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.
- SourceAvailable from: fundacion-barcelo.com.ar[show abstract] [hide abstract]
ABSTRACT: The early notion that cancer is caused by mutations in genes critical for the control of cell growth implied that genome stability is important for preventing oncogenesis. During the past decade, knowledge about the mechanisms by which genes erode and the molecular machinery designed to counteract this time-dependent genetic degeneration has increased markedly. At the same time, it has become apparent that inherited or acquired deficiencies in genome maintenance systems contribute significantly to the onset of cancer. This review summarizes the main DNA caretaking systems and their impact on genome stability and carcinogenesis.Nature 06/2001; 411(6835):366-74. · 38.60 Impact Factor
- [show abstract] [hide abstract]
ABSTRACT: The roles of genetic constitution versus environmental factors in cancer development have been a matter of debate even long before the discovery of 'oncogenes'. Evidence from epidemiological, occupational and migration studies has consistently pointed to environmental factors as the major contributing factors to cancer, so it seems reasonable to discuss the importance of chemical carcinogenesis in the present 'age of cancer genetics'.Nature reviews. Cancer 03/2005; 5(2):113-25. · 35.00 Impact Factor
- Chemical Reviews 03/2006; 106(2):420-52. · 41.30 Impact Factor
Spectroscopic and Theoretical Insights into Sequence Effects of
Aminofluorene-Induced Conformational Heterogeneity and Nucleotide Excision
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
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
* To whom correspondence should be addressed. Tel: 401 874 5024.
Fax: 401 874 5766. E-mail: firstname.lastname@example.org.
‡University of Rhode Island.
§East Tennessee State University.
⊥Institute of Biophysics and Institute of Organic Chemistry and
Biochemistry, Academy of Sciences of the Czech Republic.
Biochemistry 2007, 46, 11263-11278
10.1021/bi700858s CCC: $37.00© 2007 American Chemical Society
Published on Web 09/18/2007
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.
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
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-
CAUTION! 2-Aminofluorene deriVatiVes are mutagens
and suspected human carcinogens and must be handled with
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
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
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
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
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
Meneni et al.
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
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.
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
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
tioned flanking sequence effect, not the AF/FAF swap, which
has been demonstrated to be conformationally compatible
at various sequence contexts (8).
19F NMR Spectroscopy
Signal Assignments. The S/B19F signal assignments were
made using the general strategy described previously (28,
30, 31). Figure 4 shows dynamic19F NMR spectra of the
-AG*N- and -CG*N- duplexes recorded in the -115 to -121
ppm range between 5 and 60 °C. All eight FAF-modified
duplexes exhibited two major well-resolved19F signals at
or below 20 °C, which represented unique19F environments,
namely, the S and B conformation (see below). S/B
heterogeneity was evidenced by the varying intensities of
the imino proton signals in the1H NMR spectra (Supporting
Information Figure S8). The low intensity imino proton
signals at 10-12 ppm usually represent those of the modified
dG at the lesion site and the nearby base pairs. The slow
interconversion of the two conformers in each duplex was
confirmed by temperature dependent NOESY/exchange
spectroscopy. For example, Figures 5a and 5b show the
NOESY/exchange spectra of the -AG*T- duplex taken at 5
and 20 °C, respectively. Strong temperature dependence of
the off-diagonal cross-peaks implied a transfer of magnetiza-
tion from one peak to another, indicating that the two
conformers (S and B) originated from the same chemical
structure (28). Similar NOESY profiles were observed for
all the duplexes studied in the present study.
Specific S/B signal assignments were made based on ring
current and H/D isotope effects (28, 31). The S conformer
was more susceptible to a ring current effect, which enabled
the base-displaced FAF to be better shielded than the B
conformer. The rationale for the H/D effect was that the19F
resonance of the exposed FAF residue in a B conformer is
more susceptible to solvent-induced shielding (usually
>0.2 ppm) than the buried FAF in an S conformer (usually
<0.1 ppm) upon increasing the deuterium content from 10%
to 100% (63). The H/D isotope effects were measured at 20
and 60 °C for all the duplexes studied, and the results are
summarized in Supporting Information Table S2. In all cases,
the downfield B conformer signals exhibited consistently
greater H/D shielding effects (0.13-0.22 ppm) than the
upfield S conformer signals (0.01-0.06 ppm) at 20 °C.
