Sugar conformational effects on the photochemistry of thymidylyl(3′-5′)thymidine

Article abstract of DOI:10.1021/jo030086p

The synthesis and conformational analysis of 2′-O,5-dimethyluridylyl(3′-5′)-2′-O,5-dimethyluridine (1a), the analogue of thymidylyl(3′-5′)thymidine (TpT; 1b) in which a methoxy group replaces each 2′-α-hydrogen atom, are described. In comparison with TpT, such modification increases the population of the C3′-endo conformer of the sugar ring puckering at the 5′- and 3′-ends from 30 to 75% and from 37 to 66%, respectively. Photolyses of 1a and TpT at 254 nm are qualitatively comparable (the cis-syn cyclobutane pyrimidine dimer and the (6-4) photoproduct are formed), although it is significantly faster in the case of 1a. These results are explained by the increased propensity of the modified dinucleotide to adopt a base-stacked conformation geometry reminiscent of that for TpT.

Full text of DOI:10.1021/jo030086p

Su ga r Con for m a tion a l Effects on th e P h otoch em istr y of
Th ym id ylyl(3′-5′)th ym id in e
Tomasz Ostrowski,† J ean-Claude Maurizot,‡ Marie-The´re`se Adeline,†
J ean-Louis Fourrey,† and Pascale Clivio*,†
Institut de Chimie des Substances Naturelles, CNRS, 1 Avenue de la Terrasse,
91190 Gif sur Yvette, France, and Centre de Biophysique Mole´culaire, CNRS, Rue Charles Sadron,
45071 Orle´ans Cedex 2, France
pascale.clivio@icsn.cnrs-gif.fr
Received March 10, 2003
The synthesis and conformational analysis of 2′-O,5-dimethyluridylyl(3′-5′)-2′-O,5-dimethyluridine
(1a ), the analogue of thymidylyl(3′-5′)thymidine (TpT; 1b) in which a methoxy group replaces each
2′-R-hydrogen atom, are described. In comparison with TpT, such modification increases the
population of the C3′-endo conformer of the sugar ring puckering at the 5′- and 3′-ends from 30 to
75% and from 37 to 66%, respectively. Photolyses of 1a and TpT at 254 nm are qualitatively
comparable (the cis-syn cyclobutane pyrimidine dimer and the (6-4) photoproduct are formed),
although it is significantly faster in the case of 1a . These results are explained by the increased
propensity of the modified dinucleotide to adopt a base-stacked conformation geometry reminiscent
of that for TpT.
In tr od u ction
which take place within the DNA double helix. Conse-
quently, we have recently started a program consist-
ing of systematically comparing the photochemistry of
thymine-thymine dinucleoside monophosphate ana-
logues endowed with particular conformational properties
with that of the parent deoxydinucleotide 1b (TpT), a
good B-DNA model. Such studies have already allowed
us to show that, upon UV irradiation, peptide nucleic
acids dimers sensus stricto do not give rise to (6-4) PP
but lead exclusively to CPD.4 Conversely, in the case of
“amide-3” dimer, both (6-4) PP and CPD are formed.5
In continuing our work, we were interested in probing
the effect of the major conformational parameter that
distinguishes B-DNA from A-DNA, the sugar pucker, on
the photochemistry of TpT. Interestingly, in the oligo-
nucleotide series, the 2′-R-methoxy modification of 2′-
deoxyribose has been shown to induce a significant shift
of the conformational equilibrium of the sugar ring from
the C2′-endo toward the C3′-endo conformation, resulting
in a mimic A-conformation.6 This trend is likely to be
operative at the dinucleotide level for pyrimidines, since
the ratio of C3′-endo conformers of ribodinucleotides7 vs
Exposure of DNA to solar ultraviolet light results in
the formation of dipyrimidine photoproducts, the most
common being cyclobutane pyrimidine dimers (CPD) and
pyrimidine (6-4) pyrimidone photoproducts ((6-4)PP) with
their Dewar valence isomers (Figure 1).1 A good deal of
evidence points to these photoproducts being at the origin
of the molecular mechanism leading to skin carcinogen-
esis through their combined mutagenic and immuno-
suppressive properties.2 However, the respective contri-
bution of each class of photoproduct in skin cancer
development is not yet fully known, nor are the factors
that influence their respective formation. Indeed, under-
standing these factors is critical to further investigate
photocarcinogenesis. At present, parameters such as
photophysical properties of the targeted nucleobases,
wavelength, flanking sequence context, and conforma-
tional features, including secondary structure and DNA-
protein interaction, have been identified to play an
important role in affecting photoproduct formation.3
However, the rules that govern the relationships be-
tween DNA conformational parameters and preferential
photoproduct formation are unclear.
(3) (a) Matsunaga, T.; Hieda, K.; Nikaido, O. Photochem. Photobiol.
1991, 54, 403-410. (b) Sage, E. Photochem. Photobiol. 1993, 57, 163-
174. (c) Pfeifer, G. P. Photochem. Photobiol. 1997, 65, 270-283. (d)
Perdiz, D.; Grof, P.; Mezzina, M.; Nikaido, O.; Moustacchi, E.; Sage,
E. J . Biol. Chem. 2000, 275, 26732-26742. (e) Liu, X.; Conconi, A.;
Smerdon, M. J . Biochemistry 1997, 36, 13710-13717. (f) Aboussekhra,
A.; Thoma, F. EMBO J . 1999, 18, 433-443. (g) Ghosh, R.; Paniker,
L.; Mitchell, D. L. Photochem. Photobiol. 2001, 73, 1-5. (h) Wang, Y.;
Gross, M. L.; Taylor, J .-S. Biochemistry 2001, 40, 11785-11793.
(4) Clivio, P.; Guillaume, D. Tetrahedron Lett. 1998, 39, 6881-6884.
(5) Thomas, M.; Clivio, P.; Guillaume, D.; Fourrey, J .-L. Nucleosides
Nucleotides Nucleic Acids 2001, 20, 927-929.
