Naphthalene, phenanthrene, and pyrene as DNA base analogues: Synthesis, structure, and fluorescence in DNA

Article abstract of DOI:10.1021/ja9612763

We describe the synthesis, structures, and DNA incorporation of deoxyribonucleosides carrying polycyclic aromatic hydrocarbons as the DNA 'base' analogue. The new polycyclic compounds are 1-naphthyl, 2-naphthyl, 9-phenanthrenyl, and 1-pyrenyl deoxynucleos

Full text of DOI:10.1021/ja9612763

J. Am. Chem. Soc. 1996, 118, 7671-7678
7671
Naphthalene, Phenanthrene, and Pyrene as DNA Base
Analogues: Synthesis, Structure, and Fluorescence in DNA
Rex X.-F. Ren, Narayan C. Chaudhuri, Pamela L. Paris, Squire Rumney IV, and
Eric T. Kool*,†
Contribution from the Department of Chemistry, UniVersity of Rochester,
Rochester, New York 14627
ReceiVed April 18, 1996X
Abstract: We describe the synthesis, structures, and DNA incorporation of deoxyribonucleosides carrying polycyclic
aromatic hydrocarbons as the DNA “base” analogue. The new polycyclic compounds are 1-naphthyl, 2-naphthyl,
9-phenanthrenyl, and 1-pyrenyl deoxynucleosides. The compounds are synthesized using a recently developed
C-glycosidic bond formation method involving organocadmium derivatives of the aromatic compounds coupling
with a 1R-chlorodeoxyribose precursor. The principal products of this coupling are the R-anomers of the
deoxyribosides. An efficient method has also been developed for epimerization of the R-anomers to â-anomers by
acid-catalyzed equilibration; this isomerization is successfully carried out on the four polycyclic nucleosides as well
as two substituted phenyl nucleosides. The geometry of the anomeric substitution is derived from 1H NOE experiments
and is also correlated with a single-crystal X-ray structure of one R-isomer. Three of the polycyclic C-nucleoside
derivatives are incorporated into DNA oligonucleotides via their phosphoramidite derivatives; the pyrenyl and
phenanthrenyl derivatives are shown to be fluorescent in a DNA sequence. The results (1) broaden the scope of our
C-glycoside coupling reaction, (2) demonstrate that (using a new acid-catalyzed epimerization) both R- and â-anomers
are easily synthesized, and (3) constitute a new class of deoxynucleoside derivatives. Such nucleoside analogues
may be useful as biophysical probes for the study of noncovalent interactions such as aromatic π-stacking in DNA.
In addition, the fluorescence of the phenanthrene and pyrene nucleosides may make them especially useful as structural
probes.
Introduction
probes in biophysical and biochemical studies.8 In contrast to
placement of such reporter groups at the end of a DNA strand
using nonnucleotide linkers, the attachment of a reporter to a
DNA base allows for placement and probing even near the
middle of a stretch of DNA. Such a strategy has found
considerable practical use in fluorescence-based automated DNA
sequencing.9 An alternative approach to the conjugation of a
fluorophore to a natural DNA base is the more direct modifica-
tion of a DNA base itself to render it fluorescent. A number
of modified DNA bases with useful fluorescence properties have
been reported recently; among the most widely used nucleosides
of this type are 2-aminopurine10 and ethenoadenosine.11
A large number of nonnatural analogues of DNA nucleosides
have been synthesized in recent years. Changing the structure
of the base moiety attached to deoxyribose has been a useful
strategy for probing structure and function in DNA. For
example, a number of base analogues have been used to test
the importance of specific hydrogen bonding interactions which
may be important for function of the natural nucleic acid
bases.1-3 Using this strategy, workers have examined the
importance of hydrogen bonding in stabilizing DNA and RNA
structure,1 in protein-DNA interactions,2 and in the fidelity of
enzymatic DNA and RNA synthesis.3
We have undertaken a program to develop a new class of
DNA base analogues which are designed to serve as biophysical
probes.12 The molecules we have chosen to synthesize and
study are nucleosides having “base” moieties which are non-
polar, weakly hydrogen bonding aromatic groups. We have
described the synthesis13 and study14 of substituted benzene-
derived nucleosides as nonpolar isosteres of pyrimidine DNA
bases; these compounds were intended as ideal steric mimics
of the natural analogs but with little or no hydrogen bonding
Modified DNA bases have also been synthesized with the
purpose of serving as reporter groups in physical and biochemi-
cal studies of structure and function. Examples of reporter
groups which have been attached to DNA bases include biotin4
and digoxigenin5 groups, spin-label groups,6 and DNA-cleaving
moieties.7 Among the most prominent class of reporters used
in DNA are fluorescent-tagged DNA bases which can serve as
* Author to whom correspondence should be addressed.
† e-mail: etk@etk.chem.rochester.edu.
X Abstract published in AdVance ACS Abstracts, August 1, 1996.
(1) SantaLucia, J.; Kierzek, R.; Turner, D. H. Science 1992, 256, 217.
(2) (a) Smith, S. A.; Rajur, S. B.; McLaughlin, L. W. Nature Struct.
Biol. 1994, 1, 198. (b) Lesser, D. R.; Kurpiewski, M. R.; Jen-Jacobson, L.
Science 1990, 250, 776.
(7) Dreyer, G. B.; Dervan, P. B. Proc. Natl. Acad. Sci. U.S.A. 1985, 82,
968.
(8) Beaucage, S. L.; Iyer, R. P. Tetrahedron 1993, 49, 1925.
(9) (a) Ansorge, W.; Sproat, B. S.; Stegemann, J.; Schwager, C. J.
Biochem. Biophys. Methods 1986, 13, 315. (b) Karger, A. E.; Harris, J.
M.; Gesteland, R. F. Nucleic Acids Res. 1991, 19, 4955.
(10) (a) Ward, D. C.; Reich, E.; Stryer, L. J. Biol. Chem. 1969, 244,
1228. (b) Bloom, L. B.; Otto, M. R.; Beechem, J. M.; Goodman, M. F.
Biochemistry 1993, 32, 11247.
(11) (a) Secrist, J. A.; Barrio, J. R.; Leonard, N. J. Science 1972, 175,
646. (b) Toulme´, J. J.; He´lene, C. Biochim. Biophys. Acta 1980, 606, 95.
(12) Schweitzer, B. A.; Kool, E. T. J. Org. Chem. 1994, 59, 7238.
(13) (a) Chaudhuri, N. C.; Kool, E. T. Tetrahedron Lett. 1995, 1795.
(b) Chaudhuri, N. C.; Kool, E. T. Ibid. 1995, 4910.
(3) (a) Kornberg, A.; Baker, T. A. DNA Replication, 2nd ed.; W. H.
Freeman: New York, 1992. (b) Echols, H.; Goodman, M. F. Ann. ReV.
Biochem. 1991, 60, 477. (c) Strazewski, P.; Tamm, C. Angew. Chem., Int.
Ed. Engl. 1990, 29, 36.
(4) Kempe, T.; Sundquist, W. I.; Chow, F.; Hu, S. L. Nucleic Acids Res.
1985, 13, 45.
(5) Zischler, H.; Nanda, I.; Schafer, R.; Schmid, M.; Epplen, J. T. Hum.
Genet. 1989, 82, 227.
(6) (a) Weygand-Durasevic, I.; Susic, S. Biochim. Biophys. Acta 1990,
1048, 38. (b) Hustedt, E. J.; Spaltenstein, A.; Kirchner, J. J.; Hopkins, P.
B.; Robinson, B. H. Biochemistry 1993, 32, 1774.
(14) Schweitzer, B. A.; Kool, E. T. J. Am. Chem. Soc. 1995, 117, 1863.
