Thermal stabilisation of RNA·RNA duplexes and G-quadruplexes by phosphorothiolate linkages

Article abstract of DOI:10.1039/c2ob26940f

The effect of 3′-S-phosphorothiolate linkages on the stability of RNA·RNA duplexes and G-quadruplex structures has been studied. 3′-Thio-2′-deoxyuridine was incorporated into RNA duplexes and thermal melting studies revealed that the resulting 3′-S-phosph

Full text of DOI:10.1039/c2ob26940f

Organic &
Biomolecular
Chemistry
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Thermal stabilisation of RNA·RNA duplexes and
G-quadruplexes by phosphorothiolate linkages†‡
Michael M. Piperakis,§a James W. Gaynor,a Julie Fisherb and Richard Cosstick*a,b
Cite this: Org. Biomol. Chem., 2013, 11,
966
The effect of 3’-S-phosphorothiolate linkages on the stability of RNA·RNA duplexes and G-quadruplex
structures has been studied. 3’-Thio-2’-deoxyuridine was incorporated into RNA duplexes and thermal
melting studies revealed that the resulting 3’-S-phosphorothiolate linkages increased the stability of the
duplex to thermal denaturation. Additionally, and contrary to expectation, a similar effect on duplex
stability was observed when the same thionucleoside was incorporated into the RNA strand of a
RNA·DNA duplex. A suitably protected derivative of 3’-thio-2’-deoxyguanosine was prepared using an
oxidation–reduction strategy and this residue also increased the thermal stability the [d(TGGGGT)]4
G-quadruplex when positioned centrally. The results are discussed in terms of the influence that the
sulfur atom has on the conformation of the furanose ring and imply that the previously noted high
thermal stability of parallel RNA quadruplexes is not derived from H-bonding interactions of the
2’-hydroxyl group, but can be attributed to conformational effects.
Received 3rd October 2012,
Accepted 30th November 2012
DOI: 10.1039/c2ob26940f
www.rsc.org/obc
has proved to be an extremely useful probe for studying metal
ion-dependent phosphoryl-transfer. In particular it has con-
Introduction
The 3′-phosphorothiolate linkage (3′-SP, 1, Fig. 1), in which the tributed considerably to our understanding of the mechanism
3′-oxygen atom of the phosphodiester bond is replaced by of RNA cleavage by the group I2 and group II3 introns and the
sulfur, can formerly be considered as the result of incorporat- splicesome.4
ing a 3′-thionucleoside into nucleic acids.1 This modification
When incorporated into DNA this modification has also
been valuable in characterizing the mechanism of action of
nucleic acid processing enzymes as exemplified by restriction
endonuclease BfiI,5 and the Klenow fragment of DNA polymer-
ase.6 Interestingly, incorporating a 3′-SP linkage into DNA
shifts the conformation of the sugar to which the sulfur is
attached to the north pucker (C3′-endo), a conformation that is
usually associated with standard A-form RNA.7 As a conse-
quence of this conformational effect, the incorporation of
3′-SP linkages into a DNA strand increases the thermal stability
of a DNA·RNA duplexes, but destabilises DNA·DNA duplexes.8
The conformational preference of this modification to adopt
the C3′-endo sugar pucker also appears to be responsible for
its ability to stabilise the 4-stranded i-motif structure,9 which
is found in a number of gene promoter sequences.10
It is evident that 3′-SP linkage acts as a RNA mimic even
when it is associated with a deoxyribose sugars; it thus raises
the question as to whether this modification could also stabi-
lise other nucleic acids structures which have a preference to
adopt the C3′-endo sugar pucker. In particular, two structures
were of interest: RNA·RNA duplexes and the G-quadruplex. The
stabilisation of RNA duplexes (both thermally and in terms of
improving resistance to nucleases) is important in generating
highly active short interfering RNAs (siRNA). However, in
order to achieve efficient processing of the siRNA into the
Fig. 1 3’-S-Phosphorothiolate linkage.
aDepartment of Chemistry, University of Liverpool, Liverpool, L69 7ZD, UK.
E-mail: rcosstic@liv.ac.uk, J.W.Gaynor@liv.ac.uk; Fax: +44 (0) 151 794 3588;
Tel: +44 (0) 151 794 3514
bSchool of Chemistry, University of Leeds, Leeds, LS2 9JT, UK.
E-mail: J.Fisher@chem.leeds.ac.uk; Tel: +44 (0) 113 343 6577
†Electronic supplementary information (ESI) available: Details for the synthesis
of 5′-O-dimethoxytrityl-2′-deoxy-3′-thiouridine phosphor-amidite; oligonucleotide
synthesis; HPLC and thermal melting studies. See DOI: 10.1039/c2ob26940f
‡In memory of Professor Har Gobind Khorana who was a PhD graduate (1948) at
the University of Liverpool.
§Current address: John Innes Centre, Norwich Research Park, Norwich, NR4
7UH, UK. E-mail: Michael.Piperakis@jic.ac.uk
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ribonucleoprotein complex, known as the RNA-induced silen-
cing complex (RISC), stabilisation has to be achieved with
minimum perturbation to the standard A-form RNA duplex.11
In this respect, the advantageous conformational properties of
the 3′-SP modification are achieved without the introduction
of any steric encumbrance that is likely to interfere with either
the assembly or function of the RISC complex. In addition, as
the 3′-SP linkage is used in a DNA context, it is likely to prove
resistant to most RNases.
G-quadruplexes are four stranded structures and are
formed because of the capacity of guanine bases to self-associ-
ate through Hoogsteen hydrogen bonds, resulting in stacked
G-quartets.12 Potential quadruplex structures have been associ-
ated with the G-rich sequences of telomeres13 and are also
linked with gene promoter regions.14 Interestingly, parallel
tetrameric G-quadruplexes derived from ribonucleosides
[e.g. r(UGGGGU)4] have been shown to be thermally more
stable than those composed of 2′-deoxyribonucleosides
[d(TGGGGT)4],15,16 although it is not entirely clear as to
whether the stabilisation stems solely from conformational
effects or whether hydrogen bonding interactions involving
the 2′-hydroxyl groups also contribute.17
Scheme 1 Synthesis of 3’-thio-2’-deoxyuridine monomer. Reagents and con-
ditions: (i) NaSBz, DMAc, 110 °C, 40–74% or BzSH, pyridine, 110 °C, 48%;
(ii) NaOH, H2O/EtOH, 92%; (iii) P(NiPr2)2(OCH2CH2CN), 1H-tetrazole, MeCN,
72%. Full experimental details are available in the ESI.†
Synthesis of 3′-thio-2′-deoxyguanosine for incorporation into
DNA
Whilst the synthesis of pyrimidine 3′-thionucleosides is readily
achieved through the use of anhydronucleoside intermediates
to control stereochemistry at the C3′ position, this route is not
available for their purine counterparts. There are a number of
methods for epimerising the C3′ of purine nucleosides
and we have previously used Herdewijn’s method for pre-
paring protected derivatives of 2′-deoxy-3′-thioadenosine
(6, Scheme 2A).19 However, we were attracted to the recently
reported in situ oxidation–reduction procedure of Eisenhuth
and Richert for the preparation of xylo-purines.20 Thus,
Previous work on the incorporation of phosphorothiolate
linkages into DNA has been limited to the pyrimidine nucleo-
sides.18 In order to synthesise G-quadruplex sequences, it is
necessary to prepare a suitably protected derivative of 3′-thio-
2′-deoxyguanosine. We now report the synthesis of RNA
duplexes containing 3′-thio-2′-deoxyuridine and show that this
modification thermally stabilises RNA duplexes. In addition
3′-thio-2′-deoxyguanosine has been incorporated into DNA
strands and is demonstrated to stabilise G-quadruplex
structures.
