Spectroscopic and theoretical insights into sequence effects of aminofluorene-induced conformational heterogeneity and nucleotide excision repair.

Abstract

A systematic spectroscopic and computational study was conducted in order to probe the influence of base sequences on stacked (S) versus B-type (B) conformational heterogeneity induced by the major dG adduct derived from the model carcinogen 7-fluoro-2-aminofluorene (FAF). We prepared and characterized eight 12-mer DNA duplexes (-AG*N- series, d[CTTCTAG*NCCTC]; -CG*N- series, d[CTTCTCG*NCCTC]), in which the central guanines (G*) were site-specifically modified with FAF with varying flanking bases (N = G, A, C, T). S/B heterogeneity was examined by CD, UV, and dynamic 19F NMR spectroscopy. All the modified duplexes studied followed a typical dynamic exchange between the S and B conformers in a sequence dependent manner. Specifically, purine bases at the 3'-flanking site promoted the S conformation (G > A > C > T). Simulation analysis showed that the S/B energy barriers were in the 14-16 kcal/mol range. The correlation times (tau = 1/kappa) were found to be in the millisecond range at 20 degrees C. The van der Waals energy force field calculations indicated the importance of the stacking interaction between the carcinogen and neighboring base pairs. Quantum mechanics calculations showed the existence of correlations between the total interaction energies (including electrostatic and solvation effects) and the S/B population ratios. The S/B equilibrium seems to modulate the efficiency of Escherichia coli UvrABC-based nucleotide excision repair in a conformation-specific manner: i.e., greater repair susceptibility for the S over B conformation and for the -AG*N- over the -CG*N- series. The results indicate a novel structure-function relationship, which provides insights into how bulky DNA adducts are accommodated by UvrABC proteins.

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Spectroscopic and Theoretical Insights into Sequence Effects of
Aminofluorene-Induced Conformational Heterogeneity and Nucleotide Excision
Repair†,∇
Srinivasa Rao Meneni,‡Steven M. Shell,§Lan Gao,|Petr Jurecka,⊥,#Wang Lee,‡Jiri Sponer,⊥Yue Zou,§
M. Paul Chiarelli,|and Bongsup P. Cho*,‡
Department of Biomedical and Pharmaceutical Sciences, College of Pharmacy, UniVersity of Rhode Island,
41 Lower College Road, Kingston, Rhode Island 02881, Department of Biochemistry and Molecular Biology,
Quillen College of Medicine, East Tennessee State UniVersity, Johnson City, Tennessee 37614, Department of Chemistry,
Loyola UniVersity, Chicago, Illinois 60626, Institute of Biophysics, Academy of Sciences of the Czech Republic,
V.V.i. KraloVopolska 135, 612 65 Brno, Czech Republic, Institute of Organic Chemistry and Biochemistry, Academy of Sciences
of the Czech Republic, FlemingoVo nam. 2, 166 10 Prague, Czech Republic, and Department of Physical Chemistry,
Palacky UniVersity, Tr. SVobody 26, 77146 Olomouc, Czech Republic
ReceiVed May 6, 2007; ReVised Manuscript ReceiVed July 11, 2007
ABSTRACT: A systematic spectroscopic and computational study was conducted in order to probe the
influence of base sequences on stacked (S) versus B-type (B) conformational heterogeneity induced by
the major dG adduct derived from the model carcinogen 7-fluoro-2-aminofluorene (FAF). We prepared
and characterized eight 12-mer DNA duplexes (-AG*N- series, d[CTTCTAG*NCCTC]; -CG*N- series,
d[CTTCTCG*NCCTC]), in which the central guanines (G*) were site-specifically modified with FAF
with varying flanking bases (N ) G, A, C, T). S/B heterogeneity was examined by CD, UV, and dynamic
19F NMR spectroscopy. All the modified duplexes studied followed a typical dynamic exchange between
the S and B conformers in a sequence dependent manner. Specifically, purine bases at the 3′-flanking site
promoted the S conformation (G > A > C > T). Simulation analysis showed that the S/B energy barriers
were in the 14-16 kcal/mol range. The correlation times (τ ) 1/κ) were found to be in the millisecond
range at 20 °C. The van der Waals energy force field calculations indicated the importance of the stacking
interaction between the carcinogen and neighboring base pairs. Quantum mechanics calculations showed
the existence of correlations between the total interaction energies (including electrostatic and solvation
effects) and the S/B population ratios. The S/B equilibrium seems to modulate the efficiency of Escherichia
coli UvrABC-based nucleotide excision repair in a conformation-specific manner: i.e., greater repair
susceptibility for the S over B conformation and for the -AG*N- over the -CG*N- series. The results
indicate a novel structure-function relationship, which provides insights into how bulky DNA adducts
are accommodated by UvrABC proteins.
