if it were true, I would expect Eaf1 to exist as a stable component of picNuA4. Nevertheless, it is clear that Eaf1 appears to be protecting the Epl1 C-terminus from proteolytic degradation; perhaps Eaf1 regulates picNuA4 stability indirectly via NuA4’s acetylation activity on non-histone targets. It is tempting to speculate that one such target is the ubiquitin-mediated degradation pathway. Indeed, the mammalian homolog of NuA4 is targeted for degradation by the ubiquitin ligase Mdm2 and a recent screen identified subunits of the proteasome as Esa1 acetylation targets (Legube et al. 2002; Mitchell et al. 2013). Furthermore, genes encoding for NuA4 subunits genetically interact with genes in ubiquitin-dependent catabolic pathways (Lu et al. 2009)( Chapter 4). Thus, it seems likely that NuA4 stability could be regulated through a ubiquitin-mediated degradation pathway. Elucidating the molecular mechanism and functional connection between NuA4 and the degradation     128 pathway will shed light on how NuA4 activity is regulated in the cell. Identification of ubiquitination targets and sites within the NuA4 complex serves as a crucial first step in teasing apart this potential regulatory pathway. It is likely that the Epl1 C-terminus is a target for ubiquitin-mediated degradation.    We are beginning to appreciate a broader role for HATs in acetylating many non-histone targets through large-scale proteomic screens aimed at identifying acetyl lysine residues in S. cerevisiae (Henriksen et al. 2012; Lin et al. 2009; Mitchell et al. 2013). A growing list of literature describing NuA4’s role in auto-acetylation provides a potential mechanism for how NuA4 may be regulating its own structure and function. To date, 42 sites of auto-acetylation have been identified on 10 out of the 13 NuA4 subunits (Mitchell et al. 2013; Yuan et al. 2012; Lin et al. 2008). The Epl1 EPcA domain appears to be a key target of NuA4 acetylation and these modifications on Epl1 contribute to the catalytic activity of NuA4 (Mitchell et al. 2013). Interestingly, it appears that picNuA4 is sufficient for auto-acetylation of Epl1 in vitro, raising the need to examine the contributions of NuA4 and picNuA4 for general lysine acetylation (Berndsen et al. 2007). Systematic mutation of sites of NuA4 auto-acetylation would not only uncover acetylation sites needed for NuA4 HAT activity, but also provide novel insight into the regulatory potential of these sites in NuA4 complex stability.   From our findings in Chapter 4, I predicted that severe growth defect in cells lacking the Epl1 C-terminus may have resulted from increased picNuA4 activity as a general lysine acetyltransferase. Indeed, although epl1-CΔ resulted in a dramatic increase in picNuA4 levels in the cell, the level of total H4 acetylation did not increase proportionally. Furthermore, a     129 substantial fraction of picNuA4 in epl1-CΔ cells is found in the supernatant instead of being primarily chromatin-bound suggesting that a significant portion of picNuA4 acts on non-chromatin targets. A recent large-scale screen identified over 80 proteins as Esa1 targets, with the majority of proteins containing more than one acetylation site (Mitchell et al. 2013). The transfer of an acetyl group onto the lysine residues of proteins can alter their localization, activity, stability and interactions with their target proteins (Spange et al. 2009; Yang 2004). Despite a tendency for metabolic enzymes and multi-subunit complexes to be highly acetylated, the biological consequences of the majority of lysine acetylation events identified thus far remain unclear. To fully understand the functional consequence of truncating the Epl1 C-terminus, we also need to examine the acetylation status of non-histone proteins in epl1-CΔ cells. Furthermore, this approach will also allow us to differentiate between non-histone targets of picNuA4 and NuA4.   The various domains of Eaf1 are central to its fundamental role as a scaffolding platform for NuA4. The N-terminal region of Eaf1 associates with the Eaf3/5/7 submodule that is linked to the elongating RNA polymerase II to promote DNA accessibility (Auger et al. 2008; Rossetto et al. 2014). Previous studies show that the SANT domain is required for binding to Tra1, a shared subunit of NuA4 and SAGA that has a regulatory role in targeting the catalytic activity of both HATs (Auger et al. 2008; Grant et al. 1998; Knutson and Hahn 2011; Helmlinger et al. 2011). However, the lack of growth defect in an eaf1-SANTΔ is more in line with data I presented in Chapter 4 demonstrating that Tra1 binding does not require the SANT domain (Auger et al. 2008). Lastly, while previous studies reported that the shared module docked onto the HSA domain of Eaf1, our more precise excision of the HSA domain     130 resulted in the complete disassociation of NuA4. While unexpected, our finding highlighted a key role in the Eaf1 HSA domain in NuA4 complex integrity and function. Intriguingly, the mammalian homologue of NuA4, TIP60, contains an Eaf1-like protein as the key scaffolding subunit, p400. Since TIP60 complex is a functional and structural merge of yeast SWR1-C and NuA4, it does not come as a surprise that p400 appears to be an Eaf1-Swr1 fusion protein (Auger et al. 2008; Doyon et al. 2004; Lu et al. 2009). p400 contains SANT, HSA, and ATPase domains, whereas neither of the yeast homologues contain all three domains. The mammalian p400 appears to have the ATPase domain of Swr1 inserted into Eaf1 between the HSA and SANT domain (Auger et al. 2008; Lu et al. 2009). Indeed, sequence analysis of HSA domain across eukaryotic evolution suggests that the HSA domain of p400 originated from Eaf1, highlighting the functional importance and conservation of the Eaf1 HSA domain (Lu et al. 2009).   In closing, the interplay between the various types of chromatin modifications is emerging as a central mechanism by which cells regulate fundamental biological processes. While S. cerevisiae has proven to be an effective model organism to tease apart the crosstalk between chromatin-modifying factors, there remains a need to examine similar interactions in higher eukaryotes. Emerging evidence suggests that the human homologues, TIP60 and SRCAP, play critical roles in choreographing intricate developmental steps and cancer progression. Thus, increasing the need to examine the crosstalk between these chromatin-remodelers and the “message” contained within distinct chromatin neighbourhoods by way of post-translational modifications. The work presented in this thesis makes important contributions     131 towards understanding the interplay between various chromatin-modifying mechanisms in the cell and provides guidance for further investigation in higher eukaryotes.        132 References    Abbott, D. W., V. S. Ivanova, X. Wang, W. M. Bonner, and J. 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