Solid-phase synthesis and structural characterisation of phosphoroselenolate-modified DNA: A backbone analogue which does not impose conformational bias and facilitates SAD X-ray crystallography

Article abstract of DOI:10.1039/c9sc04098f

Oligodeoxynucleotides incorporating internucleotide phosphoroselenolate linkages have been prepared under solid-phase synthesis conditions using dimer phosphoramidites. These dimers were constructed following the high yielding Michaelis-Arbuzov (M-A) reaction of nucleoside H-phosphonate derivatives with 5′-deoxythymidine-5′-selenocyanate and subsequent phosphitylation. Efficient coupling of the dimer phosphoramidites to solid-supported substrates was observed under both manual and automated conditions and required only minor modifications to the standard DNA synthesis cycle. In a further demonstration of the utility of M-A chemistry, the support-bound selenonucleoside was reacted with an H-phosphonate and then chain extended using phosphoramidite chemistry. Following initial unmasking of methyl-protected phosphoroselenolate diesters, pure oligodeoxynucleotides were isolated using standard deprotection and purification procedures and subsequently characterised by mass spectrometry and circular dichroism. The CD spectra of both modified and native duplexes derived from self-complementary sequences with A-form, B-form or mixed conformational preferences were essentially superimposable. These sequences were also used to study the effect of the modification upon duplex stability which showed context-dependent destabilisation (-0.4 to-3.1 °C per phosphoroselenolate) when introduced at the 5′-Termini of A-form or mixed duplexes or at juxtaposed central loci within a B-form duplex (-1.0 °C per modification). As found with other nucleic acids incorporating selenium, expeditious crystallisation of a modified decanucleotide A-form duplex was observed and the structure solved to a resolution of 1.45 ?. The DNA structure adjacent to the modification was not significantly perturbed. The phosphoroselenolate linkage was found to impart resistance to nuclease activity.

Full text of DOI:10.1039/c9sc04098f

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Solid-phase synthesis and structural
characterisation of phosphoroselenolate-modified
DNA: a backbone analogue which does not impose
conformational bias and facilitates SAD X-ray
crystallography†
Cite this: Chem. Sci., 2019, 10, 10948
All publication charges for this article
have been paid for by the Royal Society
of Chemistry
a
Patrick F. Conlon, a Olga Eguaogie, a Jordan J. Wilson, a Jamie S. T. Sweet,
Julian Steinhoegl, b Klaudia Englert,c Oliver G. A. Hancox, b Christopher J. Law,d
b
c
be
*
Sarah A. Allman,
James H. R. Tucker,
James P. Hall
a
*
and Joseph S. Vyle
Oligodeoxynucleotides incorporating internucleotide phosphoroselenolate linkages have been prepared
under solid-phase synthesis conditions using dimer phosphoramidites. These dimers were constructed
following the high yielding Michaelis–Arbuzov (M–A) reaction of nucleoside H-phosphonate derivatives
with 50-deoxythymidine-50-selenocyanate and subsequent phosphitylation. Efficient coupling of the
dimer phosphoramidites to solid-supported substrates was observed under both manual and automated
conditions and required only minor modifications to the standard DNA synthesis cycle. In a further
demonstration of the utility of M–A chemistry, the support-bound selenonucleoside was reacted with an
H-phosphonate and then chain extended using phosphoramidite chemistry. Following initial unmasking
of methyl-protected phosphoroselenolate diesters, pure oligodeoxynucleotides were isolated using
standard deprotection and purification procedures and subsequently characterised by mass
spectrometry and circular dichroism. The CD spectra of both modified and native duplexes derived from
self-complementary sequences with A-form, B-form or mixed conformational preferences were
essentially superimposable. These sequences were also used to study the effect of the modification
upon duplex stability which showed context-dependent destabilisation (ꢀ0.4 to ꢀ3.1 ꢁC per
phosphoroselenolate) when introduced at the 50-termini of A-form or mixed duplexes or at juxtaposed
central loci within a B-form duplex (ꢀ1.0 ꢁC per modification). As found with other nucleic acids
incorporating selenium, expeditious crystallisation of a modified decanucleotide A-form duplex was
observed and the structure solved to a resolution of 1.45 A˚. The DNA structure adjacent to the
modification was not significantly perturbed. The phosphoroselenolate linkage was found to impart
resistance to nuclease activity.
Received 16th August 2019
Accepted 11th October 2019
DOI: 10.1039/c9sc04098f
rsc.li/chemical-science
including the measurement3,4 and manipulation5,6 of gene
expression, DNA nanotechnology7,8 and data storage.9
Introduction
Whilst simple Watson–Crick base-pairing rules can be used
to programme both intramolecular and intermolecular
assembly of three-dimensional structures from simple
duplexes,10,11 the tertiary interactions of folded nucleic acids
derived from more complex assemblies and their recognition,
especially by nucleic acid binding proteins or small molecule
effectors, typically requires structural elucidation at the atomic
level.12,13 Characterisation of novel structural motifs including
such complexes by X-ray crystallography is augmented by the
introduction of an anomalous heavy atom scattering centre.14,15
Anomalous scattering of X-rays at the Se–K edge has been of
considerable value in facilitating structural biology studies of
proteins following the pioneering work of Hendrickson and co-
The automated solid-phase synthesis of oligonucleotides using
phosphoramidite chemistry1 (Fig. 1A) is a platform technology
which has been transformative in a wide range of applications2
aSchool of Chemistry and Chemical Engineering, Queen's University Belfast, David Keir
Building, Stranmillis Road, Belfast, BT9 5AG, UK. E-mail: j.vyle@qub.ac.uk
bReading School of Pharmacy, University of Reading, Whiteknights, Reading RG6 6AP,
UK. E-mail: james.hall@reading.ac.uk
cSchool of Chemistry, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK
dSchool of Biological Sciences, Queen's University Belfast, 15 Chlorine Gardens, Belfast
BT9 5AH, UK
eDiamond Light Source, Chilton, Didcot, Oxfordshire, OX11 0DE, UK
† Electronic supplementary information (ESI) available: PDB ID: 6S7D. See DOI:
10.1039/c9sc04098f
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Fig. 1 (A) General scheme for the solid-phase synthesis of oligonucleotides using phosphoramidite chemistry. (B) Examples of selenium-
modified nucleic acid analogues prepared using this chemistry.
