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Histone acetylation, not stoichiometry, regulates linker histone binding in Saccharomyces cerevisiae Lawrence, Mackenzie Brie Dorothea 2019

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       HISTONE ACETYLATION, NOT STOICHIOMETRY, REGULATES LINKER HISTONE BINDING IN SACCHAROMYCES CEREVISIAE  by Mackenzie Brie Dorothea Lawrence  BSc., The University of Guelph, 2015   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF   MASTER OF SCIENCE  in   FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Biochemistry and Molecular Biology)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  April 2019  © Mackenzie Brie Dorothea Lawrence, 2019         ii COMMITTEE PAGE The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, a thesis/dissertation entitled:  Histone Acetylation, Not Stoichiometry, Regulates Linker Histone Binding in S. cerevisiae   submitted by Mackenzie B. D. Lawrence  in partial fulfillment of the requirements for the degree of Master of Science in Biochemistry and Molecular Biology   Examining Committee: Dr. LeAnn Howe Supervisor  Dr. Vivien Measday  Supervisory Committee Member   Supervisory Committee Member Dr. Peter Stirling  Additional Examiner     Additional Supervisory Committee Members:  Supervisory Committee Member  Supervisory Committee Member         iii ABSTRACT Linker histones play a fundamental role in shaping chromatin structure, but how their interaction with chromatin is regulated is not well understood. A combination of genetic and genomic approaches were used to explore the regulation of linker histone binding in the yeast, Saccharomyces cerevisiae. We found that increased expression of Hho1, the yeast linker histone, resulted in a severe growth defect, despite only subtle changes in chromatin structure. Further, this growth defect was rescued by mutations that increase histone acetylation. Consistent with this, genome-wide analysis of linker histone occupancy revealed an inverse correlation with histone tail acetylation in both yeast and mouse embryonic stem cells. Collectively, these results suggest that histone acetylation negatively regulates linker histone binding in S. cerevisiae and other organisms and provides important insight into how chromatin structure is regulated and maintained to both facilitate and repress transcription.                 iv LAY SUMMARY Nearly every cell in our body contains a copy of our genome. The genome is encoded by DNA which serves as a blue-print responsible for directing all cellular activities, including growth and proliferation. In order to allow for the packaging of the genome into the nucleus of each cell, DNA is wrapped around proteins referred to as histones. Additional proteins, including another family of histones known as linker histones, bind to the spooled DNA and facilitate compaction of DNA within the cell. Histone proteins can be modified by enzymes which can alter their chemical properties and impact DNA compaction. In this study, we investigated how the binding of linker histones is regulated, specifically examining the role of the histone modification acetylation in mediating this interaction.                   v PREFACE This thesis is based on the following first author publication: Lawrence, M. B. D., Coutin, N., Choi, J. K., Martin, B. J. E, Irwin, N. A. T., Young, B., Loewen, C. and Howe, L. J. (2017). Histone Acetylation, Not Stoichiometry, Regulates Linker Histone Binding in Saccharomyces cerevisiae. Genetics, 207(1), 347-355. Experiments were designed by Dr. LeAnn Howe, Dr. Jennifer Choi, Nicolas Coutin and Mackenzie Lawrence. Dr. Jennifer Choi conducted the SGA screen and Nicolas Coutin conducted the Hho1 ChIP-sequencing experiments. The subsequent data analysis and all other experiments were conducted by Mackenzie Lawrence.                     vi TABLE OF CONTENTS ABSTRACT ................................................................................................................................................................... iii LAY SUMMARY .......................................................................................................................................................... iv PREFACE ....................................................................................................................................................................... v TABLE OF CONTENTS ............................................................................................................................................ vi LIST OF TABLES ..................................................................................................................................................... viii LIST OF FIGURES ...................................................................................................................................................... ix LIST OF ABBREVIATIONS .......................................................................................................................................x  – INTRODUCTION ............................................................................................................................ 1 1.1 Chromatin .............................................................................................................................................. 1 1.2 Histone Post-Translational Modifications ............................................................................... 5 1.3 Histone Acetylation ........................................................................................................................... 6 1.4 Linker Histones ................................................................................................................................ 12 1.5 Summary of Research Hypotheses .......................................................................................... 21  – MATERIALS AND METHODS .................................................................................................. 24 2.1 Yeast Strains and Plasmids ......................................................................................................... 24 2.2 Quantitative Immunoblot Analysis .......................................................................................... 26 2.3 Chromatin Immunoprecipitation-Quantitative PCR Analysis ..................................... 27 2.4 Micrococcal Nuclease Digestion of Yeast Chromatin ....................................................... 28 2.5 Synthetic Dosage Lethality Screen ........................................................................................... 29 2.6 ChIP-Sequencing Analysis ........................................................................................................... 29 2.7 Data Availability ............................................................................................................................... 31  – RESULTS ......................................................................................................................................... 32        vii 3.1 Refining linker histone stoichiometry in S. cerevisiae .................................................... 32 3.2 Increased linker histone stoichiometry is toxic in S. cerevisiae ................................. 35 3.3 Histone acetylation negatively regulates linker histone binding in S. cerevisiae 40  – DISCUSSION .................................................................................................................................. 49 REFERENCES ............................................................................................................................................................ 54                          viii LIST OF TABLES Table 1.1: Histone lysine acetyltransferases in S. cerevisiae and known substrates .................. 8 Table 1.2: Histone lysine deacetylases in S. cerevisiae and known substrates ............................. 8 Table 2.1: Yeast strains used in this study .................................................................................................. 25 Table 2.2: Primers used to generate knock-out strains in this study .............................................. 25 Table 2.3: Primers used to verify knock-out strains in this study .................................................... 26 Table 2.4: Plasmids used in this study .......................................................................................................... 26 Table 2.5: Antibodies used in this study ...................................................................................................... 27 Table 2.6: ChIP-QPCR primers used in this study .................................................................................... 28                                  ix LIST OF FIGURES Figure 1.1: Structural comparison of the globular domains of human H1.0 and yeast Hho1 ....................................................................................................................................................................................... 14 Figure 3.1: Quantifying endogenous Hho1 levels in S. cerevisiae .................................................... 33 Figure 3.2: Refining linker histone stoichiometry in S. cerevisiae ................................................... 34 Figure 3.3: Increased linker histone stoichiometry is toxic in S. cerevisiae ................................ 37 Figure 3.4: Altered histone levels exacerbate toxicity due to increased linker histone stoichiometry in S. cerevisiae ........................................................................................................................... 39 Figure 3.5: Histone acetylation negatively regulates linker histone binding in S. cerevisiae42 Figure 3.6: Histone acetylation negatively correlates with linker histone binding in S. cerevisiae .................................................................................................................................................................. 45 Figure 3.7: Histone acetylation negatively correlates with linker histone binding in S. cerevisiae .................................................................................................................................................................. 