Phenotypic screens, which focus on measuring and quantifying discrete cellular changes rather than affinity for individual recombinant proteins, have recently attracted renewed interest as an efficient strategy for drug discovery. In this article, we describe the discovery of a new chemical probe, bisamide (CCT251236), identified using an unbiased phenotypic screen to detect inhibitors of the HSF1 stress pathway. The chemical probe is orally bioavailable and displays efficacy in a human ovarian carcinoma xenograft model. By developing cell-based SAR and using chemical proteomics, we identified pirin as a high affinity molecular target, which was confirmed by SPR and crystallography.
Phenotypic screens, which focus on measuring and quantifying discrete cellular changes rather than affinity for individual recombinant proteins, have recently attracted renewed interest as an efficient strategy for drug discovery. In this article, we describe the discovery of a new chemical probe, bisamide (CCT251236), identified using an unbiased phenotypic screen to detect inhibitors of the HSF1 stress pathway. The chemical probe is orally bioavailable and displays efficacy in a human ovarian carcinoma xenograft model. By developing cell-based SAR and using chemical proteomics, we identified pirin as a high affinity molecular target, which was confirmed by SPR and crystallography.
Despite the recent
extraordinary progress seen in cancer therapy
using molecularly targeted drugs, the disease commonly remains resistant
to effective long-term treatment. Even when excellent responses to
drugs are initially observed, resistance is almost inevitable and
patients are then left with few treatment options[1] as the discovery of targeted therapies in oncology has
focused on relatively few protein families.[2] To break this cycle and expand the treatment options for cancer
patients, new approaches are needed to discover novel druggable protein
targets.[3]In phenotypic screens,
small molecules undergo high-throughput
screening against intact cells, rather than recombinant proteins,
and discrete phenotypic changes in the cell are measured and quantified.[4] Interest in phenotypic screens has increased
significantly in recent years due to their potential to effectively
discover new drugs.[5] Phenotypic screens
have several advantages over screens using recombinant proteins. Hits
from a phenotypic screen will, by definition, be cell permeable and
have cellular activity, potentially reducing optimization cycles and
timelines. Also, because the screening approach is unbiased, established
knowledge of the biology of molecular targets is not required. Finally,
polypharmacology is often observed with small molecules and structurally
related protein families; this can be crucial for efficacy and is
perfectly compatible with a phenotypic screening approach.[6]In contrast, progressing hits from a phenotypic
screen can generate
a number of unique challenges. Discovering pharmacodynamic (PD) biomarkers
in vivo for use in animals can be difficult when developing hits from
phenotypic screens, as the pathways commonly need to be activated
with an external stimulus.[7] Cell-based
screens are typically more expensive and time-consuming and so may
require a greater commitment prior to the screening campaign.[8] Furthermore, molecular target identification,
deconvolution, and validation are crucial steps if new chemical probes[9] and drugs are to be discovered. These are often
a bottleneck in phenotypic screening.[10] Even with successful target deconvolution, the target discovered
may not be of interest; for example, the target may already be drugged
or be a known antitarget. Finally, polypharmacology may be a serious
impediment to compound progression because the interaction with multiple
structurally related protein targets may prove impossible to deconvolute.[11] These challenges alter the balance between prioritizing
druglike properties of compounds and an efficient target identification
strategy.[12]To execute a successful
phenotypic screening campaign, it is critical
to select an appropriate phenotype for small molecule intervention.
HSF1 is a transcription factor and the master regulator of the ancient,
canonical heat shock response.[13] A large
body of work has verified the importance of HSF1 to tumorigenesis
and cancer progression.[14] HSF1 has been
proposed to be activated by various elements of the cancer state,
potentially reprogramming the transcriptome in a way that is overlapping
with, but distinct from, the heat shock response.[15] Also, a strong correlation has been reported between the
expression of activated HSF1 in tumors and adverse clinical outcomes.[16] This evidence indicates that the inhibition
of HSF1-mediated transcription could be a viable strategy in cancer
treatment.[17] Moreover, inhibiting the HSF1
stress pathway would represent an attempt at targeting non-oncogene
addiction and proteotoxic stress, which has been proposed to be advantageous.[18] However, HSF1 is a ligandless transcription
factor and so is unlikely to be amenable to standard drug discovery
strategies and direct inhibition with small molecules. Therefore,
we proposed that an inhibitor of HSF1-mediated transcription, which
antagonized the HSF1 pathway but without necessarily binding directly
to HSF1, could be discovered and developed via a cell-based phenotypic
screen.
Results
HSF1 Phenotypic Assay
To observe
HSF1-mediated transcription
in an in vitro setting, the HSF1 pathway is activated by a validated
heat shock protein 90 (HSP90) inhibitor,[19] or another form of external stress,[20] which initiates the heat shock response. Commonly, the output of
the heat-shock response is quantified by measuring the induction of
heat shock 70 kDa protein 1 (HSP72) expression, the stress-inducible
HSP70 isoform.[19] HSF1 pathway inhibitors
are then defined by their ability to block the induction of HSP72.
Several HSF1-mediated HSP72 induction inhibitors have been discovered
via this method with different proposed molecular mechanisms of action
(Figure ).[21,28]
Figure 1
Inhibitors
of HSF1-mediated HSP72 induction.
Inhibitors
of HSF1-mediated HSP72 induction.With the aim of discovering novel hit-matter that inhibits
HSF1-mediated
transcription, we previously carried out a cell-based high-throughput
phenotypic screen in U2OS human osteosarcoma cells of ∼200000
compounds, including ∼35000 compounds from a kinase-focused
library.[22] The screen quantified the inhibition
of HSP72 induction using the Arrayscan assay[28] following treatment with the HSP90 inhibitor tanespimycin (17-AAG).[23]
The Bisamide Series
Using this screen,
we identified
a potent hit from the kinase-focused deck, bisamide 1 (CCT245232) (Figure ).[24] Following resynthesis, the screening
hit was confirmed and displayed a pIC50 = 8.55 ± 0.09
(IC50 = 2.8 nM, n = 49)[25] in our HSP72 cell-based enzyme-linked immunosorbent assay
(ELISA) in U2OS cells. The HSP72 cell-based ELISA assay is an alternative
assay format to the Arrayscan assay for quantifying the induced expression
of HSP72 and was used as our primary phenotypic pathway assay throughout
the study.[26] The IC50 was defined
as the concentration that inhibited the signal to 50% of the 17-AAG
(250 nM) induced HSP72 expression, relative to the control 17-AAG
alone (see Supporting Information for details).
The clear structural feature defining this chemotype was the N,N′-4-methyl-1,3-phenylenediamide
core (Figure ).
Figure 2
HSF1 pathway
inhibitor, bisamide 1.
HSF1 pathway
inhibitor, bisamide 1.We had previously demonstrated that pan cyclin-dependent
kinase
(CDK) inhibitors, and potent CDK9 inhibitors in particular that act
as transcription antagonists,[27] can inhibit
the HSF1-mediated HSP72 induction phenotype; we initially suspected
a similar mechanism for this chemotype.[28] However, upon biochemical screening of bisamide 1 against
CDK2 and CDK9, no inhibition was observed (<10% inhibition at 1
μM, see Supporting Information).
Therefore, we hypothesized that bisamide 1 was acting
through a different mechanism of action and consequently 1 was submitted for further characterization.
Hit Characterization: Kinase
Activity
HSF1 is regulated
by multiple post-translational phosphorylations,[29] so we hypothesized that kinase inhibition, other than CDK2
and CDK9, was causing the observed HSF1-mediated HSP72 induction inhibition.
Bisamide 1 was screened against a broad kinase set using
the KINOMEscan biochemical screening platform, a binding assay which
gave single-point percentage inhibition values at 1 μM (DiscoverX, http://www.discoverx.com).[30] Analysis of this data set for 442 kinases revealed
that bisamide 1 inhibited only nine kinases >50% and
four kinases >90% (see Supporting Information for details). To validate the kinase hits, orthogonal functional
assays were carried out on the four kinases that displayed >90%
inhibition.
Of these, KIT, PDGFRA, and PDGFRB failed to confirm in their respective
functional assays, all returning pIC50 < 5 (IC50 > 10000 nM). Bisamide 1 did display modest activity
against BRAF, returning a pIC50 = 6.38 (IC50 = 420 nM, n = 1).[31] Although
it seemed unlikely that the modest biochemical BRAF activity would
translate into cellular activity, seven commercially available[32] potent BRAF inhibitors of differing chemotypes
were tested in our cellular HSF1-mediated HSP72 induction inhibition
assay for comparison and none displayed pIC50 > 5 (IC50 < 10000 nM).[33] This indicated
that BRAF inhibition was not an important contributor to the observed
HSF1 transcription inhibition phenotype (see Supporting
Information).
Hit Characterization: Cellular
To
establish whether
bisamide 1 displays cellular activity without activation
of the heat shock response using the HSP90 inhibitor 17-AAG, the growth
inhibitory effects were assayed in the U2OS cell line using the CellTiter
Blue (CTB) assay (Promega). Following 4 days of treatment, bisamide 1 displayed a highly potent pGI50 = 7.74 ±
0.08 (GI50 = 18 nM, n = 13), when compared
to vehicle. To assess the broader single agent cellular activity of 1, the compound was assayed for growth inhibition against
a large, genetically diverse panel of human cancer cell lines (Genomics
of Drug Sensitivity in Cancer, www.cancerrxgene.org).[34] Of the
635 cell lines assayed, 628 (99%) displayed a pGI50 >
6
(GI50 < 1000 nM), 455 (72%) a pGI50 >
7 (GI50 < 100 nM), and 18 (2.8%) a pGI50 >
8 (GI50 < 10 nM), with no clear selectivity against
any specific
tissue type (see Supporting Information for details). These results suggested that bisamide 1 had wide-ranging anticancer activity and that activation of the
HSF1 pathway with an HSP90 inhibitor was not required for 1 to inhibit cancer cell growth.
Cell-Based SAR
Given the complexity of factors underlying
cell-based structure-activity relationships (SAR), we focused on matched
pair changes to establish which features were crucial for the cellular
activity of the bisamide chemotype. There were two primary aims to
our medicinal chemistry strategy. First, because our biochemical kinase
screening had failed to reveal any potential kinase targets, a broader
target identification strategy would need to be developed. Second,
although bisamide 1 had excellent cellular potency, the
compound had poor aqueous solubility (kinetic solubility <2 μM),
which would need to be addressed to validate HSF1 pathway inhibition
as an anticancer strategy in an in vivo human tumor xenograft model.
