NOVEL ASPECTS TO THE ROLE OF RAD9A DURING THE DNA DAMAGE RESPONSE

by

Megan Lee Sierant

A thesis submitted to the Department of Biochemistry, in conformity with the requirements for the degree of Doctor of Philosophy

Queen’s University Kingston, Ontario, Canada

(March, 2010)

Copyright ©Megan Lee Sierant, 2010

Abstract

The human Rad9A checkpoint protein is required for genomic stability and proper execution of the DNA damage checkpoint. Previous work has shown Rad9A to be the key member of a heterotrimeric toroidal structure known as the 911 complex, along with Hus1A and Rad1, which is similar in structure to PCNA. Recent literature suggests Rad9A is a novel oncogene, found to be upregulated in a number of cancers and high mRNA levels are thought to have a protective effect on tumour growth. This thesis describes two novel functions for the Rad9A protein. The first is as a facilitator for the nuclear translocation of the Claspin adaptor protein, required for successful Chk1‐mediated checkpoint activation. The second is as part of a novel nuclear structure containing important members of the homologous recombination

DNA repair pathway. Work described herein also confirms the existence of a Rad9A paralogue, Rad9B, that partially rescues defects associated with Rad9A‐deficiency and is expressed in both tumour and undifferentiated embryonic stem cell lines.

This work discusses these findings in the context of current literature and provides future experiments to continue investigations into the function of this vital checkpoint protein.

ii Co‐Authourship

All of the work described in this thesis was performed by the authour with the following exception: the cellular differentiation, mRNA extractions, cDNA synthesis, and qRT‐PCR reactions were performed by Nicole Archer, under the direction of the authour.

iii Acknowledgements

Firstly, I would like to thank Scott for providing the environment in which I have stumbled my way along to become the scientist I am today. The road was long and winding but I made it! Cheers!

I would also like to thank my labmates, both past and present. Thank you for endless mirth, ranting, and opportunities to procrastinate! I do not thank you for the Boney M Christmas CD, which should be outlawed.

I would also like to thank certain members of the other labs on the third floor of Botterell Hall. You know who you are. This trip has been infinitely more enjoyable with you guys, even if it took infinitely longer than it should have because of you. I forgive you guys though, no hard feelings.

There are two venues which also require my thanks: the Grad Club and the

Sports. My two favourite watering holes in this city. Grad Club, you are smelly, over‐crowded, and often the wrong temperature but without you I would be 20 pounds lighter and left without a decent place for Trivia. Sports, you are the best place to watch sports (good name!) and eat turkey sammiches even if the local wildlife also prefers you as a watering hole, they are nothing if not entertaining. I will miss both of you.

My family should also be acknowledged for both their emotional and financial support. This endeavour would not have been possible without you.

iv Thank you. I will finally be moving out of the “guest” bedroom. At 30. You must be so proud.

Finally, I would like to thank Glenn, my Rock. Things are always better when you’re around, even though you smell like nachos.

v Table of Contents

Abstract...... ii Co­Authourship ...... iii Acknowledgements...... iv Table of Contents ...... vi List of Figures...... ix List of Tables ...... xi List of Abbreviations, Symbols, and Chemical Formulas ...... xii Chapter 1: General Introduction ...... 1 1.1 The molecular basis of cancer...... 5

1.2 The Rad9A checkpoint protein ...... 9

Chapter 2: Literature Review ...... 10 2.1 Chromatin structure...... 11

2.1.1 Histone modifications...... 12 2.1.2 DNA modifications ...... 13 2.2 DNA damage detection ...... 14

2.2.1 ATM...... 14 2.2.2 ATR/ATRIP...... 18 2.2.3 DNA‐PK ...... 19 2.2.4 The 911 Complex and the Rad17/RFC clamp loading complex ...... 20 2.2.5 The MRN complex ...... 28 2.2.6 Summary of DNA damage sensors ...... 30 2.3 Checkpoint transducers ...... 30

2.3.1 ATM, ATR/ATRIP, and DNA‐PK...... 31 2.3.2 γH2AX ...... 33 2.3.3 Claspin ...... 34 2.3.4 BRCA1 and BARD1...... 35 2.3.5 TopBP1...... 37 2.3.6 Summary of DNA damage transducers ...... 38 2.4 Checkpoint effectors ...... 38

2.4.1 Chk1...... 39 2.4.2 Chk2...... 42 2.4.3 The Cdc25 Phosphatases...... 45 2.4.4 Wee1/Myt1 ...... 47 2.4.5 Cyclin/CDK complexes ...... 48 2.4.6 Summary of the DNA damage effectors ...... 50 2.5 Double­strand break repair ...... 50

vi 2.5.1 Homologous recombination...... 51 2.5.2 Non‐homologous end joining ...... 54 2.5.3 Alternative non‐homologous end joining ...... 57 2.6 Checkpoint Release ...... 58

2.7 Hypotheses and specific aims ...... 59

Chapter 3: Materials and Methods...... 61 3.1 Cell Lines and Culture Conditions...... 61

3.2 Cell Synchronization, Transfections and Treatments...... 62

3.3 Immunofluorescence and Confocal Microscopy...... 65

3.4 Immunoprecipitations and Immunoblotting ...... 68

3.5 mRNA extraction, cDNA synthesis, and qRT­PCR ...... 70

Chapter 4: The Rad9A checkpoint protein is required for nuclear localization of the Claspin adaptor protein ...... 71 4.1 Rad9A and Claspin interact constitutively ...... 71

4.1.1 The Rad9A‐Claspin interaction is not cell cycle dependent...... 71 4.1.2 Modification of the Rad9A protein does not affect the interaction with Claspin ...... 74 4.2 Rad9A affects the nuclear localization of Claspin...... 78

4.2.1 Over‐expression of a non‐nuclear form of Rad9A alters the subcellular localization of Claspin ...... 78 4.2.2 Extraction‐resistant Rad9A forms large nuclear foci with Claspin...... 85 4.2.3 Rad9A molecules are able to multimerize...... 90 4.3 Rad9A/B are responsible for Claspin localization ...... 93

4.3.1 rad9A­null mES cells express Rad9B...... 93 4.3.2 Differentiation of Rad9A‐null mES cells alters the subcellular localization of Claspin but has no affect on WT mES cells...... 96 4.3.3 Conclusions about the interaction between Rad9A and Claspin ...... 96 Chapter 5: The identification and characterization of a novel nuclear structure containing members of the homologous recombination DNA damage response pathway...... 98 5.1 The RDF closely associate with Xi...... 98

5.1.1 Rad9A closely associates with the histone variant macroH2A1...... 98 5.1.2 Rad9A colocalizes with the facultative heterochromatin marker H3trimK9..101 5.1.3 RDF‐macroH2A1 foci overlap with BRCA1 foci...... 101 5.1.4 RDF colocalize with γ‐H2AX independent of exogenous DNA damage...... 104 5.2 The RDF colocalize with members of the homologous recombination pathway...... 113

vii 5.2.1 The RDF does not colocalize with TopBP1, Claspin, or the telomeric TRF2 protein ...... 113 5.2.2 The RDF colocalizes with Mre11, Rad51, ATM, and Rad17...... 118 5.2.3 Members of the RDF are not static...... 123 5.2.4 Summary of the RDF...... 131 5.3 The RDF is altered after perturbations to the cell cycle...... 131

5.3.1 Cell synchronization reduces the number of RDF...... 132 5.3.2 Treatment of asynchronous HeLa cells with either cisplatin, pentoxifylline, or both affects foci formation...... 132 5.3.3 Conclusions about the RDF...... 135 Chapter 6: Discussion...... 136 6.1 Rad9A and Claspin ...... 136

6.2 The RDF contains members of the HR DNA repair pathway...... 141

6.3 Exploring the role of Rad9A in tumour cells...... 152

6.4 Concluding Remarks...... 153

References...... 154 Appendix A: Generation of a conditional rad9A­null cell line ...... 178 A.1 hTERT­RPE1 cells and genomic rad9A and Cre recombinase­mediated excision ...... 178

A.2 Constructs and methods...... 179

A.3 PCR confirmation strategy...... 181

A.4 Current status ...... 181

A.5 Appendix A­specific References ...... 182

viii List of Figures

1.1 Overview of checkpoint function...... 4 1.2 Progression of a cancerous lesion from normal tissues ...... 7 2.1 Schematic of the checkpoint response ...... 16

2.2 The protein sequence of the human homologue of S. pombe Rad9 ...... 25 2.3 The Chk1‐activation complex...... 41 4.1 Rad9A and Claspin interact during all phases of the cell cycle in HeLa cells..73 4.2 Mutations in the Rad9A C‐terminal tail do not affect Rad9A‐Claspin interactions regardless of the presence of DNA damage...... 77 4.3 Over‐expression of non‐nuclear Rad9A protein changes the subcellular localization of Claspin in asynchronous HeLa cells...... 80 4.4 Over‐expression of non‐nuclear Rad9A protein changes the subcellular localization of Claspin in asynchronous hTERT‐RPE1 cells...... 82 4.5 Over‐expression of non‐nuclear Rad9A protein changes the subcellular localization of Claspin in undamaged, asynchronous murine embryonic stem cells ...... 84

4.6 Extraction of soluble proteins reveals large, nuclear Rad9A‐Claspin foci in asynchronous Rad9A‐transfected HeLa cells...... 87

4.7 Extraction of soluble proteins reveals large, nuclear Rad9A‐Claspin foci in asynchronous Rad9A‐transfected hTERT‐RPE1 cells...... 89 4.8 Immunoprecipitation against epitope tagged Rad9A protein reveals the presence of endogenous Rad9A after immunoblot...... 92 4.9 Differentiation of murine embryonic stems cells alters the localization of Claspin in the absence of Rad9A which can be restored by over‐expression of wild‐type Rad9A...... 95 5.1 Rad9A‐GFP fusion protein closely associates with macroH2A1 in asynchronous HeLa cells ...... 100 5.2 Rad9A‐macroH2A1 foci and Rad9A‐H3trimK9 foci do not directly overlap in asynchronous HeLa cells ...... 103

5.3 The RDF colocalizes with the Xi and BRCA1...... 106 5.4 Rad9A colocalizes with macroH2A1 in HeLa cells but not in other cell types108

ix 5.5 The RDF colocalizes with BRCA1 in multiple cell types...... 110 5.6 The RDF and the Xi colocalize with a large γH2AX foci...... 112

5.7 Rad9A and γH2AX colocalize in a large body in multiple cell types...... 115 5.8 The RDF does not contain TopBP1, Claspin, TRF2, or CenpA ...... 117 5.9 Rad9A forms large foci with Mre11 in multiple cell types ...... 120 5.10 Rad9A forms large foci with Rad51 in multiple cell types...... 122

5.11 Rad9A forms large foci with ATM in multiple cell types ...... 125 5.12 RDF colocalize with Rad17 and γH2AX in a large body in HeLa cells...... 128 5.13 The RDF are variable structures ...... 130

5.14 The RDF disappears after attempts at synchronization via exposure to thymidine and hydroxyurea but is enriched after exposure to pentoxifylline134 6.1 Models describing the order of recruitment events at the RDF...... 148

x List of Tables

3.1 Tissue culture conditions for the different cell lines used during the course of this thesis ...... 63

3.2 Primary antibodies used in immunoblotting and immunofluorescence experiments ...... 66

3.3 Alexa secondary antibodies used in immunofluorescence experiments ...... 67 5.1 Colocalization between members of the HR pathway and Rad9A ...... 126

xi List of Abbreviations, Symbols, and Chemical Formulas

911 Rad9A‐Hus1‐Rad1 A alanine ATM ataxia telangiectasia mutated

ATR atm and rad3‐related ATRIP ATR interacting protein BASC BRCA1‐associated surveillance complex BRCT BRCA1 carboxy terminus

BrdU 5‐bromo‐2‐deoxyuridine CDK cyclin‐dependent kinase coIP coimmunoprecipitation

CP cisplatin D‐MEM Dulbecco’s Modified Eagle media DAPI 4',6‐diamidino‐2‐phenylindole DDR DNA damage response

DNA deoxyribonucleic acid DNA‐PK DNA‐dependent protein kinase DSB(s) double‐strand break(s)

FBS fetal bovine serum GFP green fluorescent protein Gy gray (1 Gy = 107.2 R) HR homologous recombination

HRP horseradish peroxidase hTERT human telomerase reverse transcriptase

HU hydroxyurea

xii Hus1 hydroxyurea sensitivity gene 1 IB immunoblot

IF immunofluorescence IP immunoprecipitation IR ionizing radiation LIF leukemia inhibitory factor mES cell murine embryonic stem cell Mre11 meiotic recombination 11 MRN Mre11‐Rad50‐NBS1 complex

MW molecular weight NBS Nijmegen breakage syndrome NES nuclear export signal NHEJ non‐homologous end joining

NLS nuclear localization sequence PAGE polyacrylamide gel electrophoresis PARP poly(ADP‐ribose) polymerase

PBS phosphate buffered saline PCNA proliferating cellular nuclear antigen PCR polymerase chain reaction PFA paraformaldehyde

PIKK phosphatidylinositol‐3‐OH kinase related kinase PTX pentoxifylline RA all­trans retinoic acid

RDF Rad9A‐dense foci RFC replication factor C

RNA ribonucleic acid

xiii RPA replication protein A RPE retinal pigmented epithelium

RT room temperature S serine ssDNA single‐strand DNA T threonine

TopBP1 topoisomerase IIβ binding protein 1 UV ultraviolet WCE whole cell extract

Xi inactive X chromosome XiST Xi‐specific transcript XLF XRCC4‐like factor XP Xeroderma Pigmentosum

xiv Chapter 1: General Introduction

A cell is a complex structure capable of startling feats of biological engineering every cell cycle, most notably the replication of its DNA. DNA is a biochemical code for producing proteins, comprised of a series of building blocks known as nucleotides. A nucleotide is made up of three parts: a simple sugar, a phosphate, and one of four possible bases (adenine [A], guanine [G], cytosine [C], or thymine [T]). It is these DNA molecules that control the functions occurring inside a single cell, and it is groups of cells that define the architecture of an entire organism.

A human cell contains approximately one meter of DNA, which is coiled, compacted, and supercoiled into chromatids in order to fit inside the nucleus, a structure roughly 5,000,000 times smaller. The chromatids are constantly being rearranged, wound and unwound as the cellular processes necessary for cell survival require access to the genetic information stored in the sequence of the DNA molecules; therefore, the detection and correction of any mistakes in the DNA sequences are vital for the health of replicating cells and, in turn, for the entire organism.

In order to maintain healthy cellular function, it is necessary to ensure the

DNA inside the cell is undamaged and that gene sequences resulting from the combinations of the bases remains relatively unchanged from generation to generation. Changes in the DNA sequence, more commonly known as mutations, can be either base substitutions, insertions, or deletions. Furthermore, mutations can be silent, producing no change in the corresponding protein, or result in coding for a different protein, or in a truncated version of the protein. This can lead to

1 aberrant cell growth and the development of diseases, such as cancer, or can trigger a process known as apoptosis in which the cell undergoes programmed cell death.

This can also produce changes to an organism as mutations occur over time and can result in adaption to changing conditions, such as environmental stresses, and is the mechanism by which organisms evolve. However, the cell has evolved a number of quality control mechanisms to ensure the number of mutations passed on is minimized, these are known as checkpoints [1].

The checkpoint was first defined in 1989 by Hartwell and Weinert as a mechanism for ensuring the completion of one stage of the cell cycle before the onset of the next stage [1]. Continued work on the nature of checkpoints reveals that the archetypal checkpoint can be thought of as a biochemical pathway consisting of three main components: a sensor, a transducer, and an effector. The sensor is responsible for detecting aberrations during cellular processes and relaying this signal to the transducer. The transducer amplifies the signal by activating one or more effectors. The effector then initiates another signaling cascade to transiently halt the cell cycle until the error has been repaired, after which the cell is released to continue through the cell cycle or, if the damage is catastrophic, to initiate apoptosis (Figure 1.1). DNA checkpoints are particularly important during DNA replication as a method of ensuring the daughter cell receives a complete copy of the parent cell’s genome and for preventing prolonged cell cycle arrest associated with compromised replication fork procession, which can result in premature cellular division. Checkpoints are also required to detect and correct

DNA that is damaged during normal cellular processes (referred to as endogenous

2

Figure 1.1 ­ Overview of checkpoint function. A schematic diagram depicting the order of events after damage, represented by the lightening bolt, is detected and triggers checkpoint‐mediated arrest of the cell cycle. The transducer is responsible for amplifying and relaying the signal to the effector, which facilitates cellular arrest. After resolution of the damage, the cell is released back into the cell cycle.

3

Normal Cellular Progression

Resolution Sensors • Continuation • Detect of the cell cycle aberration

Effector Transducer • Arrests cell • Relays signal cycle

Apoptosis • Programmed cell death

4 damage). One example being breaks caused by chromosome separation during mitosis or by replication fork stalling. DNA can also be damaged by exogenous sources, such as oxygen free radicals, ultraviolet light (UV) or gamma‐irradiation

(IR), which react with the DNA molecule to arrest DNA replication or induce breakages, and are also detected by checkpoints. Aberrant checkpoint function, or the ability to bypass a particular checkpoint, is a common trait among tumour cells and is believed to be a key step during the development of cancer.

1.1 The molecular basis of cancer

Cancer is an umbrella term given to wide range of diseases which arise from cells in any tissue in the body. The progression from a normal cell to a cancerous cell is termed “transformation” and begins when a normal cell undergoes mutations which, during subsequent rounds of replication, result in a loss of replicative control. This allows the daughter cells to evolve into a population of dysplastic cells with only some resemblance to the tissue of origin. This population of cells may undergo apoptosis, thus removing themselves from the tissue, or may continue the transformation process becoming a neoplastic tissue. The term “neoplastic” refers to a “new” population of cells with little resemblance to the parent tissue and exhibits novel behaviors, such as by having a faster growth rate or different nutritional requirements. A neoplasm may be benign or malignant and often develops into a cancerous lesion. A summary of the process of transformation is depicted in Figure 1.2. Benign lesions are usually of little threat to the host

5

Figure 1.2 – Flowchart depicting the progression of a cancerous lesion from normal tissues. Cancerous tissue progresses from normal tissue through the dysplastic stage. During this stage cells have some resemblance to the tissue of origin and retain the ability to undergo programmed cell death, known as apoptosis. These tissues may also either regress back to a normal tissue, via apoptosis of the affected cells, or progress into neoplastic tissue by expansion of the dysplastic population. A neoplasm can be either benign or malignant promoting either survival or death of the host organism, respectively.

6 Malignant Death Neoplastic Normal Dysplatic Cells Benign Survival Cells Cells Apoptosis

7 organism whereas a malignant lesion has the ability to spread throughout the body, a process known as metastasis, and can result in the death of the host. All this can result from a simple mistake during DNA replication that is then passed on to subsequent generations of daughter cells.

The mechanism behind the uncontrolled replication, a hallmark of the neoplastic cell, is a series of genetic mutations resulting in the deregulation of two key cellular functions: the first being that the process by which a cell can undergo apoptosis, or commit cellular suicide, is deactivated and the second being that DNA replication/damage checkpoints are also rendered inactive [2]. Together, these two changes result in a population of cells that are unable to remove themselves from the surrounding tissue and are able to continue to proliferate in the presence of increasing genomic instability. Genes that are capable of controlling cell growth are known as tumour suppressors and are often inactivated via a series of mutations in neoplastic cells. These genes are also frequently members of the checkpoint pathway, involved in the transient arrest of the cell cycle after DNA damage has been detected. In contrast, genes which are able in increase cellular proliferation are often over‐expressed, or “turned on”, and are referred to as oncogenes. By inactivating tumour suppressors and/or activating oncogenes a cell becomes able to ignore its original programming, replicating continuously, which can eventually result in tumour formation. In fact, Knudson’s work first noted that two mutations inactivating both copies of the retinoblastoma gene were required to initiate the disease, leading to the theory that as few as two mutations are required to initiate cancer [3]. This highlights the importance in understanding the mechanisms behind

8 DNA damage recognition as well as checkpoint function and cellular replication, as these are all affected in cancerous cells.

1.2 The Rad9A checkpoint protein

A checkpoint protein of significant importance is Rad9A, currently believed to be responsible for sensing DNA damage and initiating the DNA damage response

(DDR). Rad9A exists in a complex with two other proteins: Hus1A and Rad1, which are collectively known as the 911 complex and all three proteins are similar in sequence and shape [4, 5]. The 911 complex is structurally analogous to the proliferating cellular nuclear antigen (PCNA) DNA replication protein, a sliding clamp responsible for increasing processivity of the DNA polymerases during DNA replication [6‐9]. Unique to Rad9A is a large C‐terminal tail, which possesses regulatory sequences important for the function of the 911 complex [10‐13].

Currently, the 911 complex is believed to link the checkpoint signaling cascade and

DNA repair machinery. This is accomplished through signal transduction transmitted by post‐translational modifications of the Rad9A C‐terminal tail [11‐

15]. To provide a complete picture of the DDR, the next section provides an overview of the DNA damage checkpoint response and DNA repair starting with basic DNA structure through the three components of the checkpoint response and ending with a brief description of the mechanisms involved in checkpoint release.

9 Chapter 2: Literature Review

Checkpoints are biochemical pathways responsible for detecting and initiating the resolution of DNA damage acquired during cellular processes

(endogenous) and from environmental sources (exogenous) [16]. Due to the importance of genomic integrity on the overall survival of the organism, it is vital to ensure the DNA is under constant surveillance from DNA damaging agents; this is the primary role of the DNA damage and DNA replication checkpoints, which are the focus of the work described in this thesis.

