1521-0103/353/3/446–457$25.00
THE JOURNAL OF PHARMACOLOGY AND EXPERIMENTAL THERAPEUTICS
Copyright ª 2015 by The American Society for Pharmacology and Experimental Therapeutics
http://dx.doi.org/10.1124/jpet.114.222448
J Pharmacol Exp Ther 353:446–457, June 2015
Minireview
Trapping Poly(ADP-Ribose) Polymerase
Yuqiao Shen, Mika Aoyagi-Scharber, and Bing Wang
BioMarin Pharmaceutical Inc., Novato, California
Received December 29, 2014; accepted March 9, 2015
Introduction
Poly(ADP-ribose) polymerase (PARP)-1 and its close relative PARP2 play essential roles in the repair of DNA singlestrand breaks (SSBs) (Rouleau et al., 2010; Curtin and Szabo,
2013). PARP1 is an abundant nuclear protein that detects
broken DNA strands and binds damaged DNA through its
N-terminal zinc-finger domain (Zn), leading to activation of
the C-terminal catalytic domain (CAT) that hydrolyzes NAD1
to produce linear and branched poly(ADP-ribose) polymers
that extend hundreds of ADP-ribose units (Krishnakumar
and Kraus, 2010; Langelier et al., 2011). This post-translational
modification, referred to as PARylation, of multiple protein
substrates, including PARP1 itself, promotes the recruitment of
DNA repair proteins (such as X-ray repair cross-complementing
protein 1 and MRE11A) to the site of SSBs, allowing subsequent
repair (Rouleau et al., 2010; Curtin, 2012). Auto-PARylation of
PARP1 occurs at its regulatory domain; due to the high negative
charge of poly(ADP-ribose), extensive auto-PARylation of
PARP1 facilitates its dissociation from DNA, which is necessary
for the completion of DNA repair (Satoh and Lindahl, 1992;
Langelier et al., 2011; Luo and Kraus, 2012). This process is
critical for the resealing of DNA SSBs during base excision
repair (BER) (Juarez-Salinas et al., 1979; Benjamin and Gill,
1980; Durkacz et al., 1980), as well as repairing topoisomerase
dx.doi.org/10.1124/jpet.114.222448.
trap PARP differ by several orders of magnitude, with the ability to
trap PARP closely correlating with each drug’s ability to kill cancer
cells. In this article, we review the available data on molecular
interactions between these clinical-stage PARP inhibitors and
PARP proteins, and discuss how their biologic differences might
be explained by the trapping mechanism. We also discuss how to
use the PARP-trapping mechanism to guide the development of
PARP inhibitors as a new class of cancer therapy, both for singleagent and combination treatments.
(Top) 1 cleavage complexes (Zhang et al., 2011) and DNA doublestrand breaks (DSBs) (Audebert et al., 2004; Haince et al., 2008).
PARP2 functions in a similar manner to PARP1, but is comparatively less abundant, contributing only 5–10% of the total
PARP activity (Ame et al., 1999; Schreiber et al., 2006).
Small-molecule inhibitors of PARP1 and PARP2 are in development as a new class of anticancer agents (Lord and
Ashworth, 2012; Curtin and Szabo, 2013). Several selective
and potent PARP inhibitors have been reported, all of which
are analogs of nicotinamide (Curtin and Szabo, 2013; Ekblad
et al., 2013; Lupo and Trusolino, 2014). These inhibitors compete with NAD1 at the catalytic pocket of PARP, inhibiting
PARylation (Curtin and Szabo, 2013; Ekblad et al., 2013).
PARP inhibitors were originally developed to potentiate the
tumor-killing activity of ionizing irradiation and genotoxic
agents, such as temozolomide (TMZ) and topotecan. Indeed,
all PARP inhibitors tested thus far have demonstrated radioor chemopotentiation (Curtin and Szabo, 2013), consistent
with the key role of PARP1 and PARP2 in DNA damage repair
(Yelamos et al., 2011). An additional use for PARP inhibitors
was reported by both Bryant et al. (2005) and Farmer et al.
(2005), demonstrating that inhibition of PARP1/2 is synthetically lethal with the loss of function of either the BRCA1 or
BRCA2 tumor suppressor gene. Loss of BRCA1/BRCA2 gene
function is believed to cause a deficiency in homologous
recombination (HR)–mediated double-strand DNA break repair, making these cells highly susceptible to the DNA lesions
ABBREVIATIONS: ARTD, ADP-ribosyltransferase subdomain; BER, base excision repair; CAT, C-terminal catalytic domain; CPT, camptothecin;
D-loop, donor-site loop; DSB, double-strand break; HD, helical regulatory subdomain; HR, homologous recombination; N3mA, N3-methyladenine;
N7mG, N7-methylguanine; PARP, poly(ADP-ribose) polymerase; siRNA, small interfering RNA; SSB, single-strand break; TMZ, temozolomide; Top,
topoisomerase; Zn, Zinc-finger domain.
446
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
ABSTRACT
Recent findings indicate that a major mechanism by which poly
(ADP-ribose) polymerase (PARP) inhibitors kill cancer cells is by
trapping PARP1 and PARP2 to the sites of DNA damage. The
PARP enzyme-inhibitor complex “locks” onto damaged DNA and
prevents DNA repair, replication, and transcription, leading to cell
death. Several clinical-stage PARP inhibitors, including veliparib,
rucaparib, olaparib, niraparib, and talazoparib, have been evaluated for their PARP-trapping activity. Although they display similar
capacity to inhibit PARP catalytic activity, their relative abilities to
Trapping PARP
Evidence for PARP Trapping
Many clinically investigated PARP inhibitors, including
talazoparib (BMN 673; BioMarin Pharmaceutical Inc., Novato,
CA), niraparib (MK-4827; Tesaro Inc., Waltham, MA), olaparib
(AZD-2281; AstraZeneca, London, UK), rucaparib (AG-014699;
Clovis Oncology, Boulder, CO), and veliparib (ABT-888; AbbVie
Inc., North Chicago, IL), are highly efficacious at inhibiting
PARP catalytic activity, with very similar IC50 values in the
single-digit nanomolar range (Fig. 1A) (Rouleau et al., 2010;
Murai et al., 2012, 2014a; Shen et al., 2013). PARP inhibitors
were initially thought to exert their antitumor activity solely by
inhibiting the catalytic activity of PARP1 and PARP2. Based on
this theory, the inability of PARP to auto-PARylate prevents its
dissociation from DNA, resulting in “trapped” PARP-DNA
complexes that interfere with the repair of SSBs, and accumulation of unrepaired SSBs leads to DSBs and cell death. This
has led to an early version of the PARP trapping theory
(Lindahl, 1993; Helleday, 2011; Kedar et al., 2012), which
predicted that the cytotoxicity in BRCA-deficient cells and the
capacity for chemopotentiation are determined solely by
catalytic inhibition, implying that these clinical PARP inhibitors would exhibit similar cytotoxic potency given their
comparable PARP-inhibitory activities. However, we and others
have shown that the ability of these PARP inhibitors to kill
BRCA-deficient cancer cells and to potentiate the cytotoxicity
of TMZ differs by several orders of magnitude (Fig. 1A) (Shen
et al., 2013). Whereas rucaparib, olaparib, and niraparib have
comparable cancer cytotoxicity, talazoparib is approximately
25- to 100-fold more potent in these assays. In contrast, veliparib
is ∼1000- to 10,000-fold less potent than talazoparib in cytotoxicity assays. Similar observations were obtained in other laboratories (Murai et al., 2012, 2014a; Stewart et al., 2014). This
magnitude of differences in the cytotoxicity cannot be explained
simply by differences in catalytic inhibition. These differences in
cytotoxicity also cannot be explained by differential off-target
activity of the PARP inhibitors, as olaparib and veliparib have
similar selectivity profiles against 13 of the 17 human PARP
family members (Wahlberg et al., 2012). Talazoparib also
shows a similar selectivity profile against the PARP family
members (unpublished data). In addition, a recent report
demonstrated that PARP1 knockout cells are highly resistant
to talazoparib, indicating that the potent cytotoxicity of
talazoparib is mediated primarily by PARP1 rather than offtarget activities (Murai et al., 2014a).
A revised PARP trapping concept was proposed based on
analyses of several lines of experimental data (Murai et al.,
2012, 2014a). This new concept suggests that, in addition to
catalytic inhibition, some PARP inhibitors may induce an
allosteric conformational change in PARP1 and PARP2, thereby
stabilizing their associations with DNA. These studies have
provided direct evidence that PARP inhibitors exert their cytotoxicity primarily by this trapping mechanism rather than by
classic enzyme inhibition. The capacity to trap PARP-DNA
complexes varies widely across different PARP inhibitors and is
not correlated with their PARP catalytic inhibition. As
described below, several lines of evidence support this PARPDNA–trapping model.
First, if PARP catalytic inhibition were the sole mechanism of
PARP inhibitor cytotoxicity, one would expect the antitumor
cytotoxicity of an inhibitor to be proportional to its catalytic
inhibition. As discussed above, the cytotoxicities of veliparib,
rucaparib, olaparib, niraparib, and talazoparib differ by several
orders of magnitude, both in terms of single-agent activity in
BRCA-deficient cells and in the sensitization of cancer cells to
alkylating agents. This strongly suggests that some of the PARP
inhibitors may possess additional mechanisms of cytotoxic
action in addition to the inhibition of PARP enzymatic activity.
Second, if PARP catalytic inhibition were the sole mechanism
of PARP inhibitor cytotoxicity, depletion of PARP would be
expected to have equivalent or greater effect on the cytotoxicity,
as compared with the PARP inhibition by small-molecule inhibitors. However, treating cancer cells that harbor wild-type
PARP gene with PARP inhibitors, such as talazoparib and
olaparib, induces far superior cytotoxicity than genetic depletion
of PARP (Murai et al., 2012, 2014a). Similarly, knockdown of
PARP gene expression by small interfering RNA (siRNA) is far
less effective than PARP inhibitor treatment in inducing
apoptosis in BRCA-deficient cells (Bryant et al., 2005; Farmer
et al., 2005; Strom et al., 2011). Consistent with this cytotoxicity
data, SSB repair is delayed to a greater extent by smallmolecule inhibitors, as compared with PARP gene knockdown
by siRNA (Strom et al., 2011), with the level of DSBs considerably higher in olaparib-treated wild-type cells than in
PARP gene knockout cells (Murai et al., 2012).