We consider the elucidation of the factors which drive
the photochemistry occurring at the dinucleotide level to
be a prerequisite step in understanding the phenomena
* To whom correspondence should be addressed. Fax: 33 1 69 07
72 47.
† Institut de Chimie des Substances Naturelles.
‡ Centre de Biophysique Mole´culaire.
(1) (a) Cadet, J .; Vigny, P. In Bioorganic Photochemistry: The
Photochemistry of Nucleic Acids; Morrison, H., Ed.; Wiley: New York,
1990; Vol. 1, pp 1-272.
(2) (a) Cleaver, J . E.; Crowley, E. Front. Biosci. 2002, 7, d1024-
1043. (b) Ullrich, S. E. Front. Biosci. 2002, 7, d684-703.
(6) Lesnik, E. A.; Freier, S. M. Biochemistry 1998, 37, 6991-6997
and references therein.
(7) Lee, C.-H.; Ezra, F. S.; Kondo, N. S.; Sarma, R. H.; Danyluk, S.
S. Biochemistry 1976, 15, 3627-3638.
10.1021/jo030086p CCC: $25.00 © 2003 American Chemical Society
Published on Web 08/01/2003
6502
J . Org. Chem. 2003, 68, 6502-6510

Photochemistry of Thymidyl(3′-5′)thymidine
F IGURE 1. Major DNA photoproducts formed at adjacent thymine sites.
SCHEME 1. Syn th esis of 1a a
2′-deoxyribodinucleotides8 is increased and since 2′-O-
methylation of ribodinucleotides induces an additional
stabilization of the C3′-endo form.9 Therefore, we propose
to probe, at the single-stranded dinucleotide level, the
effect of the C3′-endo conformational preference induced
by 2′-methoxylation on the photochemistry of thymine.
We herein report the synthesis of 2′-O,5-dimethyl-
uridylyl(3′-5′)-2′-O,5-dimethyluridine (or 2′-R-methoxy-
thymidylyl(3′-5′)-2′-R-methoxythymidine (TmopTmo)) 1a ,
together with its sugar conformational analysis and
photochemical behavior. In addition, to rationalize the
observed photochemical results, the stacking properties
of 1a and 1b are compared.
Resu lts
Syn th esis of 2′-O,5-Dim eth ylu r id yl (3′-5′)-2′-O,5-
d im eth ylu r id in e (1a ). Whereas the synthesis of oligo-
nucleotides containing 2′-O,5-dimethyluridine (2a ) has
been reported,10 as far as we know, dinucleotide 1a has
never been prepared. We have obtained this compound
as outlined in Scheme 1.
a
Legend: (a) ref 17; (b) (i) DmtCl, pyridine, (ii) Mg(OCH3)2,
DMF, 68% yield from 4; (c) refs 11 and 12; (d) (i) Ac2O, pyridine,
(ii) TFA/CH2Cl2 3%, 71% yield from 5; (e) (i) tetrazole, CH3CN,
(ii) I2, THF/H2O/2,6-lutidine; (f) (i) NH4OH/MeOH, (ii) 80% aque-
ous AcOH, 26% yield from 7.
Two general strategies have been developed to syn-
thesize 2′-O,5-dimethyluridine nucleosides. One route
uses a 2-O-methyl ribosylation procedure of thymine,11
whereas the other one makes use of 5-methyluridine
derivatives. In the latter case, the 2′-O-methyl group is
obtained either by methylation of the 2′-hydroxyl group10,12
or, to circumvent multiple blocking/deblocking steps, by
a 3′-assisted regiospecific opening of a 2,2′-anhydro
intermediate.13 We decided to follow such an approach,
starting from 5-methyluridine and employing Mg(OMe)2
as a source of methoxide for the 3′-assisted regiospecific
opening of its corresponding 2,2′-anhydro intermediate
as described for the synthesis of 2′-O-methyluridine
(Scheme 1).14 Thus, 5-methyluridine (3)15 was treated
with diphenyl carbonate under basic catalysis conditions
to give the anhydro derivative 416 according to the
procedure described for the synthesis of 2,2′-cyclo-
uridine.17 Then, compound 4 was dimethoxytritylated and
the 2,2′-anhydro linkage was regio- and stereoselectively
(8) Cheng, D. M.; Sarma, R. H. J . Am. Chem. Soc. 1977, 99, 7333-
7348.
(9) (a) Cheng, D. M.; Sarma, R. H. Biopolymers 1977, 16, 1687-
1711. (b) Kawai, G.; Yamamoto, Y.; Kamimura, T.; Masegi, T.; Sekine,
M.; Hata, T.; Iimori, T.; Watanabe, T.; Miyazawa, T.; Yokoyama, S.
Biochemistry 1992, 31, 1040-1046.
(13) Ross, B. S.; Springer, R. H.; Tortorici, Z.; Dimock, S. Nucleosides
Nucleotides 1997, 16, 1641-1643.
(10) (a) Ohtsuka, E.; Inoue, H. J apanese Patent 02 264 792, 1990;
Chem. Abstr. 1991, 114, 164721j. (b) Martin, P. Helv. Chim. Acta 1995,
78, 486-504.
(14) McGee, D. P. C.; Zhai, Y. Nucleosides Nucleotides 1996, 15,
1797-1803.
(11) Chanteloup, L.; Thuong, N. T. Tetrahedron Lett. 1994, 35, 877-
(15) J ones, S. S.; Reese, C. B.; Ubasawa, A. Synthesis 1982, 259-
260.
880.
(12) Lewis, A. F.; Revankar, G. R.; Hogan, M. E. J . Heterocycl. Chem.
1993, 30, 1309-1315.
(16) Fraser, A.; Wheeler, P.; Dan Cook, P.; Sanghvi, Y. S. J .
Heterocycl. Chem. 1993, 30, 1277-1287.
J . Org. Chem, Vol. 68, No. 17, 2003 6503

Ostrowski et al.