workers using either selenomethionine16 or a high-affinity class” structures of proteins and nucleic acids, where new
ligand.17 The molecular biology techniques required to engi- structural folds or ligand-induced distortions exclude using
neer polypeptide sequences containing a mutated selenome- other approaches to structure solution such as molecular
thionine residue have now become routine and as this residue replacement (which relies upon prior knowledge of a structure
does not disrupt protein folding, it is widely used in MAD based upon its sequence). However, the true impact of using Se-
(multiple wavelength anomalous dispersion) or SAD (single SAD and Se-MAD may never fully be known as once the structure
wavelength anomalous diffraction) applications.18 Concurrent of a new fold or motif has been determined, this can then be
increased access to third generation synchrotron sources which used as the basis for solving future structures without requiring
generate more focussed, tunable X-rays of high brightness and anomalous data. SAD and MAD phasing is therefore a great
stability coupled with automation of the screening of crystal- facilitator of structural science and is used in drug development
lisation conditions has engendered a considerable upsurge in programmes worldwide, due to the expeditious ability to
protein crystal structure solution such that in the year 1990, 132 determine pharmacophore–target interactions. However, for
protein structures were deposited in the protein databank a structure to be biologically “valid” the incorporation of a heavy
whereas in 2018 this number had risen to 10 306. Of the 137 000 atom (such as Se or Br) into the crystal needs to have little or no
protein crystal structures in the PDB repository by the end of effect on the conformation or topology of the molecule, whilst
2018, over 10% had been solved using SAD or MAD methods. also not modifying the molecular arrangement of any potential
These methodologies are particularly important for “rst in binding sites. This is a particular problem when working with
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nucleic acids as the most commonly used nucleotide analogues, Subsequently, Xu and Kool exploited related chemistry for the
which contain heavy atom groups, either alter the topology of synthesis of oligomers incorporating this analogue following
the nucleic acid (shiing its conformation) or change the template-directed chemical ligation. However, consistent with
groups present in either the major or minor groove, both of the extreme sensitivity of phosphoroselenoate monoesters
which are binding sites for most DNA-targeting drugs.
towards ariel oxidation in aqueous solution,54 incomplete
Methodology for the synthesis of selenium-modied nucleic reaction was observed. In contrast, Michaelis–Arbuzov (M–A)
acid (Fig. 1B) sequences under solid-phase conditions was rst reaction of a nucleoside 30-H-phosphonate with a 50-selenocya-
described three decades ago.19 In this report, selenisation of nate
enabled
synthesis
of
a
fully-characterised
support-bound H-phosphonate precursors required treatment phosphoroselenolate-bridged dimer (TpSedT) with high
every 3–4 hours over three or four days and gave mixtures of the efficiency.49
diastereomeric phosphoroselenoates. Subsequently, more effi-
Herein, we describe the further developments to this meth-
cient selenisation protocols20,21 and stereoselective synthesis22 odology enabling the isolation of pure dinucleoside phosphor-
have been developed and this modication has enabled the oselenolate triesters in good to excellent yields, their
solution of refractory structures such as homo-DNA.23 However, subsequent phosphitylation and efficient introduction into
the sensitivity of intermediates during oligomerisation and oligodeoxynucleotides under solid-phase conditions. Further-
short half-life of the nal product (ca. 30 days under ambient more, we report a model DNA crystal structure incorporating
conditions in aqueous solution) has limited their application, this modication which evinces an A-form morphology with
especially in vivo. Likewise, nucleobase analogues which minimal perturbation of the backbone.
incorporate a selenone function can be accessed using concise
synthetic pathways24–28 but are sensitive towards oxidation,
hydrolysis and photochemical degradation.
A third class of selenium-modied oligonucleotides in which
selenoether functions have been introduced at the 20,29–32 40,33,34
Results and discussion
Preparation of dinucleotide phosphoramidites
or terminal 50-positions35 or appended to a nucleobase36 exhibit The protected M–A product from the synthesis of TpSedT49
greater resilience towards the conditions typically employed appeared to be an attractive target as efficient dimer coupling
during standard solid-phase oligonucleotide synthesis using has been reported during the solid-phase construction of
phosphoramidite chemistry although mitigation of side- oligonucleotides containing sulfur.55,56 Furthermore, this
reactions arising from selenoether oxidation is required37,38 strategy would avoid the preparation and use of intermediates
and replacement of an oxygen at a stereogenic centre provides containing a labile chalcogen–P(III) bond.57,58 We therefore
additional synthetic demands, e.g., up to 14-steps are required sought to optimise the synthesis and isolation of the protected
to prepare 40-seleno-40-deoxynucleosides from D-ribose.39,40 The dinucleoside phosphoroselenolate triester from the corre-
20-selenomethyl ether analogues (especially pyrimidine deriva- sponding selenocyanate.
tives) are considerably more accessible (requiring only six steps
50-Deoxythymidine-50-selenocyanate has been prepared on
to prepare on large scale)41 and can also promote crystallisation small scales in solution over 24 hours 59 or using liquid assisted
of modied oligomers.29,42–45 However, although the furanoside grinding over 9–11 hours.60,61 In order to prepare this material
ring of individual nucleosides modied in this fashion adopt more efficiently, we therefore examined the application of
a conformation typically found in B-form DNA,46 when incor- microwaves which has been reported to enhance the rate of S-
porated into oligodeoxynucleotide duplexes, the location of the selective alkylation of the thiocyanate anion.62 In the presence
20-selenomethyl function in the minor groove provides confor- of excess potassium selenocyanate, clean transformation of 50-
mational steering towards the formation of RNA-like A-form tosylthymidine (1) into the corresponding 50-selenocyanate (2)
duplexes. Although the epimeric arabino-20-selenomethyl was ꢁobserved within 90 minutes following irradiation (20 W,
analogue reverses this trend,31 full details of its preparation 100 C) of the stirred reaction mixture in acetonitrile (Scheme
have not yet been reported and the steric bulk of both isomers 1). As previously observed,60 quenching of the excess seleno-
can interfere with groove-binding interactions.46
cyanate with benzyl bromide greatly facilitated subsequent
Replacement of a bridging oxygen (30 or 50) within an inter- purication by silica gel chromatography and pure 50-deoxy-
nucleotide phosphate diester by selenium gives rise to phos- thymidine 50-selenocyanate (2) was isolated in 75% yield.
