Investigating the Role of Rad51 in Mammalian Ectopic Homologous Recombination

Jennifer Knapp

A Thesis presented to The University of Guelph

In partial fulfilment of requirements for the degree of Master of Science in Molecular and Cellular Biology

Guelph, Ontario, Canada

©Jennifer Knapp, July, 2013

ABSTRACT

Investigating the Role of Rad51 in Mammalian Ectopic Homologous Recombination

Jennifer Knapp Advisor: University of Guelph, 2013 Dr. Mark D. Baker

DNA damage occurs through endogenous and exogenous sources, and can lead to stalled replication forks, genetic disorders, cancer, and cell death. Homologous recombination (HR) is a relatively fast and error-free repair pathway for damaged DNA, which can occur through a gene conversion event or through a crossing-over event with the exchange of genetic material. Homologous recombination occurs most frequently in the G2 phase of the cell cycle and utilizes the sister chromatid as the repair template. When the sister chromatid is unavailable, the homologous chromosome or a homologous sequence in an ectopic location can be used to repair the lesion; the latter of which is referred to as ectopic homologous recombination (EHR). Rad51 is a key protein involved in HR, and to test its role in EHR, variant Rad51 proteins were expressed in murine hybridoma cells. These Rad51 variants were assayed for their effects on EHR. Excess wild-type Rad51 as well as a deficiency of wild- type Rad51 decreased EHR from the background level found in these cell lines. Thus, Rad51 is necessary for EHR, but there may be an optimal amount of Rad51 required for efficient EHR.

Expression of the Rad51 catalytic mutants Rad51K133A and Rad51K133R was found to have an inhibitory effect on EHR, as expected based on the loss of ATP binding and ATP hydrolysis, respectively, in these variants. Excess wild-type Rad51 was verified in this study to increase HR via a gene targeting assay.

MMC treatment, but not ionizing radiation, leads to an increase in EHR in the presence of excess wild- type Rad51. Thus, endogenous levels of Rad51 are sufficient to maintain EHR, but in the presence of excess wild-type Rad51, the level of EHR can increase in response to certain DNA damaging agents and in response to gene targeting.

Acknowledgements

I would like to thank first and foremost my supervisor Dr. Mark Baker for the opportunity to work in his lab as a Master’s student. Thank you for all the support, help and feedback throughout this project. Thank you to my advisory committee members Dr. Ray Lu and Dr. David Josephy for their helpful guidance and advice through committee meetings, as well as editing this manuscript. Thank you to Alissa Magwood for her help in training, helpful consultations about experimental design, and troubleshooting tips. Thank you to all the members of the Baker lab, past and present (Alissa Magwood, Maureen Mundia, Iulia Cealic,

Vatsal Desai, and Taryn Athey), for their helpful comments and friendliness that made the lab a great place to work. Thank you to my family for being extremely supportive of me throughout this project and encouraging me every step of the way. Thank you to Hugo Toste for being extremely understanding and supportive throughout my Master’s degree, but especially through the writing process. A big thank you to all my friends for keeping me grounded and not allowing me to be too overwhelmed by this project and thesis.

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Table of Contents Acknowledgements ...... iii List of Figures ...... vi List of Tables ...... vii List of Abbreviations ...... viii Chapter 1: Introduction and Literature Review ...... 1 1.1 DNA Damage ...... 1 1.2 DNA Damage Repair Pathways ...... 3 1.3 Repair Pathways for DNA damage...... 4 Direct Reversal of DNA Damage ...... 4 Base Excision Repair (BER) ...... 5 Nucleotide Excision Repair (NER) ...... 6 Mismatch Repair (MMR) ...... 7 Non-homologous End Joining (NHEJ) ...... 8 Homologous Recombination (HR) ...... 11 Double-Strand Break Repair (DSBR) ...... 15 Synthesis-Dependent Strand Annealing (SDSA) ...... 17 Break-induced Repair (BIR) ...... 17 Single-Strand Annealing (SSA) ...... 20 1.4 Homologous Recombination Proteins ...... 22 Replication Protein A (RPA) ...... 22 Mre11/Rad50/NBS1 Complex (MRN complex) ...... 23 Breast Cancer Susceptibility 1 (BRCA1) ...... 24 Breast Cancer Susceptibility 2 (BRCA2) ...... 24 Rad52 ...... 25 Rad54 ...... 26 1.5 Rad51 and its Paralogs ...... 27 1.6 Mouse Hybridoma Cells and HR ...... 30 1.7 Hypotheses and Objectives ...... 33 Chapter 2: Materials and Methods ...... 34 2.1 Hybridoma Cell Lines ...... 34

Sp6/HL ...... 34 igm482 ...... 34 pRCµRI #4 and pRCµRI #9 ...... 35 pRCµRI #4 & #9 Derived Expressors of Rad51 Variants ...... 35 2.2 Cell Culture Conditions ...... 39 2.3 Plasmids ...... 39 2.4 Electroporation of Hybridoma Cells ...... 40 2.5 Western Blot Analysis ...... 41 2.6 TNP-Specific Plaque Assay ...... 43 2.7 DNA Damage ...... 47 Mitomycin C (MMC) ...... 47 Ionizing Gamma Radiation (IR) ...... 47 2.8 Statistical Analysis ...... 48 Chapter 3: Results ...... 49 3.1 Stable Expression of FLAG-tagged Wild-type Rad51 and Rad51 Catalytic Variants ..... 49 3.2 Effect of Wild-type Rad51 Expression on Ectopic Homologous Recombination ...... 54 3.3 Effect of Rad51 Depletion on EHR ...... 58 3.4 Effect of Rad51 Catalytic Variants on EHR ...... 59 3.5 Gene Targeting ...... 64 3.6 Gene Targeting after 24h MMC Treatment ...... 66 3.7 Exposure to MMC ...... 68 3.8 Exposure to Ionizing Radiation ...... 70 Chapter 4: Discussion ...... 72 Future Directions ...... 77 References ...... 79

List of Figures

Figure 1 – Non-homologous End Joining (NHEJ) ...... 10 Figure 2 – Donor Sequence Locations for HR and EHR ...... 12 Figure 3 - Double Strand Break Repair (DSBR) ...... 16 Figure 4 - Synthesis-dependent Strand Annealing (SDSA) ...... 18 Figure 5 – Break-Induced Repair (BIR) ...... 19 Figure 6 - Single-strand Annealing (SSA) ...... 21 Figure 7 - pRCμ Vector and Immunoglobulin Locus Structure ...... 31 Figure 8 – Plasmid Maps for Generation of Rad51 Variant Expressors ...... 37 Figure 9 – TNP-Specific Complement Dependent Plaque Assay ...... 45 Figure 10 – Subculture Generation ...... 46 Figure 11 - Western Analysis of Rad51 Expression ...... 52 Figure 12 - Western Analysis of Rad51 Knockdowns ...... 53 Figure 13 – EHR in Parental Cell Lines #4 and #9 ...... 55 Figure 14 – Effect of FLAG-tagged Wild-type Rad51 Expression on EHR ...... 57 Figure 15 – Effect of Rad51 Depletion on EHR ...... 59

Figure 16 – Effect of Expression of Rad51K133A on EHR ...... 61 Figure 17 – Effect of Rad51K133R Expression on EHR...... 63 Figure 18 – Effect of MMC Treatment on Gene Targeting ...... 67 Figure 19 – Effects of 6h and 24h of MMC Treatments on EHR ...... 69 Figure 20 – Effect of 4 Gy Ionizing Radiation on EHR ...... 71

List of Tables Table 1 - Summary of Cell Lines...... 38 Table 2 - Diagnostic Antibodies...... 42 Table 3 - Gene Targeting Results ...... 65

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List of Abbreviations

ANOVA One-way analysis of variance

BER Base excision repair

BIR Break-induced repair bp Base pair

BRCA1 Breast cancer susceptibility 1

BRCA2 Breast cancer susceptibility 2

BSA Bovine serum albumin

Cµ µ gene constant region

DMEM Dulbecco’s modified Eagle medium

DSB DNA double-strand break

DSBR Double-strand break repair dsDNA Double-stranded DNA

EHR Ectopic homologous recombination

GG-NER Global genomic nucleotide excision repair

HR Homologous recombination

HRP Horse-radish peroxidase hyg Hygromycin B phosphotransferase gene

IgM Immunoglobulin M

IR Ionizing radiation kb Kilobase kDa Kilodalton

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MMR DNA mismatch repair neo Neomycin phosphotransferase gene

NER Nucleotide excision repair

NHEJ Non-homologous end joining

NLS Nuclear localization signal

PFC Plaque-forming cells

RPA Replication protein A

SDSA Synthesis-dependent strand annealing

SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis siRNA Small interfering RNA

SSA Single-strand annealing

SSB Bacterial single-strand binding protein ssDNA Single-stranded DNA

TC-NER Transcription-coupled nucleotide excision repair

TNP 2,4,6-Trinitrophenyl

TNP-SRBC TNP-coupled sheep red blood cells

UV Ultraviolet light

WB Western immunoblotting

XRCC3 X-ray repair cross complementing 3

Chapter 1: Introduction and Literature Review

1.1 DNA Damage

An organism’s DNA encodes all the information needed for that organism to survive.

Cells must replicate and maintain their genomes with incredible accuracy to avoid deleterious, cancer-causing mutations and to ensure the faithful transmission of genetic information to the next generation (Preston et al., 2010). Genetic mutations are the basis for many disorders and diseases, including cancer, but mutations are also necessary to drive evolution. DNA damage is associated with premature ageing and predisposition to certain cancers and inherited diseases

(Aguilera and Gómez-González, 2008). There is a low level of inherent error in the DNA replication process that is caused by polymerase errors, such as incorrect base addition, primer slipping and polymerase slipping; however, DNA damage from exogenous sources is also a factor. A variety of chemicals and substances in the environment can damage DNA and cause problems with replication. Many different DNA lesions, such as point mutations, deletions, translocations and breaks, can cause adverse effects within the genome. Whole chromosome additions or deletions, as well as the shortening or lengthening of repetitive DNA sequences caused by microsatellite instability can also occur and cause adverse effects (Draviam et al.,

2004). Replication stress, followed by DNA double-strand breaks, genomic instability, and consequently, a favoured p53 deficiency, is associated with the early development of cancer

(Gorgoulis et al., 2005). It is critical that these errors are caught by the cell during cell cycle checkpoints and subsequently repaired (Gorgoulis et al., 2005).

The replication of DNA during S phase of the cell cycle provides many opportunities for errors to occur. As the cell progresses through a series of checkpoints to ensure that DNA damage is caught before the cell can continue to the next stage in the cell cycle, many

processes ensure that if there is a problem with the DNA replication, this problem is not passed on to daughter cells. When a replication fork encounters an area of damaged DNA the replication fork can become stalled or even collapse (Petermann and Helleday, 2010).

Although there are many active forks throughout replication in eukaryotes, a stalled fork can leave an area of un-replicated DNA, or result in a single-strand break or a more serious double- strand break. If DNA replication is hindered by a stalled fork, the cell must stop its progression through the cell cycle until replication is complete and the problem resolved. Stalled forks are able to re-start after the blocking agent is removed; however, if the fork has collapsed, it is more complicated to restart replication. A collapsed replication fork can lead to the dissociation of the replication machinery and/or double-strand breaks in the DNA (Petermann and Helleday, 2010). Replication forks can become stalled due to endogenous single strand breaks (Saleh-Gohari et al., 2005), or through the addition of exogenous agents such as mitomycin C (MMC), cisplatin, hydroxyurea, or ionizing gamma radiation, which introduce double strand breaks in DNA. These fork-stalling agents have been used to study replication fork restart and DNA repair through homologous recombination. Homologous recombination can be used to restart stalled replication forks through the use of the sister chromatids

(reviewed in Cox 2002). Fork restart through recombination-dependent pathways has been shown in Escherichia coli (Heller and Marians, 2006), but the details of mammalian fork restart are slightly more hazy, due to the involvement of more proteins than simply the homologs of the bacterial proteins, and a larger variety of DNA damage repair pathways.

1.2 DNA Damage Repair Pathways

Many DNA lesions in eukaryotes are converted into ssDNA breaks and ultimately into dsDNA breaks prior to the repair process (Zou and Elledge, 2003). Any kind of DNA break is able to initiate a range of signal cascades that end either in DNA repair or apoptosis of the affected cell. Single-stranded DNA, present at the sites of DNA lesions, attracts replication protein A (RPA) which has a high affinity for ssDNA (Wyka et al., 2003). The complex of

RPA bound to ssDNA is then able to recruit ATR/ATRIP (Ataxia telangiectasia and Rad3

Related/ATR interacting protein) dimers, Rad17 and 9-1-1 (Rad9, Hus1, and Rad1) complexes which stimulate the kinase activity of ATR (Zou and Elledge, 2003). The MRN complex, which is similar to the MRX complex in yeast, is composed of Mre11, Rad50, and Nbs1, and recognizes DNA double-strand breaks (DSBs). The MRN complex is involved in the recruitment of protein kinase ATM (Ataxia telangiectasia-mutated), which is a key step in the propagation of the DNA damage response signal (Dupré et al., 2006). Once ATR and ATM are activated, they phosphorylate many proteins linked to DNA repair, including p53 (a tumor suppressor gene), BRCA1, Nbs1 (part of the MRN complex), and H2AX (Histone H2A protein) (Matsuoka et al., 2007). However, the most important phosphorylation targets of

ATR/ATM are Chk2 and Chk1, which are kinases that when active are able to inactivate

Cdc25 (Cell division cycle 25) through phosphorylation. In the absence of Cdc25 functionality,

CDK1 (Cyclin dependent kinase 1) is unable to be dephosphorylated and so initiates cell cycle arrest at any of the cell cycle checkpoints (Reviewed in Jackson and Bartek, 2009). Arrest of the cell cycle allows the DNA lesions to be repaired before continuation onto the next phase.

ATM/ATR signalling also increases the transcription of many DNA repair proteins and allows for their recruitment and activation at the sites of the DNA lesions. The ATM/ATR

signalling stops when the DNA lesions have been repaired and this enables the cell cycle to resume. Cellular senescence or apoptosis is triggered by ATR/ATM signalling if the DNA lesions cannot be repaired (Cimprich and Cortez, 2008).

1.3 Repair Pathways for DNA damage

A brief introduction to several DNA repair processes is given below. It is important to note that none of these processes occurs in isolation, but many use some of the same proteins and even complement or compete with each other. Some types of DNA damage can be repaired by a simple reversion of the damage but others require a more complicated process with the involvement of multiple proteins.

Direct Reversal of DNA Damage

Cells have developed some simple DNA repair pathways that are able to correct damaged bases without their excision or the excision of several nucleotides around the damaged base (Reviewed in Eker et al., 2009). These pathways involve single proteins with high substrate specificity that are able to correct certain types of base damage, UV-induced

DNA damage, and DNA alkylation damage. Alkyltransferases and dioxygenases are the two types of repair proteins that are able to repair alkylated bases. Alkyltransferases act by irreversibly transferring the alkyl group to themselves, and are then subjected to ubiquitin- mediated degradation since they cannot be recovered after this process (Coulter et al., 2007).

Dioxygenases use oxidative demethylaton to reverse alkyl base damage, and the oxidized product is released as fomaldehyde (Duncan et al., 2002). UV light causes two main types of

DNA lesions, 6-4 photoproducts (6-4 PP) and cyclobutane pyrimidine dimers (CPD) (Yang,

2011). In prokaryotes, a simple pathway is able to repair both of these lesions through the

action of a photolyase specific to either 6-4 PP or CPD. Visible blue light is used by photolyases to cleave the covalent bond linking the pyrimidine dimer, restoring the bases to their original configuration (Kao et al., 2005). Photolyases are found in organisms from all three kingdoms of life.

Base Excision Repair (BER)

Unlike the direct reversal pathways, base excision repair requires the action of multiple proteins in concert, but it is one of the most important pathways in eukaryotes. Knocking out the BER pathway in mice has been shown to cause embryonic lethality (Cabelof et al., 2003) and in humans, mutations in this pathway have been shown to lead to several types of cancer, including one type of heritable colon cancer (Lefevre et al., 2006). The BER pathway is utilized to remove hydrolysed, oxidized or alkylated bases, which could be caused through exogenous sources or endogenous metabolic processes.

The first step of BER is the recognition of the DNA lesion by a DNA glycosylase that cleaves the glycosidic bond that attaches the base to the DNA backbone; this creates an abasic site with an intact backbone (Reviewed in Prasad et al., 2011). AP endonuclease 1 is then recruited to the site to cut the phosphodiester bond, which leaves a single nucleotide gap in that

DNA strand. DNA polymerase β fills this gap using the opposite strand as a template. Ligase I or XRCC1/DNA ligase III complex are then recruited to seal the nick left by DNA polymerase

β.

Nucleotide Excision Repair (NER)

Unlike BER, NER acts on a wider variety of lesions and is able to remove larger DNA lesions that cause helix distortions. NER is the main pathway for the repair of UV light- induced DNA lesions (CPD and 6-4 PP) in all organisms (Yang, 2011). It has been shown in humans that mutations in this pathway cause UV sensitivity and an elevated risk of skin cancer along with severe developmental and neurological abnormalities (Setlow et al., 1969). There are two pathways that are considered NER, global genomic NER (GG-NER) and transcription- coupled NER (TC-NER). These two pathways differ only in their recognition of the DNA lesion, and for the remainder of the repair process they are the same, utilizing the same proteins

(Reviewed in Rechkunova et al., 2011). Rather than directly recognizing the lesion itself, GG-

NER is thought to recognize a helix distortion or a loss of DNA rigidity around the lesion through the action of the XPC/Rad23B/Centrin-2 complex, and then, with UV-DBB, is able to initiate the assembly of the repair complex (Yang, 2007). The first step in TC-NER occurs when RNA polymerase becomes stalled at a lesion during transcription; it then attracts the proteins needed to initiate the NER process (Selby et al., 1997). After the recognition of the

DNA lesion, both pathways converge, and two helicases (XPB and XPD) are recruited to unwind the DNA. The XPG 3’ nuclease and the XPF-ERCC1 5’ nuclease cleave the DNA around the lesion and an oligonucleotide (~25-30 bp) containing the lesion is removed. DNA polymerases δ and ε fill in the gap left over and DNA ligase III seals the resulting nick, completing the NER process.

