Discovery and characterization of small-molecule inhibitors of 8-oxoguanine
DNA glycosylase 1
Mari Eknes Ytre-Arne
Thesis for the degree of Philosophiae Doctor (PhD)
Department of Microbiology Oslo University Hospital
University of Oslo 2017
© Mari Eknes Ytre-Arne, 2018 Series of dissertations submitted to the Faculty of Medicine, University of Oslo ISBN 978-82-8377-217-3 All rights reserved. No part of this publication may be reproduced or transmitted, in any form or by any means, without permission. Cover: Hanne Baadsgaard Utigard. Print production: Reprosentralen, University of Oslo.
ACKNOWLEDGEMENTS
The work presented in this thesis was carried out from July 2013 to December 2017 at
the Department of Medical Biochemistry and the Department of Microbiology,
Rikshospitalet, Oslo University Hospital. In addition, six months were spent at the
European Molecular Biology Laboratory/European Synchrotron Radiation Facility,
Grenoble, France. Financial support for the project was provided by the Norwegian
Research Council (SYNKNØYT).
First of all I would like to express my sincere gratitude to my main supervisor Dr.
Bjørn Dalhus. Thank you for giving me the opportunity to take a PhD, and for
introducing me to the field of structural biology. Your never-ending patience and
fabulous pedagogic skills are highly appreciated. Thank you for all the Grenoble trips,
all the pizza dinners at Le Petit Rochers, all your stories and for critical reading of the
manuscripts and my thesis.
This degree would not have been possible without my co-supervisor, Professor Magnar
Bjørås. I feel very fortunate that I ended up in your lab. Not just because you are a
brilliant scientist, but also because you are a fabulous motivator. "Æ skjøinn itj
krisestæmninga" and "vi kainn itj bestrid nattur'n" pretty much sums it up. Thank you
for always seeing the jar as half-full, for your enthusiasm and always being available
for questions and scientific discussions. Finally, thank you for your valuable feedback
on my manuscripts and thesis and for all the support throughout this work.
I am also indebted to Dr. José Antonio Márquez for hosting me at EMBL, Grenoble,
and all the members of the HTX-team. Thank you for welcoming me into the group
and making me feel I belonged there from Day 1. I am especially grateful to Guillaume
Hoffmann and Dr. Irina Cornaciu for putting down numerous hours in the lab working
on optimizing the soaking and harvesting of the OGG1 crystals.
Further I would like to thank all the collaborators and co-authors for their valuable
work and contributions. A special thanks to Dr. Øyvind Jacobsen, my third supervisor,
for the scientific discussions, for always keeping an open mind, answering all my
stupid questions about chemistry and interactions, and for your brilliant pedagogic
skills. I am very grateful for your thorough reading of and contributions to the
manuscripts. Pernille Blicher, thank you for outstanding technical assistance, for being
the perfect room-mate on countless conferences, for all the fun (and sometimes
completely shameless) lunch topics, and most of all for being a really good friend.
Thank you for taking care of everyone in the lab, and basically running it. When you
make it to the olympics, this lab is going down.
I would also like to thank Dr. Mari Kaarbø for valuable feedback and critical reading
of my thesis and for being able to answer almost every scientific and non-scientific
question, and Professor Lars Mørkrid for statistical assistance, feedback on my thesis,
for your kindness and for making sure I always knew when there would be homemade
bread and jam in the lunch room.
Present and former co-workers, thank you for creating such a friendly and inspiring
work environment. I will truly miss you. Thanks to Dr. Paul Hoff Backe for helpful
discussions about structure biology, Dr. Alexander Rowe for introducing me to R, for
letting me borrow your iMac while writing my thesis, and for always being supportive,
and to Anna, Emma, Tine, Lene and Alexander Ø for sharing ups and downs and office
the last four years. Henning, thank you for numerous cups of coffee and for being a
verbal punching bag whenever I needed one ;)
To Janne, Ina and the rest of Sosietuppene: Thank you for all the encouragement and
support, and for making my lab life and social life so much more enjoyable!
Camilla & Tore, thank you for being around, and for always helping out. I am forever
grateful.
I would like to thank my family for always believing in me, and especially my parents
for the nerdy genes. A special thanks to my mother for always coming to our rescue.
My dear Hallvar, I am deeply grateful for your patience and support. Thank you for
making me coffee every morning, and the world's best pizza every Friday, for cooking
just about every meal for me the last four years, and for making sure we are never out
of wine or Parma ham or nice cheeses (although occasionally out of bread and other
everyday necessities). Most of all, thank you for taking such good care of our son
while I was working. I love you!
To my sweet little boy. The hardest part was being away from you. Nå kan vi bygge
togbane :)
Oslo, December 2017
Mari Eknes Ytre-Arne
TABLE OF CONTENTS LIST OF PAPERS .............................................................................................................. 1
ABBREVIATIONS ............................................................................................................ 2 SUMMARY ........................................................................................................................ 5
1. INTRODUCTION ....................................................................................................... 7 1.1. DNA damage and repair .................................................................................................... 7
1.1.1. The BER pathway ...................................................................................................... 8 1.2. Human 8-oxoguanine glycosylase 1 (OGG1) ................................................................. 12
1.2.1. OGG1 structure and lesion recognition .................................................................... 12 1.2.2. OGG1's mechanism of catalysis ............................................................................... 15 1.2.3. OGG1 phenotypes .................................................................................................... 15 1.2.4. OGG1 is involved in epigenetic regulation of gene transcription ............................ 15 1.2.5. OGG1 as a drug target for cancer treatment ............................................................. 17
1.3. PARP1 inhibitors ............................................................................................................ 18 2. AIMS ......................................................................................................................... 19
3. ABSTRACTS OF PAPERS ...................................................................................... 20 4. DISCUSSION ............................................................................................................ 22
4.1. Drug Discovery ............................................................................................................... 22 4.1.1. Docking - Paper I ..................................................................................................... 23 4.1.2. Differential scanning fluorimetry (DSF) - Paper II .................................................. 25 4.1.3. Crystallographic screening - The CrystalDirect Technology - Paper I and II .......... 26 4.1.4. Chemical synthesis of 8-oxoG analogs - Paper III ................................................... 29
4.2. Assessment of findings ................................................................................................... 30 4.2.1. Brief evaluation of the pipelines/approaches ........................................................... 30 4.2.2. We have obtained the first experimental structures of OGG1 in complex with an inhibitor .................................................................................................................................. 31 4.2.3. Several OGG1 inhibitors had a modest potentiating effect towards bleomycin - Paper II 32
4.3. Expanding the horizon for OGG1 as a drug target ......................................................... 33 5. CONCLUDING REMARKS AND FUTURE PERSPECTIVES ............................. 35
REFERENCES ................................................................................................................. 37
1
LIST OF PAPERS This thesis is based on the following papers, which will be referred to by their Roman
numbers.
I Identification of a small-molecule inhibitor of OGG1 by structure-based
methods.
Ytre-Arne ME, Cornaciu I, Hoffmann G, Blicher P, Jacobsen Ø, Leonard G,
Bowler MW, Bjørås M, Márquez JA, Dalhus B.
Manuscript
II Inhibitors of OGG1 identified by differential scanning fluorimetry and
crystallographic screening.
Ytre-Arne ME, Underhaug J, Hoffmann G, Cornaciu I, Wang W, Hajjar E,
Jacobsen Ø, Blicher P, Bowler MW, Martinez A, Márquez JA, Bjørås M, Dalhus
B.
Manuscript
III Synthetic Routes to N-9 Alkylated 8-Oxoguanines; Weak Inhibitors of the
Human DNA Glycosylase OGG1.
Mahajan TR, Ytre-Arne ME, Strøm-Andersen P, Dalhus B, Gundersen LL.
Molecules, 20, 15944-65 (2015)
2
ABBREVIATIONS
5-hmC 5-hydroxymethylcytosine
5-hmU 5-hydroxymethyluracil
5-ohU 5-hydroxyuracil
8-oxoG 8-oxoguanine
A Adenine
ADP Adenosine diphosphate
AP Apurinic/apyrimidinic
BCL-2 B-cell lymphoma 2
BER Base-excision repair
C Cytosine
CRIMS Crystallization information management system
DDR DNA damage response
DNA Deoxyribonucleic acid
dRP Deoxyribose phosphate
DSB Double-strand break
dsDNA Double-stranded DNA
DSF Differential scanning fluorimetry
EMBL European molecular biology laboratory
ESR1 Estrogen receptor 1
FaPy Formamidopyrimidine
G Guanine
Gh Guanidinohydantoin
HR Homologous repair
3
HTS High-throughput screening
Kd Dissociation constant
KO Knockout
LSD1 Lysine specific demethylase
MAL Methyl aminolevulinate
MBD4 Methyl-CpG-binding domain protein 4
Mm Molecular mass
MPG N-methylpurine DNA glycosylase
MUTYH MutY homolog DNA glycosylase
NEIL1/2/3 Endonuclease VIII-like 1/2/3
NER Nucleotide excision repair
NHEJ Non-homologous end joining
NMR Nuclear magnetic resonance
NTH1 Endonuclease III homolog 1
OGG1 8-oxoguanine glycosylase 1
PARP1 Poly [ADP-ribose] polymerase 1
PDB Protein data bank
PDT Photodynamic therapy
Phe Phenylalanine
Pol ! DNA polymerase !
PQS Potential G-quadruplex-forming sequence
RNA Ribonucleic acid
ROS Reactive oxygen species
SMUG1 Single-strand selective monofunctional uracil-DNA glycosylase
Sp Spiroiminohydantoin
SPR Surface plasmon resonance
4
SSB Single-strand break
ssDNA Single-stranded DNA
T Thymine
TDG Thymine-DNA glycosylase
Tm Melting temperature
Tyr Tyrosine
U Uracil
UNG Uracil N-glycosylase
UV Ultra violet
VEGF Vascular endothelial growth factor
WT Wildtype
5
SUMMARY Every single day each of our 1013 cells experience tens of thousands of DNA damage
events. If not repaired, DNA lesions can lead to mutations during replication, and give
rise to cancer, premature aging and neurodegenerative disease. Mammalian cells possess
a remarkable DNA repair machinery which maintains genome integrity. Nevertheless, if
the damage load exceeds the repair capacity, the cell will undergo apoptosis
(programmed cell death). This is utilized in the treatment of cancer. Ionizing radiation
and chemotherapy drugs such as cisplatin and bleomycin are just a few examples of anti-
cancer agents that exert their therapeutic cytotoxicity by causing DNA damage. During
treatment with a DNA damage-inducing agent, members of the DNA repair machinery
may act as resistance factors. Thus, inhibition of enzymes or pathways involved in DNA
repair should sensitize the cells towards the treatment.
The DNA repair enzyme 8-oxoguanine glycosylase 1 (OGG1) is one of 11 known human
DNA glycosylases. The DNA glycosylases catalyze the first step of the base excision
repair (BER) pathway, and OGG1 is the primary enzyme for recognition and removal of
8-oxoguanine (8-oxoG), which is a major mutagenic oxidative base lesion. 8-oxoG is
flipped into the lesion-specific pocket of OGG1, where the catalytic cleavage of the N-
glycosidic bond takes place. If not repaired, 8-oxoG can mispair with adenine during
replication and thus give rise to a G:C to T:A transversion mutation.
In this work we have applied a combination of screening methods, as well as a synthetic
chemistry approach, to identify putative inhibitors of OGG1. The screening methods
include molecular docking, differential scanning fluorimetry, biochemical assays and
crystallography. The emphasis has been on the development and application of automated
crystallographic screening, using the CrystalDirect technology at EMBL, Grenoble. We
have collected nearly 1,000 datasets and screened 60 ligands using this technology,
including both soaking and co-crystallization trials. This resulted in the first five known
structures of OGG1 in complex with a ligand. The structures of the OGG1/ligand
complexes reveal that the ligands bind in the lesion-specific pocket of OGG1 and mimic
several of the interactions that OGG1 makes with 8-oxoG. The ligands will thereby likely
6
prevent the correct positioning of 8-oxoG for the catalytic removal of the base to take
place. Indeed, three of these ligands have been confirmed as OGG1 inhibitors. The
structures of the OGG1/ligand complexes provide insight into interactions and binding
modes that could be utilized in development of potent and specific OGG1 inhibitors.
In addition to the ligands confirmed by crystallographic screening, we have identified
several OGG1 ligands and inhibitors with binding affinities and IC50 values in the low
!M range. The effect of five of the OGG1 inhibitors in combination with the
chemotherapy drugs cisplatin and bleomycin were also studied in cell viability assays.
Four inhibitors demonstrated modest potentiating effect of bleomycin-induced cell death
in a lung cancer cell line, and two of these inhibitors also sensitized a bone cancer cell
line towards the same drug.
The majority of OGG1 ligands and inhibitors presented in this thesis have low molecular
masses (<300 Da). Combined with relatively good affinity and inhibitory effect on
OGG1, several of them may serve as starting points for development of more potent and
specific OGG1 inhibitors.
A synthetic approach to design substrate-analog inhibitors of OGG1 was also applied.
Non-hydrolyzable analogs of 8-oxoguanine were synthesized in order to obtain molecules
that would inhibit OGG1 activity. Unfortunately, the synthesized molecules had only
modest inhibitory effect even at high concentrations.
Altogether the findings presented in this thesis provide a significant contribution to the
understanding of how small-molecules can bind to and inhibit the catalytic activity of
OGG1.
7
1. INTRODUCTION
1.1. DNA damage and repair
DNA is a surprisingly fragile molecule considering its importance for cell integrity. It is
estimated that a single human cell experiences tens of thousands DNA damage events per
day.1 The most common DNA lesion is caused by the spontaneous loss of nitrogen bases,
forming apurinic/apyrimidinic (AP) sites.2 These sites can further fragment
spontaneously giving rise to cytotoxic single-strand breaks (SSB). Deamination of
adenine, cytosine and guanine also occur frequently and results in bases with miscoding
properties.2 Moreover, normal cellular metabolism produces vast amounts of reactive
oxygen species (ROS), which can introduce oxidative DNA lesions.2–4 External sources
such as UV light, chemicals, tobacco smoke and radiation also introduce DNA damage.5,6
Some DNA lesions cause the inactivation of genes by blocking transcription, while others
can give rise to mutations during replication, and DNA damage can lead to aging, cancer
and/or neurodegenerative diseases.2,4,7 Maintenance of genome integrity is thus crucial
for our health and survival.
Upon sensing DNA damage, our cells can elicit a DNA damage response (DDR), which
activates a plethora of mechanisms to deal with the damage.6,8–10 For instance,
mammalian cells possess an extensive DNA repair machinery which counteract the
detrimental effects of DNA lesions, and the DDR leads to increased transcription of
enzymes involved in DNA repair and thereby up-regulation of DNA repair pathways. In
addition, cell cycle checkpoints are activated in order to stall the cell cycle, thereby
allowing time for repair of the lesions before replication of the DNA and cell division
take place. If the damage load is too severe for sufficient DNA repair to take place, the
cell may undergo apoptosis. Sometimes, the DDR can also activate damage tolerance
processes, such as translesion synthesis, as a temporary solution to surpass the stalled
replication machinery.6,8,10–13
The nature of the lesion determines which DNA repair pathway will be utilized.
Intrastrand cross-links and bulky lesions, such as pyrimidine dimers induced by UV light,
8
are repaired via the nucleotide excision repair (NER) pathway,14–17 whereas non-bulky
base lesions, such as deamination, oxidation and alkylation of bases, are repaired via the
base excision repair (BER) pathway.2,17,19–24 This pathway also repairs single-strand
breaks (SSB), while double-strand breaks (DSB) are repaired by the homologous repair
(HR) pathway during replication or the error-prone non-homologous end joining (NHEJ)
pathway.25,26
In the work presented here, human 8-oxoguanine DNA glycosylase 1 (OGG1) is the
target enzyme. BER in human cells is thus the most relevant DNA repair pathway and
will be described in more detail in the following section.
1.1.1. The BER pathway DNA damage arising from chemical modifications of single bases, such as oxidation,
alkylation or deamination, is repaired via the multi-step base-excision repair (BER)
pathway (Figure 1).2,17,19–24 The BER pathway is initiated by DNA glycosylases
recognizing and removing a chemically modified nitrogen base by cleaving the N-
glycosidic bond.27–30 The BER can proceed via a long-patch or a short-patch pathway.
Briefly, in long-patch BER a repair tract of two or more nucleotides is generated, whereas
in short-patch BER a single nucleotide repair tract is produced.31 Removal of 8-oxoG by
OGG1 has been demonstrated to mainly proceed via short-patch BER, thus only this
pathway will be discussed further in this thesis.32
The mechanism underlying the glycosylase activity depends on whether the glycosylase
is monofunctional (Figure 1a) or bifunctional (Figure 1b). Bifunctional DNA
glycosylases possess AP lyase activity in addition to the glycosylase activity.24,33,34 The
mechanisms for the downstream events of the BER pathway also depend on the nature of
the DNA glycosylase.