A mutation is defined as a heritable change in genome
sequence and may occur as a result of an imbalance between
repair and replication (1). Most mutagens and carcinogens
are metabolized in ViVo into reactive electrophiles, which
subsequently react with cellular DNA to produce DNA
adducts (2). Relating adduct formation with a specific
mutation at the molecular genetic level is a challenging task
(3-7). Earlier efforts to simply connect the types and extent
of DNA adduct formation with mutations have largely been
unsuccessful. This is due in part to an oversimplified view
that damaged DNA adopts a single major structure that is
responsible for a specific mutation (6, 8). Certain adducts
derived from bulky carcinogens adopt multiple DNA con-
formations, and the resulting heterogeneities are modulated
by the nature of the carcinogen as well as the base sequence
contexts surrounding the lesion (8-12). Small energy
differences among conformers could shift the adduct popula-
tion balance, consequently influencing the choice of incoming
dNTP within the active site of a polymerase (8). Since
mutation is inherently an infrequent biological event, ob-
served mutations could happen due to replication of certain
minor adduct conformers that have escaped the repair process
(6).
Arylamines are an important group of bulky mutagens,
which include heterocyclic amines found in overcooked foods
and cigarettes (13). The prototype arylamine, 2-aminofluo-
†We are grateful to the NIH (R01CA098296 to B.P.C. and
R01CA86927 to Y.Z.) for their financial support for this work. This
research was made possible in part by the use of the RI-INBRE
Research Core Facility supported by the NCRR/NIH (P20 RR016457).
J.S. was supported by Grants LC06030, MSM0021622413, and
AVOZ50040507 by Ministry of Education of the Czech Republic. P.J.
was supported by grants (MSM6198959216 and LC512) from the
Ministry of Education of the Czech Republic.
∇Dedicated to Professor Chang Kiu Lee on the occasion of his 60th
birthday.
* To whom correspondence should be addressed. Tel: 401 874 5024.
Fax: 401 874 5766. E-mail: bcho@uri.edu.
‡University of Rhode Island.
§East Tennessee State University.
|Loyola University.
⊥Institute of Biophysics and Institute of Organic Chemistry and
Biochemistry, Academy of Sciences of the Czech Republic.
#Palacky University.
11263
Biochemistry 2007, 46, 11263-11278
10.1021/bi700858s CCC: $37.00 © 2007 American Chemical Society
Published on Web 09/18/2007

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rene, and its derivatives produce two major C8-substituted
dG adducts: AF1and AAF in ViVo (Figure 1a) (13, 14).
Despite structural similarities, they exhibit unique mutation
and repair activities and have been subjected to extensive
structure-activity studies (13). Translesion synthesis of AF
adducts is achieved with high fidelity polymerases, whereas
replication of AAF adducts requires specialized bypass
polymerases. Accurate nucleotide incorporation has been
shown to be feasible, albeit slow, opposite the AF lesion in
the active site of a DNA polymerase I Bacillus fragment,
but not opposite the rigid AAF adduct which blocks
nucleotide incorporation (15). In a similar study involving
T7 DNA polymerase, unlike the AAF adduct, low electron
density was observed around the AF moiety in the poly-
merase active site, suggesting conformational flexibility even
in the solid state (16). Solution structures of the N-
deacetylated AF adduct have been studied extensively with
various DNA sequence contexts (17-31). The AF in fully
paired DNA duplexes is in sequence dependent equilibrium
between the external B-type (B) and stacked (S) conforma-
tions, as defined by its location (major groove and base-
displaced, respectively) or the glycosidyl (?) configurations
(anti- and syn-, respectively) of the modified dG (Figure 1b).
