Synthesis, biological evaluation, and structure-activity relationships of a novel class of apurinic/apyrimidinic endonuclease 1 inhibitors

Article abstract of DOI:10.1021/jm201537d

APE1 is an essential protein that operates in the base excision repair (BER) pathway and is responsible for a‰￥ 95% of the total apurinic/apyrimidinic (AP) endonuclease activity in human cells. BER is a major pathway that copes with DNA damage induced by several anticancer agents, including ionizing radiation and temozolomide. Overexpression of APE1 and enhanced AP endonuclease activity have been linked to increased resistance of tumor cells to treatment with monofunctional alkylators, implicating inhibition of APE1 as a valid strategy for cancer therapy. We report herein the results of a focused medicinal chemistry effort around a novel APE1 inhibitor, N-(3-(benzo[d]thiazol-2-yl)-6-isopropyl-4,5,6,7-tetrahydrothieno[2,3-c] pyridin-2-yl)acetamide (3). Compound 3 and related analogues exhibit single-digit micromolar activity against the purified APE1 enzyme and comparable activity in HeLa whole cell extract assays and potentiate the cytotoxicity of the alkylating agents methylmethane sulfonate and temozolomide. Moreover, this class of compounds possesses a generally favorable in vitro ADME profile, along with good exposure levels in plasma and brain following intraperitoneal dosing (30 mg/kg body weight) in mice. This article not subject to U.S. Copyright. Published 2012 by the American Chemical Society.

Full text of DOI:10.1021/jm201537d

Article
pubs.acs.org/jmc
Synthesis, Biological Evaluation, and Structure−Activity
Relationships of a Novel Class of Apurinic/Apyrimidinic
Endonuclease 1 Inhibitors
Ganesha Rai,†,§ Vaddadi N. Vyjayanti,‡,§ Dorjbal Dorjsuren,† Anton Simeonov,† Ajit Jadhav,†
David M. Wilson, III,*,‡ and David J. Maloney*,†
†NIH Chemical Genomics Center, National Center for Advancing Translational Sciences, National Institutes of Health, Bethesda,
Maryland 20892-3370, United States
‡Laboratory of Molecular Gerontology, National Institute on Aging, National Institutes of Health, Baltimore, Maryland 21224, United
States
S
* Supporting Information
ABSTRACT: APE1 is an essential protein that operates in the base excision repair (BER) pathway and is
responsible for ≥95% of the total apurinic/apyrimidinic (AP) endonuclease activity in human cells. BER is a
major pathway that copes with DNA damage induced by several anticancer agents, including ionizing
radiation and temozolomide. Overexpression of APE1 and enhanced AP endonuclease activity have been
linked to increased resistance of tumor cells to treatment with monofunctional alkylators, implicating
inhibition of APE1 as a valid strategy for cancer therapy. We report herein the results of a focused medicinal
chemistry effort around a novel APE1 inhibitor, N-(3-(benzo[d]thiazol-2-yl)-6-isopropyl-4,5,6,7-
tetrahydrothieno[2,3-c]pyridin-2-yl)acetamide (3). Compound 3 and related analogues exhibit single-digit
micromolar activity against the purified APE1 enzyme and comparable activity in HeLa whole cell extract assays and potentiate
the cytotoxicity of the alkylating agents methylmethane sulfonate and temozolomide. Moreover, this class of compounds
possesses a generally favorable in vitro ADME profile, along with good exposure levels in plasma and brain following
intraperitoneal dosing (30 mg/kg body weight) in mice.
temozolomide (TMZ).8,9 TMZ is a promising drug recently
added to the arsenal of alkylating agents for the adjuvant
INTRODUCTION
Apurinic/apyrimidinic (AP) endonuclease 1 (APE1) is the
■
chemotherapy of brain cancers because of its ability to readily
primary mammalian enzyme responsible for the removal of
cross the blood−brain barrier.10 It has therefore been
postulated that APE1 would be an attractive target in anticancer
treatment paradigms involving coadministration with certain
DNA-interactive drugs, where strategic regulation of its repair
activity would improve the therapeutic efficacy and clinical
outcome.
abasic (or AP) sites in DNA.1,2 Such lesions can arise as
products of the repair activity of DNA glycosylases, which carry
out the first step of base excision repair (BER), or as products
of spontaneous or damage-induced hydrolysis of the N-
glycosidic bond that links the base to the sugar moiety of the
phosphodiester DNA backbone. APE1 initiates AP site repair
by catalyzing a Mg2+-facilitated strand incision event immedi-
ately 5′ to the lesion, leaving behind a single-strand break with a
normal 3′-hydroxyl residue and a 5′-abasic fragment. Other
proteins of BER, such as DNA polymerase β and DNA ligase 3,
complete the repair response by removing the remaining abasic
residue, replacing the missing nucleotide, and sealing the nick.3
Besides being the predominant repair enzyme for AP sites in
DNA, APE1 also possesses repair nuclease activities against
various nonconventional 3′-blocking terminal groups, such as
phosphoglycolates, phosphates, and chain-terminating nucleo-
side analogues.4 In addition, APE1 has roles in regulating gene
expression, most notably through its capacity to modulate the
DNA binding activity of several transcription factors, e.g., AP-1,
p53, and NF-κB, via a less well understood redox
mechanism.5−7
Targeting DNA repair enzymes as single-agent cancer
therapy has been validated as a viable strategy by the discovery
and clinical evaluation of poly ADP-ribose polymerase (PARP)
inhibitors.11 PARP1 is an enzyme that facilitates efficient repair
of single-strand breaks in DNA. Thus, inhibition of PARP1
leads to the accumulation of one-ended double-strand DNA
breaks upon replication fork collapse that are ultimately
repaired via homologous recombination (HR).12 BRCA1/2
are proteins involved in the HR pathway, and consequently,
treatment of BRCA-deficient cancer cells (e.g., ∼10−20% of
triple negative breast cancers) with PARP1 inhibitors leads to
irreparable DNA damage and ultimately cell death.13 This
synthetic lethal relationship offers the prospect of selective
targeting of cancer cells, since normal cells would maintain the
ability to repair DNA double-strand breaks. However, despite
APE1 protein levels and intracellular distribution have been
correlated with cancer type and stage, as well as responsiveness
to clinical DNA-damaging agents, such as the alkylator
Received: November 14, 2011
Published: March 28, 2012
This article not subject to U.S. Copyright.
Published 2012 by the American Chemical
Society
3101
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
continued efforts and promising results in this area of research,
the use of PARP1 inhibitors has not been without its recent
setbacks in the clinic.14 While synthetically lethal combinations
involving APE1 inhibitors have not yet been established, it is
not unreasonable to postulate the use of APE1 inhibitors as
single agent therapy by such a mechanism.
As a result of the promising therapeutic potential of this
target, several reports have described the identification and
characterization of small molecules that inhibit APE1 repair
endonuclease activity.15 Kelley and co-workers described the
identification of 2,4,9-trimethylbenzo[b][1,8]naphthyridin-5-
amine, 1 (AR03) through a fluorescence-based high-throughput
screen (HTS) of 60 000 compounds (Figure 1).16 1 was found
lucanthone (also a topoisomerase inhibitor),21 methoxy-
amine,22 and various arylstibonic acids.23
We recently reported on the development of a 1536-well
fluorescence-based, quantitative HTS (qHTS) assay, which was
used to screen the Library of Pharmacologically Active
Compounds (LOPAC1280) for novel APE1 endonuclease
inhibitors.24 This library is a collection of well characterized
bioactive molecules that is often used to help validate an assay
platform before screening of an entire small molecule
collection. Our initial efforts identified several compounds,
including 6-hydroxy-DL-DOPA, reactive blue, and myricetin,
which inhibit AP site cleavage activity of purified APE1, as well
as HEK293T and HeLa whole cell extracts, and exhibit
potentiating effects of MMS toxicity. These prior studies
established general methods for identification and validation of
APE1 inhibitors. While encouraged by these results, we were
eager to identify novel small molecule inhibitors that would
represent an improved starting point for medicinal chemistry
optimization, as the three LOPAC1280 compounds mentioned
above appear to be generally promiscuous given the broad
publications on their various modes of action.