Third, there is now mounting evidence demonstrating that
PARP itself is required for the cytotoxic effects of the PARP
inhibitors. Depletion of PARP renders cells resistant to both
talazoparib and olaparib (Murai et al., 2012, 2014a). Likewise,
cells with PARP1 loss-of-function mutations are 100-fold more
resistant to olaparib and talazoparib than wild-type cells, and
restoring PARP1 expression causes these cells to revert to wildtype sensitivity (Pettitt et al., 2013). These data indicate that
PARP1 protein is the critical mediator of the cytotoxic effect of
PARP inhibitors, arguing against the suggestion that these
differences in cytotoxicity result from off-target activities. PARP
is also required for sensitizing tumor cells to the alkylating
agents TMZ and methyl methanesulfonate (Liu et al., 2009;
Murai et al., 2012, 2014a). Other data have also confirmed that
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
caused by PARP inhibition (Ashworth, 2008; Helleday, 2011).
This concept has gained widespread support from both preclinical research and clinical investigations, providing the basis
for the first clinical attempt to exploit synthetic lethality in
cancer treatment. Subsequent work has shown that, in addition
to BRCA1/BRCA2 mutations, deficiency in other HR genes
(e.g., Fanconi anemia genes) may also cause cancer cells to
become sensitive to PARP inhibitors (McCabe et al., 2006; Peng
and Lin, 2011; Turner and Ashworth, 2011).
Until recently, PARP inhibitors were assumed to work solely
by inhibiting the catalytic activity of PARP1/PARP2. However,
Pommier and colleagues recently demonstrated that catalytic
inhibition is not the only mechanism by which PARP inhibitors
exert cytotoxic effects (Murai et al., 2012, 2014a). Selected PARP
inhibitors may also trap PARP1 and PARP2 on damaged DNA
by way of a poisonous allosteric effect (Murai et al., 2012, 2014a).
Trapped PARP-DNA complexes prevent DNA replication and
transcription, killing cancer cells more effectively than catalytic
inhibition. Not all PARP inhibitors have equivalent PARPtrapping activity. In fact, the capacity to trap PARP varies
significantly among several clinical-stage PARP inhibitors. In
this review, we discuss the evidence for the PARP trapping
concept, the chemical structure-activity relationship for PARP
trapping, and the clinical implications of PARP trapping.
447
448
Shen et al.
wild-type cells are more sensitive to PARP inhibitor cytotoxicity
than PARP12/2 mouse cells (Horton et al., 2005; Kedar et al.,
2012). The effects of PARP1 ablation are not cell type specific, as
it was observed in chicken cells (Murai et al., 2012, 2014a),
mouse cells (Horton et al., 2005; Kedar et al., 2012; Pettitt et al.,
2013), and human cancer cells (Liu et al., 2009; Murai et al.,
2012, 2014a; Pettitt et al., 2013; Das et al., 2014). Although the
PARP1 loss-of-function mutant cell lines also showed resistance
to veliparib, this effect was observed only at very high drug
concentrations, consistent with the relatively weak trapping
effects of veliparib (Murai et al., 2012).
Finally, PARP-DNA complexes have been demonstrated
both in intact cells and in cell-free systems (Kedar et al., 2012;
Murai et al., 2012, 2014a). The presence of PARP inhibitors
significantly stabilizes the association of PARP with DNA in
a reversible manner. Importantly, the potency to trap PARPDNA complexes varies considerably among different PARP
inhibitors. Of the five PARP inhibitors tested, veliparib is
primarily a catalytic inhibitor with little trapping activity,
while olaparib, niraparib, and rucaparib trap PARP ∼100-fold
more efficiently than veliparib. Talazoparib is the most potent
PARP trapper studied to date, promoting PARP-DNA complexes ∼100-fold more efficiently than olaparib, niraparib, and
rucaparib (Murai et al., 2012, 2014a). These data suggest that
the anticancer cytotoxicities of PARP inhibitors do not have
a linear relationship with catalytic inhibition. In contrast, the
cytotoxicities correlate very well with trapping potency.
In summary, current data support a model in which PARP
inhibitors use a dual mechanism to induce cell killing: one by
directly inhibiting PARP catalytic activity and the other by
allosterically affecting the conformational flexibility and dynamics of PARP to enhance its affinity for the SSB repair
intermediate. This second mechanism is often referred to as
PARP trapping or PARP “poisoning.” Trapped PARP-DNA
complexes promote cytotoxicity to a much greater degree than
the unrepaired DNA SSBs that are caused by the loss of PARP
activity, with the PARP inhibitors varying considerably in
their trapping capacities. The clinical efficacy data for PARP
inhibitors in cancer patients with deleterious BRCA gene
mutations are consistent with the in vitro cytotoxicity levels of
the PARP inhibitors, nicely correlating the clinical activity with
the ability to trap PARP. For completeness of this discussion,
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Fig. 1. Structure, molecular weight, and biologic activities of PARP inhibitors undergoing clinical (A) or preclinical (B) development. MW, molecular
weight; PARP1 and PARP2, enzymatic assays; Cell PARylation, an assay that measures poly(ADP-ribose) formation in human LoVo cells; Cytotoxicity,
an assay that measures single-agent cytotoxicity in human Capan-1 cell line that is BRCA2-deficient.
Trapping PARP
we note that an alternative conclusion has been presented at
meetings, but not yet published (Hopkins et al., 2014; Solomon
et al., 2014). These investigators argue that the inhibition of
catalytic activity alone is responsible for the observed PARP
trapping. Further work will be required to resolve the differences in methodology and interpretations that are raised by
this work. In particular, it remains to be established whether
the alternative mechanism can explain both the requirement of
PARP for the cytotoxic effects of the PARP inhibitors and the
large differences in cytotoxicity among the PARP inhibitors with
similar catalytic inhibition activities. At this point, cocrystal
structures that demonstrate allosteric change of the PARP
protein in the presence of the inhibitors are not available. As
described in the following section, intriguing hypotheses
based on the current knowledge of the binding mechanism
between PARP and PARP inhibitors may be postulated.
Comparative analyses of the inhibitor-binding modes may
provide insight into potential structural attributes for the differential levels of PARP-trapping activity reported for the
selected set of inhibitors (Fig. 1). Although other structural
aspects of the PARP superfamily proteins, including inhibitor
selectivity, have been extensively studied and thoroughly reviewed elsewhere (Ferraris, 2010; Ekblad et al., 2013; Papeo
et al., 2013; Steffen et al., 2013), the scope of this review will
be limited to the structural data on PARP1, the most-studied
DNA-dependent enzyme of the PARP superfamily, focusing
on those inhibitors for which PARP-trapping activity has been
evaluated or their closely related analogs for which cocrystal
structures are publicly available for analyses (Fig. 1).
Inhibitor Binding in PARP1 CAT Domain
The CAT domain of PARP1 (Fig. 2A), containing the helical
regulatory subdomain (HD) and the ADP-ribosyltransferase
subdomain (ARTD), has been structurally well characterized,
offering considerable information about ligand-binding interactions. In particular, the highly conserved NAD1 -binding
(“donor”) site in the ARTD (Fig. 2A) has been exploited
extensively in the development of PARP inhibitors, including
those in late-stage clinical development (Fig. 1) (Ferraris,
2010; Ekblad et al., 2013; Papeo et al., 2013; Steffen et al.,
2013). Within the relatively large NAD1-binding site, these
inhibitors exhibit a range of diverse binding modes targeting
distinct binding regions (Fig. 2B).
Nicotinamide-Binding Pocket. All known PARP1 inhibitors (Fig. 1) are anchored in the nicotinamide pocket within
the NAD1-binding site (Fig. 2). The core carboxamide moiety,
shared among these PARP inhibitors, forms critical hydrogen
bonds with conserved Gly863 and Ser904 residues (Fig. 2B)
(Ferraris, 2010; Ekblad et al., 2013; Papeo et al., 2013; Steffen
et al., 2013). These inhibitors also contain an aryl (and sometimes a bicyclic) ring system forming p-stacking interactions
with conserved Tyr896 and Tyr907 residues (Fig. 2B). Other
conserved residues bordering the nicotinamide pocket, such as
pocket-forming Ala898 and Lys903, and a predicted catalytic
residue (Glu988) (Ruf et al., 1998) have also been exploited in
the design of inhibitors (Fig. 2B) (Ferraris, 2010; Ekblad et al.,
2013; Papeo et al., 2013; Steffen et al., 2013). Veliparib, the
smallest PARP inhibitor in clinical development, binds within
the NAD1-binding site primarily through critical interactions
with or near the nicotinamide-binding residues. The carboxamide
benzimidazole core maintains canonical interactions with conserved Gly863, Ser904, and Tyr907 residues in the nicotinamide pocket (Fig. 2B) (PDB ID: 2RD6) (Penning et al., 2009),
while the benzimidazole nitrogen forms a water-mediated
hydrogen bond with Glu988 (Ruf et al., 1998). The comparable binding mode (PDB ID: 3GN7) and enzymatic potency
reported for its enantiomer (Penning et al., 2009) suggest that
interactions of the carboxamide-benzimidazole scaffold at the
nicotinamide subsite (Fig. 2) play a significant role in overall
binding potency.
Adenine-Ribose–Binding Pocket. Located immediately
adjacent to the nicotinamide pocket of PARP is a large hydrophobic pocket (known as the adenine-ribose–binding site),
representing another potential druggable subsite (Fig. 2A).
Many known PARP1 inhibitors leverage the presence of this
adenine-ribose subsite by incorporating a large hydrophobic
group onto the core nicotinamide pharmacophore (Ferraris,
2010; Ekblad et al., 2013; Papeo et al., 2013; Steffen et al., 2013).
An early preclinical PARP1 inhibitor, FR257517 (IC50, 14 nM)
(Fig. 1B), binds to the nicotinamide subsite via a quinazolinone
moiety, while the remaining part of the inhibitor (including the
dihydropyridinyl and fluorophenyl moieties) reaches into the
adenine-ribose–binding pocket, creating a new hydrophobic
subsite with Leu769, Arg878, Ile879, and Pro881 residues
(Fig. 2B) (PDB ID: 1UK0) (Kinoshita et al., 2004). Several other
preclinical inhibitors, for which structural data are available,
also exhibit a similar binding mode that involves secondary
contacts in the adenine-ribose–binding subsite (Hattori et al.,
2004; Kinoshita et al., 2004; Miyashiro et al., 2009; Gangloff
et al., 2013; Ye et al., 2013; Patel et al., 2014). From the
chemical scaffold and binding mode reported in the tankyrase 2
cocrystal structure (Narwal et al., 2012), olaparib, while bound
to the nicotinamide pocket via a phthalazinone core, also likely
has both diacylpiperazine and cyclopropyl moieties reaching
deeply into the adenine-ribose–binding pocket.