TABLE 1. J 1′2′ a n d J 3′4′ Cou p lin g Con sta n ts (Hz)a a n d
tively). Thus, in comparison to TpT (1b) (30% of C3′-endo
for Tp- and 37% of C3′-endo for -pT),8 2′-R-methoxylation
increases, as expected, significantly the population of the
C3′-endo pucker of each sugar, from 30 to 75% and from
37 to 66% for the Tmop- and -pTmo residues, respectively.
The influence of bis-methoxylation on the sugar pucker
of a dinucleoside monophosphate was unknown. In fact,
only the effect of 2′-O-methylation of the 5′-unit of
pyrimidine ribodinucleotides on puckering had been
studied.9 Considering that the conformation of a sugar
is not critically influenced by the nature of the adjacent
sugar,20 the effect of the 2′-methoxylation of TpT at the
5′-end (∆% ) 45%) is in accordance with the observed
stabilization of the C3′-endo form when comparing the
5′-sugar puckers of dCpdC8 and CmopC9a (∆% ) 47%).
In contrast, the effect of C2′-methoxylation on the C3′-
endo conformational equilibrium is less pronounced at
the 3′-end sugar of TpT (∆% ) 29%). This trend clearly
demonstrates the effect of the phosphate group in sta-
bilizing the C3′-endo conformation of the 5′-end sugar.
Previously observed at the monomer level, this behavior
is due to the repulsive steric effect of the phosphate group
in the C2′-endo form and the methoxy group.9b Otherwise,
the C3′-endo conformational preference is due to the
electronegativity of the 2′-substituent21 (gauche stereo-
electronic effect of the O4′-C1′-C2′-O2 fragment)22 and
to repulsive steric effects between the 2′-methoxy group
and the 2-carbonyl group of the pyrimidine base.9b The
combination of these factors is undoubtedly responsible
for the observed predominance of the C3′-endo sugar
puckering of 1a .
P op u la tion Distr ibu tion (%) of Con for m er s of 1a
Tmop-
-pTmo
J 1′2′
2.4
3.2
J 3′4′
7.4
6.2
C2′-endob
C3′-endo
25
34
78,c 75b
65,c 66b
a
b
Coupling constants are accurate to (0.1 Hz. Calculated from
1′2′/(J 1′2′ + J 3′4′). c Calculated from J 3′4′/(J 1′2′ + J 3′4′).
J
opened at the sugar 2′-position by magnesium methoxide,
affording 512 in 68% yield (from 4, two steps). Phos-
phitylation of 5 with 2-cyanoethyl-N,N-diisopropyl-
chlorophosphoramidite in the presence of Hu¨nig’s base
led to phosphoramidite 611,12 in 80% yield. Acetylation of
5 (Ac2O, pyridine) followed by detritylation (TFA/CH2Cl2
3%) gave 3′-O-acetyl-2′-O,5-dimethyluridine 7 in 71%
yield. Finally, phosphoramidite 6 was condensed with 7
in the presence of tetrazole in acetonitrile, and subse-
quent oxidation with iodine in THF/H2O/2,6-lutidine
afforded the corresponding phosphotriester 8. Dimer 1a
was obtained after treatment of 8 with aqueous am-
monia-methanol and then 80% aqueous acetic acid. The
overall yield of the condensation/oxidation/deprotection
steps was 26% (based on 7, four steps).
NMR -Ba sed Con for m a t ion of 2′-O,5-Dim et h yl-
u r id yl(3′-5′)-2′-O,5-d im eth ylu r id in e (1a ). The 2′-O-
methylribose protons of 1a were assigned using the 2D
COSY technique. Signals of the 3′- and 5′-end sugar
protons were differentiated from the multiplicity of the
H3′ signal of the 5′-end sugar and confirmed by a 31P
decoupling experiment.
P h otoch em ica l Beh a vior of 2′-O,5-Dim eth ylu r id yl
(3′-5′)-2′-O,5-d im eth ylu r id in e (1a ). The photochemical
behavior of 1a was studied in aqueous solution at 254
1
nm. Examination of the H NMR spectrum of the crude
irradiation mixture of 1a revealed the formation of two
photoproducts (9a and 10a ) whose respective yield varied
with the photolysis time (Scheme 2).
Signals of the base portion of the photoproducts could
be easily observed. Photoproduct 9a displayed two sin-
glets (one proton each) at δ 7.92 and 5.04 and two singlets
(three protons each) at δ 2.27 and 1.75. Comparison of
these signal chemical shifts with those of key protons of
9b, the (6-4) PP of TpT (H6 -pT, δ 8.00; H6 Tp-, δ 5.10;
CH3 -pT, δ 2.32; CH3 Tp-, δ 1.76)23 suggested that 9a was
the 2′-dimethoxy (6-4) analogue of TpT. In addition,
HPLC analysis using a photodiode detector showed, for
9a , a UV chromophore of the (6-4) type (λmax 329 nm).
Compound 10a displayed a UV spectrum reminiscent of
the cyclobutane class (low absorption above 254 nm). This
was confirmed by the presence of two shielded methyl
signals at δ 1.49 and 1.44 and the lack of two vinylic H6
protons compared with 1a . Unfortunately, comparison of
the methyl or the H1′ signal chemical shifts of 10a with
those of the cis-syn (10b) and trans-syn I CPD of TpT24
In aqueous solution, the sugar pucker of nucleosides
and nucleotides exists as an equilibrium mixture of C2′-
endo and C3′-endo conformers, whose respective popula-
1
tions can be determined by H NMR using vicinal spin-
coupling constants of the sugar protons.18 The population
of the C3′-endo conformers can be estimated using the
sum of J 1′2′ + J 3′4′ and the observed J 1′2′ and J 3′4′ values
(Table 1).18,19 We used a value of J 1′2′ + J 3′4′ ) 9.5 Hz
corresponding to the ribodinucleotide series to compute
the population of the C3′-endo conformer of 1a .7
The maximum observed deviation in the values com-
puted separately from J 1′2′ and J 3′4′ was 3%. We consid-
ered only the J 1′2′-derived value, since its determination
was more accurate than that of J 3′4′ due to complex
multiplicity. Calculations revealed that both the 5′- and
3′-terminal 2′-O-methylribose units of 1a preferentially
adopt a C3′-endo conformation (75% and 66%, respec-
(20) Peeling, J .; Hruska, F. E.; Loewen, P. C. Can. J . Chem. 1978,
56, 522-529.
