Structure-guided design of anti-cancer ribonucleotide reductase inhibitors

Article abstract of DOI:10.1080/14756366.2018.1545226

Ribonucleotide reductase (RR) catalyses the rate-limiting step of dNTP synthesis, establishing it as an important cancer target. While RR is traditionally inhibited by nucleoside-based antimetabolites, we recently discovered a naphthyl salicyl acyl hydrazone-based inhibitor (NSAH) that binds reversibly to the catalytic site (C-site). Here we report the synthesis and in vitro evaluation of 13 distinct compounds (TP1-13) with improved binding to hRR over NSAH (TP8), with lower K_D’s and more predicted residue interactions. Moreover, TP6 displayed the greatest growth inhibiting effect in the Panc1 pancreatic cancer cell line with an IC₅₀ of 0.393 μM. This represents more than a 2-fold improvement over NSAH, making TP6 the most potent compound against pancreatic cancer emerging from the hydrazone inhibitors. NSAH was optimised by the addition of cyclic and polar groups replacing the naphthyl moiety, which occupies the phosphate-binding pocket in the C-site, establishing a new direction in inhibitor design.

Full text of DOI:10.1080/14756366.2018.1545226

Journal of Enzyme Inhibition and Medicinal Chemistry
ISSN: 1475-6366 (Print) 1475-6374 (Online) Journal homepage: https://www.tandfonline.com/loi/ienz20
Structure-guided design of anti-cancer
ribonucleotide reductase inhibitors
Tessianna A. Misko, Yi-Ting Liu, Michael E. Harris, Nancy L. Oleinick, John
Pink, Hsueh-Yun Lee & Chris G. Dealwis
To cite this article: Tessianna A. Misko, Yi-Ting Liu, Michael E. Harris, Nancy L. Oleinick,
John Pink, Hsueh-Yun Lee & Chris G. Dealwis (2019) Structure-guided design of anti-cancer
ribonucleotide reductase inhibitors, Journal of Enzyme Inhibition and Medicinal Chemistry, 34:1,
438-450, DOI: 10.1080/14756366.2018.1545226
To link to this article: https://doi.org/10.1080/14756366.2018.1545226
© 2019 The Author(s). Published by Informa
UK Limited, trading as Taylor & Francis
Group.
View supplementary material
Published online: 07 Jan 2019.
Submit your article to this journal
Article views: 74
View Crossmark data
Full Terms & Conditions of access and use can be found at
https://www.tandfonline.com/action/journalInformation?journalCode=ienz20

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
2019, VOL. 34, NO. 1, 438–450
https://doi.org/10.1080/14756366.2018.1545226
RESEARCH PAPER
Structure-guided design of anti-cancer ribonucleotide reductase inhibitors
Tessianna A. Miskoa,# , Yi-Ting Liub,#, Michael E. Harrisc, Nancy L. Oleinickd, John Pinke, Hsueh-Yun Leeb,f,* and
Chris G. Dealwisa,g,*
b
aDepartment of Pharmacology, School of Medicine, Case Western Reserve University, Cleveland, OH, USA; School of Pharmacy, College of
c
d
Pharmacy, Taipei Medical University, Taipei, Taiwan; Department of Chemistry, University of Florida, Gainesville, FL, United States; Department
e
of Radiation Oncology, School of Medicine, Case Western Reserve University, Cleveland, OH, USA; Case Comprehensive Cancer Center, School
f
of Medicine, Case Western Reserve University, Cleveland, OH, USA; Ph.D Program in Biotechnology Research and Development, College of
g
Pharmacy, Taipei Medical University, Taipei, Taiwan; Department of Chemistry, Center for Proteomics, Case Western Reserve University,
Cleveland, OH, USA
ABSTRACT
ARTICLE HISTORY
Received 25 July 2018
Revised 26 September 2018
Accepted 2 November 2018
Ribonucleotide reductase (RR) catalyses the rate-limiting step of dNTP synthesis, establishing it as an
important cancer target. While RR is traditionally inhibited by nucleoside-based antimetabolites, we
recently discovered a naphthyl salicyl acyl hydrazone-based inhibitor (NSAH) that binds reversibly to
the catalytic site (C-site). Here we report the synthesis and in vitro evaluation of 13 distinct compounds
(TP1-13) with improved binding to hRR over NSAH (TP8), with lower KD’s and more predicted residue
interactions. Moreover, TP6 displayed the greatest growth inhibiting effect in the Panc1 pancreatic cancer
cell line with an IC50 of 0.393mM. This represents more than a 2-fold improvement over NSAH, making
TP6 the most potent compound against pancreatic cancer emerging from the hydrazone inhibitors. NSAH
was optimised by the addition of cyclic and polar groups replacing the naphthyl moiety, which occupies
the phosphate-binding pocket in the C-site, establishing a new direction in inhibitor design.
KEYWORDS
Ribonucleotide reductase;
cancer; inhibitor; pancreatic;
phosphate-binding
Introduction
approach for developing inhibitors in the past few decades has
involved modifying natural nucleoside substrates, resulting in the
production of the clinically used drugs such as gemcitabine,
fludarabine, clofarabine, and cladrabine17–23. Gemcitabine is a
billion-dollar drug used as a front line treatment of pancreatic
cancer24. Gemcitabine derives its main cytotoxicity through DNA
chain termination25,26, irreversible inhibition of hRR27,28 and its
interactions with several phosphate-binding proteins, such as
deoxycytidine deaminase (dCMP deaminase)29, thymidylate
synthase30, CTP-synthase31,32, and topoisomerase-133,34. The chain
termination activity and lack of specificity exhibited by gemcita-
bine and other nucleoside-based drugs contribute to the toxic
side effects that patients endure17–22. Thus, we have proposed
that the development of a selective RR-targeted inhibitor may
reduce toxic side effects and expand the therapeutic window35.
Our lab has set out to discover a new class of inhibitors
that has higher selectivity for hRRM1, moving away from the
nucleoside-based analogues and towards reversible, competitive,
non-nucleoside inhibitors. An in silico drug screen of 350,000 com-
pounds from the Cincinnati chemical library (formerly the Proctor
and Gamble chemical library) was performed using a human RR
structure complexed with the effector TTP and the substrate GDP
(PDB code 3HND)35,36. From this screen came the discovery of our
lead inhibitor, NSAH35. Crystallographic and kinetic studies indi-
Human ribonucleotide reductase (hRR) is a ubiquitous multi-
subunit enzyme that is crucial for cell division and DNA repair1–4
.
RR catalyses the rate-determining step of dNTP synthesis by
removing the 20 hydroxyl of the ribose to generate a deoxyri-
bose1. RR is critical for the maintenance of a balanced nucleotide
pool in cells where imbalances lead to mutator phenotypes1,5,6
.
RR activity is maintained under tight regulation at the levels of
transcription7, allostery1, cellular localisation8, and enzyme inhib-
ition by Sml1 in Saccharomyces cerevisiae9. hRR is a multi-subunit
enzyme consisting of a large subunit hRRM1, containing a cata-
lytic site (C-site) and two allosteric sites known as the specificity
site (S-site) and the activity site (A-site)1 (Figure 1). hRRM1 associ-
ates with the small subunit hRRM2 that houses a free radical
essential for catalysis to form the holoenzyme10. ATP and dATP
bind the A-site, inducing active and inactive hexamers, respect-
ively11. Binding of dGTP, dTTP, dATP, or ATP to the S-site not only
induces dimerisation of hRRM1 but also acts as a selection gate,
regulating the relative Kcat/Km for the four NDP substrates: ADP,
GDP, CDP, or UDP, respectively1.
Due to the crucial role hRR plays in replication, it is a major
target for cancer chemotherapy12–14. One of the first described
hRR inhibitors was hydroxyurea, which targets the di-iron cluster
of the hRRM2 subunit to block catalysis15,16. The most common cated that NSAH binds reversibly to the C-site of hRR and acts as
CONTACT Chris G. Dealwis
Cleveland, OH 44106, USA, Hsueh-Yun Lee
cxd114@case.edu
Department of Pharmacology, School of Medicine, Case Western Reserve University, 10900 Euclid Avenue,
hyl@tmu.edu.tw School of Pharmacy, College of Pharmacy, Taipei Medical University, 250 Wuxing St. Taipei
11031, Taiwan.
#These authors contributed equally to this work.
*These authors contributed equally to this work.
Supplemental data for this article can be accessed here.
ß 2019 The Author(s). Published by Informa UK Limited, trading as Taylor & Francis Group.
This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (http://creativecommons.org/licenses/by-nc/4.0/), which permits
unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
439
Figure 1. Structure of hRRM1 dimer.
a competitive inhibitor. The x-ray crystal structure of NSAH bound or diacylhydrazine to explore the structure-activity relationship. As a
to hRRM1 was determined to 2.7 Å resolution (PDB code 5TUS)35
result, compounds TP1-13 (Figure 2(B)) were synthesised and rele-
.
Through chemical mimicry, NSAH occupies the space where the vant assays to characterise their interaction with hRRM1 were con-
diphosphate ribose and base of the substrate bind in the C-site. ducted in this study. Using docking studies to explore possible
NSAH adopts an S-shaped conformation, thermodynamically interactions with hRRM1, it was determined that this library of com-
pounds displayed an increase in the interactions with the phos-
phate-binding region of the C-site, and the best binding
compounds make use of strong interactions to either the phos-
phate-binding region or residues near loop 2. Cancer cell studies
indicated that group 1 compounds showed the greatest potency in
favouring the E-isomer. The crystal structure reveals that stabilisa-
tion of this conformation is due to a strong hydrogen bond
between the carbonyl of NSAH and residues Ser217 and Cys218
and a hydrogen bond between the hydroxyl of the phenol of
NSAH and residues Cys218 and Ser217.
This study also revealed that the salicyl acyl hydrazine moiety cells, where polar substituents incorporating electronegative ele-
has a significant contributory role in the inhibitory activity against ments distinguished the more cytotoxic compounds. In fact, the
most potent inhibitors from this class demonstrated up to a two-
fold improvement in potency against the growth inhibition of pan-
creatic cancer cells (Panc1) relative to NSAH. The results of this
study will lead to the design of future generations of compounds
that further improve on target hRR inhibition and cytotoxic efficacy.
RR. NSAH inhibits cancer cell growth with IC50s within two-fold of
gemcitabine and leads to the depletion of deoxypurine pools, a
hallmark of cellular hRR inhibition. Unlike gemcitabine, NSAH
demonstrated little measurable toxicity against normal mobilised
peripheral blood progenitor cells, giving NSAH a higher thera-
peutic index than found for gemcitabine35. Nevertheless, medi-
cinal chemistry and synthetic approaches can be used to improve
the potency of NSAH and its target selectivity towards hRR.
Indeed, a recent paper from our laboratory reported on a library
of 25 NSAH analogues whose modifications were designed to
target residues within the C-site of hRR that are important for
interactions with natural substrates37. These results indicated that
Materials and methods
Synthesis and characterisation of TP1-13
Nuclear magnetic resonance (1H NMR and 13C NMR) spectra were
recorded with Bruker Fourier 500 NMR spectrometers, with chem-
ical shifts in parts per million (d) downfield from tetramethylsilane
(TMS), the internal standard (Supplemental pages S10–35). High-
resolution mass spectra (HRMS) were recorded with a JEOL (JMS-
700) mass spectrometer (Supplemental pages S36–48). The puri-
ties of the final compounds were determined using an Agilent
1100 series HPLC system with a C-18 column (Agilent ZORBAX
Eclipse XDB-C18 5 lm, 4.6 mm ꢀ 150 mm) and were found to be
ꢁ95%. Flash column chromatography was conducted using silica
gel (Merck Kieselgel 60, No. 9385, 230 ꢂ 400 mesh ASTM). All reac-
tions were conducted under an atmosphere of dry N2.
those analogues which demonstrated
a 2–4 fold improved
potency of cell-free hRR over NSAH all showed hydrogen bonding
with Ser606, Thr607, and Ser217, residues that are known to
hydrogen bond with natural substrates.
The present study investigated the structure activity relationship
of a new library of compounds to explore the chemical diversity
that can target a compound to the phosphate- and ribose-binding
domains within the C-site. One structure-guided approach to this
goal was to modify the naphthalene ring of NSAH by dissociating
the fused ring, providing a biphenyl moiety with different types of
substitutions (Group 1). Furthermore, the phenyl ring was replaced
by thiophene and furan using a bioisosterism strategy (Figure 2(A)).
(E)-2-Hydroxy-N’-(2-oxoindolin-3-ylidene)benzohydrazide (TP1)
To understand the effect of the linker on RR modulation, the hydra- A mixture of methyl salicylate (1.0 eq), N2H4 (1.1 eq), and EtOH was
zide group was replaced by diacylhydrazine or thiazole (shown in heated to reflux until the reaction complete. The reaction mixture
purple, Figure 2(A), Group 2). Meanwhile, some polar rings such as was filtrated and washed with EtOH to afford a white solid. The
pyridine, adenine, isatin and 2-pyridone were linked by hydrazide resultant was re-suspended in EtOH, and then isatin (1.1 eq) was