phoroselenolate analogues. These are achiral and potentially
During optimisation of both the reaction conditions and
more stable than the corresponding phosphoroselenoates but subsequent purication of the M–A product derived from 2 and
have been documented in only a very limited number of cyanoethyl-protected nucleoside 30-H-phosphonate diesters,
communications.47–49
several issues became apparent. Specically, the neutral sily-
Alkylation of the lead salt of O,O-diethylphosphoroselenoate lating agent, N,O-bis(trimethylsilyl)acetamide (BSA), which
was rst reported in 1911 50 and this strategy has been applied served both to render the selenonucleoside soluble in chloro-
to the preparation of internucleotide phosphoroselenolate form and enhance the rate of the M–A reaction also promoted
linkages using 50-halothymidine derivatives either in DMF47 or decyanoethylation of both the starting material and the
under aqueous conditions.48 In the former case, character- product. Furthermore, subsequent quenching of the reaction
isation of the dimer product was equivocal indicating that following addition of water resulted in detritylation in the
perhaps both O- and Se-alkylation had occurred.51–53 absence of a strong organic base but if such a base was used
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Scheme 1 Preparation of dinucleotide phosphoramidites 5a–d. Reagents and conditions: (a) (i) KSeCN (1.5 eq.), MeCN, MW power 20 W, 100 ꢁC,
90 min; (ii) BnBr (0.6 eq.), MeOH, rt, 60 min. (b) 3a–e (1.5 eq.), 2,6-lutidine (5 eq.), MeCN, rt, 30–60 min. (c) (i) ClP(OCE)NiPr2 (2 eq.), iPr2NEt (3 eq.),
DCM, rt, 45 min; (ii) MeOH (1.5 eq.), rt, 15 min. *in MeCN (4d) or DCM, ‡in DMSO-d6. Key. BasePG. AdePAc: 9-(N6-phenoxyacetyladeninyl). CytAc: 1-
(N4-acetylcytosinyl). GuaiBu: 9-(N2-isobutyrylguaninyl). Thy: 1-thyminyl. DMTr: 4,40-dimethoxytrityl. MMTr: 4-methoxytrityl CE: 2-cyanoethyl. p:
P(O)OMe. Ts: p-toluenesulfonyl.
during work-up, further decyanoethylation of the phosphor- (5a), dC (5b) or T (5d) and 50-monomethoxytrityl-protected dG (5c)
oselenolate was evident.63
in greater than 70% yields. During attempted processing of 4e
In order to ameliorate issues associated with b-elimination of (DMTrdGiBupSedT) under the same conditions, detritylation was
the cyanoethyl function, methyl-protection of the phosphate was evident and therefore further reaction to the corresponding
adopted. This provides greater resilience towards phosphate phosphoramidite was not attempted.
deprotection in the presence of stronger organic bases such as
Analysis of the dimer phosphoramidites by 31P NMR showed
triethylamine and has recently been utilised for the preparation of expected resonances associated with two separate diastereo-
di- and trinucleotide phosphoramidites.63–65 The methyl-protected meric P(III) and P(V) phosphorus centres (e.g., Fig. 2). Low levels
30-H-phosphonates (3a–e) were prepared from the corresponding (<4%) of putative H-phosphonate derivatives were also apparent
phosphoramidites under modied literature conditions.66 and as this can be associated with autocatalytic degradation,68,69
Following extractive work-up, the materials were pure by 31P NMR. the stability of a dimer phosphoramidite in solution was
However, in our hands, these H-phosphonates were less stable examined by 31P NMR. At 4 ꢁC, 5a (0.1 M in acetonitrile)
towards dephosphonylation than those derived from the
cyanoethyl-protected precursors and low levels of the nucleoside
congeners were identied by TLC as contaminants. Thus, once
isolated, the solids were stored at ꢀ20 ꢁC and used within 24
hours. To accommodate the presence of these impurities, excess
H-phosphonate (1.5 eq.) was used in the M–A reaction. In
a further change to the previous published procedure, acetoni-
trile was used as solvent and in the presence of 2,6-lutidine,
nucleoside selenocyanate 2 could be dissolved in this mixture
under gentle heating without silylation. Addition of this solution
to thymidine H-phosphonate (3d) in acetonitrile at room
temperature gave complete and clean M–A reaction within 30
minutes. We note that these conditions contrast with those
typically employed to promote such reactions with chalcogen
electrophiles following conversion of the H-phosphonate into the
corresponding phosphite tautomer in the presence of a silylating
agent or a strong base. The protected dimers (4a–e) were isolated
as mixtures of diastereoisomers following silica gel column
chromatography. Phosphitylation of the dimers under modied
Fig. 2 31P{1H} NMR of phosphoramidite 5a (prepared from DMTrdA-
literature conditions67 enabled isolation of the corresponding
PAcpSedT – 4a) in MeCN with (inset) expansions of resonances asso-
ciated with P(III) and P(V) nuclei.
phosphoramidites derived from 50-dimethoxytrityl-protected dA
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remained untransformed over 24 days. A separately prepared
solution of the same composition was stored on the DNA syn-
thesiser at ambient temperature. Aer 24 days, solid deposition
was observed and 31P NMR analysis of the combined materials
(solid and liquid) showed over 80% degradation of the phos-
phoramidite although no transformation of the phosphor-
oselenolate function was evident.
Preparation of phosphoroselenolate-modied
oligodeoxynucleotides under solid-phase conditions via
coupling of dimer phosphoramidites
Optimisation of solid-phase synthesis conditions was per-
formed with the dimer phosphoramidite 5d (derived from
DMTrdTpSedT) using manual syringe addition of all reagents
and acetonitrile washes to prepare a model homothymidine-
derived pentamer incorporating two phosphoroselenolate
linkages (ODN 1 – Table 1). Under optimised conditions,
quantitative coupling (by trityl release) of 5d in the presence of
the activator 5-S-benzylthiotetrazole was observed over ve
minutes with subsequent oxidation using 0.02 M iodine.
Duplicate, one mmol-scale syntheses of the trityl-on, model
pentamer were performed. Removal of phosphate diester pro-
tecting groups and cleavage from the controlled pore glass
(CPG) support required a two-step procedure. Initial demethy-
lation of the phosphoroselenolate triesters was effected
following treatment with a solution of 0.2 M sodium dieth-
yldithiocarbamate (NaDEC) in acetonitrile for 30 minutes at
room temperature. This method was found to be equally effi-
cient in a side-by-side comparison with standard literature
conditions using 1 : 2 : 2 (v/v/v) thiophenol/triethylamine/
dioxane.55 Excess reagents were removed following washing
with acetonitrile, the CPG dried and suspended in 1 : 1 (v/v)
35% (w/v) NH3 (aq)/40% (w/v) MeNH2 (aq) (AMA) at room
temperature for 60 minutes. Following removal of volatile
amines, the crude tritylated oligomer ODN 1 was puried using
reversed phase HPLC. Finally, treatment with aqueous acetic
acid under standard conditions effected cleavage of the 50-DMTr
function and following desalting, pure ODN 1 was isolated in
33% yield. Analysis of ODN 1 by 31P NMR (Fig. 3) showed four
resonances associated with the two internucleotide phosphate
diesters (dP ¼ ꢀ1.19, ꢀ1.25) and two internucleotide phos-
phoroselenolate diesters (dP ¼ 10.56, 10.48).
