NMR Characterization of a DNA Duplex Containing the Major
Acrolein-derived Deoxyguanosine Adduct
-OH-1,-N2-Propano-2'-deoxyguanosine*
Carlos
de los Santos
,
Tanya
Zaliznyak, and
Francis
Johnson
From the Department of Pharmacological Sciences, State University
of New York at Stony Brook, New York 11794-8651
Received for publication, October 3, 2000
 |
ABSTRACT |
The environmental and endogenous mutagen
acrolein reacts with cellular DNA to produce several isomeric
1,N2-propanodeoxyguanosine adducts. High
resolution NMR spectroscopy was used to establish the structural
features of the major acrolein-derived adduct,
-OH-1,N2-propano-2'-deoxyguanosine. In
aqueous solution, this adduct was shown to assume a ring-closed form.
In contrast, when
-OH-1,N2-propano-2'-deoxyguanosine pairs
with dC at the center of an 11-mer oligodeoxynucleotide duplex, the
exocyclic ring opens, enabling the modified base to participate in a
standard Watson-Crick base pairing alignment. Analysis of the duplex
spectra reveals a regular right-handed helical structure with all
residues adopting an anti orientation around the glycosidic
torsion angle and Watson-Crick alignments for all base pairs. We
conclude from this study that formation of duplex DNA triggers the
hydrolytic conversion of
-OH-1,N2-propano-2'-deoxyguanosine to an
open chain form, a structure that facilitates pairing with dC during
DNA replication and accounts for the surprising lack of mutagenicity
associated with this DNA adduct.
 |
INTRODUCTION |
Acrolein is a ubiquitous environmental pollutant formed by
incomplete combustion of organic materials, including wood, food, tobacco, and fuels. This
,
-unsaturated aldehyde reacts to form hydroxylated 1,N2-propano-2'-deoxyguanosine
adducts in DNA (1-3). Acrolein also is formed endogenously during the
metabolic oxidation of polyamines (4) and is an end product of lipid
peroxidation (5-7).
-OH-1,N2-propano-2'-deoxyguanosine
(
-OH-PdG)1 adducts were
detected in DNA extracted from rat and human liver (8-10) and from
lymphocyte DNA in patients undergoing treatment with cyclophosphamide
(11, 12).
The mutagenic properties of acrolein have been explored in prokaryotic
and eukaryotic cells (13-16). However, until recently, site-specific
mutagenesis studies were not feasible due to the chemical lability of
-OH-PdG under conditions required for solid phase DNA synthesis.
Accordingly, a structural analog,
1,N2-propano-2'-deoxyguanosine (PdG), which
shares the exocyclic ring but lacks the hydroxyl group of naturally
occurring acrolein-derived adducts (see Fig. 1), was adopted as a model
for structural and biological studies of exocyclic DNA adducts
(17-27).
Primer extension studies on templates containing a single PdG residue
revealed that this adduct induces targeted base substitutions and
frameshift mutations in vitro (17-19). In bacteria, PdG
induced frameshift mutations when the lesion was embedded in a CG
repeat (20). In other sequence contexts, frameshift mutations were not
observed and the principal mutagenic events in bacteria and mammalian
cells were G
T transversions and G
A transitions (21, 22).
NMR studies of PdG embedded in duplex DNA showed that, under acidic
conditions, the modified base tends to adopt a syn conformation around
the glycosyl bond, forming PdG(syn) ·dA+(anti) and
PdG(syn)·dC+(anti) alignments. Also, a pH-independent
PdG(syn)·dG(anti) base pair was observed in solution (23-25). In
each of these alignments, PdG was inside the helix and hydrogen bonds
formed across the base pair involved the Hoogsteen edge of the adduct.
An alternative PdG(anti)·dA(anti) alignment, observed at basic
pH, showed an adduct exposed to solvent, displaced into the major
groove of the helix and unstacked from the flanking bases (26). The
transition between the PdG(syn)·dA+(anti) and
PdG(anti)·dA(anti) forms was reversible with a
pKa of ~7.0, indicating that both forms are
present and in equilibrium under physiological conditions (26, 27). PdG
also was used to evaluate the thermodynamic impact of acrolein-derived
lesions in DNA duplexes. The adduct reduced thermal stability,
transition enthalpy, and transition free energy of the duplex; thermal
destabilization was insensitive to the base opposite the adduct
(28).
Recent advances in the chemical synthesis of acrolein-derived adducts
(29, 30) have made it possible to incorporate
-OH-PdG into
oligodeoxynucleotides, enabling the mutagenic properties of this adduct
in bacteria to be assessed (see accompanying articles (39, 40)).