In all phases of the cell cycle, the checkpoint culminates with cell cycle arrest via inhibition of the cyclin/CDK holoenzyme by the checkpoint effector [1]. This complex is required for progression from one phase of the cell cycle to the next and by initiating the inactivation of this complex, the checkpoint machinery is able to transiently arrest the cell cycle providing time for DNA repair [1, 17, 18]. After repair is successfully completed, the cyclin/CDK complex is reactivated, resulting in resumption of the cell cycle [19]. In cases in which the DNA damage cannot be successfully resolved, the checkpoint machinery is able to initiate apoptosis, destroying the cell before the damaged DNA can be passed onto the next generation of cells [20].

Traditionally, the checkpoint response was divided into two pathways, each responsible for detecting a particular type of DNA damage signal: the single‐strand

DNA (ssDNA) checkpoint pathway and the double‐strand break (DSB) pathway. All

DNA damage is processed into one of these two structures during checkpoint

10 activation [21], and by setting up pathways dedicated to the resolution of these structures, the cell is able to promote repair. Initially it was believed that there was very little overlap between these two branches, however evidence for extensive interactions between these pathways is increasing.

This chapter will describe the biochemical pathways utilized by the eukaryotic checkpoint system as well as provide background information on different methods of DNA repair during the late S and G2 phases of the cell cycle. It will begin by discussing chromatin structure then an introduction of the three components of the archetypal checkpoint, the sensor, transducer, and effector, followed by a brief review of the mechanism behind cell cycle arrest. An overview of three of the DSB repair pathways will be addressed as well: homologous recombination, non‐homologous end joining and alternative non‐homologous end joining. Particular emphasis will be on the 911 complex, Rad9A, and their roles in the DDR.

2.1 Chromatin structure

Each of the 46 chromosomes in a human cell consists of a single molecule of

DNA. In order to minimize tangling of these molecules, also known as catenation, and in order to have active genes available while limiting access to inactive genes, the DNA molecule undergoes a series of compactions. Compaction begins at the nucleosome level, in which the DNA strand is wound around an octamer of histone proteins [22]. There are four families of histone protein: H1, H2, H3, and H4, the H2

11 family can be further divided into the H2A and H2B subfamilies. A histone complex, the core of the nucleosome, consists of two H2A‐H2B dimers and an H3‐H4 tetramer

[23, 24]. This forms a barrel‐like structure around which 146 base‐pairs of DNA is wrapped, forming a complete nucleosome [22]. A commonly used descriptive analogy for the nucleosomes is the “bead on a string” image, in which nucleosomes form beads along a strand of DNA. To accomplish higher order structures, the H1 histone, which both secures the DNA in the nucleosome while also providing a link for adjacent nucleosomes, coil into structures known as “the 30 nm fiber”, which further compact, with the help of scaffold proteins, to form chromosomes (reviewed in Horn and Peterson [25]). Areas of the genome undergoing active transcription exhibit a less dense packing pattern and are referred to as euchromatin. Areas that are densely packed and silenced are known as heterochromatin. Packing density is a method of controlling access to genetic information and corresponds to transcriptional activity.

2.1.1 Histone modifications

The core histone proteins, H2A, H2B, H3, and H4, contain N‐terminal regions that are excluded from the nucleosome. Modifications to these tails can affect higher order chromatin structure as well as act as flags, regulating DNA‐interacting proteins. Phosphorylation of the H2A histone subtype, discussed below, is a well‐ known marker of DNA damage, acting like a flag for the DNA repair machinery [26].

Histone methylation is often indicative of regions of the genome currently silenced;

12 for example, the inactive X (Xi) chromosome is heavily enriched for trimethylation lysine 9 on the H3 subunit (H3trimK9) [27]. In contrast, histone acetylation is often associated with areas undergoing active transcription and correspond to a more relaxed chromosome structure [28‐30]. However, recent work has revealed that the combination of acetylation and methylation modifications combine in a complex manner to regulate chromatin structure and transcriptional activity and is currently the basis for the field of epigenetics [31].

As previously mentioned, histone modifications can act as flags for DNA‐ interacting proteins. The H3trimK9 modification found on the Xi chromosome acts as a loading dock for the heterochromatin protein 1 (HP1) family of proteins [32,

33]. The HP1 family of proteins aid in silencing and the packaging of the Xi body

[34, 35], and have recently been implicated in a form of heterochromatin commonly found in cancer cells [36]. Additionally, the HP1 proteins are required for repairing defects in the epigenetic code of acetylation and methylation on histone proteins

[37].

2.1.2 DNA modifications

In addition to histone modifications, DNA is also able to undergo modifications that affect how tightly it binds to the core histones. Methylation on the cytosine of CpG dinucleotides results in decreased transcription [38].

Furthermore, methylated DNA acts to recruit histone deacetylases, which further repress transcription by increasing the affinity between DNA and the core histone

13 proteins, limiting the access of the transcriptional machinery to promoter regions

[39].

2.2 DNA damage detection

DNA damage results in physical changes to the structure of the DNA strand, it is these changes that are detected by the DNA damage sensors during the checkpoint response. For example, after exposure to UV light, adjacent pyrimidine bases crosslink producing pyrimidine dimers. The resulting kink in the DNA backbone disrupts DNA synthesis, stalling replication forks, which results in ssDNA breaks. There is no single protein required for damage detection; instead, damage detection depends on multiple proteins and/or protein complexes and varies by the type of DNA damage sustained. The list of sensor proteins is under continual revision as new research is published and the mechanisms of this process are revealed. Nonetheless, several key players with defined roles, including several phosphatidylinositol‐3‐OH kinase related kinases (PIKKs), and the 911 and MRN complexes, are described here. A schematic overview is presented in Figure 2.1.

2.2.1 ATM

Ataxia telangiectasia is an autosomal recessive disorder typically affecting multiple organ systems and characterized by cerebellar ataxia, oculocutaneous telangiectasia, and lymphoid tumours [40]. Mice in which the atm gene has

14

Figure 2.1 – Schematic of the checkpoint response. Diagram depicting the proteins involved in checkpoint initiation for the non‐homologous end joining, ssDNA damage response, and homologous recombination pathways with emphasis on the proteins discussed in Chapter 2. The non‐homologous end joining pathway begins by detection of the DSB by DNA‐PK. In the ssDNA damage response, detection is accomplished by the 911 complex, Rad17/RFC, Claspin, TopBP1 and ATR/ATRIP. The MRN complex acts as the sensor for the homologous recombination pathway. From there the DNA damage signal is relayed to the appropriate transducers and effectors. Cell cycle arrest occurs at the G2/M boundary via inhibition of the cyclin B/Cdc2 complex.

15 been selectively disrupted display a phenotype mirroring the symptoms observed in human ataxia telangiectasia patients [41]. These mice were found to be sensitive to small doses (0.5 Gy) of IR and exhibit numerous chromosomal abnormalities suggesting that ATM is required for the G1 checkpoint response to IR and is also involved in maintaining genomic integrity [41].

Ataxia telangiectasia‐mutated (ATM) is the protein absent in patients of the disease Ataxia telangiectasia and is a kinase in the family of PIKKs, with a mass of approximately 370 kDa [42]. ATM exists as a homodimer in its inactivated form but after DNA damage ATM is able to undergo autophosphorylation on serine (S)1981, which coincides with dimer dissociation [43]. Bakkenist and Kastan hypothesized that changes in the structure of DNA, resulting from damage, disrupted the ATM homodimers, and were able to initiate the DDR [43]. In this way it is easy to assign

ATM the function of a DNA damage sensor; however, it is also known that ATM is responsible for activating and amplifying the checkpoint response suggesting it also functions as a DNA transducer, which is addressed in the next section. Once phosphorylated, ATM activates several downstream members of the DNA checkpoint pathway such as Chk1 [44], Chk2 [45], Rad9A [11], and BRCA1 [46], as well as providing signaling the location of DNA damage via H2AX phosphorylation

[47].

As previously mentioned, the DDR is roughly divided into two arms: the DSB repair pathway and the ssDNA damage pathway; ATM is primarily involved in the

17 DSB repair pathway [48, 49]. In fact, Suzuki et al. demonstrated that ATM homodimers are constitutively bound to DNA and this binding increases as a result of irradiation, which is consistent with regulation of the DSB arm of the DDR [50].

Furthermore, it seems the amount of ATM protein activated is proportional to the number of DSBs present, reinforcing the role of ATM as a sensor [51]. Together, these data provide a strong argument for the role of ATM as a sensor.

2.2.2 ATR/ATRIP

Atm and rad3‐related kinase (ATR) is also a member of the PIKK family of kinases responsible for initiating the DDR. The ATR‐mediated checkpoint preferentially responds to ssDNA signals [52]. However, during the DDR, crossover between the ATM‐mediated and the ATR‐mediated repair pathways occurs ensuring checkpoint activation [44, 53]. ATR exists in a heterodimer with the ATR‐ interacting protein (ATRIP) and this interaction is required for ATR kinase activity

[54]. This heterodimerization is also conserved in eukaryotes signifying its importance in proper ATR function [54‐56].

In contrast to atm‐/‐ mice, ATR‐deficient mice are nonviable, unable to progress past day 7.5 post‐conception [57]. The embryos show evidence of elevated levels of chromosomal breakages, and increased levels of apoptosis [57]. A conditional mouse line, in which atr can be excised using the Cre/lox system, showed that ATR was required for the checkpoint response during G2 and after exposure to IR [58]. Further analysis of ATR function reveals an important role in

18 moderating the rate of replication fork progression during S phase, along with Chk1 and Claspin, and by assisting Chk1 in regulating origin density during DNA replication [59‐61]. One group has also shown that ATR is loaded onto chromatin during normal, unperturbed S phase, possibly to monitor DNA replication [62].

Additionally, ATR mutants lacking kinase activity show sensitivity to both UV and IR as well as show increased levels of cell death and abrogated checkpoint function

[63]. As previously mentioned, ATR acts upstream of Chk1, activating the effector kinase during the DDR [52]. Yoshioka et al. used a human cell‐free system to show that ATR is recruited to ssDNA damage before both the PCNA clamp loading replication factor C (RFC) complex and the mismatch repair machinery, supporting the role of ATR as a DNA damage sensor [64]. Combined, these data prove a vital role for ATR in DNA replication and repair.

2.2.3 DNA‐PK

DNA‐PK was originally described as an activator of the nonhomologous end joining (NHEJ) DSB repair pathway (discussed below) and as a moderator of V(D)J recombination, the process of creating antibody diversity, during the maturation of

B and T cells [65‐67]. DNA‐PKcs is related to ATM and a member of the PIKK family of kinases [68, 69]. The exact mechanism of DNA‐PK activation by DNA remains unknown but data suggests the orientation of strand termini may play a key role in stimulating DNA‐PK activity, with the 3’ strand required for annealing and the 5’ strand required for DNA‐PK activtion [70].

19 The DNA‐PK holoenzyme is a complex consisting of the 350 kDa DNA‐PK catalytic subunit (cs) and the Ku heterodimer, comprised of the Ku70 and Ku80 proteins [71]. DNA‐PKcs interacts with the Ku heterodimer via the C‐terminal domain of Ku80 and, since DNA‐PKcs is unable to bind DNA, this interaction is required for the association of the holoenzyme with DNA, and activation as previously mentioned [71‐73]. The Ku heterodimer binds preferentially to dsDNA ends, conveniently locating DNA‐PKcs at the site of DSBs conferring DNA damage sensor function to the DNA‐PK holoenzyme [74].

2.2.4 The 911 Complex and the Rad17/RFC clamp loading complex

Members of the 911 complex, Rad9A, Hus1A, and Rad1, interact to form a toroidal heterotrimer capable of encircling the DNA strand [4, 5, 7, 75, 76]. All three proteins are required for 911 complex formation, interacting in a head‐to‐tail manner, and this complex is structurally similar to the homotrimeric PCNA required for DNA replication [6, 7, 77‐82]. The similarities between 911 and PCNA extend to

DNA loading as well: PCNA is loaded onto DNA by the RFC complex whereas 911 is loaded by the Rad17/RFC complex in which Rad17 replaces the large subunit, RFC1, existing in a complex with proteins RFC2‐5 [75]. The exact function of the 911 complex is not yet well understood though it is known to be required for DNA replication, as well as the DNA damage and the DNA replication checkpoints [13, 14,

86‐90]. Older hypotheses have the 911 complex using its sliding clamp abilities to scan the DNA molecule for aberrations, signaling to the rest of the DNA checkpoint

20 proteins, and acting as a loading platform for the DNA repair machinery. However, there is also evidence suggesting that Rad17 and ATR are required to localize the

911 complex to damaged DNA which challenges that hypothesis [91]. Recent crystallization work suggests 911 does act as a sensor but that it does not scan along the DNA molecule; instead, there is the possibility that the 911 complex may interact with specific bases during the DDR [81]. Ample evidence supporting the

911 complex as a regulator of the ssDNA machinery, able to interact with several repair proteins such as FEN1 and AP endonuclease 1 [92], DNA ligase I [93, 94], one of the glycosylases [95], and DNA polymerases associated with DNA repair [92, 96].

There is also evidence to support a role for the interaction between the 911 complex and the histone deacetylases 1 (HDAC1) during the G2/M checkpoint suggesting the

911 complex may also be involved in the maintenance or silencing of chromatin following DNA damage [77].

Of the three members of the 911 complex, Rad9A is arguably the key component due to the presence of a nuclear localization sequence (NLS) [97], a consensus region for interacting with the large subunit of the RPA complex [98], and an interacting motif for the apoptotic control proteins Bcl‐2/Bcl‐XL [10]. These motifs have not yet been identified in Hus1A and Rad1. In addition, there are extensive phosphorylational events on the Rad9A C‐terminal tail, an extra 120 amino acids sharing no homology to either Hus1A, Rad1, or PCNA [4, 13, 89].

Rad9A was first discovered in Schizosaccharomyces pombe as a plasmid able to restore resistance to UV and IR in the rad9­192 radiosensitive strain [99, 100]. A

21 structural homologue was discovered in a human infant brain library, which consisted of 391 amino acids with an approximate molecular weight (MW) of 42 kDa. This homologue was able to restore resistance to both IR and HU as well as repair some checkpoint defects when expressed in a rad9::ura4+ S. pombe mutant

[101]. A murine homologue, Rad9A, was also identified that was able to partially restore function in the rad9::ura4+ S. pombe mutant. Ablation of the murine homologue results in embryonic lethality at E9.5, a key point during embryogenesis, as well as severe defects in chromosomal stability and checkpoint function the rad9‐ null murine embryonic stem cells (mES) [87, 102]. During embryogenesis, day 9.5 post‐conception corresponds to the first major wave of cellular differentiation, in which the progenitors for the major organ systems are established (Manipulating the Mouse Embryo: A Laboratory Manual, 2003 Ed.).

Rad9A has been found to contain a number of phosphorylation sites in a number of different contexts: constitutive, damage‐dependent and cell‐cycle dependent [4, 11, 13, 89]. Some of these phosphorylation sites have also been shown to have interdependencies and regulate interactions with other proteins, such as the topoisomerase IIβ binding protein 1 (TopBP1) [11‐13]. Five constitutive phosphorylation sites have been identified thus far: S277, S328, S336, T355 and

S387 [11, 13]. A DNA damage‐dependent phosphorylation site was identified as

S272 and is modified by ATM following DNA damage by IR, UV and HU [11].

Another damage‐dependent phosphorylation site, referred to as the σ‐site in the literature, exists and requires prior phosphorylation at S387 [13]. The final phosphorylation site to be identified and confirmed is the T292 cell‐cycle dependent

22 site. Prior phosphorylation at the S277 residue is required in order for phosphorylation at the T292 residue to occur and is restricted to the M phase of the cell cycle [11]. Three additional phosphorylation sites, S341, S375, and S380 were identified using mass spectrometry and Edman degradation but these remain to be confirmed by additional studies [89]. Phosphorylation of the C‐terminal tail does not regulate the formation of the 911 complex or the interaction with the

Rad17/RFC clamp loader [4]. The features of the Rad9A protein are summarized in

Figure 2.2.

Rad9B is a recently discovered paralogue of Rad9A, expressed primarily in the testes that is able to interact with Rad1, Hus1A, and Rad17 [103, 104].

Paralogues have yet to be discovered in yeast and could be unique to metazoans.

Both groups also reported the presence of Rad9B mRNA in tumour cells, underlining the importance of Rad9 for cell survival [103, 104]. During the course of this thesis, it was also revealed that undifferentiated mES cells also express Rad9B and that levels decrease after exposure to retinoic acid (RA) [105], a potent initiator of differentiation. Data also shows that Rad9B is not as heavily phosphorylated as

Rad9A nor did the C‐terminal tails share significant homology, suggesting differing roles in checkpoint function [103]. More work is required in this area to determine the role of Rad9B and the reason for its tissue specificity.

Rad1 and Hus1A, the other members of the 911 complex, are also phosphorylated but these events do not regulate the formation, activity, or interactions of the 911 complex [76, 79]. However, recent evidence shows that

23

Figure 2.2 – The protein sequence of the human homologue of S. pombe Rad9. Rad9A is 391 amino acids long, containing a BH3 domain (purple) near the N‐terminus [97], an NLS (pink) near the C‐terminus [10], the CRD required for interaction with RPA70 (yellow) [98], and shares homology with PCNA (green) [77]. Residues corresponding to the C‐terminal tail are in black. Phosphorylation sites identified by the Davey lab are in larger type and are damage‐dependent (red), constitutive (blue), and cell cycle‐dependent (orange). Phosphorylation sites unconfirmed by the Davey lab, mapped by the Karnitz lab are highlighted in yellow [89], while the Y28 site, identified by the Kufe lab and also unconfirmed, is underlined [106]. PubMed accession number NP_004575.

24

1 MKCLVTGGNV KVLGKAVHSL SRIGDELYLE PLEDGLSLRT VNSSRSAYAC

51 FLFAPLFFQQ YQAATPGQDL LRCKILMKSF LSVFRSLAML EKTVEKCCIS

101 LNGRSSRLVV QLHCKFGVRK THNLSFQDCE SLQAVFDPAS CPHMLRAPAR

151 VLGEAVLPFS PALAEVTLGI GRGRRVILRS YHEEEADSTA KAMVTEMCLG

201 EEDFQQLQAQ EGVAITFCLK EFRGLLSFAE SANLNLSIHF DAPGRPAIFT

251 IKDSLLDGHF VLATLSDTDS HSQDLGSPER HQPVPQLQAH STPHPDDFAN

301 DDIDSYMIAM ETTIGNEGSR VLPSISLSPG PQPPKSPGPH SEEEDEAEPS

351 TVPGTPPPKK FRSLFFGSIL APVRSPQGPS PVLAEDSEGE G

25 ubiquitination of Rad1 may regulate the interaction between the 911 complex and

Rad17/RFC [81]. This is confirmed in the crystallization data by the alignment of

Rad1 and Rad17 during loading [81]. Depletion of Rad1 leads to disruption in ATR‐ mediated signaling, but not ATM‐mediated signaling, and disruption of the 911 sliding clamp as well as resulting in a radio‐resistant DNA synthesis phenotype

[107]. This phenotype occurs when cells continue DNA synthesis after exposure to radiation, a sign of compromised checkpoint function [107]. Deletion of hus1A in mice leads to embryonic lethality which can be rescued by co‐expression in a p21‐/‐ background, which allows for bypass of the Hus1A‐independent checkpoint event triggering apoptosis in the Hus1A‐null mice [108]. These compound mice exhibit defects in genomic stability and sensitivity to genotoxins such as UV and HU; interestingly, they show only moderate increased sensitivity to IR [108]. The hus1A+/‐ single mutants displayed widespread apoptosis and an increase in chromosomal abnormalities, which is similar to the rad9A‐/‐ mES cells and consistent with compromised checkpoint function [87, 108]. There is also evidence derived from the chicken B lymphocyte model showing the requirement for Hus1A in the resolution of DSB following drug‐induced replication fork collapse and resulting from exposure to IR [109, 110]. Interestingly, these same works show that

Chk1 activation does not require Hus1A, as it was unaffected in Hus1A‐null cells

[109, 110]. These depletion studies reveal the importance of the components of the

911 complex in genomic maintenance, proper checkpoint function as well as embryonic development.

26 Like Rad9A, Hus1A was also found to have a paralogue, Hus1B, however, conflicting data exists on whether Hus1B interacts with Rad9A. Hang et al. report that Hus1B interacts with Rad1 but not Rad9A whereas Dufault et al. reported

Hus1B was able to complete 911 complexes with both Rad9A and Rad9B, creating several possible versions of the 911 complex [103, 111]. It was also reported that

Hus1B shared similar tissue‐specific expression as Rad9B, showing high level expression in the testes and low level expression in other tissues [111]. Further research is needed before definite conclusions about the roles of these paralogues during genomic maintenance can be made.

There is a growing body of data concerning the role of Rad9A and the 911 complex in tumour biology. Higher levels of Rad9A protein have been found in a number of non‐small cell lung cancer tumour samples. In addition, a single nucleotide polymorphism in the Rad9A coding sequence, changing histidine 239 to an arginine, is associated in 16 % of cases with the development of lung adenocarcinoma [112, 113]. A recent paper exploring the incidence of these polymorphisms in families susceptible to breast cancer identified a locus in the

11q13 region, a region corresponding to several genes, notably Rad9A, and suggested that Rad9A may be altered in breast cancer as well [114]. Amplification of this same region, with confirmed increases in Rad9A mRNA levels, was correlated with increased tumour size in breast cancer patients, further confirming the link between Rad9A expression and breast cancer [115]. Targeted deletion of Rad9A in the skin of mice exposed to the carcinogen 7,12‐dimethylbenzanthracene, commonly used to induce skin lesions, was associated with higher incidences of

27 tumours, analysis of these skin tumours revealed a high level of DNA DSBs cementing the role for Rad9A and 911 in the resolution of DSBs and tumour development [116]. Based on the data presented here, low levels of Rad9A protein are associated with genomic instability whereas higher levels of Rad9A are associated with cell survival in tumour samples demonstrating an important role for

Rad9A, and the 911 complex, in maintaining cellular integrity.