As such, we conducted a large-scale qHTS in a 1536-well
concentration response format on our complete in-house
collection, which at the time contained 352 498 small
molecules25 as part of the NIH Molecular Libraries Small
Molecule Repository (MLSMR). This effort represents the
largest reported screening campaign against APE1 thus far and
led to the identification of several novel chemotypes displaying
potent inhibition of APE1 activity. In this report, we discuss the
synthesis, biological characterization, and structure−activity
relationship (SAR) of one of the molecules, N-(3-(benzo[d]-
thiazol-2-yl)-6-isopropyl-4,5,6,7-tetrahydrothieno[2,3-c]-
pyridin-2-yl)acetamide (3) (Figure 1). This compound, along
with related analogues, displayed low micromolar APE1
inhibition and activity in HeLa whole cell extract assays,
potentiated the cytotoxicity of MMS and TMZ, and led to a
hyperaccumulation of AP sites in HeLa cells treated with MMS.
Figure 1. Previously reported APE1 endonuclease inhibitors (1 and 2)
and the lead chemotype (3).
to have low micromolar in vitro potency against purified human
APE1 and inhibited AP site incision activity of whole cell
extracts and the repair of AP sites in SF767 glioblastoma cells.
Moreover, 1 potentiated the cytotoxicity of methyl methansul-
fonate (MMS) and TMZ in SF767 cells. 7-Nitro-1H-indole-2-
carboxylic acid, 2 (CRT0044876) (Figure 1), was identified by
Madhusudan et al. in 2005, and they described specific
inhibition of the exonuclease III family of AP endonucleases
and the induction of AP sites in HT1080 fibrosarcoma cells.17
While a synergistic cell killing effect was seen with the inhibitor
when combined with MMS or TMZ, other subsequent studies
have been unable to reproduce the potentiating effect of 2.18
More recently, Madhusudan and colleagues described the
results of a virtual screen of 2.6 million compounds from which
several low micromolar APE1 inhibitors were found.19 Other
reported APE1 inhibitors include the bis-carboxylic acid
containing small molecules described by Zawahir et al.,20
CHEMISTRY
■
The synthesis of the lead chemotype 3 from the qHTS
campaign commenced with treatment of commercially available
a
Scheme 1. Synthesis of qHTS “Hit” 3 and Related Analogues 4−8
a
Reagents and conditions: (a) S, morpholine, EtOH, reflux, 2 h (87%); (b) AcCl, Hunig’s base, CH2Cl2, rt, 1 h (91%); (c) TFA, CH2Cl2, rt, 1 h
(99%). (d) 3: acetone, NaCNBH3, MeOH/THF, rt, 6 h (75%). 6: paraformaldehyde (aq), NaCNBH3, MeOH/THF, rt, 6 h. 7: PhCHO, NaCNBH3,
MeOH/THF, 6 h. 8: AcCl, Hunig’s base, CH2Cl2, rt.
3102
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
a
Scheme 2. Synthesis of Analogues 9−24
a
Reagents and conditions: (a) S, morpholine, EtOH, reflux, 2 h (84%); (b) AcCl, Hunig’s base, CH2Cl2, rt, 1 h (91%); (c) LiOH, THF/H2O, reflux,
24 h (65%); (d) TFA, CH2Cl2, rt, 0.5 h; (e) dimethylacetamide, 170 °C, microwave, 2 h (77%); (f) Br2, CHCl3, −10 °C, 1 h (70%); (g) R-B(OH)2,
Pd(PPh3)4, Na2CO3, DME, 150 °C, microwave, 0.5−1.5 h; (h) acetone, NaCNBH3, MeOH/THF, rt, 6−10 h.
a
Scheme 3. Synthesis of Analogues 25−46
a
Reagents and conditions: (a) TFA, CH2Cl2, rt, 0.5 h; (b) acetone, NaCNBH3, MeOH, rt, 6 h; (c) RC(O)Cl, (i-Pr)2NEt, CH2Cl2, rt. 31, 33, 36:
RC(O)OH, EDC, DMAP, DMF, rt, 1 h, (85−90%). (d) NaNO2, HCl−H2SO4, 0 °C, 2 h; (e) H3PO2, rt, 1 h; (f) t-BuNO2, CuBr2, MeCN, 0 °C →
rt, 4 h; (g) Pd(OAc)2, dppp, CO(g) (1 atm), NEt3, DMSO−MeOH (1:1), 60 °C, 24 h, (69%); (h) LiOH, THF/MeOH/H2O (6:2:1), rt, 3 h (91%);
(i) acetohydrazide, EDC, DMF, rt, 2 h (97%); (j) Burgess reagent, THF, 100 °C, microwave, 0.5 h (73%), then TFA, CH2Cl2, rt, 0.5 h; (k) N′-
hydroxyacetimidamide, HATU, (i-Pr)2NEt, 50 °C for 15 min, then 150 °C for 15 min, microwave, 1 h, then TFA, CH2Cl2, rt, 0.5 h; (l) 5-methyl-
1,3,4-oxadiazol-2-amine, Pd2(dba)3, xantphos, Cs2CO3, dioxane, 125 °C, microwave, 2 h, then TFA, CH2Cl2, rt, 0.5 h; (m) 3-methyl-1,2,4-oxadiazol-
5-amine, Pd2(dba)3, xantphos, Cs2CO3, dioxane, 125 °C, microwave, 2 h, then TFA, CH2Cl2, rt, 0.5 h.
N-(Boc)-4-piperidone with 2-benzothiazoleacetonitrile in the
presence of elemental sulfur and morpholine to afford the key
intermediate 3a as shown in Scheme 1.26 Acetylation of the 2-
amino group, followed by Boc deprotection with trifluoroacetic
acid (TFA) and reductive amination with acetone and
NaCNBH3, gave the desired compound 3 in good yield.
Starting with intermediate 3a, several different N-substituted
analogues were synthesized, including R = Me (6, via reductive
amination with paraformaldehyde), R = Bn (7) (via reductive
amination with benzaldehyde), and R = Ac (8, via acylation
with AcCl).
The synthesis of analogues 9−24, which involved mod-
ification to the C-3 benzothiazole ring, required the develop-
ment of a more convergent route. While the original synthesis
of lead compound 3 only involved four steps, diversity of the
benzothiazole ring occurs in the first step and thus requires the
availability of the necessary acetonitrile derivative. Moreover,
each modified benzothiazole analogue would involve a
3103
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
a
Scheme 4. Synthesis of Analogues 47−55
a
Reagents and conditions: (a) m-CPBA, CH2Cl2, 0 °C, 1 h, (87%); (b) MnO2, toluene, 120 °C, microwave, 10 min (49%).
minimum of four steps, making library synthesis quite
laborious. As such, we envisaged a late-stage Suzuki coupling
of commercially available boronic acids to the 3-bromothio-
phene intermediate (11) allowing for rapid access to a variety
of analogues at the 3-position as shown in Scheme 2. Reaction
of N-(Boc)-4-piperidone with ethyl 2-cyanoacetate using the
conditions described above (S, morpholine), followed by
acetylation and subsequent saponification of the formed ethyl
ester, gave the desired 3-COOH intermediate 9a. Decarbox-
ylation using dimethylacetamide at 170 °C for 2 h in a
microwave gave the des-carboxy product 10a after optimization
of the reaction conditions. Typically, this transformation is
carried out with copper using quinoline as the solvent or oxalic
acid in isopropanol, both at elevated temperatures. Our
conditions are quite mild (albeit at elevated temperatures),
and a wide range of functional groups should be tolerated.
Subsequent bromination of 10a using Br2 in chloroform at −10
°C followed by Boc deprotection gave 11 in good yield.
Importantly, we found that the late-stage Suzuki coupling could
be accomplished using the unprotected piperidine moiety via
treatment of 11 with various commercially available boronic
acids to provide 12−20 in good yields. Reductive amination
with acetone, NaCNBH3, and MeOH/THF as the solvent gave
the N-isoproyl analogues 22−24.