47 Figure 3.8: Histone acetylation negatively correlates with linker histone binding in mouse embryonic stem cells ........................................................................................................................................... 48                    x LIST OF ABBREVIATIONS α   Alpha; signifies anti Δ   Delta; signifies deletion C   Degree Celsius ac   Acetyl  ATP   Adenosine triphosphate bp   base pair BSA   Bovine serum albumin  ChIP   Chromatin immunoprecipitation ChIP-QPCR  ChIP-quantitative PCR ChIP-seq   ChIP-sequencing  CTD   C-terminal domain C-terminal  Carboxy-terminal D. melanogaster Drosophila melanogaster DNA    Deoxyribonucleic acid  Esa1   Essential SAS2-related acetyltransferase 1 EDTA   Ethylenediaminetetraacetic acid  Gcn5   General control non-derepressible 5  GEO   Gene Expression Omnibus H2AS129  Histone H2A serine 129  H3K4   Histone H3 lysine 4 H3K9   Histone H3 lysine 9 H3K14  Histone H3 lysine 14 H3K18  Histone H3 lysine 18 H3K20  Histone H3 lysine 20 H3K23  Histone H3 lysine 23  H3K36  Histone H3 lysine 36  H3K56  Histone H3 lysine 56        xi H3K79  Histone H3 lysine 79 H3S10   Histone H3 serine 10 H4K5   Histone H4 lysine 5  H4K8   Histone H4 lysine 8  H4K12  Histone H4 lysine 12 H4K16  Histone H4 lysine 16 H4R3   Histone H4 arginine 3 HA   Hemagglutinin   HAT   Histone acetyltransferase HDAC   Histone deacetylase complex  Hho1   Histone H1   K   Lysine  me   Methyl MNase   Micrococcal nuclease NET-seq  Native elongation transcript-seq NCP   Nucleosome core particle   NFR   Nucleosome free region  NTD   N-terminal domain NuA3    Nucleosome acetyltransferase for histone H3  NuA4    Nucleosome acetyltransferase for histone H4 OD600   Optical density, at 600nm ORF   Open reading frame  PCR   Polymerase chain reaction  PHD   Plant homeodomain  PBS-T   Phosphate buffered saline with tween PHD   Plant homeodomain  PTM   Post-translational modification  Q   Glutamine  R   Arginine         xii rDNA   Ribosomal DNA RNA   Ribonucleic acid  RNAPII  RNA polymerase II Rpd3   Reduced potassium dependency 3 Rpd3L   Rpd3 large complex  Rpd3S   Rpd3 small complex rpm   Rotations per minute RSC   Remodels structure of chromatin  Rtt109  Regulator of Ty1 transposition 109 S. cerevisiae  Saccharomyces cerevisiae  SAGA   Spt-Ada-Gcn5-acetyltransferase   SAM   S-adenosyl methionine  SDS   Sodium dodecyl sulfate  SDS-PAGE  Sodium dodecyl sulfate-polyacrylamide gel electrophoresis SGA   Synthetic genetic array  STORM  Stochastic optical reconstruction microscopy SWI/SNF  Switching/sucrose non-fermenting  TE   Tris-EDTA TSS   Transcription start site  YEATS   Yaf9-ENL-AF9-Taf14-Sas5              1  – INTRODUCTION 1.1 Chromatin  DNA All living cells contain genetic information in the form of deoxyribonucleic acids (DNA), necessary for growth and proliferation. DNA molecules are long polymers composed of covalently linked nucleotides. Within cells, DNA does not usually exist as a single molecule, rather two strands of anti-parallel DNA form a double helical structure that can be copied, allowing for genome duplication. These DNA molecules contain distinct series of nucleotides called genes, which encode for proteins. The haploid human genome is composed of approximately 3.2 billion base pairs (bp) and encodes more than 20,000 protein-coding genes (Human Genome Sequencing Consortium 2004). In order to facilitate the compaction of DNA into the nucleus of the cell, DNA is packaged into chromatin.  Histones Chromatin is a dynamic structure composed of DNA, histones and non-histone proteins. The basic repeating unit of chromatin is the nucleosome core particle (NCP), which is composed of 147 bp of DNA wrapped 1.7 times around an octamer of core histones H2A, H2B, H3 and H4 (van Holde 1989; Luger et al. 1997). A fifth protein, termed the “linker histone”, binds the nucleosome dyad, interacting with the DNA entering and exiting the nucleosome (Syed et al. 2010; Meyer et al. 2011; Zhou et al. 2013; 2015b; Bednar et al. 2017). The binding of the linker histone wraps an additional 20 bp of DNA around the NCP,        2 completing the super-helical turns forming a structure known as the chromatosome (Allan et al. 1980; Thoma and Koller 1981; Zhou et al. 1998). Nucleosomal arrays are folded into higher-order chromatin fibres, compacting DNA further (Luger and Richmond 1998). In order to regulate gene expression and other DNA-templated processes, chromatin is dynamically and reversibly altered to allow access of the cellular machinery to the underlying DNA.  The core histones are highly conserved, basic proteins that comprise the major architectural proteins of chromatin. They contain three distinct structural domains, a long, hydrophilic N-terminal tail, a globular hydrophobic core and a short, hydrophobic C-terminus (Arents and Moudrianakis 1995; Luger and Richmond 1998). The bulk of the histone protein mass is comprised of the globular core, which contains the highly conserved structural motif, termed “the histone fold” (Arents and Moudrianakis 1995). The histone fold, which consists of 3 -helices separated by 2 loops (Luger et al. 1997), forms an extensive protein-protein interface important for histone heterodimerization (Arents et al. 1991; Arents and Moudrianakis 1993). The N-terminal tails of the core histones protrude from the surface of the nucleosome and are targets for post-translational modifications (PTMs) (Luger et al. 1997; Morales and Richard-Foy 2000). H2A is the only canonical core histone with a flexible C-terminal tail. The C-terminal tail of H2A protrudes from the NCP at the dyad axis (Luger et al. 1997) and is primarily responsible for the interactions at the interface between the H3-H4 tetramer and H2A-H2B dimers (Suto et al. 2000).        3  The core histones are synthesized primarily during S phase of the cell cycle, and incorporated into chromatin in a replication-dependent manner (Spalding et al. 1966; Borun et al. 1967). In humans this includes H2A, H2B, H3.1, H3.2 and H4, while the budding yeast, Saccharomyces cerevisiae (S. cerevisiae) has a single canonical H3, in addition to H2A, H2B and H4. Multiple copies of the core histone genes are located in several histone gene clusters in the mammalian and yeast genomes (Marzluff et al. 2002; Eriksson et al. 2012). In addition to the core histones that form the canonical nucleosome, non-allelic histone variants also exist. Histone variants are synthesized throughout the cell cycle and are incorporated into chromatin in a replication-independent manner. In most cases, only single copies of variant genes are found throughout an organisms’ genome (Marzluff et al. 2002; Eriksson et al. 2012). In humans, 13 core histone variants, including 2 testis-specific variants have been identified (for reviews see: Kamakaka and Biggins 2005; Sarma and Reinberg 2005; Talbert and Henikoff 2010; Maze et al. 2014). However, S. cerevisiae only contains two core histone variants, the H2A variant HTZ1 and the centromeric H3 variant, CSE4.   Higher order packing and DNA accessibility    Different levels of chromatin compaction exist within the nucleus. Highly compact chromatin is often referred to as heterochromatin. Chromosomal regions that contain a high density of repetitive DNA elements, including clusters of satellite sequences and transposable elements, are targets for heterochromatin formation (Lohe et al. 1993). In        4 many organisms, heterochromatin is found in the telomeres and in the domains surrounding centromeres of chromosomes, referred to as centromeric heterochromatin. These regions often remain condensed throughout the cell cycle. In contrast, euchromatin is a lightly packed form of chromatin that encompasses all non-heterochromatic regions of the genome. Euchromatin decondenses during interphase of the cell cycle and contains the majority of genes that encode for cellular proteins. Unlike mammals, the majority of the genome of the budding yeast, S. cerevisiae is composed of euchromatin. However, several regions of the genome are silenced or can be silenced, including the two silent mating-type loci, HMR and HML, telomeres and ribosomal RNA-encoding DNA (rDNA) (Weiss and Simpson 1998; Ravindra et al. 1999; Rusche et al. 2003). The formation of chromatin facilitates the packaging of DNA into the nucleus of cells, however, it also represents a significant physical barrier to transcription, DNA replication and repair.   Several mechanisms exist to regulate chromatin structure within the cell. Firstly, sliding and restructuring of nucleosomes is accomplished by chromatin remodelers, large multi-subunit, ATP-dependent enzymes. Secondly, the post-translational modification (PTM) of DNAs and histones can alter chromatin dynamics. Lastly, the incorporation of non-canonical histone variants can also modify chromatin structure by altering nucleosome stability and existing patterns of histone PTMs. These mechanisms allow for regulation of chromatin condensation and DNA accessibility.           5 1.2 Histone Post-Translational Modifications The post-translation modification (PTM) of histone proteins is one mechanism by which chromatin structure can be dynamically regulated. Histone PTMs can directly alter chromatin structure by disrupting histone-DNA contacts, or indirectly by modulating the recruitment and binding of chromatin modifying factors (for review see: Bannister and Kouzarides 2011). The majority of histone PTMs occur on the highly conserved histone N-terminal tails which protrude away from the nucleosome core (Luger et al. 1997; Strahl and Allis 2000). To date, a wide variety of histone modifications and sites have been identified including lysine acetylation, lysine and arginine methylation, serine phosphorylation and lysine ubiquitylation (Murray 1964; Allfrey et al. 1964; Stevely and Stocken 1966; Hunt and Dayhoff 1977; Zhao and Garcia 2015). Numerous studies have shown that histone PTMs are correlated with diverse cellular processes, including gene activation (Cheung et al., 2000), heterochromatic gene silencing (Rea et al., 2000), DNA replication (Vogelauer et al., 2002) and DNA damage responses (van Attikum and Gasser, 2005). However, despite the potential for combinatorial complexity of histone modifications, only a small number of states are actually observed in vivo (Liu et al. 2005a; Wang et al. 2008). Understanding how histone PTM mediate chromatin structure and dictate biological outcomes remains a major of focus in the field.           