Sequence-Dependent S/B Heterogeneity. S/B heterogeneity
was markedly sequence-dependent, especially on the 3′-
flanking bases. Supporting Information Figure S5 compares
the19F NMR spectra of the -AG*N- and -CG*N- duplex
series acquired at 20 °C in a stacked format. Specifically,
the population of S conformer for the -AG*N- series
decreased in order of 3′-dG (68%) > dA (61%) . dC (40%)
FIGURE 3: CD spectral overlays of the FAF-modified duplexes in the 200-400 nm region at 15 °C for (a) the -AG*N- and (b) -CG*N-
Table 1: Effects of FAF-modification on the Thermal and Thermodynamic Stability of DNA Duplexes
-AG*G- 9.8 (12.7) 75.8 (101.0)
-AG*A- 8.2 (11.1)63.5 (87.8)
-AG*C- 9.1 (11.0) 63.4 (74.4)
-AG*T- 8.4 (10.2)68.6 (76.8)
-CG*G-10.2 (13.5) 73.8 (87.8)
-CG*A- 9.4 (11.7)75.2 (77.7)
-CG*C- 10.0 (12.7) 67.2 (74.2)
-CG*T-9.3 (11.7) 72.3 (76.0)
0.20 51.0 (56.2)
45.6 (54.2) 2.924.3
50.4 (56.6) 1.911.0
45.6 (52.8)1.8 8.2
49.2 (59.8) 2.32.6
54.5 (64.4)2.7 7.0
aThe central trimer portion of the 12-mer duplex (G* ) FAF adduct, see Figure 1).bThe results of curve fit and Tm-lnCt dependence were
within (15% of each other, and therefore these numbers are averages of the two methods. The average standard deviations for -∆G°, -∆H°, and
Tmare (0.22, (6.33, and (0.4, respectively. The values in parentheses are for the unmodified control duplexes.cTmvalues at 14 µM taken from
the 1/Tm- lnCt/4 Meltwin plots .d∆∆G ) ∆G° (FAF-modified duplex) - ∆G° (control duplex).e∆∆H ) ∆H° (FAF-modified duplex) - ∆H°
(control duplex).f∆Tm ) Tm (FAF-modified duplex) - Tm (control duplex).gRatio of the negative over the positive areas in the 250-360 nm
region (29) (see Figure 3).
11268 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.
> dT (36%). A similar trend was obtained for the -CG*N-
series, 3′-dG (66%) > dA (56%) ∼ dC (53%) . dT (37%).
Sequence-Dependent19F NMR Chemical Shifts. The19F
signals for the B conformer in both the -AG*N- and -CG*N-
series appeared at less than 0.13 ppm (Supporting Informa-
tion Figure S9 and SI Table S2). This was in contrast to the
S conformer signals in the same contexts, which exhibited
significant deshielding (0.3-0.7 ppm) going from 3′-T to A
or G. The results clearly indicated the sensitivity of the S
conformer19F chemical shifts to the nature of the 3′-flanking
base. The stacking efficiencies between the carcinogen and
the 3′-flanking base play an important role (see below). The
largest chemical shift difference between the two signals was
observed for the -AG*T- duplex.
Thermodynamics of the FAF-Modified Duplex System. An
overall thermodynamic picture of the FAF-modified duplex
system can be characterized by two different equilibriums:
first, the equilibrium between the S and B conformers in
the duplex, and second, a melting equilibrium between the
duplex and single strands. Dynamic19F NMR data (Figure
4) provided valuable kinetic and thermodynamic information
on both equilibria (Table 3, see below). Description of the
thermodynamic diagram and assumptions undertaken in the
present study are given in Supporting Information Figure
S12. The dynamic NMR results showed that the S signal
increased with increasing temperatures, i.e., ∆∆H(S-B) was
positive. The S conformer was enthalpically disfavored, but
entropically favored with respect to the B conformer. Using
the van’t Hoff equation, the values of ∆∆H(S-B) were
estimated to be about 2-6 kcal/mol, which depended on the
sequence contexts. These values may be inaccurate since the
temperature dependence of the S signal was obscured by an
unknown heat capacity difference between the S and B
conformers, and melting of the duplex at higher temperatures.