Phosphate-Binding Residues. A phosphate-binding region formed by HD residues at the NAD1 site (Fig. 2A) (Ruf
et al., 1998) has also been explored for novel inhibitor-scaffold
designs. According to a publicly available cocrystal structure
(PDB ID: 3L3M), the compound referred to as A927929 (IC50,
6 nM) (Fig. 1B) binds the CAT domain by interacting with the
Asp766 residue, which is predicted to be involved in phosphate
binding, in addition to key interactions with nicotinamidebinding Gly863 and Ser904 residues (Fig. 2B) (Penning et al.,
2010). Recent studies also report crystal structure data (PDB
ID: 2OQA; compound 59) that describe a similar inhibitorbinding pose directed toward the phosphate-binding region
(Patel et al., 2014). Based on the chemical scaffolds in the
absence of PARP1 cocrystallographic structure data, rucaparib
and niraparib may also be directed toward the phosphatebinding residues within the NAD1 site. Compound A906894
(Fig. 1B) also projects toward the phosphate-binding residues
(PDB ID: 3L3L). However, the relatively small size of this
compound may limit a direct contact with the HD phosphatebinding residues (Gandhi et al., 2010). Instead, this inhibitor,
via its piperidine nitrogen, forms a hydrogen bond with Gly888
located on a donor-site loop (D-loop: Gly876–Gly894) that,
together with the N-terminal HD, borders the NAD1-binding site (Fig. 2A).
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Current View of PARP-Trapping StructureActivity Relationship
449
Shen et al.
450
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Trapping PARP
Structural Basis for PARP-DNA Trapping
DNA-Dependent PARP1 Interdomain Communication.
The distinct binding modes in the CAT domain observed among
the PARP1 inhibitors (Fig. 2B) combined with newly available
structural data on noncatalytic domains (Langelier et al., 2008,
2011, 2012; Loeffler et al., 2011; Ali et al., 2012; Hassler and
Ladurner, 2012) may suggest key structural insights into the
DNA-trapping activity of PARP inhibitors. The crystal structure
of the “near” full-length PARP1 enzyme containing 1) zincfinger domains Zn1 and Zn3, 2) WGR domain named after
a conserved Trp-Gly-Arg sequence, and 3) CAT domain consisting of HD-ARTD subdomains bound to a DNA DSB (PDB ID:
4DQY) demonstrates that DNA binding by Zn1, Zn3, and WGR
domains destabilizes the CAT domain, thereby activating the
PARP enzymatic activity (Langelier et al., 2012). Specifically,
structural changes upon DNA binding in the noncatalytic
domains lead to displacements of Leu698 and Leu701 residues
from the hydrophobic core of the HD (Fig. 3A), decreasing the
stability of ARTD in the CAT domain; an increase in the
protein dynamics of the ARTD is suggested to stimulate PARP
catalysis (Langelier et al., 2012). The proposed mechanism is
consistent with a previously identified gain-of-function mutation at Leu713 within the HD hydrophobic core that leads to
DNA-independent PARP1 activity (Fig. 3A) (Miranda et al.,
1995). The importance of interdomain communication in DNAdependent activation is also demonstrated by Trp318 mutations at the HD-WGR-Zn3 domain interfaces, resulting in
a catalytically inactive PARP1 without affecting DNA binding
(Steffen et al., 2014). Notably, the HD subdomain, implicated
as a focal point for interdomain communication, is an important CAT domain structural element defining the size of the
inhibitor-binding pocket, and sometimes directly interacting
with bound inhibitors (Iwashita et al., 2005; Penning et al.,
2010).
Inhibitor-Dependent PARP1 Interdomain Communication. An improved understanding of the structural basis for
DNA-dependent PARP1 activation (Hassler and Ladurner,
2012; Langelier and Pascal, 2013) has led to a new hypothesis
that inhibitor-induced reverse-allosteric signaling may drive
PARP-trapping activity (Murai et al., 2012). Using a fluorescence anisotropy-based DNA-binding assay, 10- to 40-fold
differences in the biochemical trapping potency have been
reported among the PARP inhibitors with comparable potencies in inhibiting catalysis, further suggesting possible
inhibitor-specific allosteric effects (Murai et al., 2012, 2014a).
The differential binding modes among the inhibitors may
explain why those equally potent PARP1 catalytic inhibitors
exhibit significant differences in their abilities to trap PARP
on DNA (Murai et al., 2012, 2014a). As discussed earlier and
further elaborated on in the following section, a substantial
body of structural data indicates that PARP inhibitors differ
considerably in overall size/scaffold (Fig. 1) and consequently
in binding modes (Fig. 2B).
The size or bulk of a PARP inhibitor may be correlated with
the level of reverse-allosteric signaling, which may be related
to the degree of PARP trapping (Marchand et al., 2014). Among
the five clinical inhibitors evaluated for PARP-trapping ability
(Fig. 1A), the smallest inhibitor, veliparib (mol. wt., 244), was
the least potent in trapping PARP relative to the bulkier PARP
inhibitors, such as olaparib (mol. wt., 434), niraparib (mol. wt.,
320), rucaparib (mol. wt., 323), and talazoparib (mol. wt., 380)
(Murai et al., 2012, 2014a). The low trapping activity of
veliparib correlated with the absence of inhibitor-induced
PARP1 conformational changes during microsecond molecular
dynamic simulations (Marchand et al., 2014). Under the same
experimental conditions, the binding of comparatively bulkier
olaparib to the CAT domain caused steric bumps on the HD
subdomain, triggering conformational rearrangements in the
DNA-binding domains to stabilize the PARP1-DNA complex
(Marchand et al., 2014). A lack of inhibitor-induced steric
Fig. 2. Inhibitor binding to PARP1 CAT domain. (A) PARP1 inhibitors bind the NAD+ cosubstrate-binding site on the C-terminal CAT domain of the
multidomain PARP1 protein. Crystal structures of the near full-length PARP1 in complex with DNA have been determined (PDB ID: 4DQY) (Langelier
et al., 2012). Based on a homologous structure of diphtheria toxin (PDB ID: 1TOX) (Bell and Eisenberg, 1996), a NAD+ molecule was modeled in the
PARP1 substrate-binding site, which consists of multiple druggable subpockets. (B) PARP1 inhibitors are targeted to distinct binding regions within the
relatively large NAD+-binding site: (1) nicotinamide-binding pocket (Penning et al., 2009), (2) adenine-ribose–binding pocket (Kinoshita et al., 2004), (3)
phosphate-binding region (Penning et al., 2010), and (4) outer borders of the NAD+ site (Aoyagi-Scharber et al., 2014). Inhibitor-binding interactions in
each subsite are illustrated by molecular surface (left) and stick model (right) representations. MOE (Molecular Operating Environment, Version
2014.09; Chemical Computing Group Inc., Montreal, QC, Canada) and PyMOL (The PyMOL Molecular Graphics System, Version 1.7.4; Schrödinger,
LLC., New York, NY) were used for structural analyses and figure generation.
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Outer Borders of the NAD1 Site. The majority of known
PARP1 inhibitors target the nicotinamide pocket and one of the
two other (adenine-ribose or phosphate) subsites within the
relatively large NAD1 cosubstrate-binding site (Fig. 2, A and B)
(Wiholm and Myrhed, 1993; Ekblad et al., 2013; Papeo et al.,
2013; Steffen et al., 2013). However, based on recent cocrystal
structural data, talazoparib (PDB ID: 4PJT) (Fig. 2B) adds
another dimension to the already diverse range of inhibitorbinding modes (Aoyagi-Scharber et al., 2014). Talazoparib represents a new tripartite chemical scaffold consisting of a core
tricyclic group with stereospecific disubstituents (Fig. 1A)
(Shen et al., 2013). While the cyclic amide moiety is tethered
to the nicotinamide subsite and the unsaturated six-membered
ring system forms a water-mediated interaction with the catalytic
residue, Glu988 (Ruf et al., 1998), the 8S-(4-fluorophenyl) and
9R-methyltriazole moieties are directed toward the outer edges
of the NAD1-binding pocket (Fig. 2B) (Aoyagi-Scharber et al.,
2014). The first substituent, the 4-fluorophenyl group, reaches
out to the unique ligand-binding space near the D-loop
(Gly876–Gly894), which, as mentioned above, forms the lid of
the NAD1-binding site (Fig. 2B). Two D-loop residues, Tyr889
and Met890, interact with this fluorophenyl group by forming
p-stacking and water-mediated hydrogen-p/hydrogenbonding interactions, respectively (Fig. 2B). The second
substituent, the methyltriazole group, is involved in a watermediated hydrogen-bonding interaction to the backbone amide
of the highly conserved aromatic Tyr896 residue within the
nicotinamide pocket (Fig. 2B). The methyl group of the triazole
moiety, when bound to PARP1, points toward the HD, which
not only defines the size of the NAD1-binding site within the
CAT domain, but also has critical regulatory roles in the catalytic function of PARP1.
451
452
Shen et al.
bumps in the HD subdomain by small inhibitors can be explained by the limited extent of the binding interactions on the
CAT domain. The methylpyrrolidine moiety of veliparib appears to be too small to make significant contacts with the rest
of the NAD1 site (Fig. 2B) (Penning et al., 2009), including the
HD subdomain implicated as a crucial structural element in
the proposed reverse-allosteric mechanism (Marchand et al.,
2014). HYDAMTIQ (mol. wt., 274) (Fig. 1B), another potent
inhibitor similar to veliparib in size and binding mode, also
shows low levels of PARP trapping and cytotoxicity, further
supporting a potential relationship between the molecular size
of the inhibitors and the degree of trapping activity (Pellicciari
et al., 2011; Marchand et al., 2014).
The molecular size of the inhibitor alone, however, seems
insufficient to describe the differential trapping of PARP1 reported for PARP inhibitors. Among the bulky PARP1 inhibitors, no correlation can be firmly established between inhibitor
size and degree of PARP trapping. For instance, talazoparib
has been shown to be the most potent PARP-trapping inhibitor
tested to date, despite its small size relative to olaparib (Fig.
1A) (Murai et al., 2014a). Recent studies suggest that the shape
and flexibility of inhibitors may contribute to the efficiency of
trapping PARP-DNA complexes (Murai et al., 2014a). Compared with other bulky yet flexible inhibitors, talazoparib has
unique tripartite chemical scaffolds that are rigid and stereospecific (Fig. 1A) (Shen et al., 2013; Aoyagi-Scharber et al.,
2014). The shape and flexibility of the inhibitor dictate the
extent and type of binding interactions, which vary significantly among these clinical inhibitors and may explain marked
differences in PARP-trapping activities.