S0002-7863(96)01276-0 CCC: $12.00 © 1996 American Chemical Society

7672 J. Am. Chem. Soc., Vol. 118, No. 33, 1996
Ren et al.
potential. Similarly, we have synthesized indole- and benz-
imidazole-derived nucleosides as purine isosteric mimics.12
We have recently begun to focus on base stacking interactions
as an important noncovalent interaction which has received very
little experimental attention in the context of DNA. Since
surface area, electrostatics, polarizability, and hydrophobicity
are all factors which may influence π-stacking interactions in
aqueous solution,15 we felt that a useful test of these effects
might be probed with a simple series of aromatic compounds
standing in for DNA bases in nucleic acid structures.
Scheme 1
With those aims in mind we describe here the synthesis of a
series of DNA nucleoside analogues which contain polycyclic
aromatic hydrocarbons as the DNA “base” equivalents. New
analogues containing pyrene, phenanthrene, and two naphthalene
isomers are described. The aromatic rings are coupled to the
sugar using a modification of our recently developed orga-
nocadmium strategy.14 Also described is a new method for
isomerization of such C-nucleosides from R- to naturally-
configured â-epimers. The title compounds are synthesized in
a concise manner and in relatively good yields, thus establishing
the relatively broad scope of both the coupling and epimerization
methods for C-nucleoside synthesis in general. We further
demonstrate the incorporation into DNA of naphthalene,
phenanthrene, and pyrene nucleosides using standard automated
methods. In addition to the potential utility of the analogues
in aromatic stacking studies, it is found that the pyrene and
phenanthrene nucleosides in particular show fluorescence emis-
sion properties in DNA which may enhance their utility as
probes of structure and dynamics.
derivatives,13 this coupling with the larger polycyclic aromatics
yields a mixture of R- and â-anomers in good overall yields
(54-81% isolated yields). The major isomer in all four cases
is formed with retention of configuration; thus, R-anomeric
C-nucleosides (the p-toluoyl esters of 1a, 2a, 3a, and 4a) are
the primary products. Measured ratios of the two isomers (by
NMR integration) ranged from 5:1 (R:â) for the 1-naphthyl
derivative to 10:1 for the 9-phenanthrenyl derivative. The
configuration at the C-1′ carbons of all isomers was determined
by analysis of H1′-H2′ coupling constants for the protected
nucleosides, by 1H nuclear Overhauser experiments on the
deprotected nucleosides, and by correlation with an X-ray crystal
structure of one of the R-anomeric compounds (see below).
Although the desired â-anomers (the toluoyl esters of 1-4)
could also be isolated from this coupling reaction, the yields
were less than ideal. Studies were thus undertaken to find
conditions under which the predominant R-anomers could be
converted to the desired â-configuration. It was anticipated that
acidic conditions might allow epimerization at the C-1 position
by reversible ring-opening. Experimentation with several sets
of conditions revealed that benzenesulfonic acid in refluxing
xylene, in the presence of a small amount of water, did indeed
result in ready equilibration of the R-anomers to mixtures of â-
and R-isomers after several hours. Addition of a small amount
of water was found to be necessary for the isomerization. The
equilibration was then carried out for all four R-isomers (1a,
2a, 3a, and 4a) as their bis-toluoyl esters. We also tested the
isomerization on two previously reported substituted benzene
nucleosides (the toluoyl esters of 5a and 6a) to test the scope
of the reaction. Significantly, the major component of each
mixture after equilibration was in all cases the desired â-anomer
(1-6). The ratios of â- to R-isomers ranged from 4:1 for the
trimethylbenzene derivative to 2.5:1 for the 1-naphthyl deriva-
tive. Isolated yields of the desired â-anomers after column
chromatography ranged from 28 to 54%. The R-anomers and
mixed fractions could be reisolated and recycled in the isomer-
ization if desired. Interestingly, the deprotected free nucleosides
themselves did not undergo any observable isomerization under
these conditions, even at extended reaction times.
Results
Unsubstituted polycyclic aromatic hydrocarbons have not
commonly been incorporated into deoxyribonucleosides as the
directly coupled C-nucleosides, although substituted16a and
unsubstituted16b naphthyl ribonucleosides have been reported.16
A naphthyl deoxynucleoside was synthesized for use in poly-
merase chain reaction experiments, although its structure was
not characterized in detail.17 The smallest monocyclic member
of the series, benzene, has been incorporated previously as a
ribonucleoside18 and also as a deoxyribonucleoside,19 and the
latter was briefly studied when incorporated into a DNA strand.
We recently described a relatively high-yield strategy for
coupling of aromatic benzene-derived compounds to the gly-
cosidic position of deoxyribose.13 Subsequent studies showed
that the primary products derived from these reactions are
R-configured isomers, with the â-isomers appearing as minor
products.13b However, recent work has shown (see below) that
it is possible to equilibrate these R-C-nucleosides to â-isomers
in good yield, and so the coupling method can now be applied
generally to the synthesis of both anomers of deoxynucleosides.
Synthesis. The previously described method of C-nucleoside
coupling13 was utilized to generate the new aromatic nucleosides
1-4 as their bis-toluoyl esters (Scheme 1). The method
involves the reaction of organocadmium derivatives of the
aromatic species with the well-known R-chlorosugar synthon
of Hoffer20 (Scheme 1). As seen in reactions with benzene
With the new method for epimerization to â-anomeric
configuration in hand, the synthetic scheme made possible the
facile generation of the six aromatic C-nucleosides (1-6) in
generally good yields (Scheme 2). The toluoyl protecting
groups were removed in methanolic base with yields ranging
from 50 to 78%. Following this overall scheme, the free
unprotected nucleosides were produced in a total of only three
steps (aromatic coupling, isomerization, ester deprotection).
(15) Hunter, C. A. Angew. Chem., Int. Ed. Engl. 1993, 32, 1584.
(16) (a) Matsumoto, T.; Katsuki, M.; Suzuki, K. Tetrahedron Lett. 1988,
29, 6935. (b) Ohrui, H.; Kuzuhara, H.; Emoto, S. Agr. Biol. Chem. (Tokyo)
1972, 36, 1651-1653.
(17) Newton, C. R.; Holland, D.; Hepinstall, L. E.; Hodgson, I.; Edge,
M. D.; Markham, A. F.; McLean, M. J. Nucleic Acids Res. 1993, 21, 1155.
(18) Klein, R. S.; Kotick, M. P.; Watanabe, K. A.; Fox, J. J. J. Org.
Chem. 1971, 36, 4113.
(19) Millican, T. A.; Mock, G. A.; Chauncey, M. A.; Patel, T. P.; Eaton,
M. A. W.; Gunning, J.; Cutbush, S. D.; Neidle, S.; Mann, J. Nucleic Acids
Res. 1984, 12, 7435.
Structural Assignments. The structural assignment of
anomeric configuration for compounds 1-6 (and 1a-6a) was
made by 1H NOE studies of all compounds, by examination of
coupling constants for H1′ and H2′ protons, and by correlation
with an X-ray crystal structure of the R-1-naphthyl nucleoside
3a (below). In addition, the compounds were characterized by
(20) Hoffer, M. Chem. Ber. 1960, 93, 2777.

Polycyclic Aromatic DNA Bases
J. Am. Chem. Soc., Vol. 118, No. 33, 1996 7673
Figure 1. Illustration of qualitative differences in nuclear Overhauser
enhancements observed for R- (left) and â-anomers (right) of com-
pounds 1-6. In R-anomers, irradiation of the C-2′-â proton gives
enhancements in both the C-1′ and C-3′ protons, while irradiation of
the C-2′-R proton gives little or no enhancement for either. In the
â-anomers, irradiation of the C-2′-â proton gives enhancement only in
the C-3′ proton, while irradiation of the C-2′-R proton gives enhance-
ment only in the C-1′ proton.
Table 1. H1′-H2′ Coupling Constants and Proton NOE Data for
â-Isomers of Aryl Nucleosides 1-6 in CD3OD
1
their H and 13C NMR spectra and by high-resolution mass
spectrometry.
Proton nuclear Overhauser effects were used to examine the
geometries of the anomeric isomers of compounds 1-6. The
approach used was to separately irradiate the H-2′ proton
resonances situated at δ 1.7-2.7 and observe enhancements at
vicinal 1′ and 3′ protons (see Figure 1 and Tables 1 and 2).