exhibited a typical two-site chemical exchange behavior
(Figure 4). The S/B19F signals were in slow exchange at or
below 5 °C, but upon raising the temperature they broadened
and moved closer together, and eventually coalesced between
40 and 55 °C. Sharp signals at 60 °C were due to the fast
rotating modified G* FAF residues of denaturing duplexes
(see below), and substantial H/D isotope effects (0.16-
0.20 ppm) were observed (SI Table S2). The B conformer
signals of certain duplexes (i.e., -AG*C-, -AG*T-, and
19F NMR Spectra. All eight FAF duplexes
FIGURE 4: Dynamic19F NMR spectra (-115 to -121 ppm) of the -AG*N- and -CG*N- duplexes (G* ) FAF adduct, N ) G, A, C, T).
Spectra of all sequences were obtained at seven standard temperatures (labeled as 5, 10, 20, 30, 40, 50, and 60 °C). Additional temperatures
were recorded for -AG*G (25, 35, 45, 47 °C), -AG*A- (35, 43 °C), -AG*C- (37 °C), -AG*T- (15, 25, 33, 35 °C), -CG*G- (45 °C), -CG*A-
(25, 35 °C), -CG*C- (45), and -CG*T- (35 °C).
FIGURE 5: Temperature dependent19F NMR NOESY spectra of
the -AG*T- duplex recorded in 10% D2O/90% H2O pH 7.0 buffer
at (a) 5 °C and (b) 20 °C. B ) B-type conformer; S ) stacked
conformer (see Figure 1). Off diagonal cross-peaks are shown in
the dotted circles.
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11269
-CG*T-) were split between 20 and 30 °C, suggesting the
existence of additional B-like conformer intermediates (B*).
A similar observation was made in the primer-template
sequence contexts (31).
It should be noted that for all sequences the intensity of
the downfield fluorine signal initially decreased with increas-
ing temperatures and then increased again. A key to
understanding this behavior is to realize that the downfield
signal belonging to the19F exposed to solvent is a sum of
the signals coming from the B conformer and denatured
modified single strands. This is depicted in Figure 6, which
captures the S/B (XS, red curve) and melting equilibria
(melting curve, black line) for the -AG*G- duplex as modeled
by the van’t Hoff equation. The diagram is only an
illustration of principle, and the thermodynamic character-
istics used to generate it are approximate and were chosen
to resemble the steepness of the XScurve at low temperatures
(e. g., ∆∆H(S-B) ∼5 kcal/mol); the real behavior in19F
NMR (Figure 4) is somewhat obscured by nonideal DNA
melting etc. The blue and orange curves represent signals
of19F exposed to solvent (XB) and shielded (XS) by the
flanking base pairs, respectively. At low temperatures, the
ratio of the two signals corresponded primarily to the S/B
equilibrium, whereas at high temperatures it represented
exclusively the melting process.
We conducted lineshape analysis by assuming two-site
exchange for all the dynamic19F NMR spectra (Figure 4,
see Supporting Information Figure S10) (46). The resulting
kinetic and thermodynamic parameters are summarized in
Table 2. The S/B interconversion energy barriers (∆Gq)
obtained from the Eyring equation, ∆GqC ) 4.58TC(10.32
+ log TC/kC) cal/mol, at 30 °C was found to be 14-15 kcal/
mol. The energy differences (∆G° ) -RT ln Keq) of the
two conformers were relatively small (<500 cal/mol). The
coalescence temperature (TC) is defined as the temperature
at which the two signals are merged into a single, flat-topped
peak (46). Determination of exact TCvalues for each duplex
was difficult since sufficient numbers of dynamic spectra
were not obtained. A close inspection of emerging signals,
however, indicated that TCdecreased in the order of -CG*C-
> -CG*G- > -AG*G-, -CG*A- > -AG*A-, -AG*C-,
-AG*T-, -CG*T-. These results were consistent with the Tm
trend (Table 1) and appeared to be in line with the number
of Watson-Crick hydrogen bonds surrounding the lesion
site, i.e., the -CG*C- and -CG*G- sequences have three
hydrogen bonds on both sides of the lesion (Table 1). The
exchange rate constants at TC (kC ) 2.22∆ν, ∆ν ) the
separation in frequency in Hz between the two conformer
signals at 5 °C, when dynamic exchange is minimal) were
determined to be in the range of 739-1715 s-1. The
correlation times (τ ) 1/κ) of the modified duplexes were
found to be in the narrow range of 2.2-5.2 ms at 20 °C and
signify the amount of time the lesion spends in one
conformation before changing to another conformation.