A minor groove binding wedged (W) conformer has also
been observed in duplexes in which the lesion is mismatched
with purine bases (23, 24, 30). The dynamics of the AF-
induced B/S/W heterogeneity have been shown to be
modulated by both the base sequence contexts and the length
of primers, and contribute to polymerase activity through a
long-range effect (31).
Probing the base sequence effects is complicated due to a
delicate balance among various contributing chemical forces.
Aromatic base stacking is the most likely factor for deter-
mining the local sequence dependent conformational vari-
ability within the double helical DNA; however, the rela-
tionship is complex (32). Intrinsic base stacking (the neat,
direct base-base forces) is a relatively simple interaction
dominated by common van der Waals terms modulated by
electrostatics and steric factors (33). Nucleobases possess
dipolar electrostatic distribution and thus prefer large van
der Waals overlaps between coplanar aromatic rings. The
nature of intercalator-base stacking is very similar to base-
base stacking (34). However, capturing selectively the role
of DNA stacking is very difficult since the stacking associa-
tion arises not just from the intrinsic base stacking energy
terms, but from their complex interplay with many other
forces including solvation effects (35, 36). The overall
balance of forces varies with environment changes and
nucleic acid structure (37, 38).
Nucleotide excision repair (NER) is the major cellular
pathway for removal of a wide variety of bulky lesions,
otherwise, mutagenesis occurs (39-41). Sequence depen-
dence of the efficiencies of DNA repair and replication may
account for the presence of mutational “hot spots”, such as
the Escherichia coli NarI exonuclease sequence (5′-CG1G2-
CG3C-3′) (11). Extensive efforts have been directed toward
understanding the detailed molecular mechanisms by which
bulky adducts are recognized in both bacterial and mam-
malian NER systems. Although the human NER proteins
show no homology to the prokaryotic proteins, the overall
strategy is the same: recognition, helix unwinding, incision,
and patch (41). The hallmark of NER is its ability to repair
a wide variety of lesions. In E. coli NER, the DNA damage
1Abbreviations: NER, nucleotide excision repair; AF, aminofluorene
adduct (N-[deoxyguanosin-8-yl]-2-aminofluorene); B conformer, B-type
external groove-bound conformer; CD, circular dichroism; DFT, density
functional theory; FAF, fluoroaminofluorene adduct (N-[deoxyguanosin-
8-yl]-7-fluoro-2-aminofluorene);19F NMR,19F nuclear magnetic reso-
nance spectroscopy; QM, quantum mechanics; LC-TOF-MS, liquid
chromatography time-of-flight mass spectrometry; S conformer, stacked
base-displaced conformer; VDW, van der Waals.
FIGURE 1: (a) Chemical structures of C8-substituted dG-aminofluorene adducts. AF adduct, N-(2′-deoxyguanosin-8-yl)-2-aminofluorene;
AAF adduct, N-(2′-deoxyguanosin-8-yl)-2-acetylaminofluorene; FAF adduct, N-(2′-deoxyguanosin-8-yl)-7-fluororo-2-aminofluorene. (b)
The major groove views of the central trimer segments of the B and S conformers of the AF-modified -AG*A- duplex, as an example. The
modified dG and the complementary dC are shown in red and green, respectively, and the AF moiety is highlighted with gray CPK. In the
B conformer, anti-[AF]dG maintains Watson-Crick hydrogen bonds, thereby placing the AF ring in the major groove. The AF moiety of
the S conformer stacks into the helix with the modified dG in the syn conformation. (c) Sequence contexts of the -AG*N- and -CG*N-
duplex series (G* ) FAF adduct, N ) G, A, C, T).