2.2.5 The MRN complex

The MRN complex is composed of meiotic recombination 11 (Mre11), Rad50, and the product of the Nijmegen breakage syndrome (NBS) gene Nbs1, and is a key player in initiating repair of DSBs by both the HR and NHEJ pathways as well as in meiosis, telomere maintenance, and sister chromatid association [117‐128]. Each member of the MRN complex plays a different role in the resolution of DSBs. Briefly,

Rad50 acts as a dimer to bind the free DNA ends, Mre11 resects the ends via endo/exonuclease activity, ensuring bluntness, before ligation by DNA ligase IV, all of which is moderated by Nbs1. Both AT‐like disorder and NBS have been attributed to decreased or abolished expression of members of the MRN complex, specifically Mre11 and Nbs1 respectively [129‐132]. The MRN complex has also been shown to have sensor functions upstream of ATM by activating ATM during the DDR [133]. MRN is also required for ATR‐mediated activation of RPA2, which coats ssDNA during the checkpoint response [134].

28 Mre11 was originally identified in Saccharomyces cerevisiae as a mutant unable to undergo meiosis during sporulation, sensitive to the genotoxin methyl methanesulfonate, which triggers the ssDNA checkpoint pathway, and is conserved in humans and S. pombe [135‐137]. As mentioned above, mutations in the Mre11 protein result in ATLD and complete ablation results in embryonic lethality [131,

138]. Mre11 possesses DNA binding activity as well as 5’‐3’ and 3’‐5’ exonuclease activity, which is required for resolution of DSBs [139‐141]. Mre11 binds DNA free ends and recent work has revealed that it acts as a dimer to facilitate DNA‐end bridging during DNA repair [142].

The homodimer Rad50 contains two domains essential for function: a globular domain for interacting with DNA and Mre11 at DNA ends, and a zinc‐hook domain for tethering the DNA molecules together [119, 143‐147]. Elimination of this hook mirrors depletion of Rad50 characterized by increased sensitivity to genotoxins as well as accumulation of cells in anaphase [148]. Deletion of Rad50 in mice is embryonic lethal and work on hypomorphic alleles (rad50S/S) shows growth defects, due to decreased cellular proliferation, and cancer predisposition [149]. mre11‐null/rad50‐null Drosophilia melanogaster show chromosomal breaks and telomeric fusions, illustrating the role of Rad50 (and subsequently Mre11) in the maintenance of genomic integrity providing an explanation for the accumulation of cells in anaphase [150].

Nbs1, or Nibrin, is the final component of the MRN complex, and is ablated in

Nijmegen Breakage Syndrome [132]. Nbs1 interacts with the C‐terminal tail of

29 Mre11 and plays a role in initiating apoptosis after DNA damage in both ATM‐ dependent and –independent pathways [151‐153]. Localization of the p53‐ moderator Mdm2 after IR is dependent on Nbs1 and provides another mechanism for regulation of post‐IR apoptosis via the p53 pathway [154]. However, since Nbs1, but not Mre11 or Rad50, is dispensable for embryonic development, it suggests that the function of Nbs1 is redundant in some manner. The three proteins of the MRN complex, with their different functions, combine to create an effective sensor of DNA

DSBs capable of both recognizing DNA damage and initiating repair.

2.2.6 Summary of DNA damage sensors

Sensing DNA damage is the first step during the checkpoint response. Early sensors such as ATM, ATR/ATRIP, and MRN detect aberrations in DNA structure by mechanisms that are not fully understood. The 911 complex is slowly being revealed to be either a late‐acting sensor or an early transducer, only additional research will clarify its proper function. The common trait to all of these proteins is that they all form close associations with the DNA molecule and can transmit the

DNA damage signal down the checkpoint signaling cascade.

2.3 Checkpoint transducers

Checkpoint transducers relay and amplify the DNA damage signal to the effectors. Initially it was believed that the sensor and transducer categories of checkpoint protein were separate but it is now known that many of the proteins

30 involved in the initial detection of DNA damage also possess transducer function. In this way, there is much overlap between these two categories with many sensor proteins also possessing transducer abilities.

2.3.1 ATM, ATR/ATRIP, and DNA‐PK

As previously mentioned, the PIKK family of proteins play a vital role in the checkpoint cascade as sensors of DNA damage. However, these proteins also have transducer functions due to the number of downstream checkpoint proteins requiring ATM, ATR, and/or DNA‐PK for activation, and by the ability of these proteins to amplify the DNA damage signal [155].

ATM is capable of activating downstream effectors, such as the checkpoint kinases Chk1 and Chk2, confirming its transducer functions [44, 45]. These functions are not restricted to the checkpoint kinases since ATM also activates other important proteins such as the clamp loader complex Rad17/RFC [156], Rad9A [11],

BRCA1 [157], and ATR [158] during the DSB response pathway. Further confirming the role of ATM as a transducer, ATM is activated by the C‐terminal of Nbs1 of the

MRN sensor complex after DSBs [133, 159‐161]. The creation of small oligonucleotides by MRN‐mediated end processing also stimulates ATM activity and there is also a growing line of evidence suggesting ATM regulates ATR activation after exposure to IR via TopBP1 [158, 162‐164]. These data combine to provide evidence for the transducer side of ATM.

31 In addition to its role as a checkpoint sensor, ATR also has transducer functions evidenced by the fact that TopBP1 is capable of activating ATR/ATRIP along with the 911 complex in response to checkpoint activation [165‐167]. The

MRN complex has also been shown to activate ATR, requiring ATM and the MRN complex [134, 168]. New research shows that the circadian rhythm protein CLK2 is also required for proper ATR activation [169]. In addition, as previously mentioned,

ATR/ATRIP is required for mediating the Chk1 signaling pathway via phosphorylation on S317 and S345 in response to checkpoint activation [52]. ATR is also required for successful activation of Rad17/RFC, the clamp‐loader complex for the 911 complex implying ATR is an important regulator of 911‐mediated repair

[75, 156, 170]. By being both the receiver and possessing the capability to relay the

DDR signal, ATR demonstrates transducer function.

The final member of the PIKK family possessing transducer functions is the

DNA‐dependent protein kinase. DNA‐PK is capable of phosphorylating both itself and downstream targets, making it both a sensor (as previously discussed) and a transducer [171‐173]. This autophosphorylation is required for increased activity by repair proteins during the DDR [174]. Evidence also suggests that DNA‐PK suppresses HR via phosphorylation of replication protein A (RPA) during S phase

[175]. Mice lacking functional DNA‐PK, usually via targeted deletion of DNA‐PKcs, display sensitivity to IR as well as showing defects in V(D)J recombination, which result in a phenotype similar to that of severe combined immunodeficiency mice and chromosomal instability via telomeric fusions, also characteristic of disruption of MRN, the downstream target of DNA‐PK [150, 176‐179]. DNA‐PK also plays a key

32 role in initiating apoptosis via the p53‐mediated pathway and suppresses p53‐ independent apoptotic pathways [180‐182]. Receiving signals, controlling repair pathways, and moderating apoptosis suggest DNA‐PK possesses transducer functions as well as sensor functions.

2.3.2 γH2AX

The histone subtype H2AX is phosphorylated on the carboxy terminal S139 after treatment with a DSB‐inducing agent, becoming the variant commonly known as γH2AX [26]. This occurs within minutes after DNA damage and spans hundreds of kilobases both up‐ and downstream of the damage [183‐185]. γH2AX is a commonly used signal of DNA damage as it is well conserved between organisms, known to occur in both yeast and higher eukaryotes, and persists after damage until repair is complete, approximately 2 hours post‐damage [184]. This phosphorylation is mediated by the members of the PIKK family, in humans γH2AX is phosphorylated by ATM but not ATR [47, 186]. There is also evidence that DNA‐PK also redundantly phosphorylates H2AX after IR and this is enhanced in the presence of histone acetylation [186, 187]. Cells in which H2AX phosphorylation is reduced show impaired DDR and recruitment of checkpoint proteins [188, 189]. In this way, it is an effective marker of affected regions and is possibly the mechanism by which

DNA damage sensors recruit downstream effectors to the correct area of the genome.

33 Further evidence for γH2AX as a transducer is supported by the fact that

γH2AX can influence the choice of repair pathway, subsequently regulating genomic stability. Cells lacking γH2AX use the more error‐prone single‐strand annealing pathway (not addressed in this work) instead of utilizing the less error‐prone HR‐ mediated repair [190]. Cell cycle arrest via the p53 pathway has also been shown to require γH2AX, suggesting versatility for γH2AX as a signal of DNA damage [191].

2.3.3 Claspin

The Claspin adaptor protein was first identified in Xenopus laevis as a protein required for successful activation of the Chk1‐mediated checkpoint response [192].

Based on this, homologues were discovered in mammals and yeast that shared similar functions with xClaspin [192‐194]. Furthermore, work on Claspin revealed that it behaves in analogous manner to Rad9A during TopBP1 recruitment: Claspin is currently hypothesized to act as a loading dock for the proteins required for Chk1 activation, namely the 911 complex, Rad17/RFC, ATR/ATRIP, and TopBP1 [52, 56,

86, 89, 98, 107, 193, 195‐198]. Claspin also regulates fork stability during DNA replication in a manner that is separate from its ability to regulate Chk1 activation

[199].

Claspin was shown to interact with members of the circadian rhythm pathways such as Timeless (Tim) and Tim‐interacting protein (TIPIN) [200]. These two proteins regulate DNA synthesis via the Chk1‐mediated checkpoint response

[200‐202]. Work by Yoshizawa‐Sugata et al. also suggests that Tim/TIPIN may be

34 required for nuclear localization of Claspin [200]. Between its role in Chk1 activation and its role during DNA replication, the Claspin adaptor protein is slowly gaining recognition as an important part of the DDR.

2.3.4 BRCA1 and BARD1

The breast cancer susceptibility genes were discovered in the 90’s as loci that were often lost or altered in breast and ovarian cancer patients. These loci were found on chromosomes 17q and 13q and the gene products of these loci were named BRCA1 and BRCA2 respectively [203, 204]. The implications of the chromosomal alterations associated with these two genes were grasped immediately, and over the next 15 years a flood of papers exploiting these genes in screens for breast and ovarian cancers were published. Mutations in the BRCA1 protein are being investigated as prognostic indicators, whether breast cancer will develop, and how a specific cancer will respond to therapy. It was also found that

BRCA1 and BRCA2 mRNA levels could be influenced by the estrogen steroid hormone, solidifying the link between breast and ovarian cancers with estrogen as well as being regulated in a cell cycle specific manner with mRNA levels peaking in the G1 and S phases [205‐208].

The breast cancer susceptibility gene BRCA1 is arguably one of the most active transducer proteins in the DNA checkpoint pathway. It exists as a heterodimer with its interacting protein, the BRCA1‐associated ring domain

(BARD1), and together act as an E3 ubiquitin ligase [209‐212]. The BRCA1/BARD1

35 complex has been implicated in a number of important cellular processes such as transcriptional activation of repair genes, such as Rad51 [213‐215], moderation of centromeres during S phase, an important regulator of chromosome stability [216,

217], and regulation of the DNA DSB repair machinery [218, 219]. This could be one of the functions for the BRCA1‐associated surveillance complex (BASC) [220]. The

BASC is a higher order protein complex comprised of a number of DNA repair and

DNA checkpoint proteins such as the MRN complex, the mismatch repair proteins

MSH2 and MSH6, ATM, the BLM helicase, as well as the PCNA clamp loader RFC complex [220]. BRCA1 is differentially phosphorylated in response to various genotoxins. In response to UV exposure BRCA1 is phosphorylated by ATM on S1457 whereas S1387 along with S1423 and S1524 are phosphorylated in response to IR, though the implications of these post‐translation modifications are not yet known

[221]. Evidence also exists for the role of BRCA1 in regulating the choice of repair pathways promoting recombinational repair via the MRN complex in response to signals from the Chk2 effector kinase [222]. The role of BRCA1 in moderating genome surveillance is dependent upon its interaction with Rad51, an important component for HR‐mediated repair, and not on the BARD1‐mediated E3 ubiquitin ligase activity [223]. However, despite all that is published about BRCA1, the mechanism of its tumour suppressor activity remains elusive and is the subject of many hypotheses.

BARD1 is currently hypothesized to regulate the transcriptional activity of

BRCA1 via its E3 ubiquitin ligase activity [224]. However, there is also evidence that

BARD1 has both BRCA1‐dependent and ‐independent roles during the cell cycle.

36 During mitosis, it was found that BARD1 is able to bind to the midbody, the site at which cytokinesis begins, without BRCA1 and that BARD1 is able to moderate ubiquitin‐mediated degradation of the Aurora B kinase, a kinase required for chromosomal rearrangements during mitosis [225]. Both BRCA1 and BARD1 play key roles during the DDR however there is still much that needs to be learned about these two key players.

2.3.5 TopBP1

TopBP1 is the human homologue of the fission yeast protein Cut5 and was identified via a yeast two hybrid screen against topoisomerase IIβ [226]. It contains several BRCT repeats characteristic of BRCA1‐interacting proteins [227], and interacts with the 911 complex [12, 13, 197], Claspin [228], and BRCA1 [226].

TopBP1 is required for the successful completion of S phase as well as for successful activation of the Chk1‐mediated ssDNA DDR [229‐232]. The TopBP1 protein also interacts with the ATR/ATRIP complex and mediates ATM‐dependent activation of

ATR following DNA damage [164, 165]. The interactions with other sensor/transducer proteins and the role of TopBP1 in regulation of the cell cycle and DDR suggest that TopBP1 is an important transducer of the DNA damage signal.

TopBP1 has also been suggested to be a new target for breast cancer screens: it is altered in a subset of breast cancers, further confirming its importance in maintaining genomic integrity [233, 234].

37 2.3.6 Summary of DNA damage transducers

The family of DNA checkpoint signal transducers has much more variability than the DNA damage sensors, which reflect their role as amplifiers of the DNA damage signal. Proteins in this category are associated with signal transduction; for example, members of the PIKK family have kinase function and BRCA1/BARD1 is an ubiquitin ligase. TopBP1 and the 911 complex, even though they possess no intrinsic signal transduction activity or their own, are vital components of downstream activation complexes. The ability to amplify and relay the DNA damage signal, increasing the number of proteins involved in the DDR is the key characteristic of a DNA damage transducer.

2.4 Checkpoint effectors

The effector is the final component in the archetypal checkpoint pathway. In actuality, the effector is a short signaling cascade consisting of three steps with the final step being the inactivation of the CDK/cyclin complex, which is the mechanism responsible for cell cycle arrest [235]. In this cascade, Chk1/Chk2 act to inactivate the Cdc25 phosphatase, preventing the cyclin/CDK from initiating the next phase of the cell cycle, as well as activating the Wee1 kinase, which adds inhibitory phosphorylations to the cyclin/CDK complex promoting arrest. The double actions of Chk1/Chk2 ensure the cyclin/CDK complex is inactivated, halting cell cycle progression; this signaling pathway is summarized in Figure 2.1.

38 2.4.1 Chk1

Chk1 is a serine/threonine kinase activated in response to the ssDNA damage pathway via ATR/ATRIP [236]. It is through Chk1 that the ssDNA damage checkpoint pathway connects to the cell cycle arrest mechanism. Significant evidence exists showing Chk1 is also activated in response to DSBs as well, acting as a backup to Chk2‐mediated cell cycle arrest (addressed in the next section) [44,

237]. Deletion of Chk1 is embryonic lethal in higher metazoans but Chk1‐deficient cells are viable in the chicken B lymphocyte cell line, DT‐40 [238]. These cells were found to be sensitive to genotoxins and Chk1 was essential for the G2/M DNA damage checkpoint arrest. These cells also demonstrate that Chk1 is dispensable in regulating the normal progression from G2 to mitosis as these cells were able to enter mitosis in the absence of a DNA damage signal [238]. However, this data has recently been challenged by the discovery that Chk1 was required during mitosis for proper chromosome segregation in heterozygous mice [239]. This group also reported that disrupting Chk1 activity resulted in nondisjunction and binucleation, a result of interrupted cytokinesis confirming a role for Chk1 in mitosis [239].

As previously mentioned, activation of Chk1 in response to the DNA damage signal requires more than just the ATR/ATRIP complex [90]. Claspin [193, 196],

TopBP1 [197], the 911 complex [89, 193], and Rad17/RFC [193, 240] are all required for Chk1 activation. This Chk1‐activation complex is depicted in Figure

2.3. In addition to the requirement for the initiation of arrest, Chk1 is also involved in maintaining cell cycle arrest, preventing premature entry into mitosis [241].

Phosphorylation, and thereby activation, of Chk1 occurs on S345 and is the product

39

Figure 2.3 – Schematic of theChk1 activation complex. Diagram depicting the proteins involved in Chk1‐mediated checkpoint initiation during the DDR.

40 of ATR/ATRIP [236]. ATM‐mediated Chk1 activation occurs via phosphorylation on

S317 in response to IR and requires the DSB repair protein Nbs1, a component of the MRN complex [44]. Upon activation, Chk1 is also able to inactivate the Cdc25 phosphatase via phosphorylation [242]. Chk1 is responsible for phosphorylating the Cdc25A phosphatase on T504, deactivating it, resulting in the persistence of the inhibitory phosphorylation on the Cdk1/cyclin B complex, ultimately resulting in cell cycle arrest [243, 244]. Chk1 also phosphorylates Cdc25B on S230 and S563 in the absence of DNA damage [245]. This isoform of Cdc25 is involved in initiating early mitotic events and providing a mechanism for Chk1‐mediated mitotic activation [246]. Chk1 also phosphorylates the Wee1 kinase, increasing its activity towards Y15 of Cdc2, required for maintaining checkpoint arrest [247]. Stating the role of Chk1 during the DDR as briefly as possible: Chk1 works by deactivating the cyclin/CDK activator Cdc25 and activation of the cyclin/CDK inhibitor Wee1.

2.4.2 Chk2

The Chk2 serine/threonine kinase, also known as Cds1 in yeast, is the key effector to the DNA DSB pathway, moderated via ATM. Activation of the Chk2 kinase after exposure to radiation occurs via phosphorylation of the T68 residue by

ATM [45, 248]. As is the case with Chk1, there is significant overlap between the ssDNA response pathway and the DSB response pathways in that ATR/ATRIP is also able to activate Chk2 via phosphorylation of the T68, S50, and T26 residues [249].

Once activated, Chk2 phosphorylates Cdc25 on S123, targeting it for destruction

42 [242, 250]. In fact, evidence exists showing that Cdc25 can be stimulated even in the absence of T68 phosphorylation, demonstrating the importance of this action [251].

Unlike either ATR or Chk1, deletion of Chk2 in mice does not result in embryonic lethality nor do these mice develop spontaneous tumours like the ATM‐/‐ mice [41, 53, 57, 238]. Chk2‐deficient murine cells displayed defects in triggering

IR‐dependent apoptosis as well as defects in the G1/S checkpoint [53]. IR‐ dependent apoptosis was found to be restored in deficient cells by expressing complementary Chk2 in both an ATM‐dependent and an ATM‐independent manner; this indicates that Chk2 is activated by more than just the ATM‐moderated branch of the checkpoint response pathway [53]. ATR activates Chk2 in response to UV and

IR, providing additional evidence supporting crosstalk between the DSB and ssDNA checkpoint pathways [252]. Alterations in Chk2 activity have been implicated in several human cancers including breast [253, 254], colon [253, 255], ovarian [253], lung [256], prostate [257], and Hodgkin’s lymphoma [258].

Chk2 is also able to increase transcription of genes moderated by the E2F‐1 family of transcription factors [259]. By expressing a dominant negative form of

Chk2, E2F‐1‐mediated apoptosis was abolished [259]. Confirming the role of Chk2 in moderating E2F transcription, it was found that Chk2 was able to phosphorylate, and activate, the RNA polII‐binding protein Che1, a known regulator of E2F‐ mediated transcription [260]. It is also possible that Chk2 could affect mRNA stability of a subset genes, this is based on the finding that Chk2 was responsible for phosphorylating the HuR mRNA stability protein, leading to increased translation of

43 the stress response gene SIRT1 [261]. Increased transcription of XRCC1 and BRCA2 via the FoxM1 transcription factor have also been reported as well as the ability of

Chk2 to activate transcription via another family of the Fox transcription factors, the

FoxO1 family [262, 263]. Collectively, these data provide strong evidence for Chk2 as a moderator of transcription of DNA repair genes, further solidifying the role of

Chk2 as an effector of the DSB DDR pathway.

Chk2 was also predicted to activate Mre11, a key component in the DSB resolution pathway [264]. Evidence also exists for the role of Chk2 in regulating the activities of the breast cancer susceptibility gene product, BRCA1. It was found that

Chk2‐dependent phosphorylation of S988 on BRCA1 aided in the promotion of HR vs. NHEJ during recombinational repair, which is supported by the Chk2‐dependent activation of Mre11 mentioned above [222]. Mutation of this site abolished HR‐ mediated repair and suppression of the NHEJ pathway [222]. Similar to Chk1, Chk2 phosphorylation of BRCA1 S988 also seems to regulate spindle assembly formation, implicating Chk2 as a requirement for successful DNA segregation during mitosis

[265]. Additionally, evidence exists for the role of Chk2 in moderating the fidelity of

NHEJ through activation of BRCA1 [266, 267]. Chk2 also seems to be able to promote base‐excision repair by phosphorylating XRCC1 [268]. Based on these data, Chk2 is clearly a key member of the DDR with a variety of functions pertaining to the maintenance of genomic integrity.

