Probing the: Hras -1Y i-motif with small molecules

Article abstract of DOI:10.1039/c8md00311d

Non-B DNA structures represent intriguing and challenging targets for small molecules. For example, the promoter of the HRAS oncogene contains multiple G-quadruplex and i-motif structures, atypical globular folds that serve as molecular switches for gene

Full text of DOI:10.1039/c8md00311d

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RESEARCH ARTICLE
Y
Probing the hras-1 i-motif with small molecules†
a
a
a
b
Cite this: Med. Chem. Commun.,
018, 9, 2000
Sara N. Journey, Stephanie L. Alden, Will M. Hewitt, Megan L. Peach,
a
a
2
Marc C. Nicklaus
and John S. Schneekloth Jr
*
Non-B DNA structures represent intriguing and challenging targets for small molecules. For example, the
promoter of the HRAS oncogene contains multiple G-quadruplex and i-motif structures, atypical globular
folds that serve as molecular switches for gene expression. Of the two, i-motif structures are far less stud-
Y
ied. Here, we report the first example of small organic compounds that directly interact with the hras-1
i-motif. We use a small molecule microarray screen to identify drug-like small molecules that bind to the
Y
hras-1 i-motif but not to several other DNA or RNA secondary structures. Two different lead compounds,
1
and 2, were discovered to have 7.4 ± 5.3 μM and 5.9 ± 3.7 μM binding affinity by surface plasmon reso-
Received 25th June 2018,
Accepted 19th September 2018
nance and similar affinity by fluorescence titration. A structure–activity relationship (SAR) was developed
and two improved analogues of 2 demonstrated submicromolar binding affinities. Both compounds display
pH-dependent binding, indicating that they interact with the DNA only when the i-motif is properly folded.
Chemical shift perturbation shows that 1 alters the structure of the i-motif, while 2 has no effect on the
i-motif conformation, indicating different modes of interaction.
DOI: 10.1039/c8md00311d
rsc.li/medchemcomm
by monovalent cations such as potassium, i-motifs are stabi-
lized by multiple [C·CH] base pairing events. The [C·CH]
Introduction
+
+
Genomic research has demonstrated that noncanonical DNA
structures occur throughout the mammalian genome. In ad-
base pair depends on one of the cytosine bases being proton-
ated; as a consequence, the formation of an i-motif is highly
pH dependent. While the specific pH at which unfolding oc-
curs is sequence dependent, folded i-motifs generally exist
below pH ∼7 while unfolded structures predominate at
higher pH. From a thermodynamic perspective, i-motifs are
less stable than the corresponding G4 structures and typically
1
dition to the classical B-DNA structure, DNA can adopt diverse
secondary and tertiary structures including hairpins, triplexes,
G-quadruplexes (G4s), i-motifs (also known as C-
quadruplexes), three-way junctions, four-way junctions (exem-
plified by Holliday Junctions), and repeat expansions, among
2
other structures. Alternatively folded DNA is often associated
with changes in gene regulation and is therefore associated
with many diseases, including cancer. Many DNA structures
have roles in regulating the expression of specific genes by
forming in promoter regions. One prominent example of this
is G4 structures, which are globular, folded nucleic acid struc-
3
tures that occur in G-rich regions of gene promoters. G4s
have been broadly demonstrated to regulate the expression of
many oncogenes and are well-studied as targets for small
molecules. A less studied, but likely equally important struc-
4
ture is the i-motif, or C-quadruplex.
Folded i-motif structures occur in C-rich regions of the ge-
nome, typically on the complementary strand to a G4-
4
a
forming sequence. In contrast to G4s, which are stabilized
a
Chemical Biology Laboratory, National Cancer Institute, Frederick, MD, USA.
E-mail: schneeklothjs@mail.nih.gov
b
Chemical Biology Laboratory, Basic Science Program, Frederick National
Laboratory for Cancer Research, Leidos Biomedical Research Inc., Frederick, MD,
USA
Fig. 1 (A) Structure and pH dependence of the HRAS i-motif. (B) Illus-
tration of the [C·C] basepair. (C) Sequence of the HRAS i-motif and
+
†
Electronic supplementary information (ESI) available. See DOI: 10.1039/
fluorescently labeled screening construct used in this manuscript.