Surprisingly, synthesis past the adduct was essentially error-free. To
better understand the striking differences between the model adduct,
PdG, and the naturally occurring acrolein-derived adduct,
-OH-PdG,
we used NMR spectroscopy to determine the solution structure of
-OH-PdG both as a free nucleoside and in duplex DNA. For the latter
structure, the adduct was incorporated opposite dC at the center of an
11-mer oligodeoxynucleotide duplex (referred to as the acr-dG·dC
duplex). Our data establish that
-OH-PdG nucleoside exists in a
closed form in solution but undergoes complete conversion to an open
structure in duplex DNA. The chemical structure of
-OH-PdG and the
duplex sequence employed in this study are shown in Fig.
1.

View larger version (19K):
[in this window]
[in a new window]
|
Fig. 1.
A, chemical structure of
-(OH)-1,N2-propano-2'-deoxyguanosine. Optical
isomers with the hydroxyl group in (R) or (S)
configuration are possible. B, sequence and numbering scheme
of the acr-dG·dC duplex.
|
|
 |
EXPERIMENTAL PROCEDURES |
Synthesis and Purification of Oligodeoxynucleotide
Duplexes--
The oligodeoxynucleotide strand containing
-OH-PdG
was synthesized following methods recently described (29). Briefly, the
N2-dihydroxybutyl derivative of dG was
introduced into oligomeric DNA by standard phosphoramidite chemical
procedures. Sequences containing a terminal
O-5'-dimethoxytrityl group were isolated by treatment of the
crude synthetic product with concentrated ammonia for 46 h at room
temperature and purified by reverse phase HPLC. The mobile phase
consisted of solvent A (0.1 M triethylamine acetic acid
buffer, pH 6.8) and solvent B (acetonitrile). Using a linear gradient
of 0% to 50% of B over 50 min, the desired sequence was eluted as a
main fraction at ~34 min. The O-5'-dimethoxytrityl group
was removed by treatment with 80% acetic acid for 30 min, and the
solution was extracted with ether three times. The
O-5'-dimethoxytrityl off-products were then purified
by HPLC. Subsequent treatment of the oligomer containing the
N2-dihydroxybutyl-dG residue with an excess of
an aqueous solution of sodium periodate (0.1 M) at room
temperature, until all the starting material disappeared, yielded the
desired product. After an additional round of HPLC purification,
oligodeoxynucleotide sequences were desalted by passing them through a
Sephadex G-25 column and subsequently converted to the sodium salt
using a Dowex 50W cation exchange column. Unmodified
oligodeoxynucleotide sequences were prepared and purified by standard
methods. Electrospray mass spectrometry was used to confirm correct
mass/charge ratio of both oligomers.
Duplex Formation and Sample Preparation--
A 1:1 stoichiometry
of the duplex was obtained by monitoring the intensity of individual
NMR proton signals during gradual addition of the unmodified strand to
the
-(OH)-PdG-containing strand. NMR samples consisted of 130 A260 of the duplex dissolved in 0.6 ml of 10 mM phosphate buffer (pH 6.5) containing 50 mM NaCl and 1 mM EDTA in either 99.96% D2O or
90% H2O-10% D2O (v/v), corresponding to a
concentration of ~1.8 mM. Samples of the monomeric
-OH-PdG nucleoside were dissolved in a similar buffered solution at
a final concentration of 0.2 mM. Samples were degassed
before collection of the NMR data.
NMR Experiments--
One- and two-dimensional NMR spectra were
recorded on Varian Inova spectrometers operating at 11.75- and
14.1-Tesla field strengths. Proton chemical shifts were referenced
relative to TSP at 0.0 ppm. Phase-sensitive (31) NOESY (120, 200, and
300 ms mixing times), COSY, double quantum filtered-COSY,
COSY45, and TOCSY (70- and 120-ms isotropic mixing time) spectra in
D2O buffer were collected with a repetition delay of
1.5 s, during which the residual water signal was suppressed by
saturation. NOESY spectra (120- and 220-ms mixing time) in 10%
D2O buffer were recorded using a jump-return reading pulse
(32). Time domain data sets consisted of 2048 by 300 complex data
points in the t2 and t1
dimensions, respectively. For the COSY45 spectrum, 4096 complex points
were used in the t2 dimension. NMR data were
processed and analyzed using the Felix program (Biosym Technologies,
Inc.) running on Silicon Graphics computers. Time domain data sets were multiplied by shifted sinebell window functions prior to Fourier transformation. In the spectra of the free nucleoside, the residual water signal present in the time domain date was eliminated further by
subtraction of a fitted polynomial function. No base line correction was applied to the transformed spectra. A three-dimensional model of
the acr-dG·dC duplex was built using INSIGHTII (Biosym Technologies, Inc.) by replacing the nonhydrogen-bonded amino proton of a
deoxyguanosine residue at the sixth position of a B-form 11-mer duplex
for the
,
-dihydroxypropyl moiety. Using the conjugate gradient
method, this model was energy-minimized to ensure that distances
between H
/H
' and H
/H
' protons of
-OH-PdG and the H1'
protons of C7 and G18 residues were within the observable NOE range
(see text). Energy minimization was performed on Silicon Graphics
computers using the program X-PLOR 3.851 (33).
