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Theses & Dissertations Graduate Studies

Fall 12-20-2019

Characterizing the Combination of RPA Inhibitors with PARP Characterizing the Combination of RPA Inhibitors with PARP

Inhibitors in High-Grade Serous Ovarian Cancer Inhibitors in High-Grade Serous Ovarian Cancer

Yat Tang University of Nebraska Medical Center

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CHARACTERIZING THE COMBINATION OF RPA INHIBITORS WITH PARP

INHIBITORS IN HIGH-GRADE SEROUS OVARIAN CANCER

by

Yat T. Tang

A THESIS

Presented to the Faculty of

the University of Nebraska Graduate College

in Partial Fulfillment of the Requirements

for the Degree of Master of Science

Medical Sciences Interdepartmental Area

Oral Biology

Under the Supervision of Professor Gregory G. Oakley

University of Nebraska Medical Center

Omaha, Nebraska

December 2019

Advisory Committee:

Peter J. Giannini, D.D.S., M.S.

Sundaralingam Premaraj, B.D.S., M.S., Ph.D., FRCD(C)

ii

ACKNOWLEDGEMENTS

I would like to thank my adviser, Dr. Gregory Oakley, for his knowledge,

guidance, and willingness to help me throughout this project. Participating in this project

has reminded me of the unforeseen challenges to overcome in the laboratory setting,

and the joy of obtaining meaningful results.

Thank you to my other committee members, Dr. Peter Giannini and Dr. Prem

Premaraj. Dr. Giannini has made himself available for meetings and was always willing

to offer a helping hand. Dr. Premaraj for asking questions to remind me of the

fundamental of experimental techniques.

Finally, I would like to thank all those that have supported my project by offering

technical advice and support. Much gratitude to Brendan Byrne for his assistance with

growing cells, drafting figures, and being accommodating with his time.

iii

CHARACTERIZING THE COMBINATION OF RPA INHIBITORS WITH PARP

INHIBITORS IN HIGH-GRADE SEROUS OVARIAN CANCER

Yat T. Tang, D.D.S., Ph.D., M.S.

University of Nebraska, 2019

Advisor: Gregory G. Oakley, Ph.D.

High-grade serous ovarian cancer (HGSC) is the most common and deadly

gynecologic malignancy. HGSC patients with BRCA1/2 mutations have homologous

recombination deficiency (HRD), requiring parallel pathways to maintain genome

integrity (e.g., PARP1, PARP2). Approximately 50% of ovarian carcinomas are

estimated to exhibit HRD. For the remaining 50% and the large percentage of HRD

patients with acquired or innate resistance to single-agent PARP inhibitors, there is a

need to develop alternative therapeutic strategies.

Replication Protein A (RPA) is a heterotrimeric protein crucial for genome

maintenance. Phosphorylation of RPA in DNA damage response (DDR) is a negative

regulator of DNA end resection. RPA interacts with multiple proteins at the N-terminus

of RPA1 (DBD-F) to function. A novel strategy is to specifically inhibit DBD-F with an

inhibitor that blocks RPA protein-protein interactions, and combined with PARP

inhibition, has the potential to increase replication stress and DNA damage while

selectively inducing cell death in HGSC cells containing both wild-type and mutant

BRCA1/2.

This thesis characterized PARP inhibitors with RPA inhibitors in HGSC cells.

Examination includes the analysis ssDNA binding, dsDNA unwinding, cell viability, and

detection of phosphorylation in biochemical markers via immunofluorescence (IF)

studies. An enhanced effect was observed with RPA and PARP inhibition in cell viability

assays, and markers of replication stress and DNA damage was observed in IF studies.

Future studies will include in vivo characterization using xenografts in mouse models.