9
Simple outline of the BER pathway when initiated by a a) monofunctional or b) bifunctional Figure 1.glycosylase a) The monofunctional glycosylase cleaves the glycosidic bond by hydrolysis and leaves
an AP site that is further processed by APE 1 which cleaves the DNA backbone 5' of the lesion. In the
next step Pol ! utilizes its 5'-dRP lyase activity to create a single nucleotide gap before inserting the
correct nucleotide. Finally DNA ligase seals the nick in the backbone. b) The bifunctional glycosylase
forms a Schiff base intermediate when removing the damaged base. The intrinsic AP lyase activity of
the bifunctional glycosylase cleaves the DNA backbone 3' of the lesion site, resulting in a blocked 3'
end that must be processed by APE1 to yield a single nucleotide gap. Pol ! inserts the correct
nucleotide and DNA ligase seals the nick in the DNA backbone in the final steps of the BER pathway.
10
Briefly, a monofunctional DNA glycosylase cleaves the N-glycosidic bond by hydrolysis,
the damaged base is released and the resulting apurinic/apyrimidinic (AP) site is further
processed by AP endonuclease 1 (APE1) which cleaves the DNA backbone 5' to the AP
site.35,36 The resulting nick in the DNA backbone has a 3'-OH end and a 5'-dRP end.
DNA Polymerase ! (Pol !) possesses dRP lyase activity and is thus capable of removing
the 5'-dRP end and subsequently inserting a normal nucleotide in the vacant space using
the complementary strand as a template.37–41 Finally DNA ligase seals the nick in the last
step of the BER pathway.42,43
When a bifunctional glycosylase cleaves the N-glycosidic bond, a Schiff base
intermediate is formed.24 The enzyme then utilizes its AP lyase activity to cleave the
DNA backbone at the 3' side of the lesion forming an AP site with a blocked 3' end.33
This is a substrate for APE1 which cleaves the backbone on the 5' side of the lesion
leaving a single nucleotide gap. DNA Pol ! and DNA ligase inserts a normal nucleotide
and seals the backbone, respectively.39–43
To date, 11 human DNA glycosylases have been described. Six are classified as
monofunctional, four as bifunctional and OGG1 has a dual reaction mode acting both as a
mono- and bifunctional DNA glycosylase (Table 1). Each DNA glycosylase is specific
for one or several types of DNA lesions (Table 1). Only OGG1 will be described in
detail in this thesis.
11
Table 1 - The Human DNA Glycosylases
Protein Mechanistic class Substrates*
MPG Monofunctional Alkylated purines, ethenopurines
UNG Monofunctional U, 5-ohU and other uracil derivatives in ssDNA or
dsDNA
SMUG1 Monofunctional Many of the same substrates as UNG
TDG Monofunctional T,U or 5-hmU mispaired with G in dsDNA
MBD4 Monofunctional T,U or 5-hmU mispaired with G in CpG
dinucleotides
MUTYH Monofunctional A mispaired with 8-oxoG or FaPy-G
OGG1 Mono-/Bifunctional 8-oxoG and FaPy-G paired with C
NTH1 Bifunctional Oxidized pyrimidines and formamidopyrimidines
NEIL1 Bifunctional Sp, Gh, Oxidized pyrimidines and FaPys in
ssDNA and dsDNA
NEIL2 Bifunctional Sp, Gh, Oxidized pyrimidines and FaPys, prefers
ssDNA
NEIL3 Bifunctional Sp, Gh, Oxidized pyrimidines and FaPys in
ssDNA and dsDNA, weak glycosylase and AP
lyase activity
Based on Wallace, S. S. Base excision repair: A critical player in many games. DNA Repair 19, 14–26
(2014).44
*U = uracil, T = thymine, C = cytosine, G = guanine, A = adenine, ssDNA = single-strand DNA, dsDNA =
double-strand DNA, 5-ohU = 5-hydroxyuracil, 8-oxoG = 8-oxoguanine, 5-hmU = 5-hydroxymethyluracil,
5-hmC = 5 hydroxymethylcytosine, FaPy = formamidopyrimidine, Sp = spiroiminodihydantoin, Gh =
guanidinohydantoin.
12
1.2. Human 8-oxoguanine glycosylase 1 (OGG1)
Guanine has the lowest oxidation potential of the four nitrogen bases in DNA, and 8-
oxoguanine (8-oxoG) is the most frequent base lesion induced by ROS (Figure 2).2 Due
to its propensity to mispair with adenine during replication, this lesion is highly
mutagenic and can give rise to G:C to T:A transversion mutations if not repaired before
the cell divides.45–47 This is a common mutation in somatic cancers.48
Guanine and 8-Oxoguanine. The two bases differ at two positions. In 8-oxoG, an oxygen has Figure 2.replaced a hydrogen at C8, and a hydrogen has replaced a lone pair electron at N7.
Human 8-oxoguanine glycosylase 1 (OGG1) is the primary enzyme for recognition and
removal of oxidized guanine lesions in our DNA, specifically 8-oxoguanine base
pairing with cytosine, and ring-opened structures such as 2,6-diamino-4-hydroxy-5-
formamidopyrimidine (FaPyG).24,49–54
1.2.1. OGG1 structure and lesion recogntion OGG1 (Figure 3) belongs to the Helix-hairpin-Helix (HhH) structural superfamily, and
its HhH motif is followed by a Gly/Pro rich loop. In addition, OGG1 has an anti-parallel
!-sheet domain.55 Unlike most other DNA binding proteins, the cleft where OGG1
interacts with the DNA backbone is nearly charge-neutral, containing only a single
basic residue: His270, which hydrogen bonds with 8-oxoguanine's 5' phosphate.55,56
DNA binding proteins typically form salt bridges with the DNA but this does not apply
to OGG1. The enzyme is thought to interact with the DNA mainly through dipolar
electrostatic interactions, made up by the large number of !-helices oriented with their
positively charged N-terminal ends toward the negatively charged DNA (Figure 3).55
13
Structure of human OGG1 in complex with DNA containing 8-oxoG. The protein and DNA Figure 3.are shown in cartoon mode. ! helices are shown in blue, ! sheets are shown in green and loops are
shown in pink. The DNA backbone is shown in orange with bases indicated in yellow and blue sticks.
(PDB:1YQR)55
Another peculiar feature of OGG1 is that the enzyme interacts only with the sugar-
phosphate backbone of the 8-oxoG-containing strand, while making no contact to the
backbone of the complementary strand. The conserved HhH motif is thought to be
involved in positioning the DNA duplex for presentation of the lesion into the active site
pocket. Upon recognition of an oxidized guanine, OGG1 flips the damaged base out of
the helix and into its lesion specific pocket. Tyr203 wedges into the helix below the
estranged cytosine. This wedging aids in introducing a sharp kink in the DNA helix while
also unstacking the cytosine from its neighboring bases (Figure 4a). The cytosine is
coordinated by hydrogen bonds with Asn149, Arg154 and Arg204 (Figure 4a).55
When flipped into OGG1's lesion specific pocket, 8-oxoG's !-faces are sandwiched
between Phe319 and Cys253 (Figure 4b). The side chain carbonyl of Gln315 hydrogen
bonds with N1H and N2H of 8-oxoguanine, while the lesion's O6 atom is coordinated by
two tightly bound water molecules of which one is forming a hydrogen bonding bridge to
the side chain amide NH2 of Gln315. Gln315 is thus important for ‘reading’ and
interacting with the Watson-Crick base-pairing groups of 8-oxoG.
14
Figure 4. Close-up of key interactions between OGG1 and 8oxoG/C-containing DNA. a) Key Figure 4.interactions of the estranged cytosine (light red) with OGG1. Hydrogen bonds are shown as blue
dashes. The wedging residue Tyr203 is also depicted. 8-oxoG is shown in yellow (PDB:1EBM),55 b)
Key interactions of 8-oxoG (yellow) with central OGG1 residues for recognition, binding and
catalysis. Hydrogen bonds are represented by blue dashes, water molecules by cyan spheres
(PDB:1EBM).55
8-OxoG differs from guanine solely at two positions: an oxygen has replaced the
hydrogen at C8, and a hydrogen has replaced a lone electron pair at N7. All the
interactions between OGG1 and 8-oxoguanine mentioned above would also be possible
with a normal guanine. However, the enzyme discriminates between the two bases solely
by a single hydrogen bond from the carbonyl group of Gly42 to the N7 H of 8-
oxoguanine, an interaction that cannot be matched by guanine.55,56 In fact, undamaged
guanine does not appear to be fully flipped into the active site pocket but rather resides at
an exo site, about 5 Å away.56 This discrimination between the two bases is remarkable,
especially considering that the concentration of 8-oxoguanine is roughly one million-fold
lower than the undamaged base.56
15
1.2.2. OGG1's mechanism of catalysis While OGG1 traditionally has been classified as a bifunctional glycosylase with Lys249
implicated as the key residue for both glycosylase and AP lyase activity,57 several studies
have suggested that the enzyme mainly functions as a monofunctional glycosylase in
vivo.58–62 Specifically, in presence of free 8-oxoG under physiological concentrations of
Mg2+, the AP lyase activity of OGG1 is inhibited.61 A previously published study by our
group points at Asp268 as the catalytic residue for the glycosylase activity and further
supports that Lys249 is crucial for the specific recognition and positioning of 8-
oxoguanine for the hydrolysis reaction.56,62 The presence of Lys249's !-amino group in
the active site is needed for the recognition of 8-oxoG, and it is speculated that the
observed in vitro AP lyase activity is merely a result of this amino group's proximity to
the anomeric C1' in 8-oxoguanosine or the corresponding AP site.62
1.2.3. OGG1 phenotypes
The OGG1 gene is conserved in multiple species, and valuable information about OGG1
phenotypes has been gained by studying mice and the murine homolog Ogg1.
Mice deficient in Ogg1 are both viable and fertile.63 They appear healthy into adulthood
and are often considered as having no distinct phenotype. Nevertheless, mice lacking
Ogg1 do have a higher level of 8-oxoG lesions in their liver cells compared to their
wildtype counterparts, and this leads to an elevated spontaneous mutation rate, primarily
of G:C to T:A transversions.63,64 Compared to wildtype mice, Ogg1 knockout mice are
also more sensitive to UVB radiation, and as a consequence are more likely to develop
skin cancer.65
1.2.4. OGG1 is involved in epigenetic regulation of gene transcription
In the last few years, several studies have implicated a role for OGG1 in the epigenetic
regulation of gene transcription.66–68 While the presence of 8-oxoG in the DNA generally
has been regarded as detrimental to cellular processes, evidence now points at ROS-
induced oxidative DNA damage as a signal for gene activation.
A recent study shows that gene expression is enhanced when OGG1 initiates repair of 8-
oxoG in the promoter sequence of vascular endothelial growth factor (VEGF).68 VEGF
16
contains a guanine-rich, potential G-quadruplex-forming sequence (PQS) in its promoter.
The activity of OGG1 on 8-oxoG lesions in the PQS causes the formation of AP sites,
which enables melting of the duplex. This leads to unmasking of the G-quadruplex fold
which again leads to transcriptional activation of the downstream VEGF gene.68
OGG1 has previously been demonstrated to play a role in estrogen-induced expression of
genes involved in cell progression and apoptosis in the breast cancer cell line MCF7.66
Estrogens induce expression of genes by binding to estrogen receptors, which then bind
to estrogen responsive elements in gene promoters and trigger demethylation of the
histone tails within the nucleosomes by lysine specific demethylase 1 (LSD1). The
demethylation generates hydrogen peroxide which forms 8-oxoG lesions in the adjacent
DNA, and thereby the recruitement of OGG1 and Topoisomerase II! to the promoter.
This causes a conformational change in the chromatin that is required for the estrogen-
induced transcription of the downstream gene.66 The promoter of the studied gene, BCL-
2, contains a PQS in the region that is oxidized during the demethylation process, thus
one could speculate that the same mechanism as demonstrated for the VEGF promoter
applies here.68 Furthermore, PQS is found in promoter sequences throughout the genome,
and OGG1 could thus play a pivotal role in regulation of gene expression in response to
ROS.
Ogg1 has been reported to have a role in the modulation of anxiety-like behavior in mice
together with Mutyh.67 Behavioral studies show that the double knockout
(Ogg1-/-/Mutyh-/-) mice are less anxious than wildtype mice, but appear to have impaired
learning abilities. The cognitive phenotype is thought to be linked to the epigenetic
regulation of estrogen receptor 1 (ESR1), but in this case Ogg1 appears to repress gene
transcription. It is speculated that the observed increase in expression of ESR1 target
genes in the double mutant mice could be due to dysregulation of epigenetic and
transcriptional states, caused by the accumulation of 8-oxoG at promoter sites.67
OGG1 has also been implicated as a modulator of the inflammatory response, based on
the resistance of Ogg1 knockout mice to induced inflammation.69 This activity seems to
be linked to OGG1's role in the LSD1-dependent pathway described for the estrogen-
induced gene expression of BCL-2.66,70
17
1.2.5. OGG1 as a drug target for cancer treatment Considering that OGG1, in addition to its underlying DNA repair activity, is involved in
the modulation of various cellular processes, one might expect that targeting this enzyme
could have more implications than desired. Nevertheless, the normal lifespan and absence
of any severe phenotypes of the Ogg1 knockout mice supports that OGG1 could be a
promising drug target, especially for temporary treatment by adjuvants and synthetic
lethality approaches against cancer.
Cancer cells are fast dividing and many of the cancer therapies used in clinics today
target this characteristic. One way of doing so is by inducing DNA damage. Cancer cells
often have altered redox states and reduced DNA repair capacity.71,72 They are therefore
prone to accumulate DNA damage and thus likely to be more sensitive to the DNA
damaging agents than healthy cells. As mentioned above, our cells activate cell cycle
checkpoints in response to DNA damage and undergo apoptosis if the damage load is too
excessive for sufficient repair to take place. Ionizing radiation and several chemotherapy
drugs such as cisplatin, cyclophosphamide and bleomycin are just a few examples of
treatments that exert their therapeutic cytotoxicity by inducing DNA damage.73–78
During cancer treatment, OGG1 and other members of the DNA repair machinery can act
as resistance factors for the treatment. Moreover, if the tumor cells up-regulate the
targeted DNA repair pathway in response to the treatment, therapeutic resistance may
occur. Several studies have specifically shown that up-regulation of OGG1 contributes to
cellular resistance to cisplatin, and that suppression of OGG1 sensitizes cancer cells
towards bleomycin and ionizing radiation.75,79,80 Inhibition of OGG1 thus holds great
promise as an anti-cancer strategy particularly in combination with established anti-
cancer agents that induce an increase in ROS.
Only a few inhibitors of DNA glycosylases have been described in the literature so far.81–
85 The first inhibitors of OGG1 were reported in 2015 when Donley et al. presented 13
OGG1 inhibitors with potencies in the low !M range.85 With the exception of one, the
inhibitors were demonstrated to inhibit the bifunctional catalytic mechanism of OGG1,
namely the formation of a Schiff base intermediate.85 Since OGG1 appears to mainly
18
function as a monofunctional glycosylase in vivo, these inhibitors, or inhibitors based on
the same scaffold, might not be ideal for use in treatment.
1.3. PARP1 inhibitors
The BER pathway contains one member that is already well established as an anti-cancer
target, namely Poly(ADP-ribose) polymerase 1 (PARP1). PARP1 binds to SSBs in the
DNA and catalyzes, as the name implies, the formation of poly-ADP ribose.86,87 This is
thought to be a signal for recruitment of the DDR machinery to sites of DNA damage88.
PARP1 is involved in multiple DNA repair pathways including BER, but the full
function of this enzyme in DNA repair is not yet fully understood. Nevertheless, PARP1
inhibitors are a prime example of the principle of synthetic lethality. The breast cancer
genes BRCA1 and BRCA2 are tumor suppressor genes. Both BRCA1 and BRCA2 have
roles in double-strand break repair by HR, and BRCA1 is also involved in facilitating the
DNA damage response, including both checkpoint activation and multiple DNA repair
pathways.89 Tumor cells deficient in BRCA1 or BRCA2, are extremely sensitive to
PARP1-inhibitors.90–92
The first PARP1-inhibitor, olaparib, was launched in 2014 and was the first DNA repair
enzyme inhibitor to be marketed as a drug.93,94 Rucaparib is another PARP1-inhibitor
already in clinical use, and several others are in clinical trials.93–95
OGG1 has been shown to interact with PARP1 during oxidative stress and to stimulate
the poly(ADP-ribosyl)ation activity of PARP-1. Furthermore, PARP1 has been shown to
modulate OGG1 activity, and loss of OGG1 sensitizes cells to PARP-1 inhibitors.96
Exactly how the interaction between OGG1 and PARP1 takes place is unknown.
Nevertheless one could speculate that combining PARP1 inhibitors and OGG1-inhibitors
could have a synergistic effect that would prove beneficial in the treatment of BRCA-
mutant cancers.
19
2. AIMS The overall goal of this project was to identify inhibitors of OGG1 that could have
potential as adjuvants in cancer treatment. We applied two separate drug-screening
pipelines, as well as a synthetic chemistry approach in our search for OGG1 inhibitors.