440
T.A. MISKO ET AL.
Figure 2. Schematic and structures of synthesised compounds.
added. The resulting mixture was heated to reflux until the reac- preparation of TP1 (reaction time: 12 h): mp ¼ 273.5 ꢂ 274.9 ꢃC;
tion was complete (12 h). The reaction was quenched by adding 1H NMR (500 MHz, DMSO-d6)
d 6.96–7.01 (m, 2H), 7.05 (d,
H2O and the resulting suspension was filtered and washed by
J ¼ 8.5 Hz, 1H), 7.32 (t, J ¼ 7.5 Hz, 1H), 7.43–7.48 (m, 3H), 7.62–7.65
(m, 3H), 7.88 (d, J ¼ 2.0 Hz, 1H), 7.91 (dd, J ¼ 1.0, 8.0 Hz, 1H), 8.76
(s, 1H), 11.34 (s, 1H), 11.80–11.86 (m, 1H), 12.11 (s, 1H). 13C NMR
(125 MHz, DMSO-d6) d 116.15, 117.54, 117.78, 119.49, 126.62,
127.34, 127.77, 129.09, 129.37, 130.37, 132.00, 134.45, 139.90,
149.23, 157.61, 159.50, 165.07. HRMS (ESI) for C20H17N2O3 [M þ H]þ
calculated 333.1239, found 333.1241.
1
EtOH to obtain TP1 (42%). mp ¼ 320.4 ꢃC (decomposed); H NMR
(500 MHz, DMSO-d6) d 6.92 (d, J ¼ 8.0 Hz, 1H), 6.97 (t, J ¼ 8.0 Hz,
1H), 7.01 (d, J ¼ 8.5 Hz, 1H), 7.08 (t, J ¼ 8.0 Hz, 1H), 7.35 (dt, J ¼ 1.0,
8.0 Hz, 1H), 7.44 (dt, J ¼ 1.5, 8.5 Hz, 1H), 7.57 (d, J ¼ 7.5 Hz, 1H), 7.97
(dd, 1.0, 7.5 Hz, 1H), 11.13 (s, 1H), 11.64 (s, 1H), 14.35 (s, 1H). 13C
NMR (125 MHz, DMSO-d6) d 111.34, 111.59, 115.75, 117.28, 117.51,
117.87, 120.14, 120.71, 120.95, 121.23, 122.35, 122.86, 124.17,
131.82, 131.85, 131.94, 133.26, 134.70, 134.93, 137.49, 139.03,
142.86, 144.35, 156.51, 157.14, 162.03, 163.36, 165.28. HRMS (ESI)
for C15H12N3O3 [M þ H]þ calculated 282.0879, found 282.0880.
(E)-2-Hydroxy-N’-(2-hydroxy-5-(thiophen-3-yl)-benzylidene)-
benzohydrazide (TP3)
The title compound was obtained in 70% overall yield from com-
(E)-2-Hydroxy-N’-((4-hydroxy-[1,1’-biphenyl]-3-yl)methylene)-
benzohydrazide (TP2)
pound 14 in a manner similar to that described for the prepar-
1
ation of TP1 (reaction time: 12 h): mp ¼ 282.9 ꢂ 284.6 ꢃC; H NMR
The title compound was obtained in 63% overall yield from com- (500 MHz, DMSO-d6) d 6.96–7.01 (m,3H), 7.46 (dt, J ¼ 1.0, 8.0 Hz,
pound 14 in
a
manner similar to that described for the 1H), 7.52 (d, J ¼ 5.0 Hz, 1H), 7.61 (dd, J ¼ 3.0, 4.5 Hz, 1H), 7.67 (dd,

