Analysis of the structure and stability of a backbone-modified oligonucleotide: Implications for avoiding product inhibition in catalytic template-directed synthesis

Article abstract of DOI:10.1021/ja972869w

The structural and thermodynamic origins of the destabilization of a backbone-modified DN duplex 1, formed between d(CpGpT(N)TpGpC), containing a single aminoethyl group (-CH₂-CH₂-NH₂⁺-) in place of the phosphod

Full text of DOI:10.1021/ja972869w

J. Am. Chem. Soc. 1998, 120, 3019-3031
3019
Analysis of the Structure and Stability of a Backbone-Modified
Oligonucleotide: Implications for Avoiding Product Inhibition in
Catalytic Template-Directed Synthesis
Peizhi Luo, John C. Leitzel, Zheng-Yun J. Zhan, and David G. Lynn*
Contribution from the Searle Chemistry Laboratory, 5735 Ellis AVenue, Department of Chemistry,
The UniVersity of Chicago, Chicago, Illinois 60637
ReceiVed August 14, 1997
Abstract: The structural and thermodynamic origins of the destabilization of a backbone-modified DNA duplex
1, formed between d(CpGpTNTpGpC), containing a single aminoethyl group (-CH2-CH2-NH2+-) in place
of the phosphodiester (-O-PO2--O-) linkage of the central TT dimer, and d(GpCpApApCpG) are inves-
tigated. Analyses for the corresponding native duplex and two other related structural analogues of duplex 1
have been compared. Duplex 1 shows a cooperative thermal melting transition that is consistent with a two-
state process. At a 2 mM concentration, the melting temperature of duplex 1 is reduced by 17 °C from the
native duplex, and this decrease in stability is further assigned to an unfavorable decrease in enthalpy of 7
kcal mol-1 and a favorable increase in entropy of 15 eu mol-1. NMR structural analysis shows that the
modified duplex 1 still adopts a canonical B-DNA conformation with Watson-Crick base pairing preserved;
however, the CH2 group that replaces the native PO2- group in the modified backbone is flexible and free to
collapse onto a hydrophobic core formed by the base edges and sugar rings of the flanking TT/AA nucleosides
of the duplex. This conformation is significantly different from the maximally solvent-exposed orientation of
the native phosphate in DNA. The entropic origin of the 15 eu mol-1 difference between the native and the
modified duplex 1 is attributed to the hydrophobic interaction between the collapsed ethylamine linkage with
the hydrophobic core of duplex 1. This assignment is further supported by a favorable comparison between
the observed change in entropy and the estimated value for the hydrophobic interaction around the modified
region. This estimated value is based on recent experimental measurement of the hydrophobic interaction
between aliphatic groups and nucleic acids as well as ethylamine solvent transfer data. The overall decrease
in the stability of duplex 1 results from a decrease in base stacking and hydrogen-bonding interactions between
the base pairs. A model is, therefore, proposed to explain how the change in the local backbone conformation
could disrupt the long-range cooperativity of DNA duplex formation upon backbone modifications. These
studies provide an approach for identifying the factors that control the stability of the nucleic acid duplex
structures containing backbone modifications, with direct implication for designing antisense oligonucleotides
and template-directed reactions containing non-native phosphodiester linkages.
Introduction
with structural variation of the backbone linkage. Product
inhibition severely limits template-directed reactions,4 and this
insight into the control of duplex stability has recently made it
possible to design reactions that catalytically translate the infor-
mation encoded in DNA into different polymer backbones.5
This catalytic reaction involves the template-directed reductive
ligation of an imine to a product, d(CpGpTNTpGpC), containing
a single aminoethyl group (-CH2-CH2-NH2+-) replacing the
normal TT phosphodiester linkage. The catalytic cycle exploited
the observation that the duplex formed between the backbone-
modified product and its complementary d(GpCpApApCpG)
template, 1, is substantially destabilized relative to the substrate
Zamecnik and Stephenson1 first showed that synthetic oli-
gonucleotides could be used to block gene expression by binding
specifically with complementary sequences of target mRNAs.
Over the past decade the efforts to modify native phosphodiester
linkages to improve nuclease resistance and membrane perme-
ation of naturally occurring oligonucleotides for therapeutic ap-
plications have been extensive.2 These neutral backbone-modi-
fied oligonucleotides generally show attenuated binding affinity
to their complementary RNA strand relative to the corresponding
nucleic acid.2e,f,3 These studies have therefore clearly demon-
strated that it is possible to tune nucleic acid duplex stability
(3) Egli, M. Angew. Chem., Int. Ed. Engl. 1996, 35, 1894.
(4) (a) Naylor, R.; Gilham, P. T. Biochemistry 1966, 5, 2722. (b) Orgel,
L. E. Nature 1992, 358, 203. (c) Hong, J.-I.; Feng, Q.; Rotello, V.; Rebek,
J. Science 1992, 255, 848. (d) Li, T.; Nicolaou, K. C. Nature 1994, 369,
218. (e) Ertem, G.; Ferris, J. P. Nature 1996, 379, 238. (f) Sievers, D.; von
Kiedrowski, G. Nature 1994, 369, 221.
(5) (a) Goodwin, J. T.; Lynn, D. G. J. Am. Chem. Soc. 1992, 114, 9197.
(b) Goodwin, J. T.; Luo, P. Z.; Leitzel, J. C.; Lynn, D. G. InSelf-Production
of Supramolecular Structures: From Synthetic Structures to Models of
Minimal LiVing Systems, Fleischaker, G. R., Colonna, S., Luisi, P. L., Eds.;
Kluwer Academic Publishers: Dordrecht, The Netherlands, 1994; pp 99-
104. (c) Zhan, Z.-Y. J.; Lynn, D. G. J. Am. Chem. Chem. Soc. 1997, 119,
12420-12421.
* Voice mail and fax: (312) 702-7063. E-mail: d-lynn@uchicago.edu.
(1) (a) Zamecnik, P. C.; Stephenson, M. L. Proc. Natl. Acad. Sci. U.S.A.
1978, 75, 280. (b) Stephenson, M. L.; Zamecnik, P. C. Proc. Natl. Acad.
Sci. U.S.A. 1978, 75, 285.
(2) (a) De Mesmaeker, A.; Waldner, A.; Lebreton, J.; Hoffmann, P.;
Fritsch, V.; Wolf, R.; Miller, P. S. In Oligonucleotides: Antisense Inhibitors
of Gene Expression; Cohen, J. S., ed.; CRC Press: Boca Raton, FL, 1989,
pp 79-93. (b)Uhlmann, E.; Peyman, A. Chem. ReV. 1990,90, 544. (c)
Carruthers, M. H. Acc. Chem. Res. 1991, 24, 278. (d) Varma, R. S. Synlett
1993, 621. (e) De Mesmaeker, A.; Altmann, K.-H.; Waldner, A.; Wende-
born, S. Curr. Opin. Struct. Biol. 1995, 5, 343. (f) De Mesmaeker, A.; Ha¨ner,
R.; Martin, P.; Moser, H. E. Acc. Chem. Res. 1995, 28, 366.
S0002-7863(97)02869-2 CCC: $15.00 © 1998 American Chemical Society
Published on Web 03/21/1998

3020 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Scheme 1. Synthesis of the Modified DNA Dimersa
Luo et al.
by a change in the duplex structure, the modified single strand,
or both? Although the comparisons between a backbone-
modified duplex with its native analogue have generally assumed
that only the duplex stability, rather than the single strand, has
been changed upon backbone modification, the experimental
basis for such an implicit assumption is not yet clear. (4) What
is the overall structure in the modified duplex as compared to
its native analogue? Is it still nativelike, retaining the general
features characteristic of the native duplex, or is the structure
significantly altered? If the overall structure of the native duplex
is preserved upon backbone modification, is it possible to detect
structural changes around the modified region? (5) Finally, can
any global and local structural changes be related to changes
in ∆S and ∆H values?
As a specific example to illustrate the points listed above,
we have studied the short DNA duplex, 1, formed between
d(CpGpTNTpGpC) and its complementary d(GpCpApApCpG)
strand. A single phosphodiester is replaced by the aminoethyl
group,5 and the approach taken is similar in spirit to studies of
point mutations on protein stability. A comparison is made of
the melting curves obtained for duplex 1, and its analogues,
with the native DNA sequence. Both UV and NMR analyses,
under differing salt and sample concentration conditions, are
employed, and the thermal unfolding curves are dissected into
changes in enthalpy and entropy. For the native DNA duplex,
quantitative comparisons are made among values derived from
UV and NMR measurements as well as from calculated values
from the literature. NMR measurements show that the structure
of duplex 1 is nativelike and maintains the Watson-Crick base
pairs as exhibited by similar cross-peak patterns characteristic
of the native duplex. However, significant perturbation in the
modified TT region is detected using the NOE cross-peaks
between the two methylenes of the aminoethyl linkage and the
flanking sugar protons. This result allows for a rationalization
of the changes in enthalpy and entropy of the backbone-modified
duplex to the observed local and global structural changes. These
results have allowed a model to be proposed which attempts to
explain the observed destabilization of backbone-modified
oligonucleotides with neutral linkers. One important implication
underlying this model, as well as others proposed recently,6 is
the role that the polyanionic phosphate backbone plays in
conferring duplex stability. The comprehensive approach that
we have taken should be useful for analyzing other backbone-
modified nucleic acids and allow the factors that control duplex
stability to be better defined.
a Dimers shown were subsequently converted to phosphoramidates
and incorporated into DNA oligomers.
