Structurally Diverse Histone Deacetylase Photoreactive Probes: Design, Synthesis, and Photolabeling Studies in Live Cells and Tissue

Article abstract of DOI:10.1002/cmdc.201900114

Histone deacetylase (HDAC) activity is modulated in vivo by post-translational modifications and formation of multiprotein complexes. Novel chemical tools to study how these factors affect engagement of HDAC isoforms by HDAC inhibitors (HDACi) in cells and tissues are needed. In this study, a synthetic strategy to access chemically diverse photoreactive probes (PRPs) was developed and used to prepare seven novel HDAC PRPs 9–15. The class I HDAC isoform engagement by PRPs was determined in biochemical assays and photolabeling experiments in live SET-2, HepG2, HuH7, and HEK293T cell lines and in mouse liver tissue. Unlike the HDAC protein abundance and biochemical activity against recombinant HDACs, the chemotype of the PRPs and the type of cells were key in defining the engagement of HDAC isoforms in live cells. Our findings suggest that engagement of HDAC isoforms by HDACi in vivo may be substantially modulated in a cell- and tissue-type-dependent manner.

Full text of DOI:10.1002/cmdc.201900114

Accepted Article
Title: Structurally Diverse Histone Deacetylase Photoreactive Probes:
Design, Synthesis, and Photolabeling Studies in Live Cells and
Tissue
Authors: Shaimaa M. Aboukhatwa, Thomas W. Hanigan, Taha Y.
Taha, Jayaprakash Neerasa, Rajeev Ranjan, Eman E. El-
Bastawissy, Mohamed A. Elkersh, Tarek F. El-Moselhy,
Jonna Frasor, Nadim Mahmud, Alan McLachlan, and Pavel A.
Petukhov
This manuscript has been accepted after peer review and appears as an
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of the final Version of Record (VoR). This work is currently citable by
using the Digital Object Identifier (DOI) given below. The VoR will be
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to this Accepted Article as a result of editing. Readers should obtain
the VoR from the journal website shown below when it is published
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content of this Accepted Article.
To be cited as: ChemMedChem 10.1002/cmdc.201900114
Link to VoR: http://dx.doi.org/10.1002/cmdc.201900114
A Journal of
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10.1002/cmdc.201900114
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FULL PAPER
Structurally Diverse Histone Deacetylase Photoreactive Probes:
Design, Synthesis, and Photolabeling Studies in Live Cells and
Tissue
Shaimaa M. Aboukhatwa, [a, b] Dr. Thomas W. Hanigan, [a] Taha Y. Taha, [a] Dr. Jayaprakash Neerasa, [a]
Dr. Rajeev Ranjan, [c] Dr. Eman E. El-Bastawissy, [b] Dr. Mohamed A. Elkersh, [b, d] Prof. Dr. Tarek F. El-
Moselhy, [b] Prof. Dr. Jonna Frasor, [e] Prof. Dr. Nadim Mahmud, [c] Prof. Dr. Alan McLachlan, [f] Prof. Dr.
Pavel A. Petukhov[a]
[a]
S. M. Aboukhatwa https://orcid.org/0000-0002-5013-7158, Dr. T. W. Hanigan, T. Y. Taha http://orcid.org/0000-0002-7344-7490, Dr. J. Neerasa, Prof. Dr.
P. A. Petukhov http://orcid.org/0000-0003-1554-242X.
Department of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, University of Illinois at Chicago, 833 South Wood Street, Chicago, IL
60612 (USA), E-mail: pap4@uic.edu.
[b]
[c]
[d]
[e]
[f]
S. M. Aboukhatwa https://orcid.org/0000-0002-5013-7158, Dr. E. E. El-Bastawissy, Dr. M. A. Elkersh, Prof. Dr. T. F. El-Moselhy,
Department of Pharmaceutical Chemistry, Faculty of Pharmacy, Tanta University, Tanta, 31527 (Egypt).
Dr. R. Ranjan, Prof. Dr. N. Mahmud http://orcid.org/0000-0003-3113-9155.
Section of Hematology/Oncology, College of Medicine, University of Illinois at Chicago, Chicago, IL 60612 (USA).
Dr. M. A. Elkersh,
Department of Pharmaceutical Chemistry, Faculty of Pharmacy, Pharos University, Alexandria, 21311 (Egypt).
Prof. Dr. J. Frasor
Department of Physiology and Biophysics, College of Medicine, University of Illinois at Chicago, Chicago, IL 60612.
Prof. Dr. A. McLachlan https://orcid.org/0000-0002-4168-2365
Department of Microbiology and Immunology, College of Medicine, University of Illinois at Chicago, Chicago, IL, 60612.
Supporting information for this article is given via a link at the end of the document.
Abstract: Histone deacetylase (HDAC) activity is modulated in vivo
by post-translational modifications and formation of multiprotein
complexes. Novel chemical tools to study how these factors affect
engagement of HDAC isoforms by HDAC inhibitors (HDACi) in cells
and tissues are needed. In this study, a synthetic strategy to access
chemically diverse photoreactive probes (PRPs) was developed and
used to prepare seven novel HDAC PRPs 9–15. The class I HDAC
isoform engagement by PRPs was determined in the biochemical
assays and photolabeling experiments in live SET-2, HepG2, HuH7,
and HEK293T cell lines and in mouse liver tissue. Unlike the HDAC
protein abundance and biochemical activity against recombinant
HDACs, the chemotype of the PRPs and the type of cells were key in
defining engagement of HDAC isoforms in live cells. Our findings
suggest that engagement of HDAC isoforms by HDACi in vivo may be
substantially modulated in a cell and tissue type dependent manner.
However, substrate specificity and HDAC catalytic activity, and
hence biological function, of HDACs are often modulated in vivo
via formation of protein-protein complexes and post-translation
modifications,[4] which these assays do not recapitulate. It was
also demonstrated that the catalytic activity of at least some of the
HDACs is dispensable and HDACs can play a non-enzymatic
structural (scaffolding) role.[2b] These modulatory mechanisms
have been shown to be cell-state dependent and can be
dysregulated in diseases and conditions.[5] A typical alternative
approach in this case would be to use cellular phenotypic
readouts. With few exceptions, HDAC isoform substrate
specificity in live cells and in vivo is poorly understood, and these
biochemical assays almost always remain the only avenue for
measuring potency and selectivity of HDACi. HDACi are non-
hydrolysable HDAC substrate analogues, and, therefore, the
same mechanisms that control substrate specificity of HDACs
should also affect binding of HDACi. Since cellular context is lost
in the biochemical inhibitory assays, HDACi potency and
selectivity may not accurately reflect the in situ target engagement
of HDACi. Several experimental approaches, including HDAC
photoreactive probes (PRPs), have emerged to assess HDACi
target engagement within complex systems.[6] The importance of
cellular context to study engagement of HDACs was further
highlighted by a recent study by Bantscheff et al [6a] who showed
that HDACi tethered to polymer beads exhibited chemotype- and
deacetylase complex-dependent binding to HDAC isoforms. Cell
permeable HDAC PRPs were reported by Cravatt et al,[6c, 7] Storer
et al,[8] and our laboratory.[6f, 9] Despite the overall progress in
applying PRPs to study engagement of HDACs in situ, synthetic
accessibility to drug-like cell permeable HDAC PRPs with
photoreactive groups able to generate short lived reactive
intermediates to accurately report on target engagement is limited.
Introduction
Histone deacetylases (HDACs) are promising therapeutic
targets.[1] Four HDAC inhibitors (HDACi) of multiple class I and II
HDACs are FDA approved for the treatment of T-cell lymphoma
and multiple myeloma. Since HDAC isoforms deacetylate both
histone and non-histone proteins and generally play unique non-
redundant functions,[2] it is hypothesized that selective HDACi will
offer a desired therapeutic effect while minimizing toxicity. To
investigate this hypothesis, many isoform-selective HDACi have
been synthesized and tested (Figure 1).[3]
HDACi potency and isoform selectivity are typically evaluated in
biochemical assays under pseudo-equilibrium utilizing individual
purified recombinant HDAC isoforms and a synthetic substrate.