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
441
J ¼ 2.0, 8.5 Hz, 1H), 7.75 (d, J ¼ 1.5 Hz, 1H), 7.88–7.92 (m, 2H), 8.72 134.29, 137.00, 139.87, 148.14, 149.60, 153.73, 156.86, 159.38,
(s, 1H), 11.33 (s, 1H), 11.86–12.08 (m, 2H). 13C NMR (125 MHz, 165.21. HRMS (ESI) for C19H16N7O2 [M þ H]þ calculated 374.1365,
DMSO-d6) d 116.17, 117.43, 117.78, 119.27, 119.49, 120.01, 126.47, found 374.1367.
127.31, 127.45, 129.10, 129.97, 134.45, 141.22, 149.35, 157.24,
159.46, 165.05. HRMS (ESI) for C18H15N2O3S [M þ H]þ calculated
(E)-2-Hydroxy-N’-((2-hydroxynaphthalen-1-yl)methylene)-
benzohydrazide (TP8)
339.0803, found 339.0804.
The title compound was obtained in 78% overall yield from com-
(E)-N’-(6-(furan-3-yl)-2-hydroxy-3-methoxybenzylidene)-2-
pound 14 in a manner similar to that described for the prepar-
hydroxybenzohydrazide (TP4)
ation of TP1 (reaction time: 12 h): mp ¼ 223.8 ꢃC; 1H NMR
The title compound was obtained in 61% overall yield from com- (500 MHz, DMSO-d6) d 6.97–7.04 (m, 2H), 7.23 (d, J ¼ 9.0 Hz, 1H),
pound 14 in a manner similar to that described for the preparation 7.38–7.42 (m, 1H), 7.46 (t, J ¼ 7.5 Hz, 1H), 7.59 (t, J ¼ 8.0 Hz, 1H),
1
of TP1 (reaction time: 12 h): mp ¼ 228.3 ꢃC (decomposed); H NMR 7.87 (d, J ¼ 8.0 Hz, 1H), 7.91–7.94 (m, 2H), 8.30 (d, J ¼ 8.5 Hz, 1H),
(500 MHz, DMSO-d6) d 3.83 (s, 3H), 6.70 (s, 1H), 6.84 (d, J ¼ 8.0 Hz, 9.54 (s, 1H), 11.93 (s, 2H), 12.73 (s, 1H). 13C NMR (125 MHz, DMSO-
1H), 6.94–6.99 (m, 2H), 7.09 (d, J ¼ 8.5 Hz, 1H), 7.44–7.48 (m, 1H), d6) d 109.08, 116.17, 117.79, 119.37, 119.61, 121.43, 124.03, 128.21,
7.81–7.85 (m, 3H), 8.79 (s, 1H), 11.71 (s, 1H), 12.18 (s, 1H),12.51 (s, 128.29, 129.24, 129.40, 132.18, 133.39, 134.49, 148.17, 158.62,
1H). 13C NMR (125 MHz, DMSO-d6) d 56.37, 112.98, 114.89, 115.62, 159.29, 164.48. HRMS (ESI) for C18H15N2O3 [M þ H]þ calculated
115.69, 117.84, 119.38, 120.65, 123.60, 126.44, 128.73, 134.61, 307.1083, found 307.1085.
141.32, 143.96, 147.82, 149.33, 150.38, 159.79, 165.18. HRMS (ESI)
for C19H17N2O5 [M þ H]þ calculated 353.1137, found 353.1139.
2–(5-(4-Methoxyphenyl)thiazol-2-yl)-phenol (TP9)
A mixture of 2-hydroxybenzonitrile (1.0 eq), O,O-diethyl dithio-
(E)-2-Hydroxy-N’-((3-hydroxy-4-methoxy-[1,1’-biphenyl]-2-
phosphate (2.0 eq), and H2O was stirred at 80 ꢃC until reaction
yl)methylene)-benzohydrazide (TP5)
was complete (4 h). The reaction mixture was cooled, quenched
The title compound was obtained in 58% overall yield from com- with saturated NaHCO3, and extracted by ethyl acetate (3 times).
pound 14 in a manner similar to that described for the prepar- The organic layer was collected and dried under reduced pressure.
1
ation of TP1 (reaction time: 12 h): mp ¼ 257.9 ꢂ 259.4 ꢃC; H NMR The resulting residue was dissolved in EtOH, and then 4-methoxy-
(500 MHz, DMSO-d6) d 3.85 (s, 3H), 6.77 (d, J ¼ 8.0 Hz, 1H), 6.91 (t, phenacyl bromide (1.0 eq) was added. The mixture was heated to
J ¼ 7.5 Hz, 1H), 6.94 (d, J ¼ 8.5 Hz, 1H), 7.12 (d, J ¼ 8.5 Hz, 1H), reflux until the reaction was complete (4 h). The solvent was
7.34–7.37 (m, 2H), 7.42–7.51 (m, 4H), 7.78 (dd, J ¼ 1.0, 8.0 Hz, 1H), removed under reduced pressure. The resulting residue was puri-
8.54 (s, 1H), 11.64 (s, 1H), 12.16 (s, 1H), 12.56 (s, 1H). 13C NMR fied by recrystallization with hexane and MeOH to afford a white
1
(125 MHz, DMSO-d6)
d
56.39, 114.70, 115.34, 115.61, 117.82, solid TP9 (63%). mp ¼ 95.2 ꢂ 96.8 ꢃC; H NMR (500 MHz, DMSO-d6)
119.31, 120.82, 127.77, 128.67, 128.91, 130.36, 134.59, 135.97, d 3.84 (s, 3H), 6.94 (dt, J ¼ 1.0, 8.0 Hz, 1H), 7.00–7.03 (m, 3H), 7.33
139.53, 147.90, 149.22, 150.16, 159.84, 165.30. HRMS (ESI) for (dt, J ¼ 1.0, 7.5 Hz, 1H), 7.64 (s, 1H), 7.77 (dd, J ¼ 1.0, 7.5 Hz, 1H),
C21H19N2O4 [M þ H]þ calculated 363.1345, found 363.1347.
7.84 (d, J ¼ 9.0 Hz, 2H), 11.30 (s, 1H). 13C NMR (125 MHz, DMSO-d6)
d 54.41, 109.78, 113.91, 116.96, 117.10, 119.32, 126.18, 126.94,
127.10, 131.36, 154.01, 156.33, 160.15, 168.19. HRMS (ESI) for
C16H14NO2S [M þ H]þ calculated 284.0745, found 284.0748.
(E)-2-Hydroxy-N’-(2-hydroxy-3-methoxy-6-(thiophen-3-
yl)benzylidene)-benzohydrazide (TP6)
The title compound was obtained in 75% overall yield from com-
pound 14 in a manner similar to that described for the prepar-
ation of TP1 (reaction time: 12 h): mp ¼ 229.5 ꢃC; 1H NMR
(E)-N’-((2-Hydroxynaphthalen-1-yl)methylene)-4–(2-oxopyridin-
1(2H)-yl)-benzohydrazide (TP10)
(500 MHz, DMSO-d6) d 3.83 (s, 3H), 6.82 (d, J ¼ 8.5 Hz, 1H), The title compound was obtained in 40% overall yield from com-
6.91–6.97 (m, 2H), 7.08 (d, J ¼ 8.5 Hz, 1H), 7.20 (d, J ¼ 5.0 Hz, 1H), pound 14 in a manner similar to that described for the prepar-
7.44 (dt, J ¼ 1.5, 8.5 Hz, 1H), 7.52(d, J ¼ 1.5 Hz, 1H), 7.67 (dd, J ¼ 3.0, ation of TP1 (reaction time: 12 h): mp ¼ 258.6 ꢃC; 1H NMR
5.0 Hz, 1H), 7.81 (d, J ¼ 8.0 Hz, 1H), 8.67 (s, 1H), 11.69 (s, 1H), 12.19 (500 MHz, DMSO-d6) d 6.36 (t, J ¼ 6.5 Hz, 1H), 6.52 (d, J ¼ 9.0 Hz,
(s, 1H), 12.55 (s, 1H). 13C NMR (125 MHz, DMSO-d6) d 56.39, 114.72, 1H), 7.25 (d, J ¼ 9.0 Hz, 1H), 7.42 (t, J ¼ 7.5 Hz, 1H), 7.54 (t,
115.60, 115.64, 117.84, 119.35, 120.78, 124.66, 126.79, 128.71, J ¼ 7.5 Hz, 1H), 7.60–7.73 (m, 4H), 7.90 (d, J ¼ 8.0 Hz, 1H), 7.94 (d,
130.09, 130.63, 134.60, 139.82, 147.87, 149.24, 150.29, 159.83, J ¼ 9.0 Hz, 1H), 8.12 (d, J ¼ 8.0 Hz, 2H), 8.26 (d, J ¼ 8.0 Hz, 1H), 9.54
165.27. HRMS (ESI) for C19H16N2NaO4S [M þ Na]þ calculated (s, 1H), 12.36 (s, 1H), 12.73 (s, 1H). 13C NMR (125 MHz, DMSO-d6) d
391.0728, found 391.0731.
106.34, 109.02, 119.37, 121.11, 121.18, 124.04, 127.53, 128.28,
128.31, 128.89, 129.46, 132.11, 132.69, 133.31, 139.12, 141.29,
144.13, 147.63, 158.55, 161.51, 162.20. HRMS (ESI) for C23H18N3O3
[M þ H]þ calculated 384.1348, found 384.1351.
(E)-N’-(4–(6-amino-9H-purin-9-yl)benzylidene)-2-
hydroxybenzohydrazide (TP7)
The title compound was obtained in 33% overall yield from com-
pound 14 in a manner similar to that described for the prepar-
N’-(2-Hydroxy-1-naphthoyl)-nicotinohydrazide (TP11)
ation of TP1 (reaction time: 12 h): mp ¼ 229.5 ꢃC; 1H NMR The title compound was obtained in 55% overall yield from com-
(500 MHz, DMSO-d6) d 6.95–7.01 (m, 2H), 7.41–7.47 (m, 3H), pound 14 in a manner similar to that described for the prepar-
7.90–7.97 (m, 3H), 8.08 (d, J ¼ 8.5 Hz, 2H), 8.25 (s, 1H), 8.54 (s, 1H), ation of TP1 (reaction time: 3 h): mp ¼ 227.1 ꢂ 228.6 ꢃC; 1H NMR
8.68 (s, 1H), 11.79 (s, 1H), 11.91 (s, 1H). 13C NMR (125 MHz, DMSO- (500 MHz, DMSO-d6) d 7.04 (d, J ¼ 7.5 Hz, 1H), 7.13 (d, J ¼ 8.5 Hz,
d6) d 116.55, 117.74, 119.46, 119.88, 123.27, 128.74, 129.13, 133.39, 1H), 7.35 (d, J ¼ 9.0 Hz, 1H), 7.41 (t, J ¼ 7.5 Hz, 1H), 7.49 (dt, J ¼ 1.5,