imine. The difference in the thermal melting temperature was
as high as 15 °C,5c but the structural and energetic origins of
the altered stability have not been defined.2-5
In contrast with the explosive growth in synthesizing backbone-
modified oligonucleotides, only limited progress has been made
in determining the structural and energetic factors which control
the stability of the duplexes.3 De Mesmaeker et al.2e,f have
reviewed the chemical modifications and the stability of the
second generation antisense oligonucleotides that contain no
phosphorus atom in the modified linkages. The hybridization
stability of these duplexes, as measured by the thermal melting
temperature, is rationalized in terms of the change in charge,
conformational rigidity, structural preorganization, hydrophobic-
ity, and steric hindrance differences (see Figures 5 and 6 and
Table 1 in ref 2f). In principle, structural studies by NMR and
X-ray can provide important insight into the origins of the
changes in enthalpy and entropy resulting from backbone
modifications. However, the thermodynamic analyses depend
heavily on the thermal melting temperature,2d-f which involves
a complex interplay between the enthalpic and entropic com-
ponents of the duplexes and/or the single strands.3 We saw
the following issues relating to the structure and stability of
the backbone-modified oligonucleotides as critical to any such
analysis: (1) Since the formation of a nucleic acid duplex is
highly cooperative, could a chemical modification in the middle
of one strand change the coopertivity of duplex formation,
deviating, for example, from the usual two-state thermal
transition? (2) Can the thermodynamic stability be dissected
into changes in enthalpy and entropy of the modified duplex
relative to its native analogue? (3) Is the difference between
the stability of the modified duplex and the native duplex caused
Results
Synthesis of Modified Nucleosides. The hexameric tem-
plate, d(GpCpApApCpG), was designed to be small, palindro-
mic, and non-self-complementary, and to have a high G/C
content to enhance association.3 The central template adenosine
residues simplified the synthetic chemistry to modifications of
thymidine (Scheme 1). The ease of radical allylation at C-3′
(60% recrystallized yield over three steps) set the stereochemical
position of the aldehyde that was ultimately generated by
oxidative cleavage.3a Only the generation of the 5′-amine was
required on the second thymidine derivative. For bulk systhesis,
reductive amination of the 3′-carboxyaldehyde 5 with the 5′-
amine 7 gave the amine-linked dimer 8 in >95% purified
yield.3a,7 Acetylation of 8 gave cleanly the N-acyl dimer 9, and
further oxidation of 5 to 6 and condensation with 7 gave the
amide dimer 10. Each of these dimers was converted into the
(6) Richert, C.; Roughton, A. L.; Benner, S. A. J. Am. Chem. Soc. 1996,
118. 4518.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3021
Scheme 2. Modified DNA Duplex with the TT Dimer
Region As Indicateda
Figure 1. ln(CT/4) vs 1/Tm for the native duplex in 500 mM NaCl, 10
mM NaH2PO4, 2 mM NaN3, pH 6 buffer.
to eq 2a,8a,b or alternatively from a nonlinear least-squares fit
to eq 2b, where Cd is the duplex concentration. The nonlinear
ln(CT/4) ) (∆H/R)(1/Tm) - (∆S/R)
Cd ) 2 exp((∆H/R)(1/Tm) - (∆S/R))
(2a)
(2b)
least-squares analysis has the advantage of fitting the thermo-
dynamic functions and baseline parameters simultaneously to
the primary data. This approach avoids the empirical assump-
tion of the linear baseline regions at the low- and high-tem-
perature limits. Both approaches gave identical values of ∆H
and ∆S (Table 1); however, as expected, the errors are much
larger with the least-squares fit. At a 67% confidence level,
the error was 7 times larger. A similar increase in error was
previously noted by Santoro and Bolen9 in determinations of
the ∆G of protein unfolding.
a Three linkages in the TT dimer are shown: 1, aminoethyl; 2,
N-acetylaminoethyl; 3, amide.
NMR Analyses. The design of the template placed two
adenine bases in the center of the duplex. The two aromatic
methyls of the thymidine bases in the complementary strand,
T3-CH3 and T4-CH3, reside in the center of the duplex and flank
the site of the structural change that is under consideration. The
chemical shifts of these methyl protons in the native and the
modified duplex 1 show cooperative thermal transitions with
temperature only when measured in the presence of the template
strand (Figure 2a). In addition to being able to monitor the
center of the duplex, these NMR analyses have several other
advantages. First, the higher concentrations of the nucleic acids
used in the NMR experiments move the Tm for these short
strands above room temperature in H2O. Second, the baselines
for the duplex, δd, and the single strands, δs, are well defined
(Figure 2a). Finally, the exchange rate between the two states
is fast on the NMR time scale. When the observed chemical
shift, δobs, is modeled as a function of these two states, eq 3,10
the resulting van’t Hoff plots from eq 4, Figure 2b, give ∆H
appropriate phosphoramidite for incorporation into the hexa-
nucleotide via solid-phase synthesis. This approach permitted
the ready synthesis of 1 as well as the modified hexamers 2
and 3 shown in Scheme 2.
Thermodynamic Measurements. Strand association was
analyzed with both UV and NMR spectroscopies, and different
methods for extracting the thermodynamic values were explored.
In all cases, attempts were made to fit the data to a simple
equilibrium between the DNA duplex and the single strands.
In this two-state model, the equilibrium constant, K, for
association of non-self-complementary strands is expressed by
eq 1, where CT is the total concentration of oligonucleotide
strands and f is the fraction of strands in the duplex form.8a,b
K ) 2f/[(1 - f)2CT]
(1)
UV Analyses. The melting temperature, Tm, of the native
duplex was determined via the hypochromic shift of the base
transition at five different concentrations in 500 mM NaCl. The
∆H and ∆S values were derived from the slope and intercept,
respectively, of a plot of ln(CT/4) vs 1/Tm in Figure 1, according
δobs ) (1 - f)δs+ fδd
(3)
(4)
ln K ) -(∆H/R)(1/T) + (∆S/R)
(7) Caulfield, T. J.; Prasad, C. V. C.; Prouty, C. P.; Saha, A. K.; Sardaro,
M. P.; Schairer, W. C.; Yawman, A.; Upson, D. A., & Kruse, L. I. Bioorg.
Med. Chem. Lett. 1993, 3, 2771.
(8) (a) Borer, P. N.; Dengler, B.; Tinoco Jr, I.; Uhlenbeck, O. C. J. Mol.
Biol. 1974, 86, 843. (b) Albergo, D. D., Marky, L. A., Breslauer, K. J.;
Turner, D. H. Biochemistry 1981, 20, 1409. (c) Albergo, D. D.; Turner, D.
H. Biochemistry 1981, 20, 1413.
and ∆S values for both the T3-CH3 and T4-CH3 groups that
(9) Santoro, M. M.; Bolen, D. W. Biochemistry 1988, 27, 8063.
(10) (a) Tran-Dinh, S.; Neumann, J.-M.; Huynh-Dinh, T.; Genissel, B.;
Igolen, J.; Simonnot, G. Eur. J. Biochem. 1982, 124, 415. (b) Pieter, J.;
Mellema, J.-R.; Van Den Elst, H.; Van Der Marel, G.; Van Boom, J.; Altona,
C. Biopolymers 1989, 28, 741.

3022 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Luo et al.
Table 1. Thermodynamic Parameters of Thermal Unfolding from the Native Duplex and the Backbone-Modified Duplex 1 to Coiled Strandsa
predictedb
(native)
UV meltc
(native)
T3CH3
(native)
T3CH3
(duplex 1)
T4CH3
(native)
T4CH3
(duplex 1)
d
d
d
d
thermodynamic parameters
-∆H (kcal mol-1
)
44.4
43.6 ((1.8) 47.3 ((1.3) 40.2 ((0.9) 46.1 ((2.5) 39.1 ((1.7)
7.1 7.0
120.2 ((5.8) 136.9 ((4.0) 121.4 ((3.0) 131.9 ((8.2) 117.1 ((5.7)
15.5 14.8
difference in -∆H between native and modified duplex 1
-∆S (eu mol-1
)
126.1
difference in -∆S between native and modified duplex 1
a The results are derived from the van’t Hoff plots in Figures 1b and 2b. b Predicted values were calculated using the nearest-neighbor thermodynamic
data11a at 1 M NaCl, pH 7. The 126.1 eu mol-1 was derived from the predicted ∆H and ∆G using the nearest-neighbor approximation. c UV melting
at different DNA concentrations in 500 mM NaCl, 10 mM NaH2PO4, 2 mM NaN3, pH 6. d The change in chemical shift with temperature as
monitored by T3 and T4 methyl groups at 2 mM total oligonucleotide concentration in 100 mM NaCl, 10 mM NaH2PO4, 2 mM NaN3, pH 6.
Justifying the Analyses. The calculated ∆H value for this
sequence, using the nearest-neighbor approximation of Breslauer
et al.,11a is -44.4 kcal mol-1. These estimated ∆H values for
short oligonucleotide duplexes give excellent agreement with
calorimetry measurements.8a,b,11a The ratio of the UV-deter-
mined ∆H to this calculated value is 0.98, justifying the use of
the two-state assumption for duplex melting.8b
Although UV analyses have been used extensively for
measuring the global unfolding of DNA duplexes, the ∆δ of
the thymidine methyl resonances may be diagnostic of a more
localized perturbation. These methyls, which reside in the center
of the duplex, yield ∆H values that are of the same order, -47.3
kcal mol-1 from T3-CH3 and -46.1 kcal mol-1 from T4-CH3,
but slightly more negative than those from calculations (Table
1). The more negative values may result from the cooperative
transition monitored only at the center of the duplex, free from
the fraying of base pairs at the ends, or from the isotope effect
in D2O.8b
The ∆S value was also calculated from the predicted ∆G and
∆H of Breslauer et al.11a to be -126 eu mol-1 in 1 M NaCl.
