Aberrant cyclization affords a C-6 modified cyclic adenosine 5′-diphosphoribose analogue with biological activity in Jurkat T cells

Article abstract of DOI:10.1021/jm201127y

Two nicotinamide adenine dinucleotide (NAD⁺) analogues modified at the 6 position of the purine ring were synthesized, and their substrate properties toward Aplysia californica ADP-ribosyl cyclase were investigated. 6-N-Methyl NAD⁺ (6-N-methyl nicotinamide adenosine 5′-dinucleotide 10) hydrolyzes to give the linear 6-N-methyl ADPR (adenosine 5′-diphosphoribose, 11), whereas 6-thio NHD⁺ (nicotinamide 6-mercaptopurine 5′-dinucleotide, 17) generates a cyclic dinucleotide. Surprisingly, NMR correlation spectra confirm this compound to be the N1 cyclic product 6-thio N1-cIDPR (6-thio cyclic inosine 5′-diphosphoribose, 3), although the corresponding 6-oxo analogue is well-known to cyclize at N7. In Jurkat T cells, unlike the parent cyclic inosine 5′-diphosphoribose N1-cIDPR 2, 6-thio N1-cIDPR antagonizes both cADPR- and N1-cIDPR-induced Ca²⁺ release but possesses weak agonist activity at higher concentration. 3 is thus identified as the first C-6 modified cADPR (cyclic adenosine 5′-diphosphoribose) analogue antagonist; it represents the first example of a fluorescent N1-cyclized cADPR analogue and is a new pharmacological tool for intervention in the cADPR pathway of cellular signaling.

Full text of DOI:10.1021/jm201127y

Article
pubs.acs.org/jmc
Aberrant Cyclization Affords a C-6 Modified Cyclic Adenosine
5′-Diphosphoribose Analogue with Biological Activity in Jurkat T Cells
Christelle Moreau,† Tanja Kirchberger,‡ Bo Zhang,† Mark P. Thomas,† Karin Weber,‡ Andreas H. Guse,‡
and Barry V. L. Potter*,†
†Wolfson Laboratory of Medicinal Chemistry, Department of Pharmacy and Pharmacology, University of Bath, Claverton Down,
Bath, BA2 7AY, United Kingdom
‡Calcium Signalling Group, Department of Biochemistry and Signal Transduction, Center of Experimental Medicine,
University Medical Center Hamburg-Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany
S
* Supporting Information
ABSTRACT: Two nicotinamide adenine dinucleotide (NAD+)
analogues modified at the 6 position of the purine ring were
synthesized, and their substrate properties toward Aplysia
californica ADP-ribosyl cyclase were investigated. 6-N-Methyl
NAD+ (6-N-methyl nicotinamide adenosine 5′-dinucleotide 10)
hydrolyzes to give the linear 6-N-methyl ADPR (adenosine
5′-diphosphoribose, 11), whereas 6-thio NHD+ (nicotinamide
6-mercaptopurine 5′-dinucleotide, 17) generates a cyclic
dinucleotide. Surprisingly, NMR correlation spectra confirm this compound to be the N1 cyclic product 6-thio N1-cIDPR
(6-thio cyclic inosine 5′-diphosphoribose, 3), although the corresponding 6-oxo analogue is well-known to cyclize at N7. In Jurkat
T cells, unlike the parent cyclic inosine 5′-diphosphoribose N1-cIDPR 2, 6-thio N1-cIDPR antagonizes both cADPR- and N1-
cIDPR-induced Ca2+ release but possesses weak agonist activity at higher concentration. 3 is thus identified as the first C-6
modified cADPR (cyclic adenosine 5′-diphosphoribose) analogue antagonist; it represents the first example of a fluorescent N1-
cyclized cADPR analogue and is a new pharmacological tool for intervention in the cADPR pathway of cellular signaling.
cADPR is readily hydrolyzable at the N1 glycosidic bond
linkage to give ADPR in both neutral aqueous solution and
under physiological conditions,12,13 thus rendering chemical
synthesis of analogues challenging. The main choice has been
between total chemical synthesis14 and a chemoenzymatic ap-
proach15,16 developed earlier and modeled on the biosynthesis
of cADPR from NAD+. By combination of both approaches, a
large number of cADPR analogues have been synthesized over
the years and their Ca2+ release activities examined in several
systems, such as sea urchin egg homogenate (SUH) and T cells
inter alia. Modification at the 8-position with 8-amino and
8-bromo groups in particular has been shown to convert cADPR
from an agonist to an antagonist,16−19 although at high concen-
trations this does not apparently hold.20 However, 8-substituted
cyclic adenosine 5′-diphosphocarbocyclic ribose (8-X cADPcR)
analogues developed by Shuto21 were reported to be agonists in
sea urchin egg homogenates, indicating that 8-substitution alone
may not be responsible for antagonist activity. 3′-O-Methyl
cADPR was also found to have antagonist properties, at least in
SUH.13 The hydrolysis-resistant 7-deaza 8-bromo cADPR has
found several biological applications22,23 as a cell permeant
competitive antagonist.17,24,25 Analogues with structural mod-
ifications of the “northern” ribose12 showed agonist activities in
T cells, including those with more radical changes.26,27
INTRODUCTION
Cyclic adenosine 5′-diphosphoribose (cADPR, 1, Figure 1),
discovered by Lee et al. in 1987,1 is one of the principal second
■
Figure 1. C-6 modified cADPR analogues and numbering system.
messenger molecules that mobilize intracellular Ca2+ in a different
way to the well-established D-myo-inositol 1,4,5-trisphosphate
(Ins(1,4,5)P3),2 by gating the ryanodine receptor.3,4 The cADPR/
Ca2+ signaling system is active in diverse mammalian cellular
systems such as cardiac muscle, acinar cells, and plant cells.5
cADPR is a metabolite of nicotinamide adenine dinucleotide
(NAD+) and is produced enzymatically by ADP-ribosyl cyclases
(ADPRC). Its structure was fully characterized by Lee et al.6 as
a cyclic 18-membered dinucleotide featuring two glycosidic bonds
and a pyrophosphate linkage. Several excellent reviews dealing
with the chemistry of cADPR and the cADPR/Ca2+ signaling
system have appeared in recent years.3,5,7−11
Received: May 12, 2011
Published: January 16, 2012
© 2012 American Chemical Society
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Recently, we have also pioneered a synthesis of novel cADPR
derivatives that are highly stable both chemically and enzymati-
cally. The 8-bromo N1-cyclic inosine 5′-diphosphoribose (8-Br-
N1-cIDPR) thus obtained proved to be a novel agonist of Ca2+
release in intact cells.28,29 The opposed bioactivities of the
classical antagonist 8-Br-cADPR and 8-Br-cIDPR thus show
that the smallest of structural changes (i.e., NH2 → CO at
C-6, as with changes at C-8 and C-3′) can transform a cADPR
antagonist into an agonist. Furthermore, the enhanced stability of
8-Br-cIDPR allows direct chemical modification at the 8-position,
including to the parent N1-cIDPR, (2, Figure 1), which is
roughly equipotent to cADPR in permeabilized T cells.30 This
stable analogue was recently cocrystallized with the wild-type
ectoenzyme CD38, a multifunctional enzyme responsible for
the formation and metabolism of cADPR, providing insight into
substrate binding and the catalytic process.31
The chemoenzymatic approach relies on the selectivity of
Aplysia cyclase that, while promiscuous, may not always recog-
nize the required NAD+ analogue as substrate. Moreover, while
NAD+ analogues generally cyclize at N1, some are hydrolyzed
to the corresponding linear ADPR analogue. Another exception,
ethenonicotinamide adenine 5′-dinucleotide, in which the N1 posi-
tion is substituted, cyclizes through the N7 nitrogen by ADPRC.32
NHD+, a close analogue of NAD+ but bearing a 6-keto group on
the purine ring rather than an amino group, cyclizes at N7, giving
the fluorescent, biologically inactive N7-cIDPR, as indeed does the
nicotinamide adenosine 5′-dinucleotide (6-NMe-NAD+) has,
however, not yet been investigated. N1-cIDPR was prepared via
both 8-Br-NHD+ and 8-Br-N1-cIDPR to bypass the problem
encountered with N7 cyclization.30 To the best of our knowledge,
even a total synthesis was never attempted for such a class of
compound, presumably because of the difficulty of incorporating
an intact “northern” ribose into the molecule. In order to further
investigate the requirements for N1 versus N7 cyclization inter
alia, we report here syntheses of both 6-NMe-NAD+ and 6-thio
NHD+ and an investigation of their cyclization behavior.
