View of Histone Modifications As Epigenetic Marker

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2011 Role of Histone Gene Dosage in DNA Damage Repair Dun Liang

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COLLEGE OF MEDICINE

ROLE OF HISTONE GENE DOSAGE IN DNA DAMAGE REPAIR

DUN LIANG

A dissertation submitted to the Department of Biomedical Sciences in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Degree Awarded: Summer Semester, 2011

The members of the committee approve the Dissertation of Dun Liang defended on July 1st, 2011.

______Akash Gunjan Professor Directing Dissertation

______David Gilbert University Representative

______Myra Hurt Committee Member

______Johanna Paik Committee Member

______Yanchang Wang Committee Member

Approved: ______Richard Nowakowski, Chair, Department of Biomedical Sciences

______John Fogarty, Dean, College of Medicine

The Graduate School has verified and approved the above-named committee members.

To be as firm and resolute in my actions as I was able, and not to adhere less steadfastly to the most doubtful opinions, when once adopted, than if they had been highly certain… To endeavor always to conquer myself rather than fortune, and change my desires rather than the order of the world, and in general, accustom myself to the persuasion that, except our own thoughts, there is nothing absolutely in our power; so that when we have done our best in things external to us, all wherein we fail of success is to be held, as regards us, absolutely impossible…

--Discourse on the Method, René Descartes, 1637

iii

ACKNOWLEDGEMENTS

My PhD training is an odyssey both in science and life. I am full of gratitude to people who helped me all the way along.

First and foremost I would like to thank my advisor Dr. Akash Gunjan, who is not only a good teacher but also a good friend to me, for his immeasurable support during the past six years. He taught me to do science, to think like a scientist, and most importantly, to live an exploring, creative and productive life. He has always been generous towards my confusions and mistakes; he provided me with freedom and space for development, and with prompt guidance or directions when needed; he showed me the appropriate bearing towards daily frustration and disappointments. He is a true role model in motivating me to pursue in science. I would not be who I am today without his mentoring.

Secondly, I would like to acknowledge my other committee members: Drs. Myra Hurt, Yanchang Wang, David Gilbert and Johanna Paik. They offered me invaluable support, encouragement and advice towards my research. I deeply appreciate the extra effort of Dr. Wang put in to help me to adapt to the cultural differences as a foreign student and provide me with many yeast strains and other materials. Special thanks are due to Dr. Hurt for her consistent financial support and understanding. She will be remembered by my family as serving on the PhD committees for both my wife and me. Additionally thanks to Dr. Gilbert, who has always been the source of insight about the beauty of science and to Dr. Paik, who provided me countless support and care. I take this opportunity to thank them sincerely and deeply.

As a member of Gunjan lab family, I express my appreciation to my lab colleagues. It has been a great pleasure to work and share my life with them in the lab.

iv Special acknowledgement goes to Dr. Rakesh Singh, who helped to open my eyes and mind in science, as well as to provide critical inputs for my projects. He will be also thanked for his persistence in asking all those “forbidden to graduate students” questions to me over the last few years to keep me awake. Many thanks due to Ugander Reddy for the help in optimizing my experiments and for being a great companion in the lab. I would like to give a special thank to Marie-Helene Kabbaj for her endless support in lab management and material preparation. Apart from the work, I also learned to be a healthy and eco-friendly scientist from them all.

Here within, I would like to acknowledge other faculties, staffs, postdocs and graduate students in the Department of Biomedical Sciences, who have created such a friendly, dynamic, supportive and inspiring research environment. For those not specifically mentioned, please accept my sincerest appreciation for your help and assistance.

I am also filled with gratitude to my parents for providing me the best education opportunities available and understanding of the path that I chose in my life. Their love and support granted me the opportunity to move forward without any hesitation or concern. Finally, words of thanks are due to my wife, who I met, fell in love with and married here in FSU. She makes my life different, she makes me home. I could not finish this work without her company and comfort. Thank you.

TABLE OF CONTENTS

List of Tables...... ix List of Figures...... x Abstract ...... xii 1. DNA DAMAGE AND REPAIR...... 1 1.1 Various Types of DNA Damage ...... 1 1.2 Different Types of DNA Damage Repair ...... 4 1.2.1 Direct repair...... 5 1.2.2 Mismatch repair...... 6 1.2.3 Excision repair...... 6 1.2.4 Break repair...... 10 1.3 Choice of DSB repair pathway: HR vs. NHEJ ...... 26 1.3.1 Nature of DNA DSB breaks ...... 27 1.3.2 Effect of cell cycle regulation on the choice of DSB repair pathway ... 29 1.3.3 NHEJ factor’s binding to the break ends ...... 30 1.3.4 DSB repair pathway choice in different species ...... 31 1.4 DNA Damage Signaling in Budding Yeast ...... 33

2. THE ROLE OF HISTONES, CHROMATIN STRUCTURE, AND EPIGENETIC MODIFICATIONS IN DNA REPAIR ...... 38 2.1 Histones Serve as Primary Protein Components of Chromatin...... 38 2.2 Definition of Epigenetics...... 42 2.3 Overview of Histone Modifications as Epigenetic Marker...... 43 2.4 Role of Histone Modifications in DSB Repair ...... 45 2.4.1 H2A phosphorylation ...... 45 2.4.2 Histone ubiquitination ...... 46 2.4.3 Methylation of H3K79 and H4K20 ...... 48 2.4.4 Histone acetylation ...... 49 2.5 Chromatin Remodeling and DNA Repair ...... 52 2.6 Role of Histone Chaperones in DSB Repair...... 58 2.7 DNA methylation and DNA damage repair...... 62

3. HISTONES: THE ENEMY WITHIN...... 64 3.1 Histones: Enemy or Friend?...... 64 3.2 Sources of Excess Histones...... 68 3.2.1 Multiple histone genes...... 69 3.2.2 Replication arrest...... 72 3.2.3 Histones evicted during transcription or DNA damage repair ...... 73 3.3 Regulation of Histone Level ...... 75 3.3.1 Transcriptional control of histone gene expression ...... 75

vi 3.3.2 Posttranscriptional regulation of histone gene expression...... 78 3.3.3 Posttranslational regulation of histone proteins ...... 79 3.3.4 Histone chaperones in regulating histone proteins...... 80

4. EXCESS HISTONE LEVELS MEDIATE CYTOTOXICITY VIA MULTIPLE MECHANISMS...... 83 4.1 Abstract...... 83 4.2 Introduction ...... 84 4.3 Results ...... 85 4.3.1 Budding yeast mutants lacking certain histone modifying enzyme subunits are sensitive to the presence of excess histones...... 85 4.3.2 Excess histones do not appear to alter the bulk nucleosomal structure but lead to subtle alterations in the fine structure of chromatin...... 86 4.3.3 Overexpression of histone gene pairs, but not individual histones, results in the alteration of transcript levels of numerous genes...... 91 4.3.4 Excess histones can bind to RNA ...... 95 4.4 Discussion...... 97 4.5 Materials and Methods...... 102

5. HISTONE GENE DOSAGE MODULATES DNA REPAIR VIA THE HOMOLOGOUS RECOMBINATION PATHWAY ...... 107 5.1 Abstract...... 107 5.2 Introduction ...... 108 5.3 Results ...... 112 5.3.1 Histone gene dosage has a major influence on the sensitivity of budding yeast cells to DNA damaging agents...... 112 5.3.2 Changes in histone gene dosage have little effect on the bulk chromatin structure or global gene expression...... 114 5.3.3 The DNA damage checkpoint is efficiently activated in cells with reduced histone dosage...... 117 5.3.4 Reduction in histone gene dosage enhances survival of yeast cells following a DNA double strand break (DSB) in a checkpoint independent manner...... 122 5.3.5 Improved survival of strains carrying a reduced histone dosage following DSB induction depends on homologous recombination and not on non-homologous end joining ...... 129 5.3.6 Reduction in histone gene dosage enhances spontaneous recombination rates...... 130 5.3.7 Histones compete with homologous recombination factors for binding to damaged DNA...... 132 5.4 Discussion...... 137 5.5 Materials and Methods...... 142

vii 6. CONCLUSIONS ...... 146

REFERENCES...... 151 APPENDIX A...... 180

APPENDIX B...... 193

BIOGRAPHICAL SKETCH...... 195

viii LIST OF TABLES

1.1 Homologs of DSB repair factors in different species...... 14

1.2 Homologs of DNA damage checkpoint proteins in different species...... 37

4.1 List of strains used in the study described in chapter 4 ...... 106

5.1 List of budding yeast strains used in the study described in chapter 5...... 144

A.1 List of genes affected 2-fold or more by the simultaneous overexpression of the histone H3-H4 gene pair...... 180

A.2 List of genes affected 2-fold or more by the simultaneous overexpression of all the four core histones...... 185

A.3 List of genes that are upregulated or downregulated in response to overexpression of the H3-H4 gene pair as well as upon overexpression of all the core histones...... 192

B.1 List of genes affected 2-fold or more upon hht2-hhf2 deletion...... 193

LIST OF FIGURES

1.1 Different forms of DNA damage and the multiple DNA repair pathways...... 3

1.2 Scheme for homologous recombination during DSB repair ...... 20

1.3 Scheme for Non-Homologous End Joining during DSB repair in budding yeast ... 25

1.4 The DNA damage response pathway in budding yeast...... 36

2.1 Nucleosome and basic chromatin structure...... 41

2.2 “Access, Repair, Restore” model for the role of histones during DSB...... 61

3.1 Consequences of excess histones ...... 67

3.2 Sources of excess histones ...... 70

3.3 Regulation of Histone Levels ...... 76

4.1 Yeast cells lacking histone modifying enzyme subunits are sensitive to histone overexpression...... 87

4.2 Yeast cells lacking histone modifying enzyme subunits are sensitive to co- overexpression of the histone H3-H4 gene pair...... 88

4.3 Histone overexpression does not affect the bulk chromatin structure but alters the fine structure of chromatin...... 90

4.4 Indirect end-labelling analysis of chromatin from histone overexpression cells following in vivo DNase I cleavage...... 92

4.5 Genome wide comparison of gene expression in wild type yeast cells upon overexpression of individual or pairs of histone genes...... 93

4.6 Excess histones can bind to RNA...... 96

5.1 Histone gene dosage influences the DNA damage sensitivity of wild type and DNA damage checkpoint deficient yeast strains...... 113

5.2 Global chromatin structure remains unchanged upon deletion of the hht2-hhf2 histone gene pair...... 116

x 5.3 Genome wide comparison of gene expression in wild type yeast cells to yeast cells carrying a deletion of the hht2-hhf2 gene pair ...... 118

5.4 The DNA damage checkpoint remains intact and responds normally to replication inhibition upon deletion of the hht2-hhf2 histone gene pair ...... 119

5.5 Deletion of the histone gene pair hht2-hhf2 does not alter the intra-S checkpoint.120

5.6 Flow cytometry analysis of the progress of replication in response to low doses of the methylating agent Methylmethane Sulfonate (MMS)...... 121

5.7 Galactose inducible DSB at the MAT locus ...... 124

5.8 A reduction in histone gene dosage enhances survival of budding yeast cells following HR mediated repair of an induced DSB ...... 125

5.9 Change of damage signaling and HR dependent repair upon hht2-hhf2 deletion.128

5.10 Reduction in histone gene dosage affects the efficiency of DSB repair via the Homologous recombination (HR) but not the Non- Homologous End Joining (NHEJ) pathway...... 131

5.11 Reduction in histone gene dosage results in elevated rates of spontaneous homologous recombination...... 134

5.12 Histones compete with HR factors for binding to DNA repair sites ...... 135

5.13 Recruitment of RPA1-TAP to a HO mediated DSB at the MAT locus...... 136

5.14 Model to illustrate the potential effect of excess histones on HR or NHEJ mediated repair of a DSB...... 141

ABSTRACT

In eukaryotes, each individual chromosome is one large DNA molecule packed by histone proteins into a compact nucleoprotein filament. Two molecules each of core histone proteins H2A, H2B, H3 and H4 assemble to form an octamer protein core around which 147 base pairs of DNA is wrapped to form the nucleosome core particle and this structure is repeated to form chromatin. Histones are essential proteins as they package the genomic DNA to fit it inside the relatively tiny nucleus and regulate DNA accessibility. However, when present in excess, the positively charged histones can bind non-specifically to negatively charged DNA and affect all forms of DNA metabolism such as transcription, replication, repair and recombination.

The DNA of all organisms is under constant threat of damage from both exogenous and endogenous agents that can contribute to genomic instability, which is characterized by the increased rate of acquisition of alterations in the genome and is a hallmark of cancer cells. Hence, cells have evolved multiple mechanisms to ensure genomic stability. Since DNA damage and repair occurs in a chromatin context in eukaryotes, chromatin structure and histones may affect genomic stability. Not surprisingly, scarcity of histones during DNA replication results in spontaneous DNA damage. On the other hand, accumulation of excess histones leads to genomic instability in the form of excessive chromosome loss, enhanced sensitivity to DNA damaging agents and cytotoxicity. Therefore, histone synthesis is tightly regulated at transcriptional, posttranscriptional, as well as posttranslational levels.

Here, we have investigated the mechanism/s via which excess histones exert their deleterious effects in vivo in the budding yeast. We find that the presence of excess histones saturates certain histone modifying enzymes, potentially interfering with their activities. Additionally, excess histones appear to bind non-specifically to DNA as

xii well as RNA, which can adversely affect their metabolism. Microarray analysis revealed that upon overexpression of the histone H3 and H4 gene pair or all four core histones but not individual histones, about 240 genes were either up or downregulated by 2-fold or more. Interestingly, histone overexpression does not affect the bulk chromatin structure, but alters the fine structure of chromatin. Overall, we present evidence that excess histones are likely to mediate their cytotoxic effects via multiple mechanisms that are primarily dependent on inappropriate electrostatic interactions between the positively charged histones and diverse negatively charged molecules in the cell.

We have also investigated how changes in histone gene dosage affects the DNA damage sensitivity of budding yeast cells that have two copies of each histone gene when only one copy is needed for survival. We found that overexpression of histones led to an increase in DNA damage sensitivity. Next, we deleted the second copy of the gene pair (HHT2-HHF2) encoding histones H3 and H4 that contributes 6-8 fold more histone mRNA than the first gene pair (HHT1-HHF1), to create an experimental system to study the effects of reduced histone levels in vivo . A reduction in the dosage of histone H3-H4 resulted in a significant decrease in DNA damage sensitivity. By taking the advantage of a HO endonuclease induced DNA double stand break (DSB) at the budding yeast mating type (MAT) locus, we were able to study the DSB repair process in detail in strains with a reduced histone gene dosage. We found that the efficiency of Homologous Recombination (HR) at a DSB, as well as genome wide HR, was elevated in hht2-hhf2 deletion strain, while Non-Homologous End Joining remained unchanged. These effects were not associated with global changes in the expression of DNA repair genes or DNA damage checkpoint responses. We also found that there was no alteration of gross chromatin structure in response to changes in histone gene dosage. One mechanism by which reduced histone dosage leads to elevated HR mediated repair of a DSB at the MAT locus is through enhanced recruitment of the HR factors, as determined by the Chromatin Immunoprecipitation (ChIP) assay. Concomitant with this, cells experience a greater histone loss around this DSB upon a reduction in histone gene dosage. We propose that high levels of

xiii endogenous histones generated by multiple genes in eukaryotes compete with HR factors, thereby reducing HR efficiency and may normally function to restrain potentially excessive HR activities during S-phase.

Overall, our findings help to explain the basis for the existence of multiple mechanisms that regulate histone levels and highlight their role in maintaining genomic stability and cell viability. Our findings could have major implications for DNA repair, genomic stability, carcinogenesis and aging in human cells that have dozens of histone genes.

xiv CHAPTER 1

DNA DAMAGE AND REPAIR

Being the primary carrier of genetic information, the DNA needs to be faithfully passed from one generation to the next. The survival and fitness of a cell, tissue type, organ as well the individual as a whole is highly dependent on the fidelity of DNA transmission and its maintenance. In terminally differentiated cells, the DNA is unique in that it is the largest macromolecule that is not renewed following its initial synthesis and remains the very same molecule throughout the cell’s life. As such, this DNA is likely to suffer from routine wear and tear over time. Additionally, a typical DNA molecule inside a cell may stretch for millions of nucleotides, every single one of which has to be placed in the correct position during replication. This can be a very challenging task for the cell and failure to achieve this may result in errors with deleterious consequences. Further, although DNA is considered to be a reasonably stable molecule, its chemical nature makes it labile and susceptible to damage by chemical and physical agents generated from internal sources within the cell as well as external sources. Hence, cells have evolved numerous strategies to detect and repair damage caused to their DNA by a variety of agents.

1.1 Various Types of DNA Damage

There are three major categories of DNA damage: errors that are incorporated into the genome during DNA replication; lesions that caused by cell metabolism; damage that are caused by agents in the external environment. The various forms of DNA damage are illustrated in Fig1.1 (Fleck and Nielsen, 2004).

1 During DNA replication, a crucial property of the DNA polymerase is to insert the right nucleotide complementary to the template base with kinetic selectivity (Joyce and Benkovic, 2004). To ensure accuracy, the major replicative polymerases possess a 3’ exonuclease dependent proofreading mechanism to degrade the incorrectly base paired nucleotides (Shevelev and Hubscher, 2002). Nevertheless, some errors do escape from this proofreading and form base pair mismatches between the newly synthesized strand and template strand. This kind of error is insidious because there is no actual lesion or break on the DNA, as normal nucleotides are incorporated. If these are not detected and corrected before the second round DNA replication, these errors could pass on to daughter cells as legitimate mutations that will persist in the genome and can never be noticed.

Apart from replication errors, the DNA itself is vulnerable and under constant attack of from both inside the cell and outside. The mitochondrial respiratory chain generates Adenosine-5'-triphosphate (ATP) using electron transport starting from NADH (nicotinamide Adenine dinucleotide H) but at the same time produces various reactive oxygen species (ROS) in every cell. Routine cellular metabolism reactions byproducts containing ROS such as hydrogen peroxide and superoxide anions, as well as reactive nitrogen species (RNS) such as Nitric oxide (NO) and other alkylating species. Mutations caused by oxidative DNA damage include a range of specifically oxidized purines and pyrimidines, alkali labile sites and single strand breaks (Cooke et al., 2003; Waris and Ahsan, 2006). One of the most frequently studied base lesion caused by an oxidative agent is 7,8-dihydro-8-oxoguanine or 8-oxoG, which is the oxidation product of guanine. The 8-oxoG adduct is highly mutagenic because it can pair with A (Adenine) as well as C (Cytosine) residues, leading to a greatly increased frequency of spontaneous G/C to T/A transversion mutations in the absence of timely repair (Cheng et al., 1992). Hence, cells have evolved a strong antioxidant system to counteract the effects of the free radicals generated during energy production and other metabolic activities.

Figure1.1 Different forms of DNA damage and the multiple DNA repair pathways. Figure is adapted from (Fleck and Nielsen, 2004).

Finally, there are numerous dangerous mutagens present in the environment. Ionizing radiation (IR) such as X-ray and gamma rays, as well as particles such as alpha/beta particles and neutrons are all capable of inflicting the most hazardous kinds of DNA damage. Not only are a variety of chemical modifications of DNA induced by ionizing radiation derived free radicals, but they also cause DNA double strand breaks (DSBs) by directly attacking the deoxyribose backbone of DNA. Additionally, UV radiation from daily sun exposure is strongly absorbed by the DNA bases causing pyrimidine dimers and pyrimidine 6-4 photoproducts (Rastogi et al., 2010). Moreover, alkylating chemicals present in drugs and environmental toxins display potent reactivity towards DNA. Methyl or ethyl groups can be easily transferred to reactive sites either on the bases or to the phosphates in the DNA backbone (Gates, 2009). A good example is the alkylation of oxygen-6 of guanine yielding O6-methylguanine (O6-meG), which is often mismatched with thymine (T) resulting in G to A transitions. Oxidizing materials in the external environment could be much more harmful compared to those from inside. Under certain conditions the DNA can suffer hydrolysis damage as well. For instance, deamination of a cytosine base will generate a uracil and spontaneous hydrolytic cleavage of the glycosidic bonds in DNA often yields an apurinic site (AP site) on the DNA, an event referred to as depurination (Gates, 2009).

1.2 Different Types of DNA Damage Repair

Different types of DNA damage can lead to distinct cellular outcomes that are highly variable dependent on the dosage, location, cell cycle timing, number of lesions and the cell type. There are two major consequences of DNA lesions. One possibility is that the damage causes physical breaks, nicks or covalent obstacles that will block replication or transcription of critical genes. In this case, there will be inevitable cell cycle arrest or cell death if the lesion persists long

4 enough, and as such these lesions are cytostatic or cytotoxic. On the other hand, damage such as base transversions and transitions could be harmless initially and will not stop cell from dividing. However, after the mispaired DNA replicated, permanent alterations in the primary sequence of DNA will occur in the form of mutations or chromosomal aberrations which is believed to be intimately linked to cancer (Hoeijmakers, 2009). Moreover, the chance that a cell undergoes damage under normal conditions is estimated to be close to 105 lesions per cell per day (Hoeijmakers, 2009). Facing such a constant high risk of DNA damage, cells have evolved elaborate mechanisms to detect and repair their damaged DNA before they cause further problems or form mutations as illustrated in the lower half of Fig1.1 (Fleck and Nielsen, 2004). Although different repair systems are not exclusively responsible for one type of DNA damage, one can distinguish them by the nature of repair. Except for the repair of mismatched bases, most lesion repairs fall into three categories: direct repair, excision repair and break repair.

1.2.1 Direct repair

Direct repair is a simple reaction in which one or two enzymes detect and reverse the damage at the same site of lesion. For instance, the ultra violet (UV) irradiation -induced pyrimidine dimer is repaired by simple reversal reaction called photo-reactivation. DNA photolyase captures energy from light and uses it to break the covalent bonds linking adjacent pyrimidine to restore the two individual bases (Beukers et al., 2008). Similarly, O6-methylguanine-DNA methyltransferase (MGMT) repairs O6-methylguanine by a unique lesion reversal mechanism which recognizes the methyl group and transfers it to one of its own cysteine residues (Hall et al., 1990). This is a suicide reaction and as such the same methyltransferase molecule can be used just once and is inactivated afterwards.

5 1.2.2 Mismatch repair

To correct errors of DNA polymerases that escape their 3’→ 5’ exonucleolytic proofreading, a mechanism called mismatch repair (MMR) has evolved. The mismatch can be identified in DNA because they fail to from canonical Watson– Crick base pairs. However, since the base and phosphate backbone is not modified or damaged, it is not easy to distinguish which strand carries the correct genetic information. Thus, a mismatch cannot be repaired by an excision mechanism which simply excises the damaged base, or a short DNA fragment containing the damage respectively (Stojic et al., 2004). In humans, the mismatch is recognized by heterodimer of MutS Homolog 2/6 (MSH2/MSH6), which binds to the DNA and forms an ATP-dependent repair clamp. By binding with another heterodimer MLH1/PMS2, this tetrahetromeric complex can translocate in either direction along the DNA in search of a strand discontinuity (Gradia et al., 1999). When this tetrahetromeric complex encounters a strand discontinuity (such as a gap between Okazaki fragments), it may be bound by Proliferating Cell Nuclear Antigen (PCNA) leading to the loading of Exonuclease1 (EXO1), which in turn initiates degradation of the nicked strand towards the mismatch. Polymeraseδ (Polδ), PCNA and Replication factor C (RFC) as well as DNA ligase1 complete the repair by re-synthesizing the degraded DNA strand and sealing the ends, while Replication protein A (RPA) proteins protect the template strand (Stojic et al., 2004). The function of the MMR system is not just restricted to the mitotic cell cycle, but it also plays critical roles in meiotic recombination and chromosome pairing. Since replication errors are very common in all organisms, the MMR proteins are highly conserved across different species.

1.2.3 Excision repair

1.2.3.1 Base Excision repair

6 The easiest way to clear out damaged bases is to remove and replace the altered bases. There are mainly two excision repair pathways: base excision repair (BER) and nucleotide excision repair (NER). Base lesions without helix distortions that are usually caused by cellular metabolism derived oxidative stress, are mainly repaired by BER (Barnes and Lindahl, 2004). For example, smaller lesions such as oxidation productss (8-oxoG), spontaneous hydrolysis sites (AP site), deaminations (cytosine to uracil), non-enzymatic alkylation (3- methyladenine), as well as some single strand breaks are likely to be corrected by BER (Wilson and Bohr, 2007). The major steps of BERare as follows. First, the damaged base is excised to generate an abasic site. Then the phosphodiester backbone at the abasic site is cleaved to generate a single strand break (SSB). In the third step, the excised nucleotide is replaced and this followed by theclean up of the terminal end(s) if required. In the last step, DNA ligase seals the nicked DNA strand (Wilson and Bohr, 2007).

One of the most crucial enzymes involved in initiating BER are DNA glycosylases, which are not only responsible for recognizing the lesion, but also for excising the inappropriate substrate base. DNA glycosylases are lesion specific and cells have multiple glycosylases with different specificities. For example, there are special glycosylases that deal with uracil, while a different one deals with 8-oxoG. However, individual glycosylases may recognize more than one type of damage, and each specific modification may be recognized by more than one type of glycosylase, giving a degree of redundancy in the process (Maynard et al., 2009). Glycosylases flip out the damaged base from DNA helix and cleave the N-glycosidic bond between deoxyribose and the base, leaving behind an AP site (Stivers and Jiang, 2003). Some glycosylases, such as 8- oxoguanine glycosylase (OGG1), are bifunctional and function in excising the damaged base and incising DNA backbone at the same time (Hazra et al., 2001). The latter function overlaps with AP (apurinic) endonuclease, which cleaves an AP site to yield a 3' hydroxyl adjacent to a 5' deoxyribosephosphate (dRP). In order for ligation to occur, a DNA strand break must have a hydroxyl on its 3' end

7 and a phosphate on its 5' end. In humans, polynucleotide kinase-phosphatase (PNKP) promotes formation of these ends during BER. This protein has a kinase domain, which phosphorylates 5' hydroxyl ends, and a phosphatase domain, which removes phosphates from 3' ends (Wiederhold et al., 2004). DNA polymerase β is then recruited for inserting the correct single nucleotide to the AP site, which is referred as short patch BER. On the other hand, during long- patch BER, DNA synthesis is thought to be mediated by pol and pol along with the PCNA, the same factors that carry out the bulk of DNA replication (Wilson and Bohr, 2007). Thus, with the cooperation of glycosylases, AP endonuclease, terminal processing enzymes and polymerase, the damaged nucleotide is replaced with the correct one and a “perfect” single strand break is created. The remaining gap is filled by DNA ligase3α/XRCC1 (X-Ray cross- complementing) complex or by DNA ligase 1 alone (Tomkinson et al., 2001).

1.2.3.2 Nucleotide Excision repair

Unlike BER, the nucleotide excision repair (NER) enzymes do not recognize any particular lesion. Rather, this system functions as recognizing bulky distortions of DNA double helix, such as thymine dimer/ 6-4 photoproducts caused by UV, bulky chemical adduct on the base and some DNA intrastrand/interstrand crosslinks (ICLs) (Gillet and Scharer, 2006). NER is a versatile repair pathway, such that mutants associated with NER pathway cause a wide range of clinical symptoms, from mild solar sensitivity to severe skin cancers, developmental disorders and neurodegenerative diseases (Cleaver et al., 2009). In fact, diseases such as Xeroderma pigmentosum (XP) and Cockayne syndrome (CS) belong to few cases of cancer or aging related disorders whose etiology is directly linked to improper DNA damage repair at the molecular level (Hoeijmakers, 2009). The basic mechanism of NER is referred to as “cut and patch”. Recognition of helix distortion leads to the removal of a short single- stranded DNA patch that includes the lesion, creating a single-strand gap in the DNA, which is subsequently filled in by DNA polymerase using the undamaged

8 strand as a template. NER has two branches or sub-pathways: global genome repair (GGR), which repairs damage in both transcribed and untranscribed DNA strands in active and inactive genes; while transcription-coupled repair (TCR) removes distorting lesions that block elongating RNA polymerases from the transcribed DNA strand (Hoeijmakers, 2001). In prokaryotic organisms, such as E. coli, four proteins involved in NER are UvrA (UV repair protein A), UvrB, UvrC and UvrD. Two UvrA and one UvrB form an ATP-dependent complex to scan and recognize distortions. Upon encountering a distortion, the UvrAs load UvrB onto the damaged DNA and subsequently dissociate from the complex.On the other hand, during TCR, the Mfd (mutation frequency decline) protein, also called the transcription repair coupling factor (TRCF), recruits UvrA to sites of damaged DNA associated with stalled RNA polymerase (Guo et al., 2010). UvrB “melts” the DNA helix to form a bubble like pre-incision DNA-protein complex and recruits UvrC, which cleaves both side of the lesion creating a 12~13bp long single-stranded oligonucleotides that is displaced by DNA helicase UvrD. DNA polymerase I (Pol I) fills the gap generated in the DNA duplex and triggers the release of UvrB. Newly synthesized DNA is then joined to the extant DNA by DNA ligase (Guo et al., 2010). The principle mechanism of NER in eukaryotic cells is similar to that in bacteria, although the specific functions of the molecules involved are much more complicated. Two independent complexes XPC (Xeroderma pigmentosum C)/HR23B/Centrin2 and DDB1/DDB2(DNA Binding protein1/2) heterodimer are believed to be involved in the early steps of base damage recognition during NER (Nouspikel, 2009). A multiprotein transcription/repair complex, designated transcription factor IIH (TFIIH), is recruited to the damage site and later on functions along with other factors such as various XP proteins, ERCC1 (Excision repair cross-complementing), RPA, etc. Incision product is 24~32bp in length. The resulting gap is then filled in by the combined actions of DNA polymerase or , PCNA, RPA, and the final nick is sealed by DNA ligase. For TCR in eukaryotes, the stalled RNA polymerase complexes recruit CSB (Cockayne Syndrome B) and CSA (Cockayne Syndrome A) proteins, which belong to SWI/SNF (SWItch/Sucrose NonFermentable)

9 chromatin remodeling factor family (Fousteri et al., 2006). A relaxed region near the damage site is generated by CSB/CSA and other chromatin remodeling factors to increase accessibility and transcription/repair machinery. However, the detailed molecular mechanism of NER in eukaryotes is not fully understood yet.

1.2.4 Break repair

Among the all the various kinds of DNA damage, a double strand break (DSB) where the phosphate backbones of the two complementary DNA strands are broken simultaneously is probably the most lethal kind of damage. Not only does this cause a discontinuity in the genetic code but broken DNA ends are also vulnerable to further chemical attack resulting in damaged bases, the formation of abnormal DNA structures or even the total lost of large fragments of the chromosomes. The physical discontinuity of DSB presents a serious challenge during cell division because equal segregation of replicated genome relies on the intact chromosome carrying a single centromere. As such, unrepaired or misrepaired DSBs often lead to loss or translocation of genetic information and can have significant consequences, such as cell transformation or cell death (Khanna and Jackson, 2001). As little as one DSB can be sufficient to kill a cell if it inactivates an essential gene or triggers apoptosis (Rich et al., 2000).

1.2.4.1 DSBs are generated by various means

DSB can be produced in a variety of ways. In fact, most kinds of DNA damage mentioned previously in this chapter can eventually lead to DSBs. The most prominent cause and also the major exogenous agent for DSBs is ionizing radiation (IR) whose energy can directly rupture the DNA phosphodiester backbone by brute force.

A DSB can be considered to be a cluster of single-strand breaks (SSBs) with at least one SSB on each strand within a short distance. Certain chemotherapeutic

10 drug such as bleomycin and tirapazamine that act via the production of free radicals mimic the function of IR (Banath and Olive, 2003).

A second cause of DSBs are strong reactive oxygen species, such as a high concentration of hydrogen peroxide is also able to generate DSBs (Dahm-Daphi et al., 2000). During the course of normal oxidative respiration, mitochondria convert a small portion (0.1% to 1%) of the oxygen to superoxide. Superoxide dismutase (SOD) in the mitochondria or cytosol can convert this to hydroxyl free radicals, which may react with DNA to cause SSBs or DSBs (Termini, 2000).

The third and most important cause of DSBs in mitotic cells is replication of DNA. DSB can be produced when DNA replication forks encounter DNA single strand break or other lesions. For example, DNA alkylating agents such as methyl methanesulfonate (MMS) results in BER intermediates or unrepaired methyl damage which form DSB upon encountering the replication machinery (Wyatt and Pittman, 2006). Camptothecin (CPT), a topoisomerase I inhibitor is a commonly used anti-cancer drug. Topoisomerase-I is covalently linked to nicked DNA, and CPT blocks the release of the enzyme from DNA. Replication fork stalls at Topoisomerase-I (Top1) associated SSBs resulting in the formation of DSBs. Moreover, treatment with drugs like hydroxyurea (HU), an inhibitor of ribonucleotide reductase (RNR) and dNTP synthesis, will lead to stalled or collapsed replication forks followed by replication-coupled DSB (Shimizu et al., 2007). DNA interstrand crosslink (ICL) agents such as cisplatin and mitomycin C (MMC) covalently link two strands of DNA, thereby blocking vital DNA metabolism like replication. Upon the stalling of a replication fork at these lesions, a DSB is introduced (Scharer, 2005). Under these conditions, DSBs are more likely to be generated during S phase. When the DNA replication machinery encounters a replication-blocking lesion, the DNA polymerase stalls at the blocked site. This may result in the formation of a Y-shaped DNA structure if the blockage persists, which may be recognized by a specific endonuclease. This in

11 turn could produce a nick in the template strand resulting in the induction of a DSB near the replication-blocking lesion.

A fourth cause of DSBs is the inadvertent action of nuclear enzymes on DNA. These include failures of type II topoisomerases, which transiently break both strands of the duplex. If the topoisomerase fails to rejoin the strands, then a DSB results (Adachi et al., 2003). Drugs that inhibit topoisomerase function cause DSB, for example, doxorubicin which prevents topoisomerase II (Top2)-mediated DNA religation can promote DSBs (Hammond et al., 2003). Moreover, inadvertent action by nuclear enzymes in lymphoid cells, such as the recombination-activating gene (RAG) complex (composed of RAG1/2) and activation-induced deaminase (AID) are responsible for initiating physiologic breaks for antigen receptor gene rearrangement. However, they sometimes accidentally act at off-target sites outside the antigen receptor gene loci causing collateral damage (Mahowald et al., 2008). In humans, these account for about half of all of the chromosomal translocations that result in lymphoma. Nevertheless, during the course of normal biological events, such as homologous recombination during meiosis or in V(D)J recombination and immunoglobulin chain class switching or apoptotic DNA fragmentation, most DSBs are maintained in manageable condition.

Finally, physical or mechanical stress on the DNA duplex is a relevant cause of DSBs. In prokaryotes, this arises in the context of desiccation, which is quite important for the survival in the wild (Pitcher et al., 2007). In eukaryotes, telomere failures can result in chromosomal fusions that have two centromeres, and this results in physical stress by the mitotic spindle (breakage/fusion/bridge cycles) with DSBs (Murnane, 2006).

There are two distinct and complementary repair mechanisms for DSB repair, homologous recombination (HR) and non-homologous end joining (NHEJ). Briefly, Homologous recombination repair utilizes the intact homologous DNA

12 sequence, in most cases the sister chromatid, as a template for accurate repair (Krogh and Symington, 2004; San Filippo et al., 2008). Non-homologous end joining, on the other hand, ligates the two broken ends together with minimal processing at the site. It is more efficient but prone to generating mutations at the sites of DSBs. Furthermore, because there is no apparent mechanism to ensure that the two ends being joined were originally contiguous, NHEJ could produce chromosomal rearrangements such as inversions and translocations (Lieber, 2010).

1.2.4.2 DSB repair by Homologous Recombination

The molecular mechanisms and factors involved in HR were first discovered using genetic screens in the budding yeast. Many members of the group of genes involved in recombinational repair were first identified as mutants sensitive to ionizing radiation, so they all start with rad for radiation sensitivity. They include Rad50, Rad51, Rad52, Rad54, Rad55, Rad57, Rad59 Xrs2,Mre11, etc., and are referred to as the Rad52 epistasis group (Symington, 2002). These genes are highly conserved among eukaryotes, highlighting their importance for cell survival. The role of major yeast HR repair proteins in DSB repair is described below (See Table 1.1 for homologs of other organisms).

Rad50 is a 153kD protein serves as a subunit of the Mre11-Rad50-Xrs2 (MRX) complex. Rad50 is a highly conserved member of the family of Structural Maintenance of Chromosome (SMC) proteins (Strunnikov and Jessberger, 1999). Sharing a common feature of SMC family, Rad50 contains two globular ATPase domains at it N terminal and C terminal, linked by α-helical coiled-coil domain. Two Rad50 proteins can form a homodimer in the presence of ATP, creating a DNA-binding interface. Mre11 tends to bind to the base of the coiled-coil to form

the (Mre11/Rad50)2 hetero-tetramer, whose center is the DNA-binding interface. Within the coiled-coil region is a conserved zinc-binding Cys-x-x-Cys motif which

Table 1.1 Homologs of DSB repair factors in different species

S. Human E. Features cerevisiae coli Mre11 Mre11 SbcD Ø ssDNA endonuclease, 3’-5’ exonuclease Rad50 Rad50 SbcC Ø DNA binding and unwinding Xrs2 Nbs1 - Ø Structure-specific DNA end binding Rad51 hRad51 RecA Ø DNA identification HR sequence, pairing and strand exchange,ATPase Rad52 hRad52 RecO Ø Filament formation and ssDNA Annealing RPA RPA SSB Ø ssDNA binding - BRAC2 - Ø Rad51 loading Sae2 CtIP RecQ Ø Endonuclease involved in end processing Sgs1 BLM/WRN - Ø Helicase for end resection Exo1 Exo1 RecJ Ø 5'-3' exonuclease for end resection Yku70 Ku70 - Ø Form Ku70–Ku80 heterodimer facilitate NHEJ Yku80 Ku80 - and suppress other repair pathway Dnl4 Ligase IV - Ø ATP-dependent DNA ligase Lif1 XRCC4 - Ø Bind to Dnl4 and stimulate ligase activity Nej1 XLF - Ø Similar to Lif1 - DNA-PK - Ø Activated upon binding with Ku/DNA complex Pol4 Polm/l - Ø Polymerase for gap filling ? Artemis Ø Nuclease participates in end processing

14 functions as a hook to mediate intermolecular interaction between two Rad50 molecules through the coiled-coil region (Hopfner et al., 2002). It is commonly believed that the function of Rad50 is DNA binding which can bring the ends of DSBs together and facilitate Mre11 nuclease activity.

Mre11 is 78kD subunit of MRX complex that bears nuclease activities including 3’-5’ exonuclease activity, endonuclease activity on circular and linear ssDNA and endonuclease cleavage of hairpin ends or 3’ssDNA overhangs(Moreau et al., 1999). The nuclease activity of Mre11 is manganese-dependent and localized to the N-terminus of the protein (Paull and Gellert, 1998). The N-terminal portion of Mre11, which contains motifs shared by the phosphoesterase family, is also important for maintaining interaction with the other MRX complex subunits, Xrs2 and Rad50 (Johzuka and Ogawa, 1995). The C-terminus is responsible for mediating DNA binding to both single- and double-stranded DNA in a structure- and sequence-specific manner.

Xrs2 is another subunit of the MRX complex whose homolog in vertebrates is Nbs1, named after “Nijmegen breakage syndrome”. Xrs2 binds DNA in a structure-specific manner and has been shown to be important for targeting the MRX complex to DNA ends (Featherstone and Jackson, 1998). The C-terminal domain contains both Mre11 and Tel1 binding sites that mediate translocation of Mre11 to the nucleus and Tel1 phosphorylation of Xrs2. Xrs2 is also able to stimulate Mre11 exonuclease activity and facilitates MRX complex association with DNA ligase (D'Amours and Jackson, 2001).

The Mre11/Rad50/Xrs2 (MRX) is a stable complex with a predicted stoichiometry of 2:2:1, however, Rad50 and Xrs2 are believed to interact through Mre11. Functions of this complex include DNA binding, exonuclease and endonuclease activities, DNA unwinding, and DNA end recognition (Grenon et al., 2001; Lee et al., 1998). In addition to the repair functions listed above which are mostly

15 involved in homologous recombination, the MRX complex also facilitates DSB repair via nonhomologous end-joining as well as introduction of DSBs in meiosis, detection of damaged DNA, DNA damage checkpoint activation, telomerase recruitment, and suppression of gross chromosomal rearrangements.