Analogues 25−46 were synthesized in an effort to further
understand the SAR of the amide moiety as shown in Scheme
3. Analogue 25, in which the 2-amino group was left
unacetylated, was prepared in two steps from the common
intermediate 3a via Boc deprotection and regioselective
reductive amination of the piperidine moiety with acetone
and NaCNBH3. Analogues 26−33, involving either reaction
with the requisite acid chloride or EDC-mediated peptide
coupling conditions (for 31, 33, and 36), were utilized to afford
the desired products in good yields. In addition to the various
amide analogues, we wanted to investigate amide bioisosteres
such as oxadiazoles 43−46. The synthesis of oxadiazole
analogues 43 and 44 commenced with conversion of the 2-
amino group to the corresponding bromide using tert-butyl
nitrite and copper bromide in acetonitrile. Palladium-catalyzed
carboxylation was achieved using catalyst Pd(OAc)2, catalyst
dppp, in DMSO−MeOH under an atmosphere of CO(g) to
afford the desired methyl ester derivative 41a in 69% yield.
Saponification of 41a with lithium hydroxide in a THF/
MeOH/H2O mixture gave carboxylic acid 42a in high yield.
Formation of the acylhydrazide was accomplished using
acetohydrazide and EDC in DMF. Dehydrative cyclization
was achieved using methyl N-(triethylammoniumsulfonyl)-
carbamate (Burgess reagent) in THF at 100 °C in the
microwave to give, after Boc deprotection (TFA, CH2Cl2), the
1,3,4-substituted oxadiazole 43 in good yield. 1,2,4-Oxadiazole
derivative 44 was prepared in three steps from intermediate 42a
via treatment with N′-hydroxyacetimidamide, HATU, Hunig’s
base at (50 °C → 150 °C) to afford the cyclized product, which
after Boc deprotection gave the desired product 44. Access to
2-aminooxadiazole analogues 45 and 46 was achieved via
Buchwald−Hartwig-type cross-couplings of the requisite
commercially available aminooxadiazole (5-methyl-1,3,4-oxa-
diazol-2-amine (45) and 3-methyl-1,2,4-oxadiazol-5-amine
(46)) using Pd2(dba)3, xantphos, and cesium carbonate in
the microwave for 2 h followed by Boc deprotection.
As shown in Scheme 4, we were eager to investigate various
heteroatoms in place of the piperidine nitrogen of lead
compound 3 (e.g., O, 47; S, 48; SO2, 49). Accordingly, the
syntheses of 47 and 48 were carried out in a manner similar to
that shown in Scheme 1, except tetrahydropyran-4-one and
tetrahydrothiopyran-4-one were used for the preparation of 47
and 48, respectively. Synthesis of sulfone derivative 49 was
accomplished via treatment of 48 with m-CPBA in methylene
chloride at 0 °C, and pyridine analogue 50 was obtained via
MnO2 oxidation of intermediate 4. Five-membered ring
analogues 51 and 52 were prepared using the route described
in Scheme 1 except N-Boc-3-pyrrolidinone was used as the
starting material, whereas for 53, N-(Boc)-3-piperidone was
utilized. Analogues 54 and 55 were synthesized from 1-Boc-4-
azepanone, which upon cyclization (S, morpholine) gave two
1
separable regioisomers that were easily distinguishable by H
NMR; both reactions were carried through using the described
sequence to provide the desired products.
RESULTS AND DISCUSSION
■
SAR Analysis of Compound 3 Using the Fluorescence-
Based HTS Assay. As a benchmark, we included some of the
previously reported APE1 inhibitors. Specifically, 1 was already
part of the MLSMR collection and 2 was purchased from Sigma
3104
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
Chemical Company. As shown in Table 1, compound 2 showed
only modest inhibition (40%) at concentrations up to 57 μM,
Table 2. SAR of Analogues 9−24
Table 1. Activity of Previously Reported APE1 Inhibitors (1
and 2) and SAR of Analogues 3−8
a
IC50 values were determined using the qHTS protocol and represent
a
b
IC50 values were determined using the qHTS protocol and represent
the average of three separate experiments. For compounds that did
b
the average of three separate experiments. For compounds that did
not achieve full inhibition (>85%), the % inhibition at the highest
not achieve full inhibition (>85%), the % inhibition at the highest
tested concentration (57 μM) is noted. Compounds noted as >57 μM
are considered inactive.
tested concentration (57 μM) is noted. Compounds noted as >57 μM
c
are considered inactive. IC50 of this compound reflects data from
primary screen; follow-up studies with a powder sample were not
further pursued.
benzothiophene (15), benzofuran (17), oxazole (19), and
thiazole (20), resulted in greatly diminished activity. Similar
results were obtained when that position was modified with a
simple phenyl group (22) or numerous other substituted
phenyl derivatives (data not shown). Interestingly, even the
structurally similar benzoxazole analogue (23) had a 10-fold
loss in activity; however, the 2-(4-phenylthiazole) analogue
(24) displayed potent inhibition with activity comparable to
that of the lead compound. The combination of the reduced
activity for the benzoxazole derivative (23) and the
maintenance of activity for compound 24 suggests that the
thiazole motif is involved in an essential interaction with APE1
and should be maintained in some capacity in future analogues.
However, additional substitution is required, since the thiazole
alone (compound 20) displayed minimal activity. As such,
additional modifications of the pendent aryl ring in analogue 24
may prove fruitful.
in agreement with previous reports that showed an inability to
reproduce the activity of this small molecule.18 1 did show
APE1 inhibition, albeit lower than the reported IC50 value (1.5
μM reported16 vs 32 μM observed). Since assay variability is to
be expected, this observation emphasizes the need to include
prior art as internal comparative controls. This result is from
the primary screen and not from a synthesized or purified
commercial powder source, so the source and effective
concentration of the sample could also contribute to the
discrepancy. Our lead compound 3 exhibited an IC50 of 2 μM
in the qHTS assay (Table 1) and an IC50 of 12 μM in a
radiotracer incision assay (RIA). The confirmed activity,
chemical tractability, and preliminary cell-based data, which
showed that compound 3 potentiated the cytotoxicity of MMS
in HeLa cells (vide infra), led us to proceed with optimization
and SAR exploration of this chemotype.
The goal of analogues 25−46 (Table 3) was to gain a better
understanding of the SAR associated with the N-acetyl moiety
of the lead compound 3. Thus, a variety of different compounds
were prepared, including a des-acetyl, numerous amides,
carbamates, carboxylic acids, esters, and amide bioisosteres
(1,2,4-oxadiazoles and 1,3,4-oxadiazoles). Our first interest was
to look at the des-acetyl analogue 25; this modification was
clearly unfavorable with only 40% inhibition at 57 μM being
observed. Interestingly, compound 25 was less potent than
analogue 40, which has the amino group removed altogether.
Many of the amide analogues resulted in a decrease in potency
[e.g., tert-butyl (26), inactive; cyclopentane (28), 55% at 57
μM; Bn (29), 53% at 57 μM; (CH2)2Ph (30), inactive].
However, other amide analogues exhibited a less pronounced
drop in activity [e.g., cyclopropane (27), 6 μM; cyclohexane
(37), 8.3 μM; CHF2 (33 and 36), 3.8 and 5.3 μM,
respectively]. Replacing the amide with other functional groups,
such as carboxylic acid (42) or ester (41), resulted in complete
loss of activity; however, methyl carbamate (32) was marginally
active with an IC50 of 11 μM. Given the possibility of hydrolysis
of the amide by intracellular amidases, we were eager to explore
the tolerance of amide bioisosteres such as 1,2,4-oxadiazoles
and 1,3,4-oxadiazoles. To this end, both analogues that lack the
2-amino group (43 and 44) and those that maintained this
moiety (45 and 46) were investigated. The studies revealed the
necessity of the 2-amino group for maintaining activity for these
Our first SAR investigations involved modification of the
piperidine nitrogen moiety, which revealed that acylation was
not well tolerated, as introduction of both a Boc group (5) and
an acetyl group (8) led to a significant loss of activity.