 |
RESULTS |
NMR Characterization of the acr-dG·dC Duplex: Nonexchangeable
Protons--
At pH values over 6.5, the one-dimensional proton
spectrum of the acr-dG·dC duplex displays a main set of sharp signals
manageable for NMR characterization. Below this pH value, a second
conformation of the duplex in solution is evident by the presence of
minor resonances that become stronger as the pH is reduced (see Fig. 5
below). Therefore, assignment of the proton signals follows the
examination of NOESY and COSY spectra collected at pH 6.5 using
standard analysis procedures (34, 35). Fig.
2 shows an expanded region of a NOESY
spectrum (300-ms mixing time) recorded in 100% D2O buffer
at 30 °C, depicting interactions between the base and the H1' proton
regions. Indicative of a right-handed helix, each base proton (purine
H8 or pyrimidine H6) shows NOE cross-peaks to the H1' proton of the
ipso and 5'-flanking sugar residues. At the center of the duplex these
NOE interactions can be traced without interruption, suggesting that
the presence of
-OH-PdG does not cause large perturbations of the
double-helix structure. In addition, the intensity of intra-residue
base-H1' NOE peaks is much weaker than that of the H5-H6 cross-peaks of cytosine residues suggesting an anti-conformation around the
glycosydic torsion angle for all residues of the acr-dG·dC
duplex (35). Additional evidence of a regular right-handed helix is the
observation of NOE peaks between each cytosine (H5) and the base proton
of its 5'-side neighbor (Fig. 2, peaks A-F). Analogous
directionality of NOE interactions is present between the base and
sugar H3', H2', H2" protons in other regions of the same spectrum
(regions not shown). Similarly, nonexchangeable protons of the central (A4 C5 acr-dG C7 A8)·(T15 G16 C17 G18 T19) segment have
chemical shift values almost identical to those of the corresponding
unmodified control duplex, indicating only a minor deviation from the
canonical DNA conformation. Chemical shifts of the nonexchangeable
protons of the acr-dG·dC duplex measured at 30 °C are listed in
Table I.

View larger version (24K):
[in this window]
[in a new window]
|
Fig. 2.
Duplicate contour plots of a portion of the
NOESY (300-ms mixing time) spectrum recorded in 100% D2O
buffer, pH 6.5, 30 °C. The figure shows distance connectivities
between base and H1' sugar protons in the (left) modified
and (right) unmodified strands of the acr-dG·dC duplex.
Solid lines connect each base proton (purine H8/pyrimidine
H6) to its own (peaks labeled on the figure) and 5'-flanking
H1' sugar protons. Labeled peaks are assigned as follows: A,
A4(H8)-C5(H5); B, acr-dG(H8)-C7(H5); C,
G10(H8)-C11(H5); D, G12(H8)-C13(H5); E,
G16(H8)-C7(H5); F, A20(H8)-C21(H5).
|
|
View this table:
[in this window]
[in a new window]
|
Table I
Proton chemical shifts of the acr-dG·dC duplex
Values are given in parts per million (ppm) relative to TSP. Chemical
shifts are recorded in phosphate buffer (10 mM), pH 6.5, containing 50 mM NaCl. Nonexchangeable protons are at
30 °C; exchangeable protons are at 5 °C.
|
|
Identification of the proton signals of the propyl bridge follows from
the analysis of COSY, TOCSY, and NOESY spectra collected in 100%
D2O buffer solutions. In the 300-ms mixing time NOESY spectrum, a proton signal at 4.93 ppm, assigned to H
, displays NOE
cross-peaks to the overlapping H
/H
' protons as well as the H
/H
' protons within the propyl moiety (Fig.
3A, peaks A and B, respectively). Accordingly, in a TOCSY (120-ms mixing
time) spectrum recorded under identical temperature and pH conditions, cross-peaks are present between these same proton signals (Fig. 3B, peaks A and B, respectively), and
among the H
, H
', and H
/H
' protons of
-OH-PdG (region not
shown). An intriguing observation is the simultaneous presence of NOE
peaks between the H
/H
' of the adduct and the H1' protons of G18
and C7 residues located in opposite strands of the duplex (Fig.