Rad51 is a 43-kD protein and shares a 30% homology with E. coli RecA protein and 59% homology with human and mouse homologs. Rad51 is highly conserved in a wide variety of eukaryotic organisms from yeast to humans, reflecting its significance in homologous recombination. Like the RecA protein, Rad51 has a catalytic ability of nucleotide binding and hydrolysis. Rad51 binds with higher affinity to DNA duplexes with ss-tails than to duplex or single stranded oligonucleotides. After binding, Rad51 forms right-handed helical nucleoprotein DNA filaments and then activates its ATPase activity (Shin et al., 2003). Certain mediator proteins are required for Rad51 filament assembly, including Rad52, Rad55/Rad57 and Rad54, whereas RPA has to be displaced from the filament before strand exchange can occur (Kiianitsa et al., 2002). Once assembled, Rad51 nucleoprotein filament is capable of interacting with a second DNA molecule (single or double stranded) to initiate strand exchange. Rad51 along with Rad52 may be also responsible for homology search.

RPA is a ssDNA binding protein with high ssDNA binding affinity and provides stability to the unwound DNA. The RPA complex consists of three subunits of 70, 34 and 14kD, which are encoded by RFA1, RFA2, RFA3 genes respectively (Brill and Stillman, 1991). After resection of 5’ nucleotides at a DSB, the 3’ ssDNA extension is immediately coated by RPA proteins. The primary role of RPA in HR is to remove secondary structure in ssDNA in order to allow more efficient binding by Rad51.

Rad52 is a protein of 504 amino acids, within which the N-terminal 200 residues share homology with other member of Rad52 epistasis group. The less conserved C-terminals of Rad52 contains the Rad51 interaction region. The

16 interaction between Rad52 and Rad51 is thought to stimulate Rad51 protein- mediated DNA strand exchange. Both yeast and human Rad52 are multimeric and electron microscopy studies show they can form a ring structure with a central channel (Ranatunga et al., 2001). The central channel is thought to be a potential DNA binding site. In vitro experiments indicate Rad52 promotes annealing of complementary ssDNA, which is stimulated by RPA. Rad52 tends to preferentially bind to the ends of ssDNA of tailed duplex molecules, especially RPA coated ssDNA (Mortensen et al., 1996). Additionally, in strains lacking RAD52, ssDNA is quickly resected suggesting that Rad52 also plays a structural role in protecting DNA strands form exonucleolytic degradation. Rad52 protein alone cannot displace RPA from ssDNA. So, the presumptive function of Rad52 is to facilitate the formation of Rad51-ssDNA nucleoprotein filaments by recruiting Rad51 protein while forming a co-complex with the RPA-ssDNA structure.

Another recombination factor Rad59 is believed to share some functions of Rad52 and provide assistance for strand invasion (Symington, 2002). Rad59 is a 26-kDa protein with homology to the N-terminus of Rad52, which includes the self-interaction and DNA-binding domains of Rad52. It was identified in a screen for Rad51-independent spontaneous mitotic recombination (Bai and Symington, 1996). RAD59 is important for SSA between chromosomal direct repeats, especially as the repeat length decreases (Sugawara et al., 2000). Overexpression of Rad52 suppresses radiation sensitivity of rad59 mutant, however, Rad59 can neither substitute for the annealing function of Rad52 or Rad51-independent recombination (Krogh and Symington, 2004).

Rad54 belongs to the Swi/Snf protein family whose members are DNA- dependent ATPases capable of chromatin remodeling (Eisen et al., 1995). The dsDNA dependent ATPase of Rad54 is believed to be necessary for homologous DNA pairing and chromatin remodeling during Rad51-ssDNA nucleoprotein filament strand invasion of dsDNA. Rad54 has been demonstrated

17 to translocate along duplex DNA and redistribute the associated nucleosomes. Rad54 also mediates an alteration in duplex DNA conformation that results in a DNA linking number change. In addition, Rad54 facilitates Rad51 binding to ssDNA, stabilizes Rad51 nucleoprotein complexes, and stimulates Rad51- mediated D-loop formation (Klein, 1997).

Rad55 and Rad57 can form a heterodimer that functions as mediator in strand exchange. Both of them share some homology to RecA and Rad51, particularly in the sequence motifs involved in the binding of nucleoside triphosphates (Sung, 1997). Accordingly, they all have ssDNA binding ability. Like Rad52, the concentration of Rad55/Rad57 complex in cell is much lower than Rad51, but it is sufficient for them to carry out a mediator function, which is presumably different from Rad52.

Although the roles of individual HR repair factors is far from being fully understood, numerous sophisticated genetic and biochemical studies do provide an overall view of the HR mechanism (Barzel and Kupiec, 2008; Wyman and Kanaar, 2004)(San Filippo et al., 2008). The current model of homologous recombination is illustrated in Fig 1.2 (Barzel and Kupiec, 2008). The elaborate process of HR is initiated by processing the broken ends of DNA at the site of a DSB to yield 3’-single strand overhangs, which invades a homologous double strand. The end-processing starts with the recognition of the DSB by the MRX complex and is followed by the recruitment of the endonuclease Sae2 (mammalian ortholog CtIP) by MRX complex. More extensive processing requires exonuclease Exo1, Dna2 and helicase Sgs1 (Huertas, 2010). It has been shown that BRCA1 (breast cancer type 1), the protein of breast cancer susceptible gene, is also involved in end resection by forming a complex with MRN(Mre11/Rad50/Nbs1) and CtIP in mammalian cells, although its exact role remains unclear (Chen et al., 2008). DNA ends resected by Exo1 and Sgs1 are a fundamental step of HR because it provides long stretches of ssDNA for the subsequent steps. The 3’ssDNA overhangs exposed after end processing is

18 bound and protected immediately by RPA proteins. Rad51 is then recruited to form nucleoprotein filaments by replacing RPA. In mammalian cells, the recruitment of Rad51 is facilitated by breast cancer susceptibility protein BRCA2 (breast cancer type 2), which is a large protein that binds Rad51 through a series of conserved BRCT (Breast Cancer Gene 1 carboxyl-terminal) repeat motifs (Wong et al., 1997). The BRCT repeat is believed to stimulate Rad51 nucleoprotein filament formation on ssDNA in the presence of ATP (Carreira et al., 2009). In S. cerevisiae, BRCA2 is absent but other proteins such as Rad52 function as an alternative for loading Rad51 onto ssDNA.

Once the Rad51 nucleoprotein filament is formed, it catalyzes the strand exchange reaction in which ssDNA invades homologous duplex DNA forming a D-loop structure. The invading 3' ends serve as primer for DNA synthesis. The invasion process has been well illustrated by the crystal structure of recombination-deficient A (RecA) filament in E. Coli, and is ATP-dependent (Chen et al., 2008b). Based on RecA studies, it is expected that the homology search process occurs by way of random collisions between the presynaptic filament and the duplex molecule (Bianco et al., 1998). RecA-bound ssDNA is stretched globally but maintains a B-DNA like conformation locally in base triplets. This unusual structure favors Watson-Crick type base pairing during homology sampling with the complementary strand in a destabilized donor duplex (Kass and Jasin, 2010). In addition, as mentioned above, Rad54 is a multi-functional factor that is involved in opening the chromatin configuration to allow strand invasion, dissociating Rad51 and migration of the branch point.

After the formation of D-loop and DNA synthesis dependent extension of the invading strand, there are two distinct pathways for DNA synthesis and branch resolution. For DSBs in mitotic cells, the main pathway used is called synthesis- dependent strand annealing (SDSA). In the SDSA pathway, the D-loop is unwound and the elongated (repaired) ssDNA strand is released from the loop.

Figure1.2 Scheme for homologous recombination during DSB repair. Figure is adapted from (Barzel and Kupiec, 2008). See text for details.

20 Then, it subsequently anneals with the complementary ssDNA strand from the other end of break where the DSB originally occurred. The reaction is completed by gap-filling DNA synthesis and ligation. Only non-crossover products are formed in this pathway. On the other hand, if the other end of the break is also captured along with gap-filling DNA synthesis and ligation, it will result in two crossed DNA strands or Holliday junctions. This pathway is called the double- strand break-repair (DSBR), and despite its name, it can explain much of the meiotic segregation in fungi and links crossing over and gene conversion as different outcomes of DSB repair (San Filippo et al., 2008). The resolution of Holliday junctions is carried out by specialized endonucleases, such as human GEN1 (yeast Yen1) and SLX1/SLX4 (Klein and Symington, 2009) and can result in either non-crossover or crossover products.

Sometimes a DSB is closely flanked by direct repeats. This DNA organization provides the opportunity to repair the DSB by a deletion process using the repeated DNA sequences, called single-strand annealing (SSA). In the SSA pathway, the DSB ends are resected, but then instead of engaging in a homologous DNA sequence search for strand invasion, the resected ends anneal to each other. The process is finished by nucleolytic removal of the protruding single-stranded tails and results in deletion of the sequences between the direct repeats and also one (or more) of the intervening repeats (Prado et al., 2003).

Some DSBs, such as those that can occur at telomeres or at broken replication forks, are single-ended (Hackett and Greider, 2003). These too can participate in HR, through a single-ended invasion process called break-induced replication (BIR). In BIR, the DSB end is also processed similar to the resection that occurs in other recombination at repair pathways (Davis and Symington, 2004; Henson et al., 2002; McEachern and Haber, 2006). The single-stranded tail then invades a homologous DNA sequence, often the sister chromatid or the homolog chromosome, but sometimes a repeated sequence on a different chromosome.

21 The invading end is used to copy information from the invaded donor chromosome by DNA synthesis. When the sister chromatid or homolog chromosome is used, the repair is accurate. When a repeated sequence on a non-homologous chromosome is engaged to initiate repair, the result is a nonreciprocal translocation. Most BIR events are dependent on the HR factors used in DSBR and SDSA, but a small fraction can occur independently of these factors, including Rad51. BIR is often used to repair broken or shortened telomeres (Lydeard et al., 2007).

1.2.4.3 DSB repair by Non-Homologous End Joining

While HR appears to be the more accurate method of repair because genetic information is faithfully copied from an intact homologous sequence, it also requires more effort to do so. NHEJ is an alternative DSB repair process requiring less processing. NHEJ specific repair factors were identified and characterized by assessing DSB repair under conditions where homologous recombination is not possible. This was accomplished by using a classic method of analyzing the rejoining of broken DNA ends that do not share any homology in the genome of the cell and so recombination is not readily available (Boulton and Jackson, 1996; Schar et al., 1997). The most common NHEJ assay is to transform exogenous linearized plasmid which requires recircularization by NHEJ in vivo due to lack of any homologous sequence in the cells. Major NHEJ factors include Ku70/80, Dnl4/Lif1/Nej1 (DNA ligaseIV/XRCC4/XLF in higher eukaryotes), DNA-dependent protein kinase (higher eukaryotes only) (See Table 1.1 for homologs in other organisms).

Ku70 and Ku80 are subunits of the Ku heterodimer that are named after their molecular mass (70kD and 83kD respectively). Ku is an abundant nuclear protein complex that binds with high affinity to duplex DNA ends independent of

22 the end sequence or precise structure (Mimori and Hardin, 1986). Once bound to double-stranded DNA (dsDNA), Ku can translocate along the molecule in a manner that does not require ATP, allowing multiple subunits of Ku to bind to linear DNA (de Vries et al., 1989). The observed crystal structure of Ku indicates that there is a beta-barrel ring structure in the center which may serve as the end binding site (Walker et al., 2001). Binding of Ku70/80 to a DSB presumably protects the broken DNA ends from nucleolytic degradation and marks the damaged site in order to continue future processing. Once bound to DNA, Ku can interact with MRX complex to form an end bridging complex. Then, Ku may stabilize ends and recruit Dnl4/Lif1 to the break site (Downs and Jackson, 2004). Besides NHEJ, Ku heterodimer also participates in telomere maintenance pathways (Hediger et al., 2002).

Dnl4 is an ATP-dependent DNA ligase (mammalian homolog is DNA ligase IV) which physically interacts with Lif1 in vivo. So Dnl4’s main function is to fulfill the last step of NHEJ by connecting both ends of the break. In the presence of Lif1 and Nej1, Dnl4 is capable of ligating not only nicks, but even compatible (4-nt overhang) ends, as well as ends with 2bp microhomology, 1nt gaps. With the addition of Ku, it can ligate incompatible DNA ends (Gu et al., 2007). Lif1 does not have any enzymatic ability but is required for stability and activity of the DNA ligase. It can bind to DNA cooperatively and target Dnl4 to repair site (Grawunder et al., 1998). Dnl4, Lif1 and Nej1 form a complex with stoichiometry of 1:2:2 as assayed by gel filtration experiments (Ahnesorg et al., 2006). Recently, Dnl4/Lif1 complex has been shown to not only have an essential structural role in the assembly of a functional NHEJ complex on DNA ends, but also acts in combination with Ku to suppress HR. Ku purified from both S. cerevisiae and mammalian cells is able to improve the binding of Dnl4/Lif at DNA ends.

DNA-PKcs (DNA-dependent protein kinase catalytic subunit) is a key player in the mammalian NHEJ pathway, and with a molecular weight of around 470kD, it

23 is the largest kinase in the cell. DNA-PKcs is recruited by Ku proteins and is specifically activated by binding to duplex DNA ends of a wide variety of end configurations (Lieber, 2010). Activated DNA-PK is a serine/threonine kinase with S/TQ site specificity, which phosphorylates other proteins as well as itself. DNA-PK can possibly tether the broken ends to facilitate rejoining and recruits as well as activates proteins for end processing and ligation (Chen et al., 2005). Although yeast lacks a homolog of DNA-PKcs, there is compelling genetic and biochemical evidence that the MRX complex is the end processing factor in yeast NHEJ (Chen et al., 2001). The MRX (Mre11/Rad50/Xrs2) complex is clearly required for NHEJ since mutants of mre11, rad50 and xrs2 are as defective in plasmid end joining as ku70 and ku80 mutants. The real function of MRX in NHEJ is not clear so far, nevertheless, the nuclease activity of Mre11 is probably required for end alignment and end processing at the DSB site. Rad50 might function through its zinc-binding hook to hold broken DNA ends together. Since Xrs2 has been found to interact with Lif1, it might function to recruit Lif1 to the ends held together by the MRX complex(Lewis and Resnick, 2000).

In contrast to HR, the overall mechanism of NHEJ is much simpler. The current model of NHEJ is illustrated in Fig 1.3 (Downs et al., 2007). Briefly, the ring-like heterodimer Ku70/Ku80 binds to DSB ends and interacts with DNA-PKcs (in mammalian cells). Together, they function to protect and synapse the two ends. Then the DNA ends are joined by the Dnl4/Lif1 complexes (DNA ligase 4/XRCC4 complex in mammals). Despite the straightforward mechanism, most interesting part of NHEJ lies in its unique capability of rejoining highly variable DNA ends with many different structures. This pathway makes use of a number of processing steps that include cleavage and gap filling before ligation. In the mammalian system, the nuclease Artemis is recruited via its interaction with DNA-PKcs. Together, the Artemis/DNA-PKcs complex is able to cleave a variety of damaged DNA overhangs. Cleavage of DNA ends that leaves DNA gaps is

Figure 1.3 Scheme for Non-Homologous End Joining during DSB repair in budding yeast. Figure is adapted from (Aylon and Kupiec, 2004). Following DSB creation (A), the Ku heterodimer is recruited to the broken ends (B). The Mre11/Rad50/Xrs2 (MRX) complex is then recruited (C). Both complexes may play roles in holding the broken ends together and participate in end-processing. The Lif1/Dnl4 complex is then recruited. The MRX and Lif1/Dnl4 complexes promote activity of Lif1/Dnl4, resulting in ligation of the broken ends.

25 followed by gap filling by PolX family polymerase such as polymerase μ and polymerase λ. They are believed to interact with Ku/DNA intermediate via their BRCT domain (Ma et al., 2005). In general, diversity of end processing arises from the following variations: (1) Loading flexibility of different polymerases, nucleases and ligases, such as a variable loading sequence or repetitive loading of components; (2)Independent processing of individual break ends as opposed to identical processing of both ends; (3) Different extent of end processing, for example, a range of 0 to 25 base pairs nucleolytic resection is possible, as is the nucleotide addition reaction (Gauss and Lieber, 1996); (4) Last but not least, the biochemical activities of each of the enzymes (polymerases, nucleases and ligases) itself offer great variations (Lieber, 2010). This highly flexible mechanism is essential to permit NHEJ to handle a very diverse array of DSB end configurations for rejoining. On the other hand, processing of DNA ends prior to joining by these multiple steps can lead to unpredictable deletions and insertions accounting for the more error-prone nature of repair by NHEJ compared to HR.

Besides the canonical NHEJ pathway described above, end rejoining also occurs when some NHEJ factors are not available. This is a distinct pathway termed alternative NHEJ (alt-NHEJ), in which ligase IV, Ku or DNA-PKcs is not present, whereas the end joining is relatively successful (Weinstock et al., 2007; Yan et al., 2007). Although this mechanism is not fully understood, in most cases, substantial terminal microhomology seems to be required to help align the broken stands of DNA and a number of other enzymes are involved in this process (Lieber, 2010; Zhang and Jasin, 2011).

1.3 Choice of DSB repair pathway: HR vs. NHEJ

While both HR and NHEJ are effective repair mechanisms for recovery from a potentially lethal DSB, they compete with each other in the cells. Undoubtedly, the choice and balance between these two pathways is critical for the

26 maintenance of genomic integrity. The choice between HR and NHEJ differs widely among different species, varies with the nature of the DSB and its occurrence during different stages of the cell cycle, as elaborated below.

1.3.1 Nature of DNA DSB breaks

DSBs are induced by a variety of treatments that differ in the amount as well the “quality” of the break ends. DSBs can be produced by ionizing radiation, X-ray machines or lasers, which also cause other types of lesion including SSBs, sugar damage and base modifications, etc. (Lukas et al., 2005; Walter et al., 2003). DSBs generated by these methods are considered having “dirty” ends, which often can not be ligated directly. These ends have different structural requirements that need to be processed specifically before the actual repair can take place. On the other hand, restriction enzymes such I-SceI and HO induce DSBs with complementary single-strand overhangs that can be easily ligated and do not require otherwise essential steps for processing authentic damage from physical sources such as radiation. These ends are considered as “clean”. For example, both IR-induced HR and meiotic HR are markedly reduced in mre11 mutants(Moreau et al., 1999). In contrast, mre11 mutants show relatively mild (less than 2-fold) defects in HR stimulated by “clean” DSB produced by HO endonuclease (Krishna et al., 2007). The difference indicates that the lower level of IR-induced HR in mre11 mutants represent defects in end processing rather than a recombination defect. Indeed, Mre11 is required for the repair of I-SceI endonuclease induced DSBs via NHEJ repair and its exonuclease activity essential (Zhuang et al., 2009). When HR is blocked and yeast cells with mre11 mutation are forced to repair nuclease derived DSBs by NHEJ, the repair efficiency decreases by ~100 fold (Moore and Haber, 1996).

Radio-sensitivity of HR deficient mutants is at times comparable to that of NHEJ deficient cells. In chicken DT40 cells, HR is preferentially used over NHEJ in G2

27 phase as shown by the hypersensitivity of rad54 cells but not ku70 cells to X-rays (Sonoda et al., 2006). Thus, irrespective of other factors that might influence the choice of a DSB repair pathway, it is quite likely that dirty ends may need extensive processing and may be poor substrates for NHEJ, whereas clean ends are readily ligated through fast NHEJ. However, this does not mean that dirty ends are better repaired by HR.

DNA breaks occurring indirectly as a result of damage or discontinuity during replication are clearly different from those mentioned in previous paragraph. They are unconventional DSBs, that often have one free end (Wyman and Kanaar, 2006). Yeast HR mutants as well as chicken DT40 mutants in the RAD52 epistasis group are highly sensitive to Camptothecin (CPT), a topoisomerase I (Top1) inhibitor that causes stalled replication forks and subsequently DSB (Pommier et al., 2003). In contrast, mutants lacking either of the NHEJ factors, namely Ku70, DNA-PKcs, or DNA ligase IV, are resistant to killing by CPT (Adachi et al., 2004). This is simply a reflection of the nature of the break ends. In the replication block scenario, NHEJ simply cannot perform effective repair because there is only one free end. On the other hand, if a break is created directly and both ends are in close proximity, this would favor their repair by NHEJ. Moreover, recent research has shown a detrimental effect of Ku proteins and other NHEJ factors in the Fanconi anemia (FA) repair pathway which is responsible for the repair of lesion caused by DNA crosslinking agents (Adamo et al., 2010; Pace et al., 2010). In mitotic cells, it has been shown that DNA repair defects of C. elegans FANC mutants and FA-deficient human cells are significantly suppressed by eliminating NHEJ (Adamo et al., 2010). Loss of Ku in FANC mutant cells greatly relieved their cisplatin sensitivity (Pace et al., 2010). These findings indicate a potential function for the FA pathway in processing DNA ends, thereby diverting double-strand break repair away from abortive NHEJ and toward homologous recombination; in other words, NHEJ is sometimes not only undesirable, but can also be deleterious.

28 1.3.2 Effect of cell cycle regulation on the choice of DSB repair pathway

A more comprehensive understanding of the choice between HR/NHEJ is from the view point of cell cycle regulation and end resection. NHEJ substrates are double-stranded ends with limited processing whereas HR substrates are 3’ single-stranded tails produced by extensive 5’-end resection. This 5’-ended DSB resection is irreversible, which makes it a good stage to commit to the repair pathway that channels DSB repair to HR (Frank-Vaillant and Marcand, 2002). DSB-end resection has been demonstrated to be tightly regulated during the cell cycle.

G1-arrested cells fail to perform efficient DSB-induced HR, mostly due to inefficient end resection; on the other hand, they exhibit an increased efficiency of non-homologous end joining (Ira et al., 2004). In G1 cells, few RPA and Rad52 foci are observed after lower dose of IR irradiation (Lisby et al., 2004). Meanwhile, increased IR sensitivity is detected in Ku deficient cells (Takata et al., 1998). These findings clearly implicate that cells favor NHEJ above HR in G1 phase, when homologous sequences are not readily available. DSB-end resection is active in S and G2 phases, thus activating HR. The biological significance for this up regulation of resection is related to the presence of a sister chromatid only during S and G2. The sister chromatid is the preferred HR template because it promotes an error-free repair, while use of the homologous chromosome or an ectopic region can compromise genome stability. Cohesins are among the factors that determine the choice of the sister as the main template to repair DSBs by HR in S and G2. After DSB formation γH2A.X recruits cohesins de novo, thus maintaining the two sister chromatids in close proximity to facilitate sister- chromatid recombination (Strom et al., 2004). Consistently, cohesins have been observed to co-localize with sites of IR-induced DNA damage in mammals (Kim et al., 2002). In addition, as pointed out before, HR appears to be the predominant DSB repair for stalled or collapsed replication forks during S phase.

29 Cdc28, the budding yeast CDK that is active in S and G2 phases, has been shown to promote DSB resection, channeling repair to HR. Several reports have demonstrated that endonuclease Sae2 (ortholog of human CtIP and S. pombe Ctp1) is a key regulator of the DSB-repair pathway choice at the molecular level. Ctp1/CtIP amounts are low in G1 and peak during S phase in fission yeast and humans (Limbo et al., 2007; Yu and Baer, 2000). Human CtIP has been shown to functionally interact with the MRN complex and regulate DSB resection in mammalian cells (Sartori et al., 2007). Human CtIP, as well as yeast Sae2, is also a target of the checkpoint transducer kinases. Phosphorylated CtIP interacts with several factors in human cells, such as BRCA1 (Li et al., 2000). BRCA1 is recruited to DNA damage sites by MRN and can form a complex with MRN and CtIP in S and G2. BRCA1-mediated recruitment of CtIP to DNA damage sites would be important to facilitate DSB resection and subsequent HR repair. An important role of end resection is the generation of RPA coated ssDNA, which acts as a crucial recognition signal for ATR activation and initiates cell cycle checkpoint signaling (Zou and Elledge, 2003). By doing so, it appears that the end processing initiates a feedback regulation by coupling DSB repair with cell cycle arrest, so that cells can ensure that repair is finished before the cell divides.

Apart from end resection, HR is also regulated during the cell cycle through HR factor recruitment. BRCA2 modulates the assembly of Rad51-filament after resection by interacting with and stabilizing of Rad51. It has been shown that BRAC2 is phosphorylated by CDKs and dissociates from Rad51 outside of S phase. In other words, BRAC2 serves as S phase facilitator of homologous recombination by recruiting and stabilizing Rad51.

1.3.3 NHEJ factor’s binding to the break ends

An enhanced rate of end resection has been demonstrated in yeast Ku mutants using direct molecular analysis (Lee et al., 1998). Since Ku binds DNA ends, it presumably physically blocks access of the end resection machinery. More

30 recently, the homolog of DNA ligase IV/XRCC4 in yeast, Dnl4-Lif1, has also been shown to have an inhibitory effect on end resection, possibly by stabilizing Ku at DNA ends, although the effect is somewhat less pronounced than with Ku (Zhang et al., 2007). Moreover, in chicken cells, Ku mutation actually leads to increased resistance to ionizing radiation during late S/G2, both in otherwise wild-type cells as well as in DNA-PKcs mutant cells this has been interpreted as Ku dependent interference of HR during these phases of the cell cycle (Fukushima et al., 2001). On the other hand, the binding of Ku proteins to the breaks protects the ends from irreversible resection that leads to HR repair. In most cases, this mechanism acts prior to HR repair, which could possibly be an attempt to repair by NHEJ before HR starts its function (Kim et al., 2005). This means that cells are provided with repair options in a stepwise manner, with NHEJ usually preceding HR, but with potential for collaboration between HR and NHEJ as well. Evidence from double mutant analysis has shown that loss of one HR and one NHEJ factor results in more severe phenotype than loss of any single factor. Mice with mutations in both Rad54 and Ku80 show seriously decreased survival and extreme sensitivity to IR compared to single mutants (Couedel et al., 2004). This “double insurance” provided by NHEJ and HR pathways in the repair of a DSB is of great benefit for cells in maintaining genomic integrity.

1.3.4 DSB repair pathway choice in different species

The choice of NHEJ and HR varies greatly between budding yeast and mammalian cells. In yeast, HR plays a dominant role in DSB repair following ionizing radiation in the G1 as well as G2 phase (Aylon and Kupiec, 2004). In contrast, NHEJ repairs over 60% of exogenously induced DSBs in mouse ES cells (Liang et al., 1998). Furthermore, genes involved in HR, e.g. RAD51 or RAD54, are not expressed in resting mammalian cells even after exogenous genotoxic stresses (Tan et al., 1999). Thus, it seems that NHEJ predominates

31 under most conditions in vertebrates. It is not fully understood why there is different preference for DSB repair pathways between yeast and higher eukaryotes. One reason could be that yeast lacks three important NHEJ factors that are present in mammalian cells: DNA-PKcs, Artemis and PNK (Polynucleotide Kinase). DNA-PKcs could be the key missing protein because it facilitates alignment of non-complementary ends and regulates end-processing during NHEJ (Budman and Chu, 2005). Artemis has DNA-PK dependent endonuclease activity on DNA hairpin structures, and DNA-PK dependent endonuclease processing of single-strand overhangs, with preferential cleavage at the dsDNA/ssDNA junction (Pawelczak and Turchi, 2010). ATM hyper- phosphorylates Artemis following IR treatment and Artemis nuclease activity is required for DSB rejoining (Riballo et al., 2004). Mammalian PNK is both a kinase for adding a phosphate to 5’OH and a phosphatase for removing 3’ phosphate groups, which are found in some oxidative damage or upon end processing. PNK creates 5'-phosphate/3'-hydroxyl termini, which are a necessary prerequisite for ligation during repair. PNK is recruited to XRCC4 through interactions between its N-terminal FHA (Forkhead-associated) domain (Bernstein et al., 2005).

The greater use of NHEJ in higher eukaryotes could be related to their large genome size, low gene density and extensively condensed chromosomes. Yeast genome size (12Mbp) is much smaller than the human genome (3.2Gbp). Compared to yeast genome, whose coding regions comprises more than 70% of the whole sequence, the coding sequences in mammalian genome only accounts for less than 3%. The more complex and condensed genomes of mammalian cells not only presents a great challenge of searching and locating homologous template for homologous recombination, but also provides much higher tolerance of small scale deletions or insertions introduced by NHEJ (Stark and Jasin, 2003). This is supported by the observation that gene targeting is inefficient in cells of higher eukaryotes, while random integration very efficient and more frequent than targeted integration by over three orders of magnitude (Fattah et al., 2008). The

32 chances of deletions and insertions altering coding sequences in a mammalian cell are much lower than that in yeast. In addition, in budding yeast, silent heterochromatin only occurs at the silent mating type loci, rDNA repeats and subtelomeric regions (Oberdoerffer and Sinclair, 2007). However, in higher eukaryotes, the abundance and complexity of heterochromatin and higher order structure pose greater challenge for HR compared to NHEJ (Misteli and Soutoglou, 2009; Woodcock and Ghosh, 2010). Additional discussion of how chromatin structure and components affect choice of HR and NHEJ will be described in the following chapters.

1.4 DNA Damage Signaling in Budding Yeast

In order to detect and transmit the DNA damage signal, cells have evolved a sophisticated DNA damage checkpoint system to coordinate various repair related activities. Activation of this pathway results in a number of downstream biological consequences, including cell cycle arrest, transcription regulation, efficient DNA damage repair or even apoptosis if the repair becomes overwhelming. In short, similar to all signal transduction pathways, the DNA damage signaling pathway consists with three major components: sensors, transducers/mediators and effectors (Harper and Elledge, 2007). The sensors monitor DNA for structural abnormalities and then initiate the checkpoint signal. Transducers/ mediators further transmit and amplify this signal. Effectors control the biological consequences of triggering this pathway. The DNA damage checkpoint pathways also share with other signal transduction pathways the phenomenon of downregulation of the signal, or adaptation, in the continuous presence of the initiating signal (Figure1.4) (D'Amours and Jackson, 2002). Checkpoint proteins are well conserved from yeast to human cells, indicating

33 that the basic functions of these pathways have been preserved throughout evolution.

Here, I will primarily refer to the factors that are present in the budding yeast, although their homologs have been clearly identified in other species (see Table 1.2 for comparison). Undoubtedly, PI3(phosphatidylinositol 3’)-kinases Mec1 and Tel1 are two key players in the DNA damage signaling(Harrison and Haber, 2006). Mec1, the central DNA damage response kinase in the budding yeast belongs to the conserved family of PI3-kinases and is related to fission yeast Rad3, and human ATR (Ataxia Telangiectasia-Rad3-related) (Lowndes and Murguia, 2000). A Mec1 related but non-essential yeast PI3-kinase known as Tel1 is homologous to human ATM (Ataxia Telangiectasia Mutated) and carries out numerous functions similar to Mec1 (Sanchez et al., 1996). In the budding yeast, the initial lesion may be processed by the MRX (Mre11, Rad50 and Xrs2) complex (D'Amours and Jackson, 2002). MRX complex acts as damage sensor and recruits Tel1 to damaged DNA ends, which can then phosphorylate histone H2A in chromatin to create an amplified damage signal. The ssDNA generated by MRX and other nucleases binds to RPA proteins and recruits another kinase Mec1 along with its regulatory partner Ddc2. Mec1 then recruits and activates Rad24 containing RFC (Replication Factor C)-like clamp loader to load the PCNA like repair clamp complex Rad17-Mec3-Ddc1 complex (homologous to the mammalian 9-1-1 complex) onto DNA (Harper and Elledge, 2007). Activated Mec1/Tel1 phosphorylates master effector kinase Rad53 and this leads to a dramatic increase its kinase activity. Another checkpoint mediator Rad9 undergoes Mec1 dependent phosphorylation in response to DNA damage and binds to Rad53. This binding of phosphorylated Rad9 may facilitate the hyperphosphorylation and activation of Rad53 during early stages of the DNA damage response (Gilbert et al., 2001). Rad53, the main transducer kinase in the budding yeast DNA damage response pathway is homologous to the fission yeast Cds1 and human CHK2 tumor suppressor. A Rad53-related transducer

34 kinase Chk1 plays a minor role in the budding yeast DNA damage checkpoint by acting to prevent the degradation of the anaphase inhibitor Pds1, thus arresting the cells at G2/M (Sanchez et al., 1999). Upon activation in response to DNA damage, Rad53 triggers a phosphorylation mediated cascade of activities such as cell cycle arrest, activation of downstream effectors, expression of DNA repair genes and promotes efficient DNA repair. Mec1 and Rad53 protect cells against DNA damage via a number of other functions. DNA damage during S-phase slows down the rates of replication fork elongation independently of checkpoint kinases (Tercero and Diffley, 2001) and triggers a Rad53/Mec1-dependent block in the firing of late origins (Santocanale and Diffley, 1998). As a result, DNA damage leads to an abrupt decrease in DNA synthesis (Paulovich and Hartwell, 1995). In addition, Mec1 and Rad53 are required to prevent both spontaneous and DNA damage-induced collapse of replication forks (Tercero and Diffley, 2001), presumably through their ability to phosphorylate proteins at stalled replication forks.

Interestingly, Mec1 and Rad53 are essential for viability in the budding yeast, while Tel1 and Chk1 are dispensable. In mammals, ATR and CHK1 are essential for viability, while ATM and CHK2 are non-essential. The essential function of Mec1 and Rad53 in the budding yeast is to promote deoxyribonucleotide triphosphate (dNTP) production during S-phase to coincide with DNA replication. This is achieved via phosphorylation and degradation of Sml1 (Zhao et al., 2001), a stoichiometric inhibitor of ribonucleotide reductase (Rnr). The lethality, but not the DNA damage sensitivity of mec1 and rad53 mutants is rescued by elevating dNTP levels through disruption of the sml1 gene or overexpression of Rnr large subunits (Desany et al., 1998; Zhao et al., 2001). The essential functions of ATR and CHK1 are not clear at present, but they are likely to be related to their functions in responding to endogenous replication stress.

Figure 1.4 The DNA damage response pathway in budding yeast. Figure is adapted from (D'Amours and Jackson, 2002). See text for more details.

36 Table 1.2 Homologs of DNA damage checkpoint proteins in different species

S. cerevisiae Human S. pombe Features Mec1 ATR Rad3 Ø PIKK Tel1 ATM Tel1 Ø PIKK Lcd1/Ddc2 ATRIP Rad26 Ø PIKK subuint Mre11/Rad50/Xrs2 MRE11/RAD50/NBS1 Mre11/Rad50/Nbs1 Ø MRX complex Rad24 RAD17 Rad17 Ø RFC-like clamp loader Ddc1/Rad17/Mec3 RAD9/RAD1/HUS1 Rad9/Rad1/Hus1 Ø PCNA-like clamp Rad53 CHK2 Cds1 Ø Effector kinase Chk1 CHK1 Chk1 Ø Effector kinase Rad9 53BP1 or MDC1 or Crb2 Ø Mediator BRCA1

37 CHAPTER 2

THE ROLE OF HISTONES, CHROMATIN STRUCTURE, AND EPIGENETIC MODIFICATIONS IN DNA REPAIR

2.1 Histones Serve as Primary Protein Components of Chromatin

In eukaryotes, chromosomes consist of long genomic DNA molecules packaged in association with proteins. Chromosomes are 50,000 times more compact than the extended form of DNA. Histones are the main packaging protein component of chromosomes. Histones were discovered over a century ago (in 1884) by German scientist Albrecht Kossel while processing of acid extracts of bird erythrocyte nuclei (Kossel, 1911). Histones were found to be the most abundant proteins in the nucleus, with extremely high binding affinity for nucleus acids. Numerous studies over time have suggested that histones are crucial proteins because they not only package the DNA into the relatively small nucleus, but also regulate access to genetic information, thereby modulating processes such as replication, transcription, recombination, DNA damage repair and cell division (Wu and Grunstein, 2000). In eukaryotes, there are 5 types of histone proteins: core histones H2A, H2B, H3, H4 and linker histone H1. Two molecules each of core histones H2A, H2B, H3 and H4 form an octameric protein core around which 147 base pairs of DNA is wrapped around in 1.65 left-handed super- helical turns to form the nucleosome core particle (Luger et al., 1997) (see Figure 2.1 right lower corner). All four core histones are low molecular weight proteins ranging from 11kD to 15 kD and highly conserved across all eukaryotic species.

38 They are present in similar amounts in almost all the cells of an organism, which is consistent with the fixed stoichiometry of the nucleosome core particle composition (Van Holde, 1989). In most eukaryotes, a fifth kind of histone, namely histone H1 or linker histone binds close to the entry and exit sites of the DNA around the nucleosome core particle, thereby sealing two full turns of the DNA by incorporating an additional ~20bp of DNA (Thomas, 1999), resulting in the formation of the nucleosome, the basic repeating unit of chromosomes. Nearly the entire genomic DNA inside the nucleus is packaged into nucleosomes to give rise to nucleoprotein filaments termed chromatin. Histone H1 is larger than core histones with a size of 20-25kD. The central region of histone H1 is conserved, while its N- and C- terminal regions are highly variable. Consistent with their high affinity binding to the negative charged DNA molecules, all histones are positively charged. Histones contain as much as 20% positively charged amino acids such as lysines and arginines, as well as serines and threonines residues than can be phosphorylated. The charged residues are mainly localized in the N- and C- termini of the proteins.

All four core histones share a conserved structure called histone-fold domain, which consists of three α-helices and two short unstructured loops in between (Arents and Moudrianakis, 1995). H2A and H2B have an additional α-helix at their C-termini, while H3 has an extra α-helix at its N-terminus. The histone-fold domain is important for the protein-protein interactions and heterodimer formation. A histone heterodimer is formed by the head-to-tail association of a specific histone pair, either H2A with H2B or H3 with H4, through the histone-fold domain. The heterodimer of H2A/H2B and H3/H4 serve as the basic assembly units during nucleosome assembly (Park and Luger, 2006). The N-terminal tail of each histone is normally unstructured both in isolation in solution and in the nucleosome, which indicates that the formation of the nucleosome does not require the N-terminal tails (Ausio et al., 1989). Moreover, the lack of a defined structure and exposure to the milieu outside of the nucleosome make the histone

39 N-terminal tails a perfect substrate for extensive post-translational modifications that can alter the over all structure and function of chromatin.

The crystal structure of the nucleosome core particle was solved in 1997 by the Richmond group (Luger et al., 1997). Two H3/H4 heterodimers form a tetramer through a single major contact between the two H3 molecules. The association between two H2A/H2B heterodimers and (H3/H4)2 tetramer is achieved by two major contacts between H4 and H2B in the octamer. The entire structure of nucleosome resembles a thick, flat disc (resembling a cheese cake) with the histone octamer occupying the center and the DNA wrapped in 1.65 turns around its lateral side (see Figure 2.1, lower right corner).

The nucleosomal structure is repeated on continuous DNA to form a “beads on a string” filamentous structure of 10nm diameter which is readily visible under the electron microscope. The assembly of chromatin into more compact higher order structures requires the aid of histone H1. Histone H1 serves as a linker to stabilize and condense nucleosomes into higher order structures by presumably binding to the nucleosomal dyad axis and interacting with the linker DNA (Thomas, 1999). The next level of compaction of the chromatin fiber is the highly controversial 30nm fiber, which is believed to be a common structure in the higher order folding of interphase fibers. Nevertheless, its existence in vivo and formation are still in debate (Li and Reinberg, 2011; Maeshima et al., 2010; Tremethick, 2007). Chromatin fibers are believed to be further organized into massive tertiary structure by self assembly into 100-400nm thick filaments by various hypothetical schemes (see Figure 2.1). The packaging of DNA with the help of the histone proteins into chromatin is of great significance for the functioning of the multitude of cellular machineries that have to read the information encoded in the DNA. As such, the packaging of DNA into chromatin may provide a physical barrier for access to DNA, thereby restricting access to factors or machinery involved in DNA replication, transcription, repair and

Figure 2.1 Nucleosome and basic chromatin structure. The crystal structure of the nucleosome core particle consists of two molecules each of H2A (in yellow), H2B (in red), H3 (in blue) and H4 (in green) core histones and 147 bp of DNA wrapped around them. The hypothetical higher order structure of chromatin is shown in schematic representations. Figure is adapted from (Chakravarthy et al., 2005).

recombination. Thus, histones not only serve to package the DNA, but also provide the basis for multiple layers of regulation of chromatin structure and function.