6 1.3 Histone Acetylation  Histone acetylation is one of the most widely studied histone PTMs. It was first described in 1963 and was subsequently shown to be important for regulating ribonucleic acid (RNA) expression in vitro (Phillips 1963; Allfrey et al. 1964). This led to the proposal that histone acetylation was a dynamic and reversible modification, important for regulating gene expression. Indeed, various studies have demonstrated a clear correlation between histone acetylation and gene expression in vivo (Gershey et al. 1968; Hebbes et al. 1988). Since its initial discovery, numerous acetylation sites have been identified and a tremendous amount of work has been done to elucidate the regulation and functional consequences of histone acetylation.   HATs and HDACs  Histone acetylation is maintained by the competing activities of two groups of enzymes. Histone acetyltransferases (HATs) catalyze the addition of a single acetyl moiety to the ε-amino group of lysine residues (DeLange et al. 1969; Millar and Grunstein 2006) and histone deacetylases (HDACs) catalyze their removal (Taunton et al. 1996). Acetylation occurs primarily on histone N-terminal tails, however, the globular domains and C-terminal domain of H2A can also be acetylated (Zhang et al. 2003; Valero et al. 2016). Numerous HATs have been identified, many of which are components of large multi-subunit complexes that contain auxiliary domains important for enzymatic activity and targeting (Table 1.1) (Struhl 1998). Although many recombinant HAT proteins can acetylate free histones, often nucleosomal acetylation occurs only in the context of the in vivo HAT        7 complexes. Moreover, incorporation of HATs into complexes often alters substrate specificity. For example, Gcn5 which was first identified in yeast as a transcriptional co-activator (Georgakopoulos and Thireos 1992) and later shown to have HAT activity (Brownell et al. 1996), is a component of the Ada, SLIK/SALSA, HAT-A2 and SAGA complexes (Grant et al. 1997; Sendra et al. 2000; Pray-Grant et al. 2002). While recombinant Gcn5 preferentially acetylates histone H3K14, it can also acetylate residues on histone H4 (Kuo et al. 1996). However, within the Ada and SAGA complexes, Gcn5 acetylates nucleosomal H3 and H2B, but not histone H4 (Grant et al. 1997; 1999; Suka et al. 2001; Jiang et al. 2007). Similarly, Sas3 alone acetylates free histones, while the Sas3-containing NuA3 HAT complex acetylates nucleosomal histone H3 at some of the same residues as the Gcn5-containing HATs, in addition to distinct H3 targets (Takechi and Nakayama 1999; John et al. 2000; Howe et al. 2001). Like HATs, many HDACs are also catalytic subunits of multi-protein complexes (Table 1.2). Some of these complexes have broad and overlapping substrate specificities, while others act at specific lysine residues. For example, Rpd3 can exist in both the Rpd3S and RpdL complexes, which together deacetylate nearly all sites on histones H2A, H2B, H3 and H4 (Rundlett et al. 1996; Zhou et al. 2009). In contrast, Hda1 deacetylates lysine residues on histones H2B and H3 (Wu et al. 2001). Together, HATs and HDACs reversibly modify histones and play a role in mediating chromatein dynamics.             8 Table 1.1: Histone lysine acetyltransferases in S. cerevisiae and known substrates   Catalytic Enzyme    Complex   Substrate   Gcn5  SAGA  H2B, H3 tails Gcn5 SLIK H2B, H3 tails Gcn5 ADA H3 tails Gcn5 HAT-A2 H3 tails Hat1 HATB H2A, H4 tails Esa1 NuA4 H2A, H4 tails Esa1 Pic NuA4 H2A, H4 tails Sas2 SAS H4 tails Sas3 NuA3 H3 tails Rtt109 Rtt109-Vps75 H3(K56), tails       Table 1.2: Histone lysine deacetylases in S. cerevisiae and known substrates  Catalytic Enzyme    Substrate   Hda1  H2B, H3 tails Hos1 H2B tails Hos2 H4 tails Hos3 H2B, H4 tails Hst1 H3 tails Hst3 H3(K56) Hst4 H3(K56) Rpd3 H2A, H2B, H3, H4 tails   Sir2 H3, H4 tails          9 Targeting  Histone acetylation is typically associated with actively transcribed genes. Acetylation occurs primarily at promoter regions, however, some acetylation marks are localized throughout the bodies of transcribed genes (Pokholok et al. 2005; Liu et al. 2005b; Weiner et al. 2015). Promoter-targeted acetylation plays a role in transcription initiation. In yeast, this is accomplished by the interaction of the HAT complexes, SAGA (Gcn5) and NuA4 (Esa1) with transcriptional activators, which promotes the recruitment of the basal transcription machinery required for transcription initiation (Bhaumik and Green 2002). Acetylation is also targeted to gene bodies by several different mechanisms. Firstly, various HATs are localized to transcribed regions through interactions with the phosphorylated CTD of RNAPII, including NuA4 (Esa1) (Ginsburg et al. 2009) and SAGA (Gcn5) (Govind et al. 2007) in yeast and PCAF in humans (Obrdlik et al. 2008). HATs are also recruited to gene bodies through H3K4 and K36 methylation. The histone methyltransferases responsible for modifying these histone residues associate with elongating RNAPII, creating a trail of methylation which can serve as docking sites for HATs (Shi et al. 2006; Martin et al. 2006; Shi et al. 2007). Lastly, acetylated histones can also be incorporated into chromatin at promoter and transcribed regions via histone turnover (Dion et al. 2007; Rufiange et al. 2007).  Histone hypoacetylation is often associated with repressed chromatin and inactive genes. In yeast, at least 9 HDACs have been identified, which preferentially deacetylate particular sets of genes and regions of the genome (Table 1.2) (Robert et al. 2004; Li et al. 2013).        10 Several of these HDACs are recruited to promoter regions, where they are thought to play a role in repressing transcription. For example, Rpd3L is targeted to promoter regions via the transcriptional repressor, Ume6 (Kadosh and Struhl 1997; Rundlett et al. 1998). Another way that HDACs are thought to be targeted to promoter regions is via association with the Tup1-Ssn6 co-repressor complex, which interacts with sequence specific transcriptional repressors (Davie et al. 2003; Malavé and Dent 2006). In addition to preferentially deacetylating promoters, HDACs also function throughout gene bodies. For example, Rpd3S is targeted to actively transcribed genes through association with di- and tri-methylated H3K36, resulting in deacetylation from the mid to 3’ end of genes (Carrozza et al. 2005; Joshi and Struhl 2005; Li et al. 2009; Drouin et al. 2010). This form of HDAC targeting is thought to restore chromatin structure in the wake of elongating RNAPII, preventing cryptic transcription along gene bodies (Carrozza et al. 2005; Li et al. 2009).   Impact on chromatin structure Histone acetylation is thought to contribute to the opening of chromatin structure through a variety of mechanisms. For example, histone acetylation neutralizes the positively charged lysine residues in the N-terminal tails, reducing their affinity for the negatively charged phosphate backbone of DNA (Hong et al. 1993; Lee et al. 1993). Although acetylated nucleosomes do not undergo large structural changes, they exhibit decreased thermal and salt stability and increased unwrapping of the outer-turn of nucleosomal DNA, particularly the linker DNA (Ausió and Van Holde 1986; Li et al. 1993; Gansen et al. 2009; Wakamori et al. 2015). Numerous studies have also shown that histone acetylation results        11 in the decompaction of higher-order chromatin fibres, with H2B and H4 acetylation playing the largest role in unfolding (Norton et al. 1989; Tse et al. 1998; Shogren-Knaak 2006; Wang and Hayes 2008; Robinson et al. 2008; Allahverdi et al. 2011; Wakamori et al. 2015). Thus, histone acetylation can directly regulate both the structure of individual nucleosomes and chromatin fibers, increasing the accessibility of regulatory proteins to chromatin templates.  Histone acetylation can also regulate chromatin structure indirectly by mediating the recruitment and binding of chromatin modifying factors. For example, several families of protein domains have been identified with specificity for acetylated lysine residues, including bromodomains (Dhalluin et al. 1999), YEATS (Le Masson et al. 2003) (Li et al. 2014), double PHD (Zeng et al. 2010) and double PH domains (Su et al. 2012). The most well-characterized of these are the bromodomains, which are found in a wide range of nuclear proteins, including many HATs, transcriptional regulators and chromatin remodelers (Dhalluin et al. 1999; Filippakopoulos et al. 2012). For example, the Gcn5 bromodomain can bind to various acetyl-lysine residues on histone H3 and H4 and is thought to be important for stimulating HAT activity (Owen 2000; Hudson et al. 2000; Cieniewicz et al. 2014). Many chromatin remodelers, including RSC and SWI/SNF in yeast contain numerous bromodomains that have been shown to promote efficient anchoring and remodeling activity of acetylated nucleosomes (Kasten et al. 2004; Hassan et al. 2007). Lastly, histone acetylation can also block the activity of various chromatin-associated proteins. For example, H3K14ac has been shown to negatively regulate the activity of the        12 chromatin remodeler ACF (Shogren-Knaak et al. 2006) and also block the activity of the H3K4 demethylase, Jhd2 (Maltby et al. 2012a). Therefore, histone acetylation can indirectly regulate chromatin structure and transcription through interactions with numerous chromatin modifying proteins.    1.4 Linker Histones   Metazoan linker histones  Linker histones belong to a distinct histone protein family and are present in most eukaryotic cells. Unlike the highly conserved core histones, linker histones exhibit much greater sequence variability. Despite this, metazoan linker histones share a three-domain structure consisting of a short N-terminal tail, a central globular domain and a long, basic C-terminal tail (Allan et al. 1980). The globular domain has a winged-helix fold structure that is conserved from yeast to humans (Ramakrishnan et al. 1993) and is necessary for generating the 168 bp chromatosome particle, resulting from micrococcal nuclease (MNase) digestion of native chromatin (Figure 1.1) (Singer and Singer 1976; Allan et al. 1980; Patterton et al. 1998; Zhou et al. 2013). The N- and C-terminal domains are less conserved and account for the majority of sequence heterogeneity between the H1 subtypes (Ponte 2003). No significant role in chromatin binding and condensation has been identified for the N-terminal domain (NTD), however, it has been proposed that the NTD may assist in targeting and anchoring the globular domain to the nucleosome (Allan et al. 1980; 1986). Further, extensive PTM sites on the NTD suggest that it serves as a binding        13 platform for other proteins (Wiśniewski et al. 2007; Review: Godde and Ura 2008). The highly basic CTD is intrinsically disordered in aqueous solution, however, in the presence of tetrahedral anions and DNA it adopts considerable α-helical character (Clark et al. 1988; Ali et al. 2004a). Concomitant with the ability of the CTD to bind negatively charged species, its primary role is to stabilize the folding of nucleosomal arrays into chromatin fibers via neutralizing the charge of the linker DNA (Allan et al. 1986; Clark and Kimura 1990; Carruthers et al. 1998; Misteli et al. 2000; Hendzel et al. 2004). Further, the CTD is required for higher-affinity binding of H1 to chromatin (Zhou et al. 2013). This three-domain structure is found in complex eukaryotes, however, some protists have a linker histone that more closely resemble only the C-terminal tail of the H1 linker histones (for review see: Kasinsky et al. 2001).            