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University of Nebraska Medical Center University of Nebraska Medical Center
DigitalCommons@UNMC DigitalCommons@UNMC
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
Follow this and additional works at: https://digitalcommons.unmc.edu/etd
Part of the Cancer Biology Commons, and the Female Urogenital Diseases and Pregnancy
Complications Commons
Recommended Citation Recommended Citation Tang, Yat, "Characterizing the Combination of RPA Inhibitors with PARP Inhibitors in High-Grade Serous Ovarian Cancer" (2019). Theses & Dissertations. 416. https://digitalcommons.unmc.edu/etd/416
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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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