Although specific assignment of which resonance corresponds
to 2′-R and which to 2′-â could not be made a priori, analysis
of predicted NOE effects makes possible a simple approach to
assignment of stereochemistry at the 1′ position. Examination
of the structures of R- and â-nucleosides (Figure 1) shows that
for R-anomers the 2′-â proton is in close proximity to both the
1′ and 3′ protons, while the C-2′-R proton is not near either
one of these protons. In â-anomers, on the other hand, the 2′-â
proton is near only the 3′ proton, while the 2′-R proton is only
near the 1′ proton. Thus, in an R-anomer, separate irradiation
of each of the C-2′ protons should lead to two and zero NOE
enhancements at the vicinal protons, while in a â-anomer these
two irradiations would lead to one significant enhancement for
each irradiation.
To test this prediction we carried out these experiments on
naphthyl nucleoside isomers 3 and 3a (Tables 1 and 2). The
diester of 3a is the principal product of the glycosidic coupling
reaction (with the diester of 3 being a minor product). Irradia-
tion of one of the 2′ protons of the nucleoside gave significant
nuclear Overhauser enhancements of 8 and 7% at the 1′ and 3′
protons; however, irradiation of the other 2′ proton gave no
significant enhancement at either 1′ or 3′ positions. Using the
analysis above, this indicates that this compound is an R anomer.
This assignment was confirmed by a single-crystal X-ray
structure obtained for the compound (below). To complete the
analysis of the two isomers, we carried out the same experiments
on the isomeric nucleoside 3, which is the major product after
Scheme 2

7674 J. Am. Chem. Soc., Vol. 118, No. 33, 1996
Ren et al.
Table 2. H1′-H2′ Coupling Constants and NOE Data for
R-Isomers of Aryl Nucleosides (compounds 3a, 5a, and 6a) in
CD3OD
Figure 2. ORTEP drawing of 1-naphthyl nucleoside nucleoside 3a
from single-crystal X-ray structure. The structure has the R-anomeric
configuration and a C-3′-exo S-type sugar conformation.
relationship empirically adjusted for nucleosides21 predicts J )
9.2 and 2.8 Hz, respectively, indicating a small change in ring
conformation in solution relative to that in the crystal, or else
a nonideal match of the empirical relationship for this C-
nucleoside. Interestingly, although this compound clearly is an
R-anomer, the experimentally measured coupling constants are
more consistent with those commonly seen for â, rather than
R, anomers of natural nucleosides (see below).
epimerization of 3a. Irradiation of one of the 2′ protons gave
an 8% enhancement of the 1′ proton (and none at the 3′ proton),
while irradiation of the other 2′ proton gave a 5% enhancement
of the 3′ proton (and none for the 1′ proton). Thus, this
compound is assigned to the â configuration. Also consistent
with this assignment is a separate experiment in which the 1′
proton was irradiated; here we observed a 6% enhancement at
the 4′ proton (Table 1), while the R-isomer shows no such
enhancement (data not shown).
The NOE experiments were then carried out for the isomers
of 1, 2, and 4-6. The results are shown in Tables 1 and 2.
The results were all consistent with the model, in that one isomer
of each pair gave two and ∼zero enhancements of the vicinal
protons for the two H-2′ irradiations while the other clearly gave
one significant enhancement for each of the two irradiations.
The isomers which gave two and zero enhancements were
assigned as R-anomers, and those with one and one enhance-
ments were assigned to be â-anomers. Also consistent with
these assignments were NOE enhancements in the H-4′ positions
on irradiation of the H-1′ protons for the â-isomers (Table 1)
which were absent in the R-isomers.
These assignments were also internally consistent in that the
major isomers obtained from the glycosidic coupling reaction
were all of the same anomeric configuration (R). Similarly,
the major isomers isolated from the epimerization were all of
the same anomeric configuration (â). In addition, all the isomers
assigned as R had H-1′ resonances which qualitatively appeared
as pseudotriplets (they are actually doublets of doublets), having
both coupling constants near 6.0-8.0 Hz. The isomers assigned
to the â configuration had H-1′ resonances which appeared as
nearly evenly spaced doublets of doublets (J ≈ 5 and 10 Hz).
These H-1′-H-2′ coupling constant trends are consistent with
a literature report of similar coupling constants for a related
â-C-nucleoside19 (although they are reversed relative to obser-
vations for â-N-nucleosides; see Discussion).
Also useful in confirmation of these structural assignments
were X-ray structural data obtained for 1-naphthyl compound
3a (Figure 2). A crystal suitable for analysis was obtained by
recrystallization from methylene chloride/hexane. The structure
shows the R configuration and in analogy to natural nucleosides
the naphthalene is in an anti conformation, with the aromatic
group oriented away from the sugar. The deoxyribose ring is
in a C-3′-exo (S-type) conformation.
Incorporation into DNA. The â-C-deoxynucleosides 1, 2,
and 3 were then carried on with the aim of incorporating them
into DNA oligonucleotides by automated solid-phase methods
(Scheme 2). The incorporation into DNA of substituted benzene
nucleosides 5 and 6 has recently been reported elsewhere.22
Standard methods were used to convert the unprotected nucleo-
sides to 5′-dimethoxytrityl-protected derivatives in yields ranging
from 59 to 92% after purification. These were then converted
into cyanoethyl phosphoramidite derivatives, which were ob-
tained in 50-89% yields after purification by column chroma-
tography. Incorporation into oligodeoxynucleotides was carried
out with standard coupling chemistry but with lengthened
coupling times, and stepwise yields for coupling of these
compounds were >95% by trityl monitoring. To test for intact
incorporation, we synthesized trinucleotides having the sequence
T-X-T and examined them by proton NMR (Figure 3); the
spectra of the crude unpurified oligonucleotides show clear
resonances very similar to those of the free nucleosides and
having the expected aromatic integrations relative to anomeric
C-1′ protons and thymine C-6 protons and C-5 methyl groups.
This confirms both the presence of the intact structures (as
expected for unreactive aromatic hydrocarbons) and the high
coupling yields, since di- and mononucleotides which would
result from incomplete coupling are not seen.
Fluorescence Properties in DNA. Since polycyclic aromat-
ics such those in the nucleosides 1, 2, and 3 have been studied
in other contexts as fluorescent probes,23 we undertook the
examination of possible fluorescence properties of oligonucle-
otides containing these structures in aqueous buffer. We
synthesized heptamer oligodeoxynucleotides having the se-
quence 5′-dXCGCGCG, (where X ) 1, 2, and 3) which are
self-complementary and form duplexes with the polycyclic
aromatic nucleoside situated at the 5′ ends. These were purified
by preparative denaturing gel electrophoresis.
Emission spectra were measured for the three sequences in a
pH 7.0 buffer (10 mM PIPES buffer, 100 mM NaCl, 10 mM
MgCl2) at 25 °C, conditions under which they likely form
(21) Davies, D. B. Prog. Nucl. Magn. Reson. Spectrosc. 1978, 12, 135.
(22) Ren, X.-F.; Schweitzer, B. A.; Sheils, C. J.; Kool, E. T. Angew.
Chem., Int. Ed. Engl. 1996, 35, 743.
(23) (a) Schulman, S. G. Molecular Luminescence Spectroscopy;
Wiley: New York, 1990; pp 1-27. (b) Slavik, J. Fluorescent Probes in
Cellular and Molecular Biology; CRC Press: Boca Raton, 1990; p 1-36.