Mismatch Repair (MMR)

During DNA replication, errors are made by the DNA polymerase that may be missed by its proofreading ability and these may be repaired by MMR (Martin et al., 2010). These replication errors can include single base mismatches as well as insertions or deletions caused by a misalignment of the newly-synthesized DNA and the template strand due to repetitive sequences in proximity (Streisinger and Bruce, 1960). DNA-damaging agents can produce base adducts which are also recognized by MMR proteins which then mediate the damage response to these lesions (Yoshioka et al., 2006). MMR proteins also prevent gross rearrangements of the genome through their negative regulation of homologous recombination. These proteins are also responsible for mediating the conversion of ssDNA breaks into dsDNA breaks during somatic hypermutation and class-switch recombination of the immunoglobulin genes in mammalian B cells (Stavnezer et al., 2008).

Because MMR is an essential pathway in DNA repair, the proteins involved in this process are highly conserved throughout the different taxa of life, although, as expected, the eukaryotic proteins are more complex. Cells deficient in MMR show a higher frequency of frameshifts, codon alterations, and microsatellite instability (Martin et al., 2010). In humans, mutations in the MMR genes dramatically increase the predisposition to hereditary colon cancer and are implicated in the development of 5-15% of other cancers (Hewish et al., 2010).

The MMR pathway must be able to distinguish between the nascent DNA strand which contains the replication errors and the template strand in order to prevent the wrong correction of the lesion. In gram-negative prokaryotes, the hemi-methylation status of the DNA distinguishes the newly-synthesized DNA (Bae et al., 2003). In gram-positive prokaryotes and

eukaryotes, strand discontinuities associated with DNA replication can serve as a signal for the newly-synthesized DNA (Fukui, 2010).

A MutS homodimer recognizes the mismatched base in prokaryotes and then a MutL homodimer is recruited. This MutL homodimer activates MutH, which nicks the unmethylated strand of DNA. DNA polymerase III repairs the gap left by mismatch repair and DNA ligase seals the resulting nick (Fukui et al. 2010). In eukaryotes, the helix distortion is recognized by two heterodimers, MSH2/6 and MSH2/3, which are homologous to the MutS homodimer

(reviewed in Martin et al., 2010). After binding to the dsDNA at the site of the lesion, the MSH recruits MLH heterodimers (homologous to MutL). The MSH/MLH complex travels along the

DNA in an ATP-hydrolysis dependent manner until it encounters a ssDNA break bound by some proteins, including replication factor (RFC) and proliferating cell nuclear antigen

(PCNA). In cooperation with PCNA, RFC, and RPA, the exonuclease EXO1 removes nucleotides from the newly synthesized DNA toward and beyond the lesion. Once the removal of the lesion is complete, DNA polymerase δ is recruited to fill the gap and DNA ligase I is able to seal the nick (Martin et al., 2010).

Non-homologous End Joining (NHEJ)

NHEJ is one of the major pathways for the repair of DNA double-strand breaks (DSB).

This pathway does not require a homologous template from which to repair the DSB, unlike homologous recombination (HR) which is another main pathway for the repair of DSBs

(reviewed in the following section). IR-induced DSBs as well as those induced by chemical mutagens are repaired through NHEJ, but NHEJ also helps with the resolution of V(D)J recombination during the differentiation of B- and T- lymphocytes in vertebrates (Lieber et al.,

2004). NHEJ is the main pathway used in G0, G1 and M phases of the cell cycle, since the homologous template required for HR is not available during these stages (Sonoda et al.,

2006). This method of repair of DNA DSBs is considered to be error-prone, since no homology is required and insertions or deletions could occur during the repair. It is important to note that two ends of the same DSB can be repaired by NHEJ with high fidelity due to the complementary nature of their sequences after resection occurs (Wilson and Lieber, 1999;

Durant and Nickoloff, 2005).

Figure 1 shows the process of NHEJ, which begins with the binding of Ku70 and Ku80 to both ends of the DSB (forming the Ku70/80 heterodimer) shortly after the break happens, since these proteins are at a high concentration in the nucleus and have high affinity for dsDNA ends (Mari et al., 2006). The binding of these proteins prevents the DNA ends from being resected, a process which is necessary for the HR pathway, which means Ku70/80 is instrumental in guiding a DSB to NHEJ for repair instead of HR pathways. DNA-dependent protein kinase catalytic subunit (DNA-PKCS) is recruited by Ku70/80 to the DSB and acts with the heterodimers to bind the DNA ends together in a synaptic complex (Bennett et al., 2012). A conformational change occurs in the DNA-PKCS activating its kinase functions which recruits nucleases, polymerases and ligases to the DNA ends (Uematsu et al., 2007). Artemis is an example of a nuclease which is able to remove damaged bases from the DNA ends prior to ligation, or resect ends which do not fit together (Reviewed in De Ioannes et al., 2012). These activities could result in gaps that require the action of DNA polymerase before ligation can occur, stimulated by ligase IV and XRCC4 (Wu et al., 2007). This process allows for the ligation of the DNA ends, successfully repairing the DSB lesion.

DSB

Binding of NHEJ proteins Ku 70/80

Synaptic complex forms DNA-PK CS with DNA ends

Artemis

Resection and gap filling Ligase at DNA ends IV/XRCC4/XLF

Ligation and dissociation of complex

Figure 1 – Non-homologous End Joining (NHEJ). The Ku70/80 heterodimer binds the DSB shortly after its formation, preventing the resection of the DNA ends. DNA-PKCS are recruited to the DSB and help with the recruitment of the nuclease Artemis which resects the broken DNA ends in preparation for ligation. Ligase IV ligates the broken ends together with the help of XRCC4 and XLF.

Homologous Recombination (HR)

Homologous recombination (HR) and other HR-mediated processes, such as unequal sister-chromatid exchange and ectopic HR, are important pathways for the repair of DNA double-strand breaks (Kanaar et al., 1998) and for restarting replication forks (Aguilera and

Gómez-González, 2008). HR is an essential pathway in all forms of life as it helps maintain genomic integrity and suppresses tumors (Wyman and Kanaar, 2006; Moynahan and Jasin,

2010). Because this is a homology-driven pathway for repair of DSBs there is a requirement for a nearly identical sequence of DNA (preferably the sister chromatid) from which the lesion can be repaired. This use of a homologous sequence of DNA means that the original sequence is conserved with a high degree of fidelity, an advantageous trait in a DNA repair pathway.

Ectopic homologous recombination (EHR) involves a homologous donor sequence that is located in the genome in a position other than parallel to the endogenous locus on the homologous chromosome or sister chromatid (Figure 2). Ectopic homologous recombination differs from homologous recombination in that the homology search to find an ectopic donor must be much broader than the local search required to find the sister chromatid or homologous chromosome. Ectopic recombination could require different proteins or different stoichiometric ratios of the homologous recombination proteins due to this requirement for a homology search with a wider range. Since Rad51 is the protein implicated in the homology search for a donor sequence for recombination (West, 2003), EHR may have different requirements for this protein from HR.

1 2 3

5 4

Figure 2 – Donor Sequence Locations for HR and EHR. Depicted here are a pair of homologous sequences (open rectangles) located on either replicated homologs (thin lines) or replicated heterologs (thick lines). Pathways 1 and 2 are considered homologous recombination since both of these donor sequences are in the same place as the endogenous locus, but on the homolog or sister chromatid, respectively. Pathways 3-6 are considered ectopic homologous recombination since the donor sequences are not in the same place as the endogenous locus and could even be on a completely different chromosome, i.e. not the homologous chromosome.

In eukaryotes, HR is instrumental in replication fork restart and, in particular, in dealing with single-ended DSBs that occur from a stalled or collapsed replication fork. In support of the role of HR in repairing single-ended DSBs caused by stalled or collapsed replication forks, several replication inhibitors (mimosine, ciclopirox olamine, aphidicolin, and hydroxyurea) were shown to promote homologous recombination (Saintigny et al., 2001;

Lundin et al., 2002). Many DNA damaging agents introduce double ended DSBs in DNA, including IR and MMC; these IR- or MMC-induced lesions are also a main substrate of the HR pathways. Interstrand crosslinks (ICLs) block strand separation during replication and transcription and are another substrate for HR-mediated repair (Hinz, 2010). Aside from the role of HR in repairing DNA lesions, it also contributes to the genetic variability of a population by allowing meiotic recombination between homologous chromosomes to take place (Shirleen Roeder, 1990; Kleckner, 1996; Rosu et al., 2011). Homologous recombination is a relatively fast and error-free process due to the use of a homologous sequence to complete the repair conservatively (Dudáš and Chovanec, 2004). However, small differences in the sequence or an area of repetitive DNA could lead to non-conservation of the sequence. Thus, upregulation of HR could lead to tumorigenesis as it could lead to genomic rearrangements and genomic instability. It has been shown that cancer cells often have a higher frequency of HR

(Halazonetis et al., 2008). It is important then, that this process be tightly regulated and controlled in properly functioning cells.

HR and EHR can take place through many pathways in mammals, all of which are interconnected and diverge only after the initial steps have been carried out. The broken ends of the DNA DSB are resected by a 5’ to 3’ exonuclease so that they have single-stranded DNA tails with free 3’-OH groups. These ssDNA tails are required for the homology search and

subsequent strand invasion into the homologous sequence (Yeeles and Dillingham, 2010). The resection of the DNA ends involves a number of proteins, including the nuclease Dna2, Sgs1 (a

RecQ-family helicase), and RPA (a ssDNA binding protein) (Cejka et al., 2010). The resection of the DNA ends in yeast also involves 5’ to 3’ exonuclease Exo1, and Sae2 (EXO1 and CtIP in mammals), which probably interact with the MR(X)N complex (Kousholt et al., 2012;

Tomimatsu et al., 2012). DNA unwinding near the double strand break is carried out by the

MRX complex acting in concert with topoisomerase 3 and the Rmi1 complex (Cejka et al.,

2010).

After the resection of the DNA ends, the ssDNA overhangs are bound by RPA to ensure no secondary structures form within the DNA and that the strands remain intact while

RPA recruits the rest of the proteins necessary for HR (Blackwell and Borowiec, 1994). Rad51 displaces the RPA and binds to the ssDNA converting it into a nucleoprotein filament that is able to invade dsDNA in search of a homologous sequence to use as a template for the repair.

A brief description of the homology search and strand invasion steps and some of their mediators (Rad52, Rad54, BRCA2) is provided below.

Strand invasion takes place into the homologous donor region, which creates a displacement loop (D-loop) of DNA when initial base-pairing of the invading strand occurs

(Raynard and Sung, 2009). The continued base pairing of the invading strand along the homologous template is mediated through Rad51 association and dissociation along the DNA filament and is reviewed in Holthausen et al. (2010). After this point there are at least three ways in which the D-loop can be resolved, each of which is described briefly below. A fourth

HR mechanism called single-strand annealing (SSA), which does not involve D-loop resolution, is also described.

Double-Strand Break Repair (DSBR)

In the DSBR pathway (Figure 3), after the Rad51 nucleoprotein filament has invaded the homologous region of duplex DNA and formed a D-loop structure, DNA synthesis begins, extending the D-loop so that it can be captured by the other end of the DSB (Pardo et al.,

2009). DNA synthesis then continues at both original ssDNA ends, working off the D-loop as a template. An intermediate structure is formed consisting of a double Holliday junction after gap filling and ligation has occurred on both strands. The resolution of this double Holliday junction can yield crossover products or non-crossover products as shown in Figure 3 (San

Filippo et al., 2008). Crossing over results in a reciprocal exchange of genetic material between the two DNA molecules and is desirable particularly in meiosis where crossover events are necessary to hold homologous chromosomes together in bivalents (Rosu et al., 2011). In mitotic cells, however, crossing over is not desirable, since the homologous donor region could be in an ectopic location, in which case crossing over could result in genetic rearrangements that could be harmful to the cell. For this reason, crossing over during the repair of DNA DSBs is repressed in mitotic cells (Shulman et al., 1995; Johnson et al., 2001; Dayani et al., 2011).

The double Holliday junction can be dissolved with the help of Bloom’s syndrome helicase

(BLM) in conjunction with type I topoisomerase (TOPOIIIα), reducing the risk of any chromosomal rearrangements (Wu et al., 2006).

DSB

Resection of DNA ends 3’ 3’

D-loop formation

Branch migration + second end capture

Dissolution of double Holliday Double Holliday junction junction formation C D

A A B B

C D

Crossover (A+D or B+C) Non-crossover (A+B or C+D)

Figure 3 - Double Strand Break Repair (DSBR). After the resection of the DSB ends, filament formation, strand invasion, D-loop formation, and branch migration occurs along with the capture of the second DSB end by the D-loop. This results in a double Holliday junction structure that can be resolved into crossover products (cleaving at A+D or B+C) or non-crossover products (cleaving at A+B or C+D). The double Holliday junction structure can also be dissolved, yielding the same products as synthesis-dependent strand-annealing (SDSA).

Synthesis-Dependent Strand Annealing (SDSA)

SDSA is a pathway of D-loop resolution, shown in Figure 4, which happens when the

D-loop is unable to capture the second 3’ ssDNA end (Heyer et al., 2010). The invading strand is synthesized regularly within the D-loop, until the dissolution of the D-loop and its displacement from the homologous template region. This newly-synthesized DNA is able to anneal to the free 3’ ssDNA from the other end of the DNA DSB and then DNA synthesis of the resulting ssDNA gap is initiated. SDSA yields only non-crossover products since no

Holliday junctions were formed in this pathway. For this reason, it is thought to be the primary pathway in somatic cells. SDSA produces the same products as the dissolution of the double

Holliday junction structure formed in DSBR (Figure 3).

Break-induced Repair (BIR)

BIR, the third homology dependent pathway for DSB repair (Figure 5), occurs when only one end of the DSB can be found or when there is only one end to start (i.e. the end of a telomere) (Lydeard et al., 2010). The BIR pathway involves most of the same steps as discussed above, including the invasion of a Rad51 nucleoprotein filament into a region of homology. DNA synthesis within the D-loop can continue until the end of the homologous template chromosome, and after dissociation, this newly synthesized DNA is used as a template for the second strand of the repaired molecule. It is important to note that any genetic information that was present on the second end of the DSB is lost after this repair pathway occurs, potentially leading to tumorigenesis (Smith et al., 2007). Even with this flaw, BIR is an important pathway for the repair of DSBs in telomeres and cells deficient in BIR also have decreased viability.

DSB

Resection of DNA 3’ ends 3’

D-loop formation

D-loop dissociation

Annealing and DNA synthesis

Figure 4 - Synthesis-dependent Strand Annealing (SDSA). This pathway for DSB repair occurs after the D-loop forms, but is then dissociated after some DNA synthesis has occurred. The newly synthesized DNA is able to anneal to complementary sequences on the second end of the DSB, and the gap is filled in by DNA polymerase.

DSB

Resection of DNA ends and loss of one chromatid arm 3’

D-loop formation

D-loop migration and DNA synthesis

Figure 5 – Break-Induced Repair (BIR). When a DSB occurs and the second end is lost or non-existent (i.e. a telomere end) BIR occurs. Resection, filament formation, D-loop formation and strand invasion occur but there is no second end for the D-loop to capture. The D-loop migrates along the template as DNA synthesis continues, potentially until the end of the template. Synthesis of the second strand occurs using the newly synthesized DNA as a template.

Single-Strand Annealing (SSA)

The final homology-based repair pathway to be discussed here, SSA, does not involve the resolution of a D-loop structure or the invasion of a Rad51 nucleoprotein filament. In fact, in SSA, the Rad51 nucleoprotein filament is not formed. In yeast, SSA is a Rad52-dependent process, but is independent of most of the other HR proteins including Rad51 (Ivanov et al.,

1996). Rad52 has strong single-strand annealing activity and is able to overcome the RPA- ssDNA complex which poses a barrier to SSA (Sugiyama and Kowalczykowski, 2002; Mott and Symington, 2011). If there are direct repeats at the locus of the DSB, SSA can occur if the resected ends of the DSB result in complementary sequences being exposed (Schildkraut et al.,

2005). These complementary ends anneal to each other and any overhanging 5’ sequence is removed. As a result of this, one or more of the direct repeats around the DSB are lost due to the nature of the repair mechanism; therefore, this is a non-conservative method of DSB repair.

Complementary DNA repeats DSB

Resection of DNA ends 3’ 3’

Annealing

Cleavage of overhangs, gap filling and ligation

Figure 6 - Single-strand Annealing (SSA). This pathway does not involve Rad51 filament formation, strand invasion or D-loop formation. In regions of repetitive DNA, when the DSB ends are resected, if complementary sequences result they are able to anneal. Overhanging sequences are removed and gaps are filled to complete the repair.

1.4 Homologous Recombination Proteins

Homologous recombination requires a host of different proteins, all acting in concert, to perform the DNA repair process effectively. Theoretically, EHR might require the same proteins as HR due to the similarity of the two pathways; however, there may be some differences in requirements due to the wider homology search required by EHR to find a donor sequence. A brief overview of the major HR proteins is given here and a more detailed description of Rad51 and its functions is also provided, as it is the protein of interest in the current study.