iv

TABLE OF CONTENTS

ACKNOWLEDGEMENTS ....……………………………………………………………...…… ii

ABSTRACT …………………………………………………………………………………...… iii

TABLE OF CONTENTS ..……………………………………………………………………… iv

LIST OF FIGURES .……………….…………………………………………………………… vi

LIST OF ABBREVIATIONS ..………………………………………………………………… viii

CHAPTER 1: INTRODUCTION ..………………………………………………………………1

High-Grade Serous Ovarian Cancer ….…………………………………………………1

Poly (ADP-ribose) Polymerase (PARP) …..…………………………………………… 2

PARP Inhibitors ...………………………………………………………………………… 4

Replication Protein A (RPA) Structure, Phosphorylation, and Function ....………… 6

Role of RPA in Homologous Recombination and DNA Resection .....……………… 9

RPA DBD-F as a novel small-molecule therapeutic target ....……………………… 10

Hypothesis and Specific Aims ....……………………………………………………… 13

CHAPTER 2: MATERIAL AND METHODS ………………………………………………… 14

Compounds and Antibodies ...…………………………………………………..…….. 14

Electrophoretic Mobility Shift Assay (EMSA) ..………………………………………. 14

Cell Viability Assay...……………………………………………………………………. 15

Cell Lines ......……………………………………………………………………………. 15

Immunofluorescence Detection ...…………………………………………..………… 16

CHAPTER 3: RESULTS ……………………………………………………………………… 17

PAME does not inhibit RPA ssDNA binding .………………………………………… 17

PAME inhibits the ability of RPA to unwind dsDNA ………………………………… 17

Effects of RPA and PARP inhibitors on cell viability ...……………………………… 18

Immunofluorescence detection of RPA S4/S8 and Histone H2AX Phosphorylation

by RPAi and PARPi ..…………………………………………………………………… 19

v

FTE282 (Control) evaluated with RPAi and PARPi .………………………………… 20

OVCAR5 (BRCA1-) examined with RPAi and PARPi .……………………………… 21

OVCAR8 (BRCA1-) examined with RPAi and PARPi .……………………………… 22

OVCAR4 (BRCA1/2+) examined with RPAi and PARPi .…………………………… 23

Immunofluorescence detection of pH2AX after a 30-hour recovery period .……… 24

CHAPTER 4: DISCUSSION .………………………………………………………………… 27

RPAi effects on ssDNA Binding and dsDNA unwinding ..…………………………… 27

RPAi and PARPi effects on cell viability ..……..……………………………………… 27

RPAi and PARPi effects on RPA S4/S8 and H2AX phosphorylation ..…………… 27

RPAi and PARPi effects on pH2AX after a 30-hour recovery period ....…………… 28

CHAPTER 5: CONCLUSIONS ………………………………………………………………. 29

LITERATURE CITED .………………………………………………………………………… 30

vi

LIST OF FIGURES

1. Mechanism of PARP Action ..…………………………………..…………………………. 3

2. Chemical structures of six small-molecule PARP inhibitors available in the clinic or in

clinical trials .…………………………………………………..……………………………. 4

3. Dual cytotoxic mechanisms of PARP inhibitors ....………………………..…………….. 5

4. Cartoon structure of the RPA heterotrimer ....………………………………..………….. 6

5. Schematic diagram of the RPA heterotrimer subunits with domain functions and

consensus PIKK/CDK site, DNA-binding domains (DBD), and phosphorylation targets

………..………………………………………………………………………………………. 8

6. Schematic diagram of Ser4/Ser8 RPA2 hyperphosphorylation in the G2 phase of the

cell cycle following DNA Damage ...…………………………………………..………….. 8

7. Schematic of RPA showing key phosphorylation sites on RPA32 .………………….. 10

8. Schematic illustrating during DNA damage response, phosphorylated RPA is a

negative regulator of BLM helicase and DNA resection ....………………..………….. 10

9. Chemical structures of PARP and RPA inhibitors examined ...………………………. 13

10. PAME was evaluated for its RPA inhibitory property of ssDNA binding ....………….. 17

11. PAME was examined for its ability to inhibit dsDNA unwinding ..…………………….. 17

12. Effects of RPAi/PARPi in HGSC OVCAR cells examined by cell viability assay …..…19

13. Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no

drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in FTE282

cells .……………………………………………………………………………………….. 21

14. Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no

drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR5

(BRCA1-) cells ...………………………………………………………………………….. 22

vii

15. Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no

drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR8

(BRCA1-) cells ..………………………………………………………………………….. 23

16. Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no

drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR4

(BRCA1/2+) cells .……………………………………………………………………….. 24

17. Immunofluorescence detection of pH2AX with no drug treatment (DMSO), RPAi

(PAME 25µM), PARPi (75nM TZ), and both drugs (75nM TZ+25µM PAME) after 30h

recovery …….…………………………………………………………………………….. 26

viii

LIST OF ABBREVIATIONS

ATM ataxia-telangiectasia mutated kinase

ATR ATM and Rad3-related kinase

BRCA breast cancer gene

BLM Bloom Syndrome protein

CDK cyclin-dependent kinase

DBD DNA binding domains

DDR DNA damage response

DNA deoxyribonucleic acid

dsDNA double-stranded DNA

ssDNA single-stranded DNA

DNA-PK DNA-dependent protein kinase

DRS DNA replication stress

EMSA electrophorectic mobility shift assay

FDA Food and Drug Administration

FTE fallopian tube epithelium

H2AX histone H2A variant

HGSC high-grade serous ovarian cancer

HR homologous recombination

HRD homologous recombination deficiency

OVCAR high-grade ovarian adenocarcinoma epithelial

PARP poly (ADP-ribose) polymerase

PARPi PARP inhibition

PAME (name of RPA inhibitor)

PIKK phosphoinositide 3-kinase related kinase

RPA replication protein A

ix

RPA S4/S8 RPA serine 4 and serine 8

RPAi RPA inhibition

TZ talazoparib (PARP inhibitor)

WH winged helix-turn-helix

1

CHAPTER 1: INTRODUCTION

High-Grade Serous Ovarian Cancer

High-grade serous ovarian cancer (HGSC) is the most common and deadly

gynecologic malignancy. The World Health Organization (WHO) estimated 225,500

cases of ovarian cancer will be diagnosed and 140,200 patients will succumb to this

disease, representing the seventh most common form of cancer and the eighth leading

cause of cancer-related death among women worldwide (1,2). In the western nations,

ovarian cancer is the fifth most frequent cause of cancer-related death in women (3).

The current 5-year overall survival rate in the US is approximately 47.6% (4). Majority of

HGSC cases have been found to originate in the fallopian tube and exhibiting mutations

in the BRCA1/2 genes (5). Due to their origin in the fallopian tube, HGSC spreads early

in the course of disease, and by the time they become symptomatic, they are usually

high stage tumors, resulting in poor outcomes.

HGSC patients with BRCA1/2 mutations become deficient in homologous

recombination (HR). HR is a conserved DNA double-strand break (DSB) repair pathway

that uses information stored in a sister chromatid to repair damaged genome.

Homologous recombination deficiency (HRD) allows parallel pathways to maintain

genome integrity, in particular pathways involving poly (ADP-ribose) polymerases

(PARP), PARP1 and PARP2, thus are sensitive to PARP inhibition. This has led to a

rapid clinical development of PARP1/2 inhibitors. HRD also occurs due to other

mechanisms, such as epigenetic modifications and mutations of other genes involved in

the HR pathway. Ovarian cancers with these alterations behave similarly to those with

BRCA mutations, and this behavior is termed the‘‘BRCAness’’ phenotype (6,7).

Approximately 41-50% of ovarian carcinomas are estimated to exhibit HRD (8).

Unfortunately, for the remaining 50-59% and the HRD patients with acquired or innate

2

resistance to single-agent PARP inhibitors which is frequently observed, there is a

pressing need to develop additional therapeutic strategies.

Poly (ADP-ribose) Polymerase (PARP)

Poly (ADP-ribose) polymerase (PARP) is a family of nuclear enzymes that plays

critical roles in signaling the presence of DNA damage and facilitating DNA repair.

PARP catalyzes the addition of ADP-ribose units to DNA, histones, and various DNA

repair enzymes, which affects cellular processes such as replication, transcription, gene

regulation, and protein degradation. Deletion of PARP1 in mouse models weaken DNA

repair but is not lethal (9). The residual PARP activity, estimated to be 10%, is due to

PARP2. Mice with double knockout of PARP1 and PARP2 die during embryogenesis,

suggesting that PARP2 plays a critical role in absence of PARP1 (10) and that only

PARP1 and PARP2 need to be inhibited to inhibit DNA repair (11,12).