a
5′-TBDMS-protected derivative of deoxyguanosine (8a,
Scheme 2B) was converted to the corresponding xylo derivative
(9a, 79% yield) using this oxidation–reduction strategy. Intro-
duction of the masked thiol was accomplished in 65% through
a Mitsunobu reaction using thiobenzoic acid to give the benzo-
thioate ester and exchange of the TBDMS group for the
dimethoxytrityl group was achieved in 85% for these two
further steps.
In an attempt to avoid the protecting group exchange, the
oxidation–reduction procedure was also performed directly on
the dimethoxytritylated substrate (8b), but in this case the xylo
derivative (9b) was obtained in only 20% yield. Likewise, a
Mitsunobu reaction on 9b gave the DMT-protected benzothio-
ate ester (10b) in very low yield (5%). In both cases the yields
were reduced largely due to loss of the DMT group. The DMT-
protected thioester (10b) was subsequently hydrolysed to give
the protected 3′-thio-2′-deoxyguanosine 11 under conditions
similar to those previously used for hydrolysis of the corres-
ponding cytidine derivative.18 Finally, phosphitylation of 11
gave the required 3′-thio-2′-deoxyguanosine phosphorothio-
amidite (12) in an overall yield of 21% starting from deoxy-
guanosine. In addition, the phosphitylation 2′-deoxy-
3′-thioadenosine19 (6) was also accomplished under similar
conditions to give the phosphorothioamidite (7).
Results and discussion
Synthesis of 3′-thio-2′-deoxyuridine for incorporation into RNA
As outlined above, the incorporation of 3′-SP linkages derived
from deoxyribose sugars was expected to confer favourable
conformational properties and nuclease resistance to RNA
with minimal alteration to the overall structure. 3′-Thio-
2′-deoxyuridine is the synthetically most convenient “RNA-
compatible” deoxyribonucleoside for this purpose. Although
this nucleoside has previously been incorporated into RNA
substrates to study RNA splicing,2a,4 no details of the synthesis
of the required deoxyuridine monomer have previously been
reported. The synthesis of the 5′-O-dimethoxytrityl-2′-deoxy-
3′-thiouridine phosphoramidite (5) is shown in Scheme 1 and
it follows the route previously described for the analogous thy-
midine derivative.18 (Full experimental details are available in
the ESI.†) Interestingly, opening of the anhydronucleoside (2)
to give the benzothioate ester (3) can be done either using pre-
prepared sodium thiobenzoate (40–74% yield), as previously
reported,18 or by using thiobenzoic acid in pyridine (48%). The
latter procedure is more straightforward and gives much more
consistent yields.
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Fig. 2 Reverse-phase HPLC trace for a mixture of ORNs 1–4 (see Table 1)
showing the retention time increasing with increasing number of phosphoro-
thiolate linkages. HPLC elution gradient = 0–16% MeCN over 30 min in 0.1 M
triethylammonium bicarbonate (pH 7.6).
DNA and in that context was previously shown to give smooth
thermal melting curves.8 Oligoribonucleotides were prepared
using 2′-O-TBDMS-protected monomers and 5-ethylmercapto-
1H-tetrazole (ETT) (0.25 M) as the activator. For incorporation
of the 2′-deoxy-3′-thiouridine phosphorothioamidite (5) the
concentration of ETT was increased to 1 M and a 15 min coup-
ling was used.18 Oligoribonucleotides were synthesised
without the 5′-terminal DMT group attached and following
removal of the 2′-O-TBDMS groups, the oligoribonucleotides
were purified by ion-exchange HPLC (see Table 1 for sequences
prepared).
The oligoribonucleotides (ORNs) were characterised by elec-
trospray mass spectrometry and their purity established by
reverse-phase HPLC. The incorporation of 3′-thio-2′-deoxyuri-
dine into the oligoribonucleotides increased hydrophobicity as
clearly indicated by the increasing retention times on reverse-
phase HPLC as more phosphorothiolate linkages are incorpor-
ated (Fig. 2).
Scheme 2 Synthesis of 3’-thio-2’-deoxyguanosine and 3’-thio-2’-deoxyadeno-
sine monomers. Reagents and conditions: (i) P(NiPr2)2(OCH2CH2CN), 5-ethylmer-
capto-1H-tetrazole, CH2Cl2, 82%; (ii) Dess–Martin periodinane, CH2Cl2, 0 °C to
25 °C, then NaBH4, −60 °C, 79% and 20% yield for 9a and 9b, respectively;
(iii) PPh3, diisopropyl azodicarboxylate, BzSH, THF, 0 °C, 65% and 5% yield for
10a and 10b, respectively; (iv) AcOH, THF, TBAF; (v) DMTCl, py, 85% over (iv)
and (v); (vi) NaOH, THF/MeOH/H2O; (vii) P(NiPr2)2(OCH2CH2CN), 5-ethylmer-
capto-1H-tetrazole, CH2Cl2, 81% yield over (vi) and (vii).
Synthesis and characterisation of oligodeoxynucleotides
Oligodeoxyribonucleotides containing phosphorothiolate lin-
kages were prepared as previously described, with the 5′-termi-
nal DMT group attached and they were purified by reverse-
phase HPLC (see Table 2 for sequences prepared).18 As pre-
viously noted, the HPLC traces for the hexamer sequences all
showed slower eluting peaks corresponding to the G-quadru-
plex structure.21
Table 1 List of RNA sequences, together with their theoretical and experimen-
tal average masses
Average masses
ORN sequence
ORN no
Theoretical
Expt
r(GCGUUUUUUUUUUGCG)
1
2
3
4
4990.93
4990.99
4991.06
4991.13
5141.28
4990.65
4991.03
4990.77
4990.77
5141.29
Table 2 List of DNA sequences, together with the theoretical and experimen-
r(GCGUUUUU
r(GCGUUUU
r(GCG UUUU
r(CGCAAAAAAAAAACGC)
̲
̲
tal average masses
̲
̲
̲
s
̲
̲
̲
̲
Average masses
—
ODN sequence
Theoretical
Experimental
All sequences written 5′–3′. U̲s̲ = indicates a 3′-SP linkage associated
with a 2′-deoxy-3′-thiouridine residue.