A647 = AlexaFluor 647.
c8md00311d
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unfold at lower temperatures, even at low pH. A second factor
that governs i-motif folding is molecular crowding, and bio-
ture. Thus, multiple distinct chemotypes bind to the HRAS
i-motif with varying affinity and effects on the structure and
are likely suitable for further development into chemical
probes.
5
chemical conditions that mimic crowding cause increased
folding of i-motif sequences, even in cases of higher pH. A
third important contributing factor is torsional stress. Under
negative superhelicity (induced by transcription), both G- and
C-quadruplexes form to eliminate disfavored single-stranded
sequences (Fig. 1).
Results and discussion
To identify small molecules that bind to the HRAS i-motif, we
1
2
There is considerable evidence that i-motifs serve as regu-
used an SMM screening approach. This approach has been
used previously in our laboratory to identify small molecules
that bind to other folded nucleic acid structures such as RNA
6
latory structures within cells. A recent study revealed that
7
i-motifs are folded and can be visualized in living cells. Pro-
1
2c,13
teins have also been characterized that bind and unfold i-mo-
hairpins and DNA G4s.
We designed a 5′-AlexaFluor 647-
8
Y
tifs, as is the case with hnRNP LL and the BCl2 i-motif. Fur-
labeled hras-1 i-motif construct for use in the SMM screen
1
4
thermore, while G4 structures typically suppress gene
expression, i-motif-forming sequences are often associated
with increased expression of the associated gene. To further
support this observation, several small molecules that bind
to i-motifs have been shown to control gene expression as
(Fig. 2). Briefly, SMM slides containing 7042 compounds
were incubated for one hour at room temperature with 150
nM labeled i-motif sequence in a Tris buffer. Higher concen-
trations of DNA resulted in high background and did not pro-
vide usable data. After washing, slides were dried and im-
aged. A composite Z-score was generated for each compound
on the array, and compounds with a Z-score greater than
three were selected as hits. 61 compounds were identified for
further study after visual inspection of hit spots and compari-
son to 15 other structured oligonucleotides screened using
4
a,c
well.
One example of an oncogene containing an i-motif is
9
the HRAS gene. HRAS encodes for a small GTPase in the RAS
family of oncoproteins that stimulate proliferation by signal-
ing to the nucleus. Driver mutations that render the HRAS
enzyme constitutively active are found in many cancers, for
example in 29% of young adult patients with bladder can-
1
2a
the SMM platform. We selected 13 compounds to purchase
from this list for further study on the basis of availability and
representation from different structural classes. Of the 13
compounds that were purchased, many were found to be im-
1
0
cer. However, it has proven difficult to develop small mole-
cules that target the HRAS protein due to the high affinity of
the protein for its GTP cofactor (coupled with millimolar con-
centrations of GTP in the cell) as well as limited sites for
small molecule ligands to bind on the enzyme.
1
pure by LC/MS and H NMR analysis upon receipt, and we
obtained pure samples of eight by HPLC purification.
Next, each of the eight compounds was qualitatively ana-
To overcome the difficulties associated with targeting
HRAS at the protein level, controlling HRAS expression with
small molecules that bind to the i-motif is an attractive strat-
egy. For example, a small molecule might either stabilize or
destabilize the i-motif structure, or interfere with protein/
DNA interactions, any of which might have an impact on
Y
lyzed for binding to the hras-1 i-motif by surface plasmon
resonance (SPR). We designed a 5′-biotin-labeled i-motif se-
quence and immobilized the oligonucleotide to a streptavidin
surface. Each compound was injected at a concentration of
100 μM to assess binding to the folded DNA. Of the eight
compounds, six displayed binding to the DNA. We next used
1
1
gene expression. However, there are relatively few examples
of small molecules that bind to i-motifs in general. A major
challenge in establishing these important structures as po-
tential drug targets is to characterize druglike compounds
that specifically bind to the i-motif fold. More broadly, the
discovery of small molecules that specifically recognize
folded nucleic acid structures is of significant interest to both
the chemical biology and drug discovery communities. For
SPR to measure equilibrium dissociation constants (K ) for
D
each of the six compounds. Two compounds, 1 and 2, were
found to have K values in the single digit micromolar range
D
(7.4 ± 5.3 μM and 5.9 ± 3.7 μM, respectively). Other com-
pounds showed weaker binding or data that was difficult to
accurately fit to binding models (Table 1).