44 2.4.3 The Cdc25 Phosphatases

The Cdc25 phosphatases are dual specificity phosphatases, able to dephosphorylate both phospho‐tyrosine and phospho‐threonine residues, consisting of three isoforms in mammalian cells [19, 269‐274]. Each isoform has a unique function, Cdc25A is required for the G1/S transition while Cdc25C is required for the transition from G2 to mitosis [275, 276]. Cdc25B is required for the transition into mitosis but also appears to have separate function from Cdc25C [246,

277]. One group found that both Cdc25B and Cdc25C were able to restore function in cdc25­null yeast cells but that these two isoforms responded to different checkpoint signals during the G2/M transition [278]. This group also found that activation of Cdc25C in response to both HU and IR required both of the checkpoint kinases, Chk1 and Chk2, but that activation of Cdc25B in response to these insults only required Chk1 [278]. They also found that Cdc25B and Cdc25C were equally active during the response to DNA damage from UV. Evidence also exists showing that Cdc25A, in conjunction with Cdc25B, is required for successful initiation of mitosis [279]. This group suggested that Cdc25C alone was not able to initiate mitosis, but required Cdc25A to catalyze chromosome condensation and Cdc25B to activate the Cdk1/cyclin B1 complex [279]. New evidence also exists demonstrating the requirement of Cdc25B for proper centriole formation during chromosome replication [280]. It is clear from the work on these phosphatases that initiation of mitosis is complicated and checkpoint initiation at this transition is dependent upon the type of DNA damage sustained, be it processed into ssDNA or DSBs.

45 Another method of regulating Cdc25 activity is via phosphorylation, which is required for cytoplasmic sequestration by the 14‐3‐3 family of proteins. Cdc25C is cytoplasmic during interphase, a result of 14‐3‐3 binding to the phosphorylated

S216 residue and that binding of 14‐3‐3 results in nuclear export of Cdc25C [275,

281]. This phosphorylation is moderated by the Chk1 kinase; however, work has also shown that elimination of either of the nuclear export signals (NES) in the N‐ terminus of Cdc25C results in nuclear retention, leading to mitotic initiation, and is not required for 14‐3‐3 binding [282‐284]. Combined, the NES and 14‐3‐3 binding moderate the activity of the Cdc25C phosphatase during the DDR [282]. For yet another layer of regulation, the NES of Cdc25C are activated via phosphorylation of

S193 by Plk3 [285]. To summarize, Cdc25C activity is moderated by Plk3‐mediated nuclear exclusion and Chk1‐mediated 14‐3‐3 binding, both of which act synergistically to regulate the phosphatase activity of Cdc25C on the Cdk/cyclin complex. Confliciting data about 14‐3‐3‐mediated regulation of Cdc25B exists in the literature. Davezac et al. reported Cdc25B is also regulated via 14‐3‐3 binding to phospho‐S323 while Mils et al. reported that binding of Cdc25B and 14‐3‐3 did not require phosphorylation [286, 287]. Another group reported that amino acid residues on the C‐terminal end of the 14‐3‐3 binding motif were able to regulate the efficacy of binding [288]; these data suggest this process requires further study. In keeping with this pattern, Cdc25A is also regulated via Chk1 phosphorylation of two residues, S178 and T507, in 14‐3‐3 binding domains, regulating proteolysis and preventing mitotic initiation before DNA replication is complete [289].

46 Over‐expression of the different Cdc25 isoforms is associated with poor prognosis in a number of cancers including head and neck [290], gastric [291], non‐ small cell lung [292], breast and ovarian [293], and colorectal [294]. Higher levels of

Cdc25A are associated with poor prognosis, higher Gleason score, and metastasis in prostate cancer; additionally, Cdc25A is also able to physically interact with the androgen receptor, thereby decreasing its transcriptional activity, which plays a role in the development of prostate cancer [295]. In esophageal squamous cell carcinoma, autoantibodies against Cdc25B were detected in 36% of patients tested and 0% in healthy test subjects [296]. This suggests that the Cdc25 phosphatase isoforms could be targets for either screening or treatment in specific carcinomas.

2.4.4 Wee1/Myt1

The Wee1 kinase has the opposite function on the Cdk/cyclin complexes to the Cdc25 phosphatase. Whereas Cdc25 takes away the phosphorylation on Cdk1, allowing normal cellular progression, Wee1, with its partner Myt1, also known as

Mik1, add the inhibitory tyrosine phosphorylations, T14 and tyrosine 15, to Cdk1 preventing progression [297‐301]. In addition to inactivating Cdc25, Chk1 and

Chk2 are also capable of activating Wee1/Mik1 [242, 247]. Abrogation of Chk1 via the Chk1‐inhibitor UCN‐01 results in decreased activation of Wee1 [302, 303]. The antagonistic actions of the Wee1 kinase and the Cdc25 phosphatases provide for a more exact control over the progression from the G2 phase to mitosis. Further control is executed over Wee1 via increased transcription after DNA damage, which

47 is stimulated by BRCA1 [219]. Wee1 is also required for the embryonic development of both mice and X. laevis, and regulates the size of tomato fruit, this evolutionary conservation highlights the importance of Wee1 in the cell cycle [304‐

306]. Phosphorylation of Wee1 results in 14‐3‐3‐mediated proteosomal degradation and results in cell cycle progression [307, 308].

2.4.5 Cyclin/CDK complexes

The cyclin/CDK complexes are the main regulatory proteins required for cellular progression, the entire checkpoint process funnels down to these complexes. Each phase of the cell cycle is associated with a particular cyclin/CDK combination and it is upon these that the effectors of the cell cycle checkpoints, such as Cdc25 and Wee1, act to enforce cell cycle arrest. Since this subject has been covered thoroughly in the literature, the following section will very briefly summarize the role of the cyclin/CDKs during the DNA damage response.

The family of cyclin proteins were initially identified by their changing levels as the different phases of the cell cycle progress, hence the name cyclins, more recently they were identified by sequence homology via the “cyclin box” [309, 310].

For example, as the DNA synthesis phase progresses, levels of cyclin A decrease while levels of cyclin B increase in preparation from the shift from S phase to the G2 phase. Though there are 29 members of the cyclin family, only some are known to interact and activate CDKs. There are 4 cyclin families associated with cell cycle progression with each family temporally defined. The A cyclins are associated with

48 the shift from S phase to G2, the B cyclins with G2 to M progression, the D and E families of cyclins are associated with G1 progression and passage through the restriction point respectively.

The other half of the cyclin/CDK complex is the serine/threonine kinase subunit, which is only active when bound to its cyclin partner (reviewed in [311‐

315]). The number of cyclin/CDK targets is staggering and includes such proteins as

Rb, the retinoblastoma protein, BRCA1, BRCA2, BARD1, Cdc25A and C, p53, Ku70, and members of the MCM DNA replication complex. CDKs affect the transcription of a number of genes required for that particular phase of the cell cycle. Like cyclins, the number of CDKs is greater than the number involved with the cell cycle. There are 20 CDK and CDK‐like proteins, of which only a few are associated with the cell cycle. CDK1, often referred to as Cdc2, is best known for being the CDK involved in the progression from G2 to M. The G1 to S progression is governed by CDK2 along with CDK4 and CDK 6, while CDK3 is involved in re‐initiation of the cell cycle following quiescence, also known as G0.

Due to their importance in regulating the cell cycle, the cyclin/CDK complex has become a target for cancer treatment and a number of new drugs are looking to exploit any difference between cyclin/CDK regulation in normal versus cancerous cells. Evidence exists suggesting that cancerous cells use specific cyclin/CDK combinations to regulate their cell cycle, combinations that are different from normally proliferating cells. Selective CDK inhibitors targeting a single cyclin/CDK combination may help to target actively proliferating tissues, such as neoplastic

49 tissue. This new class of drugs is currently under investigation and is a promising addition to the palate of anti‐cancer therapeutics.

2.4.6 Summary of the DNA damage effectors

The proteins in this category represent the end‐stage for the DNA damage signal following checkpoint activation. It is these proteins that are responsible for halting cellular progression allowing time for DNA repair to take place and represent the culmination of an unperturbed checkpoint signaling cascade.

Disruptions to the checkpoint cascade will manifest as cell cycle arrest failure via the mechanisms described here. Like the transducers, these proteins possess either kinase or phosphatase function, allowing them to directly influence the activity levels of other proteins, which is necessary for proper checkpoint function.

2.5 Double­strand break repair

The DSB is a lethal form of DNA damage and is often to blame for the loss of large regions of DNA information; therefore, it is crucial that any DSB present are resolved in timely and efficient manner. There are currently three known methods of DSB repair employed by the cell, the method chosen mostly depends on which phase of the cell cycle the cell is presently in but the exact method of choice of repair pathway is not yet fully understood. Cells in the G2 phase, containing two full copies of the genome, use the HR method, as it is the most error‐free method. Cells in the

G1 do not contain an extra copy of the genome and so must rely on less exact

50 methods of repair, the NHEJ and alt‐NHEJ pathways. Depending on the amount of the genome duplicated in S phase helps to determine which pathway is chosen and evidence exists suggesting that the HR and NHEJ pathways compete for DNA free ends during this phase. Proteins involved in these three pathways overlap and often contribute to each pathway in some way.

2.5.1 Homologous recombination

Of the three major DSB repair pathways utilized by the cell, HR is the most precise due to the presence of sister chromatids, which are used a template to fill in sequence gaps resulting from the break or DSB processing. In addition its role in

DNA repair, HR is also used by the cell to create genetic variability during the process of meiosis and was recently discovered to play a role in creating immunoglobulin variability during V(D)J recombination, a process thought to be restricted to the NHEJ pathway, though these aspects of HR will not be discussed any further [316‐318].

The DSB is detected by the MRN complex, which plays a major role in, not only detecting, but also in signaling and resolving DSBs. Each component of the

MRN complex plays a key role in repairing the DSB, the zinc hook domain of Rad50 acts to pull the ends of the damaged DNA closer together in preparation for repair

[145]. After Rad50 links the ends together the dimerization activity of Mre11 act to further join and stabilize the DNA free ends as well as utilizing its nuclease activity to resect the DNA ends producing the 3’ overhang required for HR [140, 319]. Both

51 Rad50 and Mre11 are capable of dimerizing to form M2R2 heterotetrameric complexes, a key step in repair [145, 320]. Ablation of this ability results in a similar phenotype to the deletion mutants [142, 148]. Nbs1, or Nibrin, is utilized to initiate apoptosis in the case the damage cannot be resolved as well as catalyzing the effects of Mre11 and unwinding DNA at the break site in preparation for the next step of HR

– strand invasion [123, 152]. Around this time the homologous strands are aligned so the undamaged strand can act as a template to the damaged one, promoted by the actions of Rad50 and cohesin [118].

In addition to the repair aspects of MRN, there is also the signaling aspect.

Evidence exists suggesting MRN is responsible for activating the PIKK ATM in response to DSB, ATM is a key transducer during DNA repair in part due to the alteration of the H2AX histone subtype to γH2AX, and that this recruitment is dependent upon parts the Nbs1 protein [47, 159, 161, 321‐324]. Once activated

ATM is also responsible for activating the serine/threonine checkpoint kinases Chk1 and Chk2 as well as phosphorylating members of MRN, such as Nbs1 [44, 325, 326].

MRN signaling also extends to the p53‐mediated pathways, recruitment of the p53‐ moderator Mdm2 is accomplished by Nbs1 [154].

The 911 complex has also been shown to play a role in the detection of DSBs and in localizing the TopBP1 transducer the sites of DNA damage [12]. The BRCT domains of TopBP1 provide another link to the BRCA1‐mediated branch of DSB repair as well as to the E2F1 transcription factor, via Chk2 [226, 327, 328]. TopBP1 also binds to Nbs1, further linking the 911 and MRN complexes to the transducer

52 machinery [329]. As the Chk1 arm of the DDR is activated this means that ATR is activated as well as ATM, further increasing the DNA damage signal. Evidence also suggests than ATM and members of MRN are also capable of activating ATR in response to IR ensuring signal amplification and damage resolution [168].

The BRCA1 protein is brought in to the DSB repair pathway by the interaction with TopBP1 but also by being the key member of the BASC, which, as previously mentioned, includes the MRN complex as well as Rad51 and BRCA2

[220]. Interestingly, it seems that BRCA1 negatively affects the activities of the MRN complex, perhaps ensuring this complex does not begin randomly attacking adjacent regions, and that its role during HR does not depend on its E3 ligase activity [223,

330]. Rad51 and BRCA2 are well known members of the HR pathway, Rad51 has been shown to coat ssDNA, stabilizing it during the repair process and catalyzing strand invasion between the sister chromatids while BRCA2 modulates the binding specificity of Rad51 [331, 332]. BRCA2 is closely involved in the Rad51‐mediated

HR pathway and deletion of mouse BRCA2 results in Rad51‐mediated cell death and embryonic lethality [205, 333, 334]. BRCA2 interacts with Rad51 via the BRCA1 carboxy‐terminal (BRCT) repeats in BRCA2 and, in turn, protects Rad51 from proteosomal degradation [335]. Over‐expressing a proteosome‐resistant form of

Rad51 fully complements the BRCA2‐deficient phenotype in mouse and human cells suggesting the effects of BRCA2 are executed via Rad51 exclusively [335]. BRCA2 also modulates the activity of the Rad51 protein both by catalyzing Rad51 binding to ssDNA and preventing binding to dsDNA, ensuring the specificity of Rad51 to sites of DSB repair, in addition to the proteosomal protection mentioned above

53 [331]. Unwinding by Nbs1 and strand invasion by Rad50 and Rad51 are key steps in preparing the damaged DNA strands for repair, after these steps a DNA polymerase, either δ or η, evidence exists supporting both, fills in the missing bases using the sister chromatid as a template [118, 336‐338]. The structure formed by the four strands of DNA after strand invasion is referred to as a Holliday junction, named for the scientist who first proposed them, is a key step in resynthesizing the invading, damaged strand. After resynthesis, the Holliday junction is resolved enzymatically, a process still not well understood in humans but known to involve the Gen1 protein [339], and the repair is complete.

2.5.2 Non‐homologous end joining

During phases of the cell cycle in which there is only a single copy of genome present, during G1 and the S phases for instance, a different method of DSB repair is used, the NHEJ pathway. Many of the same players are involved in both the HR and

NHEJ pathways and evidence points at BRCA1 in determining which pathway will be utilized, in that small window during the cell cycle in which either pathway could be used [222]. The NHEJ pathway is also known as microhomology‐mediated end joining, is moderated by the PIKK DNA‐PK rather than ATM or ATR and evidence suggests DNA‐PK suppresses other methods of DSB repair in favour of NHEJ [175].

Unfortunately, this method does not ensure that the ends joined are of the same chromosome and this is mechanism behind many different chromosomal translocations characteristic of specific cancers [340]. For example, Burkitt’s

54 lymphoma is a well‐characterized translocation between chromosomes 8 and 14

[341], and the Philadelphia chromosome, a translocation between chromosomes 9 and 22, found in chronic myeloid leukemia [342]. As previously mentioned, this is method primarily responsible for the V(D)J recombination utilized during immunoglobulin diversification and is also a key component in telomere maintenance [66, 343‐345]. Defects in any of the major components of the NHEJ pathway result in the severe combined immunodeficiency phenotype [346].

In HR the MRN complex is responsible for detecting DSBs but in NHEJ the Ku hetertodimer, made up of Ku70 and Ku80, binds to the damaged DNA ends and acts to recruit the other members of the NHEJ pathway to the sites of damage [71]. Ku, in addition to its role as the activating component of DNA‐PK, is a ring‐shaped complex that encircles the DNA strand, contains a putative DNA binding domain on the Ku70 subunit, and a large C‐terminal tail on Ku80, capable of binding DNA‐PKcs as well as other checkpoint proteins such as BRCA1 [72, 347‐350]. Despite its importance during the NHEJ repair pathway, mice devoid of Ku80 protein do not die during embryogenesis [351]. They do however exhibit a variety of defects including premature aging, most likely linked to the role of NHEJ in telomeric maintenance, and, interestingly, defects in homologous recombination [351‐353]. Curiously, deletion of Ku70 in mice results in a different phenotype suggesting the Ku components may have roles outside the Ku heterodimer. Ku70‐/‐ mice show increased levels of thymic lymphomas whereas there is no increased risk of cancer in the Ku80‐/‐ mice [354].

55 After detection by Ku, recruitment of the downstream proteins to the DSB site begins. Ku attracts DNA‐PK and the nuclease Artemis, which bind to the DNA free ends, where DNA‐PK autophosphorylation moderates the nucleolytic activity of

Artemis, ensuring Artemis does not remove more bases than is necessary for proper

DNA end processing [174, 355, 356]. Autophosphorylation of DNA‐PK is believed to play an important role in the regulation of assembly and disassembly of the DNA‐ protein complexes formed during NHEJ as well as repressing repair via other pathways [175, 348]. There is conflicting evidence whether DNA‐PK is required for recruitment of the DNA ligase IV/XRCC4 complex, one group found that DNA‐PK was required for the interaction with Ku while another found the opposite to be true and it was Ku that was required for recruitment, additional research is required to clear up this uncertainty [357, 358]. DNA ligase IV/XRCC4 is the component responsible for reconnecting the DNA ends together during the completion of the

DNA repair process with XRCC4 moderating the activity of DNA ligase IV [359‐361].

There are two DNA polymerases responsible for filling in gaps created during the nuclease‐dependent end processing step, DNA pol λ and pol μ, both of which have been implicated in the repair process [362, 363].

In addition to DNA ligase IV and XRCC4, another factor has been identified as a key player in the NHEJ pathway: the XRCC4‐like factor (XLF), also known as

Cernunnos, the human homologue to the yeast NHEJ protein Nej1 [364]. XLF binds directly to the DNA ligase IV/XRCC4 complex and promotes ligation of the DSB

[365]. Cells deficient for XLF show increased sensitivity to genotoxins and defects in

56 V(D)J recombination, similar to deletion mutants of the other NHEJ components, and has been associated with clinical symptoms in a human patient [366‐369].

2.5.3 Alternative non‐homologous end joining

The alternative NHEJ, alt‐NHEJ, repair pathway utilizes the poly(ADP‐ribose) polymerase (PARP)‐1 as the sensor of DSBs, competing with Ku to determine whether the classic NHEJ or alt‐NHEJ repair pathways will be used for repair [370].

Interestingly, there is also a new line of evidence suggests that choice of DSB repair pathway may also be dependent upon DSB microhomology sequences [371]. In fact, it seems PARP‐1 plays a role in protecting cells from NHEJ‐induced death during

DSB repair by promoting HR repair [372]. PARP‐1 is a ADP‐ribosylase, capable of utilizing NAD to add poly(ADP‐ribose) to DNA‐binding proteins, particularly the H1 histone subunit, at sites of damaged DNA and is thought to flag damaged DNA for repair [373, 374]. There is also evidence suggesting a role for PARP‐1 in V(D)J recombination, PARP‐1 deficient mice show abnormal antibody responses, and in sister chromatid exchange during HR [375, 376]. To further confirm the role of

PARP‐1 in DSB repair, one group reported an interaction between TopBP1 and

PARP‐1, where PARP‐1 was able to influence the activity of TopBP1‐protein interactions [377, 378]. Another group reported an increased Chk1‐dependent checkpoint arrest in the absence of PARP‐1, indicating a role for PARP‐1 in moderating the DSB repair pathway after IR exposure [379]. There is also growing evidence supporting a role of PARP‐1 in regulating the ATM‐mediated DDR pathway

57 via moderation of ATM‐dependent phosphorylation of downstream targets such as

H2AX [380, 381]. PARP‐1 is responsible for recruiting the XRCC1/DNA ligase III complex to stabilize and join the DNA free ends respectively, mirroring the

XRCC4/DNA ligase IV complex in NHEJ [382, 383].

These three pathways, HR, NHEJ, and alt‐NHEJ, are the three main ways in which DSBs are resolved in the cell. DNA damage involving only a single strand of the DNA duplex utilizes a different set of DNA repair pathways.

2.6 Checkpoint Release

Despite the importance of checkpoints in cellular survival, less is known about how they are released, allowing the cell to resume cell cycle progression, which is surprising given that prolonged checkpoint arrest is capable of triggering apoptosis. In order to reinitiate the cell cycle, inactivation of the effector kinases and reactivation of the cyclin/CDK complexes are important steps. The S. pombe homologue of protein phosphatase 1 (PP1), Dis2, dephosphorylates Chk1, reducing its inhibitory effects on Cdk1 [384, 385]. In addition to this, a feedback loop initiated by Cdk1, a target of the Wee1 kinase, phosphorylating Wee1 thereby maintaining the inactivated state of Cdk1 and regulating its own activity via its target [386]. In budding yeast, it appears that prolonged G1 arrest causes the cell to adapt to the checkpoint signal and results in eventual release from G1 arrest activated by the S. cerevisiae BRCA1 homologue and moderated by Pho85, a CDK required for normal progression from G1 to S and downregulated in the presence of

58 DNA damage [382]. Resolution of the spindle checkpoint during mitosis is dependent on ubiquitination of checkpoint proteins, resulting in their degradation, by the anaphase promoting complex [388]. These data suggest that alleviation of the checkpoint response may be mediated by inactivation, either via inhibitory post‐ translational modifications or by degradation, of key checkpoint proteins but much more work in this area is needed to confirm this conclusion.

2.7 Hypotheses and specific aims

Given the requirement for both Rad9A and Claspin in regulating Chk1 activation [192, 193], it was hypothesized that the interaction between those two proteins would be important for downstream checkpoint signal transduction. This prompted investigations into the possible role of the post‐translational modifications of the Rad9A C‐terminal tail during the Rad9A‐Claspin interaction.

Specifically, the role of Rad9A phosphorylation in moderating this interaction was to be determined as well as investigating a functional role for this interaction during the DDR. The work resulting from these studies is reported in Chapter 4.