Quantum Mechanics (QM) Calculations
Full computational evaluation of the free energy changes
is unfeasible, due to major methodological uncertainties, even
when resorting to the force field description (36, 38). In
addition, the variation of % S conformer corresponded to a
few tenths of a kcal/mol of the free energy change between
S and B (Table 2). Therefore, instead of evaluating the ∆G
directly we focused on some of the key contributions to the
free energies, with the aim to obtain theoretical insight into
the forces governing the S/B equilibrium. Fairly accurate
quantum mechanical calculations can be performed to
evaluate interaction (base stacking and base pairing) and
solvation energies, which can be correlated with the interac-
tion enthalpy or other measurable quantities.
The NMR S/B Ratios and QM Calculations. First, we
determined whether the ∆∆H(S-B) calculated using the QM
methods correlates with the S/B ratio (more precisely, with
ln(S/B)) from the NMR experiment. The S/B ratio was
determined by the free energy difference between S and B.
The underlying assumption for ∆∆H(S-B) to correlate with
the S/B ratio is that the entropy change between S and B is
similar for all four sequences in a series. If this were the
case, the ∆∆G(S-B) trend would be given primarily by the
enthalpy change ∆∆H(S-B). More details about the model
are given in Supporting Information Figure S12. The results
of our QM calculations and the enthalpy difference between
S and B are summarized in Table 3. The correlation
coefficients (R2) of the calculated interaction energy differ-
ences with ln(S/B) were 0.86 and 0.44 for the -AG*N- and
-CG*N- series, respectively. With inclusion of the continuum
solvent correction, R2were 0.63 and 0.57 for the -AG*N-
and -CG*N- series, respectively. Thus, the S/B ratio is only
marginally correlated with the base interaction energies.
The in Vacuo interaction energy loss (∆∆Eint(S-B)) of B
to S conversion was clearly due to the loss of hydrogen
bonding at the lesion site, which ranged between 17.6 and
Table 2: Dynamic NMR and Thermodynamic Parameters for the
FAF-modified 12-mer Duplexes
sequencea% Bb% Sb
6040 263 3.81240 38815.4
36 64 4352.31097
6337 4462.21321 30914.1
aThe trimer core sequence of the FAF-modified 12-mer duplexes
used in this study. See Figure 1c for full sequence contexts. G* )
FAF adduct.b% population of B and S conformers by integration of
19F NMR signals at 20 °C (SD ) (3%).cRate constant. (SD ( 25%.)
The data were obtained by complete line shape analysis of temperature
dependent19F NMR results using the WINDNMR-Pro (see Experi-
mental Procedures).dExchange time (1/k) indicates the amount of time
the adduct spends in one conformation before jumping into another
conformation.eRate constants at a coalescence temperature, a lower
limit on the exchange rate between the two conformers. kC) 2.22∆V
(the chemical shift difference between the two signals in Hz at slow
exchange, i.e., at 5 °C).fThe energy difference between the two
conformers: ∆G° ) -RT ln K [Keq) S/B].gThe S/B interconversion
energy barrier @ 30 °C obtained from Eyring equation (46): ∆GqC)
4.58TC(10.32 + log TC/kC) cal/mol. SD ( 0.2 kcal/mol.
11270 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.