Of the five clinical-stage PARP inhibitors tested (Fig. 1),
those inhibitors with noticeable ability to stabilize the PARPDNA complexes appear to interact with the D-loop, though to
differing extents, at the outer border of the NAD1 site (Fig. 2, A
and B). Using a structural analogy with A927929 (Penning
et al., 2010), the potent PARP trappers rucaparib and niraparib
(Fig. 1A) are likely bound close to the D-loop (Fig. 2B). Interestingly, a recent molecular dynamics simulation study
reported an alternative bound conformation of olaparib,
another PARP trapper, forming a hydrogen bond with the
D-loop residue Met890 (Marchand et al., 2014). Talazoparib,
the most potent PARP trapper evaluated to date, has a unique
stereospecific 4-fluorophenyl substituent favorably oriented to
interact with the Met890 backbone via a water molecule and
the Tyr889 side chain on the D-loop (Figs. 2B and 3B). The
degree of PARP-trapping capacity may be associated with the
extent to which the inhibitor interacts with the D-loop residues.
The D-loop is structurally linked to the HD subdomain (Fig.
3B), suggesting that the inhibitor interactions with the D-loop
residues may influence conformational flexibility of the CAT
domain. The PARP1 D-loop has been described as a relatively
rigid structure, unlike the flexible counterpart in other PARP
superfamily proteins (Wahlberg et al., 2012). However, recently
determined PARP1 inhibitor cocrystal structures (PDB IDs:
4HHY, 4HHZ, and 4GV7) (Lindgren et al., 2013; Ye et al., 2013)
reveal that the Tyr889 side chain on the D-loop can adopt
multiple rotameric positions among noncrystallographic
symmetry-related molecules (Fig. 3C). When bound to
talazoparib, these alternative side-chain conformations would
not be permitted due to steric constraints imposed by the
4-fluorophenyl substituent (Figs. 2B and 3C). Additional studies
are required to elucidate the structural consequences of the
decreased side-chain flexibility of the D-loop residues, such as
Tyr889, on the conformational stability of the HD or entire CAT
domain and subsequent interdomain communication that has
been proposed to be critical for the allosteric signaling of PARP1
(Langelier et al., 2012).
Newly available structural data on noncatalytic domains
(Hassler and Ladurner, 2012; Langelier and Pascal, 2013),
combined with a wealth of cocrystal structure data available
for the inhibitor-bound PARP1 CAT domain (Ferraris, 2010;
Ekblad et al., 2013; Papeo et al., 2013; Steffen et al., 2013),
signify an important first step toward elucidating the molecular basis for the differential PARP trapping among the
inhibitors (Murai et al., 2012, 2014a). The D-loop, known for
its involvement in recognizing the acceptor adenine moiety
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Fig. 3. The N-terminal HD and D-loop implicated in both inhibitor binding and interdomain communications of PARP1. (A) Mutations of the HD core
hydrophobic Leu residues increase DNA-independent activity, suggesting that the HD distortion is critical for DNA-dependent PARP activation (PDB
IDs: 4DQY and 1TOX) (Bell and Eisenberg, 1996; Langelier et al., 2012). (B) Talazoparib forms unique interactions with residues on the D-loop, which is
structurally linked to the HD domain, implicated in allosteric signaling (PDB ID: 4PJT) (Aoyagi-Scharber et al., 2014). Dotted lines represent interresidue contacts formed across the HD and D-loop interface. (C) Superpositions of the 33 PARP1 CAT domain structures from PDB indicate that the
D-loop residue, Tyr889, can assume multiple side-chain conformations. Binding of a rigid stereospecific inhibitor (e.g., talazoparib) may sterically
restrict the side-chain flexibility (PDB IDs: 4PJT, 4GV7, 4HHY, 4HHZ, 4L6S, 4DQY, 3GN7, 3L3L, 3L3M, 3GJW, 2RD6, 1WOK, 1UK0, and 1UK1)
(Hattori et al., 2004; Kinoshita et al., 2004; Iwashita et al., 2005; Miyashiro et al., 2009; Gandhi et al., 2010; Penning et al., 2010; Langelier et al., 2012;
Gangloff et al., 2013; Lindgren et al., 2013; Ye et al., 2013; Aoyagi-Scharber et al., 2014).
Trapping PARP
(Ruf et al., 1998) and well characterized as a sequence-diverse
element in the PARP superfamily (Ekblad et al., 2013; Papeo
et al., 2013; Steffen et al., 2013), may also contribute to the
protein dynamics of the CAT domain that are implicated in
the putative inhibitor-induced reverse-allosteric signaling.
Further investigations of the interdomain interactions and
allostery of the full-length or near full-length enzyme using
diverse biophysical techniques, such as small-angle X-ray
scattering, molecular dynamics simulations, and X-ray crystallography (Mansoorabadi et al., 2014; Marchand et al., 2014;
Patel et al., 2014), may increase our understanding of the
PARP-trapping mechanisms, providing new avenues to develop improved anticancer therapeutic agents and treatment
approaches.
Clinical Implications
2009, 2012). However, as noted in a review by Pommier (2013),
“there is no simple linear correlation between topoisomerase
levels and drug response.” Other factors that influence the
sensitivity to Top inhibitors, such as the protein kinase TAK1
and SLFN11, have been identified via two main approaches: 1)
identification of alterations in the DNA repair and stress
response of cells to Top inhibitors, and 2) genomic analyses of
cancer cells (e.g., siRNA screens and gene expression correlations) (Pommier, 2013). Similar approaches may help identify
biomarkers associated with the response to PARP inhibitors.
Can PARP Expression Level and PARP Activity
Serve as Biomarkers for Sensitivity to PARP Inhibitors? HR deficiency has emerged as the key determinant of
the cancer cell response to PARP inhibitors; other factors affecting the sensitivity to PARP inhibitors (e.g., P-glycoprotein
overexpression and loss of 53BP1) have also been described
(Lord and Ashworth, 2012; Bouwman and Jonkers, 2014). In
addition to these mechanisms, an intriguing question that derives from the PARP trapping concept is whether the tumor
response to the PARP inhibitors may be associated with the
PARP expression level and/or PARP activity in the cells. Though
inconclusive at present, several lines of evidence appear to be
consistent with this hypothesis, as discussed below.
A study by Pettitt et al. (2013) has shown that either complete
loss or reduction of PARP1 expression is associated with
acquired resistance to PARP inhibitors. Cells with complete
loss of PARP1 expression are ∼100-fold more resistant to
olaparib and talazoparib than the parental cell line, while
reduction of PARP1 expression by siRNA treatment is associated with partial resistances to the PARP inhibitors (Pettitt
et al., 2013). Similarly, Singh et al. (2013) reported that
a common characteristic in several Ewing’s sarcoma cell lines
that have developed resistance to talazoparib (500- to 1000-foldhigher IC50 as compared with the parental cell lines) is the loss
or reduction of PARP1 expression. In addition, human cancer
cell lines that have acquired resistance to a combined treatment
of veliparib plus TMZ also show greatly reduced PARP1 levels
(Liu et al., 2009). Michels and colleagues analyzed several
cisplatin-resistant cancer cell lines and found that a subset of
these lines overexpress PARP1 and display hyperactivity of
PARylation. Furthermore, a positive correlation was observed
between the cellular PARylation and the sensitivity to PARP
inhibitors (Michels et al., 2013). It should be noted that clinical
experience thus far has shown that platinum-sensitive tumors
are more likely to respond to PARP inhibitors than are platinumresistant or platinum-refractory diseases (Fong et al., 2010), so
the clinical prospect of this observation remains to be evaluated.
Nonetheless, this study lends additional support to the idea that
cellular PARylation activity may be an important factor that
influences the sensitivity to PARP inhibitors.
HR-defective cells may manifest hyper-PARylation activity.
Disruption of BRCA2 and other HR genes leads to PARP hyperactivation, which correlates with an increased sensitivity to
PARP inhibitors (Gottipati et al., 2010). Restoring BRCA2
activity (either by genetic reversion of the BRCA2 gene or by
stable expression of a wild-type BRCA2 gene) led to reduced
PARP activity and corresponding resistance to PARP inhibitors
(Gottipati et al., 2010). It appears that cells with DSB repair
deficiency may become highly dependent on PARP-mediated
DNA repair, raising the possibility that PARylation activity
may serve as a surrogate biomarker for DSB repair deficiency
and a relevant predictor of response to PARP inhibitors.
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Understanding the mechanisms of action of PARP inhibitors may assist physicians in selecting the most appropriate
drug for treating cancer. The effectiveness in PARP trapping
may be an important factor to consider when formulating
a therapeutic regimen involving a PARP inhibitor (singleagent or in combination). The PARP trapping concept may
also shed new light on how to identify patients who will (or
will not) be likely to respond to PARP inhibitor therapies.
Lessons from Topoisomerase Inhibitors. Top inhibitors may provide useful insights into the development of PARP
inhibitors as anticancer agents. Inhibitors of Top1 [e.g., camptothecin (CPT), topotecan, and irinotecan] and Top2 (e.g., doxorubicin and etoposide) are among the most effective and
commonly used anticancer drugs. These drugs kill cancer cells
by trapping their respective protein targets (Top1 or Top2) on
DNA rather than by classic enzymatic inhibition (Pommier
et al., 2010; Pommier, 2013). Depletion of Top1 causes cells to
become resistant to CPT (Eng et al., 1988; Nitiss and Wang,
1988; Pommier et al., 1999; Miao et al., 2007), and reductions
in Top1 and Top2 levels in tumors diminishes the effects of the
Top inhibitors (Rasheed and Rubin, 2003; Beretta et al., 2006;
Burgess et al., 2008), highlighting the Top requirements in
the cytotoxic action of these drugs. X-ray crystallographic
studies have revealed the ternary complexes of Top1-DNACPT (PDB ID: 1T8I) and Top2-DNA-etoposide (PDB ID:
3QX3), in which the drugs simultaneously interact with both
the DNA and the enzymes (Redinbo et al., 1998; Ioanoviciu
et al., 2005; Wendorff et al., 2012), providing direct evidence
for the Top-trapping model (Pommier and Marchand, 2011).