2.2 Definition of Epigenetics

In a literal sense, “epigenetics” means the regulation above or in addition to (Greek prefix epi-) genetics. In the past few decades, epigenetics has been a rapidly developing field, yet it is vaguely defined, and is broadly applied to many aspects of biology. First introduced by Conrad Waddington in 1939 by adapting the Greek word epigenesis, epigenetics was merely a linkage between genetic programming and embryonic development (Waddington, 1939). Parallel to the identification of DNA as the molecular basis of inheritance, scientists realized that in order to address some fundamental questions, they needed to go beyond a genetic explanation. One such question why all genes are either not expressed, or expressed at the same magnitude in a variety of differentiated cells, even though all of them are derived from same genetic origin and have the same underlying DNA sequence. Moreover, in most somatic cells, the phenotype attributed to the pattern of gene expression is permanent and inheritable mitotically. In 1975, Holliday and Pugh, as well as Riggs, published two papers independently speculating on the molecular model of gene activation or inactivation by DNA modifications other than base sequence and its inheritance (Holliday and Pugh, 1975; Riggs, 1975). These two visionary papers suggested that DNA methylation could have strong effects on gene expression, and that changes in DNA methylation might therefore explain the switching on and off of genes during development (Holliday, 2006). DNA methylation has been intensively investigated since then (Suzuki and Bird, 2008). Holliday later on gave the following definition of epigenetics: "the study of the changes in gene expression which occur in organisms with differentiated cells and the mitotic

42 inheritance of given patterns of gene expression” (Holliday, 1994). However, researchers in 1990s and 2000s accumulated conclusive evidence that epigenetic regulation can be also mediated by histones, the architectural proteins that package DNA into nucleosomes. It is now clear that histones are an indispensable partner of DNA in regulating its activities and play an increasingly important role in the epigenetic control of cellular processes. Thus, the definition of epigenetics was broadened further as follows: “an epigenetic trait is a stably heritable phenotype resulting from changes in a chromosome without alterations in the DNA sequence” (Berger et al., 2009). The key processes that are responsible for epigenetic regulation are DNA methylation, histone modifications (as well as histone variants, chaperones and dosage control), chromatin remodeling, and posttranscriptional gene regulation by non-coding RNA (micro- RNAs). Although it is still not clear whether epigenetic traits that result from DNA or histone and chromatin modifications can be stably passed on to the next generation, it definitely poses a challenge to the traditional understanding of inheritance and evolution. From a practical viewpoint, great progress in the field of epigenetics has revealed a complex and delicate regulation of all the important aspects of DNA metabolism, such as DNA damage repair and associated disease development, such as cancer. This chapter will mainly focus on the epigenetic regulation of chromatin functions related to DNA damage repair.

2.3 Overview of Histone Modifications as Epigenetic Marker

Histone posttranslational modifications (PTMs) include acetylation of lysine (K), methylation of lysine or arginine (R), phosphorylation of serine (S) or threonine (T), ubiquitination and sumoylation of lysine, as well as ADP-ribosylation of glutamate (E) residues (Peterson and Laniel, 2004). Adding to the complexity is the fact that each lysine residue can accept one, two or even three methyl groups, and an arginine can be either mono- or di-methylated with the latter being symmetric or asymmetric. These histone modifications are predominantly found

43 in the flexible N-termini of the core histones, which extends out from the lateral surface of the nucleosome, although modifications within C-terminal and the histone-fold domains have also been identified (Mersfelder and Parthun, 2006). Several dozens of histone modifying enzymes are involved in adding or removing the modifications, including histone acetyltransferases (HATs), histone deacetylases (HDACs), histone methyltransferases (HMTs) and histone demethylases (HDMs), etc. Each member of a specific family shows a strict specificity for individual histone tails and for specific histone residues (Kouzarides, 2007). Generally, differences in electrostatic properties between the modified and unmodified forms of these residues can significantly affect interactions between histone core and the DNA wrapped around it. Histone modifications regulate DNA metabolism through three ways. The first is the disruption of interactions between histone octamer core particle and DNA; the second is the influence of higher order chromatin structure by affecting contact between adjacent nucleosomes; the third is recruitment of factors, particularly transcriptional regulators. In general, acetylation of lysine residues is associated with improved chromatin accessibility and enhanced transcription. Since acetylation of lysines will partially neutralize the positive charge of the histone, this will potentially weaken its binding to the negatively charged phosphodiester backbone of DNA. In addition, lysine acetylation can also recruit bromodomain containing proteins with specialized functions in transcriptional control. On the other hand, methylation can function as either repressive or active modification depending on the location of the particular lysine or arginine in the N-terminal tail. Methylation is recognized by chromodomain, Tudor or PHD (Plant HomeoDomain) domain containing proteins, which perform a variety of different tasks in modulating chromatin function. Similar to the genetic code, it has been proposed that the different histone modifications also form a “histone code” that direct the fine- tuning of chromatin functions in highly combinatorial and complex ways (Strahl and Allis, 2000). A particular modification mark or set of marks could have different, even opposite biological consequences. For example, H3K4me (methylation of lysine 4 in histone H3) and H3K36me methylated lysine are

44 found in active chromatin, whereas methylation of H3K9 or H3K27 are generally associated with genes whose transcription is repressed (Kouzarides, 2007). Thus, similar to DNA methylation, rather than being defined absolutely by individual modifications, the epigenetic regulation via histone modification lies in the pattern of modifications spreading over certain genes and the trans-effector proteins recruited by these modifications. Here, I am going to focus on those modifications that are related to repair of DNA damage.

2.4 Role of Histone Modifications in DSB Repair

2.4.1 H2A phosphorylation

Among the known histone modifications, H2A(X) phosphorylation is probably the most studied in the context of the DNA damage response. In mammals, phosphorylation occurs at the serine 139 of the histone H2A variant H2A.X in the C-terminal SQ motif (Rogakou et al., 1998). In the budding yeast, where there is no H2A.x variant, phosphorylation occurs at serine 129 (Downs et al., 2000). This unique modification is often referred to as gamma H2A or γH2A(X). This phosphorylation is catalyzed by phosphoinositide-3 kinases (PI3K), including ATM (Ataxia Telangiectasia Mutated), ATR (Ataxia Telangiectasia-Rad3-related) and DNA-PKcs (DNA-dependent protein kinase catalytic subunit). It has been suggested that ATM is redundant with DNA-PKcs for H2AX phosphorylation after ionizing radiation, whereas ATR is more important during the response to single strand breaks and stalled replication forks (Bonner et al., 2008). In yeast, H2A phosphorylation can be detected as far as 50Kb on either side of DSB by using by chromatin immunoprecipitation (ChIP) (Shroff et al., 2004). In mammalian cells, γH2AX spreads even further, surrounding a region as big as 2Mb on either side of the break (Rogakou et al., 1998).

45 The spreading of γH2A(X) into a large area around DNA lesion suggests the important role of γH2A(X) in amplifying DNA damage signaling. Although H2A.X consists only about 10% of total mammalian H2A, the rapidity of its phosphorylation suggests that at least some of the phosphorylation is present in chromatin before damage and exchange of H2A for H2A.X could occur after damage (De Koning et al., 2007). Consistent with this property, the detection of γH2A(X) is relatively easy (Pilch et al., 2003). Numerous researchers have taken the advantage of γH2A(X) nuclear foci visualized under imunofluorescence microscope using an antibody directed against phosphorylated H2A(X). Furthermore, γH2A(X) is believed to be a very early marker of the DNA damage response and is responsible for the initiation of the signaling cascade in response to DSBs. Upon the formation of a DSB, ATM undergoes autophosphorylation and becomes activated and then phosphorylates H2A.X. The phosphate on serine 139 is recognized by the BRCT (BRCA1 C-terminal) domain of the adaptor protein MDC1 (Mediator of DNA-Damage Checkpoint 1), which also interacts with ATM (Stucki et al., 2005). This allows ATM to accumulate at the break site and phosphorylate more H2A.X. γH2A(X) and MDC1 then recruits MRN complex via the interaction between the MRN complex subunit NBS1 and MDC1. The MRN complex in turn recruits more mediators and subsequent repair effectors (Chapman and Jackson, 2008). A two-stage recruitment model has been suggested, where the initial migration of repair proteins to DSBs does not require γH2A(X), but the subsequent association of DNA damage response factors with chromatin regions distal to the lesion is dependent on γH2A(X) (Huertas et al., 2009). In addition, MDC1 also recruits E3 ubiquitin ligase RNF8 (RING finger protein 8) in a phosphorylation-dependent manner and leads to the accumulation of 53BP1 (p53 Binding Protein 1) and BRCA1 (Breast Cancer 1) repair proteins. RNF8 in turn can ubiquitylate histone H2A and H2A.X (Kolas et al., 2007; Mailand et al., 2007). Thus, γH2A(X) also cooperates with other histone modifications during the response to DNA damage. The role of γH2A(X) in

46 chromatin remodeling will be discussed in detail later in section 2.5 of this chapter.

2.4.2 Histone ubiquitination

Ubiquitination is a common form of post-translational modification that covalently attaches ubiquitin, a highly conserved 76 amino acid polypeptide, to a lysine residue on a target protein (Pickart and Eddins, 2004). As mentioned above, H2A.X or H2A around the DSB is ubiquitinated by UBC13/RNF8 ubiquitin ligase complex following their recruitment. The ubiquitination is initiated by UBC13/RNF8 complex and then the ubiquitin conjugates are amplified by another ubiquitin ligase RNF168 (RING finger protein 8) by catalyzing the formation of lysine 63(K63)-linked ubiquitin conjugates (Doil et al., 2009). RNF8 is recruited via interaction of its FHA domain with phosphorylated MDC1 in a γH2A.X dependent manner. Knocking down of RNF8 impairs the G2/M checkpoint and results in cellular sensitivity to low doses of IR. Thus, MDC1 and RNF8 dependent H2A.X or H2A ubiquitination is believed to be a critical event for the downstream repair response (Panier and Durocher, 2009; Stewart et al., 2009). In addition to RNF8 mediated ubiquitination, a UV-induced mono- ubiquitination of histone 2A in the vicinity of DNA lesions has been described (Bergink et al., 2006). The UV-induced ubiquitination requires functional NER, and coincides with the sequestration of PCNA (proliferating cell nuclear antigen), suggesting that the ubiquitination is linked to DNA repair. Since ubiquitin mutations that eliminate all lysine residues do not affect the ubiquitination, mono- ubiquitination is the most probable outcome of this NER-dependent process. H2A is the primary target and the ubiquitination occurs at K119. In contrast to RNF8 mediated ubiquitination, this mono-ubiquitination process is γH2A.X independent. Additional studies suggest that the observed H2A ubiquitination may be a post- repair process related to chromatin restoration (Zhu et al., 2009). Apart from H2A, a H3 and H4 ubiquitin ligase complex called CUL4-DDB-ROC1 (Damaged DNA Binding protein, Cullin 4, RING finger protein) has also been reported (Wang et

47 al., 2006). Reduction of histone H3 and H4 ubiquitination by knockdown of CUL4A impairs recruitment of the repair protein XPC (Xeroderma Pigmentosum group C) to the damaged foci following UV irradiation and inhibits the repair process. Biochemical studies indicate that CUL4-DDB-ROC1-mediated histone ubiquitination weakens the interaction between histones and DNA and facilitates the recruitment of repair proteins to damaged DNA (Wang et al., 2006).

2.4.3 Methylation of H3K79 and H4K20

Methylation of H3K79 and H4K20 is involved in DNA damage repair (Dinant et al., 2008; Huyen et al., 2004; Sanders et al., 2004). Unlike γH2A(X), H3K79me and H4K20me are not induced by DNA damage; instead, they are constitutively present on chromatin. Both modifications are located on the outside of the nucleosome but are buried in higher order chromatin structure. The common feature of these two methylations is that they serve as docking sites for recruiting DNA damage signal transducer proteins in different species. All these proteins contain the Tudor domain which belongs to the chromodomain methyl-lysine binding family. Once recruited to the lesion, they in turn recruit other proteins to activate repair and stimulate the checkpoint response.

It has been shown in fission yeast that H4K20me is required for the recruitment for Crb2 (homolog of human 53BP1, budding yeast Rad9). The methyltransferase responsible for this modification, Set9, is necessary for Crb2 foci formation and its subsequent phosphorylation (Sanders et al., 2004). This modification is independent of H2A phosphorylation. Interestingly, Crb2 is able to recognize H4K20me and γH2A simultaneously through the Tudor domain and the BRCT domain, respectively (Nakamura et al., 2005). Further more, methyltransferase SUV4-20 exists in fruit flies and humans is responsible for the ubiquitous methylation of H4K20. Depletion of SUV4-20H in HeLa cells impaired the formation of 53BP1 foci, suggesting H4K20me is required for a proper DNA damage response (Yang et al., 2008).

A recent report showed that 53BP1 functions in an XRCC4 (X-Ray cross- complementing)-dependent NHEJ repair, most likely mediated by its interaction with H4K20 (Botuyan et al., 2006). On the other hand, MDC1 mediated HR repair is strictly dependent on its interaction with γH2AX. These mutually exclusive scenarios suggest a specialized adaptation of the "histone code" in the engagement of distinct DSB repair pathways (Xie et al., 2007). Similarly, H3K79me is responsible for recruiting human 53BP1 and budding yeast Rad9 to DSB sites via their Tudor domain (Huyen et al., 2004). Dot1, the HMT responsible for this modification has been well conserved during evolution. It is responsible for methylation of H3K79 in the euchromatin regions of human cells and 90% of the chromatin in budding yeast. This constitutive modification suggests that it can serve as ready-made marker for DNA damage: whenever a DSB occurs, it will lead to the passive relaxation of chromatin higher order structure, allowing 53BP1 access to the exposed H3K79me and promote rapid sensing of the DSB and robust repair response (Costelloe et al., 2006).

2.4.4 Histone acetylation

As mentioned earlier, acetylation of lysine neutralizes the positive charge on histones, which in turn leads to the relaxation of chromatin structure, thereby promoting transcriptional activity in the region. This is also true in the case of DNA damage repair. The modulation of chromatin structure by acetylation is thought to facilitate access of DSB repair proteins to the lesion (Downey and Durocher, 2006). Acetylation of conserved lysine residues is important for normal cell growth and efficient repair following treatment with MMS (methyl methanesulfonate) or endonucleases (Bird et al., 2002; Qin and Parthun, 2002). HATs and HDACs have been reported to be recruited to a DSB induced by the HO endonuclease in budding yeast (Downs et al., 2004; Tamburini and Tyler, 2005). Five lysine residues on histone H3 at positions K9, K14, K18, K23, K27 and four residues on H4 at positions K5, K8, K12, and K16 have been observed

49 to undergo reversible acetylation upon DSB formation (Tamburini and Tyler, 2005). Different HAT complexes are recruited to the DNA damage sites, where they provoke a transient increase in the acetylation of specific histone residues during repair (Bird et al., 2002; Downs et al., 2004; Murr et al., 2006; Qin and Parthun, 2002). Moreover, mutants of the HATs responsible for some of these acetylations, such as Esa1, NuA4, Gcn5 and Hat1, are sensitive to DSBs. In fact, the generation of a single DSB induces hyperacetylation of histone H4 in the surrounding chromatin. Interestingly, some HAT such as Tip60 not only acetylates histone, but also acetylates and activates ATM. Suppression of Tip60 blocks the activation of ATM's kinase activity and prevents the ATM-dependent phosphorylation of p53 and Chk2 (Sun et al., 2005). In addition, treatment of HAT deficient cells with agents that relax chromatin rescues the DSB repair by facilitating the recruitment of HR repair factors such as Rad51 around the lesion. Results from these studies suggest that histone acetylation induced chromatin relaxation is required for efficient recruitment of DNA repair proteins involved in HR repair. On the other hand, the choice of repair pathway might also depend directly on the regulation of the enzymatic activity of HATs and HDACs. There is evidence that DNA-PK, a PI-3K with functions in NHEJ, can inactivate the Gcn5 HAT via phosphorylation (Barlev et al., 1998). As maximal acetylation levels have been shown to occur during HR mediated repair (Tamburini and Tyler, 2005), this suggests that the inhibition of this HAT may direct cells towards NHEJ rather than HR (Costelloe et al., 2006). This is consistent with the findings that I am going to present in the later chapters: compared to HR repair, NHEJ appears to require less relaxed chromatin for efficient DSB repair.

Among all the histone acetylations involved in DNA repair, H3K56ac appears to be growing in significance and has been intensively studied recently. First reported by the Verreault group in budding yeast, acetylation of H3K56 has been shown to be required for DSB repair, especially for lesions formed during DNA replication, such as following camptothecin (CPT) treatment (Masumoto et al., 2005). In unperturbed cells, H3K56ac is cell cycle dependent and acetylation of

50 H3K56 occurs predominantly on the newly synthesized histone H3 during S phase prior to their deposition onto the chromatin during replication. In the absence of DNA damage, this acetylation disappears when cells progress through G2 phase. However, in the presence of DNA damage, such as following treatment with CPT or MMS, H3K56 acetylation is retained in a Rad9/Mec1 dependent manner (Hyland et al., 2005; Masumoto et al., 2005; Thaminy et al., 2007). Further studies identified Rtt109 at the H3K56 HAT which works with the aid of histone chaperone Asf1 for this acetylation (Driscoll et al., 2007; Han et al., 2007). Two HDACs, Sir2 homologues Hst3 and Hst4, that were cell cycle regulated and could deacetylate H3K56 were also discovered (Celic et al., 2006; Maas et al., 2006). In the absence of exogenous DNA damaging agents, Hst3 is maximally expressed in the late S and G2 phases to remove the H3K56ac mark. Hst4, which is maximally expressed in M and G1 phases of the cell cycle, then maintains the deacetylated state until new histones are synthesized in late G1 and S-phase. However, in the presence of DNA damage, Hst3/Hst4 are phosphorylated in a Mec1-dependent manner, and subsequently ubiquitylated and targeted for degradation by the proteosome. Thus, in the absence of sufficient histone deacetylase activity, high levels of the normally transient H3K56ac mark are maintained upon DNA damage (Costelloe and Lowndes, 2010). In addition, it was found in yeast that H3K56 acetylation drives Asf1- dependent re-assembly of chromatin after DSB repair (Chen et al., 2008a). Mutation of K56 to glutamine, mimicking permanent acetylation, partially bypasses the requirement of Asf1 in resistance to DSB-generating agents. Nevertheless, DSB repair itself was operational in Asf1 mutants. On the other hand, lack of H3K56 acetylation in HAT Rtt109 mutants leads to persistent activation of checkpoint protein Rad53. These observations indicate that restoration of chromatin, driven by acetylated H3K56, is a signal for the completion of DSB repair. The histone chaperone CAF-1 (Chromatin Assembly Factor-1) is involved in acetylated H3K56-driven chromatin restoration (Li et al., 2008). H3K56 acetylation promotes the association of histone H3 with CAF-1 and Rtt109, and facilitates CAF-1-dependent nucleosome assembly. Thus, acetylated

51 H3K56 coordinates the function of H3–H4 chaperones in nucleosome assembly. Similar to yeast, H3K56ac was found recently in human cell as well (Das et al., 2009; Tjeertes et al., 2009; Yuan et al., 2009). These studies also showed the H3K56ac increases in response to a variety of DNA damaging agents, such as MMS, IR, HU or CPT. Tyler group has proposed a model of “access, repair and restore” for the roles of H3K56ac during DNA repair based on recent findings(Ransom et al., 2010) (see Figure 2.2), although there is conflicting data from the Jackson group which shows H3K56ac is rapidly and reversibly reduced in response to DNA damage(Tjeertes et al., 2009).

Studies of histone acetylation during DNA repair are of great clinical significance and are drawing more and more attention recently (Portela and Esteller, 2010). The most prominent alteration in a histone modification in cancer cells is a global reduction in the mono-acetylation of H4K16 (Fraga et al., 2005). Loss of acetylation is mediated by HDACs, which have been shown to be overexpressed or mutated in different tumor types (Ropero et al., 2006; Zhu et al., 2004). In addition, mutations or deletions in HAT genes are found to occur more frequently in several different cancer cells compared to normal cells (Portela and Esteller, 2010). Hypoacetylation is also observed in many neurological disorders, such as Parkinson’s disease and Huntington’s disease (Urdinguio et al., 2009). Although the underlying mechanism of how loss of histone acetylation contributes to cancer and other diseases is still unknown, histone acetylation is becoming an important potential biomarker and target for therapeutic development, with the best example being the use of HDAC inhibitors (HDACi) to treat certain cancers and neurological disorders, while numerous clinical trials are ongoing using newer HDACi (Gunjan and Singh, 2010; Shankar and Srivastava, 2008).

2.5 Chromatin Remodeling and DNA Repair

52 It is believed that the chromatin structure must be modified in order to enable the access of repair factors (machinery) at the site of the DNA lesion (Groth et al., 2007). This process may start with the occurrence of histone modifications as described above, and then maybe followed by recognition and interaction of histone modifications by chromatin remodeling complexes. In addition to covalent histone modifications, the structure of chromatin can also be directly manipulated by ATP-dependent chromatin remodeling complexes. These remodeling complexes utilize the energy of ATP hydrolysis to mobilize nucleosomes in the chromatin to create a relatively relaxed local structure and increased DNA accessibility. The main changes remodeling complexes bring to the chromatin include dissolution of the DNA-histone contacts (looping), translocation of nucleosome along the DNA (sliding), nucleosome eviction /disruption, as well histone variant replacement (Wang et al., 2007). Chromatin remodeling promotes DNA damage repair by not only facilitating the access of repair proteins to the lesion, but also by promoting the activation of PI-3 kinases through γH2A(X).

Chromatin remodeling complexes are highly conserved from budding yeast to mammalian cells with common catalytic ATPase subunits. Based on the different domain structure and interactions with histone modifications, chromatin remodeling complexes can be roughly divided into four groups: SWI/SNF (SWItch/Sucrose Non Fermentable) family, ISWI (imitation SWI) family, NuRD (Nucleosome Remodeling and Deacetylation)/Mi-2/CHD (chromodomain, helicase, DNA binding) family and INO80 (Inositol requiring 80) family (Wang et al., 2007). SWI/SNF family contains a C-terminal bromodomain that binds to acetylated histone tails (Hassan et al., 2002). ISWI family contains a SANT (SWI3, ADA2, NCOR and TFIIIB) and a SLIDE (SANT-like ISWI) domain that mediate interactions with histone and linker DNA(Boyer et al., 2004). NuRD/Mi- 2/CHD family members bear unique tandem chromodomains that specifically recognize histone methylations (Sims et al., 2005). INO80/SWR1 (Sick With Rat) family members have split ATPase domains (Bao and Shen, 2007). Different

53 complexes carry out their ATP-dependent remodeling roles through different chromatin interactions and recognition of distinct histone modifications. Additionally, the requirement of multiple subunits for the actions of all the complexes also strongly indicates a delicate collaboration in their function and regulation. Insights into the role of chromatin remodeling in DNA damage repair largely comes from chromatin immunoprecipitation (ChIP) studies conducted in budding yeast, taking advantage of its simpler chromatin structure and ready availability of defined DSB produced by a galactose inducible HO endonuclease. Five yeast remodeling complexes have been identified with distinct role in the repair of DSBs: INO80, SWR1, SWI/SNF, RSC and Rad54 (Osley et al., 2007). In this chapter, I will mainly focus on INO80, SWR1, SWI/SNF and RSC in chromatin remodeling complexes in relation to DSB repair.

The most compelling evidence for the involvement of chromatin remodeling in DSB repair comes from intensive studies of INO80. Yeast INO80 complex contains approximately 12 subunits with ATPase activity (Ino80) and helicase activity (Osley and Shen, 2006). Several lines of evidence directly link INO80 dependent chromatin remodeling to DSB repair. First, three groups simultaneously showed that the INO80 complex is recruited to a HO endonuclease-induced DSB through a specific interaction with the DNA damage- induced γH2A (Downs et al., 2004; Morrison et al., 2004; van Attikum et al., 2004). Recruitment of Ino80 is compromised in cells lacking the H2A S129. Second, INO80 is also required for histone eviction from chromatin near the DSB. Mutant arp8, which is defective in the INO80 ATPase, displays delayed histone eviction, and similar delay in nucleosome disruption assayed by carefully designed ChIP and qPCR experiments (Tsukuda et al., 2005). Third, the loss of INO80 catalytic activity by either deletion of the Ino80 ATPase or ATPase- stimulating subunits Arp5 or Arp8, results in sensitivity to agents causing DSBs (Osley et al., 2007). Fourth, the INO80 remodeling activity may be required to enable access to end-processing enzymes such as MRX or other factors to the DSB site. INO80 mutations impair the binding of Mre11, Ku80 and Mec1 at DSBs,

54 resulting in defective end-processing and reduced checkpoint activation (van Attikum et al., 2007). This also suggests that INO80 is not only involved in HR repair, but also required for NHEJ pathway. Last but not least, the Les4 subunit of the INO80 complex is phosphorylated by the Mec1/Tel1 kinases following exposure to DNA-damaging agents (Morrison et al., 2007). Mutation of les4's phosphorylation sites does not significantly affect DNA repair processes, but does influence DNA damage checkpoint responses. This finding indicates that INO80 as a chromatin remodeller can be integrated in Mec1/Tel1 DNA damage signaling pathway (Morrison et al., 2007). Although INO80 can interact with DNA, histones as well as nucleosomes in vitro (Shen et al., 2000), the mechanism that it adopts to alter chromatin is unknown. There is also controversy about the real chromatin situation around a DSB lesion. Earlier ChIP data showed that histone H2B was not lost from DSB proximal chromatin (Shroff et al., 2004). However, there is suspicion that the efficiency of crosslinking by formaldehyde used in the ChIP procedure between histones and dsDNA when compared to histone and ssDNA is different, which could lead to an underestimation of histone loss (Sinha and Peterson, 2009). Most interestingly, although the initial recruitment of INO80 to DSB is dependent on γH2A, its chromatin remodeling and nucleosome eviction activity can occur independently of H2A phosphorylation (Morrison et al., 2004; van Attikum et al., 2004). It is possible the γH2A.X may be initially dispensable for remodeling but it is important for maintaining the remodeled state of chromatin that enables complete repair. Since the process of de-phosphorylation takes place on non-chromatin bound H2A.X, another use of INO80 is to remove γH2AX from the chromatin (Chowdhury et al., 2005; Keogh et al., 2006). It is tempting to speculate that INO80 could displace γH2A.X after completion of repair and serve as terminator of the DNA damage signaling. Further, this process might need certain assistance from histone chaperones such as Asf1 to deal with regions further than 5kb from the break site where INO80 is absent (Osley et al., 2007). Interestingly, this speculation correlates well with the H3K56ac “access, repair, restore” model (Chen and Tyler, 2008), especially regarding the intriguing role of Asf1 in both the cases (see Figure 2.2).

SWR1 is a conserved complex that was initially identified in yeast as Swi2/Sfn2 related ATPase subunit (Krogan et al., 2003). Similar to INO80, recruitment of SWR1 requires γH2A.X and it shares the Arp4 subunit that interacts directly with γH2A.X in vitro. The SWR1 complex functions along with NuA4 HAT to replace γH2A.X with the H2AZ (Htz1 in budding yeast) variant, which cannot be phosphorylated (Kusch et al., 2004; Papamichos-Chronakis et al., 2006). The exact role of this variant exchange is unknown but it is probably important for DNA damage-response survival because loss of H2A.Z and SWR1 function causes sensitivity to genotoxic agents. In the budding yeast, the SWR1 complex shares four subunits (Act1, Arp4, Yaf9, and Swc4) with the NuA4 complex (Morrison and Shen, 2009). These two complexes function together to regulate the deposition of H2A.Z into nucleosomes. Current models suggest that SWR1 recruitment to chromatin depends on those shared subunits between the SWR1 and NuA4 complexes. The Nu4A complex acetylates the deposited H2A.Z and this modification may play a role in blocking the spread of heterochromatin into euchromatin (Babiarz et al., 2006). In DNA repair, the acetylation of H2A.Z may serve as an important regulatory step during these histone exchanges. The role of the SWR1 complex could be also related to the exchange of modified histones when repair is completed. In this regard, Domino/p400, the Drosophila homolog of SWR1, is required for the exchange of phospho-H2Av (the equivalent of mammalian γH2A.X) by unmodified H2Av following its acetylation by Tip60. However, after DSB induction, H2A.Z in yeast does not become more abundant in the damaged chromatin. Instead, γH2A.X and all the other core histones are removed near the breaks in a coordinated and SWR1-independent fashion, and only the loading of Ku80 is impaired in the Swr1 mutant strain (van Attikum et al., 2007). This suggests that SWR1 activity may be restricted to repair of DNA lesions by specific pathways.

In budding yeast, SWI/SNF complex consists of 11 subunits, among which the catalytic subunit Swi2/Snf2 is conserved throughout evolution (Mohrmann and Verrijzer, 2005). Although SWI/SNF is mainly considered important in transcription, it also plays a role in DSB repair. Mutants of SWI/SNF subunits are sensitive to DSB inducing agents and are recruited to the MAT locus after HO expression (Chai et al., 2005). SWI/SNF associates with both the MATa recipient locus and HR donor locus HMLα, while it does not appear to have a crucial function in NHEJ. In vitro biochemical studies indicated that a successful homology search on chromatin fibers does not require remodeling complexes; however, SWI/SNF might play an essential role in synapsis between the invading ssDNA and donor locus in heterochromatic regions (Sinha and Peterson, 2008). Indeed, in the absence of the Swi2 or Snf5 subunits, the invading MATa presynaptic filament is unable to locate and synapse with the heterochromatic HMLα (Chai et al., 2005). Rad51 fails to mediate successful homology search in the presence of reconstituted heterochromatin in the donor locus using recombinant nucleosomes and Sir proteins, while addition of SWI/SNF could restore its function (Sinha et al., 2009). On the contrary, addition of INO80, SWR1, RSC or Rad54 could not perform same function as SWI/SNF, indicating SWI/SNF may have a specific role in regulating HR by facilitating synapsis especially when the donor locus is located in a heterochromatic region.

The RSC (Remodel the Structure of Chromatin) complex contains about 15 subunits, sharing two identical and at least four homologous subunits with the SWI/SNF complex, though there are several important functional differences between the two(Wang, 2003). Similar to the SWI/SNF complex, RSC plays an important role in transcription, but is more abundant than SWI/SNF and essential for viability. Also, RSC mutants are sensitive to DSB but the defects seem to be attributed to both HR and NHEJ (Shim et al., 2005). RSC binds to both recipient and donor loci during HR repair. Depletion of the ATPase subunit Sth1 or Rsc2 severely reduces chromatin remodeling and loading of Mre11 and Ku at the DSB

57 (Shim et al., 2007). RSC is rapidly recruited to a DSB at MAT locus in at the same time window as MRX recruitment (Chai et al., 2005). These observations suggest that the involvement of the RSC in chromatin remodeling around DSB is probably through the MRX complex. MRX is required for histone eviction around DSB (Tsukuda et al., 2005). Thus, given the well known role of MRX in damage sensing, end resection and recruitment of downstream proteins, remodeling complexes such as RSC could cooperate in the eviction of histones and recruitment of more repair factors. In addition, RSC may be involved in cohesin loading at the break due to its analogous role in loading cohesin on chromosome arms during the cell cycle (Huang et al., 2004; Shim et al., 2005). Cohesins facilitate the repair of DSB by maintaining the two sister chromatids in close proximity for repair by HR and they are also recruited to breaks in a γH2A.X and Mre11 dependent way (Huertas et al., 2009; Unal et al., 2004). Therefore, RSC could channel DSB repair into different pathways: when homologous sequence in sister chromatid is available, HR is promoted by RSC through Mre11 mediated cohesion loading; whereas, in the absence of donor sequence, NHEJ kicks in for alternative repair.

2.6 Role of Histone Chaperones in DSB Repair

In order to render damaged DNA accessible, chromatin remodeling around the DNA lesion will inevitably result in the eviction of some histones from the chromatin. These DNA-free histones exist only transiently in the cells, although they can also last long. It has been shown in Xenopus that large quantities of histones are synthesized and stored during oogenesis for the rapid cell cycles during early development (Woodland and Adamson, 1977). Due to the nature of their composition, the positively charged histones have very high affinity for any negatively charged molecule in the cells, such as nucleic acids. Uncontrolled wandering of histones in the cells could lead to deleterious effects (Singh et al.,

58 2009b). Therefore, most free histones are escorted by specific proteins called histone chaperones. When DNA is repaired, histones have to be re-deposited on the newly repaired DNA, by a process known as chromatin assembly. In the process of chromatin assembly, all the histones to be deposited are presumably escorted by histone chaperones. So histone chaperones are not only preventing unwanted binding of free histones to other negatively charged molecules, but also controlling histone supply and incorporation into the chromatin. As such, histone chaperones are bound to be important players in DNA damage repair (De Koning et al., 2007).

Until recently, not much evidence has been reported for the involvement of H2A/H2B chaperones in DNA damage repair except for FACT (Facilitates Chromatin Transcription) complex in exchanging H2A.X (Avvakumov et al., 2011). Most research done in this area was focused on histone H3/H4 chaperonees. Histone H3/H4 exists as a heterodimer when bound to some histone chaperones, including Asf1, HIRA (Histone Regulatory homolog A) and CAF-1 (Tagami et al., 2004). Asf1 is considered as a donor for handing over H3/H4 heterodimer to HIRA and CAF-1, which are the two chaperones responsible for replication- independent and replication-coupled chromatin assembly respectively (De Koning et al., 2007). Currently, there is no direct evidence that histone chaperones play an active role other than taking free histones passively during chromatin disassembly around a DSB. Asf1 and CAF-1 appear to be dispensable for chromatin disassembly, while their mutants fail to re-assemble chromatin after DNA repair (Chen et al., 2008a; Kim and Haber, 2009). DNA repair related chromatin assembly shares many factors and similar mechanism with replication- coupled chromatin assembly (Verreault, 2000). Thus, the most crucial histone chaperones related to DNA damage are Asf1 and CAF-1 (see Figure 2.2). Indeed, genetic studies in yeast, as well as biochemical analyses of human histone chaperones showed that both Asf1 and CAF-1 are involved in repair of different DNA lesions. Mutants of either Asf1 or CAF-1 are sensitive to different

59 DNA damaging agents, such as ionizing radiation, UV, radiomimetic agents and MMS (Linger and Tyler, 2007).

Very similar to the process of replication coupled chromatin assembly (Verreault, 2000), CAF-1 is recruited by PCNA to the sites of DNA lesions (Nabatiyan et al., 2006). CAF-1 is required for the incorporation of newly synthesized, replication- coupled histone H3.1, which is first captured by Asf1 and then handed to CAF-1 for deposition, onto the repaired DNA (Green and Almouzni, 2003; Polo et al., 2006). To further distinguish the role of CAF-1 in DNA replication and DNA repair, a DNA damage study was conducted in quiescent cells and CAF-1 recruitment to damage sites induced by various agents was observed. Depletion of CAF-1 by RNA interference in bleocin-treated quiescent cells in vivo results in a significant loss of cell viability and an accumulation of DSBs (Nabatiyan et al., 2006). These results indicate that the CAF-1 contributes to chromatin assembly after DNA damage repair in a mechanism similar to its role during DNA replication, but is independent of major DNA synthesis. The function of CAF-1 during DSB is likely to be subjected to tight regulation, presumably via interaction with the DNA synthesis/end resection related DNA helicase, such as Sgs1(human homolog of BLM/WRN)(Ransom et al., 2010).

Apart from the cooperative functions of Asf1 with CAF-1during chromatin assembly, Asf1 has the important function of acting as a protein cofactor in the acetylation of newly synthesized histone H3K56, which has been elucidated in recent years (Chen et al., 2008a; Li et al., 2008). As described in detail previously, Asf1 is required to activate the HAT Rtt109 to acetylate H3K56, which plays critical roles in targeting the histones to the sites of DSB repair in order to achieve chromatin assembly. Based on these findings, the original model of “Access-Repair-Restore” proposed by Smerdon (Smerdon, 1991) has been developed further (Avvakumov et al., 2011) (see Figure 2.2). The requirement ofAsf1, Rtt109 or H3K56ac to deactivate the checkpoint response suggests a model in which the stretch of chromatin bearing acetylated H3K56 deposited onto

Figure 2.2 “Access, Repair, Restore” model for the role of histones during DSB repair. The variant histone H2AX is phosphorylated on S139 (S129 of H2A in S. cerevisiae) in the chromatin surrounding the site of damage. Once DNA resection commences, displaced histones are likely captured by chaperones. However, this has not been demonstrated, and identities of chaperones that may be involved in this process are unknown. Repair: While the repair machinery assembles and mends the break, PARP1, which is upregulated in response to damage, mediates poly-ADP-ribosylation of FACT, thereby rendering it inactive. Restore: With DNA damage repaired, histone chaperones participate in restoration of the chromatin structure. Asf1/Rtt109 and CAF-1 deposit newly synthesized histones onto DNA to replace those lost during the repair process. Meanwhile, the FACT chaperone functions to exchange phosphorylated H2A with an unmodified version in order to release any repair proteins retained by this mark and to permit resetting of the cell-cycle checkpoint triggered by damage. Figure is adapted from (Avvakumov et al., 2011). recently repaired DNA signals that DNA repair is complete(Chen and Tyler, 2008).

2.7 DNA methylation and DNA damage repair

In contrast to a great variety of histone modifications, DNA methylation is just a simple addition of a single methyl (CH3-) group to cytosine at position 5. In mammals, it happens almost exclusively on cytosines preceding guanine (CpG) in the DNA sequence. The majority of CpG sites in human DNA are methylated. Methyl groups flag approximately 5-6% of cytosines in healthy cells(Bird, 2002). In general, CpG methylation is associated with gene silencing. This may have developed as a defense against expression of parasitic DNA elements(Yoder et al., 1997). DNA methylation is mediated by DNA methyltransferases (DNMT) that can catalyze the transfer of methyl groups. In humans, the de novo DNA methyltransferases DNMT3a and DNMT3b methylate the genome during embryonic development, whereas the maintenance DNA methyltransferase DNMT1 methylates hemi-methylated DNA following mitosis as a “photocopier” (Robertson, 2001). However, although there are plenty of reports implicating DNA methylation in cancer, there is very little direct evidence of its role in DNA damage repair (Portela and Esteller, 2010). In contrast, 5-methylcytosine is prone to spontaneous deamination and point mutation to thymine. Although 5- methylcytosine represents only 1% of the bases in the human genome, its potential mutagenic hazard is well illustrated by the fact that CpG dinucleotides are involved in one third of point mutations causing human genetic disorders (Cooper and Youssoufian, 1988). Yet, several studies have shown that DNA methylation via DNA methyltransferases might be involved in the restoration of epigenetic information during DNA repair. DNMT1 forms a complex with PCNA and is recruited to DNA repair sites (Mortusewicz et al., 2005).Defective DNMT1 results in significantly elevated mutation rates and genomic instability in mouse cells (Chen et al., 1998; Okano et al., 1999), as well as in human cell lines (Xu et al., 1999). Moreover, de novo methylation of cytosines flanking a DSB is observed following HR mediated repair. As a direct consequence, the gene in these repaired molecules was silenced. This indicates that DNA methylation in

62 eukaryotes could serve as a marker for homologous recombination (Cuozzo et al., 2007). Recently, DNMT1 has been shown to interact with the PCNA, Chk1 and the “9-1-1” (Rad9-Rad1-Hus1) complex, which may contribute to its localization, and as such DNMT1 status may influence the rate of DSB repair following gamma-irradiation (Ha et al., 2011). Nevertheless, it is always difficult to unequivocally exclude the influence of transcriptional alterations in certain genomic areas that could lead to these findings as well. Clearly, more work, especially using embryonic stem cells is needed to elucidate the detailed role of DNA methylation in DNA damage repair.

63 CHAPTER 3

HISTONES: THE ENEMY WITHIN

Despite being the main packaging protein for DNA, histones are not necessarily continuously associated with DNA through the whole cell cycle. During DNA replication, the parental nucleosomes must be disrupted to allow processing related to its duplication by the replication machinery. Newly synthesized histones have to be transported to the nucleus and deposited onto the nascent DNA, while the parental histones also need to be transferred onto either the leading or the lagging strand (Verreault, 2000). In this transient process of histone eviction and re-deposition, cells encounter two opposing challenges related to histone protein levels as elaborated below.

3.1 Histones: Enemy or Friend?

The first challenge cells have to meet is the high demand of histones during the formation of new chromatin. During S phase, not only the DNA but also chromatin structure is replicated, so the cell needs to produce a large number of histones (two-fold increase) in a relative short period of time to assemble new DNA into chromatin. This is actually quite a challenge because the formation of new nucleosomes has to occur as soon as enough new DNA for nucleosome assembly emerges from the replication machinery. More precisely, the distance between where histones are assembled and the branch point of replication fork is approximately 100-400bps (Sogo et al., 1986). Therefore, histones are

64 synthesized mainly during S-phase when newly replicated DNA needs to be rapidly packaged into chromatin.

Timely nucleosome assembly is very crucial for cells as delays between DNA synthesis and histone deposition lead to a loss of viability in yeast (Han et al., 1987). Inhibition of histone synthesis leads to spontaneous DNA damage and S- phase arrest in human cells (Nelson et al., 2002; Ye et al., 2003a). Moreover, it has been shown that repression of either histone H2B or H4 results in the cell becoming incapable of chromosome segregation (Saunders et al., 1990). A similar phenomenon was observed during meiosis where the partial depletion of H2A/H2B dimers in yeast blocks progression into the first meiotic division (Tsui et al., 1997). The reasonable hypothesis in these studies is that a shortage of any one core histone or perhaps all four core histones will result in chromatin assembly being unable to keep pace with the progressing replication fork. This will result in aberrant chromosome structures that might interfere with mitotic/meiotic divisions.