442
T.A. MISKO ET AL.
9.0 Hz, 1H), 7.54 (dt, J ¼ 1.0, 8.0 Hz, 1H), 7.88 (dd, J ¼ 1.5, 7.5 Hz, having precipitates that did not pellet were excluded from the list
1H), 7.94 (d, J ¼ 8.0 Hz, 1H), 7.97 (d, J ¼ 8.5 Hz, 1H), 8.08 (d,
of 13 compounds examined in this manuscript.
J ¼ 9.0 Hz, 1H), 10.41 (s, 1H), 10.80 (s, 1H). 13C NMR (125 MHz,
DMSO-d6) d 103.07, 110.14, 117.64, 118.82, 120.29, 123.72, 124.17,
Protein expression and purification of hRRM1
128.05, 128.58, 128.97, 129.26, 132.65, 133.94, 134.20, 156.83,
157.04, 161.88, 164.22. HRMS (ESI) for C17H13N3NaO3 [M þ Na]þ
The hRRM1 protein was expressed in Escherichia coli Bl21- codon
calculated 330.0855, found 330.0850.
plus (DE3)-RIL cells and purified using a peptide affinity column,
as previously described in Fairman et al.38. The homogenous pro-
tein was pooled and concentrated to 0.2 mg/mL, quantified using
UV spectroscopy.
(E)-2-Hydroxy-N’-(pyridin-3-ylmethylene)-1-
naphthohydrazide (TP12)
The title compound was obtained in 45% overall yield from com-
KD determination by fluorescence quenching
pound 14 in a manner similar to that described for the prepar-
ation of TP1 (reaction time: 6 h): mp ¼ 202.8 ꢂ 204.5 ꢃC; 1H NMR
(500 MHz, DMSO-d6) d 7.18 (d, J ¼ 9.0 Hz, 1H), 7.31 (d, J ¼ 7.5 Hz,
1H), 7.44–7.50 (m, 2H), 7.78 (d, J ¼ 8.0 Hz, 1H), 7.82–7.86 (m, 2H),
8.22 (s, 1H), 8.35 (d, J ¼ 8.0 Hz, 1H), 8.54 (d, J ¼ 4.0 Hz, 1H), 8.85
(s, 1H). 13C NMR (125 MHz, DMSO-d6) d 114.48, 117.49, 123.11,
123.21, 124.14, 127.15, 127.92, 128.22, 130.92, 131.65, 132.00,
134.76, 145.03, 148.45, 149.99, 153.14, 166.39. HRMS (ESI) for
C17H14N3O2 [M þ H]þ calculated 292.1086, found 292.1088.
The dissociation constant (KD) was measured for each of the com-
pounds using a tryptophan florescence quenching assay, as
described previously with hRRM1 at 0.2 mg/mL36. Tryptophan flor-
escence spectra of hRRM1 were measured using
a Horiba
Fluoromax-4, 1155D-3113 FM spectrophotometer after exciting the
sample at 295 nm. A background spectrum was recorded with
protein in KD buffer followed by the incremental addition of com-
pounds (0–150 mM) at room temperature. The data were fitted in
Origin graphing software using a quadratic form of the equilib-
rium binding equation
N’-(2-Hydroxybenzoyl)-nicotinohydrazide (TP13)
ꢀ
ꢀ
ꢁ ꢁ
A mixture of methyl salicylate (1.0 eq), N2H4 (1.1 eq), and EtOH
was heated to reflux until the reaction was complete (2 h). The
reaction mixture was filtered and washed with EtOH to afford a
white solid. To a mixture of the resulting product (1.0 eq), K2CO3
(1.2 eq) and p-dioxane nicotinoyl chloride (1.2 eq) was added and
the resultant mixture was heated at 40 ꢃC until the reaction was
complete (4 h). The reaction mixture was quenched with H2O and
extracted by ethyl acetate (3 times). The organic layer was col-
lected, dried over MgSO4, and filtered. The resulting filtrate was
dried under reduced pressure to obtain TP13 as a white solid
pffiffiffiffiffiffiffi
4cx
2
ð
Þ
ð
Þ
ꢂ
A
c þ x þ K ꢂ c þ x þ K
y ¼
;
2c
where A is the amplitude of the reaction, c is the concentration of
enzyme, x is the substrate concentration, y is the percent of fluor-
escence quenching and K is the KD. Measurements were made in
duplicate to estimate error. KD’s were corrected for the measured
concentration of soluble compound from the concentration of
total added compound. Error of the corrected KD’s were calculated
(75%). mp ¼ 216.6 ꢂ 217.7 ꢃC; 1H NMR (500 MHz, DMSO-d6) d using the error propagation equation
6.95–7.00 (m, 2H), 7.46 (t, J ¼ 7.5 Hz, 1H), 7.57 (dd, J ¼ 5.0, 8.0 Hz,
sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
ꢂ
ꢃ
ꢂ
ꢃ
1H), 7.92 (d, J ¼ 7.0 Hz, 1H), 8.26 (d, J ¼ 8.0 Hz, 1H), 8.78 (d,
J ¼ 4.0 Hz, 1H), 9.08 (d, J ¼ 1.0 Hz, 1H), 10.73 (s, 1H), 10.90 (s, 1H),
11.83 (s, 1H). 13C NMR (125 MHz, DMSO-d6) d 115.18, 117.84,
119.59, 124.19, 128.51, 128.94, 134.66, 135.75, 148.93, 153.06,
159.53, 164.62, 167.91. HRMS (ESI) for C13H12N3O3 [M þ H]þ calcd
258.0879, found 258.0880.
2
rx
ra
rb
¼
2 þ
x
a
b
Where rx is the error of the corrected KD, x is the corrected KD,
ra is the error of the slope of the drug concentration plot, a is the
slope of the drug concentration plot, rb is the error of the meas-
ured KD and b is the measured KD.
Compound preparation and solubility determination
For each compound, stock solutions were made in 100% DMSO to
assure full solubility. Spectra of standard solutions in 100% aceto-
nitrile were taken using UV spectroscopy. An extinction coefficient
was calculated from the plot of the absorbance vs drug concen-
tration using Beer’s law (Supplemental Table 1).
Cancer cell line growth inhibition assay
Growth inhibition assays were performed in the Translational
Research Shared Resource of the Case Comprehensive Cancer
Centre. Human pancreatic cancer cells (Panc1) were maintained in
standard growth medium consisting of (RPMI1640 þ 10% FBS þ
2 mM glutamine þ 100 U/mL penicillin, and 100 mg/mL strepto-
mycin). Cells were monitored and shown to be negative for myco-
plasma contamination using the Mycoplasma Detection kit
(MycoAlertTM, Lonza, Basel, Switzerland). For growth inhibition
assays, cells were harvested by trypsinisation and seeded into 96-
well tissue culture plates at 2500 cells/mL. The following day, trip-
licate wells were treated with an appropriate volume of 5x-inhibi-
tor-containing medium. The cells were cultured for 3 additional
days at 37˚C in a 5% CO2 humidified incubator. Cell growth was
assessed by measuring total DNA content per well using an adap-
KD values were determined in KD buffer [50 mM TRIS, pH 8.0,
5 mM MgCl2, 5% glycerol and 10 mM DTT]. Because the solubility
of the compounds was limited in this buffer, the solubility was
empirically determined as follows. Increasing amounts of a com-
pound were dissolved in 1.5 ml of KD buffer. The solutions were
mixed by vortex and centrifuged at 5000ꢀg for 10 min, and the
supernatant was promptly removed and absorbance measured by
UV/Vis spectrophotometry. Using the extinction coefficients, the
concentration of soluble drug was calculated for each total con-
centration (Supplemental Figure 1). Drug solubility measurements
were carried out in duplicate. Plots of measured vs predicted con-
centrations were plotted in Origin graphing software. Samples tation of the method of Labarca and Paigen39.