RESULTS AND DISCUSSION
■
Synthesis of 6-N-Methyl NAD+. 6-NMe adenosine 7 was
earlier synthesized from 1-N-methyladenosine via a Dimroth
rearrangement that involves an opening and recyclization of the
adenine ring.34 We synthesized 7 by a different synthetic route
that could provide wider flexibility for other modifications longer
term. 6-Chloropurine 4 was first activated with TMS-triflate,
then condensed with tetraacetyl ribose 5 to produce the 6-chloro
protected nucleoside 6 in its β configuration (Scheme 1). Treat-
ment with methylamine hydrochloride followed by acetate re-
moval with methanolic ammonia produced 6-NMe adenosine 7
in good yield. Phosphorylation was achieved by using the esta-
blished POCl3/TEP method but, in contrast to other nucleosides,
both the 5′-monophosphate 8a and 3′,5′-bisphosphate 8b were
formed as a mixture. Subsequent treatment with triphenyl-
phosphine, morpholine, and dipyridyl disulfide produced two
morpholidates. The desired monomorpholidate 9a was then
easily isolated by ion-exchange chromatography and condensed
with β-NMN+ to form the target 6-NMe-NAD+ 10.
Incubation of 6-NMe-NAD+ 10 with Aplysia cyclase gene-
rated only the hydrolyzed product 6-NMe-ADPR 11, confirmed
1
by H NMR spectroscopy and a molecular ion of 572 in the
electrospray mass spectrum. The latter is in accord with the
structure of a linear nucleotide rather than the cyclic 6-NMe-
cADPR, as it differs by 17 mass units for the additional hy-
droxyl group. Moreover, the presence of two NMR doublets at
5.24 and 5.14 ppm represents a typical anomeric proton pattern
for the terminal ribose hydroxyl in the α and β configurations.
This result is perhaps not too surprising based upon another
report of the hydrolysis of C-6 substituted NAD+ analogues,
but these had more radical changes.7
It was initially assumed that cyclization of 6-NMe-NAD+ is
most likely blocked by the steric hindrance brought about by
the free rotation of the methyl group. However, an early review7
reported that the compound with an H at C-6 rather than an
amino group is apparently also unable to cyclize, indicating
that the cyclization catalyzed by Aplysia cyclase might (not
surprisingly) involve the participation of the C-6 amino group.
It may be possible that the steric hindrance of the methyl group
disturbs the electrophilic attack at the N1 position and there-
fore blocks the cyclization. It seems likely that the steric bulk
of the NMe might interfere with the approach to the known
covalent E179 enzyme−ribose intermediate for cyclization.35
For hydrolysis to occur requires the formation of the 6-NMe-
NAD+−enzyme covalent complex, which allows water to attack
the ribosyl C-1″ rather than N1. Another explanation could be
interference with the actual binding of 6-NMe-NAD+ to Aplysia
cyclase by reducing the H-bonding sites in the active site.
Synthesis of 6-Thio NHD+. 6-Thio NHD+ was first re-
ported by Atkinson et al.36 and synthesized by coupling 6-thio
IMP (14) with nicotinamide mononucleotide with dicyclohex-
ylcarbodiimide, a method developed by Todd et al.37 The low
Figure 2. N7 cyclization of NHD+ by Aplysia cyclase.
guanosine congener NGD+ (Figure 2).33 The reasons for this are
not fully understood. 6-Thio NHD+ (where a 6-mercaptopurine
replaces the hypoxanthine ring of NHD+) has yet to be explored
as a potential substrate for ADP-ribosyl cyclase. Since the only
modifications at position 6 in the cyclic nucleotide explored
to date (NH2 and CO) result in major changes in biological
activity, we anticipated that the 6-thio (CS) derivative in
particular would be of significant interest.
The synthesis of C-6 modified cADPR analogues has never
been straightforward. NAD+ analogues with substituents at C-6
such as SMe, NH(CH2)6NH2 or H all fail to cyclize when incu-
bated with ADPRC.7 The corresponding hydrolyzed product is
formed instead. The simple N-methylated analogue, 6-N-methyl
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a
Scheme 1. Synthesis of 6-NMe NAD+
a
Reagents and conditions: (i) TMSOTf, DBU, MeCN, 60 °C, 1 h; (ii) methylamine hydrochloride, DCM/EtOH/Et3N, 60 °C, overnight;
(iii) NH3/MeOH, 0 °C, 3 h; (iv) POCl3, TEP, H2O, 0 °C, 3 h; (v) PPh3, dipyridyl disulfide, morpholine, room temp, 2 h; (vi) β-NMN+, MnCl2 in
formamide, room temp, 48 h; (vii) Aplysia californica, 25 mM HEPES (pH 7.4), room temp.
a
Scheme 2. Synthesis of 6-Thio NHD+
a
Reagents and conditions: (i) dimethoxypropane, acetone, HClO4, 20 min, room temp; (ii) POCl3, TEP, H2O, 0 °C, 1 h; (iii) dinitrofluorobenzene,
Et3N, MeCN, room temp, 1 h; (iv) diisopropyl-di-tert-butylphosphoramidite, tetrazole, DCM, room temp, 1 h, then mCPBA, −78 °C, 10 min;
(v) 10% mercaptoethanol in MeCN + 1% DIPEA, room temp, 1 h; (vi) 50% aq TFA, room temp, overnight; (vii) morpholine, PPh3, dipyridyl
disulfide, DMSO, room temp, 1 h; (viii) β-NMN+, 0.2 M MnCl2 in formamide, room temp, overnight.
yield and lack of structural data in the initial report prompted
us to investigate a new, reliable and more modern, route to
6-thio NHD+. The key compound is 14, which was previously
synthesized mostly in the 1960s more or less successfully.38,39
We initially reasoned that the synthesis of 14 should be easily
accomplished by selective phosphorylation of 6-thioinosine 12
by adaptation of a published method40 that we have used very
successfully on a wide range of nucleosides. However, treat-
ment of 6-thioinosine with POCl3 in TEP was unsuccessful
because of the poor solubility of the starting nucleoside in TEP
(Scheme 2). An isopropylidene protecting group at the 2′,3′-
hydroxyls was then considered to have the double advantage
of solving the solubility issue and being removable during the
phosphorylation procedure, which is carried out under acidic
conditions. Phosphorylation at the 5′-position proved to be very
difficult. Indeed, HPLC analysis of the quenched reaction
showed a complex mixture of products that could not be iso-
lated by reverse-phase chromatography. We reasoned that there
was a mixture of protected and deprotected material as well as
products phosphorylated at the 5′-OH, the sulfur atom, or both,
since the sulfur is more nucleophilic than the oxygen.
Phosphoramidite chemistry has been reported on 2′,3′-O-
isopropylidene-6-thioinosine 1341 using N,N-diisopropyl-di-
tert-butylphosphoramidite as a phosphitylating reagent.42 Both
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tert-butyl and isopropylidene protecting groups could then be
cleaved under acidic conditions. However, in our hands the
phosphorylation step could not be repeated, as the starting
material is insoluble in most organic solvents except DMF and
DMSO, and therefore only the starting material was recovered
after 24 h with no sign of product.
Thio-substituted nucleotides are particularly useful in molec-
ular biology, as they have been incorporated into oligonucleo-
tides by chemical methods and used for postsynthetic modi-
fication.43−46 During the former process, the sulfur is always
protected to avoid unnecessary side reactions during phosphi-
tylation and oxidation of the phosphite.47 Since the synthesis of
6-thio IMP is hampered by the lack of solubility of 6-thioino-
sine, protecting the sulfur should further solve the solubility
issue encountered. Various protecting groups have been used
such as cyanoethyl, which can be removed by treatment with
DBU. Although the cyanoethyl group has been used extensively,
we decided to utilize the 2,4-dinitrophenyl (DNP) group, as
previous studies have shown that it can easily be removed with
mercaptoethanol under very mild alkaline conditions.48
cation step is avoided during which, from our experience, some
product often gets unavoidably lost. 14 was thus obtained in a
satisfying 48% yield after a four-step procedure from 2′,3′-O-
isopropylidene-6-thioinosine.
With 6-thio IMP in hand, we proceeded to synthesize the key
intermediate 6-thio NHD+ 17. The synthesis of the pyrophos-
phate was achieved using a procedure initially reported by
Moffatt50 and later improved by Lee et al.51 that relies on the
coupling of a sugar phosphate with a nucleotide phosphoro-
morpholidate in the presence of a Lewis acid. We have previously
used this method to successfully generate various NAD+ and
NHD+ analogues in relatively high yield.30,52 6-Thio IMP was first
activated using a combination of morpholine/dipyridyl disulfide
and triphenylphosphine to yield the morpholidate that was then
condensed with β-NMN+ with MnCl2 as Lewis acid. A 31P NMR
shift of around −10 ppm clearly indicated the formation of a
pyrophosphate linkage. Purification on reverse-phase chromatog-
raphy afforded the desired 6-thio NHD+ 17 as the sole product.