Both goals required us to improve our understanding of the cellular
SAR, particularly to identify a vector-to-solvent through which either
a chemical probe linker or a solubilizing group could be attached,
without disrupting the ability of the compound to bind to the primary
pharmacological target or targets.To develop the cell-based
SAR, we switched to the human ovarian carcinoma cell line SK-OV-3,
which was relatively sensitive to growth inhibition and is commonly
used in translational drug discovery both in vitro[35] and in vivo.[36] When the activity
of bisamide 1 in the SK-OV-3 cell line was assayed, it
gave a pIC50 = 7.17 ± 0.07 (IC50 = 68 nM, n = 4) using the HSP72 cell-based ELISA assay induced with
250 nM 17-AAG. Bisamide 1 also inhibited the proliferation
of SK-OV-3 cells in the CTB assay with a pGI50 = 8.08 ±
0.12 (GI50 = 8.4 nM, n = 12).We
first focused the exploration on the benzodioxane motif (Scheme ). The left-hand
ring system analogues (Table , entries 1–6) were synthesized via a three-step procedure
from the commercially available 2-methyl-5-nitroaniline 2. Following amide bond formation, through the reaction of 2-methyl-5-nitroaniline 2 with 2-methylquinoline-6-carboxylic acid 3 and
subsequent reduction of the nitro-group using iron, the second amide
bond was generated using the corresponding carboxylic acid in a HATU-mediated
coupling to afford the various benzodioxane analogues in good to moderate
yields.
Scheme 1
Synthesis of the Benzodioxane Bisamide Replacements
Reagents and conditions: (i)
oxalyl chloride, DMF, DCM, then, pyridine, DCM; (ii) Fe, NH4Cl, EtOH/H2O; (iii) RPhCO2H, HATU, DIPEA, DMF.
Table 1
Bisamide Analogues
All results
are quoted as the geometric
mean ± SEM, pIC50/pGI50/pKD = −log IC50/GI50/KD (M). The number of repeats, n, are described in parentheses. ND = not determined.
Cell-based ELISA assay for inhibition
of HSP72 induction; SK-OV-3 cells were pretreated with compound at
the relevant concentration for 1 h before the addition of 250 nM 17-AAG.
HSP72 levels were then quantified after 18 h.
Growth inhibition was measured after
96 h of treatment and compared to vehicle control.
pKD values
were measured through analysis of the sensorgram at equilibrium where
possible. Values were then fitted to a one-site specific binding model
using Graphpad Prism Version 6.
Synthesis of the Benzodioxane Bisamide Replacements
Reagents and conditions: (i)
oxalyl chloride, DMF, DCM, then, pyridine, DCM; (ii) Fe, NH4Cl, EtOH/H2O; (iii) RPhCO2H, HATU, DIPEA, DMF.All results
are quoted as the geometric
mean ± SEM, pIC50/pGI50/pKD = −log IC50/GI50/KD (M). The number of repeats, n, are described in parentheses. ND = not determined.Cell-based ELISA assay for inhibition
of HSP72 induction; SK-OV-3 cells were pretreated with compound at
the relevant concentration for 1 h before the addition of 250 nM 17-AAG.
HSP72 levels were then quantified after 18 h.Growth inhibition was measured after
96 h of treatment and compared to vehicle control.pKD values
were measured through analysis of the sensorgram at equilibrium where
possible. Values were then fitted to a one-site specific binding model
using Graphpad Prism Version 6.Changing the 5-substituted dioxane isomer 1 (Table , entry 1) to the
6-substituted dioxane isomer 6 (Table , entry 2) resulted in a complete loss of
cellular activity (pIC50 < 5), indicating that the isomer
of the benzodioxane was crucial. The acyclic dimethoxy analogue 7 (Table ,
entry 3) also displayed a complete loss of cellular activity. Next,
we investigated the role of the two oxygen atoms of the dioxane ring.
Removing the para-oxygen of the dioxane to give the meta-chroman 8 (Table , entry 4) resulted in a modest 5-fold decrease
in cellular HSF1-mediated HSP72 induction inhibition and a 7-fold
decrease in growth inhibition compared to 1; while the para-chroman isomer 9 (Table , entry 5) displayed a further 6-fold decrease
in HSF1-mediated HSP72 induction inhibition and an 8-fold decrease
in growth inhibition compared to 8, which represents
a 37- and 51-fold decrease in cellular activity respectively compared
to the original hit, bisamide 1. Surprisingly, isochroman 10 (Table , entry 6) displayed no measurable cellular activity. From these
results, it was clear that the benzodioxane moiety was crucial for
the cellular activity of the bisamide series. Because the SAR surrounding
the benzodioxane was steep and complex, we decided that this region
was unlikely to be solvent exposed and so would be incompatible with
linker and solubilizing group attachment.Second, we investigated
the role of the central ring and amide
moieties in the cellular activity of the bisamide chemotype. We hypothesized
that the amide groups could play an important role in hydrogen bonding
with a potential target and in controlling the overall shape of the
ligand, while substitution at the central ring could lead to the discovery
of solvent-exposed vectors. The synthesis of these analogues (Table , entries 7–10)
was similar to that described in Scheme . Substitution at both the 6-position (Table , entry 7) and the
4-position (Table , entry 8) of the central benzene ring resulted in a complete loss
of cellular activity, suggesting that neither of these vectors represented
a viable route to solvent. Owing to the potential for the toluene
methyl group to affect the conformation of the right-hand amide, we
decided to leave this group unchanged. N-Methyl substitution
of the left-hand amide (Table , entry 9) and an attempted sulfonamide bioisosteric replacement
(Table , entry 10)
also resulted in a complete loss of cellular activity. Again, owing
to the steep SAR in this region of the chemotype, it was clearly incompatible
with linker and solubilizing group attachment.Finally, we examined
the role of the 2-methylquinoline ring in
the cellular activity of the bisamide series. All analogues (Table , entries 11–13)
were synthesized via a method similar to that described in Scheme . First, we removed
the 2-methyl group, which is both lipophilic and has the potential
to be a weak hydrogen bond donor, to give quinoline 15 (Table , entry 11).
Removal of this group resulted in no significant change in the cellular
activity when compared to 1 (Table , entry 1). Next we moved the quinoline nitrogen
from the 1- to the 3-position to give isoquinoline 16 (Table , entry 12).
Again, no significant change in cellular activity was observed when
compared to bisamide 1. Finally, we partially reduced
the quinoline ring to give tetrahydroquinoline 17 (Table , entry 13). By changing
to tetrahydroquinoline 17, the nitrogen atom now acts
as a hydrogen bond donor rather than an acceptor. Despite this reversal
in anticipated binding properties, only a 3.5-fold decrease in HSF1-mediated
HSP72 induction inhibition and no significant change in antiproliferative
activity was observed when compared to bisamide 1. The
broad SAR of the quinoline region of the molecule suggested that this
moiety is solvent exposed and so could be exploited for solubilizing
group and linker attachment.
Exploiting Vectors-to-Solvent: Solubilizing
Group Optimization
Owing to the complexities of the cellular
SAR, attaching a solubilizing
group to the solvent-exposed region of the ligand was considered a
more expeditious strategy to generate an in vivo chemical probe suitable
for use in animal models rather than attempting to optimize the intrinsic
solubility of the bisamide series without insight from structural
information. To address this objective, we adopted an iterative strategy
to exploit the rapid synthesis of analogues that would exhibit improved
physicochemical properties and demonstrate the appropriate mouse pharmacokinetic
(PK) parameters for in vivo study (Scheme ).
Scheme 2
Synthesis of Solubilizing Group Analogues
Reagents and conditions: (i)
oxalyl chloride, DMF, 1,4-benzodioxane-6-carboxylic acid, DCM, then,
4-methyl-3-nitroaniline, pyridine, DCM, RT, then Pd/C (10%), H2 (1 atm), EtOH; (ii) NaH, RCH2OH, THF, 0 °C
to reflux; (iii) n-BuLi, CO2(s), THF,
−78 °C; (iv) HATU, DIPEA, DMF, RT.
Synthesis of Solubilizing Group Analogues
Reagents and conditions: (i)
oxalyl chloride, DMF, 1,4-benzodioxane-6-carboxylic acid, DCM, then,
4-methyl-3-nitroaniline, pyridine, DCM, RT, then Pd/C (10%), H2 (1 atm), EtOH; (ii) NaH, RCH2OH, THF, 0 °C
to reflux; (iii) n-BuLi, CO2(s), THF,
−78 °C; (iv) HATU, DIPEA, DMF, RT.The optimal route to the synthesis of these analogues would be
to introduce the solubilizing group via nucleophilic aromatic substitution
on the quinoline as the final step; however, this gave consistently
low yields. Therefore, the solubilizing group was introduced at the
start of the synthesis. Formation of the precursor carboxylic acid 19 was achieved via lithium halogen exchange of the corresponding
aryl bromide 20. The resulting bisamides were then assayed
for cellular activity and in vitro PK parameters (Table ).
Table 2
Optimization
of the Bisamide Solubilizing
Group
pGI50 = −log GI50 (M); geometric mean ±
SEM, n = number
of biological repeats in parentheses. For the corresponding SK-OV-3
HSP72 cell-based ELISA data See Supporting Information.
Mouse liver microsome
(MLM) assay
was carried out at Cyprotex, arithmetic mean of n = 2. In vitro Clint is calculated from the half-life
using standard procedures and assumes the fraction unbound in the
assay is 1.[37]
Kinetic solubility (KS) measured
via an in-house HPLC method from phosphate buffer at pH 7.4, all values
quoted to 1 SF, the dynamic range of the assay is 1–100 μM,
arithmetic mean of n = 2.
Measured via an in-house HPLC method,
arithmetic mean of n = 2, all values quoted to 2
SF.[38]
MoKa version 2.5.2, all values quoted
to 2 SF. See Supporting Information for
details.
pGI50 = −log GI50 (M); geometric mean ±
SEM, n = number
of biological repeats in parentheses. For the corresponding SK-OV-3
HSP72 cell-based ELISA data See Supporting Information.Mouse liver microsome
(MLM) assay
was carried out at Cyprotex, arithmetic mean of n = 2. In vitro Clint is calculated from the half-life
using standard procedures and assumes the fraction unbound in the
assay is 1.[37]Kinetic solubility (KS) measured
via an in-house HPLC method from phosphate buffer at pH 7.4, all values
quoted to 1 SF, the dynamic range of the assay is 1–100 μM,
arithmetic mean of n = 2.Measured via an in-house HPLC method,
arithmetic mean of n = 2, all values quoted to 2
SF.[38]MoKa version 2.5.2, all values quoted
to 2 SF. See Supporting Information for
details.We first attached
the oxygen-linked ether chain to give glycol 21 (Table , entry 2). No significant
decrease in either HSF1-mediated HSP72
induction inhibition (see Supporting Information) or the antiproliferative activity was observed when compared to
lead bisamide 1 (Table , entry 1). This suggested first that the oxygen linker
had no detrimental effect on activity and, second, that this region
of the molecule was indeed solvent exposed. To carry out multiparameter
optimization of the solubilizing group, we used four in vitro properties
to assess each compound’s potential for in vivo mouse PK: microsomal
stability, kinetic solubility at pH 7.4, lipophilicity as measured
by LogD7.4, and the predicted basicity
of the solubilizing group. The glycol 21 displayed an
increase in lipophilicity compared to lead bisamide 1; this increase was reflected in the 2.4-fold decrease in microsomal
stability with a modest increase in kinetic solubility. However, glycol 21 remained a low solubility compound, which is inconsistent
with good oral bioavailability. From these data, we concluded that
the compound would need to be charged to deliver the correct balance
of properties. To introduce a charged moiety, we first focused on
the dimethylamine group with a 3-carbon linker. Dimethylamine 22 (Table , entry 3) displayed no significant change in cellular activity despite
this analogue being predominately charged at physiological pH, according
to the calculated pKa. Consistent with
its cationic character, dimethylamine 22 displayed a
1.4 log unit decrease in lipophilicity when compared to 21, which was accompanied by an 18-fold improvement in kinetic solubility.