Knowledge about how a hit/lead molecule interacts with its target molecule is key for any
rational ligand optimization. Therefore we sought to identify ligands of OGG1 by
crystallographic screening. At the beginning of this project no inhibitors of OGG1 had
been reported in the literature, and thus we wanted to sample a large variety of different
molecules. We applied two separate drug-screening pipelines. One where the
crystallographic screening was preceded by molecular docking (Paper I), and one
employing a biophysical DSF screening followed by further filtering of the initial hits by
biochemical assays. The prescreening allowed us to screen a larger number of molecules
than would have been possible with crystallographic screening alone.
In addition to the screening approaches, we wanted to investigate if non-hydrolyzable
substrate analogs of 8-oxoG could inhibit OGG1 activity. Various 9-alkyl-8-oxopurines
were synthesized from commercially available 2-amino-6-chloropurine and the
synthetically easily accessible compound O-carbamoylguanine.
In order to further characterize the ligands identified by the two screening pipelines, as
well as the synthesized substrate analogs, various methods for determining binding
affinity, thermostabilizing effects and inhibitory effect were applied. Finally, selected
OGG1 inhibitors were tested for their ability to potentiate chemotherapy drug-induced
cell death in various cancer cell lines.
20
3. ABSTRACTS OF PAPERS
Paper I: Identification of a small-molecule inhibitor of OGG1 by structure- based
methods.
The DNA glycosylases remove base lesions from our DNA as the first responders in the
Base Excision Repair (BER) pathway. Since ionizing radiation and certain chemotherapy
drugs used in the treatment of cancer and other conditions, exert their cytotoxic effect by
inducing DNA damage, the DNA glycosylases, as well as other members of the DNA
repair machinery, have the potential to act as treatment resistance factors. By inhibiting
these enzymes it is envisioned that cells will be sensitized, allowing administration of
lower doses, thereby reducing the severity of side-effects and/or improving clinical
outcomes for patients. We have used structure- based methods including virtual and
crystallographic screening to identify inhibitors of human 8-oxoguanine DNA
glycosylase 1 (OGG1) – a key enzyme for removal of oxidized base lesions from DNA.
Here, we present a series of novel ligands/inhibitors of OGG1 that share a common
scaffold consisting of an aminopyrimidine moiety connected via an amino bridge to a
substituted phenyl group. The molecules identified have binding affinities ranging from
1.7 µM to 0.43 mM. Furthermore, we present crystallographic X-ray structures of OGG1
in complex with three of these low- molecular weight ligands. These show that the
ligands bind in various conformations and orientations in the lesion specific pocket of the
enzyme, where they interact with several residues key to the recognition and removal of
8-oxoguanine. One of the ligands, denoted NCI-6, inhibits the glycosylase activity of
OGG1 with an IC50 value of 78 µM. The series of OGG1 ligands presented here could
serve as interesting scaffolds for the development of more potent and specific OGG1
inhibitors for use in cancer treatment.
Paper II:Inhibitors of OGG1 identified by differential scanning fluorimetry and
crystallographic screening.
Ionizing radiation and a range of chemotherapy drugs used in cancer treatment induce
21
DNA damage in order to exert their therapeutic cytotoxicity. In this respect, the members
of our cells’ DNA repair machinery serve as resistance factors to the treatment. Inhibitors
of DNA repair enzymes may thus have clinical potential as adjuvants potentiating the
effect of the primary cancer therapy. In this study we focus on the enzyme 8-oxoguanine
DNA glycosylase 1 (OGG1), which recognizes and removes oxidized guanines as the
first step of the base excision repair (BER) pathway. We aimed to identify inhibitors of
OGG1 using a combination of biophysical, biochemical and crystallographic screening
methods. Here we present five novel inhibitors of OGG1, several of which have both Kd
and IC50-values in the low µM range. The inhibitors were tested for their ability to
increase the cytotoxicity of two established cancer drugs in a cell viability assay, and four
inhibitors potentiated bleomycin-induced cell death in a human lung carcinoma cell line.
Moreover we have obtained experimental crystal structures of OGG1 in complex with
two of the inhibitors. Both inhibitors bind in the active site lesion-specific pocket of
OGG1, and thereby prevent correct positioning of 8-oxoguanine for the glycosylase
activity of the enzyme.
Paper III: Synthetic Routes to N-9 Alkylated 8-Oxoguanines; Weak Inhibitors of the
Human DNA Glycosylase OGG1.
The human 8-oxoguanine DNA glycosylase OGG1 is involved in base excision repair
(BER), one of several DNA repair mechanisms that may counteract the effects of chemo-
and radiation therapy for the treatment of cancer. We envisage that potent inhibitors of
OGG1 may be found among the 9-alkyl-8-oxoguanines. Thus we explored synthetic
routes to 8-oxoguanines and examined these as OGG1 inhibitors. The best reaction
sequence started from 6-chloroguanine and involved N-9 alkylation, C-8 bromination,
and finally simultaneous hydrolysis of both halides. Bromination before N-alkylation
should only be considered when the N-substituent is not compatible with bromination
conditions. The 8-oxoguanines were found to be weak inhibitors of OGG1. 6-Chloro-8-
oxopurines, byproducts in the hydrolysis of 2,6-halopurines, turned out to be slightly
better inhibitors than the corresponding 8-oxoguanines.
22
4. DISCUSSION
4.1. Drug Discovery
The development of a new drug is a challenging, lengthy and costly process. In order to
reach a lead compound for clinical trials as efficiently as possible, a continous
development and improvement of the screening methods is needed.97 In this project we
have used a combination of screening methods including docking (Paper I), differential
scanning fluorimetry (Paper II) and crystallographic screening (Paper I and II). In
addition, we have utilized a synthetic chemistry approach (Paper III). In this section I
will briefly present the methods we have utilized and evaluated for screening for ligands
of OGG1.
Before drug screening was an option, synthetic organic chemistry had a leading role in
drug development.98 In paper III, the synthetic approach was used to synthesize analogs
of 8-oxoguanine based on a hypothesis that compounds mimicking the natural substrate
might inhibit the activity of OGG1.
In the early 1990s, experimental high-throughput screening (HTS) of compound libraries
was established as the main screening method applied by the pharmaceutical
companies.99–101 During a HTS approach, libraries containing 100,000s to millions of
drug-like molecules ranging in molecular mass (Mm) from 250 to 600 Da are screened.
This method is costly and while the potency of the hit molecules is typically below 30
!M, the attrition rate of compounds when going from hit to lead is high.102 The method is
still in use, but during the last decades several alternatives for drug screening have
emerged, some of which have great advantages over the classical high-throughput drug
screening.
For instance, during the last two decades, fragment-based lead discovery has been
established as a widely used method for drug discovery.101 The term 'fragments' is here
used to describe low molecular weight compounds, less than 250-300 Da, and the idea is
that screening of smaller molecules allows a more efficient sampling of chemical space.
Using this fragment-based approach, the emphasis is based on ligand efficiency rather
23
than ligand potency.101–103 Hits from a fragment screening typically have affinities in mM
down to 30 !M, but they possess several advantages over the typical HTS hits when it
comes to developmental potential.102 The contribution of each atom in the fragment to the
binding efficiency is typically much higher for a fragment hit than for a HTS hit.101–
103Also, since the molecular mass of a hit compound almost inevitably increases during
lead optimization, the smaller size of a fragment is beneficial by allowing a size increase
without necessarily compromising typical drug-like properties. Also, identifying several
fragments binding in adjacent sites in the target protein allows for design of highly potent
ligands through fragment joining or fragment extension. Due to the fragments' lower
binding affinities, biophysical methods such as X-ray crystallography and nuclear
magnetic resonance (NMR) spectroscopy are particularly applicable for fragment-based
screening approaches, as these methods can detect low affinity-ligands.101–103 Moreover,
the ligand-target complex structures obtained by these methods are useful for guiding the
lead optimization process.101,102 Surface plasmon resonance (SPR), microscale
thermophoresis (MST) and differential scanning flurimetry (DSF) can also be applied for
fragment-based screening.101
Several of the OGG1 ligands presented in this work can be classified as fragments. With
the exception of TFA23 and TFA37 presented in Paper II, all the ligands regarded as hits
in this work have molecular masses below 300 Da.
4.1.1. Docking - Paper I
In paper I, docking was used as the primary screening method. Docking is an inexpensive
and efficient screening method that allows one to screen millions of compounds if
desired. The only pre-requisite is a 3D-structure of the target protein, preferably
determined by X-ray crystallography or NMR.105 The docking programs are getting more
and more advanced and accurate, and with substantial knowledge about the target
molecule, it can be an asset to any drug discovery process. Nevertheless, docking is still
an in silico experiment and the results will depend on the scoring function(s) of the
program used as well as the many choices one has to make both in preparing your target
molecule, when selecting docking parameters, and, if applicable, the manual inspection
and selection of hits to include in further experiments.
24
When preparing our target molecule structure (the apo structure of OGG1, PDB:1ko9),
we removed all water molecules from the structure, added hydrogens and performed a
restrained energy minimization. The docking itself was performed using the extra-
precision mode in Glide (Schrödinger). Based on comparison of free and DNA-bound
crystal structures of OGG1, OGG1 does not appear to undergo any substantial
rearrangement when binding DNA. We therefore decided to treat OGG1 as a rigid
receptor during the docking.
A compound library that contained 1596 compounds was screened. This might seem
small for a virtual screening approach. However, the library was designed in a way that
included a wide range of pharmacophores. Also, the molecules range in size from 120 to
700 Da, but have a mean Mm of 271 Da. ~1400 of the compounds have a Mm of <300
Da, hence the majority of the compounds can be classified as fragments. Thus, the
chemical space sampled may not be so small considering the size of the molecules
screened. Finally, the compounds in this library are available in mg amounts free of
charge from NCI.
During the manual inspection process, we prioritized molecules that we considered to
have drug-like properties, such as comprising an appropriate number of hydrogen bond
donors and acceptors. We discarded some high-scoring hits containing features such as
complex fused ring structures, polysaccharide derivatives and highly symmetric
molecules, among others. Smaller molecules were also preferred over larger ones. We
were quite conservative when considering the possible interactions between the docking
hits and OGG1. We prioritized the fulfillment of hydrogen bond interactions for
hydrogen bond donors and acceptors, and the presence of hydrophobic interactions rather
than seemingly less obvious contributions to binding affinity such as halogen bonds. As
in any selection process we might have omitted some ligands for further testing that could
in fact be high affinity binders and potent inhibitors of OGG1. Moreover, considering
that several of our experimental structures of OGG1/ligand complexes revealed
interactions via water molecules, it would be interesting to perform a docking where the
most conserved water molecules in the active site are included.
25
The docking model of the inhibitor identified by the virtual screening, NCI-6, in complex
with OGG1 positions the ligand in roughly the same region as confirmed by our
experimental structure. However the docked binding mode of NCI-6 is upside-down
compared to our experimental structure, and so the predicted interactions differ from the
ones revealed in the experimental structure. Moreover, some of the residues in the active
site of the docking model are oriented differently than in the experimental structure with
NCI-6. Our experimental structure reveals that one of these residues, Asn149, is involved
in a halogen bond interaction with NCI-6. Perhaps allowing some flexibility in the
docking site could improve the hit rate from the virtual screening.
4.1.2. Differential scanning fluorimetry (DSF) - Paper II Ligand binding can affect the thermal stability of a protein. This is utilized in Differential
Scanning Fluorimetry (DSF) where the apparent melting temperature (Tm) of a protein is
monitored during increasing temperature by using a special dye whose signal is quenched
in water, but that elicits fluorescence when binding to hydrophobic residues. As the
protein structure is transforming via the molten globule state towards full denaturation by
the increasing temperature, the hydrophobic residues usually residing on the inside of the
protein are exposed so that the dye can bind and emit its fluorescence. The Tm is
calculated from the inflection point of the melting curve, and molecules causing an
increase in Tm relative to the free protein are presumed to bind and stabilize the protein.
The Tm of a protein is defined as the temperature point where 50% of the protein is
denatured and 50% is in a native state. The method does not require any former
knowledge about the structure (or even the function) of the target, and DSF can be used
to screen for any conditions stabilizing (or destabilizing) your protein of interest. This can
be used to gain information about ideal buffer conditions or crystallization conditions or
to search for ligands for a target molecule.106,107
DSF was used as the primary screening method in Paper II. Our collaborators at the
University of Bergen performed the high-throughput screening of OGG1 using the
MyriaScreen Diversity Library from Sigma Aldrich. This compound library consists of
10 000 compounds with drug-like properties ranging in Mm from 120 to 500 Da, and the
library is designed to contain molecules covering a large structural diversity. All
26
compounds were tested at a concentration of 2 mg/mL in the initial DSF screening,
making the test concentration of the smaller compounds higher than for the larger
compounds. This may have introduced a bias favoring the smaller molecules as hits, but
at the same time "balanced" the molecules' chance of being regarded as hits considering
the expected lower binding affinity (and presumably thermostabilizing effect) of smaller
molecules compared to larger molecules.
DSF is inexpensive and relatively quick to set up and run, allowing the screening of a
substantial number of molecules experimentally in short time. The hits from the initial
screening were validated by concentration dependent stabilization in order to omit false
positives. The results from the concentration dependent stabilization were also used to
calculate Kd values for the ligands. The DSF screening discovered 136 OGG1 ligands
from a selection of 10,000 molecules. However, since DSF does not give any information
about where a hit from the screening binds to the target protein, the recorded "binders"
could in principle bind anywhere on the target molecule. Therefore it was not surprising
that not all 136 hits turned out to be OGG1 inhibitors. Nevertheless, DSF provided a
valuable set of molecules experimentally demonstrated to bind to our target molecule, and
did indeed result in the experimental structure of two OGG1/ligand complexes and three
inhibitors with Kd values in the low !M range.
4.1.3. Crystallographic screening - The CrystalDirect Technology - Paper I and II When going from hit to lead or from lead to drug, an experimental structure of the
hit/lead molecule in complex with the target molecule is invaluable in guiding the further
development towards a potent and specific drug candidate. Crystallographic screening is a
labor intensive, time consuming process but during the last decade there has been a
tremendous development in technology that has reduced the workload, risk of crystal
breaking, and the time from start to finish considerably. With the development of cryst-
allization robots and pipetting machines the time and effort needed for the first steps of a
crystallographic trial (establishing crystallization conditions and setting up crystallization
plates) were considerably reduced. Furthermore, only a decade ago, it typically took 1,5-2
hours to collect a single dataset at a synchrotron beamline. It was sufficient to take simple
27
notes to keep track of the crystals and datasets. During the last years the introduction of
fast pixel array detectors at beamlines, and the launch of automated data collection and
processing have also shortened the time needed for data collection and processing
immensely and in 2013, the time required to collect a diffraction dataset had been much
shortened. With the continuing increasing capacity of beamlines, as well as the
introduction of highly automated beamlines, it was not long until data from hundreds of
crystals could be collected during a 24-hours shift. This called for a more sophisticated
way to keep track of all the crystals and datasets.
In addition, when we started this project in 2013, the harvesting and cryo-protection of
crystals still had to be done manually in most academic labs, and had thus become the
main rate limiting step in the processing of crystals. The handling of crystals during
harvesting is a delicate, time consuming process that both requires skill and exposes the
crystals to mechanical stress. Furthermore, since the crystallization solutions contain
water, the addition of a cryo-protectant to the crystal-containing drops before flash-
freezing in liquid nitrogen is crucial, creating both an additional workload and
introducing yet another critical crystal handling step for the fragile protein crystals.
The crystallographic screening part of this project has been a pilot project for the
development and implementation of the CrystalDirect technology at EMBL,
Grenoble.108,109 The CrystalDirect technology utilizes specially designed crystallization
plates to automate the crystal harvesting step. The CrystalDirect plates are 96-well plates
with a thin, low X-ray background film as the bottom. This thin film can be cut by laser
photoablation during harvesting and this feature is also convenient for ligand delivery,
and cryo-protection.
The crystallographic screening presented in Paper I and Paper II contributed to the
development of an automated method for ligand delivery by an in situ soaking approach.
Instead of fishing crystals and transferring them to a ligand solution drop for soaking, the
ligand solution is added to the crystal drop of interest and the soaking takes place by
diffusion. This saves the crystals of the mechanical stress of transfer. Moreover, diffusion
is a more gentle way to expose the crystals to the ligands/organic solvents, and thereby
allows higher ligand concentrations to be used in soaking without risking crystals
28
cracking and breaking or dissolving. The details for how this was done differed for the
NCI-ligands (Paper I) and the TFA-ligands (Paper II). For Paper I the addition of ligand
solution was carried out by first using the laser of the harvesting robot prototype to drill a
small hole in the film under the crystal drop of interest. We mounted a 0.5 !L precision
syringe (Hamilton) on a micromanipulator of the harvesting robot prototype to position
the syringe to the tiny crystallization drops, and then added the ligand manually using the
syringe.For Paper II the Cartesian PixSys crystallization robot (Cartesian Technologies)
was programmed to deliver the ligand solutions (50 nL) on top of the crystallization
drops in a fully automated mode.
The CrystalDirect technology also offers the possibility to use liquid-removal by
aspiration as a cryo-protectant, thereby eliminating the extra workload of formulating
special cryo-solutions for different crystallization solutions used. Moreover the
harvesting of crystals by laser photoablation also facilitates harvesting of small crystals
that would be difficult to fetch with the classical "fishing" approach. All our crystal
harvesting experiments at EMBL utilized liquid removal by aspiration, and the harvested
crystals were immediately transferred by the robot into a gaseous stream of 100 K cold
nitrogen before manual transfer to pucks in liquid nitrogen.