The second challenge that cells face is that histones are highly positively charged proteins which can bind very avidly and non-specifically to negatively charged DNA. Indeed, despite the huge demand for histones for chromatin assembly, significant amount of free histones are not found in cells under normal growth conditions. Non-chromatin bound free histones can be extremely harmful to cells (Fig 3.1). This opinion is supported by several lines of evidence from diverse areas of study. First, biochemically, the non-specific association of histones with DNA does not result in a proper nucleosome structure; instead, it results in the formation of insoluble aggregates (Carruthers et al., 1999). In vitro studies involving reconstitution of nucleosomes from recombinant histones and DNA have shown that even a slight stoichiometric excess of histones over DNA is sufficient to promote chromatin aggregation (Carruthers et al., 1999; Dyer et al., 2004). Second, in vitro reconstitution experiments have shown that not only the

65 core histone octamer but also the H3/H4 tetramer alone can provide an nearly absolute block to transcript elongation by RNA polymerase II (Chang and Luse, 1997). Moreover, greater transcriptional repression was observed in reconstituted nucleosomal template that contains a greater ratio of histones to DNA (Steger and Workman, 1999).The level of nucleosome occupancy is inversely proportional to the transcriptional initiation rate at the promoter. Loss of histone H3/H4 is observed in the coding regions of the most heavily transcribed genes (Lee et al., 2004). Hence, histones arising from the soluble histone pool could re-assemble onto gene promoters and promote repression (Schermer et al., 2005). Moreover, if a large quantity of free histone accumulates in the cells, the machinery responsible for the transcription-related histone eviction could get clogged up with this excess histone burden. Consistent with this hypothesis, yeast cells are found to accumulate excess histones when normal metabolism of transcriptionally evicted histones was abolished (Morillo-Huesca et al., 2010). There results indicate that chromatin-free histones probably pose a significant threat to ongoing transcription in a manner that is dependent on the proper control of histone disassembly and reassembly. Third, free histones have been also shown play a role in genomic instability. In yeast, overexpression of histone genes exhibited an increased frequency of mitotic chromosome loss (Meeks- Wagner and Hartwell, 1986) and an increase in DNA damage sensitivity (Gunjan and Verreault, 2003). Mutants in Drosophila that produce stable histone mRNAs showed late embryonic lethality during development, due to the accumulation of histones in non-replicating cells (Sullivan et al., 2001). Overexpression of the centromere specific histone H3 variant CENP-A (homolog of budding yeast Cse4) leads to chromosome mis-segregation (Au et al., 2008). Excess histone accumulation causes loss of viability in budding yeast mutants deficient in the posttranslational degradation of histones (Singh et al., 2009a). In addition, excess histones are likely to have more profound effects at the organism level in multicellular organisms. Mutations causing accumulation of histone mRNAs are

Figure 3.1 Consequences of excess histones. See text for details.

67 believed to contribute to apoptosis and chronological aging in budding yeast (Mazzoni et al., 2003; Palermo et al., 2010).

Further, extracellular histones are cytotoxic toward endothelium cells and lead death of mice within one hour of injection. Mice challenged with a lower dose of histones showed many characteristics of sepsis, whereas introduction of antiapoptotic factor APC (short for activated protein C) can rescue the deleterious effects of histones (Xu et al., 2009). Although these studies fail to demonstrate the detailed mechanism underlying the ill effects of excess histones, they do highlight the importance of histone regulatory pathways.

Hence, because of the dual nature of the challenges potentially generated by changes in histone levels as described above, a delicate balance must be maintained between histone levels and DNA synthesis during S phase. Such a fine balance will assure production of enough histones for incorporation into new chromatin and prevent excessive accumulation of histones, which would be hazardous for dividing cells (Singh et al., 2009b).

3.2 Sources of Excess Histones

Histones are such vital proteins for the packaging DNA into chromatin, regulating access to the genetic information, maintaining epigenetic marks and ensuring the proper segregation of DNA, that they are produced in excess to ensure the availability of sufficient histones (Gunjan et al., 2006). On the other hand, despite the huge demand for histones for chromatin assembly, significant pools of free histone are not found in the cells under growth conditions as their non-specific interactions with DNA and other negatively charge cellular components can be extremely harmful to various aspects of DNA metabolism. Yet, genetic evidence strongly points to the existence of excess histones in the cell, either in a diseased condition, or transiently. For clarity, we define the free or excess histones

68 referred to here as non-chromatin-associated histone, although they are not necessarily free of any physical interactions, and in some cases are carried on histone chaperones (De Koning et al., 2007). Using the analogy of supply and demand, our laboratory has attempted to illustrate the sources of excess histones (Singh et al., 2010). Whenever there is an increase in histone supply or a decrease in cellular histone demand, excess histones are generated. In terms of the origin of excess histones, they can either come from newly synthesized histone pool or chromatin bound histones evicted during transcription or DNA damage repair (Fig. 3.2).

3.2.1 Multiple histone genes

Eukaryotes have multiple genes encoding each histone protein, some of which are non-allelic variants (Stein et al., 1984).Generally speaking, there are three major patterns of multiple histone gene arrangements. In a unicellular eukaryote, such as budding yeast, there are two nonallelic copies of genes for each of the four core histones(Osley, 1991). However, they are not distributed randomly and H2A genes are paired with H2B, while H3 genes are paired with H4. Each of these pairs of core histone genes are driven by a common divergent promoter. Both H3 and H4 genes encode identical histone H3 proteins and H4 proteins respectively, while the two H2A and H2B genes encode slightly different proteins. Compared to yeast histone genes, a more complex histone gene organization in metazoa reflects a long course of evolution. The predominant pattern in fruit fly or sea urchin is a tandem repeat where all five histone genes are placed one after another with intergenic spacer DNA to form a repeat unit. Identical units are repeated in tandem tens to hundreds of times, with the repeats being extremely homogeneous, containing only minor differences (Maxson et al., 1983). In mammals, histone genes are clustered on certain chromosomes, with one major locus and two minor loci, each of which contains multiple genes

Figure 3.2 Sources of excess histones. See text for detail.

70 (Marzluff et al., 2002). There are 10~20 functional copies of each core histone gene in mice and humans. Compared to homogenous tandem repeats observed in lower metazoans, in the each mammalian cluster, histone genes are distributed in an erratic manner. Histone genes generally carry strong promoters and are transcriptionally active mainly during S-phase of the cell cycle. By virtue of the dozens of histone genes driven by strong promoters, cells tend to produce large amounts of histones beyond their need during S phase. By comparing nucleosome occupancy and histone expression levels (Ghaemmaghami et al., 2003); one can theoretically predict the gap between histone demand and supply in the budding yeast.

We have estimated that approximately 8 fold more of some core histones are produced than the actual amount needed for chromatin packaging in yeast. Then, why do eukaryotes carry multiple histone genes? Since histones are essential for survival, is it possible that extra copies serve as backup? This is unlikely because most essential proteins do have backup gene copies; plus, even if backup copies exist, they do not need to be transcriptionally active. A second possibility is that the multiple histone genes may have distinct functions. This appears to be true for non-allelic core histone variants, especially in the case of multiple types of H2A/H2B or H3 variants and DNA replication independent histone variants (Banaszynski et al., 2010; Talbert and Henikoff, 2010). Yet it does not fully explain whythere is not a single variant of H4 and each of 14 H4 genes in humans encodes identical proteins (Marzluff et al., 2002). The third possibility is that multiple histone genes will cope better with the challenge of the high demand for histones during chromatin assembly in S phase. As pointed out previously, histones are primary synthesized in S phase and deposited by chromatin assembly factors or histone chaperones on to the replicating DNA to form chromatin. Multiple histone genes enable abundant histone synthesis during DNA replication, which in most cases occurs within a relatively short time window. This is especially crucial for higher eukaryotes with extensive genomes and those with very short embryonic cell cycles as in Xenopus and Drosophila.

71 During the early stages of fly embryogenesis, the DNA replicates every 10~15 minutes, whereas transcriptional activity is low. Embryonic cells utilize stored histone mRNA that synthesizes histones for a short period of time prior to the start of replication to meet the demands of rapid chromatin assembly (Maxson et al., 1983). Hence, it is quite possible that cells produce histones in excess of the amount required for rapid chromatin assembly during replication.

3.2.2 Replication arrest

In order to meet the huge demand of histone deposition during DNA replication without making an excess of histones, cells apply tight controls on histone production. In fact, histone synthesis is tightly coordinated with DNA replication and the majority of histones are exclusively synthesized during S phase of the cell cycle, a phenomenon that is evolutionary conserved from yeast to metazoans (Osley, 1991). The mutual influence of DNA and histone synthesis on each other when one of the two is affected provides the best evidence that the two are coupled. Inhibition of DNA synthesis with hydroxyurea or cytosine arabinonucleoside leads to a decline in the steady-state concentrations of histone mRNA, as well as the transcription rate of histone genes (Heintz et al., 1983; Sittman et al., 1983). On the other hand, repression of histone transcription in turn triggers a concerted block of DNA synthesis (Nelson et al., 2002). The S- phase specific production of histones ensures that superfluous histones are not produced during the rest of the cell cycle. However, the actual extent of inhibited histone production may not always match the inhibition of DNA replication. As a reflection of DNA synthesis inhibition, histone synthesis down-regulation takes some time, which inevitably leads to accumulation of non-chromatin bound histones (Bonner et al., 1988). Replication arrest mostly comes from DNA damage during S phase, which results in DNA replication either slowing down or coming to a complete halt. This is either due to the physical impediment posed by the lesions to the passage of DNA polymerase, or due to the activation of the S phase DNA damage checkpoint that prevents the firing of new origins

72 (Paulovich and Hartwell, 1995; Tercero and Diffley, 2001). The accumulation of free histones on histone chaperones in response to both DNA damage and replication arrest has been demonstrated in the budding yeast (Gunjan and Verreault, 2003), as well as in cultured mammalian cells in response to replication inhibition (Groth et al., 2005).

3.2.3 Histones evicted during transcription or DNA damage repair

It has been known for decades that the chromatin structure is perturbed near actively transcribed regions as assayed by nuclease hypersensitivity experiments (Weintraub and Groudine, 1976; Wu et al., 1979). The transcriptional machinery appears to prefer promoters lacking histones than promoters fully packaged into chromatin (Boeger et al., 2003; Fedor and Kornberg, 1989). In fact, there is an inverse correlation between nucleosome occupancy at the promoters and the transcription rate of downstream genes. This is also true for open reading frames (ORFs) (Bernstein et al., 2004; Lee et al., 2004). Thus, the temporary displacement of histones, or a persistent absence of histones in the case of a heavily transcribed region, suggests that the presence of histone proteins on the DNA is likely to serve as a barrier for transcription. However, the fate of the histones evicted during transcription is not clear. Transcriptionally evicted histones could be recycled via a mechanism similar to the recycling of older histones during chromatin assembly associated with DNA replication. First, the chromatin structure is loosened up to allow access to the transcriptional machinery, and then the displaced histones are restored after the passage of the RNA polymerase. Alternatively, the evicted histones may be targeted for degradation and replaced with histone variants which specifically mark transcriptionally active regions. A good example of this would be the enrichment of H3.3 in the interbands of Drosophila salivary gland polytene chromosomes where genes are actively transcribed and canonical H3 is removed (Schwartz and Ahmad, 2005). Factors such as Spt16 (part of histone chaperone FACT

73 complex) function both as transcription elongation factors and nucleosome assembly proteins (Belotserkovskaya et al., 2003; Kaplan et al., 2003). In fact, histone H3 carries the majority of histone modifications, especially the marks for heterochromatin, suggesting that the replication-independent pathway is involved in the regulation of heterochromatin silencing and transcriptional repression (Workman and Abmayr, 2004). Moreover, recent evidence suggests a role of Spt16 in preventing the accumulation of transcriptionally evicted histones, as a deficiency of spt16 leads to deleterious effects, including loss of viability and G1 delay (Morillo-Huesca et al., 2010). Results from this study demonstrate a crucial requirement for the excess histone degradation machinery for dealing with free histones during transcription in the absence of spt16.

Apart from transcription, histones are evicted from chromatin undergoing DNA repair as well. As mentioned in the previous chapter, the accessibility of chromatin is crucial when DNA damage takes place in tightly packaged chromatin and repair factors are trying to gain access to the DNA lesion. Relaxation of chromatin structure is achieved by the cooperative action of different mechanisms, such as the DNA damage sensors for detecting lesions, damage specific histone modifications for marking the target regions and the removal of nucleosomes around the lesions by chromatin remodeling complexes.. By using an inducible site specific endonuclease such as HO or I-SceI, nucleosome dynamics and DSB repair factor recruitment at flanking regions of a DSB was assayed by ChIP and Southern blotting (Tsukuda et al., 2009b). An increase in the accessibility to micrococcal nuclease digestion and eviction of histones at the regions flanking the DSB was observed (Tsukuda et al., 2005; van Attikum et al., 2007). Further confirmation for histone eviction from the ssDNA following resection of DSBs comes from the fact that the histones are replaced by the ssDNA binding protein RPA and Rad51 (Dubrana et al., 2007). ATP-driven remodeling complexes INO80 as well as SWI/SNF are also involved in nucleotide excision repair for UV-induced photo lesions, suggesting histone displacement and exchange may occur in DNA damage other than DSBs as well

74 (Hara and Sancar, 2002; Jiang et al., 2010). Although the extent and exact mechanism of histone disassembly around DSBs or other damage are still in debate, the free histones evicted from damaged chromatin could be substantial due the high frequency of DNA damage the cell faces (Hoeijmakers, 2009), and the 2Mb chromatin domain that gets modified around a DSB in mammalian cells.

3.3 Regulation of Histone Levels

To achieve a delicate control of histone levels, the cells have developed multiple layers of regulation including transcriptional, posttranscriptional, posttranslational mechanisms as well as regulation via histone chaperones (Fig. 3.3).

3.3.1 Transcriptional control of histone gene expression

The expression of the majority of histone genes is transcriptionally regulated in strict correlation with DNA replication. When cells enter S-phase, nascent histone mRNA synthesis increases 3~5-fold over the basal level of G1 phase cells (Harris et al., 1991). Transcription of the four histones genes is restricted to late G1 to early S phase.

Much of study in histone transcriptional control was done in budding yeast due to the relative simplicity of its histone gene structure (Osley, 1991). As mentioned earlier, histone genes are present in pairs in the budding yeast: H3 is paired with H4, whereas H2A paired with H2B. Each histone gene pair is transcribed divergently from a common promoter. There are two cis-acting factors within the histone gene pair promoters: Upstream activation sequences (UAS) and Negative cis-acting element (NEG), which play different roles in their cell cycle regulation (Osley, 1991). As the name suggests, UAS serves as an activator that

Figure 3.3 Regulation of Histone Levels. See text for details.

76 is responsible for the coordinated transcriptional activation of histone genes at the G1-S boundary. NEG is responsible for histone gene repression (NEG is absent from one of four histone gene pairs, HTA2-HTB2 in the budding yeast) when cells are not in S phase or when DNA replication is inhibited. Deletion of the NEG element leads abolishes the cell cycle dependent transcriptional control of histone genes, leading to the accumulation of histone mRNA throughout the cell cycle. Some trans-acting proteins are also involved in this regulation. The Histone Regulation (Hir) complex comprising of Hir1, Hir2, Hir3 and Hpc2 proteins, in cooperation with Asf1 (Anti Silencing Function 1), are believed to be co-repressors of histone transcription through the NEG elements (Spector et al., 1997; Sutton et al., 2001). The HIR complex in association with Asf1 is believed to function not only as a histone gene repressor outside of S phase, but also as a nucleosome assembly factor (Green et al., 2005; Gunjan et al., 2005). However, the molecular mechanism of its transcriptional repression is still not clear. RSC (Remodels the Structure of Chromatin), an abundant nucleosome remodeling complex, is also recruited to histone promoters in a Hir complex and NEG dependent manner (Ng et al., 2002). In addition, recent research from our laboratory has demonstrated that the DNA damage checkpoint kinase Mec1/Tel1-Rad53 pathway in the budding yeast is responsible for the repression of histone genes when DNA replication is inhibited by genotoxic agents (Paik et al. unpublished data).

In mammals, histone genes also contain specific elements that are required for cell cycle activation. It is believed that the G1-S histone gene expression is regulated by Cyclin E-Cdk2 substrate NPAT (nuclear protein, Ataxia- Telangiectasia locus), presumably, with the aid of Hir complex homologue known as HIRA (Hall et al., 2001; Ye et al., 2003b; Zhao et al., 2000). Additionally, transcription factor such YY1 may also play a role in human histone gene control (Palko et al., 2004). Interestingly, the subnuclear or nuclear localization pattern of NPAT, YY1 and HIRA are all altered in response to replication arrest, suggesting a potential role in same pathway of histone gene repression when DNA

77 replication stalls. Nevertheless, the details of this regulatory mechanism are poorly understood.

3.3.2 Posttranscriptional regulation of histone gene expression

The second strategy that cells use to regulate histone levels is by regulating histone pre-mRNA and mRNA stability, which is believed to play a more important role than the transcriptional control of histone genes in metazoans, particularly in response to inhibition of replication. In comparison to activated transcription (3~5 fold), the posttranscriptional regulation accounts for 8~10 fold increase of histone mRNA during the transition from G1 to S phase (Harris et al., 1991). Since there are no introns in most histone genes, the histone mRNAs only require processing by an endonucleolytic cleavage to become mature mRNA, which is then transported to the cytoplasm and is translated (Osley, 1991). Three decades of efforts by Marzluff and colleagues have revealed fascinating details of histone mRNA regulation in metazoans (Marzluff et al., 2008; Sittman et al., 1983). Replication-dependent histone mRNAs are the only metazoan mRNAs that lack polyadenylated tails, ending instead in a conserved stem-loop. Metazoan histone transcripts contain a conserved stem-loop structure at the 3’ end of the histone mRNAs, as opposed to the polyadenylated transcripts of most other proteins and yeast histones. The 3’ end processing requires binding of the 31 kDa Stem-loop binding protein (SLBP) and recruitment of Histone downstream element (HDE), U7 small nuclear ribonucleoprotein (U7 snRNP) and other factors or endonucleases (Dominski and Marzluff, 2007). The processing of histone pre-mRNA is cell cycle regulated through the SLBP, whose cellular concentration increases 10-fold before the entry of cells into S phase, remains high during S phase and abruptly declines at the end of S phase (Marzluff and Duronio, 2002; Whitfield et al., 2000). SLBP also tends to protect the 3’ end of mRNA from degradation by exonuclease 3’hExo, which is probably responsible for the elimination of histone mRNAs at the end of S phase or when cells encounter replication inhibitors (Dominski et al., 2003). Moreover, similar to the

78 poly (A) tail of other eukaryotic mRNA, the stem loop structure along with SLBP are required for efficient translation of histone mRNA in vivo and in vitro (Sanchez and Marzluff, 2002). Similar to transcriptional control, the SLBP dependent posttranscriptional histone mRNA regulation is also cell cycle regulated. SLBP is found to be recruited to the sites of histone gene transcription (Abbott et al., 1999), and therefore is an ideal candidate for the cell cycle control of histone mRNA abundance. Indeed, translation of SLBP mRNA is regulated during the cell cycle, accumulating just prior to entry into S phase and being rapidly degraded by the proteasome at the end of S phase, similar to the timing of degradation of histone mRNAs (Whitfield et al., 2000). Moreover, consensus cyclin dependent kinase (CDK) phosphorylation site and a consensus cyclin binding site were discovered in the C-terminal of SLBP (Zheng et al., 2003). In addition, downregulation of histone mRNA is also observed when replication is inhibited and various DNA damage checkpoint kinases have been reported to be involved in different studies (Kaygun and Marzluff, 2005a; Muller et al., 2007; Su et al., 2004). Results from these studies indicate that multiple kinases may be functioning in redundant manner to ensure a tight correlation between DNA replication and histone production (possibly via a combination of both transcriptional and post-transcriptional mechanisms).

In yeast, histone mRNAs are polyadenylated, and this stem-loop dependent processing mechanism is absent. However, recent studies suggests that yeast histone mRNA levels might be regulated posttranscriptionally by the poly(A) polymerases Trf4/5 (Reis and Campbell, 2007) and via the Lsm1-7-Pat1 mRNA degradation complex (Herrero and Moreno, 2011).

3.3.3 Posttranslational regulation of histone proteins

The third strategy for the regulation of histone protein levels is histone proteolysis (Gunjan et al., 2006). Generally, histone proteins are considered very stable with half-life up to 4-5 months (Commerford et al., 1982; Tsvetkov et al., 1989). The

79 stability of histone proteins not only ensures the stable packaging of chromatin, but also preserves the epigenetic PTMs on histones. However, in these early studies, the half-lives only reflect the stability of chromatin bound histones. It has been shown that “excess” non-chromatin associated histones are rapidly degraded with a half-life of around 30-40 minutes in budding yeast (Gunjan and Verreault, 2003). In the same study, a post-translational surveillance mechanism that monitors the accumulation of excess histones and triggers histone degradation through the DNA damage checkpoint kinase Rad53 is described (Gunjan and Verreault, 2003). Surprisingly, Rad53 mediated regulation of histone protein level is Mec1/Tel1 independent, which means it can function in the absence of the DNA damage checkpoint pathway. This study reveals that this unique role of Rad53 in dealing with histone degradation on a routine basis is likely to be a house keeping function of Rad53 compared to the “life or death” checkpoint response. Furthermore, recent research from our laboratory indicates that this Rad53-dependent degradation of histones involves modifications of the excess histones including phosphorylation and polyubiquitylation before their proteolysis by the proteasome (Singh et al., 2009a). Rad53 associated histones are phosphorylated in vivo; and the tyrosine 99 residue of histone H3 is critical for the efficient ubiquitylation and degradation of this histone. Ubc4 and Ubc5 have been identified as the ubiquitin conjugating E2 enzymes responsible for histone degradation, whereas Tom1 is believed to be the E3 ligase. Although not all the details of the molecular mechanism involved in this pathway are clear, it provides an excellent method for degrading unneeded histones after their synthesis. The degradation appears to be responsible for not only the removal of histones leftover at the end of S phase, but also for eliminating excess histones when cells encounter DNA damage or replication arrest. However, a similar posttranslational regulation of mammalian histones has not been described yet, but given the highly conserved nature of chromatin regulatory mechanisms, such regulation is very likely to exist in mammalian cells.

3.3.4 Histone chaperones in regulating histone proteins

Last but not least, histones that have been either recently synthesized or displaced from certain regions of the chromatin, need to be dealt with at least until they are either marked for degradation or reassembled on to chromatin. The accumulation of these histones is likely to form a pool of free histones. However, despite the large amounts of histones synthesized to meet the huge demand for histones during their rapid deposition in replication-coupled chromatin assembly, a large pool of free histones is not detected in normal cells. Where do histones go when they are not associated with DNA? In order to manage non-chromatin bound histones more rigorously and keep them from associating non-specifically with negatively charged molecules, cells have also evolved mechanisms that enable histones to be chaperoned by regulatory proteins. Instead of assembling onto chromatin by themselves, histones are assembled through a series of nucleosome assembly factors or histone chaperones in a highly ordered manner (Polo and Almouzni, 2006). In both yeast and higher eukaryotes, H3-H4 heterodimer serves as an assembly unit and is deposited by Chromatin assembly factor1 (CAF1), presumably with the help of another chaperone Anti silencing function1 (Asf1), in a reaction coupled to DNA replication (Smith and Stillman, 1989; Tagami et al., 2004; Tyler et al., 1999). The binding of CAF1 to PCNA (proliferating cell nuclear antigen) promotes the deposition of newly synthesized or parental histones onto the nascent DNA strands and Asf1 is believed to transfer histones rapidly to CAF-1 for immediate deposition (Groth et al., 2005; Sharp et al., 2001). The escort function of these histone chaperones is vital and cells with defects in chaperones CAF1 or Asf1 exhibit increased genomic instability (Myung et al., 2003; Prado et al., 2004). During transcription, chaperones such as the FACT (Facilitates chromatin transcription) complex facilitate transcription by destabilizing nucleosomal structure, thus letting RNA polymerase II (Pol II) access DNA during elongation, and probably accepting and then re-depositing histones back onto the chromatin when transcription is finished (Belotserkovskaya et al., 2003; Morillo-Huesca et al., 2010). Therefore, the escort function of chaperones is not limited to transport and deposition of

81 histones to the places where they are needed, but also to sequester them when histones are not needed and serve as a very small reserve pool of histones as well.

82 CHAPTER 4

EXCESS HISTONE LEVELS MEDIATE CYTOTOXICITY VIA MULTIPLE MECHANISMS

4.1 Abstract

The accumulation of excess histone proteins in cells has deleterious consequences such as genomic instability in the form of excessive chromosome loss, enhanced sensitivity to DNA damaging agents and cytotoxicity. Hence, the synthesis of histone proteins is tightly regulated at multiple steps and transcriptional as well as posttranscriptional regulation of histone proteins is well established. Additionally, we have recently demonstrated that histone protein levels are regulated posttranslationally by the DNA damage checkpoint kinase Rad53 and ubiquitin-proteasome dependent proteolysis in the budding yeast. However, the underlying mechanism/s via which excess histones exert their deleterious effects in vivo are not clear. Here we have investigated the mechanistic basis for the deleterious effects of excess histones in budding yeast. We find that the presence of excess histones saturates certain histone modifying enzymes, potentially interfering with their activities. Additionally, excess histones appear to bind non-specifically to DNA as well as RNA, which can adversely affect their metabolism. Microarray analysis revealed that upon overexpression of histone gene pairs, about 240 genes were either up or downregulated by 2-fold or more. Overall, we present evidence that excess histones are likely to mediate their cytotoxic effects via multiple mechanisms that are primarily dependent on inappropriate electrostatic interactions between the positively charged histones and diverse negatively charged molecules in the cell. Our findings help explain the basis for the existence of multiple distinct mechanisms that contribute to the

83 tight control of histone protein levels in cells and highlight their importance in maintaining genomic stability and cell viability.

4.2 Introduction

Histone proteins are essential for viability and cells need to achieve a very delicate balance between histone and DNA synthesis during the packaging of its genome into chromatin. Histone proteins are regulated transcriptionally(Osley, 1991; Stein et al., 1984), posttranscriptionally (Kaygun and Marzluff, 2005a; Marzluff et al., 2008; Reis and Campbell, 2007), translationally (Borun et al., 1975) and posttranslationally (Gunjan et al., 2006; Gunjan and Verreault, 2003; Singh et al., 2009a). Why are the histone proteins subjected to such a high degree of regulation? This is probably because on one hand scarcity of histones results in inviability (Han et al., 1987), while on the other hand the presence of excess histones has been shown to result in excessive mitotic chromosome loss (Meeks-Wagner and Hartwell, 1986), increased DNA damage sensitivity and cytotoxicity (Gunjan and Verreault, 2003; Singh et al., 2009a). However, the mechanisms underlying these deleterious effects of excess histone accumulation are unclear and so we were interested in investigating them. Our main hypothesis is that due to their high positive charge, histones may exhibit non- specific electrostatic interactions with many negatively charged molecules in the cells, including nucleic acids such as DNA and RNA, as well as negatively charged proteins. This postulate predicts that excess histones could potentially exert their deleterious effects via at least four main mechanisms: (1) Excess histones can compete with and prevent the appropriate assembly of variant histones. This has already been shown to be the case upon overexpression of histone H3 that interferes with the correct deposition of the centromere specific histone H3 variant CENP-A, resulting in a chromosome loss phenotype (Au et al., 2008; Glowczewski et al., 2000). (2) Excess histones can potentially swamp histone chaperones and histone modifying enzymes by binding to them and

84 perhaps tying them up in futile catalytic cycles. We have previously obtained some evidence supporting this possibility by demonstrating that overexpression of certain histone chaperones can alleviate the toxicity due to histone overexpression (Gunjan and Verreault, 2003). (3) Excess histones can stick non- specifically to the DNA, thus affecting chromatin structure and thereby potentially altering gene expression. (4) Similarly, when present in excess, histones can stick to coding as well as non-coding structural and regulatory RNAs, presumably altering translation and other activities of RNA. It should be noted that the four potential mechanisms listed above are not mutually exclusive and it is very likely that one or more or all of these contribute to the deleterious effects of excess histones, although the relative contribution of each mechanism may vary. Here we have systematically explored the last three of the four mechanisms listed above and find that excess histones mediate their deleterious effects via multiple mechanisms in the budding yeast, largely by virtue of inappropriate electrostatic interactions with cellular macromolecules carrying the opposite charge.

4.3 Results

4.3.1 Budding yeast mutants lacking certain histone modifying enzyme subunits are sensitive to the presence of excess histones.

There are many histone modifying and chromatin remodelling enzymes that often function as multi-subunit complexes. Further, many such activities have overlapping roles and function redundantly. We reasoned that some of these non-essential histone modifying activities might be sensitive to the presence of excess histones if one or more of these redundant activities were deleted from the cells. Hence, we screened several yeast deletion strains from the genome deletion collection (Open Biosystems) lacking subunits of non-essential chromatin modifying factors for sensitivity to histone overexpression as described previously (Singh et al., 2009a). We found that deletion strains corresponding to

85 set2 (histone methyltransferase), hda2 (histone deacetylase), sas3 (catalytic subunit of the NuA4 histone acetyltransferase complex) and taf14 (a subunit of INO80 and SWI/SNF chromatin remodelling complexes, NuA4 histone acetyltransferase complex, as well as TFIID and TFIIF transcription factor complexes) were sensitive to overexpression of histone H3 (Fig. 4.1 A), consistent with the idea that in these strains the absence of one histone modifying factor places undue burden on the remaining histone modifying activities still available in the cell, which are quickly overwhelmed upon histone overexpression, which was verified by Western blotting (Fig. 4.1 B). A caveat in our studies is that most of our experiments involve overexpression of just one core histone, while situations involving the generation of excess endogenous histones presumably include all four core histones in excess and may not be fully comparable to our experiments. However, in all the experiments that we have repeated using co-overexpression of two or four core histones (Fig. 4.2 A and data not shown), we have obtained similar results.

4.3.2 Excess histones do not appear to alter the bulk nucleosomal structure but lead to subtle alterations in the fine structure of chromatin.

If excess histones could stick non-specifically to the DNA, it is possible that they can alter the gross and fine structure of chromatin. In vitro, even a slight stoichiometric excess of histones over DNA is sufficient to trigger chromatin aggregation and block transcription (Steger and Workman, 1999). Hence, we next tested if overexpression of histone H3 affects chromatin structure in vivo. For this we analyzed the chromatin structure of cells with or without histone overexpression following cleavage with micrococcal nuclease that cleaves in between nucleosomes and provides low resolution information regarding the underlying bulk chromatin structure (Martens and Winston, 2002). We found that overexpression of histone H3 did not result in any significant changes in the bulk chromatin structure and the nucleosomal ladder generated by micrococcal

Figure 4.1 Yeast cells lacking histone modifying enzyme subunits are sensitive to histone overexpression. (A) Sensitivity of mutant yeast cells lacking histone modifying enzyme subunits to histone overexpression. Wild type (WT) or the indicated mutant strains were transformed with a plasmid carrying galactose inducible HA-epitope tagged H3 (pYES2-HTH-HHT2) or the empty vector (pYES2-HTH) (Singh et al., 2009a). The rad53Δ strain carries the crt1Δ as well to suppress the loss of viability due to the essential nature of Rad53 (Huang et al., 1998). 10-fold serial dilutions of the indicated strains were plated on glucose or galactose media and incubated for 3 days at 30 ℃ prior to being photographed. (B) Histone H3 overexpression in yeast cells lacking histone modifying enzyme subunits. Four independently isolated transformants for each of the indicated mutant strains carrying the plasmid pYES2-HTH-HHT2 were grown overnight in minimal media lacking uracil and with raffinose as the carbon source. Cells were then treated with galactose for 90 minutes prior to harvesting 10 million cells per sample. Whole cell lysates were prepared by the alkaline lysis method (Kushnirov, 2000) and processed for Western blotting to detect both the endogenous and overexpressed epitope-tagged histone H3 using the H3-C polyclonal antibody as described previously (Gunjan and Verreault, 2003; Singh et al., 2009a). Asterisks indicate the transformants used in the experiment shown above in (A).

Figure 4.2 Yeast cells lacking histone modifying enzyme subunits are sensitive to co-overexpression of the histone H3-H4 gene pair. (A.) Sensitivity of mutant yeast cells lacking histone modifying enzyme subunits to co- overexpression of the histone H3-H4 gene pair. Wild type (WT) or the indicated mutant strains were transformed with plasmids carrying galactose inducible HA-epitope tagged H3 (pYES2- HTH-HHT2) and H4 (pYES6/CT-HA3-HHF2) or the corresponding empty vectors (pYES2-HTH) and (pYES6/CT). The rad53Δ strain carries the crt1Δ to suppress the loss of viability due to the essential nature of Rad53 (Huang et al., 1998). 10-fold serial dilutions of the indicated strains were plated on glucose or galactose media and incubated for 3 days at 30°C prior to being photographed. (B.) Co-overexpression of the histone H3-H4 gene pair in yeast cells lacking histone modifying enzyme subunits. The indicated strains used above in panel (A.) that carry the galactose inducible HA-epitope tagged H3 (pYES2-HTH-HHT2) and H4 (pYES6/CT-HA3-HHF2) were grown overnight in minimal media lacking uracil but with raffinose and Blasticidin. Cells were then treated with or without 2% galactose for 90 minutes as indicated prior to harvesting 10 million cells per sample. Whole cell lysates were prepared by the alkaline lysis method47 and processed for Western blotting with HA antibodies to detect the overexpressed HA tagged H3 and H4 histones, while the endogenous histone H3 was detected using the H3-C polyclonal antibody as describe previously (Gunjan and Verreault, 2003) and serves as a loading control.

88 nuclease cleavage was intact (Fig. 4.3A). However, we did notice a subtle but reproducible decrease in the nucleosomal repeat length by ~10–15 bp upon histone H3 overexpression, suggesting that the nucleosomes were closer together and that the internucleosomal linker length was reduced. We also analyzed the micrococcal nuclease cleavage pattern in detail at several loci using indirect end-labeling analysis using primer extension (Axelrod and Majors, 1989). As opposed to the largely unaltered bulk nucleosomal ladder, end-labeled primers specific for the mating type MATa locus and the RDN (ribosomal DNA) locus revealed significant alterations in the micrococcal nuclease cleavage patterns at both these loci (Fig. 4.3B). Similar results were obtained for the actin (ACT1) locus (data not shown). Several micrococcal nuclease cleavage sites are blocked while a few new cleavage sites are generated following histone H3 overexpression, suggesting that although the bulk chromatin structure is unaffected, subtle alterations in the fine structure of chromatin may be widespread in the presence of excess histones.

We decided to confirm the alterations in the fine structure of chromatin upon histone overexpression using high resolution in vivo cleavage of the chromosomal DNA upon the galactose inducible expression of DNase I that cleaves in the minor groove of DNA and hence exhibits a ~10 bp periodicity of cleavage on naked DNA, although DNA bound to proteins shows considerable protection from cleavage (Wang and Simpson, 2001). Indirect end-labeling analysis using primer extension of DNA cleaved with DNase I in vivo confirmed our results obtained with micrococcal nuclease and clearly showed that the fine structure of chromatin at all the three loci (MATa, ACT1 and RDN) analyzed was significantly altered upon histone H3 overexpression (Fig. 4.4). Control experiments using microscopy and immunoprecipitation techniques clearly showed that the overexpressed histone was present in the nucleus and was largely associated with the chromatin (data not shown). Although we have not yet attempted to deduce the exact nature of the changes in chromatin structure that are detected by the altered nuclease cleavage patterns upon histone

Figure 4.3 Histone overexpression does not affect the bulk chromatin structure but alters the fine structure of chromatin. (A) Micrococcal nuclease digestion of bulk chromatin. MNase cleaved DNA isolated from cells in the presence or absence of histone overexpression was purified as described in the Materials and Methods section. This DNA was then resolved on a 2% agarose gel and stained with ethidium bromide to visualize the nucleosomal ladder. The positions of mono-, di-, tri- and tetra- nucleosomes are indicated. (B) Analysis of chromatin structure at specific loci using MNase cleavage. The MNase digested genomic DNA purified above in (A) was used as a template for analysis by indirect end-labeling with multiple cycle primer extension (Axelrod and Majors, 1989) using 32P-end-labeled primers specific to the MATa and RDN loci to perform linear PCR. The reaction products were resolved on a 5% denaturing polyacrylamide gel which was processed for autoradiography. On the right hand side of the parts, a “-“sign indicates loss of a MNase cleavage site while a “+” sign indicates the gain of a site upon GAL-HTH-H3 overexpression.

90 overexpression, it is possible that any such alterations in the fine structure of chromatin could potentially affect DNA transactions such as transcription.

4.3.3 Overexpression of histone gene pairs, but not individual histones, results in the alteration of transcript levels of numerous genes.

To directly assay if the binding of excess histones to chromatin alters the levels of yeast transcripts, we used a microarray based analysis of genome-wide gene expression (Horak and Snyder, 2002). Transcripts corresponding to 5,849 budding yeast genes were analyzed before and after histone overexpression using Nimblegen’s Saccharomyces cerevisiae 4 x 72 K array as described in the Materials and Methods section (Fig. 4.5). Surprisingly, although overexpression of histone H3 alone results in significant toxicity in cell viability assays (Fig. 4.1A) (Gunjan and Verreault, 2003; Singh et al., 2009a) and alters the fine structure of the chromatin (Figs. 4.3B and 4.4), it did not alter transcript levels beyond the 2- fold threshold for significant changes that we had arbitrarily chosen for our microarray experiments (Fig. 4.5A and D). Nevertheless, overexpression of the histone H3 and H4 gene pair resulted in a more than 2-fold change in the transcript levels of ~225 genes (Fig. 4.5B and D and Appendix A Table1), while overexpression of all four core histones resulted in a significant alteration of ~240 transcripts (Fig. 4.5C and D and Appendix A Table 2). About 40% of the transcripts significantly upregulated upon the overexpression of all core histones were also upregulated upon the overexpression of the H3-H4 gene pair alone (Appendix A Table 3). Several of the transcripts altered upon histone gene pair overexpression correspond to essential genes and these may have a significant negative impact on cell viability and thus contribute to the cytotoxic effects of excess histones. However, histone overexpression results in the alteration of only ~4% of the budding yeast transcriptome, which is different compared to the results obtained from the converse microarray experiment involving depletion of histone H4, where a substantial fraction (~25%) of the yeast transcriptome was found to be affected (Wyrick et al., 1999).

Figure 4.4 Indirect end-labelling analysis of chromatin from histone overexpression cells following in vivo DNase I cleavage. Galactose inducible DNase I was expressed in wild type cells either expressing or not expressing GAL-HA-H3 as described previously (Wang and Simpson, 2001). The DNase I cleaved genomic DNA was purified and analyzed by indirect end-labeling with multiple cycle primer extension(Axelrod and Majors, 1989) using a 32P-end-labeled primer specific to the ACT1 locus in addition to the primers for MATa and RDN loci described above in Figure 4.3. The reaction products were resolved on a 5% denaturing polyacrylamide gel which was processed for autoradiography. On the right hand side of the parts, a “-“sign indicates loss of a DNase I hypersensitive site while a “+” sign indicates the gain of a site upon GAL-HTH-H3 overexpression.

Figure 4.5 Genome wide comparison of gene expression in wild type yeast cells upon overexpression of individual or pairs of histone genes. (A) Overexpression of histone H3 alone does not result in appreciable changes in gene expression. Microarray analysis was carried out on total RNA isolated from the wild type W303-1A strains carrying either the empty vector or a galactose inducible histone H3 gene as described in the Materials and Methods section. A scatter plot is shown and illustrates the expression level of genes along the X- and Y-axes representing transcript levels reflecting gene expression values on a log2 scale for the strains indicated. The three solid green lines represent fold-changes in transcript levels from the baseline, which is represented by the green line in the middle and indicates identical transcript levels in the two strains being compared. The upper solid green line represents a 2-fold increase in transcript levels, while the lower green line represents a 2-fold reduction in transcript levels. The dashed purple line is the linear regression with the coefficient of correlation R2 indicated on the bottom right corner. Each transcript is represented by a color coded data point to reflect where it is in comparison to the middle green line, with red color for upregulated transcripts and blue color for downregulated transcripts (the darker the color, the greater the fold-change in the transcript levels). Only the transcript levels of the overexpressed histone H3 (HHT2) was significantly elevated in this experiment and the corresponding data point is indicated by the arrow. (B) Changes in gene expression upon co-overexpression of the histone H3-H4 gene pair. Scatter plot for the indicated strains as in (A). (C) Changes in gene expression upon simultaneous overexpression of all four core histones. Scatter plot for the indicated strains as in (A). (D) Summary of the gene expression data derived from microarray analyses. The number of transcripts altered 2-fold or more upon overexpression of different histones is summarized in a tabular form.