Unfortunately, the decrease in lipophilicity was not reflected in
an improvement in microsomal stability. We hypothesized that this
was due to oxidation of the N-methyl groups, so we
moved to cyclic amines to reduce the CYP450 mediated degradation.
Morpholine 23 (Table , entry 4) showed little improvement in microsomal
stability or kinetic solubility when compared to 21.
Analysis of the LogD7.4 and calculated
pKa for morpholine 23 suggested
that the solubilizing group was not sufficiently basic to balance
the physicochemical properties of the compound. Moving to piperidine 24 (Table , entry 5), we observed a better balance in physicochemical properties,
with the reduction in lipophilicity reflected in improvements in both
microsomal stability and kinetic solubility. However, piperidine 24 was predicted to be highly basic and we were concerned
that this would have a detrimental effect on permeability and therefore
on oral bioavailability. To reduce the basicity of the solubilizing
group without increasing lipophilicity, we reduced the linker length
to give 25 (Table , entry 6), thereby exploiting the inductive effect of the
oxygen. Despite the 0.9 log unit decrease in predicted basicity, the
kinetic solubility and lipophilicity were unchanged. Unfortunately, 25 displayed a decrease in microsomal stability, so we reduced
the ring size to give pyrrolidine 26 (CCT251236, Table , entry 7),[24] which displayed the desired balance of in vitro
properties, while maintaining excellent cellular activity with a pIC50 = 7.73 ± 0.07 (IC50 = 19 nM, n = 15, see Supporting Information, Table S3 for details) for inhibition of HSF1-mediated HSP72 induction. The
free GI50 in SK-OV-3 cells was then calculated from the
free fraction in the cell assay, which gave a free GI50 = 1.1 nM.[39] Western blotting confirmed
that pyrrolidine 26 blocked the HSF1-mediated induction
of both HSP72 and HSP27 as representative heat shock proteins, following
treatment with the HSP90 inhibitor 17-AAG. Also, qPCR analysis demonstrated
that 26 inhibited the induction of HSP72 at the mRNA
level, clearly blocking the induction of HSPA1A gene
expression with a pIC50 = 7.40 (IC50 = 40 nM, n = 1, see Supporting Information). Taking these results together, pyrrolidine 26 was
selected for in vivo study.
Mouse Pharmacokinetics (PK)
To assess
the potential
of bisamide 26 as an in vivo chemical tool to study HSF1-mediated
transcriptional activity inhibition, the compound was dosed in BALB/c
mice at 5 mg/kg as an oral solution and iv bolus. Blood concentrations
were then measured over a 24 h period (Table ).
Table 3
Mouse Blood PK Parameters
for Bisamide 26a
mouse
dose po/iv (mg/kg)
AUC0–24hPO (h·nM)b
blood Cl (mL/min/kg)b
Vss (L/kg)b
half-life
(h)
%F
fubc
AUCu,0–24hPO (h·nM)f
Clu (mL/min/kg)g
Vdu (L/kg)h
BALB/c
5/5
5800 (8100–4200)
9.2 (12–7.3)
4.2 (6.3–2.8)
5.3
39
0.0083d
48
1100
510
athymic
20/0
1900 (2800–1400)
NA
NA
NA
NA
0.015e
29
NA
NA
All values are quoted to 2 SF. The
90% confidence intervals (CI) are in parentheses. NA = not applicable.
All values are quoted to 2 SF. The
90% confidence intervals (CI) are in parentheses. NA = not applicable.The geometric mean of n = 3 individual mice.fub = fup/B:P.fup =
0.010 (0.011–0.0097), B:P@1 μM = 1.2:1 (1.4–1.0).fup =
0.025 (0.030–0.020), B:P@1 μM = 1.7:1 (1.7–1.7).AUCu = AUC·fub.Clu = Cl/fub.Vdu = Vss/fub.Analysis of the mouse PK data revealed that bisamide 26 possessed low total blood clearance (10% hepatic blood flow)[40] and moderate oral bioavailability, with a half-life
sufficient to allow once-daily dosing. In vitro assessment of the
plasma protein binding and the blood to plasma ratio revealed that 26 was highly bound to plasma proteins (∼99%); therefore,
the unbound clearance was high, with a low free exposure from the
5 mg/kg oral dose, equivalent to a free Cav0–24h = 2.0 nM.[41,42] The high unbound
volume of distribution indicates that 26 readily binds
to tissues, consistent with the basicity of the compound.[43]
Bisamide 26 Displays Efficacy
in a Human Ovarian
Carcinoma Xenograft Model
Despite the high unbound clearance
of bisamide 26 in mouse, the good oral bioavailability,
half-life, and excellent in vitro cellular activity (free GI50 = 1.1 nM) encouraged us to test the potential efficacy of 26 in an in vivo human tumor xenograft model. On the basis
of the free Cav0–24h = 2.0 nM observed in nontumor bearing immunocompetent BALB/c mice
following the 5 mg/kg po qd dose, a 20 mg/kg po qd dose in immunodeprived
athymic mice was selected to cover ∼10 times the in vitro free
GI50 in SK-OV-3 cells.[44] To
our surprise, following this 20 mg/kg po dose, the free exposure was
actually a disappointing AUCu0–24h =
29 h·nM, equivalent to a free Cav0–24h = 1.2 nM (Table , see Supporting Information for details). Despite the lower than expected free exposure of bisamide 26, this dose still represented coverage of the in vitro free
GI50 in SK-OV-3 cells and was well tolerated in a mouse
multidose tolerability study;[45] therefore,
the 20 mg/kg po qd dose was selected for further investigation.[46]SK-OV-3 cells were injected subcutaneously
into athymic mice for tumor formation. Once tumors were established,
the mice were randomized into treatment and control groups and were
dosed orally once-a-day with either vehicle or 20 mg/kg of bisamide 26. Tumor volumes and mouse body weights were measured throughout
and tumor weights were measured at the end of the study, while total
tumor concentrations were measured 2 and 6 h post final dose (Figure and Supporting Information).
Figure 3
Efficacy of bisamide 26 against SK-OV-3 human ovarian
carcinoma xenograft model. Blue, vehicle control, n = 8; red, 26, 20 mg/kg po qd, n =
8 (vehicle = 10% DMSO, 90% of a 25% (2-hydroxypropyl)-β-cyclodextrin
in 50 mM citrate buffer pH 5). Error bars: arithmetic mean ±
SEM. Dosing breaks were carried out on days 5–12, 14, 16, 18,
20, 22, 24, 26, 29, 31.
Efficacy of bisamide 26 against SK-OV-3 human ovarian
carcinoma xenograft model. Blue, vehicle control, n = 8; red, 26, 20 mg/kg po qd, n =
8 (vehicle = 10% DMSO, 90% of a 25% (2-hydroxypropyl)-β-cyclodextrin
in 50 mM citrate buffer pH 5). Error bars: arithmetic mean ±
SEM. Dosing breaks were carried out on days 5–12, 14, 16, 18,
20, 22, 24, 26, 29, 31.The mice were dosed intermittently throughout the study with 26 to maintain their condition, as assessed through monitoring
body weights (see Supporting Information). Clear therapeutic efficacy was observed with bisamide 26, with a tumor growth inhibition (%TGI)[47] of 70% based on final tumor volumes. The study was terminated after
33 days, and comparison of the control and treated arms indicated
a 64% reduction in mean tumor weights (p = 0.015)[48] with total tumor concentrations of 26 as high as 940 nM, consistent with the compound’s basicity
and high volume of distribution (see Supporting
Information).
Target Identification
Following
the successful efficacy
study, the potential for bisamide 26 as a chemical probe
to study the effects of HSF1 transcription inhibition, both in vitro
and in vivo, were clear. However, the variety of HSF1 transcription
inhibitors in the literature[49] suggests
there are multiple mechanisms through which this phenotype may be
observed, each with differing potential for drug discovery.To decipher the molecular mechanism of the bisamide series, we needed
to discover their protein targets. The potency of the bisamide 26 in cell-based assays suggests that only high affinity efficacy
and epistatic targets would be of interest.[11] Our biochemical kinase screening had indicated that there were no
high affinity kinase targets. We therefore expanded our biochemical
screening to include other protein families. Bisamide 26 was submitted to the Cerep Diversity Screen (Cerep, http://www.cerep.fr) comprising 98
molecular targets, including receptors and enzymes, measured at 10
μM.[50] Analysis of these screening
data revealed that bisamide 26 displays a good selectivity
profile. Only six targets (the receptors: adenosine A2A and A3, histamine
H2 and H3, muscarinic, and the enzyme acetylcholine esterase) displayed
>80% inhibition, and these generally represented the highly promiscuous
receptor protein targets (see Supporting Information for details). No hits displayed sufficient activity to clearly relate
the molecular target to the efficacious free concentrations achieved
in vitro or in vivo.Because no clear protein families had been
revealed from our biochemical
screening, a different approach was necessary. We decided to exploit
a chemical proteomics strategy and using our knowledge of the cellular
SAR, we designed a protein pull-down chemical probe to identify high
affinity molecular targets from a human cancer cell lysate (Figure ).
Figure 4
Tool compounds for target
identification. All quoted cellular activities
are in the SK-OV-3 cell line, the numbers of repeats are in parentheses.