11264 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.

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recognition has been shown to be modulated by the adduct
size and structure, as well as the neighboring base sequences.
This has led to several intriguing recognition hypotheses such
as “multipartite” and “indirect read” models (42, 43).
Similarly dramatic sequence effects have also been observed
on mutational outcomes in both bacterial and mammalian
cells (44). These results clearly reinforce the importance of
understanding flanking sequence effects and the delicate
balance between repair and mutation. The inherent sequence
dependence of the AF-induced S/B/W heterogeneity could
be a key driving force to various repair and mutational
outcomes.
In the present study, we have prepared a series of 12-mer
DNA duplexes in which the fluorine probe 7′-fluoro-2-
aminofluorine (FAF) (Figure 1) is site-specifically incorpo-
rated at the central guanines and the immediate flanking bases
are varied systemically. The utility of FAF as a useful
fluorine structure probe has been well documented (28-31).
Incorporation of a fluorine atom at the remote C7 position
of AF has been shown to maintain similar carcinogenicity
in various rat tissues including the liver (8). The profiles of
DNA adduct formation and optical behavior of AF and FAF
are also similar. The S/B conformational heterogeneities of
these modified duplexes were examined by CD and UV
melting curve experiments, as well as dynamic19F NMR
spectroscopy. The van der Waals (VDW) energy and
quantum mechanics (QM) calculations were carried out in
order to gain theoretical insight into the observed sequence
effects on S/B heterogeneities. We also measured the NER
activities of the modified sequences in the E. coli UvrABC
system, and the results were examined in terms of the S/B
population ratios obtained from the dynamic19F NMR data.
Our results showed that both the base-carcinogen stacking
interaction and solvation play a critical role in the sequence
effect on the AF heterogeneity. The19F NMR/NER correla-
tions suggested that the S conformer is more susceptible to
repair than the B conformer, and that excision efficiency is
preferred for those with dC over dA on the 5′-side of the
lesion. Taken together, the results provide novel insights into
how bulky carcinogen-DNA adducts are accommodated by
NER proteins in a conformationally specific manner. A
preliminary account of the present work has been com-
municated (45).
EXPERIMENTAL PROCEDURES
CAUTION! 2-Aminofluorene deriVatiVes are mutagens
and suspected human carcinogens and must be handled with
caution.
Crude oligodeoxynucleotides in 10-15 µmol scales in
desalted form were obtained from Sigma-Genosys (The
Woodlands, TX). All HPLC solvents were purchased from
Fisher Inc. (Pittsburgh, PA).
Synthesis and Purification of FAF-Modified Oligodeoxy-
nucleotides. We prepared two sets of 12-mer oligonucleotides
(namely, -AG*N- series, d[CTTCTAG*NCCTC]; -CG*N-
series, d[CTTCTCG*NCCTC]), in which the FAF-modified
guanines (G*) are flanked by four natural bases (N ) G, A,
C, T] (Figure 1c). Additional FAF adduction occurred when
N was G as in the -AGG*- and -CGG*- sequences.
Preparation of FAF-modified oligodeoxynucleotides was
conducted using the procedures described previously (28,
29). Briefly, unmodified oligodeoxynucleotides were treated
with N-acetoxy-N-(trifluoroacetyl)-7-fluoro-2-aminofluorene
in a pH 6.0 sodium citrate buffer and placed in a shaker for
18-24 h at 37 °C (Supporting Information (SI) Figure S1).