By UV in 500 mM NaCl, ∆S ) -120 ( 5.8 eu mol-1; by
NMR in 100 mM NaCl, ∆S ) -137 ( 4.0 and -132 ( 8.2 eu
mol-1 from T3-CH3 and T4-CH3, respectively. These differ-
ences in ∆S are accounted for quantitatively by the difference
in salt concentrations used in the experiments. According to
the polyelectrolyte theory of nucleic acids,12 the salt effect is
primarily of entropic origin that arises from counterion con-
densation on the phosphate backbones of the duplex, and this
assignment is supported by precise scanning microcalorimetry
measurements.13 By substituting the concentration of NaCl into
eq 6, where C is the molar salt concentration and R is the gas
constant, the change in entropy due to the salt effect on duplex
Figure 2. (a, top) Chemical shift change of the T3 methyl groups in
the native (b) and the modified duplex 1 (O) as a function of
temperature. The curves through data points are fitted to eq 5 using
the nonlinear least-squares method. Lines through each melting profile
are fit with either floating or fixed baseline parameters in δs and δd.
The difference in Tm between native and modified duplex 1 is ca. 17
°C in 2 mM solutions in 9:1 (v/v) H2O-D2O, 100 mM NaCl, 10 mM
NaH2PO4, 2 mM NaN3, pH 6 buffer. (b, bottom) van’t Hoff plots of
the thermal unfolding profiles for the native ([) and modified duplex
1 (]).
∆S ) (-1.11 + 0.36 log C)R eu mol-1 of phosphate (6)
melting is predicted as 2.9, 2.4, and 2.2 eu mol-1 of phosphate
at 100 mM, 500 mM, and 1 M NaCl, respectively. If the NaCl
concentration is changed from 100 mM to 1 M, the increase in
∆S upon duplex formation is predicted to be 0.7 eu mol-1 of
phosphate or 7 eu mol-1 of a hexamer duplex containing 10
phosphate groups. The predicted increase of 7 eu mol-1 in ∆S
upon duplex formation going from 100 mM to 1 M NaCl lies
within the observed difference range of 11 eu mol-1 from T3-
CH3 and 5.8 eu mol-1 from T4-CH3 by NMR and the calculated
∆S at 1 M NaCl (Table 1). The values measured by UV differ
slightly from the predicted values. This general agreement
among the predicted and UV- and NMR-determined ∆H and
are in close agreement with those determined from UV analysis
(Table 1). By combining eqs 1 and 3, the change in chemical
shift can be fit to eq 5, where T is the temperature in Kelvin, R
δobs ) δs + (δd - δs)(1 + c - ((1 + c)2 - 1)1/2)
c ) 1/(CTK) ) (1/CT) exp(∆H/(RT) - ∆S/R)
(5)
is the gas constant, and δs, δd, ∆H, and ∆S are the parameters
in the nonlinear least-squares analysis. By this method, the T3-
CH3 gives ∆H ) -45.5 kcal mol-1 and ∆S ) -131 eu mol-1
,
(11) (a) Breslauer, K. J.; Frank, R.; Blo¨cker, H.; Marky, L. A. Proc.
Natl. Acad. Sci. U.S.A. 1986, 83, 3746. (b) Vesnaver, G.; Breslauer, K. J.
Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 3569.
in general agreement with the direct van’t Hoff analysis, but
again the error is 8-9 times greater. Holding the baseline
parameters of δd and δs at the values used in the van’t Hoff
analysis, this error is reduced 3-fold, consistent with the floating
baselines being largely responsible for the errors.9
(12) (a) Manning, G. S. Q. ReV. Biophys. 1978, 11, 179. (b) Record, Jr.
M. T.; Mazur, S. J.; Melancon, P.; Roe, J.-H.; Shaner, S. L.; Unger, L.
Annu. ReV. Biochem. 1981, 50, 997.
(13) Privalov, P. L.; Filimonov, V. V. J. Mol. Biol. 1978, 122, 447.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3023
∆S values for the native duplex suggests that the errors
associated with these measurements are too small to account
for the large difference in the enthalpy and entropy between
the native and the modified duplex 1 (see the discussion below).
The Modified Duplex. By UV analyses, the modified duplex
1 shows a cooperative melting transition at both 10 and 100
µM; however, the melting temperature is reduced ∼17 °C
relative to the native duplex. At 2 mM, the NMR-detected
thermal transitions of both duplex 1 and the native duplex are
more cooperative, and the Tm of duplex 1 is 17 °C lower than
that of the native duplex (Figure 2a). Each thymidine methyl
reported the thermal transition, and more significantly, the
difference in the thermodynamic values determined from T3-
CH3 and T4-CH3 for the native and the modified duplex 1 were
the same (Table 1). By this analysis, the overall decrease in
stability of the modified duplex is seen to originate from two
opposing factors: (i) a large decrease in enthalpy, 7 kcal mol-1
and (ii) a favorable change in entropy of 15 eu mol-1
,
.
The positive charge in the backbone is expected to contribute
enthalpic stabilization to duplex 1 due to the more favorable
Coulombic interaction along the oligonucleotide backbone.14a
The entropic contribution resulting from the counterion con-
densation on the modified backbone is expected to be unfavor-
able relative to its native analogue. Therefore, the observed
changes in both ∆S and ∆H values between native and modified
duplex 1 are in the opposite direction from those expected by
electrostatic considerations. The N-acyl- and amide-linked
hexamers of duplexes 2 and 3 were prepared as shown in
Scheme 1 to remove the positive charge in the backbone. At a
5 µM duplex concentration in 1 M NaCl, the neutral backbone
of the N-acyl duplex 2 showed no cooperative thermal melting
transition. The N-acyl group in duplex 2 alters both the steric
bulk of the linkage and the hybridization of the N atom;
however, duplex stability was tolerant to substituents on the
nitrogen atom at the corresponding position of an amide linkage
when the group was changed from H to methyl to isopropyl.2f,14b
Given the increased conformational flexibility of the ethylamine
linkage of duplex 1 in comparison with the corresponding amide
linkage in 3 (amide analogues 14a to 14c in Table 1 of ref 2f),
the acyl substituent is expected to be well tolerated in 2. Amide
duplex 3 was prepared also, and its Tm of 14 °C, only slightly
less than the Tm (18 °C) of the native DNA duplex, was
consistent with the observation made by De Mesmaeker et
al.2f,14b These data suggest that the positive charge is not the
major factor contributing to duplex stability, and other factors
should be sought to explain the overall destabilization of duplex
1 relative to the native structure.
Figure 3. Temperature dependence of exchangeable imino proton
resonances for (a, top) the native and (b, bottom). the modified duplex
1. The exchange-out temperature of the hydrogen-bonded imino protons
is higher by ca. 20 °C in the native duplex relative to the modified
duplex 1. The same samples as in Figure 2 are used, and the results
are consistent with a ca. 17 °C difference in Tm between both duplexes
shown in Figure 2a.
Structural Analyses. Figure 3 shows the resonances of the
exchangeable imino protons in 9:1 H2O-D2O for both the native
DNA and the modified duplex 1. All six of the expected protons
are present in both structures, consistent with their participation
in hydrogen bonding with the complementary strand. As the
temperature is increased, the protons on the modified strand
begin to exchange with solvent approximately 20 °C lower than
do those in the native DNA. In both cases, fraying of the ends
is followed by an apparent cooperative disruption of the internal
base pairs.
signal/noise ratio by accumulating many scans at the specific
resonances (see the Materials and Methods). Figure 4 shows
that irradiation of the imino proton of T4 results in two sharp
H2 NOEs; a strong H2 NOE with its base-pairing partner A3
and a weaker H2 NOE with A4. Only one strong H2 NOE
with A4 is observed when the imino proton of T3 is irradiated
(Figure 4). These are characteristic of the Watson-Crick base-
pairing motif in the B-form DNA duplex for both the native
duplex (Figure 4a) and the modified duplex 1 (Figure 4b).15a
To the extent that the Watson-Crick base-pairing motif is
These protons are assigned using the standard approach that
follows the sequential connectivities involving exchangeable
imino, amino, and adenine-H2 protons by 1D NOE difference
spectra.15 The 1D spectra made it possible to obtain a sufficient
(14) (a) Dempcy, R. O.; Browne, K. A.; Bruice, T. C. J. Am. Chem.
Soc. 1995, 117, 6140. (b) De Mesmaeker, A.; Waldner, A.; Lebreton, J.;
Hoffmann, P.; Fritsch, V.; Wolf, R.; Freier, S. Angew. Chem., Int. Ed. Engl.
1994, 33, 226.
(15) (a) Chou, S. H.; Hare, D. R.; Wemmer, D. E.; Reid, B. R.
Biochemistry 1983, 22, 3037. (b) Chou, S. H.; Flynn, P.; Reid, B. R.
Biochemistry 1989, 28, 2435. (c) Weiss, M. A.; Patel, D. J.; Sauer, R. T.;
Karplus, M. Nucleic Acids Res. 1984, 12, 4035.

3024 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Luo et al.
Figure 4. 1D NOE difference spectra showing the imino and AH2
proton connectivities for (a, top) the native and (b, bottom) the modified
1 DNA duplexes in the T3T4/A4A3 base pairs. Irradiation of the imino
proton assigned to T4 results in two sharp H2 NOEs: a strong H2
NOE with its pairing partner A3 and a weaker H2 NOE with A4. Only
one strong H2 NOE with A4 is observed when the imino proton of T3
is irradiated, consistent with a right-handed duplex. The NOE connec-
tivities for imino and adenine H2 protons in the T3T4/A4A3 base pairs
indicate that the Watson-Crick base-pairing motif is preserved in the
backbone-modified DNA duplex 1.