Experimental H-1′ to H-2′ coupling constants for the ester
of this compound (3a) in CDCl3 were J ) 8.0 and 6.0 Hz. The
corresponding dihedral angles generated from the X-ray structure
are found to be 8.1° and 124.5°. Application of the Karplus

Polycyclic Aromatic DNA Bases
J. Am. Chem. Soc., Vol. 118, No. 33, 1996 7675
Figure 3. 400 MHz proton NMR spectra for trinucleotides (sequence T-X-T) containing (a) pyrenyl, (b) phenanthryl, and (c) naphthyl nucleosides
1-3 at the X position.
duplexes (data not shown). The naphthalene-containing se-
quence showed no emission detectable above background. The
other compounds showed structured fluorescence emission
profiles consistent with published spectra for the polycyclic
aromatic parent structures.23 The phenanthrene-containing
oligonucleotide had the most intense emission (not shown), with
the strongest peak at 370 nm. The pyrene-modified sequence
showed a similar emission profile but with an emission
maximum at 395 nm and with peak intensity considerably lower
(by ca. 700-fold in peak height) than that for the phenanthrene
case, suggesting considerable quenching by the DNA under
these conditions and in this sequence.
the epimerization will convert these initial adducts into the
â-isomers, also in generally good yield.
It is interesting that the cadmium- or zinc-mediated reactions
give primarily R-anomers. This corresponds to retention of
configuration since the starting sugar synthon contains only the
R-chloro isomer with no â-isomer observable by proton NMR.
One possible explanation for this result is that the chloride is
lost in a dissociative process and that the subsequent bond
formation occurs with selective R-face attack by the organo-
metallic species. Further studies will be required to better
elucidate this mechanism.
The Anomeric Effect and Deoxyribose Conformation.
Proton NMR studies of the natural N-nucleosides found in DNA
have shown that the 2′-deoxynucleosides in normal anomeric
â-orientation generally have very similar coupling constants
between the H-1′ proton and the vicinal H-2′ protons.21,24 The
two coupling constants are of the same magnitude; for example,
those of â-thymidine are 7.0 and 7.0 Hz,24 and thus the H-1′
resonance usually appears as a pseudotriplet. This has also held
true for other N-nucleosides containing base moieties not found
in DNA.12 By comparison, the R-anomers of the natural DNA
N-nucleosides have considerably different coupling constants
for this same H1′ resonance; for R-thymidine these are found
to be 8.0 and 3.0 Hz, and the resonance appears as a doublet of
doublets.24 Virtually the same coupling constants are found for
â- and R-deoxyadenosine as well.25 This difference between
the two isomers arises from the different dihedral angles which
are present in the two anomeric isomers.
Discussion
Utility of the C-Nucleoside Synthesis and Epimerization
Methods. The transition metal-mediated coupling reaction used
here to generate aromatic C-nucleosides has been previously
found to be useful in the generation of R-deoxynucleosides (and
small amounts of â-deoxynucleosides) of substituted benzene
derivatives.13 The present results establish that the reactions
can easily be extended to larger aromatic hydrocarbons without
a penalty in yield. Since the only apparent requirement appears
to be the ability for a given aromatic halide to be converted to
the corresponding Grignard species, it seems likely that the
method may well be generally useful in the synthesis of many
different C-deoxynucleosides.
With the addition of the present method for epimerization at
the anomeric center, the scheme for nucleoside assembly
becomes more generally useful. After this equilibration all
nonpolar aromatic nucleosides studied to date (benzenes,
naphthalenes, phenanthrene, and pyrene) give predominantly the
â-anomer as the more stable isomer. Thus, if R-anomers of
C-nucleosides are desired, the coupling reaction will generate
them directly in good yield; however, if â-anomers are desired,
A few reports on C-nucleosides have described quite different
H1′-H2′ coupling behavior for the two anomers.26 These
compounds often appear to have coupling constants which show
(24) Cleve, G.; Hoyer, G.; Schulz, G.; Vorbruggen, H. Chem. Ber. 1973,
106, 3062.
(25) Robins, M. J.; Robins, R. K. J. Am. Chem. Soc. 1965, 87, 4934.

7676 J. Am. Chem. Soc., Vol. 118, No. 33, 1996
Ren et al.
different trends for R- and â-isomers relative to N-nucleosides.
Such differences may arise in part from the presence or absence
of the electronegative nitrogen at the C-1 position. However,
the comparisons between C- and N-nucleosides have generally
been made between pairs of nucleosides which differ in more
than just the presence or absence of polar nitrogens in the rings.
It is possible that steric differences between in a series of
compounds might affect ring geometry as well.
In that light it is useful to compare these H1′-H2′ coupling
constants (and thus ring conformations) for two compoundssone
an N-nucleoside and one a C-nucleosidesin which steric
differences are very small. This helps to eliminate steric effects
in the analysis of anomeric effects. Comparison of â- and
R-thymidine and â- and R-difluorotoluene nucleosides has
shown that they are sterically virtually identical in the base
moieties.12,27 Comparison of the H1′-H2′ coupling constants
shows considerable differences, however. The â-thymidine
values (7.0 and 7.0 Hz)24 are similar to those for the R-anomer
of the difluorotoluene nucleoside (7.6 and 7.6 Hz), and both
resonances appear as pseudotriplets. Conversely, the R-anomer
of thymidine has coupling constants (8.0 and 3.0 Hz) which
are similar to those for the â-anomer of the difluorotoluene
nucleoside (10.4 and 4.6 Hz), and these appear as doublets of
doublets. Thus, our comparison underscores this inverse
relationship and also points out the strong influence of the polar
nitrogen at C-1′ in natural nucleosides on influencing deoxyri-
bose ring geometry.
Indeed, every C-nucleoside we have synthesized to date
(including more than ten different base analogues derived from
aromatic hydrocarbons) has shown the same inverse relationship
for coupling constants relative to N-nucleosides. The 1H NOE
experiments and the X-ray crystal data described herein have
since clarified and solidified the inverse relationship.
Potential Utility of Polycyclic Aromatic Nucleosides as
Probes in DNA. While a good deal of experimental data have
been examined on the relative effects of base stacking and
hydrogen bonding on the stability of RNA helices,28 much less
data have been generated on the topic of base stacking in DNA.
“Dangling end” effects have been studied as measures of
stacking of all four bases in RNA helices,28 but in DNA only
thymine has been briefly examined using this approach.29 Since
it is likely that electrostatics, van der Waals effects, and
hydrophobicity may all play a role in base stacking in DNA,
we synthesized the polycyclic aromatic compounds described
here as possible probes of some of those effects. A detailed
study of base stacking in DNA using both natural and nonnatural
nucleosides will be reported in due course.
dynamics of nucleic acid helices. Experiments directed to this
possibility are now underway.
Conclusions
We have developed a convenient synthesis of nucleoside
analogues containing aromatic hydrocarbons as the “base”
moiety. The synthetic approach involves coupling of orga-
nocadmium derivatives of aromatic hydrocarbons to a widely-
used R-chlorosugar synthon. This coupling works efficiently
and generates predominantly R-anomers for compounds 1-6.
â-Anomers can be obtained as the major product by a new acid-
catalyzed isomerization of the R-compounds. The overall
scheme thus allows generation of either R- or â-anomer of
C-nucleosides, as desired, in good yields. The polycyclic
compounds are quite stable and can be incorporated into DNA
oligonucleotides using standard procedures. Such compounds
may be useful as probes in DNA, and the pyrene and
phenanthrene nucleosides in particular have fluorescence emis-
sion properties which make them potentially useful as reporter
groups in nucleic acids.
Experimental Section
1H, 13C, and 31P NMR spectra were recorded with a 300 MHz
spectrometer unless otherwise noted, chemical shifts are given in δ
(ppm) using solvent as internal reference, and the coupling constants
are in hertz (Hz). NOE difference spectra were also performed on a
300 MHz instrument. The mass spectra were performed using electron
impact or chemical ionization. All reactions were monitored by thin-
layer chromatography (TLC) using EM Reagents plates with fluores-
cence indicator (SiO2-60, F-254). Flash column chromatography was
conducted using EM Science silica gel 60 (230-400 mesh). Mass
spectral analyses were performed by the University of California,
Riverside Mass Spectrometry Facility, Riverside, CA. All reactions
were carried out under a nitrogen atmosphere in dry, freshly distilled
solvents under anhydrous conditions unless otherwise specified. THF
was distilled from sodium metal/benzophenone, methylene chloride was
distilled from NaH, and pyridine was distilled from BaO prior to use.