PARP1 has a zinc-finger DNA binding domain which binds to ssDNA breaks (SSB),

cleaves NAD+ and attaches multiple ADP-ribose units to the target protein as a form of

post-translational modification (11-13). This leads to a negatively charged target, and

the subsequent unwinding and repair of the damaged DNA through the base excision

repair pathway. PARP1 can also bind to dsDNA breaks (DSB). PARP1 activates

several proteins involved in homologous recombination repair (14,15), and is believed to

prevent accidental recombination of homologous DNA. PARP1 has been implicated in

BRCA1- and BRCA2-dependent homologous recombination repair (16-18). When

PARP1 is inhibited, SSBs persist and result in stalled replication forks and DSBs. In

BRCA1- and BRCA2-deficient cells, these lesions are not repaired through homologous

recombination repair, leading to cell cycle arrest and apoptosis. Less is known about the

role of PARP-2 in DNA repair. It is believed that both PARP1 and PARP2 participate in

overlapping DNA damage signaling processes and may partially compensate for one

another (19).

3

The primary activity of PARP1/2 is the poly-ADP ribosylation (PARylation) of key

components of chromatin and the DNA damage response (DDR) (Figure 1). DNA-

damaging agents activate PARP, resulting in poly(ADP-ribose)-branched chains

attached to DNA, recruiting associated repair proteins and cell cycle checkpoint

mediators. This cascade may lead to cell cycle arrest while the cell commits to either

DNA repair or apoptosis. Overactivation of PARP will lead to NAD+ depletion and

necrotic cell death. PARP inhibition is thought to impair DNA repair function, leading to

cellular dysfunction and death, and may also affect other PARP mediated DNA

modulating effects. PARP1 functions to open up chromatin and facilitate recruitment of

downstream DNA repair factors to damaged sites (20). After completing this recruitment

role, PARP auto-PARylation, the negative charge that PAR chains impart upon PAR1,

triggers the release of bound PARP from DNA to allow access for other DNA repair

proteins to complete repair.

Figure 1: Mechanism of PARP action. DNA-damaging agents activate PARP, resulting in poly(ADP-ribose)-branched chains attached to DNA, recruiting associated repair proteins and cell cycle checkpoint mediators. This cascade may lead to cell cycle arrest while the cell commits to either DNA repair or apoptosis. Overactivation of PARP will lead to NAD+ depletion and necrotic cell death. PARP inhibition is thought to impair DNA repair function, leading to cellular dysfunction and death, and may also affect other PARP-mediated DNA modulating effects. Figure taken from Ratnam and Low (21).

4

PARP Inhibitors

PARP inhibitors represent the first FDA approved DDR-targeted medicines and have

transformed treatment paradigms for subgroups of patients with ovarian and breast

cancers. DDR deficiencies are common in cancer and represent an Achilles’ heel that

can be targeted. PARP inhibitors all interact with the binding site of the PARP enzyme

cofactor, β nicotinamide adenine dinucleotide (β-NAD+), in the catalytic domain of

PARP1 and PARP2.

Currently, there are six small-molecule PARP inhibitors available in the clinic or in

clinical trials: rucaparib, olaparib, niraparib, talazoparib, veliparib, and pamiparib.

Olaparib, rucaparib, and niraparib have obtained FDA approval in ovarian cancer in

different settings and talazoparib, approved for metastatic breast cancer (22), is in phase

3 clinical trials for ovarian cancer.

Figure 2: Chemical structures of the six small-molecule PARP inhibitors available in the clinic or in clinical trails: rucaparib, olaparib, niraparib, talazoparib, veliparib, and pamiparib. Talazoparib is the largest in size and possesses a more rigid structure, enhancing its binding ability to create the “PARP trapping” effect.

While there are mechanistic similarities, the six PARP inhibitors differ in their

chemical structure, potency, and clinical doses used for patients (23). Talazoparib, a

second-generation drug, is the largest in size and possesses a more rigid structure

5

compared with the other PARP inhibitors (Figure 2) (24). These differences in size and

rigidity are thought to be the basis for distinct capacity of each drug to prevent the

release of bound PARP1/2 from chromatin; a phenomenon known as "PARP trapping"

(24). Talazoparib is able to bind chromatin and create these trapped PARP–DNA

complexes to an approximately 100-fold greater degree than rucaparib, niraparib, or

olaparib, whereas veliparib displays negligible PARP-trapping ability (23,25). PARP

trapping is one mechanism that PARP inhibitors induce DNA replication stress (Figure

3). By generating an obstacle for the replication machinery, PARPi increase replication

stress in cancer cells (26). Other replication stress-inducing mechanisms include

impairing single-strand break repair (16) and accelerating replication fork progression

(27), which all combined, amplify the replication stress in tumor cells. However, the

stronger PARP trappers often have to be used at lower doses in the clinic due to lower

maximum tolerated dose (MTD) achieved (24). The appeal of combination therapy with

PARP inhibitors exploits the action of PARP inhibition while potentially sparing patients

the toxicities related to higher doses.

Figure 3: Dual cytotoxic mechanisms of PARP inhibitors. In the upper pathway, inhibition of PARP enzyme activity interferes with the repair SSBs, leading to replication fork damage that requires HR repair. In the lower pathway, trapping of PARP-DNA complexes also leads to replication fork damage but uses additional repair pathways and lethality is not restricted to HR deficiency. Figure adapted from Murai et al. (23)

6

Replication Protein A (RPA) Structure, Phosphorylation, and Function

RPA is a heterotrimeric, single-stranded DNA-binding protein with subunits RPA1

(RPA70, 70kDa), RPA2 (RPA32, 32kDa), and RPA3 (RPA14, 14kDa) that performs

critical functions for genome maintenance and cell proliferation. One structural feature

that provides RPA with its functional versatility are the oligonucleotide binding (OB)

folds, which are beta-barrel structures capable of wrapping around ssDNA (28). These

OB folds form 6 DNA-Binding domains (DBD’s), DBD-A, B, C, and F on RPA1, DBD-D

on RPA2, and DBD-E on RPA3 (29). RPA binds ssDNA with high affinity through

multiple DNA binding domains (DBDs). RPA can alter its affinity for ssDNA through

variable contact of these DBD’s with ssDNA, resulting in four distinct binding modes

(28). DBD-A, B and C are located on RPA1, while DBD-D and the C-terminal winged

helix-turn-helix domain (WH) are located on the N-terminus of RPA2 (Figure 4).