Fig. 3 31P{1H} NMR of ODN 1 in 60 : 40 (v/v) H2O : D2O and (inset)
ESI-mass spectrum showing isotope pattern associated with molec-
ular ion.
393 Hz and 390 Hz)70 although somewhat weaker than that
found in the corresponding triester.
ODN 1 was further characterised using ESI-MS (Fig. 3 – inset)
which gave a molecular ion peak ([M ꢀ H]ꢀ) displaying the ex-
pected isotope distribution pattern.
In order to further examine the scope of the manual dimer
phosphoramidite coupling and deprotection protocol devel-
oped for ODN 1, a series of self-complementary 10-mer
sequences designed to form A-form duplexes (ODN 2–5) were
prepared. Chain extension of support-bound octanucleotides
using phosphoramidite 5d was performed using the same cycle
as previously described. However, less efficient couplings were
observed using 5a–c and therefore an extended double
coupling71 was employed with these phosphoramidites.
Following oxidation, the columns were washed with aceto-
nitrile and dried under argon. Deprotection using NaDEC fol-
lowed
by
AMA
liberated
the
crude,
tritylated
oligodeoxynucleotides which were puried by RP-HPLC and
reduced in vacuo. Detritylation of the puried oligomers was
accomplished under literature conditions using an extended
treatment to fully remove the monomethoxytrityl function from
ODN 4.
The modied Dickerson–Drew dodecamer72 sequence ODN 6
was prepared under automated conditions using the
phosphoroselenolate-linked dimer phosphoramidite 5a which
was incorporated in place of the central AT motif. In addition to
the extended coupling of 5a, the standard cycle was modied to
Satellite peaks associated with 31P–77Se coupling were
consistent with a phosphorus–selenium single bond (1JPSe
¼
Table 1 Phosphoroselenolate-modified oligodeoxynucleotide sequences prepared on solid support using dinucleotides phosphoramidites 5a–d
Phosphoramidite (coupling time)
Oligodeoxynucleotide sequence 50/30
Yielda (%)
Mol. ionb (calcd)
5d (5 min)
ODN 1 d(TSeTTSeTT)
657 nmol (33%)
541 nmol (27%)
251 nmol (13%)
484 nmol (24%)
337 nmol (17%)
198 nmol (20%)*
1585.070 (1585.112)
3090.370 (3090.472)
3091.365 (3091.468)
3091.351 (3091.468)
3090.360 (3090.472)
3708.450 (3708.450)
5a (2 ꢂ 7.5 min)
5b (2 ꢂ 7.5 min)
5c (2 ꢂ 7.5 min)
5d (5 min)
ODN 2 d(ASeTCCCGGGAT)
ODN 3 d(CSeTCCCGGGAG)
ODN 4 d(GSeTCCCGGGAC)
ODN 5 d(TSeTCCCGGGAA)
ODN 6 d(CGCGAASeTTCGCG)
5a (2 ꢂ 7.5 min)
a
Determined using 3260 nm of the corresponding native sequence on a 2 mmol scale (* from 1 mmol). b m/z [M ꢀ H]ꢀ
(
80Se).
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include a wash using 10% (v/v) pyridine/MeCN following the precipitated DCU, the mixture of activated esters reacted with
oxidation step. In the absence of this wash, the CPG retained amino-functionalised controlled pore glass. The CPG-bound
iodine residues which were found to engender lower yields of selenonucleoside (7) thus obtained was reacted with the
the modied oligomers. The subsequent ve monomer nucleoside H-phosphonate 3f. Aer 60 minutes, trityl assay
coupling cycles were performed under standard conditions and indicated that the support loading (39 mmol gꢀ1) was consistent
gave efficient nucleotide incorporation according to trityl yields. with that typically used in solid-phase DNA synthesis indicating
Once complete, the support-bound dodecamer was that efficient M–A reaction had occurred.
treated with a solution of 150 mM dithiothreitol (DTT) in 1 : 1 (v/v)
Using this material (1.75 mmol), chain extension was per-
H2O : absolute ethanol to remove potential oxidised selenium formed under manual conditions using standard monomer
species37,45 and further demethylated using NaDEC as previously phosphoramidites using the cycle developed for ODN 1.
described. Further deprotection with AMA enabled the pure trityl- Subsequent deprotection with AMA liberated the crude trity-
on oligomer to be isolated following RP-HPLC purication. This lated tetramer which was puried by RP-HPLC, detritylated and
material was then detritylated and desalted under standard desalted to afford pure ODN 7.
conditions to afford pure ODN 6 in 20% yield. Under the same
conditions the corresponding native sequence was isolated in
45% yield. All selenium-containing oligodeoxynucleotides were
Enzymatic digestion
Further conrmation of the integrity of the internucleotide
phosphoroselenolate linkage during both coupling and down-
stream processing was provided by enzymatic digestion of both
characterised by mass spectrometry in which the m/z ratios
exhibited the expected isotope distribution patterns and did not
show any peaks associated with selenoxide formation.
ODN
5 and the corresponding native sequence. Using
commercially-sourced snake venom with both phosphodies-
terase and phosphatase activities, complete digestion of the
native 10-mer to the corresponding nucleosides was found
Preparation of a model phosphoroselenolate-modied
oligodeoxynucleotide under solid-phase conditions via
Michaelis–Arbuzov chemistry
ꢁ
within eight hours at 37 C (Fig. S2†). Under the same condi-
In order to demonstrate the general utility of the M–A reaction
for constructing internucleotide phosphoroselenolates under
solid-phase conditions, this chemistry was used to prepare
a tetranucleotide sequence which, in its native form, crystallises
in the four-stranded, i-motif conformation.73,74 Thus, ODN 7 was
prepared from the corresponding support-bound nucleoside
selenocyanate (Scheme 2). Succinylation of selenonucleoside 2
under standard conditions75 gave complete consumption of
starting material. Following extraction with cold citric acid, the
succinyl ester (6) was isolated contaminated with the corre-
sponding symmetrical diselenide (30 mol%). This crude mate-
rial was treated with excess condensing agent (DCC) in the
presence of 4-nitrophenol76 and following removal of
tions, ODN 5 liberated the nucleosides dA, dC and dG in the
expected ratios but less than 1 mol% of T (tR ¼ 18.1 min) was
observed (Fig. 4). However, a single major peak with signi-
cantly longer retention time (tR ¼ 38.7 min) was identied as the
dimer TpSedT following coinjection with an authentic sample.49
Previously, Stec and coworkers examined nuclease digestion of
putative TpSedT and described using ten times the quantity of
enzyme required to effect cleavage of the corresponding phos-
phorothioate substrate. During this treatment, formation of the
corresponding diselenide; (SedT)2, was observed and this
compound is assumed to be the origin of a novel hydrophobic
material (tR ¼ 45.1 min) which was also apparent during
extended treatment of pure TpSedT with snake venom. Early
studies on the digestion of dimers incorporating a 50-thio-
thymidylate (e.g., TpSdT) also reported slow cleavage activi-
ties77,78 although Xu and Kool subsequently described snake
venom digestion of a 20-mer containing a single internal
modication which was shown to proceed at essentially the
Scheme 2 Preparation of ODN 7. Reagents and conditions: (a) suc-
cinic anhydride (1 eq.), DMAP (0.8 eq.), pyridine, rt, overnight; (b) (i) p-
nitrophenol (1 eq.), DCC (2.5 eq.), DCM, rt, 2 h; (ii) amino SynBase 500/
110 CPG (500 mg), DMF, Et3N (4 eq.), rt, 5 h. (c) 3f, MeCN, rt, 1 h; (d)
solid-phase DNA synthesis using manual syringe addition. (e) (i) AMA,
rt, 2 h. (ii) 80% AcOH, rt, 1 h. See ESI† for full details.