Figure 5. Expanded regions of the NOESY spectrum (D2O, pH 6, 1
°C, 200-ms mixing time) showing (a, top) the sequence-specific
connectivities between nonexchangeable base H8/H6 and sugar H2′/
2′′ protons for the modified DNA duplex 1. The sequential connec-
tivities for the d(CpGpTNTpGpC) or TT strand and its complementary
d(GpCpApApCpG) or AA strand are shown in solid and dashed lines,
respectively, with the cross-peaks designated by the base and its
sequence number. (b, bottom) NOE connectivities of the intranucleotide
sugars H1′ and H3′ with their H2′/2′′ protons in the modified DNA
duplex 1. The reversal in the chemical shift of H2′/2′′ protons in T3
are revealed by a stronger cross-peak of the high-field signal with the
T3-H1′ proton. The T3-H3′ proton is shifted upfield (see Figure 6)
due to the chemical modification in the T3 sugar 3′ position.
preserved in the native duplex, it is similarly maintained in the
backbone-modified duplex 1.
Characterization of the Modified Duplex Conformation
via Nonexchangeable Protons. Figure 5a shows the charac-
teristic NOESY cross-peaks for the sequence-specific connec-
tivities between nonexchangeable base H8/H6 and sugar H2′/
2′′ protons in the modified DNA duplex 1.16 The solid lines
show the sequence-specific assignments of the backbone-
modified TNT strand. Each base proton H8/H6 displays NOEs
with its own and 5′-flanking sugar H2′/2′′ protons and/or 3′-
flanking methyl groups involving thymidine. Starting from the
5′-terminal nucleotide with only two NOEs of the 5′-base proton
with its own sugar H2′/2′′ protons, the sequential connectivities
from base H8/H6 to sugar H2′/2′′ protons were obtained across
the entire strand without disruption. The dashed lines in Figure
5a show the corresponding assignments for the complementary
native DNA strand in the modified duplex 1.
peak intensity of H1′ with H2′′ is stronger than with H2′ because
the H1′-H2′′ distance is usually shorter under all sugar pucker
conformations. For the same reason, the corresponding NOE
cross-peak intensity of the H3′ with H2′′ is weaker than that of
H3′ with H2′. On the basis of these criteria, the chemical shifts
of H2′ and H2′′ protons in the 3′-modified T3 sugar were
assigned unambiguously even though their resonance frequen-
cies are reversed in comparison with the usual order of H2′
and H2′′ (Figure 5), except for the H2′ and H2′′ protons of 3′-
terminal G6 with a free 3′-OH group (Figure 5b). These
assignments are important in defining the orientation of the
methylene groups in the modified backbone with respect to the
The intranucleotide NOEs of H1′ and H3′ with H2′/2′′ are
utilized to distinguish the H2′ from H2′′ protons on the basis
of the variations in NOE cross-peak intensities. The NOE cross-
(16) (a) Scheek, R. M.; Boelens, R.; Russo, N.; van Boom, J. H.; Kaptein,
R. Biochemistry 1984, 23, 1371. (b) Wu¨thrich, K. NMR of Proteins and
Nucleic Acids; Wiley & Sons: New York, 1984.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3025
Table 2. Proton Chemical Shift Assignments for the Modified DNA Duplex 1 at pH 6, 1 °Ca
residue
H6/H8
H5/CH3
5.43
H1′
H2′
H2′′
H3′
H4′
H5′/H5′′
other
G1
C2
A3
A4
C5
G6
8.01
7.51
8.30
8.28
7.30
7.93
5.99
5.48
5.96
6.12
5.59
6.17
2.63
2.13
2.81
2.65
1.89
2.62
2.79
2.40
2.97
2.84
2.30
2.36
4.87
4.90
5.10
5.07
4.80
4.68
4.27
4.18
4.35
4.42
4.16
4.21
3.73/3.73
4.12/4.10
4.15
4.23
4.26
AH2 7.43
AH2 7.74
5.29
4.08
C1
G2
T3
T4
G5
C6
7.73
8.09
7.46
7.53
8.04
7.49
5.92
5.79
6.11
5.86
5.97
5.99
6.22
2.15
2.78
2.27
1.72
2.65
2.22
2.51
2.91
1.86
2.37
2.77
2.22
4.75
5.01
2.78
4.58
5.07
4.53
4.11
4.26
4.05
4.35
4.45
4.27
3.76
4.14/4.06
4.42/4.27
3.71/3.56
1.49
1.72
C6′ 2.02; C7′ 3.39
5.98
4.08
a Referenced to TSP at 0.00 ppm.
ring. In contrast to the corresponding PO2- group that lies on
the outside of the native duplex with maximal solvent acces-
sibility, the C7′ CH2 group in the modified duplex 1 can main-
tain close contact with the hydrophobic duplex interior and the
T3 sugar ring. This orientation is supported by the stronger
NOE cross-peak of T3-H1′-C7′ (H1′-C7′; ca. 2.7 Å) as com-
pared to the T3-H1′-C6′ cross-peak (H1′-C6′; ca. 3.2 Å) and
the NOE cross-peak of C7′ with T3-H2′′ in Figure 6. In com-
parison, the corresponding distances of the O1P and O2P atoms
-
of a PO2 group to the T3-H1′ proton, or H1′(i)-O1P(i + 1)
and H1′(i)-O2P(i + 1), are 5.2-5.5 and 4.8-5.4 Å, respec-
tively, in a canonical B-DNA conformation,17,18 exceeding the
detectable distance limit of the NOE (ca. 4.5-5.0 Å). An NOE
cross-peak between the H1′ proton and a C7′ methylene has
been observed in one other backbone modification (see Figure
9 in ref 17a), although it was suggested that the magnetiza-
tion exchange may have occurred by spin diffusion in this
case. The T3-H1′-C7′ cross-peak is indicative of an altered
backbone orientation in duplex 1 as the presence of this cross-
peak was confirmed in an NOESY spectrum collected at a 50-
ms mixing time (see the Supporting Information), making spin
diffusion unlikely as a major contributor to the observed cross-
peaks.19
Table 3 tabulates proton-proton distances derived from NOE
cross-peaks that are characteristic of the backbone conformation
in the modified region. For comparison, the phosphate oxygen-
proton distances in a native B-form DNA, but with a T3 sugar
pucker in either C2′-endo or C3′-endo forms, are also listed.
The difference between the distances of T3-H1′ to C7′ meth-
ylene of 1 and T3-H1′ to OP* of the native duplex is too large
to be adequately accounted for by changes in the connecting
sugar puckers (see Table 3). These data argue that the modified
backbone of duplex 1 deviates significantly from the backbone
conformation of the canonical DNA ensembles.
Figure 6. Expanded region of the NOESY spectrum (D2O, pH 6, 1
°C, 200-ms mixing time) which highlights the cross-peaks of C7′ and
C6′ methylene protons on the modified linkage with flanking sugar
protons. Notice the stronger C7′-T3-H1′ cross-peak relative to those
of C6′-T3-H1′ and even T3-H1′/H2′′. The cross-peaks are assigned
as the sugar protons and methylene groups are in Scheme 1.
T3 sugar ring. Figure 5b also indicates the chemical shift
overlap for H2′ and H2′′ in C6 as shown by a single strong
cross-peak between H1′ and H3′ with H2′/2′′, respectively.
The base-pairing scheme and sequential connectivities, with
the uniformly much stronger NOEs of an intraresidue base
proton with its own H2′/2′′ protons than the corresponding NOEs
with the 5′-flanking interresidue H2′/2′′ protons across both
strands, establish a canonical right-handed B-DNA conforma-
tion for the backbone-modified DNA duplex 1.16 The chemical
shift assignments of the modified DNA duplex 1 are listed in
Table 2.
NOEs of the Modified Linkage with Flanking Sugar
Protons. The orientation of the phosphodiester linkage relative
to the flanking nucleosides has eluded direct NOE observation
in the DNA duplex due to the lack of protons and severe overlap
of the 5′,5′′ resonances.17 The presence of the methylene units
designated as C6′ and C7′ in the modified linkage of 1 permitted
detailed analysis of the backbone geometry using the corre-
sponding NOEs of these protons with the 5′-flanking sugar
protons. These data are shown in a well-resolved region of the
NOESY spectrum in Figure 6 and are summarized in Table 3
for the NOE cross-peaks of the C6′ and C7′ protons with the
T3-H1′, -H2′′, and -H4′ sugar protons underneath the T3 sugar
Structural Model. A preliminary structural model was
derived by computer simulations with ca. 180 constraints derived
from NMR experiments. A semiquantitative approach was taken
to generate a 3D model using the Biosym software package.20
The initial structure was built and modified from a B-DNA
duplex using the Biopolymer and Builder modules in InsightII.21
The energy minimization and dynamic simulations were carried
out using the Discover module in InsightII with the CVFF force
(18) (a) Chandrasekaran, R.; Birdasall, D. L.; Leslie, A. G. W.; Ratcliff,
R. L. Nature 1980, 283, 743. (b) Dickerson, R. E. J. Mol. Biol. 1983, 166,
419.
(19) Reid, B. R.; Banks, K.; Flynn, P.; Nerdal, W. Biochemistry 1989,
28, 10001.
(17) (a) Gao, X.; Brown, F. K.; Jeffs, P.; Bischofberger, N.; Lin, K.-Y.;
Pipe, A. J.; Noble, S. A. Biochemistry 1992, 31, 6228. (b) Gao, X.; Jeffs,
P. W. J. Biomol. NMR 1994, 4, 17.
(20) Dwyer, T. J.; Geierstanger, B. H.; Mrksich, M.; Dervan, P. B.;
Wemmer, D. E. J. Am. Chem. Soc. 1993, 115, 9900.
(21) Arnott, S.; Hukins, D. W. L. J. Mol. Biol. 1973, 81, 93.

3026 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Luo et al.