Figure 4: Cartoon structure of the RPA heterotrimer. The RPA heterotrimer, RPA1 (RPA70), RPA2 (RPA32), and RPA3 (RPA14), is depicted with the oligonucleotide binding (OB) folds DBD-A through DBD-F, and an acidic alpha-helix binding to the basic cleft of the N-terminal of RPA1 (DBD-F).

RPA exhibits a variable binding affinity to ssDNA while simultaneously interacting

with multiple proteins on multiple protein interaction domains. Thus, RPA activity needs

to be modulated to accommodate the metabolic needs of the cell (28). This is achieved

via phosphoinositide 3-kinase related kinase (PIKK)-dependent phosphorylation at a

number of potential sites, primarily on the N-terminus of RPA2 (29). For instance, RPA2

7

is phosphorylated at certain sites throughout stages of the cell cycle such as Ser23 and

Ser29 in G1/S transition, G2, and M-phase (30). Substitution mutations of Ser23 and

Ser29 with alanine and exposure of M-phase cells to DNA damage lead to delayed

mitotic exit, suggesting a role of RPA in mitotic checkpoint recovery (31).

In DNA damage response (DDR), RPA2 is phosphorylated on at least 9 sites: Ser4,

Ser8, Ser11, Ser12, Ser13, Thr21, Ser23, Ser29 and Ser33 (Figure 5) (32). Different

RPA2 target sites are phosphorylated by different PIKK’s, such as DNA-dependent

protein kinase (DNA-PK) and ataxia-telangiectasia mutated kinase (ATM)

phosphorylating S4/S8, and ATM and Rad3-related kinase (ATR) phosphorylating S33

(33). Studies show that phosphorylation of certain RPA2 sites occurs in a sequential

manner and phosphorylation of one site may be necessary to enable phosphorylation of

other sites. For example, blocking phosphorylation of residues such as T21, S23, or

S33 via a mutation prevents S29 and S4/S8 phosphorylation in response to

camptothecin (34). Additionally, reciprocal priming relationships may exist between

sites, as evidenced by S4/S8 showing mutual priming activity towards T21 (33).

Phosphorylation of S4/S8 is considered to be a specific and sensitive marker of DNA

damage (Figure 6) (35). Mutant forms of RPA containing alanine residues in place of S4

and S8 position of RPA2 have been shown to suppress the apoptosis response, possibly

through impaired detection of DNA damage and appropriate DDR signaling (36).

8

Figure 5: Schematic diagram of the RPA heterotrimer subunits with domain functions and consensus PIKK/CDK site, DNA-binding domains (DBD), and phosphorylation targets. The RPA heterotrimer, RPA1 (RPA70), RPA2 (RPA32), and RPA3 (RPA14), has the ability to differentially modulate its affinity for DNA through different combinations of DBD contact while also modulating its role in DNA metabolism via different phosphorylation sites. Figure adapted from Borgstahl et al. (29)

Figure 6: Schematic diagram of Ser4/Ser8 RPA2 hyperphosphorylation in the G2 phase of the cell cycle following DNA Damage. Phosphorylation of S4S8 at T0 following a DNA-damage event is followed by an increase in S4S8 hyperphosphorylation levels at T1. Figure adapted from Borgstahl et al. (29)

9

Role of RPA in Homologous Recombination and DNA Resection

Homologous recombination is initiated by the MRE11-RAD50-NBS1 (MRN) complex,

which rapidly localizes to DSBs (37). MRN initiates HR by removing adducts from the

DNA ends and by loading the Bloom’s syndrome (BLM) helicase along the exonuclease

1 (EXO1) or DNA2 nuclease and helicase (38,39). DNA resection occurs when ssDNA

is generated by nucleolytic degradation of one of the two DNA strands. Replication

protein A (RPA) rapidly coats the ssDNA that is generated during DNA resection. RPA-

ssDNA filaments are phosphorylated by ATR, together with ATM, cyclin-dependent

kinase (CDK), and DNA-dependent protein kinase, catalytic subunit (DNAPKcs).

RPA is phosphorylated as part of DNA damage response (DDR) and is a negative

regulator of DNA resection, stopping resection at the proper length, in which helicases

and nucleases process the free DNA ends to expose ssDNA. RPA phosphorylation has

been found to be a critical regulator of resection on chromatin and other processes that

involve BLM helicase. Phosphorylated RPA (pRPA) induces conformational changes

within the RPA1 subunit and inhibits DNA resection via inhibition of BLM helicase (40).

The N-terminus of RPA1 interacts with both BLM and the phosphorylated N-terminus of

RPA2 (Figure 7) to regulate DNA resection. pRPA suppresses BLM initiation at DNA

ends and triggers the intrinsic helicase strand-switching activity (Figure 8). Thus, pRPA

provides a feedback loop between DNA resection and DDR.

10

Figure 7: Schematic of RPA showing key phosphorylation sites on RPA2 (RPA32) (left on top). RPA2 phosphorylation induces physical interactions with RPA1 (RPA70N) (left on bottom). Schematic depiction of BLM interacting with the N-terminus of RPA1 (RPA70N) via at least two N-terminal acidic patches (red) (right). Figure adapted from Soniat et al. (41)

Figure 8: Schematic illustrating during DNA damage response, phosphorylated RPA is a negative regulator of BLM helicase and DNA resection. Termination of DNA resection is necessary for maintaining genome stability during DNA break repair. As phosphorylated RPA accumulates during resection, a negative feedback loop occurs between DNA resection and its termination. Figure taken from Soniat et al. (41)

RPA DBD-F as a novel small-molecule therapeutic target

RPA interacts with multiple proteins and exchanges protein-binding partners in an

organized and controlled process primarily through interactions with the N-terminus of

RPA1 (DBD-F). The N-terminus of RPA1 is connected to the rest of RPA1 through a

long flexible linker, which allows for it to function as a protein-protein interaction domain

and contribute to roles in DNA damage signaling (42). The N-terminus of RPA1 is less

involved in ssDNA binding activity of the RPA complex compared to the other DBDs.