6-Thio NHD+ was then incubated with Aplysia californica
cyclase. There are several possible outcomes for this reaction
(Figure 3), i.e., formation of 6-thio N1-cIDPR, 6-thio N7-cIDPR
as for the oxo congener, 6-thio IDPR (6-thioinosine 5′-diphos-
phate ribose), and it is possible to envisage cyclization on to the
sulfur to give the 6-thio-S-cIDPR. The new product displayed a
molecular ion of 557 in the ES− mass spectrum, which ruled out
the formation of both the hydrolyzed product 6-thio IDPR and
surprisingly also the N7 cyclized product 6-thio N7-cIDPR, for
which a mass of 575 and 558, respectively, would be expected.
1H NMR spectroscopy showed a chemical shift of 5.2 ppm for
H-2′, a typical value for a nucleotide in the syn conformation,
which also ruled out the formation of 6-thio N7-cIDPR. Addi-
tionally, a C-6 13C chemical shift of 175 ppm is typical of a thione
form (thiol, ∼155 ppm), therefore ruling out the putative sulfur-
cyclized product 6-thio S-cIDPR as a possibility. Crucially, a
gHMBC spectrum showed cross-peaks between the H-2 proton
of the purine ring (δ 9.32 ppm) and the anomeric carbon C-1″ of
the “northern” ribose (δ 94.9 ppm) and between the anomeric
The synthesis of 6-thio NHD+ 10 is outlined in Scheme 2.
2′,3′-O-Isopropylidene-6-thioinosine 13 was prepared in a very
high yield (86% over five steps) using published methods start-
ing from inosine 12.41,49 Protection of the sulfur was achieved
by treatment of 2′,3′-O-isopropylidene-6-thioinosine with tri-
ethylamine and 2,4-dinitrofluorobenzene. The yellow product
15 obtained after flash chromatography could then be selec-
tively phosphitylated at the 5′-hydroxyl using the tert-butyl
protected phosphitylating reagent in high yield to give 16.
Removal of the DNP protecting group was accomplished by
treatment with mercaptoethanol under mild alkaline conditions.
Careful purification by flash chromatography led to a very clean
product in 75% yield. Next, both the isopropylidene and tert-
butyl protecting groups were removed very cleanly by treat-
ment with 50% aqueous TFA to generate the desired 6-thio
IMP 14 in 86% yield, without further purification. This clearly
is the advantage of the DNP group, as an ion-exchange purifi-
Figure 3. Four possible outcomes from the incubation of 6-thio NHD+ with ADPRC.
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Table 1. Conformational Analysis of NAD+ Analogues and
Their Respective Cyclic Dinucleotides
proton H-1″ (δ 6.65 ppm) with the carbon C-2 (δ 144.9 ppm)
and C-6 (δ 175.5 ppm) of the nucleobase (Figure 3). Finally,
the 31P−31P coupling constant for the pyrophosphate linkage in
3 showed J = 11.8 Hz, similar to that of cADPR and cIDPR
(13.5 53 and 12.5 30 Hz, respectively), whereas the 31P−31P cou-
pling of the linear IDPR generally has a much higher frequency
(∼20 Hz). These analytical data thus provide evidence of a
successful and surprising cyclization of 6-thio NHD+ into the
corresponding 6-thio N1 cIDPR 3 (Figure 4).
C2′ endo
(%)
a
a
b
c
a
a
H-1′
H-2′
Δ1′‑2′ conf J1′,2′
J3′,4′
cADPR
5.80
6.19
5.89
6.07
5.20
3.90
5.18
4.63
5.2
0.6
syn
anti
syn
anti
syn
5.6
3.2
nd
nd
nd
2.2
64
30
61
30
74
N7-cIDPR
N1-cIDPR
N7-cGDPR
2.29
0.71
1.44
0.8
3.0
6.1
2.8
6.3
6-thio N1-cIDPR 6.0
a
Data obtained from various sources: cADPR;59 N7-cIDPR and N7-
cGDPR;59−61 N1-cIDPR.30 Difference in chemical shifts between
H-1′ and H-2′. Preferred glycosidic bond conformation.
b
c
to be correctly aligned in the active site but the substrate
is hydrolyzed rather than cyclized at N7. The enzyme can
also cyclize NHD+, NGD+, and NXD+ to their respective
N7 cyclized dinucleotide N7-cIDPR, N7-cGDPR, and N7-
cXDPR.33 In those cases, the most nucleophilic N7 nitrogen in
guanine/hypoxanthine is often invoked to rationalize cycliza-
tion at this position.33
Figure 4. 1H/13C correlations based on gHMBC spectrum supporting
the formation of the N1−C1″ bond.
However, nucleophilicity is not the only factor to consider.
We previously made correlations showing that the conforma-
tion about the glycosidic bond of NHD+ analogues appears to
play a role in their cyclization.52 We argued that a syn confor-
mation about the glycosidic bond is key for a successful
cyclization at N1, i.e., that the enzyme seems to utilize the linear
precursor in its prearranged conformation (at least in the case
of the hypoxanthine series and presumably the guanosine series
as well). Indeed, NHD+ (anti) cyclizes at N7 (product also in
anti conformation), 8-X-NHD+ (syn) cyclizes at N1 (products
in syn conformation), and 7-deaza NHD+ (anti and no
possibility to cyclize at N7) hydrolyzes to the linear 7-deaza
IDPR.52 However, the NAD+ analogues cyclize at N1 regardless
of their conformation; NAD+ (anti), 8-X NAD+ (syn), and
7-deaza NAD+ (anti) cyclize at N1, all forming products in their
syn conformation. When 6-thio NHD+ is used as a substrate,
the enzyme converts it into 6-thio N1-cIDPR, therefore sug-
gesting that the enzyme utilizes this substrate in the same way
as NAD+ but clearly differently from NHD+, although NHD+
and 6-thio NHD+ are structurally very similar.
Lee et al. have intensively investigated the structures of both
Aplysia ADP-ribosyl cyclase and human CD38 to unravel the
catalytic mechanism of the NAD+ cyclization and cADPR hy-
drolysis reactions. Recently, they suggested that the cyclization
reaction of NAD+ analogues occurs through a four-step sequence
where both residues Tyr-81 and Phe-174 play an instrumental
role by stabilizing the nucleobase through π-stacking interac-
tions in a folded conformation so that cyclization can occur.63,64
Mutagenesis confirmed that Phe-174 was likely to be respon-
sible for folding the linear substrate in the correct confor-
mation. However, this is only true for adenine-based substrates;
mutagenesis did not affect the cyclization of NGD+ to N7-
cGDPR. Therefore, it is likely that two residues are responsible
for the different base cyclization sites (N1 versus N7) and that
residue Phe-174 may also be involved in the cyclization of
6-thio NHD+ to 6-thio N1-cIDPR.
The photochemical properties of 6-thio N1-cIDPR 3 were
also examined. While adenine and guanosine based nucleotides
have an absorbance maximum of around 260 nm, replacement
of the oxygen by a sulfur atom shifts the UV absorbance spec-
trum to 320−340 nm. 6-Thio cIDPR seems to be no exception.
The UV spectrum in water exhibited intense absorption at
320 nm (λmax) with an extinction coefficient (ε) of 18 600
mol−1 dm3 cm−1 (data not shown). A smaller peak at 267 nm
was also observed. More interestingly, 6-thio cIDPR displays
fluorescent properties in water at room temperature. When it is
excited at 335 nm (just above the maximum UV absorption),
an emission spectrum with a single peak at 415 nm was ob-
served (see Supporting Information). In contrast, no fluores-
cence was observed by the related N1-cIDPR (data not shown).
Thus far, only N7-cyclized dinucleotides such as N7-cIDPR,
N7-cGDPR, and etheno cyclic ADP diphosphoribose have been
demonstrated to be fluorescent, and this property was used to
rule out the formation of N7-cyclic product when no fluores-
cence was observed.33 To the best of our knowledge, 6-thio
N1-cIDPR is therefore the first fluorescent N1-cyclized cADPR
analogue and this property may find applications in cADPR
binding protein biochemistry.