Biochemically, the formation of Top1 cleavage complex and Top2
cleavage complex has been demonstrated in cells treated with
Top1 and Top2 inhibitors, respectively. Thus, Top inhibitors
trap Tops to DNA much like PARP inhibitors trap PARP1/2
(Pommier et al., 1999; Pommier, 2013). Despite the analogous
interactions between the Top inhibitors and Top1/2 and between
the PARP inhibitors and PARP1/2, there are important differences. Top inhibitors directly target the DNA-protein interface
and form covalent bonds with Top1 or Top2, whereas PARP
inhibitors bind noncovalently with PARP1/2 at the catalytic
active site away from the associated DNA (Patel et al., 2012).
Because the cytotoxicity of Top inhibitors appears to be
positively correlated with the expression levels and activities of
Top1 and Top2 (Burgess et al., 2008; O’Malley et al., 2009;
Pfister et al., 2009), assays are being developed to measure the
expression levels and activities of Top1 and Top2 (Pfister et al.,
453
454
Shen et al.
examples, see Delaney et al. (2000) and Calabrese et al. (2004)],
even before the PARP-trapping mechanism was elucidated. In
fact, the first clinical trial of a PARP inhibitor was a combination
study of rucaparib and TMZ (Plummer et al., 2013). An
improved understanding of the PARP-trapping mechanism
leads to completely novel combination strategies; rather than
PARP inhibitors acting as the potentiator of the TMZ
cytotoxicity, PARP inhibitors should be considered to be the
cytotoxic agents by inducing PARP-DNA–trapping complexes,
and TMZ as a potentiator that induces DNA lesions. As the
creation of the N7mG and N3mA adducts is efficient (Sarkaria
et al., 2008), TMZ may be effective at reduced dosages (to
minimize the toxicity), allowing for maximal antitumor efficacy
by dosing PARP inhibitors at levels approaching their maximum
tolerated doses. We envision that distinctive mechanisms of
sensitivity/resistance will exist for the combination of PARP
inhibitors plus TMZ, as compared with TMZ immunotherapies.
The traditional mechanisms of TMZ resistance related to
O6-methylguanine adducts may not be relevant, as cancer cells
expressing O6-methylguanine methyltransferase and those
having defective mismatch repair maintain their sensitivity
to PARP inhibitors plus TMZ. It should be noted that the
concept at the present time is applicable only to combinations
of PARP inhibitors with TMZ (and other DNA alkylating
agents). Whether this combination strategy can apply to other
DNA-damaging agents has yet to be evaluated. This novel
combination approach with DNA alkylating agents is currently
being tested both preclinically (Feng et al., 2014; von Euw et al.,
2014; Smith et al., 2015) and in the clinic (for examples, see
NCT02116777 and NCT02049593 at www.clinicaltrials.gov).
In the second category, synergistic combinations that do not
use PARP trapping but require PARP catalytic inhibition are
considered (Murai et al., 2014b). Combination with the Top1
inhibitor CPT is an example of this category. Olaparib and
veliparib sensitize tumor cells to CPT to a similar degree, and
PARP depletion itself enhances the cytotoxic effect of CPT,
which is not further augmented by olaparib or veliparib (Murai
et al., 2014b). These results suggest that the synergistic effect of
CPT with PARP inhibitors is due to PARP catalytic inhibition
rather than PARP trapping (Murai et al., 2014b). This synergy
is consistent with recent findings demonstrating that PARP is
coupled with tyrosyl-DNA phosphodiesterase-1 (Das et al.,
2014), and with X-ray repair cross-complementing protein 1,
polynucleotide kinase, and ligase III, which repairs the DNA
lesions caused by Top1 inhibitors (Pommier et al., 2006; Das
et al., 2014). Based on this mechanism, PARP inhibitors may
preferentially potentiate Top1 inhibitors either in tyrosyl-DNA
phosphodiesterase-1–deficient cells or in cells harboring ERCC1
mutations (Pommier et al., 2010; Zhang et al., 2011; Pommier,
2013).
In the third category, combinations of additive but mutually
independent mechanisms are considered (Murai et al.,
2014b). Examples for this category include Top2 inhibitors
(e.g., etoposide) and cross-linking agents (e.g., cisplatin). The
rationale for combining PARP inhibitors with these agents is
not based on either PARP-DNA trapping or PARP catalytic
inhibition. Therefore, additive as opposed to synergistic effects may be expected when PARP inhibitors are combined
with these agents. Cisplatin and etoposide cause DNA lesions
whose repair does not involve PARP-mediated pathways
(Bowman et al., 2001; Zhang et al., 2011; Pommier, 2013),
whereas PARP inhibitors kill cells by acting on DNA lesions
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
It should be emphasized that PARP expression level/PARP
activity is unlikely to be the sole determinant of the tumor
response to PARP inhibitors. To date, sensitivity to single-agent
PARP inhibitors in the clinical setting appears to be closely tied
to HR defects, whereas PARP expression/activity may just be
one of the additional factors that influence tumor response. As
an example of the complexity of relating PARP levels to sensitivity to PARP inhibitors, a recent report shows a relationship
between increased PARP expression in HCC1937-derived stem
cells and resistance to olaparib (Gilabert et al., 2014). However,
given that the HCC1937 cell line is highly resistant to olaparib
despite its BRCA1 mutation status (Lehmann et al., 2011)
the interpretation of this result is complicated. In any case,
testing the possible link between PARP expression/activity
and sensitivity to PARP inhibitors in the clinical setting is
warranted.
Designing Drug Combination Strategies for PARP
Inhibitors with Genotoxic Agents. Combinations of PARP
inhibitors with DNA-damaging agents are undergoing active
clinical investigations (Curtin and Szabo, 2013; Murai et al.,
2014b). As PARP inhibitors are not equal in their ability to trap
PARP, careful selection of the appropriate PARP inhibitor(s)
for each combination therapy is critical for maximizing the
chance of clinical success. The choice of PARP inhibitor should
be based not only on whether PARP is involved in the repair of
the DNA lesions produced by the genotoxic agent, but also on
whether the lesions trap PARP-DNA complexes. According to
a recent publication (Murai et al., 2014b), three categories of
combination strategies for use with PARP inhibitors and genotoxic agents are envisioned.
In the first category, highly synergistic combinations taking
advantage of PARP trapping are considered (Murai et al.,
2014b). This is exemplified by the combination of PARP inhibitors with the alkylating agent TMZ. TMZ induces two
types of DNA lesions: 1) O6-methylguanine adducts, which
cause DNA DSB and apoptosis—cells with defective mismatch repair pathway or cells with functional O6-methylguanine methyltransferase are resistant to this mechanism; and 2)
N7-methylguanine (N7mG) and N3-methyladenine (N3mA)
DNA adducts, which account for ∼90% of all methylation
events caused by TMZ but are nonlethal in most normal
cells and tumor cells, because they are readily repaired by
BER (Sarkaria et al., 2008; Zhang et al., 2012). The repair of
N7mG and N3mA adducts by BER requires PARP to recognize
and bind the repair intermediate 59-deoxyribose phosphate
(Cistulli et al., 2004; Horton and Wilson, 2013a,b; Horton et al.,
2014). In fact, 59-deoxyribose phosphate is a preferred substrate for PARP trapping (Murai et al., 2014b; Smith et al.,
2015). Thus, PARP inhibitors turn the nonlethal N7mG and
N3mA adducts into cytotoxic lesions by trapping PARP at
59-deoxyribose phosphate. This ability to potentiate TMZ cytotoxicity varies greatly among the various PARP inhibitors and
aligns well with the PARP-trapping capacity of these molecules
(Shen et al., 2013; Murai et al., 2014b). Optimal potentiation of
TMZ by olaparib requires PARP. In PARP-depleted cells,
olaparib and veliparib showed similar (weak) potentiation of
TMZ (Murai et al., 2014b). These data indicate that combining
TMZ with a potent PARP-trapping agent is a more rational
approach than combining TMZ with a PARP inhibitor that
primarily acts by catalytic inhibition.
The potent combination effects with PARP inhibitors and
TMZ have been demonstrated in numerous studies [for
Trapping PARP
Acknowledgments
The authors thank Dr. L. E. Post (BioMarin Pharmaceutical Inc.) for
guiding this review and providing valuable suggestions and to
Drs. Y. Pommier (National Cancer Institute, Bethesda, MD) and J. Murai
(Kyoto University, Kyoto, Japan) for reviewing the manuscript and
sharing their expert insights. The authors thank E. Johnson for editing
the manuscript and P. Fitzpatrick, G. Vehar, R. Srivastava, K. Krishnan,
M. Henderson, E. Wang, and K. Yu (BioMarin Pharmaceutical Inc.) for
their suggestions, all of which markedly improved the manuscript. The
authors also appreciate the professional and competent assistance from
ApotheCom in the preparation of the manuscript.
Authorship Contributions
Performed data analysis: Shen, Aoyagi-Scharber, Wang.
Wrote or contributed to the writing of the manuscript: Shen, AoyagiScharber, Wang.
References
Ali AA, Timinszky G, Arribas-Bosacoma R, Kozlowski M, Hassa PO, Hassler M,
Ladurner AG, Pearl LH, and Oliver AW (2012) The zinc-finger domains of PARP1
cooperate to recognize DNA strand breaks. Nat Struct Mol Biol 19:685–692.
Amé JC, Rolli V, Schreiber V, Niedergang C, Apiou F, Decker P, Muller S, Höger T,
Ménissier-de Murcia J, and de Murcia G (1999) PARP-2, a novel mammalian DNA
damage-dependent poly(ADP-ribose) polymerase. J Biol Chem 274:17860–17868.
Aoyagi-Scharber M, Gardberg AS, Yip BK, Wang B, Shen Y, and Fitzpatrick PA
(2014) Structural basis for the inhibition of poly(ADP-ribose) polymerases 1 and 2
by BMN 673, a potent inhibitor derived from dihydropyridophthalazinone. Acta
Crystallogr F Struct Biol Commun 70:1143–1149.
Ashworth A (2008) A synthetic lethal therapeutic approach: poly(ADP) ribose polymerase inhibitors for the treatment of cancers deficient in DNA double-strand
break repair. J Clin Oncol 26:3785–3790.
Audebert M, Salles B, and Calsou P (2004) Involvement of poly(ADP-ribose)
polymerase-1 and XRCC1/DNA ligase III in an alternative route for DNA
double-strand breaks rejoining. J Biol Chem 279:55117–55126.
Bell CE and Eisenberg D (1996) Crystal structure of diphtheria toxin bound to nicotinamide adenine dinucleotide. Biochemistry 35:1137–1149.
Benjamin RC and Gill DM (1980) Poly(ADP-ribose) synthesis in vitro programmed by
damaged DNA. A comparison of DNA molecules containing different types of
strand breaks. J Biol Chem 255:10502–10508.