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
443
Scheme 1. Synthetic approach to compounds TP1–13.
In silico docking studies
heterocycles were also designed. Their syntheses are illustrated in
Scheme 1A and 1B. Methyl salicylate (14) was reacted with hydra-
zine to obtain 2-hydroxybenzohydrazide (15) which was subjected
to reaction with various aldehydes or isatin to give the corre-
sponding products TP1-8 and TP10 (Figure 2(B)). Methyl 2-
hydroxy-1-naphthoate (16) underwent a similar synthetic route to
afford compound TP12 (Figure 2(B)). Meanwhile, the reaction of
15 and 17 with nicotinoyl chloride yielded, respectively, com-
pounds TP13 and TP11, which possess a diacylhydrazine linkage
(Figure 2(B)). The reaction of 2-hydroxybenzonitrile (18) with O,O-
diethyl dithiophosphate, which generated the corresponding thio-
amide product, was followed by cyclisation with 4-methoxyphe-
nacyl bromide to furnish compound TP9. These analogues were
designed to enhance interactions with the C-site (Figure 1).
In silico docking of the compounds was performed using the
Glide module of the Schrodinger 2017–3 modelling software suite
€
as previously described36,40–42. The docking site for the C-site was
defined as a 5 Å box centred on the lead compound NSAH (TP8)
bound to the hRRM1 (PDB code 5TUS)37. Compounds were
docked to the C-site using Glide SP. Hits were scored by a glide
scoring function and examined for their interactions with the C-
site residues using Maestro.
Results
Rationale for compound design and synthesis
Our previous study identified NSAH (TP8), containing a 2-hydroxy-
benzohydrazide moiety, as an RR modulator. Its structure contains
three parts, namely: 2-hydroxybenzoyl group, hydrazide linkage,
and naphthalene ring. In order to understand structure-activity
relationships, specifically the influence of the hydrazide moiety,
we designed diacylhydrazine-containing TP11 and TP13 and thia-
zole derivative TP9 (Figure 2(B)). Furthermore, hydrazide deriva-
In silico compound docking interactions with hRRM1
Potential interactions of TP1-13 with hRRM1 were investigated
using the Schrodinger docking software suite glide40–42. A hRRM1
€
dimer with lead compound NSAH bound to the C-site (PDB ID:
5TUS) was used as a model for the docking of the new com-
pounds35. It was assumed that the primary site of binding for these
tives (TP1-8, TP10 and TP12) linked with biaryls or various compounds would be the C-site, because NSAH and all previously

444
T.A. MISKO ET AL.
Figure 3. Predicted binding interactions of TP compounds at the C-site of hRRM1.
studied derivatives of NSAH have demonstrated a preference for that ligands quenching at least 25% of the fluorescence have suffi-
the C-site35,37. Our studies showed that compounds TP2–7, 9–10 cient affinity for hRRM1, TP1-13 were characterised as binders of
are longer and therefore extend beyond NSAH, allowing for the hRRM1. TP1, 4, 6, 7, 10, and 12 quenched at least 90% of the tryp-
possibility of more interactions (Figure 3(A–D) and Supplemental tophan fluorescence (Figure 4(A–B)). TP2, 5, 8, 9, 11, and 13 each
Figure 1(B–F)). TP3 was not predicted to bind well to the phos- quenched approximately 80% (Figure 4(C)), and TP3 quenched
phate-binding site in comparison to other new compounds approximately 70% of tryptophan fluorescence. While percentage
(Figure 3(A), Supplemental Figure 2(C)). However, docking pre- of quenching indicates the ability of a compound to bind to the
dicted it to have an increase in the number of strong interactions enzyme, measuring the concentration of a compound needed to
closer to the loop 2 region. TP7 was unique in that it was pre- quench the fluorescence by half provides the KD, a measure of the
dicted to interact strongly in the phosphate-binding site with the affinity of a compound for the enzyme. Corrected KD’s (see
residues: Thr607, Ser448, Ser606, and near the loop 2 region with Methods) ranged from 2.9 0.39 – 22.0 5.67 mM, where all of the
residues: Pro294, Ala245, as well as to a residue to which NSAH new compounds have a lower KD than NSAH (22.0 5.67 mM)
exhibits strong hydrogen bonding (Ser217) (Figure 3(C), (Table 1). The reduction in the KD indicates that the chemical modi-
Supplemental Figure 2(G)). This is partly because TP7 extends to fications targeting the phosphate-binding site and loop 2 resulted
the outer extremities of the catalytic site (C-site). Thus, for all com- in better binding to hRRM1 as compared to NSAH (Table 1).
pounds except TP3, there was either an increase in the number of
predicted C-site interactions and/or an increase in the number of
strong binding interactions to the phosphate-binding site of the C-
site, in comparison to the interactions that NSAH makes (Figures
3(A–D), Supplemental Figure 1(A–I), Supplemental Figure 2(A–M)).
Inhibition of growth of Panc1 pancreatic cancer cells
Since gemcitabine is a core component of the current standard of
care for pancreatic cancer, and we hoped to improve upon the
therapeutic index for drugs treating this disease, we selected
Panc1, a cell line derived from pancreatic cancer, for the study of
the new compounds. Panc1 cells growing in 96-well plates were
exposed to each compound continuously for three days, after
Investigation of compound binding to hRRM1 using intrinsic
tryptophan fluorescence
TP1-13 were analysed for their ability to bind to hRRM1 and there- which DNA content per well was measured as an indicator of cell
fore quench its intrinsic tryptophan fluorescence. Using the criterion growth. As shown in Figure 5 and Table 1, TP1, 7, 9, 11, 12, and