93 Interestingly, further analysis of our microarray data revealed that about 40% of the open reading frames (ORFs) whose transcript levels are significantly affected by histone overexpression are located in “mini clusters” of mainly 2, but occasionally up to 3 – 4 ORFs that lie adjacent to each other either on the Watson or the Crick strand and appear to be coordinately regulated (Appendix A Tables 1–3). Further, the majority (~70%) of these mini clusters are located relatively close to the telomeres and sometimes the centromeres (i.e., the distance between the mini cluster and the telomere or centromere is less than a third of the total length of the chromosome arm on which the cluster is located). One such mini cluster comprises of ORFs (YKR079C, YKR080C, YKR081C) that are upregulated 2~3 fold upon the overexpression of the H3-H4 gene pair and is located on chromosome XI relatively close to the telomere. A mini cluster that is downregulated 2~3 fold upon the overexpression of the H3-H4 gene pair is located on chromosome XIV relatively close to the telomere and comprises of genes (YNR056C, YNR057C, YNR058W) that are involved in biotin biosynthesis and whose expression maybe sensitive to iron levels. Another telomere proximal mini cluster can be found on chromosome XV and comprises of genes that appear to be involved in iron metabolism (YOR382W, YOR383C, and YOR385W) and are downregulated more than 3-fold upon overexpression of the H3-H4 gene pair as well as the overexpression of all core histones. These telomere proximal mini clusters of ORFs that appear to be either coordinately up or downregulated upon histone overexpression may represent loci with certain inherent chromatin structural features such as specific nucleosome positioning or occupancy that may be crucial for their regulated expression under normal conditions. As such, these loci would be particularly sensitive to changes in histone levels as these chromatin structural features could be readily altered upon histone overexpression.

94 4.3.4 Excess histones can bind to RNA.

Alteration in the transcript levels of essential genes is unlikely to explain all of the cytotoxic effects of histone overexpression, since H3 overexpression results in cytotoxicity without affecting transcript levels and as such must be mediating its cytotoxic effects via other mechanisms. Due to the potentially high affinity of the positively charged histones for negatively charged molecules, it is possible that apart from sticking to DNA, excess histones can also stick to RNA. To detect if this was indeed occurring in vivo, particularly in the case of H3 overexpression alone, we performed a RNA Immunoprecipitation (RIP) assay (Gilbert et al., 2004; Gilbert and Svejstrup, 2006) similar to the popular Chromatin Immunoprecipitation (ChIP) assay in the presence and absence of galactose induced epitope tagged HA3-HHT2 (HA-H3) overexpression in a strain carrying a FLAG-tagged chromosomal HHT1 gene encoding FLAG-H3 as well. Using real time quantitative Polymerase Chain Reaction (qPCR), we clearly detected enhanced binding of the overexpressed histone HA-H3 to transcripts from three loci (ACT1, MATa, endogenous histone HHT1 as well as a mixture of endogenous and exogenous histone HHT2) that we assayed for (Fig. 4.6), suggesting that excess histones are capable of binding to RNAs in vivo and potentially altering their activities. Further, no signal was obtained in control qPCR carried out following RNaseA treatment of the immunoprecipitated material, as well as when the Reverse Transcriptase was left out of the reaction, clearly demonstrating that the signals obtained are the result of the genuine binding of histones to RNA. In the same experiment, the binding of endogenous FLAG-H3 (that was not overexpressed) to RNA was barely above the background, suggesting that under normal conditions endogenous histones are not very likely to bind to RNA and potentially alter its functions.

Figure 4.6 Excess histones can bind to RNA. An exponentially growing yeast strain with a FLAG epitope on the chromosomal HHT2 gene (FLAG-H3) and carrying the plasmid pYES6/CT-HA3-HHT2 for galactose inducible HA3-H3 overexpression was treated with or without galactose prior to performing RNA immunoprecipitation (RIP) using FLAG or HA antibodies as described in Materials and Methods. The amount of ACT1, MATa, HHT1 and HHT2 transcripts co-immunoprecipitated using the FLAG and HA antibody beads in the absence of galactose mediated histone overexpression (i.e., the background signal) was arbitrarily set to “1” and the relative binding of HA-H3 to these RNAs upon the addition of galactose is shown here. No signal above background was obtained in control qPCR carried out following RNaseA treatment of the immunoprecipitated material, as well as when the Reverse Transcriptase enzyme was left out of the reverse transcription reaction used to generate the cDNA for qPCR analysis. Error bars represent standard error of the mean from three experiments.

96 4.4 Discussion

We have presented evidence here to suggest that the presence of excess histones negatively impacts numerous cellular processes via multiple mechanisms based largely on potential electrostatic interactions between the highly positively charged histones and negatively charged molecules in the cell. Mutant yeast cells such as rad53Δ, tom1Δ and ubc4Δ ubc5Δ that are defective in the regulation of histone protein levels and harbor excess endogenous histones, are very sensitive to exogenous histone overexpression which is highly toxic for these cells (Gunjan and Verreault, 2003; Singh et al., 2009a). All these mutant strains are also sensitive to DNA damaging agents and exhibit genomic instability in the form of elevated chromosome loss rates. The data presented here allows us to finally explain the deleterious effects of excess histones not just in these mutants, but also in wild type cells. Excess histones can swamp the binding sites available on histone chaperones (Gunjan and Verreault, 2003) and then overload histone modifying enzymes. This can have serious consequences for the normal regulation of gene expression and the formation/maintenance of epigenetic marks on histones. Not surprisingly, cells lacking certain histone modifying enzymes were sensitive to histone overexpression (Fig.4.1A). Further, the stability of histone posttranslational modifications would depend not only on the enzymes that put them on and remove them, but also on the stability of the histone proteins themselves. For example, until the discovery of the first histone lysine demethylase a few years ago (Shi et al., 2004), histone methylation marks were considered to be as stable as histone themselves (Byvoet et al., 1972; Thomas et al., 1972). There is a constant exchange of histones between the chromatin bound and free states as a result of transcriptional eviction , the action of chromatin remodelling factors (Shivaswamy et al., 2008) and the opposing effects of chromatin assembly and disassembly (Kim et al., 2007; Takahata et al., 2009). We have recently obtained evidence that the tyrosine 99 residue of histone H3 may be phosphorylated only when histone H3 is not bound to the

97 chromatin and this modification may serve as a mark to target this histone for degradation in the budding yeast (Singh et al., 2009a). Hence, our findings regarding the degradation of non-chromatin bound histones may have wide- ranging ramifications for the stability and maintenance of epigenetic marks on histones in chromatin by providing an additional layer of regulation.

We also found that excess histones can alter the fine structure of chromatin. This could potentially impact all metabolic activities that require access to the DNA. Hence, we have now systematically analyzed the genome wide effects of excess histones on gene expression using microarrays. We found that although overexpression of histone H3 alone is toxic to the cells (Fig. 4.1A) (Gunjan and Verreault, 2003; Singh et al., 2009a) and alters chromatin fine structure, it did not result in an appreciable alteration of gene expression (Fig. 4.5A). However, overexpression of the H3-H4 gene pair or simultaneous overexpression of all four core histones lead to a significant alteration in the expression levels of 4% of the budding yeast genome, including several genes that are essential for viability. This difference between overexpression of H3 alone or in combination with H4 may be due to the fact that histones are deposited on to the DNA as pairs of H3- H4 and H2A-H2B dimers. Hence, although overexpression of H3 alone may lead to its non-specific association with the DNA, this association may not be very strong in vivo and as such it may not prove to be a significant obstacle for the

transcriptional machinery. On the other hand, (H3-H4)2 tetramers and the core histone octamer are known to associate tightly with the DNA and so upon the overexpression of the H3-H4 gene pair or all the core histones, it is likely that these excess histones will form chromatin structures that may present a significant challenge to the transcriptional machinery (Chang and Luse, 1997) and our microarray data appears to support this notion. Overall, our microarray data suggests that although excess histones are capable of altering the expression of a small fraction of yeast genes that probably contributes to their deleterious effects in vivo, the fact that H3 overexpression does not alter gene expression and is still highly toxic strongly argues that the other mechanisms

98 discussed here are likely to be mediating most of the cytotoxic effects of excess histones.

Numerous studies in recent years have revealed the dynamic nature of chromatin structure; particularly in the context of transcription (Kim et al., 2007; Shivaswamy et al., 2008; Takahata et al., 2009). Our microarray based analysis of yeast genes with altered expression patterns upon histone overexpression revealed that a significant number of them lie in small clusters of 2~4 genes that are coordinately regulated (Appendix A Tables1–3). These clusters could have unique chromatin features that may facilitate the normal regulated expression of genes within them. Such special chromatin features may include promoter regions with highly positioned nucleosomes or promoters and/ or coding sequences that are largely devoid of nucleosomes. A cursory survey of published data on nucleosome occupancy in yeast (Dion et al., 2007)suggests that this may be the case indeed with at least a few of the identified mini clusters. For example, the YNR056C, YNR057C, YNR058W cluster which is downregulated upon histone overexpression (Appendix A Table 1) appears to have poor nucleosome occupancy in yeast based on published data (Dion et al., 2007). It is possible that histone overexpression results in a higher nucleosome occupancy in this region, thereby resulting in lower levels of transcription consistent with our microarray data. On the other hand, the YOR382W, YOR383C, YOR385W cluster that is also downregulated upon histone overexpression appears to have well- positioned nucleosomes based on published genome wide nucleosome occupancy data (Dion et al., 2007). How exactly histone overexpression brings about a reduction in the expression levels of ORFs within mini clusters that may have either poorly positioned or highly positioned nucleosomes is unclear. Future studies involving chromatin fine structure mapping will confirm if there are indeed common chromatin structural features within the identified mini clusters and reveal the exact nature of any chromatin structure changes at these loci upon histone overexpression.

99 Another intriguing feature of the identified mini clusters is their telomere (and occasionally centromere) proximal localization. This is reminiscent of the effect of histone H4 depletion where ~50% of the genes within 20Kb of the telomeres were preferentially derepressed, compared to de-repression of just 15% of the genes genome wide (Wyrick et al., 1999). Any special chromatin features in these clusters may be related to their relative proximity to the telomeres (or the centromeres) that are known to have specialized chromatin structures, although these are not normally known to spread over ~20 Kb from the telomeres in the budding yeast and the closest mini clusters are at least that far away from the telomere (Buhler and Gasser, 2009). Another possibility is that histone overexpression may result in their excessive association with certain loci where they may not be present normally and this in turn may lead to their silencing by the recruitment of Sir proteins, perhaps via subsequent localization of these loci to the nuclear periphery (Taddei et al., 2009). Future studies will reveal whether the relative telomere proximal location of the mini clusters affected by histone overexpression has any functional significance.

As opposed to the potential alterations in transcription caused by excess histones, the process of transcription itself may generate excess histones if the reassembly of the transcriptionally evicted histones onto the chromatin is blocked, as in conditional spt16 mutants, where the evicted histones are presumably subjected to Rad53 mediated histone proteolysis (Morillo-Huesca et al., 2010). In fact, during the G1 phase of the cell cycle, the presence of excess transcriptionally evicted histones in the spt16 mutants or exogenously expressed histones in wild type cells triggers the downregulation of the G1 cyclin CLN3, leading to a delay in S-phase entry. Contrary to a situation of histone excess, it has also been suggested that a lack of adequate histones due to sub-optimal histone pre-mRNA processing may also trigger a similar G1 arrest (Marzluff et al., 2008). Hence, eukaryotic cells may have evolved surveillance mechanisms similar to known cell cycle checkpoints to protect them from the harmful effects of both excess histone accumulation as well as a scarcity of histones. These

100 mechanisms delay S-phase entry by prolonging the G1 phase of the cell cycle, presumably to provide cells with additional time to either degrade excess histones or increase histone mRNA production as needed prior to S-phase entry and start of replication, when excess or inadequate amounts of histones could potentially be highly detrimental for the cells.

Additional problems due to the presence of excess histones may arise from the translational defects occurring as a result of these histones binding to coding RNAs (Fig. 4.6) and interfering with their translation. Further, since the majority of the RNAs in the cell are structural ribosomal RNAs (rRNAs) that are crucial for ribosome function, their non-specific interaction with excess histones would serve as a double setback for ongoing protein synthesis in the cell. Hence, it is not surprising that overexpression of histones is highly toxic and confers lethality in mutants such as rad53Δ that cannot efficiently degrade excess histones (Gunjan and Verreault, 2003). This lethality probably represents the cumulative effects of excess histones by the various mechanisms discussed above. In fact, even wild type yeast cells experience ~20% lethality upon histone overexpression (data not shown) and exhibit genomic instability in the form of enhanced rates of chromosome loss (Gunjan and Verreault, 2003; Meeks-Wagner and Hartwell, 1986; Singh et al., 2009a). Genomic instability is characterized by the increased rate of acquisition of alterations in the genome and is associated with most human cancers (Hoeijmakers, 2009). Taken together, these data suggest that improper histone stoichiometry and aberrant chromatin structure may contribute to genomic instability and carcinogenesis. As such, our studies highlight the potentially crucial role of proper histone stoichiometry and chromatin structure as well as multiple histone regulatory mechanisms in maintaining viability and genomic stability.

101 4.5 Materials and Methods

Yeast strains, plasmids and Western blotting. Yeast strains are listed in Table 4.1. Plasmid pYES2-HTH-HHT2 has been described previously (Singh et al., 2009a) and was used for the galactose induced overexpression of epitope- tagged histone H3 to assay the sensitivity of yeast mutants to histone overexpression in Figure 4.1, as well as for histone H3 overexpression to monitor the effects on chromatin structure in Figure 2. Plasmid pSUN1 has been described elsewhere(Wang and Simpson, 2001) and was used for galactose mediated DNaseI expression in the in vivo footprinting experiment described in Figure 3. Plasmids pYES6/CT-HA3-HHT2 and pYES6/CT-HA3- HHF2 carry galactose inducible H3 and H4 genes respectively sub-cloned between the BamHI and XbaI sites in the multiple cloning site of the high copy 2μ based plasmid with a Blasticidin selectable marker (pYES6/CT from Clontech). Plasmid pHM90 was a gift from Dr. Hiroshi Masumoto and carries a construct for the galactose inducible overexpression of the H3-H4 gene pair (GAL1-10-FLAG- HHT1-HHF1) upon integration at the TRP1 locus, as in the microarray experiments in Figure 4.5. Dr. Mary Ann Osley generously provided p67 (CEN- HIS3-GAL1-10-FLAGHTA2-HTB2), a low copy plasmid for the galactose inducible overexpression of the H2A-H2B gene pair, which was used in our microarray experiments shown in Figure 4.5. Our antibodies and Western blotting procedure have been described in detail elsewhere (Gunjan and Verreault, 2003; Singh et al., 2009a).

Chromatin structure analysis by nuclease digestion and primer extension. Wild type W303-1A cells carrying the empty vector pYES2-HTH or the pYES2- HTH-HHT2 plasmid were grown overnight in minimal media lacking uracil and with raffinose as the carbon source. Equal amounts of cells were then treated with 2% galactose for 4 hours to induce histone H3 overexpression (GAL-HTH- H3) following which they were fixed with 1% formaldehyde for 15 minutes. For each sample, nuclei isolated from 250 million cells were digested with 1 unit

102 of micrococcal nuclease (MNase) for 10 minutes at 37ºC in the presence of 1 mM

CaCl2 as described previously(Martens and Winston, 2002). Following MNase cleavage, the formaldehyde crosslinks were reversed and the cleaved DNA was purified by phenol/chloroform extraction followed by ethanol precipitation. The purified DNA was used as such for analysis by agarose gel electrophoresis (Fig. 4.3A) and primer extension (Fig. 4.3B). Primer extension of the MNaseI digested DNA was carried out essentially as described previously(Axelrod and Majors, 1989) using linear polymerase chain reaction (PCR). For the analysis of chromatin fine structure upon histone overexpression using in vivo footprinting(Wang and Simpson, 2001), wild type W303-1A cells carrying either the pSUN1 plasmid alone or in combination with the pYES6/CT-HA3-HHT2 plasmid were used. The cells were grown in minimal media lacking the appropriate selection markers and raffinose as the carbon source, prior to overnight treatment with 2% galactose. 250 million cells were then harvested for each sample and the DNaseI cleaved DNA was purified by phenol/chloroform extraction followed by ethanol precipitation. The DNA was then digested to completion with the restriction endonuclease StyI and re-purified as before prior to use in primer extension reactions as described previously(Axelrod and Majors, 1989).

Microarray analysis Wild type W303-1a yeast cells harboring an empty vector (pYES2-HTH)(Singh et al., 2009a); a high copy plasmid (pYES2-HTH-HHT2)11 for galactose inducible histone H3 overexpression (GAL-H3); an integrated construct (pHM90) for galactose inducible H3-H4 gene pair (GAL-H3-H4); or GALH3-H4 plus a low copy plasmid (p67) for the galactose inducible expression of H2A-H2B gene pair (GAL-H2A-H2B-H3-H4) were grown to a density of 10 million cells per ml in minimal media lacking appropriate selection markers. Galactose was then added to a final concentration of 2% for 4 hours to induce histone overexpression prior to harvesting 50 million cells for extraction of total RNA using the RNeasy kit from

103 Qiagen following the manufacturer’s instructions. After appropriate quality control analysis of the extracted RNA (on BioRad’s Experian Automated Electrophoresis system), the RNA was reverse transcribed using random hexamers to generate cDNA. The cDNA was then labeled and hybridized in duplicate to the Nimblegen Saccharomyces cerevisiae 4 x 72 K array following instructions provided in the Nimblegen Arrays User’s Guide. The array was scanned and hybridization signals from the histone overexpression samples were normalized to the signals obtained from the empty vector sample. The raw data was analyzed using ArrayStar, before exporting it to Microsoft Excel for presentation.

RNA immunoprecipitation (RIP). The wild type yeast strain (YAG1021) carrying a FLAG-tagged endogenous H3 gene (FLAG-HHT2) at its normal chromosomal location along with the pYES6/CT-HA3-HHT2 plasmid for galactose inducible HA3-H3 overexpression was used for the RIP experiments. This strain allows us to detect the binding of endogenous levels of epitope tagged H3 (FLAG-H3) to RNA in the absence of galactose, while in the presence of galactose it permits the detection of the binding of overexpressed HA3-H3 to RNA. Cells were grown overnight in 100 ml of rich media with raffinose as the carbon source and Blasticidin as the selection antibiotic until they reached a density of 10 million cells/ml. Equal amounts of the culture was then treated with or without 2% galactose for 4 hours prior to addition of 1% formaldehyde to the culture media to allow crosslinking of proteins to nucleic acids for 15 minutes. Crosslinked cells were harvested to prepare whole cell extracts that were sonicated and treated with RNase-free DNase I to digest all DNA. Then RNA immunoprecipitation (RIP) was carried out using equal amounts of whole cell extracts essentially as described previously(Gilbert and Svejstrup, 2006) with a minor modification. To absolutely ensure complete removal of any contaminating DNA in our RIP experiments, we initially treated the whole cell extracts with RNase-free DNaseI and followed this up with a second round of DNaseI treatment of the material immunoprecipitated by the antibody beads as well. FLAG antibody beads were used to immunoprecipitate

104 (IP) endogenous FLAG-H3 (FLAG-IP), while HA antibody beads were used to IP the exogenously overexpressed HA3-H3 (HA IP). The RNAs immunoprecipitated with the antibody beads were recovered after reversing the formaldehyde crosslinks and reverse transcribed to give cDNAs that were quantified by real time quantitative PCR (qPCR) using primers and probes specific for transcripts arising from the ACT1, MATa, HHT1 and HHT2 loci.

105 Table 4.1 List of strains used in the study described in chapter 4

Name Genotype Reference W303-1A MAT a ho ade2-1 trp1-1 can1-100 leu2- (Thomas and 3,112 his3-11,15 ura3-1 Rothstein, 1989) YAG1021 MAT a W303 hht2::FLAG-HHT2-URA3 This Study BY4741 MAT a his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 Open Biosystems YRS216 MAT a BY4741 rad53::HIS3 crt1::KanMx This Study YLD283 MAT a BY4741 sas3::KanMx Open Biosystems YLD295 MAT a BY4741 taf14::KanMx ,, YLD300 MAT a BY4741 hda2::KanMx ,, YLD312 MAT a BY4741 set2::KanMx ,,

106 CHAPTER 5

HISTONE GENE DOSAGE MODULATES DNA REPAIR VIA THE HOMOLOGOUS RECOMBINATION PATHWAY

5.1 Abstract

In eukaryotes, multiple genes encode histone proteins that package the genomic DNA into chromatin and regulate DNA accessibility. When present in excess, the positively charged histones can also bind non-specifically to the negatively charged DNA and affect metabolic processes that require access to the DNA, such as replication, recombination and repair. We have investigated the effect of altered histone gene dosage on the DNA damage sensitivity of budding yeast, Saccharomyces cerevisiae, which has two genes encoding each core histone. Histone overexpression resulted in enhanced sensitivity to a variety of DNA damaging agents, while a reduction in histone H3/H4 gene dosage suppressed the DNA damage sensitivity, even in mutants lacking the DNA damage checkpoint, suggesting that this effect was independent of the DNA damage checkpoint or the damaging agent. Investigations of the HO endonuclease mediated DNA double stand break (DSB) at the MAT locus revealed an increase in DNA repair via Homologous Recombination (HR) upon a reduction in histone gene dosage, while Non-Homologous End Joining (NHEJ) remained unaffected. Further, cells experience greater histone loss around this DSB upon a reduction in histone gene dosage, while the recruitment of HR factors is concomitantly enhanced. We propose that high levels of endogenous histones generated by multiple genes compete with HR factors for binding to DNA, thereby reducing HR efficiency and may normally function to restrain excessive HR activities during S- phase. Our findings could have major implications for DNA repair, genomic

107 stability, carcinogenesis and aging in human cells that have dozens of histone genes.

5.2 Introduction

Histones are essential proteins encoded by multiple genes and help package the DNA into the relatively small nucleus, as well as regulate DNA accessibility(Stein et al., 1984; Wolffe, 1995). However, since histones are positively charged, when present in excess they can bind non-specifically to negatively charged DNA and affect all aspects of DNA metabolism, including DNA repair (Gunjan and Verreault, 2003; Singh et al., 2009a; Singh et al., 2010). Not surprisingly, the accumulation of excess histones is harmful for the cells and results in genomic instability and enhances the sensitivity of budding yeast to DNA damaging agents (Gunjan and Verreault, 2003; Meeks-Wagner and Hartwell, 1986). All eukaryotes carry multiple genes encoding each core histone protein, ranging from two genes for each core histone in the budding yeast to several hundred in other popular model systems like Drosophila and Xenopus (Stein et al., 1984). Each diploid human cell has 28 copies of histone H4 gene alone that encode identical H4 proteins (Marzluff et al., 2002). Of the multiple genes encoding each histone protein in eukaryotes, some are nonallelic variants that may have specialized functions (Stein et al., 1984; Wolffe, 1995). However, the rationale behind the existence of multiple genes encoding the same histone protein is unclear. Since histones are essential for viability, one possibility is that the multiple histone genes simply serve as a backup in case of inactivating mutations in one or more genes. However, this is unlikely as the majority of genes essential for viability in eukaryotes have no additional gene copies to serve as backup. A second possibility is that multiple histone genes may be required to synthesize the enormous quantities of histones that are required for chromatin assembly during DNA replication. However, several studies over the past two decades have clearly shown that the full complement of the multitude of histone genes are

108 not all required for maintaining bulk chromatin structure or viability in several species, including the budding yeast (Norris et al., 1988; Norris and Osley, 1987), fission yeast (Castillo et al., 2007), chicken (Takami and Nakayama, 1997a, b; Takami et al., 1997) and mice(Fan et al., 2005; Fan et al., 2001; Lin et al., 2000; Sirotkin et al., 1995) . A potential problem that may arise with the presence of multiple copies of histone genes being driven by very strong promoters is that more histones may end up being synthesized than what is required for chromatin assembly and maintenance (Singh et al., 2009b). To protect the cells from the deleterious effects of excess histone accumulation, histone levels are tightly regulated transcriptionally (Kaygun and Marzluff, 2005b; Osley, 1991), posttranscriptionally (Dominski et al., 2003; Kaygun and Marzluff, 2005a; Marzluff and Duronio, 2002; Reis and Campbell, 2007), translationally (Borun et al., 1975; Graves et al., 1987) as well as posttranslationally (Gunjan and Verreault, 2003; Singh et al., 2009a). These mechanisms ensure a tight coupling between the levels of DNA and histone synthesis and downregulate histones in response to replication arrest or DNA damage during S-phase (Kaygun and Marzluff, 2005a; Lycan et al., 1987; Singh et al., 2009b; Sittman et al., 1983).

To preserve genomic integrity, cells have evolved a number of highly efficient DNA surveillance and repair mechanisms to detect and repair DNA damage caused by both external and endogenous genotoxic agents. Defects in these mechanisms increase the incidence of mutations and genome instability, which are implicated in oncogenesis (Kastan and Bartek, 2004). Upon DNA damage or replication stress, checkpoint responses arrest the cell cycle to provide additional time for efficient repair (Weinert et al., 1994). In the budding yeast, two essential protein kinases, Mec1 and Rad53, play multiple roles in the DNA damage and replication arrest response (Harper and Elledge, 2007; Lowndes and Murguia, 2000). DNA damage in the budding yeast leads to the Mec1/Tel1 dependent hyperphosphorylation of Rad53 and a dramatic increase in its kinase activity (Sanchez et al., 1996) . Activation of Rad53 triggers a phosphorylation mediated

109 cascade of events that bring about all the known responses to DNA damage. Mec1 and Rad53 are also required to prevent both spontaneous and DNA damage-induced collapse of replication forks (Desany et al., 1998; Tercero and Diffley, 2001) and block the firing of late origins (Santocanale and Diffley, 1998). As a result, DNA damage leads to an abrupt decrease in DNA synthesis (Paulovich and Hartwell, 1995). Hence, it is not surprising that both mec1 and rad53 mutants are exquisitely sensitive to DNA damaging agents and strong genetic suppressors of their DNA damage sensitivity are not known. The contribution of histones in regulating DNA repair has only recently begun to be evaluated. Using a tetracycline regulated histone H4 gene, it was reported that a reduction in the level of histone H4 results in replication fork collapse, elevated recombination and genomic instability in budding yeast cells (Clemente-Ruiz and Prado, 2009; Prado and Aguilera, 2005). However, the application of these findings to wild type cells is complicated by that fact that the substantial depletion of histones in these studies resulted in pleiotropic effects on cell physiology including chromatin structure alterations, slow growth, spontaneous replication fork collapse and DNA damage. In fact, the results obtained upon the depletion of histone H4 in yeast cells are very reminiscent of the spontaneous DNA damage and S-phase arrest observed in human cells upon inhibition of chromatin assembly (Ye et al., 2003a). On the other hand, murine embryonic stem cells with reduced histone H1 levels were reported to be resistant to DNA damaging agents, although the DNA repair pathways mediating these effects were not evaluated (Murga et al., 2007). In recent years we have uncovered a novel role for the budding yeast checkpoint kinase Rad53 in histone metabolism (Gunjan and Verreault, 2003; Singh et al., 2009a). Rad53, but not Mec1, is required for degradation of excess histones that are not packaged into chromatin. As a consequence, rad53 mutants accumulate abnormally high amounts of soluble histones and are sensitive to histone overexpression. The DNA damage sensitivity, slow growth and chromosome loss phenotypes of rad53 mutants can be significantly suppressed by disrupting one of the two loci encoding histones H3/H4, arguing that these phenotypes are partly due to the presence of excess

110 histones. Taken together, these data strongly suggest that there may be an intimate connection between DNA damage sensitivity, DNA repair processes and histone dosage.

In this study we have investigated how changes in histone gene dosage affect the DNA damage sensitivity of budding yeast cells. We discovered that a reduction in histone H3 and H4 gene dosage in results in a significant resistance to all common DNA damaging agents tested. This appears to be largely due to an increase in the efficiency of repair via the HR pathway (Aylon and Kupiec, 2004) upon a reduction in histone gene dosage, while NHEJ remains unaffected. The observed effects on DNA damage sensitivity upon changes in histone gene dosage are not associated with global changes in the expression of DNA repair genes, alterations in the gross chromatin structure or the DNA damage checkpoint. Using the GAL-HO system (Haber, 2002; Holmes and Haber, 1999) in budding yeast strains with intact silent mating type loci, we found that cells with reduced histone gene dosage experience greater histone loss around the lesion, while the recruitment of several key repair and HR factors to the lesion was concomitantly enhanced. Based on these data, we propose that excess histones compete with the HR machinery and prevent it from accessing the lesion, thus reducing the efficiency of repair by HR. As such, even a moderate reduction in histone gene dosage enhances the efficiency of DNA repair and makes cells more resistance to DNA damaging agents. Further, although the reasons behind the existence of multiple histone genes in eukaryotes are unclear, our results suggest the possibility that they may be required to generate high levels of histones to potentially suppress HR, thereby protecting cells against the deleterious effects of excessive recombination, particularly during S-phase (Gangloff et al., 2000; Kerrest et al., 2009; McMurray and Gottschling, 2003; Schmidt and Kolodner, 2006; Yamagata et al., 1998). Overall, our findings may have major implications for DNA repair, genomic stability and carcinogenesis in human cells that have dozens of histone genes (Marzluff et al., 2002).

111

5.3 Results

5.3.1 Histone gene dosage has a major influence on the sensitivity of budding yeast cells to DNA damaging agents

The budding yeast Saccharomyces cerevisiae has two copies of each core histone gene (Fig. 5.1a) and only one copy is required for survival (Osley, 1991). Of the two gene pairs (HHT1-HHF1 and HHT2-HHF2) encoding histone H3 and H4, the second gene pair HHT2-HHF2 contributes 6-8 times more histone mRNA during S-phase than the first gene pair HHT1-HHF1 (Cross and Smith, 1988). However, any one of the two H3-H4 gene pairs is sufficient for viability, despite the lack of dosage compensation by the remaining gene pair. These characteristics of the budding yeast histone H3 and H4 genes make it an ideal system to study the relevance of multiple genes encoding the same histone protein in eukaryotes. We have previously reported that histone overexpression increases the DNA damage sensitivity of the checkpoint mutant rad53 (that is also defective in degrading excess histones), while a reduction in histone gene dosage results in resistance to DNA damaging agents (Gunjan and Verreault, 2003; Singh et al., 2009a). However, any potential influence of histone dosage on DNA repair in wild type cells has not been evaluated previously. Hence, we first assayed the effect of histone overexpression on the DNA damage sensitivity of wild type W303 and isogenic DNA damage checkpoint mutant yeast strains as controls (Fig. 5.1b). Consistent with our previous findings (Gunjan and Verreault, 2003; Singh et al., 2009a), rad53 mutants were extremely sensitive to the strand- break reagent Bleocin (Chen and Stubbe, 2005), particularly in the presence of histone overexpression. Even the wild type cells were more sensitive to DNA damage in the presence of histone overexpression, suggesting that excess histones can presumably block efficient DNA repair in wild type cells as well. Since histone overexpression resulted in an increase in the DNA damage sensitivity of all strains tested, we wondered if a reduction in histone gene

112

Fig. 5.1 Histone gene dosage influences the DNA damage sensitivity of wild type and DNA damage checkpoint deficient yeast strains. (a) Organization of histone H3 and H4 genes in the budding yeast. The paired histone genes are transcribed divergently from a common promoter. Regulatory elements in the promoter are indicated. uas = Upstream activating sequence; neg = Negative element; tata = TATA box; HHT = Histone H3 gene; HHF = Histone H4 gene. The relative levels of total histone mRNA for H3 and H4 contributed by each gene pair is indicated. (b) Overexpression of histone H3 increases DNA damage sensitivity of yeast strains.Indicated strains transformed with either a vector encoding galactose inducible histone H3 (pYES2- HTH-HHT2) (Singh et al., 2009a) or the empty vector (pYES2) were plated in 10-fold serial dilutions on glucose or galactose media with or without Bleocin. The plates were incubated for 4 days at 30°C prior to being photographed. (c) Deletion of one of the hht2-hhf2 histone gene pair encoding histone H3 and H4 decreases DNA damage sensitivity of yeast strains. Ten-fold serial dilutions of exponentially growing yeast strains of the indicated genotypes were plated and treated with the indicated amounts of various DNA damaging agents. UV = Ultra Violet light; HU = Hydroxyurea; MMS = Methyl Methane Sulfonate. The plates were incubated for 4 days at 30°C prior to being photographed.

113 dosage could reduce DNA damage sensitivity of yeast strains. To test this idea, we next examined the effect of deletion of the hht2-hhf2 gene pair on the DNA damage sensitivity of wild type as well as the rad53 and mec1 DNA damage checkpoint mutant strains to a variety of common DNA damaging agents (Fig. 5.1c). Compared to the wild type strain, there is only 14~20% histone H3-H4 mRNA in the strains carrying hht2-hhf2 deletions (Cross and Smith, 1988). Mec1 is the homolog of the essential human checkpoint kinase ATR and regulates all aspects of the DNA damage response in budding yeast (Harper and Elledge, 2007). Surprisingly, deletion of the hht2-hhf2 gene pair resulted in a substantial decrease in the sensitivity of not just the rad53 mutant strains that accumulate excess histones, but mec1 mutant strains as well that do not accumulate excess histones (Gunjan and Verreault, 2003; Singh et al., 2009a). In fact, the mec1 mutant was only as sensitive as the wild type W303 cells to DNA damage with significantly high amounts of Methylmethane Sulfonate (MMS) and Bleocin (Fig. 5.1c). Such a dramatic suppression of the DNA damage sensitivity of DNA checkpoint mutants such as mec1 towards a variety of different DNA damaging agents is unprecedented and highlights the enormous influence that histone levels can have on DNA damage sensitivity. Even wild type W303 cells carrying the hht2-hhf2 deletion were more resistant to DNA damaging agents than their wild type counterparts with the full complement of histone genes and this effect was more prominent at higher does of the damaging agent (Fig. 5.1c, compare the sensitivity to 200mM hydroxyurea, HU and 0.0125% MMS). These results imply that histone gene dosage are a major influence on the DNA damage sensitivity of budding yeast cells and even normal wild type levels of histones can potentially interfere with efficient repair.

5.3.2 Changes in histone gene dosage have little effect on the bulk chromatin structure or global gene expression

We next sought to understand the mechanism/s via which histone gene dosage may influence the DNA damage sensitivity of yeast cells. Since we have recently

114 published the effects of histone overexpression on yeast cells in extensive detail (Singh et al., 2010), we will focus mainly on the effects of histone gene deletion on yeast cells for the rest of the paper. As histones are a key component of the chromatin, it is reasonable to hypothesize that an alteration of histone gene dosage may change chromatin structure. To determine whether global chromatin structure is altered in strains carrying histone gene deletions, we performed micrococcal nuclease (MNase) digestion analysis of chromatin from wild type and hht2-hhf2 deletion strains (Figure 5.2). Consistent with previous reports (Clark-Adams et al., 1988; Norris et al., 1988), the kinetics and pattern of MNase digestion was identical between wild type and hht2-hhf2Δ deletion strains, suggesting that the bulk chromatin structure is not altered upon histone gene deletion. Chromatin structure is generally considered as a barrier to gene expression by inhibiting the binding and function of the transcriptional machinery (Workman and Kingston, 1998). We have recently reported changes in the fine structure of chromatin as well as the gene expression patterns of ~4% of the genome upon an increase in histone dosage (Singh et al., 2010). Thus, another possible effect of a reduction in histone gene dosage could be changes in the expression of genes (Clark-Adams et al., 1988), especially those genes known to be involved in DNA damage response (DDR) (Jelinsky and Samson, 1999). We investigated this possibility using genome wide gene expression analysis using microarrays (Horak and Snyder, 2002). Compared to the wild type cells, the expression of ~50 genes were significantly affected (expression changes of more than 2-fold) in the hht2-hhf2 deletion strain Figure 5.3; Appendix B), none of which are known to be involved in the DDR (Jelinsky and Samson, 1999). These results suggest that yeast cells can form an intact and functional chromatin structure even if there are fewer histones available in hht2-hhf2Δ strains. This is not surprising given that yeast cells normally appear to produce more histone proteins than is necessary for packaging their genome into chromatin (Singh et al., 2009b).

115

Fig. 5.2 Global chromatin structure remains unchanged upon deletion of the hht2-hhf2 histone gene pair. Yeast nuclei extracted from the indicated strains were treated with 1 unit of MNase for increasing time periods (5’ and 10’) and purified using phenol:chloroform extraction. The purified DNA was then separated on a 2% agarose gel and visualized by staining with ethidium bromide. Bands corresponding to mono-,di-, tri-, tetra-nucleosomes are indicated by arrows.

116 5.3.3 The DNA damage checkpoint is efficiently activated in cells with reduced histone dosage

The observed effect of a reduction in histone gene dosage on the DNA damage sensitivity of yeast cells (Fig. 5.1c) could potentially be due to differences in the DNA damage checkpoint activation. Hence, we also investigated if there were any differences in DNA damage checkpoint activation between wild type and hht2-hhf2Δ strains. To evaluate the activation of the DNA damage checkpoint, we first measured the budding index of these strains following the release of cells from alpha-factor mediated G1 arrest into HU. In the presence of HU, both the wild type and hht2-hhf2Δ strains arrested as large budded cells with similar kinetics and remained arrested for at least an hour after removal of HU, suggesting that their checkpoint response to HU was intact (Figure 5.4). In the same experiment, the checkpoint defective mec1 cells progressed through mitosis and into the next cell cycle within an hour of HU removal. We also used Pulsed Field Gel Electrophoresis (PFGE) (Schwartz and Cantor, 1984) to visualize the presence of ongoing replication in HU treated cells (Figure 5.5) as well as flow cytometry to follow the extent of replication in the presence of MMS (Figure 5.6). However, we were unable to detect any significant differences in the activation of the DNA damage cell cycle checkpoint between wild type and hht2- hhf2Δ using these assays. Further, we did not detect any significant differences in the rate of S-phase progression between the wild type and hht2-hhf2Δ strains.

117

Fig. 5.3 Genome wide comparison of gene expression in wild type yeast cells to yeast cells carrying a deletion of the hht2-hhf2 gene pair. Microarray analysis was carried out on total RNA isolated from exponentially growing wild type W303-1a or hht2-hhf2Δ cells following overnight growth in YP-galactose media essentially as described in Singh et al., 2010. A scatter plot is shown and illustrates the expression level of genes along the X- and Y-axes representing transcript levels reflecting gene expression values on a log2 scale for the strains indicated. The three solid green lines represent fold-changes in transcript levels from the baseline, which is represented by the green line in the middle and indicates identical transcript levels in the two strains being compared. The upper solid green line represents a 2-fold increase in transcript levels, while the lower green line represents a 2-fold reduction in transcript levels. The dashed purple line is the linear regression with the coefficient of correlation R2 indicated on the bottom right corner. Each transcript is represented by a color coded data point to reflect where it is in comparison to the middle green line, with red color for upregulated transcripts and blue color for downregulated transcripts (the darker the color, the greater the fold-change in the transcript levels). For clarity, transcripts that show a two-fold or greater change are represented by white data points.

118

Fig. 5.4 The DNA damage checkpoint remains intact and responds normally to replication inhibition upon deletion of the hht2-hhf2 histone gene pair. Budding index of the indicated yeast strains was calculated after release from alpha- factor mediated G1 arrest into media with or without the replication inhibitor hydroxyurea (HU, 200mM). Cells were collected at 20 minute intervals. The percentage of large budded cells (i.e., cells arrested at G2/M) is plotted against time to monitor the progression of cells through the cell cycle. All strains complete replication and cell division to enter the next cell cycle in the absence of HU. However, in the presence of HU, all cells arrest as large budded cells by 120 minutes, at which time the HU was washed away. Following the removal of HU, wild type W303 and hht2-hhf2 deletion strains remain arrested in G2/M as large budded cells, while many of the mec1 mutant cells lacking a functional DNA damage checkpoint proceed undergo cell division to enter the next cell cycle within 60 minutes of HU removal.