In addition this project contributed data and feedback for the further development of the
Crystallization Information Management System (CRIMS) at EMBL, Grenoble. In 2014,
we could use CRIMS to track our crystals from crystal setup, via growth in the
crystallization hotel, to harvest. Then we could manually name our samples and create
files that we could transfer to the data collection software at the beamline when we
assigned the pucks to the sample changer, and in this way the datasets were automatically
named correctly.
All the methods applied on the OGG1 crystals have contributed to the development of the
CrystalDirect Automatic Crystal Harvester from Arinax (https://www.arinax.com). The
harvester spends about one minute per harvest and has a success rate of 99%. The method
for soaking of crystals is now an automated variant of the approach used in Paper I: the
CrystalDirect harvesting robot makes a small cut in the plate membrane using its laser,
then places a drop of liquid to be soaked in on the aperture and the liquid then diffuses
29
through the small opening into the crystal drop. In addition, a range of features have been
implemented since the OGG1 project was part of the testing material - including the
ability to transfer the cryo-cooled samples into a dewar by a sample-changer robot.
CRIMS can now automatically name and track crystals all the way from setup to
beamline. One can use CRIMS to select which crystals to harvest, as well as cut shape,
and then CRIMS will transfer the information to the CrystalDirect Automatic Crystal
Harvester, which will perform the harvesting job for you while CRIMS will keep track of
all the events. In general, most of the features we used are still being utilized, but the
interspersed manual operations have been replaced by automatic procedures, which was
the ultimate goal from the beginning. Briefly summarized, this technology makes it
possible to efficiently screen a large number of compounds using X-ray crystallography.
We have collected nearly 1000 datasets for this project and screened 60 ligands by
crystallography, including both soaking and co-crystallization trials. The co-
crystallization approach did not result in any protein/ligand structures, while soaking of
preformed crystals resulted in the experimental structure of five novel OGG1/ligand
complexes. Since the OGG1 crystals were quite densely packed with relatively small
water channels available for diffusion, large ligands may not have been possible to soak
into the crystals even with long soaking times. Despite repeated efforts, we did not obtain
complexes of all our identified ligands/inhibitors. In particular, the lack of a crystal
structure of the potent OGG1 inhibitor TFA-7 in complex with OGG1 is puzzling since
the small size of this ligand should not be an obstacle in soaking. Unfortunately, no co-
crystallization experiments for TFA-7 succeeded either, despite numerous attempts.
4.1.4. Chemical synthesis of 8-oxoG analogs - Paper III
For paper III, we chose a more classical approach to drug discovery. Instead of screening,
we applied synthetic chemistry to synthesize potential inhibitors of OGG1. The
hypothesis was that non-hydrolyzable alkylated analogs of 8-oxoG might inhibit enzyme
activity. The molecules synthesized contained either a Cl or a OH-group at C6 (X-group
in Figure 5) and various alkylated groups (CH2-c-hexyl, c-hexyl, c-pentyl, or c-pent-2-
enyl) at N9 (R-group in Figure 5).
30
General structure of the compounds synthesized for Paper III. Figure 5.
Even at high concentration (500 mM), the compounds had only a modest effect on the
activity of OGG1. The chlorinated compounds appeared to be better inhibitors of OGG1
than their 6-oxo counterparts, with two of the compounds (6b and 6c) inhibiting the
glycosylase activity by about 30% at this concentration. To study the specificity of these
compounds, they were also tested against the DNA glycosylase NTH1, another
glycosylase that belongs to the same structural family as OGG1 and recognizes and
removes oxidized pyrimidines in DNA. Compound 6b also reduced the cleavage activity
of NTH1 by 30%, whereas compound 6c appeared to be more specific towards OGG1.
4.2. Assessment of findings
4.2.1. Brief evaluation of the pipelines/approaches Three different, but somewhat overlapping, approaches to identify and confirm inhibitors
or ligands of OGG1 were explored in this work.
While the screening pipeline (Docking ! crystallographic screening) described in Paper I
initially only resulted in a single OGG1 inhibitor, NCI-6, the search for and following
assessment of the NCI-6 analogs allowed us to gain a better understanding of which
features of these molecules that were important for binding and possibly inhibiting
OGG1. We have yet to obtain activity data for the analogs, and this should be prioritized
in the future as some of these analogs may compete with NCI-6 as the best OGG1
inhibitor from this ligand series.
The screening pipeline for Paper II (DSF ! activity assays ! crystallographic screening)
was in many ways the most successful in this project, and the path pursued more
intensively. We obtained the experimental structure of two ligands in complex with
OGG1 (TFA15 and TFA19), and three additional compounds (TFA7, TFA23 and
N
N N
HN
H2NR
O
X
31
TFA37) were also studied further as they demonstrated good inhibitory properties of
OGG1's enzyme activity. Cell based assays showed that four of the five inhibitors
sensitized cancer cell lines to bleomycin-induced cell death, and thus may have a
potential for further development.
Due to the low inhibitory effect of the molecules synthesized in Paper III, we did not do
any follow up work on these inhibitors. In retrospect it would have been interesting to at
least obtain Kd values for the compounds in order to compare them with the other OGG1
ligands identified. The synthetic approach was the least successful of the three paths
chosen for this project. However, synthetic chemistry would be crucial for lead
optimization of the established OGG1 inhibitors and ligands.
4.2.2. We have obtained the first experimental structures of OGG1 in complex with an inhibitor To date, 13 potent inhibitors of OGG1 have been described in the literature.85 However,
no experimental structure of OGG1 in complex with a ligand/inhibitor has been
published, hence the binding modes and mechanisms of inhibition of these inhibitors are
unknown. Furthermore, none of the inhibitors are potent enough to be used as a drug,
which necessitates further development of the inhibitors. Without knowledge about how
the inhibitors bind, any rational ligand expansion is challenging. If structures of smaller
molecules binding to the target molecule are available, they might be used to guide the
design of more potent and specific inhibitors, for instance by using tools such as
OpenGrowth.110
In this thesis the experimental structures of OGG1 in complex with five novel ligands is
presented, and these complexes contribute to the understanding of desirable or beneficial
features of an OGG1 inhibitor.
Our OGG1/ligand complexes revealed a high degree of repeating and overlapping
interactions between the flipped base in the original 8-oxoG-containing DNA substrate or
ligands and target enzyme. In nearly all our ligand-enzyme structures, the crucial
hydrogen bond from an amino group of the ligand to the carbonyl group of Gly42 is
32
present. This carbonyl group is the key protein signature that enables OGG1 to
discriminate between undamaged and oxidized guanine.55,56 The presence of a !-stacking
interactions with Phe319 in all our structures also indicates that this interaction
contributes substantially to binding stabilization. In addition, several other OGG1/8-oxoG
interactions are found in our OGG1/ligand structures, including water-mediated hydrogen
bonds. If an OGG1 inhibitor is to compete with 8-oxoG for the binding site, it needs to
bind with higher affinity. The fact that our ligands already fulfill some of the interactions
8-oxoG participates in, is promising for the further development into potent OGG1
inhibitors.
4.2.3. Several OGG1 inhibitors had a modest potentiating effect towards bleomycin - Paper II The OGG1 inhibitors discovered and analyzed in Paper II were tested in cell based assays
in combination with the chemotherapy drugs cisplatin and bleomycin. The drugs and cell
lines tested (A549 human lung carcinoma, U2OS human osteosarcoma and T98G human
glioblastoma) were chosen based on a combination of published data indicating a
promising effect of inhibition of OGG1, and in-house availability.75,80 This resulted in the
combination of some drugs/cancer types that might not be relevant for clinical use.
We could observe a modest effect of the OGG1 inhibitors in potentiating bleomycin-
induced cell death in the lung carcinoma and osteosarcoma cells. We did not check the
expression of OGG1 in the different cell types before conducting the cell survival assays.
Although the level of OGG1 varies from cell type to cell type, OGG1 seems to be
expressed in all cancers and thus makes it reasonable to presume that our target enzyme
was expressed in the cell lines tested.111,112 We have yet to check the specificity of our
inhibitors for OGG1 versus other DNA glycosylases. Further, we cannot exclude that the
compounds may have other targets in the cells besides OGG1.
In retrospect, considering the large variation in the sensitivity of cells towards different
types of treatment, testing the combination of inhibitor and cancer drug in other cell
types, and with other anti-cancer agents, might have revealed a stronger potentiating
effect of the OGG1 inhibitors. One particularly interesting approach could be in
combination with topical photodynamic therapy (PDT) used in treatment of non-
33
melanoma skin cancers.113 The drug Metvix contains a photosensitizing agent (methyl
aminolevulinate - MAL) and is used for PDT. MAL is a prodrug that is metabolized into
phototoxic compounds inside cells. Upon light-activation these compounds lead to the
production of ROS, which again generate oxidative damage to the cell.113 Thus OGG1
inhibitors should be especially pertinent as adjuvants for PDT-treatment.
4.3. Expanding the horizon for OGG1 as a drug target
When we first started the quest to identify inhibitors of OGG1 the primary goal was to
find inhibitors of OGG1 that could be used in combination with other cancer drugs to
sensitize and aid in killing cancer cells. However, OGG1 may have more potential as a
drug target than first anticipated. In light of OGG1's importance for cognitive function,
involvement in epigenetic regulation of a range of targets, and observed pre-disposition
to some cancer types in KO animals,65–68 temporary inhibition or stimulation of OGG1
may be fruitful approaches for the treatment of a range of conditions or diseases.
Given the role of OGG1 as an activator of transcription of VEGF and BCL-2,66,68
administration of an OGG1 inhibitor during cancer treatment could in fact serve a dual
effect on tumor cell growth and survival. Angiogenesis stimulated by the expression of
VEGF is a prerequisite for tumor growth and BCL-2 is an anti-apoptotic protein.66,114
Thus, OGG1 inhibitors may both sensitize the cancer cells to the conventional treatment
(chemotherapy or radiation) and slow down tumor growth and induce apoptosis by
repressing transcription of VEGF and BCL-2.
During an inflammatory response, ROS production is increased and OGG1 is
activated.115 As briefly mentioned in the introduction, studies on mice have shown that
disruption of Ogg1 reduces the inflammatory response in endotoxic shock, type I diabetes
and contact hypersensitivity.69 In addition, Ogg1 deficiency down-regulates an allergen-
induced airway inflammatory response in mice.116 These results suggest that OGG1
inhibitors have potential as anti-inflammatory agents.
While this project has focused on identifying OGG1 inhibitors, stimulation of OGG1 in a
therapeutic context should not be ignored. Ogg1 has proven to protect neurons against
34
oxidative DNA damage and cell death under ischemic conditions in mice117, suggesting
that stimulation of OGG1 shortly after a stroke could the improve clinical outcome. In
addition to increased repair of 8-oxoG, stimulation of OGG1 may increase the
transcription of VEGF and BCL-2, both of which codes for gene products that would be
beneficial for recovery after a stroke. Moreover, another mice study revealed that the
Ogg1 knockout mice have higher glycogen content in the liver and display
hyperglycemia and elevated insulin levels compared to wildtype mice. This indicates that
OGG1 is involved in repressing gluconeogenesis in liver cells, and stimulation of OGG1
may thus be utilized to treat diabetes type II.118 The involvement of OGG1 in the
regulation of cognitive function could also be targeted.67 The behavioral studies by
Bjørge et al. suggest that Ogg1 has a dominant and antagonistic effect to Mutyh,
regarding learning abilities. The formation of memories and learning is probably a fine-
tuned process, however one could speculate if the stimulation and stabilization of OGG1
could improve learning and memory formation.
Drugs that exert their action by stimulation typically target receptors. However, in
principle it should be possible to stimulate the activity of an enzyme as well, for instance
by administering a ligand or cofactor that stimulates enzyme activity directly, or
indirectly by stabilizing the enzyme thereby prolonging its lifetime and/or improving the
catalytic performance.
35
5. CONCLUDING REMARKS AND FUTURE PERSPECTIVES
We have identified several OGG1 inhibitors and ligands that have the potential to be
optimized into potent and specific inhibitors of OGG1. We have yet to test the specificity
of our ligands and inhibitors towards other DNA glycosylases, but given the strict
substrate recognition of the different glycosylases, it should be possible to design potent
and selective inhibitors to target OGG1 specifically. The most promising of our ligands
for OGG1 inhibition are quite small (<280 Da) and should thus have plenty of
possibilities for expansion without compromising typical drug-like properties, as
described by Lipinski's rule of five; a Mm less than 500 Da, logP below 5, less than 5
hydrogen bond donors, less than 10 hydrogen bond acceptors and less than 10 rotatable
bonds.119
The identified OGG1 ligands are quite diverse, both in structure and size, but they all
possess at least one ring system, being either a substituted phenyl ring or a purine analog,
or combinations of these and other ring systems. Our ligands also typically contain
hydroxy-, methyl and/or amino groups, and several of the molecules identified as
potential OGG1 inhibitors also contain one or more halogen atoms.
Since we lack activity data for the NCI-6 analogs from Paper I, we cannot refer to them
as inhibitors. A good Kd value does not guarantee an inhibitory effect and vice versa, as
proven by the discrepancy between Kd and IC50 values for one of the ligands in Paper II.
Moreover, where on the target molecule the ligand binds is crucial in order to affect the
activity of an enzyme. This can be revealed by an experimental structure, like the ones
we have obtained for several of the ligands. Based on the data collected so far, NCI-6 is
the most obvious lead candidate from Paper I, however several of the NCI-analogs could
also be interesting to study further, including NCI-606, NCI-608, NCI-609 and NCI-610.
The fact that they differ so little structurally and show similar binding affinities and
thermostabilizing effects could indicate that they also bind in the lesion-specific pocket.
From paper II, TFA7 and TFA19 stand out as the most interesting lead candidates to
pursue further. TFA7 has the lowest Kd (1 !M) and IC50 (0.5 !M) values of all our
36
OGG1 ligands, and was also shown to sensitize A549 lung cancer cells towards
bleomycin. However, we have not been able to obtain the structure of the OGG1/TFA7
complex yet and its SwissAME profile gives a PAINS (Pan-assay interference
compound) alert, meaning that caution must be taken if deciding to pursue this ligand
further. Its small size and low synthetic accessibility score, combined with good binding
affinity and inhibitory properties supports including this ligand in a lead optimization
process. TFA19 also holds great promise for further development, as demonstrated by its
inhibitory effect on OGG1, low !M affinity and ability to sensitize cancer cell lines to
bleomycin. The experimental structure of the OGG1/TFA19 complex structure will of
course be an asset for further ligand growth, and the low synthetic accessibility score
indicates that this ligand is particularly pertinent for optimization by synthetic chemistry
approaches.
By combining methods for screening and evaluating ligand or inhibitor potential, we
have gained broad knowledge about our ligands and inhibitors. As we also experienced,
not all methods can be applied for all molecules, making the scope of methods available
even more crucial. Now that screening by crystallography can be carried out much more
efficiently, it would be interesting to screen a larger number of compounds by this
method. However, a simple and fast pre-screening, such as DSF used in the study in
Paper II, should probably be maintained to reduce cost and workload. If docking is used
as a pre-screening, one could imagine skipping the manual selection step and rather
include the top 100-200 ranked hits in the following X-ray crystallographic screening.
This may result in the discovery of less obvious ligands that can have great potential for
further development into drugs. Careful preparation of the target molecule would be
crucial for successful docking, as well as the selection of parameters for the docking
itself. DSF could be applied either before or after the crystallographic screening, to
identify ligands that stabilized the target molecule. This could be utilized both for the
development of potential inhibitors, but also for the stabilization aspect itself if
identifying stimulators of enzyme activity. The fact that this method also can be used to
establish Kd values is of course advantageous. However, as mentioned above, keeping a
range of methods available will be an asset to every drug discovery program.
37
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Molecules 2015, 20, 15944-15965; doi:10.3390/molecules200915944
molecules ISSN 1420-3049
www.mdpi.com/journal/molecules Article
Synthetic Routes to N-9 Alkylated 8-Oxoguanines; Weak Inhibitors of the Human DNA Glycosylase OGG1
Tushar R. Mahajan 1, Mari Eknes Ytre-Arne 2,3, Pernille Strøm-Andersen 3, Bjørn Dalhus 2,3 and Lise-Lotte Gundersen 1,*
1 Department of Chemistry, University of Oslo, P. O. Box 1033, Blindern, N-0315 Oslo, Norway; E-Mail: [email protected]
2 Department of Microbiology, Oslo University Hospital, P. O. Box 4950, Nydalen, N-0424 Oslo, Norway; E-Mails: [email protected] (M.E.Y.-A.); [email protected] (B.D.)
3 Department of Medical Biochemistry, Institute of Clinical Medicine, University of Oslo, P. O. Box 4950, Nydalen, N-0424 Oslo, Norway; E-Mail: [email protected]
* Author to whom correspondence should be addressed; E-Mail: [email protected]; Tel.: +47-228-570-19.