Fig. 4 RP-HPLC chromatogram of the snake venom digest of ODN 5
(after 8 hours at 37 ꢁC). For full conditions see ESI.†
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same rate as the unmodied sequence and cleaved the internal
phosphorothiolate linkage.79
Although the scissile bond (30O–P) is not modied in the
phosphoroselenolate analogues examined in this study, it is
assumed that distortion in the phosphoryl moiety arising out of
the extended bond lengths and compressed bond angle asso-
ciated with the P–Se–C50 bridge (vide infra) inhibits productive
binding of the dimer.
UV thermal denaturation studies
The effect of the phosphoroselenolate linkage upon duplex
stability was investigated by comparing thermal denaturation of
the modied oligomers with the corresponding native
sequences (Table 2). Sequence-dependent destabilisation of
duplexes designed to exhibit A-form conformations was
observed following phosphoroselenolate substitution of the 50-
terminal phosphodiester linkage (entries 1–4). The largest
reductions in melting temperatures were observed for
sequences in which the native oligomers had the greatest
stabilities by virtue of the presence of a terminal C–G (entry 2) or
G–C (entry 3) base pair and in these circumstances, DTm values
of ꢀ6.2 ꢁC and ꢀ4.9 ꢁC (respectively) were observed. In contrast,
introduction of a phosphoroselenolate-linkage downstream
from a terminal A–T base pair (entry 1) had a minor effect upon
Fig.
5
CD spectra comparing the conformation of phosphor-
oselenolate-modified sequences ( ) and the corresponding native
(all phosphodiester) sequences ( ). (A) ODN 4; (B) ODN 6.
ꢁ
duplex stability (DTm ¼ ꢀ0.7 C) although the effect upon the
ꢁ
T–A (entry 4) pairing was more signicant (DTm ¼ ꢀ4.0 C).
The effect of introducing juxtaposed phosphoroselenolate
bridges into the more extended backbone typical of B-form
helices was tested using the Dickerson–Drew dodecamer
duplexes displayed a high level of congruence, with only small
changes in band intensity observed (Fig. S5–S9†). For example,
conservation of the mixed A/B conformation adopted in solu-
tion by ODN 4 (Fig. 5A) highlights that the modication is not
perturbing or directing the conformation of the duplex in
a boarderline case.
ꢁ
sequence ODN 6 (entry 5). A decrease in stability of 1.9 C was
observed in comparison with the native oligomer. Xu and Kool
reported the effect upon duplex stability of substituting a single
internal phosphate diester-linkage within a 17-mer (B-form)
sequence with either
a phosphorothiolate or phosphor-
The advantage of the phosphoroselenolate modication over
ribose modications, such as replacement of O with Se in the ring,
which typically leads to formation of an A-form duplex,33 is clear. It
should also be noted that both the native and modied dodeca-
mer (Fig. 5B) exhibited the classical B-form again illustrating that
the modication has had little effect on the solution conforma-
tion of the oligonucleotide. In all cases the incorporation of the Se
into the DNA backbone means that the environment in the major
and minor grooves is unaffected, unlike with modications such
as 5-bromo-dU or most selenium modied DNA. Whilst some of
these modications (such as a 20-SeMe analogues)42 may not
signicantly affect the native conformation if this is A-form DNA,
oselenolate analogue.48 Duplexes formed between these
chalcogen-modied oligomers and the native compliment dis-
played DTm values of ꢀ3.0 ꢁC (Se) and ꢀ2.0 ꢁC (S). These values
are consistent with those obtained in the current study in which
two modications are introduced per duplex.
Solution phase conformation studies
The effects of introducing the phosphoroselenolate modica-
tion upon the conformations of self-complementary duplex
DNA sequences were investigated using circular dichroism. The
spectral shape and peak positions of native and modied
Table 2 Melting temperatures (Tm) of self-complementary phosphoroselenolate-modified and native oligodeoxynucleotide duplexes
a
Entry
Sequence 50/30
X ¼
Tm/ꢁC
X ¼
Tm/ꢁC
DTm /ꢁC
1
2
3
4
5
d(AXCCCGGGAT)
d(CXCCCGGGAG)
d(GXCCCGGGAC)
d(TXCCCGGGAA)
d(CGCGAAXTCGCG)
SedT (ODN 2)
SedT (ODN 3)
SedT (ODN 4)
SedT (ODN 5)
SedT (ODN 6)
46.9
48.1
50.7
43.0
55.9
T
T
T
T
T
47.6
54.3
55.6
47.0
57.8
ꢀ0.7
ꢀ6.2
ꢀ4.9
ꢀ4.0
ꢀ1.9
a
DTm ¼ Tm(X ¼ SedT) ꢀ Tm(X ¼ T). Conditions: ssDNA (10 mM), NaPi (10 mM, pH 7.0), NaCl (100 mM).
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not the case, with the nucleotide possessing a C30-endo sugar
pucker, consistent with that of A-DNA.
The nucleotide 30 to the modied base also possesses a C30-
endo pucker. The base 50 to the modication site possesses
a non-standard pucker, C20-exo, which is attributed to the
proximity of a neighbouring strand, forming a crystal contact
within the lattice. This structure therefore conrms that the
introduction of the modication does not signicantly perturb
the solid-state conformation of the duplex.