3A, peaks E and C, respectively). Besides this, the presence of a sharp nonexchangeable proton signal is
evident at 9.58 ppm, at 30 °C, and slightly upfield at 5 °C (see
Fig. 5 below), in a region of the spectrum that is normally devoid of
proton signals associated with the duplex. This minor signal shows no
cross-peak to any exchangeable or nonexchangeable proton of the duplex
and, based on its chemical shift, is assigned to a small percentage of
the aldehydic open form of
-OH-PdG (see Fig. 6 below).

View larger version (28K):
[in this window]
[in a new window]
|
Fig. 3.
A, contour plot of a portion of a
NEOSY (300-ms mixing time) spectrum recorded in 100% D2O
buffer, pH 6.5, 30 °C. The figure shows interactions between the
H1'/H3' and H2'/H2" sugar proton regions. Labeled peaks are assigned as
follows: A, acr-dG(H )-acr-dG(H /H '); B,
acr-dG(H )-acr-dG(H /H '); C,
C7(H1')-acr-dG(H /H '); D, C7(H1')-acr-dG(H /H ');
E, G18(H1')-acr-dG(H /H '). B, contour plot
of a portion of a TOCSY (120-ms mixing time) spectrum recorded in 100%
D2O buffer, pH 6.5, 30 °C, showing the same
expanded region as in A. Labeled peaks are assigned as
follows: A, acr-dG(H )-acr-dG(H /H '); B,
acr-dG(H )-acr-dG(H /H ').
|
|
Exchangeable Protons--
In the sample dissolved in 10%
D2O buffer, the 1D proton spectrum shows 11 imino proton
signals resonating between 12.0 and 14.0 ppm, in the Watson-Crick
region (see Fig. 5 below). Sequence-specific assignment of the
exchangeable proton signals results from the analysis of a NOESY
(220-ms mixing time) spectrum collected at 2 °C (pH
6.5). Fig. 4 shows expanded contour plots
depicting NOE interactions between the imino and the amino/base proton
regions of this spectrum. Each thymine imino proton displays a
strong NOE interaction to the H2 proton of the corresponding adenine partner, thus establishing the formation of Watson-Crick alignments for
all A·T base pairs of the duplex (Fig. 4, peaks A-D).
Similarly, the presence of NOE cross-peaks between the guanine imino
and the amino protons of the cytosine partner indicates the formation of Watson-Crick alignments in all nonlesion-containing G·C base pairs
of the duplex (Fig. 4, peaks E, E', I,
and I'). Surprisingly, a remaining imino proton
signal at 12.64 ppm, which is originated at the acr-dG·dC pair of the
duplex, displays strong NOE cross-peaks with three different amino
proton signals. Based on interactions to the previously assigned
C17(H5) proton and their strong NOE connectivity, which is only
observed in 10% D2O buffer, two of these signals are
readily assigned to the amino protons of the lesion-partner C17
residue. Thus, peaks J and J' in Fig. 4 originate from NOE interactions between acr-dG(N1H) imino and C17(N4H2) protons.
The third NOE cross-peak originates from the interaction between
acr-dG(N1H) and acr-dG(N2H) protons of the adduct (Fig. 4, peak
K). These connectivities are only possible when the adduct exists
in a ring-opened state so that the lesion-containing base pair adopts
the standard Watson-Crick alignment. Consistent with these assignments
and supporting the open form of
-OH-PdG, N1H and N2H display NOE
cross-peaks to the H
/H
' and H
/H
' protons of the propyl
chain (Fig. 4, peaks Q-T).

View larger version (36K):
[in this window]
[in a new window]
|
Fig. 4.
Contour plot of a portion of a NOESY (220-ms
mixing time) spectrum recorded in 10% D2O buffer, pH 6.8, 2 °C, showing distance connectivities for the exchangeable protons
of the acr-dG·dC duplex. Labeled peaks are assigned as follows:
A, T9(N3H)-A14(H2); B, T19(N3H)-A4(H2);
C, T3(N3H)-A20(H2); D, T15(N3H)-A8(H2);
E, G12(N1H)-C11(N4H)hb; E',
G12(N1H)-C11(N4H)nhb; F,
G2(N1H)-C21(N4H)hb; F',
G2(N1H)-C21(N4H)nhb; G,
G10(N1H)-C13(N4H)hb; G',
G10(N1H)-C13(N4H)nhb; H,
G18(N1H)-C5(N4H)hb; H',
G18(N1H)-C5(N4H)nhb; I,
G16(N1H)-C7(N4H)hb; I',
G16(N1H)-C7(N4H)nhb; J,
acr-dG(N1H)-C17(N4H)hb; J',
acr-dG(N1H)-C17(N4H)nhb; K, acr-dG(N1H)-
acr-dG(N2H); L, G16(N1H)-acr-dG(N2H); M,
G2(N1H)-A20(H2); N, G10(N1H)-A14(H2); O,
G18(N1H)-A4(H2); P, G16(N1H)-A8(H2);
Q/Q', acr-dG(N1H)-acr-dG(H /H ');
R, acr-dG(N1H)-acr-dG(H /H ');
S/S', acr-dG(N2H)-acr-dG(H /H ');
T, acr-dG(N2H)-acr-dG(H /H '). hb and
nhb denote hydrogen-bonded and nonhydrogen-bonded,
respectively.