Conformational Analysis. There have been reports that
the Ca2+-release activity and antagonism54,55 of cADPR ana-
logues may be linked to their conformation in solution. The
major puckering mode of cADPR is C2′ endo in the N9 ribose
moiety with a syn conformation about the N9 glycosidic bond.6
The furanose ring is in equilibrium between C2′ endo/C3′ endo
1
forms, and the ratio can be calculated from H NMR data
following the equation [C2′ endo] = [J1′,2′/(J1′,2′ + J3′,4′)] × 100.56
Also, the H-2′ chemical shift can be used as an indicator of
glycosidic bond conformation.57,58 Therefore, from our NMR
data, 6-thio cIDPR exhibits a 74% C2′ endo puckering with a syn
conformation about the glycosidic bond (Table 1), which is
consistent with that observed by both cADPR and N1-cIDPR.
Mechanistic Study. ADP-ribosyl cyclase can use different
substrates to produce structurally distinct products. The
cyclization of NAD+ analogues usually takes place at the N1
position of the adenine ring. However, NAD+ analogues, which
have their N1 position blocked (e.g., etheno NAD+), cyclize at
N7.32 If N1 is free but is electronically deactivated (e.g., in 2-
fluoro NAD+62) and the purine is still adenine, then this seems
With the help of molecular modeling, we investigated further
how NAD+ cyclization to cADPR may occur. The ligand in the
ribo-2′-F-NAD+·ADPRC structure (PDB code 3I9O)63 was
manually manipulated by rotating individual bonds to approxi-
mate the position the adenine would have to be in to attack
C-1″ and form the N1-cyclized product (Figure 5). In order
for NAD+ to cyclize at N1, the adenine must rotate around the
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Figure 5. Two views of a model of the position of the adenine of the intermediate from ribo-2′-F-NAD+·ADPRC complex immediately prior ring
closure. The hydrogen bond from the base to Glu-98 is shown. The model was built by rotating individual bonds in the PDB 3I9O structure (ribo-2′-
F-NAD+·ADPRC) using the Schrodinger software running under Maestro, version 9.0.111. The color scheme for the ligand is as follows: pink,
̈
carbon; blue, nitrogen; red, oxygen; white, hydrogen; light blue, fluorine; orange, phosphorus. The color scheme for the residues is the same as for
the ligand except the carbons are green.
N9/C-1′ bond to adopt the syn configuration. It can then
approach the C-1″ by stacking with Trp-140 and having the
6-amino group forming at least one hydrogen bond to Glu-98
to further orient and hold the adenine in position. This pro-
posed mechanism has some support from the kinetic data for
Glu-98 mutants, as indeed mutation of this residue was shown
to reduce cyclase activity.65 NHD+ (or NGD+) would be unable
to form the hydrogen bonds to Glu-98, and the lack of stabili-
zation may partly explain the lack of cyclization at N-1.
pH there will be a mixture of four tautomeric forms for
the oxygen-containing compound (Figure 7) while there will
Exactly why 6-thio NHD+ cyclizes at N1 is unknown at
present. It is intuitively clear that in order for 6-thio NHD+ to
cyclize, the enzyme should be capable of stabilizing it in its thiol
form, although the mercaptopurine base normally exists pre-
dominantly in its thione form in solution.66 In the case where
the protonation state plays a role in the reaction mechanism, the
pKa values of N1, N7, and X1 (NH2, OH, SH) and X2 (O,
NH, S) were calculated computationally (Figure 6; see
Figure 7. Presumed different forms of the oxygen containing com-
pound that may be present at physiological pH.
be only two forms of the nitrogen- and sulfur-containing
compound.
Previously, we argued that the deciding interaction through
the adenine ring was via hydrogen bond donation from the
base, since the hypoxanthine base has only hydrogen acceptor
character in its keto form. However, the 6-thiopurine base has
virtually no hydrogen bond acceptor or donor character, which
may imply that hydrogen bonding is unimportant at C-6 and
that cyclization is influenced by other factors such as the size of
the substituent (the sulfur atom is larger than the nitrogen and
oxygen atoms; see Supporting Information), the conformation
about the glycosidic bond (for the oxygen containing nucleo-
tides), electronic properties, or a combination of these. It may
be that an H-bonding interaction with the enzyme locks the
purine of NHD+ in the anti form, leading to N7 cyclization.
However, this interaction could be very weak or nonexistent
with the 6-thiopurine, allowing it to rotate to facilitate N1
cyclization. In the case of 8-Br NHD+ the strong orienting
effect of the bromo group may be enough to overcome such
H-bonding. Protein crystallography with such a ligand may
reveal those interactions as important for catalysis to occur.
Pharmacology. The Ca2+ release activity of the newly
synthesized 6-thio cIDPR 3 was evaluated fluorimetrically in
Figure 6. Tautomeric form of purines.
also Supporting Information). The calculations were done on
the nucleoside (adenosine, inosine, and 6-thioinosine) and each
respective tautomer; we presume that it is reasonable to pro-
pose that these values would follow the same trend for the
corresponding NAD/NHD analogue.
For three of the possible ionization states, the program was
unable to calculate a pKa, presumably because they are too
extreme. At physiological pH, most of these ionizable groups
are going to be either fully protonated or deprotonated, the
exception being for the oxygen-containing compound (see
Supporting Information). In the A tautomer the X1 ionization
(OH → O−) has a pKa of 6.56, and in the B tautomer the N1
ionization (NH → N−) has a pKa of 8.67. These two values are
within the range of physiological pH, which means that at this
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permeabilized Jurkat T cells, and the results are shown in
Figure 8. Because of the intrinsic fluorescence of 6-thio cIDPR,
induced Ca2+ release was observed (Figure 8c). While the IC50
for cADPR induced Ca2+ release was between 30 and 100 μM, a
somewhat better inhibition of Ca2+ release induced by N1-
cIDPR 2 was obtained (IC50 ≈ 3 μM). We thus demonstrate
here that a simple modification from a CO to CS bond at
C-6 weakens agonist activity but enhances antagonist activity in
Jurkat T cells. Antagonist activity was specifically strong when
N1-cIDPR 2 was used to trigger Ca2+ release, indicating that
the oxygen group in cIDPR can be replaced at lower 6-thio
cIDPR 3 concentrations compared to the situation where Ca2+
release was induced by cADPR (Figure 8c). The most likely
explanation for this differential effect of 6-thio cIDPR 3 is differ-
ences in protonation pattern and hydrogen bonding interactions.
The amino group in cADPR can be a hydrogen bond donor or
acceptor, whereas the oxygen group in cIDPR is a good hy-
drogen bond acceptor. Obviously, 6-thio cIDPR 3 competes
more effectively with cIDPR for binding to the cADPR receptor.
Taken together, these results show that replacement of the
keto moiety by a more hydrophobic thione does interfere with
the functional consequences of binding of the ligand to its
receptor. There is an obvious role in the C-6 group in antago-
nizing Ca2+ release, if not directly then through subtle confor-
mational effects or other. It may well be that a change from a
hypoxanthine to a mercaptopurine results in a different proto-
nation pattern and hydrogen bonding interactions. The amino
group in cADPR can be a hydrogen bond donor or acceptor,
whereas the oxygen group in cIDPR is a good hydrogen bond
acceptor, as opposed to the sulfur atom. A C2′-endo/syn con-
formation is obviously not the sole factor that governs agonist/
antagonist activity, since all three nucleotides cADPR, cIDPR,
and 6-thio cIDPR have the same global conformation, although
one can suggest that a C3′-endo/anti conformation (as in N7-
cIDPR and N7-cGDPR, Table 1) would most likely lead to
inactive compounds. It is also well-known that the hydrophobicity
of sulfur leads to differences in the associated water shell. Thus, it
may be that differential arrangements of additional water mole-
cules in the receptor binding site can influence the receptor
machinery.
Figure 8. 6-Thio-cIDPR-induced Ca2+ release and effect of 6-thio-
cIDPR on cADPR and cIDPR-induced Ca2+ release in permeabilized
Jurkat T cells. Jurkat T cells were permeabilized, and [Ca2+] was
measured in the presence of Fluo-3, ATP, and an ATP regenerating
system as detailed in the Experimental Section. (a) Ca2+ release
induced by addition of 6-thio-cIDPR. Characteristic tracings from
representative experiments are shown. (b) Concentration−response
curve of 6-thio-cIDPR induced Ca2+ release. Results represent Ca2+
increase over baseline (Δ values) expressed as mean SD (n = 4−7)
of single tracings. (c) The inhibitory effect of 6-thio-cIDPR was esti-
mated by previous addition of 6-SH-cIDPR and subsequent addition
of either cIDPR or cADPR. Since 6-thio-cIDPR also elicited a weak
agonist effect on its own (a, b), calculation of its antagonist effect on
cADPR (30 μM) or cIDPR (30 μM) was carried out by subtracting
the corresponding agonist data (data in part b). Concentration−
response curves represent the mean SD (n = 3) of single tracings. A
one-phase exponential decay was used to fit curves; r2 was 0.6486 for
cADPR and 0.7107 for cIDPR.