Replication Protein A (RPA)

RPA is a eukaryotic protein made up of three subunits (70, 30 and 14 kDa). These subunits are highly conserved across a wide range of taxa and bind with high affinity to ssDNA

(Wold, 1997). RPA is the “first responder” to regions of ssDNA surrounding a DSB, binding tightly to regions of ssDNA and protecting these regions from nuclease attack and from forming unwanted secondary structures. In DNA damage signalling, RPA is involved in the recruitment of the DNA damage checkpoint kinases, which facilitate checkpoint activation

(Liu et al., 2012). RPA is also involved in DNA repair where its role is the recruitment of many of the DNA repair proteins and aiding in cleavage at the DNA damage site, removal of mismatched bases and gap filling (Prakash and Borgstahl, 2012). In HR-mediated pathways of

DNA repair, RPA is involved in the recruitment of Rad52 and BRCA1 while keeping the ssDNA unwound to aid Rad51 nucleoprotein filament formation (Blackwell and Borowiec,

1994). Yeast Rad52 is recruited to double-strand breaks by the action of RPA, probably through a direct physical binding or interaction (Hays et al., 1995; Lisby et al., 2004), but this

is not found for human Rad52. RPA is also able to bind to the displaced strand of the D-loop, stabilizing this complex that was initiated by the Rad51 nucleoprotein filament. Interestingly,

RPA inhibits the nucleation of the Rad51 nucleoprotein filament in vitro, but this effect is reversed when the reaction is supplemented by Rad52 in yeast or BRCA2 in humans which act as mediators to help displace the RPA from the ssDNA in favour of the binding of Rad51

(Sugiyama and Kowalczykowski, 2002).

Mre11/Rad50/NBS1 Complex (MRN complex)

The MRN complex in mammals (Mre11/Rad50/XRS2 (MRX) in yeast) is involved in cell cycle signalling but is also important in the DNA damage response of the cell. The MRN complex plays a role in other pathways as well, including telomere maintenance and DNA replication resumption and repair (Lamarche et al., 2010). The importance of this complex can be demonstrated in mice, where knocking out any of the three genes that make up this complex results in embryonic lethality, and in humans causes genomic instability syndromes (Luo et al.,

1999). In yeast, null mutations in any of these three genes results in an increased sensitivity to

DNA damaging agents and a slower resection of programmed meiotic DSBs (Ivanov et al.,

1996). Mre11 exhibits 3’-5’ exonuclease activity which is highly conserved among taxa and is required for the resection of DSB ends; the first step in HR (Reviewed in Rupnik et al., 2010).

Rad50 has a zinc-hook motif which it uses to bind the DNA ends and bridge them together. A disruption in this functionality in yeast results in the same phenotype as if one of these three genes is knocked out. NBS1 is the component of this complex that is responsible for its recruitment to the DSB as its C-terminal domain is able to bind to γH2AX, which helps in the recruitment (Zhang et al., 2006). NBS1 forms the complex with Mre11 and Rad50, recruiting them to the DSB, along with ATM which it is also able to activate (Rupnik et al., 2010). Once

ATM is activated, it is able to initiate a signal cascade by phosphorylating various target proteins, which leads to cell cycle arrest.

Breast Cancer Susceptibility 1 (BRCA1)

Inherited BRCA1 mutations have been linked to the predisposition for breast and ovarian cancers as well as pancreatic and prostate cancers (Roy et al., 2012). BRCA1 forms a heterodimer with BARD1 (BRCA1-associated RING domain 1), and this complex is involved in the formation of polyubiquitin chains that signal downstream events, but do not signal protein degradation. This heterodimer is able to ubiquitinate proteins involved in many pathways, including transcriptional regulation, cell cycle control, DNA repair, and chromatin remodelling (Starita and Parvin, 2006). BRCA1 interacts with the MRN complex, and through this interaction is involved in HR-mediated DNA repair, as well as SSA. BRCA1 is also a mediator of DSB end resection involving the action of CtIP. BRCA1 directly interacts with

PALB2, which locates and binds to BRCA2, and is involved with the recruitment of Rad51 to the DSB through this interaction (Zhang et al., 2009).

Breast Cancer Susceptibility 2 (BRCA2)

Several mutations in BRCA2 are implicated in familial cases of breast cancer. One of these mutations causes a truncation of the BRCA2 protein. Cells carrying this BRCA2 truncation are defective for Rad51 transport into the nucleus suggesting that full length BRCA2 allows Rad51 to be translocated into the nucleus upon DNA damage (Davies et al., 2001). IR- induced Rad51 foci formation in these cells was impaired by truncated BRCA2, however, normal S-phase Rad51 foci formation occurs in the presence of truncated BRCA2 (Tarsounas et al., 2003). A crystal structure of a BRCA2 domain (~90 kDa) was demonstrated to be able to

bind to single stranded DNA in an experiment in which it interacted with oligo(dT), oligo(dC) and mixed sequence ssDNA. In this experiment the BRCA2 domain did not interact with dsDNA, supporting the possibility that it is able to recruit itself to the double-strand break, where there are resected ends and exposed regions of ssDNA (Yang et al., 2002).

BRCA2 recruits Rad51 to dsDNA:ssDNA junctions and functions as a recombination mediator that facilitates the assembly of the Rad51 nucleoprotein filament, thus aiding homology-directed DSB repair (Jensen et al., 2010). It has been shown that BRCA2 is able to regulate the DNA binding preference and localization of Rad51 by stabilizing Rad51 filament formation on ssDNA, but blocking Rad51 filament formation on dsDNA (Carreira et al., 2009).

It is thought that a single molecule of BRCA2 is able to bind up to six molecules of Rad51, and was strongly biased towards binding to ssDNA versus dsDNA (Jensen et al., 2010; Thorslund et al., 2010). BRCA2 was also shown to aid Rad51 in binding to RPA-coated ssDNA, which is an essential step in HR-mediated repair pathways (Liu et al., 2010).

Rad52

Rad52 is important in both SSA and DSBR pathways of HR-mediated repair. In yeast,

Rad52 is the recombination mediator that is responsible for the recruitment of Rad51 to the

DSB and for aiding Rad51 nucleoprotein filament formation by removing RPA from ssDNA

(Shinohara and Ogawa, 1998). Rad52 is able to recruit Rad59 to the DNA DSB through direct or indirect interactions and the nuclear localization of Rad59 is dependent on Rad52 (Lisby et al., 2004). Strand annealing associated with capture of the D-loop in DSBR is stimulated by

Rad52 (McIlwraith and West, 2008). Rad52 forms heptameric rings and binds ssDNA in a sequence-independent manner. When Rad52 is bound to ssDNA, the bases point away from the

protein, allowing for complementary sequences to anneal (Singleton et al., 2002). In yeast knocking out Rad52 results in high sensitivity to DSB-inducing agents and causes defects in many DNA repair pathways including HR, SSA, and BIR. In vertebrates, however, it is suggested that Rad52 is functionally redundant with some other protein since null Rad52 mutants show no heightened sensitivity to DSB-inducing agents and show only minor defects in HR-mediated repair (Rijkers et al., 1998). Mouse cells deficient for both Rad52 and Rad54 show heightened sensitivity to DNA damage induced by MMC or IR exposure (de Vries et al.,

2005). Fujimori et al. (2001) showed that chicken DT40 B-cells reduced for Rad52 did not show increased sensitivity to DSB-inducing agents. However, a double knockdown of Rad52 and the Rad51 paralog XRCC3 (X-ray Repair Complementing 3) is not viable. These findings suggest that Rad52 and XRCC3 act together, or perhaps Rad52 acts with several Rad51 paralogs.

Rad54

Rad54 is present in a wide range of taxa and is a highly conserved protein (Heyer et al.,

2006). Rad54 mutants are viable in yeast and mammals, but the absence of Rad54 renders them highly sensitive to DSB-inducing DNA damaging agents such as IR and DNA cross-linking agents. Rad54 functions as a dsDNA-dependent ATPase, using the hydrolysis of ATP to power translocation along the DNA strand, however, unlike other helicases, this translocation is not accompanied by strand unwinding (Mazin et al., 2010). This translocation along the DNA coupled with the ability of Rad54 to stimulate chromatin remodelling and aid in the removal of proteins from dsDNA, makes it clear that Rad54 has a role in providing accessibility to other repair proteins. In yeast, Rad54 is recruited to the DSB only in the presence of Rad52, Rad51, and Rad55-57 (Lisby et al., 2004). Rad55-Rad57, along with Rad52, are key proteins in the

assembly of Rad51 nucleoprotein filaments in yeast (Sung, 1997; Lisby et al., 2004). Rad54 is instrumental in linking various stages of HR together, and stimulates D-loop formation and

Rad51-mediated strand exchange (Heyer et al., 2006). In a somewhat reciprocal relationship,

Rad51 stimulates the activities of Rad54 such as ATP hydrolysis, translocation along the DNA, and chromatin remodelling (Kiianitsa et al., 2002). Rad51 dissociation from the DNA repair complex is facilitated by the action of Rad54. Through its DNA translocation and ATP hydrolysis functions, Rad54 is able to drive branch migration of a double Holliday junction while interacting with Mus81-Mms4 (the Holliday junction resolvase), stimulating its DNA cleavage action (Matulova et al., 2009).

1.5 Rad51 and its Paralogs

The current study focuses on Rad51 as it is one of the key proteins in homologous recombination. The primary functions of Rad51 are to form the Rad51 nucleoprotein filament on ssDNA, perform the homology search, and initiate strand invasion and initial base-pairing with template DNA. Rad51 null mutants in yeast are viable, but show an increased sensitivity to DNA-damaging agents and recombination defects (Sonoda et al., 1998; Symington, 2002).

Although yeast is able to survive without Rad51, knocking out Rad51 results in early embryonic lethality in mice (Lim and Hasty, 1996; Tsuzuki et al., 1996), further demonstrating how important Rad51 is to mammalian systems. Rad51 is a highly conserved protein in eukaryotes and a functional homolog of the Escherichia coli protein RecA (Symington, 2002).

Rad51 contains a central ATPase domain that contains Walker A and Walker B motifs that are necessary for ATP binding and hydrolysis (Benson et al., 1994). Two variants of

Rad51 have been used to study the function of these motifs, Rad51K133A and Rad51K133R, which

are deficient in ATP binding and ATP hydrolysis respectively (Morrison et al., 1999; Chi et al.,

2006). Rad51K133A and Rad51K133R correspond to Rad51K191A and Rad51K191R in

Saccharomyces cerevisiae (Sung and Stratton, 1996), and can be used to assess the requirement for ATP binding and hydrolysis by Rad51 during homologous recombination or ectopic homologous recombination.

The N-terminus of Rad51 is the most highly conserved portion of Rad51 across eukaryotes and is involved in DNA binding. It is essential that this DNA binding domain of

Rad51 be conserved, as one of the main functions of Rad51 monomers is assembly onto ssDNA to form a right-handed helical nucleoprotein filament involved in homology searching and strand invasion. Rad51 has equal affinity for both dsDNA and ssDNA (Sung and

Robberson, 1995), and as it needs to bind to ssDNA preferentially, it is aided by Rad52 and

BRCA2 which recruit it to ssDNA specifically and help displace RPA from the ssDNA (Liu et al., 2010; Thorslund et al., 2010). During homologous recombination, Rad51 is recruited to the single stranded DNA (ssDNA) and forms the nucleoprotein filament after the displacement of

RPA (Yu et al., 2001; Mazin et al., 2003). Rad52 and BRCA2 also help stabilize the Rad51 nucleoprotein filament during its formation on the ssDNA (Jensen et al., 2010).

Rad51 monomers must be bound to ATP, inducing a conformational change in the protein to its active form, in order to assemble into the nucleoprotein filament (Pellegrini et al.,

2002). ATP hydrolysis is not needed for the assembly of the nucleoprotein filament (Bugreev and Mazin, 2004; Chi et al., 2006), but is required for its disassembly once the homology search is complete and base pairing begins (Modesti et al., 2007). The search for a homologous donor sequence is carried out by this ATP-bound Rad51-ssDNA nucleoprotein filament (West,

2003). The nucleoprotein filament is then able to undergo strand invasion to form a

heteroduplex with the homologous donor segment of DNA (Baumann et al., 1996; Gupta et al.,

1997). The ATPase function of Rad51 is important for strand invasion and homologous pairing between the invading strand and the region of homology (Sung, 1994). Rad51 can be dissociated from ssDNA through an increase in its ATPase function brought on by several helicases, namely Srs2 in yeast or BLM in humans (Antony et al., 2009).

There are several proteins in both yeast and mammals which share 20-30% of their sequence with Rad51, identifying them as paralogs of Rad51, which act as accessory proteins in HR. Suwaki et al. (2011) reviewed the known roles of Rad51 paralogs and state that complexes of these proteins are involved in genome maintenance and HR pathways. In yeast there are two Rad51 paralogs, Rad55 and Rad57, whereas in mammals there are five Rad51 paralogs, Rad51B, Rad51C, Rad51D, XRCC2, and XRCC3 (Daboussi et al., 2002; Symington,

2002). Rad55 and Rad57 form a heterodimer that interacts with Rad51 and ssDNA (in vitro and in vivo). Weak ATPase activity is detected in both of these proteins and they both contain a conserved ATPase domain (Sung, 1997). Hays et al. (1995) showed that a deficiency of Rad55 and Rad57 caused cells to become sensitive to IR damage but this phenotype can be rescued by the overexpression of Rad51 and Rad52. The mammalian Rad51 paralogs also interact with one another to form stable complexes, Rad51B interacts with Rad51C, Rad51C interacts with

XRCC3, and Rad51D interacts with XRCC2 (Sung et al., 2003). These complexes have been shown to bind to many structures involved in HR in vivo such as ssDNA, dsDNA (with or without a 3’ tail), and Holliday junctions (Sung et al., 2003). Knocking out these proteins in chicken DT40 cells and hamster cell lines lead to an increase in sensitivity to both DNA crosslinking agents and IR; these cells were also hindered with respect to Rad51 foci formation and HR events (Takata et al., 2001). Although chicken DT40 cells and hamster cell lines are

still viable when Rad51 paralogs are knocked out, knocking these genes out in mice results in embryonic lethality indicating that these proteins are critical for development (Takata et al.,

2001).

1.6 Mouse Hybridoma Cells and HR

Cell lines derived from the mouse Sp6/HL hybridoma (Kӧhler and Shulman, 1980) were used in the current study. Sp6/HL cells produce a functional IgM antibody that specifically binds to the hapten 2,4,6-trinitrophenyl (TNP). The production of this antibody can be used to detect these cells via a TNP-specific plaque assay, described in detail in Materials and Methods of this thesis (Section 2.6). The Sp6/HL hybridoma has lost the expression of the gamma and kappa chains from the myeloma cells and has retained a single copy of the TNP- specific μ gene originating in the splenic B cells. The single copy of the TNP-specific µ heavy chain gene in the Sp6/HL hybridoma is a particularly useful feature, since, if that one copy is disrupted, the cells can no longer produce this functional antibody (Kӧhler and Shulman,

1980). Several derivatives from the Sp6-HL line were isolated that bear deletions in the Cμ region, ranging from 2bp to 4kb in size (Kӧhler et al., 1982). Due to these Cμ deletions, the cells produce a non-functional, truncated version of the TNP-specific antibody (Kӧhler et al.,

1982). The structure of the immunoglobulin µ locus in the Sp6/HL-derived igm482 hybridoma is shown in Figure 7. The igm482 hybridoma bears a 2bp Cμ3 deletion as indicated by the triangle representation in Figure 7B.

Homologous recombination and ectopic homologous recombination have been extensively studied in these and other cell lines derived from Sp6-HL. Ectopic homologous recombination between the endogenous Cμ region and a wild-type Cμ region located elsewhere

Sp6/HL Cµ

1 2 3 4 A) B) igm482 Cµ Endogenous µ 1 2 3 4 locus V TNP Sµ H Cµ exons pRCµ pRC C) Sp6/HL Cµ amp Ectopic wild- 1 2 3 4 type Cµ donor µ igm482 Cµ neo Endogenous µ 1 2 3 4 locus V TNP Sµ H Cµ exons

Figure 7 - pRCμ Vector and Immunoglobulin Locus Structure. A) The pRCμ vector containing a wild-type Cµ region was electroporated into the igm482 hybridoma cell line and incorporated in an ectopic location, resulting in the cell lines pRCμR1 #4 and pRCμR1 #9. B) The structure of the immunoglobulin locus in igm482 cells bearing a 2bp deletion in the Cμ3 region, represented by a solid triangle. C) The structure of the immunoglobulin locus in parental cell lines #4 and #9, along with the ectopic donor incorporated in these cell lines.

in the genome has been shown to restore functional IgM production in hybridomas (Baker and

Read, 1992). Furthermore, it was shown that deletions in the Cμ region of 600bp and 4kb were able to be repaired by ectopic homologous recombination, albeit with a lower efficiency than the repair of a 2bp deletion. The amount of homology required to sustain ectopic homologous recombination was assessed to be between 1.9 and 4.3kb (Baker et al., 1996). It was also shown that a donor sequence on the same chromosome as the endogenous locus resulted in a frequency of ectopic homologous recombination that was as much as 4 orders of magnitude higher than if the donor sequence is located on a separate chromosome (homologous or otherwise). Thus, the homology search to find a donor sequence for recombination can distinguish between sequences on the same chromosome versus sequences on separate chromosomes and locates them preferentially (Shulman et al., 1995; Baker et al., 1996).

A well-established gene targeting assay was also used to study homologous recombination in these cell lines. This assay is based on the ability of a plasmid bearing a wild- type Cμ region to correct the 2bp Cμ3 deletion present in the igm482 chromosomal µ gene and restore functional IgM production in the cells (Baker et al., 1988). Through these experiments, it was established that an excess of wild-type Rad51 enhances gene targeting and cell growth

(Rukść et al., 2007). The stable transgenic expression of the ATP binding and hydrolysis deficient mutants of Rad51 (Rad51K133A & Rad51K133R) significantly reduced gene targeting efficiency (Rukść et al., 2007).