119

Fig. 5.5 Deletion of the histone gene pair hht2-hhf2 does not alter the intra- S checkpoint. Wild type and hht2-hhf2 deletion strains were arrested with α-factor for 120’ before releasing in the presence or absence of 200mM HU. Cells released from α-factor into HU were incubated in HU for 90’ before washing it away. Cells were harvested at indicated time points. Spheroplasts were generated by treating the sample cells with 5mg/ml Zymolyase for 1hr at 30℃ with gentle shaking and embedded in agarose plugs. Pulsed-Field Gel Electrophoresis (PFGE) was performed to resolve the fully replicated budding yeast chromosomes as described previously (Schwartz and Cantor, 1984). In the presence of HU, replication stalls in both wild type and hht2-hhf2 deletion strains and this is indicated by the lack of the chromosomal bands in the gel, since replication intermediates are not resolved by PFGE as they do not migrate out of the wells. It takes up to 60 minutes for both these strains to complete DNA replication after the removal of HU before the full length chromosomes are resolved again. This indicates the presence of a functional intra-S phase checkpoint in both these strains. Further, no differences were observed in the kinetics of DNA replication or checkpoint activation between these two strains.

120

Fig. 5.6 Flow cytometry analysis of the progress of replication in response to low doses of the methylating agent Methylmethane Sulfonate (MMS). Exponentially growing wild type and hht2-hhf2 deletion strains were arrested with α-factor for 2hr before releasing the cells to progress through S-phase in the presence or absence of a low dose of MMS (0.033%) which greatly slows down replication, but does not stop it completely (Tercero and Diffley, 2001). Cells were collected at the indicated time points and resuspended in 70% ethanol. Cells were removed from ethanol, rehydrated, treated with RNaseA and stained with 30μg/ml propidium iodide (PI) before analyzing their DNA content by flow cytometry. Although both the strains completed DNA replication within 60 minutes of release from α -factor in the absence of MMS, neither of the two strains had completed DNA replication in the presence of MMS even after 80 minutes of release from α -factor, suggesting that the normal DNA damage checkpoints are functional in both these strains.

121 5.3.4 Reduction in histone gene dosage enhances survival of yeast cells following a DNA double strand break (DSB) in a checkpoint independent manner

Since a reduction in histone gene dosage results in resistance to a variety of DNA damaging agents that cause different kinds of DNA lesions and are often repaired by different repair pathways (Fig. 5.1c), we next turned our attention to potential DNA repair pathway/s that may be utilized in common for the repair of a variety of different DNA lesions. Several different kinds of DNA lesions may be processed by different repair machineries to ultimately generate DSBs (McKinnon and Caldecott, 2007) that can be repaired by the two competing pathways of homologous recombination (HR) or Non-Homologous End Joining (NHEJ) (Aylon and Kupiec, 2004). A DSB is perhaps the most harmful kind of DNA damage that a cell could incur and this event could potentially lead to loss of the entire chromosome arm. In fact, even a single unrepaired DSB can cause lethality (Bennett et al., 1993). An in vivo model system (Fig. 5.7) has been developed in budding yeast cells using galactose inducible-HO endonuclease (GAL-HO) expression (Haber, 1998, 2002; Hicks et al., 2011; Holmes and Haber, 1999) for the study of DSB repair. In this system, a single DSB at the naturally occurring HO cleavage site in the mating type (MAT) locus on chromosome III can be created upon GAL-HO expression in budding yeast cells lacking a functional endogenous HO gene (Fig. 5.7). This DSB at the MAT locus is predominantly repaired via the multi-step and multi-factorial process of HR (Aylon and Kupiec, 2004), where the broken ends are first resected in a 5’ to 3’ direction, giving rise to 3’ overhangs with single stranded DNA (Huertas, 2010). Then, one of the two silent mating loci (HMLα and HMRa) adjacent to the MAT locus is used as a donor template and copied via new DNA synthesis to accurately repair the DSB at the MAT locus (Aylon and Kupiec, 2004). NHEJ involves the simple ligation of the broken ends and normally repairs the DSB at MAT locus at a much lower frequency than HR in the presence of the silent mating loci, but becomes

122 the only way to repair the DSB at MAT locus when the silent mating type loci are deleted.

Next we decided to test whether DSB repair was indeed affected upon deletion of the hht2-hhf2 gene pair by inducing a single defined DSB in the yeast genome using GAL-HO. For this we first stably integrated the GAL-HO construct at the ADE3 locus in wild type, hht1-hhf1Δ and hht2-hhf2Δ strains with intact silent mating type loci. Five-fold serial dilutions of these strains were plated on glucose or galactose containing plates. Compared to wild type, the hht2-hhf2Δ strain showed a dramatic improvement of viability on galactose containing plates, while hht1-hhf1Δ strain appeared to exhibit a small but reproducible improvement in survival on galactose media (Fig. 5.8a). This result was confirmed by a viability assay to quantify the ratio of viable colonies formed on galactose versus glucose plates, which revealed that the survival of the hht2-hhf2Δ strain following the GAL-HO mediated DSB at the MAT locus was roughly ~5000-fold better than the wild type strain (Fig. 5.8b). This effect on viability of the hht1-hhf1Δ and hht2- hhf2Δ strains following GAL-HO cleavage mirrors the relative transcriptional output of the HHT1-HHF1 and HHT2-HHF2 gene pairs. Since the HHT2-HHF2 gene pair contributes 80~86% of the histone H3-H4 mRNAs in the cell (Cross and Smith, 1988), it strongly argues that the observed effect of the hht2- hhf2Δ strain on viability in response to the GAL-HO break is due to the differences in endogenous histone levels in the cells carrying histone gene deletions. Due to the much milder effect of the hht1-hhf1 deletion on the DNA damage sensitivity compared to the dramatic effects observed upon hht2-hhf2 deletion (Fig. 5.8 a, b), we decided to focus mainly on the hht2-hhf2Δ strains for the rest of our studies.

123

Figure 5.7 Galactose inducible DSB at the MAT locus. Galactose inducible endonuclease HO will generate a specific double strand break (DSB), which will be preferably repaired by homologous sequences HML (HMR for MATa).

124

Fig. 5.8 A reduction in histone gene dosage enhances survival of budding yeast cells following HR mediated repair of an induced DSB. (a) Survival of strains following induction of HO mediated DSB. The indicated yeast strains carrying galactose inducible HO endonuclease (GAL-HO) were grown overnight in YP-raffinose media to reach a concentration of 1~2x107 cells/ml. Then 5-fold serial dilutions of the indicated strains were plated on glucose or galactose plates. The plates were incubated for 3 days at 30°C prior to being photographed. (b) Quantitation of the survival of yeast strains following induction of HO mediated DSB. The viability of the strains was tested by plating 1000 cells from each strain in (a) on YP-glucose media or 100,000 cells on YP-galactose media in duplicate. The viability was quantitated by normalizing the number of colonies formed on galactose to the number of colonies formed on glucose media. The error bars represent standard error of the mean from three independent repeats of the experiments under identical conditions. (c) Rad53 hyperphosphorylation following induction of GAL-HO mediated DSB. Exponentially growing wild type and hht2-hhf2Δ strains carrying the GAL-HO construct were treated with 2% galactose and cells were harvested every hour thereafter. Whole cell extracts were prepared as described (Kushnirov, 2000) and Western blotting to detect phosphorylated Rad53 was carried out as described previously (Gunjan and Verreault, 2003). (d) Survival of rad53Δ strains following induction of HO mediated DSB. Cell growth was assayed as described above in (a). (e) Survival of mec1Δ strains following induction of HO mediated DSB. 10-fold serial dilutions of the indicated strains were plated on glucose or galactose plates and cell growth was assayed as described above in (a).

125 Lethality due to the GAL-HO mediated DSB in the continuous presence of galactose has been linked to the persistent activation of the DNA damage checkpoint, resulting in the prolonged arrest of the cells in G2/M (Lee et al., 1998; Sandell and Zakian, 1993; Toczyski et al., 1997). Although we did not find any differences in checkpoint activation between the wild type and hht2-hhf2Δ strains (Figures 5.4, 5.5, 5.6), it still possible that the hht2-hhf2Δ strain somehow fails to activate the DNA damage checkpoint appropriately in response to a single DSB. To test this possibility, we monitored phosphorylation of checkpoint protein Rad53 following the addition of galactose to wild type and hht2-hhf2Δ strains carrying the GAL-HO construct (Fig. 5.8c). We could detect Rad53 phosphorylation 2 hours after galactose addition in both wild type and hht2- hhf2Δ strains and this persisted for several hours. Importantly, we did not observe any significant differences in Rad53 phosphorylation upon galactose addition in these two strains, suggesting that both the strains activate the DNA damage checkpoint appropriately. Further, the rad53 deletion strain that is defective in the DNA damage checkpoint also showed much better survival of GAL-HO mediated DSB upon the deletion of hht2-hhf2 (Fig. 5.8d). Similar results were also observed for the mec1 deletion strain that lacks a functional DNA damage checkpoint but nevertheless exhibited a remarkably improved survival of the GAL-HO mediated DSB upon the deletion of hht2-hhf2 (Fig. 5.8e). This data strongly argues that the better survival of yeast strains carrying the GAL-HO construct on galactose media is independent of a functional DNA damage checkpoint. However, we did observe a consistent difference in H2A phosphorylation between wild type and hht2-hhf2Δ strain upon MMS treatment in S phase. H2A phosphorylation has been recognized as one of the earliest markers of DSBs (Chambers and Downs, 2007; Downs et al., 2000). Interestingly, after releasing alpha-factor arrested yeast cells into S-phase, when compare to the wild type strain, the H2A phosphorylation appeared earlier in the hht2- hhf2Δ strain and was maintained at lower levels for the rest time points in response of MMS (Fig. 5.9a). It seems that hht2-hhf2Δ strain is primed for H2A

126 phosphorylation when cells just enter S phase and this may result in more efficient checkpoint signaling leading to faster repair kinetics, resulting in an overall lower accumulation of H2A phosphorylation following DNA damage. Moreover, H3K56 acetylation has been shown to be required for efficient DSB repair, especially for DSBs that occur during DNA replication (Masumoto et al., 2005). Recent evidence suggests that this modification plays an important role in chromatin dynamics during DNA replication and DNA damage repair (Chen et al., 2008a; Li et al., 2008). Histones with this modification are found in association with histone chaperones Asf1 and CAF-1, presumably marking newly synthesized or free histones evicted from the lesion, which are subsequently deposited onto the chromatin following repair. By monitoring H3K56 acetylation after MMS treatment in S-phase, we observed attenuation of this epigenetic mark in the hht2-hhf2Δ strain compared to wildtype cells (Fig. 5.9a), which is also consistent with faster repair kinetics in these cells.

Another potential reason for the dramatic increase in the survival of the hht2- hhf2Δ strain carrying the GAL-HO construct on galactose media compared to wild type cells could be that this strain mutates the HO cleavage site at the MAT locus making it refractory to further cleavage. We investigated this directly by sequencing the HO cleavage site in 17 colonies of the hht2-hhf2Δ strain isolated from galactose plates and found that they all had identical wild type HO cleavage sites. Further, the hht2-hhf2Δ strain did not show any increase in mutation frequency compared to wild types cells as assayed by the reversion rates of the ade2-1 mutation in these strains. Hence, all our results are consistent with the hht2-hhf2Δ strain repairing the GAL-HO mediated DSB at the MAT locus with

high fidelity and efficiency.

127 Fig. 5.9 Change of damage signaling and HR dependent repair upon hht2- hhf2 deletion (a) Damage signaling is activated earlier but maintained at lower levels due to more efficient repair in hht2-hhf2 deletion strains. Exponentially showing cells were arrested in G1 with α-factor for 2 hours then released into rich media to progress through S-phase. 0.033% MMS was added 15’ after the release from G1 arrest and samples containing 1x107 cells were taken at the indicated time points. Protein extracts from all samples were prepared and the proteins were resolved through SDS 4-12% polyacrylamide gradient gel and processed for Western blotting with either anti-phospho-H2A, anti-H3 K56 or H3 antibody. (b) Survival of the strains with rad52 deletion upon GAL-HO induction. 10-fold serial dilutions of the indicated strains were plated on glucose or galactose plates and cell growth was assayed as described above in Fig.5.8 (a).

128 5.3.5 Improved survival of strains carrying a reduced histone dosage following DSB induction depends on homologous recombination and not on non-homologous end joining

The HO-mediated DSB at the MAT locus is predominantly repaired by the HR pathway in the presence of the homologous donor sequences, but NHEJ may also play a role (Aylon and Kupiec, 2004; Haber, 1998, 2002; Hicks et al., 2011; Holmes and Haber, 1999). Hence, we next tested whether the improved survival of strains lacking hht2-hhf2 to a GAL-HO mediated DSB was dependent upon the HR or NHEJ mediated repair. For this we used a strategy similar to the one used in Fig. 5.8 a and b, except that the GAL-HO carrying strains now had the HMRa and HMLα sequences deleted from the silent mating type loci to prevent HR mediated repair of the HO mediated DSB at the MAT locus and force the repair to occur exclusively via the NHEJ pathway. Ten-fold serial dilutions of these strains were plated on glucose or galactose containing media and their growth was monitored over several days (Fig. 5.10a). Although the growth on galactose media was very poor presumably due to the repeated cycles of DSB repair and cleavage by HO in the continuous presence of galactose, the improved survival of strains lacking the hht2-hhf2 gene pair was completely abolished upon the deletion of HMRa and HMLα sequences, and no significant differences were observed between the different strains. To obtain more quantitative data, a survival assay was also carried out using these cells and the repair efficiency was estimated by comparing the number of colonies formed on galactose versus glucose plates, but once again no significant differences were observed in the survival of the strains (Fig. 5.10 b). Additionally, the deletion of HR factor Rad52 in the hht2-hhf2Δ strain also abolished most of the improved survival of the hht2-hhf2Δ strain observed on galactose plates (Fig. 5.9b), although a slight improvement in the survival of rad52 deletion strain on GAL plates upon hht2-hhf2 deletion was also observed and probably reflects the minor effect of hht2-hhf2Δ on Rad52 independent repair pathways (Coic et al., 2008). To further rule out any significant contribution of NHEJ, we also used a

129 plasmid recircularization assay to study NHEJ as described previously (Bird, 2002). Consistent with the NHEJ mediated repair of the DSB at the MAT locus in the absence of HMRa and HMLα sequences (Fig. 5.10 a, b), no significant differences in NHEJ repair efficiency were observed in strains with altered histone gene dosage using the plasmid recircularization assay (Fig. 5.10 c). Together, these results clearly show that NHEJ efficiency is not affected by a reduction in histone gene dosage and that the improved survival of a GAL-HO mediated DSB in strains carrying reduced histone dosage is primarily dependent upon HR mediated repair.

5.3.6 Reduction in histone gene dosage enhances spontaneous recombination rates The recombination mediated repair of a HO mediated DSB at the MAT locus is a highly specialized, site-specific and natural event in the budding yeast life cycle and the results obtained in Fig. 5.8 a, b may not be generally applicable to all HR mediated events occurring in yeast cells (Haber, 1998). To ascertain whether the observed effect of hht2-hhf2 deletion on the repair of a DSB at the MAT locus was applicable to HR at other sites in the yeast genome, we next investigated the effect of hht2-hhf2 deletion on HR between direct repeats using the previously described LU system (Fig. 5.11 a) (Prado and Aguilera, 1995). Using the LU system integrated at the HIS3 locus we found that spontaneous HR between direct repeats is significantly elevated in the hht2-hhf2 deletion strain (Fig. 5.11 b) and nearly identical results were obtained with the LU system maintained on a plasmid or a linear mini-chromosome. Hence, we conclude that a reduction in histone gene dosage leads to an increase in the efficiency of HR throughout the genome and may explain the lower sensitivity of hht2-hhf2 deletion strains to DNA damaging agents.

130

Fig. 5.10 Reduction in histone gene dosage affects the efficiency of DSB repair via the Homologous recombination (HR) but not the Non- Homologous End Joining (NHEJ) pathway. (a) Survival of strains lacking the HMRa and HMLα donor sequences upon GAL- HO mediated DSB induction. Ten-fold serial dilutions of the indicated GAL-HO strains carrying deletions of hml and hmr sequences were plated to determine their relative viability on glucose and galactose media. The plates were incubated for 4 days at 30°C prior to being photographed. (b) Quantitation of the survival of strains lacking the HMRa and HMLα donor sequences following a DSB created by HO endonuclease. Quantitative representation of the result shown in (a) exactly as described for Fig. 5.8 b. (c) Plasmid recircularization assay for NHEJ. Equal amounts of HindIII linearized or uncut supercoiled plasmids carrying a KanR marker for G418 resistance were transformed into the same number of yeast cells from each strain and plated on media containing the selection antibiotic G418. Viable colonies were counted and normalized to wild type cells transformed with uncut supercoiled plasmid. The error bars represent standard error of the mean from three independent repeats of the experiments under identical conditions.

131 5.3.7 Histones compete with homologous recombination factors for binding to damaged DNA

Both HR and NHEJ pathways require that a number of repair factors have direct access to the DNA at the site of the lesion (Aylon and Kupiec, 2004). Since the major DNA binding proteins in eukaryotic cells are histones, accessibility to the DSB would be nearly guaranteed if the region around the lesion was largely histone free during DNA repair (Linger and Tyler, 2007). This is indeed the case during repair of the HO mediated DSB at the MAT locus from which histones are displaced presumably due to the actions of chromatin remodeling factors (Tsukuda et al., 2005; Tsukuda et al., 2009a). However, if excess histones are present in the vicinity of the DNA lesion, they can potentially compete with repair proteins for binding to DNA repair sites, thereby inhibiting or slowing down the repair process. The converse of this scenario is also possible such that if histone levels are low, the repair factors will not have to compete with the histones for binding to the DNA at the site of the lesion and thus carry out the repair more efficiently. To test if this is true in vivo, we introduced Tandem Affinity Purification (TAP) tagged HR (Rad51, Rad52 and RPA1) and NHEJ factors (Ku80) in the wild type or hht2-hhf2Δ strains carrying the GAL-HO construct. The association of the TAP tagged proteins as well as histone H4 with the DNA in the vicinity of the DSB at the MAT locus following cleavage by the HO endonuclease was assayed using Chromatin Immunoprecipitation (ChIP) as described previously (Kuo and Allis, 1999). The DNA coimmunoprecipitated by the HR, NHEJ or histone H4 was quantitated by quantitative real-time Polymerase Chain Reaction (PCR) (Wong and Medrano, 2005) using fluorescently labeled primers to specifically amplify sequences adjacent to the HO cutting site in the MATa locus. ChIP experiments that have been carried out on the MAT locus in the context of DSB repair so far have largely utilized strains lacking the HMRa and HMLα donor sequences that are incapable of repairing the HO break via HR, which is likely to result in the accumulation of long stretches of single stranded DNA in the continuous presence of galactose (Downs et al., 2004; Morrison et al., 2004;

132 Shim et al., 2007; Shroff et al., 2004; Tsukuda et al., 2005; van Attikum et al., 2007; van Attikum et al., 2004). Our experiments have been carried out using strains that are more physiologically relevant for the study of HR as they have intact HMRa and HMLα sequences that allow high efficiency repair of the HO break via HR. Using these strains, we consistently observed a greater enrichment of the HR factors Rad51 (Fig. 5.12a), Rad52 (Fig. 5.12 b) and RPA (Figure 5.13) at the HO mediated DSB in the MAT locus, particularly at the later time points. This result is consistent with the idea that a reduced histone dosage facilitates HR by increasing the probability of the association of the HR repair factors with DNA in the vicinity of the DSB. In contrast, no significant differences were observed in the recruitment of the NHEJ factor Ku80 between wild type and hht2-hhf2Δ strains (Fig. 5.12 c), consistent with our finding that NHEJ is not affected by a reduction in histone dosage. Moreover, we also consistently observed a greater loss of histone H4 from the vicinity of the DSB in the hht2- hhf2 deletion strain compared to the wild type cells (Fig. 5.12 d). Together, these results strongly suggest that there is competition between DNA repair factors and histones for binding to damaged DNA in wild type cells and a reduction in histone gene dosage minimizes this competition, thereby facilitating DNA repair.

133

Fig. 5.11 Reduction in histone gene dosage results in elevated rates of spontaneous homologous recombination.

(a) Scheme for the LU direct repeat recombination system. In the LU system (Prado and Aguilera, 1995), two 600bp nonfunctional internal fragments of the LEU2 gene are interrupted by a 2.5kb fragment. Successful homologous recombination will excise the 2.5kb fragment and generate a functional LEU2 gene, allowing recombinants to be scored as Leu+ colonies on synthetic media lacking leucine. The LU recombination system can be integrated in different locations in the yeast genome, or placed on a centromeric plasmid and as such it can be used to obtain a clear picture of a general effect on HR. (b) Reduction in histone gene dosage elevates HR rates between direct repeats. A LU system was inserted into the HIS3 locus on chromosome 15 in the indicated strains. Wild type W303, hht1- hhf1Δ and hht2-hhf2Δ strains carrying the LU direct repeat system were plated on YPD or synthetic media lacking leucine (-LEU). The frequencies of Leu+ recombinants in the different strains were calculated as described previously (Prado and Aguilera, 1995) to calculate the recombination frequencies. The error bars represent standard error of the mean from three independent repeats of the experiments under identical conditions.

134

Fig. 5.12 Histones compete with HR factors for binding to DNA repair sites. (a) Recruitment of Rad51-TAP to a HO mediated DSB at the MAT locus. Galactose was added to induce a HO mediated DSB at the MATa locus in wild type or hht2-hhf2Δ yeast strains carrying TAP-tagged Rad51. Cells were harvested every 30 minutes following the addition of galactose and subjected to ChIP analysis as described in the Methods section. Real-time PCR signal obtained from the MATa locus was normalized to the background signal obtained from the ACT1 locus, which serves as an internal control to ensure that equal amounts of DNA was used in each reaction. All data are shown relative to the Ct (cycle threshold) of the 0’ time point which was given an arbitrary value of 1. The error bars represent standard error of the mean from three independent repeats of the experiments under identical conditions. (b) Recruitment of Rad52-TAP to a HO mediated DSB at the MAT locus. The recruitment of Rad52-TAP was studied as described above in a. (c) Recruitment of Ku80-TAP to a HO mediated DSB at the MAT locus. The recruitment of Ku80-TAP was studied as described above in a. (d) Loss of histone H4 from a HO mediated DSB at the MAT locus. The loss of histone H4 was studied as described above in a.

135

Fig. 5.13 Recruitment of RPA1-TAP to a HO mediated DSB at the MAT locus. Recruitment of RPA1-TAP was studied as described in legend for Fig. 5.10

136 5.4 Discussion

In this study we have shown that the histone gene dosage has a dramatic influence on the DNA damage sensitivity of budding yeast cells (Fig. 5.1). Our results suggest that even the normal histone gene dosage in wild type yeast cells appears to generate an excess of histones, which in turn appears to interfere with efficient DNA repair by competing with the repair factors for binding to the DNA lesion (Fig. 5.8, 5.11, 5.12). Further, deletion of the hht2-hhf2 gene pair that contributes most of the histone H3 and H4 mRNA in the cell results in a significant reduction in the DNA damage sensitivity of wild type as well as DNA damage checkpoint mutant yeast strains (Fig. 5.1, 5.8). The effect of a reduced histone gene dosage on the DNA damage sensitivity of yeast cells is primarily mediated by an increase in the efficiency of DNA repair by the HR pathway (Fig. 5.8, 5.10, 5.11, 5.12), while the NHEJ pathway is unaffected (Fig. 5.10). This is not surprising as repair via the HR pathway involves intimate DNA and protein transactions over long stretches of chromatin, while repair via the NHEJ pathway essentially involves the religation of the broken ends with limited amount of end- processing (Aylon and Kupiec, 2004)( Fig. 5.14). In fact, the resection of the broken ends is the first step in HR mediated DSB repair (Huertas, 2010) and involves the generation of single-stranded DNA with concomitant loss of histones (Tsukuda et al., 2005). Additionally, the donor loci also exhibit substantial chromatin remodeling that facilitates repair of the DSB at the MAT locus via HR (Tsukuda et al., 2009a). Since histones can interact efficiently with single stranded DNA and may even form nucleosome like structures (Caffarelli et al., 1983; Palter and Alberts, 1979; Palter et al., 1979), the presence of appreciable quantities of free histones in the vicinity of such HR repair intermediates is likely to present a significant competition for the HR factors for binding to the exposed DNA (Fig. 5.14). Further, DSBs in haploid yeast are repaired via HR primarily during the S and G2 phases of the cell cycle when homologous sequences are readily available for HR mediated repair. Since the yeast histone genes are predominantly expressed during early S-phase (Osley, 1991), histones are also

137 likely to be present at their highest concentrations in the cell during S and G2 phases of the cell cycle and thus have their maximal effect on HR mediated repair, rather than NHEJ.

Our results raise the important question of why eukaryotes have evolved with multiple copies of histone genes when they are clearly not required for viability (Castillo et al., 2007; Osley, 1991; Takami and Nakayama, 1997a) and also appear to make a huge excess of histone proteins over what is required for chromatin assembly (Ghaemmaghami et al., 2003; Singh et al., 2009b), potentially leading to deleterious consequences (Singh et al., 2010). Assuming a nucleosome every ~160bp and 100% recycling of the pre-existing histones, about ~150,000 newly synthesized molecules of each core histone will be needed to package the newly replicated haploid budding yeast genome of 12.5 x 106 bp during each S-phase. However, the two copies of histone H4 genes 6 (HHF1 and HHF2) together have been estimated to produce ~1.2 x 10 histone H4 molecules per cell (Ghaemmaghami et al., 2003), which would correspond to a ~8-fold excess over what is needed for chromatin assembly during each S- phase. Even the HHF1 gene with the lower transcriptional output is estimated to produce ~524,000 molecules of histone H4, which would correspond to a ~3-fold excess. Hence, it is not surprising that either copy of the two genes encoding histones H3 and H4 in the budding yeast is sufficient for viability and the gross chromatin structure is unaffected upon deletion of any one of the two gene copies. Even if the data regarding the number of molecules of individual proteins in these large scale studies (Ghaemmaghami et al., 2003) is off by 2-3 folds, it is very likely that all the four core histones in the budding yeast are produced in quantities much higher than their requirement for chromatin assembly during S- phase. So, despite the tight transcriptional and posttranscriptional control of histone genes (Osley, 1991), why do yeast cells make such a huge excess of histone proteins? It is known that delays between DNA replication and chromatin assembly during S-phase results in spontaneous DNA damage, genomic instability and inviability (Clemente-Ruiz and Prado, 2009; Han et al., 1987;

138 Myung et al., 2003; Prado and Aguilera, 2005; Prado et al., 2004; Ye et al., 2003a). Hence, to avoid the deleterious consequences of delayed or insufficient histone deposition during DNA replication, one possibility is that high levels of histones are synthesized during S-phase to ensure a sufficiently high histone concentration for rapid chromatin assembly by a relatively limited number of histone chaperone molecules. For the assembly of 300,000 new histone H3 and H4 molecules, the estimated number of all the known histone H3 and H4 chaperone molecules available are as follows: Asf1 ~6230, Cac1 ~ 1590, Hir1 ~ 846 (Note: histone chaperones that work as multi-subunit complexes have similar or lower number of molecules per cell for their other subunits than shown here) (Ghaemmaghami et al., 2003). So, essentially less than 10,000 histone chaperone molecules have the challenging task of assembling 300,000 histone H3 and H4 molecules during the budding yeast S-phase which typically lasts around 20 minutes at 30°C. It is possible that this task can only be achieved at high effective histone protein concentrations that would allow the histone chaperones to have a quick turn-around time after each round of histone deposition, such that they do not have to spend too much time looking for the next histone molecule to deposit.

Mutant yeast strains such as those lacking the sgs1, srs2 or rrm3 helicases exhibit greatly elevated rates of mitotic recombination that is highly deleterious for the cells as it results in genomic instability (Kerrest et al., 2009; Schmidt and Kolodner, 2006; Yamagata et al., 1998). Based on our current results, another role for the high levels of histones produced by multiple histone genes may be to keep the HR machinery in check and prevent it from initiating spurious recombination events using DNA replication intermediates as templates, thereby buffering HR activity in cells during the S and G2 phases of the cell cycle and contributing to the overall genomic stability of the cells. This idea would also be consistent with the finding that partial depletion of histone H4 results in genomic instability due to elevated recombination (Prado and Aguilera, 2005). More

139 importantly, combined with the exciting recent finding that aging yeast cells have reduced levels of histones (Feser et al., 2010), our results could help explain in part the hyper-recombinational state of aging yeast cells (McMurray and Gottschling, 2003). Not surprisingly, histone overexpression increases the life- span of yeast cells (Feser et al., 2010) and our findings strongly suggest that this is likely to be in part due to the dampening of HR activity, thereby limiting genomic instability which is likely to aggravate the aging process in these cells.

Irrespective of what the underlying reasons for the presence of multiple histone genes in all eukaryotes prove to be eventually, due to the high degree of conservation of DNA repair and chromatin processes, the presence of multiple histone genes is likely to have major ramifications for these processes in all eukaryotes, including humans. For example, many current anti-cancer drugs rely upon their ability to cause DNA damage for their effectiveness and based on our studies in yeast, it is possible that the loss of histone genes from genetically unstable cancer cells may make it relatively easy for them to develop resistance to such drugs. Hence, our results could have serious implications for genomic stability of human cells with their dozens of histone genes (Marzluff et al., 2002) that can significantly influence numerous aspects of human health and disease. Future studies will reveal if histone gene copies are indeed lost from histone gene clusters in human cancer cells that are refractory to DNA damaging chemotherapeutic drugs, and if the mechanism of resistance is dependent upon elevated HR, then the HR machinery could be targeted in an attempt to destroy such refractory cancers.

140

Fig. 5.14 Model to illustrate the potential effect of excess histones on HR or NHEJ mediated repair of a DSB. (a) Effect of excess histones on HR mediated DSB repair. Due to the presence of long stretches of nucleosome deficient single stranded DNA as intermediates at both the recipient and donor loci during HR mediated DSB repair, the availability of significant amounts of histones even in wild type cells results in competition between the HR factors and histones for binding to the exposed DNA. This can lead to a reduction in the efficiency of HR mediated repair. For simplicity, only a few HR factors and one nuclease complex (MRX) are indicated. (b) Effect of excess histones on NHEJ mediated DSB repair. Significant stretches of DNA are not exposed at the DSB site during NHEJ mediated repair and as such this repair process is unlikely to be affected by the presence of significant quantities of histones. NHEJ factors are indicated.

141 5.5 Materials and Methods

Yeast strains and plasmids The yeast strains used are listed in Table 5.1. Standard techniques for growing and transforming yeast were used(Burke et al., 2000). Plasmids used for histone overexpression have been described in detail elsewhere (Gunjan and Verreault, 2003; Singh et al., 2009a). For the galactose induced expression of HO endonuclease, cells were routinely grown in minimal media supplemented with “yeast synthetic drop-out medium supplement” (Sigma) containing 2% raffinose prior to the addition of 2% galactose. Galactose inducible genes were repressed by addition of 2% glucose. α-factor was used at a concentration of 5μg/ml for 2–3 hours to arrest cells in G1. For longer experiments with G1-arrested cells, more α-factor was added at regular intervals. The hht1-hhf1Δ and hht2-hhf2Δ gene pairs were deleted from various strains using a PCR strategy described previously (Longtine et al., 1998).

Plasmid recircularization assay for NHEJ efficiency The plasmid recircularization assay to study NHEJ was carried out as described previously (Bird et al., 2002). In this assay, a plasmid containing Kanr marker is first digested using the restriction endonuclease HindIII in the middle of the marker to mimic a DSB. Then 0.2μg of the digested (linearized) or undigested (supercoiled) plasmid was transformed into 3x107 cells from each strain and the transformants were selected on G418 plates. The Kanr marker encodes resistance to the antibiotic G418 and lacks any significant homology within the budding yeast genome, so the Hind III generated DSB can only be repaired via NHEJ. Only those cells that have successfully repaired the linearized plasmid and generated a functional Kanr gene will survive on media containing G418. NHEJ repair efficiency was calculated by comparing the number of colonies formed by strains transformed with the digested plasmid versus undigested plasmid.

142 Chromatin Immunoprecipitation (ChIP) Cells were grown overnight to reach a concentration 1~2x107 cell/ml in YP- Raffinose media. 2% galactose was added to the culture at the 0’ time point and then 10ml aliquots of the cultures were collected every 30 minutes and fixed with 1% formaldehyde for 15 minutes. Fixed cells were disrupted using glass beads in lysis buffer and cell lysate was sonicated on ice to shear the chromatin into 300~1000bp fragments. Equal amounts of whole cell extract were incubated at 4℃ for 1.5 hrs with 20’ of IgG-coupled sepharose beads (GE healthcare) to immunoprecipitate TAP tagged proteins, or histone H4 antibody bound protein-A beads to pull down histone H4. Then protein-DNA complexes were eluted and protein-DNA crosslinks were reversed at 65℃ overnight. The immunoprecipitated DNA was purified by phenol-chloroform extraction and then was quantitated by real-time PCR (Applied Biosystems) using primers and fluorescently labeled probes (TaqMan) to specifically amplify sequences near the HO cleavage site (70bp downstream of HO site) at the MATa locus. The ACT1 locus was also assayed to serve as internal control. Real-time PCR reactions were performed in triplicate for each DNA sample and the data from three independent experiments were averaged.

143 Table 5.1 List of strains used in the study described in chapter 4.

Strain Genotype Reference MATa ho ade2-1 trp1-1 can1-100 leu2-3,112 his3- (Thomas and Rothstein, W303-1A 11,15 ura3-1 1989) CY2034 W303 MATa rad53K227A::KanMX4 (Pellicioli et al., 1999) U960-5C W303 MATa rad53::HIS3 sml1-1 RAD5+ (Zhao et al., 1998) U953- W303 MATa mec1::TRP1 sml1::HIS3 (Zhao et al., 1998) 61A W303 MATa rad53::HIS3 sml1::TRP1 hht2- (Gunjan and Verreault, YAG713 hhf2::KanMx 2003) W303 MATa mec1::TRP1 sml1::HIS3 hht2- (Gunjan and Verreault, YAG722 hhf2::KanMx 2003) YAG116 W303 MATa hht1-hhf1::KanMx This study YAG107 W303 MATa hht2-hhf2::TRP1 " YLD1 W303 MATa ade3::GAL-HO-URA3 " W303 MATa hht1-hhf1::KanMx ade3::GAL-HO- YLD10 " URA3

YLD9 W303 MATa hht2-hhf2::TRP1 ade3::GAL-HO-URA3 "

YLD87 W303 MATa his3::pRS314-LU-HIS3 "

YLD99 W303 MATa hht1-hhf1::Kan his3::pRS314-LU-HIS3 "

W303 MATa rad53::HIS3 sml1-1 ade3::GALHO- YAG717 " URA3-ADE3 W303 MATa rad53::HIS3 sml1-1 hht2- YAG810 " hhf2::Kan ade3::GAL-HO-URA3-ADE3 W303 MATa rad53::HIS3 sml1-1 hht1- YAG844 " hhf1::Kan ade3::GAL-HO-URA3-ADE3 W303 MATa hht2-hhf2::TRP1 his3::pRS314-LU- YLD111 " HIS3 YAG883 W303 MATa hmr::TRP1 hml::(URA3)5-FOA " W303 MATa hmr::TRP1 hml::(URA3)5-FOA YAG938 " ade3::GAL-HO-URA3 W303 MATa hht1-hhf1::Kan hmr::TRP1 YAG959 " hml::(URA3)5-FOA ade3::GAL-HO-URA3 W303 MATa hht2-hhf2::Kan hmr::TRP1 YAG960 " hml::(URA3)5-FOA ade3::GAL-HO-URA3 BY4741 MATa RAD51-TAP-HIS3 ade3::GAL-HO- YLD34 " URA3 BY4741 MATa RAD51-TAP-HIS3 ade3::GAL-HO- YLD145 " URA3 hht2-hhf2::Kan

144 Table 5.1 continued… BY4741 MATa RAD52-TAP-HIS3 ade3::GAL-HO- YLD35 " URA3 BY4741 MATa RAD52-TAP-HIS3 ade3::GAL-HO- YLD156 " URA3 hht2-hhf2::Kan BY4741 MATa RPA1-TAP-HIS3 ade3::GAL-HO- YLD37 " URA3 BY4741 MATa RPA1-TAP-HIS3 ade3::GAL-HO- YLD152 " URA3 hht2-hhf2::Kan BY4741 MATa KU80-TAP-HIS3 ade3::GAL-HO- YLD36 " URA3 BY4741 MATa KU80-TAP-HIS3 ade3::GAL-HO- YLD150 " URA3 hht2-hhf2::Kan

145 CHAPTER 6

CONCLUSIONS

Histones are essential proteins since they not only package DNA into the relatively small nucleus, but also regulate DNA accessibility and chromatin stability. However, since histones are positively charged, they can bind non- specifically to negatively charged DNA and affect DNA metabolism such as transcription, DNA replication, damage repair and recombination. Genomic instability is a hallmark of cancer cells. To ensure the high fidelity transmission of genetic information, cells have evolved multiple mechanisms for maintaining genomic integrity. Current understanding is that epigenetic regulation plays indispensable roles in DNA damage repair and the coordination of repair and epigenetic processes is likely to be crucial for cell viability (Groth et al., 2007). Previous work done by our group have promoted the novel concept that excess histones influence DNA metabolism, especially DNA damage repair (Gunjan and Verreault, 2003; Singh et al., 2009a). Therefore, understanding the mechanism of histone dosage in regulating DNA damage repair will enable us to better define the role of epigenetic regulation in maintaining genomic stability.

146 presence of excess histones may saturate the remaining histone modifying enzymes, either in their chromatin associated or free forms, potentially interfering with their enzyme activities. Subsequently, we observed that histone overexpression does not affect the bulk chromatin structure as assayed by nucleosomal ladder generated by MNase digestion. However, indirect end- labeling analysis using primer extension at specific loci following MNase or DNaseI cleavage indicated subtle alterations in the fine structure of chromatin in response to histone overexpression. To further investigate whether chromatin structure alteration would have any effect on transcription, a genome wide microarray analysis was performed. Microarray analysis revealed that upon the simultaneous overexpression of the histone H3 and H4 gene pair or all the four core histones, about 240 genes were either up or downregulated by 2-fold or more. Surprisingly, although overexpression of histone H3 alone results in significant toxicity and in viability(Gunjan and Verreault, 2003) and alters the chromatin fine structure, it does not have any effect on transcription. This is probably due to the deposition of histone dimers during chromatin assembly/disassembly (Tagami et al., 2004), such that a single histone may not serve as substrate in this process, or a significant impediment to the progress of RNA polymerases. Due to their positive charge, histones are likely to have high affinity for negatively charged molecules, and as such it is possible that excess histones can also bind to RNA. Indeed, we observed this in vivo by immunoprecipitating overexpressed histone H3 along with several mRNAs, suggesting that excess histones are capable of binding to RNAs and potentially altering their activities. Overall, we present evidence that excess histones are likely to mediate their cytotoxic effects via multiple mechanisms that are primarily dependent on inappropriate electrostatic interactions between the positively charged histones and diverse negatively charged molecules in the cell.

Furthermore, we have also investigated how changes in histone gene dosage affect the DNA damage sensitivity of budding yeast cells. Budding yeast have two copies of each histone gene when only one copy is needed for survival. In

147 this study, the second copy of H3-H4 gene (HHT2-HHF2), which contributes 6-8 fold more histone mRNAs than the first gene copy (HHT1-HHF1) was deleted, which provides us with a robust system to reduce the endogenous levels of histones without causing the collapse of the chromatin structure. Here, we report that overexpression of histones leads to an increase in the DNA damage sensitivity in response to variety of damaging agents. On the other hand, the reduction of histone H3 and H4 gene dosage resulted in a significant decrease in the DNA damage sensitivity. By taking advantage of the HO endonuclease mediated DNA double stand break (DSB) at the MAT locus, we were able to study the process of DSB repair in detail. The efficiency of Homologous Recombination (HR) at a DSB, as well as genome wide HR, was elevated in hht2-hhf2 deletion strain; whereas Non-Homologous End Joining (NHEJ) remained unchanged. These effects were not associated with global changes in the expression of DNA repair genes or DNA damage checkpoint responses. We also found that there was no alteration of gross chromatin structure in response to changes in histone gene dosage. The most likely mechanism by which reduced histone dosage leads to elevated HR is through enhanced recruitment of the HR factors, as determined by the Chromatin Immunoprecipitation (ChIP) assay. Concomitant with this, cells also experience greater histone loss around this DSB upon a reduction in histone gene dosage. We propose that high levels of endogenous histones generated by multiple genes compete with HR factors for binding to DNA, thereby reducing HR efficiency and may normally function to restrain excessive HR activities during S-phase.