1
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10.1002/cmdc.201900114
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Linker
ZBG
SBG
H
O
N
HN
H
O
N
N
O
N
H
OH
H
N
H
N
O
N
H
OH
OH
O
O
N
S
O
panobinostat (2)
potent pan HDACi
FDA approved 2015
vorinostat (1)
givinostat (3)
pan HDACi
Phase III
prototype pan HDACi
FDA approved 2006
N
H
O
O
N
N
H
O
O
O
N
N
H
N
H
OH
F
N
O
N
NH2
N
N
H
O
F
F
H
O
NH2
O
PCI-34051 (4)
HDAC8 selective
preclinical
entinostat (5)
HDAC1, HDAC3 selective
Phase III
PDA-106 (6)
HDAC1, HDAC3 selective
preclinical
TMP-269 (7)
class IIa selective
preclinical
Figure 1. Representative HDACi 1–7 of diverse chemotypes in different stages of clinical development. The pharmacophore of HDACi is annotated on the structure
of vorinostat (1). SBG: Surface Binding Group, ZBG: Zinc Binding Group.
We have recently demonstrated that incorporation of
a
study is to design and synthesize PRPs based on 1 and other
common HDACi 2–7 (Figure 1) and validate them in target
engagement studies in live cells in culture and in tissues in situ.
To accomplish this goal, we developed a general synthetic
strategy that incorporates a TFPA and an alkyne reporter groups
as an integral part of the SBGs of HDACi, designed and
synthesized seven novel PRPs 9–15 (Figure 2), evaluated their
biochemical potency and selectivity for individual recombinant
HDAC isoforms, and conducted photolabeling studies in four cell
lines in culture and in mouse liver tissue.
photoreactive tetrafluorophenyl azide (TFPA) group and an
alkyne reporter into the surface binding group (SBG) resulted in a
potent, relatively non-selective, and cell permeable photoreactive
probe (8, Figure 2) that structurally resembles an FDA approved
HDACi (1, Figure 1).[6f] Photolabeling with this probe showed that
engagement of HDAC isoforms is dependent on the
posttranslational modification state of HDACs, which is cell-type
dependent and generally does not correlate with the selectivity
profile measured in biochemical assays. The goal of the current
F
R
H
N
N3
F
F
R
O
R
H
N
H
N
H
N
OH
R =
R
OH
N
H
OH
OH
O
O
O
O
F
N
8
9
10
11
N
O
O
H
N
H
N
F
N
O
H
R
N
N
F
N
R
N
H
H
OH
O
N
F
R
NH2
R
O
NH2
O
TFPA
12
13
14
15
16
Figure 2. Tetrafluorophenylazide (TFPA) photoreactive probes 8–15 designed based on diverse chemotype HDACi and TFPA control
compound 16.
and stable in biological systems, yet reactive under copper(I)-
catalyzed “click reaction” conditions. Given the extensive
structure-activity relationships (SAR) available for most of the
Results
parent compounds, the TFPA and the alkyne moieties were
placed at positions deemed to tolerate the same size substituents
PRP Design and Chemical Synthesis
when possible. The ZBG and the linker regions were unaltered to
maintain similarity to the parent HDACi.[12] All the parent HDACi
and the corresponding PRPs were docked to the binding site of
HDAC isoforms as described previously[13] to ensure that the
TFPA and the reporter alkyne moieties do not interfere with the
binding of the PRPs. The in silico physicochemical properties;
octanol/water partition coefficient (SlogP), solubility (logS), and
topological polar surface area (TPSA) of PRPs and the parent
HDACi were calculated and compared to those of the parent
HDACi (Figure S1 and Table S1, Supporting information).
Generally, the differences in SlogP, logS, and TPSA between the
parent HDACi and PRPs were found to be comparable or smaller
than those for the previously published PRPs.
The PRPs synthesized in this work are based on diverse parent
HDACi with varying ZBG, linker and SBG. Parent HDACi
containing three ZBGs were chosen to synthesize PRPs 8–15.
These include either hydroxamic acid (8–12), o-aminoanilide (13
and 14), or trifluoromethyloxadiazole (15) (Figure 2). Linker
regions of the PRPs contained both saturated and unsaturated
aliphatic and aromatic moieties similar to those in the reported
HDACi (Figure 1). Although several photoreactive groups are
accessible synthetically, the TFPA was chosen based on its
similarity to aryl-based moieties present in the SBG of most
HDACi, short half-life of the corresponding nitrene to capture
interactions with HDACs (and possibly other proteins) while PRPs
are still in the binding site of HDACs,[10] and fewer side-reactions
that could decrease the yield of the PRP-protein adducts.[11] The
alkyne handle was chosen because it is small, electronically inert,
PRPs 8 and 9 were designed based on the chemotype of 1
(Figure 1). PRP 8 isosterically replaces the phenyl ring of 1 with
2
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10.1002/cmdc.201900114
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the TFPA and alkyne moieties so that only a single phenyl ring is
present in the SBG. PRP 9 includes the TFPA and alkyne handle
extended off the phenyl ring of 1. PRP 10 was designed based on
the chemotype of the potent pan HDACi panobinostat, 2 (Figure
1) and its published SAR.[14] PRP 11 represents aromatic
hydroxamic acid class of HDACi similar to givinostat (3). The
chemotype of HDAC8 selective compound PCI-34051 (4) was
used to design PRP 12. Replacement of the p-methoxy phenyl
substituent at the indole nitrogen with the TFPA group and
attachment of the alkyne handle at the indole C-3 were chosen as
both positions could tolerate substituents of similar electronic and
steric properties while retaining HDAC8 selectivity.[15] PRPs 13
and 14 were based on the o-aminoanilide-based inhibitors
selective for HDAC1 and HDAC3,[16] entinostat (5) and PDA-106
(6), respectively. PRP 15 was designed by replacing the SBG of
TMP-269 (7), a class IIa selective inhibitor,[17] with TFPA and
alkyne moieties. Compounds 16 (Figure 2 and Scheme S1,
Supporting information) and S20 (Scheme S2, Supporting
information) were designed to serve as a control for non-specific
photolabeling and as a competitor lacking the alkyne tag,
respectively.
To incorporate the TFPA and alkyne moieties into a structurally
diverse set of HDACi chemotypes, the synthetic strategy needed
to be amenable to the reaction conditions used in the synthesis of
the parent compounds. For example, most parent compounds
utilize reductive amination for amine synthesis, hydroxylamine
hydrochloride reaction with esters to form hydroxamic acid ZBG,
or acid catalyzed Boc-deprotection to form benzamide ZBG. We
found that the TFPA was not stable in most of the standard
reaction conditions used for the synthesis of the parent
compounds (Scheme S3, Supporting information). To circumvent
this issue, our optimized general strategy installs the TFPA in the
final steps of the synthesis from a perfluorobenzyl (PFB) group, in
the presence of a protected ZBG. Further deprotection of the
ZBG in mild conditions allows to retain all the other moieties intact.
Our optimized general strategy for HDAC PRPs synthesis is
outlined in Scheme 1 and detailed for PRPs 9–15 in Schemes 2–
6. The synthetic schemes for the non-HDACi control 16 and the
competitor S20 are provided in Schemes S1 and S2, respectively.
F
F
F
F
F
F
F
F
F
N3
F
F
O
PG
ZBG
PG
ZBG
F
F
N
F
N
ZBG
linker
linker
linker
H N
+
+
2
Br
Scheme 1. General synthetic strategy for PRPs 8–15. PG: protecting group.
The synthesis of PRPs 9 and 14 is shown in Scheme 2. A
reductive amination reaction of pentafluorobenzaldehyde with p-
(aminomethyl)aniline 17 gave secondary amine 18. Coupling of
suberic acid monomethyl ester with either O-tritylhydroxylamine
19 or N-boc-1,2-phenylenediamine 20 followed by ester
hydrolysis gave acids 21 or 22, respectively. Coupling of 18 with
alkylated with propargyl bromide resulting in 25 or 26, respectively.