Table 3. Comparisons of the Proton-Proton Distance Derived from NOE Cross-Peaks Characteristic of the Backbone Conformation in the
Modified Region with the Corresponding Phosphate Oxygen-Proton Distances in B-Form DNA but with the T3 Sugar Pucker in Either the
C2′-endo or C3′-endo Conformation
i to i + 1a
H1′-OP* b
H2′-OP *
H2′′-OP*
H3′-OP*
H4′-OP*
distances in B-DNAc
4.8-5.5
4.7
H1′-C7′
2.7
3.7-5.3
3.8
2.9-4.8
2.5
H2′′-C7′
3.2
2.4-3.8
3.9
4.7
2.8
H4′-C7′
2.7
distances in twisted B-DNAd
T3 sugar to C7′ methylene
NOE derived distancese
NOE cross-peaks at 50 ms
H2′-C7′
H3′-C7′
∼4.5
>3.5
medium
null
weak
very weak
medium
a i and i + 1 refer to two consecutive residues. b OP* refers to the pseudo atom for two O1P and O2P atoms. c Distance ranges for B-DNA were
taken from Table 4 in ref 17a. d The twisted B-DNA represents the extreme case of the T3 sugar pucker switching from normal C2′-endo to
C3′-endo. e The NOE-derived distances were obtained using the integration of the NOE cross-peak at 50-ms and 200-ms mixing times; the NOE
cross-peak intensities at 50-ms mixing time provided the control on the distance estimates.
Figure 7. Comparison of structures with their solvent-accessible surfaces (dots) in the TT region of (a) the ethylamine-containing strand, and (b)
its native counterpart in B-form DNA. Both strands were cut out of the full duplexes, generated from crystal structure data for (b) and from
molecular dynamics simulations incorporating NMR constraints for (a). The intruding C7′ methylene with the flat solvent-accessible surface in the
modified linkage 1 contrasts markedly with the maximally solvent-accessible phosphate of the phosphodiester linkage.
field parameters. The hydrogen bonds in the Watson-Crick
base pairs were constrained as supported by the imino proton
experiments to prevent the two strands from breaking apart
during simulations. The simulations were started by using the
simulated annealing protocol to satisfy local NOE constraints
in the TT dimer region while the rest of the structure was
constrained. Once the NOE constraints in the TT dimer region
were satisfied, the whole structure was refined to remove steric
clashes with all experimental constraints using dynamic calcula-
tions and energy minimization to generate the preliminary 3D
model. It should be pointed out that DNA is an extended,
nonglobular molecule, and NMR can only provide short-range
NOE distance constraints (<4.5 Å). Given no direct long-range
experimental constraints, we refrain from inferring any changes
in the global structural features of the derived DNA model,
focusing instead on the local structure in the modified TT dimer
region where the extensive NOEs are available for constraining
structures. Figure 7a highlights the TT dimer region from the
calculated structure model next to the corresponding region in
the native B-DNA21 duplex in stick model representation, Figure
7b. The solvation surfaces were created by rolling a water
molecule of 1.4 Å radius over the van der Waals surfaces of
the molecule.22 As constrained by the NMR data, the hydro-
phobic C7′ methylene unit has swung from the usual solvent-
-
exposed position of the PO2 group into the interior of the
double helix, in close contact with a core formed by the base
edges and sugar rings of the flanking TT/AA nucleosides. We
simply refer to this region as the hydrophobic core of a DNA
(22) Alden, C. J.; Kim, S.-H. J. Mol. Biol. 1979, 132, 411.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3027
duplex although it is certainly more extended than the core of
a globular protein.
difference in ∆S exclusively as a function of conformational
differences would require a difference of approximately seven
bonds along the DNA backbone between the native and modi-
fied duplex, or 7 × 1.987 ln 3 ) 15 eu mol-1. Although the
ethylamine linker may be more flexible than the phosphodiester
linkage,2e,f the number of backbone bonds in duplex 1 has not
been changed and the observed difference in ∆S is too large to
be explained by the change in the conformational entropy of
the modified backbone.
Discussion
The structural and thermodynamic analyses of duplex 1 have
been compared to those of an unmodified DNA duplex of the
same sequence. The destabilizing effect of the modification
may be the result of duplex destabilization, stabilization of the
backbone-modified single strand, or both. Because the nearest-
neighbor data are derived from measurements in which the
single-stranded oligonucleotides are in the random coiled state,
the good agreement between the observed and predicted values
of ∆H for the native duplex (Table 1) argues that there is no
single-stranded structure that contributes to the observed ∆H
and ∆S values on duplex formation.11b Indeed, a direct
calorimetric measurement11b has shown that the thermodynamic
contribution of the structured single-stranded DNA makes the
observed ∆H value of duplex formation significantly lower than
the predicted value using nearest-neighbor data.11a
The NMR data provide complementary evidence supporting
the observed change in stability being attributable to the duplex.
The difference between the ∆δ values of the T3-CH3 groups in
duplex 1 and its native analogue is 0.04 ppm, and the ∆δ values
converge in the thermally unfolded strands. This convergence
of the chemical shift values suggests that the T3-CH3 probe is
responsive to the local change in duplex 1 rather than to the
chemical modification, and therefore, duplex destabilization,
rather than a structured single strand, is the primary contributor
to the observed thermodynamic difference between duplex 1
and the native DNA.
Although an overall B-DNA structure with base stacking and
interstrand H-bonds was clearly present in duplex 1, a global
destabilization of duplex 1 relative to its native duplex is
detected by both UV and NMR monitored thermal unfolding
experiments. This destabilization is reflected in an unfavorable
change in ∆H of 7 kcal mol-1 (Table 1).
A significant conformational change in the backbone of
duplex 1 was detected by the NMR experiments. This structural
change is of particular interest in relation to the significant
entropic stabilization. A 15 eu mol-1 change in entropy is
significant; sequence-dependent variations in ∆S are typically
less than 3 eu mol-1 per base pair.11a Modifying the phos-
phodiester into the neutral ethyl phosphotriester gives a similar
positive entropic change, but only of ∼7 eu mol-1 per modi-
fication.23 Given the extensive cooperativity in forming a DNA
duplex, it is likely that this local change near the modified
backbone could cause a global weakening in the stability of
duplex 1. Therefore, both the direction and magnitude of the
changes in ∆H and ∆S and their correlation with the structural
change in the backbone of the duplex 1 deserve further
consideration.
With respect to ion condensation, the replacement of a phos-
phate group with a positively charged ethylamine should reduce
the net charge density along the modified backbone. The
entropic driving force resulting from the counterion condensation
on the modified DNA backbone would become less favorable
to duplex formation rather than more favorable. With respect
to hydration, high-resolution X-ray diffraction data from Dick-
erson and co-workers reveal the presence of a spine of H2O
molecules in the minor groove of A-T base pairs25 and the
dT‚dA duplex appears to have an even higher degree of
hydration.26 Breslauer and co-workers27 have attempted to inter-
pret the thermodynamic data for DNA bending drug interactions
in terms of the disruption of levels of DNA hydration beyond
the minor groove. The increase in entropy appeared to correlate
with the release of the tightly bound water molecules into the
bulk medium resulting from the structural transition and/or
ligand binding. We have no direct experimental evidence that
identifies the change in water binding in the modified backbone
even though such a disruption is expected in the modified dAdT
region and might contribute to the observed increase in ∆S.5
On the other hand, an entropic factor has been identified by
studying the duplex formation of a DNA hexamer in 10 mol %
alcohol-water mixtures.8c The overall destabilization upon
introduction of alcoholic cosolvents to the DNA solution is
attributed to an unfavorable change in entropy that ranges from
7 (in MeOH) to 20 (in PrOH) eu mol-1 (see Table 3 in ref 8c).
This change is consistent with a contribution from nonspecifc
hydrophobic interactions to stabilizing the DNA duplex; the
entropic driving force is disrupted upon adding alcohols. In-
cidently, this change in ∆S is of the same magnitude as we
have observed here, suggesting that changes in the hydrophobic
interaction from the native to the modified duplex 1 might be
invoked to explain the significant and favorable change in
∆S, even though only one backbone linkage is modified in
duplex 1.
Collapse of the Modified Backbone onto the Hydrophobic
Interior of the DNA Duplex. The NMR analysis showed that
the C7′ methylene replacing the PO2- group can reside in close
contact with a core formed by the base edges and sugar rings
of the connecting nucleotides. The structural change shown in
Figure 7 is consistent with a partitioning of the flexible
ethylamine onto the hydrophobic core of this duplex. Hydro-
phobic collapse is a term frequently invoked to describe the
folding of proteins,28 but is not commonly applied to nucleic
acids. This interpretation nevertheless seems valid in this case
for several reasons.
Factors Contributing to the Observed Difference in ∆S.
The factors that contribute to the change in entropy can be
expressed by eq 7. The ∆Sconformation has been estimated by the
∆Sobserve ) ∆Sconformation + ∆Sion-condensation + ∆Sother
(7)
The structures of nucleic acids display the protein folding
pattern of “polar out, nonpolar in”. The polyanionic phosphate
groups are exposed to water, and the hydrophobic moieties of
expression nR ln 3 where R is the gas constant and n is the
number of bonds per residue along the backbone.24 Each bond
can populate three conformers in the random strand, but only
one upon duplex formation. To account for the observed
(25) Drew, H. R.; Dickerson, R. E. J. Mol. Biol. 1981, 151, 535.
(26) Pilet, J.; Blicharski, J.; Brahms, J. Biochemistry 1975, 14, 1869.
(27) (a) Breslauer, K. J.; Remeta, D. P.; Chou, W.-Y.; Ferrante, R.; Curry,
J.; Zaunczkowski, D.; Snyder, J. G.; Marky, L. A. Proc. Natl. Acad. Sci.
U.S.A. 1987, 84, 8922. (b) Marky, L. A.; Breslauer, K. J. Proc. Natl. Acad.
Sci. U. S.A. 1987, 84, 4359.
(23) Pless, R. C.; Ts’o, P. O. P. Biochemistry 1977, 16, 1239.