The involvement of DNA damage response proteins ATM, ATR and BRCA1 in

Xi maintenance supports the idea that checkpoint function is somehow required for maintenance of the inactive X chromosome [389‐391]. Previous work in our laboratory demonstrated the presence of large, Rad9A‐containing bodies at the Xi

[12], and led us to hypothesize that Rad9A was playing a role in chromatin maintenance or silencing at the Xi. The second aim of this work was to test this

59 hypothesis by determining how Rad9A was involved at Xi via microscopic methods.

The findings of these investigations are displayed in Chapter 5.

Combined, these two aims will further understanding of the role played by the Rad9A checkpoint protein during the DDR.

60 Chapter 3: Materials and Methods

3.1 Cell Lines and Culture Conditions

The cervical carcinoma cell line HeLa (CCL‐2), a human telomerase‐reverse transcriptase‐transformed retinal pigment epithelial cell line (hTERT‐RPE1, CRL‐

4000), and fibroblasts taken from the skin of a patient suffering from Klinefelter’s syndrome, known as the Dempsey cell line (CCL‐28), were obtained from the ATCC cell repository (Manassas, VA). An immortalized and undifferentiated breast tissue cell line, 184‐hTERT, was a generous gift from Dr. Chris Mueller at Queen’s

University in Kingston ON, Canada. Murine embryonic stem cells (mES), both wild‐ type and containing a genomic deletion of the rad9A gene were a gift from Dr.

Howard Lieberman of Columbia University of New York NY.

HeLa cells were maintained in Dulbecco’s Modified Eagle’s medium (DMEM)

(Invitrogen, Burlington, Canada) supplemented with 10 % heat inactivated fetal bovine serum (FBS) (Sigma Aldrich, Oakville, Canada) and 1x antibiotic‐antimycotic

(Invitrogen). Heat inactivation of FBS was performed via incubation at 55 oC for 40 min. HeLa cells stably expressing a Rad9A‐GFP fusion protein were cultured in the presence of 100 μg/mL G418 (Sigma) [12]. hTERT‐RPE1 cells were maintained in

DMEM F12 HAM media (Invitrogen), supplemented with 10 % heat inactivated FBS and 5 mM L‐glutamine (Sigma). Dempsey cells were maintained in McCoy’s modified 5a media supplemented with 1.5 mM L‐glutamine, 2.2 g/L sodium bicarbonate, and 20 % FBS. The immortalized, undifferentiated breast tissue cell line, 184‐hTERT, was maintained in mammary epithelial cell growth medium

61 (MEGM) bullet kit (Cedarlane Laboratories, Burlington, Canada), which contains 10 ng/mL hEGF, 5 µg/mL insulin, 0.5 µg/mL hydrocortisone, 50 µg/mL gentamicin, 50 ng/mL amphotericin‐B, and 52 µg/mL bovine pituitary extract. A summary of these conditions is displayed in Table 3.1.

mES were maintained in gelatinized culture dishes in KnockOut DMEM

(Invitrogen) supplemented with the following: 15% FBS (ES qualified – Invitrogen),

0.1 mM β‐mercaptoethanol (Bioshop, Burlington, Canada), 0.1 mM nonessential amino acids (Invitrogen), 2 mM L‐glutamine, 50 ug/mL penicillin/streptomycin

(Sigma), and 103 units/mL leukemia inhibitory factor (LIF) (ESGRO ‐ Millipore,

Etobicoke, Canada). In order to induce differentiation in mES cells, they were grown in media in which LIF was omitted and supplemented with 10 μM all trans‐retinoic acid (RA). Cells were either harvested as normal for mRNA or replated on coverslips (described below), as the experiment required. All cells were cultured at

37 °C and 5 % CO2 environment in 100 mm, 60 mm, or 6‐well tissue culture dishes

(Sarstedt, Montreal, Canada) as required.

3.2 Cell Synchronization, Transfections and Treatments

HeLa cells were synchronized via double thymidine block as previously described [11, 12]. Briefly, cells were treated with 2 mM thymidine (Bioshop) for 18 hrs and released by washing once with PBS followed by incubation in fresh media for 8 hrs. After this recovery period, cells are treated with 2 mM thymidine for a further 18 hrs and released in the same manner. Following the second release, the

62 Table 3.1 – Tissue culture conditions for the different cell lines used during the course of this thesis.

Cell Line Source Media Supplements HeLa (CCL‐2) ATCC DMEM 10 % FBS; 1x antibiotic/antimycotic hTERT‐RPE1 DMEM‐ ATCC 10 % FBS; 5 mM L‐glutamine (CCL‐4000) F12 HAM 10 ng/mL hEGF; 5 μg/mL insulin; 0.5 Dr. Chris μg/mL hydrocortisone; 50 μg/mL 184‐hTERT MEGM Mueller gentamicin; 50 μg/mL amphotericin‐B; 52 μg/mL bovine pituitary extract Dempsey McCoy’s 20 % FBS; 1.5 mM L‐glutamine; 2.2 g/L ATCC (CCL‐28) 5a sodium bicarbonate 15 % FBS; 0.1 mM β‐mercaptoethanol; 0.1 Dr. mM non‐essential amino acids; 2 mM L‐ KnockOut mES Howard glutamine; 50 μg/mL DMEM Leiberman penicllin/streptomycin; 103 units/mL ES‐ GRO

63 peaks of S phase, G2, and M are observed at approximately 2‐4 hours, 6 hours, and

8‐10 hours, respectively. Approximately 45 % of cells have progressed to G1 12 hrs following release.

Transfections were performed with the FuGENE 6 transfection reagent

(Roche Diagnostics, Mississauga, Canada) according to the manufacturer’s instructions. Unless otherwise indicated, cells were cultured for 48 hrs post‐ transfection and treated as indicated. The ratio of reagent (μL) to DNA (μg) for all reactions was 4:2. Transfections were also carried out via electroporation, otherwise known as either electrotransfection or electropermeablization, where indicated, using the method outlined in van den Hoff et al. [392]. Briefly, cells were harvested via trypsinization, washed twice in PBS pre‐warmed to 37°C and

7 resuspended at 1x10 cells / mL in ice‐cold cytomix (120 mM KCl, 0.15 mM CaCl2, 10 mM K2HPO4/KH2PO4 [pH 7.6], 25 mM HEPES [pH 7.6], 2 mM EGTA [pH 7.6], 5 mM

MgCl2, supplemented with freshly made 2 mM ATP [pH 7.6] and 5 mM GSH).

Approximately 5 μg of DNA was mixed with 400 μL (4x106 cells) of the cell‐cytomix slurry and added slowly to an ice‐cold 4 mm electroporation cuvette, to prevent formation of bubbles, and shocked using a Gene Pulser (Bio‐Rad, Mississauga,

Canada) set to 0.40 kV and 500 μF. Cuvettes were placed immediately on ice then the contents plated in pre‐warmed DMEM in 10 cm dishes. This growth medium was replaced the following day to remove any cells that did not survive the electroporation procedure. Cells were harvested 48 hrs post‐electroporation.

64 Irradiation was carried out using a Victoreen Electrometer (Atomic Energy of

Canada, Mississauga, Canada) 137Cs γ‐irradiator with an approximate dose rate of

0.5 Gy / min for the required time. Unirradiated cells were transported and treated similarly as the irradiated samples but were not exposed to the γ‐ray source.

3.3 Immunofluorescence and Confocal Microscopy

Cells were seeded at 20 % confluence on glass coverslips, and cultured for 48 hrs in either 6‐well or 6 cm dishes. mES cells were plated on coverslips coated in

o fibronectin that had been allowed to dry for 30 min at 37 C, at 5 % CO2 in a humidified environment. Cells undergoing transfection were allowed to grow for 24 hrs before transfection.

Cell fixation was achieved via incubation in 4 % paraformaldehyde (PFA) for

15 min at room temperature (RT). Permeablization was performed in phosphate buffered saline (PBS) / 0.5 % Triton X‐100 for 10 min at RT followed by incubation in ice‐cold blocking solution (PBS, 0.1 % Triton‐X100, 5 % normal goat serum) for at least one hour at 4 °C. Primary antibodies (Table 3.2) were diluted in ice‐cold blocking solution, then were added drop‐wise to the cover slips and incubated for one hour at RT. Following incubation, the cover slips were washed three times for 5 min per wash in PBS containing 0.1 % Triton‐X100. Alexa fluorochrome‐conjugated secondary antibodies (Table 3.3) were diluted in ice‐cold blocking solution at a ratio of 1 / 200, added drop‐wise to the coverslip and incubated for 30 min in the dark.

Due to the light‐sensitivity of the secondary antibodies, manipulations were

65 Table 3.2 – Primary antibodies used in immunoblotting and immunofluorescence experiments. Antibody Dilution Dilution Description Company (IB) (IF) polyclonal, RCH Antibodies, Rad9A 1/2000 1/200 described in [4] Kingston, ON Santa Cruz c‐Myc monoclonal, cat.# 1/2500 1/200 Biotechnology, Santa (9E10) sc‐40 Cruz, CA Calbiochem (now EMD BRCA1 (Ab‐ monoclonal, cat.# 1/1000 1/100 Biosciences), La Jolla, 1) MS110 CA Claspin (H‐ polyclonal, cat.# Santa Cruz 1/500 1/100 300) sc‐48771 Biotechnology Upstate (now polyclonal, cat.# macroH2A1 N/A 1/200 Millipore), Temecula, 07‐219 CA γH2AX (Ser monoclonal, cat.# 139) (clone N/A 1/200 Upstate 05‐636 JBW301) polyclonal, cat.# H3trimK9 N/A 1/200 Upstate 07‐442 TRF2 (clone monoclonal, cat.# N/A 1/200 Upstate 4A794) 05‐521 Mre11 (Ab‐ polyclonal, cat.# N/A 1/100 Calbiochem 1) PC‐388 Rad51 (Ab‐ polyclonal, cat.# N/A 1/100 Calbiochem 1) PC130 polyclonal, cat .# Santa Cruz TopBP1 N/A 1/100 sc‐32923 Biotechnology polyclonal, cat.# Santa Cruz ATM N/A 1/100 sc‐7230 Biotechnology Rad17 (H‐ polyclonal, cat.# Santa Cruz N/A 1/100 300) sc‐5613 Biotechnology polyclonal, cat.# Oct4 1/500 N/A Abcam (Cambridge MA) 19857 polyclonal, cat.# Santa Cruz CenpA N/A 1/100 sc‐22787 Biotechnology

66 Table 3.3 – Alexa secondary antibodies used in immunofluorescence experiments. Excitation Emission Antibody* Cat. # Wavelength Wavelength (nm)** (nm)** goat anti‐chicken 488 A11039 495 519 goat anti‐rabbit 555, A21429 555 565 highly cross‐absorbed goat anti‐mouse 555, A21424 555 565 highly cross‐absorbed goat anti‐rabbit 633, A21071 632 647 highly cross‐absorbed goat anti‐mouse 633, A21052 632 647 highly cross‐absorbed *All secondary antibodies were used at a dilution of 1/200 and were purchased from Invitrogen (formerly Molecular Bioprobes) of Burlington, Canada. **Values are approximations based on the peak excitation/emission wavelengths as published in the Handbook of Fluorescent Probes and Research Products, Ninth Edition by Richard P. Haugland. Actual emissions data were recorded over a range optimized to minimize signal crossover and bleed‐through during image acquisition.

67 performed under low light conditions to minimize photobleaching. The cover slips were washed three times for 5 min per wash in PBS containing 0.1 % Triton‐X100., mounted in Antifade Gold supplemented with 4',6‐diamidino‐2‐phenylindole (DAPI;

Invitrogen) on clean glass slides, allowed to cure overnight, and sealed using nail polish. Rad9A‐GFP expressing cells were prepped in low light conditions during the entire slide preparation in order to minimize signal loss. Where indicated, coverslips were treated with Extraction Buffer (EB ‐ 50 mM HEPES [pH 7.5], 150 mM NaCl, 1 mM EDTA, 0.5 % NP‐40, Complete Mini protease inhibitors ‐ Roche), for the removal of soluble proteins, for 30 min at 4 °C prior to fixation.

Images were acquired using a Leica TCS SP2 MP inverted confocal microscope (Leica, Richmond Hill, Canada) and an 100 X oil‐immersion objective.

Images with signals from two or more different fluorochromes were captured sequentially to minimize spectral overlap. Raw TIFF images were processed and overlays of data from different channels were created in Adobe Photoshop (Adobe

Systems Canada, Etobicoke, Canada). Images presented in Figures 4.3‐4.7 and 5.1‐

5.13 are representative of the cellular population, confirmed via visual examination of at least 20 fields of view.

3.4 Immunoprecipitations and Immunoblotting

To harvest cells for immunoprecipitation (IP), cells were washed twice with ice‐cold PBS and collected via scraping on ice in 1 mL per 10 cm dish NETN buffer

(250 mM NaCl, 20 mM Tris‐HCl [pH 8.0], 0.5 % NP‐40 and 10 % glycerol)

68 supplemented with Complete Mini protease inhibitors (Roche). To shear genomic

DNA, lysates were passed sequentially through three syringes of lessening gauges.

The lysates were then precleared for at least two hours at 4 °C with mixing, with one of protein A sepharose (BioVision, Mountainview, CA), protein G sepharose

(BioVision), or IgY agarose (Aves Labs, Tigard, OR) beads, for primary antibodies raised in rabbit, mouse, or chicken, respectively. The beads were then removed via centrifugation at 228 g for 5 min at 4 °C and the pre‐cleared lysate transferred to a fresh tube. Approximately 1‐2 μg of primary antibody was added and the reactions left to incubate on a nutator at 4 °C overnight. IP was performed by addition of the appropriate bead‐conjugated secondary antibody and continuing the incubation for a further two hours at 4 °C with agitation. Samples were then washed three times by rinsing the beads with 500 μL NETN buffer and collection via centrifugation at

228 g for 5 min at 4 °C, resuspended in 60 μL NETN and 30 μL SDS‐PAGE sample buffer and subjected to SDS‐PAGE.

After SDS‐PAGE, proteins were transferred to Hybond nitrocellulose membrane (G.E. Healthcare, Baie d’Urfé, Canada) using either a semi‐dry transfer apparatus (Bio‐Rad) for 40 min at 18 V or a wet transfer in a BioRad Criterion plate transfer apparatus 18 hrs at 20 V, as indicated. Membranes were then prepared for immunoblot (IB) as follows: blots were blocked in 5 % non‐fat milk powder

(Bioshop) and 0.5 % albumin (Bioshop) in PBST (PBS containing 0.5 % Tween 20) for at least one hour at RT with rocking. Blots were then incubated in primary antibody diluted in PBS as indicated (Table 3.2), overnight at 4 °C with mixing and the following day were washed with PBST 6 times for 10 min each at RT, with

69 rocking. The appropriate horse radish peroxidase (HRP)‐conjugated secondary antibody, α‐chicken (Cedarlane Laboratories), α‐mouse (Cedarlane Laboratories) or

α‐rabbit (Biorad), was diluted in PBS and incubated for at least one hour at RT with rocking. Blots were then washed a further 6 times before a 5 min incubation in

Pierce Biolynx ECL reagent (Fisher Scientific, Ottawa, Canada) and exposure to

Kodak X‐OMAT LS film (Mandel Scientific Company, Guelph, Canada).

3.5 mRNA extraction, cDNA synthesis, and qRT­PCR

Cells were harvested and mRNA extractions performed via the RNEasy kit

(Qiagen, Mississauga, Canada) according to manufacturer’s instructions. Extracted

RNA was analyzed via spectrometry using an Eppendorf BioPhotometer (Eppendorf

Canada, Mississauga, Canada) using the Factor 10, LP 1 mm Hellma TrayCell.

Approximately 1 μg of RNA was used as a template for cDNA synthesis via the

Promega Reverse Transcription system (Fisher) according to the manufacturer’s instructions. For the qRT‐PCR reactions, Taqman probes (Applied Biosystems,

Streetsville, Canada) against murine 18S RNA, Rad9A, Rad9B were used. One hundred nanograms of cDNA was used to quantify gene expression on an Eppendorf

‐ΔΔC LightCycler and raw data was analyzed according to the 2 T method described in

Livak and Schmittgen [393].

70 Chapter 4: The Rad9A checkpoint protein is required for nuclear

localization of the Claspin adaptor protein

4.1 Rad9A and Claspin interact constitutively

It has previously been shown, via biochemical techniques, that Claspin and

Rad9A of the 911 complex interact and that this interaction is required for execution of the checkpoint arrest [193]. In order to further define this interaction, a number of different conditions were examined via IP followed by IB and IF. These included experiments to determine whether this interaction was cell cycle‐dependent, damage‐dependent, or whether elements present in the C‐terminal tail of Rad9A were required. The bulk of the following findings are reported in Sierant et al.

[105].

4.1.1 The Rad9A‐Claspin interaction is not cell cycle dependent

Given that the levels of Claspin oscillate during the cell cycle, peaking in the S and G2 phases [193], the possibility that the interaction between Rad9A and Claspin was cell cycle‐dependent was examined. HeLa cells were synchronized, lysed, and

IPs performed. Co‐IP reactions against endogenous Rad9A produced bands consistent with the reported size of the Claspin protein after probing IBs with antibodies directed against Claspin (Figure 4.1A). The reciprocal reaction, in which the co‐IP reaction against Claspin was probed with α‐Rad9A antibodies, produced the characteristic banding pattern associated with Rad9A [4, 11‐13], visible as a

71

Figure 4.1 – Rad9A and Claspin interact during all phases of the cell cycle in HeLa cells. HeLa cells were synchronized via double thymidine block and released for the indicated number of hours before being harvested in NETN buffer. A. α‐Rad9 IP followed by α‐Claspin IB (top). B. α‐Claspin IP followed by α‐Rad9A IB (bottom). Relative quantity of protein per well was analyzed relative to the IP target via densitometry and were normalized against the IP target. Values are presented as percentage of α‐Claspin signal relative to the α‐Rad9A signal for each lane.

72 series of four bands between the 48 and 63 kDa markers (Figure 4.1B). Slight variations in intensity of the bands of the secondary IBs in both panels A and B were observed and densitometry values were obtained. This was to determine if these variations were the result of cell cycle‐dependent changes in the Rad9A‐Claspin interaction or if they were the result of small differences during loading. When the densitometry values obtained from panel A where compared to the values obtained in panel B no significant variations were observed. This led to the conclusion that the slight differences in band signal strength were the result of differences in loading and not of a significant cell cycle‐dependent change in the interaction between Rad9A and Claspin.

4.1.2 Modification of the Rad9A protein does not affect the interaction with Claspin

The interaction between Rad9A and TopBP1 is dependent upon phosphorylation at the S387 constitutive site [12]. To examine if this site or if other elements in the C‐terminal tail are involved in the Claspin‐Rad9A interaction, a series of Rad9A C‐terminal truncation mutants were over‐expressed and used in IP reactions. Previously constructed C‐terminal truncation mutants in which the last

17 (Δ17), 36 (Δ36), or 59 (Δ59) amino acids have been deleted were transfected into HeLa cells along with a full‐length Rad9A construct (WT). The Rad9A NLS

(Figure 2.2) is absent in both the Δ36 and Δ59 truncation mutants [4, 10, 11, 13]. IP reactions against the N‐terminal myc epitope tag present on these constructs were performed and followed by α‐Claspin IB. Claspin‐reactive bands were detected

74 under all conditions (Figure 4.2A), indicating the Rad9A‐Claspin interaction is not dependent upon the last 59 amino acids of the Rad9A C‐terminal tail. Densitometry revealed a slight increase in band signal in the Δ59 mutant compared to loading and was attributed to both the dominant negative effects demonstrated by these mutants, resulting from the elimination of the S387 phosphorylation site, and from the deletion of the NLS [10, 12, 80].

One of the key residues in the Rad9A tail is the S272 residue, which is phosphorylated in response to IR‐induced DNA damage [11]. To investigate whether the damage‐dependent phosphorylation site at S272 was involved in

Claspin binding, a S272A point mutant was immunoprecipitated in a co‐IP reaction against the myc‐tag, while endogenous Rad9A was immunoprecipitated with polyclonal α‐Rad9 antibodies (Figure 4.2B). Expression of Rad9A‐S272A did not affect the interaction between Rad9A and Claspin (Figure 4.2B). In addition, the presence of IR‐induced DNA damage was also examined to see whether this affected the interaction between Rad9A and Claspin. In both the presence of DNA damage, there was no effect on Claspin binding when compared to the undamaged control (Figure 4.2B). Combined, the data presented in both Figures 4.1 and 4.2 suggest the Rad9A‐Claspin interaction is constitutive in at least a subset of the total

Rad9A and Claspin in a cell and does not depend on phosphorylation at either the

S272 or S387 residues.

75

Figure 4.2 – Mutations in the Rad9A C­terminal tail do not affect Rad9A­ Claspin interactions regardless of the presence of DNA damage. Asynchronously growing HeLa cells were transfected with different Rad9A mutant constructs and IP performed against either myc or endogenous Rad9A, as indicated. A. Expression of either full‐length or different truncation mutants of Rad9A results in an α‐Claspin reactive band after IB. B. Cells were transfected with different point mutations or wild‐type Rad9A and treated as indicated with 10 Gy IR followed by a 30 min recovery period. IPs were against either endogenous Rad9A (end) or the myc tag of different Rad9A constructs (WT or S272A). All samples show α‐Claspin reactive bands above the 175 kDa marker. Relative quantity of protein per well was analyzed relative to the IP target via densitometry and were normalized against the IP target. Values are presented as percentage of α‐Claspin signal relative to the α‐Rad9A signal for each lane.

76

4.2 Rad9A affects the nuclear localization of Claspin

Amino acids 356‐364 in the Rad9A C‐terminal code for an NLS (Figure 2.1), characterized by Hirai and Wang [10], and previous work has demonstrated that abolishing the Rad9A‐NLS results in aberrant protein expression [12]. The protein sequence of Claspin was obtained from the National Center for Biotechnology

Information (NCBI) database, accession number NP_071394, and entered into the

PredictNLS Online NLS database (http://cubic.bioc.columbia.edu) from the

University of Columbia’s CUBIC (Columbia University Bioinformatics Center) facility.