The cell lines that will be used in the present study were derived from the igm482 hybridoma (the cell line with the 2bp Cμ3 deletion) through the transfection and integration of a pSV2neo-derived (Southern et al., 1981) vector that includes a wild-type (Sp6) Cμ region as well as a neomycin resistance gene (pRCµ) (Figure 7A). These cell lines are referred to as

ectopic parent lines (pRCμR1 #4 and pRCμR1 #9) because they have a wild-type donor Cμ sequence stably incorporated in an ectopic position relative to the endogenous igm482 μ gene.

Ectopic homologous recombination can correct the 2bp Cµ3 deletion restoring functional TNP- specific antibody production in the cells (Baker and Read, 1992; Baker et al., 1996). This feature makes these cells an ideal system in which to study EHR.

1.7 Hypotheses and Objectives

In the current study, the role of Rad51 in ectopic homologous recombination is investigated through the expression of wild-type Rad51 and several Rad51 catalytic variants in the ectopic parent cell lines (#4 and #9). Ectopic homologous recombination was tested in these cell lines in the presence and absence of DNA damage. Due to the global homology search necessary for EHR, it is hypothesized that the requirements for Rad51 in ectopic homologous recombination may differ from the requirements in homologous recombination. It was hypothesized that an increase in wild-type Rad51 would be beneficial to EHR while a knockdown of endogenous Rad51 would hinder EHR, as similar results were seen with a gene targeting assay (Rukść et al., 2007). Expression of catalytic mutants of Rad51 were shown to reduce the frequency of gene targeting (Rukść et al., 2007); it was therefore hypothesized that expression of these catalytic mutants would also reduce the frequency of EHR.

It is also hypothesized that inducing DNA damage in these cells would increase opportunities for EHR repair and thus increase the observed frequency of EHR. DNA damage was induced with both MMC treatments and ionizing radiation treatments and both were hypothesized to increase the frequency of EHR since both cause DNA DSBs among other lesions.

Chapter 2: Materials and Methods

2.1 Hybridoma Cell Lines

Sp6/HL

All cell lines used in the experiments presented in this thesis are derived from Sp6/HL, a murine hybridoma cell line obtained from the fusion of splenic B cells from a 2,4,6- trinitrophenol (TNP)-immunized Balb/c mouse and IgG-secreting X63-Ag8 myeloma cells.

The Sp6/HL hybridoma has lost the expression of the gamma and kappa chains from the myeloma cells and has retained a single copy of the TNP-specific μ gene originating in the splenic B cells. Consequently, Sp6/HL secretes an immunoglobulin IgM (κ) that is specific to

TNP (Kӧhler and Shulman, 1980). igm482

The cell line igm482 is a derivative of Sp6/HL that bears a 2 base-pair (bp) deletion in the μ heavy chain gene, specifically in the third exon of the µ gene constant region (Cµ3). The

Cµ3 deletion results in a premature stop codon that results in the formation of a truncated μ heavy chain lacking exon Cµ4. This truncation yields IgM which is unable to polymerize or activate complement. The secreted level of anti-TNP IgM in this variant bearing the 2bp deletion is comparable to that of Sp6/HL (Baumann et al., 1985; Baker et al., 1988). This 2bp deletion can be corrected via HR in an assay where a vector bearing a wild-type Cµ region is electroporated into the cell. This wild-type donor fragment can be used during HR to restore the endogenous Cµ locus and enable the cells in which this occurs to produce full-length, functional, TNP-specific IgM. This mechanism provides a useful method to study HR, as the

frequency of this HR event can be assessed by monitoring the number of cells regaining the ability to produce functional IgM. A complement-dependent lysis assay involving TNP- coupled sheep red blood cells can be used to detect cells in which HR has corrected the 2bp deletion, as plaque forming cells (PFC). The frequency of HR events can be estimated by taking the ratio of PFC to the total number of cells plated. pRCµRI #4 and pRCµRI #9

The hybridoma cell line igm482 was transfected with a vector (pRCµ) (Figure 7A) bearing a wild-type Cµ region and G418R transformant cell lines were obtained and checked by

Southern blot analysis to verify stable integration of one or a few Cµ region copies at a single position (Baker et al., 1996). It was determined that these cells were able to use the ectopic wild-type Cµ region as a donor to correct the 2bp deletion in the endogenous igm482 Cµ region (Baker et al., 1996). Cells that have undergone this EHR event regained the ability to produce functional IgM, and thus were detected as PFC in a TNP-specific plaque assay (Baker et al., 1988). The cell lines pRCµRI #4 and #9 are identical to igm482 with the exception of the single vector integration site bearing approximately two ectopic wild-type Cµ region(s) and a neomycin resistance gene (Lee and Baker, 2007). Both pRCµRI #4 and #9 were shown to have a low vector copy number (~2), via Southern blot and densitometric analysis (Lee and Baker,

2007). pRCµRI #4 & #9 Derived Expressors of Rad51 Variants

The hybridoma cell lines pRCµRI #4 and #9 were transfected with a pSV2hyg-derived plasmid containing each of the following modified sequences of mouse Rad51: (i) wild-type,

FLAG-tagged Rad51; (ii) FLAG-tagged Rad51K133A; (iii) FLAG-tagged Rad51K133R; and (iv)

siRNAs against Rad51 mRNA. These plasmids are shown in Figure 8, and described further in section 2.4. The stable expression of each of these Rad51 variants was achieved in each of the ectopic parent cell lines. Between 30 and 50 HYGR clones from each electroporation were screened for Rad51 expression by Western analysis and the level of expression was assessed via densitometry in comparison with a β-actin loading control (Section 2.5). A summary of all cell lines is presented in Table 1.

Wt Rad51 Rad51K133A

3x FLAG 3x FLAG

800 801

- XmnI E- XmnI E

amp amp Hyg Hyg

Rad51K133R Rad51 siRNA

EcoRI

3x FLAG

906, 908, 812 XmnI E- & 779 XmnI Hyg amp amp Hyg EcoRI

Figure 8 – Plasmid Maps for Generation of Rad51 Variant Expressors. This figure shows the plasmids used to achieve stable transgenic expression of Rad51 variants in the ectopic parent cell lines (vectors 800, 801, and 812). These vectors all contain a FLAG-tag that is expressed along with the Rad51 variant gene. Vector 800 is pFLAGE-Rad51hygwt, vector 801 is pFLAGE-Rad51K133Ahyg, and vector 812 is pFLAGE-hygK133R. This figure also shows the three vectors used to generate Rad51 knockdowns in these cell lines (906, 908, 779), each of which contains a different siRNA sequence. All the vectors contain a single XmnI site by which they can be linearized, as well as resistance genes for selection in mammalian cells (hyg) and bacteria (amp).

Table 1 - Summary of Cell Lines.

Cell line Rad51 expression

Sp6/HL Endogenous Rad51 igm482 Endogenous Rad51 pRCµRI #4 and #9 Endogenous Rad51

4-800- & 9-800- Endogenous Rad51 + wild-type FLAG-Rad51

4-801- & 9-801- Endogenous Rad51 + FLAG-Rad51K133A

4-812- & 9-812- Endogenous Rad51 + FLAG-Rad51K133R

#4 kd & #9 kd Knocked down for endogenous Rad51

2.2 Cell Culture Conditions

All hybridoma cell lines were grown in Dulbecco’s modified Eagle’s medium (DMEM;

Gibco, Grand Island, NY) supplemented with 100 units/ml penicillin and streptomycin,

50 µM 2-mercaptoethanol, and 15% heat-inactivated fetal bovine serum at 37°C with 7% CO2 atmosphere (Kӧhler and Shulman, 1980). For identification and selection of transformants, this media was supplemented with one of two selectable agents: aminoglycoside G418 (600 µg/ml) or hygromycin (700 µg/ml). The cell lines pRCµRI #4 and #9 are G418 resistant due to a neomycin phosphotransferase (neo) gene inserted along with the ectopic donor wild-type Cµ region. The transfected plasmids for the expression of Rad51 variants in these cell lines contained a hygromycin resistance gene which encodes hygromycin B phosphotransferase

(hyg). Both of these resistance genes, neo and hyg, encode kinases that phosphorylate and inactivate each of the chemicals, respectively. Expression of these genes permits selection of

HYG- and G418-resistant transformants (HYGR and G418R) in media supplemented with these chemicals. After generation of each cell line, many copies were frozen from the original culture, so that cells of the same passage number could be used for each experiment.

2.3 Plasmids

All plasmids used were maintained in E. coli strain DH5α that was grown in Luria-

Bertani (LB) broth supplemented with 50 µg/ml ampicillin at 37°C. These plasmids were purified by maxi-prep using PureLink HiPure Plasmid Filter Purification Kits (Invitrogen).

Purified plasmid DNA was then subjected to phenol:chloroform treatment and ethanol precipitation. Plasmids were linearized through a restriction digest with XmnI prior to

electroporation into ectopic parent cell lines pRCµRI #4 and #9. Several plasmids were used in the generation of cell lines for this thesis and are summarized in Figure 8. These plasmids are all derived from the same vector backbone (pFLAGhygE-) and thus contain many of the same features. All the plasmids contain resistance genes for selection in both bacteria (amp) and mammalian (hyg) cell systems. Each plasmid also contains a single XmnI site that is used to linearize the plasmid DNA before it is transfected into hybridoma cells. Each of plasmids 800,

801, and 812 contain a gene for a separate Rad51 variant, namely wild-type Rad51,

Rad51K133A, and Rad51K133R respectively. These plasmids also include an in-frame 3×FLAG- tag at the 5’ end of the Rad51 gene so that it becomes an N-terminal FLAG-tag after translation. The plasmids 906, 908, and 779 each contain an siRNA complementary to a separate region of mouse Rad51 mRNA (one at the beginning, middle, and end).

2.4 Electroporation of Hybridoma Cells

Cell cultures were grown to an appropriate density (~2-4 × 105 viable cells/ml) and a volume of medium containing 2 × 107 viable cells (determined by counting viable cells using trypan blue exclusion) was harvested by centrifugation at 800 rpm for 10 minutes at 4°C. The cells were washed once with 1 × PBS before being resuspended in 0.75 ml permeabilization buffer (140mM KCl, 1mM MgCl2, 1mM ATP, 10mM glucose, 1mM EGTA, 0.2mM CaCl2,

10mM Hepes). Cells suspended in permeabilization buffer were transferred to a cuvette with a

4 mm electrode gap (BioRad Inc.) with 50 µg linearized, phenol/chloroform-treated and ethanol-precipitated plasmid. The cells were then pulsed twice in a BioRad Gene Pulser

(BioRad Inc.) at 700 V, 25 µF and incubated on ice for 10 minutes. One ml of DMEM was then added to the cuvette and the cuvette was transferred to a 37°C, 7% CO2 incubator for a

recovery period of 20 minutes. The cells were then suspended in an appropriate volume of

DMEM in order to be plated into 96-well microtitre plates at limiting dilution to permit the recovery of individual HYGR clones (Baker, 2004). Selective medium supplemented with hygromycin was added to these plates 24h after the electroporation for a final concentration of

700 µg/ml. Colonies of HYGR transformants became visible 9-12 days after the electroporation; individual colonies were transferred to 24-well plates and saved by cryopreservation for later analysis.

2.5 Western Blot Analysis

Western immunoblotting was used to detect stable transgenic expression of the Rad51 variants used in this study, and to assess the level of expression via densitometry.

Approximately 5×106 hybridoma cells were harvested via centrifugation and lysed with non- denaturing buffer (1% NP-40, 137mM NaCl, 20mM Tris-Cl pH 8.0, 2mM EDTA, 10% glycerol, 1 tablet protease inhibitor per 10ml (Roche)). Total protein concentration was determined via bicinchoninic acid (BCA) assay. An equal volumes (50 µg) of each whole cell extract was loaded onto a 10% Sodium Dodecyl Sulphate Polyacrylamide Gel (SDS-PAGE).

Proteins were then Western transferred to polyvinylidene fluoride (PVDF) membrane at 30 V and 4°C overnight in transfer buffer (Volume 1L: 200 ml methanol, 3.03g Tris base, 14.4g glycine). The PVDF membrane was then blocked for 1.5h at room temperature with Tris-

Phosphate Buffer and 3% non-fat milk proteins. Next, the membrane was incubated for 1-2h with a primary antibody specific to the protein under investigation. This incubation was followed by incubation for 1h with an appropriate secondary antibody conjugated with horse- radish peroxidase (HRP). Table 2 gives a summary of the antibodies used in this set of

Table 2 - Diagnostic Antibodies.

Target Protein Primary Antibody Secondary Antibody

Mouse Rad51 Mouse monoclonal anti-human HRP-conjugated goat anti-mouse Rad51 14B4 (Abcam) IgG (Southern Biotech)

Mouse β-actin Mouse monoclonal anti-human HRP-conjugated goat anti-mouse β-actin AC-15 (Sigma) IgG (Southern Biotech)

FLAG-tag Mouse monoclonal anti-FLAG HRP-conjugated goat anti-mouse M2 (Sigma) IgG (Southern Biotech)

experiments and their targets. ECL Plus reagent (GE Healthcare) is then added to the HRP- coupled protein bands, which results in chemiluminescence that was detected using X-ray film.

2.6 TNP-Specific Plaque Assay

This assay is used to detect correction of the 2bp deletion in the igm482 Cµ3 region and the production of full-length TNP-specific IgM in the recombinant cells. This correction can take place via EHR using the ectopic donor wild-type Cµ region in the pRCµRI #4 and #9 derived cell lines, or via HR, through a wild-type Cµ region supplied on a vector in a gene targeting assay. Both of these methods for correcting the endogenous Cµ region produce plaque forming cells (PFC) by restoring the ability of these cells to initiate the complement- dependent lysis of TNP-SRBCs (Baker et al., 1988).

When measuring EHR in a cell line, the cultures are grown to a density of ~2-4×105 cells/ml, and a volume containing 1×107 cells per plate was harvested by centrifugation. The cells were washed once with 1× PBS and then added to 2.7 ml of 0.5% agarose along with 100

µl TNP-SRBCs (1:2.5 with PBS). The agarose, TNP-SRBCs and hybridoma cells were mixed thoroughly and plated. Plates were then incubated at 37°C in 7% atmospheric CO2 for 1-1.5h.

After this incubation the plates were flooded with 2 ml each of a 1:10 guinea pig complement:PBS dilution. Plates were then returned to the same incubator for another 1-1.5h incubation. Plaques become visible during this time and are counted. Plates are returned to the incubator overnight and the count of PFCs is verified the next morning. Two to three plates are assayed for each cell line. A schematic representation of this assay is shown in Figure 9.

Due to fluctuations in EHR frequencies in parental cell line #4 (see Results Section 3.3), subcultures of all cell lines were generated (Luria and Delbrück, 1943). All cell lines described here are originally obtained from single cells after limited dilution plating from an electroporation (described above, section 2.4). A schematic representation of subculture generation is shown in Figure 10. Subcultures (15 of parent #4 and 3 of all other cell lines) were made from each cell line by depositing 100 cells into individual wells of a 24-well plate.

Once each subculture grew up to the appropriate density (2-4 × 107 cells/ml) they were assayed for EHR by the TNP-specific plaque assay described above.

The gene targeting assay (Baker et al., 1988) involves the transfection of a gene targeting vector (pRCµ) into the cells. This gene targeting vector contains a 9.7 kb wild-type Cµ fragment (Figure 7) which is released through an EcoRI digest before being electroporated into

2×107 recipient cells. A recovery period of 48-72h is allowed before the measurement of PFCs is conducted as described above. The survival of the cells was assayed 24h after electroporation by trypan blue exclusion, compared to a non-electroporated control culture and ranged from 15% to 60%.

TNP-SRBCs Washed + hybridomas Agarose

Plate + Allow agarose to set

1) Incubate 1h 2) Add complement 3) Incubate 1h

Plaques from IgM- secreting cells Number of plaques = number of PFC

Figure 9 – TNP-Specific Complement Dependent Plaque Assay. Hybridoma cells (1×107 per plate) are collected by centrifugation, washed with 1×PBS, added to TNP-coupled SRBCs and agarose, and plated. After a 1h incubation at 37°C in 7% atmospheric CO2 the plates are flooded with 2 ml of guinea pig complement diluted 1:10 with PBS. Plaques become visible after a second 1h incubation and are counted. Each plaque represents one PFC.

Figure 10 – Subculture Generation. Following electroporation, cells are plated into 96-well tissue culture plates at a density of 6000 cells/well with an appropriate selection agent so that the colonies that will grow have started from a single cell (A). Several of these clones are screened for transgene expression through western blotting. Transgene expressors are then cultured in flasks (B). For each cell line that is chosen as an appropriate transgene expressor, subcultures are made starting with 100 cells each to ensure a more accurate picture of the frequency of recombination (C). Once these subcultures reach an appropriate density ~2-4 x 105 cells/ml, they are plaque assayed in triplicate with each subculture of each cell line having 3 replicate plates, each with 1 x 107 cells.

2.7 DNA Damage

DNA damage was induced through the addition of MMC to the cell cultures or by exposing them to ionizing radiation (IR). MMC is a cross-linking DNA damaging agent that may cause double strand breaks in DNA. Ionizing gamma radiation causes a variety of DNA lesions including double strand breaks.

Mitomycin C (MMC)

An untreated control flask was also included for every cell line tested. After either a 6h or 24h incubation in the DMEM supplemented with MMC the cells were re-harvested by centrifugation, washed once with 1× PBS and then resuspended in DMEM for a recovery period of 48-72h. After the recovery period, the PFCs were detected using the TNP-specific plaque assay as above.