To independently measure KD values using an orthogonal
biophysical technique, we developed a fluorescence titration
Y
the specific case of the hras-1 i-motif, to date there have
1
3a
been no small molecule ligands reported.
assay.
In this assay, we used the 5′-AlexaFluor 647 con-
In this study, we report the discovery of new classes of
small molecules that bind to the hras-1 i-motif. We use a
struct designed for screening. KD values were derived by mea-
suring ligand-induced changes in the fluorescence of the
tagged oligonucleotide construct. In this assay, both com-
pounds were validated to bind to the i-motif. Compound 1
was found to have a KD of 1.6 ± 0.3 μM, in excellent agree-
Y
small molecule microarray (SMM) screening strategy to rap-
idly identify small organic compounds that interact with the
Y
hras-1 i-motif but do not bind to other folded oligonucleo-
tides. Multiple orthogonal affinity measurement techniques
reveal that compounds from these series have affinities rang-
ing from the high nanomolar to low micromolar range and
are readily chemically synthesized. Furthermore, circular di-
chroism, NMR, and fluorescence resonant energy transfer
D
ment with SPR experiments. In contrast, 2 had a K of 36.6 ±
13.7 μM, higher than the value recorded by SPR. As com-
pound 1 was the tightest binding compound, we chemically
synthesized this compound and demonstrated that it had
equivalent binding by both SPR and fluorescence titration
and was used for all future experiments (Fig. 3).
(FRET) assays reveal ligand-induced effects on i-motif struc-
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Fig. 2 (A) Structures and raw SMM screening data for two hit compounds that bind to the HRAS i-motif. (B) Selectivity for hit compounds across
multiple SMM screens with different oligonucleotides. Data are reported as composite Z scores taking into account two replicate SMM spots.
To evaluate whether 1 was well dispersed (not aggregating)
and bound to unlabeled hras-1 i-motif DNA in solution, we
ence of DNA, compounds that bind are protected from mag-
netization transfer and ligand peaks exhibit positive phasing
(indicated as up in Fig. 4). Importantly, compounds that ag-
gregate in aqueous solution (a common source of false posi-
tives in biochemical assays) can also be easily identified in
this experiment by positive phasing in the waterLOGSY spec-
trum in the absence of DNA. Gratifyingly, 1 did not aggregate
Y
next performed a ligand-observed gradient spectroscopy
1
5
(
waterLOGSY) NMR experiment. In this experiment, reso-
nant energy transfer between bulk water and the ligand is
measured. In the absence of DNA, ligand peaks are phased
negatively (indicated as down in Fig. 4). However, in the pres-
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Table 1 Chemical structures and affinity measurements of compounds
to the HRAS i-motif
folded. In contrast, when the pH was raised to 8.02, where
the DNA exists predominantly as a hairpin, no binding was
observed. Thus, 1 binds to DNA in a fold-dependent manner.
Similar effects were observed with 2, which only bound to the
HRAS DNA in the i-motif form.
a
b
Entry
1
Compound structure
SPR K
D
(μM)
FIA K
D
(μM)
7.4 ± 5.3
1.6 ± 0.3
Next, we evaluated the effects of compound binding on
the chemical shifts of the i-motif protons in NMR (Fig. 5). Al-
Y
though there is no structure of the hras-1 i-motif available,
2
3
5.9 ± 3.7
0.2 ± 0.2
35.6 ± 13.7
0.6 ± 0.5
characteristic peaks for i-motif imino protons are clearly visi-
1
6,17
ble at δppm ∼15–16.
In the presence of increasing concen-
trations of 1, the characteristic i-motif protons broaden con-
siderably (Fig. 5A and B). However, no significant changes
are observed elsewhere in the spectrum. Furthermore, no cor-
responding increase of Watson–Crick imino protons was ob-
served at δppm ∼12.5–14.
To further probe the effects of ligands on the conforma-
tion of the i-motif, we used circular dichroism (CD) spectro-
scopy. An unlabeled hras-1 i-motif sequence displayed a
4
5
0.3 ± 0.3
3.0 ± 2.3
11.6 ± 4.8
16.0 ± 18.1
Y
peak minimum at 248 nm and a maximum at 289 nm, char-
4
a,9
acteristic of a properly folded i-motif.