Tool compounds for target
identification. All quoted cellular activities
are in the SK-OV-3 cell line, the numbers of repeats are in parentheses.To carry out the chemical proteomics
pull-down strategy, the bisamide
warhead would need to be attached via a linker to a solid-phase bead,
without disrupting binding to the molecular target or targets. The
solubilizing group vector was the obvious choice for the linker attachment,
but to test whether cellular activity was maintained, amide 27 was designed as a cell-permeable mimic of the solid-phase
probe. Pleasingly, amide 27 maintained excellent cellular
potency (SK-OV-3 pGI50 = 8.04 ± 0.06, GI50 = 9.1 nM, n = 4), so amine 29 was
used for attachment to the bead via amide bond formation. In addition
to bisamide 26, tetrahydroquinoline 17 was
selected as a second positive control due to its distinct structural
difference. These two compounds would be used to displace specific
molecular targets from the solid-phase probe. Also, to help distinguish
false positives from the pull-down experiment, a physicochemically
matched negative control was designed. The dioxane isomer 6 (Table , entry 2)
had previously been shown to lose all cellular activity. To physicochemically
match the dioxane isomer 6 to bisamide 26, the same ethoxy pyrrolidine solubilizing group was attached to
give isomer 28, which, as expected, displayed no measurable
cellular activity.With all four probe molecules in hand, the
compounds were submitted
to the stable isotope labeling by amino acids in cell culture (SILAC)
mass-spectrometry based pull-down assay (Evotec, https:www.evotec.com) to identify
and quantify molecular targets of the bisamide series from the lysate
of SK-OV-3 cells. Each protein captured by the bead, whether specific
or nonspecific, was analyzed in this methodology by quantitative mass
spectrometry. By using SILAC to quantify the relative amounts of protein
captured from the lysate, this approach can, in principle, determine
the apparent affinity of every stable protein in the lysate for the
surface-bound bisamide probe, although particularly low abundance
proteins may not be discovered via this method and proteins that are
unstable to the lysis conditions would not be detected. Displacement
of the captured proteins with the free active bisamides 26 and 17 then allowed for estimation of their apparent
affinities for specific protein targets. Proteins that were apparently
displaced from the bead-bound probe by the inactive isomer 28 were considered nonspecific or irrelevant for the cellular activity
and discarded. The putative protein targets could then be ranked based
on their affinity for further investigation (Table , see Supporting Information for details).[51]
Table 4
Molecular
Targets from the Pull-down
Assay Using the Bisamide Probes in SK-OV-3 Cell Lysatea
protein
protein
EC50 (μM)b
26 AC50 (μM)c
26Kiapp (μM)d
17 AC50 (μM)c
17 Kiapp (μM)d
28 AC50 (μM)c
GSK3β
1.3
1.2
NA
10
NAe
NAb
Pirin
1.3
1.5
0.028
10
0.19
NAb
(A) SK-OV-3 cell lysate protein
binding curves for immobilized 29. (B) Protein displacement
curves for pyrrolidine 26. Curves are an average of two
different mixing conditions. EC50 and AC50 are
determined from analysis of the curves without limits.
EC50 is the apparent
immobilized bisamide probe concentration at which 50% of the available
protein is bound. The probe bisamide is assumed to be >10-fold
molar
excess over proteins in the lysate.
AC50 is the apparent
bisamide concentration at which 50% of the protein is displaced from
the immobilized bisamide probe. At least 2-fold enrichment of bound
proteins when compared to the matrix.
Kiapp values are
estimated using the Cheng–Prusoff equation,
apparent [29-immobilized] = 67 μM,[52] assuming no mass transport limitation, interactions are
purely competitive and all interactions are direct.
Not applicable (NA). Kiapp cannot be calculated if the interaction
with the probe is indirect.
(A) SK-OV-3 cell lysate protein
binding curves for immobilized 29. (B) Protein displacement
curves for pyrrolidine 26. Curves are an average of two
different mixing conditions. EC50 and AC50 are
determined from analysis of the curves without limits.EC50 is the apparent
immobilized bisamide probe concentration at which 50% of the available
protein is bound. The probe bisamide is assumed to be >10-fold
molar
excess over proteins in the lysate.AC50 is the apparent
bisamide concentration at which 50% of the protein is displaced from
the immobilized bisamide probe. At least 2-fold enrichment of bound
proteins when compared to the matrix.Kiapp values are
estimated using the Cheng–Prusoff equation,
apparent [29-immobilized] = 67 μM,[52] assuming no mass transport limitation, interactions are
purely competitive and all interactions are direct.Not applicable (NA). Kiapp cannot be calculated if the interaction
with the probe is indirect.Analysis of these data from the quantified chemical proteomics
pull-down experiment revealed very few molecular targets of the bisamides,
consistent with the selectivity observed with our biochemical screening.
After excluding highly promiscuous proteins commonly observed in pull-down
experiments,[53] three putative targets were
identified. PDE6D displayed only weak apparent affinity for the bead-bound
probe and little selectivity between active (26 and 17) and inactive (28) probes and so was not considered
further. GSK3β displayed apparent high affinity for both active
analogues, with no apparent affinity for the inactive analogue. However,
GSK3β had previously been assayed in our biochemical kinase
screening and both binding and functional assays (see Supporting Information) had demonstrated that
the bisamide series have no measurable affinity for this kinase; we
therefore concluded that GSK3β was potentially an indirect target
of the bisamide series. The final putative protein target was pirin.
The binding curve for pirin to the bead-bound bisamide probe was quite
shallow; however, the displacement curves for pirin, which were very
similar to the displacement curves for GSK3β with both cell
active probe ligands, gave robust data with an apparent Ki = 28 nM for 26 and no affinity was observed
with the inactive control. Using this method on a SK-OV-3 cell lysate,
pirin emerged as the only remaining putative target of the bisamide
series.[54]
Pirin Validation
Very little is known about pirin,
and few papers have been published discussing its possible function.
Pirin was first identified in 1997 by Wendler et al. from a yeast
two-hybrid screen to discover interactors of the transcription factor
NFI/CTF1. Pirin was described as an iron-binding, metal-dependent
protein, predominately localized within the nucleus, highly conserved
across species, and ubiquitously expressed in all tissue types.[55] Following this initial discovery, Scheidereit
et al. reported that human pirin interacts with the proto-oncoprotein,
BCL3, linking pirin with the NFκB pathway.[56] The role of pirin in the NFκB pathway was further
demonstrated by Lui et al.; using SPR, they revealed the metal-dependent
formation of a pirin/p65/DNA complex and hypothesized the role of
human pirin to be a redox-sensing transcription factor regulator.[57] The redox activity of pirin has previously been
discussed, as the expression level of pirin was proposed to be under
the control of the transcription factor NRF2 through changes in cellular
oxidative stress.[58] Wang et al. previously
reported that human pirin interacts with the tumor suppressor protein
EAF2/U19 and that exogenous overexpression of pirin increased colony
formation in LNCaP human prostate cancer cells.[59] In the human melanoma cell line WM266.4, Alcalay et al.
demonstrated that shRNA knockdown of pirin could induce a senescence
phenotype and suppress colony formation.[60] Also in the WM266.4 cell-line, Osada et al. demonstrated that siRNA
knockdown of pirin could suppress cell migration.[61,62] These results are consistent with the observation by Simizu et al.
that siRNA knockdown of pirin suppressed migration of human adenocarcinoma
HeLa cells and that pirin was important for epithelial–mesenchymal
transition (EMT).[63] However, little evidence
has emerged of an antiproliferative phenotype from genetic inhibition
of pirin expression and our in-house data using siRNA in SK-OV-3 cells
was consistent with this finding.[64] Nonetheless,
because our original screening paradigm was designed to discover inhibitors
of transcription, the proposed role of pirin as a redox-sensitive
transcription factor regulator was certainly intriguing.To
confirm pirin as a high affinity molecular target of the bisamide
series, we needed to assess the affinity of the compounds and establish
SAR in an orthogonal assay format. Pirin has no known catalytic function
and no known endogenous ligand in mammalian cells, consequently we
decided to focus on surface plasmon resonance (SPR) to measure the
affinity of the bisamide series for pirin. Purified recombinant human
pirin was attached to the SPR chip through a standard amide coupling.
The affinity of the bisamide ligand was assessed by equilibrium analysis
of the resulting sensorgram (Figure ).
Figure 5
Representative SPR sensorgram and binding isotherm of
bisamide 26 bound to recombinant pirin. The dotted-line
represents
the time-point the equilibrium response was measured. The binding
isotherm was fitted to a one-site specific binding model using Graphpad
Prism 6.07.
Representative SPR sensorgram and binding isotherm of
bisamide 26 bound to recombinant pirin. The dotted-line
represents
the time-point the equilibrium response was measured. The binding
isotherm was fitted to a one-site specific binding model using Graphpad
Prism 6.07.The known pirin ligand
TPh A[61] was used
as a positive control and to protect the binding site during amine
coupling to the SPR chip but displayed only modest affinity in our
hands (pKD = 5.77 ± 0.04, KD = 1700 nM, n = 4).[65] The SPR sensorgram showed that bisamide 26 is indeed a tight-binding ligand of pirin. Equilibrium
analysis revealed that bisamide 26 had a pKD = 7.36 ± 0.01 (KD =
44 nM, n = 3). However, the ratio of theoretical Rmax to measured Rmax from the binding isotherm was 0.40, indicating apparent substoichiometric
binding. This could be due to the coupling of pirin to the SPR chip
impairing protein folding or because pirin is a metal-dependent protein,
and the metal was leaching out into the running buffer; both effects
could negatively impact the apparent binding affinity and stoichiometry.To establish the SAR surrounding pirin binding, several bisamide
analogues were selected for further study. We first focused on the
two other analogues used in target identification. The cell active
analogue tetrahydroquinoline 17 (Table , entry 13) was also found to display tight-binding
affinity for pirin; in contrast, the negative control dioxane isomer 28 (pIC50 < 5.00, IC50 > 10000
nM, n = 2) displayed no measurable affinity. These
data suggested
that pirin SAR and the cellular SAR were linked. Increasing the steric
bulk of the solubilizing group, as exemplified by piperidine 24 (Table , entry 4), again returned a high affinity pirin ligand (pKD = 7.52 ± 0.02, KD = 30 nM, n = 3), consistent with the potent
cellular activity. However, structural changes to the benzodioxane
amide motif proved more complex. The original hit, bisamide 1 (Table ,
entry 1), was a tight-binding pirin ligand; whereas the cell inactive
analogues, methylated amide 13 (Table , entry 9) and dimethoxy 7 (Table , entry 3), displayed
no affinity for pirin. However, the two chroman isomers 8 and 9 (Table , Entries 4 and 5) were tight-binding ligands for pirin but
their target affinity did not reflect their decrease in cellular activity
when compared to bisamide 1. Finally, the isochroman
analogue 10 (Table , entry 6) was still a high affinity ligand for pirin,
despite displaying no cellular activity (see Supporting Information, Table S11 for details).Pirin is a member
of the cupin super family of proteins, which
are so-called because they all possess a conserved β-barrel.[66] However, due to minimal sequence homology between
members of the cupin family, crystallography is often needed to identify
them. The cupins display a huge array of functionality, including
oxidases and isomerases.[67] To better understand
pirin SAR, we determined the crystal structure of efficacious bisamide 26 bound to pirin (Figure ).
Figure 6
X-ray crystal structure of bisamide 26. PDB 5JCT, Pymol image of
pirin (light-blue cartoon representation) in complex with bisamide 26 (cyan stick representation), 2Fo – Fc map contoured at 1.0 σ
(blue mesh) and Pymol image of pirin (magenta, blue stick representation)
in complex with bisamide 26 (cyan stick representation),
distances shown are in Å, 2Fo – Fc contoured at 1.0 σ (blue mesh). Crystals
belonged to the space group P212121 and diffracted to a resolution of 1.73 Å.