Moreover, the benzyl substituted analogue (7) showed greatly
reduced activity with only 41% inhibition at the highest
concentration tested (57 μM). In contrast, removal of the
isopropyl group (4) and introduction of the smaller alkyl group
(R = Me, 6) had little effect on potency with IC50 values of 2.9
and 3.8 μM, respectively. These data suggest that potency is
greatly affected by the absence of a basic nitrogen (analogues 5
and 8). However, there is also not much tolerance for larger
hydrophobic groups given that the benzyl substituted analogue,
which maintains the basic character of the nitrogen, is weakly
active.
The next area of SAR exploration involved modification of
the benzothiazole moiety at the 3-position of the thiophene.
Given the results obtained from our initial round of SAR
(Table 1), where we found analogue 4 (lacking isopropyl
group) to have potency comparable to that of derivative 3,
most analogues were submitted for testing as the free amine to
eliminate one step in the analogue synthesis. As shown in Table
2, many of the changes were not productive, with many
compounds being inactive or only having modest activity at the
highest concentration. Replacing the benzotriazole motif with
other heterocyles, including furan (13), thiophene (14),
3105
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
Table 3. SAR of Analogues 25−46
Table 4. SAR of Analogues 47−55
a
IC50 values were determined using the qHTS protocol and represent
b
the average of three separate experiments. For compounds that did
a
not achieve full inhibition (>85%), the % inhibition at the highest
tested concentration (57 μM) is noted. Compounds noted as >57 μM
are considered inactive.
IC50 values were determined using the qHTS protocol and represent
b
the average of three separate experiments. For compounds that did
not achieve full inhibition (>85%), the % inhibition at the highest
tested concentration (57 μM) is noted. Compounds noted as >57 μM
are considered inactive.
Table 5. APE1 Inhibition of Select Compounds in
Radiotracer Incision, HTS, and HeLa Whole Cell Extracts
Assays
bioisosteres, as compounds 43 and 44 were completely inactive
while 45 and 46 had modest activity. While these last two
compounds have approximately 8-fold less activity than the
corresponding amide analogue (4), it does suggest the
possibility of utilizing such modifications on this class of
compounds in later rounds of optimization if deemed necessary
by pharmacokinetic (PK) studies.
a
RIA IC50
(μM)
HTS IC50
(μM)
HeLa cell extract, % incision
b
compd
activity
3
12
5
2.0
0.6 0.11
1.4 1.1
26 3.7
12 1.3
15 0.5
38 7.7
15 14
4
2.9
6
5
3.8
23
24
33
51
52
54
55
13
3
20.4
2.9
Our last foray into the SAR characterization described herein
involved modification of the piperidine ring, where we replaced
the nitrogen with other heteroatoms. In the absence of
structural information to guide our medicinal chemistry efforts,
we aimed to explore the effect (i.e., altering key hydrogen bond
contacts) of moving the nitrogen to different positions on the
ring and varying the ring size (e.g., n = 5, 6, and 7). As shown in
Table 4, most of these changes were met with limited success;
however, a few interesting lessons were learned. Replacing the
nitrogen with oxygen (analogue 47) or sulfur (analogue 48) led
to compounds that were still active against APE1, albeit only at
the highest concentration. Sulfone derivative 49 had very
minimal efficacy (29%), and pyridine analogue 50 was
completely inactive. Varying the ring size with pyrrolidine
analogues 51 and 52 had a limited effect on activity, with IC50
values of 3 and 3.1 μM, respectively. Differentially substituted
piperidine analogue 53 had only modest efficacy at 57 μM,
while the seven-membered ring derivatives 54 and 55 displayed
activities of 12.9 and 17.6 μM, respectively.
APE1 Inhibition in Radiotracer Incision and HeLa
Whole Cell Extract Assays. Having tested all of the
synthesized analogues using the fluorescence-based HTS
assay to establish a rank order of potency, we next aimed to
validate their activity by testing in the RIA. This assay is
considered the “golden standard”, but because of its low
throughput nature, only our top analogues were analyzed via
this method (Table 5). Most analogues had comparable activity
across both experimental platforms, with 52 emerging as the
most potent compound in the RIA.
4
3.8
4
3.1
1
3.3
0.1 0.03
14 2.8
1.1 0.3
14
13
12.9
17.6
a
IC50 values were determined for select compounds using the RIA (see
Experimental Methods for details); assays were run in duplicate at 0, 1,
b
3, 10, 30, 50, and 100 μM inhibitor concentration. 100 μM of
inhibitor was used in these experiments, and the values represent the
average and standard deviation of three independent measurements
relative to the no inhibitor control.
Since APE1 comprises ≥95% of the total AP endonuclease
activity in mammalian cells, most, if not all, of the AP site
incision activity of human whole cell extracts is APE1-
dependent.27 Thus, as a means of assessing the specificity of
candidate APE1 inhibitors and their potential biological
potency, we determined the effect of the most promising
actives on total AP site cleavage activity of HeLa whole cell
extracts. This experiment was initially done using a single
concentration of inhibitor (100 μM) to quickly establish
relative activities of the selected analogues. The results showed
that several compounds exhibited near 100% inhibition of the
incision activity of HeLa extracts (analogues 3, 4, 52, and 55;
Table 5). Taking into account the activity observed in the HTS,
radiotracer, and whole cell extract assays (Table 5), we decided
to pursue compounds 52 and 3 further. As such, these two
compounds were tested again in the HeLa whole cell extract
assay at five different concentrations (0, 1, 3, 10, 30, and 100
3106
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
μM; Figure 2). Notably, each compound was found to inhibit
total AP site incision activity with IC50 values comparable to the
(determined from Lineweaver − Burk plots), of these
experiments. In both cases, the kcat decreases less than 6-fold,
and only at the high inhibitor concentration. In the case of 52,
KM increases substantially (up to 10-fold), while the KM
increases less dramatically with 3 (up to ∼4-fold). Such trends
would suggest that these compounds act as competitive
inhibitors of APE1 activity and, thus, presumably bind the
active site of the enzyme.
To further explore the potential mechanism of action of the
inhibitor compounds, a previously established electrophoretic
mobility shift assay (EMSA) was employed to determine the
consequence of 3 and 52 on APE1 and 32P radiolabeled AP-
DNA complex formation and stability. The results with both
inhibitors, which in the initial experiments were preincubated
with the protein prior to substrate addition, showed that the
percentage of APE1-DNA complex (C) decreased in a
compound-dose-dependent manner (Figure 4). In particular,
the binary complex was essentially absent when the protein was
pretreated with 30 μM inhibitor, with compound 52 exhibiting
a slightly more reproducible, greater effect. If we preincubated
either of the inhibitors with the radiolabeled DNA substrate
prior to the addition of APE1, we similarly saw a significant
reduction in protein−DNA complex formation (Supporting
Information Figure 1). Not surprisingly, a >30-fold excess of
cold (unlabeled) AP-DNA essentially abolished the formation
of radiolabeled substrate complex (Supporting Information
Figure 1). These data suggest that the small molecules bind the
same site on APE1 as the DNA substrate (albeit apparently
with lesser affinity than AP-DNA itself), thereby acting as
competitive inhibitors, although an allosteric effect cannot be
ruled out in the absence of high resolution complex structural
information. Moreover, it is currently unknown whether these
inhibitors affect other activities of APE1, such as its redox
function.
Figure 2. Inhibition of HeLa whole cell extract AP site incision activity
with 3 or 52. HeLa whole cell extract (300 ng) was incubated with 0,
1, 3, 10, 30, or 100 μM indicated inhibitor at room temperature for 15
min, prior to the addition of 0.5 pmol of radiolabeled AP-DNA
substrate. The reaction was then carried out at 37 °C and analyzed as
described in Experimental Methods. Shown is a bar graph reporting
the relative percent incision activity in comparison to the no inhibitor
control, arbitrarily set at 100%. The values represent the averages and
standard deviation of three independent experimental data points.