5′-d(CGCAAAAAAAAAACGC)
4885.24
4933.44
1863.35
1879.31
1879.31
1879.31
1879.31
4884.72
4932.88
1862.42
1878.42
1878.42
1878.37
1878.27
5′-d(CGCAAAAA
5′-d(TGGGGT)
5′-d(TG GGGT)
5′-d(TGG GGT)
5′-d(TGGG GT)
5′-d(TGGGG T)
̲s̲A̲s̲A̲s̲AAACGC)
̲s̲
Synthesis and characterisation of oligoribonucleotides
̲s̲
The sequence of the RNA strand chosen for incorporation of
the 2′-deoxy-3′-thiouridine is shown in Table 1. It is based on
an equivalent DNA sequence that has previously been used to
study the incorporation of phosphorothiolate linkages into
̲s̲
̲s̲
A̲s̲ or G̲s̲ indicates a 3′-SP linkage associated with either 2′-deoxy-
3′-thioadenosine or 2′-deoxy-3′-thioguanosine, respectively.
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that the deoxyribose sugars associated with the 3′-SP linkages
should favour the 3′-endo conformation, thus increasing base
stacking interactions. However, other factors are likely to affect
the structure. For example in RNA duplexes, the 2′-hydroxyl
groups line the minor groove and show a hydration pattern
similar to that of the negatively charged phosphate group.23
This significantly contributes to the thermodynamic stability
of RNA. By increasing the number of 3′-thio-2′-deoxyuridine
residues in the RNA duplex, the number of 2′-OH groups in
the minor groove is correspondingly reduced resulting in a
drop of hydration. In the absence of additional information it
is difficult to predict the how the reduced hydration at the
central base pairs will affect the duplex structure and stability.
Interesting results were obtained for the thermal melting
studies performed on DNA·RNA duplexes between the 5′-r-
(GCGUUUUUUUUUUGCG) and the complementary DNA
strand (duplexes 5–8, Table 3). In DNA·RNA duplexes, the RNA
strand has been shown to adopt the A-type conformation,
whilst in the DNA strand the nucleotide conformations are
closer to that of B-DNA and globally, these hybrid duplexes
show closer resemblance to the A-form rather than the
B-form.24 As expected, based on the previously discussed con-
formational preference of the 3′-thiosugar, incorporation of
3′-SP linkages into the DNA strand of an RNA·DNA duplex
increased the Tm by about 2.0 °C per modification (duplex 6,
Table 3). More surprisingly, a similar increase in Tm was
observed when the 3′-SP linkages were incorporated into the
RNA strand (duplex 7). This was contrary to expectation as in
this case the 3′-SP modification was predicted to enhance the
mismatch in sugar conformations between the two strands
and thereby decrease the Tm. When 3′-SP linkages were incor-
porated into both strands and in direct opposition (duplex 8),
the Tm was increased by just over 11 °C (ΔTm = 1.8 °C per
modification). Thus, the 3′-SP linkages increase the thermal
stability of a RNA·DNA duplex independent of which strand is
modified. The reason for this is unclear, although previous
studies have shown that the global structure of a RNA·DNA
duplex in which the RNA strand is pyrimidine rich (as in the
case of these duplexes) has less A-type helical character and is
thermodynamically less stable than when the RNA strand is
purine rich.25 This may mean that for the duplex under investi-
gation here, the incorporation of 3′-SP linkages into either
strand globally increases the A-type character of the duplex
and this results in the higher Tm values. In this respect it
would be interesting to examine phosphorothiolate substi-
tution into an analogous duplex in which the RNA strand was
purine rich.
Fig. 3 Melting curves for RNA duplexes 1–4 (see Table 3). Melting studies were
performed on samples containing 1.5 μM duplex in sodium phosphate buffer
(10 mM, pH 7.0) containing 100 mM NaCl. Duplex 1 = red, duplex 2 = black,
duplex 3 = blue, duplex 4 = green.
Table 3
Tm data for RNA·RNA duplexes (duplexes 1–4) and DNA·RNA duplexes
(duplexes 5–8)
Duplex
number
Sequence
Tm (°C) ΔTm (°C)
1
50.9
51.9
53.2
54.3
30.0
35.9
36.5
41.1
—
2
3
4
5
6
7
8
1.0a
2.3
3.4
—
5.9
6.5
11.1
U̲s̲ or A̲s̲ indicates a 3′-SP linkage associated with either 2′-deoxy-
3′-thiouridine or 2′-deoxy-3′-thioadenosine, respectively. Samples were
prepared as described in the legend to Fig. 3. a Whilst a ΔTm of 1 °C is
at the limits of experimental error, it is consistent with the general
trend, which is revealed more clearly in the more substituted duplexes.
Thermal melting studies on RNA·RNA and RNA·DNA duplexes
All RNA·RNA duplexes produced melting curves with sharp
monophasic transitions (Fig. 3) and show a shift to higher
Tm values for increasing numbers of 3′-SP linkages incorpor-
ated (approximately 1 °C increase per modification, see
Table 3). NMR studies on simple dinucleotides have previously
shown22 that 3′-thio-2′-deoxyribose has a greater tendency to
adopt the 3′-endo conformation than the corresponding ribose
sugar and in this respect incorporation of the 3′-SP linkages
would be expected to provide greater preorganisation of the
modified strand for binding to its RNA complement; this is
likely to be the cause of the increased Tm values for duplexes
2, 3 and 4. An interesting observation is the trend in decreas-
ing hypochromicity with increasing number of 3′-SP modifi-
cations (Fig. 3). A drop in hypochromicity implies a reduction
in base stacking interactions which is an important factor in
the overall stability of the duplex. As noted above, it is expected
Spectrophotomeric studies on G-quadruplex structures
Tetrameric G-quadruplex structures are well suited to analogue
replacement studies and in particular [5′-d(TGGGGT)]4 and
simple variants of this quadruplex, have previously been
investigated.15–17 For this study, formation of G-quadruplexes
was confirmed by a thermal difference spectrum (TDS), which
is simply the difference between UV spectra recorded for the
unfolded and folded states. The base stacking interactions
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with regard to the sugar conformations, with the DNA quadru-
plexes adopting the C2′-endo pucker whereas the RNA equival-
ent adopts a mixture of C2′ and C3′-endo conformations.15,29
Parallel RNA tetrameric quadruplexes are known to be con-
siderably more thermally stable than their DNA counterparts.
This is a result of both a much faster association and slower
dissociation for the RNA quadruplexes.16 A potential expla-
nation for the difference between these RNA and DNA quadru-
plexes relates to the effect that the sugar conformation has on
the syn/anti equilibrium; the anti-conformation, which favours
parallel quadruplexes, is more accessible from the C3′-endo
pucker.30 The results presented here are also consistent with
this interpretation, given that a 3′-SP linkage acts as a confor-
mational mimic of a ribonucleoside. Additionally, it has been
established that the analogous, fully LNA-modified tetramole-
cular quadruplexes, in which the sugar is locked into the
C3′-endo pucker, also shows a similar stabilisation effect.17
There has been speculation that stabilisation of the tetrameric
G-quadruplexes by 2′-OH and 2′-OMe substituents,31 as well as
LNA residues,17 could be due to the ability of these modifi-
cations to participate in H-bonding interactions. However,
the results reported here, with a modification that has no
H-bonding functionality at the 2′-position, suggest that
H-bonding interactions do not play a major role in stabilising
the structure.