Although surprising, this is not the first time that Ca2+ release
has been observed at higher concentrations. Indeed, we pre-
viously reported that the classical and widely used antagonists
such as 8-bromo cADPR and 8-amino cADPR also show such
unexpected agonist activity at high concentration.20 One of the
reasons invoked to explain this earlier unobserved effect is that
there may be two previously unrecognized different sites of
action, such as an additional lower affinity nucleotide binding
site, which causes Ca2+ mobilization distinct from the high affi-
nity cADPR site. This would not have been noticed with the
parent cIDPR, since it only acts as a potent agonist, but is again
revealed with the structurally related 6-thio cIDPR.
Fluo3 was used as a Ca2+ indicator to avoid any possible inter-
ference. At low concentration, 6-thio cIDPR 3 did not stimulate
Ca2+ release in T cells. Ca2+ release was, however, observed at
significantly higher concentration (100 μM and above, Figure 8a
and Figure 8b). In comparison with N1-cIDPR 2, weaker agonist
activity was observed for 6-thio cIDPR 3; however, in the cur-
rent series of experiments the permeabilized cell preparations
responded also somewhat more weakly to the naturally occur-
ring cADPR 1. Regarding antagonism by 6-thio cIDPR 3 a
concentration-dependent effect on cADPR and N1-cIDPR 2
The C-6 position in cADPR may therefore be revealed as an
attractive new site to imbue antagonistic properties in a cADPR
analogue. Its proximity to the crucial N1−C1″ linkage makes
it likely that structural modifications could lead to modulation
of activity of cADPR binding proteins. As such, it will be of
interest to develop new ways to approach the synthetic prob-
lem of C-6 modification. Although the present result has been
achieved somewhat unexpectedly via a chemoenzymatic route,
it seems most likely that greater flexibility in this regard will be
offered by total synthetic approaches in the future.
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fixed wavelength UV-1 optical unit (280 nm). The following puri-
fication methods were employed: LiChroprep RP-18 equilibrated with
0.05 M TEAB buffer (pH 6.0−6.4), gradient of 0.05 M TEAB buffer
against MeCN at 5 mL/min, Q-Sepharose washed with H2O, gradient
of 1 M TEAB buffer (pH 7.1−7.6) against H2O at 5 mL/min, and
AG MP-1 washed with H2O, gradient 150 mM TFA against H2O at
3 mL/min. Synthetic phosphates were assayed by an adaptation of the
Briggs phosphate test.68
Computational Details. cADPR, N1-cIDPR, and 6-thio N1-cIDPR
were built using Sybyl-X 1.1.1, and the volumes were calculated. The
pKa values were calculated using the Sparc online calculator http://
sparc.chem.uga.edu/sparc/.
Pharmacology. Materials. Fluo-3 was purchased from Molecular
Probes. Saponin and KH2PO4 were obtained from Fluka. ATP, crea-
tine phosphate, EGTA, Tris, and NaCl were provided from Sigma
Aldrich. MgCI2, CaCl2, and KCl were procured from Merck Chemicals.
HEPES was purchased from Biomol. Creatine kinase was obtained
from Roche. Culture medium reagents were supplied by Invitrogen or
Biochrom.
Cell Culture. Jurkat T-lymphocytes (subclone JMP) were cultured
as described previously69 at 37 °C in the presence of 5% CO2 in RPMI
1640 medium containing Glutamax I and HEPES (25 mM) and
supplemented with 7.5% (v/v) NCS (newborn calf serum), 100 units/
mL penicillin, and 100 μg/mL streptomycin.
CONCLUSION
■
In summary, we present a synthetic focus upon enzymatic C-6
structural modifications with a view to their enzymatic incor-
poration into cADPR. The substituted N6-NAD+ analogue 6-
NMe-NAD+, upon incubation with Aplysia cyclase, hydrolyzes
to the linear product 6-NMe-ADPR, presumably because of steric
hindrance and/or reduction of hydrogen bonding interaction with
the enzyme. A high yielding route also generated another C-6
modified analogue, 6-thio NHD+. Surprisingly, this analogue
cyclizes at N1 in a similar manner to NAD+, to give the novel
fluorescent cADPR analogue 6-thio N1-cIDPR, although the
parent oxo-congener NHD+ cyclizes at N7. A mechanistic study
implies a more complex explanation than simply a hydrogen
bonding interaction, as the enzyme recognizes this thio deriva-
tive as a normal substrate, although it is structurally closer to
NHD+ than it is to NAD+. Biological evaluation in permeabi-
lized Jurkat T cells reveals that this compound is decreased
in its agonist activity relative to cIDPR but possesses new anta-
gonist activity against both cADPR- and cIDPR-induced Ca2+
release, showing that substitution of the oxygen of the hypo-
xanthine ring for sulfur interferes with the functional con-
sequences of ligand binding to its receptor. Like two known
classical antagonists, 6-thio cIDPR shows agonist activity at
high concentrations. Antagonist activity in cADPR analogues
has thus now been demonstrated in compounds substituted
at the 8-position of the purine, the 3′-hydroxyl group of the
“southern” ribose, and now for the first time in a compound
with a C-6 structural modification. Both the fundamental causes
of antagonism and the switch to agonism at higher concen-
tration in some cases thus seem highly complex and seem to
defy a simple explanation. The present study demonstrates,
however, that future synthetic efforts to facilitate further C-6-
structural modification should be a fruitful enterprise in eluci-
dating cADPR SAR and defining useful new tools for dissection
of its signaling pathway.
Ca2+ Release Experiments in Permeabilized Cells. Permeabi-
lized cells were prepared as described,70 and the Ca2+ concentration
was measured by the use of Fluo-3. In brief, cells were permeabilized in
the presence of saponin (55 μg/mL) for 20 min in an intracellular
buffer (20 mM HEPES, 110 mM KCI, 2 mM MgCI2, 5 mM KH2PO4,
10 mM NaCI, pH 7.2) at 37 °C. An aliquot containing 1 × 108 cells
was transferred to a cuvette, and fluorescence was measured in a Hitachi
F-2000 spectrofluorometer (excitation 504 nm, emission 524 nm) at
37 °C in the presence of Fluo-3 (1 μM) with continuous stirring.
Reuptake of Ca2+ into stores was achieved by addition of ATP (1 mM),
creatine phosphate (20 mM), and creatine kinase (20 U/mL). At the
end of each experiment, the free Ca2+ concentration was calibrated
by addition of CaCl2 and subsequently by addition of EGTA/Tris
and calculated by using the following equation: [Ca2+] = Kd(F − Fmin)/
(Fmax − F) where Fmin is the fluorescence intensity in the absence of
Ca2+, Fmax is the fluorescence intensity of the Ca2+-saturated indicator, F
is the fluorescence during the measurement, and Kd is the dissociation
constant of Fluo-3. Kd of Fluo-3 (503 nM) was determined on the basis
of the calcium calibration buffer kit (Molecular Probes) for intracellular
buffer containing 1 mM ATP, 20 mM creatine phosphate, and 20 U/mL
creatine kinase at 37 °C, pH 7.06.
EXPERIMENTAL SECTION
■
General. All reagents and solvents were of commercial quality and
were used without further purification unless described otherwise.
Triethylamine and morpholine were dried over potassium hydroxide,
distilled, and then stored over potassium hydroxide pellets. ADP-
ribosyl cyclase was purified from the ovotestis of Aplysia californica.67
H2O was of Milli-Q quality. All 1H, 13C, and 31P NMR spectra of final
compounds were collected in D2O on a JEOL Delta machine at 270
MHz (1H) or 109 MHz (31P) or on a Varian Mercury-vx system at 400
6-Chloroadenosine 2′,3′,5′-Triacetate (6). Compound 6 was
synthesized according to a modified Vorbruggen condensation. To a
̈
vigorously stirred solution of 6-chloropurine (500 mg, 3.23 mmol), β-
D-ribofuranose 1,2,3,5-tetraacetate (926 mg, 2.91 mmol), and DBU
(1.3 mL, 8.70 mmol) in dry MeCN (22 mL) was added TMSOTf
(2.12 mL) at 0 °C under an argon atmosphere. The reaction mixture
was heated at 60 °C for 1 h and carefully quenched by addition of a
saturated solution of NaHCO3 (100 mL). The crude compound was
extracted with DCM (3 × 100 mL), and the organic layers were
combined, dried over MgSO4, and evaporated under reduced pressure.