(24) (a) Privalov, P. L. AdV. Protein Chem. 1979, 33, 167. (b) Baldwin,
R. L. Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 8069. (c) Luo, P.; Lynn, D.
G. Submitted for publication.
(28) Kim, P. S.; Baldwin, R. L. Annu. ReV. Biochem. 1990, 59, 631.

3028 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Luo et al.
Scheme 3. A Model for the Altered Stability of Modified Duplex 1a
a The duplex with the negatively charged phosphate removed (A) initiates the collapse of the ethyl amine onto the hydrophobic interior of the
duplex (B). The two flanking phosphates are forced to reduce the distance separating them, causing backbone curvature and loss of the optimal
geometry for base-stacking and H-bonding interactions (C).
the bases and sugars are buried inside. Estimation of the
solvent-accessible surface has shown that two-thirds of the
hydrophobic surfaces are buried upon duplex formation, while
the phosphate groups account for almost half of the surface
exposed to the solvent22 and are maximally partitioned into
water.25,29 Further, examination of the DNA duplex structure
shows that the positions occupied by the PO2- and the 5′-O on
the backbone linkage are flexible, being able to rotate without
causing undue constraints on base pairing and sugar pucker
geometry. In contrast, conformational degrees of freedom in
both the 3′-O and 5′-methylene positions along the backbone
are primarily determined by the sugar pucker of the connecting
nucleotides. Indeed, the X-ray structure of a single backbone-
modified DNA with the O atom at the 3′-O position replaced
by a methylene group is essentially identical to the B-form
conformation of the native DNA duplex.30 It follows that the
C6′ methylene in the modified duplex 1 could be accommodated
in a conformation similar to that of the corresponding O atom
in the native DNA duplex (Figure 7a). Therefore, the spatial
orientation of any chemical group at the unrestricted position
of the phosphate, or the C7′ methylene in duplex 1, should be
determined by partitioning between the solvent-exposed surface
and the solvent-shielded hydrophobic interior.
Estimates of the Magnitude of the Hydrophobic Interac-
tion. Using in vitro selection experiments, the magnitude and
specificity of the hydrophobic interaction between aliphatic
groups with the interior of RNA have been directly estimated.31
The binding free energy per methylene is up to 1.5 kcal mol-1
at 25 °C, comparable to the similar interaction in proteins.
Because the hydrophobic interaction is entropy-driven at room
temperature,24b ∆S values of transferring one CH2 group from
a water-exposed site into the hydrophobic interior of nucleic
acids can be 5 eu mol-1, or up to 10 eu mol-1 for the two CH2
groups in duplex 1, a significant percentage of the observed
difference in ∆S between the native and the modified duplex 1
(Table 1).
solutions;32a the change in ∆S is -42.2 eu mol-1 for ethylamine,
-39.8 eu mol-1 for ethanol, and -33.6 eu mol-1 for ethane at
25 °C, respectively.32 After the change in entropy due to
vaporization at 25 °C has been corrected, the change in ∆S from
the liquid state to an infinitely dilute aqueous solution for ethanol
is -7 eu mol-1. Alternatively, using the liquid hydrocarbon
model,24b,c the ∆Cp ) 33.9 eu mol-1 for transfer of pure ethanol
to an aqueous solution at infinite dilution at 25 °C,33 the
corresponding change in ∆S is equal to ∆Cp ln(T/Ts) or -9 eu
mol-1, where Ts ) 386 K for the reference temperature in the
liquid hydrocarbon model.24b Both of these estimates are of
magnitude and direction similar to those seen with the RNA
measurements and close to the observed 15 eu mol-1 attributed
to the hydrophobic collapse of the ethylamine group onto the
interior of the DNA duplex. The hydrophobic interaction
between the ethylamine linkage and the hydrophobic core
formed by the sugar rings and base edges appears to be a
significant contributor to the change in ∆S of duplex 1.
Local Backbone Distortion and Structural Stability of the
Modified Duplex. Due to the intrinsic difficulty in describing
cooperativity involving long-range interactions underlying the
folding properties of biopolymers such as proteins,28 a model
based on the cooperativity of the DNA duplex stability is
required to relate the local backbone distortion, which is detected
by NMR and assigned to a favorable change in ∆S, to the overall
destabilization of the modified duplex 1. Scheme 3 suggests
that a duplex containing a more hydrophobic linkage (A) initiates
the conformational partitioning onto the hydrophobic interior
of the duplex (B). This movement accentuates the gap between
the two flanking phosphates, forcing a movement to shorten
their intervening distance and destabilizing the optimal geometry
for duplex stability (C).
The driving force for the movement of these phosphates could
have several origins. It has been demonstrated recently that
asymmetric charge neutralization of DNA phosphate groups can
result in the bending of the DNA duplex around the site of
neutralization.34 We suggest that another mechanism, namely,
the collapse of a hydrophobic linkage such as C7′ methylene
of duplex 1, can also contribute to the bending of the DNA
duplex. The positive charge could also act in concert with the
Despite their positive charge, amines, like alcohols, are
considered soluble hydrocarbons because their thermodynamic
properties in dilute aqueous solution are controlled predomi-
nantly by the aliphatic groups. A large and positive change in
heat capacities and negative change in entropy are observed
upon transferring from the vapor state to infinitely dilute aqueous
(32) (a) Franks, F., Reid, D. S. In Water; Franks, F., Ed.; Plenum Press:
New York, 1973; Vol 2, Chapter 5, pp 323-404. (b) Frank, H. S.; Evans,
M. W. J. Chem. Phys. 1945, 13, 507.
(33) Alexander, D. M.; Hill, D. J. T. Aust. J. Chem. 1969, 22, 347.
(34) (a) Strauss, J. K.; Maher III, L. J. Science 1994, 266, 1829. (b)
Strauss, J. K.; Roberts, C.; Nelson, M. G.; Switzer, C.; Maher, L. J., III.
Proc. Natl. Acad. Sci. U.S.A. 1994, 93, 9515.
(29) Saenger, W. Principles of Nucleic Acid Structure; Springer-
Verlag: New York, 1984.
(30) Heinemann, U.; Rudolph, L.-N.; Claudis, A.; Morr, M.; Heikens,
R. F.; Blocker, H. Nucleic Acids Res. 1991, 19, 427.
(31) Majerfeld, I.; Yarus, M. Nature Struct. Biol. 1994, 1, 287.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3029
Materials and Methods
hydrophobic collapse by pulling the two flanking phosphates
of the same strand together, although the N-acyl duplex 2 argues
that the positive charge in the duplex 1 might stabilize duplex
1 relative to 2, possibly by a Coulombic attraction with the
interstrand phosphates. The neutral amide 3 is significantly
stabilized, and this stability can be assigned to the reduced
flexibility of the amide and the resulting inability of this linker
to partition onto the hydrophobic core. The local distortion is
proposed to disrupt the cooperativity in forming a duplex,
causing backbone curvature, a loss of the optimal geometry for
base-stacking and H-bonding interactions, and resulting in the
general loss in the enthalpic stabilization of duplex 1.
Synthetic Procedures. General methods as well as detailed
procedures for the construction of 4-7 are provided in the Supporting
Information.35-40
Bis-O-t-BDMS-5′-thymidinylaminoethylthymidine (8). 3′-Car-
boxyaldehyde-5′-OTBDMS-3′-deoxythymidine (5) (281 mg, 740 µmol)-
and 5′-amino-3′-OTBDMS-5′-deoxythymidine (7) (314 mg, 880 µmol,
1.2 equiv) were dissolved in 3 mL of freshly distilled THF, and 233
mg (1.1 mmol, 1.5 equiv) of sodium cyanoborohydride41 was added
with stirring at room temperature. TLC indicated that reaction was
complete within approximately 2 h (TLC; 9:1 CHCl3-MeOH; Rf )
0.25). The reaction solution was concentrated and chromatographed
on silica gel (9:1 CHCl3-MeOH) to yield a white foam, 504 mg (700
µmol, 95%). 1H NMR (300 MHz, CDCl3/TMS): δ ) 7.58 (d, J ) 1
Hz, 1H, H-6); 7.17 (d, J ) 1 Hz, 1H, H-6); 6.12 (t, J ) 6.2 Hz, 1H,
H-1′); 6.06 (dd, J ) 4.1 Hz, 6.3 Hz, 1H, H-1′); 4.30 (m, 1H, H-3′b);
4.00 (dd, J ) 2.9 Hz, 12.3 Hz, 1H, H-5′a); 3.92 (m, 1H, H-4′b), 3.73
(m, 2H, H-4′a, H-5′′a); 2.8-2.9 (m, 2H, H-5′/5′′b) 2.72 (t, J ) 7.3 Hz,
2H, H-7′/7′′a); 2.1-2.3 (m, 7H, H-6′/6′′a, H-2′/2′′a, H-2′/2′′b, H-3′a);
1.93 (d, J ) 1 Hz, 6H, thymine-CH3); 0.93, 0.89 (s, 18H, (CH3)3CSi);
0.11, 0.08 (s, 12H, (CH3)2Si). 13C NMR (75 MHz, CDCl3): δ ) 164.4,
164.1 (C-4a,b); 150.7, 150.6 (C-2a,b); 136.1, 135.8 (C-6); 111.3, 110.3
(C-5a,b); 86.6, 85.4 (C-4′a,b); 85.4, 85.3 (C-1′a,b); 72.8 (C-3′a); 63.1
(C-5′a); 51.3 (C-7′a); 48.7 (C-5′); 40.5, 39.4 (C-3′a, C-2′b); 35.7 (C-
6′a); 32.8 (C-2′a); 26.2, 25.9 ((CH3)3CSi a,b); 18.7, 18.1 ((CH3)3CSi,
a,b); 12.8 (thymine-CH3 a,b); -4.5, -4.6, -5.1 ((CH3)2Si a,b).
Bis-O-t-BDMS-5′-thymidinyl-N-acetylaminoethylthymidine (9).