(21) Guschlbauer, W.; J ankowski, K. Nucleic Acids Res. 1980, 8,
1421-1433.
(17) McGee, D. P. C.; Vargeese, C.; Zhai, Y.; Kirschenheuter, G. P.;
Settle, A.; Siedem, C. R.; Pieken, W. A. Nucleosides Nucleotides 1995,
(22) Plavec, J .; Tong, W.; Chattopadhyaya, J . J . Am. Chem. Soc.
1993, 115, 9734-9746.
14, 1329-1339.
(18) Altona, C.; Sundaralingam, M. J . Am. Chem. Soc. 1973, 95,
(23) Rycyna, R. E.; Alderfer, J . L. Nucleic Acids Res. 1985, 13, 5949-
2333-2344.
5963.
(19) Davies, D. B.; Danyluk, S. S. Biochemistry 1974, 13, 4417-
(24) Koning, T. M. G.; van Soest, J . J . G.; Kaptein, R. Eur. J .
Biochem. 1991, 195, 29-40.
4434.
6504 J . Org. Chem., Vol. 68, No. 17, 2003

Photochemistry of Thymidyl(3′-5′)thymidine
SCHEME 2. P h otop r od u cts F or m ed u p on 254 n m P h otolysis of 1a a n d 1b
could not ascertain 10a stereochemistry at the C5 and
C6 positions. This ambiguity was solved by considering
the stereochemical pathway leading to photoproducts.
Indeed, photolysis of 1b has been shown to lead to two
major adducts, the (6-4) adduct 9b and the cis-syn cyclo-
butane isomer 10b,23,25 both resulting from a photochemi-
cal process between two bases in an anti glycosidic bond
conformation. Since the (6-4) adduct 9a results from a
photochemical process between two bases in an anti
glycosidic bond conformation, it is reasonable to propose
that 10a also results from a similar stereochemical
pathway and consequently has the same stereochemistry
as the cis-syn cyclobutane photoproduct 10b. Prolonged
irradiation of 1a led exclusively to 9a , an observation
consistent with the known photoreversal of the cis-syn
CPD upon UV C exposure that led to accumulation of
the (6-4) photoproduct.25 Hence, this photochemical study
demonstrated that increasing the ratio of the C3′-endo
pucker from 30 to 75% and from 37 to 66% for the 5′-
and 3′-residues, respectively, does not qualitatively modify
the photochemical behavior of the dimer, since (6-4) and
cyclobutane photoproducts 9a and 10a were formed.
To compare more closely the photochemical behavior
of 1a and 1b, we photolyzed a solution prepared by
mixing volume/volume solutions of these compounds
having the same optical density and monitored the
reaction by HPLC.26 After 1 h of irradiation, ca. 58% of
1b was consumed and the peak area ratio of 1b/1a was
1.5. Considering that both CPDs have a similar photo-
reversal rate, which is quite reasonable for the photo-
splitting reaction involving the same cyclobutane, this
means that the photolysis of 1a is 33% more rapid than
that of 1b after 1 h. Assuming similar molar extinction
coefficients for the corresponding adducts in the 2′-
deoxyribose and 2′-O-methylribose series, UV detection
could be used for determining the formation efficiency
of photoproducts in each series. After 1 h of UV exposure,
the yield of CPD in the 2′-O-methylribose series was 1.5
times higher than that in the TpT series and the (6-4)
PP formation was found to be 1.3 times more efficient in
the 2′-O-methylribose series. To determine if the forma-
tion of one class of adduct was favored within a series,
we then compared the (6-4)/CPD ratio in each class after
1 h. The small difference obtained (8%) indicated that
none of the (6-4) or CPD photochemical pathways were
significantly favored.
These photochemical data indicate that 2′-R-methox-
ylation of 1b results in an enhancement of photoproduct
formation rate. Most probably, the C3′-endo conforma-
tional preference derived from 2′-R-methoxylation induces
other conformational changes to be responsible for the
observed photolysis difference. This may be an increased
population of stacked species, since 2′-O-methylation of
ribodinucleotides can enhance intramolecular base stack-
ing.27 Such stacked species should have a higher propen-
sity to undergo photoproduct formation. To investigate
the hypothesis that 1a exhibits an increased stacking
ability, in comparison to 1b, we performed circular
dichroism (CD) studies.
Cir cu la r Dich r oism Stu d ies. CD spectra of 1b and
1a were obtained in the 225-330 nm region over the
temperature range 20-80 °C (Figures 2 and 3, respec-
tively). The CD spectrum of TpT (1b) was consistent with
previously published results.28 At 20 °C, the CD ampli-
tude of 1a was found to be much larger than that of 1b
(a factor of 3 at about 280 nm).
It is generally accepted that the CD spectra of dinucleo-
tides result from two contributions: one from each mono-
nucleoside and a second due to the asymmetric interac-
tion between the two bases. To examine the possible
monomer contribution in the observed magnitude en-
hancement of the CD of 1a with respect to 1b, we
recorded the CD spectrum of the respective mononucleo-
side constituents (2′-O,5-dimethyluridine 2a and thymi-
dine 2b) (Figures 2 and 3). The intensity of the positive
band of 2a was found to be larger than that of 2b.
However, if methoxy substitution per se contributes, in
part, to the CD difference between 1a and 1b, it is not
totally responsible for this difference and other factors,
such as the geometry of interaction between the two
chromophores and/or the fraction of stacked conforma-
tion, must be involved (see below).
For compounds 1a and 1b, comparison of the CD
spectrum of the dimer with that of its corresponding
mononucleoside (Figures 2 and 3) revealed large differ-
ences, as already reported for 1b and 2b.28 The differ-
ences between monomer and dimer spectra are clear
evidence of a nonrandom conformation of bases in the
dimer and arise from the presence of base-stacked
conformations.28 Assuming a two-state model, where a
stacked and an unstacked form of the dimer are in
equilibrium, it has been proposed that the CD difference
(25) J ohns, H. E.; Pearson, M. L.; LeBlanc, J . C.; Helleiner, C. W.