Academic Editor: Roman Dembinski
Received: 2 June 2015 / Accepted: 26 August 2015 / Published: 2 September 2015
Abstract: The human 8-oxoguanine DNA glycosylase OGG1 is involved in base excision repair (BER), one of several DNA repair mechanisms that may counteract the effects of chemo- and radiation therapy for the treatment of cancer. We envisage that potent inhibitors of OGG1 may be found among the 9-alkyl-8-oxoguanines. Thus we explored synthetic routes to 8-oxoguanines and examined these as OGG1 inhibitors. The best reaction sequence started from 6-chloroguanine and involved N-9 alkylation, C-8 bromination, and finally simultaneous hydrolysis of both halides. Bromination before N-alkylation should only be considered when the N-substituent is not compatible with bromination conditions. The 8-oxoguanines were found to be weak inhibitors of OGG1. 6-Chloro-8-oxopurines, byproducts in the hydrolysis of 2,6-halopurines, turned out to be slightly better inhibitors than the corresponding 8-oxoguanines.
Keywords: alkylation; cancer; DNA; enzyme inhibitors; guanine; halogenation
OPEN ACCESS
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1. Introduction
Chemo- and radiotherapy are, in addition to surgery for removal of solid tumors, the two main treatment protocols currently available to improve the outcome of cancer patients in general, but treatment-related toxicity, the risk of secondary cancers, and the emergence of resistance limit their effectiveness [1]. Some chemotherapeutic drugs and radiotherapy work partly by imposing high concentrations of DNA damage on the genome of cancer cells, beyond the repair capacity of those cells. The drug-exposed cancer cells are heavily dependent on efficient DNA repair to survive. Consequently, inhibitors that reduce DNA repair activities should sensitize cancer cells to chemo- and/or radiotherapy [2–5].
Several DNA repair mechanisms counteract exogenous and endogenous processes that destabilize or directly damage genomes. The processes include, among others, base excision repair (BER), a mechanism that depends on enzymes that recognize small modifications in the native bases in DNA, resulting from alkylation, oxidation, deamination, or hydrolysis of the DNA bases. The pathway is initiated by a damage-specific DNA glycosylase that removes the altered base [6]. Some of these enzymes mainly remove oxidized bases, such as the human 8-oxoguanine DNA glycosylase (OGG1) that removes guanines that have been oxidized at the C8-position. The 8-oxoguanine base in the DNA is flipped into a lesion recognition pocket on the enzyme surface, exposing the Watson–Crick signature of guanine and the oxidized C8 position (Figure 1).
Figure 1. Structural details of 8oxoG base flipped into the lesion recognition pocket of OGG1 (Protein Data Bank deposition 1EBM [7]). The protein backbone is shown as a blue ribbon/helix. Selected amino acid side chains and the 8oxoG base are shown as ball-and-stick. Hydrogen bonds between the protein and 8oxoG are shown as dashed lines. Asp268 is the catalytic residue in OGG1. Symbols 5′ and 3′ indicate the position of the 5′ and 3′ phosphodiester links in the DNA.
We envisage that potent inhibitors of OGG1 may be found among the 9-alkyl-8-oxoguanines. The 8-oxo derivatives of guanosine or deoxyguanosine are probably not inhibitors of the glycosylases since they themselves may be substrates for the enzymes that cleave N,O-acetals in nucleic acids. As a continuance of our synthetic studies directed towards 9-substituted 8-oxoadenines [8,9], we herein present strategies for the synthesis of N-9 substituted 8-oxoguanines. Previous routes include rather tedious constructions of the guanine ring system [10–12], and hydrolysis of purine precursors; hydrolysis of
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8-halopurines [13–16], or less conveniently hydrolysis of N-7 functionalizedpurines [11,17–19]. Results regarding inhibitory activity against the human DNA glycosylase OGG1 are also presented.
2. Results and Discussion
2.1. Chemistry
We found it most convenient to start the synthesis of 9-alkyl-8-oxopurines from commercially available purines, and in our opinion the best way to introduce the 8-oxo group would be by hydrolysis of an 8-halopurine. However, there still was the question of whether the halogen or the N-9 substituent should be introduced first and which protection/activation groups should be employed in the synthesis. Ideally, such groups should also be removed in the final hydrolysis step. Regioselectivity in N-alkylation of guanine derivatives was also an issue [20–27]. We chose to start from two guanine precursors, commercially available 2-amino-6-chloropurine (1a) and the O-carbamoylguanine 1b, easily available from guanine [28,29]. The synthetic routes explored are all summarized in Scheme 1.
Reagents and conditions: (a) See Table 1; (b) See Table 2; (c) 1. Ac2O, NaOAc, AcOH, 2. NaOH(aq), ∆; (d) 1. LDA, 2. (CCl2Br)2, THF, −78 °C; (e) Br2, CHCl3; (f) See [30]; (g) HCl(aq), EtOH.
Scheme 1. Synthetic routes to 8-oxoguanines 5.
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Table 1. N-alkylation of guanine precursors 1a and 1b.
Entry R2 R6 R Reagents and
Conditions Ratio 2:3:1 a Yield (%) 2 b Yield (%) 3 b
1 Cl NH2 CH2-c-hexyl RBr, K2CO3, DMF, rt, 72 h
80:20:0 67, 2a 10, 3a
2 Cl NH2 CH2-c-hexyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
93:7:0 76, 2a 5, 3a
3 OCONPh2 NHAc CH2-c-hexyl RBr, K2CO3, DMF, rt, 72 h
81:19:0 45, 2e 7, 3e
4 OCONPh2 NHAc CH2-c-hexyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
82:18:0 70, 2e 3, 3e
5 Cl NH2 c-hexyl RI, K2CO3, DMF, rt, 72 h
15:0:85 – c –
6 Cl NH2 c-hexyl ROTs, K2CO3, DMF, rt, 72 h
– d 33, 2b – c
7 Cl NH2 c-hexyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
8:4:88 – c – c
8 Cl NH2 c-hexyl ROH, DIAD, PPh3, THF, ultrasound, 14 h
27:0:73 20, 2b –
9 Cl NH2 c-hexyl ROH, DIAD, PPh3, DMF, 150 °C, μW, 2 h
41:8:51 – c – c
10 OCONPh2 NHAc c-hexyl ROTs, K2CO3, THF, rt, 72 h
– d 30, 2f – c
11 OCONPh2 NHAc c-hexyl ROTs, K2CO3, DMF, 80 °C, 72 h
– d,e – c – c
12 OCONPh2 NHAc c-hexyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
– d 22, 2f – c
13 Cl NH2 c-pentyl RBr, K2CO3, DMF, rt, 72 h
86:14:0 71, 2c 5, 3c
14 Cl NH2 c-pentyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
91:9:0 72, 2c 6, 3c
15 OCONPh2 NHAc c-pentyl RBr, K2CO3, DMF, rt, 72 h
76:15:09 52, 2g – c
16 OCONPh2 NHAc c-pentyl ROH, DIAD, PPh3, THF, 70 °C, 14 h
90:10:0 58, 2g – c
17 Cl NH2 c-pent-2-enyl RBr, K2CO3, DMF, rt, 24 h
23:16:61 18, 2d – c
18 Cl NH2 c-pent-2-enyl ROH, DIAD, PPh3, THF, 70 °C, 42 h
55:18:27 40, 2d – c
19 Cl NH2 c-pent-2-enyl ROAc, Pd(PPh3)4,NaH, DMSO, f 50 °C, 48 h
75:25:0 53, 2d 18, 3d
a From 1H-NMR spectra of the crude products, the signals from H-8 in compounds 1, 2, and 3 were integrated; b Isolated yields; c Not isolated in pure form; d Difficult to determine due to overlapping signals in the 1H-NMR spectra; e A complex mixture was formed; f Comparable results were obtained in DMF.
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Table 2. Synthesis of 8-bromopurines 4.
Entry Starting Material a Reagents and Conditions Yield (%) 4 a,b
1 2a Br2, H2O 79%, 4a 2 10 RBr, K2CO3, DMF 34%, 4a 3 10 ROH, DIAD, PPh3, THF, 70 °C 56%, 4a 4 2b Br2, H2O 70%, 4b 5 2c Br2, H2O 81%, 4c 6 2d 1. LDA, 2. CCl2BrCCl2Br, THF, −78 °C 32%, 4d 7 10 ROH, DEAD, PPh3, THF, 70 °C 42%, 4d 8 10 ROAc, Pd(PPh3)4, NaH, DMF, 50 °C 29%, 4d
a The structures are shown in Scheme 1; b Isolated yields.
First we chose to N-alkylate the substrates 1 before C-8 halogenation and hydrolysis. Alkylations were conducted by various methodologies in order to find the conditions that gave the desired N-9 alkylated isomer 2 with high selectivity and in a good isolated yield (Scheme 1, Table 1). Relatively simple alkylating agents were chosen for the model reactions and we focused on alkylation with alkyl halides in the presence of base, Mitsunobu reactions, and Pd-catalyzed allylic alkylation.
The cyclohexylmethyl substituent could be introduced at N-9 either by reaction with alkyl bromide in the presence of a base [31,32] (Table 1, Entries 1 and 3) or with cyclohexylmethanol under Mitsunobu conditions (Table 1, Entries 2 and 4). The latter is often claimed to be more N-9 selective compared to classical alkylations of purines [33–35]. In all cases a mixture of the N-9 and N-7 alkylated isomers (2 and 3) was formed with good selectivity for the desired isomer 2. The isomers were identified from HMQC and HMBC-NMR, as described before [31].
The guanine precursor 1b, carrying a bulky substituent at C-6 that may sterically block N-7, is reported to react with high N-9 selectivity in other N-functionalization reactions [28,29,36–41]. Nevertheless, we found the regioselectivity in N-alkylation of purine 1b equal or slightly poorer compared to 6-chloroguanine 1a in all reactions performed in this study. In the alkylation of compound 1b, minor amounts of other relatively polar products were formed under both reaction conditions. These often made purification of the N-7 alkylated isomer 3 difficult. The identity of the byproducts could not be determined, but they may be formed as a result of cleavage of the O6-protecting group. Alkylation of N2, as observed by others [41], was not seen.
Introduction of the cyclohexyl group at N-9 turned out to be quite difficult (Table 1, Entries 5–12). Both starting materials (1a and 1b) did not react with cyclohexyl bromide (data not shown) and reacted slowly with cyclohexyl iodide or the corresponding tosylate, but compounds 2b and 2f could be isolated in modest yields (Table 1, Entries 5, 6, 10 and 11). It is, however, well known that cyclohexyl halides or pseudo halides may react sluggishly in substitution reactions [42]. The results were not significantly improved when the Mitsunobu reaction was employed (Table 1; Entries 7, 8, and 12), not even under ultrasound (Table 1, Entry 8) or microwave conditions (Table 1, Entry 9).
The cyclopentyl group could easily be installed at N-9 on both starting materials 1a and 1b by reaction with cyclopentyl bromide and base (Table 1, Entries 13 and 15) or by alkylation under Mitsunobu conditions (Table 1, Entries 14 and 16). The selectivity for N-9 was higher in the Mitsunobu reactions, but the isolated yields were comparable due to more tedious purification when Mitsunobu conditions, also producing phosphine oxides and reduced azodicarboxylates, were employed.
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Finally we introduced the cyclopent-2-enyl group at N-9 (Table 1, Entries 17–19). These reactions were only conducted at the guanine precursor 1a, since we so far had not observed any significant improvement in regioselectivity when compound 1b was employed and we had observed problems with compounds derived from purine 1b later in the planned synthetic sequence. In addition to alkylation with the halide and Mitsunobu reaction with the alcohol, we also attempted palladium catalyzed alkylation with the allylic acetate [43]. 3-Bromocyclopentene could only be generated as a 15% solution in CCl4 and the reagent had a limited stability, probably partly due to traces of the radical initiator used in the synthesis left in the solution [44], which may explain the low yield of product 2d (Table 1, Entry 17). The Mitsunobu reaction between purine 1a and cyclopenten-2-ol was surprisingly slow, and full conversion was not achieved even after several days. Furthermore, the N-9/N-7 selectivity was only ca. 4:1 (Table 1, Entry 18). Pd-catalyzed allylic alkylation of purine 1a went to completion and gave the isomers 2d and 3d in a 4:1 ratio (Table 1, Entry 19).
The 6-chloropurines 2a, 2b, and 2c were readily brominated on C-8 simply by treatment of bromine in water (Scheme 1; Table 2; Entries 1, 4, and 5). For compound 2d, which has an alkene function, the bromide was introduced by C-8 lithiation followed by trapping with CCl2BrCCl2Br (Table 2, Entry 6) [9,32,45,46]. However, the yield was surprisingly low and also another route to bromide 4d was examined (see below). Finally hydrolysis of the dihalopurines 4, employing conditions used for hydrolysis of other 8-bromopurines [13–16,47], gave the 8-oxoguanines 5. Complete conversion was achieved in the hydrolysis compound 4a, whereas small amounts of the partly hydrolyzed chlorides 6 where present after hydrolysis of purines 4b–d even after prolonged reaction times.
Attempts to brominate the O-carbamoylguanine 2e failed (Scheme 1). Treatment with bromine or lithiation followed by trapping with CCl2BrCCl2Br only resulted in cleavage of the carbamoyl protecting group to give the guanine derivative 7. When compound 2e was treated with NBS, no reaction took place at all. Thus, no attempts were made to brominate the carbamoyl protected guanines 2f and 2g.
Since bromination of the cyclopentenylpurine 2d turned out to be a challenge, we also examined the possibility for introducing the 8-halo substituent before the N-9 alkyl group (Scheme 1). We chose to brominate the THP protected compound 8 [30] and removed the protection group under mild acidic condition, but direct bromination of purine 1a in a moderate yield has also been reported [48].
Alkylation of 8-bromo-6-chloropurin-2-amine (10) by bromomethylcyclohexane in the presence of K2CO3/DMF (Table 2, Entry 2) occurred slowly compared to alkylation of 2-amino-6-chloropurine (1a) under the same set of reaction conditions (for alkylation of compound 1a see Table 1). NMR analysis showed that approximately 50% of the starting material was intact even after 96 h reaction time and the desired product was isolated in a low yield. Also, ca. 4% of N-7 alkylated isomer was formed, as judged by NMR. When compound 10 was reacted under Mitsunobu (Table 2, Entry 3) conditions, high conversion (ca. 95%) and almost full selectivity towards the desired N-9 alkylated isomer 4a was achieved, as judged by 1H-NMR. However, the product 4a was isolated only in 56% due to tedious separation from reduced DIAD. Since compound 10 reacted slower (conventional alkylation) or comparably (Mitsunobu alkylation) to compound 1a, it was concluded that there were no benefits associated with introducing the bromide before the N-alkyl group for the synthesis of 8-bromopurines 4a–c.
Also, synthesis of the 9-cyclopentenylpurine 4d by N-alkylation of compound 10 was examined (Table 2, Entries 7 and 8) since bromination of 2-amino-6-chloro-9-cyclopentenylpurine 2d turned out to give only a low yield of the desired product. Again, isolation of the desired product from alkylation
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under Mitsunobu conditions turned out to be troublesome. We tried this Mitsunobu alkylation using the water-soluble azodicarboxylate DMEAD (di-2-methoxyethyl azodicarboxylate) as well as DIAD [49]. Purification of the product was less complicated, but the conversion was low and ca. 40% of starting material 10 was recovered. Also, Pd-catalyzed allylation turned out to be a very slow reaction and even after six days only 29% of the desired compound 4d could be isolated, together with 32% unconverted starting material 10.
2.2. Biology
As previously mentioned, our hypothesis was that N-alkyl-8-oxoguanines may inhibit the human 8-oxoguanine DNA glycosylase (OGG1). Other substituents in the purine 8-position are probably not tolerated, for instance 8-bromo- and 8-aminoguanines are reported to be enhancers for OGG1 activity [50]. Thus, the 8-oxoguanines 5 as well as the partly hydrolyzed 6-chloro-8-oxopurines 6 were tested against human DNA glycosylases OGG1 and NTH1. A general structure of the tested compounds is shown in Figure 2 and the results are presented in Tables 3 and 4, and Supplementary Figure S19.
Figure 2. General structure of the compounds shown in Table 3.
Table 3. % Activity of OGG1 in the presence of compounds 5 or 6 at 0.2 mM concentration.
Compound X R % Activity5a OH a CH2-c-hexyl 89 ± 5 5b OH a c-hexyl 92 ± 2 6b Cl c-hexyl 70 ± 11 5c OH c-pentyl 101 ± 12 6c Cl c-pentyl 72 ± 9 5d OH c-pent-2-enyl 92 ± 7 6d Cl c-pent-2-enyl 84 ± 3
a The predominant 6-oxo tautomer of compounds 5 is shown in Scheme 1.
Table 4. % Activity of NTH1 in the presence of compounds 5 or 6 at 0.5 mM concentration.
Compound X R % Activity5a OH a CH2-c-hexyl 96 ± 3 5b OH a c-hexyl 123 ± 20 6b Cl c-hexyl 73 ± 37 5c OH c-pentyl 102 ± 16 6c Cl c-pentyl 108 ± 18 5d OH c-pent-2-enyl 104 ± 21 6d Cl c-pent-2-enyl 89 ± 13
a The predominant 6-oxo tautomer of compounds 5 is shown in Scheme 1.