Ionizing Gamma Radiation (IR)

Hybridoma cell cultures were grown to a density of 2-4×105 cells/ml and an appropriate volume containing 2×107 cells was harvested by centrifugation. These cells were resuspended in DMEM media at 2×106 cells/ml and subjected to 4 Gy ionizing radiation. This value was chosen since it is intermediate in the range (3-5 Gy) used by Rukść et al. (2007). An untreated control flask was also included for every cell line tested. After irradiation treatment the cells

underwent a recovery period of 48-72h. After the recovery period, the PFC were detected using the TNP-specific plaque assay as above.

2.8 Statistical Analysis

Student’s t-tests for one and two samples were performed using VassarStats software which is available online at www.vassarstats.net. Significance was assessed at the p < 0.05 level.

Chapter 3: Results

The goal of the experiments in this study was to gain information about the role of

Rad51 in ectopic homologous recombination. In order to achieve this goal, several variants of

Rad51 were stably expressed in ectopic cell lines, and these cell lines were put through several different assays including TNP-specific plaque assays, gene targeting and DNA damage treatments.

3.1 Stable Expression of FLAG-tagged Wild-type Rad51 and Rad51 Catalytic Variants

The cell lines generated for this set of experiments ranged in the type and amount of

Rad51 that they express. Cell lines with higher than endogenous and lower than endogenous wild-type Rad51 levels were generated, as well as cell lines expressing two variants of Rad51:

Rad51K133A and Rad51K133R. The methods used to generate these cell lines are described in detail in Sections 2.3-2.5 of this thesis. Briefly, the parent cell lines (pRCµRI #4 and #9) were grown to a density of ~2-4×105cells/ml and 2×107 cells were electroporated with the appropriate linearized vector (or group of vectors for Rad51 knockdowns), as described in

Figure 8. After electroporation the treated cells were plated at limiting concentration in DMEM supplemented with hygromycin (700 µg/ml). This allowed for single clonal populations to arise in the plates. Thirty to fifty clonal populations per electroporation were screened by

Western blot analysis to determine putative expressors of the Rad51 variant of interest, or a level of knockdown of Rad51. After initial Western screening, several clones from each electroporation were picked and re-screened to confirm the levels of Rad51 variant expression.

Westerns showing expression levels of excess wild-type Rad51, Rad51K133A and

Rad51K133R in both parent lines are presented in Figure 11. The top sections in all the Western blots shown in this figure are visualized with a mouse Rad51-specific antibody (14B4). These sections show both the endogenous Rad51 (~37 kDa) band and the transgenic FLAG-tagged

Rad51 band (~42 kDa) for all cell lines. The middle bands in all the Western blots shown in this figure are visualized with an anti-FLAG antibody, and these bands run at ~42 kDa, the predicted size of Rad51 with the addition of the 3× N-terminal FLAG tag. The bottom bands represent the β-actin loading control for each of these Western blots (each lane received 50 µg of whole cell extract). The Western blots are separated between the two different original parent lines (#4 in Figure 11A and #9 in Figure 11B) as well as by which Rad51 variant these cells are expressing. Densitometric analysis was performed and the ratio of the FLAG-tagged

Rad51 band (visualized with an anti-FLAG antibody) to that of β-actin is presented below each lane. A higher FLAG-tagged Rad51:β-actin ratio indicates a higher expression of the particular

Rad51 protein in each cell line.

Westerns showing the levels of Rad51 knockdowns in several clones are presented in

Figure 12. The top bands in these Westerns represent levels of endogenous 37 kDa Rad51 and are visualized with an anti-Rad51 antibody (14B4). The bottom bands represent β-actin loading controls for each of these Western blots (each lane represents 10 µg of whole cell extract in order to more clearly see any knockdown effect). The Westerns present Rad51 knockdowns for the two original parent lines (#4 and #9). Densitometry was used to measure the intensity of each of the bands and the values are presented as ratios relative to the respective parental cell lines. The values presented under each lane represent the portion of endogenous Rad51 that remains in the cells as a result of the siRNA knockdown. The Rad51 knockdowns in the parent

cell line #4 were generated previously in our lab (Magwood, unpublished results), but the

Rad51 knockdowns in the parent cell line #9 were generated for this thesis.

A) Wild-type Rad51 Rad51 Rad51 K133A K133R

8 9 1

4 25

16 7 1

35 17

2 4

- - -

- -

- - -

- -

- RI RI #4

801

800 800 801

801 812

800 800 812

801

801 801

800 800

812

- - -

- -

- - -

- -

4 4 4

4 pRC 4

4 4 4

4 4

4 4 4 FLAG-Rad51 (42 kDa) Endogenous Rad51 (37 kDa)

FLAG-Rad51 (42 kDa)

β-actin

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

0 1.5 3.3 2.5 1.2 2.7 2.7 1.9 1.3 1.4 1.5 1.0 0.7 7.7 1.2 0.8 Rad51/β-actin

B) Wild-type Rad51 Rad51 Rad51

K133A K133R

8 6

7 19 21

34 4 6 17 23

- -

- - -

- - - - -

RI #9 RI

800 801

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800 801

800 812 812

801

812 800 800 800 812

812

- -

- - -

- - - - -

pRC 9 9

9 9

9 9 9

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9 9

FLAG-Rad51 (42 kDa) Endogenous Rad51 (37 kDa) FLAG-Rad51 (42 kDa)

β-actin 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

0 0.7 0.8 2.5 1.8 1.4 1.6 0.9 0.9 0.9 0.9 2.9 6.6 0.9 0.6 0.8 Rad51/β-actin

Figure 11 - Western Analysis of Rad51 Expression. A) Westerns showing the expression levels of Rad51 variants in the parent cell line pRCμRI #4. B) Westerns showing the expression levels of Rad51 variants in the parent cell line pRCμRI #9. The band at 37 kDa is the endogenous Rad51 visualized with a Rad51-specific antibody (14B4). The bands at 42 kDa are transgenic Rad51 present in these cells and are visualized by incubation with 14B4 in the top panel of A and B, and by an anti-FLAG antibody in the middle panels of A and B. All Westerns are shown alongside their β-actin loading controls. The ratios under each lane represent the densitometric ratio of the FLAG-tagged Rad51 band (visualized with an anti-FLAG antibody) to the β-actin band (higher ratios indicate higher transgenic Rad51 expression).

RI RI #9 RI #4

μ μ

#9 Rad51 #9 kd24

#9 Rad51 #9 kd12 Rad51 #9 kd16 Rad51 #4 kd15

#4 Rad51 #4 kd1

#4 Rad51 #4 kd34

pRC pRC

14B4 - Rad51 (37 kDa)

β-actin

1 2 3 4 5 6 7 8

1 0.46 0.46 0.37 1 0.60 0.55 0.63 Rad51/β-actin

Figure 12 - Western Analysis of Rad51 Knockdowns. Westerns showing the expression levels of Rad51 in knockdowns derived from the parent cell lines pRCμRI #4 and #9 alongside the respective parent lines. Each lane received 10 µg of whole cell extract protein. Wild-type Rad51 runs at 37 kDa and is visualized here with a Rad51-specific antibody (14B4). β-actin loading controls are shown for each Western. The ratios under each lane represent the densitometric ratio of the 14B4 band to the β-actin band (lower ratios indicate more effective knockdown).

3.2 Effect of Wild-type Rad51 Expression on Ectopic Homologous Recombination

The effect of varied expression of wild-type Rad51 and the Rad51K133A and Rad51K133R catalytic variants on EHR was of interest in this study. The first step in this process was to assess the level of ectopic recombination in the parent cell lines pRCµR1 #9 and pRCµR1 #4, as well as in the highest expressors of FLAG-tagged wild-type Rad51 and in the best knockdowns of endogenous Rad51 that were obtained. Figure 13 shows the levels of ectopic recombination in the parent cell lines. Values are reported as the number of TNP-specific plaque forming cells (PFCs) per 107 cells plated. PFCs are determined by counting the plaques that form on a plate of TNP-coupled sheep red blood cells as outlined in Section 2.3 of this thesis. Briefly, only cells in which the 2bp deletion in the endogenous igm482 Cµ3 region has been corrected are able to form functional TNP-specific IgM and thus appear as plaque forming cells in this assay. Therefore, each plaque that is observed represents a single recombinant cell. The parent cell line pRCµR1 #4 (or parent #4) revealed a mean EHR frequency of ~100 PFC/107 cells plated. In order to validate the frequency of EHR in this cell line, a fluctuation analysis (Luria and Delbrück, 1943) was performed using 15 subcultures each generated from 100 cells so that they would not initially contain any PFCs. The subcultures were assayed for EHR and found to have a mean frequency of EHR of 25 PFC/107 cells plated. This suggests that the higher frequency of PFC in the parent #4 cell line is due to recombinants that were generated during expansion of this cell line (i.e., a “jackpot”). Similar studies were conducted with the parent #9 cell line. However, as shown in Figure 13, these subcultures were not different in EHR frequency

120

100

cells

7 80

40 NumberofPFC/10 20

0 pRCuR1 #4 pRCuR1 #4 pRCuR1 #9 pRCuR1 #9 subclones subclones

Figure 13 – EHR in Parental Cell Lines #4 and #9. Original parent cell lines are shown beside the value obtained through assaying subcultures (each started from 100 cells) of each parent line. For parent #4, 15 subcultures were assayed, compared to 3 subcultures for parent #9. The value for pRCμR1 #4 and the subcultures of this cell line are statistically different from each other (p<0.01) in a Student’s t-test. None of the other values are significantly different from each other using the same test. Error bars represent the mean +/- the standard error of the mean. Number of independent measurements varies in this data (pRCµRI #4: 5 experiments, 3 plates per experiment, n=15; #4 subcultures: 15 subcultures, 2 plates per subculture, n=30: pRCµRI #9: 4 experiments, 3 plates per experiment, n=12; #9 subculture: 3 subcultures, 3 plates per subculture, n=9).

from the original #9 parent line. All further experiments were done with subcultures of both cell lines.

The transgenic expression of Rad51 variants in each of the parent cell lines #4 and #9 could lead to changes in the frequency of EHR in these cell lines. Therefore, levels of EHR were measured using the same TNP-specific plaque assay as described above. In order to more clearly show the difference in plaquing frequency of the Rad51-expressing cell lines, EHR is reported as fold changes in plaquing frequency compared to the respective parent lines. Figure

14 shows the fold changes in EHR for the expressors of FLAG-tagged wild-type Rad51 in each of the parental cell lines. The cell lines 4-800-4 and 9-800-8 are the highest expressors of wild- type Rad51 in their respective groupings and the values for these cell lines are significantly lower than their respective parent cell lines. Another expressor, 4-800-2, is also significantly lower than the parental #4 cell line. The other cell lines shown here are not statistically different from their respective parent lines. The data suggest that an increase in the amount of wild-type Rad51 does not increase EHR and in fact, with higher levels of wild-type Rad51 expression, may actually decrease it.

A) B) 1.2 1.2

1 1

0.8 0.8

0.6 0.6

0.4 0.4 Foldchange in EHR

Foldchange in EHR

0.2 0.2

0 0 pRCuR1 4-800-2 4-800-8 4-800-4 pRCuR1 9-800-6 9-800-7 9-800-8 #4 #9

Figure 14 – Effect of FLAG-tagged Wild-type Rad51 Expression on EHR. These graphs show the fold change in EHR in cells over-expressing wild-type Rad51 compared to each of the parent cell lines. Both of these graphs present EHR in the Rad51-expressors as a fold-difference compared to the respective parent line. The expressors are arranged in the order of lowest to highest Rad51 expression as per the FLAG/β-actin ratios given in Figure 11. The highest Rad51 expressers are 4-800-4 and 9-800-8, these cell lines (along with 4-800-2) are significantly reduced for EHR from their parent lines (p<0.05). The other cell lines shown here (4-800-9, 9-800-6 and 9-800-7) are not statistically different from the parent lines. Significance was determined by a Student’s T-test. These values are based on 2 (parent #4, A) or 3 (parent #9, B) separate experiments, and for each experiment, 3 plates per cell line (n=6 in A, n=9 in B).

3.3 Effect of Rad51 Depletion on EHR

Rad51 knockdowns (Figure 12) were also assessed for any changes in their EHR frequencies. With a decreased amount of wild-type Rad51, it was expected that Rad51 depleted cells would be deficient in EHR. Figure 15 shows three knockdown cell lines per parental cell line, and the EHR frequencies are presented as fold changes relative to the parent cell lines.

The data shows that all of the Rad51 knockdown cell lines have lower EHR frequencies

(p<0.05) than their respective parent lines with the exception of knockdown cell line #15 which is not significantly different from parental #4 cells. In the parental cell line #4, knockdown #15 had a much higher EHR frequency in one of the two experiments conducted. As such, the higher frequency (a potential outlier) experiment was eliminated to present the other experiment as the grey bar in Figure 12A. This shows that the EHR frequency in knockdown

#15 in the parental cell line #15 is probably lower than the value presented for the mean of the two experiments. As indicated by the densitometric analysis presented in Figure 12, these cell lines have similar levels of endogenous Rad51 which accounts for the reduction in EHR in these cell lines. The Rad51 knockdown cell lines in the parental cell line #9 retain 37-46% of endogenous Rad51, whereas the Rad51 knockdown cell lines in the parental cell line #4 retain

55-63% of endogenous Rad51.

Fold change in EHR of Rad51 Fold change in EHR of Rad51 knockdowns in pRCuR1 #4 knockdowns in pRCuR1 #9 1.2 1.2

1 1

0.8 0.8

0.6 0.6

0.4 0.4

Foldchange in EHR Foldchange in EHR 0.2 0.2

0 0 pRCuR1 #4 kd1 #4 kd15 #4 kd34 pRCuR1 #9 kd12 #9 kd16 #9 kd24 #4 #9

Figure 15 – Effect of Rad51 Depletion on EHR. Fold change in ectopic homologous recombination frequencies in Rad51 knockdowns compared to the parent cell lines #4 and #9. All knockdowns with the exception of knockdown #15 in the parent cell line #4 are significantly lower (p<0.05) than their respective parent lines (Student’s t-test). For knockdown #15 in the parental cell line #4, one of the experiments yielded a much higher EHR frequency, as such it is eliminated to present the other experiment as the grey bar. There is no error bar for this bar as it is from a single experiment. Error bars shown represent the mean +/- the standard error of the mean. Fold differences in EHR were calculated per experiment and the ratios for two different experiments (3 plates per cell line, per experiment, n=6) were averaged to yield the values shown here.

3.4 Effect of Rad51 Catalytic Variants on EHR

In each of the parent cell lines #4 and #9, Rad51 variants deficient in ATP hydrolysis and binding were also expressed. Rad51K133A is a variant of Rad51 that has been shown to be deficient in ATP binding and hydrolysis and Rad51K133R is able to bind, but cannot hydrolyse

ATP (Chi et al., 2006). Figure 16 shows the fold changes in EHR when the variant expressed is

Rad51K133A. The cell lines in the first graph are in order from least transgene expression (4-

801-17) to most transgene expression (4-801-4) based on the densitometric ratios presented in

Figure 11, although the levels of expression do not vary greatly. The cell lines presented in the second graph all have very similar, if not the same, levels of Rad51K133A expression. All cell lines expressing this Rad51 variant are significantly reduced (p<0.05) for EHR as compared to their respective parent lines. This result is expected since the Rad51 variant being expressed is missing the ATP hydrolysis functionality, which is hypothesised to be necessary for EHR.

These cell lines retain a normal level of endogenous wild-type Rad51 and so are still able to undergo some EHR. It is interesting to note that the level of EHR depression in these cell lines does not seem to correlate with the level of transgenic expression of Rad51K133A, although for the various cell lines, the Rad51K133A expression levels do not vary enormously, if at all Figure

11.

Fold change in EHR of Fold change in EHR of

Rad51K133A expressors in Rad51K133A expressors in pRCuR1 #4 pRCuR1 #9 1.2 1.2

1 1

0.8 0.8

0.6 0.6

0.4 0.4

Foldchange in EHR Foldchange in EHR

0.2 0.2

0 0 pRCuR1 4-801-17 4-801-35 4-801-4 pRCuR1 9-801-6 9-801-27 9-801-36 #4 #9

Figure 16 – Effect of Expression of Rad51K133A on EHR. Fold change in ectopic homologous recombination frequencies in expressors of Rad51K133A compared to the parent cell lines #4 and #9. EHR in all of these cell lines is significantly lower (p<0.05) than in the respective parent lines (Student’s t-test). The mean +/- the standard error of the mean is presented. Fold differences in EHR were calculated per experiment (n=3 per experiment) and then two experiments were averaged to yield the values shown here.

Figure 17 shows the fold changes in EHR when the variant expressed is deficient in

ATP hydrolysis but is still able to bind ATP (Rad51K133R). The only two cell lines which are significantly lower than their respective parent lines are 4-812-25 and 9-812-21 (p<0.05). The other cell lines are not statistically different from their respective parent cell lines. For 4-812-3 and 4-812-1 in the first experiment, one of the subcultures for each of these cell lines was much higher than the others. These outlying subcultures were eliminated in the grey bars presented in Figure 17A, and these revised bars show the same data excluding the subcultures that were higher. These revised bars are also not significantly different from the parental cell line #4. It was expected that expression of Rad51K133R would decrease EHR in these cell lines, perhaps for the same reasons stated above for Rad51K133A. It is important to note that these cell lines all still retain a normal endogenous level of wild-type Rad51 so the effect is not as severe as it could be if more of the Rad51 in the cell were of this variant. The Rad51K133R variant retains the ability to bind ATP. Thus, ATP-bound Rad51 may still be able to function somewhat in the context of endogenous Rad51 in promoting EHR. However, it is not as efficient as endogenous wild-type Rad51 in this respect. As with the Rad51K133A expressors above, the level of expression of this variant of Rad51 does not appear to correlate to the level of reduction in EHR, in fact, it is the cell lines with lower transgenic expression which show a significant decrease in EHR. Although, there is some evidence that over-expression of Rad51 catalytic variants can actually serve to increase HR; it is possible that this over-expression compensates somewhat for the ATPase deficiency (Forget et al., 2007).