Beretta GL, Perego P, and Zunino F (2006) Mechanisms of cellular resistance to
camptothecins. Curr Med Chem 13:3291–3305.
Booth L, Cruickshanks N, Ridder T, Dai Y, Grant S, and Dent P (2013) PARP and
CHK inhibitors interact to cause DNA damage and cell death in mammary carcinoma cells. Cancer Biol Ther 14:458–465.
Bouwman P and Jonkers J (2014) Molecular pathways: how can BRCA-mutated
tumors become resistant to PARP inhibitors? Clin Cancer Res 20:540–547.
Bowman KJ, Newell DR, Calvert AH, and Curtin NJ (2001) Differential effects of the
poly (ADP-ribose) polymerase (PARP) inhibitor NU1025 on topoisomerase I and II
inhibitor cytotoxicity in L1210 cells in vitro. Br J Cancer 84:106–112.
Bryant HE and Helleday T (2006) Inhibition of poly (ADP-ribose) polymerase activates ATM which is required for subsequent homologous recombination repair.
Nucleic Acids Res 34:1685–1691.
Bryant HE, Schultz N, Thomas HD, Parker KM, Flower D, Lopez E, Kyle S, Meuth
M, Curtin NJ, and Helleday T (2005) Specific killing of BRCA2-deficient tumours
with inhibitors of poly(ADP-ribose) polymerase. Nature 434:913–917.
Burgess DJ, Doles J, Zender L, Xue W, Ma B, McCombie WR, Hannon GJ, Lowe SW,
and Hemann MT (2008) Topoisomerase levels determine chemotherapy response
in vitro and in vivo. Proc Natl Acad Sci USA 105:9053–9058.
Calabrese CR, Almassy R, Barton S, Batey MA, Calvert AH, Canan-Koch S, Durkacz
BW, Hostomsky Z, Kumpf RA, Kyle S, et al. (2004) Anticancer chemosensitization
and radiosensitization by the novel poly(ADP-ribose) polymerase-1 inhibitor
AG14361. J Natl Cancer Inst 96:56–67.
Cardnell RJ, Feng Y, Diao L, Fan YH, Masrorpour F, Wang J, Shen Y, Mills GB,
Minna JD, Heymach JV, et al. (2013) Proteomic markers of DNA repair and PI3K
pathway activation predict response to the PARP inhibitor BMN 673 in small cell
lung cancer. Clin Cancer Res 19:6322–6328.
Cistulli C, Lavrik OI, Prasad R, Hou E, and Wilson SH (2004) AP endonuclease and
poly(ADP-ribose) polymerase-1 interact with the same base excision repair intermediate. DNA Repair (Amst) 3:581–591.
Curtin NJ (2012) DNA repair dysregulation from cancer driver to therapeutic target.
Nat Rev Cancer 12:801–817.
Curtin NJ and Szabo C (2013) Therapeutic applications of PARP inhibitors: anticancer therapy and beyond. Mol Aspects Med 34:1217–1256.
Das BB, Huang SY, Murai J, Rehman I, Amé JC, Sengupta S, Das SK, Majumdar P,
Zhang H, Biard D, et al. (2014) PARP1-TDP1 coupling for the repair of topoisomerase
I-induced DNA damage. Nucleic Acids Res 42:4435–4449.
Delaney CA, Wang LZ, Kyle S, White AW, Calvert AH, Curtin NJ, Durkacz BW,
Hostomsky Z, and Newell DR (2000) Potentiation of temozolomide and topotecan
growth inhibition and cytotoxicity by novel poly(adenosine diphosphoribose) polymerase inhibitors in a panel of human tumor cell lines. Clin Cancer Res 6:
2860–2867.
Durkacz BW, Omidiji O, Gray DA, and Shall S (1980) (ADP-ribose)n participates in
DNA excision repair. Nature 283:593–596.
Ekblad T, Camaioni E, Schüler H, and Macchiarulo A (2013) PARP inhibitors:
polypharmacology versus selective inhibition. FEBS J 280:3563–3575.
Eng WK, Faucette L, Johnson RK, and Sternglanz R (1988) Evidence that DNA
topoisomerase I is necessary for the cytotoxic effects of camptothecin. Mol Pharmacol 34:755–760.
Farmer H, McCabe N, Lord CJ, Tutt AN, Johnson DA, Richardson TB, Santarosa M,
Dillon KJ, Hickson I, Knights C, et al. (2005) Targeting the DNA repair defect in
BRCA mutant cells as a therapeutic strategy. Nature 434:917–921.
Feng Y, Shen Y, and Wang E (2014) Talazoparib (BMN-673) as single agent and in
combination with temozolomide or PI3K pathway inhibitors in small cell lung
cancer and gastric cancer models. 26th EORTC-NCI-AACR Symposium on Molecular Targets and Cancer Therapeutics; 2014 Nov 18–21; Barcelona, Spain.
Ferraris DV (2010) Evolution of poly(ADP-ribose) polymerase-1 (PARP-1) inhibitors.
From concept to clinic. J Med Chem 53:4561–4584.
Fong PC, Yap TA, Boss DS, Carden CP, Mergui-Roelvink M, Gourley C, De Greve J,
Lubinski J, Shanley S, Messiou C, et al. (2010) Poly(ADP)-ribose polymerase inhibition: frequent durable responses in BRCA carrier ovarian cancer correlating
with platinum-free interval. J Clin Oncol 28:2512–2519.
Gandhi VB, Luo Y, Liu X, Shi Y, Klinghofer V, Johnson EF, Park C, Giranda VL,
Penning TD, and Zhu GD (2010) Discovery and SAR of substituted 3-oxoisoindoline4-carboxamides as potent inhibitors of poly(ADP-ribose) polymerase (PARP) for the
treatment of cancer. Bioorg Med Chem Lett 20:1023–1026.
Gangloff AR, Brown J, de Jong R, Dougan DR, Grimshaw CE, Hixon M, Jennings A,
Kamran R, Kiryanov A, O’Connell S, et al. (2013) Discovery of novel benzo[b][1,4]
oxazin-3(4H)-ones as poly(ADP-ribose)polymerase inhibitors. Bioorg Med Chem
Lett 23:4501–4505.
Gilabert M, Launay S, Ginestier C, Bertucci F, Audebert S, Pophillat M, Toiron Y,
Baudelet E, Finetti P, Noguchi T, et al. (2014) Poly(ADP-ribose) polymerase 1
(PARP1) overexpression in human breast cancer stem cells and resistance to olaparib.
PLoS One 9:e104302.
Gottipati P, Vischioni B, Schultz N, Solomons J, Bryant HE, Djureinovic T, Issaeva N,
Sleeth K, Sharma RA, and Helleday T (2010) Poly(ADP-ribose) polymerase is hyperactivated in homologous recombination-defective cells. Cancer Res 70:5389–5398.
Ha K, Fiskus W, Choi DS, Bhaskara S, Cerchietti L, Devaraj SG, Shah B, Sharma S,
Chang JC, Melnick AM, et al. (2014) Histone deacetylase inhibitor treatment
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
that spontaneously accumulate during the cell cycle. In this
scenario, two independent cell-killing mechanisms act in
parallel.
There are likely more examples of genotoxic agents that use
these three mechanisms. In addition, PARP inhibitors may also
be effective in combination with other agents that interfere
with DNA repair, such as CHK1/2 inhibitors and ATM inhibitors. From a mechanistic point of view, these combinations
may likely have synergistic effects (Bryant and Helleday, 2006;
Rehman et al., 2012; Tang et al., 2012; Booth et al., 2013;
We˛ sierska-Ga˛ dek and Heinzl, 2014), but whether they use the
trapping property of a PARP inhibitor has yet to be examined.
Finally, other targeted cancer therapeutic agents, such as
histone deacetylase inhibitors and phosphoinositide 3-kinase
inhibitors, may be considered for combination with PARP inhibitors to achieve improved anticancer activity. The mechanisms of these combinations are beyond the scope of this
review, but many interesting reports are available (Johnson
et al., 2011; Nowsheen et al., 2012; Cardnell et al., 2013; Ha
et al., 2014).
In summary, trapping of PARP on DNA has emerged as
a major mechanism by which PARP inhibitors exert antitumor
activity. The ability to induce PARP trapping varies considerably among the various PARP inhibitors. Their differential
PARP-trapping activities may be attributed to the differences
in their chemical scaffolds and hence binding modes in the CAT
domain. The distinct binding modes among these inhibitors
may serve as a structural basis for their varying abilities to
modulate interdomain communication and allosteric regulation, which have been postulated to underpin PARP trapping.
Recent advances in our understanding of DNA-induced PARP1
catalytic activation provide initial structural insight into potential molecular mechanisms for inhibitor-induced DNA
trapping. Understanding the molecular mechanism of PARP
trapping will assist in developing existing and new PARP inhibitors, as well as identifying optimal clinical applications of
various PARP inhibitors, allowing the rational selection of
appropriate combination therapies.
455
456
Shen et al.
McCabe N, Turner NC, Lord CJ, Kluzek K, Bialkowska A, Swift S, Giavara S,
O’Connor MJ, Tutt AN, Zdzienicka MZ, et al. (2006) Deficiency in the repair of
DNA damage by homologous recombination and sensitivity to poly(ADP-ribose)
polymerase inhibition. Cancer Res 66:8109–8115.
Miao ZH, Player A, Shankavaram U, Wang YH, Zimonjic DB, Lorenzi PL, Liao ZY,
Liu H, Shimura T, Zhang HL, et al. (2007) Nonclassic functions of human
topoisomerase I: genome-wide and pharmacologic analyses. Cancer Res 67:
8752–8761.
Michels J, Vitale I, Galluzzi L, Adam J, Olaussen KA, Kepp O, Senovilla L, Talhaoui
I, Guegan J, Enot DP, et al. (2013) Cisplatin resistance associated with PARP
hyperactivation. Cancer Res 73:2271–2280.
Miranda EA, Dantzer F, O’Farrell M, de Murcia G, and de Murcia JM (1995)
Characterisation of a gain-of-function mutant of poly(ADP-ribose) polymerase.
Biochem Biophys Res Commun 212:317–325.
Miyashiro J, Woods KW, Park CH, Liu X, Shi Y, Johnson EF, Bouska JJ, Olson AM,
Luo Y, Fry EH, et al. (2009) Synthesis and SAR of novel tricyclic quinoxalinone
inhibitors of poly(ADP-ribose)polymerase-1 (PARP-1). Bioorg Med Chem Lett 19:
4050–4054.