Table 4 T1/2 data for G-quadruplexes. G
̲s̲ indicates a 3’-SP linkage associated
with 2’-deoxy-3’-thioguanosine
Quadruplex no
G-quadruplex
T1/2/°C
ΔT1/2/°C
9
[5′-d(TGGGGT)]4
[5′-d(TG GGGT)]4
[5′-d(TGG GGT)]4
[5′-d(TGGG GT)]4
[5′-d(TGGGG T)]4
61.1
60.1
70.0
68.7
61.2
N/A
−1.0
8.9
7.6
0.1
10
11
12
13
̲s̲
̲s̲
̲
̲
̲
̲
Melting studies were performed on samples containing 10 μM single
strand in sodium cacodylate buffer (10 mM, pH 7.0) containing
100 mM NaCl. The heating rate was 0.5 °C min−1
.
As already noted above, the current results show that incor-
poration of the 3′-SP linkage into the terminal dG residues has
only a small effect on quadruplex stability (quadruplexes 10
and 13). This positional effect is consistent with NMR studies
which show that these outer tetrads are more flexible than
those in the centre and are therefore less likely to contribute to
the stability of the structure as a whole.15,32 The marginal drop
in T1/2 observed here for incorporation of the phosphorothio-
late linkage at the 5′-terminal G residue (quadruplex 10,
Table 4) can also be rationalised based on previous studies. It
has been shown that T1/2 is increased when guanosine ana-
logues that preferentially adopt the syn-conformation are incor-
Fig. 4 Thermal difference spectra in the 220–340 nm region for the five quad-
ruplexes studied (9–13, Table 4). Pink = quadruplex 9; green = quadruplex 10;
brown = quadruplex 11; blue = quadruplex 12 and orange = quadruplex 13.
Thermal difference spectra were performed on samples containing 10 μM single
strand in sodium cacodylate buffer (10 mM, pH 7.0) containing 100 mM NaCl.
present in each type of nucleic acid structure is reflected in the
TDS and it thus has a specific shape which is unique to that
particular structure.26 The TDS for the unmodified and the
3′-SP substituted quadruplexes (sequences shown in Table 4)
all showed positive (∼243 and 273 nm) and negative
(∼295 nm) peaks that are characteristic of a G-quadruplex and
the global shape of all the spectra was very similar (Fig. 4).26
The results of thermal melting studies performed on the
[5′-d(TGGGGT)]4 quadruplexes containing 3′-SP linkages are
shown in Table 4. The measured Tm for a G-quadruplex gener-
ally depends on the heating rate and is not therefore the true
porated at the 5′-terminal
G residue of a tetrameric
quadruplex, suggesting that a 5′-syn residue might, in fact,
stabilise the structure.33 In this case the lower T1/2 for quadru-
plex 10 is likely to result from the syn conformation being less
accessible due to the C3′-endo sugar pucker induced by the
3′-SP linkage.
thermodynamic Tm; this apparent Tm is often referred to as
16,27
T1/2
.
A sequence specific effect on the T1/2 of the modified
quadruplexes is evident from the data and a significant
enhancement in the T1/2 is observed when the single modified
residue is located at either of the central positions (Table 4,
quadruplexes 11 and 12). In comparison, the effect is much
smaller when either of the terminal dG residues is modified
and a marginal destabilising effect results from incorporation
at the 5′-dG residue (quadruplex 10).
Previous studies have shown that DNA tetrameric quadru-
plexes such as [d(TGGGGT)]4 and its RNA equivalent
[r(UGGGGU)]4 adopt similar structures; both have a parallel
arrangement of the four strands with all nucleosides in the
anti conformation.15,28 There are however, some differences
Conclusions
Previous studies have shown that 3′-S-phosphorothiolate lin-
kages are able to stabilise RNA·DNA duplexes8 (when the modi-
fication is in the DNA strand) and the I-motif.9 The current
study now reveals that both RNA·RNA and G-quadruplex
structures are also thermally stabilised by the 3′-SP linkage. In
addition, it is now revealed that the 3′-SP linkage stabilises
RNA·DNA duplexes irrespective of which strand contains the
modification. Thus of all the structures so far investigated it
is only DNA·DNA duplexes that are thermally destabilised by
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3′-SP linkages.8 In all cases this stabilisation appears to stem powdered NaBH4 (0.81 g, 21.3 mmol) was subsequently added
from the influence that the sulfur atom has on the confor- and the mixture allowed to stir at −60 °C overnight. Acetone
mation of the furanose ring. In particular, results presented (35 mL) was next added and the slurry was allowed to warm to
here for the G-quadruplexes imply that the high thermal stabi- room temperature. Half of the volume of solvent was removed
lity of parallel RNA quadruplexes is not derived from by evaporation before the mixture was taken up in EtOAc
H-bonding interactions of the 2′-hydroxyl group, but can be (200 mL) and washed successively with equal volumes of H2O,
attributed to conformational effects.
NaHCO3 and brine. The collected organic layer was then dried
over Na2SO4, filtered and concentrated under reduced pressure
to afford the crude product as an off-white solid. Purification
by flash column chromatography, eluting with a gradient of
1% MeOH in CHCl3–3% MeOH in CHCl3, yielded the title
compound as a white amorphous solid (3.80 g, 8.41 mmol,
Experimental
General
Unless otherwise stated, all general reagents were purchased 79%).
from either Aldrich or BDH and used as supplied. 2-Cya-
1H NMR (400 MHz, DMSO) δ 11.69 (br s, 2 H, NH), 8.14
noethyl-N,N,N′,N′-tetraisopropylphosphorodiamidite was pur- (s, 1 H, H8), 6.12 (dd, J = 1.5, 8.0 Hz, 1 H, H1′), 5.35 (d, J = 3.8
chased from Chemgenes. Solvents were purchased from BDH Hz, 1 H, 3′-OH), 4.37–4.32 (m, 1 H, H3′), 4.00–3.88 (m, 2 H,
or Fisher. HPLC grade acetonitrile and triethylamine were H4′, H5′), 3.79–3.71 (m, 1 H, H5′), 2.80–2.62 (m, 2 H, CH-
used to prepare RP-HPLC buffers. Milli-Q water and HPLC (CH3)2, H2′), 2.25–2.18 (m, 1 H, H2′), 1.09 (d, J = 6.8 Hz, 6 H,
methanol were used to prepare ion-exchange HPLC buffers. CH(CH3)2), 0.83 (s, 9 H, C(CH3)3), 0.00 (s, 6 H, Si(CH3)2.