The intermediates 25 and 26 were subject to regioselective
azidation to yield the TFPA derivatives 27 or 28, respectively.
Detritylation of 27 was carried out under mild Lewis acid catalyzed
conditions to give PRP 9. Deprotection of 28 under acidic
conditions gave the o-aminoanilide PRP 14.
either 21 or 22 gave the secondary amines 23 or 24 that were
F
NH2
F
F
F
NH2
a
H
H2N
N
F
F
H
H
F
F
F
N
N
17
F
18
c
R
6
H
N
O
O
NH2
Boc
H
Ph
b
HO
N
Ph
Ph
23: R = O-Trt
R
6
NH2 or
O
24
: R = Ph-2-NHBoc
N
O
O
H
20
21: R = O-Trt
22:
19
R = Ph-2-NHBoc
F
F
F
F
F
N3
F
H
N
H
N
H
H
N
R
d
e
N
R
F
6
6
F
N
O
O
F
N
O
O
f
9
:
R = OH
27
28
: R = O-Trt
: R = Ph-2-NHBoc
25: R = O-Trt
26
g
14
: R = Ph-2-NHBoc
: R = Ph-2-NH2
Scheme 2. Synthesis of PRPs 9 and 14. Reagents and Conditions: a) 1. perfluorobenzaldehyde, DCM,18 h, 2. NaBH4, CH3OH, 0 °C→rt, 4 h; b) 1. octanedioic
acid monomethyl ester, EDC•HCl, HOBt, DMAP, Et3N, CHCl3, 18 h, 2. NaOH, 2:1 MeOH/H2O, 8 h; c) EDC•HCl, HOBt, Et3N, CHCl3, 12 h; d) propargyl bromide,
K2CO3, CH3CN, 16 h; e) NaN3, Bu4NN3, DMF, 75–80 °C, 18 h; f) MgBr2, DCM, 30 min; g) HCl, 1,4-dioxane, 0 °C, 3 h.
In Scheme 3, the synthesis of PRP 10 started with the procedure
reported for its parent compound 2.[14] The resulting aldehyde 31
was reacted with propargylamine under reductive conditions to
give 32 that was alkylated with PFB bromide to give the amino
ester 33. A sequence of hydrolysis, tritylation, azidation, and
deprotection yielded the final PRP 10. The synthesis of PRPs 11
3
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and 13 is shown in Scheme 4. A reductive amination reaction of
methyl 4-formylbenzoate 37 with propargyl amine gave the
secondary amine 38 that was alkylated with PFB bromide and
hydrolyzed to the corresponding acid 40. Coupling of acid 40 with
either 19 or 20 followed by azidation and deprotection gave PRPs
11 or 13, respectively. PRP 12 was synthesized as shown in
Scheme 5. Alkylation of N-1 nitrogen in commercially available
indole 45 followed by reductive amination with N-methyl propargyl
amine gave ester 47. Hydrolysis of 47 followed by reaction with
19 resulted in ester 49 that was subjected to azidation and
deprotection to give PRP 12. The synthesis of PRP 15 is shown
in Scheme 6. Trifluoromethyloxadiazole 53 was synthesized
following previously reported procedure.[17] The perfluorophenyl
group and the alkyne moieties were reacted with ethylene
diamine 54 via reductive amination followed by alkylation to give
58. Next, coupling of 53 and 58, followed by azidation gave the
hydroxamic acid formation from esters. Several rounds of
experimentation with different conditions and synthetic routes
showed that coupling of the corresponding carboxylic acid with O-
trityl hydroxylamine permitted introduction and maintenance of
the masked hydroxamic acid ZBG during all the steps of the
synthesis (Scheme S3, A). Azidation reaction of the PFB group
with trimethylsilyl azide resulted in low yields of the desired TFPA
product irrespective of the temperature, solvent, and the duration
of the reaction. It was found that the reaction between PFB-based
intermediates and stochiometric amount of sodium azide and
catalytic amount of tetrabutylammonium azide, which was
adopted from previously reported procedure,[18] gave 25–95%,
yields. The temperature and reaction time needed optimization for
each chemotype to ensure complete conversion of PFB to TFPA.
It was also important to monitor completion of the reaction using
liquid chromatography mass-spectrometry analysis (LC-MS) and
19F NMR as the starting PFB derivatives often exhibited identical
retention times and were inseparable by chromatography.
Lowering reaction temperature required longer reaction times
resulting in formation of additional side products and a reduced
yield. The optimal temperature and duration of this reaction were
ca. 80 °C and ca. 18 h, respectively, except for intermediate 50,
for which this reaction had to be conducted at 50 °C for 22 h to
final product 15.
F
O
O
X
F
F
O
Br
R
O
c
F
N
a
29
+
R
O
33
: R = OCH3, X = F
obtain acceptable (≥ 25%) reaction yields (Scheme S3, C).
d
e
31
: R = CHO
O
F
b
34: R = OH, X = F
35: R = NH-OTrt, X = F
36
47
: R = OCH3, X = F
30
X
F
F
32
: R =
HN
c
O
48
49
: R = OH, X = F
d
f
O
O
F
N
: R = NH-OTrt, X = F
: R = NH-OTrt, X = N3
10: R = NH-OH, X = N3
b
g
e
f
R
50: R = NH-OTrt, X = N3
12: R = NH-OH, X = N3
N
FB
Scheme 3. Synthesis of PRP 10. Reagents and conditions: a) Pd(OAc)2,
NaOAc, NMP, 120 °C, 1 h; b) propargylamine, STAB-H, DCE, 16 h; c)
perfluorobenzylbromide, K2CO3, CH3CN, 14 h; d) LiOH, 2:1 THF/H2O, 20 h; e)
19, EDC•HCl, HOBt, DMAP, Et3N, CHCl3, 6 h; f) NaN3, Bu4NN3, DMF,75–80 °C,
18 h; g) MgBr2, DCM, 3 h.
Scheme 5. Synthesis of PRP 12. Reagents and conditions: a)
perfluorobenzylbromide, NaH, DMF, 14 h; b) N-methylpropargylamine, STAB-
H, DCE, 5 h; c) LiOH, 6:3:1 THF/H2O/dioxane, 12 h; d) 19, EDC•HCl, HOBt,
DMAP, Et3N, CHCl3, 6 h; e) NaN3, Bu4NN3, DMF, 50 °C, 22 h; f) MgBr2, DCM,
3 h. PFB: perfluorobenzyl.
F
X
F
F
O
O
R
O
O
O
N
R1
N
R2
H
OH
O
N
O
F
N
O
O
F
F
F
N
N
NH2
N
HO
a
d
HO
HO
a
d
b
38: R1 = H,
: X = F,
52
F
53
51
41
R = O-Trt
43
: X = F,
R = Ph-2-NHBoc
b
c
e
f
e
g
R
39
R
2 = OCH3
F
H
: R1 = PFB,
: X = N3
,
,
: X = N3,
R = Ph-2-NHBoc
37
42
R = O-Trt
44
56: R1 = Boc, R2 = H
N
or
F
F
2 = OCH3
H2
N
R
e
: R1 = PFB,
40
R
11: X = N3
R = OH
13: X = N3,
R = Ph-2-NH2
: R1 = Boc, R2
: R1 = H, R2
=
=
57
2 = OH
F
54
55
: R = H
H
f
N
c
58
N
R1
: R = Boc
Scheme 4. Synthesis of PRPs 11 and 13. Reagents and Conditions: a) 1.
propargylamine, DCM, 16 h, 2. NaBH4, CH3OH, °C→rt, h; b)
R2
0
4
F
pentafluorobenzyl bromide, K2CO3, CH3CN, 16 h; c) LiOH, 6:3:1
THF/H2O/dioxane, 24 h; d) 19 for 41 or 20 for 43, EDC•HCl, HOBt, DMAP, Et3N,
CHCl3, 12 h; e) NaN3, Bu4NN3, DMF, 75–80 °C, 18 h; f) MgBr2, DCM, 3 h; g)
HCl, 1,4-dioxane, 0 °C, 6 h. PFB = perfluorobenzyl.