Murai J, Huang SY, Das BB, Renaud A, Zhang Y, Doroshow JH, Ji J, Takeda S,
and Pommier Y (2012) Trapping of PARP1 and PARP2 by clinical PARP inhibitors.
Cancer Res 72:5588–5599.
Murai J, Huang SY, Renaud A, Zhang Y, Ji J, Takeda S, Morris J, Teicher B, Doroshow
JH, and Pommier Y (2014a) Stereospecific PARP trapping by BMN 673 and comparison with olaparib and rucaparib. Mol Cancer Ther 13:433–443.
Murai J, Zhang Y, Morris J, Ji J, Takeda S, Doroshow JH, and Pommier Y (2014b)
Rationale for poly(ADP-ribose) polymerase (PARP) inhibitors in combination
therapy with camptothecins or temozolomide based on PARP trapping versus
catalytic inhibition. J Pharmacol Exp Ther 349:408–416.
Narwal M, Venkannagari H, and Lehtiö L (2012) Structural basis of selective inhibition of human tankyrases. J Med Chem 55:1360–1367.
Nitiss J and Wang JC (1988) DNA topoisomerase-targeting antitumor drugs can be
studied in yeast. Proc Natl Acad Sci USA 85:7501–7505.
Nowsheen S, Cooper T, Stanley JA, and Yang ES (2012) Synthetic lethal interactions
between EGFR and PARP inhibition in human triple negative breast cancer cells.
PLoS One 7:e46614.
O’Malley FP, Chia S, Tu D, Shepherd LE, Levine MN, Bramwell VH, Andrulis IL,
and Pritchard KI (2009) Topoisomerase II alpha and responsiveness of breast
cancer to adjuvant chemotherapy. J Natl Cancer Inst 101:644–650.
Papeo G, Casale E, Montagnoli A, and Cirla A (2013) PARP inhibitors in cancer
therapy: an update. Expert Opin Ther Pat 23:503–514.
Patel AG, Flatten KS, Schneider PA, Dai NT, McDonald JS, Poirier GG,
and Kaufmann SH (2012) Enhanced killing of cancer cells by poly(ADP-ribose)
polymerase inhibitors and topoisomerase I inhibitors reflects poisoning of both
enzymes. J Biol Chem 287:4198–4210.
Patel MR, Bhatt A, Steffen JD, Chergui A, Murai J, Pommier Y, Pascal JM, Trombetta
LD, Fronczek FR, and Talele TT (2014) Discovery and structure-activity relationship
of novel 2,3-dihydrobenzofuran-7-carboxamide and 2,3-dihydrobenzofuran-3(2H)-one-7carboxamide derivatives as poly(ADP-ribose)polymerase-1 inhibitors. J Med Chem 57:
5579–5601.
Pellicciari R, Camaioni E, Gilbert AM, Macchiarulo A, Bikker JA, Shah F, Bard J,
Constantino G, Gioiello A, Robertson GM, et al. (2011) Discovery and characterization of novel potent PARP-1 inhibitors endowed with neuroprotective properties:
from TIQ-A to HYDAMTIQ. Med Chem Commun 2:559–565.
Peng G and Lin SY (2011) Exploiting the homologous recombination DNA repair
network for targeted cancer therapy. World J Clin Oncol 2:73–79.
Penning TD, Zhu GD, Gandhi VB, Gong J, Liu X, Shi Y, Klinghofer V, Johnson EF,
Donawho CK, Frost DJ, et al. (2009) Discovery of the Poly(ADP-ribose) polymerase
(PARP) inhibitor 2-[(R)-2-methylpyrrolidin-2-yl]-1H-benzimidazole-4-carboxamide
(ABT-888) for the treatment of cancer. J Med Chem 52:514–523.
Penning TD, Zhu GD, Gong J, Thomas S, Gandhi VB, Liu X, Shi Y, Klinghofer V,
Johnson EF, Park CH, et al. (2010) Optimization of phenyl-substituted benzimidazole carboxamide poly(ADP-ribose) polymerase inhibitors: identification of
(S)-2-(2-fluoro-4-(pyrrolidin-2-yl)phenyl)-1H-benzimidazole-4-carboxamide (A-966492), a
highly potent and efficacious inhibitor. J Med Chem 53:3142–3153.
Pettitt SJ, Rehman FL, Bajrami I, Brough R, Wallberg F, Kozarewa I, Fenwick K,
Assiotis I, Chen L, Campbell J, et al. (2013) A genetic screen using the PiggyBac
transposon in haploid cells identifies Parp1 as a mediator of olaparib toxicity. PLoS
One 8:e61520.
Pfister TD, Hollingshead M, Kinders RJ, Zhang Y, Evrard YA, Ji J, Khin SA, Borgel
S, Stotler H, Carter J, et al. (2012) Development and validation of an immunoassay
for quantification of topoisomerase I in solid tumor tissues. PLoS One 7:e50494.
Pfister TD, Reinhold WC, Agama K, Gupta S, Khin SA, Kinders RJ, Parchment RE,
Tomaszewski JE, Doroshow JH, and Pommier Y (2009) Topoisomerase I levels in
the NCI-60 cancer cell line panel determined by validated ELISA and microarray
analysis and correlation with indenoisoquinoline sensitivity. Mol Cancer Ther 8:
1878–1884.
Plummer R, Lorigan P, Steven N, Scott L, Middleton MR, Wilson RH, Mulligan E,
Curtin N, Wang D, Dewji R, et al. (2013) A phase II study of the potent PARP
inhibitor, rucaparib (PF-01367338, AG014699), with temozolomide in patients
with metastatic melanoma demonstrating evidence of chemopotentiation. Cancer
Chemother Pharmacol 71:1191–1199.
Pommier Y (2013) Drugging topoisomerases: lessons and challenges. ACS Chem Biol
8:82–95.
Pommier Y, Barcelo JM, Rao VA, Sordet O, Jobson AG, Thibaut L, Miao ZH, Seiler
JA, Zhang H, Marchand C, et al. (2006) Repair of topoisomerase I-mediated DNA
damage. Prog Nucleic Acid Res Mol Biol 81:179–229.
Pommier Y, Leo E, Zhang H, and Marchand C (2010) DNA topoisomerases and their
poisoning by anticancer and antibacterial drugs. Chem Biol 17:421–433.
Pommier Y and Marchand C (2011) Interfacial inhibitors: targeting macromolecular
complexes. Nat Rev Drug Discov 11:25–36.
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
induces ‘BRCAness’ and synergistic lethality with PARP inhibitor and cisplatin
against human triple negative breast cancer cells. Oncotarget 5:5637–5650.
Haince JF, McDonald D, Rodrigue A, Déry U, Masson JY, Hendzel MJ, and Poirier
GG (2008) PARP1-dependent kinetics of recruitment of MRE11 and NBS1 proteins
to multiple DNA damage sites. J Biol Chem 283:1197–1208.
Hassler M and Ladurner AG (2012) Towards a structural understanding of PARP1
activation and related signalling ADP-ribosyl-transferases. Curr Opin Struct Biol
22:721–729.
Hattori K, Kido Y, Yamamoto H, Ishida J, Kamijo K, Murano K, Ohkubo M,
Kinoshita T, Iwashita A, Mihara K, et al. (2004) Rational approaches to discovery
of orally active and brain-penetrable quinazolinone inhibitors of poly(ADP-ribose)
polymerase. J Med Chem 47:4151–4154.
Helleday T (2011) The underlying mechanism for the PARP and BRCA synthetic
lethality: clearing up the misunderstandings. Mol Oncol 5:387–393.
Hopkins T, Solomon L, Shi YH, Rodriguez L, Donawho C, DiGiammorino E, Panchal
S, Olson A, Stolarik D, Osterling D, et al. (2014) PARP inhibitors trap PARP1 onto
damaged DNA via catalytic inhibition and not by an allosteric mechanism
(Abstract). Eur J Cancer 50:82–83.
Horton JK, Stefanick DF, Naron JM, Kedar PS, and Wilson SH (2005) Poly(ADPribose) polymerase activity prevents signaling pathways for cell cycle arrest after
DNA methylating agent exposure. J Biol Chem 280:15773–15785.
Horton JK, Stefanick DF, Prasad R, Gassman NR, Kedar PS, and Wilson SH (2014)
Base excision repair defects invoke hypersensitivity to PARP inhibition. Mol
Cancer Res 12:1128–1139.
Horton JK and Wilson SH (2013a) Predicting enhanced cell killing through PARP
inhibition. Mol Cancer Res 11:13–18.
Horton JK and Wilson SH (2013b) Strategic combination of DNA-damaging agent
and PARP inhibitor results in enhanced cytotoxicity. Front Oncol 3:257.
Ioanoviciu A, Antony S, Pommier Y, Staker BL, Stewart L, and Cushman M (2005)
Synthesis and mechanism of action studies of a series of norindenoisoquinoline
topoisomerase I poisons reveal an inhibitor with a flipped orientation in the ternary DNA-enzyme-inhibitor complex as determined by X-ray crystallographic
analysis. J Med Chem 48:4803–4814.
Iwashita A, Hattori K, Yamamoto H, Ishida J, Kido Y, Kamijo K, Murano K, Miyake
H, Kinoshita T, Warizaya M, et al. (2005) Discovery of quinazolinone and quinoxaline derivatives as potent and selective poly(ADP-ribose) polymerase-1/2
inhibitors. FEBS Lett 579:1389–1393.
Johnson N, Li YC, Walton ZE, Cheng KA, Li D, Rodig SJ, Moreau LA, Unitt C,
Bronson RT, Thomas HD, et al. (2011) Compromised CDK1 activity sensitizes
BRCA-proficient cancers to PARP inhibition. Nat Med 17:875–882.
Juarez-Salinas H, Sims JL, and Jacobson MK (1979) Poly(ADP-ribose) levels in
carcinogen-treated cells. Nature 282:740–741.
Kedar PS, Stefanick DF, Horton JK, and Wilson SH (2012) Increased PARP-1 association with DNA in alkylation damaged, PARP-inhibited mouse fibroblasts. Mol
Cancer Res 10:360–368.
Kinoshita T, Nakanishi I, Warizaya M, Iwashita A, Kido Y, Hattori K, and Fujii T
(2004) Inhibitor-induced structural change of the active site of human poly(ADPribose) polymerase. FEBS Lett 556:43–46.
Krishnakumar R and Kraus WL (2010) The PARP side of the nucleus: molecular
actions, physiological outcomes, and clinical targets. Mol Cell 39:8–24.