5-Ethylmercapto-1H-tetrazole and all reagents for DNA/RNA 13C NMR (101 MHz, DMSO) δ 180.5 (NHibu CvO), 155.2 (C6),
synthesis were supplied by Link Technologies. DNA synthesis 148.6 and 148.4 (C2, C4), 138.4 (C8), 120.2 (C5), 85.8 (C4′), 82.9
grade acetonitrile was used to prepare phosphoroamidite and (C1′), 69.2 (C3′), 62.5 (C5′), 41.4 (C2′), 35.1 (CH(CH3)2), 26.2
activator solutions. Analytical thin layer chromatography was (C(CH3)3), 19.2 (CH(CH3)2), 18.4 (C(CH3)3), −4.9 (SiCH3).
performed on UV254 sensitive, silica gel 60 coated, aluminium ES-HRMS; C20H33N5O5SiNa requires 474.2149; [M + Na]+
=
tlc plates purchased from Merck. Ellman’s reagent was used to 474.2152 (0.7 ppm).
distinguish compounds containing a 3′-thiol group.19 Flash
N2-Isobutyryl-5′-O-dimethoxytrityl-2′-deoxyxyloguanosine
column chromatography was performed using silica (particle (9b). Reaction conditions as for 9a, performed on N2-
size 40–63 μM, supplied by BDH).
isobutyryl-5′-O-dimethoxytrityl-2′-deoxyguanosine34 (8b, 2.89 g,
All NMR spectra were recorded on a Bruker 400 MHz spec- 4.52 mmol). Purification by flash column chromatography,
1
trometer; operating at 400 MHz for H, 101 MHz for 13C NMR eluting with a gradient of 2% MeOH in CH2Cl2–4% MeOH in
and 162 MHz for 31P. All spectra were recorded in either CH2Cl2, yielded the title compound as a pale yellow amor-
d6-DMSO or CDCl3, relative to an internal standard of tetra- phous solid (0.58 g, 0.91 mmol, 20%).
methylsilane (1H NMR and 13C) or an external standard of
1H NMR (400 MHz, DMSO) δ 11.77 (br s, 2 H, NH), 8.01
85% H3PO4 (31P). All chemical shifts are reported in ppm and (s, 1 H, H8), 7.42–7.37 (m, 2 H, m-Ph), 7.29–7.18 (m, 7 H, o-Ph,
coupling constants (J) in Hertz. 13C and 31P spectra were p-Ph, o-Ar2), 6.87–6.78 (m, 4 H, m-Ar2), 6.24–6.18 (m, 1 H, H1′),
1H decoupled and were singlets unless otherwise stated. Standard 5.30 (d, J = 3.0 Hz, 1 H, 3′-OH), 4.34 (m, 1 H, H3′), 4.25–4.19
nucleoside numbering systems have been used with additional (m, 1 H, H4′), 3.73 (s, 3 H, OMe), 3.72 (s, 3 H, OMe), 3.39–3.35
abbreviations as follows: Ar2 = 4-methoxyphenyl on DMT; Ph = (m, 1 H, H5′), 3.18 (dd, J = 2.7, 10.2 Hz, 1 H, H5′), 2.84–2.73
phenyl on DMT, OCH2 = cyanoethyl. Mass spectra were recorded (m, 1 H, CH(CH3)2), 2.69 (ddd, J = 5.3, 8.4, 14.4 Hz, 1 H, H2′),
on a MicroMass LCT mass spectrometer using electrospray ionis- 2.31–2.25 (m, 1 H, H2′), 1.13 (d, J = 1.9 Hz, 3 H, CH(CH3)2),
ation (EI) and direct infusion syringe pump sampling. All mass 1.11 (d, J = 1.9 Hz, 3 H, CH(CH3)2). 13C NMR (101 MHz, DMSO)
spectra were visualised as a series of multicharged ions.
δ 180.6 (NHibu CvO), 158.4 (p-Ar2), 158.3 (C6), 148.6 and
For details regarding DNA/RNA synthesis, HPLC chromato- 148.3 (C2 and C4), 145.4 (ipso-Ph), 138.2 (C8), 136.1 (ipso-Ar),
graphy and thermal melting studies see ESI.† Details of the 130.1–127.0 (o-Ph, m-Ph, p-Ph, o-Ar2), 121.0 (C5), 113.4 (m-Ar2),
sample preparation for thermal melting studies are given in 85.7 (CPh(Ar)2), 84.4 (C4′), 83.4 (C1′), 69.6 (C3′), 63.5 (C5′), 55.3
the legends to the figures and tables.
(OMe), 41.1 (C2′), 35.1 (CH(CH3)2), 19.3 (CH(CH3)2). ES-HRMS:
N2-Isobutyryl-5′-O-tert-butyldimethylsilyl-2′-deoxyxylo-gua- C35H37N5O7Na requires 662.2591; [M + Na]+ = 662.2598
nosine (9a). A solution of N2-isobutyryl-5′-O-tert-butyldi- (1.1 ppm).
methylsilyl-2′-deoxyguanosine20 (8a, 4.81 g, 10.65 mmol)
N2-Isobutyryl-5′-O-tert-butyldimethylsilyl-3′-S-benzoyl-2′-deoxy-
dissolved in anhydrous CH2Cl2 (20 mL) was slowly added over 3′-thioguanosine (10a). Diisopropyl azodicarboxylate (0.66 mL,
10 min to a chilled to 0 °C solution of Dess–Martin periodi- 3.33 mmol) was added dropwise to a stirred solution of triphe-
nane in CH2Cl2 (15% wt, 45.2 mL, 15.98 mmol) under N2. The nylphosphine (0.87 g, 3.33 mmol) in anhydrous THF (8 mL)
reaction was allowed to stir at 0 °C for 1 hour. The cooling that had been chilled to 0 °C. The yellow mixture was allowed
bath was then removed and the reaction let stir at room temp- to stir at 0 °C for 30 min under N2. A solution of the xylo
erature for 4 further hours. 2-Propanol (35 mL) was next added nucleoside 9a (0.5 g, 1.11 mmol) dissolved in anhydrous THF
and the resulting white slurry cooled to −60 °C. Freshly (3 mL) followed by a solution of thiobenzoic acid (0.46 mL,
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3.86 mmol) dissolved in anhydrous THF (2 mL) were next
1H NMR (400 MHz, CDCl3) δ 11.84 (s, 1 H, NH), 8.97
added dropwise to the yellow mixture at 0 °C. The resulting (s, 1 H, NH), 7.87–7.83 (m, 2 H, o-SBz), 7.74 (s, 1 H, H8),
reaction was left to stir at room temperature for 4 h under N2. 7.61–7.57 (m, 1 H, p-SBz), 7.47–7.42 (m, 2 H, m-SBz), 7.23–7.20
Solvent was then evaporated and the recovered yellow residue (m, 2 H, o-Ph), 7.13–7.03 (m, 7 H, p-Ph, m-Ph, o-Ar2), 6.61–6.55
was dissolved in CH2Cl2 and washed successively with H2O (m, 4 H, m-Ar2), 6.02 (dd, J = 2.2, 7.7 Hz, 1 H, H1′), 5.59–5.50
and NaHCO3. The collected organic layer was then dried over (m, 1 H, H3′), 4.09–4.03 (m, 1 H, H4′), 3.62 (s, 6 H, OMe), 3.33
Na2SO4, filtered and concentrated under reduced pressure to (dd, J = 3.2, 10.6 Hz, 1 H, H5′), 3.26–3.17 (m, 2 H, H5′, H2′),
afford the crude product as a yellow oil. Purification by flash 2.54–2.36 (m, 2 H, H2′, CH(CH3)2), 1.18 (d, J = 6.8 Hz, 3 H, CH-
column chromatography, eluting with 1% MeOH in CHCl3, (CH3)2), 1.11 (d, J = 7.0 Hz, 3 H, CH(CH3)2).13C NMR (101 MHz,
yielded the title compound as a white amorphous solid CDCl3) δ 192.2 (SBz CvO), 179.0 (NHibu CvO), 158.8 (p-Ar2),
(0.41 g, 0.72 mmol, 65%).