Triethylamine (112 mg, 1.1 mmol, 20 equiv), acetic anhydride (22.5
mg, 220 µmol, 4 equiv), and bis-O-t-BDMS-5′-thymidinylaminoeth-
ylthymidine (8) (40 mg, 55 µmol) were dissolved in 2 mL of
dichloromethane. The reaction was complete in 1 h as determined by
TLC (EtOAc, Rf ) 0.18; 9:1 CHCl3-MeOH, Rf ) 0.43). Concentration
of the solvent gave 9 as a colorless oil in quantitative yield. NMR
analysis indicated the presence of the two N-acetate rotamers in
approximately equal amounts. 1H NMR (400 MHz, CDCl3/TMS): δ
) 9.66, 9.38, 9.31 (3s, 2H, NHa,b); 7.61, 7.56, 7.23, 7.00 (4s, 2H,
H-6′); 6.15 (t, J ) 6.7 Hz, 0.5H, H-1′a); 6.07 (m, 1H, H-1′b); 5.87
(dd, J ) 5 Hz, 8 Hz, 0.5H, H-1′a); 4.31 (m, 1H, H-3′b); 3.99 (m, 2H,
H-4′a, H-5′a); 3.75 (m, 2H, H-4′b, H-5′′a), 3.43 (m, 4H, H-7′/7′′a, H-5′/
5′′b); 2.5 (m, 1H, H-3′a); 2.25 (m, 4H, H-6′/6′′a, H-2′a, H-2′b); 2.1
(m, 2H, H-2′′a, H-2′′b); 2.10, 2.13 (s, 3H, (CH3)C(O)N); 2.06, 1.96,
1.94, 1.93 (s, 6H, thymine-CH3); 0.94, 0.93 (s, 9H, (CH3)3CSi); 0.91
(s, 9H, (CH3)3CSi); 0.14, 0.13, 0.12, 0.11, 0.10, 0.91 (s, 12H, (CH3)2-
Si). 13C NMR (75 MHz, CDCl3): δ ) 170.8, 170.7 ((CH3)C(O)N);
164.4, 164.3, 164.2 (C-4a,b); 150.9, 150.7, 150.6, 150.2 (C-2a,b); 137.6,
137.5, 136.5, 135.9 (C-6a,b); 111.4, 110.5, 110.4 (C-5a,b); 88.3-85.3
(8s, C-4′a,b; C-1′a,b); 73.8, 72.9 (C-3′b); 63.2, 62.8 (C-5′a); 51.2 (C-
7′a); 47.8, 47.3 (C-5′a); 39.7, 39.6, 39.5, 39.1 (C-3′a, C-2′b); 35.4, 35.2
(C-6′a); 31.5, 30.4 (C-2′a); 26.1, 25.9 ((CH3)3CSi a,b); 22.0, 21.5
((CH3)C(O)N 18.6, 18.1 ((CH3)3CSi, a,b); 12.8, 12.7 (thymine-CH3 a,b);
-4.3, -4.5, -4.7, -5.2 ((CH3)2Si a,b).
Conclusions
In this work we have analyzed the structural and thermody-
namic properties of the backbone modified DNA duplex 1 by
comparing the native duplex with 1, 2, and 3. A concern in
making this comparison is that the physical factors which control
the stability of a modified duplex may vary from one modifica-
tion to another and chemical modifications, such as removal of
a single phosphate, can introduce factors other than those
intrinsic to the native duplex. We posed five questions that
we saw as central to the studies of backbone-modified oligo-
nucleotide analogues and have developed them for this com-
parison.
Both structural and thermodynamic evidence supports the use
of a two-state model in analyzing the thermal unfolding of a
duplex with a single structural change in the center of one strand.
This structural change destabilizes the duplex. While the effect
of charge is analyzed qualitatively by referring to the properties
of chemical analogues 2 and 3 as well as data from the
literature,2e,f a more quantitative or semiquantitave approach is
taken in the analysis of the change in entropy due to solvation
and the hydrophobic nature of the modification. The favorable
change in ∆S has been assigned to the hydrophobic nature of
the ethylamine linkage, and the unfavorable change in ∆H is
due to the more global disruption of H-bonding and stacking
interactions. This effect is best summarized in a model that
takes into account how a local backbone distortion might couple
with long-range interactions in determining DNA duplex
formation.34 This model highlights a fundamental property
underlying the folding of biopolymers: a single modification
in the DNA backbone can result in favorable changes in entropy
and, at the same time, alter the overall duplex stability by
disrupting the cooperativity of the folding process. This study
of a single modification provides the groundwork and prospec-
tive necessary for understanding and designing the more
extensively modified structures necessary for metabolically
stable antisense reagents.
Amide Dimer 10. 5′-OTBDMS-3′-deoxythymidine-3′-acetic acid
(6) (100 mg, 250 µmol) and 5′-amino-3′-OTBDMS-5′-deoxythymidine
(7) (100 mg, 270 µmol, 1.1 equiv) were dissolved in 5 mL of
dichloromethane. 4-(Dimethylamino)pyridine (Fisher; 16 mg, 130
µmol, 0.5 equiv) and p-toluenesulfonic acid (15 mg, 80 µmol, 0.3 equiv)
were added, followed by 1-(3-(dimethylamino)propyl)-3-ethylcarbo-
diimide‚HCl (58 mg, 300 µmol, 1.2 equiv). The reaction solution was
This model has proven to be of value in the design and
characterization of systems for template-directed synthesis. The
backbone-modified structures have increasingly underscored the
role played by the phosphates in preorganizing the nucleic acids
into an extended conformation. This preorganization appears
to be an important structural contributor in driving efficient
strand recognition and duplex formation.6 The rigidity of the
imine precursor to duplex 1 has now been exploited in preparing
the smallest catalytic nucleic acid that efficiently gives catalytic
turnover and reduces production inhibition.5 It is now possible
to better consider other ligation and polymerization reactions
that can be used to extend this general concept of translating
information from a DNA polymer into other polymeric ana-
logues.
(35) Ogilvie, K. K. Can. J. Chem. 1973, 51, 3799.
(36) (a) Prisbe, E. J.; Martin, J. C. Synth. Commun. 1985, 15, 401. (b)
Fiandor, J.; Tam, S. Y. Tetrahedron Lett. 1990, 31, 597.
(37) Keck, G. E.; Enholm, E. J.; Wiley: M. R. Tetrahedron 1985, 41,
4079.
(38) Chu, C. K.; Doboszewski, B.; Schmidt, W.; Ullas, G. V. J. Org.
Chem. 1989, 54, 2767.
(39) Demuth, M.; Ritterskamp, P.; Weigt, E.; Schaffner, K. J. Am. Chem.
Soc. 1986, 108, 4149.
(40) Yamamoto, I.; Sekine, M.; Hata, T. J. Chem. Soc. Perkin Trans. 1
1980, 306.
(41) Borch, R. F.; Bernstein, M. D.; Durst, H. D. J. Am. Chem. Soc.
1971, 93, 2897.

3030 J. Am. Chem. Soc., Vol. 120, No. 13, 1998
Luo et al.
stirred at room temperature for 2 h, whereupon TLC indicated the
reaction was complete. Dilution with chloroform and washing with
water were followed by concentration of the organic layer and flash
chromatography (EtOAc; Rf ) 0.21) to yield 125 mg (170 µmol, 68%)
as a slightly yellow oil. See ref 42 for independent syntheses of this
linkage. 1H NMR (400 MHz, CDCl3/TMS): δ ) 9.47, 9.29 (2 br s,
2H, thymine-NH a,b); 7.59 (d, J ) 1 Hz, 1H, H-6); 7.07 (m, 1H, C(O)-
NH); 7.02 (s, 1H, H-6); 6.25 (t, J ) 6.5 Hz, 1H, H-1′a); 5.68 (t, J )
6.8 Hz, 1H, H-1′b); 4.40 (dt, J ) 4.5 Hz, 7.1 Hz, 1H, H-3′b); 3.93 (m,
2H, H-4′b, H-5′a); 3.81 (m, 1H, H-4′a); 3.75 (dd, J ) 2.8 Hz, 11.3 Hz,
H-5′′a); 3.67 (ddd, J ) 4.8 Hz, 6.6 Hz, 14.2 Hz, 1H, H-5′b); 3.46 (dt,
J ) 3.5 Hz, 14.2 Hz, 1H, H-5′′b); 2.83 (m, 1H, H-3′a); 2.64 (dt, J )
6.7 Hz, 13.5 Hz, 1H, H-2′b); 2.4 (m, 2H, H-6′/6′′a); 2.1-2.25 (m, 3H,
H-2′′b, H-2′/2′′a); 1.93, 1.91 (s, 6H, thymine-CH3); 0.93, 0.88 (s, 18H,
(CH3)3CSi); 0.13, 0.12, 0.07, 0.08 (s, 12H, (CH3)2Si). 13C NMR (75
MHz, CDCl3): δ ) 171.5 (C-7′a); 164.4 (C-4a,b); 151.2, 150.6 (C-
2a,b); 138.7, 135.9 (C-6a,b); 111.2 (C-5a,b); 89.8 (C-4′a,b); 85.6, 84.8
(C-1′a,b); 72.4 (C-3′b); 64.5 (C-5′a); 40.5 (C-5′b); 39.7, 39.5 (C-3′a,
C-2′b); 37.8 (C-6′a); 36.1 (C-2′a); 26.1, 25.9 ((CH3)3CSi a,b); 18.0
((CH3)2Si a,b); 12.7, 12.5 (thymine-CH3 a,b); -4.6, -4.7, -5.2 ((CH3)2-
Si a,b).