Typical HPLC profiles and their on-line UV spectra are
shown for the -CGG- sequence (Supporting Information
Figures S2, S3). The early eluting peak at 19.5 min is that
of an unreacted oligodeoxynucleotide. Two late eluting ones
at 32.8 and 36.7 min exhibited shoulder absorptions in the
300-350 nm range, which are characteristic for modification
at the C8 of dG. Purification of the modified oligodeoxy-
nucleotides up to >97% purity was accomplished by repeated
injections onto reverse phase HPLC (SI Figure S2). The
HPLC system consisted of a Hitachi EZChrom Elite unit
with a L2450 diode array as a detector and employed a
Waters XTerra MS C18 column (10 × 50 mm, 2.5 µm) with
a 60 min gradient system involving 3 to 15% acetonitrile in
pH 7.0 ammonium acetate buffer (0.10 M) with a flow rate
of 2.0 mL/min. A typical yield for modification was in the
range of 20-50% depending on sequences.
Enzyme Digestion/HPLC. The FAF-modified 12-mer oli-
gonucleotides were determined to be C8-substituted guanine
adducts by a standard sequential enzyme digestion procedure
to 2′-deoxynucleosides and cochromatography with the
standard FAF adduct, N-[deoxyguanosin-8-yl]-7-fluoro-2-
aminofluorene (28). Each sample (∼1 OD) was dissolved
in 3 mL of Bis-Tris- EDTA (pH 7.0) buffer and 0.5 mg of
DNase and incubated at 37 °C with shaking for 3-4 h. Snake
venom phosphodiesterase I (100 µL, ∼0.05 unit) and alkaline
phosphatase (∼2.5 unit) were then added, and the incubation
was continued for 1-2 days. The sample was loaded on a
Millipore Centricon YM-3 centrifugal filter (Yellow, MW
cutoff ) 3000) and centrifuged, and the filtrate was extracted
three times with H2O-saturated 1-butanol. The butanol
fractions were combined, back-washed with water, and then
evaporated to dryness using a SpeedVac (Thermo Savant).
The sample was dissolved in methanol and subjected to
reverse phase HPLC. Supporting Information Figure S3
shows a typical HPLC chromatogram (-CG*G- sequence)
of the resulting digests and the HPLC conditions.
Sequence Analyses by Enzyme Digest/LC-TOF-MS. The
FAF-modified oligonucleotides were sequenced using a
Waters ESI-time-of-flight-mass spectrometer (LC-TOF-MS)
in the negative ion mode based on the exonuclease (3′-5′ or
5′-3′) strategies described previously (29). Briefly, 5-10 µg
of FAF-modified oligodeoxynucleotides was combined with
ca. 0.01 unit of a 3′-5′- or a 5′-3′-exonuclease in a 1 mM
solution of MgCl2and incubated for different times up to 2
h. Enzyme digests were separated online using a 10 min
water/acetonitrile gradient. The aqueous phase was 5 mM
in both ammonium acetate and dimethylbutylamine (pH 5.6),
and the acetonitrile was made 0.1% in formic acid. The spray
voltage was 3.4 kV. Illustrative assignment details are given
in Supporting Information Figures S4-S7 for the FAF-
modified -CG*G- and -CGG*- sequences. The molecular ion
results of all the eight FAF-modified sequences used in the
present study are summarized in SI Table S1.
UV-Melting Experiments. UV-melting data were obtained
using a Beckman DU 800 UV/vis spectrophotometer equipped
with a 6-chamber, 1 cm path length Tmcell, and a Peltier
temperature controller, according to the procedure described
previously (29). Duplex solutions (0.2-14 µM) were pre-
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11265

Page 4

pared in a pH 7.0 buffer containing 0.2 M NaCl, 10 mM
sodium phosphate, and 0.2 mM EDTA. Thermodynamic
parameters were calculated using the program MELTWIN
version 3.5.
Circular Dichroism (CD) Spectra. CD measurements were
conducted on a Jasco J-810 spectropolarimeter equipped with
a variable Peltier temperature controller (29). Typically, 2
ODS of each strand was annealed with an equimolar amount
of a complementary sequence in 400 µL of a neutral buffer
containing 0.2 M NaCl, 10 mM sodium phosphate, 0.2 mM
EDTA. Spectra were acquired using a 1 mm path length cell.