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Compounds 6b and 6c inhibit the OGG1 enzyme by ca. 30%, followed by compounds 5a, 5b, and 6d at ca. 10%–15%, all at 0.2 mM ligand concentration. Interestingly, the halogenated compounds seem in general to be better inhibitors than their 6-oxo derivatives. To check enzyme specificity, we tested the same seven compounds at the higher concentration of 0.5 mM against NTH1, a structural but not functional homolog of OGG1. Both enzymes have a deep pocket for binding of oxidized bases; in general, OGG1 repairs oxidized purines while NTH1 is involved in repair of oxidized pyrimidines. Compound 6b reduced the NTH1 activity by around 25% at 0.5 mM ligand concentration. An effect of varying the N-9 substituent is not so evident from the few compounds examined.
3. Experimental Section
3.1. General Information
1H-NMR spectra were recorded at 300 MHz with a Bruker DPX 300, at 400 MHz with a Bruker DPX 400 or at 600 with a Bruker AVI 600 instrument (Bruker BioSpin AG, Fällanden, Switzerland). The 13C-NMR spectra were recorded at 75, 100, or 150 MHz with the Bruker instruments listed above. Assignments of 1H and 13C resonances are inferred from 1D 1H-NMR, 1D 13C-NMR, DEPT, or APT, and 2D NMR (HMQC, HMBC) spectroscopical data. 1H- and 13C-NMR spectra of all novel compounds can be found in the Supplementary Material (Figures S1–S18). HRMS (EI) was performed with a double-focusing magnetic sector VG Prospec Q instrument (Waters, Manchester, UK) and HRMS (ESI) with a TOF quadrupole Micromass QTOF 2 W instrument (Waters). Melting points were determined with a Büchi Melting point B-545 apparatus (Büchi Labortechnik AG, Flawil, Switzerland) and are uncorrected. Dry DMF and THF were obtained from a solvent purification system, MB SPS-800 (MBraun, Garching, Germany). Acetic anhydride and diisopropylamine were distilled over CaH2. DMSO was dried over activated 3 Å molecular sieves for four days. Potassium carbonate was oven dried at 150 °C under high vacuum for 12 h. A saturated aqueous solution of Br2 was prepared by stirring water (20 mL) with Br2 (0.200 mL) in a closed container for 15 min at ambient temperature. Sodium hydride (ca. 60% in mineral oil) was washed with dry pentane under inert atmosphere prior to use. All other reagents were commercially available and used as received. The following compounds were available by literature methods: Cyclohexyl tosylate [51], cyclopentenyl bromide [44], cyclopent-2-enol [52], cyclopentenyl acetate [53], 1b [29], 8 [30].
3.2. Synthesis
3.2.1. 2-Amino-6-chloro-9-(cyclohexylmethyl)-9H-purine (2a) and 2-Amino-6-chloro-7-(cyclohexylmethyl)-7H-purine (3a)
Method A: K2CO3 (1.63 g, 11.8 mmol) was added to a stirring solution of compound 1a (1.00 g, 5.90 mmol) in dry DMF (30 mL) at ambient temperature under N2. After 20 min bromomethylcyclohexane (0.905 mL, 6.49 mmol) was added and the resulting mixture was stirred for 72 h, filtered. and evaporated. The isomers were separated by flash chromatography on silica gel, eluting with MeOH–CH2Cl2 (1:9) to yield 2a (1.05 g, 67%) and 3a (150 mg, 10%).
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2a: colorless solid; mp 148–150 °C (lit. [54], 154–155 °C); 1H-NMR (DMSO-d6, 400 MHz) δ 8.10 (s, 1H, H-8), 6.91 (s, 2H, NH2), 3.88 (d, J = 7.4 Hz, 2H, NCH2), 1.88–1.72 (m, 1H, H-1 in c-hex), 1.68–1.52 (m, 3H, c-hex), 1.51–1.42 (m, 2H, c-hex), 1.19–1.02 (m, 3H, c-hex) 1.00–0.85 (m, 2H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 159.8 (C, C-2), 154.3 (C, C-4), 149.3 (C, C-6), 143.7 (CH, C-8), 123.3 (C, C-5), 48.8 (CH2, NCH2), 37.1 (CH, C-1 in c-hex), 29.9 (CH2, C-3 and C-5 in c-hex), 25.8 (CH2, C-4 in c-hex), 25.0 (CH2, C-2 and C-6 in c-hex); HREIMS m/z 265.1092 (calcd for C12H16ClN5, 265.1094). Spectral data were in good agreement with those reported before [54].
3a: colorless solid mp 228–231 °C. 1H-NMR (DMSO-d6, 400 MHz) δ 8.32 (s, 1H, H-8), 6.62 (s, 2H, NH2), 4.10 (d, J = 7.2 Hz, 2H, NCH2), 1.82–1.70 (m, 1H, H-1 in c-hex), 1.69–1.54 (m, 3H, c-hex), 1.50–1.41 (m, 2H, c-hex), 1.24–1.06 (m, 3H, c-hex), 1.03–0.89 (m, 2H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 164.2 (C, C-4), 159.9 (C, C-2), 149.8 (CH, C-8), 142.3 (C, C-6), 114.9 (C, C-5), 51.8 (CH2, NCH2), 38.6 (CH, C-1 in c-hex), 29.6 (CH2, C-3 and C-5 in c-hex), 25.8 (CH2, C-4 in c-hex), 25.1 (CH2, C-2 and C-3 in c-hex); HREIMS m/z 265.1096 (calcd for C12H16ClN5, 265.1094).
Method B: Compound 1a (200 mg, 1.18 mmol) was added to a solution of cyclohexylmethanol (141 mg, 1.24 mmol) and PPh3 (325 mg, 1.24 mmol) in dry THF (10 mL) under N2. The resulting suspension was treated with diisopropyl azodicarboxylate (DIAD) (0.244 mL, 1.24 mmol) and the reaction mixture was stirred at 70 °C for 7 h before cyclohexylmethanol (141 mg, 1.24 mmol), PPh3 (325 mg, 1.24 mmol), and DIAD (0.244 mL, 1.24 mmol) were added. The mixture was stirred for another 7 h at 70 °C, cooled, treated with brine (10 mL), and extracted with CH2Cl2 (3 × 75 mL). The combined organic layers were washed with water (50 mL), dried (Na2SO4) and evaporated in vacuo. The isomers were separated by flash chromatography on silica gel eluting with EtOAc–Hexane (gradient; 70%–100% EtOAc) followed by MeOH–EtOAc (1:9) to yield 2a (240 mg, 76%) and 3a (16 mg, 5%).
3.2.2. 2-Amino-6-chloro-9-(cyclohexyl)-9H-purine (2b)
Method A: The title compound was prepared from compound 1a (200 mg, 1.18 mmol), K2CO3 (326 mg, 2.36 mmol) and cyclohexyl tosylate (450 mg, 1.77 mmol) in DMF (15 mL) as described for the synthesis of compounds 2a above. MeOH–EtOAc (1:19) was used for flash chromatography to yield 2b (98 mg, 33%). Colorless needles; mp 163–165 °C (lit. [55], 165 °C); 1H-NMR (DMSO-d6, 400 MHz) δ 8.23 (s, 1H, H-8), 6.88 (s, 2H, NH2), 4.28–4.12 (m, 1H, H-1 in c-hex), 2.01–1.75 (m, 7H, c-hex), 1.45–1.17 (m, 3H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 159.5 (C, C-6), 153.5 (C, C-4), 149.3 (C, C-2), 141.2 (CH, C-8), 123.5 (C, C-5), 53.5 (CH, C-1 in c-hex), 31.9 (CH2, c-hex), 25.1 (CH2, c-hex), 24.7 (CH2, c-hex); HREIMS m/z 251.0934 (calcd for C11H14ClN5, 251.0938). Spectral data were in good agreement with those reported before [55].
Method B: The title compound was prepared from compound 1a (1.00 g, 5.90 mmol), cyclohexanol [2 × (620 mg, 6.19 mmol)], PPh3 [2 × (1.62 g, 6.19 mmol)] and DIAD [2 × (1.22 mL, 6.19 mmol] in THF (50 mL) as described for the synthesis of compounds 2a above. After each addition of cyclohexanol the mixture was subjected to sonication for 20 min using a sonicator probe. EtOAc–Hexane (gradient; 30%–100% EtOAc) was used for flash chromatography to yield 2b (295 mg, 20%).
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3.2.3. 2-Amino-6-chloro-9-(cyclopentyl)-9H-purine (2c) and 2-Amino-6-chloro-7-(cyclopentyl)-7H-purine (3c)
Method A: The title compounds were prepared from compound 1a (1.00 g, 5.90 mmol), K2CO3 (1.63 g, 11.8 mmol) and bromocyclopentane (0.696 mL, 6.49 mmol) in DMF (50 mL) as described for the synthesis of compounds 2a and 3a above. MeOH–EtOAc (1:19) was used for flash chromatography to yield 2c (994 mg, 71%) and 3c (75 mg, 5%).
2c: colorless solid; mp 137–140 °C (lit. [55], 142 °C); 1H-NMR (DMSO-d6, 400 MHz) δ 8.20 (s, 1H, H-8), 6.86 (s, 2H, NH2), 4.77–4.65 (m, 1H, H-1 in c-pent), 2.16–2.02 (m, 2H, c-pent) 2.00–1.75 (m, 4H, c-pent), 1.72–1.60 (m, 2H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 159.3 (C, C-2), 153.7 (C, C-4), 149.1 (C, C-6), 141.4 (CH, C-8), 123.5 (C, C-5), 55.1 (CH, C-1 in c-pent), 31.4 (CH2, C-3 and C-4 in c-pent), 23.2 (CH2,C-2 and C-5 in c-pent); HREIMS m/z 237.0777 (calcd for C10H12ClN5, 237.0781). Spectral data were in good agreement with those reported before [34,55,56].
3c: colorless solid; mp >230 °C (dec.); 1H-NMR (DMSO-d6, 400 MHz) δ 8.46 (s, 1H, H-8), 6.59 (s, 2H, NH2), 5.11–5.01 (m, 1H, H-1 in c-pent), 2.24–2.10 (m, 2H, c-pent), 2.02–1.90 (m, 2H, c-pent), 1.86–1.62 (m, 4H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 164.3 (C, C-4), 159.7 (C, C-2), 146.6 (CH, C-8), 142.2 (C, C-6), 115.1 (C, C-5), 58.0 (CH, C-1 in c-pent), 32.6 (CH2, C-3 and C-4 in c-pent), 23.1 (CH2, C-2 and C-5 in c-pent); HREIMS m/z 237.0776 (calcd for C10H12ClN5, 237.0781). Spectral data were in good agreement with those reported before [34,55].
Method B: The title compounds were prepared from compound 1a (200 mg, 1.18 mmol), cyclopentanol [2 × (107 mg, 1.24 mmol)], PPh3 [2 × (325 mg, 1.24 mmol)] and DIAD [2 × (244 µL, 1.24 mmol)] in THF (10 mL) as described for the synthesis of compounds 2a and 3a above. EtOAc–hexane (gradient; 70%–100% EtOAc) followed by MeOH–EtOAc (1:9) were used for flash chromatography to 2c (202 mg, 72%) and 3c (6 mg, 6%).
3.2.4. 2-Amino-6-chloro-9-(cyclopent-2-enyl)-9H-purine (2d) and 2-Amino-6-chloro-7-(cyclopent-2-enyl)-7H-purine (3d)
Method A: The title compound 2d was prepared from compound 1a (200 mg, 1.18 mmol), K2CO3 (490 mg, 3.54 mmol) and 3-bromocyclopentene (0.29 mL, ca. 80% pure, ca. 2.4 mmol) in DMF (20 mL) as described for the synthesis of compounds 2a and 3a above, except that the reaction time was 24 h. EtOAc–hexane (gradient; 50%–100% EtOAc) followed by MeOH–EtOAc (1:9) were used for flash chromatography to yield 2d (49 mg, 18%). Colorless solid; mp 154–154.5 °C (lit., [57] 166.0–166.7 °C); 1H-NMR (DMSO-d6, 400 MHz) δ 7.96 (s, 1H, H-8), 6.88 (s, 2H, NH2), 6.26–6.18 (m, 1H, c-pent), 5.93–5.84 (m, 1H, c-pent), 5.51–5.41 (m, 1H, c-pent), 2.73–2.61 (m, 1H, c-pent), 2.47–2.36 (m, 2H, c-pent), 2.00–1.87 (m, 1H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 159.6 (C, C-6), 153.6 (C, C-4), 149.3 (C, C-2), 141.1 (CH, C-8), 137.3 (CH, C-2 in c-pent), 128.6 (CH, C-3 in c-pent), 123.6 (C, C-5), 59.4 (CH, C-1 in c-pent), 31.2 (CH2, C-5 in c-pent), 30.4 (CH2, C-4 in c-pent); HREIMS m/z 235.0624 (calcd for C10H10ClN5 235.0625). Spectral data were in good agreement with those reported before [57].
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Method B: The title compound 2d was prepared from compound 1a (340 mg, 2.01 mmol), cyclopent-2-enol [2 × (0.180 mL, 2.03 mmol)], PPh3 [2 × (531 mg, 2.03 mmol)] and DIAD [2 × (0.442 mL, 2.03 mmol)] in THF (20 mL) as described for the synthesis of compounds 2a and 3a above. EtOAc–Hexane (gradient; 70%–100% EtOAc) followed by MeOH–EtOAc (1:9) were used for flash chromatography to yield 2d (187 mg, 40%).
Method C: A solution of compound 1a (100 mg, 0.590 mmol) and NaH (18 mg, 0.77 mmol) in dry DMSO (5 mL) was stirred at room temperature for 20 min under Ar atmosphere. The mixture was added to a solution of cyclopent-2-en-1-yl acetate (0.070 mL, 0.77 mmol) and Pd(PPh3)4 (103 mg, 0.0890 mmol) in dry DMSO (5 mL) and the resulting mixture was stirred at 50 °C for 48 h under Ar and evaporated in vacuo. The product was purified by flash chromatography as described in Method B to yield 2d (73 mg, 53%) and 3d (25 mg, 18%).
3d: colorless solid; mp 155–157 °C (dec.); 1H-NMR (DMSO-d6, 400 MHz) δ 8.15 (s, 1H, H-8), 6.61 (s, 2H, NH2), 6.34–6.28 (m, 1H, c-pent), 6.03–5.96 (m, 1H, c-pent), 5.82–5.75 (m, 1H, c-pent) 2.61–2.34 (m, 3H, c-pent) 1.96–1.83 (m, 1H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 164.4 (C, C-4), 159.8 (C, C-2), 146.2 (CH, C-8), 142.3 (C, C-6), 138.3 (CH, C-2 in c-pent), 127.9 (CH, C-3 in c-pent), 114.8 (C, C-5), 62.5 (CH, C-1 in c-pent), 32.1 (CH2, C-5 in c-pent), 31.0 (CH2, C-4 in c-pent); HREIMS m/z 235.0631 (calcd for C10H10ClN5, 235.0625). Spectral data were in good agreement with those reported before [57].
3.2.5. 2-Acetamido-9-(cyclohexylmethyl)-9H-purin-6-yl diphenylcarbamate (2e) and 2-Acetamido-7-(cyclohexylmethyl)-7H-purin-6-yl diphenylcarbamate (3e)
Method A: The title compounds were prepared from compound 1b (200 mg, 0.515 mmol), K2CO3 (142 mg, 1.03 mmol) and bromomethylcyclohexane (0.144 mL, 1.03 mmol) in DMF (7 mL) as described for the synthesis of compounds 2a and 3a above. MeOH–CH2Cl2 (1:9) followed by MeOH–CH2Cl2 (1:4) were used for flash chromatography to yield 2e (111 mg, 45%) and 3e (18 mg, 7%).
2e: colorless solid; mp. 192–194 °C; 1H-NMR (CDCl3, 400 MHz) δ 7.97 (s, 1H, NH), 7.87 (s, 1H, H-8), 7.47–7.24 (m, 10H, Ph), 3.99 (d, J = 7.0 Hz, 2H, NCH2), 2.58 (s, 3H, CH3), 1.85 (m, 1H, H-1 in c-hex), 1.78–1.58 (m, 5H, c-hex), 1.30–1.07 (m, 3H, c-hex), 1.07–0.92 (m, 2H, c-hex); 13C-NMR (CDCl3, 100 MHz) δ 171.1 (C, CONH), 156.2 (C, OCON), 155.4 (C, C-4), 152.2 (C, C-2), 150.6 (C, C-6), 144.4 (CH, C-8), 141.9 (C, Ph), 129.3 (CH, Ph), 127.2 (br, 2 × CH2, Ph), 120.6 (C, C-5), 50.5 (CH2, NCH2), 38.4 (CH, C-1 in c-hex), 30.8 (CH2, C-3 and C-5 in c-hex), 26.1 (CH2, C-4 in c-hex), 25.6 (CH2, C-2 and C-3 in c-hex), 25.3 (CH3); HRESIMS m/z 485.2311 (calcd for C27H29N6O3 + 1, 485.2301).