|
|
Evidence of base stacking is seen in the connectivities between the
adenine H2 protons and the imino protons of the flanking base pairs
(Fig. 4, peaks M-O) and those among the imino protons of
the duplex (region not shown). Likewise, the strong NOE peak between
the amino proton of the adduct and G16(N1H) at the 3'-flanking base
pairs indicates proper stacking of
-OH-PdG inside the duplex (Fig.
4, peak L). Chemical shifts of the exchangeable protons of
the acr-dG·dC duplex measured at 2 °C are listed in Table I.
Proton Spectra of the
-OH-PdG Nucleoside--
The unexpected
observation that
-OH-PdG exists in an open form in the duplex
prompted us to investigate its state at the nucleoside level. In
contrast to observations made with the duplex sample, no proton signals
are observed around 9.60 and 4.90 ppm (Fig.
5B). The analysis of a TOCSY
spectrum of the nucleoside dissolved in 100% D2O buffer,
pH 6.5, 30 °C, reveals that the H
/H
', H
/H
', and H
protons resonate at 3.52/3.48, 2.22/1.92, and 6.36 ppm, respectively
(data not shown). These chemical shift values are slightly downfield
from those previously reported for the adduct dissolved in dimethyl
sulfoxide (30, 36) and suggest a prevalent closed state for the
-OH-PdG nucleoside dissolved in water. In addition, the exocyclic
form of the adduct is insensitive to pH changes and only the
ring-closed state is observed under a wide range of values (Fig.
5B).

View larger version (26K):
[in this window]
[in a new window]
|
Fig. 5.
A, pH dependence of the exchangeable
proton spectrum of the acr-dG·dC duplex dissolved in 90%
H2O buffer at 2 °C. The assignment of imino proton
signals at the lesion site is given in the figure. Asterisks
denote an alternative conformation of the duplex present at acidic pH
values. B, pH dependence of the nonexchangeable proton
spectrum of -OH-PdG nucleoside dissolved in 100% D2O
buffer at 30 °C.
|
|
Upon duplex formation, the chemical shifts of protons on the propyl
chain move significantly upfield, especially H
, that changes from
6.36 ppm in the nucleoside to 4.93 ppm in the duplex. This chemical
shift value, which is inconsistent with the aldehydic proton of the
lesion that resonates at 9.58 ppm, is ascribed to the H
proton of
the propyl chain in which the carbonyl group is present in the hydrated
form (dihydroxy) of the adduct (Fig. 6).
The relative population of these two forms is dependent on the pH of
the sample, the aldehydic form being favored by basic conditions. Apart
from these states of
-(OH)-PdG, an alternative conformation of the
acr-dG·dC duplex, which may involve protonated cytosine residues,
becomes evident at pH 6.4 and lower values (Fig. 5A). Proton
chemical shifts of the
-(OH)-1,N2-PdG
nucleoside are listed in Table II.

View larger version (12K):
[in this window]
[in a new window]
|
Fig. 6.
Chemical rearrangement exerted by
-(OH)-PdG. The exocyclic form present on the
free nucleoside can add a water molecule to afford the hydrated open
conformation observed in the acr-dG·dC duplex. Alternatively,
chemical rearrangement of -OH-PdG produces the
N2-( -oxopropyl) configuration of the adduct.
The two open forms of the adduct reach equilibrium with the hydrated
structure favored at neutral basic solutions.
|
|
View this table:
[in this window]
[in a new window]
|
Table II
Proton chemical shifts of -(HO)-PdG
Values are given in ppm relative to TSP. Chemical shifts were recorded
in phosphate buffer (10 mM), pH 6.5, containing 50 mM NaCl. Nonexchangeable protons are at 30 °C;
exchangeable protons are at 5 °C.
|
|
 |
DISCUSSION |
Solution Conformations of the acr-dG·dC Duplex--
Early in the
course of these studies it became evident that the acr-dG·dC duplex
adopts a single conformation only at neutral or basic pH (Fig. 5).