The binding polarity of RPA directs the proper location of the initial protein complexes

recruited by RPA. Many important genome maintenance proteins containing an acidic

11

alpha-helical domain have been identified that bind to the N-terminus of RPA1 through

electrostatic contributions.

Recent studies have revealed additional interacting proteins with DBD-F (43). One

example is the Bloom syndrome protein (BLM), a helicase that contributes to genomic

stability through its activity in DNA replication and HR repair of double-strand breaks

(DSBs), binds specifically to DBD-F (44). An additional number of DNA repair and

checkpoint response proteins have also been shown to bind to RPA1 in response to

replication stress (28).

A novel therapeutic strategy is to develop an inhibitor that binds specifically to the

DBD-F protein-binding domain and block its interaction with DNA processing proteins,

which would allow RPA to bind ssDNA, while selectively and simultaneously inhibiting

several critical DNA damage response pathways (e.g., pathways involving ATR, p53,

and Rad9) to regulate DNA resection. This therapeutic strategy should result in a

different phenotype from the full protein knockout, since, ideally, the protective ssDNA

binding and other functions of RPA would not be affected in normal cells not

experiencing high constitutive replication stress. Conventional genetic loss-of-function

experiments and RNA-mediated silencing of a protein can exhibit different effects

compared to small-molecule inhibition of protein function. With small-molecule inhibition,

one may be able to achieve a therapeutic window, as initiation of the DDR in cancer

cells would be inhibited. As many cancer cells are constitutively undergoing higher

replication stress than normal cells, this inhibition would be cytotoxic. Blocking DBD-F

without affecting the ssDNA binding function and other functions of RPA can be

accomplished by identifying a small molecule that inhibits RPA binding to double-

stranded DNA (dsDNA) but not ssDNA binding. Under specific experimental conditions,

RPA binds dsDNA with lower affinity than ssDNA via a highly efficient helix

destabilization process (45,46). This unwinding ability is greatly reduced in the absence

12

of the DBD-F of RPA (40). However, deletion of DBD-F does not significantly alter

ssDNA binding affinity. The basic cleft of DBD-F has been shown to contribute to the

nucleation of dsDNA, the initial and rate limiting step in DNA unwinding by RPA. In the

nucleation process, RPA binds to transient ssDNA regions in the dsDNA contributing to

the helix destabilization of dsDNA (47). Taking advantage of this unique property of

RPA, the Oakley laboratory has developed a high-throughput assay to identify small

molecules that bind specifically to DBD-F and not the DBDs involved in ssDNA binding,

leading to the identification of PAME as a candidate inhibitor targeting DBD-F.

By targeting the cellular response to replication stress via RPA, this leads to an

increase in replication stress and simultaneously decreases the replication stress

response in cancer cells that have constitutively high DNA replication stress (48). An

inhibitor that blocks RPA protein-protein interactions combined with PARP inhibition has

the potential to increase replication stress and DNA damage while selectively inducing

cell death in HGSC cells containing both wild-type and mutant BRCA1/2 (Figure 9). This

strategy has been demonstrated in a study where inhibition of the DBD-F domain of RPA

increased replication stress in head and neck squamous cell carcinoma cells as

determined by pan-nuclear H2AX phosphorylation in S-phase (48). This type of

phosphorylation has been determined to be replication stress specific as previously

demonstrated with Chk1 inhibitors (24).

13

Hypothesis

The hypothesis of this thesis is the combination of RPA inhibition and PARP

inhibition will exhibit an enhanced effect and inhibit RPA’s response to DNA damage,

reducing cell viability in HGSC OVCAR cells. This is examined using the electrophoretic

mobility assay, cell viability assay, and immunofluorescence detection.

Figure 9: Chemical structures of RPA and PARP inhibitors examined. PAME has been identified from an initial screen as a RPA DBD-F inhibitor. Talazoparib has been in use in the clinic as a PARP second-generation inhibitor.

Specific Aims

1. Evaluate effects of PAME (RPAi) on RPA ssDNA binding and dsDNA unwinding.

2. Evaluate effects of PAME (RPAi) and Talazoparib (PARPi) on cell viability.

3. Evaluate effects of PAME (RPAi) and Talazoparib (PARPi) on DNA damage

response and replication stress.

14

CHAPTER 2: MATERIALS AND METHODS

Compounds and Antibodies

Talazoparib (BMN-673), a PARP1/2 inhibitor, was purchased from Med Chem

Express. PAME (RPA inhibitor, NSC149526) was acquired from the National Cancer

Institute Developmental Therapeutics Program.

Primary antibodies used for immunohistochemistry (IHC) are mouse anti-RPA32

raised against RPA32 of human origin (9H8, Santa Cruz Biotechnology, Dallas, TX) and

rabbit anti-S4S8 Phospho RPA32 (Bethyl Laboratories Inc, Montgomery, TX). Rabbit-

anti-H2AX (Upstate) was raised against H2AX of human origin.

Secondary antibodies and IHC for detection of RPA32 was carried out using Anti-

mouse Ig HRP Detection Kit (BD Biosciences, San Jose, CA). For pS4S8-RPA32

detection, anti-rabbit Bethyl IHC Accessory Kit was used

Electrophoretic Mobility Shift Assay (EMSA)

All EMSA incubations were performed in 10 μl binding buffer (15 mM NaCl was used

with dsDNA analyses) with 0.4 pmol RPA at RT. Reactions were loaded on 1% agarose

gels in 40 mM Tris acetate, and run for 15 min at 100 V. Gels were scanned and

visualized on the Odyssey infrared scanner.

For screening of compounds that disrupt ssDNA bound RPA, RPA was pre-bound to

2 pmol of polyT(30) oligonucleotide labeled with 5′ IRDYE-700 (Integrated DNA

Technologies) for 5 min, then exposed to 2 nmol of inhibitor for 30 min. For

measurement of the inhibition of RPA/ssDNA binding, RPA was exposed to varying

amounts of inhibitor for 5 min followed by the addition of 2 pmol of ssDNA for 30 min.

For dsDNA binding studies, a fluorescent dsDNA substrate was generated by

annealing the fluorescent polyT (30) oligonucleotide to an unlabeled polyA(30)

oligonucleotide. The duplex DNA was treated with mung bean nuclease to remove any

remaining ssDNA. For detection of dsDNA binding and helix destabilization, RPA was

15

exposed to varying amounts of inhibitor for 5 min before the addition of 2 pmol of dsDNA

for 30 min.