The yellow residue obtained was further purified by flash column
chromatography, eluting with DCM−MeOH, 20:1, to give the desired
MHz (1H) or 100 MHz (13C). All H and 13C NMR assignments are
1
based on gCOSY, gHMBC, gHMQC, and DEPT experiments.
Abbreviations for splitting patterns are as follows: br, broad; s, singlet;
d, doublet; t, triplet; m, multiplet. UV spectra were collected in
aqueous solution on a Perkin-Elmer Lambda EZ 201 or Lambda 3B
spectrophotometer. High resolution time-of-flight mass spectra were
obtained on a Bruker Daltonics micrOTOF mass spectrometer using
electrospray ionization (ESI). HPLC analyses were carried out on a
Waters 2695 Alliance module equipped with a Waters 2996 photo-
diode array detector (210−350 nm). The chromatographic system
consisted of a Hichrom guard column for HPLC and a Phenomenex
Synergi 4 μm MAX-RP 80A column (150 × 4.60 mm), with elution at
1 mL/min with the following ion-pair buffer: 0.17% (m/v) cetrimide
and 45% (v/v) phosphate buffer (pH 6.4) in MeOH. The purities of
all phosphate containing compounds were determined by HPLC, and
they were in all cases over 95%. All other compounds were analyzed
using a Phenomenex Gemini column 5 μm C18 (150 mm × 4.6 mm),
with elution at 1 mL/min with a MeCN/H2O gradient (5−65% over
20 min). Preparative chromatography was performed on a Pharmacia
Biotech Gradifrac system equipped with a peristaltic P-1 pump and a
1
compound as a yellow oil (960 mg, 77%). H (CDCl3, 270 MHz) δ
8.76 (s, 1H, H-8), 8.28 (s, 1H, H-2), 6.21 (d, 1H, J1′,2′ = 5.2 Hz, H-1′),
5.92 (dd, 1H, J2′,3′ = 5.4 Hz and J2′,1′ = 5.2 Hz, H-2′), 5.62 (dd, 1H,
J3′,2′ = 5.4 Hz and J3′,4′ = 4.9 Hz, H-3′), 4.37 (m, 3H, H-4′ and H-5′),
2.14 (s, 3H, CH3), 2.10 (s, 3H, CH3), and 2.07 (s, 3H, CH3).
6-N-Methyladenosine 5′-Acetate. The title compound was
synthesized by adaptation of a literature protocol.71 To a suspension of
6-chloroadenosine triacetate 6 (800 mg, 1.94 mmol) and methylamine
hydrochloride (573 mg, 8.42 mmol) in a mixture of DCM (20 mL)
and ethanol (4.2 mL) was added triethylamine (3.8 mL). The resulting
mixture was stirred at 60 °C overnight. TLC analysis indicated that the
starting material was completely consumed and three more polar spots
were given. The reaction mixture was transferred into a pressure tube
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with triethylamine (4 mL), methylamine hydrochloride (500 mg, 7.35
mmol), and ethanol (4 mL). The mixture was heated at 60 °C for 6 h
and left at room temperature for 2 days. TLC of the mixture indicated
only one major spot at Rf = 0.25 (DCM−MeOH, 10:1). The solvent
was removed and the crude compound was purified by flash column
chromatography, eluting with DCM−MeOH, 30:1, to give a mixture
of the desired compound and the methylamine hydrochloride. Pure
title compound was obtained as a white solid by washing the resulting
mixture with Milli-Q water (420 mg, 67%). 1H (DMSO-d6, 270 MHz)
δ 8.35 (s, 1H, H-8), 8.24 (brs, 1H, H-2), 7.81 (brs, 1H, NH), 5.91 (d,
1H, J1′,2′ = 5.0 Hz, H-1′), 5.61 (m, 1H, 2′-OH), 5.42 (m, 1H, 3′-OH),
4.67 (m, 1H, H-2′), 4.31−4.18 (m, 4H, H-3′, H-4′, H-5′), 2.95 (brs,
3H, CH3N), 1.95 (s, 3H, CH3); HRMS (ES+) calcd for C13H18N5O5
324.1308 (MH)+, found 324.1302.
165 μmol), and MgSO4 (38 mg, 317 μmol) was added a solution of
MnCl2 in formamide (0.2 M, 1.18 mL). The resulting suspension was
stirred at room temperature for 48 h under a nitrogen atmosphere,
and the reaction mixture was quenched by dropwise addition of
MeCN (2 mL). The yellow precipitate was filtered, washed with
acetone, and dissolved in small amount of Milli-Q water. The aqueous
solution was treated with Chelex 100 (sodium form) to remove any
residual manganese and purified by reverse-phase chromatography,
eluting with a gradient of 0.05 M TEAB against MeCN. The title
compound was isolated as a glassy solid in the triethylammonium form
(51 mg, 64%). HPLC, tR = 3.4 min at 254 nm; UV (H2O) λmax 262.5
1
nm (ε/dm3 mol−1 cm−1 14 560); H (D2O, 400 MHz) δ 9.25 (s, 1H,
HN-2), 9.07 (d, 1H, J6,5 = 5.9 Hz, HN-6), 8.71 (d, 1H, J4,5 = 8.2 Hz,
HN-4), 8.26 (s, 1H, H-8), 8.09 (dd, 1H, J5,4 = 8.2 Hz and J5,6 = 5.9 Hz,
HN-5), 7.97 (s, 1H, H-2), 5.99 (d, 1H, J1″,2″ = 5.5 Hz, H-1″), 5.91 (d,
1H, J1′,2′ = 5.9 Hz, H-1′), 4.70 (dd, 1H, J2′,1′ = 5.9 Hz and J2′,3′ = 5.5 Hz,
H-2′), 4.48 (m, 1H, H-4″), 4.44 (m, 2H, H-3″ and H-3′), 4.37 (m, 1H,
H-2″), 4.32−4.16 (m, 5H, H-4′, H-5′, and H-5″), and 2.93 (CH3N);
31P (D2O, 109 MHz) δ −10.73 (brs); HRMS (ES−) calcd for C22H28-
6-N-Methyladenosine (7). A suspension of 6-N-methyladenosine
5′-acetate (400 mg, 1.23 mmol) in a saturated methanolic ammonia
(50 mL) was stirred at room temperature for 3 h, after which the
solvent was removed in vacuo. The residue obtained was purified by
flash column chromatography, eluting with DCM−methanol, 10:1, to
produce the title compound as a white solid (300 mg, 86%); mp 178−
N7O14P2 676.1169 [M − H]−, found 676.1158.
1
180 °C; H (DMSO-d6, 270 MHz, D2O shake) δ 8.31 (s, 1H, H-8),
6-N-Methyladenosine 5′-Diphosphate Ribose (6-N-Methyl
ADPR, 11). To a solution of 10 (20 mg, 24 μmol) in HEPES buffer
(25 mM, pH 7.4, 60 mL) was added Aplysia ADP-ribosyl cyclase
(80 μL). The resulting solution was stirred at room temperature until
RP-HPLC indicated that all the starting material had reacted. The
solution was diluted, and then product was purified by ion-exchange
chromatography, eluting with a gradient of 0−50% 1 M TEAB buffer
against Milli-Q water. The appropriate fractions were collected, evapo-
rated and excess TEAB was removed by coevaporating with MeOH
(3 times) to give the compound as a glassy solid in its triethylammo-
nium form (10 mg, 59%). HPLC, tR = 11.8 min at 254 nm; UV (H2O)
8.20 (brs, 1H, H-2), 5.85 (d, 1H, J1′,2′ = 6.4 Hz, H-1′), 4.57 (dd, 1H,
J2′,1′ = 6.4 Hz and J2′,3 = 5.2 Hz, H-2′), 4.12 (dd, 1H, J3′,2′ = 5.2 Hz and
J3′,4′ = 3.0 Hz, H-3′), 3.97 (app.q, 1H, J4′,5′a = J4′,5′b = J4′,3′ = 3.0 Hz, H-
4′), 3.64 (dd, J5′a,5′b = 12.9 Hz and J5′a,4′ = 3.0 Hz, 1H, H-5′a), 3.55 (dd,
1H, J5′b,5′a = 12.9 Hz and J5′b,4′ = 3.0 Hz, H-5′b), and 2.94 (brs, 3H,
CH3N); HRMS (ES+) calcd for [M + H]+ C11H16N5O4 282.1202
(MH+), found 282.1201.