However, adduct-containing sequences are unstable to the basic
conditions used during sample purification, which promote oligomer
polymerization (data not shown). Therefore, we chose to conduct our
studies at pH 6.5 where ~85% of the acr-dG·dC duplex is in the
conformation present at basic pH. The directionality of sequential NOE
interactions indicates that this conformation is a double-stranded
helix with residues adopting an anti orientation around the
glycosidic bond (Fig. 2). The pattern of NOE peaks observed for the
exchangeable imino protons establish that all base pairs of the
acr-dG·dC duplex have a Watson-Crick alignment (Fig. 4). At the
lesion-containing base pair, this becomes possible only if
-OH-PdG
adduct exists as an open form with the N2-propyl
chain pointing away from the helix and toward the solvent. In this
conformation, the H
/H
' and H
/H
' protons of
-OH-PdG are
found in the minor groove of the helix, close to H1' protons of
residues in both strands of the duplex (Fig. 3A,
peaks B, C, and D), and its
Watson-Crick edge remains accessible forming a fully hydrogen-bonded
acr-dG·dC base pair (Fig. 4, peaks J, J', and
K). These structural characteristics are readily fulfilled within a regular B-form helix, as shown by the energy-minimized model
of the acr-dG·dC duplex (Fig. 7).

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 7.
Left, central segment of the
energy-minimized model of the acr-dG·dC duplex having the adduct in
the hydrated conformation. The adduct stays in the minor groove
of the helix pointing the dihydroxypropyl moiety directly toward the
solvent. The model fulfills short distances detected as NOE cross-peaks
and predicts the formation of a strong hydrogen bond between
acr-dG(O H) and C17(O4') (yellow dotted lines).
Right, view from the top of the helical axis
showing Watson-Crick alignment at the acr-dG·dC base pair.
|
|
Duplex DNA Induces Ring Opening--
Spectroscopic data of the
-OH-PdG nucleoside in aqueous solutions establish a pH-independent
1,N2-closed conformation for the adduct (Fig.
5B), suggesting that duplex formation catalyzes the
rearrangement of the propyl bridge to an open form. An analogous
transformation was described recently for DNA duplexes in which the
deoxyguanosine-malondialdehyde adduct M1G is positioned
opposite dC (37). However, a "canonical" Watson-Crick base pair
forms only in the case of
-OH-PdG. This difference may explain in
part why M1G is mutagenic in bacteria (38), whereas
-OH-PdG is not (see accompanying articles (39, 40)). The role of the
partner base in promoting ring opening of
-OH-PdG adducts will be
the subject of future investigations.
Comparison with Duplexes Containing PdG--
An unsubstituted
1,N2-propano-2'-deoxyguanosine adduct has been
used extensively in biological (17-22) and structural (23-27) studies
as a model for natural acrolein lesions. PdG tends to adopt the
syn conformation when the adduct is positioned opposite dG
at neutral pH and when dA or dC residues in the complementary strand
are protonated under acidic conditions (23-25). The
syn conformation permits formation of hydrogen-bonded base
pairs through the Hoogsteen edge of the adduct while stacking with
flanking residues. Results of the present study establish a fundamental difference between
-OH-PdG and PdG in that, under appropriate conditions, the former can undergo a chemical rearrangement in aqueous
solution to assume an open chain form. Thus, when
-OH-PdG is in an
anti conformation, a fully hydrogen-bonded acr-dG·dC base
pair exists at neutral/basic pH values, which does not perturb the
duplex structure (Fig. 7). However, at acidic pH, the spectra of the
acr-dG·dC duplex show exchangeable proton signals that appear to
originate from the amino group of a C+ residue (Fig.
5A). Considering the strong tendency of PdG to adopt a
syn conformation, it is likely that, at acidic pH, the duplex contains a syn
-OH-PdG adduct paired to a
protonated cytosine residue forming an alignment similar to the one
described for PdG·dC (25). The structural characteristics of this
conformation in the acr-dG·dC duplex is currently under investigation.
Biological Implications--
Two laboratories have performed
primer extension and site-specific mutagenesis studies in bacteria
using DNA containing
-OH-PdG. Synthesis past the lesion is reduced
indicating that
-OH-PdG blocks DNA synthesis and, when translesional
synthesis occurs, dCMP is incorporated opposite the lesion almost
exclusively (see accompanying articles (39, 40)). The present study
provides structural grounds for understanding this behavior. At the
replication fork
-OH-PdG would adopt the closed
1,N2-exocyclic form described for the free
nucleoside in solution. As with PdG, this conformation of the adduct is
expected to hinder incorporation of dAMP, dGMP, and TMP, resulting in
the inhibition of DNA synthesis. However, incorporation of dCMP
opposite
-OH-PdG would trigger the chemical rearrangement from the
exocyclic closed form of the adduct to an opened conformation. The
subsequent formation of a replication structure stabilized by
Watson-Crick hydrogen bonds would facilitate rapid extension of the
-OH-PdG·dC pair resulting in error-free translesional DNA
synthesis. Thus, chemical rearrangement of
-OH-PdG to an open form
during DNA synthesis would account for the lack of mutagenicity
observed with the major acrolein-derived 2'-deoxyguanosine adduct in bacteria.