DNA Hexamers. The same procedure was followed for elaboration
of each dimer into oligonucleotides. The bis-silylated dimer was
deprotected overnight with excess tetrabutylammonium fluoride (1 M
in THF, Aldrich) followed by flash chromatography. Yields of this
deprotection were commonly in the 50-80% range. Removal of
residual tetrabutylammonium salts proved to be difficult, so other
methods of deprotection are currently under investigation.43
Desilylation was followed by dimethoxytritylation at the 5′-alcohol.
This protection was carried out overnight in anhydrous pyridine with
1.5 equiv of DMTrCl. Yields after chromatography were approximately
70%. Conversion to the phosphoramidite was performed as follows:
The tritylated dimer was dissolved in freshly distilled THF along with
diisopropylethylamine (3 equiv). 2-Cyanoethyl N,N-diisopropylchlo-
rophosphoramidite (1.5 equiv) was added and the reaction stirred under
nitrogen at room temperature. TLC indicated the reaction was complete
within approximately 1 h; this solution was rapidly chromatographed
on silica. Concentration was accomplished by repeated coevaporation
with heptane and acetonitrile. Following overnight evacuation on a
vacuum pump, the phosphoramidite dimer was used in solid-phase DNA
synthesis of the hexamer.
Solid-Phase Hexamer Synthesis. Solid-phase synthesis of the
modified hexamers was carried out on an ABI 380B automated
synthesizer by Paul Gardner of the University of Chicago Oligonucle-
otide Synthesis Facility on scales ranging from 3 to 9 µmol. Production
of the trityl cation at the deprotection step after addition of the modified
dimer indicated successful coupling to the growing chain. Synthetic
oligonucleotides were purified as follows: following cleavage from
the solid support, the fully deprotected oligonucleotide was subjected
to cation-exchange chromatography (Dowex 50 × 2-100, Na+ form).
The oligonucleotides were prepurified by low-pressure reversed-phase
chromatography (C18 Sep-Pak, Waters) and then purified for UV melting
curves by semipreparative HPLC using a gradient of ammonium acetate
in a mixture of 20% acetonitrile and 80% 20 mM aqueous sodium
phosphate on a DuPont Zorbax Oligo column. NMR quantities were
purified through a Sephadex G-10 column with 10 mM NH4HCO3
buffer solution. The volatile salts were removed by repeated lyo-
philization.
104 M-1 cm-1 for d(GCAACG) and 5.10 × 104 M-1 cm-1 for
d(CGTTGC). Due to uncertainty in the molar extinction coefficient
for the modified d(CGTTGC) strand, each DNA strand (about 1-2
mM) in D2O was titrated into its complementary strand in the NMR
tube while the intensity ratios of their characteristic base proton
resonances were monitored at 60 °C. The samples were lyophilized
and dissolved in 400 µL of 100 mM NaCl, 10 mM NaH2PO4, 2 mM
NaN3 buffer at pH 6 and then dried. For experiments in D2O, the
sample was lyophilized twice from 99.9% D2O and finally redissolved
in 400 µL of D2O (99.96% enrichment). For experiments in H2O, the
sample was redissolved in 400 µL of 9:1 (v/v) H2O-D2O. All the
chemical shifts in Table
2 are referenced to sodium 3-(tri-
methylsilyl)propionate-2,2,3,3-d4 (TSP; δ ) 0.00 ppm).
NMR Spectroscopy. All NMR spectra were recorded on a General
Electric Ω500 spectrometer. One-dimensional spectra of exchangeable
protons in H2O were recorded using the spin-echo 1-1 pulse sequence
with the carrier frequency at the solvent resonance to suppress the strong
solvent peak.44 The 1D NOE difference spectra were obtained by
subtracting a reference spectrum irradiated off-resonance by the
decoupler radio frequency (rf) from spectra irradiated at the desired
peaks by the decoupler rf, collected into 8K data points by co-averaging
32 scans for each spectrum with 3000 total scans and a spectral width
of 11 000 Hz.
Hypercomplex quadrature detection45 was used to obtain pure-phase
absorption mode DQF-COSY46 and NOESY47 spectra. The NOESY
spectra were collected with mixing times of 50 and 200 ms at 1 °C for
the backbone-modified DNA duplex and at 20 °C for its native
counterpart. Phase-sensitive TOCSY spectra were recorded with use
of the Waltz-16 spin-lock pulse to drive the coherence transfer, and a
filter was inserted to remove the unwanted coherence transfer pathway
by a novel phase cycling procedure which makes recording of the
TOCSY spectra fairly routine.48 Typical 2D data were collected with
512 FIDs with 2K data points each, 32-96 scans for each FID, a
spectral width of 5600-6000 Hz, a 1-s predelay time in D2O, and 1-1.5
s for solvent suppression. TOCSY spectra were collected with spin-
lock times of 65, 80, and 100 ms to achieve direct, single, and multiple
relayed through-bond magnetization transfer. Solvent was suppressed
by continuous irradiation or selective excitation with the carrier set at
the solvent resonance for spectra collected in H2O. The data were
apodized with a Lorentzian-Gaussian or phase-shifted sine-bell func-
tion. The first data point in t1 was multiplied by 0.5 to suppress t1
ridges49 and also zero-filled in t1 or symmetrized as necessary.
Structure Refinement. The distance constraints were derived from
integration of the NOESY cross-peaks at mixing times of 50 and 200
ms. The integrals of cross-peak intensities were obtained and converted
into distance constraints by the two-spin approximation for NOESY
spectra collected at a mixing time of 200 ms due to the optimum signal-
to-noise ratio of the spectrum for the volume integration. The
possibility of spin diffusion artifacts was checked by control calcula-
tions, comparing ratios of cross-peak intensities of H6-H5, H6-CH3,
and H2′-H2′′ to those of their corresponding distances by the two-
spin approximation. Relative errors of 10% in distance were used as
the estimates for both the upper and lower distance bounds.19 However,
particular care was taken to ensure that the NOE cross-peaks of the
methylene protons on the modified linkage with the 5′-flanking sugar
protons (Figure 6) were also present at the short, 50-ms mixing time
by minimizing spin-diffusion artifacts for distance constraints (Sup-
porting Information).
NMR Sample Preparation. DNA samples were exchanged into
their sodium form by passing through an ion-exchange column of a
Dowex 50 × 2-100 resin (Aldrich) versus NaOH. The lyophilized
samples were eluted through a Sep-Pak C18 column with a gradient
of 0-6% acetonitrile in water, and were finally purified through a
Sephadex G-10 column with 10 mM NH4HCO3 buffer solution. The
volatile salts were removed by repeated lyophilization. Concentrations
of DNA oligomers were determined by their UV absorbances at 260
nm, and the molar extinction coefficients were calculated to be 5.98 ×
The initial structures were built and modified from standard B-DNA
using the Biopolymer and Builder modules in InsightII. Energy
(44) Sklenar, V.; Bax, A. J. Magn. Reson. 1987, 74, 469.
(45) States, D. J.; Haberkorn R. A.; Ruben D. J. J. Magn. Reson. 1982,48,
286.
(46) Rance, M.; Sørensen, O. W.; Bodenhausen, G.; Wagner, G.; Ernst,
R. R.; Wu¨thrich, K. Biochem. Biophys. Res. Commun. 1983, 117, 479.
(47) Macura, S.; Ernst, R. R. Mol. Phys. 1980, 41, 95.
(48) (a) Bax, A.; Davis, D. G. J. Magn. Reson. 1985, 65, 355. (b) Rance,
M. J. Magn. Reson. 1987, 74, 557. (c) Luo, P.; Demetrios, T. B.;
Subramanian, R. M.; Meredith, S. C.; Lynn, D. G. Biochemistry 1994, 33,
12367.
(42) Idziak, I.; Just, G.; Damha, M. J.; Giannaris, P. A. Tetrahedron
Lett. 1993, 34, 5417.
(43) Pirrung, M. C.; Shuey, S. W.; Lever, D. C.; Fallon, L. Bioorg Med.
Chem. Lett. 1994, 4, 1345.
(49) Otting, G. H.; Widmer, G.; Wagner, G.; Wu¨thrich, K. J. Magn.
Reson. 1986, 66, 187.

Stability of the DNA Duplex with Backbone Modifications
J. Am. Chem. Soc., Vol. 120, No. 13, 1998 3031
minimization and dynamic simulations were carried out using the
Discover module in InsightII with the CVFF force field parameters.
The hydrogen bonds in the Watson-Crick base pairs were constrained
as supported by imino proton experiments to prevent dissociation of
the two strands during simulations. The target functions for dynamic
simulated annealing calculations are consisted of quadratic harmonic
terms for covalent geometry, square well quadratic potentials for the
NOE distance constraints and torsion angle constraints, and a quadratic
van der Waals repulsion term for the nonbonded contacts.50 The final
structures was minimized by switching back to standard potential
functions of the CVFF force field.
(Abelbeck Software). All data points were equally weighted unless
otherwise stated, and errors reported for the fitted parameters are at
the 67% confidence level.
Acknowledgment. We gratefully acknowledge the expertise
of Paul Gardner in solid-phase DNA synthesis and thank Dr.
Dean Astumian for the use of computing facilities and NIH,
Biotechnology Training Grant (GM-08369) to J.C.L. and NIH
Grant R21 RR12723 to D.G.L., for support.
Supporting Information Available: General synthetic
procedures, the NOESY spectrum of the native DNA duplex,
and the NOESY spectrum of the modified duplex showing the
characteristic C7′-T3-H1′ cross-peak recorded with a 50-ms
mixing time (7 pages). See any current masthead page for
ordering information and Web access instructions.
Thermal Melting and Data Analysis. Temperature-dependent
ultraviolet spectra were measured with stirring on a Hewlett-Packard
8542A diode array spectrophotometer equipped with an HP89090A
Peltier temperature controller. Data were analyzed using Kaleidagraph
(50) Nilges, M.; Clore, G. M.; Gronenborn, A. M. FEBS Lett. 1988, 239,
129.
JA972869W