X-ray crystal structure of bisamide 26. PDB 5JCT, Pymol image of
pirin (light-blue cartoon representation) in complex with bisamide 26 (cyan stick representation), 2Fo – Fc map contoured at 1.0 σ
(blue mesh) and Pymol image of pirin (magenta, blue stick representation)
in complex with bisamide 26 (cyan stick representation),
distances shown are in Å, 2Fo – Fc contoured at 1.0 σ (blue mesh). Crystals
belonged to the space group P212121 and diffracted to a resolution of 1.73 Å.Pirin is a metal-binding bicupin
characterized by the two β-barrels
that are clearly visible in the crystal structure.[68] Pirin binds a single metal-ion in one of the β-barrels,
which is where a deep, small molecule binding pocket is located. The
metal-ion is clearly visible in the electron density, although it
is unclear which metal-ion is actually bound. The metal-ion binding
site is formed of three histidine residues (His56, His58, and His101),
a glutamic acid (Glu103) and two water molecules, showing the metal-ion
possesses octahedral coordination. Bisamide 26 forms
no direct interactions with the metal-ion, instead a water-mediated
interaction is created by the two amide functional groups, which form
a hydrogen bonding pincer around this metal-coordinated water molecule,
revealing why the two amides are crucial for binding. The methyl-distal
amide also acts as a hydrogen-bond donor with an aspartic acid (Asp43)
at the base of the binding site, explaining why methylation of that
group is not tolerated. The methylpyridine ring of the quinoline substituent
is clearly solvent exposed and the solubilizing group cannot be resolved
due to flexibility, consistent with the broad SAR at this position
and the eventual success of the pull-down probe. The SAR around the
benzodioxane ring is less clear. From the cellular SAR, the oxygen
atoms of the dioxane are crucial for activity. The narrow binding
site is consistent with the cellularly inactive dioxane isomer 28 and the cellularly inactive dimethoxy analogue 7 having low affinity for pirin (Table and Supporting Information, Table
S11), as both these groups would cause a steric clash. However,
there are no clear interactions between the protein and the dioxane
ring oxygens, so it is unsurprising that changing the positions of
the oxygen atoms has little effect on pirin affinity, such that chromans 8, 9, and isochroman 10 (Table , entries 4–6)
still display high affinity for pirin, in contrast to their complex
cellular SAR.The apparent disconnect between the pirin affinity
SAR and the
antiproliferative cellular SAR led us to hypothesize that either simply
binding to pirin was not enough to recapitulate the cellular phenotype
or that binding to a second molecular target was also crucial. For
example, in the study by Lui et al.[57] the
formation of protein-protein interactions with pirin was under the
allosteric control of the redox-state of the iron bound in the protein.
Ligands bound in the metal-binding site are therefore likely to act
in an allosteric manner to regulate the function of pirin. It is commonly
known that allosteric modulators display complex SAR, where small
structural changes can have little effect on protein affinity but
can drastically alter the observed phenotype, switching ligands between
positive, negative, and neutral allosteric modulation.[69,70] The alternative hypothesis consistent with the observed complex
cellular SAR was that a second high affinity protein target was required
to explain the antiproliferative activity of the bisamide 26 and was not detected in our target-ID campaign due to either low
abundance or poor stability in the lysate. The dual targeting of proteins
is often observed; for example, CDK4 and CDK6, where simultaneous
inhibition of both kinases is necessary to observe an antiproliferative
phenotype[71] and owing to the similarity of their respective
binding sites, small-molecule inhibitors inevitably inhibit both proteins.[72] RNAi knockdown of pirin by several groups has
demonstrated very limited effects on cell proliferation, and as pirin
has no known enzymatic function, discovering and validating a cellular
biomarker to investigate its role in the antiproliferative phenotype
and in vivo efficacy is an ongoing challenge.One phenotype
that has previously been associated with a small-molecule
binding to pirin is cancer cell migration.[61] Miyazaki et al. demonstrated that their pirin ligand, TPh A, inhibited
the migration of the melanoma cell line, WM266.4.[61] To investigate whether our tool compound, bisamide 26, phenocopied this distinct pirin-ligand chemotype, we decided
to investigate its antimigratory activity using a scratch-wound assay.
The scratch-wound assay is commonly used to assess the effects of
small-molecules on cell migration.[73] Bisamide 26 demonstrated excellent antiproliferative activity against
the WM266.4 cell line (pGI50 = 7.92 ± 0.10, GI50 = 12 nM, n = 11). To exclude the effect
of inhibiting proliferation on migration in the scratch-wound assay,
the cells were plated at high confluency and the timeframe of the
assay was reduced to 30 h. The relative wound-density was then measured
at various concentrations and time-points. The negative control isomer 28 displayed no measurable antimigratory activity in stark
contrast to the potent pirin ligand bisamide 26, which
was able to strongly inhibit the migration of WM266.4 cells at 100
nM (Figure ).
Figure 7
Antimigratory
activity of chemical probe bisamide 26. (A) Wound healing
images of WM266.4 cells; after 30 h, the wound
has almost completely healed in the control in contrast to 100 nM 26. (B) Quantification of the relative wound density reveals
the maximum antimigratory activity is achieved at 100 nM 26. Each point represents the arithmetic mean ±SEM of the study
carried out in triplicate.
Antimigratory
activity of chemical probe bisamide 26. (A) Wound healing
images of WM266.4 cells; after 30 h, the wound
has almost completely healed in the control in contrast to 100 nM 26. (B) Quantification of the relative wound density reveals
the maximum antimigratory activity is achieved at 100 nM 26. Each point represents the arithmetic mean ±SEM of the study
carried out in triplicate.
Discussion
Bisamide 26 displays excellent
and wide-ranging in
vitro cellular potency, inhibiting both HSF1-mediated transcriptional
activity (defined by the inhibition of the induced expression of the
heat-shock proteins HSP72 and HSP27 and HSPA1A mRNA)
and cancer cell proliferation. Also, 26 has good mouse
PK and demonstrated its anticancer effects in vivo at low free exposures
with clear tumor growth inhibition at tolerable doses. Kinase screening
and broader proteome screening, in addition to the chemical proteomics
using SILAC quantified pull-down with rigorous controls and follow-up
testing, indicated that pirin was the sole specific protein target
that could be identified to date. SPR analysis of the binding of the
bisamide series to pirin confirms that these compounds are high affinity
ligands, with most aspects of the pirin SAR being consistent with
the cellular SAR. However, there was an apparent disconnect surrounding
the dioxane group of the chemotype. This requires further investigation
to determine whether pirin is crucial for the in vitro antiproliferative
activity and in vivo efficacy via an allosteric modulation mechanism
or whether binding to a second high affinity protein target is needed
to fully account for the observed phenotype. Because the SILAC quantified
pull-down assay can only identify protein targets that are stable
to cell lysis, additional in-cell target-ID methods are currently
under investigation to probe for a potential second target. Nevertheless,
the tool compound, bisamide 26, demonstrated the previously
proposed anticancer phenotype of pirin ligands by inhibiting the migration
of WM266.4 melanoma cells in vitro. The role of pirin in the bisamide
phenotype and the cellular effects of modulating the HSF1 pathway
with bisamide 26 are currently also under investigation
and will be reported subsequently.
Conclusions
Using
a high-throughput screen to identify inhibitors of the HSF1-mediated
stress pathway, we have discovered an extremely potent inhibitor of
human cancer cell proliferation in vitro from the bisamide chemotype.
By exploring the SAR from the cellular assays, we designed a chemical
probe, bisamide 26, which is highly potent and displays
an appropriate mouse pharmacokinetic profile to significantly inhibit
growth in a human ovarian carcinoma xenograft model. The chemical
probe 26 was also used to design a chemical proteomic
pull-down experiment, which identified the putative transcription
factor regulator, pirin, as a protein target. The high affinity of
chemical probe 26 for pirin was confirmed by SPR. Comparison
of the biophysical with the cellular data indicated that active molecules
bind pirin but that the cellular SAR is more complex, although 26 did display a potent inhibitory effect on the migration
of human melanoma cells, consistent with the putative pirin cancer
phenotype. Despite this, we propose that bisamide 26,
in combination with the physicochemically matched negative control
dioxane isomer 28, are promising chemical probes to investigate
the role of HSF1 pathway inhibition and pirin binding in vitro and
in vivo.
Experimental Section
Experimental Procedures
(Chemistry)
All final compounds
were screened through our in-house computational PAINS filter and
gave no structural alerts as potential assay interference compounds.[74] Unless otherwise stated, reactions were conducted
in oven-dried glassware under an atmosphere of nitrogen or argon using
anhydrous solvents. All commercially obtained reagents and solvents
were used as received. Thin layer chromatography (TLC) was performed
on precoated aluminum sheets of silica (60 F254 nm, Merck) and visualized
using short-wave UV light. Flash column chromatography was carried
out on Merck silica gel 60 (partial size 40–65 μm). Column
chromatography was also performed on a Biotage SP1 purification system
using Biotage Flash silica cartridges (SNAP KP-Sil). Ion exchange
chromatography was performed using acidic Isolute Flash SCX-II columns.
Semipreparative HPLC was performed on an Agilent 6120 system, flow
20 mL/min, eluents 0.1% acetic acid in water and 0.1% acetic acid
in methanol, gradient of 10–100% organic phase. 1H NMR spectra were recorded on Bruker AMX500 (500 MHz) spectrometers
using an internal deuterium lock. Chemical shifts are quoted in parts
per million (ppm) using the following internal references: CDCl3 (δH 7.26), MeOD (δH 3.31), and DMSO-d6 (δH 2.50). Signal multiplicities are recorded
as singlet (s), doublet (d), triplet (t), quartet (q), multiplet (m),
doublet of doublets (dd), doublet of doublet of doublets (ddd), broad
(br), or obscured (obs). Coupling constants, J, were
measured to the nearest 0.1 Hz. 13C NMR spectra were recorded
on Bruker AMX500 spectrometers at 126 MHz using an internal deuterium
lock. Chemical shifts are quoted to 0.01 ppm, unless greater accuracy
was required, using the following internal references: CDCl3 (δC 77.0), MeOD (δC 49.0), and DMSO-d6 (δC 39.5). High resolution mass spectra were recorded
on an Agilent 1200 series HPLC and diode array detector coupled to
a 6210 time-of-flight mass spectrometer with dual multimode APCI/ESI
source or on a Waters Acquity UPLC and diode array detector coupled
to a Waters G2 QToF mass spectrometer fitted with a multimode ESI/APCI
source. Analytical separation was carried out according to the methods
listed below. The mobile phase was a mixture of methanol (solvent
A) and water (solvent B), both containing formic acid at 0.1%, UV
detection was at 254 nm. Method I: Agilent 1200 series HPLC, Merck
Purospher STAR (RP-18e, 30 mm × 4 mm) column using a flow rate
of 1.5 mL/min in a 4 min gradient elution. Gradient elution was as
follows: 10:90 (A/B) to 90:10 (A/B) over 2.5 min, 90:10 (A/B) for
1 min, and then reversion back to 10:90 (A/B) over 0.3 min, finally
10:90 (A/B) for 0.2 min. Method II: Agilent 1200 series HPLC, Merck
Chromolith flash column (RP-18e, 25 mm × 2 mm) at 30 °C
using a flow rate of 0.75 mL/min in a 4 min gradient elution. Gradient
elution was as follows: 5:95 (A/B) to 100:0 (A/B) over 2.5 min, 100:0
(A/B) for 1 min, and then reversion back to 5:95 (A/B) over 0.1 min,
finally 5:95 (A/B) for 0.4 min. Method III: Waters Acquity UPLC, Phenomenex
Kinetex XB-C18 column (30 mm × 2.1 mm, 1.7 μ, 100 A) at
30 °C using flow rate of 0.3 mL/min in a 4 min gradient elution.