Conclusions
For the rst time, a phosphoroselenolate-modied oligomer
has been prepared de novo using phosphoramidite chemistry
with only minor modication to the standard solid-phase DNA
synthesis cycle. All modied oligonucleotides were charac-
terised by electrospray and/or MALDI mass spectrometry which
gave molecular ion masses (with appropriate isotope distribu-
tion patterns) consistent with the single-stranded sequence and
if self-complimentary, the corresponding duplex. No M+16
peaks were apparent in these spectra suggesting that either the
oxidation conditions did not lead to oxygen transfer to selenium
Fig. 6 (A) (left) The DNA structure reported here (ODN 4) and (right)
a
modification (cyan) compared with (C) the naturally occurring back-
standard A-form DNA 10-mer.81 (B) The phosphoroselenoate
bone at the terminal AC step. The chains are coloured according to or that downstream processing of such modied oligomers
leads to their removal. This was further conrmed by 31P NMR
analysis of a double-labelled model pentamer.
atom type, with carbon in red, green, cyan or magenta (as a function of
DNA chain), nitrogen in blue, oxygen in red, selenium in cyan, phos-
phorus in orange and hydrogen in white. PDB ID: 6S7D.†
Unlike previous generations of DNA analogues in which
selenium is introduced within the sugar moiety, the phos-
phoroselenolate modication described here is readily acces-
sible requiring only four high yielding steps and does not
require handling highly air-sensitive materials. Furthermore,
the addition of groups into possible binding sites could affect the
validity of drug–DNA binding studies, which is not an issue with
a phosphoroselenolate modication.
the furanose pseudorotation angles of the most common
modication, 20-SeMe, typically results in conformational
steering towards an A-form duplex. Using the related 20-SMe
Crystal structure
analogue, such conformational restrictions were not observed
To conrm that the structure of the nucleic acid is not signi- and both A- and B-DNA duplex crystal structures were solved82,83
cantly perturbed by the introduction of a phosphoroselenolate although longer collection times (and concomitant DNA
modication, crystallisation trials of oligodeoxynucleotide damage) are associated with the weaker anomalous scattering
sequences containing the group were performed. This resulted from sulfur at the CuKa wavelength. Furthermore, both
in the crystal structure of d(GSeTCCCGGGAC) – ODN 4 – being analogues interfere with minor groove binding interactions.
obtained, which allows for visualisation of the modication at Finally, the expeditious crystallisation processes observed using
˚
1.45 A resolution.
other selenium-modied nucleic acids was maintained with the
The structure shows that the overall conformation of the phosphoroselenolate.
duplex is that of A-DNA (Fig. 6A). The average twist of the helix is
Selenium-SAD has recently been applied to the de novo
31.8ꢁ which compares favourably to a standard value of 32.7ꢁ. phasing of data generated from an X-ray-free electron laser
However, at the modication site the T*–A base pair has a twist including serial femtosecond crystallography84,85 and therefore
of ca. 41ꢁ, which can be attributed to the increased bond lengths offers the potential to observe conformational changes within
0
˚
˚
from P–Se (2.25 A) and Se–C5 (1.94 A) compared to the oxygen- nucleic acids during folding, recognition and processing.
0
0
0
˚
˚
containing equivalent (P–O5 1.60 A, O5 –C5 1.42 A).
The resistance of the internucleoside phosphoroselenolate
Apart from this slight increase in bond lengths, and subse- linkage towards the exonuclease activity of snake venom offers
quent increase in helical twist angle, the base pair containing the potential to both facilitate in vivo activity of therapeutic
the phosphoroselenolate modication is otherwise consistent nucleic acid sequences86 and to probe biochemical processes in
with a standard A-DNA base pair (Fig. 6B and C), with param- the same fashion as have phosphorothiolate56,87 and we
eters such as roll, tilt, rise and slide all within the normal range envisage that the ability to label an oligonucleotide backbone
for this conformation (analysis performed using W3DNA).80 using 100% 77Se may provide a valuable tool for investigating
Whilst it could be expected that the increased twist or bond the biological processing of such materials.88 Towards these
lengths could affect the sugar pucker of the nucleotide, this is future goals we are currently investigating the application of
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this chemistry for the preparation of phosphoroselenolate- 14 W. A. Hendrickson, Q. Rev. Biophys., 2014, 47, 49–93.
linkages derived from other selenonucleoside analogues.
15 B. T. Wimberly, D. E. Brodersen, W. M. Clemons,
R. J. Morgan-Warren, A. P. Carter, C. Vonrheln, T. Hartsch
and V. Ramakrishnan, Nature, 2000, 407, 327–339.
16 W. A. Hendrickson, J. R. Horton and D. M. LeMaster, EMBO
J., 1990, 9, 1665–1672.
Conflicts of interest
There are no conicts to declare.
¨
17 W. A. Hendrickson, A. Pahler, J. L. Smith, Y. Satow,
E. A. Merritt and R. P. Phizackerley, Proc. Natl. Acad. Sci. U.
S. A., 1989, 86, 2190–2194.
Acknowledgements
This work was funded by: DEL (P. F. C.); EPSRC (EP/R511602/1, 18 X. D. Su, H. Zhang, T. C. Terwilliger, A. Liljas, J. Xiao and
EP/G007578/1 – O. E.); University of Reading, School of Phar-
Y. Dong, Crystallogr. Rev., 2015, 21, 122–153.
macy (O. G. A. H.); ERASMUS+ (J. S.); BBSRC equipment grant 19 K. Mori, C. Boiziau, C. Cazenave, M. Matsukura,
´
BB/M025624/1 “Next Generation DNA Synthesis” (J. H. R. T.)
C. Subasinghe, J. S. Cohen, S. Broder, J. J. Toulme and
and by the authors (J. S. V., O. E.). J. P. H. gratefully acknowl-
C. A. Stein, Nucleic Acids Res., 1989, 17, 8207–8219.
edges the provision of beamtime at Diamond Light Source 20 C. J. Wilds, R. Pattanayek, C. Pan, Z. Wawrzak and M. Egli, J.
(MX20077-1 and SM20214-1). We thank Rick Cosstick/James Am. Chem. Soc., 2002, 124, 14910–14916.
Gaynor (Liverpool) for their kind support in automated solid- 21 K. Tram, X. Wang and H. Yan, Org. Lett., 2007, 9, 5103–5106.
phase synthesis. Cameron Thorpe (Oxford) and Sabine Muller/ 22 P. Guga, A. Maciaszek and W. J. Stec, Org. Lett., 2005, 7,
Bettina Appel (Greifswald) provided constructive discussions
on the phosphitylation procedure. Andrew Cummings (Q. U. B.) 23 M. Egli, P. Lubini and P. S. Pallan, Chem. Soc. Rev., 2007, 36,
provided technical support for microwave reactions. Catherine 31–45.