J . Mol. Biol. 1964, 9, 503-524.
(27) Drake, A. F.; Mason, S. F.; Trim, A. R. J . Mol. Biol. 1974, 86,
(26) The concentration of 1b was 0.19 mM (ꢀ267 ) 1.85 × 104).25 An
independent analysis of the separate photolysis of 1a and 1b resolved
without ambiguity the origin of each photoproduct.
727-739.
(28) Cantor, C. R.; Warshaw, M. M.; Shapiro, H. Biopolymers 1970,
9, 1059-1077.
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Ostrowski et al.
F IGURE 2. Influence of the temperature on the CD spectrum of TpT 1b in a buffer containing 0.1 M Na phosphate, 1 M NaCl,
pH 7.0. The CD intensity decreases as the temperature was increased from 20 to 80 °C by 10 °C increment. The light blue line
corresponds to the monomer 2b. CD data are expressed per residue.
F IGURE 3. Influence of the temperature on the CD spectrum of TmopTmo (1a ) in the same buffer as in Figure 2. The CD
intensity decreases as the temperature was increased from 20 to 80 °C by 10 °C increments. The light blue line corresponds to
the monomer 2a . CD data are expressed per residue.
spectra between the dimer and its constituent monomers
depends only on the geometry of interaction of the two
base chromophores and on the fraction of the stacked
form.29 If two dimers with the same chromophore have
the same stacked geometry, the two experimental dif-
ference curves should have the same shape and differ
only by a constant positive multiplier. The difference
curve of 1a can be roughly considered as a simple
multiple of that of 1b (ca. 3) (Figure 4).
and 1b must be very similar, if not identical. In addition,
at 20 °C, the fraction of the stacked form for 1a is
approximately 3 times higher than that of 1b.
Furthermore, both dimers unstack with increasing
temperature (Figures 2 and 3). The effects of temperature
upon the amplitude of the CD difference spectra of 1a
and 1b were recorded in the melting curves (Figure 5).
At 80 °C, the melting curves converged toward low CD
amplitude limits corresponding to the unstacked confor-
mation of the dimers without attaining them. These
limits are known to be difficult to reach.27 The low-
temperature upper CD amplitude limit representing the
fully stacked conformation of the dimer was not observed
Then, if the two-state model of stacking applies to 1a
and 1b, the low-temperature stacked geometries of 1a
(29) Warshaw, M. M.; Cantor, C. R. Biopolymers 1970, 9, 1079-
1103.
6506 J . Org. Chem., Vol. 68, No. 17, 2003

Photochemistry of Thymidyl(3′-5′)thymidine
F IGURE 4. CD difference spectra between 1a and its corresponding monomer 2a and between 1b and its corresponding monomer
2b multiplied by a factor of 3 at 20 °C. Other conditions are those of Figure 2.
F IGURE 5. CD melting curves obtained by following the CD difference intensity between the dimer and the monomer at 279 nm
for 1a and 278 nm for 1b.
for 1a or 1b, thus precluding the determination of the
extent of stacking.30 However, if at 80 °C the dimers have
almost reached a completely unstacked conformation,
this shows that if 1a is more stacked than 1b, it does
not reach the 50% stacked population at 20 °C since the
sigmoidal part of the curve is not reached.
These CD studies indicated that, at room temperature,
the fraction of stacked species is higher in 1a than in
1b.
accompanied by a stacking enhancement of thymines. 2′-
O-methylation has been shown to enhance base stacking
of dipyrimidine ribodinucleotides.27,31 Similarly, 2′-
hydroxylation was found to stabilize the stacked con-
formers of dipyrimidine dinucleotides.31 However, no
explanation was given to rationalize the relationship
between 2′-O-methylation or hydroxylation upon stacking
preferences. In fact, this could be a consequence of the
C3′-endo sugar conformational preference, which is in-
creased in the order 2′-deoxy8 < ribo7 < 2′-O-methylribo,9
and that is the main difference between 1a and 1b.
Indeed, relationships between intramolecular base
stacking within dinucleotides and the sugar pucker have
been evidenced. For example, intramolecular base stack-
Discu ssion
In this study, we have observed, at the single-stranded
dinucleotide level, that 2′-R-methoxylation of 1b was
(30) Brahms, J .; Maurizot, J . C.; Michelson, A. M. J . Mol. Biol. 1967,
25, 481-495.
(31) Rabczenko, A.; Shugar, D. Acta Biochim. Pol. 1971, 18, 387-
402.
J . Org. Chem, Vol. 68, No. 17, 2003 6507

Ostrowski et al.
ing induced by low temperature is known to favor the
C3′-endo conformation in ribodinucleoside phosphates.7
A similar conclusion was drawn from examination of the
stacking/unstacking process of dinucleotides by molecular
modeling.32 However, to our knowledge, no precise mech-
anism has ever been truly proposed to account for these
observations, and the intimate correlation process that
links the C3′-endo conformation and the intramolecular
base stacking needs to be assessed. The torsion angles
along the sugar phosphate backbone are interdepend-
ent:33 for instance, the endocyclic torsion angle ν3 of the
sugar about the C4′-C3′ bond and the backbone torsion
angle δ are directly correlated, since they describe
torsions about the same C4′-C3′ bond. Moreover, δ is
correlated with several other torsion angles, one of which
is ú (O3′-P bond).34 Interestingly, molecular modeling
studies have proposed that ú was the main angle involved
at the onset of the stacking/destacking process of dinucleo-
tides,32,35,36 which is consistent with NMR studies.8 In
addition, calculations have predicted that the C2′-endo/
C3′-endo sugar pucker interconversion is associated with
the concomitant t/g-conformation interconversion of ú
corresponding to the unstacked/stacked state.36 Finally,
when the C3′-endo conformation is due to the presence
of a 2′-methoxy group, the latter is so oriented (favorable
interaction between the methoxy group and the adjacent
(n + 1) sugar residue) to stabilize the g-conformation of
ú37 and, therefore, the population of the stacked con-
former. Thus, we propose that promotion of the C3′-endo
pucker population in 1a is likely to be correlated with a
fractional change in the ú torsion angle that results in
enhancement of the stacked species compared to 1b.