Gradient elution was as follows: 10:90 (A/B) to 90:10 (A/B) over 3
min, 90:10 (A/B) for 0.5 min, and then reversion back to 10:90 (A/B)
over 0.3 min, finally 10:90 (A/B) for 0.2 min; Method IV: Waters Acquity
UPLC, Phenomenex Kinetex C18 column (30 mm × 2.1 mm, 2.6 μ,
100A), flow rate and gradient elution according to Method III. The
following reference masses were used for HRMS analysis: Agilent 1200
series, caffeine [M + H]+ 195.087652, hexakis(1H,1H,3H-tetrafluoropentoxy)phosphazene
[M + H]+ 922.009798, and hexakis(2,2-difluoroethoxy)phosphazene
[M + H]+ 622.02896 or reserpine [M + H]+ 609.280657.
Waters Acquity UPLC: leucine enkephalin fragment ion [M + H]+ 397.1876. All compounds were >95% purity by LCMS analysis unless
otherwise stated.
General Synthetic Procedures
Method A
Oxalyl chloride (1.2–1.7 equiv) was
added dropwise to a solution of the carboxylic acid (1.0–1.5
equiv) and DMF in anhydrous dichloromethane. The reaction mixture
was stirred at room temperature for 3–4 h and then concentrated.
The remaining residue was redissolved in dichloromethane and concentrated
again. Then, the aniline (1.0 equiv) was added to the remaining residue
and the combined solids were dissolved in anhydrous pyridine or a
mixture of anhydrous pyridine and dichloromethane. The resulting reaction
mixture was stirred at room temperature overnight.
Method B
1-[Bis(dimethylamino)methylene]-1H-1,2,3-triazolo[4,5-b]pyridinium 3-oxide hexafluorophosphate
(HATU, 1.2–1.3 equiv) was added to a solution of the carboxylic
acid (1.0–1.2 equiv) and N,N-diisopropylethylamine (2.0–5.0 equiv) in anhydrous DMF. The
reaction mixture was stirred for 10 min before the aniline (1.0 equiv)
was added. The reaction mixture was stirred at room temperature overnight.
The reaction mixture was then diluted with water. The resulting precipitate
was isolated by filtration and washed with water.
Method C
The (3-nitrophenyl)amide (1.0 equiv), iron
powder (9–11 equiv), and ammonium chloride (9–11 equiv)
in ethanol/water (4/1) were added to a round-bottom flask and heated
to reflux overnight. Then the reaction mixture was filtered over a
short pad of Celite or silica gel. The filtrate was reduced in vacuo
until dryness. The remaining residue was resuspended in dichloromethane
and washed with satd NaHCO3 (aq), water, and brine. The
organic layer was dried over sodium sulfate or magnesium sulfate and
reduced in vacuo until dryness to afford the corresponding (3-aminophenyl)amide.
Method D
Palladium (10% on activated carbon, 42–68
mg/1 mmol) was added to a suspension of the nitro compound in a 1:1
mixture of ethanol/ethyl acetate and stirred under H2 (1
atm.) at 28 °C overnight. The reaction mixture was then filtered
through Celite and the Celite pad washed with further ethyl acetate.
The filtrate was concentrated in vacuo to afford the corresponding
aniline.
Method E
NaH (60% in mineral oil,
1.1–2.2 equiv)
was added to a solution of the alcohol (1.2–2.8 equiv) in anhydrous
THF at 0 °C. The reaction was allowed to stir at 0 °C for
10 min, then at room temperature for 30 min. 6-Bromo-2-chloroquinoline
was added and the resulting suspension heated to reflux for 1–16
h. The reaction was allowed to cool to room temperature, then diluted
with water/satd NaHCO3 (aq and extracted with dichloromethane
(3×)). The organic layers were combined, washed with water, dried
over magnesium sulfate, and concentrated in vacuo to afford the crude
product.
Method F
n-BuLi
(1.2–2.2 equiv,
2.5 M in hexanes, freshly titrated before use using standard procedures)
was added dropwise to a solution of the aryl bromide in anhydrous
THF at −78 °C under nitrogen. The reaction was stirred
at −78 °C for 40 min before solid CO2 was added.
After stirring for 5 min, the reaction was allowed to warm to room
temperature. The reaction was quenched with water and concentrated
under reduced pressure to remove the THF. The aqueous layer was washed
with ethyl acetate, acidified to pH 3 by the addition of 2 M HCl (aq),
and concentrated in vacuo to afford the crude product.
7-Bromoisochroman (152 mg, 0.71 mmol), Herrmann’s
palladacycle (33.4 mg, 0.04 mmol), and tri-tert-butylphosphonium
tetrafluoroborate (41 mg, 0.14 mmol) were combined with 2-(trimethylsilyl)ethanol
(7.0 mL) in a microwave vial. Molybdenum hexacarbonyl (377 mg, 1.427
mmol) was then added, followed by DBU (1.0 M in THF, 2.57 mL, 2.57
mmol), and the vial promptly sealed. The reaction mixture was heated
to 130 °C for 1 h in a microwave. After this time, further 2-(trimethylsilyl)ethanol
(3.0 mL) was added and the reaction heated to 130 °C for 1 h
under microwave irradiation. The reaction mixture was concentrated
under high vacuum. The resulting residue was purified by column chromatography
(Biotage, gradient of 0–100% ethyl acetate in cyclohexane)
to afford 2-(trimethylsilyl)ethyl isochroman-7-carboxylate compound
as a yellow oil (108 mg, 54%). This oil was only 80% pure but was
used directly in the next step.[75]To a stirring solution of 2-(trimethylsilyl)ethyl isochroman-7-carboxylate
(105 mg, 0.302 mmol) in THF (3 mL) at room temperature under argon
was dropwise added TBAF 1.0 M in THF (0.45 mL, 0.45 mmol). The reaction
was allowed to stir at room temperature for 2 h. The reaction mixture
was then diluted with water (25 mL) and the THF removed in vacuo.
The aqueous layer was washed with dichloromethane (15 mL). TLC/LCMS
showed that there was product in both the aqueous and organic layers.
Therefore, the organic layer was extracted with a portion of 1 M NaOH
(15 mL), followed by water (15 mL). The combined aqueous layers were
acidified (to ∼ pH 2) using 1 M HCl (aq) and extracted with
dichloromethane (3 × 15 mL). The combined organic layer was concentrated
in vacuo to afford isochroman-7-carboxylic acid as a colorless amorphous
solid. This material was used in the next step without further purification.Isochroman-7-carboxylic acid (50 mg, 0.28 mmol), HATU (130 mg,
0.34 mmol), and N-(5-amino-2-methylphenyl)-2-methylquinoline-6-carboxamide 5 (87 mg, 0.30 mmol) in N,N-diisopropylethylamine (0.14 mL, 0.82 mmol) and anhydrous DMF (2.5
mL) were reacted according to method B. The crude product was purified
by column chromatography (Biotage, gradient of 0–5% methanol
in dichloromethane) to afford a pale-yellow amorphous solid (78 mg,
63%). IR (thin film): νmax 2965, 1650, 1600, 1528,
1499, 1285 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 10.19 (s, 1H), 10.14 (s, 1H), 8.61
(d, J = 2.0 Hz, 1H), 8.41 (d, J =
8.4 Hz, 1H), 8.24 (dd, J = 8.8, 2.0 Hz, 1H), 8.03
(d, J = 8.8 Hz, 1H), 7.88 (d, J =
2.1 Hz, 1H), 7.78 (dd, J = 8.0, 1.8 Hz, 1H), 7.67–7.65
(m, 1H), 7.60 (dd, J = 8.3, 2.2 Hz, 1H), 7.53 (d, J = 8.4 Hz, 1H), 7.29 (d, J = 8.0 Hz, 1H),
7.26 (d, J = 8.6 Hz, 1H), 4.76 (s, 2H), 3.91 (t, J = 5.7 Hz, 2H), 2.86 (t, J = 5.7 Hz, 2H),
2.71 (s, 3H), 2.25 (s, 3H). 13C NMR (126 MHz, DMSO-d6) δ 165.58, 165.46, 161.22, 148.91, 137.70,
137.66, 137.49, 136.76, 135.49, 132.87, 131.96, 130.65, 129.34, 129.26,
128.80, 128.65, 128.40, 125.96, 125.86, 124.37, 123.46, 119.05, 118.63,
67.49, 64.82, 28.24, 25.50, 17.95. HRMS (ESI+): calcd for
C28H26N3O3 (M + H)+, 452.1969; found, 452.1964.
2,6-Dimethylaniline (4.07 mL, 32.7 mmol) and sulfuric
acid (30 mL) were added to a round-bottom flask. The solution was
cooled on an ice bath and nitric acid (2.6 mL, 43.1 mmol, 69%) was
added dropwise to the stirred solution. The resulting reaction mixture
was warmed to room temperature and stirred for further 30 min at room
temperature. The reaction mixture was then poured onto cold water
and neutralized by addition of solid sodium hydroxide. 2,6-Dimethyl-3-nitroaniline
was isolated by filtration and washed with copious amounts of water.
The crude product was purified by column chromatography (Biotage,
gradient of 0–40% ethyl acetate in cyclohexane) to afford the
product as a brown amorphous solid (1.82 g, 33%). IR (solid): νmax 3421, 3349, 2919, 1634, 1513, 1460, 1348 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 6.99 (d, J = 8.1 Hz, 1H), 6.93 (d, J = 8.1 Hz, 1H), 5.21 (s, 2H), 2.15 (s, 3H), 2.14 (s, 3H). 13C NMR (126 MHz, DMSO-d6) δ
150.03, 146.44, 127.90, 126.09, 113.71, 111.03, 18.71, 13.41. HRMS
(ESI+): calcd for C8H11N2O2 (M + H)+, 167.0815; found, 167.0831.2-Methylquinoline-6-carboxylic acid 3 (1.27 g, 6.77
mmol), oxalyl chloride (750 μL, 8.40 mmol), and DMF (110 μL,
1.42 mmol) in anhydrous dichloromethane (15 mL) were reacted according
to method A. The resulting acid chloride and 2,6-dimethyl-3-nitroaniline
(1.5 g, 3.01 mmol) were then reacted in anhydrous pyridine (20 mL).