X
F
F
O
N
F
F
O
53
+
58
g
F
N
N
N
H
59: X = F
15
F
h
: X = N3
As mentioned above, azidation of the PFB group was performed
near the end of each synthesis as the resulting arylazido TFPA
group was found to be unstable under the reaction conditions
typically used to prepare most of the parent HDACi. Specifically,
we found that the TFPA group could not withstand conditions
necessary for reductive amination, palladium-catalyzed cross-
coupling, and reactions of hydroxylamine hydrochloride under
basic conditions including amidoxime formation from nitriles and
Scheme 6. Synthesis of PRP 15. Reagents and conditions: a) NH2OH, Na2CO3,
1:1 MeOH/H2O, Reflux, 4 h; b) trifluoroacetic anhydride, 0→50 °C, pyridine, 5
h; c) (Boc)2O, CHCl3, 0 °C→rt; d) 1. perfluorobenzaldehyde, DCM, 8 h, 2. NaBH4,
CH3OH, 0 °C–rt, 3 h; e) propargyl bromide, K2CO3, CH3CN, 48 h; f) HCl, 1,4-
dioxane, 1 h; g) EDC•HCl, HOBt, DMAP, Et3N, CHCl3, 12 h; h) NaN3, Bu4NN3,
DMF, 80 °C, 18 h.
4
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Biochemical Selectivity
For hydroxamate PRPs, detritylation of intermediates 27, 36, 42
and 50 (Schemes 2–5) was carried out using a Lewis acid
mediated deprotection.[19] Protic acids commonly used for
deprotection, such as trifluoroacetic acid and hydrochloric acid,
were found to be incompatible in deprotection of intermediates
containing TFPA moiety, resulting in a complex mixture of
products (Scheme S3, D). Deprotection of Boc intermediates 28
and 44 (Schemes 2 and 4) required an ice-cooled reaction for 3–
6 h. Longer reaction times or reactions at room temperature gave
the previously reported benzimidazole cyclization side product as
detected by LC-MS (Scheme S3, E).[20]
It is also important to mention that mild hydrolysis conditions,
LiOH in THF/water at room temperature, were used for hydrolysis
of PFB esters 33, 39, and 47, whereas methyl esters of 21 and 22
were hydrolyzed with NaOH in methanol/water. The latter
conditions in the case of 33, 39 and 47 or heating above 40 °C
resulted in para- hydroxylated PFB side products (as detected by
LC-MS and NMR) that were inseparable from the desired
products in the subsequent reaction steps (Scheme S3, B).
The PRPs and their parent compounds were tested for HDAC
inhibitory activity against recombinant class I HDACs as well as
HDAC4, as a representative member of class II isoforms, using a
standard experimental procedure with
a
competitive
fluorescence-based assay commonly used in the field.[21] Briefly,
inhibition of HDAC1, 2, and 3 was measured using the fluorescent
HDAC substrate Boc-L-Lys(Ac)-AMC and commercially available
recombinant human HDAC1, 2, and 3, whereas inhibition of
HDAC8 was measured using the BML-KI178 HDAC8 substrate
and a recombinant human HDAC8.[22] The pIC50 values for PRPs
8–15 compared to their parent HDACi 1–7, are listed in Table 1
and their HDAC isoform selectivity profile is given in Table 2. The
dose-response curves for all tested compounds are shown in
Figure S2, Supporting information. The potency of PRPs was
generally 10–100-fold lower than that of their parent compounds
(Table 1), whereas the isoform selectivity profile of PRPs was
comparable and in one case, PRP 13, HDAC3 selectivity was
superior to that of its parent HDACi 5 (Table 2).
Table 1. Biochemical pIC50 values of PRPs and their parent HDACi for HDAC1, 2, 3, 8 and 4.
pIC50 ± SE[a]
HDAC3
Compound
HDAC1
HDAC2
HDAC8
HDAC4
vorinostat (1)
PRP 8
7.67
±
0.01
6.92
±
0.02
7.74
±
0.02
6.61
5.64
5.30
6.41
5.12
±
±
0.06
0.11
0.10
0.20
0.09
NA[b]
6.26
6.13
9.52
5.56
±
0.04
0.07
0.10
0.07
6.77
5.95
9.70
5.32
±
0.04
0.08
0.46
0.05
6.96
6.46
9.20
5.69
±
0.06
0.10
0.20
0.06
4.42
±
0.07
PRP 9
±
±
±
±
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
NA[b]
±
panobinostat (2)
PRP 10
±
±
±
±
±
±
±
±
givinostat (3)[23]
PRP 11
6.70[c]
6.49[c]
6.80[c]
6.07[c]
±
5.43
5.50
3.33
±
0.09
0.08
0.14
4.77
4.60
4.19
±
0.01
0.03
0.06
4.96
4.94
5.30
±
0.02
0.04
0.03
5.10
8.48
5.67
0.06
0.06
0.08
PCI-34051 (4)
PRP 12
±
±
±
±
±
±
±
±
entinostat (5)[24]
PRP 13
6.61[c]
6.34[c]
6.61[c]
<5[c]
< 3
±
5.09
6.33
3.91
4.33
3.95
±
±
±
±
±
0.09
0.07
0.05
0.10
0.14
5.10
6.54
3.37
4.44
4.21
±
±
±
±
±
0.12
0.05
0.11
0.11
0.04
5.82
7.66
5.43
4.24
±
±
0.08
0.02
0.08
0.07
PDA-106 (6)
PRP 14
5.24
2.81
0.09
0.43
±
< 3
< 3
±
TMP-269 (7)
PRP 15
±
4.54
3.40
0.11
0.78
< 3
±
[a]pIC50 is equal to (- log IC50) and SE is standard error, both were calculated by non-linear regression analysis (enzyme-inhibitor model) using GraphPad prism
7.02. Values reported are the mean ± SE at least 2 replicate experiments; the numbers are rounded to three significant figures. [b]NA: less than 50% inhibition at
100 µM. The percent of inhibition at 10 and 100 µM is given in Table S2, Supporting information. [c]pIC50 of both 5 and 6 were calculated based on reported
biochemical inhibition data for both using similar assay in the annotated references.
PRP 8, with high similarity to the structure of its parent HDACi 1,
showed the highest potency among all the PRPs with pIC50 of 6.3,
6.8, 7, and 5.6 against HDAC1, HDAC2, HDAC3, and HDAC8,
respectively (Table 1). PRP 9 was ca. 3-fold less potent than PRP
8 against all tested isoforms with pIC50 of 6.1, 6.0, 6.5 and 5.3
against HDAC1, HDAC2, HDAC3, and HDAC8, respectively
5
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(Table 1). PRP 10, with pIC50 of ca. 5.5 for HDACs 1-3, was 10-
fold less potent than PRPs 8 and 9 and 10,000-fold less potent
than the parent HDACi 2, which exhibited pIC50 of ca. 9.5 for
HDACs 1-3. Potency of 10 against HDAC8, pIC50 of 5.1, was
comparable to those of 8 and 9 and was 19-fold lower than that
of 2, pIC50 of 6.4 (Table 1). PRP 11, with an average pIC50 of ca.
5.0, was 30-fold less potent than the parent HDACi 3, whose
average pIC50 was ca. 6.5 against class I HDAC isoforms (Table
1). PRPs 8–11 showed little to no selectivity to any of class I
HDAC isoforms (Table 2).