6-N-Methyladenosine 5′-Monophosphate (8a, 6-N-Methyl
AMP). A suspension of 7 (110 mg, 0.39 mmol, dried in vacuo at
100 °C for 2 h) in triethyl phosphate (1.4 mL), was heated strongly
with a heat gun for 5 min. To the resulting clear solution were added
POCl3 (0.3 mL, 3.20 mmol) and H2O (1 μL) at 0 °C, and the mixture
was stirred for 1 h. The reaction mixture was quenched by addition of
ice (15 mL). The resulting solution was extracted with cold ethyl
acetate (5 × 20 mL). The aqueous layer was neutralized with NaOH
(5 M) and loaded onto a reverse phase column, eluting with a gradient
of 0−30% MeCN against 0.05 M TEAB. Fractions containing the
desired compound were pooled, evaporated and excess TEAB was
coevaporated with MeOH (3×) to give a mixture of title compound 8a
1
λmax 265.1 nm (ε/dm3 mol−1 cm−1 13 780); H (D2O, 270 MHz) δ
8.37 (s, 1H, H-8), 8.10 (s, 1H, H-2), 6.02 (d, 1H, J1′,2′ = 5.9 Hz, H-1′),
5.24 (d, 0.3H, J1″,2″ = 4.0 Hz, H-1″ ), 5.14 (m, 0.7H, H-1″α), 4.70−3.94
β
(m, 10H, H-ribose), and 2.87 (m, 3H, CH3N); 31P (D2O, 109 MHz) δ
−10.6 (brs); HRMS (ES−) calcd for C16H24N5O14P2 572.0795 [M −
H]−, found 572.0792.
2′,3′-O-Isopropylidene-6-(2,4-dinitrophenyl)thioinosine, 15.
To a suspension of 13 (80 mg, 0.236 mmol) in dry MeCN (5 mL)
were added triethylamine (197 μL, 1.416 mmol) and dinitrofluoro-
benzene (36 μL, 0.283 mmol). The resulting solution was stirred
at room temperature for 1 h after the solvent was removed under
reduced pressure and the residue obtained was purified by flash
chromatography on silica gel (EtOAc/hexane, 1:1) to yield the desired
product as a yellow foam (96 mg, 83%). HPLC, tR = 14.6 min at
260 nm; 1H (270 MHz, CDCl3) δ 8.98 (d, 1H, J = 2.5 Hz, Ar-H), 8.65
(s, 1H, H-2), 8.38 (dd, 1H, J = 8.8 and 2.5 Hz, Ar-H), 8.16 (s, 1H,
H-8), 7.97 (d, 1H, J = 8.8 Hz, Ar-H), 5.94 (d, 1H, J1′,2′ = 4.7 Hz, H-1′),
5.21−5.17 (m, 1H, H-2′), 5.12−5.08 (m, 1H, H-3′), 4.55−4.54 (m,
1H, H-4′), 3.98−3.76 (m, 2H, H-5′), 1.64 (s, 3H, CH3), and 1.37 (s,
3H, CH3); HRMS (ES+) calcd for C19H19N6O8S 4910.0980 (MH+),
found 491.0957; Rf = 0.3 (EtOAc/hexane, 6:4).
1
(150 mg, 0.20 mmol, 51%) and the bis-phosphate 8b. H (D2O, 270
MHz) δ 8.17 (s, 1H, H-8), 7.71 (s, 1H, H-2), 5.83 (d, 1H, J1′,2′
=
5.7 Hz, H-1′), 4.57 (m, 1H, H-2′), 4.37 (m, 1H, H-3′), 4.25 (m, 1H,
H-4′), 4.03 (m, 2H, H-5′) and 2.78 (m, 3H, CH3N); 31P (D2O, 109
1
MHz) δ 1.12 (s). Bisphosphate 8b: H (D2O, 270 MHz) δ 8.13 (s,
1H, H-8), 7.81 (s, 1H, H-2), 6.06 (d, 1H, J1′,2′ = 4.5 Hz, H-1′), 5.25 (m,
1H, H-2′), 5.05 (m, 1H, H-3′), 4.75 (m, 1H, H-4′ half overlap with
HOD signal), 4.55 (m, 2H, H-5′), and 2.80 (m, 3H, CH3).
6-N-Methyladenosine 5′-Monophosphate Morpholidate (9).
A suspension of 6-N-methyl-AMP and the bisphosphate as above
8a/8b (0.20 mmol, calculated by 1H NMR integration) in dry DMSO
(0.9 mL) was evaporated with dry DMF (3 × 2 mL). To the residue
were added in sequence triphenylphosphine (280 mg, 1.07 mmol),
morpholine (0.15 mL, 1.72 mmol), and dipyridyl disulfide (235 mg,
1.07 mmol). The resulting yellow solution was stirred at room tem-
perature for 4 h, after which a solution of sodium iodide in acetone
(0.2 M, 15 mL) was added and the resulting precipitate was filtered
and washed with acetone. The crude product was further purified by
ion-exchange chromatography, eluting with a gradient of 0−50% 1 M
TEAB against Milli-Q water. The solvent was evaporated in vacuo and
excess TEAB was coevaporated with MeOH (3 times) to give the title
2′,3′-O-Isopropylidene-5′-O-di-tert-butylphosphoramidite-
6-(2,4-dinitrophenyl)thioinosine, 16. To a solution of 15 (70 mg,
0.138 mmol) in dry DCM (3 mL) were added tetrazole (20 mg,
0.276 mmol) and N,N-diisopropyl-di-tert-butylphosphoramidite (66 μL,
0.208 mmol). The reaction mixture was stirred at room temperature
for 1 h, after which time TLC analysis (hexane/EtOAc, 6:4) indicated
conversion of starting material to a single phosphite. The mixture was
cooled to −78 °C, and mCPBA (47 mg, 0.276 mmol) was added. After
20 min, 10% aqueous Na2SO3 (10 mL) was added and the mixture
was warmed to room temperature. The organic layer was separated
and washed with a saturated solution of NaHCO3 (15 mL) and brine
(15 mL), dried (Na2SO4), filtered, and evaporated under reduced
pressure to leave an oil which was purified by column chromatography
on silica gel (hexane/EtOAc, 1:1) to give the title compound 16 as
compound as its triethylammonium salt (93 mg, 78%). HPLC, tR
=
2.8 min at 254 nm; UV (H2O) λmax 266.4 nm; 1H (D2O, 270 MHz) δ
8.23 (s, 1H, H-8), 7.98 (s, 1H, H-2), 5.93 (d, 1H, J1′,2′ = 4.9 Hz, H-1′),
4.68 (m, 1H, H-2′), 4.42 (m, 1H, H-3′), 4.26 (m, 1H, H-4′), 3.94 (m,
2H, H-5′), 3.44 (m, 4H, 2 × CH2O), 2.91 (m, 3H, CH3N), and 2.80
(m, 4H, 2 × CH2N); 31P (D2O, 109 MHz) δ 8.10 (s); HRMS (ES+)
C15H24N6O7P 431.1444 calcd for MH+, found 431.1441.
1
a yellow oil (85 mg, 90%). HPLC, tR = 12.1 min at 260 nm; H
(270 MHz, CDCl3) δ 8.98 (d, 1H, J = 2.5 Hz, Ar-H), 8.74 (s, 1H,
H-2), 8.35 (d, 1H, J = 2.5 Hz, Ar-H), 8.33 (s, 1H, H-8), 7.92 (d, 1H,
J = 8.8 Hz, Ar-H), 6.23 (d, 1H, J1′,2′ = 2.7 Hz, H-1′), 5.33 (dd, 1H,
6-N-Methyl nicotinamide Adenine Dinucleotide (6-N-Methyl
NAD+, 10). To a mixture of 9 (56 mg, 95 μmol), β-NMN+ (55 mg,
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J2′,3′ = 6.0 Hz and J2′,1′ = 2.7 Hz, H-2′), 5.04 (dd, 1H, J3′,2′ = 6.0 Hz and
J3′,4′ = 2.4 Hz, H-3′), 4.55−4.54 (m, 1H, H-4′), 4.16−4.12 (m, 2H,
HPLC analysis showed total consumption of starting material and
formation of a new large peak at 11 min. The water was evaporated to
a minimum, and the residue was applied on a RP-18 column, eluting
with a gradient of MeCN in 0.05 M TEAB (0−65% over 300 mL).