Cell Viability Assay

All cells were grown in Dulbecco’s modified Eagle’s Ham’s F-12 medium

(DMEM/F12) supplemented with 10% heat-inactivated fetal bovine serum (FBS) and 1X

Penicillin-Streptomycin (Pen-Strep) purchased from Thermo Fisher Scientific – US. Cell

cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2. To

obtain cells primarily in S phase, asynchronous cells were treated with 6 μM aphidicolin

(Sigma-Aldrich, St. Louis, MO) for 16–20 h. The medium containing aphidicolin was

removed, and cells were washed twice in serum-free medium and then incubated in

serum-containing medium for an additional 2–4 h.

Briefly, cells were incubated for 1–4 days. Cells were seeded in 6-well plates. Cells

were treated with DMSO, 25 μM of PAME, 75 nM of talazoparib, or 25 μM of PAME and

75 nM of talazoparib for 6 hours and removed and replaced with fresh media. Cells

were allowed to grow for an additional 18 hours. The numbers of viable cells were

counted using a hemocytometer. To measure cell death, Trypan blue staining was

performed by mixing 0.4% Trypan blue in PBS with cell suspension at a 1:10 ratio. Live

or dead cells were counted using Trypan blue exclusion method and ≥100 cells were

counted.

Data were analyzed using an unpaired 2-tailed Student’s t test to determine the

statistical significance.

Cell Lines

FTE282 was used as a model for fallopian tube epithelial cells. The parental FTE

cell line has a very low level of intrinsic replication stress.

HGSC cell lines including OVCAR5 (BRCA1+), OVCAR4 (BRCA1-), and OVCAR8

(BRCA1-) was used to cell viability assays and immunofluorescence studies. The

16

HGSC cell lines vary but overall have higher levels of constitutive replication stress

based on constitutive phosphorylation of P-Chk1 (S345 and S317), P-ATR(T1989), and

P-RPA(S33).

Immunofluorescence Detection

Cells were grown on 22-mm coverslips overnight prior to treatment with RPAi and

PARPi. Cells synchronized in G1/S with aphidicolin, non-treated and treated with PAME

and talazoparib for 6 h and the inhibitors were removed and replaced with fresh media

for an additional 6 h. After an initial wash with PBS, cells were extracted with PBS

containing 0.5% Triton X-100 for 150 sec on ice, and fixed with 4% paraformaldehyde for

15 min. Next, the coverslips were blocked with 15% goat serum at room temperature,

and then incubated with primary antibodies to H2AX (Upstate), PS4/8-RPA32 (Bethyl

Laboratories), in blocking solution for 1 h. The coverslips were washed with PBS and

incubated with an appropriate Alexa-Fluor 488- or Alexa-Fluor 568-conjugated antibody

in blocking solution for 1 h. Cells were mounted in PermaFluor (Fisher) supplemented

with 0.5 µg/mL DAPI (Roche). Immunofluorescent images were captured digitally with a

Zeiss Axiovert 200M microscope. 200 nuclei were counted per sample. A minimum of

10 foci per cell were required to count as positive. Standard deviation was calculated

from three replicate experiments.

17

CHAPTER 3: RESULTS

PAME does not inhibit RPA ssDNA binding

PAME’s RPA inhibitory effect on ssDNA binding was evaluated with RPA ssDNA

binding domains using an electrophoretic mobility shift assay (EMSA). The RPAi was

tested at concentrations of 12.5, 25, 50, 100, and 200 µM. Results did not suggest a

difference in the amount of the RPA-ssDNA complex and the amount free DNA with

increasing concentrations of PAME (Figure 10), thus PAME did not inhibit RPA binding

to ssDNA.

Figure 10: PAME was examined for its RPA inhibitory property of ssDNA binding. Increasing dosage of PAME was examined using an EMSA with a RPA-ssDNA complex and did not suggest inhibition of ssDNA binding.

PAME inhibits the ability of RPA to unwind dsDNA

To examine PAME binding to the N-terminal of RPA1, the inhibitory action of the

unwinding activity of RPA on a 30nt dsDNA was evaluated at concentrations of 1.5,

3.13, 6.25, 12.5, 25 µM using an EMSA. The results suggest that PAME has an RPA

inhibitory effect on dsDNA unwinding, with an IC50 estimated at 3.125-6.25 µM (Figure

11).

Figure 11: PAME was examined for its ability to inhibit dsDNA unwinding. To examine the ability of PAME to unwind dsDNA, the RPAi was tested using an EMSA at increasing concentration (1.50, 3.13, 6.25, 12.50, 25 µM) with an RPA-DNA complex. EMSA results suggest that PAME has an inhibition effect on dsDNA unwinding with an IC50 estimated at 3-6 µM.

18

Effects of RPA and PARP inhibitors on cell viability

RPAi (PAME) and PARPi (talazoparib (TZ)), were evaluated with HGSC OVCAR

cells and FTE cells (control) using a Trypan blue cell viability assay. Drugs tested

include DMSO (control), PAME (25µM), TZ (75nM), and a combination of PAME (25µM)

and TZ (75nM). In OVCAR5 (BRCA1+) cells, both PAME and TZ individually exhibited a

mild reduction in the proportion of viable cells, while a combination of PAME and TZ

showed a significant reduction in the proportion of viable cells (Figure 12A).

In OVCAR4 (BRCA1-) cells, PAME exhibited a significant reduction in the proportion

of viable cells, TZ also exhibited a reduction, while a combination of PAME and TZ

exhibited an additive reduction in the proportion of viable cells (Figure 12B).

In OVCAR8 (BRCA1-), PAME exhibited a decrease in the proportion of viable cells,

TZ showed a greater proportion of reduction compared with PAME, while a combination

of PAME and TZ exhibited an additive reduction in the proportion of viable cells (Figure

12C).

In FTE282 cells (control group), PAME, TZ, and the combination of both drugs did

not exhibit a significant reduction in the proportion of viable cells. (Figure 12D).

Overall, the ranking of sensitivity to the drugs for each cell line was OVCAR4 >

OVCAR5 > OVCAR8 > FTE.