PRP 12 was 200-, 30-, and 2-fold more selective for HDAC8
(Table 2) with a pIC50 of 5.7 as compared to pIC50 of 3.3, 4.2, and
5.3 for HDAC 1, 2, and 3, respectively (Table 1). PRPs 13 and 14
did not inhibit HDAC8 at the maximum tested concentration, 1 mM
(Table 1). PRP 13 exhibited pIC50 of 5.1, 5.1, and 5.8 for inhibition
of HDAC1, 2, and 3, respectively (Table 1). PRP 14 had a pIC50
of 3.9, 3.4 and 5.4 against HDAC1, HDAC2 and HDAC3,
respectively (Table 1). PRP 13 was 15-, 50-, and 3-fold more
potent than 14 against HDACs 1, 2, 3, respectively. Both PRPs
13 and 14 showed selectivity for HDAC3 over HDAC1 and
HDAC2 higher than that of their respective parent HDACi 5 and 6
(Table 2). We found that both PRP 15 and its parent HDACi 7,
which was previously reported as being selective for class II
HDACs,[17] were not selective for the class II HDAC4 and were
generally weak inhibitors of all the tested isoforms, with pIC50
ranging between 3 and 4 for both compounds (Table 1). The latter
finding highlights the variability in the results of the biochemical
assays usually encountered due to differences in experimental
conditions.
Table 2. Biochemical selectivity profiles for PRPs 12–15 and their parent HDACi
(4–7).
Selectivity[a]
Compound
HDAC1
HDAC2
HDAC1 HDAC1 HDAC2
HDAC3 HDAC8 HDAC3
HDAC2 HDAC3
HDAC8 HDAC8
vorinostat (1)
PRP 8
5.6
0.31
1.5
0.85
0.20
0.47
11
4.2
6.8
0.15
0.65
0.31
2.0
13
13
21
14
In-cell Photoaffinity Labeling
PRP 9
4.5
Following our previously published procedure[6f] outlined in Figure
3, photolabeling experiments were performed in live SET-2
(Figure 4), HepG2, HuH7 and HEK293T cells (Figure 5).
Corresponding densitometric analysis of selected class I HDAC
labeling is given in Figures S3 and S4 (Supporting information).
Briefly, cells in culture were incubated with either DMSO or
compound 16 as a control, PRP or a competitor HDACi and PRP
and incubated for 40 min with PRPs 8–12 and 15 or for 3 h with
PRPs 13 and 14, irradiated with UV light at 366 nm, washed, and
lysed. The labeled lysate was subjected to “click reaction”
conditions with an azide-conjugated 800CW IRDye and
electrophoretically separated. Antibodies for individual HDAC
isoforms were added followed by incubation with 680RD-IRDye
conjugated secondary antibodies. Visualization in both 800 and
700 nm channels show PRP labeled proteins as well as HDAC
antibody bound proteins, respectively. The PRP labeled bands
(800 nm), that were absent from the controls and decreased by at
least half upon addition of a competitor, were considered specific.
These bands (800 nm) were assigned to a particular HDAC
isoform if they counter-stained with an HDAC antibody (700 nm),
Figure 3. The concentration of the PRPs was optimized to achieve
the highest signal-to-noise ratio (Figures S5 and S6, Supporting
information) and was kept within the HDACi concentration range
commonly used in cell-based studies[15, 25] and typically observed
in cancer patients undergoing HDACi based therapy.[26]
Panobinostat
0.66
2.1
1300
3.2
1900
620
(2)
PRP 10
givinostat (3)
PRP 11
1.7
1.6
0.74
0.79
3.0
2.8
4.3
0.43
0.49
1.6
2.6
3.7
5.4
4.6
2.1
0.65
0.47
0.72
PCI-34051 (4)
PRP 12
7.9
3.6
0.0010
0.0046
> 40[b]
> 120[b]
12
0.46
0.00013
0.033
> 22[b]
> 130[b]
20.0
0.00029
0.43
0.14
1.9
0.011
1.0
0.078
0.54
entinostat (5)
PRP 13
> 41[b]
> 660[b]
260
0.98
0.62
3.5
0.19
0.047
0.030
1.2
0.19
PDA-106 (6)
PRP 14
0.076
0.0087
1.6
> 8.1[b]
> 21[b]
14
> 2.3[b]
> 28[b]
25
> 270[b]
> 17[b]
1.6
TMP-269 (7)
PRP 15
0.78
0.55
> 8.9[b]
> 16[b]
[a] Selectivity ratios based on IC50 measurements; the numbers are rounded
to two significant figures. [b]An estimate based on the highest concentration
tested.
Figure 3. Experimental workflow of photoaffinity labeling in live cells and gel-based visualization of labeled proteins.
In SET-2 cells, the bands observed with TFPA control compound
16 were the same as those observed with the DMSO control
(Figures 4a and 4b), indicating that the TFPA moiety itself would
have little effect on the labeling with PRPs. Labeling of HDAC8
was difficult to interpret due to a non-specific band found in the
control lanes at the same level as HDAC8 in SET-2 cells (Figure
6
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4a). PRP 9, which is a relatively potent inhibitor of all class I
HDACs (pIC50 ca. 6), showed labeling of HDACs 1-3 higher than
that observed with 8 (Figure 4a and Figure S3a, Supporting
information). PRPs 10 and 11 showed weak labeling of HDAC 1
and 2 isoforms only marginally different from the control (Figure
4a and Figure S3a, Supporting information). The HDAC8
selective PRP 12, showed weak labeling of HDAC1 and 2 and
distinct labeling of HDAC3. Despite being a weak HDAC inhibitor
in biochemical assay, PRP 15 labeled HDAC2 and 3, but not
HDAC4 in live cells (Figure 4a and Figure S3a, Supporting
information). The absence of HDAC4 labeling is likely due to low
abundance of HDAC4 as we could not detect it by HDAC4 specific
antibody in this type of cells. Unexpectedly, both HDAC3 selective
PRPs 13 and 14 labeled mostly HDAC2, to a lesser extent
HDAC3, and (Figure 4b and Figure S3b, Supporting information).
Labeling of HDAC1 was indistinguishable from background. The
labeling bands of HDAC2 and HDAC3 by PRP 14 were further
confirmed by competition with the parent HDACi 6 (Figure 4c).
PRP 14, although being a relatively weak HDAC2 inhibitor in the
biochemical assay (pIC50 3.4), labeled HDAC2 even at
concentrations as low as 200 nM (Figure 4d). In preliminary
labeling experiments using recombinant HDAC1 (Figure S7,
Supporting information), co-incubation with the covalent
competitor S20 showed a ca. 75% decrease in labeling with PRP
8 compared to 25% decrease when co-incubated with 1. In live
SET-2 cells, however, cellular levels of HDACs 1–3 and 8 were
decreased upon co-treatment with S20 and both S20 and 1 had
a comparable 50% decrease in labeling of HDAC2 with PRP 8
(Figure S7, Supporting information).
Figure 4. Photolabeling with PRPs 8–15 in live SET-2 cells. Labeling with (a) hydroxamate PRPs 8–12 and TFMO PRP 15, (b) o-aminoanilide PRPs 13 and 14. In
each of (a) and (b), vehicle DMSO (–) and 16 were included as controls for nonspecific bands. (c) competition of 14 HDAC-labeled bands with the HDACi 6. (d)
concentration dependent labeling of HDACs by 14. In (c) and (d), densitometry of HDAC2 labeling bands normalized to HDAC2/GAPDH signal is shown to the right
of each gel.
Photolabeling of 9 and 14 was further characterized in HepG2,
HuH7, and HEK293T cells (Figure 5). In HepG2 (Figure 5a
and Figure S4a, Supporting information) and HuH7 (Figure 5b
and Figure S4b, Supporting information), labeling patterns
were similar to those observed in SET-2 cells for both 9 and
14, with higher labeling of HDAC2 for PRP 14. In HEK293T
(Figure 5c and Figure S4c, Supporting information), 14
maintained its superior labeling of HDAC2, however, to a
lesser extent than in HepG2 and HuH7 cells. Labeling of
HDAC1 in HEK293T was observed with both 9 and 14.
Immunoblotting with anti-HDAC 1 and 2 antibodies showed
only minor differences in the protein abundance in SET-2,
HepG2, HuH7 and HEK293T cells, which does not explain the
decreased labeling of HDAC2 by 14 in HEK293T cells (Figure
S8, Supporting information).