In the presence of 1,
6
7
51.0 ± 3.0
254.5 ± 124.1
30.8 ± 3.4
this peak height decreased and shifted to a shorter wave-
length, indicating that binding of 1 has a small but measur-
able effect on the conformation of the i-motif (Fig. 5). In con-
trast, 2 had minimal effects on the structure of the i-motif.
We then evaluated the effects of ligand binding on the ther-
mal denaturation of the i-motif. Neither 1 nor 2 had any sta-
tistically significant effect on the melting temperature of the
DNA.
78.0 ± 99.0
8
9
149.2 ± 15.3
90.7 ± 12.6
9
Next, we performed a FRET assay. In this assay, as previ-
ously described, an i-motif construct was purchased
containing FRET donor/acceptor pairs on the 3′ and 5′ ends.
Compounds that cause a large change in the i-motif confor-
mation (for example unfolding) cause changes in FRET effi-
ciency and can readily be monitored. Compound 1 had mini-
mal effects in this assay, in contrast with a previously
reported protein that unfolds the structure and causes a large
242.6 ± 97.5
>209
1
0
>300
106.3 ± 35.9
1
1
1
>300
50.4 ± 15.1
9
change in FRET signal (see Fig. S56†). Combined with obser-
1
vations in the H NMR, these data together suggest that the
2
3
>300
>300
>209
>209
compound directly interacts with the i-motif core, but poten-
tially not with other regions of the structure such as the
loops. In contrast to 1, evaluation of the effects of 2 on the
i-motif NMR showed no changes in the i-motif imino protons
with increasing concentrations of compound 2. However,
some changes could be observed in other regions of the spec-
trum, suggesting that the compound interacts with other re-
gions of the DNA, potentially the loop regions. Taken to-
gether, these experiments suggest that small molecules may
interact with i-motif structures by multiple distinct modes of
binding (e.g. groove or loop binding).
To investigate the structure–activity relationship (SAR) of
these compounds, we evaluated a collection of commercially
available analogs (Table 1). After purchasing, each compound
was assessed for purity by LC/MS analysis and purified by
D
HPLC if necessary. The binding affinity (K ) of each com-
pound was measured by both fluorescence intensity titration
and SPR to give orthogonal measures of affinity (Table 1).
1
a
D
K values represent the mean of two experiments performed with
b
at least four concentrations.
replicates.
D
K values represent the mean of three
in solution as observed by negatively phased peaks in
waterLOGSY spectrum in the absence of DNA. Furthermore,
upon addition of an unlabeled hras-1 i-motif sequence, the
protons from 1 were phased positively, confirming that the
compound directly interacts with the DNA in the solution
phase (Fig. 4). Next, we used waterLOGSY NMR to evaluate
the pH-dependence of this binding interaction. Direct bind-
ing to the DNA was observed at pH 6.02, where the i-motif is
Y
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Fig. 3 Synthesis of compound 1.
With respect to 1, replacement of the hydroxyethyl amino
sidechain with a methoxy substituent abolished binding, as
did replacement of this group with other, bulky substituents.
Similarly, replacement of the amino group on the pyrimidine
ring with a dimethylamino group also caused a substantial
decrease in binding. The compound tolerated other heterocy-
clic rings on the right-hand side chain, though binding was
generally weaker. Finally, an amide analog exhibited no
binding.
For 2, a strong relationship was observed between com-
pound affinity and chemical structure. Compound 2 also con-
tains an amino-substituted pyrimidine group. Replacement
of this group with a methyl or methoxy substituent led to a
marked increase in binding affinity, with KD values in the
nanomolar range. However, a dimethylamino-substituted an-
alog had no binding activity, pointing to a role for this group
in binding. Furthermore, two analogs lacking the amine em-
bedded in the sidechain also had no observable binding,
again indicating a critical role for this functionality (Fig. 2).
Because 3 and 4 had improved binding relative to 2, we vali-
dated its binding in other assays. In comparison to 2, 3 and
4
behaved similarly in circular dichroism and chemical shift
perturbation experiments, indicating that it is likely binding
by a similar mechanism (see ESI†).