IC50 obtained with the purified enzyme. In particular,
compound 52 showed almost complete inhibition at 10 μM,
suggesting selectivity for APE1 even amidst the other
nonspecific proteins of the extract.
Mechanism of APE1 Inhibition by Compounds 3 and
52. Encouraged by the potency of compound 52 in the
radiotracer incision and whole cell extract assays, we pursued
mechanism of action studies of this compound and our initial
lead molecule 3. To reveal the mode of inhibition, we
determined the kinetic parameters for APE1 incision in the
absence or presence of 5, 10, or 20 μM inhibitor at varying
concentrations of DNA substrate. Figure 3 shows the
Michaelis−Menten plots, as well as the KM and kcat values
Cellular Effects of APE1 Inhibitors 3 and 52. To better
assess the biological potency of 3 and 52, we measured the level
of genomic AP sites in MMS-treated HeLa cells with or without
exposure to these compounds (Figure 5). As one might predict,
treatment of cells with the alkylator MMS or either of the APE1
inhibitors alone resulted in a statistically significant increase in
Figure 3. Kinetic analysis for compounds 3 and 52. APE1 (∼28 pM) was incubated without or with 5, 10, or 20 μM indicated inhibitor at room
temperature in RIA buffer for 15 min. Varying concentrations of 32P radiolabeled AP-DNA substrate (i.e., 5, 10, 25, 50, or 100 nM) were then added,
and the reactions were performed and analyzed as described in Experimental Methods. Shown are the Michaelis−Menten plots of V vs [S], reporting
the averages and standard deviations of at least seven data points for each value.
3107
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
in AP sites, providing evidence that these compounds do
indeed inhibit APE1 catalyzed repair of abasic damage in cells.
Next, we assessed the ability of these compounds to potentiate
the cytotoxic effects of MMS and TMZ in a dose−response
experiment using HeLa cells and monitoring cell viability using
a high-throughput luminescence-based detection of the cells’
ATP content, as shown in Figure 6. Both compounds greatly
potentiated the activity of these agents with optimal synergy
occurring at 5−10 μM for 3 and at 10−15 μM for 52. However,
compound 3 appears more cytotoxic against HeLa cells as a
single agent than analogue 52, with a 50% reduction in cell
viability occurring at ∼15 and ∼30 μM, respectively (see
Supporting Information Figure 2 for a semilog plot of these
data). Similar potentiation trends were observed when cell
viability was monitored through a more traditional method
using staining for live cells (Supporting Information Figure 3).
PK Properties of 3 and 52. Given that 3 and 52 exhibit a
tractable SAR, on-target APE1 inhibition, and potentiation of
the toxicity of MMS and TMZ in cell culture experiments, we
were eager to assess the pharmacokinetic (PK) properties of
these top compounds (Table 6). As shown in the in vitro
absorption, distribution, metabolism, and excretion (ADME)
data, both compounds 3 and 52 have many desirable attributes,
yet a few liabilities. Specifically, analogue 52 exhibits improved
kinetic solubility and Caco-2 permeability relative to the
original lead 3. Both compounds possess favorable cell
permeability and do not appear to be susceptible to active
transport as shown by the efflux ratios of ∼1. However,
compound 52 was found to be rapidly metabolized by mouse
liver microsomes (T1/2 = 7.8 min), whereas compound 3 shows
favorable stability (T1/2 = 80 min). These results suggest that
the general core scaffold is metabolically stable, and through
additional structural modifications, the potential metabolic
liability of 52 could be addressed while maintaining potency.
Surprisingly, as described below, we found 52 had a reasonable
PK profile despite the apparent metabolic liability. Finally, both
compounds exhibited some inhibition of CYP isozymes
(CYP2D6 and/or CYP3A4) at 10 μM. Compound 3, which
possesses substituted nitrogen (isopropyl group) and is a six-
membered ring rather than five-membered, shows no inhibition
of CYP2D6 at 10 μM. This result suggests that a more
expansive look at all the analogues described herein may
uncover structure−property relationships that obviate CYP
inhibition.
Figure 4. Stability of the APE1-DNA substrate complex in the
presence of 52 or 3. (A) Representative EMSA. APE1 (∼28 nM) was
incubated without inhibitor (“−”) or with the indicated inhibitor
concentration (1, 3, 10, 30, or 100 μM) for 10 min on ice. An amount
of 100 fmol of abasic DNA substrate (10 nM) was then added, and the
binding mixture was incubated on ice for an additional 5 min. Samples
were subjected to nondenaturing polyacrylamide gel electrophoresis to
separate the APE1-DNA complex (C) from unbound radiolabeled
DNA (DNA). Inh = inhibitor. (B) Relative complex formation without
(“0”) or with the indicated inhibitor (in μM). Shown is the average
and standard deviation of three independent experimental data points,
all relative to the APE1 control, without inhibitor.
the total number of AP sites relative to the DMSO control.
More notably, combined treatment of MMS with either of the
inhibitors resulted in a greater than additive increase (∼3-fold)
The two lead compounds (52 and 3) were analyzed for their
in vivo PK properties (Table 7) via intraperitoneal (ip)
administration at 30 mg/kg body weight in 6−8 week old CD1
mice. Both compounds were well tolerated by the animals, with
no adverse effects noted after a 24 h observation. Compound
52, the more hydrophilic analogue (CLogP ≈ 1), had a
favorable plasma t(1/2) of 5 h and a drug concentration (ng/mL)
that exceeded the IC50 for over 12 h. This compound also had
reasonable blood−brain barrier (BBB) penetration, with a high
initial concentration that quickly tailed off, resulting in a brain/
plasma (B/P) ratio of 1.4. In contrast, analogue 3, which is
more lipophilic (CLogP = 2.8), crosses the BBB quite readily,
giving rise to a B/P ratio of 21. This result correlates with
expectations, as reduction of hydrogen bond donors and
increased lipophilicity often lead to improved BBB penetration.
The ability to modulate the BBB penetration capacity of these
molecules through structural modifications could prove to be
useful depending on the cancer one is targeting.
Figure 5. AP site accumulation assay. AP site counts were measured in
HeLa cells after treatment with MMS and compounds alone and in
combination. To evaluate inhibition of APE1 by selected compounds
in cells, AP site formation was measured using the ARP assay in
triplicate (see Experimental Methods). The p values were determined
for an n using the Student’s t test, where 3, 52, and MMS are
compared with 3 and 52 alone (∗∗, p ≤ 0.001) and MMS alone (∗, p
≤ 0.01).
3108
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
Figure 6. HeLa cell viability assay with 3 or 52 alone, in combination with either MMS (0.4 mM) or TMZ (1 mM). (A) Combined treatment of the
HeLa cells with 3 at concentrations shown in the plot in the presence of 0.4 mM MMS and 1 mM TMZ. (B) Combined treatment of the HeLa cells
with 52 in the presence of 0.4 mM MMS and 1 mM TMZ (see Experimental Methods). The viability values are normalized to 100% for each
alkylating agent in the absence of compound. The assay was performed in triplicate.
a
Table 6. In Vitro ADME Properties of Lead Molecules (3 and 52)
mouse liver
microsome
stability, T1/2
(min)
PBS pH 7.4
stability: %
remaining after
48 h
aq kinetic sol.