According to this database, there is no known NLS signal in the Claspin protein sequence. To investigate whether altering the Rad9A expression pattern affected

Claspin localization, a series of IF‐based microscopy experiments were carried out.

4.2.1 Over‐expression of a non‐nuclear form of Rad9A alters the subcellular localization of Claspin

HeLa cells were transfected with either WT or Δ59 Rad9A constructs and processed for IF against Rad9A, Claspin, and the myc epitope tag. In cells over‐ expressing WT protein, the majority of Claspin was observed in the nucleus (Figure

4.3A); however, in cells over‐expressing the truncated form of Rad9A, the majority of Claspin was observed to be cytoplasmic (Figure 4.3B). This did not change 30 min after treatment with 10 Gy IR (Figure 4.3C and D). Furthermore, these finding were consistent in both undamaged and damaged hTERT‐RPE1 transfected cells

(Figure 4.4) as well as in mES cells negative for endogenous Rad9A (Figures 4.5).

78

Figure 4.3 – Over­expression of non­nuclear Rad9A protein changes the subcellular localization of Claspin in asynchronous HeLa cells. Asynchronous cells were grown on coverslips, transfected with either WT (A. and C.) or Δ59 (B. and D.) myc‐tagged Rad9A constructs and processed for IF. DNA damage was induced via exposure to 10 Gy IR, and allowed to recover for 30 min prior to fixation (C. and D.). Transfected cells were identified by strong signals in the myc channel. Nuclei were stained with DAPI (white), while antibodies were used to detect Rad9A (green), Claspin (red) and myc (blue). The overlay consists of all channels except the DAPI signal. The white bar represents 10 μm.

79

Figure 4.4 ­ Over­expression of non­nuclear Rad9A protein changes the subcellular localization of Claspin in asynchronous hTERT­RPE1 cells. Asynchronous cells were grown on coverslips, transfected with either WT (A. and C.) or Δ59 (B. and D.) myc‐tagged Rad9A constructs and processed for IF. DNA damage was induced via exposure to 10 Gy IR, and allowed to recover for 30 min prior to fixation (C. and D.). Transfected cells were identified by strong signals in the myc channel. Nuclei were stained with DAPI (white), while antibodies were used to detect Rad9A (green), Claspin (red) and myc (blue). The overlay consists of all channels except the DAPI signal. The white bar represents 10 μm.

81

Figure 4.5 – Over­expression of non­nuclear Rad9A protein changes the subcellular localization of Claspin in undamaged, asynchronous murine embryonic stem cells. Asynchronous mES cells were grown on fibronectin‐ coated coverslips, transfected with either human myc‐tagged WT (A.) or Δ59 (B.) Rad9A constructs, and processed for IF. Transfected cells were identified by strong signals in the myc channel. Nuclei were stained with DAPI (white), while antibodies were used to detect Rad9A (green), Claspin (red) and myc (blue). The overlay consists of all channels except the DAPI signal. The white bar represents 10 μm.

83

These data indicate that a dominant negative form of Rad9A that prevents 911 nuclear localization also blocks the nuclear localization of Claspin. To determine if the interaction between Rad9A and Claspin is stable, capable of interacting with DNA and wasn’t a result of Rad9A over‐expression, we repeated these experiments removing soluble proteins prior to fixation.

Removal of soluble proteins leaves only chromatin bound proteins, which are presumed active.

4.2.2 Extraction‐resistant Rad9A forms large nuclear foci with Claspin

Soluble proteins were removed via extraction with a weak detergent solution prior to processing for IF. This experiment was performed in HeLa cells (Figure 4.6) and hTERT‐RPE1 cells (Figure 4.7) and in the absence (Figures 4.6A‐B, and 4.7A‐B) or presence of DNA DSBs (Figures 4.6C‐D, and 4. 7C‐D). As in the previous experiments, DNA DSBs were produced via exposure to 10 Gy IR followed by a 30 min recovery period, to ensure full checkpoint activation. In all cases it was found that over‐expressed Rad9A and endogenous Claspin formed large, nuclear foci resistant to extraction, regardless of Rad9A C‐terminal status. These data reveal that a fraction of the non‐nuclear Rad9A‐Δ59 is able to enter the nucleus with

Claspin despite lacking the NLS motif. Additionally, the nuclear foci indicate this interaction is resistant to detergent extraction, and therefore is chromatin‐bound, suggesting it is participating in the checkpoint response. There appears to be no significant differences between the panels in Figures 4.6‐4.7, indicating that the

85

Figure 4.6 – Extraction of soluble proteins reveals large, nuclear Rad9A­ Claspin foci in asynchronous Rad9A­transfected HeLa cells. Asynchronous cells were grown on coverslips, transfected with either WT (A. and C.) or Δ59 (B. and D.) myc‐tagged Rad9A constructs and processed for IF. DNA damage was induced via exposure to 10 Gy IR, and all to recover for 30 min prior to protein extraction (C. and D.). Transfected cells were identified by strong signals in the myc channel. Nuclei were stained with DAPI (white), while antibodies were used to detect Rad9A (green), Claspin (red) and myc (blue). The overlay consists of all channels except the DAPI signal. The white bar represents 10 μm.

86

Figure 4.7 – Extraction of soluble proteins reveals large, nuclear Rad9A­ Claspin foci in asynchronous Rad9A­transfected hTERT­RPE1 cells. Asynchronous cells were grown on coverslips, transfected with either WT (A. and C.) or Δ59 (B. and D.) myc‐tagged Rad9A constructs and processed for IF. DNA damage was induced via exposure to 10 Gy IR, and all to recover for 30 min prior to protein extraction (C. and D.). Transfected cells were identified by strong signals in the myc channel. Nuclei were stained with DAPI (white), while antibodies were used to detect Rad9A (green), Claspin (red) and myc (blue). The overlay consists of all channels except the DAPI signal. The white bar represents 10 μm.

88

Claspin‐Rad9A interaction is not affected by DNA DSBs, which confirms the data presented in Figure 4.2B. However, these figures also reveal that nuclear mutant

Rad9A signal is observed in the B and D panels of Figures 4.3‐4.7 indicating that non‐nuclear Rad9A (Δ59) is able to translocate into the nucleus of cells, evidenced by strong Rad9A and myc nuclear signal. It was hypothesized that the endogenous protein was capable of assisting in nuclear translocation by interacting with the exogenous protein. This was investigated by IP against the epitope tag on exogenous protein followed by IP for endogenous protein.

4.2.3 Rad9A molecules are able to multimerize

Asynchronously growing HeLa cells, transiently expressing WT myc‐tagged

Rad9A, were harvested and IP performed against the myc tag . Prior to harvest samples were either fixed for 10 or 20 min in PFA to crosslink the cellular proteins or not, as indicated prior to harvest and the crosslinks reversed after cellular lysis.

IB against Rad9A produces bands corresponding to endogenous Rad9A (Figure 4.8) showing that epitope tagged Rad9A is able to form a stable complex with endogenous Rad9A. These bands are present in both the unfixed and fixed samples, though are easier to visualize in the unfixed samples. This multimerization provides a possible mechanism for the nuclear localization of Rad9A‐Δ59 mutant construct.

Endogenous 911 complex could be acting as a shuttle for exogenous Rad9A and explains how both mutant Rad9A and Claspin are nuclear in the non‐nuclear samples. The logical method for testing this hypothesis was to analyze Claspin

90

Figure 4.8 – Immunoprecipitation against epitope tagged Rad9A protein reveals the presence of endogenous Rad9A after immunoblot. Asynchronously growing HeLa cells were transfected with WT myc‐tagged Rad9A and were either fixed in PFA, or not, for 10 or 20 min, as indicated. Samples were then harvested via NETN buffer and IP against myc performed. IB against both Rad9A and myc were performed. In later exposures of the α‐Rad9A IB, bands corresponding to the size and pattern of endogenous Rad9A can be seen (*).

91 localization in the absence of endogenous Rad9A. A murine embryonic stem cell

(mES) line in which the rad9A loci have been deleted was utilized in order to determine the extent of the involvement of Rad9A on Claspin nuclear localization

[87].

4.3 Rad9A/B are responsible for Claspin localization

The Rad9A‐null mES cell line is one of two of the only viable Rad9A knockout cell lines currently available, the other being derived from the chicken B lymphocyte line. Both the null mouse and murine embryonic fibroblasts are inviable and show extensive chromosomal defects as well as increased sensitivities to genotoxins [87].

It was hypothesized the viability of the mES line was a result of partial rescue of the

Rad9A paralogue Rad9B, previously thought to be restricted to tumour tissue and the testis [103, 104]. In order to determine if this was occurring, quantitative PCR technique were utilized.

4.3.1 rad9A­null mES cells express Rad9B

WT and Rad9A‐deficient mES cells were treated with RA as indicated and samples collected for qRT‐PCR against Rad9B. Figure 4.9C indicates that in both WT and Rad9A‐null cells, levels of Rad9B decrease compared to the GAPDH control with the minimum of Rad9B mRNA detected after 48 hrs RA treatment. Differentiation was confirmed via decreasing Oct4 expression, a well‐known marker of

93

Figure 4.9 – Differentiation of murine embryonic stems cells alters the localization of Claspin in the absence of Rad9A which can be restored by over­ expression of wildtype Rad9A. Asynchronously growing WT and Rad9A‐ deficient mES cells were differentiated, as indicated, by exposing the cells to 10 μM RA for the indicated time. Samples were collected for IF processing, mRNA harvest, or total protein extracts, as indicated. A. WT mES cells were differentiated for 72 hrs in RA then stained for IF against Claspin (red) and DNA via DAPI stain (white). B. Rad9A‐deficient mES cells were processed as described for A. C. mRNA samples were collected and subjected to qRT‐PCR utilizing a probe against Rad9B. qPCR results were normalized against GAPDH mRNA levels. Error bars represent the standard deviation from triplicate samples D. Cells from A and B, as well as Rad9AmycWT transfected WT and Rad9‐null mES cells were scored for the number of cells showing nuclear or cytoplasmic Claspin staining. E. IB against Oct4 on whole cell extracts shows decreasing Oct4 protein levels as exposure to RA increases.

94 pluripotency (Figure 4.9D). These data reveal the Rad9A‐null mES cell line express

Rad9B and that levels decrease upon differentiation by RA. It was also observed that cellular viability decrease as exposure to RA increased with total cell death occurring at approximately 96 hrs RA treatment. This led to examine the effect of complete Rad9 ablation on Claspin localization.

4.3.2 Differentiation of Rad9A‐null mES cells alters the subcellular localization of

Claspin but has no affect on WT mES cells

Rad9A‐deficient mES cells treated with RA for 72 show primarily cytoplasmic

Claspin whereas WT RA‐treated cells show primarily nuclear Claspin staining

(Figure 4.9A and 4.9B respectively). Approximately two hundred Rad9‐null and WT mES cells were scored for cytoplasmic versus nuclear Claspin localization and the results displayed in Figure 4.9D: 77% of WT mES cells showed nuclear Claspin staining whereas only 37% of the Rad9A‐deficient cells showed nuclear Claspin.

Reconstitution of WT Rad9A via transient transfection in Rad9‐null mES cells restored Claspin nuclear localization in 83% of the cells analyzed (Figure 4.9E).

4.3.3 Conclusions about the interaction between Rad9A and Claspin

The interaction between Rad9A and Claspin has been shown here to persist during all phases of the cell cycle, during the absence and presence of DNA damage, and not to be dependent on the S272, S336, or T355 phosphorylation sites or on the

NLS sequence of Rad9A. However, it does appear that Rad9A is responsible for

96 proper nuclear localization of the Claspin protein and that Rad9B is also capable of this. Work shown here also demonstrates that Rad9A is able to interact with other

Rad9A molecules.

97 Chapter 5: The identification and characterization of a novel nuclear

structure containing members of the homologous recombination DNA

damage response pathway

5.1 The RDF closely associate with Xi

Confocal microscopy has previously shown large, Rad9A‐dense foci (RDF) in the nuclei of a subset of HeLa cells [12]. These RDF were similar in appearance to

BRCA1 foci observed in the Livingston lab and shown to be the inactive X chromosome [391, 394]. Confocal microscopy was used to determine if the RDF were localizing to the Xi in HeLa cells. This work was recently submitted for publication and is currently under review.

5.1.1 Rad9A closely associates with the histone variant macroH2A1

To determine if the RDF were the Xi chromosome, colocalization experiments with the known Xi marker, macroH2A1 [395], were performed. HeLa cells stably expressing GFP‐Rad9A were grown on coverslips and processed for IF. Rad9A

(green) and macroH2A1 (red) form large, nuclear foci that are closely associated but do not directly overlap (Figure 5.1A). This interaction does not change in the presence of DNA DSBs (Figure 5.1B). This phenomenon is seen consistently in approximately 15% of Rad9A‐GFP expressing cells and in roughly half of HeLa cells

Figure 5.1 – Rad9A­GFP fusion protein closely associates with macroH2A1 in asynchronous HeLa cells. HeLa cells stably expressing a Rad9A‐GFP fusion protein were grown on coverslips, exposed to 10 Gy IR followed by 30 min recovery (B. only) and processed for IF. Rad9A‐GFP (green) colocalizes with macroH2A1 (red) but does not directly overlap, as demonstrated in two different nuclei per panel. The white bar represents 10 μm.

99 stained for Rad9A via polyclonal antibody. The possibility that this represented a cell cycle‐specific interaction was considered and is addressed in Section 5.2.

Further analysis of the RDF at Xi focused on determining if Rad9A colocalized with the opposite domain on Xi.

5.1.2 Rad9A colocalizes with the facultative heterochromatin marker H3trimK9

Since the Xi chromosome is divided into two domains, one dominated by macroH2A1 and the other by H3trimK9 [396], the possibility that Rad9A was localizing to the other domain of the Xi was examined. Rad9A shows significant colocalization with H3trimK9, a marker of silenced heterochromatin [397], in a close but not overlapping manner in asynchronous, undamaged HeLa cells (Figure

5.2A), similar to what is observed between Rad9A and macroH2A1 (Figure 5.2B).

Taken together, these data indicate that Rad9A has a close association to both the macroH2A1 and H3trimK9 regions of the Xi but is not directly associated with either. Given the similarities with the BRCA1 foci previously reported [391, 394], the degree of colocalization between the RDF, Xi and BRCA1 was investigated.

5.1.3 RDF‐macroH2A1 foci overlap with BRCA1 foci

Given that BRCA1 localizes to the Xi [390, 391, 394], the possibility the RDF localizes through BRCA1 was explored. Rad9A (green) colocalized with macroH2A1 staining (red), with BRCA1 (blue), the colocalization between Rad9A and BRCA1

101

Figure 5.2 – Rad9A­macroH2A1 foci and Rad9A­H3trimK9 foci do not directly overlap in asynchronous HeLa cells. HeLa cells were grown on coverslips and processed for IF. A. Rad9A (green) and macroH2A1 (red) do not directly overlap. The bottom images were subjected to 6x magnification. B. Rad9A (green) forms foci with H3trimkK9 (red) that do not directly overlap. The bottom images were subjected to 6x magnification. The white bar represents 10 μm.

102 appears to completely overlap whereas the overlap with macroH2A1 is less obvious

(Figure 5.3). At this point, we began these investigations in two other cell lines: the hTERT‐RPE1 and the semi‐differentiated 184‐hTERT. Rad9A (green) was found to form large foci in all three cell types but colocalization with macroH2A1 (red) occurred only in the HeLa cell line (Figure 5.4). This experiment also included a fibroblast cell line from a Klinefelter’s patient, which contains multiple Xi, and no colocalization between Rad9A and the Xi was observed (Figure 5.4D). Further analysis of the Rad9A (green) and BRCA1 (red) colocalization shows this overlap is present in all three cell lines examined (Figure 5.5). The presence of the RDF‐

BRCA1 colocalization in multiple cells prompted the investigation of whether other members of the DDR are present at the RDF.

5.1.4 RDF colocalize with γ‐H2AX independent of exogenous DNA damage

In keeping with analyzing other members of the DDR for localization at this focus, the ubiquitous DNA damage marker γH2AX was examined. Asynchronously growing HeLa cells were processed for IF, Rad9A (green) and macroH2A1 (blue) were detected via polyclonal antibodies, while γH2AX (red) was detected via monoclonal antibody (Figure 5.6). Samples were mock irradiated (A) or exposed to

10 Gy IR followed by a 30 min recovery period as indicated (B). In both samples,

Rad9A forms large, nuclear foci that overlap with γH2AX completely and partially overlaps with the macroH2A1 foci (Figure 5.6). γH2AX staining is punctate in damaged cells but not in undamaged cells, which conforms to previously published

104

Figure 5.3 – The RDF colocalizes with the Xi and BRCA1. Asynchronously growing HeLa cells were grown on coverslips and processed for IF. Rad9A (green) colocalizes with both macroH2A1 (red) and BRCA1 (blue). The white bar represents 10 μm.

105

Figure 5.4 – Rad9A colocalizes with macroH2A1 in HeLa cells but not in other cell types. Asynchronously growing cells were grown on coverslips and processed for IF. Staining was against Rad9A (green) and macroH2A1 (red) and shows colocalization in HeLa cells (A.) but not in hTERT‐RPE1 (B.), 184‐ hTERT (C.), or Dempsey cells (D.). The white bar represents 10 μm.

107

Figure 5.5 – The RDF colocalizes with BRCA1 in multiple cell types. Asynchronously growing HeLa cells (A.), hTERT‐RPE1 (B.), and 184‐hTERT (C.) were grown on coverslips and processed for IF. Rad9A (green) and BRCA1 (red) form large overlapping foci in all three cell types. The white bar represents 10 μm.

109

Figure 5.6 – The RDF and the Xi colocalize with a large γH2AX focus. Asynchronously growing HeLa cells were grown on coverslips, exposed to 10 Gy IR, and processed for IF. Rad9A (green) colocalizes with macroH2A1 (blue) and a large γH2AX foci (red) that is present in both undamaged (A.) and damaged cells (B.). The white bar represents 10 μm.

111 data on γH2AX behaviour [185]. When this experiment was repeated, it was found that this colocalization occurred in all three of the cell types studied: HeLa, hTERT‐

RPE1, and 184‐hTERT (Figure 5.7A­C). Data present in Figures 5.4, 5.5, and 5.7 suggests these foci are not a part of the Xi but instead represent a novel structure containing members of the DDR. Experimentation along this vein, investigating other members of the DDR was begun.

5.2 The RDF colocalize with members of the homologous recombination pathway

Due to the lack of colocalization between Rad9A and the Xi in both the hTERT‐RPE1 and 184‐hTERT cell lines and the presence of two well‐known members of the DDR, it was hypothesized that the RDF was not related to the Xi, but was a large protein super‐complex associated with the Xi in HeLa cells. In order to test this hypothesis confocal analysis was performed against members of the HR repair pathway: Mre11, Rad51, and ATM on all three cell lines: HeLa, hTERT‐RPE1, and 184‐hTERT.

5.2.1 The RDF does not colocalize with TopBP1, Claspin, or the telomeric TRF2 protein

In order to determine if other confirmed Rad9A‐interacting downstream proteins were involved, colocalization with TopBP1 and Claspin were examined

(Figure 5.8A and B). Asynchronously growing HeLa cells were prepped for IF against two known Rad9A‐interacting proteins, TopBP1 (Figure 5.8A), and Claspin

113

Figure 5.7 – Rad9A and γH2AX colocalize in a large body in multiple cell types. Asynchronously growing HeLa cells (A.), hTERT‐RPE1 (B.), and 184‐hTERT (C.) were grown on coverslips and processed for IF. Rad9A (green) and γH2AX (red) form large overlapping foci in all three cell types. The white bar represents 10 μm.

114

Figure 5.8 – The RDF does not contain TopBP1, Claspin, TRF2, or CenpA. Asynchronously growing HeLa cells were grown on coverslips and processed for IF. Rad9A (green) does not for large foci with either TopBP1 (A. – red), Claspin (B. – red), TRF2 (C. – red), or CenpA (D. – red). The white bar represents 10 μm.

116

(Figure 5.8B), as well as the telomeric marker TRF2 (Figure 5.8C) [398]. Telomeric

DNA undergoes significant processing during the cell cycle due to phenomenon of telomeric shortening during subsequent DNA replication cycles. TRF2 has also been recently shown to inhibit the ATM‐mediated DNA checkpoint at telomeres, to be required for the resolution of DNA DSBs by HR and has previously been shown to interact with Rad9A via IP [398‐400]. This protein seemed like a logical target for assessing whether the RDF was a result of telomeric processing. RDF are visible in all parts but there is no corresponding signal in the either the TopBP1, Claspin, or

TRF2 channels indicating that these proteins do not participate in this event.

Additionally, indirect IF against Rad9A and the centromeric CenpA protein was carried out and no colocalization occurred between the two proteins (data not shown). This eliminates the possibility that the RDF is related to either telomeres or centrosomes.

5.2.2 The RDF colocalizes with Mre11, Rad51, ATM, and Rad17

Since the presence of γH2AX and BRCA1 were shown to be present at this focus, other members of the HR repair pathway were also analyzed. Confocal analysis of asynchronously growing cells reveals Rad9A‐Mre11 colocalization

(Figure 5.9). This colocalization persists in all three of the cell lines examined: A.

HeLa, B. hTERT‐RPE1, and C. 184‐hTERT. This experiment was repeated with staining against Rad51 (Figure 5.10), an important downstream effector in the HR pathway previously shown to interact with Rad9A [400]. As seen in the previous

118

Figure 5.9 – Rad9A forms large foci with Mre11 in multiple cell types. Asynchronously growing HeLa cells (A.), hTERT‐RPE1 (B.), and 184‐hTERT (C.) were grown on coverslips and processed for IF. Rad9A (green) and Mre11 (red) form large overlapping foci in all three cell types. The white bar represents 10 μm.