14   Figure 1.1: Structural comparison of the globular domains of human H1.0 and yeast Hho1 Phyre2 was used to model human H1.0 (NP_005309) (blue) and the average NMR ensemble of the first globular domain was used to for Hho1 (PDB: 1UST) (red). Structures were superimposed using the MatchMaker tool in Chimera (Ali et al. 2004b; Pettersen et al. 2004; Kelley et al. 2015).   Linker histones form a large and diverse family of proteins. In mice and humans, 11 linker histone subtypes have been identified, including seven somatic subtypes (H1.1-H1.5, H1.10 and H1.0), three testis-specific subtypes (H1.6, H1.7 and H1.9) and one oocyte-specific subtype (H1.8) (Happel and Doenecke, 2009). Of the somatic linker histones, H1.1 to H1.5 are expressed in a replication-dependent manner, while H1.10 and H1.0 are replication-independent and can be expressed in non-proliferating cells (Happel et al. 2009). H1.1 to H1.5 and H1.10 are ubiquitously expressed, while H1.0 accumulates in terminally differentiated cells (Zlatanova and Doenecke 1994; Happel et al. 2009). In accordance with this expression pattern, H1.0 in amphibians and birds (referred to as H5) is associated with        15 highly condensed, transcriptionally inert chromatin characteristic of terminally differentiated cells, such as nucleated erythrocytes (Coles et al. 1987; Koutzamani et al. 2002). Lastly, Drosophila melanogaster expresses two linker histone subtypes, one somatic and one germline subtype, known as BigH1, named for its extended N-terminal tail (Lifton et al. 1978; Pérez-Montero et al. 2013). The presence of multiple histone-encoding genes in many metazoans has made studying the functional roles of linker histones in vivo challenging.   Yeast linker histone, Hho1 Although linker histones were first described in mammals in the early 1950s (Stedman and Stedman 1951), no bona fide linker histone had been identified in budding yeast, leading some researchers to suggest that yeast lacked a linker histone (Shen et al. 1995). However, in the 1990s shortly after the yeast genome was sequenced, a putative linker histone, termed Hho1 was discovered (Landsman 1996; Ushinsky et al. 1997; Patterton et al. 1998). The structure of Hho1 varies from that of metazoan linker histones. Hho1 contains a short NTD, globular domain and unlike metazoan linker histones, a second globular domain connected by a linker with similar sequence and composition as the C-terminal domain of canonical histone H1 (Landsman 1996; Ushinsky et al. 1997; Ali et al. 2004b). Studies have shown that both globular domains can fold into similar structures, however, despite sharing ~43% sequence identity, they have considerably different stabilities (Ono et al. 2003; Ali et al. 2004b). This lead researchers to suggest that Hho1 could simultaneously bind two nucleosomes at once (Schäfer et al. 2005). However, similar to the CTD of        16 canonical histone H1, the second globular domain of Hho1 is thought to be largely unstructured under physiological conditions and similarly adopts helical content in the presence of tetrahedral anions (Ono et al. 2003; Ali et al. 2004b). Additionally, Hho1 was shown to form a stable ternary complex with di-nucleosomes in vitro at similar ratios of linker histone-to-nucleosome found in more complex eukaryotes (Patterton et al. 1998). Moreover, while the first globular domain of Hho1 is sufficient for generating the 168 bp chromatosome upon MNase digestion (Patterton et al. 1998), the second globular domain is not (Ali and Thomas 2004). Lastly, another study investigating the role of Hho1 in transcriptional silencing found that mutant Hho1 lacking the second globular domain was able to recapitulate the phenotype of full length Hho1 under various conditions (Yu et al. 2009). Thus, Hho1 is thought to interact with nucleosomes in a similar fashion as metazoan linker histones.   Linker histone stoichiometry and nucleosome repeat length  One proposed mechanism for regulating linker histone binding is through the alteration of linker histone abundance. Linker histone abundance varies across species and between cell types. For example, vertebrate cells, which have many gene-poor regions, express approximately 1 molecule of linker histone for every nucleosome and this varies by cell type (Woodcock et al. 2006). Studies in various mouse tissues have determined ratios of 0.79 in splenocytes, 0.83 in thymocytes, 0.79 in adult hepatocytes, 0.76 in neonatal hepatocytes and 0.50 in embryonic stem (ES) cells (Fan et al. 2003; 2005b). Transcriptionally inert avian erythrocytes express 1.3 linker histones per nucleosome        17 (Bates and Thomas 1981; Coles et al. 1987). In S. cerevisiae, linker histone abundance is much lower than that observed in other eukaryotes, consistent with the gene-dense nature of this yeast’s genome. Downs et al. (2003) estimated linker histone stoichiometry in yeast to be one molecule for every 4 nucleosomes, while data from Freidkin and Katcoff (2001) suggested it is much lower, at one molecule of Hho1 for every 37 nucleosomes. Two high throughput analyses of protein expression in yeast have estimated that there are 2610 and 6560 molecules of Hho1 per haploid cell, representing ratios of 1:26 and 1:10 for the ~68,000 annotated nucleosomes in yeast (Ghaemmaghami et al. 2003; Brogaard et al. 2012; Kulak et al. 2014). Interestingly, there tends to be a linear relationship between the ratio of linker histone to nucleosome and the nucleosome repeat length (NRL) (Woodcock et al. 2006).  The NLR is the average distance between the centres of neighbouring nucleosomes. It is an important physical property of chromatin and depends on the length of the linker DNA. One of the primary roles of linker histones, which contain a high proportion of positively charged residues, is to neutralize the charge of DNA. Thus, when the amount of linker histones per nucleosome is reduced, charge homeostasis can be restored by decreasing nucleosome spacing (Woodcock et al. 2006). In eukaryotes, NRLs exhibit a wide variety of values ranging from 165 bp in S. cerevisiae, to 175–190 bp for most vertebrate cells and tissues (van Holde 1989; Woodcock et al. 2006). It is thought that the differences in NRL in yeast are due to the fact that yeast have sub-stoichiometric levels of linker histones compared to other eukaryotes. Indeed studies in mouse, where up to three somatic H1        18 subtypes were knocked out in different tissues resulting in a 50% reduction in H1 levels, showed a consistent decrease in NRL (Fan et al. 2003; 2005b). In accordance, increased linker histone stoichiometry should result in longer NRLs. This was shown by Gunjan et al. (1999) upon overexpression H1.0 and H1.3 in cultured mouse fibroblasts, resulting in a 1.2–1.4 fold increase in total H1 and an increase in NRL of ~15 bp. Moreover, the NRL of newly replicated chromatin in HeLa cells has an NRL of ~165 bp, similar to values reported for mouse embryonic stem (ES) cells and yeast (Smith et al. 1984; Woodcock et al. 2006).   Impact on chromatin Linker histones are important factors that mediate chromatin structure. The precise location of the linker histone within the chromatosome and how this contributes to higher-order folding are long-standing questions in the field. Linker histones bind to nucleosomes at the dyad, interacting with linker DNA at the entry and exit sites of the nucleosome (Syed et al. 2010; Meyer et al. 2011; Zhou et al. 2013; 2015b; Bednar et al. 2017). Binding occurs primarily through interactions of positively charged lysine and arginine residues in globular and C-terminal domains with nucleosomal and linker DNA, respectively (Allan et al. 1980; 1986; Duggan and Thomas 2000; Brown et al. 2006; Zhou et al. 2015b; Bednar et al. 2017). Various on- and off-dyad binding modes have been reported, and substitutions at key residues within the globular domain are thought to be important for determining the mode of binding of different linker histone subtypes (Zhou et al. 2015b; 2016). Moreover, different binding modes have been associated with distinct structural states in vitro, which may be related to specific chromatin functions (Song et al. 2014; Zhou et al. 2015b).        19 However, considerably less is known regarding the role of the N- and C-terminal domains of H1 within the chromatosome or how PTMs may affect the interaction and structures of these domains. Moreover, exactly how linker histones mediate higher-order chromatin folding in vivo remains unclear.  Early studies examining the role of linker histones in chromatin condensation showed that loss of linker histones resulted in morphological changes in chromatin fibres under different salt conditions (Thoma et al. 1979). Since then, linker histones have been shown to play an important role in the formation of the 30 nm fibre and higher-order structure in vitro (Finch and Klug 1976; Thoma et al. 1979; Robinson et al. 2006). However, the existence of the 30 nm fibre and its relevance in vivo is still debated (for reviews see: Robinson and Rhodes 2006; Maeshima et al. 2010). More recently, an in vivo study using stochastic optical reconstruction microscopy (STORM) to visualize chromatin structure was performed (Ricci et al. 2015). Chromatin fibres were visualized at the single-cell level in variety of cell types during interphase, with a resolution of ~20 nm. It was found that nucleosomes are arranged into discrete heterogeneous domains along the chromatin fibre, termed “nucleosome clutches” (Ricci et al. 2015). Interestingly, differentiated human fibroblast contained larger clutches compared to mouse ES cells. Additionally, larger clutches with higher nucleosome density and linker histone levels correlated well with heterochromatin markers (Ricci et al. 2015). These findings confirm the role of linker histones in chromatin condensation, while suggesting that in vivo 30 nm structures may only exist as short fragments, rather than long continuously folded fibres (Ricci et al. 2015).        