(PBS at pH
7.4) (μM)
CYP2D6,
inh at 10
μM (%)
CYP3A4,
inh at 10
μM (%)
Caco-2, Papp
10−6 m/s at
pH 7.4
mouse plasma
stability, T1/2
(min)
compd
efflux ratio (B → A)/(A → B)
52
3
51.6
20.4
39
0
50
53
6.8
5
0.8
1.1
7.8
80
100
100
213
>250
a
All experiments were conducted by Shanghai Chempartner Co. Ltd., and values are the average of two experiments.
a
Table 7. In Vivo PK data
b
c
compd t1/2 (h) [plasma] t1/2 (h) [brain] [brain/plasma]
Cmax (μM) [plasma] Cmax (μM) [brain] tmax (h) [plasma] tmax (h) [brain] CLogP
52
3
5
2.5
1
1.35
21
36
16
92
0.25
0.25
0.25
0.25
1.02
2.83
2.1
217
a
Experiments were conducted at Pharmaron Inc. The ip administration (30 mpk) of CD1 male mice (n = 3) was monitored at eight time points
(0.25, 0.5, 1, 2, 4, 8, 12, 24 h). Compound 52 was formulated as 50% PEG 200 and 10% Cremophor EL in saline solution, and compound 3 was
b
formulated as 50% PEG 400 and 10% Cremophor EL in saline suspension. Calculated based on the average B/P ratio over eight time points (24 h
c
period). CLogP values were obtained using ChemDraw Ultra 10.0.
of APE1 inhibitors, such as 3 or other lipophilic analogues of
52, that efficiently cross the BBB and can potentially be used in
combination with a drug like TMZ. Our current efforts are
focused on defining the efficacy of this class of compounds in
vivo using mouse xenograft models in combination therapy
with TMZ and other relevant DNA-damaging cancer chemo-
therapeutics.
CONCLUSION
■
While the work described herein did not lead to a tremendous
improvement in the potency of the initial “hit” molecule 3, it
does provide valuable insights into the SAR profile of this
chemotype and represents the first reported medicinal
chemistry optimization campaign toward the establishment of
a novel APE1 inhibitor. In particular, this effort led to the
development of compounds with low single-digit micromolar
potency against the purified enzyme (noticeably more potent
than prior reported inhibitors), desirable in vitro and in vivo
ADME properties, and the capacity to potentiate the
cytotoxicity of relevant DNA-damaging agents, namely, MMS
and TMZ. On-target evidence was supported by the
comparable IC50 values against the purified recombinant
APE1 protein and human whole cell extracts, as well as by
increased genomic AP site accumulation in HeLa cells treated
with inhibitors alone. Analysis of the in vivo PK properties of
52 and 3 in mice revealed that analogue 52 has a better general
cytotoxicity profile, higher exposure levels, and a more
favorable t(1/2) in the plasma, whereas compound 3 crosses
the BBB more efficiently. Thus, for tumors outside the brain
cavity, a compound like 52, which does not efficiently cross the
BBB, would be useful in avoiding potential complications
associated with this vital organ. However, APE1 has been found
to be overexpressed in adult and pediatric gliomas, with an
increase in AP endonuclease activity of between 5- and 10-
fold.28 This observation indicates the need for the development
EXPERIMENTAL METHODS
■
General Chemistry. Unless otherwise stated, all reactions were
carried out under an atmosphere of dry argon or nitrogen in dried
glassware. Indicated reaction temperatures refer to those of the
reaction bath, while room temperature (rt) is noted as 25 °C. All
solvents were of anhydrous quality purchased from Sigma Chemical
Co. and used as received. Commercially available starting materials
and reagents were purchased from Sigma and were used as received.
Analytical thin layer chromatography (TLC) was performed with
Sigma Aldrich TLC plates (5 cm × 20 cm, 60 Å, 250 μm).
Visualization was accomplished by irradiation under a 254 nm UV
lamp. Chromatography on silica gel was performed using forced flow
(liquid) of the indicated solvent system on Biotage KP-Sil prepacked
cartridges and using the Biotage SP-1 automated chromatography
1
system. H and 13C NMR spectra were recorded on a Varian Inova
400 MHz spectrometer. Chemical shifts are reported in ppm with the
solvent resonance as the internal standard (CDCl3 7.26 ppm, 77.00
ppm, DMSO-d6 2.49 ppm, 39.51 ppm for 1H, 13C, respectively). Data
are reported as follows: chemical shift, multiplicity (s = singlet, d =
doublet, t = triplet, q = quartet, br = broad, m = multiplet), coupling
constants, and number of protons. Low resolution mass spectra
(electrospray ionization) were acquired on an Agilent Technologies
3109
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
6130 quadrupole spectrometer coupled to the HPLC system. High
resolution mass spectral data were collected in-house using an Agilent
6210 time-of-flight mass spectrometer, also coupled to an Agilent
Technologies 1200 series HPLC system. If needed, products were
purified via a Waters semipreparative HPLC instrument equipped with
a Phenomenex Luna C18 reverse phase (5 μm, 30 mm × 75 mm)
column having a flow rate of 45 mL/min. The mobile phase was a
mixture of acetonitrile (0.025% TFA) and H2O (0.05% TFA), and the
temperature was maintained at 50 °C.
Samples were analyzed for purity on an Agilent 1200 series LC/MS
instrument equipped with a Luna C18 reverse phase (3 μm, 3 mm ×
75 mm) column having a flow rate of 0.8−1.0 mL/min over a 7 min
gradient and a 8.5 min run time. Purity of final compounds was
determined to be >95%, using a 3 μL injection with quantitation by
AUC at 220 and 254 nm (Agilent diode array detector).
final volume. Following incubation on ice for 5 min, samples were
subjected to nondenaturing polyacrylamide gel electrophoresis (20
mM Tris, pH 7.5, 10 mM sodium acetate, 0.5 mM EDTA, 8%
polyacrylamide, 2.5% glycerol) for 2 h at 120 V in electrophoresis
buffer (20 mM Tris, pH 7.5, 10 mM sodium acetate, 0.5 mM EDTA)
to separate the APE1-DNA complex from unbound radiolabeled
DNA.31 After electrophoresis, the gel was subjected to standard
phosphoimager analysis as above, and the percentage of substrate
DNA in complex with APE1 was determined.
Genomic AP Site Accumulation in Cells. HeLa cells with 80%
confluency in a 25 cm2 flask were treated with DMSO, 275 μM MMS,
or 7.5 μM APE1 inhibitor alone or with a combination of 275 μM
MMS and 7.5 μM inhibitor for 24 h at 37 °C. Cells were then
harvested, and genomic DNA of each sample was isolated according to
Qiagen Genomic DNA isolation kit. The concentration of genomic
DNA was measured and adjusted to 100 ng/μL. An amount of 10 μL
of purified DNA was further labeled with an aldehyde reactive probe
(ARP) reagent (N′-aminooxymethylcarbonylhydrazino-D-biotin), and
AP sites were measured using the DNA damage quantification kit from
Dojindo Molecular Technologies.
MMS and TMZ Potentiation Assay. HeLa cells were plated by
multichannel pipet or Multidrop Combi dispenser (Thermo) at 6K/25
μL per well in DMEM culture medium with 10% FBS into white solid
bottom 384-well cell culture plates. Cells were cultured at 37 °C
overnight to allow for cell attachment. The following day, the entire
cell medium in the well was replaced with fresh medium containing
serial dilutions of the compounds of interest (5−30 μM) in the
presence or absence of MMS (0.4 mM) or TMZ (1 mM). The plates
were incubated for 24 h at 37 °C. Cell viability was then evaluated via
luminescence detection by adding 15 μL of CellTiter Glo reagent
(Promega, Madison, WI) to each well and incubating at room
temperature for 30 min and subsequently measuring the luminescence
using a ViewLux reader. Percent viability was calculated for each
concentration of the tested compounds in duplicate relative to the
luminescence of the negative DMSO control.
In Vivo PK Analysis. Compound 3 was dissolved in PEG 400 and
Cremophor with vortexing and sonification. Then saline was gradually
added with vortexing and sonification to obtain a final concentration of
3 mg/mL 3 in 50% PEG 400 and 10% Cremophor. Compound 52 was
dissolved in PEG 200 Cremophor with vortexing and sonification.
Then saline was gradually added as above to obtain a final
concentration of 3 mg/mL 52 in 50% PEG 200 and 10% Cremophor.