119

Figure 5.10 – Rad9A forms large foci with Rad51 in multiple cell types. Asynchronously growing HeLa cells (A.), hTERT‐RPE1 (B.), and 184‐hTERT (C.) were grown on coverslips and processed for IF. Rad9A (green) and Rad51 (red) form large overlapping foci in the HeLa and 184‐hTERT cell lines but not the hTERT‐RPE1 cell line. The white bar represents 10 μm.

121 figure, this colocalization persists in two of the cell lines examined: A. HeLa, and C.

184‐hTERT. No colocalization was observed in the hTERT‐RPE1 cell line (Figure

5.10B). Staining was also performed against the sensor PIKK ATM (Figure 5.11), colocalization was observed between Rad9A and ATM in only two of the cell lines studied. A summary of the proteins colocalizing with Rad9A in the three different cell lines is presented in Table 5.1. From Table 5.1 the following trend is revealed:

HeLa and 184‐hTERT RDF colocalize with both halves of the HR pathway, damage detection and signal transduction. In the hTERT‐RPE1 cell line colocalization is observed only with the damage detection (γH2AX and Mre11) side of the pathway indicating a difference in the event triggering the RDF between these three cell lines.

In addition to the HR‐related proteins, colocalization between Rad9A and its clamp loader, Rad17, was assessed. Asynchronously growing HeLa cells were stained for

IF against Rad9 (green) Rad17 (red), and γH2AX (blue) (Figure 5.12).

5.2.3 Members of the RDF are not static

Colocalization between Rad9A, Mre11, and BRCA1 was carried out in order to determine the degree of colocalization between these three proteins.

Asynchronously growing HeLa cells were processed for IF against Rad9A (green),

Mre11 (red), and BRCA1 (blue). Images from four cells containing RDF show that both Mre11 and BRCA1 can be present or absent suggesting this foci is part of a dynamic process (Figure 5.13). Of the 52 RDF counted, 17 stained positive for

Rad9A alone where as 5 stained positive for both Rad9A and Mre11. There were 16

123

Figure 5.11 – Rad9A forms large foci with ATM in multiple cell types. Asynchronously growing HeLa cells (A.), hTERT‐RPE1 (B.), and 184‐hTERT (C.) were grown on coverslips and processed for IF. Rad9A (green) and ATM (red) form large overlapping foci in the HeLa and 184‐hTERT cell lines but not the hTERT‐RPE1 cell line. The white bar represents 10 μm.

124

Table 5.1 – Colocalization between members of the HR pathway and Rad9A. A summary of IF experiments in HeLa, hTERT‐RPE1, and 184‐hTERT between Rad9A and members of the HR repair pathway.

HeLa hTERT­RPE1 184­hTERT

γH2AX Yes Yes Yes

ATM Yes No Yes

Mre11 Yes Yes Yes

Rad51 Yes No Yes

BRCA1 Yes No Yes

126

Figure 5.12 – RDF colocalize with Rad17 and γH2AX in a large body in HeLa cells. Asynchronously growing HeLa cells were grown on coverslips and processed for IF. Rad9A (green), Rad17 (red) and γH2AX (blue) form large overlapping foci. The white bar represents 10 μm.

127

Figure 5.13 – The RDF are variable structures. Asynchronously growing HeLa cells were grown on coverslips and processed for IF. Rad9A (green) dense foci exist in a state of flux with both Mre11 (red) and BRCA1 (blue) as shown by the different combinations of proteins present in the large foci in separate nuclei. Four cells are shown, for each cell displayed panels show data from the individual channels (top), the different combinations of two‐ way colocalization (middle) and the overlay of all three channels (bottom). The white bar represents 10 μm. A total of 52 RDF were examined and the percentage of RDF in combination with Mre11 and BRCA1 is presented in a pie chart.

129 foci that stained for both Rad9A and BRCA1 and 14 that were positive for all three proteins. These data are displayed at the bottom of Figure 5.13 in pie chart form.

5.2.4 Summary of the RDF

These experiments have yielded the following data on the RDF: it is present in multiple cell types and appears to be distinct from the Xi. It is not affected by IR‐ induced DNA damage but contains a number of prominent members of the HR DNA repair pathway as well as the DNA damage marker γH2AX in asynchronous, undamaged cells. Preliminary evidence also suggests the RDF may be the product of a dynamic process and may be loaded onto DNA via Rad17/RFC. Further experimentation will attempt to determine if this structure is related to the cell cycle.

5.3 The RDF is altered after perturbations to the cell cycle

In asynchronously growing cells, the RDF can be found in up to 28 % of HeLa cells, 5.5 % of hTERT‐RPE1 cells, and 18 % of undifferentiated 184‐hTERT cells.

Based on this, it was hypothesized the RDF may be a result of late‐stage DNA replication and cell synchronization experiments were carried out to determine if this hypothesis was valid.

131 5.3.1 Cell synchronization reduces the number of RDF

Time course experiments with both thymidine and HU show the levels of foci decrease rapidly in the first few hours of synchronization (Figure 5.14A) and do not return to normal levels until approximately 72 hours post‐release (Figure 5.14B).

Figure 5.14A also shows that the number of RDF detected decreases within a few hours of the start of synchronization and that an 8 hour release fails to restore the number of foci present in synchronized cells. This suggests the foci are related to repair as inhibition of DNA replication via both thymidine and HU causes these foci to decrease rapidly. However, it was not possible to determine any dependency on the cell cycle as the RDF did not return until approximately 48‐72 hrs post‐release.

Inhibition of HR and the presence of DNA crosslinks, capable of producing both ssDNA and DSB DNA damage, were examined.

5.3.2 Treatment of asynchronous HeLa cells with either cisplatin, pentoxifylline, or both affects foci formation

Cisplatin (CP) is a potent DNA damaging agent capable of producing intra‐ and interstrand crosslinks, halting DNA synthesis. Pentoxifylline (PTX) is an inhibitor of HR‐mediated repair via inactivation of the PIKK kinases in a poorly understood mechanism [401]. Treatment of asynchronously growing HeLa cells with increasing doses of either CP, or PTX, or both changes the number of RDF present. As can be seen in Figure 5.14C, treatment with 1 mM PTX increases the number of foci over 200 % compared to untreated asynchronous cells. Treatment

132

Figure 5.14 – The RDF disappears after attempts at synchronization via exposure to thymidine and hydroxyurea but is enriched after exposure to pentoxifylline. A. HeLa cells were grown on coverslips and exposed to 2 mM thymidine (squares) or 2 mM HU (diamonds), as indicated. Cells were released from double thymdine block for the indicated times or released after exposure to 2 mM HU for 18 hours. The number of RDF were scored from approximately 300 cells and graphed relative to asynchronous samples. B. Cells were synchronized and processed as in A but were released for either 12, 24, 48, or 72 hours, as indicated. C. Asynchronously growing HeLa cells were grown on coverslips and treated with either 1, 2, 3 mM PTX (diamonds) or 10 μM, 25 μM, or 50 μM CP (squares) or 1 mM PTX and 10 μM CP, 2 mM and 25 μM CP, or 3 mM PTX and 50 μM CP (triangles), for 24 hours prior to fixation and processed as in A.

133 with 2 mM and 3 mM doses decreased the number of foci to 175 % and 152 % respectively. Treatment with CP at either 10 μM, 25 μM, or 50 μM resulted in a severe decrease in the number of RDF detected. Cells exposed to both CP and PTX at doses of 10 μM and 1 mM, 25 μM and 2 mM, and 50 μM and 3 mM respectively, also resulted in decreased numbers of foci compared to asynchronous cells. The induction of these foci by PTX indicates that the HR pathway is indeed involved in, at least, the creation of these foci.

5.3.3 Conclusions about the RDF

This section demonstrated the presence of Rad9A, and other key members of the HR pathway, in a large nuclear focus that is ablated during cell synchronization attempts. This focus is not affected by the presence of IR‐induced DNA damage however treatment with the DNA cross‐linker CP also causes these foci to disappear.

Treatment with a PIKK inactivator, PTX, resulted in a massive induction of these foci. The results here demonstrate that these foci are not the result of a static process and involve members of the HR pathway. Further investigations are required to fully elucidate the function of the RDF and are addressed in the next section.

135 Chapter 6: Discussion

In this section I describe two novel aspects to the Rad9A checkpoint protein: as a nuclear transport mechanism for the adaptor protein Claspin, and as a component of a large protein mega‐complex with ties to the HR repair pathway.

These new roles reinforce and further define Rad9A as a vital component in the maintenance of genomic integrity and structure.

6.1 Rad9A and Claspin

Claspin and Rad9A were previously shown to interact and this interaction is vital for the activation of the Chk1 DNA damage response kinase [86, 90, 91, 107,

193]. The investigations described here into the Rad9A‐Claspin relationship revealed that the association persists in a variety of conditions studied and can be considered constitutive in a significant fraction of the Rad9A and Claspin present in a cell (Figures 4.1 and 4.2). It was also discovered that both Rad9A and Rad9B can affect the nuclear localization of the Claspin adaptor protein, resulting exclusion of

Claspin from the nucleus upon disruption of the Rad9A NLS in a stable, damage‐ and cell line‐independent manner (Figures 4.3‐7 and 4.9). Reconstitution of WT human

Rad9A protein in a mouse Rad9A/B‐deficient background appears to reverse this shift from predominantly cytoplasmic in untransfected cells to predominantly nuclear in transfected cells, recreating the effect of endogenous Rad9A/B on Claspin localization (Figure 4.9). This finding also confirms the conclusions made by

136 Hopkins et al. demonstrating the human protein was able to complement murine

Rad9A [87].

Initial experiments with the Rad9A‐null mES cells revealed no change in

Claspin localization when compared to WT mES cells. Subsequent experimentation with these cells revealed the presence of Rad9B mRNA that decreased after treatment with RA, a potent initiator of differentiation (Figure 4.9C). One could postulate the undifferentiated mES cells are able to survive due to partial rescue by substitution of Rad9B in the 911 complex but as levels decrease during differentiation the true phenotype of Rad9 ablation is revealed. The Rad9A‐null cells have been shown to be sensitive to genotoxic agents so the rescue by Rad9B is not complete [87]. Rad9A‐deficient mES cells treated with 10 μM RA for 48 hrs show the lowest levels of Rad9B mRNA, at 72 hrs the mRNA levels are comparable to the 24 hr levels (Figure 4.14C), which is attributed to the death of cells expressing low levels of Rad9B and the survivability of a slower differentiating or differentiation‐resistant population of cells.

Defining the conditions of the Rad9A‐Claspin interaction is important for a complete understanding of the events occurring during the Chk1‐mediated DNA damage response. In this thesis the interaction between two key members of the

Chk1 activation complex was confirmed, depletion of Rad9A or Claspin ablates the

Chk1 damage‐induced phosphorylation at S345 [52, 56, 86, 89, 98, 107, 193, 195‐

198]. Cytoplasmic sequestration, via deletion of the Rad9A NLS, effectively depletes both Rad9A and Claspin in the nucleus (Figures 4.3‐7) and, according to reports,

137 prevents activation of the Chk1 effector kinase. This effectively renders the Rad9A‐ null mES cells Claspin‐null as well and explains the degree of lethality encountered during the course of working with these cells.

Both the 911 complex and Claspin are ring‐shaped molecules, one possibility concerning the structure of this interaction is that these rings stack along the damaged DNA strand, which would then act as locator signals to other members of the checkpoint and DNA repair machinery [9, 75, 80‐82, 402]. Future work on this interaction could focus on determining if Claspin is able to interact with the other members of the 911 complex, Hus1A and Rad1, and would reveal whether this interaction is based on the PCNA‐like ring‐shaped domains, which suggests stacking, or based on other elements present in the Rad9A C‐terminal that were not examined here. Other future experiments could include examination of the mechanism behind

Claspin loading as this has not been addressed in the current literature.

During the course of these experiments, the observation that non‐nuclear forms of Rad9A were detected in the nucleus was repeatedly made (Figures 4.3‐4.7).

These cells still express endogenous Rad9A protein, so it was concluded that endogenous protein was able to interact with both Claspin and the exogenous

Rad9A protein, shuttling both across the nucleus. This is supported by data displayed in Figure 4.8, showing the presence of α‐Rad9 reactive bands consistent with the size of endogenous Rad9A on IBs after α‐myc IP suggesting that the 911 complex could interact with other 911 complexes. It was observed that Rad9A eluted three times from gel filtration columns in complexes that corresponded to

138 approximately 100 kDa, 500 kDa, and 670 kDa (B. Besley, M.Sc. thesis, Queen’s

University). It was also found that the relative abundance of these complexes did not change during cell cycle progression. Combining his findings with the data displayed in Figure 4.8 strongly suggests the 911 complex is able to interact with other 911 complexes including those comprised of endogenous Rad9A protein in a

“super‐complex”. This means that even small amounts of endogenous protein are able to participate in the mutant 911 complex, restoring some 911 functionality.

Taken together, these data suggest the ability of Rad9A, and presumably the 911 complex, to interact in higher order complexes containing multiple 911 complexes.

Previous work has shown that the circadian proteins Tim and its partner

TIPIN are required for proper localization of Claspin [200]. Work described here neither supports nor refutes their conclusions and the possibility exists that

Tim/TIPIN and Rad9A interact with different pools of the Claspin protein. There is mounting evidence supporting the roles of circadian proteins during the DDR [200,

403], so the possibility of an interaction between the 911 complex and Tim/TIPIN should be examined in the future. A simple α‐Rad9A IP followed by IB against Tim and TIPIN in asynchronous cells, both in the presence and absence of DNA damage would reveal if there are pools of Rad9A interacting with these proteins. Confocal analysis of cells treated with EB, removing soluble proteins, against Rad9A, Claspin, and either Tim or TIPIN would reveal whether Rad9A is interacting with a different pool of Claspin than Tim/TIPIN or could reveal that these protein are all interacting together during the DDR.

139 The finding that Rad9A can affect Claspin localization has implications for proper checkpoint function. As previously mentioned, Claspin is required for proper activation of the Chk1 effector kinase during the DDR and so proper localization of this protein is an important requirement for maintaining genomic integrity [90, 193, 195, 196]. Rad9A and Claspin also interact during the DNA synthesis phase and it is known that Claspin is required for proper genomic replication to occur, monitoring DNA synthesis and regulating replication fork rates

[60, 62, 404]. From this it is easy to postulate that Rad9A‐mediated Claspin localization is also an important event during S phase as well. Work demonstrated in this thesis also confirms previous reports that Rad9B is able to form functional

911 complexes, which provides a mechanism for the survival of the mES cells [104].

These cells are able to bypass this lethality in a mechanism that requires further investigation.

The Rad9A‐null mES cell line is a valuable tool in this field, in spite of the presence of Rad9B mRNA [105]. As previously mentioned, this cell line is sensitive to genotoxins due to partial rescue by Rad9B [87], but mRNA levels decrease after differentiation [105]. By creating stable cell lines expressing different Rad9A point mutants, such as the damage‐dependent S272A mutant or the S387A constitutive mutant, the exact nature of these post‐translation modifications can be determined without interference from either endogenous WT protein or Rad9B. Treatment of these cell lines with RA will result in a decrease in Rad9B, as previously mentioned, but will circumvent the cell death associated with complete Rad9A ablation.

Analysis of genotoxin‐treated cells with respect to Chk1 and Chk2 activation will

140 provide data on the importance of these particular Rad9A phosphorylation sites during the DNA damage response and will reveal if the DNA damage signal can be relayed down the checkpoint pathway.

The work discussed in this section shows that the initial hypothesis, which stated that the Rad9A‐Claspin interaction was important for downstream signaling events, is true. However, data presented here demonstrates that post‐translational modifications to the Rad9A C‐terminus do not affect the Rad9A‐Claspin interaction, as previously stated. Instead, the use of the dominant negative Δ59 Rad9A mutant, combined with the data obtained by exploiting the mES cell lines points to a role for

Rad9A in Claspin localization. Future experiments involving Rad9A‐null mES cells stably expressing different Rad9A isoforms, both undifferentiated and differentiated, will confirm this result and will open the door for further experimentation involving the different phosphorylational point mutants available to the Davey lab. Understanding the details of this interaction is important for defining the function of Rad9A and the 911 complex during the DNA damage response, which is important, particularly in light of new evidence supporting the role of Rad9A as a potential oncogene, upregulated in both non‐small cell lung cancer and breast cancer [112‐115].

6.2 The RDF contains members of the HR DNA repair pathway

During work with a stable cell line expressing a Rad9A‐GFP fusion protein, a large, dense nuclear focus was observed in a subset of the cells examined. One of

141 these bodies can be seen in a previously published work [12]. We found these RDF were similar to foci reported by Ganesan et al., which have been proposed to be the result of interactions between BRCA1 and the Xi [391, 394]. Based on this, investigations into the possibility of Rad9A involvement at Xi were begun. However, as work on this focus progressed it became apparent that it was not the Xi but instead was a novel structure containing a number of proteins from the HR arm of the DDR (Figures 5.3, 5.5‐7, and 5.9‐11). Additionally, initial investigations utilized a stable HeLa cell line expressing a GFP‐Rad9A fusion protein, while later work used indirect IF and an α‐Rad9A polyclonal antibody to visualize the RDF formed by endogenous Rad9A protein. The occurrence of foci in the GFP‐Rad9A cell line was in approximately 15 % of cells whereas by utilizing indirect IF, the number of foci detected increased to almost 30 % of cells. This difference is likely the result of the massive over‐expression of the GFP‐Rad9A fusion protein, which masks the staining patterns of the endogenous protein.

Further investigation revealed that the RDF colocalized with both BRCA1 and

γH2AX in multiple cell types (Figures 5.3, 5.5‐7). After analysis of the RDF in four different cell types, it was found that colocalization with Xi was restricted to HeLa cells (Figures 5.4A­C). Furthermore, no colocalization between the Xi and Rad9A occurred in the Dempsey cell line, a fibroblast line derived from a patient with

Klinefelter’s syndrome and containing multiple Xi bodies (Figure 5.4D). It should be noted that even though RDF have yet to be identified in these cells, it is possible that this is a direct result of the low Rad9A levels in these cells resulting in weak IF signals. Due to the presence of these foci in other cell lines, this foci is most likely a

142 structure relating to DNA editing and not the Xi. Combined, this was enough to shift the analysis of the RDF away from the Xi and towards investigations involving DNA repair proteins. Future investigations could focus on determining why these foci happen to locate to the Xi in HeLa cells but not in the other cell types. The close association between Rad9A and the Xi in HeLa cells could be a result of the aberrant nature of Xi in these cells, which have been shown to lack XiST [405]. Examination of the HeLa cell Xi in comparison to the hTERT‐RPE1 Xi could reveal the reason for this close association.

Currently in the literature, there are two conflicting views on the involvement of BRCA1 at Xi. Work out of the Livingston lab provides evidence for the role of BRCA1 in XiST maintenance and gene silencing whereas work by another group, described in the 2007 paper by Xiao et al. [406], provided data that refuted these claims [391, 394, 407]. Xiao described nuclear BRCA1 foci that closely associate with XiST but did not directly overlap [406]. In addition, they go on to reveal that these foci also closely colocalize with γH2AX. Data displayed in Figures

5.5 and 5.7‐13 could be the structure reported by Xiao et al. [406]. These data show overlap between Rad9A, BRCA1 and γH2AX. One could postulate that the very close association displayed in Xiao’s work between BRCA1 and γH2AX could be bridged by Rad9A and the 911 complex and this is supported by data displayed in Figures

5.3, 5.5‐7, and 5.9‐11. Obviously, further work is needed to address the role of

BRCA1 at the Xi and determine the extent, if any, of interaction between Xi and the

RDF.

143 As previously mentioned, the RDF contain a number of proteins from the HR pathway. These include ATM, γH2AX, Mre11, Rad51, BRCA1 and Rad17 (Figures 5.7 and 5.9‐12). These proteins represent important members of the sensor and transducer parts of the archetypal checkpoint response. ATM and MRN are both well‐known sensors in addition to Rad9A and the 911 complex [43, 47, 50, 81, 160,

408]. ATM is primarily a signaling/transduction molecule most likely involved in the creation of γH2AX at the RDF as well as possibly recruiting downstream proteins capable of affecting cell cycle progression. The 911 complex is most likely involved in the localization of other members of the RDF, based on its requirement for the localization of both Claspin and TopBP1 during the DDR [12, 105]. The MRN complex could be performing a number of functions, possibly as a DSB sensor for

ATM [160], it is also capable of endonuclease function as well as tethering DNA free ends via Rad50 [319]. MRN signaling has also been shown to activate pathways that could lead to either cell cycle arrest or apoptosis [154]. Though only Mre11 was examined, the other components of MRN, Rad50 and Nbs1, are presumed present.

BRCA1 and Rad51 are well‐known members of the BASC, along with MRN, and, imply significant repair ability at the RDF as well as potential helicase activity [220].

The presence of Rad51 also implies the presence of BRCA2 as BRCA2 protects

Rad51 from degradation [335]. These two proteins, as previously mentioned, are important catalysts for strand invasion during HR [331, 332], suggesting such an event could be occurring at the RDF. Taken together, the proteins present at the

RDF represent important members of the entire DDR, from detection to resolution, and suggest a possible genomic rearrangement function at this structure.

144 Colocalization with the 911 clamp‐loader Rad17 suggests that these foci are being loaded onto DNA at these sites (Figure 5.12). Clamp loading could be an indication that these structures are the result of an active process and reinforces the hypothesis that these structures are not static. There is now ample evidence supporting the idea that these foci are the result of an active process involving members of the checkpoint and DNA repair machinery (Figure 5.8, 5.12‐13 and

Table 5.1). However, future analysis of the RDF could include confirming active loading. This could involve utilizing an antibody specific for phospho‐Rad17 in IF experiments with the RDF, since reports claim that phosphorylated Rad17 is exclusively bound to chromatin and considered active [240]. This, combined with the data displayed in Figure 5.13, suggest that this structure is dynamic and led to the conclusion that it changes over time.