20 Although there are numerous outstanding questions regarding the precise role of linker histones in chromatin organization, linker histones are thought to interact with both nucleosomes and chromatin to facilitate the formation of higher-order structures.   Impact on transcription  Regardless of the exact organization of folded H1-containing chromatin, because linker histones limit DNA accessibility and promote chromatin compaction, they are thought to be general repressors of transcription (Bustin et al. 2005; Happel and Doenecke 2009). Thus, regulating their interaction with chromatin may provide a means to control access of the transcriptional machinery to DNA. Indeed, genome-wide studies of various linker histone subtypes have shown a non-uniform distribution of linker histones across the genome, with a characteristic binding pattern in which H1 occupancy is reduced over promoter regions primarily at actively transcribed genes (Cao et al. 2013; Izzo et al. 2013; Nalabothula et al. 2014; Millán-Ariño et al. 2014). Additionally, different H1 subtypes also exhibit differential binding preferences in somatic cells and during cellular differentiation. For example, the distribution of H1.5 differs in human ES cells compared to differentiated fibroblasts, demonstrating a unique role for H1.5 in chromatin condensation in differentiated cells (Li et al. 2012). Cao et al. (2013) also show that H1.3 and H1.2 in mouse ES cells show very similar distributions, however, they also exhibit differences in binding specificities. Interestingly, altering linker histone levels in vivo results in both up and down regulation of specific genes. In yeast, deletion of non-essential Hho1 actually resulted in decreased expression at a small subset of genes (Hellauer et al. 2001). Similarly, in mouse        21 ES cells with H1 gene knock-downs resulting in a 50% reduction in total H1 levels, only a small subset of genes were up or down regulated compared to wild type cells (Fan et al. 2005a). Together, this suggests that specific linker histone subtypes may play a role in fine-tuning gene expression rather than acting as general repressors of transcription.     1.5 Summary of Research Hypotheses  Linker histones limit DNA accessibility and promote chromatin compaction, therefore, regulating their interaction with chromatin may provide a means to control access of the transcriptional machinery to DNA. One proposed mechanism for regulating linker histone binding is through the alteration of linker histone abundance. S. cerevisiae expresses one molecule of linker histone for every 4–37 nucleosomes (Freidkin and Katcoff 2001; Downs et al. 2003), consistent with the gene-dense nature of this yeast’s genome. In contrast, vertebrate cells, which have many gene-poor regions, express approximately 1 molecule of linker histone for every nucleosome and transcriptionally inert avian erythrocytes express 1.3 linker histones per nucleosome (Woodcock et al. 2006). However, several studies have demonstrated that linker histone occupancy is not uniform across the genome of a given cell type, suggesting that factors in addition to protein abundance regulate linker histone binding. For example, the yeast linker histone, Hho1, cross-links poorly to the first nucleosome relative to the transcription start site (TSS) and is instead enriched in regions with increased nucleosome spacing (Rhee et al. 2014; Ocampo et al. 2016). Further, transcriptionally active regions tend to be depleted in linker histones in multiple organisms        22 (Schafer et al. 2008; Cao et al. 2013; Izzo et al. 2013; Millán-Ariño et al. 2014; Ocampo et al. 2016).   One potential mechanism for regulating linker histone binding is via histone acetylation, which is enriched on active genes. Because linker histones bind to nucleosomes via contacts with DNA as it enters and exits the nucleosome, acetylation, which promotes DNA unwrapping, could disrupt Hho1 binding sites (Neumann et al. 2009; Syed et al. 2010; Simon et al. 2011; Meyer et al. 2011; Zhou et al. 2013; 2015a; Kim et al. 2015; Ikebe et al. 2016; Bednar et al. 2017). Indeed, others have observed increased linker histone mobility in cells treated with histone deacetylase inhibitors (Raghuram et al. 2010). Despite this evidence, however, the regulation of linker histone binding by core histone acetylation has not been thoroughly explored.   In this study, we designed a novel scheme to determine linker histone stoichiometry in yeast and investigated the impact of HHO1 overexpression on chromatin structure. Our data suggest that linker histone stoichiometry in yeast is one molecule of Hho1 for every 19 nucleosomes. Moreover, we show that increasing Hho1 levels results in a severe growth defect, despite only modestly impacting Hho1 occupancy or gross chromatin structure. Hho1 toxicity could be rescued by increased histone acetylation, consistent with the negative correlation between linker histone stoichiometry and histone acetylation in both yeast and mouse embryonic stem cells. Collectively these results suggest that factors in        23 addition to linker histone stoichiometry, including histone acetylation, dictate the impact that linker histones have on chromatin structure.                            24  – MATERIALS AND METHODS 2.1 Yeast Strains and Plasmids All strains used in this study are isogenic to S288C (Table 2.1). Yeast culture and genetic manipulations were carried out using standard protocols (Smith and Burke 2014). Genomic deletions and epitope-tag integrations were verified by PCR analysis (Table 2.2 and Table 2.3). The strains carrying the histone H3 tail mutants were derived by plasmid shuffle from FY2162 (Duina and Winston 2003). The plasmid pGAL1prHHO1 was generated by cloning the HHO1 open reading frame (ORF) into the BamHI and XhoI sites of pGAL1pr (Mumberg et al. 1994). The pHHT2prHHO1HA plasmid was created by, swapping the GAL1 promoter from pGAL1prHHO1 with a fragment containing 535 bp upstream of the HHT2 gene and adding a triple HA tag. All plasmids used in this study are listed in Table 2.4.                 25 Table 2.1: Yeast strains used in this study  Yeast Strain  Mating Type   Genotype  Y7092  Mat   can1Δ∷STE2pr-Sp-his5 lyp1Δ his3Δ1 leu2Δ0 met15Δ ura3Δ0 YLH101 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 YLH379 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 hho1::TRP YLH380 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 HHT2HA::HISMX6 YLH945 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 hda1:KAN YLH948 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 rpd3::KAN YLH950 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 rco1::KAN YLH972 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 HISMX6-HAHHO1 YLH224 Mat a his3Δ200 leu2Δ1 lys2-128δ ura3-52 trp1Δ63 (hht1-hhf1)::LEU2 (hht2-hhf2)::HIS3 Ty912Δ35::his4   Table 2.2: Primers used to generate knock-out strains in this study  Primer   Sequence (5’ to 3’)   HDA1 F1  ATGGATTCTGTAATGGTTAAGAAAGAAGTACTGGAAAATCCGGATCCCCGGGTTAATTAA HDA1 R1 TTTATTATTATTCAACTTTCATAAGGCATGAAGGTTGCCGGAATTCGAGCTCGTTTAAAC RPD3 F1 CATACAAAACATTCGTGGCTACAACTCGATATCCGTGCAGCGGATCCCCGGGTTAATTAA RPD3 R1 TCACATTATTTATATTCGTATATACTTCCAACTCTTTTTTGAATTCGAGCTCGTTTAAAC RCO1 F1 ATAAAAGACACTTCCATTACCATCTGCTAATAATAATACACGGATCCCCGGGTTAATTAA RCO1 R1 TTCACGTTCCTGATTTATTCTTTATGTATGTACGCCGTTTGAATTCGAGCTCGTTTAAAC           26 Table 2.3: Primers used to verify knock-out strains in this study  Primer   Sequence (5’ to 3’)   HDA1 s-220  TCCCGCACCGTTTGGAAATC HDA1 a+383 GAGGTTGCGGCTCTTACAG RPD3 s-181 GGCTTTCGGGAAGCAAAGTG RPD3 a+393 ATGCCCAATATTACGGCCCA RCO1 s-173 CTCGCTGCAAGATCTGCCTC RCO1 a+247  CCTTGGGTTCTTCTTCCGGG   Table 2.4: Plasmids used in this study  Plasmid   Description   p123 pRS416 p249 pGAL1.416 p964 pHHT2prHHO1HA.416 p658 pGAL1prHHO1.416 p311 pHHT2K9,14,18,23R p696  pHHT2K9,14,18,23Q   2.2 Quantitative Immunoblot Analysis  Whole cell extracts (Kushnirov 2000) were loaded onto 15% polyacrylamide gels and run at 150 V at room temperature for 1 hour. Following separation, gels were equilibrated in equilibration buffer (62.5 mM Tris pH 6.8, 2.3% SDS) at room temperature for 30 min with shaking. Proteins were transferred to nitrocellulose membranes in transfer buffer (17 mM glycine, 0.15% ethanolamine, 0.2% methanol) at 100 V at room temperature for 1 hour. Nitrocellulose membranes were blocked with 2% BSA (HyClone™) for 1 hr at room temperature with shaking. Following blocking, membranes were incubated in primary        27 antibodies (Table 2.5) with PBS-T overnight at 4C with shaking. Next, membranes were washed with PBS-T for 3 X 10 min at room temperature with shaking and incubated with secondary antibodies for 1 hr at room temperature with shaking, followed by fluorescence detection and quantification using the Licor Odyssey System.  Table 2.5: Antibodies used in this study  Antibody   Animal   Company   Catalogue Number   αH3   Rabbit  GeneScript  Raised against scH3 peptide (CKDILARRLRGERS)  αHho1  Rabbit Abcam ab71833 αHA Mouse Roche 11666606001  αHA  Rat  Roche  11867431001   2.3 Chromatin Immunoprecipitation-Quantitative PCR Analysis  Chromatin immunoprecipitation-quantitative PCR (ChIP-QPCR) analysis was performed as previously described (Martin et al. 2017). Cells were grown in 50 ml of synthetic drop-out media lacking uracil with galactose for 20 hr to an OD600 of ~0.8 and lysates were immunoprecipitated with 0.9 mg of α-Hho1 antibody (Table 2.5). QPCR was performed using the Applied Biosytems StepOnePlus Real-Time PCR System using the primers listed in Table 2.6.             