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
445
Figure 4. Quenching of tryptophan fluorescence of hRRM1 by TP ligands.
13 did not inhibit the growth of Panc1 cells, even at concentra- order to better target inhibitors for therapeutic intervention. While
numerous diverse inhibitors have been developed to target hRR
and several have been approved for clinical use, there have been
few successful attempts to identify a non-nucleoside reversible
inhibitor that targets the large subunit of hRR. In a recent paper
by Ahmad et al., we reported on a rapid moderate-throughput
screen that identified the first non-nucleoside inhibitor binding to
the C-site, NSAH35,36. In a subsequent study by Huff et al., a library
of compounds was produced, replacing the substituents on either
side of the hydrazone moiety of NSAH with polar substituents to
examine their effects on activity37. It was determined that replac-
ing the phenol with polar substituents in the ortho position
achieved the best activity against hRR. Replacement of the naph-
thalene with an indole provided similar activity. It was concluded
that these derivatives favoured the C-site, just like NSAH, and all
derivatives that showed improved activity against hRR in compari-
son to NSAH had increased interactions in the phosphate-binding
site. However, these compounds had limited cellular activity.
Using the co-crystal structure of NSAH bound at the C-site of
tions of 10 mM. Compounds TP2, 3, and 10 showed moderate
growth inhibition with IC50’s between 2 and 3 mM (Figure 5 and
Table 1). Compounds TP4–6, and 8 were the most potent against
this cell line, with IC50’s below 1.5 mM (Figure 5 and Table 1).
Overall, the results indicated that some compounds in group 1
have marked activity against the growth of Panc1 cells, whereas
group 2 is mostly ineffective (Figure 5 and Table 1). TP6 demon-
strated the greatest potency, with over a 2-fold decrease in IC50
over the lead compound NSAH (TP8) (Figure 5 and Table 1). A few
of the compounds were also studied in other cell lines (A549 and
HCT-116), showing similar potency as in Panc1 cells (Supplemental
Figure 3). This finding suggests that these compounds do not tar-
get Panc1 cells specifically, but may be universally active against
cancer cells, as would be expected since RR is found in all cells.
Discussion and conclusions
Decades of effort have been invested into elucidating the struc-
ture and physiological function of ribonucleotide reductase, in hRRM1 and its interactions (PDB code 5TUS) as a template in the

446
T.A. MISKO ET AL.

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
447
present study, a small library of compounds was designed to
increase the interactions with the C-site (Figure 2). It was antici-
pated that improved interactions would strengthen the specificity
towards hRR and reduce off-target binding. The library consists of
two distinct groups. For the Group 1 compounds, the naphthalene
of NSAH was replaced in anticipation of creating more interactions
with the residues that are important in binding the phosphates of
the natural substrates (termed phosphate-binding site). For the
Group 2 compounds, the hydrazine linker was modified and polar
groups were added to these linkers. In order to determine how
these modifications affected interaction with hRR, computational
docking and experimental assays were conducted.
To determine the extent of binding of these compounds to the
hRRM1 subunit, quenching of the fluorescence of the internal
tryptophans of hRRM1 was measured. The KD’s ranged from
2.9 0.39 – 22.0 5.67 mM, indicating that all of the modified com-
pounds bind more tightly than does NSAH (KD- 22.0 5.67 mM)
(Table 1). Examination of the docking of these compounds sug-
gests that the overall decrease in KD is influenced by the pre-
dicted increase in interactions with the phosphate-binding site
within the C-site of hRRM1 compared to the binding of NSAH.
With the exception of TP3, all other analogues have stronger
interactions with the phosphate-binding site than NSAH
(Figure 3(B–D), Supplemental Figure 1(A–I), Supplemental
Figure 2(A–M)). Apart from the increase in binding to the phos-
phate-binding region, no other single binding characteristic
explains the trends in relative affinity seen with these 13 com-
pounds; rather, a collection of interactions contribute to the
observed binding pattern. All of the compounds with a KD less
than 6.2 mM are predicted to make at least one strong hydrogen
bond that could contribute to the tight binding to hRRM1. Within
this group, TP7 and 11 are predicted to have the most hydrogen
bonds, leading to stronger binding to hRRM1 (Figure 3(C) and
Supplemental Figure 1(H)). TP7 has hydrogen bonds scattered
throughout the length of the compound, interacting with the
phosphate-binding site, at Cys218 and Ser217, and near loop 2
(Figure 3(C)). Our previous predictions for optimising interactions
at the phosphate-binding site and with loop 2 appears to be vali-
dated by this study. TP11 and 13 are the only compounds in this
library that have a linker composed of single bonds, allowing for
free rotation (Figure 2(B)). With a few exceptions, compounds with
a KD less than 9.4 mM form multiple predicted hydrogen bonds.
TP3 is one of the exceptions, making fewer strong interactions
than NSAH, and not interacting with the phosphate-binding site
(Figure 3(A) and Supplemental Figure 2(C)). TP3’s lack of interac-
tions with the phosphate-binding site is compensated by stronger
interactions near loop 2. The trend that emerges from the study
of this library is that the best hRRM1 binding compounds are
either small and flexible (TP1, 11, 13) or long enough to reach
loop 2 and/or the phosphate-binding site (TP3–7). Both of these
groups of compounds’ options contain polar substituents, but
there is a limit to the benefit of size and polarity, because some
of the larger molecules had low solubility in the aqueous medium
of the fluorescence assay.
A common challenge in drug development is that interaction
with a target and inhibitory activity against a cell-free enzyme
does not always predict how the drug will perform when tested
in a cell or animal model. Thus, another approach we employed
was to query the structures of the compounds in our library using
€
QikProp, an algorithm of the Schrodinger suite to predict the cell
permeability of these compounds in Caco-2 and MDCK cells
(Table 1)43. With some exceptions, the algorithm was useful in
predicting which compounds might be the most effective against