The reaction mixture was concentrated in vacuo. The remaining residue
was redissolved in dichloromethane and washed with satd NaHCO3 (aq), water, and brine. The organic layer was dried over
sodium sulfate and reduced in vacuo until dryness to afford N-(2,6-dimethyl-3-nitrophenyl)-2-methylquinoline-6-carboxamide
as a dark-red to brown amorphous solid (1.69 g). The crude product
was used in the next synthetic step without further purification.N-(2,6-Dimethyl-3-nitrophenyl)-2-methylquinoline-6-carboxamide
(1.50 g, 4.47 mmol), iron powder (2.50 g, 44.7 mmol), and ammonium
chloride (762 mg, 44.7 mmol) in ethanol (40 mL) and water (10 mL)
were reacted according to method C. The crude product was purified
by column chromatography (Biotage, gradient of 0–30% ethanol
in dichloromethane) to afford the product as a brown amorphous solid
(1.14 g, 84%). IR (solid): νmax 3232, 2916, 1649,
1621, 1487, 1281 cm–1. 1H NMR (500 MHz,
DMSO-d6) δ 9.84 (s, 1H), 8.60 (d, J = 1.9 Hz, 1H), 8.39 (d, J = 8.4 Hz, 1H),
8.25 (dd, J = 8.8, 2.0 Hz, 1H), 8.01 (d, J = 8.7 Hz, 1H), 7.51 (d, J = 8.4 Hz, 1H),
6.82 (d, J = 8.1 Hz, 1H), 6.56 (d, J = 8.1 Hz, 1H), 4.72 (s, 2H), 2.70 (s, 3H), 2.07 (s, 3H), 1.95 (s,
3H). 13C NMR (126 MHz, DMSO-d6) δ 165.08, 161.05, 148.86, 145.57, 137.57, 135.64, 132.08,
128.77, 128.40, 128.27, 127.51, 125.89, 123.38, 123.09, 120.01, 113.46,
25.49, 18.15, 12.84. HRMS (ESI+): calcd for C19H20N3O (M + H)+, 306.1606; found,
306.1597.2,3-Dihydrobenzo[b][1,4]dioxine-6-carboxylic
acid
(415 mg, 2.30 mmol), oxalyl chloride (350 μL, 3.92 mmol), and
DMF (50 μL, 646 mmol) in anhydrous dichloromethane (10 mL) were
reacted according to method A. N-(3-Amino-2,6-dimethylphenyl)-2-methylquinoline-6-carboxamide
(703 mg, 2.30 mmol) and the acid chloride were then reacted in anhydrous
pyridine. The reaction mixture was diluted with diethyl ether. The
product precipitated from the solution, isolated by filtration, and
washed with water and diethyl ether. The crude product was purified
by column chromatography (Biotage, gradient of 0–10% ethanol
in dichloromethane) to afford compound 11 as a beige
amorphous solid (237.4 mg, 22%). IR (thin film): νmax 1649, 1584, 1487, 1290, 1067 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 10.08
(s, 1H), 9.80 (s, 1H), 8.63 (s, 1H), 8.41 (d, J =
8.4 Hz, 1H), 8.27 (d, J = 10.8 Hz, 1H), 8.04 (d, J = 8.8 Hz, 1H), 7.55 (d, J = 2.0 Hz, 1H),
7.54–7.53 (m, 1H), 7.53–7.50 (m, 1H), 7.18 (m, 2H),
6.98 (d, J = 8.4 Hz, 1H), 4.32–4.29 (m, 4H),
2.71 (s, 3H), 2.24 (s, 3H), 2.10 (s, 3H). 13C NMR (126
MHz, DMSO-d6) δ 165.22, 164.92,
161.17, 148.93, 146.73, 143.41, 137.60, 136.13, 135.30, 133.96, 132.95,
131.78, 128.86, 128.55, 128.27, 127.84, 127.38, 126.24, 125.90, 123.44,
121.61, 117.31, 117.12, 64.85, 64.47, 25.51, 18.59, 13.94. HRMS (ESI+): calcd for C28H26N3O4 (M + H)+, 468.1923; found, 428.1932.
2,3-Dihydrobenzo[b][1,4]dioxine-6-carboxylic
acid (542 mg, 3.01 mmol), oxalyl chloride (325 μL, 4.71 mmol),
and DMF (50 μL, 0.646 mmol) in anhydrous dichloromethane (10
mL) were reacted according to method A. 2,4-Dimethyl-5-nitroaniline
(500 mg, 3.01 mmol) and the acid chloride were then reacted in anhydrous
pyridine (15 mL). The reaction mixture was concentrated in vacuo.
The remaining residue was triturated with diethyl ether and dried
in vacuo to give N-(2,4-dimethyl-5-nitrophenyl)-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide as a brown amorphous solid
(1.11 g), which was used in the next synthetic step without further
purification.N-(2,4-Dimethyl-5-nitrophenyl)-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide (988 mg, 2.08 mmol), ammonium
chloride (394 mg, 23.1 mmol), and iron powder (1.29 g, 23.1 mmol)
in ethanol (10 mL) and water (2.5 mL) were reacted according to method
C to afford N-(5-amino-2,4-dimethylphenyl)-2,3-dihydrobenz[b][1,4]dioxine-6-carboxamide as a brown amorphous solid
(353 mg, 57% over two steps). IR (solid): νmax 3352,
3265, 2918, 1634, 1613, 1489, 1288 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 9.43
(s, 1H), 7.49 (d, J = 2.0 Hz, 1H), 7.47 (dd, J = 8.3, 2.1 Hz, 1H), 6.95 (d, J = 8.3
Hz, 1H), 6.77 (s, 1H), 6.58 (s, 1H), 4.65 (s, 2H), 4.40–4.10
(m, 4H), 2.02 (s, 3H), 2.00 (s, 3H). 13C NMR (126 MHz,
DMSO-d6) δ 164.53, 146.53, 144.85,
143.36, 134.81, 131.82, 128.22, 121.45, 121.11, 119.64, 117.23, 117.00,
113.13, 64.81, 64.46, 17.41, 17.29. HRMS (ESI+): calcd
for C17H19N2O3 (M + H)+, 299.1396; found, 299.1555.2-Methylquinoline-6-carboxylic
acid 3 (188 mg, 1.00
mmol), oxalyl chloride (80 μL, 0.896 mmol), and DMF (20 μL,
0.258 mmol) in anhydrous dichloromethane (5 mL) were reacted according
to method A. The resulting acid chloride and N-(5-amino-2,4-dimethylphenyl)-2,3-dihydrobenz[b][1,4]dioxine-6-carboxamide (201 mg, 0.674 mmol) were then
reacted in anhydrous pyridine (10 mL). The reaction mixture was then
diluted with dichloromethane and washed with satd NaHCO3 (aq), water, and brine. The organic layer was dried over sodium
sulfate and reduced in vacuo until dryness. The crude product was
purified by column chromatography (Biotage, gradient of 0–5%
ethanol in dichloromethane). The product was further purified by semipreparative
HPLC, according to the method specified in the general procedures,
to afford a beige amorphous solid (30.4 mg, 9.7%). IR (solid): νmax 1664, 1620, 1531, 1497, 1291 cm–1. 1H NMR (500 MHz, DMSO-d6) δ
10.09 (s, 1H), 9.71 (s, 1H), 8.59 (s, 1H), 8.40 (d, J = 8.4 Hz, 1H), 8.29–8.16 (m, 1H), 8.02 (d, J = 8.8 Hz, 1H), 7.64–7.43 (m, 3H), 7.35 (s, 1H), 7.17 (s,
1H), 6.98 (d, J = 8.3 Hz, 1H), 4.39–4.15 (m,
4H), 2.70 (s, 3H), 2.25 (s, 3H), 2.20 (s, 3H). 13C NMR
(126 MHz, DMSO-d6) δ 165.48, 164.83,
161.18, 148.91, 146.74, 143.41, 137.60, 134.65, 134.46, 132.25, 132.01,
131.85, 131.63, 128.77, 128.61, 128.40, 127.87, 125.84, 125.00, 123.43,
121.60, 117.31, 117.11, 64.84, 64.48, 25.50, 17.97, 17.89. HRMS (ESI+): calcd for C28H26N3O4 (M + H)+, 468.1923; found, 468.1914.