28 Table 2.6: ChIP-QPCR primers used in this study  Primer   Sequence (5’ to 3’)   PMA1 a+85  CCTACGATGACGCTGCATC PMA1 s+232 CCTCTGGAACTGGTCTAGC SEC15 a+2230 GACCCATGAATTGTCTCGTCAAGG SEC15 s+2082 GTAAGGCAAGACCCGGATATCTC SEC15 a+370 GCACCATACCTTGGATGTTTGC SEC15 s+230 GGACCCCGTAATTGATGAATTGG LOS1 a+1395 CAGACTTGGGTCAATTACCACG LOS1 a+2940 GTCGTCATTATCCAAGCAGGTCC LOS1 a+230 CCATTTGGATTAGCGTTCACGC LOS1 s+69 CAAGCCATCGAGCTGCTAAATG PUT4 a+714 CACGCATAGAAAGATCGTGATCC PUT4 s+546  CTGGTCACTAGGTACGTTGAC   2.4 Micrococcal Nuclease Digestion of Yeast Chromatin  Cells were grown in synthetic drop-out media lacking uracil with dextrose until stationary phase, before being washed two times in synthetic drop-out media lacking uracil with galactose. Cells were then diluted in –uracil galactose media to an OD600 of 0.2 and grown for 20 hr at 30C. Following harvest, 25 ODs of cells were resuspended in 400 ml of 1 M sorbitol, 5 mM -mercaptoethanol and 10 mg/ml zymolyase, and incubated at 37C for 10 min. Spheroplasts were washed once in 1 M sorbitol and twice in spheroplast digestion buffer (SDB) (1 M sorbitol, 50 mM NaCl, 10 mM Tris pH 8, 5 mM MgCl2, 1 mM CaCl2, 1 mM -mercaptoethanol, 0.5 mM spermidine, 0.075% NP40) before being resuspended in 450 ml of SDB. Samples were digested with varying concentrations of micrococcal nuclease (MNase) for 2 min and digestions were stopped by addition of EDTA and SDS to final        29 concentrations of 5 mM and 1%, respectively. Crosslinks were reversed by overnight incubation at 65C and DNA was purified by digestion with proteinase K, phenol:chloroform:isoamyl extraction and ethanol precipitation. Samples were resuspended in 10 mM Tris pH 8, 1 mM EDTA and treated with RNase A prior to running on a 2% agarose gel. DNA was visualized using syto60 fluorescence detection with the Licor Odyssey System.  2.5 Synthetic Dosage Lethality Screen  The synthetic genetic array (SGA) starting strain Y7092 (MAT can1Δ∷STE2pr-Sp-his5 lyp1Δ his3Δ1 leu2Δ0 met15Δ ura3Δ0) was transformed with pGAL1prHHO1. The resulting query strain was mated to the MATa deletion mutant array. SGA methodology, previously described for a plasmid-based synthetic dosage resistance screen (Chruscicki et al. 2010), was performed in triplicate with the following modifications: (1) medium lacking uracil was used to maintain the plasmid and (2) hits were scored against strains containing pGAL1prHHO1 grown on dextrose using the Balony program (Young and Loewen 2013). Hits were verified using PCR confirmation of the deletion strain, followed by transformation and dilution plating on appropriate media.   2.6 ChIP-Sequencing Analysis  ChIP-sequencing (ChIP-seq) was performed as previously described with a few alterations (Maltby et al. 2012b; a). Briefly, cells were grown in 1 liter of yeast, peptone, dextrose        30 media to mid-log phase and crosslinked with 1% formaldehyde for 15 min at 30C. The cross-linking reaction was stopped with 125 mM glycine and cells were washed twice with cold PBS. Cells were resuspended in lysis buffer [50 mM HEPES-KOH (pH 7.5), 140 mM NaCl, 1mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate], flash frozen in liquid nitrogen and ground in a coffee grinder with dry ice for 10 X 3 min. Samples were thawed, normalized by protein content and sonicated (Diagenode Biorupter, high output for 30 X 30 sec on/off) to obtain an average DNA fragment length of 200–400 bp. The lysate was cleared at 10,000 rpm for 10 min and the supernatant was retained for the whole cell extract. Magnetic Protein-G Dynabeads were added and incubated with the whole cell extract for 1 hr and then removed. Antibodies were added (15.0 ml of the -Hho1 antibody, Table 2.5) and incubated with the whole cell extract overnight. Magnetic Protein-G Dynabeads were added and incubated with the sample for 30 min. After reversal of crosslinking and DNA purification, the library construction protocol was performed as described (Maltby et al. 2012a). Equimolar amounts of indexed, amplified libraries were pooled, adapter dimers were removed by gel purification and paired-end 100 nucleotide reads were generated using v3 sequencing reagents on the HiSeq2000 (SBS) platform. Reads were aligned to the Saccer3 genome using Bowtie2 (Langmead and Salzberg 2012) and plot2DO (https://github.com/rchereji/plot2DO) (Chereji et al. 2017), deepTools (Ramírez et al. 2014; 2016) and the JavaGenomicsToolkit (http://palpant.us/java-genomics-toolkit/) were used for all subsequent analysis as indicated. Additional data used were sourced from https://www.ncbi.nlm.nih.gov/geo/ including GSE61888 (ChIP-seq of histone post-translational modifications in S. cerevisiae), GSE38384 (RNAPII ChIP-seq in S.        31 cerevisiae), GSE46134 (ChIP-seq of linker histones in mouse embryonic stem cells) and GSE29218 (ChIP-seq of H3K9ac, H3K27ac and RNAPII in mouse embryonic stem cells).  2.7 Data Availability  The ChIP-seq data generated for this study have been deposited in the Gene Expression Omnibus (GEO) database (www.ncbi.nlm.nih.gov/geo/), GEO accession no. GSE100591.                       32  – RESULTS 3.1 Refining linker histone stoichiometry in S. cerevisiae  Most eukaryotic chromatin contains approximately one molecule of linker histone for every nucleosome. In S. cerevisiae, linker histone stoichiometry is greatly reduced, but attempts to quantify the ratio of linker histone to nucleosomes have led to conflicting results. Previous studies made use of carboxyl-terminal epitope tags to quantify Hho1 levels but, surprisingly, we found that addition of a carboxyl-terminal HA tag to the endogenous HHO1 gene reduced Hho1 abundance approximately five-fold (Figure 3.1A and B). One explanation for this effect is the native HHO1 3’-UTR, which is replaced when carboxyl-terminal epitope tagging, is required for mRNA stability or protein translation. To circumvent this problem, we sought to quantify the abundance of Hho1 expressed from an unaltered HHO1 locus relative to a core histone. To this end, we generated a yeast strain expressing two copies of HHO1 (Figure 3.2A). The first copy was the endogenous, chromosomal HHO1 locus, which was unaltered (shown in yellow). The second copy was the HHO1 ORF (yellow) fused to the histone H3 promoter (HHT2pr, shown in red) with a carboxyl-terminal HA tag (blue) on a low-copy plasmid. The yeast strain also included an identical HA tag on one (HHT2) of the two copies of the histone H3 gene. By immunoblotting whole cell extracts from this strain for Hho1, HA and H3, we could directly compare signals generated with the Hho1 and H3 antibodies, using the identical HA tags on Hho1 and H3 (Figure 3.2B). Using this approach, with three biological replicates, we        33 calculated the linker histone stoichiometry in yeast to be one molecule of linker histone generated from the endogenous HHO1 locus for every 18.9 ± 1.0 nucleosomes.     Figure 3.1: Quantifying endogenous Hho1 levels in S. cerevisiae (A) Immunoblot analysis of Hho1 levels in whole cell extracts from wild type cells (HHO1) and cells with an HA-tag integrated at the carboxyl terminus of the endogenous HHO1 gene (HHO1HA). Histone H3 was used as a loading control. A band cross-reacting with the αHho1 antibody is indicated with an asterisk. (B) Quantification of Hho1 levels determined from immunoblot of Hho1 normalized to histone H3. Bars indicate the standard error of the mean from three biological replicates.          34   Figure 3.2: Refining linker histone stoichiometry in S. cerevisiae (A) Schematic representation of the genes encoding histones Hho1 and H3 in an engineered strain of S. cerevisiae. Elements from the HHO1 locus are shown in yellow, the HHT1 (Histone H Three 1) locus are shown in white and HHT2 (Histone H Three 2) locus are shown in red. The position of triple HA tags on Hho1 and Hht2 are shown in blue. pr, promoter. (B) Representative immunoblot of whole cell extracts from the strain described in A (+) as well as an isogenic strain lacking the pHHT2prHHO1 plasmid (2). Quantification of Hho1HA with αHA and αHho1 antibodies and H3 with αHA and αH3 antibodies facilitated determination of the relative ratio        35 of Hho1 to histone H3. Bands cross-reacting with the αHho1 antibody are indicated with asterisks.   3.2 Increased linker histone stoichiometry is toxic in S. cerevisiae  Linker histones are thought to negatively regulate transcription and thus the reduced linker histone stoichiometry in S. cerevisiae is consistent with the gene-dense nature of the yeast genome. To determine the impact of increased linker histone dosage on growth and chromatin structure of S. cerevisiae, we fused the HHO1 ORF to a GAL1 promoter (GAL1pr) on a low-copy vector and transformed this plasmid into wild-type yeast. Expression of Hho1 from GAL1pr resulted in a severe growth defect (Figure 3.3A), despite increasing total Hho1 abundance only threefold relative to yeast with vector alone (Figure 3.3A, B and C).   To confirm that excess Hho1 is incorporated into chromatin, we performed chromatin immunoprecipitation at multiple loci, including the 5’ end of a highly expressed gene (PMA1), the 5’ and 3’ ends of two moderately expressed genes (LOS1 and SEC15) and the middle of PUT4, an inactive gene. These results, shown in Figure 3.3D, demonstrate that HHO1 overexpression resulted in statistically significant increases (p-value for student’s t-test, < 0.05) in Hho1 occupancy at all loci tested, with the exception of PUT4. Importantly, in no case was the increase in Hho1 occupancy proportional to the over threefold increase in total Hho1 abundance observed in Figure 3.3B and C, suggesting that in yeast, linker histone binding is not strictly dictated by Hho1 levels. Interestingly, the greatest increases        36 in Hho1 binding upon HHO1 overexpression were observed on the 5’ ends of genes. Previous work has shown that Hho1 is enriched in regions with increased nucleosome spacing (Ocampo et al. 2016). To determine whether the differential incorporation of excess Hho1 at 5’ relative to 3’ genic regions was due to increased nucleosome spacing, we calculated the average spacing of nucleosomes relative to all annotated TSSs in yeast (Weiner et al. 2015). Figure 3.3E shows that average nucleosome spacing between the +1 and +2, +2 and +3, and +3 and +4 nucleosomes is 167, 165, and 164 bp, respectively. In contrast, average nucleosome spacing between the +4 and +5, and +5 and +6 nucleosomes is 161 bp, suggesting that these regions are refractory toward Hho1 binding because they lack sufficient linker DNA.   