The dose for both compounds was administered ip. All blood samples
were collected through a cardiac puncture per sampling time point
(0.25, 0.5, 1, 2, 4, 8, 12, and 24 h postdose). Approximately 0.12 mL of
blood was collected at each time point. All blood samples were
transferred into plastic microcentrifuge tubes containing heparin and
placed at −80 °C until processed (see below). At each time point (see
above), the brain was harvested immediately after euthanasia by
carbon dioxide. The brain was rinsed with saline and wiped clean and
then weighed in a sterilized plastic tube. The tissue sample was then
homogenized in water with a brain weight (g) to water (mL) ratio of
1:4 (g:mL). The detected values were then multiplied by 5 to achieve
the final concentration of the compound in the brain. Blood samples
were processed for plasma by centrifugation at 4 °C at 4000g for 5
min. Plasma samples were then stored in tubes, quickly frozen in a
freezer, and kept at −80 °C until LC/MS/MS analysis. Plasma
concentration of compound 3 or 52 at the various time points (data
obtained from the LC/MS/MS studies) was analyzed using the
WinNonlin software program.
RIA. Recombinant wild type APE1 protein was purified as
previously described.29 An amount of 50 pg of APE1 (∼140 pM)
was incubated without (positive control containing 1% DMSO) or
with the indicated inhibitor at the indicated concentration at room
temperature in RIA buffer (50 mM Tris, pH 7.5, 25 mM NaCl, 1 mM
MgCl2, 1 mM DTT, 0.01% Tween-20) for 15 min. One-half pmol of
32P 5′-radiolabeled AP-DNA substrate (18 mer) was added to a 10 μL
final volume,30 and the mixtures were incubated at 37 °C for 5 min
and the reactions stopped by adding stop buffer (0.05% bromophenol
blue/xylene cyanol dissolved in 95% formamide, 20 mM EDTA) and
heating at 95 °C for 10 min. Intact substrate was separated from
incised product on a 15% polyacrylamide denaturing gel in a Tris/
boric acid/EDTA buffer. Following electrophoresis, the gel was
subjected to standard phosphoimager analysis using the ImageQuant
5.2 software, and the percent incision activity (amount of substrate
converted to product) was calculated. IC50 values (i.e., the
concentration of inhibitor at which 50% inactivation was observed)
were extrapolated after plotting the results of duplicate incision sets (at
inhibitor concentrations ranging from 1 μM to 100 μM) and fitting
data using the Hill equation in GraphPad Prism, version 4.0, software.
HeLa Whole Cell Extract Incision Assays. To prepare protein
extracts, HeLa cells maintained in DMEM with 10% fetal bovine
serum and 1% penicillin−streptomycin were harvested, washed with
1× PBS, and resuspended in ice cold hypotonic lysis buffer (50 mM
Tris, pH 7.4, 1 mM EDTA, 1 mM DTT, 10% glycerol, 0.5 mM
PMSF). The suspension was frozen at −80 °C for at least 30 min and
then slowly thawed at 4 °C for ∼1 h. KCl was added to the cell
suspension to a final concentration of 222 mM, followed by incubation
on ice for 30 min and clarification by centrifugation at 12000g for 15
min at 4 °C. The supernatant (whole cell extract) was retained and the
protein concentration determined using the Bio-Rad Bradford reagent.
Aliquots were stored until needed at −80 °C. For the incision assays,
300 ng of HeLa whole cell extract was incubated with 0, 1, 3, 10, 30, or
100 μM indicated inhibitor at room temperature for 15 min prior to
the addition of 0.5 pmol of 32P radiolabeled AP-DNA substrate (final
volume of 10 μL). The reaction mix was then transferred to 37 °C for
5 min to allow for incision. Following addition of stop buffer and heat
denaturation, the reaction products were analyzed as indicated above.
Enzyme Kinetic Studies. An amount of 10 pg of APE1 (∼28 pM)
was incubated without (positive control, 1% DMSO) or with 5, 10, or
20 μM indicated inhibitor at room temperature in RIA buffer (see
above) for 15 min. Varying concentrations of 32P radiolabeled AP-
DNA substrate (i.e., 5, 10, 25, 50, or 100 nM) were then added to a 10
μL final volume. The mixtures were incubated at 37 °C for 5 min and
the reactions stopped by adding stop buffer and heating at 95 °C for
10 min. The reaction velocity (nanomolar substrate incised per
minute) at each substrate concentration was calculated as described
above. Lineweaver − Burk plots of 1/V versus 1/[S] were used to
determine KM and kcat and the mode of inhibition.
ASSOCIATED CONTENT
EMSA. An amount of 10 ng of APE1 (∼28 nM) was incubated
without inhibitor (positive control, 1% DMSO) or with increasing
concentrations of inhibitor (1, 3, 10, 30, and 100 μM) in binding
buffer (50 mM Tris, pH 7.5, 25 mM NaCl, 1 mM EDTA, 1 mM DTT,
10% glycerol, 0.01% Tween 20) for 10 min on ice, and then
radiolabeled 32P AP-DNA substrate (100 fmol) was added to a 10 μL
■
S
* Supporting Information
Additional experimental procedures and 1H, LC/MS, and
HRMS data for representative compounds. This material is
available free of charge via the Internet at http://pubs.acs.org.
3110
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
(10) Mutter, N.; Stupp, R. Temozolomide: a milestone in neuro-
oncology and beyond? Expert Rev. Anticancer Ther. 2006, 6, 1187−
1204.
(11) Rouleau, M.; Patel, A.; Hendzel, M. J.; Kaufmann, S. H.; Poirier,
G. G. PARP inhibition: PARP1 and beyond. Nat. Rev. Cancer 2010, 10,
293−301.
AUTHOR INFORMATION
■
Corresponding Author
*For D.J.M.: phone, 301-217-4381; fax, 301-217-5736; e-mail,
maloneyd@mail.nih.gov. For D.M.W.: phone, 410-558-8153;
fax, 410-558-8157; e-mail, wilsonda@mail.nih.gov.
(12) Petermann, E.; Helleday, T. Pathways of mammalian replication
fork restart. Nat. Rev. Mol. Cell Biol. 2010, 11, 683−687.
(13) (a) Bryant, H. E.; Schultz, N.; Thomas, H. D.; Parker, K. M.;
Flower, D.; Lopez, E.; Kyle, S.; Meuth, M.; Curtin, N. J.; Helleday, T.
Specific killing of BRCA2-deficient tumors with inhibitors of
poly(ADP-ribose) polymerase. Nature 2005, 434, 913−917. (b) Farm-
er, H.; McCabe, N.; Lord, C. J.; Tutt, A. N. J.; Johnson, D. A.;
Richardson, T. B.; Santarosa, M.; Dillon, K. J.; Hickson, I.; Knights, C.;
Martin, N. M. B.; Jackson, S. P.; Smith, G. C. M.; Ashworth, A.
Targeting the DNA repair defect in BRCA mutant cells as a therapeutic
strategy. Nature 2005, 434, 917−921.
Author Contributions
§These authors contributed equally to this work.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
The authors thank William Leister, Paul Shinn, Danielle
VanLeer, James Bougie, and Tom Daniel for assistance with
compound management and purification. We also thank
Christina Greco for critical reading of the manuscript and
technical assistance. This research was supported by the
Intramural Research Program of the NIH, National Institute
on Aging, as well as by the NIH Grant R03 MH086444-01
(D.M.W.), and the Molecular Libraries Initiative of the
National Institutes of Health Roadmap for Medical Research.
(14) Guha, M. PARP inhibitors stumble in breast cancer. Nat.
Biotechnol. 2011, 29, 373−374.
(15) Wilson, D. M. III; Simeonov, A. Small molecule inhibitors of
DNA repair nuclease activities of APE1. Cell. Mol. Life Sci. 2010, 67,
3621−3631.
(16) Bapat, A.; Glass, L. S.; Luo, M.; Fishel, M. L.; Long, E. C.;
Georgiadis, M. M.; Kelley, M. R. Novel small molecule inhibitor of
APE1 endonuclease blocks proliferation and reduces viability of
glioblastoma cells. J. Pharmacol. Exp. Ther. 2010, 334, 988−998.
(17) Madhusudan, S.; Smart, F.; Shrimpton, P.; Parsons, J. L.;
Gardiner, L.; Houlbrook, S.; Talbot, D.; Hammonds, T.; Freemont, P.