Figure 5. Photolabeling with PRPs 9 and 14 in (a) HepG2, (b) HuH7 and (c) HEK293T live cells. DMSO (–) and TFPA 16 were included as controls. IB with class I
HDAC antibodies and GAPDH is shown to the right at the level of molecular weight for each protein. Top right part of each panel shows detection of HDAC2 labeling
by IR dye (800 nm) and HDAC2 antibody (700 nm) and overlap of both labeling band and antibody (700 + 800 nm).
7
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In-tissue Photoaffinity Labeling
to the last step, to avoid formation of side-products or losing the
target compounds completely. Although PRPs containing TFPA
and other azide-based photoreactive groups are typically stable
in dark below 0 °C from several weeks to few months, they
degrade upon prolonged storage. Ability to store a stock of non-
photoreactive intermediates that can be readily converted to
PRPs in a few straightforward experimental steps is expected to
prevent the loss of PRP during storage and to yield consistent
results in photolabeling experiments. The overall applicability of
this synthetic strategy was demonstrated with three different
ZBGs, two of which can be found in the majority of currently used
HDACi. Although further SAR studies may be needed, the
convergent approach developed here can be used to conduct the
necessary optimization as well as to extend this strategy to other
not yet explored HDACi chemotypes.
To investigate the feasibility of labeling HDACs with TFPA PRPs
in tissues, PRP 8 was incubated with mouse liver homogenate
and the PRP-labeled lysates were subject to photolabeling in a
workflow similar to that optimized for live cells in culture (Figure
3). Briefly, minced fresh mouse tissue was incubated with either
DMSO or PRP 8 or pretreated with 1 and then incubated with 8
and irradiated with 366 nm UV light. The amount of tissue was
optimized to enable detection of class I HDAC isoforms. The
labeled tissue was lysed, and the resulting lysate was reacted with
the azide-conjugated biotin tag. The PRP labeled bands were
detected by 800CW IRDye-conjugated streptavidin. HDAC-
specific labeling bands were absent from DMSO control,
decreased by competition with 1 and counterstained with HDAC-
specific antibodies (Figure
6 and Figure S9, Supporting
Incorporation of the photoreactive and reporter handle groups
resulted in a variable moderate decrease in potency of the
resulting PRPs compared to their parent HDACi. The biochemical
activity of the PRPs against HDACs 1-3 and 8 was between 125
nM and 1 mM (Table 1) and the selectivity was similar to that of
the parent HDACi (Table 2). In general, PRPs 8–15 displayed
robust labeling of class I HDAC isoforms in cells in culture. Among
all the PRPs tested, PRP 14 showed the best signal-to-noise ratio
and was able to detect HDAC2 in SET-2 cells even at 200 nM
(Figure 4d). We also demonstrated that PRP 8 can label class I
HDAC isoforms in mouse liver tissue. To the best of our
knowledge, this is the first published example of labeling HDACs
information). The band corresponding to HDAC8 and recognized
with an anti-HDAC8 was relatively low in intensity compared to
that in experiments with live cells. We attribute this to the use of
anti-HDAC8 antibody raised against human HDAC8. PRP 8
labeled HDAC1-3 and 8 with the most pronounced bands for
HDAC8 (Figure 6 and Figure S9, Supporting information).
in tissue with
a small-molecule-based PRP. Unlike other
commonly used in biology approaches to study HDAC
engagement, application of PRPs is comparably faster,
inexpensive, and, most importantly, can be done in the commonly
used biological models without modifications. Our studies were
conducted in live cells in culture without overexpression of
HDACs, suggesting that the sensitivity of the newly designed
PRPs and photolabeling approach in general are sufficient to
detect endogenous levels of HDACs in cells with unaltered
machinery. This would be particularly useful in situations where
molecular biological manipulations with the cellular machinery are
experimentally or methodologically undesirable or impossible.
The differences between the biochemical and cell-based
photolabeling selectivity are observed for all the PRPs in this
study. Our analysis shows that there are four types of differences.
In the first type, less potent in biochemical assays PRPs 9 and 14
display higher labeling of HDACs compared to more potent
congeners, 8 and 13, respectively (Figures 4a and 4b and Figure
S3, Supporting information). In the second type, the isoform
selectivity patterns of PRPs 12–15 determined in the biochemical
assays do not match those observed by photolabeling in live cells.
For instance, PRP 14, despite selectivity for HDAC3 in the
biochemical assay (Table 2), showed pronounced labeling of
HDAC2 in SET-2 (Figure 4b and Figure S3b, Supporting
information), HepG2 (Figure 5a and Figure S4a, Supporting
information), and HuH7 (Figure 5b and Figure S4b, Supporting
information). In the third type, same PRPs targeted different
HDAC isoforms in cell-type dependent manner. For example,
PRR 14 showed higher HDAC2 labeling in HepG2 and HuH7 cells
(Figures 5a and 5b) compared to that in HEK293T cells (Figure
5c), which could not be explained by the difference in isoform
abundance (Figure S8, Supporting information). Whether this is
due to higher activity of HDAC2 in HepG2 and HuH7 cell lines[27]
and higher or equal to HDAC2 activity of HDAC1 in HEK293T cell
Figure 6. (a) Photolabeling with PRP 8 in mouse liver tissue, reacted with azide-
conjugated biotin and incubated with 800CW IRDye-conjugated streptavidin (b)
left: HDAC labeling bands as visualized in (a), right: immunoblotting with
680RD-IRDye conjugated HDAC1, 2, 3, and 8 antibodies.
Discussion
HDAC isoform engagement with HDACi is an important issue in
development of HDAC-based therapeutics. It still needs a robust
and readily implemented solution in typical biological settings and
models. In this paper, we have designed and synthesized a series
of novel HDAC PRPs (Figure 2) based on known HDACi (Figure
1) with diverse chemotypes and biochemical selectivity profiles.
To ensure structural and chemical uniformity and direct
systematic comparison of different chemotypes, the same
photoreactive group and reporter handle were used for all PRPs
in this study. We also developed a synthetic strategy that is
compatible with the TFPA photoreactive moiety and other groups
in the pharmacophore of HDACi (Scheme 1) that, we anticipate,
will enable others to access even broader set of HDAC PRPs for
target engagement studies.
We found that the convergent minimalistic synthetic approach
described here resulted in the expected PRPs in overall
acceptable yields with an individual yield for each reaction
throughout the synthesis of at least 25%, which is important for
scaling-up the synthesis of the PRPs. In the optimized synthetic
scheme, unmasking of the photoreactive moiety was postponed
8
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28]
line[2a,
remains to be investigated. In the fourth type,
straightforward and that HDAC PRPs would be a useful tool to
address it. We anticipate that the differences between the
biochemical profiles and the HDAC isoform engagement in live
cells, as determined by the PRPs, should also be observed for the
non-photoreactive parent HDACi, and should be taken into
consideration when analyzing the outcomes of ongoing clinical
trials of isoform-selective HDACi.[30] Further studies of the
phenomenon observed here may lead to exciting new discoveries
in biology of HDAC and novel inhibitors with potency and
selectivity engineered to be cell type selective.
chemotype-dependent labeling profile was observed for PRPs of
different ZBGs, o-aminoanilides 13 and 14 (Figure 4b, and Figure
S3b, Supporting information) versus hydroxamates 8–12 (Figures
4a and Figure S3a, Supporting information). Regardless of
variability in the labeling pattern within each class, the overall
labeling trend is characteristic of each ZBG chemotype. The
hydroxamate based-PRPs 8–12 display variable but consistent
ability to label HDACs 1-3 and 8, whereas o-aminoanilide-based
PRPs 13 and 14 labeled mostly HDAC2, to lesser extent HDAC3,
and did not label HDAC1, which binding site is highly homologous
to those of HDAC2 and HDAC3. This chemotype dependence of
target engagement is consistent with the observations by
Bantscheff et al[6a] and may be linked to the slow koff binding
kinetics of the o-aminoanilide-based HDACi.[29] Although our
efforts in this paper were focused on validation of photolabeling
using class I HDACs, differences between the photolabeling and
the biochemical profiles are expected for class II and IV HDACs
as well. The overall discordance between the biochemical and
cell-based relative selectivity is consistent with that recently
demonstrated by us for PRP 8 in photolabeling of all zinc-
dependent HDAC isoforms in different types of breast cancer
cells.[6f] These findings further underscore the importance of the
cellular context for target engagement.