To determine whether HHO1 overexpression was associated with major changes in chromatin structure, we analyzed the effect of increased Hho1 on the sensitivity of yeast chromatin to micrococcal nuclease (MNase). Figure 3.3, F and G show that increased Hho1 levels had little effect on the length of fragments generated by MNase digestion. However, HHO1 overexpression consistently resulted in loss of fragments larger than four nucleosomes and the generation of a high molecular weight, MNase-resistant DNA band (highlighted with an asterisk in Figure 3.3F), which may suggest that increased levels of Hho1 promotes the formation of nuclease-resistant domains at the expense of longer MNase-sensitive, nucleosome arrays. Collectively, these results demonstrate that overexpression of Hho1 results in modest increases in Hho1 occupancy at the 5’ regions of genes, localized changes in chromatin structure and impaired cell growth.         37   Figure 3.3: Increased linker histone stoichiometry is toxic in S. cerevisiae  (A) Ten-fold serial dilutions of wild-type yeast containing vector alone (pGAL1pr) or expressing HHO1 from a GAL1 promoter (pGAL1prHHO1) were plated on uracil drop-out media with dextrose or galactose and grown at 30oC for 3 days. (B) Representative immunoblot for Hho1 levels in extracts from cells with vector alone (pGAL1pr) or expressing HHO1 from a GAL1 promoter (pGAL1prHHO1) grown for 20 hr in uracil drop-out media with galactose. (C) Quantification of Hho1 levels determined from immunoblot of Hho1 in three biological replicates. Error bars indicate the SE of the mean. (D) ChIP-QPCR for galactose-induced Hho1 at the indicated loci. Cells containing vector alone were set to 1. Error bars indicate SE of the mean of six biological replicates. (E) Box plot of base pair distance between nucleosome positions (Weiner et al. 2012) relative to the transcriptional start site. Notches indicate the 95%        38 confidence interval for the median. (F) Chromatin from cells containing vector alone (pGAL1pr) or expressing HHO1 from a GAL1 promoter (pGAL1prHHO1) grown for 20 hr in uracil drop-out media with galactose was digested with increasing concentrations of MNase. The DNA was purified and resolved on an agarose gel. (G) Plot of DNA fragment sizes (from F) vs. the number of nucleosomes with cells containing vector alone (blue) or expressing HHO1 from a GAL1 promoter (red). The indicated nucleosome repeat lengths were determined from the slope of the lines.   The specific incorporation of excess Hho1 at the 5’ regions of genes was surprising, considering that previously published work suggests that Hho1 fails to cross-link to the 5’ linker DNA of +1 nucleosomes (Rhee et al. 2014). This inconsistency may suggest that, when expressed from its endogenous promoter, Hho1 binding is under some form of regulation. To identify proteins or genetic pathways involved in regulating the interaction of Hho1 with chromatin, we used SGA technology to overexpress Hho1 in the ~4700 non-essential yeast deletion mutants. A major class of genes identified in the screen was those that regulate core histone gene dosage (Figure 3.4) (Kurat et al. 2014). Interestingly, decreased growth due to HHO1 overexpression was observed in mutants predicted to have both decreased and increased histone levels (chi-squared test, P = 0.00263 for a random distribution). The sensitivity of cells with increased core histone levels to Hho1 overexpression was not surprising, as the combination likely interferes with processes that use DNA as a template. In contrast, the enhanced Hho1 toxicity of cells with reduced core histone levels may be due to increased nucleosome spacing, which creates additional binding sites for Hho1, interfering with DNA access. Indeed, spt10Δ, one of the most sensitive strains to HHO1 overexpression, has been shown to exhibit increased nucleosomes spacing (van Bakel et al. 2013).         39    Figure 3.4: Altered histone levels exacerbate toxicity due to increased linker histone stoichiometry in S. cerevisiae Relative growth of ~4700 non-essential deletion strains expressing Hho1 from a GAL1 promoter. Mutants that are predicted to have decreased and increased histone levels are shown in blue and red, respectively (Kurat et al. 2014). Values above 1.0 reflect improved growth and values below 1.0 reflect decreased growth. Dashed lines denote default thresholds for rescues and lethality.             40 3.3 Histone acetylation negatively regulates linker histone binding in S. cerevisiae  A second class of genes identified in our synthetic dosage screen was those encoding HDACs (Figure 3.5A). However, contrary to mutants with altered histone dosage, mutation of HDACs rescued Hho1 toxicity. To verify that loss of an HDAC could rescue growth of cells with excess Hho1, we created an hda1Δ mutant in our laboratory strain background and confirmed resistance to Hho1 overexpression by dilution plating (Figure 3.5B). We also confirmed that rescue of growth in an hda1Δ mutant was not due to a GAL1 transcription defect by quantitative immunoblot (Figure 3.5C). To verify that the impact of HDAC loss is due to loss of deacetylation of core histones, we mutated acetylation sites in the tail of histone H3 to arginine and glutamine to mimic unacetylated and acetylated lysine residues, respectively. In strains overexpressing Hho1, glutamine substitutions in the H3 tail conferred a growth advantage (Figure 3.5D), which was not due to altered Hho1 levels (Figure 3.5E). Collectively, these results suggest that histone acetylation negatively regulates the binding of Hho1 to chromatin.         41             42 Figure 3.5: Histone acetylation negatively regulates linker histone binding in S. cerevisiae Relative growth of ~4700 non-essential deletion strains expressing Hho1 from a GAL1 promoter. Mutants with gene deletions of RPD3, HDA1, HOS1, HOS2, HOS3, HOS4, HST1, HST2, HST3 and HST4 are shown in red. (B and D) Ten-fold serial dilutions of the indicated strains carrying either vector alone (pGAL1pr) or a plasmid expressing HHO1 from a GAL1 promoter (pGAL1prHHO1) were grown on uracil drop-out media, with either dextrose or galactose as indicated, at 30oC for 4 days. (C and E) Immunoblot quantification of Hho1 levels in the indicated strains expressing HHO1 from a GAL1 promoter after growth for 20 hr in uracil drop-out media with galactose. Error bars indicate the SE of the mean from three biological replicates.  To further investigate the role of histone acetylation in regulating Hho1 binding, we performed ChIP-seq analysis of Hho1 expressed from its native promoter. To visualize the data generated, we used 2D occupancy plots, which simultaneously display DNA sequencing data as: (1) the relative sequence read abundance (heatmap), (2) sequence fragment length (y-axis) and (3) position of sequence reads relative to the dyad of the +1 nucleosome (x-axis with the white dashed line indicating the +1 dyad) (Chereji et al. 2017). This analysis, presented in Figure 3.6A, shows that the input DNA used for ChIP was slightly enriched in sequences –400 to +100 bp relative to the +1 nucleosome (left panel) and contained mononucleosome-sized DNA fragments, with the peak of distribution at ~165 bp (right panel). In contrast, Hho1 antibodies immunoprecipitated primarily larger fragments with a peak of distribution of ~270 bp (right panel). Interestingly, smaller fragments were present in the Hho1 ChIP, but few small fragments overlapped the +1 nucleosome (middle panel), despite being present in the input (left panel). In contrast, reads overlapping the +1 nucleosome were precipitated with Hho1 if they were longer and also overlapped the +2 nucleosome. Collectively, these data argue that the +1 nucleosome        43 is depleted in Hho1 such that these nucleosomes can only be precipitated with Hho1 antibodies if linked to a +2 nucleosome. This is consistent with previously published work demonstrating that Hho1 fails to cross-link to the 5’ linker of the +1 nucleosome (Rhee et al. 2014).  Depletion of Hho1 over the +1 nucleosomes is consistent with the fact that these nucleosomes tend to be highly acetylated. To determine if histone acetylation and Hho1 inversely correlate genome-wide, we quantified the levels of Hho1 and multiple histone post-translational modifications (Weiner et al. 2015) over the 67,523 annotated yeast nucleosomes (Brogaard et al. 2012) and generated a pairwise Spearman correlation matrix with hierarchical clustering. Figure 3.6B shows that Hho1 occupancy clustered with histone H4R3 mono-methylation and H2AS129 phosphorylation. Little is known about the function of H4R3 methylation in yeast, but H2AS129p is enriched at repressed protein-coding genes (Szilard et al. 2010), consistent with a role of Hho1 in negatively regulating transcription. In contrast, except for H4K16ac, all histone acetylation marks in yeast clustered away from Hho1 occupancy with inverse correlation coefficients consistent with the negative regulation of linker histone binding by histone acetylation.  An explanation for the inverse correlation between acetylation and linker histone occupancy observed in Figure 3.6B is that transcription, which is linked to acetylation, disrupts the interaction of Hho1 with chromatin. Indeed, data supporting this possibility have been published (Schafer et al. 2008). To discount a role of transcription in regulating Hho1 binding, we took advantage of the fact that the association between acetylation and        44 transcription is not absolute. For example, while RNAPII traverses the entirety of a gene, histone acetylation is primarily limited to the 5’ end of the transcribed unit. Thus, pools of nucleosomes can be identified that share similar levels of RNAPII but have different amounts of histone acetylation. To determine whether the inverse correlation between Hho1 and histone acetylation is due to the presence of RNAPII, we divided yeast nucleosomes into quartile bins based on RNAPII occupancy as determined in Hobson et al. (2012). We further divided each of the resulting bins based on histone acetylation and identified nucleosomes in each bin with high (top quartile, blue) and low (bottom quartile, red) levels of H3K23ac or other acetylation marks (Weiner et al. 2015). Figure 3.6, C and D show the amounts of RNAPII and H3K23ac in the eight resulting bins, respectively. We then calculated Hho1 occupancy for each bin and plotted it as a box plot (Figure 3.6E for H3K23ac and Figure 3.7 for other acetylation sites). The results show that nucleosomes with increased H3K23ac had reduced Hho1 occupancy when levels of RNAPII were normalized, indicating that differing RNAPII levels were not responsible for altered Hho1 occupancy. Similar trends were observed when analyzing other acetylation marks (Figure 3.7). Additionally, comparable results were obtained when analyzing linker histone H1.2, H1.3 and H1o occupancy (Cao et al. 2013) in mouse embryonic stem cells relative to H3K9ac (Figure 3.8, A–E) and H3K27ac (Figure 3.8F). Collectively, these results suggest that acetylated chromatin is refractory to linker histone binding in S. cerevisiae and other organisms.        