 |
ACKNOWLEDGEMENTS |
We thank Cecilia Torres for the synthesis and
purification of modified oligodeoxynucleotides and Arthur P. Grollman
for critical reading of this manuscript.
 |
FOOTNOTES |
*
This work was supported by National Institutes of Health
Grants CA47995 and CA77094.The costs of publication of this
article were defrayed in part by the
payment of page charges. The article must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence should be addressed: Dept. of
Pharmacological Sciences, Basic Science Tower, 7th Fl., Rm. 147, State University of New York, Stony Brook, NY 11794-8651. Tel.: 631-444-3649; Fax: 631-444-3218; E-mail: cds@pharm.sunysb.edu.
Published, JBC Papers in Press, October 27, 2000, DOI 10.1074/jbc.M009028200
 |
ABBREVIATIONS |
The abbreviations used are:
-OH-PdG,
-OH-1,N2-propano-2'-deoxyguanosine;
TSP, (2,2,3,3-d4)sodium 3-trimethylsilyl-propionate;
NOESY, nuclear Overhauser effect spectroscopy;
COSY, correlation spectroscopy;
TOCSY, total correlation spectroscopy;
NOE, nuclear Overhauser effect;
dC, 2'-deoxycytidine;
dA, 2'-deoxyadenosine;
dG, 2'-deoxyguanosine;
T, thymidine;
acr, acrolein.
 |
REFERENCES |
1.
| World Health Organization Publications (1992) The WHO
Environmental Health Criteria Series, Vol. 127
|
2.
|
Galliani, G.,
and Pantarotto, C.
(1983)
Tetrahedron Lett.
24,
4491-4492[CrossRef]
|
3.
|
Chung, F. L.,
Young, R.,
and Hecht, S. S.
(1984)
Cancer Res.
44,
990-995[Abstract]
|
4.
|
Lee, Y.,
and Sayre, L. M.
(1998)
J. Biol. Chem.
273,
19490-19494[Abstract/Free Full Text]
|
5.
|
Esterbauer, H.,
Schaur, R. J.,
and Zoller, H.
(1991)
Free Radic. Biol. Med.
11,
81-128[CrossRef][Medline]
[Order article via Infotrieve]
|
6.
|
Wu, H.-Y.,
and Lin, Y.-L.
(1995)
Anal. Chem.
76,
1603-1612
|
7.
|
Chung, F. L.,
Nath, R. G.,
Nagao, M.,
Nishikawa, A.,
Zhou, G. D.,
and Randerath, K.
(1999)
Mutat. Res.
242,
71-81
|
8.
|
Nath, R. G.,
and Chung, F.-L.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
7491-7495[Abstract]
|
9.
|
Nath, R. G.,
Ocando, J. E.,
and Chung, F.-L.
(1996)
Cancer Res.
56,
452-456[Abstract]
|
10.
|
Chung, F.-L.,
Zhang, L.,
Ocando, J. E.,
and Nath, R. G.
(1999)
IARC Sci. Publ.
150,
45-54[Medline]
[Order article via Infotrieve]
|
11.
|
Alarcon, R. A.
(1976)
Cancer Treat. Rep.
60,
327-335[Medline]
[Order article via Infotrieve]
|
12.
|
McDiarmid, M. A.,
Iype, P. T.,
Kolodner, K.,
Jacobson-Kram, D.,
and Strickland, P. T.
(1991)
Mutat. Res.
248,
93-99[Medline]
[Order article via Infotrieve]
|
13.
|
Marnett, L. J.,
Hurd, H. K.,
Hollstein, M. C.,
Levin, D. E.,
Esterbauer, H.,
and Ames, B. N.
(1985)
Mutat. Res.
148,
25-34[Medline]
[Order article via Infotrieve]
|
14.
|
Curren, R. D.,
Yang, L. L.,
Conklin, P. M.,
Grafstrom, R. C.,
and Harris, C. C.
(1988)
Mutat. Res.
209,
17-22[Medline]
[Order article via Infotrieve]
|
15.
|
Smith, R. A.,
Cohen, S. M.,
and Lawson, T. A.
(1990)
Carcinogenesis
11,
497-498[Abstract]
|
16.
|
Kawanishi, M.,
Matsuda, T.,
Nakayama, A.,
Takebe, H.,
Matsui, S.,
and Yagi, T.
(1998)
Mutat. Res.
417,
63-75
|
17.
|
Shibutani, S.,
and Grollman, A. P.
(1993)
J. Biol. Chem.
268,
11703-11710[Abstract/Free Full Text]
|
18.
|
Hashim, M. F.,
and Marnett, L. J.