156.1 (C6), 147.8 and 147.7 (C2 and C4), 144.9 (ipso-Ph), 139.3
1H NMR (400 MHz, CDCl3) δ 11.95 (br s, 1 H, NH), 9.06 (s, 1 (C8), 136.9 (ipso-SBz), 136.0 (ipso-Ar2), 134.6 (p-SBz),
H, NH), 7.98–7.95 (m, 2 H, o-SBz), 7.94 (s, 1 H, H8), 7.68–7.63 130.4–127.2 (o-Ph, m-Ph, p-Ph, o-Ar2, m-SBz, o-SBz), 122.4 (C5),
(m, 1 H, p-SBz), 7.55–7.48 (m, 2 H, m-SBz), 6.13 (dd, J = 2.8, 7.4 113.3 (m-Ar2), 86.7 (CPh(Ar)2), 84.9 (C1′), 83.8 (C4′), 62.4 (C5′),
Hz, 1 H, H1′), 5.22–5.13 (m, 1 H, H3′), 4.18–4.11 (m, 1 H, H4′), 55.5 (OMe), 40.5 (C3′), 39.1 (C2′), 36.9 (CH(CH3)2), 19.2
3.89 (dd, J = 3.0, 11.8 Hz, 1 H, H5′), 3.81 (dd, J = 3.8, 11.8 Hz, (CH(CH3)2). ES-HRMS: C42H41N5O7SNa requires 782.2624;
1 H, H5′), 3.19 (ddd, J = 2.7, 7.6, 13.3 Hz, 1 H, H2′), 2.72 (spt, [M + Na]+ = 782.2646 (2.8 ppm).
J = 7.2 Hz, 1 H, CH(CH3)2), 2.56 (ddd, J = 7.6, 9.9, 13.3 Hz, 1 H,
N2-Isobutyryl-5′-O-dimethoxytrityl-2′-deoxy-3′-thioguanosine
H2′), 1.34 (d, J = 7.2 Hz, 6 H, CH(CH3)2), 0.81 (s, 9 H, C(CH3)3), (11). N2 was bubbled through an aqueous solution of NaOH
0.01–0.02 (m, 3 H, SiCH3), −0.06 (s, 3 H, SiCH3). 13C NMR (0.5 M, 20 mL) as well as a solution of MeOH–THF–H2O
(101 MHz, CDCl3) δ 192.0 (SBz CvO), 180.0 (NHibu CvO), (4 : 6 : 1, 230 mL) for 30 min. N2-Isobutyryl-5′-dimethoxytrityl-
156.0 (C6), 148.6 and 147.9 (C2 and C4), 138.8 (C8), 136.9 3′-S-benzoyl-2′-deoxy-3′-thioguanosine (10b, 2.5 g, 3.29 mmol)
(ipso-SBz), 134.5 (p-SBz), 129.3 (m-SBz), 127.7 (o-SBz), 121.9 was added to the solution mixture of MeOH/THF/H2O and the
(C5), 85.4 (C4′), 84.8 (C1′), 62.9 (C5′), 40.3 (C3′), 39.6 (C2′), 37.1 resulting solution was then cooled to 0 °C before the degassed
(CH(CH3)2), 26.2 (C(CH3)3), 19.3 (CH(CH3)2), 18.8 (C(CH3)3), NaOH was added dropwise over 5 min. The reaction was
−5.2 (SiCH3). ES-HRMS; C27H37N5O5SSiNa requires 594.2182; allowed to stir at 0 °C for 30 min under N2. A solution of cold
[M + Na]+ = 594.2160 (−3.8 ppm).
citric acid (1 M, 49.5 mL) was next added dropwise over 10 min
N2-Isobutyryl-5′-O-dimethoxytrityl-3′-S-benzoyl-2′-deoxy-3′- to neutralise the base. The reaction solution was allowed to
thioguanosine (10b). Acetic acid (3.8 mL) was added slowly to stir at 0 °C for 15 min, then taken up in EtOAc and washed
a stirred solution of N2-isobutyryl-5′-O-tert-butyldimethylsilyl- once with an equal volume of NaHCO3. The collected organic
3′-S-benzoyl-2′-deoxy-3′-thioguanosine (10a, 4.36 g, 7.62 mmol) layer was then dried over MgSO4, filtered and concentrated
in anhydrous THF (65 mL) at 0 °C. Next, a solution of tetra- under reduced pressure to afford the crude product as an off-
butylammonium fluoride (1 M, 15.24 mL, 15.24 mmol) in THF white amorphous solid (2.14 g, 3.26 mmol, 99%). The recov-
was added dropwise over 5 min under N2. The reaction was ered material was carried on to the next reaction without any
then allowed to stir at room temperature until it went to com- further purification.
pletion as confirmed by TLC. The resulting solution was then
1H NMR (400 MHz, CDCl3) δ 11.95 (s, 1 H, NH), 8.32
taken up in CH2Cl2 (100 mL) and washed successively with (s, 1 H, NH), 7.81 (s, 1 H, H8), 7.36–7.31 (m, 2 H, o-Ph),
H2O and NaHCO3. The collected organic layer was then dried 7.25–7.09 (m, 7 H, m-Ph, p-Ph, o-Ar2), 6.75–6.70 (m, 4 H,
under Na2SO4, filtered and concentrated under reduced m-Ar2), 6.05 (dd, J = 3.4, 6.8 Hz, 1 H, H1′), 3.91–3.86 (m, 1 H,
pressure to afford the title compound as an off-white amor- H4′), 3.71 (s, 6 H, OMe-H), 3.67–3.57 (m, 1 H, H3′), 3.41 (dd,
phous solid (3.31 g, 7.23 mmol, 95%). The crude product was J = 3.0, 10.8 Hz, 1 H, H5′), 3.26 (dd, J = 4.0, 10.8 Hz, 1 H, H5′),
co-evaporated with anhydrous pyridine (3 × 15 mL) and sub- 2.83 (ddd, J = 3.4, 7.4, 13.7 Hz, 1 H, H2′), 2.43–2.24 (m, 2 H,
sequently dissolved in anhydrous pyridine (70 mL). The solu- CH(CH3)2, H2′), 1.60 (d, J = 6.6 Hz, 1 H, 3′-SH), 1.12 (d, J =
tion was next treated with 4,4′-dimethoxytrityl chloride 7.0 Hz, 3 H, CH(CH3)2), 1.07 (d, J = 6.8 Hz, 3 H, CH(CH3)2).