Conclusions
In summary, a synthetic strategy to access diverse HDAC PRPs
was developed. Novel HDAC PRPs 9–15 based on seven HDACi
with diverse chemotypes 1–7 and biochemical selectivity profiles,
were synthesized, tested in the HDAC 1-3, 4, and 8 biochemical
assays, and validated in photolabeling experiments in live SET-2
cells. In addition, photolabeling with PRPs 9 and 14 was also
validated in HepG2, HuH7, HEK293T cell lines, whereas PRP 8
was also tested for photolabeling capacity in mouse liver tissue.
The synthetic strategy should be readily amenable for synthesis
of PRPs with chemotypes of other HDACi. In general, the HDAC
biochemical selectivity profiles of PRPs were similar to those of
their parent HDACi. The cell-based photolabeling and the
biochemical profiles of PRPs 9–15 were found to be substantially
different from each other and did not correlate. The labeling by
PRPs also did not show correlation with abundance of HDAC
proteins in cells. Both the PRP chemotype and the cell type were
key in defining what HDAC isoforms were labeled. This is the first
report of cell-permeable uniform set of HDAC PRPs based on
different HDACi chemotypes to probe engagement of multiple
HDAC isoforms in live cells and tissues. Further extension of
these studies to other types of cells and animal tissues as well as
identification of the mechanisms responsible for modulation of
HDACi selectivity and target engagement are in progress.
Figure 7. Hypothesis: Unlike scaffolding HDACs, catalytically active HDACs are
able to bind substrates, small molecule HDACi, and HDAC PRPs. PTMs and
HDAC protein complex components define whether the role of HDAC isoform is
catalytic or scaffolding in cell-type, cell-cycle, and subcellular location manner.
This study shows that the observed differences between the
biochemical and photolabeling profiles is a common phenomenon
that does not depend on the abundance of the HDAC proteins and
the biochemical potency of PRPs and is primarily driven by the
chemotype of the PRP and the type of cells. By comparing the
labeling of HDAC isoforms in different cell lines by the same PRP,
we can exclude the differences in binding conformations, yields in
click-reactions, and differences in photoactivation as factors for
the discordance between the biochemical and the HDAC
photolabeling profiles. It can, therefore, be argued that the cellular
context affects photolabeling by PRPs, and more importantly
HDAC isoform target engagement by HDACis, in several ways: 1)
by altering the binding affinity and kinetics of the PRPs to HDACs
via PTMs and/or conformational/accessibility changes due to
multiprotein complex formation, and/or 2) by varying the relative
abundance of the catalytically competent and catalytically inactive,
e.g. scaffolding, HDACs (Figure 7). Both these scenarios
individually or in combination would present themselves in the
same way in the photolabeling experiments in live cells. Although
we cannot exclude either of these scenarios, this report clearly
demonstrates that the answer to the question “What HDAC
isoform(s) does this HDAC ligand inhibit in cells/in vivo?” is not
Experimental Section
1. Biochemical and Biological Procedure
1.1. Fluorogenic Enzymatic Assays for Class I HDAC Isoforms
Human recombinant HDAC1, HDAC2, HDAC3 (BPS Bioscience) and
HDAC8 (In-house purified from E-coli) were diluted with assay buffer 1 (25
mM Tris–HCl, pH 8.0, 137 mM NaCl, 2.7 mM KCl, and 1 mM MgCl2, 1
mg/mL BSA) to give 4, 5, 1, and 8.5 ng/µL stocks of each isoform,
respectively. Serial dilutions of the compounds/PRPs were made in assay
buffer 2 (25 mM Tris–HCl, pH 8.0, 137 mM NaCl, 2.7 mM KCl, and 1 mM
MgCl2) starting with the highest concentration at 1 mM. 10 µL of the
enzyme stock and 30 µL of each of the serial dilutions were mixed in a
black half-area, low protein binding 96 well plate (Corning) and pre-
incubated for 5 minutes (3 hours for the o-aminoanilide derivatives, 6, 13
and 14) at room temperature (rt). Next, 10 µL of 125 µM Boc-L-Lys(Ac)-
AMC (BLA) fluorescent substrate (Chem-Implex) for HDACs 1-3 or 10 µL
of 25 µM Fluor de Lys®, BML-KI178 (Enzo Life Sciences) for HDAC8 was
added to each well and incubated for 30 minutes at rt. The reaction was
quenched by the addition of 50 µL of 1 mg/mL trypsin and 5 µM TSA in
assay buffer
2 and incubated for an additional 30 minutes. The
fluorescence signal was read at excitation wavelength 360 nm and
emission wavelength 460 nm using a Synergy 4 hybrid microplate reader
9
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from BioTek. The statistical data analysis and IC50 values were determined
using GraphPad Prism 7.02.
Igepal, 5% glycerol, 1X PIC, 1X Phosphatase inhibitor cocktail),
homogenized by vortexing for 10 seconds at room temperature and placed
on a rotating stand overnight at 4 °C. The lysate was clarified by
centrifugation at 19,000g for 10 min at 4 °C. Protein concentration was
determined using BCA assay and diluted to 2 mg/mL. 35 µg of each lysate
was reacted a 0.1 mM azide-PEG3-biotin conjugate, Sigma Aldrich
762024 (1.7 mM stock in 1:4 DMSO:t-butanol) in the presence of 1 mM
CuSO4, 0.1 mM TBTA, 1 mM TCEP for 90 minutes at room temperature.
The reactions were then stored at -20 °C overnight. Precipitated proteins
were pelleted at 6000g for 5 min at 4 °C. Proteins were washed with 250
µL methanol by brief sonication and repelleted, then resuspended in PBS
containing 0.2% SDS by brief sonication. Proteins were then separated by
SDS-PAGE and transferred to nitrocellulose membrane (as
per 1.7. below). Membrane was blocked and incubated with IRDye®
800CW streptavidin (LI-COR) and labeled protein signal was recorded with
Odyssey Sa scanner at 800 nm. Immunoblotting with HDAC specific
antibodies was done as per 1.7.
1.2. Fluorogenic Enzymatic Assay for HDAC4
HDAC4 inhibitory activity was determined using the fluorogenic HDAC4
assay kit (BPS Bioscience, catalogue
# 50064) according to the
manufacturer procedure. The statistical data analysis and IC50 values were
determined using GraphPad Prism 7.02.
1.3. Cell Culture
The human megakaryoblastic cell line SET-2 was cultured in RPMI 1640
medium (Corning, VA, USA) supplemented with 10% heat-inactivated fetal
bovine serum, FBS (Sigma Aldrich, St. Louis, MO, USA). Cells were
seeded at the density of 1 × 105/ml of medium and were grown at 37°C
with 5% CO2. Cells were harvested after 72 hours, collected by
centrifugation, washed and viability was tested by trypan blue dye
exclusion. Collected cells were then re-suspended in RPMI 1640
supplemented with 10% heat inactivated fetal calf serum. A total of 30 ×
106 cells was spun down at 500 g for 10 min at 4 °C, re-suspended in 12
mL PBS and aliquoted into 0.5 mL on a 12-well plate for photolabeling
experiments.
1.6. Photolabeling of Recombinant Proteins
Recombinant HDAC1 (200 ng) in 10 µL whole cell lysis buffer (as per 1.4.)
was pretreated with 100 mM SAHA (1) or S20 or DMSO control for 15
minutes and then treated with PRP 8 (10 mM) or DMSO control for 40-min.