Procedure for Glycosidic Coupling Reaction and Isolation of
Major r-Epimers as Bis-p-toluoyl esters of 1-6a. Dry THF (5 mL)
was placed in a round-bottomed flask equipped with a condenser, drying
tube, and addition funnel. Magnesium turnings (0.3 g, 1.2 mmol) and
a few crystals of iodine were added. 1-Bromopyrene (0.35 g, 1.2 mmol)
was added to the mixture. Slight heating was needed (40 °C) to drive
the reaction to completion. After formation of the Grignard reagent
was complete (∼1 h), dry CdCl2 (110 mg, 0.6 mmol) was added and
the reaction mixture was continuously heated under reflux for 1 h. 1′R-
Chloro-3′,5′-di-O-toluoyl-2′-deoxyribose20 (0.51 g, 1.3 mmol) was then
added to the above mixture in one portion. The solution was stirred at
room temperature for 4 h under an atmosphere of N2. The solution
was poured into 10% ammonium chloride (2 × 50 mL) and extracted
with methylene chloride. The organic layers were washed with
saturated sodium bicarbonate and brine and dried over anhydrous
magnesium sulfate. The solution was filtered, concentrated, and
purified by flash silica gel chromatography, eluting with hexanes-
ethyl acetate (9:1). The major product 1a bis-toluoyl ester was
obtained as a pale yellow oil (R-epimer, 48% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.80 (2H, d, J ) 8.0), 8.72 (2H, d, J ) 8.0), 8.05 (1H,
s), 7.92-8.00 (2H, m), 7.72-7.60 (4H, m), 7.58 (2H, d, J ) 8.0), 7.32
(2H, d, J ) 8.0), 6.96 (2H, d, J ) 8.0), 6.15 (1H, dd, J ) 8.2, 6.0),
5.76 (1H, m), 4.98 (1H, m), 4.75-4.65 (2H, m), 3.30-3.22 (1H, m),
Both the pyrene and phenanthrene nucleosides are also of
interest for their fluorescence properties. Pyrene in particular
has been incorporated into DNA in a number of studies as a
potential reporter group.30 In most previous cases the pyrene
has been incorporated at the end of strands by a simple flexible
linker chain. This flexibility allows the pyrene chromophore
to adopt a number of different conformations. The present
approach allows pyrene or phenanthrene to be inserted within
a DNA strand at any position and potentially to remain more
rigidly stacked within the helix. This closer contact with the
DNA may give more sensitive information about structure and
(26) (a) Srivastava, P. C.; Robins, P. K.; Takusagawa, F.; Berman, H.
M. J. Heterocycl. Chem. 1981, 18, 1659. (b) Hacksell, U.; Cheng, J. C.-Y.;
Daves, G. D., Jr. J. Carbohydr. Chem. 1986, 5, 287.
(27) Schweitzer, B. A.; Sheils, C. J.; Ren, X.-F.; Chaudhuri, N. C.; Kool,
E. T. In Biological Structure and Dynamics; Sarma, R. H., Sarma, M. H.,
Eds.; Adenine Press: Albany, 1996; Vol. 2, pp 209-216.
(28) Sugimoto, N.; Kierzek, R.; Turner, D. H. Biochemistry 1987, 26,
4554.
(30) (a) Telser, J.; Cruickshank, K. A.; Morrison, L. E.; Netzel, T. L.;
Chan, C. J. Am. Chem. Soc. 1989, 111, 7226. (b) Telser, J.; Cruickshank,
K. A.; Morrison, L. E.; Netzel, T. L. J. Am. Chem. Soc. 1989, 111, 6966.
(c) Lee, H.; Hinz, M.; Stezowski, J. J.; Harvey, R. G. Tetrahedron Lett.
1990, 31, 6773. (d) Yamana, K.; Gokota, T.; Ozaki, H.; Nakano, H.; Sangen,
O.; Shimidzu, T. Nucleosides Nucleotides 1992, 11, 383. (e) Prokhorenko,
I. A.; Petrov, A. A.; Gontarev, S. V.; Berlin, Y. A. BioMed. Chem. Lett.
1995, 5, 2081. (f) Li, Y.; Bevilacqua, P. C.; Mathews, D.; Turner, D. H.
Biochemistry 1995, 34, 14394. (g) Tong, G.; Lawlor, J. M.; Tregear, G.
W.; Haralambidis, J. J. Am. Chem. Soc. 1995, 117, 12151.
(29) Senior, M.; Jones, R. A.; Breslauer, K. J. Biochemistry 1988, 27,
3879.

Polycyclic Aromatic DNA Bases
J. Am. Chem. Soc., Vol. 118, No. 33, 1996 7677
3.50-3.45 (1H, m), 3.44 (3H, s), 3.38 (3H, s); HRMS (FAB, 3-NBA
matrix) calcd for C37H31O5 (M + 1) 554.2093, found 554.2069.
2a bis-toluoyl ester (R-epimer, 43% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.80 (2H, d, J ) 8.0), 8.72 (2H, d, J ) 8.0), 8.05 (1H,
s), 7.92-8.00 (2H, m), 7.72-7.60 (4H, m), 7.58 (2H, d, J ) 8.0), 7.32
(2H, d, J ) 8.0), 6.96 (2H, d, J ) 8.0), 6.15 (1H, dd, J ) 8.2, 6.0),
5.76 (1H, m), 4.98 (1H, m), 4.75-4.65 (2H, m), 3.30-3.22 (1H, m),
3.50-3.45 (1H, m), 3.44 (3H, s), 3.38 (3H, s); HRMS (FAB, 3-NBA
matrix) calcd for C35H31O5 (M + 1) 531.2172, found 531.2174.
3a bis-toluoyl ester (R-epimer, 52% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.05 (2H, d, J ) 8.0), 7.95 (2H, m), 7.83 (2H,
overlapped d), 7.71 (2H, d, J ) 8.0), 7.55 (3H, m), 7.32 (2H, d, J )
8.0), 7.19 (2H, d, J ) 8.0), 6.10 (1H, dd, J ) 8.0, 6.0), 5.69 (1H, m),
4.90 (1H, m), 4.76-4.65 (2H, m), 3.28-3.18 (1H, m), 2.52-2.45 (1H,
m), 2.48 (3H, s), 2.42 (3H, s); HRMS ((FAB, 3-NBA matrix) calcd
for C31H29O5 (M + 1) 481.2015, found 481.2025.
s), 2.47 (3H, s), 2.39-2.37 (1H, m); HRMS (FAB, 3-NBA matrix)
calcd for C31H29O5 (M + 1) 481.2015, found 481.2025.
4 bis-toluoyl ester (â-epimer, 41% isolated yield): 1H NMR (CDCl3,
ppm) δ 8.06 (2H, d, J ) 8.0), 8.02 (2H, d, J ) 8.0), 7.91-7.77 (4H,
m), 7.57-7.48 (3H, m), 7.32 (2H, d, J ) 8.0), 7.23 (2H, d, J ) 8.0),
5.70 (1H, d, J ) 5.7), 5.46 (1H, dd, J ) 3.2, 10.9), 4.77-4.75 (2H,
m), 4.66-4.65 (1H, m), 2.66 (1H, dd, J ) 5.2, 13.8), 2.48 (3H, s),
2.42 (3H, s); HRMS (FAB, 3-NBA matrix) calcd for C31H29O5 (M +
1) 481.2043, found 481.2015.
5 bis-toluoyl ester (â-epimer, 54% isolated yield): 1H NMR (CDCl3,
ppm) δ 8.02 (4H, 2 × d, J ) 8.0), 7.35-7.23 (5H, m), 6.92 (1H, s),
5.62 (1H, d, J ) 5.6), 5.42 (1H, dd, J ) 7.0, 10.8), 4.78 (1H, dd, J )
3.8, 11.8), 4.70 (1H, dd, J ) 3.8, 11.8), 4.55 (1H, m), 2.56 (1H, dd, J
) 5.0, 14.0), 2.43 (3H, s), 2.46 (3H, s), 2.23 (1H, m); HRMS (FAB,
3-NBA matrix) calcd for C30H32O5 472.2250, found 472.2234.