3e: colorless oil; 1H-NMR (CDCl3, 400 MHz) δ 8.10 (s, 1H, NH), 7.96 (s, 1H, H-8), 7.42–7.36 (m, 8H, Ph), 7.33–7.28 (m, 2H, Ph), 3.89 (d, J = 7.2 Hz, 2H, NCH2), 2.63 (s, 3H, CH3), 1.68–1.60 (m, 3H, c-hex), 1.45–1.35 (m, 2H, c-hex), 1.13–0.98 (m, 3H, c-hex), 0.92–0.72 (m, 3H, c-hex); 13C-NMR (CDCl3, 100 MHz) δ 172.0 (C, CONH), 164.9 (C-4), 152.2 (C, C-2), 151.9 (C, OCON), 149.5 (C, C-6), 148.5 (CH, C-8), 141.5 (C, Ph), 129.6 (CH, Ph), 127.6 (br, 2 × CH, Ph), 111.9 (C, C-5), 53.8 (CH2, NCH2), 38.9 (CH, C-1 in c-hex), 30.3 (CH2, C-3 and C-5 in c-hex), 26.0 (CH2,C-4 in c-hex) 25.4 (CH2, C-2 and C-6 in c-hex ), 25.6 (CH3); HRESIMS m/z 485.2313 (calcd for C27H29N6O3 + 1, 485.2301).
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Method B: The title compounds were prepared from compound 1b (200 mg, 0.515 mmol), cyclohexylmethanol [2 × (62 mg, 0.54 mmol)], PPh3 [2 × (142 mg, 0.540 mmol)] and DIAD [2 × (0.106 mL, 0.540 mmol)] in THF (10 mL) as described for the synthesis of compounds 2a and 3a above. EtOAc–Hexane (gradient; 70%–100% EtOAc) followed by MeOH–EtOAc (1:9) were used for flash chromatography to yield 2e (175 mg, 70%) and 3e (7 mg, 3%).
3.2.6. 2-Acetamido-9-(cyclohexyl)-9H-purin-6-yl diphenylcarbamate (2f)
Method A: The title compound was prepared from compound 1b (500 mg, 1.29 mmol), K2CO3 (329 mg, 2.38 mmol) and cyclohexyl tosylate (441 mg, 1.73 mmol) in THF (15 mL) as described for the synthesis of compounds 2a above. MeOH–EtOAc (1:19) was used for flash chromatography to yield 2f (180 mg, 30%). Off-white solid; mp 189–190 °C; 1H-NMR (DMSO-d6, 300 MHz) δ 10.67 (s, 1H, NH), 8.55 (s, 1H, H-8), 7.54–7.38 (m, 8H, Ph), 7.37–7.25 (m, 2H), 4.46–4.28 (m, 1H, H-1 in c-hex), 2.20 (s, 3H, CH3), 2.06–1.80 (m, 6H, CH in c-hex), 1.76–1.65 (m, 1H, c-hex), 1.51–1.14 (m, 3H, c-hex); 13C-NMR (DMSO-d6, 75 MHz) δ 168.8 (C, CONH), 155.0 (C, OCON), 154.3 (C, C-4), 151.7 (C, C-2), 150.3 (C, C-6), 144.1 (CH, C-8), 141.6 (C, Ph), 129.4 (CH, Ph), 127.1 (CH, Ph), 120.1 (C, C-5), 54.1 (CH, C-1 in c-hex), 31.9 (CH2, C-3 and C-5 in c-hex), 25.1 (CH2, C-2 and C-6 in c-hex), 24.7 (CH2, C-4 in c-hex), 24.6 (CH3); HREIMS m/z 470.2057 (calcd for C26H26N6O3, 470.2066).
Method B: The title compound was prepared from compound 1b (400 mg, 1.03 mmol), cyclohexanol [2 × (108 mg, 1.08 mmol)], PPh3 [2 × (284 mg, 1.08 mmol)] and DIAD [2 × (0.213 mL, 1.08 mmol)] in THF (10 mL) as described for the synthesis of compound 2a above. EtOAc–Hexane (gradient; 30%–100% EtOAc) was used for flash chromatography to yield 2f (107 mg, 22%) as an off-white solid.
3.2.7. 2-Acetamido-9-(cyclopentyl)-9H-purin-6-yl diphenylcarbamate (2g)
Method A: The title compound 2g was prepared from compound 1b (389 mg, 1.00 mmol), K2CO3 (277 mg, 2.00 mmol) and bromocyclopentane (0.120 mL, 1.10 mmol) in DMF (50 mL) as described for the synthesis of compounds 2a above. MeOH–EtOAc (1:19) was used for flash chromatography to yield 2g (238 mg, 52%). Colorless solid; mp 137–140 °C; 1H-NMR (DMSO-d6, 400 MHz) δ 10.62 (s, 1H, NH), 8.51 (s, 1H, H-8), 7.53–7.40 (m, 8H, Ph), 7.36–7.27 (m, 2H, Ph), 4.79–4.62 (m, 1H, H-1 in c-pent), 2.21 (s, 3H, CH3), 2.19–2.11 (m, 2H, c-pent), 2.10–1.83 (m, 4H, c-pent), 1.77–1.62 (m, 2H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 168.8 (C, CONH), 155.0 (C, OCON), 154.6 (C, C-4), 151.7 (C, C-2), 150.2 (C, C-6), 144.4 (CH, C-8), 141.6 (C, Ph), 129.4 (CH, Ph), 127.2 (CH, Ph), 120.3 (C, C-5), 56.1 (CH, C-1 in c-pent), 31.7 (CH2, C-3 and C-4 in c-pent), 24.5 (CH3), 23.5 (CH2, C-2 and C-5 in c-pent), one Ph signal was hidden; HREIMS m/z 456.1903 (calcd for C25H24N6O3, 456.1910).
Method B: The title compound 2g was prepared from compound 1b (389 mg, 1.00 mmol), cyclopentanol [2 × (91 mg, 1.1 mmol)], PPh3 [2 × (276 mg, 1.05 mmol)] and DIAD [2 × (0.207 mL, 1.05 mmol)] in THF (10 mL) as described for the synthesis of compounds 2a above. EtOAc–Hexane (gradient; 70%–100% EtOAc) followed by MeOH–EtOAc (1:9) were used for flash chromatography to yield 2g (264 mg, 58%).
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3.2.8. 2-Amino-8-bromo-6-chloro-9-(cyclohexylmethyl)-9H-purine (4a)
Method A: Sat. aq. Br2 (50 mL) was added dropwise to a rapidly stirred suspension of 2a (1.50 g, 5.64 mmol) in water (20 mL) over 10 min at ambient temperature. The flask was closed and the reaction mixture was stirred for 74 h. The flask was left open in the hood until all Br2 was evaporated before the water was removed in vacuo. The product was purified by flash chromatography on silica gel, eluting with EtOAc–Hexane (1:1) to yield 4a (1.55 g, 79%). Yellow solid; mp 169–170 °C. 1H-NMR (DMSO-d6, 400 MHz) δ 7.07 (s, 2H, NH2), 3.86 (d, J = 8.0 Hz, 2H, NCH2), 1.96–1.81 (m, 1H, H-1 in c-hex), 1.72–1.44 (m, 5H, c-hex), 1.24–0.92 (m, 5H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 159.7 (C, C-2), 155.0 (C, C-4), 148.0 (C, C-6), 129.5 (C, C-8), 123.3 (C, C-5), 49.6 (CH2, NCH2), 36.9 (CH, c-hex), 30.0 (CH2, c-hex), 25.7 (CH2, c-hex), 25.1 (CH2, c-hex); HREIMS m/z 343.0198 (calcd for C12H15BrClN5, 343.0199).
Method B: K2CO3 (231 mg, 1.67 mmol) was added to a stirring solution of compound 10 (207 mg, 0.833 mmol) in dry DMF (15 mL) at ambient temperature under N2. After 20 min, bromomethylcyclohexane (0.175 mL, 1.25 mmol) was added and the resulting mixture was stirred for 80 h before K2CO3 (115 mg, 0.835 mmol) and bromomethylcyclohexane (0.175 mL, 1.25 mmol) was added and the mixture was stirred for additional 16 h and evaporated in vacuo. The product was purified by flash chromatography on silica gel eluting with EtOAc–Hexane (2:3) to yield 4a (98 mg, 34%).
Method C: The title compound was prepared from compound 10 (175 mg, 0.704 mmol), cyclohexylmethanol [2 × (0.091 mL, 0.74 mmol)], PPh3 [2 × (276 mg, 0.740 mmol)] and DIAD [2 × (0.146 mL, 0.740 mmol)] in THF (10 mL) as described for the synthesis of compounds 2a above. EtOAc–Hexane (gradient; 20%–100% EtOAc) was used for flash chromatography to yield 4a (136 mg, 56%).
3.2.9. 2-Amino-8-bromo-6-chloro-9-(cyclohexyl)-9H-purine (4b)
The title compound was prepared from compound 2b (250 mg, 0.993 mmol) and saturated aqueous Br2 (12 mL) in water (5 mL) as described for the synthesis of compound 4a above. EtOAc–Hexane (1:1) was used for flash chromatography to yield 4b (229 mg, 70%). Yellow solid; mp 181–183 °C; 1H-NMR (DMSO-d6, 600 MHz) δ 6.98 (br s, 2H, NH2), 4.35–4.22 (m, 1H, H-1 in c-hex), 2.46–2.27 (m, 2H, c-hex), 1.92–1.75 (m, 4H, c-hex), 1.74–1.62 (m, 1H, c-hex), 1.45–1.29 (m, 2H, c-hex), 1.28–1.12 (m, 1H, c-hex); 13C-NMR (DMSO-d6, 150 MHz) δ 159.1 (C, C-2), 154.5 (C, C-4), 148.2 (C, C-6), 128.6 (C, C-8), 123.7 (C, C-5), 57.6 (CH, C-1 in c-hex), 29.7 (CH2, C-3 and C-5 in c-hex), 25.3 (CH2, C-2 and C-6 in c-hex), 24.6 (CH2, C-4 in c-hex); HRESIMS m/z 330.0131 (calcd for C11H14BrClN5 + 1, 330.0121).
3.2.10. 2-Amino-8-bromo-6-chloro-9-(cyclopentyl)-9H-purine (4c)
The title compound was prepared from compound 2c (880 mg, 3.70 mmol) and sat. aq. Br2 (35 mL) in water (10 mL) as described for the synthesis of compound 4a above. EtOAc–Hexane (1:1) was used for flash chromatography to yield 4c (950 mg, 81%).Yellow solid; mp 172–174 °C; 1H-NMR (DMSO-d6, 600 MHz) δ 6.97 (s, 2H, NH2), 4.85–4.77 (m, 1H, H-1 in c-pent), 2.33–2.17 (m, 2H, c-pent), 2.11–1.87 (m, 4H, c-pent), 1.71–1.59 (m, 2H, c-pent); 13C-NMR (DMSO-d6, 150 MHz) δ 159.1 (C, C-2), 154.3 (C, C-4), 148.2 (C, C-6), 129.3 (C, C-8), 123.9 (C, C-5), 57.7 (CH, C-1 in c-pent), 29.7 (CH2, C-3 and C-4 in c-pent), 24.4 (CH2, C-2 and C-5 in c-pent); HREIMS m/z 314.9880 (calcd for C10H11BrClN5, 314.9886).
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3.2.11. 2-Amino-8-bromo-6-chloro-9-(cyclopent-2-enyl)-9H-purine (4d)
Method A: A solution of diisopropylamine (0.145 mL, 1.03 mmol) in dry THF (3 mL) was stirred at −78 °C under Ar. n-BuLi (0.536 mL, 1.00 mmol, 1.87 M in hexane) was added dropwise. After stirring for 30 min, a solution of compound 2d (118 mg, 0.500 mmol) in THF (1.5 mL) was added. After additional stirring for 1 h at −78 °C, a solution of CBrCl2CBrCl2 (326 mg, 1.00 mmol) in THF (1.5 mL) was added dropwise and the resulting mixture was stirred at −78 °C for 1 h, and then 10 min without cooling. Saturated aqueous NH4Cl (5 mL) was added and the resulting mixture was extracted with EtOAc (3 × 50 mL). The combined organic extracts were washed with brine (100 mL), dried (MgSO4), and evaporated in vacuo. The product was purified by flash chromatography on silica gel eluting with EtOAc–Hexane (1:1) to yield 4d (50 mg, 32%). Buff solid; mp 157–158 °C (dec.); 1H-NMR (DMSO-d6, 600 MHz) δ 6.95 (s, 2H, NH2), 6.15–6.13 (m, 1H, c-pent), 5.74–5.72 (m, 1H, c-pent), 5.69–5.60 (m, 1H, c-pent) 2.90–2.79 (m, 1H, c-pent), 2.48–2.36 (m, 2H, c-pent), 2.22–2.14 (m, 1H, c-pent); 13C-NMR (DMSO-d6, 150 MHz) δ 159.3 (C, C-2), 154.4 (C-4), 148.0 (C, C-6), 136.5 (CH, C-3 in c-pent), 128.0 (C, C-8), 127.7 (CH, C-2 in c-pent), 123.5 (C-5), 62.1 (CH, C-1 in c-pent), 32.0 (CH2, C-5 in c-pent), 27.9 (CH2, C-4 in c-pent); HREIMS m/z 312.9734 (calcd for C10H9BrClN5, 312.9730).
Method B: Compound 10 (64 mg, 0.26 mmol) was added to a cooled solution of cyclopent-2-en-1-ol (44 mg, 0.51 mmol) and PPh3 (135 mg, 0.515 mmol) in anhydrous THF (5 mL) under Ar. The resulting suspension was treated with diethyl azodicarboxylate (DEAD, 0.080 mL, 0.51 mmol) and the resulting mixture was stirred at ambient temperature for 1 h and at 70 °C for 15 h. The mixture was cooled, treated with brine (50 mL), and extracted with CH2Cl2 (3 × 50 mL). The combined organic layer was washed with water (10 mL), dried (Na2SO4), and evaporated in vacuo. The product was purified by flash chromatography on silica gel eluting with EtOAc–Hexane (3:7) to yield 4d (34 mg, 42%).
Method C: A solution of compound 10 (110 mg, 0.423 mmol) and NaH (16 mg, 0.67 mmol) in dry DMF (10 mL) was stirred at ambient temperature for 20 min under Ar. Pd(PPh3)4 (77 mg, 0.067 mmol) and cyclopent-2-en-1-yl acetate (84 mg, 0.66 mmol) were added, and the resulting mixture was stirred at 55 °C. After three days Pd(PPh3)4 (77 mg, 0.067 mmol) and cyclopent-2-en-1-yl acetate (84 mg, 0.66 mmol) were added. The reaction mixture was stirred for three more days and evaporated under in vacuo. The product was purified by flash chromatography on silica gel eluting with EtOAc–Hexane (gradient 50%–100% EtOAc) followed by MeOH–EtOAc (1:9) to yield 4d (41 mg, 29%).
3.2.12. 9-(Cyclohexylmethyl)-8-oxoguanine (5a)
A mixture of compound 4a (263 mg, 0.763 mmol), NaOAc (319 mg, 3.89 mmol), glacial AcOH (9 mL), and Ac2O (1.5 mL, 17 mmol) was stirred at reflux under N2 for 16 h, before the mixture was cooled and evaporated in vacuo. The residue was suspended in water (3 mL) and stirred at ambient temperature while the pH was adjusted to 13 by dropwise addition of 10M NaOH (aq). The resulting solution was refluxed for 20 min, cooled to 0 °C, and stirred while the pH was brought down to 7 by dropwise addition of 6M HCl (aqueous). The precipitate was collected and dried in vacuo. The product was purified by flash chromatography on silica gel eluting with MeOH–CH2Cl2 (1:4) to yield 5a (160 mg, 80%). Pinkish solid; mp 297–300 °C; 1H-NMR (DMSO-d6, 400 MHz) δ 10.57 (s, 1H, NH), 10.48 (s, 1H, NH), 6.43 (s, 2H, NH2), 3.42 (d, J = 7.4 Hz, 2H, NCH2), 1.85–1.71 (m, 1H, H-1 in c-hex), 1.69–1.48 (m, 5H, c-hex),
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1.19–1.06 (m, 3H, c-hex), 0.98–0.85 (m, 2H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 153.4 (C, C-6), 152.6 (C, C-8), 150.9 (C, C-2), 148.2 (C, C-4), 98.0 (C, C-5), 44.9 (CH2, NCH2), 36.3 (CH, C-1 in c-hex), 30.1 (CH2, C-3 and C-5 in in c-hex), 25.9 (CH2, C-4 in c-hex), 25.1 (CH2, C-2 and C-6 in c-hex); HREIMS m/z 263.1380 (calcd for C12H17N5O2, 263.1382).
3.2.13. 9-(Cyclohexyl)-8-oxoguanine (5b) and 2-Amino-6-chloro-9-cyclohexyl-7H-purin-8(9H)-one (6b)
The title compounds were prepared from compound 4b (186 mg, 0.563 mmol), NaOAc (231 mg, 2.81 mmol), glacial AcOH (7 mL), and Ac2O (1.20 mL, 12.7 mmol) as described for the synthesis of compound 5a above, except that the reflux time with NaOH was 4 h. MeOH–EtOAc (1:9) was used for flash chromatography to yield 5b (106 mg, 76%) and 6b (7 mg, 11%).