McKeen (LGC Link) provided useful discussions relating to 24 J. Salon, J. Sheng, J. Jiang, G. Chen, J. Caton-Williams and
support bound octamers. We acknowledge Darren Baskerville Z. Huang, J. Am. Chem. Soc., 2007, 129, 4862–4863.
and Conor McGrann (Q. U. B.) and The Centre for Chemical and 25 J. Salon, J. Jiang, J. Sheng, O. O. Gerlits and Z. Huang, Nucleic
Materials Analysis in the School of Chemistry at the University Acids Res., 2008, 36, 7009–7018.
of Birmingham for running mass spectra. Richard Murphy 26 A. E. A. Hassan, J. Sheng, W. Zhang and Z. Huang, J. Am.
provided NMR technical support. J. S., O. G. A. H., S. A. A. & J. P.
Chem. Soc., 2010, 132, 2120–2121.
H. would like to thank the Chemical Analysis Facility at the 27 T. Habuchi, T. Yamaguchi, H. Aoyama, M. Horiba, K. R. Ito
3901–3904.
University of Reading for access to instruments.
and S. Obika, J. Org. Chem., 2019, 84, 1430–1439.
28 C. Liczner, V. Grenier and C. J. Wilds, Tetrahedron Lett., 2018,
59, 38–41.
29 Q. Du, N. Carrasco, M. Teplova, C. J. Wilds, M. Egli and
Z. Huang, J. Am. Chem. Soc., 2002, 124, 24–25.
Notes and references
1 M. H. Caruthers, Science, 1985, 230, 281–285.
¨
2 S. Kosuri and G. M. Church, Nat. Methods, 2014, 11, 499–507. 30 C. Hobartner and R. Micura, J. Am. Chem. Soc., 2004, 126,
3 R. K. Saiki, S. Scharf, F. Faloona, K. B. Mullis, G. T. Horn,
1141–1149.
H. A. Erlich and N. Arnheim, Science, 1985, 230, 1350–1354. 31 W. Zhang, J. W. Szostak and Z. Huang, Front. Chem. Sci. Eng.,
4 C. A. Heid, J. Stevens, K. J. Livak and P. M. Williams, Genome
Res., 1996, 6, 986–994.
5 M. Smith, Annu. Rev. Genet., 1985, 19, 423–462.
2016, 10, 196–202.
32 K. Morihiro, T. Kodama, Kentefu, Y. Moai, R. N. Veedu and
S. Obika, Angew. Chem., Int. Ed., 2013, 52, 5074–5078.
6 M. Jinek, K. Chylinski, I. Fonfara, M. Hauer, J. A. Doudna and 33 J. K. Watts, B. D. Johnston, K. Jayakanthan, A. S. Wahba,
E. Charpentier, Science, 2012, 337, 816–821.
7 H. Shen, Y. Wang, J. Wang, Z. Li and Q. Yuan, ACS Appl.
Mater. Interfaces, 2019, 11, 13859–13873.
8 N. C. Seeman and H. F. Sleiman, Nat. Rev. Mater., 2017, 3,
17068.
9 L. Ceze, J. Nivala and K. Strauss, Nat. Rev. Genet., 2019, 20,
456–466.
10 P. W. K. Rothemund, Nature, 2006, 440, 297–302.
B. M. Pinto and M. J. Damha, J. Am. Chem. Soc., 2008, 130,
8578–8579.
34 Y. Inagaki, N. Minakawa and A. Matsuda, Nucleic Acids Symp.
Ser., 2008, 52, 329–330.
35 N. Carrasco, D. Ginsburg, Q. Du and Z. Huang, Nucleosides,
Nucleotides Nucleic Acids, 2001, 20, 1723–1734.
36 A. Nuthanakanti, M. A. Boerneke, T. Hermann and
S. G. Srivatsan, Angew. Chem., Int. Ed., 2017, 56, 2640–2644.
¨
11 E. S. Andersen, M. Dong, M. M. Nielsen, K. Jahn, 37 C. Hobartner, R. Rieder, C. Kreutz, B. Puffer, K. Lang,
R. Subramani, W. Mamdouh, M. M. Golas, B. Sander,
H. Stark, C. L. P. Oliveira, J. S. Pedersen, V. Birkedal,
A. Polonskaia, A. Serganov and R. Micura, J. Am. Chem.
Soc., 2005, 127, 12035–12045.
F. Besenbacher, K. V. Gothelf and J. Kjems, Nature, 2009, 38 N. Tarashima, K. Hayashi, M. Terasaki, H. Taniike,
459, 73–76.
Y. Inagaki, K. Hirose, K. Furukawa, A. Matsuda and
N. Minakawa, Org. Lett., 2014, 16, 4710–4713.
12 J. Sheng, J. Gan and Z. Huang, Med. Res. Rev., 2013, 33, 1119–
1173.
39 H. Taniike, Y. Inagaki, A. Matsuda and N. Minakawa,
Tetrahedron, 2011, 67, 7977–7982.
13 R. Satange, C. K. Chang and M. H. Hou, Nucleic Acids Res.,
2018, 46, 6416–6434.
10956 | Chem. Sci., 2019, 10, 10948–10957
This journal is © The Royal Society of Chemistry 2019

View Article Online
Edge Article
Chemical Science
40 K. Ishii, N. Saito-Tarashima, M. Ota, S. Yamamoto, 66 J. S. Vyle, N. H. Williams and J. A. Grasby, Tetrahedron Lett.,
Y. Okamoto, Y. Tanaka and N. Minakawa, Tetrahedron,
2016, 72, 6589–6594.
1998, 39, 7975–7978.
67 C. Thorpe, S. Epple, B. Woods, A. H. El-Sagheer and
T. Brown, Org. Biomol. Chem., 2019, 17, 5341–5348.
¨
41 R. Micura, C. Hobartner, R. Rieder, C. Kreutz, B. Puffer,
K. Lang and H. Moroder, Curr. Protoc. Nucleic Acid Chem., 68 J. S. Hargreaves, R. Kaiser and P. K. Wolber, Nucleosides,
2006, 27, 1.15.1–1.15.34. Nucleotides Nucleic Acids, 2015, 34, 691–707.
42 J. Salon, J. Sheng, J. Gan and Z. Huang, J. Org. Chem., 2010, 69 A. H. Krotz, C. Rentel, D. Gorman, P. Olsen, H. J. Gaus,
75, 637–641.
J. V. McArdle and A. N. Scozzari, Nucleosides, Nucleotides
Nucleic Acids, 2004, 23, 767–775.
70 C. Glidewell and E. J. Leslie, J. Chem. Soc., Dalton Trans.,
1977, 527–531.