In an attempt to further investigate potential binding
modes for these compounds, we used computational
Y
docking. Because no structure of the hras-1 i-motif is avail-
able, we considered generating a homology model based on
another reported intramolecular i-motif (PDB: 1ELN). How-
ever, modeling the loop regions proved extremely challenging
due to the complex energy landscape governing loop pairing
and stacking interactions. Given that experiments suggested
that 1 binds directly to the i-motif core we generated a mini-
mal i-motif model instead (lacking the loops) from the crystal
1
8
structure of the d(ACCCT) i-motif (PDB: 1CN0). We docked
and 2 to the major groove of this structure (see modeling
1
methods in the ESI† for details). Compound 1 docked in two
nearly equivalent poses (Fig. 6A and B), with the pyrimidine
and benzyl groups flat across the bottom of the groove,
forming amino-aromatic interactions with the projecting cy-
tosine N4 amines. The amino substituent on the pyridine,
the hydroxyl group, and the pyrrolidine (protonated at pH 6)
can all hydrogen bond to the phosphate backbone. The
amino-pyrimidine group in 2 can dock similarly to that in 1
(
Fig. S59†), but the rest of the structure does not interact
strongly with the i-motif core, suggesting that the methoxy-
containing ring may bind to the loops.
In conclusion, we have identified a series of small chemi-
Y
Fig.
4
WaterLOGSY experiments demonstrate
1
exhibits pH-
cal ligands that directly bind to the hras-1 i-motif. This
1
dependent binding to i-motif DNA. (A) H NMR and WaterLOGSY NMR
in the (B) absence and (C) presence of i-motif DNA at pH 6. (D) H NMR
and WaterLOGSY NMR in the (E) absence and (F) presence of i-motif
DNA at pH 8. Methyl peaks for N-methyl-L-valine (used as an internal
negative control) can be observed at ∼0.8 ppm. Other peaks in the
spectrum correspond to protons from 1.
structure, an important regulatory element of the HRAS onco-
gene promoter, is an unexplored target for small molecules.
Here we demonstrate through SMM screening that diverse
1
Y
small molecule ligands can bind to the hras-1 i-motif by dif-
ferent mechanisms. Furthermore, these compounds have
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1
Fig. 5 H NMR chemical shift perturbations of i-motif DNA in the presence of increasing concentrations of (A) compound 1 and (B) compound 2
in deuterated Tris-acetate buffer at pH 6. Circular dichroism spectra of i-motif DNA in the presence of increasing concentrations of (C) compound
1
and (D) compound 2 in Tris-acetate buffer at pH 6 (except the purple spectrum, which was collected at pH 8).
strong SAR and can achieve submicromolar affinity to the
i-motif. In-depth biophysical characterization demonstrates
that small changes in the chemical structure can lead to
large changes in affinity, indicating highly specific interac-
tions. Finally, modeling studies provide a compelling plausi-
ble mode of interaction for 1. The observation that the
aminopyrimidine group in both 1 and 2 adopts a similar
interaction with the cytosine and phosphates suggests that
this fragment may be a general scaffold that is suitable for
other i-motif structures. A related observation is that pro-
teins bearing unnatural amino acids containing a similar
1
9
functionality exhibit improved binding to i-motifs. The
chemical structure of several of these ligands is somewhat
reminiscent of the pyrimidine core of a cytosine residue;
this may play a role in the affinity of these compounds for
the DNA, though further experimentation will be required
to validate this observation. Future work will focus on
establishing the structural basis by which such compounds
can achieve high affinity with the i-motif structure, and to
Y
further development for ligands that bind to the hras-1 or
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computational resources of the NIH HPC Biowulf cluster
(
http://hpc.nih.gov).
References
1
2
(a) F. Kouzine, D. Wojtowicz, L. Baranello, A. Yamane, S.
Nelson, W. Resch, K. R. Kieffer-Kwon, C. J. Benham, R.
Casellas, T. M. Przytycka and D. Levens, Cell Syst., 2017, 4,
3
2
44–356; (b) A. Bacolla and R. D. Wells, J. Biol. Chem.,
004, 279, 47411–47414.