Another question that needs to be addressed concerns
the real involvement of the ground-state stacked confor-
mational preferences of 1a for the observed faster pho-
tolysis rate compared to 1b. To be the case, the stacking/
destacking process has to be sufficiently slow compared
to the photochemical reaction. According to the Franck-
Condon principle, the excitation of a molecule is ex-
tremely rapid (10-15 s) compared to nuclear motion.38 In
addition, formation of CPD and (6-4) PP from TpT
involves singlet states of the excited thymine, whose
lifetime is on the order of 10-12 s.39 Thus, excitation of
an unstacked structure would lead to deactivation before
conformational motion can reorient the bases so that the
photochemical reaction occurs. Therefore, CPD and (6-
4) PP formation is highly likely to derive from the
excitation of stacked complexes39 and the ground-state
stacked conformational preferences are indeed respon-
sible for the observed photolysis rate differences. In
accordance with our conclusions, it has been demon-
strated, through variable-temperature experiments or
with solvents that disrupt base stacking, that in single-
stranded di- and oligonucleotides, base stacking favors
photoproduct formation.40
Finally, the proposed similar stacked geometries of 1a
and 1b determined from CD studies are also supported
by a molecular modeling analysis (data not shown)41 and
explain the qualitative similarities of their photochem-
istry.
Therefore, in comparison to 1b, the C3′-endo sugar
preference of 1a imposed by 2′-methoxylation results,
through the ú torsion angle, in an enhancement of the
population of stacked species with retention of similar
geometry that in turn increases the photolysis efficiency.
Con clu sion
In this work, we have synthesized 1a , the dinucleotide
analogue of 1b containing two 2′-R-methoxy groups, and
shown, as anticipated, that the two modified sugar
substitutions induce a strong enhanced conformational
bias toward the C3′-endo pucker. As a consequence, this
conformational change increases the population of the
stacked species of 1a , which in turn significantly acceler-
ates the photochemical reactivity. Exhibition of an overall
similar geometry of the stacked conformers of 1a and 1b
explains the qualitative similarity in photoproduct for-
mation.
Interestingly, under our photolysis conditions, 1a ,
which exhibits through its C3′-endo sugar pucker a
conformation reminiscent of A-DNA, did not give rise to
the spore photoproduct (SP), whose formation occurs
when DNA adopts an A conformation. So far, in the
context of polymer by opposition to the base or nucleoside
series, SP has only been obtained by UV exposure of dry
or frozen-state DNA, whose conformation most resembles
that of A-DNA or when this required A conformation is
induced by a bacterial spore protein association.42,43 It is
noteworthy that irradiation of 1b under conditions known
to allow the formation of SP in DNA never led to this
type of photoproduct.44 This could indicate that a specific
structural context of adjacent thymines, which is allowed
in DNA and not at the dinucleotide level, might be
necessary for SP formation. On the other hand, the
absence of SP may simply be ascribed to the presence of
water molecules, since it has also been proposed that the
degree of hydration was the critical factor for SP forma-
tion instead of the A conformation.45
(40) (a) Hosszu, J . L.; Rahn, R. O. Biochem. Biophys. Res. Commun.
1967, 29, 327-330. (b) Wacker, A.; Lodemann, E. Angew. Chem. 1965,
77, 133. (c) Tramer, Z.; Wierzchowski, K. L.; Shugar, D. Acta Biochim.
Pol. 1969, 16, 83-107.
(41) Santini, G. P. H.; Cognet, J . A. H.; Clivio, P. Unpublished
results.
(42) Patrick, M. H.; Rahn, R. O. In Photochemistry and Photobiology
of Nucleic Acids: Photochemistry of DNA and Polynucleotides: Photo-
products; Wang, S. Y., Ed.; Academic Press: New York, 1976; Vol. 2,
pp 35-95.
(43) (a) Mohr, S. C.; Sokolov, N. V. H. A.; He, C.; Setlow, P. Proc.
Natl. Acad. Sci. U.S.A. 1991, 88, 77-81. (b) Nicholson, W. L.; Setlow,
B.; Setlow, P. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 8288-8292.
(44) Douki, T.; Court, M.; Cadet, J . J . Photochem. Photobiol. B: Biol.
2000, 54, 145-154.
(45) (a) Rahn, R. O.; Hosszu, J . L. Biochim. Biophys. Acta 1969, 190,
126-131. (b) Patrick, M. H.; Gray, D. M. Photochem. Photobiol. 1976,
24, 507-513.
(32) Norberg, J .; Nilsson, L. Biophys. J . 1995, 69, 2277-2285.
(33) Saenger, W. In Principles of Nucleic Acid Structure; Cantor,
C. R., Ed.; Springer-Verlag: New York, 1984.
(34) Malathi, R.; Yathindra, N. J . Biomol. Struct. Dyn. 1985, 3, 127-
144 and references therein.
(35) Norberg, J .; Nilsson, L. J . Am. Chem. Soc. 1995, 117, 10832-
10840.
(36) Malathi, R.; Yathindra, N. Int. J . Biol. Macromol. 1982, 4, 18-
24.
(37) Yathindra, N.; Sundaralingam, M. Biopolymers 1979, 18, 2721-
2731.
(38) Turro, N. J . In Molecular Photochemistry; Breslow, R., Karplus,
M., Eds.; W. A. Benjamin: New York, 1967.
(39) Ruzsicska, B. P.; Lemaire, D. G. E. In CRC Handbook of
Organic Photochemistry and Photobiology; Horspool, W. M., Song, P.-
S., Eds.; CRC Press: Boca Raton, FL, 1995; pp 1289-1317.