General Procedure for Deprotection of 1′,2′-dideoxy-1′-aryl-3′,5′-
di-O-toluoyl-â-D-ribofuranoses. To a solution of 1 bis-toluoyl ester
(360 mg, 0.65 mmol) in methanol (5 mL) was added NaOMe (in
methanol, 25%, 0.5 mL, 3 equiv). The reaction mixture was stirred
for 4-6 h. Solid ammonium chloride was added until the pH was 8.
The mixture was then poured into water and extracted with EtOAc (3
× 15 mL). The combined organic layers were dried over anhydrous
MgSO4 and evaporated. Flash column chromatography (eluent EtOAc)
of the crude mixture gave 165 mg of nucleoside 1 (â-epimer, 78%):
1H NMR (CDCl3, ppm) δ 8.35 (1H, d, J ) 8.0), 8.31-8.14 (4H, m),
8.08-8.02 (3H, m), 6.25 (1H, dd, J ) 5.6, 10.4), 4.62 (1H, m), 4.28
(1H, m), 4.02-3.98 (2H, m), 2.64 (1H, ddd, J ) 2.0, 2.6, 13.4), 2.02
(2H, broad s, 2 × OH); HRMS (FAB, 3-NBA matrix) calcd for
C23H20O3 318.1256, found 318.1251.
Nucleoside 2 (â-epimer, 74%): 1H NMR (CDCl3, ppm) δ 8.78 (1H,
d, J ) 8.0), 8.68 (1H, d, J ) 8.0), 8.12 (1H, d, J ) 8.0), 7.90 (2H, m),
7.77-7.62 (4H, m), 5.95 (1H, dd, J ) 5.6, 10.4), 4.59 (1H, m), 4.22
(1H, m), 4.0 (1H, dd, J ) 6.4, 13.2), 3.95 (1H, dd, J ) 6.2, 13.4), 2.62
(1H, ddd, J ) 2.0, 5.2, 13.4), 2.25 (1H, m), 1.6 (2H, broad s, 2 ×
OH); HRMS (FAB, 3-NBA matrix) calcd for C19H18O3 294.1256, found
294.1250.
4a bis-toluoyl ester (R-epimer, 31% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.02 (2H, d, J ) 8.0), 7.92-7.97 (4H, m), 7.83 (2H,
d, J ) 8.0), 7.52-7.60 (3H, m), 7.32 (2H, d, J ) 8.0), 7.05 (2H, d, J
) 8.0), 5.72 (1H, m), 5.62 (1H, dd, J ) 8.2, 6.0), 4.85 (1H, m), 4.76-
4.65 (2H, m), 3.12-3.02 (1H, m), 2.52-2.45 (1H, m), 2.42 (3H, s),
2.38 (3H, s); HRMS (FAB, 3-NBA matrix) calcd for C31H29O5 (M +
1) 481.2043, found 481.2015.
5a bis-toluoyl ester (R-epimer, 13% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.02 (2H, d, J ) 8.0), 7.86 (1H, d, J ) 8.0), 7.45 (1H,
s), 7.23-7.28 (5H, m), 6.95 (1H, s), 5.69 (1H, br s), 5.54 (1H, dd, J )
8.0, 6.0), 4.81 (1H, br s), 4.69-4.56 (2H, m), 3.07-2.98 (1H, m), 2.43
(6H, s), 1.35 (3H, s); HRMS (FAB, 3-NBA matrix) calcd for C30H33O5
(M + 1) 472.2250, found 472.2234.
6a bis-toluoyl ester (R-epimer, 16% isolated yield): 1H NMR
(CDCl3, ppm) δ 8.0 (2H, d, J ) 8.0), 7.72 (2H, d, J ) 8.0), 7.43 (1H,
t, J ) 8.5), 7.27 (2H, d, J ) 8.0), 7.19 (2H, d, J ) 8.0), 6.76 (1H, d,
J ) 8.0), 5.61 (1H, br s), 5.57 (1H, dd, J ) 8.0, 6.0), 4.74 (1H, br s),
4.57 (2H, t, J ) 5.0), 3.02-2.93 (1H, m), 2.43 (3H, S), 2.23 (3H, s);
HRMS (FAB, 3-NBA matrix) calcd for C28H26F2O5Na 503.1646, found
503.1636.
Procedure For Epimerization of 1′,2′-Dideoxy-1′r-aryl-3′,5′-di-
O-toluoyl-D-ribofuranoses and Isolation of â-Epimers. To a solution
of 6a bis-toluoyl ester (780 mg, 1.62 mmol) in toluene (50 mL) were
added a catalytic amount of benzenesulfonic acid (∼10%), 1 drop of
concentrated H2SO4, and 2-4 drops of H2O. The reaction mixture
was refluxed under vigorous stirring for 4-6 h. The mixture was then
poured into 5% aqueous NaHCO3 (50 mL) and extracted with EtOAc
(3 × 50 mL). The combined organic layers were dried over anhydrous
MgSO4 and evaporated. Flash column chromatography (eluent solution
8:1 to 2:1 hexanes: EtOAc) of the crude mixture gave 430 mg of 6
Nucleoside 3 (â-epimer, 50%): 1H NMR (CDCl3, ppm) δ 8.06 (1H,
d, J ) 8.0), 7.88 (1H, d, J ) 8.0), 7.80 (1H, d, J ) 8.0), 7.66 (1H, d,
J ) 8.0), 7.55-7.46 (3H, m), 5.92 (1H, dd, J ) 5.2, 10.0), 4.52 (1H,
m), 4.15 (1H, m), 3.92-3.86 (2H, m), 2.54 (1H, dd, J ) 5.6, 13.3),
2.18 (1H, m), 2.02 (2H, broad s, 2 × OH); HRMS (FAB, 3-NBA
matrix) calcd for C15H16O3 244.1099, found 244.1105.
1
Nucleoside 4 (â-epimer, 68%): H NMR (CDCl3, ppm) δ 7.85-
7.80 (4H, m), 7.50-7.42 (3H, m), 5.35 (1H, dd, J ) 5.6, 10.2), 4.43
(1H, m), 4.06 (1H, m), 3.77 (2H, m), 2.6 (2H, broad s, 2 × OH), 2.33
(1H, ddd, J ) 2.0, 5.6, 13.4), 2.02 (1H, m); HRMS (FAB, 3-NBA
matrix) calcd for C15H16O3 244.1099, found 244.1110.
1
bis-toluoyl ester (â-epimer, 46% isolated yield): H NMR (CDCl3,
ppm) δ 8.0 (4H, 2 × d, J ) 8.0), 7.35-7.25 (5H, m), 6.76 (1H, t, J )
10.0), 5.64 (1H, d, J ) 5.2), 5.46 (1H, dd, J ) 10.2, 4.6), 4.78 (1H,
dd, J ) 3.8, 11.8), 4.63 (1H, dd, J ) 3.8, 11.8), 4.54 (1H, m), 2.64
(1H, dd, J ) 5.2, 11.8), 2.43 (3H, s), 2.46 (3H, s), 2.23 (1H, m); HRMS
(FAB, 3-NBA matrix) calcd for C28H26F2O5 481.1827, found 481.1853.
1 bis-toluoyl ester (â-epimer, 38% isolated yield): 1H NMR (CDCl3,
ppm) δ 8.36 (1H, d, J ) 7.9), 8.31 (1H, d, J ) 7.9), 8.20-8.17 (3H,
m), 8.13-8.05 (6H, m), 8.02 (2H, d, J ) 8.0), 7.37 (2H, d, J ) 8.0),
7.26 (2H, d, J ) 8.0), 6.34 (1H, dd, J ) 7.2, 10.8), 5.78 (1H, d, J )
5.4), 4.84-4.88 (2H, m), 4.78-4.76 (1H, m), 2.94 (1H, dd, J ) 5.0,
13.9), 2.50 (3H, s), 2.46 (1H, m), 2.40 (3H, s); HRMS (FAB, 3-NBA
matrix) calcd for C37H31O5 (M + 1) 554.2093, found 554.2069.