Samples were cooled on ice and irradiated with 366 nm light for 30 min
then incubated with 0.1 mM azide-PEG3- biotin conjugate, Sigma Aldrich
762024 (2.5 mM stock in DMSO) in the presence of 1mM CuSO4, 0.1 mM
TBTA, 1 mM TCEP for 90 minutes at room temperature. Samples were
then diluted with loading buffer, separated by SDS-PAGE and transferred
to nitrocellulose membrane (as per 1.7. below). Membrane was blocked
and incubated with IRDye® 800CW streptavidin and labeled protein signal
was recorded with Odyssey Sa scanner at 800 nm. Immunoblotting with
HDAC specific antibodies was done as per 1.7.
The human hepatocellular carcinoma HepG2 or HuH7, or the human
embryonic kidney HEK293T cells were plated in 6-well plates (0.5 x 106
cells) and grown to 80% confluence in 2 mL culture medium (RPMI 1640
supplemented with 10% FBS, penicillin-streptomycin, and non-essential
amino acids) in a humidified atmosphere containing 5% CO2 at 37 ºC. The
medium was replaced with
experiments.
1 mL PBS right before photolabeling
1.4. Cell-based Photolabeling
Cells were pretreated with 200 µM competitor for 15 minutes (or 3 h for 5
and 6) where applicable and then treated with 20 µM PRP or DMSO control.
After incubation with the PRP at 37 °C for 30 min (or 3 h for PRPs 13 and
14), the cells were cooled on ice and irradiated with 366 nm light for 30
min. The medium was removed, and the cells gently washed twice with 2
mL PBS and then covered with 1 mL PBS. Cells were scraped/transferred
from the plate into Eppendorf tubes, spun down at 1,000 g for 5 minutes
at 4 °C, the supernatant removed, and the cells resuspended in whole cell
lysis buffer (50 mM HEPES (pH 7.5), 150 mM NaCl, 1% Igepal CA-630,
5% glycerol and 1x protease inhibitor cocktail (Roche)) and 1x
1.7. Western Blot
Samples containing 30 µg protein were diluted with 4x LDS sample buffer
containing DTT (Invitrogen), boiled for 5 minutes and separated by gel
electrophoresis at 100 volts. Gels were transferred to nitrocellulose
membranes with iBlot2 transfer system (P3 for 7 minutes) and visualized
with Odyssey Sa imager at 800 nm. Membranes were then blocked with
Odyssey blocking buffer for 2 hours at rt or overnight at 4 °C. The blots
were incubated with desired antibodies (anti-HDAC1 (Abcam ab7028),
anti-HDAC2 (Abcam, ab12169), anti-HDAC3 (Abcam, ab7030), anti-
HDAC8 (Abcam, ab187139) and anti-HDAC4,5,9 (Abcam, ab131524)) at
the recommended dilutions for each in Odyssey blocking buffer (LI-COR)
overnight at 4 °C. The blots were then washed 3 × 5 minutes with PBST,
incubated with relevant species of IRDye® 680RD secondary antibody (LI-
COR, part number: 926-68071) for 1 hour and visualized with Odyssey Sa
imager at both 700 nm and 800 nm. If additional antibody probing was
necessary, membranes were stripped with 0.2 N NaOH for 10 minutes and
re-blocked with Odyssey blocking buffer.
phosphatase inhibitor cocktail (ThermoFisher)).
Samples were
homogenized by vortexing, incubated on a rotating stand at 4 °C for 1 h,
then spun down at 19,000g for 10 minutes at 4 °C and the protein
concentrations were determined by bicinchoninic acid (BCA) assay. Next,
35 µg total lysates were incubated with IRDye® 800CW Azide Infrared dye
(LI-COR, part number 926-68070) at concentration equal to that of the PRP,
TCEP (1 mM), TBTA (0.1 mM), and CuSO4 (1 mM) for 90 minutes at room
temperature. The samples were then frozen at -20 C until analyzed by
Western blot.
1.8. Ethics statement
1.5. Mouse Tissue Photolabeling
All animal experiments were Institutional Animal Care and Use Committee
(IACUC) approved and performed according to institutional guidelines with
University of Illinois at Chicago Institutional Biosafety and Animal Care
Committee approval (ACC Number: 17-012). All animal procedures were
performed in the College of Medicine Research Building at the UIC and
adhere to the policies of the NIH Office of Laboratory Animal Welfare
(OLAW), the standards of the Animal Welfare Act, the Public Health
Service Policy, and the Guide for the Care and Use of Laboratory Animals.
129/SV mouse was exsanguinated by retro-orbital bleeding, sacrificed and
fresh liver tissue (ca. 20 mg) was harvested and immediately homogenized
in an Eppendorf tube (using a Teflon homogenizer) containing 500 µL of
PBS. The tissue homogenate was centrifuged at 13,000g for 5 min at 4 °C.
The supernatant was discarded, and the cells were resuspended with 1
mL of PBS and then centrifuged again at 13,000g for 5 min at 4 °C. The
cells were resuspended in 1 mL PBS and plated in a 6-well plate at a 1:10
dilution. Cells were treated with either DMSO or10 µM PRP, or 200 µM
SAHA then 10 µM PRP in PBS in a humidified atmosphere containing 5%
CO2 at 37 ˚C (all treatments were done on cells from the same tissue
preparation) for 40 minutes. Cells were then irradiated with UV light (366
nm) for 30 min on ice, harvested by centrifugation at 13,000g for 5 min at
4 °C, washed with 1 mL PBS and repelleted. The cells were lysed by
addition of 100 µL cold lysis buffer (150 mM NaCl, 50 mM HEPES, 1%
2. General Chemistry Methods
All the reagents and solvents were obtained from commercial sources and
used without further purification. Reactions were performed under an inert
atmosphere (nitrogen or argon) whenever anhydrous solvents were used.
Reactions were monitored by thin-layer chromatography (TLC) using
Merck 60F254 silica gel plates or by Liquid Chromatography-Mass
10
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10.1002/cmdc.201900114
ChemMedChem
FULL PAPER
Spectrometry on Shimadzu LCMS-2020 with either Waters, XSelect HSS
CYANO 3.6 µm column or Agilent XBD-C18 3.5 μm column; dimensions
2.1 × 20 mm and UV detector at 254 nM. Chromatography purification was
performed on Biotage Isolera Four instrument using pre-filled KP-Sil
(normal phase) and KP-C18-HS (reverse phase) SNAP cartridges with UV
detection at 254 and 280 nm. 1H, 13C and 19F NMR spectra were recorded
on Bruker spectrometers at 400, 100 and 376 MHz, respectively. Chemical
shifts were reported on δ scale in ppm with the deuterated solvent
indicated as the internal reference. Coupling constants were reported in
Hz and the standard abbreviations indicating multiplicity were used as
follows: s = singlet, bs = broad singlet, d = doublet, t = triplet, q = quartet,
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m = multiplet. High resolution mass spectrometry experiments
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3. Docking, Synthetic Procedures, and Compound Characterization
Detailed description of the docking, synthetic procedures, and compounds
characterization is given in the supporting information.
Acknowledgements
This study was funded by the NCI/NIH grants R01 CA131970
(PAP) and R21 CA183627 (PAP), NHLBI/NIH R01 HL130760
(NM), NIAID/NIH R01 AI125401 (AMcL), the Alzheimer’s Drug
Discovery Foundation grant ADDF #20101103 (PAP), the PhRMA
Foundation Fellowship for Pharmacology and Toxicology (TYT),
and Egyptian Government scholarship JS-2971 (SA).
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Keywords: Histone deacetylase • Inhibitors • Photoaffinity
labeling • Target engagement •
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Entry for the Table of Contents
Engaged by inhibitors? The identity of the histone deacetylase
isoforms engaged by inhibitors in vivo is one of the unresolved
mysteries in epigenetic drug discovery. Using newly designed
photoreactive probes, we show that their relative histone
deacetylase isoform engagement in live cells in culture and
mouse liver tissue is chemotype- and cell-type dependent and is
substantially different from the isoform selectivity determined in
the biochemical assays.
13
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