448
T.A. MISKO ET AL.
Panc1 cells. With the exception of TP1 and 7, group 1 had the
greatest potency in Panc1 cells with IC50’s below 2.5 mM (Figure 5
and Table 1). TP6, which had the greatest potency, had the great-
est predictable permeability for both Caco-2 and MDCK cells
within the top seven compounds (Table 1). TP4–6 and 8 have
similar predicted permeabilities, with the exception of TP6, which
has a greater permeability for MDCK cells (Table 1). The com-
pounds with IC50’s between 2 and 3 mM are predicted to be less
permeable than TP4–6 and 8 (Table 1). Out of the compounds
that had cellular IC50’s greater than 10 mM (TP1, 7, 9, 11–13) only
T9 and 12 had good predicted permeability, even greater than
that of TP6 (Table 1). These results suggest that theoretical predic-
tion of permeability does not always predict activity in cells. The
lack of activity against cells could be due to an assortment of fac-
tors, including poor cell permeability, drug metabolism in the cell
rendering them inactive, or being good substrates for
efflux pumps.
Figure 5. Cancer growth-inhibitory activity of Panc1 pancreatic cancer cell line
by TP compounds.
TP4–6, which were the most potent against growth of Panc1
cells, have very similar structures (Figure 2(B)). These compounds
maintain the phenol and the hydrazine linker of NSAH, but in
PAINS candidate, our studies have validated this compound as an
place of the naphthalene is a guaiacol methoxyphenol linked to a RR inhibitor binding to the C-site of hRR1 and inhibiting RR in
cells35,37. This validation adds confidence that TP1–13 would bind
furan, benzene, or thiophene. The docking of these compounds to
the hRRM1 C-site provides evidence that they bind similarly to
each other, revealing the importance of this modification. To fur-
ther examine this modification and its contribution to enhanced
binding and cellular potency, TP2 and 3 were compared. TP2 and
3 mimic TP5 and 6, respectively, but lack the methoxy group on
the guaiacol methoxyphenol (Figure 2(B)). The removal of the
methoxy appears to cause over a 2-fold decrease in potency in
cells as well as the loss of a similar docking pattern (Table 1,
Supplemental Figure 1(B) and Figure 3(A)). In the study by Huff
et al., a few compounds contained a methoxy group, although on
a different position, which caused a similar type of docking pat-
tern37. However, while the methoxy seems to be an important
entity and might be essential for better potency, it does not do so
alone. TP9, which contains a methoxy group on a benzene ring, is
not only ineffective in Panc1 cells but also has a completely differ-
ent predicted pattern of binding to the C-site (Figure 5, Table 1
and Supplemental Figure 1(E) and 2(I)). This suggests that methox-
ylation of the benzene ring aids in improving cell and enzyme
potency but acts in concert with other polar entities. The addition
of the hydroxyls as well as the polar rings aids in the increase in
potency. Considering the effects of TP4–6 in the cells, it might be
suggested that a polar ring with an electronegative element
might further increase the relative potency.
to the hRR1 C-site as docking to this site displayed favourable
poses and possibly inhibits RR in cells. When developing com-
pounds against RR, there is always a concern that these com-
pounds contain chelating properties that may affect the Fe and
free radical housed in the small subunit, a crucial element needed
for the activity of RR. There have been many reports of agents
that inhibit RR by this mechanism32,45–49. As this was a concern,
NSAH was tested for its ability to chelate metals37. Results indi-
cated that NSAH does not chelate Fe3þ, the iron form present in
RR, up to concentrations of 5 mM nor does it bind or sequester
Mg2þ ions, which are also critical for RR activity. As these concerns
are not an issue for the lead compound NSAH, we predict that
these will not be an issue with TP1–13 either.
It is generally understood that targeting RR for cancer thera-
peutics is a challenging task as RR is present in all cells, normal
and malignant. We have previously shown that the lead com-
pound NSAH (TP8) was much less effective against blood progeni-
tor cells than against cancer cell lines, demonstrating that NSAH
has a higher therapeutic index than gemcitabine in the same cell-
cell comparison35. This suggests that NSAH has selective activity
in malignant cells, exploiting the greater proliferation in cancer
cells50. Accordingly, the new class of inhibitors designed from
NSAH might provide
a superior, safer anticancer treatment.
It is well known that phenolics have potentials for being che-
motherapeutic agents; however, these compounds often have
poor bioavailability in vivo44. Typically, phenolics are easily oxi-
dised, leaving them vulnerable to chemical degradation or bacter-
ial decomposition in the GI tract, most commonly by
glucuronidation and sulphation. TP1–8, 13 contain phenolic acids,
which proposes a risk of poor bioavailability when administered in
vivo, eliminating an oral administration and restricting it to IV
administration. As an oral compound is more desirable for
patients for its ease of use, it might be beneficial to modify these
compounds if they exhibit poor bioavailability in mouse studies.
Modifying these compounds by converting the phenolic to esters
or ethers might produce a pro-drug and increase the bioavailabil-
ity of the phenolic metabolites.
Results from this study indicate that replacing the naphthalene
ring of NSAH with other cyclic ring structures containing methoxy
and polar constituents results in analogues with superior activity
against cancer cells. Future studies will elucidate cyclic and polar
constituents which may provide further improvements in this class
of RR inhibitors.
Acknowledgements
The authors thank Dr. William E. Harte (School of Medicine, CWRU)
for insightful comments and suggestions during the development
of this project. The authors thank Drs. J. Mieyal, R. Viswanathan, S.
Huff, M. Kumar, and D. Wald, for useful discussions. This work
made use of the High Performance Computing Resource in the
Core Facility for Advanced Research Computing at Case Western
A PAINS filter (http://cbligand.org/PAINS/), which is a now well-
defined way to identify the presence of potentially problematic
functional groups in analogue design, was used on our compound
library. Despite our lead compound, NSAH, being a potential Reserve University.

JOURNAL OF ENZYME INHIBITION AND MEDICINAL CHEMISTRY
449
13. Jordheim LP, Seve P, Tredan O, Dumontet C. The ribonu-
cleotide reductase large subunit (RRM1) as a predictive fac-
tor in patients with cancer. Lancet Oncol 2011;12:693–702.
14. Shao J, Zhou B, Chu B, Yen Y. Ribonucleotide reductase
inhibitors and future drug design. Curr Cancer Drug Targets
2006;6:409–31.
15. Stearns B, Losee KA, Bernstein J. Hydroxyurea. A new type
of potential antitumour agent. J Med Chem 1963;6:201.
16. Donehower RC. An overview of the clinical experience with
hydroxyurea. Semin Oncol 1992;19:11–9.
17. Huang P, Chubb S, Plunkett W. Termination of DNA synthe-
sis by 9-beta-D-arabinofuranosyl-2-fluoroadenine. A mechan-
ism for cytotoxicity. J Biol Chem 1990;265:16617–25.
18. Heinemann V, Xu YZ, Chubb S, et al. Inhibition of ribonu-
cleotide reduction in CCRF-CEM cells by 2’,2’-difluorodeoxy-
cytidine. Mol Pharmacol 1990;38:567–72.
Disclosure statement
The authors report no declarations of interest and also declare no
competing financial interest.
Funding
This study was supported by NIH funding to P.I.: Dr. Chris G.
Dealwis (R01GM100887), Case Comprehensive Cancer Centre CTSC
pilot grant and Council to Advance Human Health. Drs. Lee and
Dealwis were funded by the Taipei Medical School, Taiwan-CWRU
grant. This research was supported in part by the Translational
Research Shared resource of the Case Comprehensive Cancer
Centre (P30CA043703) and Center for Scientific Review.
ORCID
19. Parker WB, Shaddix SC, Chang CH, et al. Effects of 2-chloro-
Tessianna A. Misko
http://orcid.org/0000-0002-9270-7382
9-(2-deoxy-2-fluoro-beta-D-arabinofuranosyl)adenine
on
K562 cellular metabolism and the inhibition of human ribo-
nucleotide reductase and DNA polymerases by its 5’-triphos-
phate. Cancer Res 1991;51:2386–94.
References
20. Parker WB, Shaddix SC, Rose LM, et al. Comparison of the
mechanism of cytotoxicity of 2-chloro-9-(2-deoxy-2-fluoro-
beta-D-arabinofuranosyl)adenine, 2-chloro-9-(2-deoxy-2-flu-
oro- beta-D-ribofuranosyl)adenine, and 2-chloro-9-(2-deoxy-
2,2-difluoro- beta-D-ribofuranosyl)adenine in CEM cells. Mol
Pharmacol 1999;55:515–20.
21. Griffig J, Koob R, Blakley RL. Mechanisms of inhibition of
DNA Synthesis by 2-chlorodeoxyadenosine in human
lymphoblastic cells. Cancer Res 1989;49:6923–8.
22. Avery TL, Rehg JE, Lumm WC, et al. Biochemical pharmacol-
ogy of 2-chlorodeoxyadenosine in malignant human hem-
atopoietic cell lines and therapeutic effects of 2-
bromodeoxyadenosine in drug combinations in mice.
Cancer Res 1989;49:4972–8.
23. Xie C, Plunkett W. Metabolism and actions of 2-chloro-9-(2-
deoxy-2-fluoro-beta-D- arabinofuranosyl)-adenine in human
lymphoblastoid cells. Cancer Res 1995;55:2847–52.
1. Brown NC, Reichard P. Role of effector binding in allosteric
control of ribonucleoside diphosphate reductase. J Mol Biol
1969;46:39–55.
2. Stubbe J, Ge J, Yee CS. The evolution of ribonucleotide
reduction revisited. Trends Biochem Sci 2001;26:93–9.
3. Jordan A, Reichard P. Ribonucleotide reductases. Annu Rev
Biochem 1998;67:71–98.
4. Kolberg M, Strand KR, Graff P, Andersson KK. Structure, func-
tion, and mechanism of ribonucleotide reductases. Biochim
Biophys Acta 2004;1699:1–34.
5. Weinberg G, Ullman B, Martin DW. Jr,.: Mutator phenotypes
in mammalian cell mutants with distinct biochemical defects
and abnormal deoxyribonucleoside triphosphate pools. Proc
Natl Acad Sci USA 1981;78:2447–51.
6. Kunz BA, Kohalmi SE, Kunkel TA, et al. International
Commission for Protection Against Environmental Mutagens
and Carcinogens. Deoxyribonucleoside triphosphate levels: a
critical factor in the maintenance of genetic stability. Mutat
Res 1994;318:1–64.
7. Huang M, Zhou Z, Elledge SJ. The DNA replication and dam-
age checkpoint pathways induce transcription by inhibition
of the Crt1 repressor. Cell 1998;94:595–605.
24. King RS. Gemcitabine. New first-line therapy for pancreatic
cancer. Cancer Pract 1996;4:353–4.
25. Baker CH, Banzon J, Bollinger JM, et al. 2’-Deoxy-2’-methyle-
necytidine and 2’-deoxy-2’,2’-difluorocytidine 5’-diphos-
phates:
potent
mechanism-based
inhibitors
of
ribonucleotide reductase. J Med Chem 1991;34:1879–84.
26. Gandhi V, Plunkett W. Modulatory activity of 2’,2’-difluoro-
deoxycytidine on the phosphorylation and cytotoxicity of
arabinosyl nucleosides. Cancer Res 1990;50:3675–80.
27. Wang J, Lohman GJ, Stubbe J. Enhanced subunit interac-
tions with gemcitabine-5’-diphosphate inhibit ribonucleotide
reductases. Proc Natl Acad Sci USA 2007;104:14324–9.
28. Heinemann V, Schulz L, Issels RD, Plunkett W. Gemcitabine:
a modulator of intracellular nucleotide and deoxynucleotide
metabolism. Semin Oncol 1995;22:11–8.
29. Heinemann V, Xu YZ, Chubb S, et al. Cellular elimination of
2’,2’-difluorodeoxycytidine 5’-triphosphate: a mechanism of
self-potentiation. Cancer Res 1992;52:533–9.
30. Honeywell RJ, Ruiz van Haperen VW, Veerman G, et al.
Inhibition of thymidylate synthase by 2’,2’-difluoro-2’-deoxy-
cytidine (Gemcitabine) and its metabolite 2’,2’-difluoro-2’-
deoxyuridine. Int J Biochem Cell Biol 2015;60:73–81.
8. Yao R, Zhang Z, An X, et al. Subcellular localization of yeast
ribonucleotide reductase regulated by the DNA replication
and damage checkpoint pathways. Proc Natl Acad Sci USA
2003;100:6628–33.
9. Zhao X, Muller EG, Rothstein R. A suppressor of two essen-
tial checkpoint genes identifies a novel protein that nega-
tively affects dNTP pools. Mol Cell 1998;2:329–40.
10. Brown NC, Canellakis ZN, Lundin B, et al. Ribonucleoside
diphosphate reductase. Purification of the two subunits, pro-
teins B1 and B2. Eur J Biochem 1969;9:561–73.
11. Holmgren A, Reichard P, Thelander L. Enzymatic synthesis of
deoxyribonucleotides, 8. The effects of ATP and dATP in the
CDP reductase system from E. coli. Proc Natl Acad Sci USA
1965;54:830–6.
12. Jordheim LP, Guittet O, Lepoivre M, et al. Increased expres-
sion of the large subunit of ribonucleotide reductase is
involved in resistance to gemcitabine in human mammary 31. Heinemann V. Ongoing selective internal radiation therapy-
adenocarcinoma cells. Mol Cancer Ther 2005;4:1268–76. based studies in the treatment of liver-dominant metastatic