Fold change in EHR of Fold change in EHR of

Rad51K133R expressors in Rad51K133R expressors in pRCuR1 #4 pRCuR1 #9 1.4 1.4

1.2 1.2

1 1

0.8 0.8

0.6 0.6 Foldchange in EHR Foldchange in EHR 0.4 0.4

0.2 0.2

0 0 pRCuR1 4-812-25 4-812-3 4-812-1 pRCuR1 9-812-21 9-812-19 9-812-34 #4 #9

Figure 17 – Effect of Rad51K133R Expression on EHR. Fold change in ectopic homologous recombination frequencies in expressors of Rad51K133R compared to the parent cell lines #4 and #9. Only 4-812-25 and 9-812-21 are significantly lower (p<0.05) than their respective parent lines. All other expressors shown are not significantly different from their respective parent lines. For each 4-812- 3 and 4-812-1 the grey bars represent the same data with the elimination of one culture from the first experiment that was abnormally high (a potential outlier). The mean +/- the standard error of the mean is presented. Fold differences in EHR were calculated per experiment (2 subcultures per experiment, 2 plates per subculture, n=4 per experiment) and then two experiments were averaged to yield the values shown here.

3.5 Gene Targeting

Excess wild-type Rad51 has been shown to increase HR (Rukść et al., 2007). Therefore, it was of interest to examine this pathway of HR in cell lines that were tested for EHR. The data presented in section 3.3 supports a role for excess wild-type Rad51 in decreasing EHR and it was expected that gene targeting would be stimulated in these cell lines. Conversely, Rad51 knockdown cell lines, which show a reduction in EHR, would be expected to show a reduced level of gene targeting. The gene targeting assay exploits the TNP-specific plaque assay to detect plaque forming cells after the transfection of a wild-type Cµ region and following sufficient time for HR with the endogenous igm482 Cµ region to take place. Importantly, the plaque forming cells that form as a result of gene targeting are in addition to those that form by

EHR.

Table 3 shows the gene targeting results for cell lines expressing FLAG-tagged wild-type

Rad51 in the context of endogenous Rad51. The results are reported as a ratio of the number of

PFC/107 cells obtained through gene targeting versus the number obtained through EHR for each cell line. The ratios were calculated for each separate experiment and then averaged over the different experiments to get the values reported in the mean column. The next column shows the fold change in the means for each cell line compared to the parental cell line #9. All cell lines expressing FLAG-tagged wild-type Rad51 are significantly (p<0.05) increased for gene targeting compared to the parental cell line #9, with the exception of 9-800-15. These cell lines show a 2-4 fold increase in gene targeting as shown by Rukść et al. (2007). In the first column of Table 3 the cell lines are arranged from lowest to highest expression levels of

FLAG-tagged wild-type Rad51. The cell line 9-800-8 is the highest expressor of wild-type

Table 3 - Gene Targeting Results

Gene Significance Cell linea Experiment Meanc Fold Changed Targeting:EHRb from #9e 1 11.90 2 7.63 pRCµRI #9 3 8.69 8.32 1 N/A 4 5.90 5 7.47 1 21.77 9-800-4 16.23 1.95 p=0.033 2 10.70 1 (60.64)f 21.44 2.58 p= 0.0005 9-800-6 2 19.84 (34.51) (4.15) (p=0.028) 3 23.04 1 23.27 9-800-15 2 12.55 16.12 1.94 p=0.069 3 12.55 1 22.50 2 31.50 3 27.20 9-800-8 31.13 3.74 p=0.0003 4 38.86 5 23.35 6 43.36 a Cell lines assayed with gene targeting, arranged from lowest to highest FLAG-tagged wild- type Rad51, based on densitometric values presented in Figure 11 b Number of PFC obtained through gene targeting assay per number of PFC obtained through EHR assay c Average of gene targeting:EHR ratios d Fold change of each cell line relative to the parental cell line #9 e Significance was determined through individual Student’s t-tests comparing each cell line to the parental cell line #9. Significance was determined at α=0.05, thus 9-800-4, 9-800-6, and 9-800-8 are significantly higher than the parent #9 while 9-800-15 is not significantly different from the parent #9 f This value is much higher than the other two for this cell line and is thus considered to be a outlying event. Values in brackets include this value and those not in brackets do not include this value.

FLAG-tagged Rad51 and consequently it shows the greatest increase in gene targeting frequency. This result is consistent with the hypothesis that excess wild-type Rad51 is beneficial to HR by gene targeting, but hinders or is of no use to EHR.

3.6 Gene Targeting after 24h MMC Treatment

It is hypothesized that the efficiency of gene targeting depends on the amount of functional Rad51 that is present in the nucleus to help these recombination events take place. It was previously observed that exposing cells to MMC damage induced Rad51 entry into the nucleus, with maximum Rad51 in the nucleus after 24h (Magwood et al., 2012). Figure 18 shows the effects of gene targeting in the hybridoma 9-800-8 (the highest expressor of FLAG- tagged wild-type Rad51) after a 24h MMC treatment of this cell line. The values in this figure represent the ratio of PFC observed after 24h MMC treatment and gene targeting to those obtained through just EHR. The value for 9-800-8 is significantly greater than that for the parental cell line #9 (p<0.05). This demonstrates that in the presence or absence of DNA damage induced by MMC, excess Rad51 is able to stimulate gene targeting 2-3 fold as seen in the previous section. The fact that the Rad51 is in the nucleus prior to the gene targeting assay does not increase this stimulation. The values reported for the gene targeting:EHR ratios in this experiment are slightly lower than in the previous section, possibly due to the DNA repair machinery also being used to repair any MMC-induced DNA damage.

Ratio Ratio 24h of MMCto GT + EHR 4

0 pRCuR1 #9 9-800-8

Figure 18 – Effect of MMC Treatment on Gene Targeting. Cells were incubated in 600nM MMC for 24h and then electroporated with the gene targeting vector as described in the Methods section. The values are expressed here as a ratio of gene targeting values to the EHR values for each of these cell lines. Ratios were calculated for each of the two separate experiments (3 plates per cell line, per experiment, n=6) and the means are presented here +/- the standard error of the mean. These values are statistically different from each other in a t-test (p<0.05).

3.7 Exposure to MMC

Mitomycin C is a DNA damaging agent that induces cross-links in the DNA. These crosslinks can lead to double strand breaks, which are repaired by the cell through several pathways, one of which is homologous recombination. A 6h exposure to 600 nM MMC was previously shown to be a level of damage that does not significantly decrease cell viability

(Rukść et al., 2007). It was also shown that after 24h of MMC exposure, Rad51 enters the nucleus of the cells, presumably to aid in DNA repair processes (Magwood et al., 2012).

Therefore, it was hypothesized that the elevation of levels of wild-type Rad51 might be advantageous to the cell lines in promoting EHR in the presence of a greater amount of DNA damage.

To test this, parental #9 cells and cells from the wild-type Rad51 expressor (9-800-8) were exposed to 600 nM MMC for 6 and 24 hours then tested for EHR. The data in Figure 19 presents the ratio of EHR for treated versus untreated cells. After 6h of MMC treatment, #9 and 9-800-8 are not significantly different; however, after 24h of MMC treatment, there is an increase in PFCs in the wild-type Rad51 expressor 9-800-8 compared to #9 and their respective

EHR values (p<0.05). Thus, in the presence of MMC-induced DNA damage, excess Rad51 stimulates EHR. This effect is not seen after a 6h incubation but is seen after a 24h incubation.

This potentially indicates that increasing the amount of DNA damage also increases the frequency of EHR in the presence of excess Rad51.

2.5

1.5

and 24h MMC to EHR to MMC 24h and

0.5

Ratio of 6h of Ratio 0 pRCuR1 #9 9-800-8 pRCuR1 #9 9-800-8 6h MMC (600nM) 24h MMC (600nM)

Figure 19 – Effects of 6h and 24h of MMC Treatments on EHR. The values for each cell line in this figure are ratios of the number of PFC observed after 6h and 24h MMC exposures and the EHR values for each of these cell lines. At 6h MMC treatment there is no significant difference between the two cell lines. At 24h of MMC exposure 9-800-

8 yields significantly more PFCs than #9 (p<0.05), accounting for the background EHR levels in these cell lines. Statistical significance was determined through a Student’s t- test. The ratios were calculated per experiment over 2 experiments with 3 replicate flasks per experiment (2 plates per flask, n=12), and then all ratios were averaged together for the mean values shown here. The error bars represent +/- the standard error of the mean.

3.8 Exposure to Ionizing Radiation

Ionizing radiation causes a variety of DNA lesions, the most serious of which are double- stranded breaks in the DNA (Hutchinson, 1985; Goodhead, 1994), which can be repaired via homologous recombination. The number of DNA DSBs present in a mammalian cell immediately after 1 Gy of ionizing radiation was estimated to be approximately 40 (Goodhead,

1994). Previous work in our lab showed that cells expressing higher levels of wild-type Rad51 showed decreased sensitivity to ionizing radiation (Rukść et al., 2007), leading to increased survivability after 3-5 Gy of ionizing radiation. I hypothesized that increased wild-type Rad51 could also promote EHR in these cell lines after exposure to ionizing radiation, similar to the hypothesis regarding MMC induced DNA damage in the previous section.

In order to test this, parental #9 cells and cells from the highest wild-type Rad51 expressor (9-800-8) were exposed to 4 Gy of ionizing gamma radiation. Cells were allowed to recover for 48-72h and then were assayed for EHR. The data for this experiment is presented in

Figure 20 as ratios of treated versus untreated cells. In contrast to the effects of MMC (Figure

19), there is no significant difference between the parental #9 cells and the 9-800-8 cells in

EHR frequencies after exposure to 4 Gy of ionizing radiation.

2.5

1.5

0.5 Ratio of 4gy Ratio of4gy irradiationEHR to

0 pRCuR1 #9 9-800-8

Figure 20 – Effect of 4 Gy Ionizing Radiation on EHR. The figure presents the ratios of PFCs obtained after exposure to 4 Gy of irradiation to the number of PFCs obtained in untreated cultures. There is not a significant difference between #9 and 9-800-8 in this experiment (p<0.05), determined by a Student’s t-test. The ratios were calculated for each of 2 independent experiments (3 plates per experiment, n=6) and then averaged for the mean value shown here. The error bars represent the mean +/- the standard error of the mean.

Chapter 4: Discussion

Expression of FLAG-tagged wild-type Rad51 and the Rad51 catalytic variants

(Rad51K133A and Rad51K133R) was obtained successfully in both ectopic parent cell lines #4 and

#9. A range of expression levels were observed for each of the Rad51 variants, however it was much easier to find expressors of wild-type Rad51 than of either of the Rad51 variants

Rad51K133A or Rad51K133R. This phenomenon is likely due to the reduction in ATPase functionality in these Rad51 variants (Morrison et al., 1999; Chi et al., 2006). Rad51 must be

ATP-bound to assemble into the nucleoprotein filament, and must be able to hydrolyze ATP to dissociate from the DNA to promote complementary base-pairing (Morrison et al., 1999; Chi et al., 2006). These catalytic variants are expressed in a background of endogenous Rad51, but if these mutant proteins are present they could impair normal Rad51 function because they act as dominant-negatives (Stark et al., 2002; Rukść et al., 2007; Kim et al., 2012). This is likely why it is easier to recover expressors of FLAG-tagged wild-type Rad51, as it is fully functional and would not interfere with filament formation or dissociation. SiRNA knockdowns of Rad51 were also difficult to obtain as Rad51 is an essential protein in mice, as evidenced by a knockout of Rad51 resulting in early embryonic lethality (Tsuzuki et al., 1996).

When parental cell lines #4 and #9 were assayed for EHR, the parental cell line #4 had a frequency for EHR that varied between experiments. To determine the frequency of EHR in this cell line, 15 subcultures were generated starting from 100 cells each in a classic fluctuation analysis (Luria and Delbrück, 1943). There should be no recombinant cells among the 100 initial cells; therefore, an accurate picture of the EHR frequency was gleaned from these subcultures. It was determined that the EHR frequency in the parental cell line #4 is ~26 PFC

per 107 cells plated and ~6 PFC per 107 cells plated in the parental cell line #9. These cell lines were shown to have a single vector integration site for the ectopic wild-type Cµ region donor sequence, and a vector copy number of approximately two inserted at this location (Lee and

Baker, 2007). The slight difference in EHR frequency between these cell lines is likely due to the location of the vector insertion site. Although the exact location of vector insertion is unknown it is possible that in the parental cell line #4 it is in a location that interacts more frequently with the endogenous Cµ region, perhaps due to proximity (Shulman et al., 1995).

This could explain the higher frequency of EHR in the parental cell line #4 compared to the parental cell line #9.

Cell lines expressing FLAG-tagged wild-type Rad51 in addition to normal levels of endogenous Rad51 have no change in their frequency of EHR, and in the highest expressors

(9-800-8, 4-800-4, and 4-800-2), EHR is significantly reduced compared to the parent cell line.

It was expected that EHR would be increased in cells expressing increased wild-type Rad51, since that is what was observed for gene targeting in cells with increased Rad51 (Rukść et al.,

2007). However, contrary to expectations, EHR frequency does not seem to follow the same relationship with wild-type Rad51 as gene targeting. Interestingly, more wild-type Rad51 did not increase EHR frequencies in any of the cell lines tested and in the highest expressors this excess Rad51 decreased the EHR frequency compared to the parental cell line. Consequently,

Rad51 knockdowns were tested to see if they were deficient in EHR and the results showed that all knockdowns (except knockdown #15 in the #4 cell line) were decreased for EHR compared to their respective parental cell line. This is an expected result as Rad51 is a critically important protein. It is possible that the Rad51 depletion in these cell lines acts as selection for other cellular attributes that promote survival of these cells. Since increasing or

decreasing wild-type Rad51 in these cell lines had a similar effect on EHR it was speculated that either (i) endogenous Rad51 is enough to carry out EHR and extra Rad51 is not useful in the context of EHR, or (ii) EHR requires a stricter stoichiometric ratio of Rad51 to other recombination proteins.

The effect of expressing the two catalytic Rad51 variants on EHR frequencies was examined. Cell lines expressing FLAG-tagged Rad51K133A were all significantly reduced for

EHR compared to their respective parent lines. On the other hand, only two cell lines expressing FLAG-tagged Rad51K133R showed a significant decrease in EHR frequency, the other expressors showed no change in EHR frequency. Expressors of Rad51K133A were expected to display a more severe phenotype as this variant of Rad51 is deficient in ATP- binding, which renders it unable to bind to ssDNA (Morrison et al., 1999; Chi et al., 2006). The

Rad51K133R variant is able to bind to ATP but unable to complete the hydrolysis reaction, thus enabling it to bind to ssDNA forming the nucleoprotein filament, but would lead to difficulties in dissociating from the ssDNA (Morrison et al., 1999; Chi et al., 2006). Because Rad51K133R retains the ability to hydrolyze ATP, it may be able to function in some capacity in EHR which would explain the difference in EHR frequencies between the expressors of these catalytic

Rad51 variants.

Cell lines expressing FLAG-tagged wild-type Rad51 were assayed for HR via gene targeting (described in section 2.6), since previous results in our lab showed that excess wild- type Rad51 stimulated gene targeting (Rukść et al., 2007). As expected, the expressors of

FLAG-tagged wild-type Rad51 tested showed a 2-4 fold increase in gene targeting compared to the parental cell line #9, which is consistent with the results of Rukść et al. (2007). The expression level of FLAG-tagged Rad51 in these cell lines was approximately correlated with

the level of stimulation of gene targeting, and especially so with the exclusion of the potential jackpot value in the 9-800-6 cell line (Table 3). The highest expressor, 9-800-8, showed the most dramatic stimulation of gene targeting.

These gene targeting results confirm that these cell lines expressing FLAG-tagged wild- type Rad51 are increased for HR via gene targeting, but simultaneously decreased for EHR. It was thought to be unlikely that a stricter stoichiometric ratio of Rad51 to other recombination proteins is needed in EHR versus HR as they involve many of the same steps with the possible exception of differences in the genome-spanning homology search required for EHR and gene targeting. For example, the genome search for homology during EHR involves chromosomal sequences, whereas for gene targeting, it involves a transformed plasmid and a genomic sequence, and there may be some different requirements for each process. It is possible that

EHR mimics the normal events of HR that respond to stalled replication forks that arise during normal cell division. If so, then endogenous levels of Rad51 might be sufficient to permit the regular levels of HR (and EHR) that occur. If this is the case, then inducing EHR in these cells should make use of the excess Rad51 and the frequencies should increase. This was tested in these cell lines by inducing DNA damage through treatment with MMC and ionizing radiation.

It was hypothesized that with an increase in DNA damage there would also be an increase in

EHR, and a greater increase in EHR in cell lines with more Rad51 available. Consequently, the highest expressor of wild-type Rad51 as well as the parent cell line #9 were used in these experiments. Cells were incubated in 600nM MMC for 6h and 24h and then their EHR frequencies were measured and compared to their untreated EHR values. After both lengths of

MMC incubation, EHR in the parent cell line #9 was reduced; however, in 9-800-8, the EHR value after a 6h incubation period was unchanged and after 24h it was higher than the untreated

frequency of EHR. This provides support for the hypothesis that the excess Rad51 in 9-800-8 is helpful to the cells when an excess of DNA damage is present. Therefore, even though the excess Rad51 does not directly stimulate EHR, in the presence of DNA damage from MMC it is able to stimulate this process.