NMR Experiments. Approximately 50-80 ODS of pure
modified oligonucleotides were annealed with an equivalent
amount of complementary sequences to produce the corre-
sponding fully paired 12-mer duplexes. The duplex samples
were centrifugated using a Pall Microsep MF centrifugal
device (Yellow, MW cutoff ) 1000). The centrifuged
samples were dissolved in 300 µL of a neutral buffer (10%
D2O/90% H2O containing 100 mM NaCl, 10 mM sodium
phosphate, and 100 µM tetrasodium EDTA, pH 7.0) and
filtered into in a Shigemi tube through a 0.2 µm membrane
filter.
All1H and19F NMR results were recorded using a 5 mm
19F/1H dual probe on a Bruker DPX400 Avance spectrometer
operating at 400.0 and 376.5 MHz, respectively. Imino proton
spectra were obtained using phase sensitive jump-return
sequences at 5 °C and referenced relative to DSS (SI Figure
S8).19F NMR spectra were acquired in the1H-decoupled
mode and referenced to CFCl3by assigning external C6F6
in C6D6 at -164.90 ppm. Long-term acquisition with1H-
coupling did not improve signal-to-noise ratios. One-
dimensional19F NMR spectra were measured between 5 °C
and 60 °C with increments of 5 and 10 °C. Additional
temperatures were used as needed to clarify signal exchange
process (see figure legends). Temperatures were maintained
by a BRUKER-VT unit using liquid N2. Spectra were
obtained by collecting 65,536 points using a 37,664 Hz
sweep width and a recycle delay of 1.0 s. A total of 1600
scans were acquired for each dynamic NMR spectrum. High
quality spectra at 20 °C were obtained in separate acquisitions
using 4000 scans (Supporting Information Figure S9). All
FIDs were processed by zero-filling, exponential multiplica-
tion using a 20 Hz line broadening factor and Fourier
transformation. The S/B population ratios were obtained by
area integration of the base-line corrected spectra (Bruker,
Billerica, MA). NOESY/exchange19F NMR spectra were
obtained in the phase-sensitive mode using the following
parameters: sweep width 4529 Hz, number of complex data
points in t21024, number of complex FIDs in t1256, number
of scans 96, dummy scans 16, recycle delays 1.0 s, and
mixing time 400 ms. The data were apodized with sine
function using 2 Hz line broadening in both dimensions and
Fourier transformed with the 1024 × 256 data matrix.
Complete Line Shape Analysis. Complete line-shape
analysis was carried out using the simulation program
WINDNMR-Pro (version 7.1.6, J. Chem. Educ. Software
Series; Reich, H. J., University of Wisconsin, Madison, WI).
Examples of simulation are shown in Supporting Information
Figure S10 for the -AG*N- duplex series. The values of
frequencies and the S/B-population ratios were determined
at the slow exchange limit (5 °C). Subsequently, several
spectra were recorded at various temperatures between 5 and
60 °C including at coalescence and into the fast exchange
region. The samples were then cooled back to the slow
exchange limit to ensure that no irreversible process have
occurred at the higher temperatures (46).
Modeling. The NMR structures of the S- and B-conform-
eric AF-modified 11-mer duplexes (19, 20) were adjusted
to that of the target 12-mer sequence (Figure 1c). The
hydrogen atom on C7 of AF was replaced with a fluorine
atom (Supporting Information Figure S11). The additional
base pair was added with Insight II (Accelrys Software, Inc.),
using classic B-DNA conformation as a guide. Partial charges
in S conformer were obtained with Gaussian 03 (Gaussian,
Inc.) (47). Force field parameters for FAF-dG were obtained
by finding the closest analogies in the parm99 (48) and
GAFF (49) force fields, and full details are given in
Supporting Information Table S5. HF calculations with the
6-31G* basis set were used to calculate the electrostatic
potential using Gaussian 03 (50), and the restrained elec-
trostatic potential (RESP) fitting algorithm was employed
to fit the charge to each atom center (47). Full details of
RESP results are given in Supporting Information Tables
S6 and S7. Molecular dynamics (MD) simulations were
carried out with AMBER 8.0, employing the modified
Cornell et al. force field parm99 version (51).