(a) M. Kaushik, S. Kaushik, K. Roy, A. Singh, S. Mahendru,
M. Kumar, S. Chaudhary, S. Ahmed and S. Kukreti, Biochem.
Biophys. Rep., 2016, 5, 388–395; (b) T. A. Brooks, S. Kendrick
and L. Hurley, Rev. Geophys., 2010, 277, 3459–3469.
3
4
(a) S. A. Ohnmacht and S. Neidle, Bioorg. Med. Chem. Lett.,
2
014, 24, 2602–2612; (b) J. L. Huppert and S.
Balasubramanian, Nucleic Acids Res., 2007, 35, 406–413.
(a) H. A. Day, P. Pavlou and Z. A. Waller, Bioorg. Med. Chem.,
2
014, 22, 4407–4418; (b) C. E. Kaiser, N. A. Van Ert, P.
Agrawal, R. Chawla, D. Yang and L. H. Hurley, J. Am. Chem.
Soc., 2017, 139, 8522–8536; (c) M. Gueron and J. L. Leroy,
Curr. Opin. Struct. Biol., 2000, 10, 326–331.
5
6
7
T. Nguyen, C. Fraire and R. D. Sheardy, J. Phys. Chem. B,
2
017, 121, 7872–7877.
S. Jonchhe, P. Shrestha, K. Ascencio and H. Mao, Anal.
Chem., 2018, 90, 3205–3210.
M. Zeraati, D. B. Langley, P. Schofield, A. L. Moye, R. Rouet,
W. E. Hughes, T. M. Bryan, M. E. Dinger and D. Christ, Nat.
Chem., 2018, 10, 631–637.
8
(a) B. Roy, P. Talukder, H. J. Kang, S. S. Tsuen, M. P. Alam,
L. H. Hurley and S. M. Hecht, J. Am. Chem. Soc., 2016, 138,
10950–10962; (b) H. J. Kang, S. Kendrick, S. M. Hecht and
L. H. Hurley, J. Am. Chem. Soc., 2014, 136, 4172–4185; (c) M.
Debnath, S. Ghosh, A. Chauhan, R. Paul, K. Bhattacharyya
and J. Dash, Chem. Sci., 2017, 8, 7448–7456; (d) S. Kendrick,
Y. Akiyama, S. M. Hecht and L. H. Hurley, J. Am. Chem. Soc.,
Fig. 6 Representative docked pose of compound 1 with the i-motif
core (PDB: 1CN0). A) Top view looking down into the major groove.
The surface of the i-motif core is colored according to electrostatic
potential. B) Side view with transparent surface. Hydrogen bonds to
the phosphate groups are indicated with dashed orange lines and the
amino-aromatic interaction between the pyrimidine group and the cy-
tosine N4 amine is indicated with a dashed green line.
2
009, 131, 17667–17676.
9
G. Miglietta, S. Cogoi, E. B. Pedersen and L. E. Xodo, Sci.
Rep., 2015, 5, 18097.
evaluate their role in modulating gene expression in cancer
cells.
10 W. Beukers, A. Hercegovac and E. C. Zwarthoff, Eur. J. Hum.
Genet., 2014, 22, 837–839.
1
1 S. Kendrick, A. Muranyi, V. Gokhale, L. H. Hurley and L. M.
Rimsza, J. Med. Chem., 2017, 60, 6587–6597.
Conflicts of interest
1
2 (a) C. M. Connelly, F. A. Abulwerdi and J. S. Schneekloth Jr.,
Methods Mol. Biol., 2017, 1518, 157–175; (b) P. J.
Hergenrother, K. M. Depew and S. L. Schreiber, J. Am. Chem.
Soc., 2000, 122, 7849–7850; (c) J. E. Bradner, O. M.
McPherson and A. N. Koehler, Nat. Protoc., 2006, 1,
2344–2352; (d) M. D. Disney, S. Magnet, J. S. Blanchard and
P. H. Seeberger, Angew. Chem., Int. Ed., 2004, 43, 1591–1594;
(e) J. W. Puckett, K. A. Muzikar, J. Tietjen, C. L. Warren, A. Z.
Ansari and P. B. Dervan, J. Am. Chem. Soc., 2007, 129,
12310–12319; ( f ) S. P. Velagapudi, A. Pushechnikov, L. P.