The parental cell line #9 and 9-800-8 were also assayed for the frequency of EHR after exposure to 4 Gy of ionizing gamma radiation. It was found with IR that there was not a significant difference between these cell lines, which was unexpected after the result from the

MMC treatments. It was postulated from this that 4 Gy of irradiation may not induce enough

DNA damage for the effect on EHR to be visible. Unfortunately, it is not possible to use a higher dose of irradiation and still recover enough healthy cells for a plaque assay analysis.

Ionizing radiation induces many DNA lesions but approximately 40 DSBs per 1 Gy of irradiation (Goodhead, 1994). The number of cells in the 107 cells that were irradiated that should have a DSB induced in the Cµ region is ~216. This was determined by assuming ~100

DSBs per cell (a conservative estimate) and ~5.4×109 bp targets in the hybridoma genome (two times the number in a regular mouse genome). Of these targets only 1.2×103 reside in the Cµ locus, which is the only locus in which repair by EHR is detectable in these cells. This results in a 2.16×10-5 chance per cell for a DSB to land in the Cµ locus. Since 107 cells are irradiated at one time, this should mean that ~216 cells will have DSBs induced in their Cµ regions where they can be repaired by EHR and detected as PFC. Even though most of these DSBs are probably repaired through pathways other than EHR, there is still a significant number of

DSBs being introduced at this locus. Thus, it cannot be said that there might be a greater effect with more DNA damage at this locus, as this amount should be sufficient to show an increase

in EHR. This likely means that EHR is not the predominant pathway in repair of DSBs induced through IR; perhaps they are instead repaired through NHEJ.

It has been shown here that EHR does not respond well to excess wild-type Rad51, unless MMC-induced DNA damage is present. Conversely, it has also been shown that gene targeting responds well to excess wild-type Rad51 with or without DNA damage. It is therefore concluded that these processes are actually somewhat similar in that they utilize the excess

Rad51 when there is excess DNA damage present. MMC treatment leads to the formation of dsDNA breaks, which excess wild-type Rad51 can help repair through increased EHR. In the case of gene targeting, the transfected vector with broken dsDNA ends may be considered analogous to the MMC-induced DNA damage, as the cell may recognize these as dsDNA break ends which require repair. Cells utilize endogenous Rad51 to “repair” a certain amount of the transfected gene targeting vector, and when excess wild-type Rad51 is supplied to these cells a significantly greater amount of vector is able to be “repaired”. Excess wild-type Rad51 may also lead to stabilization of a gene targeting intermediate structure, leading to an increase in recombination when more wild-type Rad51 is present.

Future Directions

In order to glean more information about the role Rad51 in EHR, several other experiments can be envisaged. Since it is known that Rad51 enters the nucleus following

MMC-induced DNA damage (Magwood et al., 2012), it would be interesting to see if this is also true following IR-induced DNA damage. This experiment may be able to explain the difference in EHR stimulation in the presence of excess Rad51 between MMC-treated cells and

IR-treated cells. Another option is the treatment of cells with topoisomerase I inhibitors which

will introduce dsDNA breaks (reviewed in Tomicic and Kaina, 2012), and should also stimulate EHR in the presence of excess wild-type Rad51.

Replication inhibitors, such as aphidicolin and hydroxyurea (shown to stimulate HR by

Saintigny et al. 2001) can be tested to see whether they increase EHR in the presence of excess

Rad51. Homologous recombination has been shown to be elevated in S/G2 phases of the cell cycle in both yeast (Kadyk and Hartwell, 1992) and mammals (Johnson and Jasin, 2000) due to the availability of the sister chromatid as a template for repair during these phases. EHR might mimic HR in replication fork stalling events and might therefore be stimulated in S/G2 phases of the cell cycle, perhaps moreso in the presence of excess wild-type Rad51. It would be interesting to synchronize cultures of hybridoma cells expressing varying levels of Rad51 to determine whether this is the case.

Polymerization of Rad51 is integral to the formation of the nucleoprotein filament, and so should be important for EHR and could have implications in the homology search. The role of Rad51 polymerization in EHR can be tested with polymerization-deficient Rad51 variants

(Rad51F86E and Rad51A89E) identified by Pellegrini et al. (2002).

References Aguilera, A., and Gómez-González, B. (2008). Genome instability: a mechanistic view of its causes and consequences. Nature Reviews Genetics 9, 204–217.

Antony, E., Tomko, E. J., Xiao, Q., Krejci, L., Lohman, T. M., and Ellenberger, T. (2009). Srs2 disassembles Rad51 filaments by a protein-protein interaction triggering ATP turnover and dissociation of Rad51 from DNA. Molecular Cell 35, 105.

Bae, S.-H., Cheong, H.-K., Cheong, C., Kang, S., Hwang, D. S., and Choi, B.-S. (2003). Structure and dynamics of hemimethylated GATC sites: Implications for DNA-SeqA recognition. Journal of Biological Chemistry 278, 45987–45993.

Baker, M. D. (2004). Gene targeting at the chromosomal immunoglobulin locus. Genetic Recombination: Reviews and Protocols 262, 143.

Baker, M. D., and Read, L. (1992). Ectopic recombination within homologous immunoglobulin mu gene constant regions in a mouse hybridoma cell line. Molecular and Cellular Biology 12, 4422–4432.

Baker, M. D., Read, L. R., Beatty, B. G., and Ng, P. (1996). Requirements for ectopic homologous recombination in mammalian somatic cells. Molecular and Cellular Biology 16, 7122–7132.

Baker, M., Pennell, N., Bosnoyan, L., and Shulman, M. (1988). Homologous recombination can restore normal immunoglobulin production in a mutant hybridoma cell line. Proceedings of the National Academy of Sciences USA 85, 6432.

Baumann, B., Potash, M., and Kӧhler, G. (1985). Consequences of frameshift mutations at the immunoglobulin heavy chain locus of the mouse. EMBO Journal 4, 351.

Baumann, P., Benson, F. E., and West, S. C. (1996). Human Rad51 protein promotes ATP- dependent homologous pairing and strand transfer reactions in vitro. Cell 87, 757–766.

Bennett, S. M., Woods, D. S., Pawelczak, K. S., and Turchi, J. J. (2012). Multiple protein- protein interactions within the DNA-PK complex are mediated by the C-terminus of Ku 80. International Journal of Biochemistry and Molecular Biology 3, 36.

Benson, F. E., Stasiak, A., and West, S. C. (1994). Purification and characterization of the human Rad51 protein, an analogue of E. coli RecA. EMBO Journal 13, 5764.

Blackwell, L. J., and Borowiec, J. A. (1994). Human replication protein A binds single- stranded DNA in two distinct complexes. Molecular and Cellular Biology 14, 3993–4001.

Bugreev, D. V., and Mazin, A. V. (2004). Ca2+ activates human homologous recombination protein Rad51 by modulating its ATPase activity. Proceedings of the National Academy of

Sciences USA 101, 9988–9993.

Cabelof, D. C., Guo, Z., Raffoul, J. J., Sobol, R. W., Wilson, S. H., Richardson, A., and Heydari, A. R. (2003). Base excision repair deficiency caused by polymerase β haploinsufficiency accelerated DNA damage and increased mutational response to carcinogens. Cancer Research 63, 5799–5807.

Carreira, A., Hilario, J., Amitani, I., Baskin, R. J., Shivji, M. K., Venkitaraman, A. R., and Kowalczykowski, S. C. (2009). The BRC repeats of BRCA2 modulate the DNA binding selectivity of RAD51. Cell 136, 1032.

Cejka, P., Cannavo, E., Polaczek, P., Masuda-Sasa, T., Pokharel, S., Campbell, J. L., and Kowalczykowski, S. C. (2010). DNA end resection by Dna2-Sgs1-RPA and its stimulation by Top3-Rmi1 and Mre11-Rad50-Xrs2. Nature 467, 112–116.

Chi, P., Van Komen, S., Sehorn, M. G., Sigurdsson, S., and Sung, P. (2006). Roles of ATP binding and ATP hydrolysis in human Rad51 recombinase function. DNA Repair 5, 381–391.

Cimprich, K. A., and Cortez, D. (2008). ATR: an essential regulator of genome integrity. Nature Reviews Molecular Cell Biology 9, 616–627.

Coulter, R., Blandino, M., Tomlinson, J. M., Pauly, G. T., Krajewska, M., Moschel, R. C., Peterson, L. A., Pegg, A. E., and Spratt, T. E. (2007). Differences in the rate of repair of O 6- alkylguanines in different sequence contexts by O 6-alkylguanine-DNA alkyltransferase. Chemical Research in Toxicology 20, 1966–1971.

Cox, M. M. (2002). The nonmutagenic repair of broken replication forks via recombination. Mutation Research/Fundamental and Molecular Mechanisms of Mutagenesis 510, 107–120.

Daboussi, F., Dumay, A., Delacôte, F., and Lopez, B. S. (2002). DNA double-strand break repair signalling: the case of RAD51 post-translational regulation. Cellular Signalling 14, 969– 975.

Davies, A. A., Masson, J.-Y., McIlwraith, M. J., Stasiak, A. Z., Stasiak, A., Venkitaraman, A. R., West, S. C., and others (2001). Role of BRCA2 in control of the RAD51 recombination and DNA repair protein. Molecular Cell 7, 273.

Dayani, Y., Simchen, G., and Lichten, M. (2011). Meiotic recombination intermediates are resolved with minimal crossover formation during return-to-growth, an analogue of the mitotic cell cycle. PLoS Genetics 7, e1002083.

Draviam, V. M., Xie, S., and Sorger, P. K. (2004). Chromosome segregation and genomic stability. Current Opinion in Genetics & Development 14, 120–125.

Dudáš, A., and Chovanec, M. (2004). DNA double-strand break repair by homologous recombination. Mutation Research/Reviews in Mutation Research 566, 131–167.

Duncan, T., Trewick, S. C., Koivisto, P., Bates, P. A., Lindahl, T., and Sedgwick, B. (2002). Reversal of DNA alkylation damage by two human dioxygenases. Proceedings of the National Academy of Sciences 99, 16660–16665.

Dupré, A., Boyer-Chatenet, L., and Gautier, J. (2006). Two-step activation of ATM by DNA and the Mre11-Rad50-Nbs1 complex. Nature Structural & Molecular Biology 13, 451–457.

Durant, S. T., and Nickoloff, J. A. (2005). Good timing in the cell cycle for precise DNA repair by BRCA1. Cell Cycle 4, 1216–1222.

Eker, A., Quayle, C., Chaves, I., and van der Horst, G. (2009). DNA Repair in Mammalian Cells. Cellular and Molecular Life Sciences 66, 968–980.

Forget, A. L., Loftus, M. S., McGrew, D. A., Bennett, B. T., and Knight, K. L. (2007). The human Rad51 K133A mutant is functional for DNA double-strand break repair in human cells. Biochemistry 46, 3566–3575.

Fujimori, A., Tachiiri, S., Sonoda, E., Thompson, L. H., Dhar, P. K., Hiraoka, M., Takeda, S., Zhang, Y., Reth, M., and Takata, M. (2001). Rad52 partially substitutes for the Rad51 paralog XRCC3 in maintaining chromosomal integrity in vertebrate cells. EMBO Journal 20, 5513– 5520.

Fukui, K. (2010). DNA mismatch repair in eukaryotes and bacteria. Journal of Nucleic Acids 2010.

Goodhead, D. (1994). Initial events in the cellular effects of ionizing radiations: clustered damage in DNA. International Journal of Radiation Biology 65, 7–17.

Gorgoulis, V. G. et al. (2005). Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature 434, 907–913.

Gupta, R. C., Bazemore, L. R., Golub, E. I., and Radding, C. M. (1997). Activities of human recombination protein Rad51. Proceedings of the National Academy of Sciences USA 94, 463–468.

Halazonetis, T. D., Gorgoulis, V. G., and Bartek, J. (2008). An oncogene-induced DNA damage model for cancer development. Science 319, 1352–1355.

Hays, S. L., Firmenich, A. A., and Berg, P. (1995). Complex formation in yeast double-strand break repair: participation of Rad51, Rad52, Rad55, and Rad57 proteins. Proceedings of the National Academy of Sciences USA 92, 6925–6929.

Heller, R. C., and Marians, K. J. (2006). Replisome assembly and the direct restart of stalled replication forks. Nature Reviews Molecular Cell Biology 7, 932–943.

Hewish, M., Lord, C. J., Martin, S. A., Cunningham, D., and Ashworth, A. (2010). Mismatch repair deficient colorectal cancer in the era of personalized treatment. Nature Reviews Clinical Oncology 7, 197–208.

Heyer, W. D., Ehmsen, K. T., and Liu, J. (2010). Regulation of homologous recombination in eukaryotes. Annual Review of Genetics 44, 113–139.

Heyer, W.-D., Li, X., Rolfsmeier, M., and Zhang, X.-P. (2006). Rad54: the Swiss Army knife of homologous recombination? Nucleic Acids Research 34, 4115–4125.

Hinz, J. M. (2010). Role of homologous recombination in DNA interstrand crosslink repair. Environmental and Molecular Mutagenesis 51, 582–603.

Holthausen, J. T., Wyman, C., and Kanaar, R. (2010). Regulation of DNA strand exchange in homologous recombination. DNA Repair 9, 1264–1272.

Hutchinson, F. (1985). Chemical changes induced in DNA by ionizing radiation. Progress in Nucleic Acid Research and Molecular Biology 32, 115–154.

De Ioannes, P., Malu, S., Cortes, P., and Aggarwal, A. K. (2012). Structural Basis of DNA Ligase IV-Artemis Interaction in Nonhomologous End-Joining. Cell Reports.

Ivanov, E. L., Sugawara, N., Fishman-Lobell, J., and Haber, J. E. (1996). Genetic requirements for the single-strand annealing pathway of double-strand break repair in Saccharomyces cerevisiae. Genetics 142, 693–704.

Jackson, S. P., and Bartek, J. (2009). The DNA-damage response in human biology and disease. Nature 461, 1071–1078.

Jensen, R. B., Carreira, A., and Kowalczykowski, S. C. (2010). Purified human BRCA2 stimulates RAD51-mediated recombination. Nature 467, 678–683.

Johnson, R. D., and Jasin, M. (2000). Sister chromatid gene conversion is a prominent double- strand break repair pathway in mammalian cells. EMBO Journal 19, 3398–3407.

Johnson, R., Jasin, M., and others (2001). Double-strand-break-induced homologous recombination in mammalian cells. Biochemical Society Transactions 29, 196–200.

Kadyk, L. C., and Hartwell, L. H. (1992). Sister chromatids are preferred over homologs as substrates for recombinational repair in Saccharomyces cerevisiae. Genetics 132, 387–402.

Kanaar, R., Hoeijmakers, J. H., and van Gent, D. C. (1998). Molecular mechanisms of DNA double-strand break repair. Trends in Cell Biology 8, 483–489.

Kao, Y.-T., Saxena, C., Wang, L., Sancar, A., and Zhong, D. (2005). Direct observation of thymine dimer repair in DNA by photolyase. Proceedings of the National Academy of Sciences USA 102, 16128–16132.

Kiianitsa, K., Solinger, J. A., and Heyer, W.-D. (2002). Rad54 protein exerts diverse modes of ATPase activity on duplex DNA partially and fully covered with Rad51 protein. Journal of Biological Chemistry 277, 46205–46215.

Kim, T. M., Ko, J. H., Hu, L., Kim, S.-A., Bishop, A. J., Vijg, J., Montagna, C., and Hasty, P. (2012). RAD51 Mutants Cause Replication Defects and Chromosomal Instability. Molecular and Cellular Biology 32, 3663–3680.

Kleckner, N. (1996). Meiosis: how could it work? Proceedings of the National Academy of Sciences USA 93, 8167–8174.

Kousholt, A. N., Fugger, K., Hoffmann, S., Larsen, B. D., Menzel, T., Sartori, A. A., and Sørensen, C. S. (2012). CtIP-dependent DNA resection is required for DNA damage checkpoint maintenance but not initiation. Journal of Cell Biology 197, 869–876.

Kӧhler, G., Potash, M., Lehrach, H., and Shulman, M. (1982). Deletions in immunoglobulin mu chains. EMBO Journal 1, 555.

Kӧhler, G., and Shulman, M. J. (1980). Immunoglobulin M mutants. European Journal of Immunology 10, 467–476.

Lamarche, B. J., Orazio, N. I., and Weitzman, M. D. (2010). The MRN complex in double- strand break repair and telomere maintenance. FEBS Letters 584, 3682–3695.

Lee, S. A., and Baker, M. D. (2007). Analysis of DNA repair and recombination responses in mouse cells depleted for Brca2 by SiRNA. DNA Repair 6, 809–817.

Lefevre, J. H., Rodrigue, C. M., Mourra, N., Bennis, M., Flejou, J.-F., Parc, R., Tiret, E., Gespach, C., and Parc, Y. R. (2006). Implication of MYH in colorectal polyposis. Annals of Surgery 244, 874.

Lieber, M. R., Ma, Y., Pannicke, U., and Schwarz, K. (2004). The mechanism of vertebrate nonhomologous DNA end joining and its role in V (D) J recombination. DNA Repair 3, 817– 826.

Lim, D.-S., and Hasty, P. (1996). A mutation in mouse rad51 results in an early embryonic lethal that is suppressed by a mutation in p53. Molecular and Cellular Biology 16, 7133–7143.

Lisby, M., Barlow, J. H., Burgess, R. C., and Rothstein, R. (2004). Choreography of the DNA damage response: spatiotemporal relationships among checkpoint and repair proteins. Cell 118, 699–713.