Langelier MF and Pascal JM (2013) PARP-1 mechanism for coupling DNA damage
detection to poly(ADP-ribose) synthesis. Curr Opin Struct Biol 23:134–143.
Langelier MF, Planck JL, Roy S, and Pascal JM (2011) Crystal structures of poly
(ADP-ribose) polymerase-1 (PARP-1) zinc fingers bound to DNA: structural and
functional insights into DNA-dependent PARP-1 activity. J Biol Chem 286:
10690–10701.
Langelier MF, Planck JL, Roy S, and Pascal JM (2012) Structural basis for DNA
damage-dependent poly(ADP-ribosyl)ation by human PARP-1. Science 336:
728–732.
Langelier MF, Servent KM, Rogers EE, and Pascal JM (2008) A third zinc-binding
domain of human poly(ADP-ribose) polymerase-1 coordinates DNA-dependent
enzyme activation. J Biol Chem 283:4105–4114.
Lehmann BD, Bauer JA, Chen X, Sanders ME, Chakravarthy AB, Shyr Y,
and Pietenpol JA (2011) Identification of human triple-negative breast cancer
subtypes and preclinical models for selection of targeted therapies. J Clin Invest
121:2750–2767.
Lindahl T (1993) Instability and decay of the primary structure of DNA. Nature 362:
709–715.
Lindgren AE, Karlberg T, Thorsell AG, Hesse M, Spjut S, Ekblad T, Andersson CD,
Pinto AF, Weigelt J, Hottiger MO, et al. (2013) PARP inhibitor with selectivity
toward ADP-ribosyltransferase ARTD3/PARP3. ACS Chem Biol 8:1698–1703.
Liu X, Han EK, Anderson M, Shi Y, Semizarov D, Wang G, McGonigal T, Roberts L,
Lasko L, Palma J, et al. (2009) Acquired resistance to combination treatment with
temozolomide and ABT-888 is mediated by both base excision repair and homologous recombination DNA repair pathways. Mol Cancer Res 7:1686–1692.
Loeffler PA, Cuneo MJ, Mueller GA, DeRose EF, Gabel SA, and London RE (2011)
Structural studies of the PARP-1 BRCT domain. BMC Struct Biol 11:37.
Lord CJ and Ashworth A (2012) The DNA damage response and cancer therapy.
Nature 481:287–294.
Luo X and Kraus WL (2012) On PAR with PARP: cellular stress signaling through
poly(ADP-ribose) and PARP-1. Genes Dev 26:417–432.
Lupo B and Trusolino L (2014) Inhibition of poly(ADP-ribosyl)ation in cancer: old and
new paradigms revisited. Biochim Biophys Acta 1846:201–215.
Mansoorabadi SO, Wu M, Tao Z, Gao P, Pingali SV, Guo L, and Liu HW (2014)
Conformational activation of poly(ADP-ribose) polymerase-1 upon DNA binding
revealed by small-angle X-ray scattering. Biochemistry 53:1779–1788.
Marchand JR, Carotti A, Passeri D, Filipponi P, Liscio P, Camaioni E, Pellicciari R,
Gioiello A, and Macchiarulo A (2014) Investigating the allosteric reverse signalling
of PARP inhibitors with microsecond molecular dynamic simulations and fluorescence anisotropy. Biochim Biophys Acta 1844:1765–1772.
Trapping PARP
Stewart E, Goshorn R, Bradley C, Griffiths LM, Benavente C, Twarog NR, Miller
GM, Caufield W, Freeman BB, 3rd, Bahrami A, et al. (2014) Targeting the DNA
repair pathway in Ewing sarcoma. Cell Rep 9:829–841.
Ström CE, Johansson F, Uhlén M, Szigyarto CA, Erixon K, and Helleday T (2011) Poly
(ADP-ribose) polymerase (PARP) is not involved in base excision repair but PARP
inhibition traps a single-strand intermediate. Nucleic Acids Res 39:3166–3175.
Tang Y, Dai Y, Grant S, and Dent P (2012) Enhancing CHK1 inhibitor lethality in
glioblastoma. Cancer Biol Ther 13:379–388.
Turner NC and Ashworth A (2011) Biomarkers of PARP inhibitor sensitivity. Breast
Cancer Res Treat 127:283–286.
von Euw EM, Gong KW, Rong HM, O’Brien N, Signh A, Shen Y, Post L, Finn RS,
Chmielowski B, and Slamon DJ (2014) Combination of a novel PARP inhibitor,
BMC 673, with temozolomide for the treatment of melanoma (Abstract). Pigment
Cell Melanoma Res 26:1011.
Wahlberg E, Karlberg T, Kouznetsova E, Markova N, Macchiarulo A, Thorsell AG, Pol
E, Frostell Å, Ekblad T, Öncü D, et al. (2012) Family-wide chemical profiling and
structural analysis of PARP and tankyrase inhibitors. Nat Biotechnol 30:283–288.
Wendorff TJ, Schmidt BH, Heslop P, Austin CA, and Berger JM (2012) The structure
of DNA-bound human topoisomerase II alpha: conformational mechanisms for
coordinating inter-subunit interactions with DNA cleavage. J Mol Biol 424:109–124.
We˛ sierska-Ga˛ dek J and Heinzl S (2014) Interactions between ataxia telangiectasia
mutated kinase inhibition, poly(ADP-ribose) polymerase-1 inhibition and BRCA1
status in breast cancer cells. J Cancer Prev 19:125–136.
Wiholm BE and Myrhed M (1993) Metformin-associated lactic acidosis in Sweden
1977-1991. Eur J Clin Pharmacol 44:589–591.
Ye N, Chen CH, Chen T, Song Z, He JX, Huan XJ, Song SS, Liu Q, Chen Y, Ding J,
et al. (2013) Design, synthesis, and biological evaluation of a series of benzo[de]
[1,7]naphthyridin-7(8H)-ones bearing a functionalized longer chain appendage as
novel PARP1 inhibitors. J Med Chem 56:2885–2903.
Yelamos J, Farres J, Llacuna L, Ampurdanes C, and Martin-Caballero J (2011)
PARP-1 and PARP-2: new players in tumour development. Am J Cancer Res 1:
328–346.
Zhang J, Stevens MF, and Bradshaw TD (2012) Temozolomide: mechanisms of action, repair and resistance. Curr Mol Pharmacol 5:102–114.
Zhang YW, Regairaz M, Seiler JA, Agama KK, Doroshow JH, and Pommier Y (2011)
Poly(ADP-ribose) polymerase and XPF-ERCC1 participate in distinct pathways for
the repair of topoisomerase I-induced DNA damage in mammalian cells. Nucleic
Acids Res 39:3607–3620.
Address correspondence to: Yuqiao Shen, BioMarin Pharmaceutical Inc.,
105 Digital Drive, Novato, CA 94949. E-mail: [email protected]
Downloaded from jpet.aspetjournals.org at ASPET Journals on June 18, 2017
Pommier Y, Pourquier P, Urasaki Y, Wu J, and Laco GS (1999) Topoisomerase I
inhibitors: selectivity and cellular resistance. Drug Resist Updat 2:307–318.
Rasheed ZA and Rubin EH (2003) Mechanisms of resistance to topoisomerase
I-targeting drugs. Oncogene 22:7296–7304.
Redinbo MR, Stewart L, Kuhn P, Champoux JJ, and Hol WG (1998) Crystal structures of human topoisomerase I in covalent and noncovalent complexes with DNA.
Science 279:1504–1513.
Rehman F, Elliott R, Brough R, Frankum J, Ragab A, Lord CJ, and Ashworth A
(2012) Developing rational drug combination strategies for PARP inhibitors. J Clin
Oncol 30 (suppl; abstr 3050).
Rouleau M, Patel A, Hendzel MJ, Kaufmann SH, and Poirier GG (2010) PARP inhibition: PARP1 and beyond. Nat Rev Cancer 10:293–301.
Ruf A, Rolli V, de Murcia G, and Schulz GE (1998) The mechanism of the elongation
and branching reaction of poly(ADP-ribose) polymerase as derived from crystal
structures and mutagenesis. J Mol Biol 278:57–65.
Sarkaria JN, Kitange GJ, James CD, Plummer R, Calvert H, Weller M, and Wick W
(2008) Mechanisms of chemoresistance to alkylating agents in malignant glioma.
Clin Cancer Res 14:2900–2908.
Satoh MS and Lindahl T (1992) Role of poly(ADP-ribose) formation in DNA repair.
Nature 356:356–358.
Schreiber V, Dantzer F, Ame JC, and de Murcia G (2006) Poly(ADP-ribose): novel
functions for an old molecule. Nat Rev Mol Cell Biol 7:517–528.
Shen Y, Rehman FL, Feng Y, Boshuizen J, Bajrami I, Elliott R, Wang B, Lord CJ,
Post LE, and Ashworth A (2013) BMN 673, a novel and highly potent PARP1/2
inhibitor for the treatment of human cancers with DNA repair deficiency. Clin
Cancer Res 19:5003–5015.
Singh A, Bartosz C, Jia W, Clark K, Dering J, Anderson L, Federman N, Eilber FC,
Shen Y, Finn R, et al. (2013) Mechanisms of resistance to the PARP inhibitor BMN
673 in Ewing’s sarcoma. Connective Tissue Oncology Society 18th Annual Meeting;
2013 30 Oct–2 Nov; New York, NY.
Smith MA, Reynolds CP, Kang MH, Kolb EA, Gorlick R, Carol H, Lock RB, Keir ST,
Maris JM, Billups CA, et al. (2015) Synergistic activity of PARP inhibition by
talazoparib (BMN 673) with temozolomide in pediatric cancer models in the pediatric preclinical testing program. Clin Cancer Res 21:819–832.
Solomon L, Hopkins T, Shi Y, Ellis P, Donawho C, Johnson EF, Fang G, and Maag D
(2014) Veliparib (ABT-888) potentiates the cytotoxic activity of DNA alkylating
agents by trapping PARP onto damaged chromatin. Annual Meeting of the American Association for Cancer Research; 2014 5–9 Apr; San Diego, CA.
Steffen JD, Brody JR, Armen RS, and Pascal JM (2013) Structural implications for
selective targeting of PARPs. Front Oncol 3:301.
Steffen JD, Tholey RM, Langelier MF, Planck JL, Schiewer MJ, Lal S, Bildzukewicz
NA, Yeo CJ, Knudsen KE, Brody JR, et al. (2014) Targeting PARP-1 allosteric
regulation offers therapeutic potential against cancer. Cancer Res 74:31–37.
457