71 L. J. Yu, C. J. Wiederholt, J. N. Patro, K. Haraguchi and
M. M. Greenberg, J. Org. Chem., 2005, 70, 141–149.
72 H. R. Drew, R. M. Wing, T. Takano, C. Broka, S. Tanaka,
K. Itakura and R. E. Dickerson, Proc. Natl. Acad. Sci. U. S.
A., 1981, 78, 2179–2183.
43 J. Sheng, J. Jiang, J. Salon and Z. Huang, Org. Lett., 2007, 9,
749–752.
44 J. Sheng, J. Salon, J. Gan and Z. Huang, Sci. China: Chem.,
2010, 53, 78–85.
45 H. Moroder, C. Kreutz, K. Lang, A. Serganov and R. Micura, J.
Am. Chem. Soc., 2006, 128, 9909–9918.
46 R. A. Thompson, A. M. Spring, J. Sheng, Z. Huang and
M. W. Germann, J. Biomol. Struct. Dyn., 2015, 33, 289–297.
47 L. A. Wozniak, M. Sochacki, H. Mitsuya, S. Kageyama and 73 C. H. Kang, I. Berger, C. Lockshin, R. Ratliff, R. Moyzis and
W. J. Stec, Bioorg. Med. Chem. Lett., 1994, 4, 1033–1036. A. Rich, Proc. Natl. Acad. Sci. U. S. A., 1994, 91, 11636–11640.
48 Y. Xu and E. T. Kool, J. Am. Chem. Soc., 2000, 122, 9040–9041. 74 H. A. Assi, M. Garavıs, C. Gonzalez and M. J. Damha, Nucleic
´
´
49 O. Eguaogie, P. F. Conlon and J. S. Vyle, Tetrahedron Lett.,
2016, 57, 5000–5002.
50 P. Pistschimuka, J. Prakt. Chem., 1911, 84, 746–760.
Acids Res., 2018, 46, 8038–8056.
75 S. P. Adams, K. S. Kavka, E. J. Wykes, S. B. Holder and
G. R. Galluppi, J. Am. Chem. Soc., 1983, 105, 661–663.
´
51 G. Mielniczak and A. Lopusinski, Heteroat. Chem., 2003, 14, 76 R. Zuckermann, D. Corey and P. Schultz, Nucleic Acids Res.,
121–127. 1987, 15, 5305–5321.
52 M. del C. Michelini, N. Russo, S. Alcaro and L. A. Wozniak, 77 A. F. Cook, J. Am. Chem. Soc., 1970, 92, 190–195.
Tetrahedron, 2012, 68, 5554–5563.
78 S. Chladek and J. Nagyvary, J. Am. Chem. Soc., 1972, 94, 2079–
53 L. A. Wozniak, M. Gra, Z. Kaminski and W. J. Stec,
2085.
Phosphorus, Sulfur Silicon Relat. Elem., 2008, 183, 1076–1081. 79 Y. Xu and E. T. Kool, Nucleic Acids Res., 1998, 26, 3159–3164.
54 M. Kullberg and J. Stawinski, Nucleosides, Nucleotides Nucleic 80 S. Li, W. K. Olson and X.-J. Lu, Nucleic Acids Res., 2019, 47,
Acids, 2005, 24, 659–661.
W26–W34.
55 R. Cosstick and F. Eckstein, Biochemistry, 1985, 24, 3630– 81 C. A. Frederick, G. J. Quigley, M. K. Teng, M. Coll, G. A. van
3638.
der Marel, J. H. van Boom, A. Rich and A. H.-J. Wang, Eur. J.
Biochem., 1989, 181, 295–307.
82 P. S. Pallan, T. P. Prakash, F. Li, R. L. Eoff, M. Manoharan
and M. Egli, Chem. Commun., 2009, 2017–2019.
83 J. Harp, P. Pallan and M. Egli, Crystals, 2016, 6, 125.
56 A. Grandas, W. S. Marshall, J. Nielsen and M. H. Caruthers,
Tetrahedron Lett., 1989, 30, 543–546.
57 R. Cosstick and J. S. Vyle, Nucleic Acids Res., 1990, 18, 829–
835.
58 K. Misiura and W. J. Stec, Bioorg. Med. Chem. Lett., 1994, 4, 84 J. M. Martin-Garcia, C. E. Conrad, J. Coe, S. Roy-Chowdhury
1037–1040. and P. Fromme, Arch. Biochem. Biophys., 2016, 602, 32–47.
59 A. M. Belostotskii, J. Lexner and A. Hassner, Tetrahedron 85 C. H. Yoon, H. DeMirci, R. G. Sierra, E. H. Dao, R. Ahmadi,
Lett., 1999, 40, 1181–1184.
F. Aksit, A. L. Aquila, A. Batyuk, H. Cici, S. Guillet,
¨
60 O. Eguaogie, P. F. Conlon, F. Ravalico, J. S. T. Sweet,
T. B. Elder, L. P. Conway, M. E. Lennon, D. R. W. Hodgson
and J. S. Vyle, Beilstein J. Org. Chem., 2017, 13, 87–92.
M. J. Hayes, B. Hayes, T. J. Lane, M. Liang, U. Lundstrom,
J. E. Koglin, P. Mgbam, Y. Rao, T. Rendahl, E. Rodriguez,
L. Zhang, S. Wakatsuki, S. Boutet, J. M. Holton and
M. S. Hunter, Sci. Data, 2017, 4, 170055.
ˆ
61 O. Eguaogie, J. S. Vyle, P. F. Conlon, M. A. Gılea and Y. Liang,
Beilstein J. Org. Chem., 2018, 14, 955–970.
62 Y. Ju, D. Kumar and R. S. Varma, J. Org. Chem., 2006, 71,
6697–6700.
86 R. L. Setten, J. J. Rossi and S. ping Han, Nat. Rev. Drug
Discovery, 2019, 18, 421–446.
87 R. Cosstick and J. Gaynor, Curr. Org. Chem., 2008, 12, 291–
308.
¨
63 R. Suchsland, B. Appel, M. Janczyk and S. Muller, Appl. Sci.,
2019, 9, 2199.
88 Z. Veres, L. Tsai, T. D. Scholz, M. Politino, R. S. Balaban and
T. C. Stadtman, Proc. Natl. Acad. Sci. U. S. A., 1992, 89, 2975–
2979.
64 M. Hassler, Y. Q. Wu, N. Mallikarjuna Reddy, T. H. Chan and
M. J. Damha, Tetrahedron Lett., 2011, 52, 2575–2578.
¨
65 R. Suchsland, B. Appel and S. Muller, Beilstein J. Org. Chem.,
2018, 14, 397–406.
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