Liu, J., Doty, T., Gibson, B., and Heyer, W.-D. (2010). Human BRCA2 protein promotes RAD51 filament formation on RPA-covered single-stranded DNA. Nature Structural \& Molecular Biology 17, 1260–1262.

Liu, S., Opiyo, S. O., Manthey, K., Glanzer, J. G., Ashley, A. K., Amerin, C., Troksa, K., Shrivastav, M., Nickoloff, J. A., and Oakley, G. G. (2012). Distinct roles for DNA-PK, ATM and ATR in RPA phosphorylation and checkpoint activation in response to replication stress. Nucleic Acids Research 40, 10780–10794.

Lundin, C., Erixon, K., Arnaudeau, C., Schultz, N., Jenssen, D., Meuth, M., and Helleday, T. (2002). Different roles for nonhomologous end joining and homologous recombination following replication arrest in mammalian cells. Molecular and Cellular Biology 22, 5869– 5878.

Luo, G., Yao, M. S., Bender, C. F., Mills, M., Bladl, A. R., Bradley, A., and Petrini, J. H. (1999). Disruption of mRad50 causes embryonic stem cell lethality, abnormal embryonic development, and sensitivity to ionizing radiation. Proceedings of the National Academy of Sciences USA 96, 7376–7381.

Luria, S. E., and Delbrück, M. (1943). Mutations of bacteria from virus sensitivity to virus resistance. Genetics 28, 491.

Lydeard, J. R., Lipkin-Moore, Z., Sheu, Y.-J., Stillman, B., Burgers, P. M., and Haber, J. E. (2010). Break-induced replication requires all essential DNA replication factors except those specific for pre-RC assembly. Genes & Development 24, 1133–1144.

Magwood, A. C., Mundia, M. M., and Baker, M. D. (2012). High levels of wild-type BRCA2 suppress homologous recombination. Journal of Molecular Biology 421, 38–53.

Mari, P.-O. et al. (2006). Dynamic assembly of end-joining complexes requires interaction between Ku70/80 and XRCC4. Proceedings of the National Academy of Sciences USA 103, 18597–18602.

Martin, L. M., Marples, B., Coffey, M., Lawler, M., Lynch, T. H., Hollywood, D., and Marignol, L. (2010). DNA mismatch repair and the DNA damage response to ionizing radiation: making sense of apparently conflicting data. Cancer Treatment Reviews 36, 518– 527.

Matsuoka, S. et al. (2007). ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science Signaling 316, 1160–1166.

Matulova, P., Marini, V., Burgess, R. C., Sisakova, A., Kwon, Y., Rothstein, R., Sung, P., and Krejci, L. (2009). Cooperativity of Mus81-Mms4 with Rad54 in the resolution of recombination and replication intermediates. Journal of Biological Chemistry 284, 7733–7745.

Mazin, A. V., Alexeev, A. A., and Kowalczykowski, S. C. (2003). A novel function of Rad54 Protein. Journal of Biological Chemistry 278, 14029–14036.

Mazin, A. V., Mazina, O. M., Bugreev, D. V., and Rossi, M. J. (2010). Rad54, the motor of homologous recombination. DNA Repair 9, 286–302.

McIlwraith, M. J., and West, S. C. (2008). DNA repair synthesis facilitates RAD52-mediated second-end capture during DSB repair. Molecular Cell 29, 510–516.

Modesti, M., Ristic, D., van der Heijden, T., Dekker, C., van Mameren, J., Peterman, E. J., Wuite, G. J., Kanaar, R., and Wyman, C. (2007). Fluorescent human RAD51 reveals multiple nucleation sites and filament segments tightly associated along a single DNA molecule. Structure 15, 599–609.

Morrison, C., Shinohara, A., Sonoda, E., Yamaguchi-Iwai, Y., Takata, M., Weichselbaum, R. R., and Takeda, S. (1999). The essential functions of human Rad51 are independent of ATP hydrolysis. Molecular and Cellular Biology 19, 6891–6897.

Mott, C., and Symington, L. S. (2011). RAD51-independent inverted-repeat recombination by a strand-annealing mechanism. DNA Repair 10, 408–415.

Moynahan, M. E., and Jasin, M. (2010). Mitotic homologous recombination maintains genomic stability and suppresses tumorigenesis. Nature Reviews Molecular Cell Biology 11, 196–207.

Pardo, B., Gómez-González, B., and Aguilera, A. (2009). DNA repair in mammalian cells. Cellular and Molecular Life Sciences 66, 1039–1056.

Pellegrini, L., David, S. Y., Lo, T., Anand, S., Lee, M., Blundell, T. L., and Venkitaraman, A. R. (2002). Insights into DNA recombination from the structure of a RAD51-BRCA2 complex. Nature 420, 287–293.

Petermann, E., and Helleday, T. (2010). Pathways of mammalian replication fork restart. Nature Reviews Molecular Cell Biology 11, 683–687.

Prakash, A., and Borgstahl, G. E. (2012). The structure and function of replication protein A in DNA replication. In: The Eukaryotic Replisome: a Guide to Protein Structure and Function, Springer, 171–196.

Prasad, R., Beard, W., Batra, V., Liu, Y., Shock, D., and Wilson, S. (2011). A review of recent experiments on step-to-step “hand-off” of the DNA intermediates in mammalian base excision

repair pathways. Molecular Biology 45, 536–550.

Preston, B. D., Albertson, T. M., and Herr, A. J. (2010). DNA replication fidelity and cancer. In: Seminars in Cancer Biology, 281–293.

Raynard, S., and Sung, P. (2009). Assay for human Rad51-mediated DNA displacement loop formation. Cold Spring Harbor Protocols 4, 1–4.

Rechkunova, N., Krasikova, Y. S., and Lavrik, O. (2011). Nucleotide excision repair: DNA damage recognition and preincision complex assembly. Biochemistry (Moscow) 76, 24–35.

Rijkers, T., Van Den Ouweland, J., Morolli, B., Rolink, A. G., Baarends, W. M., Van Sloun, P. P., Lohman, P. H., and Pastink, A. (1998). Targeted inactivation of mouse RAD52reduces homologous recombination but not resistance to ionizing radiation. Molecular and Cellular Biology 18, 6423–6429.

Rosu, S., Libuda, D. E., and Villeneuve, A. M. (2011). Robust crossover assurance and regulated interhomolog access maintain meiotic crossover number. Science 334, 1286–1289.

Roy, R., Chun, J., and Powell, S. N. (2012). BRCA1 and BRCA2: important differences with common interests. Nature Reviews Cancer 12, 372–372.

Rukść, A., Birmingham, E. C., and Baker, M. D. (2007). Altered DNA repair and recombination responses in mouse cells expressing wildtype or mutant forms of RAD51. DNA Repair 6, 1876–1889.

Rupnik, A., Lowndes, N. F., and Grenon, M. (2010). MRN and the race to the break. Chromosoma 119, 115–135.

Saintigny, Y., Delacôte, F., Varès, G., Petitot, F., Lambert, S., Averbeck, D., and Lopez, B. S. (2001). Characterization of homologous recombination induced by replication inhibition in mammalian cells. EMBO Journal 20, 3861–3870.

Saleh-Gohari, N., Bryant, H. E., Schultz, N., Parker, K. M., Cassel, T. N., and Helleday, T. (2005). Spontaneous homologous recombination is induced by collapsed replication forks that are caused by endogenous DNA single-strand breaks. Molecular and Cellular Biology 25, 7158–7169.

San Filippo, J., Sung, P., and Klein, H. (2008). Mechanism of eukaryotic homologous recombination. Annu. Rev. Biochem. 77, 229–257.

Schildkraut, E., Miller, C. A., and Nickoloff, J. A. (2005). Gene conversion and deletion frequencies during double-strand break repair in human cells are controlled by the distance between direct repeats. Nucleic Acids Research 33, 1574–1580.

Selby, C. P., Drapkin, R., Reinberg, D., and Sancar, A. (1997). RNA polymerase II stalled at a thymine dimer: footprint and effect on excision repair. Nucleic Acids Research 25, 787–793.

Setlow, R., Regan, J. D., German, J., and Carrier, W. (1969). Evidence that xeroderma pigmentosum cells do not perform the first step in the repair of ultraviolet damage to their DNA. Proceedings of the National Academy of Sciences USA 64, 1035–1041.

Shinohara, A., and Ogawa, T. (1998). Stimulation by Rad52 of yeast Rad51-mediated recombination. Nature 391, 404–407.

Shirleen Roeder, G. (1990). Chromsome synapsis and genetic recombination: their roles in meiotic chromsome segregation. Trends in Genetics 6, 385–389.

Shulman, M., Collins, C., Connor, A., Read, L., and Baker, M. (1995). Interchromosomal recombination is suppressed in mammalian somatic cells. EMBO Journal 14, 4102.

Singleton, M. R., Wentzell, L. M., Liu, Y., West, S. C., and Wigley, D. B. (2002). Structure of the single-strand annealing domain of human RAD52 protein. Proceedings of the National Academy of Sciences USA 99, 13492–13497.

Smith, C. E., Llorente, B., and Symington, L. S. (2007). Template switching during break- induced replication. Nature 447, 102–105.

Sonoda, E., Hochegger, H., Saberi, A., Taniguchi, Y., Takeda, S., and others (2006). Differential usage of non-homologous end-joining and homologous recombination in double strand break repair. DNA Repair 5, 1021–1029.

Sonoda, E., Sasaki, M. S., Buerstedde, J.-M., Bezzubova, O., Shinohara, A., Ogawa, H., Takata, M., Yamaguchi-Iwai, Y., and Takeda, S. (1998). Rad51-deficient vertebrate cells accumulate chromosomal breaks prior to cell death. EMBO Journal 17, 598–608.

Southern, P., Howard, B., Berg, P., and others (1981). Construction and characterization of SV40 recombinants with beta-globin cDNA substitutions in their early regions. Journal of Molecular and Applied Genetics 1, 177.

Starita, L. M., and Parvin, J. D. (2006). Substrates of the BRCA1-dependent ubiquitin ligase. Cancer Biology & Therapy 5, 137–141.

Stark, J. M., Hu, P., Pierce, A. J., Moynahan, M. E., Ellis, N., and Jasin, M. (2002). ATP hydrolysis by mammalian RAD51 has a key role during homology-directed DNA repair. Journal of Biological Chemistry 277, 20185–20194.

Stavnezer, J., Guikema, J. E., and Schrader, C. E. (2008). Mechanism and regulation of class switch recombination. Annual Review of Immunology 26, 261.

Streisinger, G., and Bruce, V. (1960). Linkage of genetic markers in phages T2 and T4. Genetics 45, 1289.

Sugiyama, T., and Kowalczykowski, S. C. (2002). Rad52 protein associates with replication protein A (RPA)-single-stranded DNA to accelerate Rad51-mediated displacement of RPA and presynaptic complex formation. Journal of Biological Chemistry 277, 31663–31672.

Sung, P. (1994). Catalysis of ATP-dependent homologous DNA pairing and strand exchange by yeast RAD51 protein. Science 265, 1241–1243.

Sung, P. (1997). Yeast Rad55 and Rad57 proteins form a heterodimer that functions with replication protein A to promote DNA strand exchange by Rad51 recombinase. Genes & Development 11, 1111–1121.

Sung, P., Krejci, L., Van Komen, S., and Sehorn, M. G. (2003). Rad51 recombinase and recombination mediators. Journal of Biological Chemistry 278, 42729–42732.

Sung, P., and Robberson, D. L. (1995). DNA strand exchange mediated by a RAD51-ssDNA nucleoprotein filament with polarity opposite to that of RecA. Cell 82, 453–461.

Sung, P., and Stratton, S. A. (1996). Yeast Rad51 recombinase mediates polar DNA strand exchange in the absence of ATP hydrolysis. Journal of Biological Chemistry 271, 27983– 27986.

Suwaki, N., Klare, K., and Tarsounas, M. (2011). RAD51 paralogs: roles in DNA damage signalling, recombinational repair and tumorigenesis. In: Seminars in Cell & Developmental Biology, 898–905.

Symington, L. S. (2002). Role of RAD52 epistasis group genes in homologous recombination and double-strand break repair. Microbiology and Molecular Biology Reviews 66, 630–670.

Takata, M., Sasaki, M. S., Tachiiri, S., Fukushima, T., Sonoda, E., Schild, D., Thompson, L. H., and Takeda, S. (2001). Chromosome instability and defective recombinational repair in knockout mutants of the five Rad51 paralogs. Molecular and Cellular Biology 21, 2858–2866.

Tarsounas, M., Davies, D., and West, S. C. (2003). BRCA2-dependent and independent formation of RAD51 nuclear foci. Oncogene 22, 1115–1123.

Thorslund, T., McIlwraith, M. J., Compton, S. A., Lekomtsev, S., Petronczki, M., Griffith, J. D., and West, S. C. (2010). The breast cancer tumor suppressor BRCA2 promotes the specific targeting of RAD51 to single-stranded DNA. Nature Structural & Molecular Biology 17, 1263–1265.

Tomicic, M. T., and Kaina, B. (2012). Topoisomerase degradation, DSB repair, p53 and IAPs in cancer cell resistance to camptothecin-like topoisomerase I inhibitors. Biochimica Et

Biophysica Acta (BBA)-Reviews on Cancer.

Tomimatsu, N., Mukherjee, B., Deland, K., Kurimasa, A., Bolderson, E., Khanna, K. K., and Burma, S. (2012). Exo1 plays a major role in DNA end resection in humans and influences double-strand break repair and damage signaling decisions. DNA Repair 11, 441–448.

Tsuzuki, T., Fujii, Y., Sakumi, K., Tominaga, Y., Nakao, K., Sekiguchi, M., Matsushiro, A., Yoshimura, Y., and others (1996). Targeted disruption of the Rad51 gene leads to lethality in embryonic mice. Proceedings of the National Academy of Sciences USA 93, 6236–6240.

Uematsu, N., Weterings, E., Yano, K., Morotomi-Yano, K., Jakob, B., Taucher-Scholz, G., Mari, P.-O., van Gent, D. C., Chen, B. P., and Chen, D. J. (2007). Autophosphorylation of DNA-PKCS regulates its dynamics at DNA double-strand breaks. The Journal of Cell Biology 177, 219–229. de Vries, F. A. et al. (2005). Inactivation of RAD52 aggravates RAD54 defects in mice but not in Schizosaccharomyces pombe. DNA Repair 4, 1121–1128.

West, S. C. (2003). Molecular views of recombination proteins and their control. Nature Reviews Molecular Cell Biology 4, 435–445.

Wilson, T. E., and Lieber, M. R. (1999). Efficient processing of DNA ends during yeast nonhomologous end joining: Evidence for a DNA polymerase β (POL4)-dependent pathway. Journal of Biological Chemistry 274, 23599–23609.

Wold, M. S. (1997). Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annual Review of Biochemistry 66, 61–92.

Wu, L., Bachrati, C. Z., Ou, J., Xu, C., Yin, J., Chang, M., Wang, W., Li, L., Brown, G. W., and Hickson, I. D. (2006). BLAP75/RMI1 promotes the BLM-dependent dissolution of homologous recombination intermediates. Proceedings of the National Academy of Sciences USA 103, 4068–4073.

Wu, P.-Y., Frit, P., Malivert, L., Revy, P., Biard, D., Salles, B., and Calsou, P. (2007). Interplay between Cernunnos-XLF and nonhomologous end-joining proteins at DNA ends in the cell. Journal of Biological Chemistry 282, 31937–31943.

Wyka, I. M., Dhar, K., Binz, S. K., and Wold, M. S. (2003). Replication protein A interactions with DNA: differential binding of the core domains and analysis of the DNA interaction surface. Biochemistry 42, 12909–12918.

Wyman, C., and Kanaar, R. (2006). DNA double-strand break repair: all’s well that ends well. Annual Reviews Genetics 40, 363–383.

Yang, H., Jeffrey, P. D., Miller, J., Kinnucan, E., Sun, Y., Thomä, N. H., Zheng, N., Chen, P.- L., Lee, W.-H., and Pavletich, N. P. (2002). BRCA2 function in DNA binding and

recombination from a BRCA2-DSS1-ssDNA structure. Science 297, 1837–1848.

Yang, W. (2007). Structure and mechanism for DNA lesion recognition. Cell Research 18, 184–197.

Yang, W. (2011). Surviving the sun: Repair and bypass of DNA UV lesions. Protein Science 20, 1781–1789.

Yeeles, J. T., and Dillingham, M. S. (2010). The processing of double-stranded DNA breaks for recombinational repair by helicase-nuclease complexes. DNA Repair 9, 276–285.

Yoshioka, K., Yoshioka, Y., and Hsieh, P. (2006). ATR kinase activation mediated by MutSα and MutLα in response to cytotoxic O6-methylguanine adducts. Molecular Cell 22, 501.

Yu, X., Jacobs, S. A., West, S. C., Ogawa, T., and Egelman, E. H. (2001). Domain structure and dynamics in the helical filaments formed by RecA and Rad51 on DNA. Proceedings of the National Academy of Sciences USA 98, 8419–8424.

Zhang, F., Ma, J., Wu, J., Ye, L., Cai, H., Xia, B., and Yu, X. (2009). PALB2 links BRCA1 and BRCA2 in the DNA-damage response. Current Biology: CB 19, 524.

Zhang, Y., Zhou, J., and Lim, C. U. (2006). The role of NBS1 in DNA double strand break repair, telomere stability, and cell cycle checkpoint control. Cell Research 16, 45–54.

Zou, L., and Elledge, S. J. (2003). Sensing DNA damage through ATRIP recognition of RPA- ssDNA complexes. Science Signaling 300, 1542.