450
T.A. MISKO ET AL.
colorectal cancer. Future Oncol (London, England) 2014;10: 41. Halgren TA, Murphy RB, Friesner RA, et al. Glide: a new
37–9.
approach for rapid, accurate docking and scoring. 2.
Enrichment factors in database screening. J Med Chem
2004;47:1750–9.
32. Heinemann V, Plunkett W. Modulation of deoxynucleotide
metabolism by the deoxycytidylate deaminase inhibitor
3,4,5,6-tetrahydrodeoxyuridine. Biochem Pharmacol 1989;38:
4115–21.
33. Pourquier P, Gioffre C, Kohlhagen G, et al. Gemcitabine
(2’,2’-difluoro-2’-deoxycytidine), an antimetabolite that poi-
sons topoisomerase I. Clin Cancer Res 2002;8:2499–504.
34. Pourquier P, Pilon AA, Kohlhagen G, et al. Trapping of mam-
malian topoisomerase I and recombinations induced by
damaged DNA containing nicks or gaps. Importance of DNA
end phosphorylation and camptothecin effects. J Biol Chem
1997;272:26441–7.
35. Ahmad MF, Alam I, Huff SE, et al. Potent competitive inhib-
ition of human ribonucleotide reductase by a nonnucleoside
small molecule. Proc Natl Acad Sci USA 2017;114:8241–6.
36. Ahmad MF, Huff SE, Pink J, et al. Identification of non-
nucleoside human ribonucleotide reductase modulators. J
Med Chem 2015;58:9498–509.
37. Huff SE, Mohammed FA, Yang M, et al. Structure-guided
synthesis and mechanistic studies reveal sweetspots on
naphthyl salicyl hydrazone scaffold as non-nucleosidic com-
petitive, reversible inhibitors of human ribonucleotide reduc-
tase. J Med Chem 2018;61:666–80.
38. Fairman JW, Wijerathna SR, Ahmad MF, et al. Structural basis
for allosteric regulation of human ribonucleotide reductase
by nucleotide-induced oligomerization. Nat Struct Mol Biol
2011;18:316–22.
39. Labarca C, Paigen K. A simple, rapid, and sensitive DNA
assay procedure. Anal Biochem 1980;102:344–52.
40. Friesner RA, Banks JL, Murphy RB, et al. Glide: a new
approach for rapid, accurate docking and scoring. 1. Method
and assessment of docking accuracy. J Med Chem 2004;47:
1739–49.
42. Friesner RA, Murphy RB, Repasky MP, et al. Extra precision
glide: docking and scoring incorporating a model of hydro-
phobic enclosure for protein-ligand complexes. J Med Chem
2006;49:6177–96.
43. Jorgensen WL, Duffy EM. Prediction of drug solubility from
structure. Adv Drug Deliv Rev 2002;54:355–66.
44. Gao S, Hu M. Bioavailability challenges associated with
development of anti-cancer phenolics. Mini Rev Med Chem
2010;10:550–67.
45. Lepoivre M, Flaman JM, Bobe P, et al. Quenching of the
tyrosyl free radical of ribonucleotide reductase by nitric
oxide. Relationship to cytostasis induced in tumor cells by
cytotoxic macrophages. J Biol Chem 1994;269:21891–7.
46. Szalai VA, Brudvig GW. Reversible binding of nitric oxide to
tyrosyl radicals in photosystem II. Nitric oxide quenches for-
mation of the S3 EPR signal species in acetate-inhibited
photosystem II. Biochemistry 1996;35:15080–7.
47. Nyholm S, Mann GJ, Johansson AG, et al. Role of ribonucleo-
tide reductase in inhibition of mammalian cell growth by
potent iron chelators. J Biol Chem 1993;268:26200–5.
48. Hershko C. Control of disease by selective iron depletion: a
novel therapeutic strategy utilizing iron chelators. Bailliere’s
Clin Haematol 1994;7:965–1000.
49. Lundberg JH, Chitambar CR. Interaction of gallium nitrate
with fludarabine and iron chelators: effects on the prolifer-
ation of human leukemic HL60 cells. Cancer Res 1990;50:
6466–70.
50. Hakansson P, Hofer A, Thelander L. Regulation of mamma-
lian ribonucleotide reduction and dNTP pools after DNA
damage and in resting cells. J Biol Chem 2006;281:7834–41.