19

Figure 12: Effects of RPAi and PARPi in HGSC OVCAR cells examined by cell viability assay. A. Ovarian cancer cell line OVCAR5 (BRCA1/2+). B. Ovarian cancer cell line OVCAR4 (BRCA1-). C. Ovarian cancer cell line OVCAR8 (BRCA1-). D. FTE282 cells (control).

Immunofluorescence detection of RPA S4/S8 and Histone H2AX

Phosphorylation by RPAi and PARPi

Immunofluorescence detection of RPA S4/S8 and histone H2AX phosphorylation

with PAME (RPAi) and TZ (PARPi) was examined in HGSC OVCAR cells with

BRCA1/2+ and BRCA1-, and FTE282 cells as control. Phosphorylation of RPA S4/S8

(pS4/S8) (red) suggests inhibitory effects of DNA resection, thus inhibition of

homologous recombination. Phosphorylation of H2AX (pH2AX) (green) suggests DNA

20

DSB formation, or DNA damage. DAPI was used to label DNA (blue) to detect where

their location, and was used to indicate that RPA pS4/S8 and pH2AX is localized with

DNA. The “Merge” column is the merging of H2AX and DAPI immunofluorescence (blue

and green) to label pH2AX localized with DNA. The proportion of positive cells detected

for pS4/S8 and H2AX was quantified in bar graphs.

FTE282 (Control) examined with RPAi and PARPi

In FTE282 cells, treatment without any inhibitors (control) did not lead to

immunofluorescence of RPA S4/S8 (red), and a small amount was detected for H2AX

(green), indicating that phosphorylation did not occur at S4/S8, thus a lack of RPA

response to DNA damage. A small amount detected in H2AX, indicating the occurrence

of DNA damage. Treatment with RPAi (PAME) led to immunofluorescence near the

same level as treatment without any drugs in RPA S4/S8, suggesting inhibition of RPA

phosphorylation in the presence of the RPAi. With RPAi treatment, the proportion of

immunofluorescence of H2AX was greater than the control, and at about the same level

as treatment with PARPi (TZ), suggesting the formation of DNA DSB was greater than

the control, and at comparable levels as treatment with PARPi.

Treatment with PARPi (TZ) led to a significant increase in pS4/S8 compared with

treatment with RPAi (PAME), RPAi and PARPi (PAME and TZ), and the control,

indicating RPA phosphorylation in response to DNA damage. The proportion of H2AX

immunofluorescence was similar to treatment with PAME, and less than the control,

suggesting DNA DSB formation. In the “Merge” column, pH2AX is shown to be localized

with DNA.

Combination treatment with PARPi and RPAi (TZ+PAME) led to a significant

reduction of pS4/S8 compared with treatment with PARPi, and comparable to the levels

in treatment with RPAi and the control, suggesting the combination of RPAi and PARPi

can inhibit pRPA, thus inhibiting RPA’s response to DNA damage. The proportion of

21

cells detected with pH2AX was greater in the combination treatment compared with

treatment with only RPAi (PAME) or PARPi (TZ).

Figure 13: Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in FTE282 cells. (40x and 60x magnification)

OVCAR5 (BRCA1/2+) examined with RPAi and PARPi

In HGSC OVCAR5 (BRCA1/2+) cells, treatment without any inhibitors (control) did

not lead to immunofluorescence at RPA S4/S8, and a minimal amount was detected at

H2AX, indicating the lack of DNA damage. Treatment with RPAi (PAME) did not result

in immunofluorescence at RPA S4/S8, and a moderate amount of immunofluorescence,

about one-third of the cells, was detected at H2AX, suggesting RPA inhibition and the

occurrence of DNA damage. Treatment with PARPi (TZ) exhibited in an increased

amount of immunofluorescence at RPA S4/S8, and an increased amount of H2AX was

detected, suggesting RPA’s response to DNA damage and the occurrence of DNA

damage. Combination treatment of PARPi and RPAi (TZ+PAME) exhibited a significant

decrease in immunofluorescence of pS4/S8 compared with PARPi (TZ), and a

22

comparable amount compared with RPAi (PAME) and control, suggesting inhibition of

RPA’s response to DNA damage. pH2AX detection exhibited a significant increase

compared to treatment with PARPi (TZ), and significantly more compared with RPAi

(PAME), with many cells displaying pan-nuclear H2AX phosphorylation, suggesting the

occurrence of DNA damage due to replication stress.

Figure 14: Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR5 (BRCA1/2+) cells. (40x and 60x magnification)

OVCAR8 (BRCA1-) examined with RPAi and PARPi

In HGSC OVCAR8 (BRCA1-) cells, treatment without any inhibitors (control) did not

lead to immunofluorescence at RPA S4/S8, and a minimal amount was detected at

H2AX. Treatment with RPAi (PAME) did not result in immunofluorescence at RPA

S4/S8, and a moderate amount of immunofluorescence, about one-third of the cells, was

detected at H2AX, indicating RPA inhibition. Treatment with PARPi (TZ) exhibited a

slightly increased amount of immunofluorescence at RPA S4/S8 and H2AX compared to

RPAi (PAME), indicating a level of RPA DDR and occurrence of DNA damage.

23

Combination treatment of PARPi and RPAi (TZ+PAME) resulted in a similar amount of

immunofluorescence of pS4/S8 compared with PARPi (TZ) and RPAi (PAME), indicating

a minimal activation of RPA phosphorylation. pH2AX detection had a significant

increase compared with treatment with PARPi (TZ) and RPAi (PAME), indicating a

greater amount of DNA damage.

Figure 15: Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR8 (BRCA1-) cells. (40x and 60x magnification)

OVCAR4 (BRCA1-) examined with RPAi and PARPi

Results in OVCAR4 (BRCA1-) cells were similar to ones obtained in OVCAR8 cells.

Treatment without any inhibitors (control) did not lead to immunofluorescence at RPA

S4/S8, and a minimal amount was detected at H2AX. Treatment with RPAi (PAME) did

not exhibit immunofluorescence at RPA S4/S8, and a moderate amount of

immunofluorescence, about one-third of the cells, was detected at H2AX, indicating RPA

inhibition and occurrence of DNA damage. Treatment with PARPi (TZ) exhibited a

24

similar amount of immunofluorescence at RPA S4/S8 and H2AX compared to RPAi

(PAME), indicating minimal response to PARP inhibition in these BRCA1/2+ cells.