(5.18 g, 15.32 mmol) and the reaction allowed to stir at room 13C NMR (101 MHz, CDCl3) δ 179.4 (NHibu CvO), 158.9
temperature under N2 until it went to completion as indicated (p-Ar2), 156.1 (C6), 148.2 and 148.1 (C2, C4), 144.8 (ipso-Ph),
by TLC. The resulting solution was then quenched by addition 137.6 (C8), 135.9 (ipso-Ar2), 130.4–127.4 (o-Ph, m-Ph, p-Ph,
of MeOH (10 mL) and the solvent was next evaporated to o-Ar2), 122.1 (C5), 113.6 (m-Ar2), 88.6 (C4′), 87.0 (CPh(Ar)2),
furnish an orange oil. The residue was dissolved in CH2Cl2 84.2 (C1′), 62.5 (C5′), 55.7 (OMe), 42.6 (C2′), 36.7 (C3′), 35.7
and washed successively with NaHCO3 and brine. The col- (CH(CH3)2), 19.3 (CH(CH3)2). ES-HRMS: C35H37N5O6SNa
lected organic layer was then dried over Na2SO4, filtered and requires 678.2362; [M + Na]+ = 678.2393 (4.5 ppm).
concentrated under reduced pressure to afford the crude product
N2-Isobutyryl-5′-O-dimethoxytrityl-2′-deoxy-3′-thio-guano-
as an orange solid. Purification by flash column chromatography, sine-3′-S-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite
eluting with a gradient of 3% MeOH in CH2Cl2–5% MeOH in (12). A solution of 5-ethylmercapto-1H-tetrazole (0.25 M,
CH2Cl2, yielded the title compound as a white amorphous 10 mL) in MeCN was added to a stirred solution of the thio-
solid (4.92 g, 6.47 mmol, 85% over 2 steps).
guanosine derivative 11 (2.19 g, 3.34 mmol) in CH2Cl2 (20 mL)
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under N2. 2-Cyanoethyl-N,N,N′,N′-tetraisopropylphosphoro- CH2Cl2 and washed once with NaHCO3. The collected organic
diamidite (1.59 mL, 5.01 mmol) was then added dropwise over layer was dried over MgSO4, filtered and concentrated under
5 min to the stirred solution of nucleoside. The reaction was reduced pressure to afford the crude product as a pale yellow
allowed to stir at room temperature for 3 hours under N2. The solid. Purification by flash column chromatography, eluting
resulting solution was subsequently quenched by addition of with 3 : 1 EtOAc : hexane, yielded the title compound as a
MeOH (6 mL) and then taken up in CH2Cl2 and washed once white amorphous solid (0.84 g, 0.96 mmol, 62% for both
with NaHCO3. The collected organic layer was dried over diastereoisomers).
MgSO4, filtered and concentrated under reduced pressure to
afford the crude product as a pale yellow solid. Purification by
flash column chromatography, eluting with a gradient of 1 : 5
EtOAc : CH2Cl2–1 : 3 EtOAc : CH2Cl2, yielded the title com-
Fast eluting diastereoisomer. 1H NMR (400 MHz, CDCl3)
δ 8.98 (s, 1 H, NH), 8.71 (s, 1 H, H2), 8.25 (s, 1 H, H8),
7.99–7.92 (m, 2 H, o-NBz), 7.58–7.51 (m, 1 H, p-NBz), 7.50–7.42
(m, 2 H, m-NBz), 7.37–7.31 (m, 2 H, m-Ph), 7.27–7.08 (m, 7 H,
o-Ph, p-Ph, o-Ar2), 6.74–6.70 (m, 4 H, m-Ph), 6.35 (dd, J = 2.3,
6.8 Hz, 1 H, H1′), 4.20–4.12 (m, 1 H, H4′), 3.79–3.73 (m, 1 H,
pound as a white amorphous solid (2.34 g, 2.74 mmol, 82%
for both diastereoisomers).
Fast eluting diastereoisomer. 1H NMR (400 MHz, CDCl3)
H3′), 3.71 (s, 6 H, OMe), 3.63–3.50 (m, 5 H, OCH2, NCH(CH3)2,
δ 12.02 (br s, 1 H, NH), 8.91 (s, 1 H, NH), 7.79 (s, 1 H, H8),
H5′), 3.35 (dd, J = 4.6, 10.6 Hz, 1 H, H5′), 3.04 (ddd, J = 2.3, 7.2,
7.37–7.30 (m, 2 H, m-Ph), 7.26–7.07 (m, 7 H, o-Ph, p-Ph, o-Ar2),
13.7 Hz, 1 H, H2′), 2.62 (ddd, J = 7.0, 10.2, 14.0 Hz, 1 H, H2′),
6.73–6.66 (m, 4 H, m-Ar2), 6.12–6.04 (m, 1 H, H1′), 4.18–4.10
2.37 (t, J = 6.3 Hz, 2 H, CH2CN), 1.13 (d, J = 6.8 Hz, 6 H, NCH-
(m, 1 H, H4′), 3.69 (s, 6 H, OMe), 3.66–3.28 (m, 7 H, H3′,
(CH3)2), 1.07 (d, J = 6.5 Hz, 6 H, NCH(CH3)2). 31P NMR
OCH2, NCH(CH3)2, 2H5′), 2.90 (ddd, J = 3.8, 8.0, 13.9 Hz, 1 H,
(162 MHz, CDCl3) δ 164.5. 13C NMR (101 MHz, CDCl3) δ 165.3
H2′), 2.58–2.31 (m, 4 H, H2′, CH2CN, CH(CH3)2), 1.28–1.04
(NHBz CvO), 156.6 (p-Ar2), 153.0 (C2), 150.6 (C6), 147.5 (C4),
(m, 18 H, NCH(CH3)2, CH(CH3)2). 31P NMR (162 MHz, CDCl3)
144.9 (ipso-Ph), 142.1 (C8), 139.8 (ipso-Ar2), 133.7 (ipso-NBz),
δ 161.9. 13C NMR (101 MHz, CDCl3) δ 179.1 (NHibu CvO),
133.1 (p-NBz), 130.8–127.3 (o-Ph, m-Ph, p-Ph, o-Ar2, o-NBz,
158.8 (p-Ar2), 156.0 (C6), 148.0 and 147.7 (C2, C4), 145.5 (ipso-
m-NBz), 125.0 (C5), 115.1 (CN), 113.5 (m-Ar2), 87.3 (CPh(Ar)2),
Ph), 137.9 (C8), 136.1 (ipso-Ar2), 130.7–127.2 (o-Ph, m-Ph, p-Ph,
84.5 (C4′), 82.9 (C1′), 62.7 (C5′), 60.9 (OCH2), 55.6 (OMe), 47.0
o-Ar2), 122.3 (C5), 117.8 (CN), 113.6 (m-Ar2), 87.1 (CPh(Ar)2),
(NCH(CH3)2), 42.9 (C2′), 39.4 (C3′), 24.6 (NCH(CH3)2), 20.5
86.7 (C4′), 85.4 (C1′), 64.1 (C5′), 60.4 (OCH2), 55.6 (OMe), 47.3
(CH2CN).