A.; Sternberg, M. J. E.; Dianov, G. L.; Hickson, I. D. Isolation of a
small molecule inhibitor of DNA base excision repair. Nucleic Acids Res.
2005, 33, 4711−4724.
ABBREVIATIONS USED
■
APE1, apurinic/apyrimidinic endonuclease 1; HTS, high-
throughput screening; qHTS, quantitative high-througput
screening; MLSMR, Molecular Libraries Small Molecule
Repository; BER, base excision repair; ML199, N-(3-(benzo-
[d]thiazol-2-yl)-5,6-dihydro-4H-thieno[2,3-c]pyrrol-2-yl)-
acetamide; TMZ, temozolomide; RIA, radiotracer incision
assay; ADME, absorption, distribution, metabolism, and
excretion; dppp, 1,3-bis(diphenylphosphino)propane; xant-
phos, 4,5-bis(diphenylphosphino)-9,9-dimethylxantene; dba,
d i b e n z y l i d e n e a c e t o n e ; E D C , 1 - e t h y l - 3 - ( 3 -
dimethylaminopropyl)carbodiimide; HATU, 2-(1H-7-azaben-
zotriazol-1-yl)-1,1,3,3-tetramethyluronoium hexafluorphos-
phate; MMS, methylmethane sulfonate
(18) Fishel, M. L.; Kelley, M. R. The DNA. Base excision repair
protein APE1/Ref-1 as a therapeutic and chemopreventive target. Mol.
Aspects Med. 2007, 28, 375−395.
(19) Mohammed, M. Z.; Vyjayanti, V. N.; Laughton, C. A.; Dekker,
L. V.; Fischer, P. M.; Wilson, D. M. III; Abbotts, R.; Shah, S.; Patel, P.
M.; Hickson, I. D.; Madhusudan, S. Br. J. Cancer 2011, 1−11.
(20) Zawahir, Z.; Dayam, R.; Deng, J.; Pereira, C.; Neamati, N.
Pharmacophore guided discovery of small-molecule human apurinic/
apyrimidinic endonuclease APE1. J. Med. Chem. 2009, 52, 20−32.
(21) (a) Bases, R. E.; Mendez, F. Topoisomerase inhibition by
lucanthone, an adjuvant in radiation therapy. Int. J. Radiat. Oncol., Biol.,
Phys. 1997, 37, 1133−1137. (b) Luo, M.; Kelley, M. R. Inhibition of
the human apurinic/apyrimidinic endonuclease (APE1) repair activity
and sensitization of breast cancer cells to DNA alkylating agents with
lucanthone. Anticancer Res. 2004, 24, 2127−2134.
REFERENCES
■
(1) Wilson, D. M. III; Barsky, D. The major human abasic
endonuclease: formation, consequences and repair of abasic lesions in
DNA. Mutat. Res. 2001, 485, 283−307.
(22) (a) Taverna, P.; Liu, L.; Hwang, H. S.; Hanson, A. J.; Kinsella, T.
J.; Gerson, S. L. Methoxyamine potentiates DNA single strand breaks
and double strand breaks induced by temozolomide in colon cancer
cells. Mutat. Res. 2001, 485, 269−281. (b) Fishel, M. L.; He, Y.; Smith,
M. L.; Kelley, M. R. Manipulation of base excision repair to sensitize
ovarian cancer cells to alkylating agent temozolomide. Clin. Cancer Res.
2007, 13, 260−267.
(23) Seiple, L. A.; Cardellina, J. H. 2nd; Akee, R.; Stivers, J. T. Potent
inhibition of human apurinic/apyrimidinic endonuclease 1 by
arylstilbonic acids. Mol. Pharmacol. 2008, 73, 669−677.
(24) Simeonov, A.; Kulkarni, A.; Dorjsuren, D.; Jadhav, A.; Shen, M.;
McNeill, D. R.; Austin, C. P.; Wilson, D. M. 3rd. Identification and
characterization of inhibitors of human apurinic/apyrimidinic
endonuclease APE1. PLoS One 2009, 4, e5740.
(25) These compounds are a part of the NIH Small Molecule
Repository; see http://mli.nih.gov/mli/compound-repository/mlsmr-
compounds/.
(2) Demple, B.; Sung, J. S. Molecular and biological roles of Ape1
protein in mammalian base excision repair. DNA Repair 2005, 4,
1442−1449.
(3) Robertson, A. B.; Klungland, A.; Rognes, T.; Leiros, I. DNA
repair in mammalian cells: base excision repair: the long and short of
it. Cell. Mol. Life Sci. 2009, 66, 981−993.
(4) Wilson, D. M. III. Processing of nonconventional DNA strand
break ends. Environ. Mol. Mutagen. 2007, 48, 772−782.
(5) Tell, G.; Quadrifoglio, F.; Tiribelli, C.; Kelley, M. R. The many
functions of APE1/Ref-1: not only a DNA repair enzyme. Antioxid.
Redox. Signaling 2009, 11, 601−620.
(6) Bhakat, K. K.; Mantha, A. K.; Mitra, S. Transcriptional regulatory
functions of mammalian AP-endonuclease (APE1/Ref-1), an essential
multifunctional protein. Antioxid. Redox. Signaling 2009, 11, 621−638.
(7) Luo, M.; He, H.; Kelley, M. R.; Georgiadis, M. M. Redox
regulation of DNA repair: implications for human health and cancer
therapeutic development. Antioxid. Redox. Signaling 2010, 12, 1247−
1269.
(8) Bapat, A.; Fishel, M. L.; Kelley, M. R. Going ape as an approach
to cancer therapeutics. Antioxid. Redox. Signaling 2009, 11, 651−668.
(9) Abbotts, R.; Madhusudan, S. Human AP endonuclease 1 (APE1):
from mechanistic insights to druggable target in cancer. Cancer Treat.
Rev. 2010, 36, 425−435.
(26) (a) Gewald, K.; Schinke, E.; Boettcher, H. 2-Amino-thiophene
aus methylenaktiven nitrile carbonylverbindiungen und schwefel.
Chem. Ber. 1966, 99, 94−100. (b) Andersen, H. S.; Olsen, O. H.;
Iversen, L. F.; Sorensen, A. L. P.; Mortensen, S. B.; Christensen, M. S.;
Branner, S.; Hansen, T.; Lau, J. F.; Jeppesen, L.; Moran, E. J.; Su, J.;
Bakir, F.; Judge, L.; Shahbaz, M.; Collins, T.; Vo, T.; newman, M. J.;
3111
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112

Journal of Medicinal Chemistry
Article
Ripka, W. C.; Moller, N. P. H. Discovery and SAR of a novel selective
and orally bioavailable nonpeptide classical competitive inhibitor class
of protein-tyrosine phosphatase 1B. J. Med. Chem. 2002, 45, 4443−
4459.
(27) Demple, B.; Harrison, L Repair of oxidative damage to DNA:
enzymology and biology. Annu. Rev. Biochem. 1994, 63, 915−948.
(28) Bobola, M. S.; Blank, A.; Berger, M. S.; Stevens, B. A.; Silber, J.
R. Apurinic/apyrimidinic endonuclease activity is elevated in human
adult gliomas. Clin. Cancer Res. 2001, 7, 3510−3518.
(29) Erzberger, J. P.; Barsky, D.; Scharer, O. D.; Colvin, M. E.;
Wilson, D. M. III. Elements in abasic site recognition by the major
human and Escherichia coli apurinic/apyrimidinic endonucleases.
Nucleic Acids Res. 1998, 26, 2771−2778.
(30) Wilson, D. M. III; Takeshita, M.; Grollman, A. P.; Demple, B.
Incision activity of human apurinic endonuclease (Ape) at abasic site
analogs in DNA. J. Biol. Chem. 1995, 270, 16002−16007.
(31) Wilson, D. M. III; Takeshita, M.; Demple, B. Abasic site
binding by the human apurinic endonuclease, Ape, and determination
of the DNA contact sites. Nucleic Acids Res. 1997, 25, 933−939.
3112
dx.doi.org/10.1021/jm201537d | J. Med. Chem. 2012, 55, 3101−3112