6508 J . Org. Chem., Vol. 68, No. 17, 2003

Photochemistry of Thymidyl(3′-5′)thymidine
layers were dried with Na2SO4 and evaporated. The residue
was chromatographed on silica gel (particule size 6-35 µm)
using a gradient of MeOH in CH2Cl2 (1-4%) as eluent to afford
the fully protected dimer 8. Compound 8 was dissolved in
MeOH/concentrated aqueous ammonia (1/1; 4 mL), and the
solution was stirred at room temperature overnight. The
residue obtained after evaporation was dissolved in 80%
aqueous acetic acid (4 mL). The resulting solution was stirred
at room temperature for 6 h, evaporated, and coevaporated
with water. Water and CH2Cl2 were added, and the aqueous
phase was separated, washed with CH2Cl2, and evaporated.
Chromatography was performed on silica gel using a gradient
of MeOH in CH2Cl2 (20-25%) containing concentrated aqueous
ammonia (5%) as eluent to afford, after dissolution in water,
filtration through cotton, and evaporation, compound 1a as a
white glassy material (8 mg, 26% yield based on 7, four
steps): 1H NMR (D2O; 500 MHz) δ 7.93 (s, 1H, H6 -pTmoa),
7.85 (s, 1H, H6 Tmop-a), 6.09 (d, 3J (H,H) ) 3.2 Hz, 1H, H1′
Exp er im en ta l Section
5′-O-(4,4′-Dim eth oxytr ityl)-2′-O,5-d im eth ylu r id in e (5).
Dry 2,2′-O-anhydro-1-(â-D-arabinofuranosyl)-5-methyluracil
(4;16 130 mg, 0.54 mmol), obtained by repeated coevaporation
with anhydrous pyridine, was dissolved in anhydrous pyridine
(6 mL). To this solution was added dimethoxytrityl chloride
(403 mg, 1.19 mmol, 2.2 equiv) and DMAP (6 mg). The solution
was stirred at room temperature overnight and then evapo-
rated. The residue was dissolved in CH2Cl2 and washed with
5% aqueous NaHCO3. The aqueous phase was extracted four
times with CH2Cl2, and the organic layers were combined,
dried with Na2SO4, and evaporated. The oily residue was
coevaporated twice with toluene and triturated with diethyl
ether. The liquid was removed by decantation, the solid residue
was dissolved in CH2Cl2 (1 mL), and 50 mL of diethyl ether
was added. The solution was kept at 5 °C for 1 h, and then
the organic phase was decanted, affording a precipitate (292
mg) that was dried under vacuum and used as such in the
next step. To magnesium methoxide prepared from magnesium
turnings (25 mg, 1.038 mmol) in dry MeOH (1 mL) were added
the above-mentioned precipitate (94 mg) and DMF (3 mL). The
resulting suspension was heated at 100 °C for 2.5 h. After
elimination of the solvent by filtration, the residue was
suspended in CH2Cl2 and stirred at room temperature for 2
h. The residue obtained after filtration of the suspension and
concentration of the filtrate was purified by silica gel chro-
matography. Elution with 0-2% CH2Cl2/MeOH afforded in
68% yield compound 5 as a foam whose 1H NMR data were
identical with those in the literature.12
3′-O-Acetyl-2′-O,5-d im eth ylu r id in e (7). To a solution of
5′-O-(4,4′-dimethoxytrityl)-2′-O,5-dimethyluridine (5; 240 mg,
0.418 mmol) in anhydrous pyridine (30 mL) was added acetic
anhydride (3.9 mL, 41.8 mmol). The mixture was stirred at
room temperature overnight and then concentrated. The
residue was dissolved in MeOH, coevaporated with MeOH, and
concentrated to dryness to give an oil. This latter was dissolved
in anhydrous CH2Cl2 (80 mL), and trifluoroacetic acid (2.4 mL)
was added. The solution was stirred at room temperature for
30 min, treated with MeOH (3 mL), and diluted with CH2Cl2
(40 mL). After washing with 5% aqueous NaHCO3 and then
with brine, the aqueous phase was extracted five times with
CH2Cl2. The organic layers were combined, dried with Na2SO4,
and evaporated. The residue was purified by silica gel chro-
matography using a gradient of MeOH in CH2Cl2 (2-3%) as
the eluent to afford 7 as a white foam (93 mg, 71% yield based
on 5, two steps): 1H NMR (CDCl3; 250 MHz) δ 7.44 (s, 1H),
5.69 (d, 1H, 3J (H,H) ) 5.6 Hz), 5.34 (dd, 3J (H,H) ) 5.1, 4.0
Hz, 1H), 4.30 (t, 3J (H,H) ) 5.4 Hz, 1H), 4.19 (m, 1H), 3.95
3
-pTmo), 5.95 (d, J (H,H) ) 2.4 Hz, 1H, H1′ Tmop-), 4.64 (ddd,
3
3J (H,H) ) 4.9, 7.4, 8.3 Hz, 1H, H3′ Tmop-), 4.49 (dd, J (H,H)
) 5.2; 6.2 Hz, 1H, H3′ -pTmo), 4.34-4.27 (m, 3H, H4′ Tmop-,
3
H4′ -pTmo, H5′ -pTmob), 4.24 (dd, J (H,H) ) 2.4, 4.9 Hz, 1H,
H2′ Tmop-), 4.17 (dt, 2J (H,H) ) 12.0, 3J (H,H) ) 3.0 Hz, 1H,
3
2
H5" -pTmob), 4.05 (dd, J (H,H) ) 2.4, J (H,H) ) 13.3 Hz, 1H,
3
H5′ Tmop-c), 4.03 (dd, J (H,H) ) 3.2, 5.5 Hz, 1H, H2′ -pTmo),
3.91 (dd, J (H,H) ) 13.3, J (H,H) ) 3.1 Hz, 1H, H5′ Tmop-c),
3.64 (s, 3H, OCH3), 3.57 (s, 3H, OCH3), 1.90 (s, 6H, CH3 Tmop-,
CH3 -pTmo); 13C NMR (CD3OD; 75.5 MHz) δ 167.5, 167.2,
152.7, 152.5, 137.9, 137.4, 112.6, 112.1, 88.4, 88.2, 84.3, 84.0,
83.4, 82.6, 71.8, 68.9, 64.7, 60.2, 59.3, 58.7, 12.9, 12.7; 31P NMR
(D2O; 243 MHz) δ -0.61; HRMS (FAB) (M + H)+ calcd for
2
3
C
22H32N4O14P 607.1653, found 607.1654.