2 bis-toluoyl ester (â-epimer, 28% isolated yield): 1H NMR (CDCl3,
ppm) δ 8.78 (1H, d, J ) 7.9), 8.70 (1H, d, J ) 7.9), 8.13-8.09 (4H,
m), 8.03 (2H, d, J ) 8.0), 7.84 (1H, d, J ) 7.9), 7.77-7.60 (4H, m),
7.36 (2H, d, J ) 8.0), 7.18 (2H, d, J ) 8.0), 6.60 (1H, dd, J ) 7.0,
10.5), 4.90 (1H, dd, J ) 3.8, 11.8), 4,84 (1H, dd, J ) 3.8, 11.8), 2.94
(1H, dd, J ) 5.1, 13.7), 2.49 (3H, s), 2.44-2.41 (1H, m), 2.38 (3H, s);
HRMS (FAB, 3-NBA matrix) calcd for C35H31O5 (M + 1) 531.2172,
found 531.2174.
Nucleoside 5 (â-epimer, 93%): 1H NMR (CDCl3, ppm) δ 7.20 (1H,
s), 6.97 (1H, s), 5.38 (1H, dd, J ) 5.6, 10.4), 4.43 (1H, m), 4.01 (1H,
m), 3.82 (2H, m), 2.32 (3H, s), 2.26 (3H, s), 2.24 (3H, s), 1.99 (1H,
m), 1.90 (2H, broad s, 2 × OH); HRMS (FAB, 3-NBA matrix) calcd
for C14H20O3 237.1491, found 237.1484.
Nucleoside 6 (â-epimer, 89%): 1H NMR (CDCl3, ppm) δ 7.46 (1H,
t, J ) 10), 6.82 (1H, t, J ) 10), 5.31 (1H, dd, J ) 5.6, 10.4), 4.32 (1H,
m), 3.92 (1H, m), 3.68 (2H, m), 2.22 (3H, s), 1.89 (1H, m), 1.78 (2H,
broad s, 2 × OH); HRMS (FAB, 3-NBA matrix) calcd for C12H14F2O3-
Na 267.0809, found 267.0812.
General Procedure for Preparation of 5′-O-tritylated â-C-
Nucleosides. The above-synthesized nucleoside 1 (165 mg, 0.52 mmol)
was coevaporated with dry pyridine (4 mL) twice and dissolved in 5
mL of pyridine and 4 mL of methylene chloride. To the above mixture
were added catalytic amount of DMAP, diisopropylethylamine (0.14
mL, 1.5 equiv) and 4,4′-dimethoxytrityl (DMT) chloride (320 mg, 1.8
equiv). The mixture was stirred at room temperature for 8 h. Hexanes
(5 mL) was added, and the mixture was loaded onto a flash silica gel
column (pre-equilbrated with 5% triethylamine in hexanes) and eluted
(5:1 hexanes:EtOAc to 2:1 hexanes: EtOAc). The product 1 DMT
ether was obtained as a yellowish foam in 64% yield (200 mg, 0.32
3 bis-toluoyl ester (â-epimer, 37% isolated yield): 1H NMR (CDCl3,
ppm) δ 8.09-8.04 (3H, m), 7.97 (1H, d, J ) 8.0), 7.91 (1H, overlapped
d, J ) 6.3, 6.2), 7.88 (1H, d, J ) 8.0), 7.51 (2H, overlapped d, J )
6.7, 6.5), 7.46 (1H, d, J ) 7.9), 7.34 (2H, d, J ) 8.0), 7.22 (2H, d, J
) 8.0), 6.02 (1H, dd, J ) 6.4, 10.7), 5.71(1H, d, J ) 5.9), 4.78-4.78
(2H, m), 4.70-4.71 (1H, m), 2.85 (1H, dd, J ) 5.0, 13.8), 2.48 (3H,
1
mmol): H NMR (CDCl3, ppm) δ 8.34 (2H, overlapped d, J ) 8.0),
8.24-8.02 (7H, m), 7.56 (2H, overlapped d, J ) 8.0), 7.45-7.27 (7H,
m), 6.86 (4H, d, J ) 8.0), 6.52 (1H, d, J ) 6.2), 6.24 (1H, dd, J ) 5.2,
10.4), 4.60 (1H, m), 4.30 (1H, m), 3.81 (6H, s), 3.56 (2H, m), 2.64

7678 J. Am. Chem. Soc., Vol. 118, No. 33, 1996
Ren et al.
(1H, m), 2.30 (1H, m); HRMS (FAB, 3-NBA matrix) calcd for C42H36O5
620.2564, found 620.2563.
5 DMT phosphoramidite (380 mg, 89%): 1H NMR (CDCl3, ppm)
δ 7.54 (2H, overlapped d, J ) 8.0), 7.45-7.40 (4H, m), 7.36 -7.22
(3H, m), 6.98 (1H, s), 6.90-6.82 (4H, m), 5.37 (1H, dd, J ) 5.8, 10),
4.56 (1H, m), 4.22 (1H, m), 3.82 (6H, s), 3.66-3.45 (3H, m), 3.42-
3.38 (2H, m), 2.82 (2H, t, J ) 5.6), 2.48 (1H, m), 2.29, (3H, s), 2.24
(3H, s), 2.18 (3H, s), 1.99 (1H, m), 1.28 (1H, m), 1.20-1.05 (12H,
m); HRMS (FAB, 3-NBA matrix) calcd for C44H55N2O6P 739.3876,
found 739.3870.
6 DMT phosphoramidite (420 mg, 84%): 1H NMR (CDCl3, ppm)
δ 7.51 (2H, m), 7.42-7.22 (9H, m), 6.84 (4H, overlapped d, J ) 8.0),
6.78 (1H, dd, J ) 9.0, 8.6), 5.38 (1H, dd, J ) 2.9, 10), 4.54 (1H, m),
4.22 (1H, m), 3.82 (6H, s), 3.66-3.45 (3H, m), 3.42-3.38 (2H, m),
2.82 (2H, t, J ) 5.6), 2.50 (2H, t, J ) 5.6), 2.05 (1H, m), 1.20-1.05
(12H, m); HRMS (FAB, 3-NBA matrix) calcd for C42H49F2N2O6P
769.3194, found 769.3209.
2 DMT ether (280 mg, 59%): 1H NMR (CDCl3, ppm) δ 8.78 (1H,
d, J ) 8.0), 8.68 (1H, d, J ) 8.0), 8.07 (2H, m), 7.8 (1H, d, J ) 8.0),
7.80-7.24 (12H, m), 6.84 (4H, overlapped d, J ) 8.0), 5.94 (1H, dd,
J ) 5.8, 10), 4.52 (1H, m), 4.22 (1H, m), 3.8 (6H, s), 3.50 (2H, m),
2.61 (1H, ddd, J ) 2.0, 5.2, 13.4), 2.25 (1H, m); HRMS (FAB, 3-NBA
matrix) calcd for C40H36O5 596.2564, found 596.2563.
3 DMT ether (50 mg, 52%): 1H NMR (CDCl3, ppm) δ 8.15 (1H,
d, J ) 8.0), 7.9 (1H, d, J ) 8.0), 7.8 (1H, d, J ) 8.0), 7.65 (1H, d, J
) 8.0), 7.49-7.45 (3H, m), 5.94 (1H, dd, J ) 5.8, 10), 4.53 (1H, m),
4.25 (1H, m), 3.8 (3H, s), 3.42 (2H, m), 3.02 (3H, s), 2.58 (1H, ddd,
J ) 2.0, 5.2, 13.4), 2.18 (1H, m); HRMS (FAB, 3-NBA matrix) calcd
for C36H34O5 546.2407, found 546.2406.
4 DMT ether (200 mg, 66%): 1H NMR (CDCl3, ppm) δ 7.83-
7.94 (4H, m), 7.56-7.27 (11H, m), 6.87 (4H, overlapped d, J ) 8.0),
5.41 (1H, dd, J ) 5.8, 10), 4.52 (1H, m), 3.82 (6H, s), 3.42 (2H, m),
2.38 (1H, dd, J ) 5.4, 13.4), 2.21 (1H, m).