25.2 kcal/mol for the -AG*N- series and 19.4-24.2 kcal/
mol for the -CG*N- series. This roughly corresponded to
the interaction energy of the G:C pair in Vacuo. The DFT-D
interaction energy can be decomposed to the DFT part (Table
3, column 3) and the dispersion part (column 4). The former
covered the electrostatic interaction, polarization contribution,
and short-range repulsion between interacting molecules. The
latter was the attractive London dispersion interaction which
usually dominates aromatic stacking. Only small differences
in dispersion energies (Table 3, column 4) indicated that the
dispersion interaction lost in the original B duplex was almost
completely compensated by the dispersion stabilization of
stacking of the FAF moiety with its flanking pairs in the S
conformer. Therefore, the S destabilization was due almost
exclusively to the loss of hydrogen bonding.
In an aqueous environment, the energy penalty for S
conformer coming from the loss of hydrogen bonding is
largely diminished because of the hydration gain correspond-
ing to hydration of the guanine and cytosine Watson-Crick
edges exposed to solvent in the S conformation (see Table
3, column 5, and the sum of the energy differences and
hydration corrections in column 6). For instance, in the case
of -CG*T- sequence (last line) the dispersion contribution
to S destabilization in Vacuo is only 0.1 out of 24.2 kcal/
mol. Then, the difference in hydration energies of S and B
is -16.7 kcal/mol in favor of S, and the resulting S energy
destabilization thus falls to 7.5 kcal/mol only. The final
energy destabilization still originates in the lost hydrogen
bonding, but is much smaller than in Vacuo.
Sequence Dependence of the Melting Enthalpy Differences.
Next, we estimated the difference between the melting
enthalpy of the modified and unmodified duplexes and
compared them with van’t Hoff thermodynamic results. The
melting enthalpy was not calculated directly because our
calculations were hampered by our inability to estimate the
enthalpy of the melted single strands, among other issues
(38). However, as explained in Supporting Information
Figure S12, the ∆∆H of melting was approximated by the
enthalpy difference between the S and B conformers: ∆∆H
= ∆∆E(S-B)weighted ) XS∆∆E(S-B), where XS is the
molar fraction of S at the melting temperature. Because we
did not know XS, we used the values at 20 °C. The
corresponding errors should be moderate, and the resulting
∆∆E(S-B) values are given in Table 3 (last column). The
magnitude of the calculated ∆∆E(S-B) corresponded well
with the measured ∆∆H of melting (Table 1), but no
correlations were observed due to significant error with both
the experimental and theoretical values.
The standard deviation of the experimentally measured ∆H
of melting was estimated to be 6.3 kcal/mol and may be
larger for the resulting ∆∆H values. In particular, experi-
mental values above 20 kcal/mol seemed to be overestimated
as they were much larger than both the theoretical and NMR
estimates. Smaller ∆∆H were to be expected from the nearest
neighbor model, in which the ∆H associated with breaking
one base pair is about 9 kcal/mol. If we assume that there is
only one base pair broken in the S structure (i.e., the G[FAF]:
Table 3: Calculated Quantum Mechanics (QM) Stabilization Energies (kcal/mol) of the FAF-Modified DNA Duplexes
-CG*C- 23.3 23.7
-CG*T- 24.2 24.1
-14.2 11.1 4.0
-17.2 2.2 1.4
-16.7 7.5 2.8
aThe trimer core sequence of the FAF-modified 12-mer duplexes used in this study. See Figure 1c for full sequence contexts. G* ) FAF adduct.
bDifference between stabilization energies of S and B conformers calculated by TPSS/6-311++G(3df,3pd)/B-0.96-27 DFT + dispersion method.
cDFT and empirical dispersion components of the total interaction energy difference form the previous column.dDifference between solvation
energies of S and B conformers (S - B), calculated with COSMO/B3LYP/6-31G*, electrostatic contribution only.e∆∆E(S-B) ) ∆∆Eint(S-B) +
∆∆Esolv.f∆∆E(S-B) corrected for the expected population of the S structure at the melting temperature; to be compared with ∆∆H, see text.
FIGURE 6: An illustration of temperature dependence of the S/B
(XS, orange) and melting (melting curve, black) equilibria for the
-AG*G- duplex modeled by van’t Hoff equation. The F-shielded
(yellow) signal represents S conformers. The F-exposed (blue)
signal is a sum of the B conformer and denatured FAF-modified
single strand signals. See text.