HR and NHEJ are the two major pathways for the repair of DSBs. The choice of either of these pathways for eukaryotic DSB repair is a topic of great interest for mechanistic studies. Apart from the determination factors mentioned earlier, epigenetic factors that affect pathway choice are emerging recently. There is evidence that certain histone modifications might serve as indicators for the engagement of the two pathways (Xie et al., 2007). Moreover, histone hyperacetylation associated chromatin remodelling is believed to direct DSB

148 towards HR rather then NHEJ (Barlev et al., 1998; Tamburini and Tyler, 2005). In contrast, the inhibition of HAT activity might favor NHEJ over HR (Costelloe et al., 2006). In our work, we demonstrated the effect of a reduced histone gene dosage on DSB repair is primarily mediated by elevated efficiency of HR pathway, while the NHEJ pathway is unaffected. Taking existing literatures into account, our finding is not surprising as repair via HR involves intimate DNA and protein transactions over long stretches of chromatin, while repair via NHEJ takes place in a limited spatial scale for the two ends to be religated without extensive processing. The processing of DNA broken ends, which is a decisive point for cells to perform either HR or NHEJ (Huertas, 2010), is coincidentally involved in the disruption of nucleosomes leading to histone loss. We found that the extent of histone loss around the lesion might be the key reason that accounts for the elevated repair efficiency. The presence of appreciable quantities of free histone in the vicinity HR repair site is likely to present a significant competition for the HR factors for binding to the exposed DNA. Moreover, the cell cycle timing of histone expression overlaps with the existence of more readily available homologous sequences, as they all take place during S phase. Thus, the effect of histone dosage is likely more prominent in HR repair pathways than NHEJ.

Many interesting questions are still left for future investigation. What is the exact mechanism which results in enhanced repair factors recruitment and enhanced histone loss in cells with reduced histone dosage? How do free histones interact with the DNA around the lesion? What role do histone chaperones and chromatin remodeling complexes play in this process? How to directly and accurately assay for the pool of excess histones in a cell, which is likely to be less than 1% of total chromatin bound histones in the cell? Can we observe similar results in other types of repair other than DSBs, such as BER or NER? What is the role of histone gene dosage in human cells, where histone genes are present in dozens of copies? What is the significance of histone dosage in aging, either replicative or chronological? Are there differences in histone levels in cancer cells versus normal cell or stem cells versus differentiated cells?

149

In the past decade, epigenetics has been one of most active research areas aimed at obtaining a better understanding of the causes and potential treatments for cancer as well as other aging related diseases. Conventional epigenetic research stresses on DNA/histone modifications, chromatin remodeling and histone variants. Not until recently, the concept of excess histones and its role in regulating DNA metabolism were elucidated. That is the reason why we are trying to address this issue in vivo. Our findings may shed some light on the intrinsic relationship between histone dosage and DNA damage repair, which could provide a new layer of regulation for the DNA damage response.

Through further research, our findings could extend our understanding of cancer and possibly uncover better treatments via two options. On one hand, reducing histone dosage may be beneficial for normal cells in repairing DNA damage resulting from exogenous and endogenous damaging agents, thereby maintaining genomic stability. Such enhanced repair may also be adopted by adult stem cells to ensure better survival. On the other hand, transient upregulation of histones can be applied in combination with chemotherapy for cancer cells, particularly in cells that exhibit drug resistance to a variety of chemotherapy agents. The deleterious effects of inhibition of DNA repair combined with the cytotoxicity of the excess histones are likely to be effective against replicating cancer cells than quiescent normal cells.

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179 APPENDIX A

Table A1. List of genes affected 2-fold or more by the simultaneous overexpression of the histone H3-H4 gene pair.

GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YOL154W 5.590 down Zps1p Putative GPI-anchored protein; transcription is induced under low-zinc conditions, as mediated by the Zap1p t YGR260W 3.986 down Tna1p High affinity nicotinic acid plasma membrane permease, responsible for uptake of low levels of nicotinic acid; YHL047C 3.401 down Arn2p Transporter, member of the ARN family of transporters that specifically recognize siderophore-iron chelates; r YOR383C 3.140 down Fit3p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved in YOR382W 3.134 down Fit2p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved in YNR056C 3.087 down Bio5p Putative transmembrane protein involved in the biotin biosynthesis pathway; responsible for uptake of 7-keto 8 YJR025C 3.016 down Bna1p 3-hydroxyanthranilic acid dioxygenase, required for biosynthesis of nicotinic acid from tryptophan via kynuren YJR078W 2.911 down Bna2p Tryptophan 2,3-dioxygenase, required for biosynthesis of nicotinic acid from tryptophan via kynurenine pathwa YIL057C 2.866 down Yil057cp Putative protein of unknown function; expression induced under carbon limitation and repressed under high g YEL021W 2.812 down Ura3p Orotidine-5'-phosphate (OMP) decarboxylase, catalyzes the sixth enzymatic step in the de novo biosynthesis YGR174W-A 2.776 down Ygr174w-ap Putative protein of unknown function YGL121C 2.686 down Gpg1p Proposed gamma subunit of the heterotrimeric G protein that interacts with the receptor Grp1p; involved in reg YLR136C 2.630 down Tis11p mRNA-binding protein expressed during iron starvation; binds to a sequence element in the 3'-untranslated re YDR114C 2.570 down Ydr114cp Putative protein of unknown function; deletion mutant exhibits poor growth at elevated pH and calcium YER045C 2.528 down Aca1p Basic leucine zipper (bZIP) transcription factor of the ATF/CREB family, may regulate transcription of genes in YNR058W 2.508 down Bio3p 7,8-diamino-pelargonic acid aminotransferase (DAPA), catalyzes the second step in the biotin biosynthesis pa YBL108C-A 2.416 down Pau9p hypothetical protein identified by homology. See FEBS Letters [2000] 487:31-36. YGL168W 2.395 down Hur1p Protein required for hydroxyurea resistance; has possible roles in DNA replication and maintenance of proper YGL230C 2.385 down Ygl230cp Putative protein of unknown function; non-essential gene YIL176C 2.372 down Pau14p YDR542W 2.371 down Pau10p YNR034W-A 2.363 down Ynr034w-ap Putative protein of unknown function; expression is regulated by Msn2p/Msn4p YCL001W-A 2.347 down Ycl001w-ap Putative protein of unknown function; YCL001W-A is not an essential gene YGR294W 2.332 down Pau12p YGL256W 2.329 down Adh4p Alcohol dehydrogenase isoenzyme type IV, dimeric enzyme demonstrated to be zinc-dependent despite sequ YKL065W-A 2.328 down Ykl065w-ap Putative protein of unknown function YAL068C 2.321 down Pau8p YFR047C 2.314 down Bna6p Quinolinate phosphoribosyl transferase, required for biosynthesis of nicotinic acid from tryptophan via kynuren YLR456W 2.303 down Ylr456wp Putative protein of unknown function YHL046C 2.296 down Pau13p Putative protein of unknown function; not an essential gene YDL007C-A 2.284 down Ydl007c-ap Putative protein of unknown function YNL195C 2.277 down Ynl195cp Putative protein of unknown function; shares a promoter with YNL194C; the authentic, non-tagged protein is d YFR012W-A 2.273 down Yfr012w-ap Putative protein of unknown function; identified by homology YHR053C 2.268 down Cup1-1p Metallothionein, binds copper and mediates resistance to high concentrations of copper and cadmium; locus i YGR288W 2.264 down Mal13p MAL-activator protein, part of complex locus MAL1; nonfunctional in genomic reference strain S288C YDL210W 2.229 down Uga4p Permease that serves as a gamma-aminobutyrate (GABA) transport protein involved in the utilization of GABA YHR185C 2.222 down Pfs1p Sporulation protein required for prospore membrane formation at selected spindle poles, ensures functionality YKR106W 2.222 down Ykr106wp Protein of unconfirmed function; displays a topology characteristic of the Major Facilitators Superfamily of mem YNL254C 2.222 down Ynl254cp

180 Table A.1 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YMR322C 2.212 down Sno4p Possible chaperone and cysteine protease with similarity to E. coli Hsp31 and S. cerevisiae Hsp31p, Hsp32p, YCR105W 2.197 down Adh7p NADPH-dependent medium chain alcohol dehydrogenase with broad substrate specificity; member of the cinn YJL037W 2.192 down Irc18p Putative protein of unknown function; expression induced in respiratory-deficient cells and in carbon-limited ch YPR124W 2.179 down Ctr1p High-affinity copper transporter of the plasma membrane, mediates nearly all copper uptake under low copper YMR244C-A 2.177 down Ymr244c-ap Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to both the cyto YCR090C 2.166 down Ycr090cp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplasm YAL018C 2.162 down Yal018cp Putative protein of unknown function YLR461W 2.157 down Pau4p Part of 23-member seripauperin multigene family encoded mainly in subtelomeric regions, active during alcoh YIL177C 2.146 down Yil177cp Putative protein of unknown function; similarity to DNA helicases that are encoded within subtelomeric Y' elem YLL064C 2.139 down Pau18p YBR302C 2.138 down Cos2p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encoded YLR307C-A 2.134 down Ylr307c-ap Putative protein of unknown function YJR011C 2.128 down Yjr011cp Putative protein of unknown function; GFP-fusion protein expression is induced in response to the DNA-dama YOR391C 2.127 down Hsp33p Possible chaperone and cysteine protease with similarity to E. coli Hsp31 and S. cerevisiae Hsp31p, Hsp32p, YBR200W-A 2.118 down Ybr200w-ap Putative protein of unknown function; identified by fungal homology and RT-PCR YGL258W-A 2.117 down Ygl258w-ap Putative protein of unknown function YIL156W-B 2.116 down Yil156w-bp Putative protein of unknown function, originally identified based on homology to Ashbya gossypii and other re YNL336W 2.116 down Cos1p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encoded YLR070C 2.110 down Xyl2p Xylitol dehydrogenase, converts xylitol to D-xylulose in the endogenous xylose utilization pathway YNR075W 2.108 down Cos10p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encoded YNR057C 2.100 down Bio4p Dethiobiotin synthetase, catalyzes the third step in the biotin biosynthesis pathway; BIO4 is in a cluster of 3 ge YHR014W 2.094 down Spo13p Meiosis-specific protein, involved in maintaining sister chromatid cohesion during meiosis I as well as promoti YBL113C 2.091 down Ybl113cp Helicase-like protein encoded within the telomeric Y' element YHR214C-D 2.088 down Yhr214c-dp Putative protein of unknown function; identified by gene-trapping, microarray-based expression analysis, and YMR175W-A 2.084 down Ymr175w-ap Putative protein of unknown function YBR301W 2.083 down Dan3p Cell wall mannoprotein with similarity to Tir1p, Tir2p, Tir3p, and Tir4p; expressed under anaerobic conditions, YFR032C-B 2.069 down Yfr032c-bp Putative protein of unknown function; identified by gene-trapping, microarray-based expression analysis, and YDL239C 2.066 down Ady3p Protein required for spore wall formation, thought to mediate assembly of a Don1p-containing structure at the YFL017W-A 2.055 down Smx2p Core Sm protein Sm G; part of heteroheptameric complex (with Smb1p, Smd1p, Smd2p, Smd3p, Sme1p, and YAR033W 2.053 down Mst28p Putative integral membrane protein, involved in vesicle formation; forms complex with Mst27p; member of DU YIL014C-A 2.053 down Yil014c-ap Putative protein of unknown function YGR008C 2.049 down Stf2p Protein involved in regulation of the mitochondrial F1F0-ATP synthase; Stf1p and Stf2p act as stabilizing facto YPL033C 2.046 down Ypl033cp Putative protein of unknown function; may be involved in DNA metabolism; expression is induced by Kar4p YDR381C-A 2.038 down Ydr381c-ap Protein of unknown function, localized to the mitochondrial outer membrane YGL260W 2.038 down Ygl260wp Putative protein of unknown function; transcription is significantly increased in a NAP1 deletion background; d YCL073C 2.036 down Ycl073cp Protein of unconfirmed function; displays a topology characteristic of the Major Facilitators Superfamily of mem YBR090C 2.035 down Ybr090cp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplasm YAL055W 2.028 down Pex22p Putative peroxisomal membrane protein required for import of peroxisomal proteins, functionally complements YLR231C 2.023 down Bna5p Kynureninase, required for biosynthesis of nicotinic acid from tryptophan via kynurenine pathway YJR161C 2.010 down Cos5p Protein of unknown function, member the DUP380 subfamily of conserved, often subtelomerically-encoded pr YER190C-B 2.007 down Yer190c-bp Putative protein of unknown function; identified by gene-trapping, microarray-based expression analysis, and YNL332W 2.003 down Thi12p Protein involved in synthesis of the thiamine precursor hydroxymethylpyrimidine (HMP); member of a subtelom YJL133C-A 2.003 up Yjl133c-ap Putative protein of unknown function; the authentic, non-tagged protein is detected in highly purified mitochon YJR122W 2.005 up Iba57p Mitochondrial matrix protein involved in the incorporation of iron-sulfur clusters into mitochondrial aconitase-ty YLR267W 2.009 up Bop2p Protein of unknown function, overproduction suppresses a pam1 slv3 double null mutation YEL055C 2.009 up Pol5p DNA Polymerase phi; has sequence similarity to the human MybBP1A and weak sequence similarity to B-type

181 Table A.1 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YKL096W 2.010 up Cwp1p Cell wall mannoprotein, linked to a beta-1,3- and beta-1,6-glucan heteropolymer through a phosphodiester bo YLR304C 2.011 up Aco1p Aconitase, required for the tricarboxylic acid (TCA) cycle and also independently required for mitochondrial ge YHR052W 2.014 up Cic1p Essential protein that interacts with proteasome components and has a potential role in proteasome substrate YHR137W 2.016 up Aro9p Aromatic aminotransferase II, catalyzes the first step of tryptophan, phenylalanine, and tyrosine catabolism YNL036W 2.019 up Nce103p Carbonic anhydrase; poorly transcribed under aerobic conditions and at an undetectable level under anaerob YDR527W 2.020 up Rba50p Protein involved in transcription; interacts with RNA polymerase II subunits Rpb2p, Rpb3, and Rpb11p; has si YNR038W 2.021 up Dbp6p Essential protein involved in ribosome biogenesis; putative ATP-dependent RNA helicase of the DEAD-box pr YDL101C 2.023 up Dun1p Cell-cycle checkpoint serine-threonine kinase required for DNA damage-induced transcription of certain targe YJL088W 2.026 up Arg3p Ornithine carbamoyltransferase (carbamoylphosphate:L-ornithine carbamoyltransferase), catalyzes the sixth s YHR197W 2.028 up Rix1p Essential component of the Rix1 complex (Rix1p, Ipi1p, Ipi3p) that is required for processing of ITS2 sequenc YPL146C 2.033 up Nop53p Nucleolar protein; involved in biogenesis of the 60S subunit of the ribosome; interacts with rRNA processing fa YPL012W 2.039 up Rrp12p Protein required for export of the ribosomal subunits; associates with the RNA components of the pre-ribosom YKR079C 2.039 up Trz1p tRNase Z, involved in RNA processing, has two putative nucleotide triphosphate-binding motifs (P-loop) and a YGL211W 2.042 up Ncs6p Protein required for thiolation of the uridine at the wobble position of Gln, Lys, and Glu tRNAs; has a role in ur YEL036C 2.044 up Anp1p Subunit of the alpha-1,6 mannosyltransferase complex; type II membrane protein; has a role in retention of gly YNL110C 2.045 up Nop15p Constituent of 66S pre-ribosomal particles, involved in 60S ribosomal subunit biogenesis; localizes to both nu YLR003C 2.051 up Ylr003cp Putative protein of unknown function that may participate in DNA replication; green fluorescent protein (GFP)- YAL012W 2.051 up Cys3p Cystathionine gamma-lyase, catalyzes one of the two reactions involved in the transsulfuration pathway that y YGR180C 2.060 up Rnr4p Ribonucleotide-diphosphate reductase (RNR), small subunit; the RNR complex catalyzes the rate-limiting step YOL058W 2.067 up Arg1p Arginosuccinate synthetase, catalyzes the formation of L-argininosuccinate from citrulline and L-aspartate in t YLR222C 2.074 up Utp13p Nucleolar protein, component of the small subunit (SSU) processome containing the U3 snoRNA that is involv YDR365C 2.075 up Esf1p Nucleolar protein involved in pre-rRNA processing; depletion causes severely decreased 18S rRNA levels YKL029C 2.083 up Mae1p Mitochondrial malic enzyme, catalyzes the oxidative decarboxylation of malate to pyruvate, which is a key inte YPR110C 2.089 up Rpc40p RNA polymerase subunit, common to RNA polymerase I and III YKL216W 2.090 up Ura1p Dihydroorotate dehydrogenase, catalyzes the fourth enzymatic step in the de novo biosynthesis of pyrimidines YER037W 2.093 up Phm8p Protein of unknown function, expression is induced by low phosphate levels and by inactivation of Pho85p YPL093W 2.097 up Nog1p Putative GTPase that associates with free 60S ribosomal subunits in the nucleolus and is required for 60S rib YJL116C 2.099 up Nca3p Protein that functions with Nca2p to regulate mitochondrial expression of subunits 6 (Atp6p) and 8 (Atp8p ) of YBL055C 2.108 up Ybl055cp 3'-->5' exonuclease and endonuclease with a possible role in apoptosis; has similarity to mammalian and C. e YOR297C 2.109 up Tim18p Component of the mitochondrial Tim54p-Tim22p complex involved in insertion of polytopic proteins into the in YLR197W 2.112 up Sik1p Essential evolutionarily-conserved nucleolar protein component of the box C/D snoRNP complexes that direct YDR449C 2.118 up Utp6p Nucleolar protein, component of the small subunit (SSU) processome containing the U3 snoRNA that is involv YML056C 2.125 up Imd4p Inosine monophosphate dehydrogenase, catalyzes the first step of GMP biosynthesis, member of a four-gene YHR065C 2.126 up Rrp3p Protein involved in rRNA processing; required for maturation of the 35S primary transcript of pre-rRNA and for YDR257C 2.133 up Set7p Nuclear protein that contains a SET-domain, which have been shown to mediate methyltransferase activity in YLR183C 2.140 up Tos4p Transcription factor that binds to a number of promoter regions, particularly promoters of some genes involved YBR022W 2.142 up Poa1p Phosphatase that is highly specific for ADP-ribose 1''-phosphate, a tRNA splicing metabolite; may have a role YNL289W 2.157 up Pcl1p Pho85 cyclin of the Pcl1,2-like subfamily, involved in entry into the mitotic cell cycle and regulation of morphog YKL009W 2.165 up Mrt4p Protein involved in mRNA turnover and ribosome assembly, localizes to the nucleolus YHL024W 2.165 up Rim4p Putative RNA-binding protein required for the expression of early and middle sporulation genes YLR058C 2.171 up Shm2p Cytosolic serine hydroxymethyltransferase, converts serine to glycine plus 5,10 methylenetetrahydrofolate; ma YNR044W 2.172 up Aga1p Anchorage subunit of a-agglutinin of a-cells, highly O-glycosylated protein with N-terminal secretion signal and YHR035W 2.175 up Yhr035wp Putative protein of unknown function; not an essential gene YKL172W 2.184 up Ebp2p Essential protein required for the maturation of 25S rRNA and 60S ribosomal subunit assembly, localizes to th YDL182W 2.186 up Lys20p Homocitrate synthase isozyme, catalyzes the condensation of acetyl-CoA and alpha-ketoglutarate to form hom YNL024C-A 2.193 up Ynl024c-ap Putative protein of unknown function; YNL024C-A is an essential gene

182 Table A.1 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YMR303C 2.193 up Adh2p Glucose-repressible alcohol dehydrogenase II, catalyzes the conversion of ethanol to acetaldehyde; involved YOL077C 2.196 up Brx1p Nucleolar protein, constituent of 66S pre-ribosomal particles; depletion leads to defects in rRNA processing an YLR397C 2.201 up Afg2p ATPase of the CDC48/PAS1/SEC18 (AAA) family, forms a hexameric complex; may be involved in degradatio YLR196W 2.211 up Pwp1p Protein with WD-40 repeats involved in rRNA processing; associates with trans-acting ribosome biogenesis fa YOR341W 2.212 up Rpa190p RNA polymerase I subunit; largest subunit of RNA polymerase I YMR120C 2.212 up Ade17p Enzyme of 'de novo' purine biosynthesis containing both 5-aminoimidazole-4-carboxamide ribonucleotide tran YDR309C 2.213 up Gic2p Protein of unknown function involved in initiation of budding and cellular polarization, interacts with Cdc42p via YNL002C 2.214 up Rlp7p Nucleolar protein with similarity to large ribosomal subunit L7 proteins; constituent of 66S pre-ribosomal partic YPL212C 2.222 up Pus1p tRNA:pseudouridine synthase, introduces pseudouridines at positions 26-28, 34-36, 65, and 67 of tRNA; nucle YAR015W 2.224 up Ade1p N-succinyl-5-aminoimidazole-4-carboxamide ribotide (SAICAR) synthetase, required for 'de novo' purine nucle YOR206W 2.235 up Noc2p Protein that forms a nucleolar complex with Mak21p that binds to 90S and 66S pre-ribosomes, as well as a nu YKR081C 2.237 up Rpf2p Essential protein involved in the processing of pre-rRNA and the assembly of the 60S ribosomal subunit; inter YOL126C 2.239 up Mdh2p Cytoplasmic malate dehydrogenase, one of the three isozymes that catalyze interconversion of malate and ox YPL126W 2.246 up Nan1p U3 snoRNP protein, component of the small (ribosomal) subunit (SSU) processosome containing U3 snoRNA YMR128W 2.248 up Ecm16p Essential DEAH-box ATP-dependent RNA helicase specific to the U3 snoRNP, predominantly nucleolar in dis YPR002W 2.251 up Pdh1p Mitochondrial protein that participates in respiration, induced by diauxic shift; homologous to E. coli PrpD, may YDR234W 2.251 up Lys4p Homoaconitase, catalyzes the conversion of homocitrate to homoisocitrate, which is a step in the lysine biosy YLR168C 2.287 up Aim30p Putative protein of unknown function that may be involved in intramitochondrial sorting; similar to Ups1p and t YOR028C 2.295 up Cin5p Basic leucine zipper transcriptional factor of the yAP-1 family that mediates pleiotropic drug resistance and sa YCL037C 2.309 up Sro9p Cytoplasmic RNA-binding protein that associates with translating ribosomes; involved in heme regulation of H YHL026C 2.311 up Yhl026cp Putative protein of unknown function; YHL026C is not an essential gene; in 2005 the start site was moved 141 YPL043W 2.321 up Nop4p Nucleolar protein, essential for processing and maturation of 27S pre-rRNA and large ribosomal subunit bioge YLR409C 2.348 up Utp21p Possible U3 snoRNP protein involved in maturation of pre-18S rRNA, based on computational analysis of larg YMR290C 2.349 up Has1p ATP-dependent RNA helicase; localizes to both the nuclear periphery and nucleolus; highly enriched in nuclea YMR014W 2.354 up Bud22p Protein involved in bud-site selection; diploid mutants display a random budding pattern instead of the wild-typ YLL027W 2.361 up Isa1p Mitochondrial matrix protein involved in biogenesis of the iron-sulfur (Fe/S) cluster of Fe/S proteins, isa1 delet YNL251C 2.364 up Nrd1p RNA-binding protein that interacts with the C-terminal domain of the RNA polymerase II large subunit (Rpo21 YNL248C 2.365 up Rpa49p RNA polymerase I subunit A49 YNL030W 2.365 up Hhf2p One of two identical histone H4 proteins (see also HHF1); core histone required for chromatin assembly and c YIR034C 2.368 up Lys1p Saccharopine dehydrogenase (NAD+, L-lysine-forming), catalyzes the conversion of saccharopine to L-lysine YPR062W 2.371 up Fcy1p Cytosine deaminase, zinc metalloenzyme that catalyzes the hydrolytic deamination of cytosine to uracil; of bio YLR129W 2.381 up Dip2p Nucleolar protein, specifically associated with the U3 snoRNA, part of the large ribonucleoprotein complex kno YMR189W 2.403 up Gcv2p P subunit of the mitochondrial glycine decarboxylase complex, required for the catabolism of glycine to 5,10-m YDR501W 2.414 up Plm2p Protein required for partitioning of the 2-micron plasmid YOL041C 2.417 up Nop12p Nucleolar protein, required for pre-25S rRNA processing; contains an RNA recognition motif (RRM) and has s YGR145W 2.419 up Enp2p Essential nucleolar protein of unknown function; contains WD repeats, interacts with Mpp10p and Bfr2p, and h YMR300C 2.423 up Ade4p Phosphoribosylpyrophosphate amidotransferase (PRPPAT; amidophosphoribosyltransferase), catalyzes first YHR196W 2.424 up Utp9p Nucleolar protein, component of the small subunit (SSU) processome containing the U3 snoRNA that is involv YOL151W 2.430 up Gre2p NADPH-dependent methylglyoxal reductase (D-lactaldehyde dehydrogenase); stress induced (osmotic, ionic, YER006W 2.432 up Nug1p GTPase that associates with nuclear 60S pre-ribosomes, required for export of 60S ribosomal subunits from t YGR128C 2.443 up Utp8p Nucleolar protein required for export of tRNAs from the nucleus; also copurifies with the small subunit (SSU) p YLL008W 2.452 up Drs1p Nucleolar DEAD-box protein required for ribosome assembly and function, including synthesis of 60S ribosom YHR092C 2.452 up Hxt4p High-affinity glucose transporter of the major facilitator superfamily, expression is induced by low levels of gluc YDR299W 2.452 up Bfr2p Essential protein possibly involved in secretion; multicopy suppressor of sensitivity to Brefeldin A YGL136C 2.464 up Mrm2p Mitochondrial 2' O-ribose methyltransferase, required for methylation of U(2791) in 21S rRNA; MRM2 deletion

183 Table A.1 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YDL020C 2.495 up Rpn4p Transcription factor that stimulates expression of proteasome genes; Rpn4p levels are in turn regulated by the YLL034C 2.496 up Rix7p Putative ATPase of the AAA family, required for export of pre-ribosomal large subunits from the nucleus; distr YOR196C 2.528 up Lip5p Protein involved in biosynthesis of the coenzyme lipoic acid, has similarity to E. coli lipoic acid synthase YNR050C 2.530 up Lys9p Saccharopine dehydrogenase (NADP+, L-glutamate-forming); catalyzes the formation of saccharopine from a YLR401C 2.538 up Dus3p Dihydrouridine synthase, member of a widespread family of conserved proteins including Smm1p, Dus1p, and YDL003W 2.539 up Mcd1p Essential protein required for sister chromatid cohesion in mitosis and meiosis; subunit of the cohesin complex YGR103W 2.543 up Nop7p Nucleolar protein involved in rRNA processing and 60S ribosomal subunit biogenesis; constituent of several d YKL143W 2.547 up Ltv1p Component of the GSE complex, which is required for proper sorting of amino acid permease Gap1p; required YHR033W 2.553 up Yhr033wp Putative protein of unknown function; epitope-tagged protein localizes to the cytoplasm YPR190C 2.564 up Rpc82p RNA polymerase III subunit C82 YGR088W 2.566 up Ctt1p Cytosolic catalase T, has a role in protection from oxidative damage by hydrogen peroxide YNL274C 2.573 up Gor1p Glyoxylate reductase; null mutation results in increased biomass after diauxic shift; the authentic, non-tagged YNL112W 2.582 up Dbp2p Essential ATP-dependent RNA helicase of the DEAD-box protein family, involved in nonsense-mediated mRN YGL234W 2.583 up Ade5,7p Bifunctional enzyme of the 'de novo' purine nucleotide biosynthetic pathway, contains aminoimidazole ribotide YJR048W 2.619 up Cyc1p Cytochrome c, isoform 1; electron carrier of the mitochondrial intermembrane space that transfers electrons fr YBR092C 2.620 up Pho3p Constitutively expressed acid phosphatase similar to Pho5p; brought to the cell surface by transport vesicles; YOR298W 2.624 up Mum3p Protein of unknown function involved in the organization of the outer spore wall layers; has similarity to the taf YKL014C 2.647 up Urb1p Nucleolar protein required for the normal accumulation of 25S and 5.8S rRNAs, associated with the 27SA2 pr YER081W 2.670 up Ser3p 3-phosphoglycerate dehydrogenase, catalyzes the first step in serine and glycine biosynthesis; isozyme of Se YJL200C 2.720 up Aco2p Putative mitochondrial aconitase isozyme; similarity to Aco1p, an aconitase required for the TCA cycle; expres YNL132W 2.722 up Kre33p Essential protein of unknown function; heterozygous mutant shows haploinsufficiency in K1 killer toxin resista YLR174W 2.757 up Idp2p Cytosolic NADP-specific isocitrate dehydrogenase, catalyzes oxidation of isocitrate to alpha-ketoglutarate; lev YOR205C 2.768 up Lrc5p Protein of unknown function; non-tagged protein is detected in purified mitochondria in high-throughput studie YDR060W 2.778 up Mak21p Constituent of 66S pre-ribosomal particles, required for large (60S) ribosomal subunit biogenesis; involved in YDR101C 2.778 up Arx1p Shuttling pre-60S factor; involved in the biogenesis of ribosomal large subunit biogenesis; interacts directly wi YBR009C 2.797 up Hhf1p One of two identical histone H4 proteins (see also HHF2); core histone required for chromatin assembly and c YPR112C 2.811 up Mrd1p Essential conserved protein that associates with 35S precursor rRNA and is required for its initial processing a YKR080W 2.849 up Mtd1p NAD-dependent 5,10-methylenetetrahydrafolate dehydrogenase, plays a catalytic role in oxidation of cytoplas YER024W 2.882 up Yat2p Carnitine acetyltransferase; has similarity to Yat1p, which is a carnitine acetyltransferase associated with the YMR229C 2.904 up Rrp5p Protein required for the synthesis of both 18S and 5.8S rRNA; C-terminal region is crucial for the formation of YMR015C 2.989 up Erg5p C-22 sterol desaturase, a cytochrome P450 enzyme that catalyzes the formation of the C-22(23) double bond YNL231C 3.038 up Pdr16p Phosphatidylinositol transfer protein (PITP) controlled by the multiple drug resistance regulator Pdr1p, localize YNL299W 3.058 up Trf5p Poly (A) polymerase involved in nuclear RNA quality control based on: homology with Trf4p, genetic interactio YKR093W 3.090 up Ptr2p Integral membrane peptide transporter, mediates transport of di- and tri-peptides; conserved protein that conta YOR249C 3.092 up Apc5p Subunit of the Anaphase-Promoting Complex/Cyclosome (APC/C), which is a ubiquitin-protein ligase required YER110C 3.121 up Kap123p Karyopherin beta, mediates nuclear import of ribosomal proteins prior to assembly into ribosomes and import YLR082C 3.199 up Srl2p Protein of unknown function; overexpression suppresses the lethality caused by a rad53 null mutation YBL039C 3.200 up Ura7p Major CTP synthase isozyme (see also URA8), catalyzes the ATP-dependent transfer of the amide nitrogen f YLR180W 3.356 up Sam1p S-adenosylmethionine synthetase, catalyzes transfer of the adenosyl group of ATP to the sulfur atom of meth YAL054C 3.358 up Acs1p Acetyl-coA synthetase isoform which, along with Acs2p, is the nuclear source of acetyl-coA for histone acetlya YDR222W 3.831 up Ydr222wp Protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplasm in a pu YDL148C 3.850 up Nop14p Nucleolar protein, forms a complex with Noc4p that mediates maturation and nuclear export of 40S ribosomal YEL039C 3.932 up Cyc7p Cytochrome c isoform 2, expressed under hypoxic conditions; electron carrier of the mitochondrial intermemb YBR021W 4.296 up Fur4p Uracil permease, localized to the plasma membrane; expression is tightly regulated by uracil levels and enviro YGL010W 4.466 up Ygl010wp Putative protein of unknown function; YGL010W is not an essential gene

184 Table A.1 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YHR007C-A 4.527 up Yhr007c-ap Putative protein of unknown function; identified by expression profiling and mass spectrometry YNL111C 5.125 up Cyb5p Cytochrome b5, involved in the sterol and lipid biosynthesis pathways; required for sterol C5-6 and fatty acid d YBR010W 5.209 up Hht1p One of two identical histone H3 proteins (see also HHT2); core histone required for chromatin assembly, invo YHR156C 5.885 up Lin1p Non-essential component of U5 snRNP; nuclear protein; physically interacts with Irr1p of cohesin complex; ma YLR342W-A 6.188 up Ylr342w-ap Putative protein of unknown function YLR343W 6.829 up Gas2p 1,3-beta-glucanosyltransferase, involved with Gas4p in spore wall assembly; has similarity to Gas1p YDR007W 16.286 up Trp1p Phosphoribosylanthranilate isomerase that catalyzes the third step in tryptophan biosynthesis; in 2004, the se

Table A2. List of genes affected 2-fold or more by the simultaneous overexpression of all the four core histones.

GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YKL056C 4.692 down Tma19p Protein that associates with ribosomes; homolog of translationally controlled tumor protein; green fluorescen YKL180W 4.560 down Rpl17ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl17Bp and has similarity to E. c YOR063W 4.285 down Rpl3p Protein component of the large (60S) ribosomal subunit, has similarity to E. coli L3 and rat L3 ribosomal prot YOL120C 4.114 down Rpl18ap Protein component of the large (60S) ribosomal subunit, identical to Rpl18Bp and has similarity to rat L18 rib YLR340W 4.100 down Rpp0p Conserved ribosomal protein P0 similar to rat P0, human P0, and E. coli L10e; shown to be phosphorylated YLR367W 3.903 down Rps22bp Protein component of the small (40S) ribosomal subunit; nearly identical to Rps22Ap and has similarity to E. YLR354C 3.900 down Tal1p Transaldolase, enzyme in the non-oxidative pentose phosphate pathway; converts sedoheptulose 7-phospha YNL069C 3.466 down Rpl16bp N-terminally acetylated protein component of the large (60S) ribosomal subunit, binds to 5.8 S rRNA; has sim YOR312C 3.368 down Rpl20bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl20Ap and has similarity to rat YGR260W 3.341 down Tna1p High affinity nicotinic acid plasma membrane permease, responsible for uptake of low levels of nicotinic acid YEL021W 3.267 down Ura3p Orotidine-5'-phosphate (OMP) decarboxylase, catalyzes the sixth enzymatic step in the de novo biosynthesis YNL178W 3.252 down Rps3p Protein component of the small (40S) ribosomal subunit, has apurinic/apyrimidinic (AP) endonuclease activit YIL148W 3.210 down Rpl40ap Fusion protein, identical to Rpl40Bp, that is cleaved to yield ubiquitin and a ribosomal protein of the large (60 YER073W 3.204 down Ald5p Mitochondrial aldehyde dehydrogenase, involved in regulation or biosynthesis of electron transport chain com YJL177W 3.153 down Rpl17bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl17Ap and has similarity to E. c YGL189C 3.135 down Rps26ap Protein component of the small (40S) ribosomal subunit; nearly identical to Rps26Bp and has similarity to rat YGL123W 3.115 down Rps2p Protein component of the small (40S) subunit, essential for control of translational accuracy; has similarity to YDR158W 3.072 down Hom2p Aspartic beta semi-aldehyde dehydrogenase, catalyzes the second step in the common pathway for methion YGL103W 3.049 down Rpl28p Ribosomal protein of the large (60S) ribosomal subunit, has similarity to E. coli L15 and rat L27a ribosomal p YGR282C 3.043 down Bgl2p Endo-beta-1,3-glucanase, major protein of the cell wall, involved in cell wall maintenance YOR209C 3.028 down Npt1p Nicotinate phosphoribosyltransferase, acts in the salvage pathway of NAD+ biosynthesis; required for silenc YBR056W 3.026 down Ybr056wp Putative cytoplasmic protein of unknown function YMR143W 3.020 down Rps16ap Protein component of the small (40S) ribosomal subunit; identical to Rps16Bp and has similarity to E. coli S9

185 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YIL124W 2.978 down Ayr1p NADPH-dependent 1-acyl dihydroxyacetone phosphate reductase found in lipid particles, ER, and mitochond YLR441C 2.973 down Rps1ap Ribosomal protein 10 (rp10) of the small (40S) subunit; nearly identical to Rps1Bp and has similarity to rat S YMR116C 2.972 down Asc1p G-protein beta subunit and guanine nucleotide dissociation inhibitor for Gpa2p; ortholog of RACK1 that inhib YDR064W 2.953 down Rps13p Protein component of the small (40S) ribosomal subunit; has similarity to E. coli S15 and rat S13 ribosomal p YPL048W 2.952 down Cam1p Nuclear protein required for transcription of MXR1; binds the MXR1 promoter in the presence of other nuclea YGR295C 2.947 down Cos6p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encode YER117W 2.939 down Rpl23bp Protein component of the large (60S) ribosomal subunit, identical to Rpl23Ap and has similarity to E. coli L14 YKR094C 2.932 down Rpl40bp Fusion protein, identical to Rpl40Ap, that is cleaved to yield ubiquitin and a ribosomal protein of the large (60 YBR191W 2.913 down Rpl21ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl21Bp and has similarity to rat YLR029C 2.901 down Rpl15ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl15Bp and has similarity to rat YDR447C 2.871 down Rps17bp Ribosomal protein 51 (rp51) of the small (40s) subunit; nearly identical to Rps17Ap and has similarity to rat S YGL076C 2.866 down Rpl7ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl7Bp and has similarity to E. co YOR382W 2.844 down Fit2p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved YMR242C 2.835 down Rpl20ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl20Bp and has similarity to rat YDL081C 2.816 down Rpp1ap Ribosomal stalk protein P1 alpha, involved in the interaction between translational elongation factors and the YNL239W 2.795 down Lap3p Cysteine aminopeptidase with homocysteine-thiolactonase activity; protects cells against homocysteine toxic YCR031C 2.780 down Rps14ap Ribosomal protein 59 of the small subunit, required for ribosome assembly and 20S pre-rRNA processing; m YPL078C 2.776 down Atp4p Subunit b of the stator stalk of mitochondrial F1F0 ATP synthase, which is a large, evolutionarily conserved e YDR009W 2.751 down Gal3p Transcriptional regulator involved in activation of the GAL genes in response to galactose; forms a complex w YNR016C 2.750 down Acc1p Acetyl-CoA carboxylase, biotin containing enzyme that catalyzes the carboxylation of acetyl-CoA to form ma YJL136C 2.742 down Rps21bp Protein component of the small (40S) ribosomal subunit; nearly identical to Rps21Ap and has similarity to rat YDR023W 2.740 down Ses1p Cytosolic seryl-tRNA synthetase, class II aminoacyl-tRNA synthetase that aminoacylates tRNA(Ser), display YMR142C 2.718 down Rpl13bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl13Ap; not essential for viability YGR240C 2.706 down Pfk1p Alpha subunit of heterooctameric phosphofructokinase involved in glycolysis, indispensable for anaerobic gro YDL055C 2.698 down Psa1p GDP-mannose pyrophosphorylase (mannose-1-phosphate guanyltransferase), synthesizes GDP-mannose f YML063W 2.685 down Rps1bp Ribosomal protein 10 (rp10) of the small (40S) subunit; nearly identical to Rps1Ap and has similarity to rat S YDR471W 2.681 down Rpl27bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl27Ap and has similarity to rat YPL143W 2.636 down Rpl33ap N-terminally acetylated ribosomal protein L37 of the large (60S) ribosomal subunit, nearly identical to Rpl33B YMR295C 2.627 down Ymr295cp Protein of unknown function that associates with ribosomes; green fluorescent protein (GFP)-fusion protein l YJR145C 2.625 down Rps4ap Protein component of the small (40S) ribosomal subunit; mutation affects 20S pre-rRNA processing; identica YDR012W 2.623 down Rpl4bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl4Ap and has similarity to E. co YHL033C 2.617 down Rpl8ap Ribosomal protein L4 of the large (60S) ribosomal subunit, nearly identical to Rpl8Bp and has similarity to ra YDR381W 2.604 down Yra1p Nuclear protein that binds to RNA and to Mex67p, required for export of poly(A)+ mRNA from the nucleus; m YDL070W 2.596 down Bdf2p Protein involved in transcription initiation at TATA-containing promoters; associates with the basal transcripti YBR031W 2.582 down Rpl4ap N-terminally acetylated protein component of the large (60S) ribosomal subunit, nearly identical to Rpl4Bp an YGR148C 2.572 down Rpl24bp Ribosomal protein L30 of the large (60S) ribosomal subunit, nearly identical to Rpl24Ap and has similarity to YHL015W 2.572 down Rps20p Protein component of the small (40S) ribosomal subunit; overproduction suppresses mutations affecting RNA YML073C 2.571 down Rpl6ap N-terminally acetylated protein component of the large (60S) ribosomal subunit, has similarity to Rpl6Bp and YHR203C 2.569 down Rps4bp Protein component of the small (40S) ribosomal subunit; identical to Rps4Ap and has similarity to rat S4 ribo YER043C 2.540 down Sah1p S-adenosyl-L-homocysteine hydrolase, catabolizes S-adenosyl-L-homocysteine which is formed after donati YKL043W 2.538 down Phd1p Transcriptional activator that enhances pseudohyphal growth; regulates expression of FLO11, an adhesin re YJL190C 2.534 down Rps22ap Protein component of the small (40S) ribosomal subunit; nearly identical to Rps22Bp and has similarity to E. YBR067C 2.489 down Tip1p Major cell wall mannoprotein with possible lipase activity; transcription is induced by heat- and cold-shock; m