Ir r a d ia tion Con d ition s. For NMR analysis, a 1 mL D2O
solution of 1a (OD260 ) 22.9) in a quartz NMR tube was
1
exposed for 7 h 30 min to the 254 nm light (2 × 15 W). A H
NMR spectrum was recorded after 1 h 30 min, 4 h 30 min,
and 7 h 30 min. For HPLC analysis, an aqueous solution (500
µL; HPLC grade) of compound 1a or 1b (OD260 ) 6.3 and 16.8
for 1a and 1b, respectively) or a 1/1 mixture of 1a and 1b (pH
6; OD260 ) 6.3 for each solution) in a quartz cuvette was
exposed for 2 h to the 254 nm light source. An aliquot of the
solution was sampled at t ) 0, t ) 1 h, and t ) 2 h and
analyzed by reverse-phase HPLC.
HP LC An a lysis. Ten to thirty microliters of the irradiation
mixture was injected on a SYMMETRY C18 (5 µm, 100Å, 4.6
× 250 mm) column using a 50 mn, 1 mL/mn linear gradient
of 0-15% CH3CN in 0.05 M aqueous ammonium acetate (pH
6.8). A photodiode array detector was used. Calculations were
made using peak areas measured at 230 nm.
2
3
2
(dd, J (H,H) ) 12.5, J (H,H) ) 1.9 Hz, 1H), 3.79 (dd, J (H,H)
Retention time (mn): 1a , 40.1; 1b, 34.5; 10a , 22.0; 10b, 15.1;
11a , 20.8; 11b, 11.2.
3
) 12.4, J (H,H) ) 2.0 Hz, 1H), 3.43 (s, 3H), 2.16 (s, 3H), 1.92
(s, 3H); 13C NMR (CDCl3; 75.5 MHz) δ 171.1, 164.5, 151.2,
138.3, 112.0, 91.0, 83.9, 81.4, 71.3, 62.4, 59.7, 21.5, 13.1; HRMS
(CI) (M + H)+ calcd for C13H19N2O7 315.1192, found 315.1185.
A sample of 7 was deprotected (1/1 MeOH/concentrated
Cir cu la r Dich r oism . Nucleosides and dinucleoside phos-
phates were dissolved in a phosphate buffer solution (0.1 M,
pH 7, 1 M NaCl) using cells with 1 cm path lengths. The
temperature was increased in 10 °C intervals from 20 to 80
°C. CD spectra were obtained in the 225-330 nm range at
0.2 nm intervals. The buffer baseline at each temperature was
subtracted from the sample data and the resulting spectrum
smoothed and converted in molar ellipticity per residue [θ].
Concentrations were based on the UV absorbance at 267 nm
for thymidine 2b and at 268 nm for methoxythymidine 2a ,
assuming an extinction coefficient (ꢀ) of 9.65 × 103 and 9.52
103 M-1 cm-1, respectively:46,12 concentration of 2b 0.108 mM
and concentration of 2a 0.0921 mM. Concentrations of 1a and
1b were calculated using their optical density at 80 °C and
considering that, at this temperature, stacking interactions
can be neglected and, therefore, their extinction coefficients
(per residue) are the same as those for their corresponding
monomers: concentration of 1a 0.0528 mM and concentration
of 1b 0.0717 mM.
aqueous ammonia) to give 2a ,12 whose H NMR spectrum was
1
recorded in D2O: 1H NMR (D2O; 300 MHz) δ 7.7 (s, 1H), 5.99
(d, 3J (H,H) ) 4.4 Hz, 1H), 4.36 (t, 3J (H,H) ) 5.5 Hz, 1H), 4.13-
4.01 (m, 2H), 3.86 (m, 2H), 3.50 (s, 3H), 1.90 (s, 3H).
2′-O,5-Dim et h ylu r id ylyl(3′-5′)-2′-O,5-d im et h ylu r id in e
(1a ). To a solution of compound 7 (16 mg, 0.05 mmol) in
anhydrous CH3CN (1 mL) was added under argon a solution
of tetrazole (12 mg, 0.165 mmol) in anhydrous CH3CN (1 mL)
followed by a solution of 612 (58 mg, 0.075 mmol) in anhydrous
CH3CN (2 mL). The mixture was stirred under argon at room
temperature for 30 min, and then a solution of iodine (38 mg,
0.15 mmol) in THF/H2O/2,6-lutidine (2/1/1; 0.5 mL) was added.
After it was stirred for 20 min at room temperature, the
mixture was evaporated and the residue was dissolved in
CH2Cl2 (1 mL). A saturated aqueous solution of Na2S2O3 was
dropped into it until decolorization, followed by H2O to obtain
the two-phase system. The separated aqueous layer was
extracted several times with CH2Cl2. The combined organic
(46) Sober, H. A. CRC Handbook of Biochemistry; Sober, H. A., Ed.;
CRC Press: Cleveland, OH, 1968; p G-50.
J . Org. Chem, Vol. 68, No. 17, 2003 6509

Ostrowski et al.
1a , HPLC chromatograms of the photolysis of 1b, 1a , and a
mixture of 1b and 1a . and peak areas of the HPLC chromato-
grams (230 nm) of the photolysis of a mixture of 1a and 1b.
This material is available free of charge via the Internet at
http://pubs.acs.org.
Ack n ow led gm en t. We thank Professor P. Potier for
his encouragement and the CNRS for financial support
(PCV 2000 and a postdoctoral fellowship to T.O.).
Su p p or tin g In for m a tion Ava ila ble: Text giving general
methods, figures giving 1H (1D and COSY) NMR spectra of
1
1a , the H NMR spectrum of the crude irradiation mixture of
J O030086P
6510 J . Org. Chem., Vol. 68, No. 17, 2003