The results summarized in Table 5.1 provide additional evidence that the

RDF is not an absolute structure but that the members vary between cell types in addition changing over time. It is interesting to note that the proteins tested thus far can be broken down into two categories: signal detection and signal transduction. Rad9A, Mre11, and ATM are all members of the signal detection side of the DDR whereas BRCA1, γH2AX, and Rad51 are all members of the signal transduction/resolution side of the DDR. It seems that the undifferentiated or less differentiated cell lines, HeLa and 184‐hTERT, colocalize with both categories. In contrast, the fully differentiated hTERT‐RPE1 cell line only seems to colocalize with the signal transducer side of the DDR. Together this suggests there are differences in the process occurring at the RDF resulting from differences in cellular

145 differentiation. In order to determine if this indeed the case, future experiments could focus on determining whether members of the RDF change during cellular differentiation. The semi‐differentiated 184‐hTERT cell line is capable of further differentiation during culture on a 3D matrix such as Matrigel. This coupled with indirect IF at different times during culture could be used to determine if the members of the RDF change as cellular differentiation occurs.

The variability displayed in the chart in Figure 5.13 is the final piece of evidence suggesting the members of the RDF are involved in an active process. The fewest number of foci were found to be positive for Rad9A and Mre11 (5/52 foci scored) whereas the greatest number of foci were positive for Rad9A alone (17/52).

Foci positive for either Rad9A and BRCA1 or all three proteins were at similar levels to those of Rad9A alone (16/52 and 14/52, respectively). Based on this data, the following model is proposed: the initial insult is detected by MRN, based on data supporting MRN as a detector of DNA DSBs [409]. Rad9A and MRN associate early and this event is followed quickly by recruitment of BRCA1, due to the low percentage of RDF positive for only Mre11 and the higher percentage positive for

Rad9A, Mre11 and BRCA1. At this point, MRN dissociates, leaving Rad9A and

BRCA1, a relatively slow event given the high proportion of triple‐positive foci. This is followed by dissociation of BRCA1 leaving only Rad9A‐positive foci, which also occurs slowly (Figure 6.1A). However, given the data presented here, we must also consider the possibility that the Rad9A‐only positive foci could exist prior to MRN recruitment (Figure 6.1B). In this modified model, the 911 complex would exist at the RDF prior to MRN and act to recruit MRN. It is possible that ATM could act as a

146

Figure 6.1 ­ Models describing the order of recruitment events at the RDF. Based on data presented in Figure 5.13, the following two models are presented which describe the possible order of events at the RDF. A. The event triggering the RDF (star) is detected by MRN, which recruits the 911 complex. This is followed by BRCA1 recruitment and MRN dissociation. BRCA1 then dissociates leaving the 911 complex. B. In this version, the event is detected by the 911 complex, which then recruits the MRN complex followed by BRCA1. MRN is the first to dissociate followed by BRCA1. The 911 complex is left at the RDF. The percentages are derived from data presented in Figure 5.14.

147 sensor for the RDF and future experimentation could focus on determining how

ATM fits into this model [43, 47, 50, 408]. Future work could also focus on determining when Rad51 is recruited, based on existing literature, it was assumed to be recruited along with BRCA1 [410‐412]. It is known Rad9A is activated quickly after damage and remains active for up to 20 hours post‐damage [12, 13]. It’s also known that Rad9A acts as a loading dock for both TopBP1 and Claspin during DNA repair [12, 13, 105]. Taken together, these data support a role for the 911 complex as the docking component of the RDF.

Cell synchronization results in a sharp decrease and eventual abolishment of these foci within hours of the initial exposure for more than 48 hrs, limiting our ability to determine the recruitment order for the HR proteins involved (Figure

5.14A and B). Treatment of HeLa cells with either PTX, CP, or both, affected the numbers of RDF found in a population of cells (Figure 5.14C). It should be noted that the overall levels of Rad9A protein were not affected by these drugs but rather the number of large, dense foci counted in a population cells was upregulated. As shown in Figure 5.14C, treatment with 1 mM PTX resulted in a massive increase in the numbers of foci counted, increasing both the number of cells containing the RDF and, in some cases, the number of foci found per cell. Exposure to higher levels of

PTX resulted in an increase in the number of RDF but in a lesser manner than 1 mM

PTX. PTX inhibits HR via inactivation of the ATM sensor kinase and can result in G2 checkpoint bypass [401, 413]. The smaller numbers of foci counted in the 2 and 3 mM doses of PTX could be related to increased apoptosis resulting from the checkpoint‐bypass. Treatment with CP or PTX and CP together caused complete

149 ablation of these foci but did not change the intensity of the Rad9A signal observed.

CP is a known inducer of DNA crosslinks and PTX is able to abolish HR‐mediated repair via inhibition of the ATM sensor kinase [401, 413]. If treatment of cells with

CP results in cell cycle arrest, it is possible that the ablation seen could be by the same mechanism that results in the disappearance of these foci following synchronization via thymidine or HU (Figures 5.14A and B).

The massive increase in the number of RDF following PTX treatment is attributed to arrest of the HR‐like event partway into processing and preventing resolution of these structures. Future work could focus on increasing the numbers of these foci via PTX and assessing whether any of the above proteins are present and whether treatment of hTERT‐RPE1 or 184‐hTERT cells results in an increase in the number of foci. If numbers do increase, it would be interesting to see if the profile, displayed in Table 5.1, is consistent after PTX treatment or if induction of these structures results in a different set of proteins being recruited. Additionally, a series of time course‐based IF experiments following cells after exposure to PTX could reveal the order in which the HR proteins are recruited providing a timeline for the formation of these structures.

Combined the above data refute the initial hypothesis that the RDF were related to heterochromatin maintenance at the Xi. Instead this hypothesis should be revised to address the possibility that these foci are the result of a DNA processing event, recruiting members of the HR pathway that is conserved across cell lines, though with small differences. One possible hypothesis, taking the above data into

150 account, is that this foci is the result of a DNA repair process, most likely related to

HR, and that the focus is actually excised, damaged DNA and associated proteins resulting from said repair process, which is awaiting export into the cytoplasm wherein the components will be degraded or recycled. Another possible hypothesis, based on the fact that Rad9A may be involved in immunoglobulin switching with

BRCA2 and Rad51 in chicken B lymphocytes [414], is that the RDF is related to HR‐ mediated genomic rearrangement events. This is supported by data from Saberi et al. who reported that HR‐mediated resolution of replication defects caused by the creation of abasic sites required both the 911 complex and Rad17 but that HR repair was unaffected in Rad9A‐null cells [414]. This is further supported by data showing that camptothecin or IR‐induced DSB require Hus1A and the HR proteins for resolution [109, 110]. This suggests involvement of the 911 complex with HR proteins in functions that are separate from traditional HR repair and corroborate the hypothesis that the RDF is related to HR but not necessarily repair. This could also explain the elevated numbers of RDF detected in HeLa cells since these cells show high levels of genomic instability.

The work described in the thesis provides the basis for a novel direction of investigations concerning the roles Rad9A and the 911 complex during cellular replication. This novel function appears to be related to DNA editing or repair and involves members of the HR repair pathway. Given the importance of both Rad9A and HR in maintaining genomic integrity, these investigations will prove to be a vital first step in unraveling the processes associated with the maintenance of genomic integrity.

151

6.3 Exploring the role of Rad9A in tumour cells

In addition to the proposed experiments discussed above, future work should focus on unraveling the exact nature of the Rad9A‐protein interactions and their importance in tumour development, especially in light of the potential protective effect of Rad9A on tumour cells [115]. There is also evidence that Rad9A‐ haploinsufficiency increases the transformative potential of a cell [415, 416]. These two seemingly opposite aspects of Rad9A are actually related as the transformation process may require that Rad9A is first downregulated, perhaps via monoallelic inactivation [415, 416], in order to bypass checkpoint events to permit transformation. Post‐transformation, Rad9A levels would increase, perhaps via gene amplification [115, 417], to afford transformed cells the protective effect associated with the over‐expression of oncogenes. Elucidating the actual function of

Rad9A and the 911 complex will be much more difficult, due to the importance for this complex during the cell cycle, as was demonstrated in this thesis and in published literature. Clearly, investigations involving the possible protective effect of Rad9A over‐expression need to be addressed.

The growing availability of tissue arrays will make analyzing large numbers of tumour samples for Rad9A mRNA levels much more efficient as well as will reveal whether tumours from specific origins have a particular Rad9A expression pattern.

Additionally, analyzing these arrays for Rad9B levels could reveal whether the

Rad9A paralogue is playing a role in tumour development. If these experiments

152 reveal that Rad9A does indeed have a protective effect then high throughput screens for small molecular inhibitors could be implemented and lead to adjuvant therapies for cancer treatments. Decreasing Rad9A/B levels in tumours cells, based on previous work by Dr. Davey’s lab and the data derived from the murine Rad9A‐null cells [87], appears to render cells more sensitive to genotoxins and could improve the efficacy of current oncotherapeutic regimes.

6.4 Concluding Remarks

The work demonstrated in this thesis shows two novel aspects to the Rad9A protein, slowly being revealed as a key component in the checkpoint signaling cascade, and furthers the field of the DNA damage checkpoint. The first is as the nuclear localization mechanism for the Claspin adaptor protein, required for proper checkpoint execution. The second is as a focal point for a number DNA repair proteins in a novel cellular structure. Difficulties arising during the study of Rad9A are the direct result of its importance in the cell cycle. Clearly, novel conditional knock‐out and reconstitutive assays must be developed, as the typical ablation studies only result in cell death. The work described here, as well as in the literature, demonstrate the importance in unraveling the role of Rad9A and the 911 complex during the DDR and in cancer.

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177 Appendix A: Generation of a conditional rad9A‐null cell line

A.1 hTERT­RPE1 cells and genomic rad9A and Cre recombinase­mediated excision

The initial focus of this thesis project was the construction of a conditional rad9A­null stable cell line in the hTERT‐RPE1 cell line, this project was eventually dropped due to time constraints. However, this project may still have uses in the future as it provides a way to create Rad9A point or truncation mutants under the influence of the native promoter and in the complete absence of endogenous protein. The hTERT‐RPE1 cell line was chosen for its genomic stability and functional checkpoints. Possessing a normal karyotype in more than 90% of cells coupled with the fact that this cell line is not tumour derived made it an ideal cell line for this project. A conditional knockout cell line is necessary as inactivation of

Rad9A has proven lethal.

The rad9A gene is located on chromosome 11 and spans a 13kb stretch of

DNA consisting of 10 exons and 9 introns. It also possesses both 5’ and 3’ untranslated regions (UTRs), both of which were included in the targeting constructs (described in section A.2). Deletion of exons 3‐5 results in a premature stop codon at the junction of exons 2 and 6 after mRNA splicing and produces a truncated protein product 49 amino acids long, of which only the first 35 would be identical to Rad9A (Figure A.1). This truncated protein is presumed nonfunctional, unable to interact with neither Hus1A nor Rad1, as it is missing the majority of the

PCNA‐homology domain and the NLS, preventing nuclear translocation.

178 Cre recombinase (causes recombination) can be used to manipulate sections of DNA in a controlled manner in different ways, depending on the type and orientation of lox (location of X‐over) sequences, Cre recombinase recognition sites.

Two identical lox sequences in the same orientation and on the same section of DNA will cause excision of the section of DNA located between them; inversion of one of the lox sequences will cause the entire section of DNA to be reversed. Finally, two different lox sequences in the opposite orientation and on two separate pieces of

DNA will result those two sections of DNA exchanging places via recombination between like lox sequences (Figure A.2 ‐ reviewed in [1]). There are several varieties of lox sequence, including loxP and lox511, both of which are used in this project, and recombination occurring between these two lox sequences non‐ existent. Cre‐mediated excision of one allele followed by Cre‐mediated swapping of the other allele is the basis for the creation of Pop‐In cell lines and has been documented in the literature using the loxP and lox511 sequences2.

A.2 Constructs and methods

The procedure for the creation of this cell line is as follows: genomic DNA was isolated from actively growing hTERT‐RPE1 cells and was used as a template for the construction of the targeting vectors. Genomic Rad9A was amplified in three parts in order to ensure proper placing in the vector backbone, pPGKneo+LS5

(primers used are listed in Table A.1). This backbone includes a neomycin resistance (neoR) cassette, conferring resistance to the antibiotic G418, allowing for

179 selection via drug resistance. All vectors are described below and are summarized in Figure A.3. Briefly, the first targeting construct (pPop1) consists of the entire genomic sequence, including both the 5’ and 3’ UTR, with the neoR cassette, flanked by loxP sequences, in intron 2 and a third loxP site, with a unique restriction site, in the middle of intron 5. The second targeting construct (pPop2) was identical to the first with the following exception: the loxP sequence in intron 5 was replaced with a lox511 sequence. The third construct, pPop3, required for the Pop‐In of different

Rad9A mutants, consists of altered rad9A genomic sequence and a GFP cassette, powered by its own promoter, for selection of positive clones via either microscopy or flow cytometry, flanked by a loxP sequence at the 5’ end and a lox511 sequence at the 3’ end.

To create the base cell line, pPop1 would be transfected into hTERTs, selected with G418, and successful HR identified via PCR (described in detail in section A.3). The resulting cell line would be rad9A+/flox and G418 resistant, treatment of this cell line with adenovirus expressing Cre recombinase (AdCre ‐

Vector BioLabs, Philadelphia, PA) would delete exons 3‐5 and the neoR cassette rendering this cell line both rad9A+/­ and G418 sensitive. This cell line is now the base for the creation of the final cell line. The initial transfection, selection, and confirmation steps would be repeated as previously described, substituting pPop2 for pPop1 and omitting the AdCre step, producing the base cell line, rad9A­/pop

(Figure A.4).

180 A.3 PCR confirmation strategy

PCR confirmation was based on two separate primer sets, the first was for inclusion of the neoR cassette in intron 2 with the 5’ primer outside the region of homology and the 3’ primer inside the neoR cassette. This reaction will only produce product if the 5’ end of the targeting construct has successfully recombined into the gene. The second PCR reaction to confirm successful recombination at the 3’ end of the gene utilized primers spanning the lox sequence in intron 5. This reaction will occur in both wildtype and recombined cells so doubles as an internal positive control; however, only the recombined cell lines will have the unique restriction site. This double product strategy is displayed in Figure A.5.

It was not possible to screen these recombinational events via Southern blot, the products produced by restriction digest were either too small (<1 kb) or too large (>8kb), both of which are not amenable to the Southern blot procedure. A suitable combination of restriction enzymes was unable to be identified and so the

PCR strategy was developed as an alternative. The current PCR strategy has allowed room for the creation of a nested set of PCR primers as a further method of confirming the cell lines. However, primers were not designed as the project was stopped before their need arose.

A.4 Current status

The constructs for this strategy are more than 90% complete with only simple cloning and sequencing, to ensure the correctness of the plasmids, left to be

181 completed. The pPop1 vector is wholly finished as only requires sequencing of the loxP sequence to ensure the unique SnaBI restriction site is outside of the loxP sequence (opposed to internally, which abrogates recognition by Cre recombinase).

The pPop2 vector is also wholly finished and in need of sequencing to ensure successful placement of the lox511/SnaBI oligonucleotide. The final vector, pPop3, is basically complete, possessing a GFP cassette, a loxP sequence flanking the 5’ end of the multi‐cloning site (MCS) and a lox511 sequence at the 3’ end of the GFP cassette. This vector still needs to have genomic rad9A elements cloned into the

MCS (the choice of elements depends on the desired final cell line and can range from single C‐terminal point mutants to a complete null cell line) and also needs to have the placement of the lox oligonucleotides and GFP confirmed via either PCR or restriction digest mapping.

A.5 Appendix A­specific References

1. Nagy, A. Cre recombinase: the universal reagent for genome tailoring. Genesis 26, 99‐109 (2000). 2. Soukharev, S., Miller, J. L. & Sauer, B. Segmental genomic replacement in embryonic stem cells by double lox targeting. Nucleic Acids Res 27, e21 (1999).

182 rad9A protein FLOX MKCLVTGGNVKVLGKAVHSLSRIGDELYLEPLEDGGSGGGCSALLSCT------rad9A protein MKCLVTGGNVKVLGKAVHSLSRIGDELYLEPLEDGLSLRTVNSSRSAYACFLFAPLFFQQ *********************************** * . *. rad9A protein FLOX ------rad9A protein YQAATPGQDLLRCKILMKSFLSVFRSLAMLEKTVEKCCISLNGRSSRLVVQLHCKFGVRK rad9A protein FLOX ------rad9A protein THNLSFQDCESLQAVFDPASCPHMLRAPARVLGEAVLPFSPALAEVTLGIGRGRRVILRS rad9A protein FLOX ------rad9A protein YHEEEADSTAKAMVTEMCLGEEDFQQLQAQEGVAITFCLKEFRGLLSFAESANLNLSIHF rad9A protein FLOX ------rad9A protein DAPGRPAIFTIKDSLLDGHFVLATLSDTDSHSQDLGSPERHQPVPQLQAHSTPHPDDFAN rad9A protein FLOX ------rad9A protein DDIDSYMIAMETTIGNEGSRVLPSISLSPGPQPPKSPGPHSEEEDEAEPSTVPGTPPPKK rad9A protein FLOX ------rad9A protein FRSLFFGSILAPVRSPQGPSPVLAEDSEGEG

Figure A.1 – Deletions of rad9A exons 3­5 will result in a truncated rad9A protein. Clustal alignment of the wild‐type rad9A protein sequence and the sequence after Cre‐mediated recombination designated rad9A FLOX. The length of the “floxed” protein is 49 amino acids with the first 35 being identical to wild‐type.

183

Figure A.2 – Cre recombinase can be used to manipulate DNA in a variety of ways. Schematic diagram depicting the different ways Cre recombinase can be used to catalyze genomic rearrangements. A. Identical lox sequences in the same orientation results in excision of the DNA fragment between them after exposure to Cre recombinase. B. Identical lox sequences in the opposite orientation will result in inversion of the DNA fragment between them after exposure to Cre recombinase. C. Sequences of DNA flanked by two different lox sequences oriented in the opposite direction on two different DNA strands results in an exchange of the genetic material between the strands after treatment with Cre recombinase.

184

Figure A.3 – DNA constructs for creation of a conditional rad9A‐null cell line. Schematic diagram showing the components of the targeting constructs used in the construction of a conditional rad9A­null cell line. The loxP oligonucleotides are shown as lighter triangles while the lox511 oligonucleotides are shown as darker triangles. The UTRs are represented by the red squares. The black bars represent the approximate location of the rad9A exons. A. Schematic of the pPop1 plasmid. B. Schematic of the pPop2 plasmid. C. Schematic of the pPop3 plasmid, this plasmid is shown without the additional of any rad9A mutant insertions.

185

Figure A.4 – Schematic representation of the construction and final products of the rad9A‐/pop cell line. The first allele is constructed in the traditional homologous recombination method and shows the locus both before and after exposure to Cre recombinase. The second allele uses both homologous recombination and Cre recombinase‐mediated DNA rearrangements to insert mutant rad9A alleles in the endogenous rad9A locus. After treatment with Cre recombinase, the neoR cassette in the second allele will be excised and recombination between the chromosome and pPop3 will cause the mutant allele to be inserted into the genomic rad9A allele. In the scenario shown, the rad9A product will be the truncated protein illustrated in Figure A.1. The GFP cassette allows for selection of successful recombinants via microscopy or flow cytrometry.

186

Figure A.5 – Confirmational PCR to show correct placement of the 5’ and 3’ ends of the targeting construct after homologous recombination. The primers shown in blue will only produce product in cells that homologous recombination between the chromosome and the targeting construct has occurred successfully. The forward primer of this pair is outside the region of homology in the targeting vector and so will not produce product in cells where the construct has integrated randomly. The primers illustrated at the 3’ end will produce product in both wildtype and successful recombinants, which acts as an internal control for the 5’ PCR. This product will also span a unique restriction site in positive recombinants allowing for confirmation of the correct placement of the 3’‐most lox oligonucleotide.

187 Table A.1 – Primers used during the construction of the Rad9A­null conditional cell line.

PCR1‐F GGCGGCCGCTAACCACGATCTACCCTC PCR1‐R GGCGGCCGCCTTTCCTCAATTCACACAC PCR2–F CCGGAGCTTACGATAGGGCAAGTGTGT PCR2‐R CCGGATCGATCATATGTGAATGCAAGCAGTGACC PCR3‐F CCCCGTCGACGACATCTGTTTCCCCAGGTTCTTTTGAGCCA PCR3‐R GGGGCTCGAGCCCTGTGGCTCAGCCCCATGAGCCTGACC GGGGGGGAGCTCATAACTTCGTATAATGTATGCTATACGAAGTTATCCGC LOXP GGGGGGGG GGGGGGCTCGAGATAACTTCGTATAATACGTACTATACGAAGTTATGGTA LOX511 CCGGGGGG NEO‐F* GAGCAGAAGGGAAGGAGAGGGAGAAACAAA NEO‐R* CGGAGATGAGGAAGAGGAGAACAGC LOX‐F* GATGCTGGAGAAGACGGTGGAAAAATGCT LOX‐R* CTGATAGAAAGAGAAGGGGCGGGAGAGAAC EGFP‐F* CCCTCGAGCCTGATTCTGTGGATAACCGTATTACCGCC EGFP‐R* GGTCTGAGGGAACAACACTCAACCCTATCTCGGTCT POP‐F* GGGGAGGCGGGGGCGG POP‐R* GCCCCAAACAAGGAAGAGAGAGACCCG * Indicates primer is for PCR­mediated confirmation after cellular recombination

188