The product came through with 18% MeCN. The appropriate frac-
tions were collected and evaporated under reduced pressure. The
residue was passed through a small Chelex column previously washed
with Milli-Q water to bring the pH down to 8. The product was
washed off with Milli-Q water. It was then lyophilized to afford the
desired cyclic dinucleotide as its sodium salt (7 μmol, 52%). HPLC,
t
H-5′), 2.03 (s, 3H, CH3), 1.63 (s, 3H, CH3), 1.44 (s, 9H, Bu), and
t
1.40 (s, 9H, Bu); 31P (decoupled, 109 MHz, CDCl3) δ −9.3 (s);
HRMS (ES+) calcd for C27H36N6O11PS 683.1895 (MH+), found
683.1871; Rf = 0.14 (hexane/EtOAc, 6:4).
2′,3′-O-Isopropylidene-5′-O-(di-tert-butylphosphoramidite)-
6-thioinosine. 16 (80 mg, 0.116 mmol) was stirred in a solution of
MeCN (10 mL) containing 10% (v/v) 2-mercaptoethanol (1 mL) and
1% DIPEA (0.1 mL) for 1 h. Water (15 mL) and DCM (30 mL) were
then added. The organic layer was washed several times with water to
remove the excess 2-mercaptoethanol. The organic phase was dried,
filtered, and evaporated in vacuo to leave an oil which was purified by
column chromatography on silica gel (DCM/acetone, 7:3) to give the
1
tR = 10.7 min at 320 nm; H (270 MHz, D2O) δ 9.32 (s, 1H, H-2),
8.29 (s, 1H, H-8), 6.65 (br s, 1H, H-1″), 5.99 (d, 1H, J1′,2′ = 6.3 Hz,
H-1′), 5.20 (dd, 1H, J2′,1′ = 6.3 Hz and J2′,3′ = 5.0 Hz, H-2′), 4.64 (dd,
1H, J3′,2′ = 5.0 Hz and J3′,4′ = 2.2 Hz, H-3′), and 4.59−4.07 (m, 8H,
H-4′, H-5′, H-2″, H-3″, H-4″, and H-5″); 31P (decoupled, 109 MHz,
D2O) δ −9.2 (d, AB system, J = 11.8 Hz), −10.5 (d, AB system, J =
11.8 Hz); HRMS (ES−) calcd for C15H19N4O13P2S 557.0150 (MH−),
found 557.0161; UV (H2O, pH 7.2). λmax 322 nm (ε/dm3 mol−1 cm−1
18 600).
1
title compound as a white waxy solid (45 mg, 75%). H (270 MHz,
CDCl3) δ 8.38 (s, 1H, H-2), 8.36 (s, 1H, H-8), 6.14 (d, 1H, J1′,2′
=
2.2 Hz, H-1′), 5.52 (dd, 1H, J2′,3′ = 6.1 Hz and J2′,1′ = 2.1 Hz, H-2′), 5.04
(dd, 1H, J3′,2′ = 6.1 Hz and J3′,4′ = 2.5 Hz, H-3′), 4.56−4.54 (m, 1H,
H-4′), 4.23−4.17 (m, 2H, H-5′), 1.63 (s, 3H, CH3), 1.44 (s, 18H, 2 ×
tBu), and 1.42 (s, 3H, CH3); 31P (decoupled, 109 MHz, CDCl3) δ
−9.3 (s); HRMS (ES+) calcd for C21H34N4O7PS 517.1880 (MH+),
found 517.1863.
6-Thioinosine 5′-Monophosphate (6-SH IMP, 14). 2′,3′-O-
Isopropylidene-5′-O-(di-tert-butylphosphoramidite)-6-thioinosine (35 mg,
0.067 mmol) was stirred in a 50% aqueous TFA solution (2 mL) at
room temperature for 24 h. The solvent was removed under reduced
pressure and coevaporated several times with MeOH to remove any
residual TFA. The residue was dissolved in water and worked up with
EtOAc (2 × 10 mL). The aqueous layer was evaporated to dryness to
produce the desired monophosphate 14 as a glassy solid (21 mg,
86%). HPLC, tR = 2.9 min at 320 nm; 1H (270 MHz, D2O) δ 8.94 (s,
1H, H-2), 8.23 (s, 1H, H-8), 6.05 (d, 1H, J1′,2′ = 3.9 Hz, H-1′), 4.60
(dd, 1H, J2′,3′ = 4.7 Hz and J2′,1′ = 3.9 Hz, H-2′), 4.41 (app t, 1H, J =
5.0 Hz, H-3′), 4.27 (dd, 1H, J4′,3′ = 5.0 Hz and J4′,5′a = 2.5 Hz, H-4′),
4.17 (ddd, 1H, J5′a,5′b = 11.8 Hz, J5′a,P = 4.4 Hz, and J5″a,4 = 2.5 Hz, H-
ASSOCIATED CONTENT
* Supporting Information
■
S
Full NMR characterization spectra, fluorescence spectrum,
HPLC traces, computational data for target compound 3, and
13C NMR data for all compounds. This material is available free
of charge via the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
*Phone: ++44-1225-386639. Fax: ++44-1225-386114. E-mail:
B.V.L.Potter@bath.ac.uk.
■
ACKNOWLEDGMENTS
■
5′a), and 4.05 (ddd, 1H, J5′a,5′b = 11.8 Hz, J5′a,P = 5.5 Hz, and J5″a,4
=
We thank the Wellcome Trust for Project Grant 084068 (to
B.V.L.P. and A.H.G.) and Program Grant 082837 (to B.V.L.P.).
Work in the Guse lab is supported by the Deutsche
Forschungsgemeinschaft (DFG) and the Deutscher Akade-
mischer Austauschdienst (DAAD).
2.8 Hz, H-5′b); 31P (decoupled, 109 MHz, D2O) δ 0.51 (s); HRMS
(ES+) calcd for C10H14N4O7PS 365.0315 (MH+), found 365.0310.
Nicotinamide 6-Mercaptopurine 5′-Dinucleotide (6-SH
NHD+, 17). 14 (13 mg, 0.036 mmol) was dissolved in dry DMSO
(1 mL) and coevaporated with dry DMF (5 × 3 mL). The residue
was dissolved in DMSO (400 μL), to which were added morpholine
(16 μL, 0.187 mmol), dipyridyl disulfide (19 mg, 0.089 mmol), and
triphenylphosphine (24 mg, 0.089 mmol), at which point the solution
became bright yellow. It was stirred for 1 h at room temperature, after
which HPLC analysis showed completion of the reaction. Precipitation
of the product occurred by dropwise addition of a solution of NaI in
acetone (0.1M, 8 mL). The resulting precipitate was filtered, washed
with acetone, and dried (δP 6.7 ppm and HPLC, tR = 5.9 min at
320 nm). It was then reacted with β-NMN+ (17 mg, 0.050 mmol) and
MgSO4 (11 mg, 0.092 mmol) in a 0.2 M solution of MnCl2 in form-
amide (0.35 mL) at room temperature overnight, after which HPLC
analysis showed completion of the reaction (tR(β‑NMN) = 2.1 min and
tR(6‑SH‑NHD) = 7 min). Precipitation occurred by dropwise addition of
MeCN. The precipitate was filtered, dissolved in Milli-Q, and applied
to a reverse-phase column, eluting with a gradient of MeCN in 0.05 M
TEAB. Further treatment with Chelex 100 to remove any para-
magnetic particles afforded the desired dinucleotide 17 as a glassy solid
(10 mg, 15.2 μmol, 31% from 6-thio IMP). HPLC, tR = 6.9 min at
ABBREVIATIONS USED
■
cADPR, cyclic adenosine 5′-diphosphoribose; cIDPR, cyclic
inosine 5′-diphosphoribose; cADPcR, cyclic adenosine 5′-
diphosphocarbocyclic ribose; cIDPRE, N1‑[(2″-O-phosphoryl-
ethoxy)methyl]-5′-O-phosphorylinosine 2″,5′-cyclic pyrophos-
phate; NAD+, nicotinamide adenosine 5′-dinucleotide; 6-NMe
NAD+, 6-N-Methyl nicotinamide adenosine 5′-dinucleotide;
NGD+, nicotinamide guanosine 5′-dinucleotide; NHD+, nic-
otinamide hypoxanthine 5′-dinucleotide; 6-thio NHD+, nicotina-
mide 6-mercaptopurine 5′-dinucleotide; ADPRC, ADP-ribosyl
cyclase; TEP, triethylphosphate; β-NMN+, β-nicotinamide 5′-
mononucleotide; CDI, carbonyldiimidazole; TEAB, triethylammo-
nium bicarbonate; IMP, inosine 5′-monophosphate; ribo-2′-F-
NAD+, ribosyl-2′-fluoro-2′-deoxy-nicotinamide adenine dinucleotide
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320 nm; 1H (270 MHz, D2O) δ 9.17 (s, 1H, HN2), 8.96 (d, 1H, J6,5
=
■
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