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11271
C pair), then the real enthalpy loss should be even smaller,
since the stacking enthalpy of the G:C pair was not
completely lost but was replaced by stacking of the FAF
moiety. Therefore, the ∆∆H values around and below 10
kcal/mol, as those obtained from the QM calculations,
seemed to be more realistic.
Van der Waals (VDW) Base Stacking Interactions
We monitored the pure van der Waals (VDW) energies
by the Amber Lennard-Jones term as sampled in MD
simulations. In all the S conformer duplexes examined, the
G[FAF] stacked well with adjacent base pairs. Table 4 lists
the sum of total stacking stabilization of G[FAF] with 5′-
and 3′-base pairs. The duplexes with 3′-T (-AG*T-, -CG*T-)
in each series were noticeably less stabilized by the VDW
stacking energies. This contrasts the B-type conformers, in
which the VDW energies of the two 3′-T duplexes were
comparable to others, but represented a significant increase
in stacking stabilization relative to the corresponding S
conformers (by +1.43 and +3.57 kcal/mol, respectively).
The VDW results supported a general trend, i.e., the greater
the VDW stacking, the greater the S conformer population.
The full details of the VDW interactions are given in
Supporting Information Tables S3 and S4.
E. coli UVrABC Nucleotide Excision Repair
Control Experiments. The biological and spectroscopic
utilities of 7′-fluoro-2-aminofluorene (FAF) as a fluorine
structure probe have well been documented (28-31). The
CD profiles of FAF-modified DNA duplexes (this study,
Figure 3) are similar to those of the AF ones we reported
previously (29). Control repair experiments were performed
to verify that the incision efficiency was not also affected
by the hydrogen/fluorine swap (45). For that, we employed
the -AGG- sequence (5′-CTTCTAGGCCTC-3′), in which
each of the two central guanines were modified with either
AF or FAF. Incision efficiencies of the resulting four
sequences were found to be comparable under identical
experimental conditions (45). These results confirmed that
the fluorine substitution had little or no effect on the incision
efficiency of the AF adduct.
Incision of the -AG*N- and -CG*N- Series. Figure 7 shows
a plot of the E. coli UvrABC incision efficiency Versus % S
conformer for the eight FAF-modified sequences. The
-AG*N- and -CG*N- series are labeled as red and blue
symbols, respectively. An excellent correlation (R2) 0.93)
was obtained when only the -AG*N- series were considered
(45). Consistently greater incision efficiencies were observed
for duplexes that are predominantly S conformer, the -AG*G-
(68%) and -AG*A- (61%) sequences. However, no such
correlation was found for the -CG*N- series. Consequently,
when eight data points were plotted together, a poor
correlation was obtained. Nevertheless, the trend persisted:
i.e., low NER was associated with low % S conformation
(shaded cyan; -AG*C-, -AG*T-, -CG*T-), and high NER
was associated with high % S conformation (shaded orange;
-AG*G-, AG*A-, -CG*G-, -CG*A-, -CG*C-).
In other words, the lesions with higher S/B ratio appeared
to be more susceptible to NER repair than most of the lesions
with lower S/B ratio (except CG*T). Moreover, the -CG*N-
series exhibited greater incision efficiencies than the -AG*N-
The precise mechanisms by which DNA adducts exert
mutations are of immense interest from the standpoint of
structure-function relationships (2-4). It is known that
certain bulky carcinogen-DNA adducts produce complex
sequence-dependent conformational heterogeneities in DNA
(4, 8-10). A profound sequence effect on mutation has been
reported for a variety of lesions on both bacterial and
mammalian cells (64-69). The key question is how base
sequence context governs equilibria between adduct confor-
mations. Is one sequence context more prone to repair or
miscode during replication than another? Are there unique
or a combination of conformers that are more or less repair
susceptible, and/or more likely to produce mutations upon
replication? Answers to these questions can, in principle, be
gleaned from understanding the nature of chemical forces
necessary to maintain adduct structures/conformations and
their interactions with relevant proteins. The established
sequence dependence of AF-induced S/B heterogeneity make
it uniquely suited for addressing these questions.