4-Methyl-3-nitroaniline 2 (5.15 g,
33.8 mmol) was dissolved in DMF (20 mL), and iodomethane (1.8 mL,
28.9 mmol) was added. Then, satd NaHCO3 (aq) (20 mL, 28.2
mmol) was added portionwise to the stirred reaction mixture. The resulting
reaction mixture was stirred at room temperature overnight. The reaction
mixture was partitioned between dichloromethane and water. The aqueous
layer was extracted with dichloromethane. The combined organic layers
were then washed with water and brine. The crude product was purified
by column chromatography (Biotage, gradient of 0–20% ethyl
acetate in cyclohexane) to afford N,4-dimethyl-3-nitroaniline
as a red amorphous solid (1.55 g, 33% yield). IR (thin film): νmax 3401, 2926, 1690, 1526, 1493, 1321, 1286 cm–1. 1H NMR (500 MHz, CDCl3) δ 7.18 (d, J = 2.6 Hz, 1H), 7.10 (d, J = 8.3 Hz, 1H),
6.75 (dd, J = 8.3, 2.6 Hz, 1H), 3.91 (s, 1H), 2.87
(s, 3H), 2.47 (s, 3H). 13C NMR (126 MHz, CDCl3) δ 149.77, 148.06, 133.16, 121.34, 117.63, 106.97, 30.65,
19.53. HRMS (ESI+): calcd for C8H11N2O2 (M + H)+, 167.0820; found,
167.0825.2,3-Dihydrobenzo[b][1,4]dioxine-6-carboxylic
acid (1.63 g, 9.03 mmol), oxalyl chloride (970 μL, 10.86 mmol),
and DMF (140 μL, 1.81 mmol) in anhydrous dichloromethane (15
mL) were reacted according to method A. The resulting acid chloride
and N-4-dimethyl-3-nitroaniline (1.5 g, 9.03 mmol)
were then reacted in anhydrous dichloromethane (15 mL) and pyridine
(3.0 mL, 34.0 mmol). The reaction mixture was diluted with dichloromethane
and washed with satd NH4Cl (aq). The aqueous layer was
extracted with dichloromethane. The combined organic layers were washed
with satd NH4Cl (aq), water, and brine. The organic layer
was dried over sodium sulfate and reduced in vacuo until dryness to
give N-methyl-N-(4-methyl-3-nitrophenyl)-2,3-dihydrobenz[b][1,4]dioxine-6-carboxamide (2.51 g), which was used in
the next synthetic step without further purification.N-Methyl-N-(4-methyl-3-nitrophenyl)-2,3-dihydrobenz[b][1,4]dioxine-6-carboxamide (2.50 g, 7.61 mmol), iron powder
(4.56 g, 82.0 mmol), and ammonium chloride (1.36 g, 80 mmol) in ethanol
(20 mL) and water (5 mL) were reacted according to method C. The crude
product was resuspended in water. The product was isolated by filtration
and washed with water and diethyl ether. The product was isolated
as a beige amorphous solid (1.18 g, 52%). IR (solid): νmax 3447, 3354, 2934, 1600, 1579, 1509, 1426, 1282 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 6.85–6.82 (m, 1H), 6.80 (d, J =
7.8 Hz, 1H), 6.78–6.72 (m, 1H), 6.66 (d, J = 8.4 Hz, 1H), 6.40–6.32 (m, 1H), 6.23–6.13 (m, 1H),
4.92 (s, 2H), 4.18 (d, J = 8.0 Hz, 4H), 3.23 (s,
3H), 1.97 (s, 3H). 13C NMR (126 MHz, DMSO-d6) δ 168.76, 147.70, 144.89, 144.04, 142.72, 130.83,
129.67, 122.40, 119.85, 118.09, 116.54, 114.85, 112.22, 64.58, 64.32,
38.98, 17.45. HRMS (ESI+): calcd for C17H19N2O3 (M + H)+, 300.1474;
found, 299.1401.2-Methylquinoline-6-carboxylic acid 3 (345 mg, 1.84
mmol), oxalyl chloride (180 μL, 2.06 mmol), and DMF (300 μL,
3.87 mmol) in anhydrous dichloromethane (10 mL) were reacted according
to method A. The resulting acid chloride and N-(3-amino-4-methylphenyl)-N-methyl-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide
(500 mg, 1.68 mmol) were then reacted in anhydrous dichloromethane
(13 mL) and pyridine (1.2 mL, 14.89 mmol). The reaction mixture was
diluted with water. The product was isolated by filtration and washed
with water and diethyl ether. The crude product was then purified
with a cation exchange sorbent (Isolute SCX-II, washing first with
methanol, and then with 10% 2 M ammonia in methanol). The product
was further purified by semipreparative HPLC according to the method
specified in the general procedures to afford a yellow amorphous solid
(22.4 mg, 2.9%). IR (solid): νmax 2927, 1624, 1493,
1282 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 10.09 (s, 1H), 8.59 (d, J = 1.8 Hz, 1H), 8.39 (d, J = 8.4 Hz, 1H), 8.21 (dd, J = 8.8, 1.9 Hz, 1H), 8.02 (d, J = 8.8
Hz, 1H), 7.52 (d, J = 8.4 Hz, 1H), 7.36 (d, J = 2.2 Hz, 1H), 7.18 (d, J = 8.2 Hz, 1H),
6.90 (dd, J = 8.1, 2.2 Hz, 1H), 6.87 (d, J = 2.0 Hz, 1H), 6.76 (d, J = 2.0 Hz, 1H),
6.68 (d, J = 8.4 Hz, 1H), 4.27–4.14 (m, 4H),
3.17 (s, 3H), 2.70 (s, 3H), 2.23 (s, 3H). 13C NMR (126
MHz, DMSO-d6) δ 169.06, 161.29,
148.95, 144.99, 143.28, 142.94, 137.62, 137.39, 131.93, 131.75, 131.25,
129.44, 128.80, 128.75, 128.39, 125.81, 125.05, 124.52, 123.47, 122.42,
118.16, 116.71, 64.60, 64.33, 49.06, 38.94, 25.51, 18.04. HRMS (ESI+): calcd for C28H26N3O4 (M + H)+, 468.1923; found, 468.1919.
1,4-Benzodioxane-6-carboxylic
acid (2.486 g, 13.80 mmol), oxalyl chloride (1.40 mL, 16.6 mmol),
and DMF (0.027 mL, 0.34 mmol) in anhydrous dichloromethane (34 mL)
were reacted according to method A. The resulting acid chloride was
dissolved in anhydrous dichloromethane (12 mL) and added dropwise
to a solution of 4-methyl-3-nitroaniline 2 (2.100 g,
13.80 mmol) and pyridine (2.23 mL, 27.6 mmol) in anhydrous dichloromethane
(35 mL). The reaction mixture was stirred at room temperature for
2 h and then concentrated. The resulting amorphous solid was suspended
in methanol, diluted with water, and then isolated by filtration and
washed with water to afford N-(4-methyl-3-nitrophenyl)-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide (4.24 g, 98%) as a pale-tan
colored amorphous solid, which was used in the next step without further
purification.Palladium (10% on activated carbon, 0.567 g) and N-(4-methyl-3-nitrophenyl)-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide (4.237 g, 13.48 mmol) in ethanol (90
mL) and ethyl acetate (90 mL) were reacted according to method D to
afford the desired product (3.803 g, 99%) as a pale-yellow amorphous
solid. IR (thin film): νmax 1641, 1611, 1582, 1289,
1064 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 9.70 (s, 1H), 7.49 (d, J = 2.2 Hz, 1H), 7.46 (dd, J = 8.3, 2.2 Hz, 1H),
7.10 (d, J = 2.0 Hz, 1H), 6.95 (d, J = 8.4 Hz, 1H), 6.83 (d, J = 8.1 Hz, 1H), 6.79 (dd, J = 8.1, 2.0 Hz, 1H), 4.81 (s, 2H), 4.32–4.26 (m,
4H), 2.01 (s, 3H). 13C NMR (126 MHz, DMSO-d6) δ 164.48, 146.89, 146.54, 143.30, 138.09, 130.04,
128.60, 121.54, 117.18, 117.05, 109.26, 106.88, 64.82, 64.47, 17.46.
(1 carbon missing) HRMS (ESI+): calcd for C16H17N2O3 (M + H)+, 285.1234;
found, 285.1233.
NaH (60% in mineral oil, 0.198 g, 4.950 mmol)
was added to 2-methoxyethanol (4.0 mL, 50.7 mmol) at 0 °C under
inert atmosphere. The reaction mixture was stirred at 0 °C for
10 min then at room temperature for 20 min before 6-bromo-2-chloroquinoline
(0.400 g, 1.649 mmol) was added. This suspension was then gradually
heated to 70 °C, the amorphous solid dissolved as heated, and
the resulting solution was allowed to stir overnight. The reaction
mixture was cooled to room temperature and diluted with water and
the resulting precipitate isolated by filtration. The precipitate
was washed with water and dried to afford 6-bromo-2-(2-methoxyethoxy)quinoline
as a dull-pink amorphous solid (0.497 g), which was used in the next
step without further purification.n-BuLi (1.84
M in hexanes) (0.578 mL, 1.063 mmol) was added dropwise to a solution
of 6-bromo-2-(2-methoxyethoxy)quinoline (0.250 g, 0.886 mmol) in anhydrous
THF (3.0 mL) at −78 °C and the resulting brown-yellow
mixture was stirred at −78 °C for 35 min. Solid CO2 was added, and the resulting orange mixture was stirred at
−78 °C for 5 min, then it was allowed to warm to room
temperature and concentrated under reduced pressure. The residue was
dissolved in brine (5.0 mL) and washed with dichloromethane (1 ×
5.0 mL). The aqueous phase was acidified with 2M HCl (aq) to pH 3,
and the resulting precipitate was isolated by filtration and washed
with water to afford 2-(2-methoxyethoxy)quinoline-6-carboxylic acid
as a pale pink amorphous solid (0.127 g), which was used in next step
without further purification.2-(2-Methoxyethoxy)quinoline-6-carboxylic
acid (0.048 g, 0.193
mmol), HATU (0.084 g, 0.220 mmol), and N-(3-amino-4-methylphenyl)-2,3-dihydrobenzo[b][1,4]dioxine-6-carboxamide 18 (0.050 g, 0.176
mmol) in N,N-diisopropylethylamine
(0.068 mL, 0.387 mmol) and anhydrous DMF (1.25 mL) were reacted according
to method B to afford the product 21 as a white amorphous
solid (87 mg, 56% over 3 steps). IR (thin film): νmax 2925, 1642, 1602, 1582, 1526, 1502, 1345, 1286, 1261, 816, 784 cm–1. 1H NMR (500 MHz, DMSO-d6) δ 10.07 (app. d, J = 3.0 Hz,
2H), 8.57 (d, J = 2.0 Hz, 1H), 8.39 (d, J = 8.8 Hz, 1H), 8.22 (dd, J = 8.7, 2.0 Hz, 1H),
7.89–7.84 (m, 2H), 7.58 (dd, J = 8.3, 2.2
Hz, 1H), 7.54 (d, J = 2.1 Hz, 1H), 7.51 (dd, J = 8.4, 2.2 Hz, 1H), 7.24 (d, J = 8.6
Hz, 1H), 7.14 (d, J = 8.8 Hz, 1H), 6.98 (d, J = 8.4 Hz, 1H), 4.61–4.55 (m, 2H), 4.32–4.29
(m, 4H), 3.77–3.72 (m, 2H), 3.33 (s, 3H), 2.24 (s, 3H). 13C NMR (126 MHz, DMSO-d6) δ
165.39, 164.81, 163.13, 147.93, 146.80, 143.38, 140.80, 137.75, 136.78,
130.57, 129.22, 128.90, 128.58, 128.15, 127.21, 126.76, 124.52, 121.65,
119.07, 118.57, 117.29, 117.12, 114.39, 70.56, 65.40, 64.85, 64.48,
58.58, 17.94. HRMS (ESI+): calcd for C29H28N3O6 (M + H)+, 514.1973;
found, 514.1969.
Authors: Dana Faratian; Annelien J M Zweemer; Yoko Nagumo; Andrew H Sims; Morwenna Muir; Michael Dodds; Peter Mullen; Inhwa Um; Charlene Kay; Max Hasmann; David J Harrison; Simon P Langdon Journal: Clin Cancer Res Date: 2011-05-13 Impact factor: 12.531
Authors: Frankie Lam; Abdullahi Y Abbas; Hao Shao; Theodosia Teo; Julian Adams; Peng Li; Tracey D Bradshaw; Peter M Fischer; Elisabeth Walsby; Chris Pepper; Yi Chen; Jian Ding; Shudong Wang Journal: Oncotarget Date: 2014-09-15
Authors: Marc L Mendillo; Sandro Santagata; Martina Koeva; George W Bell; Rong Hu; Rulla M Tamimi; Ernest Fraenkel; Tan A Ince; Luke Whitesell; Susan Lindquist Journal: Cell Date: 2012-08-03 Impact factor: 41.582
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