5b: colorless solid; mp 367–368 °C; 1H-NMR (DMSO-d6, 400 MHz) δ 10.57 (s, 1H, NH), 10.45 (s, 1H, NH), 6.37 (s, 2H, NH2), 4.04–3.91 (m, 1H, H-1 in c-hex), 2.28–2.11 (m, 2H, c-hex), 1.86–1.71 (m, 2H, c-hex), 1.69–1.52 (m, 3H, c-hex), 1.36–1.04 (m, 3H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 152.9 (C, C-6), 151.8 (C, C-8), 150.9 (C, C-2), 147.7 (C, C-4), 98.1 (C, C-5), 51.2 (CH, C-1 in c-hex), 29.5 (CH2, C-3 and C-5 in c-hex), 25.5 (CH2, C-2 and C-6 in c-hex), 24.8 (CH2, C-4 in c-hex); HREIMS m/z 249.1222 (calcd for C11H15N5O2, 249.1226).
6b: yellow solid mp 320–321 °C; 1H-NMR (DMSO-d6, 300 MHz) δ 11.20 (br s, 1H, NH), 6.54 (s, 2H, NH2), 4.12–3.98 (m, 1H, H-1 in c-hex), 2.27–2.12 (m, 2H, c-hex), 1.88–1.75 (m, 2H, c-hex), 1.73–1.58 (m, 3H, c-hex), 1.39–1.09 (m, 3H, c-hex); 13C-NMR (DMSO-d6, 75 MHz) δ 158.3 (C, C-8), 152.5 (C, C-4), 152.3 (C, C-2), 135.5 (C, C-6), 109.8 (C, C-5), 51.7 (CH, C-1 in c-hex), 29.1 (CH2, C-3 and C-5 in c-hex), 25.4 (CH2, C-2 and C-6 in c-hex), 24.8 (CH2,C-4 in c-hex); HREIMS m/z 267.0877 (calcd for C11H14ClN5O, 267.0887).
3.2.14. 9-(Cyclopentyl)-8-oxoguanine (5c) and 2-Amino-6-chloro-9-cyclopentyl-7H-purin-8(9H)-one (6c)
The title compounds were prepared from compound 4c (250 mg, 0.790 mmol), NaOAc (325 mg, 3.96 mmol), glacial AcOH (10 mL), and Ac2O (3.00 mL, 31.6 mmol) as described for the synthesis of compound 5a above, except that the refluxing time with NaOH was 6 h. MeOH–EtOAc (1:9) was used for flash chromatography to yield 5c (130 mg, 70%) and 6c (12 mg, 6%).
5c: colorless solid; mp 309–310 °C; 1H-NMR (DMSO-d6, 400 MHz) δ 10.58 (s, 1H, NH), 10.47 (s, 1H, NH), 6.35 (s, 2H, NH2), 4.46–4.42 (m, 1H, H-1 in c-pent), 2.17–2.01 (m, 2H, c-pent), 1.94–1.73 (m, 4H, c-pent), 1.63–1.50 (m, 2H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 152.9 (C, C-6), 151.9 (C, C-8), 150.9 (C, C-2), 147.8 (C, C-4), 98.2 (C, C-5), 51.8 (CH, C-1 in c-pent), 29.0 (CH2, C-2 and C-5 in c-pent), 24.3 (CH2, C-3 and C-4 in c-pent); HREIMS m/z 235.1067 (calcd for C10H13N5O2, 235.1069).
6c: colorless solid; mp 321–322 °C (dec.); 1H-NMR (DMSO-d6, 400 MHz) δ 11.20 (s, 1H, NH), 6.52 (s, 2H, NH2), 4.70–4.46 (m, 1H, c-pent), 2.20–2.01 (m, 2H, c-pent), 1.98–1.74 (m, 4H, c-pent), 1.68–1.50 (m, 2H, c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 158.3 (C, C-4), 152.6 (C, C-8), 152.2 (C, C-6), 135.5 (C, C-2), 109.9 (C, C-5), 52.1 (CH, C-1 in c-pent), 28.7 (CH2, C-2 and C-5 in c-pent), 24.3 (CH2, C-3 and C-4 in c-pent); HREIMS m/z 253.0727 (calcd for C10H12ClN5O, 253.0734).
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3.2.15. 9-(Cyclopent-2-enyl)-8-oxoguanine (5d) and 2-Amino-6-chloro-9-(cyclopent-2-enyl)-7H-purin-8(9H)-one (6d)
The title compounds were prepared from compound 4d (210 mg, 0.668 mmol), NaOAc (274 mg, 3.34 mmol), glacial AcOH (8 mL), and Ac2O (2.78 mL, 29.4 mmol) as described for the synthesis of compound 5a above, except that the refluxing time with NaOH was 30 h and the heating bath was kept at 160 °C in the first reaction step. Glacial AcOH was used for the final neutralization and EtOAc followed by MeOH–EtOAc (1:9) were used for flash chromatography to yield 5d (119 mg, 71%) and 6d (9 mg, 5%).
5d: colorless solid; mp 322–325 °C (dec.); 1H-NMR (DMSO-d6, 400 MHz) δ 10.60 (s, 1H, NH), 10.49 (s, 1H, NH), 6.34 (s, 2H, NH2), 5.99–5.96 (m, 1H, H-3 in c-pent), 5.62–5.59 (m, 1H, H-2 in c-pent), 5.30–5.21 (m, 1H, H-1 in c-pent), 2.79–2.63 (m, 1H, H-5a in c-pent), 2.40–2.26 (m, 1H, H-5b in c-pent), 2.24–2.13 (m, 1H, H-4a in c-pent), 2.13–2.03 (m, 1H, H-4b in c-pent); 13C-NMR (DMSO-d6, 100 MHz) δ 153.0 (C, C-6), 151.8 (C, C-8), 151.0 (C, C-2), 147.7 (C, C-4), 134.5 (CH, C-3 in c-pent), 129.0 (CH, C-2 in c-pent), 98.3 (C, C-5), 56.8 (CH, C-1 in c-pent), 31.8 (CH2, C-4 in c-pent), 27.4 (CH2, C-5 in c-pent); HREIMS m/z 233.0914 (calcd for C10H11N5O2, 233.0913).
6d: yellow solid; mp 310–310.5 °C; 1H-NMR (DMSO-d6, 300 MHz) δ 11.18 (s, 1H, NH), 6.48 (s, 2H, NH2), 6.06–6.02 (m, 1H, H-3 in c-pent), 5.65–5.61 (m, 1H, H-2 in c-pent), 5.37–5.27 (m, 1H, H-1 in c-pent), 2.84–2.69 (m, 1H, H-5a in c-pent), 2.43–2.05 (m, 3H, c-pent); 13C-NMR (DMSO-d6, 75 MHz) δ 158.3 (C, C-8), 152.4 (C, C-4), 152.1 (C, C-2), 135.4 (C, C-6), 135.3 (CH, C-3 in c-pent), 128.1 (CH, C-2 in c-pent), 109.9 (C, C-5), 57.2 (CH, C-1 in c-pent), 31.8 (CH2, C-5 in c-pent), 27.0 (CH2, C-4 in c-pent); HREIMS m/z 251.0568 (calcd for C10H10ClN5O, 251.0574).
3.2.16. N-[9-(Cyclohexylmethyl)-6-oxo-6,9-dihydro-1H-purin-2-yl]acetamide (7)
Method A: Br2 (33 mg, 0.21 mmol) was added slowly to a stirred solution of compound 2e (20 mg, 0.41 mmol) in CHCl3 (4 mL) and the resulting mixture was stirred for 6 h at ambient temperature. The reaction mixture was evaporated to dryness and the product was purified by flash chromatography on silica gel eluting with MeOH–EtOAc (1:19) to yield 7 (10 mg, 84%). Off-white solid; mp 271–273 °C (dec.); 1H-NMR (DMSO-d6, 400 MHz) δ 12.01 (s, 1H, N2H) 11.63 (s, 1H, NH), 7.95 (s, 1H, H-8), 3.90 (d, J = 7.4 Hz, 2H, NCH2), 2.17 (s, 3H, CH3), 1.88–1.74 (m, 1H, c-hex), 1.70–1.54 (m, 3H, c-hex), 1.53–1.44 (m, 2H, c-hex), 1.21–1.07 (m, 3H, c-hex), 1.01–0.87 (m, 2H, c-hex); 13C-NMR (DMSO-d6, 100 MHz) δ 173.5 (C, CON2), 154.9 (C, C-6), 148.8 (C, C-4), 147.6 (C, C-2), 140.2 (CH, C-8), 120.0 (C, C-5), 48.9 (CH2, NCH2), 37.4 (CH, C-1 in c-hex), 29.9 (CH2, C-3 and C-5 in c-hex), 25.8 (CH2, C-4 in c-hex), 25.0 (CH2, C-2 and C-6 in c-hex), 23.8 (CH3); HREIMS m/z 289.1534 (calcd for C14H19N5O2, 289.1539).
Method B: The title compound was prepared from compound 2e (20 mg, 0.41 mmol), diisopropylamine (0.012 mL, 0.83 mmol), n-BuLi (0.060 mL, 0.83 mmol, 1.4 M in hexane) and CBrCl2CBrCl2 (27 mg, 0.83 mmol) in THF (tot. vol. 3 mL) as described for the synthesis of compound 4d above, except that the reaction was stirred at −78 °C for 2 h after the addition of CBrCl2CBrCl2. The product was purified by flash chromatography as described above to yield 7 (7 mg, 59%).
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3.2.17. 8-Bromo-6-chloro-N,9-bis(tetrahydro-2H-pyran-2-yl)-9H-purin-2-amine (9)
The title compound was prepared from compound 8 (1.00 g, 2.96 mmol), diisopropylamine (0.84 mL, 5.9 mmol), n-BuLi (4.23 mL, 5.20 mmol, 1.4 M in hexane), and CBrCl2CBrCl2 (1.93 g, 5.92 mmol) in THF (tot. vol. 30 mL) as described for the synthesis of compound 4d above, except that the reaction was stirred at −78 °C for 2 h after the addition of CBrCl2CBrCl2. EtOAc–Hexane (1:1) was used for flash chromatography to yield 9 (987 mg, 80%). Colorless solid; mp 145–147 °C (dec.); 1H-NMR (DMSO-d6, 300 MHz) δ 8.23 (s, 1H, NH), 5.52 (dd, J = 11.0, 2.4 Hz, 1H, CH in THP), 5.13–5.02 (m, 1H, CH in THP), 4.10–3.97 (m, 1H, OCH2 in THP), 3.89–3.77 (m, 1H, OCH2 in THP), 3.69–3.56 (m, 1H, OCH2 in THP), 3.49–3.39 (m, 1H, OCH2 in THP), 3.14–2.90 (m, 1H, THP) 2.06–1.29 (m, 11H, CH2 in THP); 13C-NMR (DMSO-d6, 75 MHz) δ 157.2 (C, C-2), 154.2 (C, C-8), 148.3 (C, C-6), 129.5 (C, C-4), 124.3 (C, C-5), 84.4 (CH, N9-THP), 80.2 (CH, THP), 68.0 (CH2, OCH2 in THP), 65.7 (CH2, OCH2 in THP), 30.1, 27.6, 24.9, 24.5, 22.6 and 22.5 (all CH2, THP); HREIMS m/z 415.0417 (calcd for C15H19BrClN5O2, 415.0411).
3.2.18. 2-Amino-8-bromo-6-chloro-1H-purine (10)
A mixture compound 9 (150 mg, 0.360 mmol), 96% EtOH (10 mL) and 9.6 M HCl (0.5 mL), was stirred at ambient temperature for 30 min and neutralized by the addition of solid KHCO3. The resulting mixture was evaporated in vacuo and the product was isolated by flash chromatography on silica gel eluting with MeOH–CHCl3 (1:50:) to yield 10 (80 mg, 90%) as a yellow solid; mp >300 °C (dec.). 1H-NMR (DMSO-d6, 400 MHz) δ 13.65 (s, 1H, NH), 6.88 (s, 2H, NH2); 13C-NMR (DMSO-d6, 100 MHz) δ 159.7, 156.2, 147.1, 126.5, 124.0; HREIMS m/z 246.9257 (calcd for C5H3BrClN5, 246.9260). Spectral data were in good agreement with those reported before [48].
3.3. DNA Glycosylase Activity Assay
The enzyme OGG1 (residues12–327) was diluted to the desired concentration (60 pM) using a protein dilution buffer (15% glycerol, 1 mM EDTA, 25 mM HEPES pH 7.9, 1 mM DTT, 0.1 μg/μL BSA). Enzyme, compound 5 or 6 (0.2 mM), and 5′-32P end-labeled duplex DNA containing an 8-oxo-G/C base pair were mixed in a 10 μL reaction volume of 50 mM MOPS pH 7.5, 1 mM EDTA, 5% glycerol, and 1 mM DTT. The sequence of the damaged strand in the DNA substrate used is 5′-GCATGCCTGCA CGG-8oxoG-CATGGCCAGATCCCCGGGTACCGAG-3′, which was annealed with a complementary strand containing a C opposite 8oxoG. The reactions were incubated for 10 min at 37 °C, followed by addition of 2.5 μL 0.5 M NaOH and incubation for 20 min at 70 °C, in order to stop the reaction and ensure complete strand cleavage. Then 0.5 M HCl/0.25 M MOPS pH 7.5 (2.5 μL) was added to each sample to neutralize the pH. Formamide DNA loading buffer (15 μL) was added to the reaction mixtures and the samples were incubated at 95 °C for 5 min to denature the DNA. The reaction products were analyzed on 20% denaturing urea gels. The gels were transferred to 3M paper and dried at 80 °C for 45 min. The dry gels were placed in a storage phosphor screen overnight, and subsequently scanned on a Typhoon 9410 Variable Mode Image. ImageQuant TL Version 2003.02 (Amersham Biosciences, Piscataway, NJ, USA) was used to analyze the results. For human NTH1, the same procedure was followed, except that the DNA substrate contained a 5-hydroxyuracil/G base pair instead of the 8oxoG/C pair in the OGG1
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substrate. The concentration of NTH1 was 3 nM to make sure the activity in the assay was within the linear range. Compounds were screened at 0.5 mM concentration.
4. Conclusions
Synthetic routes to 8-oxoguanines have been examined. The best reaction sequence from chloroguanine 1a to the target compounds was found to be N-9 alkylation, C-8 bromination, and finally simultaneous hydrolysis of both halides. Bromination before N-alkylation should only be considered in cases where the N-substituent is not compatible with bromination conditions, since a bromide in the purine 8-position lowers the reactivity in N-alkylations. In most cases, alkylation with an alkyl halide in the presence of a base compared favorably to reactions under Mitsunobu conditions. 2-Amino-6-chloropurine (1a) turned out to be a superior guanine precursor compared to the O-carbamoylguanine 1b. The latter did not result in improved N-9 selectivity in the alkylation and was not compatible with standard reaction conditions for C-8 bromination.
Enzymatic assays show that partly hydrolyzed 6-chloro-8-oxopurines 6 are somewhat better OGG1 inhibitors than the 8-oxoguanines 5. However, an inhibitory effect was only observed when using at least 0.2 mM concentration of the compounds, suggesting that the R-group should be extended even further to make more interactions with the enzyme’s substrate recognition pocket. Further, testing of the same compounds at a 2.5-fold higher concentration against human NTH1, which is a structural homolog of OGG1, showed that the synthesized compounds do not inhibit NTH1 at 0.5 mM, except possibly for a weak effect for compound 6b. To develop these compounds into more potent inhibitors of OGG1, one possibility is to try compounds with more ribose-like R-groups. In the present study, the R-group contains a cyclic hydrocarbon only, and it would also be interesting to replace this with carbocyclic 2′-deoxyribose derivatives, as in antiviral drugs like abacavir and entecavir. In these nucleoside analog drugs, the R-group is not particularly larger than the R-group in our study, but it contains 5′ and/or 3′ hydroxyl groups. Since the structure of the OGG1 enzyme is known [7], molecular modeling will be included in the search for more potent OGG1 inhibitors in the future.
Supplementary Material
Supplementary materials can be accessed at: http://www.mdpi.com/1420-3049/20/09/15944/s1.
Acknowledgments
The Molecular Life Science program at the University of Oslo MLSUiO is gratefully acknowledged for a grant to Tushar R. Mahajan and the Research Council of Norway (RCN) for partial financing of the Bruker Avance instruments used in this study. Bjørn Dalhus is supported by the South-Eastern Norway Regional Health Authority (Grant No. 2014034) and Mari Eknes Ytre-Arne is supported by RCN (project No. 228563). Support from the Anders Jahres Foundation for Medical Research to BD is also acknowledged.
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Author Contributions
LLG and BD designed the research. TRM performed the synthetic organic chemistry, and MEYA and PSA the biological experiments. All authors contributed to writing the paper and read and approved the final manuscript.
Conflicts of Interest
The authors declare no conflict of interest.
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Sample Availability: Not available.
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