(NCH(CH3)2), 42.1 (C2′), 41.1 (C3′), 36.5 (CH(CH3)2), 23.1 (NCH-
Slower eluting diastereoisomer. 1H NMR (400 MHz, CDCl3)
(CH3)2), 20.5 (CH2CN), 19.0 (CH(CH3)2).
Slower eluting diastereoisomer. 1H NMR (400 MHz, CDCl3)
δ 8.97 (s, 1 H, NH), 8.71 (s, 1 H, H2), 8.23 (s, 1 H, H8),
7.98–7.93 (m, 2 H, o-NBz), 7.55–7.52 (m, 1 H, p-NBz), 7.49–7.43
(m, 2 H, m-NBz), 7.34–7.30 (m, 2 H, m-Ph), 7.24–7.10 (m, 7 H,
o-Ph, p-Ph, o-Ar2), 6.72–6.68 (m, 4 H, m-Ph), 6.39 (dd, J = 2.4,
6.8 Hz, 1 H, H1′), 4.21–4.13 (m, 1 H, H4′), 3.77–3.71 (m, 3 H,
OCH2, H3′), 3.69 (s, 6 H, OMe), 3.59–3.44 (m, 3 H, NCH(CH3)2,
H5′), 3.33 (dd, J = 4.6, 10.6 Hz, 1 H, H5′), 3.10 (ddd, J = 2.3, 7.2,
13.7 Hz, 1 H, H2′), 2.71–2.61 (m, 1 H, H2′), 2.53 (t, J = 6.1 Hz,
2 H, CH2CN), 1.11 (d, J = 6.8 Hz, 6 H, NCH(CH3)2), 1.00 (d, J =
6.5 Hz, 6 H, NCH(CH3)2). 31P NMR (162 MHz, CDCl3) δ = 160.9.
13C NMR (101 MHz, CDCl3) δ 165.3 (NHBz CvO), 156.6
(p-Ar2), 153.1 (C2), 150.6 (C6), 147.7 (C4), 144.9 (ipso-Ph), 142.1
(C8), 139.9 (ipso-Ar2), 133.7 (ipso-NBz), 133.1 (p-NBz),
130.8–127.3 (o-Ph, m-Ph, p-Ph, o-Ar2, o-NBz, m-NBz), 125.0
(C5), 115.1 (CN), 113.5 (m-Ar2), 94.9 (CPh(Ar)2), 84.5 (C4′), 82.9
(C1′), 62.7 (C5′), 60.9 (OCH2), 55.6 (OMe), 47.0 (NCH(CH3)2),
42.9 (C2′), 39.5 (C3′), 24.6 (NCH(CH3)2), 20.5 (CH2CN).
δ 11.99 (br s, 1 H, NH), 8.86 (s, 1 H, NH), 7.79 (s, 1 H, H8),
7.37–7.31 (m, 2 H, m-Ph), 7.26–7.07 (m, 7 H, o-Ph, p-Ph, o-Ar2),
6.73–6.66 (m, 4 H, m-Ar2), 6.12–6.06 (m, 1 H, H1′), 4.19–4.10
(m, 1 H, H4′), 3.79–3.30 (m, 12 H, OCH2, OMe, H3′, NCH-
(CH3)2, H5′), 3.24 (dd, J = 5.1, 10.5 Hz, 1 H, H5′), 2.99 (ddd, J =
4.6, 7.6, 13.4 Hz, 1 H, H2′), 2.61–2.31 (m, 4 H, H2′, CH2CN, CH-
(CH3)2), 1.23–0.99 (m, 18 H, NCH(CH3)2, CH(CH3)2). 31P NMR
(162 MHz, CDCl3) δ 160.7. 13C NMR (101 MHz, CDCl3) δ 179.1
(NHibu CvO), 158.9 (p-Ar2), 156.0 (C6), 148.0 and 147.7 (C2,
C4), 145.4 (ipso-Ph), 137.9 (C8), 136.1 (ipso-Ar2), 130.6–127.3
(o-Ph, m-Ph, p-Ph, o-Ar2), 122.4 (C5), 117.9 (CN), 113.6 (m-Ar2),
87.1 (CPh(Ar)2), 86.7 (C4′), 85.4 (C1′), 64.1 (C5′), 60.6 (OCH2),
55.6 (OMe), 47.3 (NCH(CH3)2), 42.1 (C2′), 41.1 (C3′), 36.5 (CH-
(CH3)2), 23.0 (NCH(CH3)2), 20.5 (CH2CN), 19.0 (CH(CH3)2).
ES-HRMS: C44H54N7O7PSNa requires 878.3441; [M + Na]+
878.3467 (3.0 ppm).
=
ES-HRMS: C47H52N7O6PSNa requires 896.3335; [M + Na]+
896.3337 (0.2 ppm).
=
N6-Benzoyl-5′-O-dimethoxytrityl-2′-deoxy-3′-thioadenosine-3′-
S-[(2-cyanoethyl)-(N,N-diisopropyl)]-phosphoramidite (7). A solu-
tion of 5-ethylmercapto-1H-tetrazole (0.25 M, 4.62 mL,) in
MeCN was added to
a stirred solution of N6-benzoyl-
5′-dimethoxytrityl-2′-deoxy-3′-thioadenosine (6, 1.04 g, 1.54 mmol)
in CH2Cl2 (20 mL) under N2. 2-Cyanoethyl N,N,N′,N′-tetraiso-
Acknowledgements
propylphosphorodiamidite (0.73 mL, 2.31 mmol) was then The work was financially supported by the EPSRC (EP/002464/
added dropwise over 5 min to the stirred solution of nucleo- 1) to JG and an EPSRC studentship to (MMP). We would like to
side. The reaction was allowed to stir at room temperature for thank Alan Mills and Moya McCarron (Liverpool) for obtaining
2 hours under N2. The resulting solution was subsequently mass spectra and Dr Inder Bhamra and Dr John Brazier for
quenched by addition of MeOH (3 mL) and then taken up in their assistance.
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16 J.-L. Mergny, A. De Cian, A. Ghelab, B. Saccà and L. Lacroix,
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