Labuda, J. M. French and M. D. Disney, ACS Chem. Biol.,
2012, 7, 1902–1909; (g) M. D. Disney, L. P. Labuda, D. J.
Paul, S. G. Poplawski, A. Pushechnikov, T. Tran, S. P.
There are no conflicts to declare.
Acknowledgements
This work was supported by the Intramural Research Program
of the National Institutes of Health, Center for Cancer Re-
search, and the National Cancer Institute (NCI), National In-
stitutes of Health (1 ZIA BC011585 04). S. N. J. was partially
funded by the Introduction to Cancer Research Careers pro-
gram. This work was funded in part with Federal funds from
the National Cancer Institute, National Institutes of Health,
under Contract HHSN261200800001E. This work utilized the
2006 | Med. Chem. Commun., 2018, 9, 2000–2007
This journal is © The Royal Society of Chemistry 2018

View Article Online
MedChemComm
Research Article
Velagapudi, M. Wu and J. L. Childs-Disney, J. Am. Chem.
Soc., 2008, 130, 11185–11194; (h) J. L. Childs-Disney, M. L.
Wu, A. Pushechnikov, O. Aminova and M. D. Disney, ACS
Chem. Biol., 2007, 2, 745–754.
14 A. M. Fleming, Y. Ding, R. A. Rogers, J. Zhu, J. Zhu, A. D.
Burton, C. B. Carlisle and C. J. Burrows, J. Am. Chem. Soc.,
2017, 139, 4682–4689.
15 (a) C. Dalvit, P. Pevarello, M. Tato, M. Veronesi, A. Vulpetti
and M. Sundstrom, J. Biomol. NMR, 2000, 18, 65–68; (b) M.
Pellecchia, B. Becattini, K. J. Crowell, R. Fattorusso, M.
Forino, M. Fragai, D. Jung, T. Mustelin and L. Tautz, Expert
Opin. Ther. Targets, 2004, 8, 597–611.
16 I. Bertini, K. S. McGreevy and G. Parigi, NMR of biomolecules
: towards mechanistic systems biology, Wiley-VCH, Weinheim,
Germany, 2012.
17 V. V. Jolad, F. K. Murad, J. R. Arnold and J. Fisher, Org.
Biomol. Chem., 2005, 3, 2234–2236.
18 J. Weil, T. Min, C. Yang, S. Wang, C. Sutherland, N. Sinha
and C. Kang, Acta Crystallogr., Sect. D: Biol. Crystallogr.,
1999, 55, 422–429.
19 X. Bai, P. Talukder, S. M. Daskalova, B. Roy, S. Chen, Z. Li,
L. M. Dedkova and S. M. Hecht, J. Am. Chem. Soc., 2017, 139,
4611–4614.
1
3 (a) C. M. Connelly, R. E. Boer, M. H. Moon, P. Gareiss and
J. S. Schneekloth Jr., ACS Chem. Biol., 2017, 12, 435–443; (b)
K. M. Felsenstein, L. B. Saunders, J. K. Simmons, E. Leon,
D. R. Calabrese, S. Zhang, A. Michalowski, P. Gareiss, B. A.
Mock and J. S. Schneekloth Jr., ACS Chem. Biol., 2016, 11,
1
39–148; (c) J. Sztuba-Solinska, S. R. Shenoy, P. Gareiss, L. R.
Krumpe, S. F. Le Grice, B. R. O'Keefe and J. S. Schneekloth
Jr., J. Am. Chem. Soc., 2014, 136, 8402–8410; (d) D. R.
Calabrese, K. Zlotkowski, S. Alden, W. M. Hewitt, C. M.
Connelly, R. M. Wilson, S. Gaikwad, L. Chen, R. Guha, C. J.
Thomas, B. A. Mock and J. S. Schneekloth Jr., Nucleic Acids
Res., 2018, 46, 2722–2732; (e) J. A. Hong, D. V. Neel, D.
Wassaf, F. Caballero and A. N. Koehler, Curr. Opin. Chem.
Biol., 2014, 18, 21–28; ( f ) J. L. Duffner, P. A. Clemons and
A. N. Koehler, Curr. Opin. Chem. Biol., 2007, 11, 74–82.
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