Van der Waals (VDW) Energy Analysis. 5 ns MD
simulation was carried out for each system. The MD
structures stabilized after 2 ns with the 2-5 ns time frame
used for analysis. The unstacked complementary dC of the
S conformer displaced into the major groove was excluded
from the calculation, similar to the FAF moiety in the major
groove of the B conformer. The VDW stacking interactions
were computed with the program ANAL of AMBER 8. The
stacking interaction energies between G-FAF and its adjacent
Watson-Crick base pairs were evaluated for both S and B
conformers. A total of 3000 snapshots were utilized. MOIL-
view was used to do a cluster analysis of the 2-5 ns
trajectory to identify the most representative structures (52),
representing the lowest average root-mean-square deviations
(rmsd) to all of the others in the cluster. The average values
and standard deviations are given in Supporting Information
Tables S3 and S4.
Quantum Mechanics (QM) Calculations
Geometries. QM calculations were carried out based on
MD structures. The G*:C modified pair and its two flanking
base pairs were dissected for QM calculations, and all
nucleobases were terminated by hydrogen atoms at the
glycosidic positions (Figure 2). Each “AMBER” nucleobase
geometry was replaced (overlay via rms fit using heavy
atoms) with an in vacuo QM-optimized planar monomer.
Then, an additional constrained gradient optimization was
carried out for the hydrogen-bonded base pairs (each pair
separately) with the positions of the heavy atoms frozen and
the hydrogen atoms relaxed. Optimizations were carried out
in Gaussian 03 (50) using B3LYP (53) density functional
and 6-311G(d,p) basis set.
Interaction Energy Calculations. Interaction energy ∆EAB
of a dimer (the individual base pair stack or pair) A-B is the
difference between the electronic energy of the dimer EAB
and the electronic energies of the infinitely separated
monomers, i.e., ∆EAB) EAB- EA- EB(33, 36).
11266 Biochemistry, Vol. 46, No. 40, 2007
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The total interaction energy for a given conformer and
sequence was calculated as a sum of dimer interaction
energies considering all pairs of interacting monomers. Thus,
the total interaction energy of the -AG*C- sequence in its
S-conformer ∆E(S) (Figure 2) was obtained as a sum of nine
interactions: A-T, A-G*[FAF], T-G*[FAF], T-C*, G*[FAF]-
C*, G*[FAF]-C, G*[FAF]-G, C*-G, and C-G (C* stands for
C opposite the adduct). The in vacuo (intrinsic) interaction
energies were calculated using the DFT-D method with
empirical dispersion energy correction (54) with TPSS
exchange-correlation functional (55), 6-311++G(3df,3pd)
basis set and the B-0.96-27 dispersion parameters using
TurboMole 5.7 code (56). The gas phase interaction energy
calculations were followed by evaluation of the solvation
energy of the whole model (three base pairs) with continuum
solvation calculation (C-PCM method with the B3LYP/6-
31G* solute description) using the Gaussian 03 program (50).
Interactions in the corresponding B structures, ∆E(B), were
determined in the same way, and the interaction energy
difference between S and B conformers was calculated as
∆∆E(S-B) ) ∆E(S) - ∆E(B). We present only the
differences ∆∆E(S-B) while all dimer interaction terms and
solvation energies are given in SI Table S8. The main sources
of possible errors in the QM calculations are the approxima-
tions of the solvation model. However, these errors for S
and B conformers should largely cancel out to give reason-
able S-B differences, provided the MD simulations are
sufficiently representative.