(1996)
J. Biol. Chem.
271,
9160-9165[Abstract/Free Full Text]
|
19.
|
Hashim, M. F.,
Schnetz-Boutaud, N.,
and Marnett, L. J.
(1997)
J. Biol. Chem.
272,
20205-20212[Abstract/Free Full Text]
|
20.
|
Benamira, M.,
Singh, U.,
and Marnett, L. J.
(1992)
J. Biol. Chem.
267,
22392-22400[Abstract/Free Full Text]
|
21.
|
Moriya, M.,
Zhang, W.,
Johnson, F.,
and Grollman, A. P.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
11899-11903[Abstract/Free Full Text]
|
22.
|
Burcham, P. C.,
and Marnett, L. J.
(1994)
J. Biol. Chem.
269,
28844-28850[Abstract/Free Full Text]
|
23.
|
Kouchakdjian, M.,
Marinelli, E.,
Gao, X. L.,
Johnson, F.,
Grollman, A.,
and Patel, D.
(1989)
Biochemistry
28,
5647-5657[Medline]
[Order article via Infotrieve]
|
24.
|
Huang, P.,
and Eisenberg, M.
(1992)
Biochemistry
31,
6518-6532[Medline]
[Order article via Infotrieve]
|
25.
|
Singh, U. S.,
Moe, J. G.,
Reddy, G. R.,
Weisenseel, J. P.,
Marnett, L. J.,
and Stone, M. P.
(1993)
Chem. Res. Toxicol.
6,
825-836[Medline]
[Order article via Infotrieve]
|
26.
|
Kouchakdjian, M.,
Eisenberg, M.,
Live, D.,
Marinelli, E.,
Grollman, A. P.,
and Patel, D. J.
(1990)
Biochemistry.
29,
4456-4465[Medline]
[Order article via Infotrieve]
|
27.
|
Huang, P.,
Patel, D. J.,
and Eisenberg, M.
(1993)
Biochemistry
32,
3852-3866[Medline]
[Order article via Infotrieve]
|
28.
|
Plum, G. E.,
Grollman, A. P.,
Johnson, F.,
and Breslauer, K. J.
(1992)
Biochemistry
31,
12096-12102[Medline]
[Order article via Infotrieve]
|
29.
|
Khullar, S.,
Varaprasad, C. V.,
and Johnson, F.
(1999)
J. Med. Chem.
42,
947-950[CrossRef][Medline]
[Order article via Infotrieve]
|
30.
|
Nechev, L. V.,
Harris, C. M.,
and Harris, T. M.
(2000)
Chem. Res. Toxicol.
13,
421-429[CrossRef][Medline]
[Order article via Infotrieve]
|
31.
|
States, D. J.,
Habekorn, R. A.,
and Ruben, D. J.
(1982)
J. Magn. Res.
48,
286-292
|
32.
|
Plateau, P.,
and Gueron, M.
(1982)
J. Am. Chem. Soc.
104,
7310-7311
|
33.
|
Brünger, A.
(1993)
X-PLOR, Version 3.1: A system for X-Ray Crystallography and NMR
, Yale University Press, New Have, CT
|
34.
|
van de Ven, J. M.,
and Hilbers, C. W.
(1988)
Eur. J. Biochem.
178,
1-18[Medline]
[Order article via Infotrieve]
|
35.
|
de los Santos, C.
(1999)
in
Comprehensive Natural Products Chemistry, vol 7: DNA and Aspects of Molecular Biology
(Barton, D.
, Nakanishi, K.
, and Meth-Cohn, O., eds)
, pp. 55-80, Elsevier Science Ltd., Oxford, UK
|
36.
|
Boerth, D. W.,
Eder, E.,
Hussain, S.,
and Hoffman, C.
(1998)
Chem. Res. Toxicol.
11,
284-294[CrossRef][Medline]
[Order article via Infotrieve]
|
37.
|
Mao, H.,
Schnetz-Boutaud, N. C.,
Weisenseel, J. P.,
Marnett, L. J.,
and Stone, M. P.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
6615-6620[Abstract/Free Full Text]
|
38.
|
Fink, A. P.,
Reddy, G. R.,
and Marnett, L. J.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
8652-8657[Abstract/Free Full Text]
|
39.
|
Yang, I.-Y.,
Hossain, M.,
Miller, H.,
Khullar, S.,
Johnson, F.,
Grollman, A. P.,
and Moriya, M.
(2001)
J. Biol. Chem.
276,
9071-9076[Abstract/Free Full Text]
|
40.
|
VanderVeen, L. A.,
Hashim, M. F.,
Nechev, L. V.,
Harris, T. M.,
Harris, C. M.,
and Marnett, L. J.
(2001)
J. Biol. Chem.
276,
9066-9070[Abstract/Free Full Text]
|
Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.