186 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YER074W 2.481 down Rps24ap Protein component of the small (40S) ribosomal subunit; identical to Rps24Bp and has similarity to rat S24 ri YLR448W 2.478 down Rpl6bp Protein component of the large (60S) ribosomal subunit, has similarity to Rpl6Ap and to rat L6 ribosomal pro YDR033W 2.469 down Mrh1p Protein that localizes primarily to the plasma membrane, also found at the nuclear envelope; the authentic, n YDR233C 2.467 down Rtn1p ER membrane protein that interacts with exocyst subunit Sec6p and with Yip3p; also interacts with Sbh1p; n YDR377W 2.459 down Atp17p Subunit f of the F0 sector of mitochondrial F1F0 ATP synthase, which is a large, evolutionarily conserved en YIL069C 2.455 down Rps24bp Protein component of the small (40S) ribosomal subunit; identical to Rps24Ap and has similarity to rat S24 ri YGR214W 2.444 down Rps0ap Protein component of the small (40S) ribosomal subunit, nearly identical to Rps0Bp; required for maturation YCR024C-A 2.443 down Pmp1p Small single-membrane span proteolipid that functions as a regulatory subunit of the plasma membrane H(+ YGR034W 2.440 down Rpl26bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl26Ap and has similarity to E. c YOR385W 2.423 down Yor385wp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplas YGL125W 2.412 down Met13p Isozyme of methylenetetrahydrofolate reductase, catalyzes the reduction of 5,10-methylenetetrahydrofolate t YKL080W 2.404 down Vma5p Subunit C of the eight-subunit V1 peripheral membrane domain of vacuolar H+-ATPase (V-ATPase), an elec YMR099C 2.402 down Ymr099cp Glucose-6-phosphate 1-epimerase (hexose-6-phosphate mutarotase), likely involved in carbohydrate metab YMR307W 2.386 down Gas1p Beta-1,3-glucanosyltransferase, required for cell wall assembly; localizes to the cell surface via a glycosylpho YDL082W 2.385 down Rpl13ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl13Bp; not essential for viability YGR250C 2.382 down Ygr250cp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplas YBR121C 2.379 down Grs1p Cytoplasmic and mitochondrial glycyl-tRNA synthase that ligates glycine to the cognate anticodon bearing tR YPL058C 2.378 down Pdr12p Plasma membrane ATP-binding cassette (ABC) transporter, weak-acid-inducible multidrug transporter requir YBL092W 2.360 down Rpl32p Protein component of the large (60S) ribosomal subunit, has similarity to rat L32 ribosomal protein; overexpr YEL054C 2.358 down Rpl12ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl12Bp; rpl12a rpl12b double m YML024W 2.353 down Rps17ap Ribosomal protein 51 (rp51) of the small (40s) subunit; nearly identical to Rps17Bp and has similarity to rat S YEL017C-A 2.351 down Pmp2p Proteolipid associated with plasma membrane H(+)-ATPase (Pma1p); regulates plasma membrane H(+)-AT YLR075W 2.345 down Rpl10p Protein component of the large (60S) ribosomal subunit, responsible for joining the 40S and 60S subunits; re YER056C-A 2.333 down Rpl34ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl34Bp and has similarity to rat YJR123W 2.320 down Rps5p Protein component of the small (40S) ribosomal subunit, the least basic of the non-acidic ribosomal proteins YDR483W 2.313 down Kre2p Alpha1,2-mannosyltransferase of the Golgi involved in protein mannosylation YGR118W 2.301 down Rps23ap Ribosomal protein 28 (rp28) of the small (40S) ribosomal subunit, required for translational accuracy; nearly YDR168W 2.297 down Cdc37p Essential Hsp90p co-chaperone; necessary for passage through the START phase of the cell cycle; stabilize YKL073W 2.296 down Lhs1p Molecular chaperone of the endoplasmic reticulum lumen, involved in polypeptide translocation and folding; YJR016C 2.290 down Ilv3p Dihydroxyacid dehydratase, catalyzes third step in the common pathway leading to biosynthesis of branched YER133W 2.288 down Glc7p Catalytic subunit of type 1 serine/threonine protein phosphatase, involved in many processes including glyco YDR524C-B 2.288 down Ydr524c-bp Putative protein of unknown function YOR232W 2.283 down Mge1p Protein of the mitochondrial matrix involved in protein import into mitochondria; acts as a cochaperone and a YKL157W 2.273 down Ape2p Zinc-dependent metallopeptidase yscII, may have a role in obtaining leucine from dipeptide substrates; sequ YOR096W 2.273 down Rps7ap Protein component of the small (40S) ribosomal subunit, nearly identical to Rps7Bp; interacts with Kti11p; de YGR279C 2.268 down Scw4p Cell wall protein with similarity to glucanases; scw4 scw10 double mutants exhibit defects in mating YDL192W 2.254 down Arf1p ADP-ribosylation factor, GTPase of the Ras superfamily involved in regulation of coated vesicle formation in YGL135W 2.253 down Rpl1bp N-terminally acetylated protein component of the large (60S) ribosomal subunit, nearly identical to Rpl1Ap an YOL040C 2.249 down Rps15p Protein component of the small (40S) ribosomal subunit; has similarity to E. coli S19 and rat S15 ribosomal p YDR226W 2.245 down Adk1p Adenylate kinase, required for purine metabolism; localized to the cytoplasm and the mitochondria; lacks cle YBR249C 2.245 down Aro4p 3-deoxy-D-arabino-heptulosonate-7-phosphate (DAHP) synthase, catalyzes the first step in aromatic amino YNR067C 2.242 down Dse4p Daughter cell-specific secreted protein with similarity to glucanases, degrades cell wall from the daughter sid YPR156C 2.240 down Tpo3p Polyamine transport protein specific for spermine; localizes to the plasma membrane; member of the major f

187 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YJR077C 2.239 down Mir1p Mitochondrial phosphate carrier, imports inorganic phosphate into mitochondria; functionally redundant with YIL138C 2.232 down Tpm2p Minor isoform of tropomyosin, binds to and stabilizes actin cables and filaments, which direct polarized cell g YER053C 2.228 down Pic2p Mitochondrial phosphate carrier, imports inorganic phosphate into mitochondria; functionally redundant with YNL173C 2.225 down Mdg1p Plasma membrane protein involved in G-protein mediated pheromone signaling pathway; overproduction sup YCR009C 2.223 down Rvs161p Amphiphysin-like lipid raft protein; subunit of a complex (Rvs161p-Rvs167p) that regulates polarization of the YNL135C 2.222 down Fpr1p Peptidyl-prolyl cis-trans isomerase (PPIase), binds to the drugs FK506 and rapamycin; also binds to the non YER178W 2.212 down Pda1p E1 alpha subunit of the pyruvate dehydrogenase (PDH) complex, catalyzes the direct oxidative decarboxylat YGR244C 2.210 down Lsc2p Beta subunit of succinyl-CoA ligase, which is a mitochondrial enzyme of the TCA cycle that catalyzes the nu YML072C 2.208 down Tcb3p Lipid-binding protein, localized to the bud via specific mRNA transport; non-tagged protein detected in a pho YPR132W 2.208 down Rps23bp Ribosomal protein 28 (rp28) of the small (40S) ribosomal subunit, required for translational accuracy; nearly YFL005W 2.204 down Sec4p Secretory vesicle-associated Rab GTPase essential for exocytosis; associates with the exocyst component S YGR094W 2.201 down Vas1p Mitochondrial and cytoplasmic valyl-tRNA synthetase YFL031W 2.200 down Hac1p bZIP transcription factor (ATF/CREB1 homolog) that regulates the unfolded protein response, via UPRE bind YDR368W 2.199 down Ypr1p 2-methylbutyraldehyde reductase, may be involved in isoleucine catabolism YOR109W 2.199 down Inp53p Polyphosphatidylinositol phosphatase, dephosphorylates multiple phosphatidylinositols; involved in trans Go YER124C 2.190 down Dse1p Daughter cell-specific protein, may participate in pathways regulating cell wall metabolism; deletion affects c YDL137W 2.180 down Arf2p ADP-ribosylation factor, GTPase of the Ras superfamily involved in regulation of coated formation vesicles in YDL040C 2.176 down Nat1p Subunit of the N-terminal acetyltransferase NatA (Nat1p, Ard1p, Nat5p); N-terminally acetylates many protei YIL018W 2.175 down Rpl2bp Protein component of the large (60S) ribosomal subunit, identical to Rpl2Ap and has similarity to E. coli L2 a YPL131W 2.172 down Rpl5p Protein component of the large (60S) ribosomal subunit with similarity to E. coli L18 and rat L5 ribosomal pro YPL270W 2.172 down Mdl2p Mitochondrial inner membrane half-type ATP-binding cassette (ABC) transporter YDL248W 2.165 down Cos7p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encode YPL210C 2.160 down Srp72p Core component of the signal recognition particle (SRP) ribonucleoprotein (RNP) complex that functions in ta YPL195W 2.158 down Apl5p Delta adaptin-like subunit of the clathrin associated protein complex (AP-3); functions in transport of alkaline YMR200W 2.157 down Rot1p Essential ER membrane protein; may be involved in protein folding; mutation causes defects in cell wall synt YOR383C 2.156 down Fit3p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved YER136W 2.151 down Gdi1p GDP dissociation inhibitor, regulates vesicle traffic in secretory pathways by regulating the dissociation of GD YPL079W 2.140 down Rpl21bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl21Ap and has similarity to rat YMR008C 2.135 down Plb1p Phospholipase B (lysophospholipase) involved in lipid metabolism, required for deacylation of phosphatidylch YMR110C 2.135 down Hfd1p Putative fatty aldehyde dehydrogenase, located in the mitochondrial outer membrane and also in lipid particl YJL171C 2.135 down Yjl171cp GPI-anchored cell wall protein of unknown function; induced in response to cell wall damaging agents and by YPR102C 2.128 down Rpl11ap Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl11Bp; involved in ribosomal as YOR198C 2.127 down Bfr1p Component of mRNP complexes associated with polyribosomes; implicated in secretion and nuclear segreg YCR034W 2.127 down Fen1p Fatty acid elongase, involved in sphingolipid biosynthesis; acts on fatty acids of up to 24 carbons in length; m YOR293W 2.124 down Rps10ap Protein component of the small (40S) ribosomal subunit; nearly identical to Rps10Bp and has similarity to rat YMR297W 2.122 down Prc1p Vacuolar carboxypeptidase Y (proteinase C), broad-specificity C-terminal exopeptidase involved in non-spec YPL084W 2.121 down Bro1p Cytoplasmic class E vacuolar protein sorting (VPS) factor that coordinates deubiquitination in the multivesicu YDR525W-A 2.113 down Sna2p Protein of unknown function, has similarity to Pmp3p, which is involved in cation transport; green fluorescent YHR001W-A 2.109 down Qcr10p Subunit of the ubiqunol-cytochrome c oxidoreductase complex which includes Cobp, Rip1p, Cyt1p, Cor1p, Q YHL047C 2.108 down Arn2p Transporter, member of the ARN family of transporters that specifically recognize siderophore-iron chelates; YPL220W 2.107 down Rpl1ap N-terminally acetylated protein component of the large (60S) ribosomal subunit, nearly identical to Rpl1Bp an YGL031C 2.105 down Rpl24ap Ribosomal protein L30 of the large (60S) ribosomal subunit, nearly identical to Rpl24Bp and has similarity to YMR072W 2.105 down Abf2p Mitochondrial DNA-binding protein involved in mitochondrial DNA replication and recombination, member of

188 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YGR124W 2.103 down Asn2p Asparagine synthetase, isozyme of Asn1p; catalyzes the synthesis of L-asparagine from L-aspartate in the a YOR220W 2.096 down Rcn2p Protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplasm and is YMR230W 2.095 down Rps10bp Protein component of the small (40S) ribosomal subunit; nearly identical to Rps10Ap and has similarity to rat YAL003W 2.092 down Efb1p Translation elongation factor 1 beta; stimulates nucleotide exchange to regenerate EF-1 alpha-GTP for the n YLL045C 2.092 down Rpl8bp Ribosomal protein L4 of the large (60S) ribosomal subunit, nearly identical to Rpl8Ap and has similarity to ra YBR018C 2.089 down Gal7p Galactose-1-phosphate uridyl transferase, synthesizes glucose-1-phosphate and UDP-galactose from UDP- YFL004W 2.086 down Vtc2p Vacuolar membrane protein involved in vacuolar polyphosphate accumulation; functions as a regulator of va YBL072C 2.080 down Rps8ap Protein component of the small (40S) ribosomal subunit; identical to Rps8Bp and has similarity to rat S8 ribo YPL249C-A 2.075 down Rpl36bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl36Ap and has similarity to rat YER130C 2.075 down Yer130cp YDL178W 2.074 down Dld2p D-lactate dehydrogenase, located in the mitochondrial matrix YBR149W 2.071 down Ara1p NADP+ dependent arabinose dehydrogenase, involved in carbohydrate metabolism; purified as homodimer; YEL009C 2.063 down Gcn4p Transcriptional activator of amino acid biosynthetic genes in response to amino acid starvation; expression is YDR127W 2.063 down Aro1p Pentafunctional arom protein, catalyzes steps 2 through 6 in the biosynthesis of chorismate, which is a precu YDL134C 2.061 down Pph21p Catalytic subunit of protein phosphatase 2A, functionally redundant with Pph22p; methylated at C terminus; f YHR098C 2.060 down Sfb3p Member of the Sec24p family; forms a complex, with Sec23p, that is involved in sorting of Pma1p into COPII YDR468C 2.054 down Tlg1p Essential t-SNARE that forms a complex with Tlg2p and Vti1p and mediates fusion of endosome-derived ves YPR160W 2.054 down Gph1p Non-essential glycogen phosphorylase required for the mobilization of glycogen, activity is regulated by cycli YOR175C 2.054 down Ale1p Lysophospholipid acyltransferase, partially redundant with Slc1p; part of MBOAT family of membrane-bound YLR454W 2.053 down Fmp27p Putative protein of unknown function; the authentic, non-tagged protein is detected in highly purified mitocho YKR058W 2.051 down Glg1p Self-glucosylating initiator of glycogen synthesis, also glucosylates n-dodecyl-beta-D-maltoside; similar to ma YGR065C 2.044 down Vht1p High-affinity plasma membrane H+-biotin (vitamin H) symporter; mutation results in fatty acid auxotrophy; 12 YBR025C 2.041 down Ola1p P-loop ATPase with similarity to human OLA1 and bacterial YchF; identified as specifically interacting with th YNR075W 2.041 down Cos10p Protein of unknown function, member of the DUP380 subfamily of conserved, often subtelomerically-encode YHR007C 2.039 down Erg11p Lanosterol 14-alpha-demethylase, catalyzes the C-14 demethylation of lanosterol to form 4,4''-dimethyl chole YMR266W 2.038 down Rsn1p Membrane protein of unknown function; overexpression suppresses NaCl sensitivity of sro7 mutant YLR295C 2.034 down Atp14p Subunit h of the F0 sector of mitochondrial F1F0 ATP synthase, which is a large, evolutionarily conserved en YBR016W 2.033 down Ybr016wp Plasma membrane protein of unknown function; has similarity to hydrophilins, which are hydrophilic, glycine- YBR221C 2.033 down Pdb1p E1 beta subunit of the pyruvate dehydrogenase (PDH) complex, which is an evolutionarily-conserved multi-p YKL209C 2.028 down Ste6p Plasma membrane ATP-binding cassette (ABC) transporter required for the export of a-factor, catalyzes ATP YPR165W 2.027 down Rho1p GTP-binding protein of the rho subfamily of Ras-like proteins, involved in establishment of cell polarity; regul YOR316C 2.026 down Cot1p Vacuolar transporter that mediates zinc transport into the vacuole; overexpression confers resistance to coba YLR194C 2.023 down Ylr194cp Structural constituent of the cell wall attached to the plasma membrane by a GPI-anchor; expression is upreg YOR178C 2.022 down Gac1p Regulatory subunit for Glc7p type-1 protein phosphatase (PP1), tethers Glc7p to Gsy2p glycogen synthase, YGL245W 2.021 down Gus1p Glutamyl-tRNA synthetase (GluRS), forms a complex with methionyl-tRNA synthetase (Mes1p) and Arc1p; c YEL013W 2.021 down Vac8p Phosphorylated vacuolar membrane protein that interacts with Atg13p, required for the cytoplasm-to-vacuole YGL167C 2.019 down Pmr1p High affinity Ca2+/Mn2+ P-type ATPase required for Ca2+ and Mn2+ transport into Golgi; involved in Ca2+ d YAL060W 2.019 down Bdh1p NAD-dependent (R,R)-butanediol dehydrogenase, catalyzes oxidation of (R,R)-2,3-butanediol to (3R)-acetoi YDR502C 2.016 down Sam2p S-adenosylmethionine synthetase, catalyzes transfer of the adenosyl group of ATP to the sulfur atom of met YGR027C 2.016 down Rps25ap Protein component of the small (40S) ribosomal subunit; nearly identical to Rps25Bp and has similarity to rat YKL182W 2.016 down Fas1p Beta subunit of fatty acid synthetase, which catalyzes the synthesis of long-chain saturated fatty acids; conta YBR139W 2.015 down Ybr139wp Putative serine type carboxypeptidase with a role in phytochelatin synthesis; green fluorescent protein (GFP YFR031C-A 2.009 down Rpl2ap Protein component of the large (60S) ribosomal subunit, identical to Rpl2Bp and has similarity to E. coli L2 a

189 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YGL028C 2.003 down Scw11p Cell wall protein with similarity to glucanases; may play a role in conjugation during mating based on its regu YBR143C 2.003 down Sup45p Polypeptide release factor involved in translation termination; mutant form acts as a recessive omnipotent su YPL198W 2.002 down Rpl7bp Protein component of the large (60S) ribosomal subunit, nearly identical to Rpl7Ap and has similarity to E. co YPL160W 2.002 down Cdc60p Cytosolic leucyl tRNA synthetase, ligates leucine to the appropriate tRNA YLR185W 2.001 down Rpl37ap Protein component of the large (60S) ribosomal subunit, has similarity to Rpl37Bp and to rat L37 ribosomal p YBR221W-A 2.000 up Ybr221w-ap Putative protein of unknown function; identified by expression profiling and mass spectrometry YIL099W 2.001 up Sga1p Intracellular sporulation-specific glucoamylase involved in glycogen degradation; induced during starvation o YLR218C 2.004 up Ylr218cp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplas YML100W-A 2.005 up Yml100w-ap Putative protein of unknown function; identified by gene-trapping, microarray-based expression analysis, and YIR013C 2.015 up Gat4p Protein containing GATA family zinc finger motifs YMR262W 2.027 up Ymr262wp Protein of unknown function; interacts weakly with Knr4p; YMR262W is not an essential gene YKL224C 2.044 up Pau16p Putative protein of unknown function YKL037W 2.054 up Aim26p Putative protein of unknown function; null mutant is viable and displays increased frequency of mitochondria YNL299W 2.069 up Trf5p Poly (A) polymerase involved in nuclear RNA quality control based on: homology with Trf4p, genetic interact YKR007W 2.077 up Meh1p Component of the EGO complex, which is involved in the regulation of microautophagy, and of the GSE com YEL035C 2.088 up Utr5p Protein of unknown function; transcription may be regulated by Gcr1p YBR196C-A 2.098 up Ybr196c-ap Putative protein of unknown function; identified by fungal homology and RT-PCR YLR174W 2.110 up Idp2p Cytosolic NADP-specific isocitrate dehydrogenase, catalyzes oxidation of isocitrate to alpha-ketoglutarate; le YDR246W-A 2.114 up Ydr246w-ap Putative protein of unknown function; identified by fungal homology and RT-PCR YOL166W-A 2.123 up Yol166w-ap Identified by gene-trapping, microarray-based expression analysis, and genome-wide homology searching YHL024W 2.145 up Rim4p Putative RNA-binding protein required for the expression of early and middle sporulation genes YFR032C 2.165 up Yfr032cp Putative protein of unknown function; transcribed during sporulation; YFR032C is not an essential gene YML083C 2.168 up Yml083cp Putative protein of unknown function; strong increase in transcript abundance during anaerobic growth comp YJL116C 2.186 up Nca3p Protein that functions with Nca2p to regulate mitochondrial expression of subunits 6 (Atp6p) and 8 (Atp8p ) o YFR035C 2.188 up Yfr035cp Putative protein of unknown function, deletion mutant exhibits synthetic phenotype with alpha-synuclein YCL036W 2.207 up Gfd2p Protein of unknown function, identified as a high-copy suppressor of a dbp5 mutation YLR461W 2.209 up Pau4p Part of 23-member seripauperin multigene family encoded mainly in subtelomeric regions, active during alco YHR156C 2.229 up Lin1p Non-essential component of U5 snRNP; nuclear protein; physically interacts with Irr1p of cohesin complex; m YFR036W 2.233 up Cdc26p Subunit of the Anaphase-Promoting Complex/Cyclosome (APC/C), which is a ubiquitin-protein ligase require YBR090C 2.243 up Ybr090cp Putative protein of unknown function; green fluorescent protein (GFP)-fusion protein localizes to the cytoplas YFL017W-A 2.255 up Smx2p Core Sm protein Sm G; part of heteroheptameric complex (with Smb1p, Smd1p, Smd2p, Smd3p, Sme1p, an YNL030W 2.282 up Hhf2p One of two identical histone H4 proteins (see also HHF1); core histone required for chromatin assembly and YKR077W 2.311 up Msa2p Putative transcriptional activator, that interacts with G1-specific transcription factor, MBF and G1-specific pro YDL218W 2.322 up Ydl218wp Putative protein of unknown function; YDL218W transcription is regulated by Azf1p and induced by starvatio YAL054C 2.362 up Acs1p Acetyl-coA synthetase isoform which, along with Acs2p, is the nuclear source of acetyl-coA for histone acetly YMR303C 2.401 up Adh2p Glucose-repressible alcohol dehydrogenase II, catalyzes the conversion of ethanol to acetaldehyde; involved YOR202W 2.518 up His3p Imidazoleglycerol-phosphate dehydratase, catalyzes the sixth step in histidine biosynthesis; mutations cause YFR012W-A 2.620 up Yfr012w-ap Putative protein of unknown function; identified by homology YBR009C 2.667 up Hhf1p One of two identical histone H4 proteins (see also HHF2); core histone required for chromatin assembly and YGR088W 2.743 up Ctt1p Cytosolic catalase T, has a role in protection from oxidative damage by hydrogen peroxide YBR021W 3.020 up Fur4p Uracil permease, localized to the plasma membrane; expression is tightly regulated by uracil levels and envi YLR343W 3.063 up Gas2p 1,3-beta-glucanosyltransferase, involved with Gas4p in spore wall assembly; has similarity to Gas1p YLR342W-A 3.108 up Ylr342w-ap Putative protein of unknown function

190 Table A.2 continued… GENE_NAME FOLD CHANGE DESCRIPTION FUNCTION YHR007C-A 3.527 up Yhr007c-ap Putative protein of unknown function; identified by expression profiling and mass spectrometry YEL039C 4.426 up Cyc7p Cytochrome c isoform 2, expressed under hypoxic conditions; electron carrier of the mitochondrial intermem YBR010W 5.400 up Hht1p One of two identical histone H3 proteins (see also HHT2); core histone required for chromatin assembly, invo YDR007W 18.085 up Trp1p Phosphoribosylanthranilate isomerase that catalyzes the third step in tryptophan biosynthesis; in 2004, the s

191 Table A3. List of genes that are upregulated or downregulated in response to overexpression of the H3-H4 gene pair as well as upon overexpression of all the core histones.

FOLD FOLD CHANGE GENE_NAME CHANGE FUNCTION DESCRIPTION H3,H4,H2A,H2B H3,H4 YGR260W 3.986 down 3.341 down Tna1p High affinity nicotinic acid plasma membrane permease, responsible for uptake of low lev YEL021W 2.812 down 3.267 down Ura3p Orotidine-5'-phosphate (OMP) decarboxylase, catalyzes the sixth enzymatic step in the d YOR382W 3.134 down 2.844 down Fit2p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GP YKL043W 1.965 down 2.538 down Phd1p Transcriptional activator that enhances pseudohyphal growth; regulates expression of FL YOR383C 3.140 down 2.156 down Fit3p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GP YHL047C 3.401 down 2.108 down Arn2p Transporter, member of the ARN family of transporters that specifically recognize siderop YNR075W 2.108 down 2.041 down Cos10p Protein of unknown function, member of the DUP380 subfamily of conserved, often subte YJR025C 3.016 down 1.966 down Bna1p 3-hydroxyanthranilic acid dioxygenase, required for biosynthesis of nicotinic acid from try YHL048W 1.932 down 1.876 down Cos8p Nuclear membrane protein, member of the DUP380 subfamily of conserved, often subtelo YDR007W 16.286 up 18.085 up Trp1p Phosphoribosylanthranilate isomerase that catalyzes the third step in tryptophan biosynth YBR010W 5.209 up 5.400 up Hht1p One of two identical histone H3 proteins (see also HHT2); core histone required for chrom YEL039C 3.932 up 4.426 up Cyc7p Cytochrome c isoform 2, expressed under hypoxic conditions; electron carrier of the mitoc YHR007C-A 4.527 up 3.527 up Yhr007c-ap Putative protein of unknown function; identified by expression profiling and mass spectrom YLR342W-A 6.188 up 3.108 up Ylr342w-ap Putative protein of unknown function YLR343W 6.829 up 3.063 up Gas2p 1,3-beta-glucanosyltransferase, involved with Gas4p in spore wall assembly; has similarit YBR021W 4.296 up 3.020 up Fur4p Uracil permease, localized to the plasma membrane; expression is tightly regulated by ur YGR088W 2.566 up 2.743 up Ctt1p Cytosolic catalase T, has a role in protection from oxidative damage by hydrogen peroxid YBR009C 2.797 up 2.667 up Hhf1p One of two identical histone H4 proteins (see also HHF2); core histone required for chrom YMR303C 2.193 up 2.401 up Adh2p Glucose-repressible alcohol dehydrogenase II, catalyzes the conversion of ethanol to ace YAL054C 3.358 up 2.362 up Acs1p Acetyl-coA synthetase isoform which, along with Acs2p, is the nuclear source of acetyl-co YDL218W 1.978 up 2.322 up Ydl218wp Putative protein of unknown function; YDL218W transcription is regulated by Azf1p and in YKR077W 1.931 up 2.311 up Msa2p Putative transcriptional activator, that interacts with G1-specific transcription factor, MBF YNL030W 2.365 up 2.282 up Hhf2p One of two identical histone H4 proteins (see also HHF1); core histone required for chrom YHR156C 5.885 up 2.229 up Lin1p Non-essential component of U5 snRNP; nuclear protein; physically interacts with Irr1p of YJL116C 2.099 up 2.186 up Nca3p Protein that functions with Nca2p to regulate mitochondrial expression of subunits 6 (Atp6 YHL024W 2.165 up 2.145 up Rim4p Putative RNA-binding protein required for the expression of early and middle sporulation YLR174W 2.757 up 2.110 up Idp2p Cytosolic NADP-specific isocitrate dehydrogenase, catalyzes oxidation of isocitrate to alp YNL299W 3.058 up 2.069 up Trf5p Poly (A) polymerase involved in nuclear RNA quality control based on: homology with Trf YKR093W 3.090 up 1.999 up Ptr2p Integral membrane peptide transporter, mediates transport of di- and tri-peptides; conserv

192 APPENDIX B

Table B.1 List of genes affected 2-fold or more upon hht2-hhf2 deletion

GENE_NAME FOLD CHANGE FUNCTION DESCRIPTION YOR382W 14.011 down Fit2p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved i YBR072W 6.625 down Hsp26p Small heat shock protein (sHSP) with chaperone activity; forms hollow, sphere-shaped oligomers that suppre YPL223C 5.541 down Gre1p Hydrophilin of unknown function; stress induced (osmotic, ionic, oxidative, heat shock and heavy metals); reg YLR136C 4.800 down Tis11p mRNA-binding protein expressed during iron starvation; binds to a sequence element in the 3'-untranslated re YBR117C 4.581 down Tkl2p Transketolase, similar to Tkl1p; catalyzes conversion of xylulose-5-phosphate and ribose-5-phosphate to sed YOR383C 3.686 down Fit3p Mannoprotein that is incorporated into the cell wall via a glycosylphosphatidylinositol (GPI) anchor, involved i YHR096C 3.563 down Hxt5p Hexose transporter with moderate affinity for glucose, induced in the presence of non-fermentable carbon so YDL222C 3.522 down Fmp45p Integral membrane protein localized to mitochondria (untagged protein) and eisosomes, immobile patches at YMR107W 3.308 down Spg4p Protein required for survival at high temperature during stationary phase; not required for growth on nonferme YHL047C 3.240 down Arn2p Transporter, member of the ARN family of transporters that specifically recognize siderophore-iron chelates; YER103W 3.203 down Ssa4p Heat shock protein that is highly induced upon stress; plays a role in SRP-dependent cotranslational protein- YPL054W 3.086 down Lee1p Zinc-finger protein of unknown function YNL195C 3.075 down Ynl195cp Putative protein of unknown function; shares a promoter with YNL194C; the authentic, non-tagged protein is YGR256W 3.033 down Gnd2p 6-phosphogluconate dehydrogenase (decarboxylating), catalyzes an NADPH regenerating reaction in the pe YLR214W 2.975 down Fre1p Ferric reductase and cupric reductase, reduces siderophore-bound iron and oxidized copper prior to uptake b YMR169C 2.860 down Ald3p Cytoplasmic aldehyde dehydrogenase, involved in beta-alanine synthesis; uses NAD+ as the preferred coenz YFL014W 2.841 down Hsp12p Plasma membrane localized protein that protects membranes from desiccation; induced by heat shock, oxida YGL121C 2.793 down Gpg1p Proposed gamma subunit of the heterotrimeric G protein that interacts with the receptor Grp1p; involved in re YOL052C-A 2.781 down Ddr2p Multistress response protein, expression is activated by a variety of xenobiotic agents and environmental or p YML128C 2.761 down Msc1p Protein of unknown function; mutant is defective in directing meiotic recombination events to homologous chr YHR087W 2.721 down Yhr087wp Protein of unknown function involved in RNA metabolism; this single domain protein has structural similarity t YDR070C 2.721 down Fmp16p Putative protein of unknown function; the authentic, non-tagged protein is detected in highly purified mitochon YOL084W 2.675 down Phm7p Protein of unknown function, expression is regulated by phosphate levels; green fluorescent protein (GFP)-fu YDL021W 2.646 down Gpm2p Homolog of Gpm1p phosphoglycerate mutase which converts 3-phosphoglycerate to 2-phosphoglycerate in g YGR088W 2.617 down Ctt1p Cytosolic catalase T, has a role in protection from oxidative damage by hydrogen peroxide YNR034W-A 2.599 down Ynr034w-ap Putative protein of unknown function; expression is regulated by Msn2p/Msn4p YBR285W 2.575 down Ybr285wp Putative protein of unknown function; YBR285W is not an essential gene and deletion of YBR285W leads to YDL223C 2.511 down Hbt1p Substrate of the Hub1p ubiquitin-like protein that localizes to the shmoo tip (mating projection); mutants are d YGR043C 2.502 down Nqm1p Protein of unknown function; transcription is repressed by Mot1p and induced by alpha-factor and during diau YOR381W 2.420 down Fre3p Ferric reductase, reduces siderophore-bound iron prior to uptake by transporters; expression induced by low YNL194C 2.366 down Ynl194cp Integral membrane protein localized to eisosomes; sporulation and plasma membrane sphingolipid content a YPL186C 2.358 down Uip4p Protein that interacts with Ulp1p, a Ubl (ubiquitin-like protein)-specific protease for Smt3p protein conjugates; YDL085W 2.276 down Nde2p Mitochondrial external NADH dehydrogenase, catalyzes the oxidation of cytosolic NADH; Nde1p and Nde2p YJL052W 2.265 down Tdh1p Glyceraldehyde-3-phosphate dehydrogenase, isozyme 1, involved in glycolysis and gluconeogenesis; tetram YHL035C 2.264 down Vmr1p Protein of unknown function that may interact with ribosomes, based on co-purification experiments; member YDR536W 2.187 down Stl1p Glycerol proton symporter of the plasma membrane, subject to glucose-induced inactivation, strongly but tran YER054C 2.158 down Gip2p Putative regulatory subunit of the protein phosphatase Glc7p, involved in glycogen metabolism; contains a co YNL117W 2.144 down Mls1p Malate synthase, enzyme of the glyoxylate cycle, involved in utilization of non-fermentable carbon sources; e

193 Table B.1 continued… GENE_NAME FOLD CHANGE FUNCTION DESCRIPTION YGR248W 2.129 down Sol4p 6-phosphogluconolactonase with similarity to Sol3p YGR052W 2.107 down Fmp48p Putative protein of unknown function; the authentic, non-tagged protein is detected in highly purified mitochon YBR255C-A 2.097 down Ybr255c-ap Putative protein of unknown function; identified by sequence comparison with hemiascomycetous yeast spec YLR303W 2.066 down Met17p Methionine and cysteine synthase (O-acetyl homoserine-O-acetyl serine sulfhydrylase), required for sulfur am YOR387C 2.054 down Yor387cp Putative protein of unknown function; regulated by the metal-responsive Aft1p transcription factor; highly indu YDR533C 2.053 down Hsp31p Possible chaperone and cysteine protease with similarity to E. coli Hsp31; member of the DJ-1/ThiJ/PfpI supe YKL096W 2.048 down Cwp1p Cell wall mannoprotein, linked to a beta-1,3- and beta-1,6-glucan heteropolymer through a phosphodiester bo YHR215W 2.026 up Pho12p One of three repressible acid phosphatases, a glycoprotein that is transported to the cell surface by the secre YAR071W 2.100 up Pho11p One of three repressible acid phosphatases, a glycoprotein that is transported to the cell surface by the secre YOL104C 2.105 up Ndj1p Meiosis-specific telomere protein, required for bouquet formation, effective homolog pairing, ordered cross-ov YHR216W 2.463 up Imd2p Inosine monophosphate dehydrogenase, catalyzes the first step of GMP biosynthesis, expression is induced YOL166W-A 2.606 up Yol166w-ap Identified by gene-trapping, microarray-based expression analysis, and genome-wide homology searching YGR149W 3.121 up Ygr149wp Putative protein of unknown function; predicted to be an integal membrane protein YOR202W 10.548 up His3p Imidazoleglycerol-phosphate dehydratase, catalyzes the sixth step in histidine biosynthesis; mutations cause

194 BIOGRAPHICAL SKETCH

Dun Liang Education

Florida State University (2005-2011) Tallahassee, FL Ph.D. in Biomedical Sciences, College of Medicine (Degree expected in August 2011)

University of Science & Technology of China (2001-2005) Hefei, China B.S. in Molecular and Cell Biology, School of Life Sciences

Research Experience

FSU July, 2006-present Tallahassee, FL • Research under the mentorship of Dr. Akash Gunjan in the Epigenetics and Genome Stability laboratory. • Focused on comprehensive investigation of Epigenetic effect of histone gene dosage on DNA damage and repair. • Promoted the novel concept of excess histones in regulating genomic instability and optimized the experiments to assay it. The findings have important implications for processes that contribute to cancer and potential pharmaceutical use.

FSU September, 2005-June, 2006 Tallahassee, FL • Finished three rotation research projects: 1) Studied the spindle-kinetochore interaction in budding yeast. 2) Purification and characterization of Src kinase. 3) Investigated transcription factor YY1 and its DNA binding properties.

USTC December, 2003–June, 2005 Hefei, China • Participated in the National High Tech Research & Development (863) Project of China. • Medical engineering investigation of monoclonal antibody against tumor surface antigen, mainly responsible for HER2 specific antibody fragments affinity maturation by phage display.

USTC July, 2004-Febuary, 2005 Hefei, China • Completed the Undergraduate Research Project of USTC, focused on the soluble expression and characterization of disulfide bond-rich sub-domains of breast tumor surface marker HER2 under direction of Dr. Jing Liu, Laboratory of Cell Immunology.

USTC September, 2002 Hefei, China • Designed and coordinated ecological field study on Pinus taiwanensis (Taiwan Pine) in Dabie Mountains in China as team lead. • The report of this survey analyzed characteristics of community structure, interspecies relationships and the habitat’s influence on the community and population of Pinus taiwanensis.

195 USTC May, 2002-June, 2002 Hefei, China • Group organizer of seashore ecology field study in Shandong province, China. • Local marine species were observed and categorized; natural resources and environmental pollution were documented in surrounding area of Qingdao city.

Teaching Experience

Florida State University Tallahassee, FL • 2007-2009: teaching assistant of small group facilitator in Clinical Microanatomy, General Medical Pharmacology, Systemic Pathology, General Medical Microbiology, Medical Biochemistry and Clinical Physiology for first and second year medical students. • 2008-2010: Instructor of Research techniques in Biomedical Sciences (BMS 5186). Provided training in laboratory techniques and experimental approaches to junior graduate students.

Publications and Honors

1. Liang, D. and Gunjan, A. Histone gene dosage modulates DNA repair via the homologous recombination pathway. Under review in PNAS.

2. Singh, R.K., Liang, D., Reddy, G.U., Paik, J., and Gunjan, A. Excess histone levels mediate cytotoxicity via multiple mechanisms. Cell Cycle. 2010 Dec 1;9(23):4611-2.(Co–first author)

3. Morillo-Huesca M, Maya D, Muñoz-Centeno MC, Singh RK, Oreal V, Reddy GU, Liang D, Géli V, Gunjan A, Chávez S. FACT prevents the accumulation of free histones evicted from transcribed chromatin and a subsequent cell cycle delay in G1. PLoS Genet. 2010 May 20;6(5):e1000964.

4. Li, LW., Liu, HB., Hu SY., Liang D., Cheng LS., Liu J. Soluble expression and characterization of disulfide bond-rich subdomains of membrane protein p185 in Escherichia coli. Chinese Journal of Biotechnology. 2005 Jul;21(4):590-6

• October, 2009 Dissertation Research Grant, Florida State University • October, 2004 Excellent Undergraduate Research Project of USTC • September, 2003 Guang-Hua Scholarship of USTC • August, 2002 Excellent Student in the Summer Exercitation of USTC

Presentations and Abstracts • January, 2011 Presented poster at Life Science Symposium “From Molecules to Medicine” at Florida State University, Tallahassee, FL • December,2009 Presented abstract at the annual meeting of American Society for Cell Biology in San Diego, CA • March, 2009 Presented abstract at the Keystone Symposia on Genome Instability and DNA repair in Taos, NM • 2006-2011 Speaker at the Cell Cycle/ Development Biology joint seminar series at FSU

196 • 2006-2011 Speaker at the Cell Biology journal club of College of Medicine, FSU

Professional Membership

• The American Association for the Advancement of Science (AAAS) • The American Society for Cell Biology (ASCB)

Professional Skills • Solid understanding of biomedical sciences and biochemical/molecular technology. • Superior hands-on expertise in yeast genetics, gel electrophoresis, immunoprecipitation, RT-PCR, qPCR, florescence microscopy, flow cytometry and transcription microarray. • Extensive experience in protein purification, library construction, antibody phage display and ELISA. • Strong problem solving skills, detail-oriented and passionate about solutions. • Highly proficient in robotic, in silico operation and data analysis; curiosity of science

Professional Training • 2010 NimbleGen Microarray training, include sample preparation, Hybridization and scanning, Department of Biological Science, Florida State University • 2008 Seminar in Medical Science Education, Department of Family Medicine and Rural Health, Florida State University. • 2007 Radiation Safety Short Course, Department of Environmental Health and Safety, Florida State University. • 2006 Biosafety Level 2 (BSL-2) and Above Laboratory Training, Department of Environmental Health and Safety, Florida State University. • 2005 Proteomics training at Biomedical Proteomics core facility in College of Medicine, Florida State University. • 2005 Cell culture training in the lab of Dr. Myra Hurt in College of Medicine, Florida State University.

Additional Skills Proficient in multiple biological and scientific software, general statistic software, stream/photo editing software, Microsoft Office Suite and Adobe Acrobat.

197