Oligonucleotide Synthesis. DNA oligonucleotides were synthesized
on an Applied Biosystems 392 synthesizer using standard â-cyanoethyl
phosphoramidite chemistry but with extended (10 min) coupling cycles
for the nonnatural residues. Stepwise coupling yields for the nonnatural
residues were all greater than 95% as determined by trityl cation
monitoring. Oligomers were purified by preparative 20% denaturing
polyacrylamide gel electrophoresis and isolated by the crush and soak
method and were quantitated by absorbance at 260 nm. Molar
extinction coefficients were calculated by the nearest neighbor method.
Values for oligonucleotides containing nonnatural residues were
estimated in the following way: each of the new nucleosides was
measured for its extinction coefficient at 260 nm. The molar extinction
coefficients for 2 and 3 were found to be 8990 and 154, respectively,
and these values were added to the value for the core sequence
dCGCGCG. For 1 in DNA we measured the absorbance at 350 nm
and subtracted 0.48 of this value from the total absorbance at 260 nm
to get the absorbance of the core DNA alone. Oligodeoxynucleotides
were obtained after purification as the sodium salt. Intact incorporation
of residues 1-3 was confirmed by synthesis of short oligomers of
sequence T-X-T (where X ) 1-3); proton NMR (400 MHz) indicated
the presence of the intact structures with the expected integration.
5 DMT ether (311 mg, 92%): 1H NMR (CDCl3, ppm) δ 7.52 (2H,
d, J ) 8.0), 7.43-7.24 (7H, m), 6.94-6.84 (6H, m), 5.34 (1H, dd, J
) 5.8, 9.8), 4.42 (1H, m), 4.18 (1H, m), 3.80 (6H, s), 3.40 (2H, m),
2.64 (1H, m), 2.29 (3H, s), 2.23 (3H, s), 2.18 (3H, s), 2.0 (1H, m);
HRMS (FAB, 3-NBA matrix) calcd for C35H37O5 538.2719, found
538.2690.
6 DMT ether (350 mg, 88%): 1H NMR (CDCl3, ppm) δ 7.46 (1H,
d, J ) 8.0), 7.39-7.24 (5H, m), 6.83 (2H, overlapped d, J ) 8.0),
6.74 (1H, dd, J ) 9.8, 8.6), 5.38 (1H, dd, J ) 5.8, 9.9), 4.42 (1H, m),
4.06 (1H, m), 3.80 (6H, s), 3.38 (2H, m), 2.38 (1H, dd, J ) 5.0, 13.4),
2.2 (1H, m); HRMS (FAB, 3-NBA matrix) calcd for C33H32F2O5
569.2115, found 569.2131.
General Procedure for Preparation of 3′-O-Phosphoramidites.
The 5′-O-tritylated compound 1 DMT ether (200 mg, 0.32 mmol) was
dissolved in 4 mL of dry methylene chloride, and to this were added
diisopropylethylamine (0.22 mL, 1.2 mmol) and 2-cyanoethyl N,N-
diisopropylchlorophosphoramidite (0.11 mL, 0.48 mmol). The reaction
mixture was stirred at room temperature for 4 h. Hexanes (4 mL) was
added, and the mixture was loaded to the flash silica gel column (pre-
equilibrated with 5% triethylamine in hexanes) and eluted. The product
was obtained as an oil, DMT phosphoramidite 1 (210 mg, 81%): 1H
NMR (CDCl3, ppm) δ 8.44-8.33 (2H, m), 8.25-8.00 (7H, m), 7.62-
7.22 (9H, m), 6.92-6.79 (4H, m), 6.28-6.20 (1H unresolved m), 4.69
(1H, m), 4.45 (1H, m), 4.0-3.2 (12H, m), 2.80 (1H, m), 2.69 (2H, t),
2.32 (1H, m), 1.15 (12H, m); HRMS (FAB, 3-NBA matrix) calcd for
C51H54N2O6P (M + H) 821.3722, found 821.3720.
Fluorescence Measurements. Fluorescence spectra were recorded
on a SPEX-Fluorolog-2 series fluorometer. A xenon lamp was used
as the source of radiation. The fluorescence measurements were taken
in the right angle mode using 0.1-0.15 µM DNA solutions in a pH
7.0 buffer (10 mM Na‚PIPES, 10 mM MgCl2, 100 mM NaCl). Five
scans were averaged at 23 °C. The excitation slits were set to 6mm
and the emission slits to 2 mm. All emission spectra were corrected
using a reference dye (rhodamine-B) to compensate for instrument
fluctuations and also by subtraction of data for buffer alone. Excitation
wavelengths of 233, 251, and 341 nm (the absorbance maxima) were
used to excite the compounds containing naphthalene, phenanthrene,
and pyrene, respectively.
2 DMT phosphoramidite (280 mg, 77%): 1H NMR (CDCl3, ppm)
δ 8.78 (1H, d, J ) 8.0), 8.68 (1H, d, J ) 8.0), 8.07 (2H, m), 7.8 (1H,
d, J ) 8.0), 7.80-7.24 (12H, m), 6.84 (4H, overlapped d, J ) 8.0),
5.94 (1H, overlapped dd, J ) 5.8, 10), 4.62 (1H, m), 4.40 (1H, m), 3.9
(2H, m), 3.8 (6H, s), 3.50 (2H, m), 3.83 (3H, s), 3.77 (3H, s), 3.66-
3.45 (3H, m), 3.42-3.38 (2H, m), 2.82 (2H, t, J ) 5.6), 2.52 (3H, t,
J ) 5.6), 2.16 (1H, m), 1.20-1.05 (12H, m); HRMS (FAB, 3-NBA
matrix) calcd for C49H53N2O6PNa 819.3537, found 819.3539.
3 DMT phosphoramidite (48 mg, 50%): 1H NMR (CDCl3, ppm)
δ 8.10 (1H, d, J ) 8.0), 7.9 (1H, d, J ) 8.0), 7.8 (1H, d, J ) 8.0),
7.52-7.24 (9H, m), 6.82 (4H, overlapped d, J ) 8.0), 5.91 (1H,
overlapped dd, 2 × isomers), 4.6 (1H, m), 4.38 (1H, m), 3.83 (3H, s),
3.77 (3H, s), 3.66-3.45 (3H, m), 3.42-3.38 (2H, m), 2.82 (2H, t, J )
5.6), 2.52 (3H, t, J ) 5.6), 2.16 (1H, m), 1.20-1.05 (12H, m); HRMS
(FAB, 3-NBA matrix) calcd for C45H52N2O6P (M + H) 747.3565, found
747.3563.
Acknowledgment. We thank the National Institutes of
Health (GM52956) and the Army Research Office for support.
We thank Mr. George Rodriguez for obtaining the X-ray
crystallographic data. E.T.K. gratefully acknowledges an Alfred
P. Sloan Foundation Fellowship and a Dreyfus Foundation
Teacher-Scholar Award.
Supporting Information Available: Carbon-13 and phos-
phorus-31 spectral data for all compounds (where appropriate),
fluorescence emission spectra, and a description of crystal-
lographic methods (6 pages). See any current masthead page
for ordering information and Internet access instructions.
4 DMT phosphoramidite (170 mg, 65%): 1H NMR (CDCl3, ppm)
δ 7.91 (1H, s), 7.83-7.78 (4H, m), 7.56-7.27 (11H, m), 6.87 (4H,
overlapped d, J ) 8.0), 5.39 (1H, dd, J ) 5.8, 10), 4.58 (1H, m), 4.30
(1H, m), 3.90 (2H, m), 3.83 (3H, s), 3.77 (3H, s), 3.66-3.45 (3H, m),
3.42-3.38 (2H, m), 2.82 (2H, t, J ) 5.6), 2.52 (3H, t, J ) 5.6), 2.16
(1H, m), 1.20-1.05 (12H, m).
JA9612763