45         46 Figure 3.6: Histone acetylation negatively correlates with linker histone binding in S. cerevisiae (A, left and middle) Two-dimensional occupancy plots of relative sequence fragment abundance, sequence fragment length and sequence fragment position from input DNA and Hho1 ChIP, relative to the dyad axis of the +1 nucleosomes of 5770 annotated genes in S. cerevisiae. Plot was generated using plot2DO (Chereji et al. 2017) run with standard settings. The relative sequence read abundance is indicated as a heatmap, the sequence fragment length is plotted on the y-axis and the position of sequence reads relative to the +1 nucleosome is plotted on the x-axis. (Right) Plot of sequence fragment lengths from input DNA and Hho1 ChIP-seq of wild-type yeast. (B) Clustered heatmap produced by the deepTools plotCorrelation module (Ramírez et al. 2014; 2016). Shown here are the Spearman correlation coefficients of Hho1 occupancy (blue text) at all uniquely mapping yeast nucleosomes (Brogaard et al. 2012) with histone post-translational modifications (Weiner et al. 2015), including histone acetylation (red text). All data sets were normalized to respective inputs using the deepTools bigwigCompare tool (C–E). All uniquely mapping yeast nucleosomes (Brogaard et al. 2012) were binned into quartiles based on RNAPII occupancy (C) and the top and bottom quartiles of H3K23ac (Weiner et al. 2015) (D). Hho1 occupancy was then plotted for each bin (E). Notches indicate the 95% confidence interval for the median.          47  Figure 3.7: Histone acetylation negatively correlates with linker histone binding in S. cerevisiae  All uniquely mapping yeast nucleosomes (Brogaard et al. 2012) were binned into quartiles based on RNAPII occupancy (Figure 3.6C) and the top and bottom quartiles of the indicated histone acetylation marks (Weiner et al. 2015) were calculated using the Java Genomics Toolkit ngs.IntervalStats tool. Hho1 occupancy was then plotted for each bin. Notches indicate the 95% confidence interval for the median.          48    Figure 3.8: Histone acetylation negatively correlates with linker histone binding in mouse embryonic stem cells The mouse genome was divided into 1000 bp windows, stepping 500 bp. Windows were divided into four quartiles based on RNAPII (GSM723019) occupancy (A) and the top and bottom quartiles for H3K9ac (GSM1000127) occupancy (B) as determined using the Java Genomics Toolkit ngs.IntervalStats tool. H1.2, H1.3 and H1o occupancies (Cao et al. 2013) were then plotted for each bin (C–E, respectively). Notches indicate the 95% confidence interval for the median. (F) Identical analysis as in C, but windows were binned based on H3K27ac (GSM1000099) instead of H3K9ac.            49   – DISCUSSION Refining linker histone stoichiometry  In this study, we investigated the regulation of linker histone binding in the yeast, S. cerevisiae. A major factor thought to regulate linker histone levels in chromatin is the abundance of linker histone in the cell. While vertebrate cells contain approximately one linker histone for every nucleosome, S. cerevisiae exhibits reduced linker histone levels; albeit the reported stoichiometry relative to nucleosomes varies depending on the study (Woodcock et al. 2006). Using a novel approach, we determined that yeast have one molecule of Hho1 for every ~19 nucleosomes. Previous work using various approaches have estimated the stoichiometry of Hho1 to nucleosomes to be 1:4, 1:10, 1:26 and 1:37 (Freidkin and Katcoff 2001; Downs et al. 2003; Ghaemmaghami et al. 2003; Kulak et al. 2014). Interestingly, the 1:26 ratio, which is the closest to our result, is the only previous study to quantify levels of un-tagged Hho1 (Kulak et al. 2014). Regardless of the exact ratio, all studies agree that Hho1 levels are well below that of nucleosomes and because linker histones bind and compact chromatin cooperatively (Routh et al. 2008), this may be important to prevent formation of higher-order chromatin structures on the gene-rich yeast genome.  Increased linker histone expression is toxic in yeast Consistent with the importance of sub-stoichiometric linker histone levels in S. cerevisiae, increased expression resulted in a severe growth defect. Interestingly, this defect was        50 accompanied by only modest increases in linker histone levels at most loci tested. This result was initially surprising since in fibroblasts, overexpressed linker histones can bind chromatin, increasing the overall nucleosome repeat length (Gunjan et al. 1999). While we cannot exclude the fact that some of the overexpressed Hho1 is cytoplasmic and thus not available for chromatin binding, multiple lines of evidence support the hypothesis that short linker DNA in yeast chromatin excludes Hho1. First, previous work shows that Hho1 is enriched in regions with increased spacing between nucleosomes (Ocampo et al. 2016), suggesting that linker DNA availability as opposed to linker histone abundance, dictates Hho1 binding. Second, in stationary phase, genic nucleosomes increase their spacing (Zhang et al. 2011), which coincides with increased linker histone binding to chromatin (Schafer et al. 2008). Finally, chromatin reconstituted on DNA that positions nucleosomes with short linkers is resistant to linker histone binding and compaction (Routh et al. 2008). Collectively, these data suggest that histone stoichiometry does not dictate linker histone binding in yeast. Instead, overexpressed Hho1 likely localizes to limited regions with longer linker DNA, such as the 5’ ends of genes, where it interferes with the early stages of transcription.   Histone acetylation negatively regulates linker histone binding  The preference of Hho1 for binding regions with longer linker DNA, makes the +1 nucleosome, which is adjacent to a nucleosome free region (NFR), a seemingly ideal ligand for binding by this linker histone. However, our data and that of others, show Hho1 depletion from the +1 nucleosome. The molecular basis for this observation was revealed        51 in this study using both genetic and genome-wide approaches, demonstrating that histone tail acetylation negatively regulates Hho1 binding. Since the 5’ ends of transcribed genes are highly acetylated, these regions may be largely refractory to Hho1 binding. The ability of histone acetylation to hamper the binding of Hho1 is not surprising, as others have observed increased linker histone mobility in cells treated with HDAC inhibitors (Raghuram et al. 2010). Together our data support a model in which histone acetylation, not histone stoichiometry, plays a dominant role in regulating linker histone binding in S. cerevisiae. Moreover, our results shed light on the puzzling finding that despite their role in compacting chromatin, depletion of linker histones does not result in global upregulation of gene expression (Shen and Gorovsky 1996; Freidkin and Katcoff 2001; Fan et al. 2005a). Since acetylation likely excludes linker histones from regions of transcriptional activity, their depletion has little impact on steady-state transcription.   Does the GAL1 promoter induce the expected amount of Hho1 expression? It is possible that a feedback mechanism exists in yeast to degrade excess free Hho1 in the cell. If this is the case, we would expect reduced Hho1 expression compared to other proteins expressed from the same GAL1 promoter. However, deletion of HDA1 which is expected to displace Hho1, does not result in reduced Hho1 levels in the cell. To explore this further, future experiments could upregulate Hho1 expression further using one or more high-copy plasmids to reach comparable levels of linker histones compared to nucleosomes. If no feedback mechanism exists in yeast to degrade free linker histones, we would expect higher Hho1 levels to result in exacerbated cellular toxicity.          52 Why is HHO1 overexpression toxic? Clearly, increased linker histone levels are toxic in S. cerevisiae. One possible explanation could be that increased Hho1 binding over the 5’ end of genes could inhibit RNAPII promoter escape or early elongation, resulting in transcription inhibition. Indeed RNAPII has been shown to accumulate at the 5’ ends of genes (Churchman and Weissman 2011). To test this, native elongation transcript-sequencing (NET-seq) could be performed in cells overexpressing Hho1. Alternatively, toxicity could be due to ectopic Hho1 expression outside of S phase of the cell cycle. Hho1 along with the core histones are primarily expressed during S phase, and increased core histone levels results in compromised cellular fitness (Spellman et al. 1998; Kurat et al. 2014). This could be tested by expressing Hho1 from its endogenous promoter from one or more high-copy plasmids, ensuring overexpression occurs primarily during S phase. If ectopic expression of Hho1 is mediating toxicity, we would expect to see improved fitness compared to cells expressing a similar amount of Hho1 from the GAL1 promoter.   Significance  This work supports a model in which core histone acetylation, which is a consequence of transcription, reinforces active transcription. Additionally, this adds to our understanding of how core histone acetylation promotes transcription in yeast and other organisms by regulating the binding of linker histones at transcribed genes. Moreover, if histone PTMs are heritable markers of gene expression states, this work provides a mechanism for the        53 epigenetic inheritance of linker histone levels and their associated transcriptional states via histone acetylation.                             54 REFERENCES Ali T., Thomas J. O., 2004 Distinct Properties of the Two Putative “Globular Domains” of the Yeast Linker Histone, Hho1p. Journal of Molecular Biology 337: 1123–1135. Ali T., Coles P., Stevens T. 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