Combination treatment of PARPi and RPAi (TZ+PAME) resulted in a similar amount of

immunofluorescence of pS4/S8 compared with PARPi (TZ) and RPAi (PAME),

suggesting RPA inhibition. pH2AX detection had a significant increase compared with

treatment with PARPi (TZ) and RPAi (PAME), indicating the occurrence of DNA

damage.

Figure 16: Immunofluorescence detection of RPA S4/S8 and H2AX phosphorylation with no drugs (control), RPAi (PAME), PARPi (TZ), and both drugs (TZ+PAME) in OVCAR4 (BRCA1-) cells. (40x and 60x magnification)

Immunofluorescence detection of pH2AX after a 30-hour recovery period

Immunofluorescence detection of pH2AX with and without drug treatment after a

30-hour recovery period was examined in each of the cell lines to assess the degree of

DNA damage remaining after allowing time for cells to repair DNA damage.

25

In FTE282 cells, treatment without drugs (DMSO), RPAi (PAME), PARPi (TZ), and

both RPAi and PARPi (TZ and PAME) did not exhibit a significant difference in pH2AX

after a 30-hour recovery period, indicating minimal DNA damage.

In OVCAR8 (BRCA1-) cells, treatment with RPAi and PARPi (TZ and PAME)

exhibited approximately 50% of the cells to be detected with pH2AX, which was

significantly greater than the 20% of cells detected with PARPi (TZ), 10% detected with

RPAi (PAME), and the control (DMSO), indicating the occurrence of DNA damage.

In OVCAR5 (BRCA1/2+) cells, treatment with RPAi (PAME) led to approximately

15% of cells detected, which was significantly greater than treatment with PARPi (TZ),

PARPi and RPAi (TZ and PAME), and the control (DMSO), indicating the occurrence of

DNA damage.

In OVCAR4 (BRCA1-) cells, treatment with RPAi and PARPi (TZ and PAME) led to

approximately 50% of the cells to be detected with pH2AX, which was significantly

greater than the 20% of cells detected with PARPi (TZ), 10% detected with RPAi

(PAME), and the control (DMSO). These results were similar to the ones detected for

OVCAR8 cells.

26

Figure 17: Immunofluorescence detection of pH2AX with no drug treatment (DMSO), RPAi (PAME 25µM), PARPi (75nM TZ), and both drugs (75nM TZ+25µM PAME) after 30h recovery in FTE282 (control), OVCAR8 (BRCA1-), OVCAR5 (BRCA1/2+), and OVCAR4 (BRCA1-) cells.

27

CHAPTER 4: DISCUSSION

The therapeutic strategy to target the cellular response to replication stress by

inhibiting RPA, thus leading to an increase in replication stress and simultaneously

decreasing the replication stress response in cancer cells, was evaluated in this study

with RPAi and PARPi in the context of HGSC OVCAR cells.

RPAi effects on ssDNA Binding and dsDNA unwinding

PAME did not inhibit RPA ssDNA binding, which is a desired RPAi effect, to not

interfere with RPA binding and function with ssDNA. This suggests that in the presence

of an RPAi, RPA can continue to bind other protein partners and function with ssDNA,

while inhibiting critical DDR pathways to regulate DNA resection.

PAME did exhibit an inhibitory effect on dsDNA unwinding in vitro, indicative of

inhibiting RPA’s ability to bind dsDNA.

Together, these results suggests that PAME is binding to DBD-F domain to inhibit

RPA’s function, since it did not affect the ssDNA binding function but did displayed

inhibition of RPA binding to dsDNA.

RPAi and PARPi effects on cell viability

Combination of RPAi and PARPi displayed an enhanced effect on inhibiting the

viability of HGSC OVCAR5 (BRCA1/2+), OVCAR4 (BRCA1-), and OVCAR8 (BRCA1-)

cells. The presumed mechanism is that replication stress induced by RPAi enhances

the PARPi effect, more effectively suppressing DNA damage response.

RPAi and PARPi effects on RPA S4/S8 and H2AX phosphorylation

Combination treatment with RPAi and PARPi displayed enhanced DNA damage in

all HGSC cells and FTE cells as compared with each of the drug treatment by itself.

Results observed across cell lines are indicative of an enhanced effect of RPAi and

PARPi.

28

The cytotoxic mechanism of action of RPA and PARP inhibition is potentially

multifactorial, with contributions from an overreliance on alternative DDR pathways in

PARP inhibitor resistant cells, as well as catastrophic DNA damage and replication

stress when PARP inhibitors are combined with RPA inhibitors. This selectively induces

cell death in HGSC cells.

Combination therapy has the potential to can overcome PARPi resistance and target

cells with or without BRCA deficiency to inhibit RPA’s role in regulating DNA resection

and homologous recombination.

RPAi and PARPi effects on pH2AX after a 30-hour recovery period

In the pH2AX 30-hour recovery immunofluorescence detection, combination

treatment of RPAi and PARPi led to approximately 50% of positive cells in the OVCAR4

and OVCAR8 BRCA1- cells, whereas in FTE282 and OVCAR5 (BRCA1+) cells, there

was only a small proportion of positive cells detected. This suggests that BRCA1- cells

that were deficient in homologous recombination were not able to repair the DNA

damage. FTE282 and OVCAR5 cells with the ability to perform homologous

recombination was able to repair their DNA damage over the recovery period. The

combination drug treatment in OVCAR4 and OVCAR8 cells also led to a greater

proportion of positive cells compared with treatment with RPAi and PARPi individually,

indicating an enhanced effect is observed over the recovery period. An enhanced effect

was not observed in the FTE282 and OVCAR5 cells, which again may be explained by

the ability of these cells to repair DNA damage.

29

CHAPTER 5: CONCLUSIONS

Results from this study suggest that the combination of RPAi (PAME) and PARPi

(talazoparib) provided an enhanced effect in reducing the proportion of cell viability and

suppressing RPA DNA damage response in immunofluorescence studies in FTE and

HGSC OVCAR cell lines. Additional studies will be needed to further examine the

effects of the combination of RPA inhibition and PARP inhibition in the context of high-

grade serous ovarian cancer, such as in mouse with an ovarian cancer model.

30

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