Nucleotide Excision Repair Assays
NER assays were conducted using the E. coli UvrABC
nuclease system. DNA duplex substrates were constructed
by ligating an AF- or FAF-12-mer (CTTCTAG*NCCTC),
with a flanking 20-mer (GACTACGTACTGTTACGGCT)
and a 19-mer (GCAATCAGGCCAGATCTGC) oligonucle-
otide on the 5′ and 3′ sides, respectively (57-59). The 20-
mer was 5′-terminally labeled with32P. The 5′-terminally
labeled DNA substrates (2 nM) were incised by UvrABC
(UvrA, 15 nM, UvrB, 250 nM, and UvrC, 100 nM) in the
UvrABC buffer (50 mM Tris-HCl, pH 7.5, 50 mM KCl,
10 mM MgCl2, 5 mM DTT, 1 mM ATP) at 37 °C in a time-
course dependent manner (60).
Quantitative data of radioactivity were obtained using Fuji
FLA-5000 Image Scanner. The amount of DNA incised (in
pmol) by UvrABC was calculated based on the total molar
amount of DNA used in each reaction and the percentage of
radioactivity in the incision products as compared to the total
radioactivity. At least three independent experiments were
performed for determination of the rates of incision.
RESULTS
Circular Dichroism
Circular dichroism (CD) results show sequence-dependent
differences in the S/B population ratios. Figures 3a and 3b
show overlays of CD spectra of the FAF-modified -AG*N-
and -CG*N- duplex series, respectively. The profiles of FAF-
induced ellipticities in the 290-360 nm range (ICD290-360nm)
were uniquely sequence dependent, similar to those observed
for the non-fluoro AF duplexes reported previously (29).
Table 1 summarizes the CD patterns in terms of the CD-/+
values, which have been defined as the ratio of the negative
over the positive areas between 250 and 360 nm (29). The
results indicated that all the FAF duplexes adopted global
B-DNA conformations. However, they differed in the extent
of the carcinogen-induced ellipticity between 290 and
360 nm: i.e., the greater the CD-/+values, the greater the
population of B-type conformer (30). In particular, the
sequences with a thymine base 3′ to the lesion exhibited the
largest negative induced shifts, 0.85 and 0.39, for -AG*T-
and -CG*T- duplexes, respectively. These CD results were
in accordance with the19F NMR and van der Waals energy
(VDW) data (see below). The general similarity of the CD
patterns between the FAF and AF duplexes indicated that
the replacement of the remote C7 hydrogen with fluorine
did not change the overall adduct conformation. Thus, FAF
was an excellent probe for conformational analysis of AF
by19F spectroscopy.
UV-Thermomelting Data
As in the CD experiments, the thermodynamic profiles
(Table 1) of the FAF-modified duplexes were found to be
similar to those of the AF-analogues. The effects of the FAF
lesions on the thermal (∆Tm) and thermodynamic (-∆∆G°
and -∆∆H°) stabilities were clearly sensitive to the im-
mediate flanking base sequences (61). An excellent correla-
tion (R2) 0.90) (not shown) between -∆G° and Tm
indicated that the modified duplexes followed an adduct-
induced destabilization of the DNA secondary structure at
or near the adduct site due to the loss of base stacking and
base pairing at or near the lesion site. As expected, the duplex
stability was typically sensitive to the nature and the polarity
of base pairs neighboring the lesion site (29, 62). Therefore,
the Tmand -∆G° values of duplexes with a 3′-flanking G:C
base pair were consistently higher than those with an A:T
pair (i.e., -AG*A-, -AG*T-, -CG*A-, and -CG*T-). Conse-
quently, the -CG*N- duplexes exhibited consistently greater
Tmand -∆G° stabilities than the -AG*N- series (Table 1).
These results indicated the importance of the total number
of hydrogen bonds available at the flanking region for duplex
stabilization. In the case of the AF analogues, we reported a
general inverse trend (R2) 0.675-0.826) between Tm/-
∆G° and % B conformer and suggested the importance of
the carcinogen stacking interaction in the enhanced stability
of the S conformer (29). No such correlations were observed
for the FAF-modified duplexes in the present study. The
reason for this discrepancy is not clear and could be complex.
It seems that a major contributing factor is the aforemen-
FIGURE 2: A model for quantum mechanic calculations of the S
conformer -AG*C- duplex. G*-FAF, FAF-modified G; C*, comple-
mentary C opposite the lesion.
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11267