Sequence Effects on S/B Conformational Heterogeneity.
We employed eight of the 16 possible flanking sequence
contexts that are modified by the fluorine probe FAF (Figure
1). The19F NMR results (Figure 4) showed that all the fully
Table 4: Van der Waals Base Stacking Interaction Energies
-30.05 ( 2.41
-32.04 ( 2.46
-30.89 ( 2.29
-28.66 ( 3.39
-29.06 ( 2.28
-29.21 ( 2.23
-29.61 ( 2.13
-26.04 ( 3.88
-29.72 ( 1.92
-30.03 ( 1.93
-29.92 ( 2.02
-30.09 ( 1.90
-29.37 ( 2.15
-29.70 ( 2.18
-29.64 ( 2.20
-29.61 ( 2.27
aThe trimer core sequence of the FAF-modified 12-mer duplexes
used in this study. See Figure 1c for full sequence contexts. G* )
FAF adduct.bThe sum of van der Waals interaction energies in kcal/
mol between the trimer containing FAF-G and 3′- and 5′-base pairs.
cTaken from Table 2.
FIGURE 7: Incision efficiencies (% DNA incision/min) of the
-AG*N- (red) and -CG*N- (blue) sequences versus % S conformer
by Escherichia coli UvrABC nuclease.
11272 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.
complementary 12-mer duplexes existed as a mixture of S
and B conformers and their population balances were
modulated by the nature of flanking base sequence contexts.
When measured at 20 °C, the population of S conformer for
both the -AG*N- and -CG*N- series decreased in order of
3′-dG > dA > dC > dT. The results clearly indicated the
importance of a purine base flanking 3′ to the lesion site for
promoting the S conformation. Although data are limited,
the 5′-flanking base did not seem to exert a similar effect
on S/B heterogeneity. For example, the population of S
conformer was similar for -AG*G- and -CG*G-, and for
-AG*A- and -CG*A-.
We carried out three distinct computational analyses to
determine which energy terms correlate with the NMR-based
S/B population ratios. We first determined the pure van der
Waals (VDW) energy (Table 4), which neglects the elec-
trostatic portion of stacking. Not surprisingly, correlations
between the VDW energy difference and the S/B ratios were
found to be poor. Nevertheless, analysis of the individual
VDW contributions suggested that purine bases at the 3′-
flanking position increased stacking energies, thereby favor-
ing the S conformation. Conversely, pyrimidine bases
reduced stacking which promoted the B conformation. The
results were in agreement with the experimental observa-
tions: i.e., the nature of 3′-flanking bases was important for
promoting (i.e., G and A) or reducing (T) stacking energy,
thus yielding greater populations of S and B conformers,
respectively. These results were also in accordance with the
CD data which showed largest negative CD ellipticities for
the -AG*T- and -CG*T- duplexes.
We then evaluated the full intrinsic (direct) base stacking
(or base-base) forces. For this purpose we utilized complete
QM calculations using the latest dispersion-augmented DFT
method (Table 3) (53), which provide a full description of
the stacking (including also electrostatics, induction, and
charge-transfer forces) (33-36). The magnitude of the S/B
energy difference was closer to the expected enthalpy
difference than the VDW analysis, and the correlations
represent improvement over the pure VDW description
discussed above. Finally, we added solvent screening effects
to the QM stacking data. The solvent was approximated by
the QM-based continuum solvent approach because we were
unable to account for the exact shape of the whole DNA
dodecamer (i.e., only the trimer segment was considered).
Upon inclusion of solvation, we achieved an overall agree-
ment between the magnitude of the calculated interaction
and the expected values (Table 1), although the correlation
coefficients were still relatively poor.
Next, let us comment on the inaccuracies in the QM
calculations. Although evaluation of the interaction energy
itself was fairly accurate, especially when considering ∆∆E
(major error cancellation), there were at least two additional
sources of error in our theoretical model. First, the calculated
interaction was critically dependent on the geometries taken
from MD simulations, which can be biased by force field
and limited sampling of the MD, especially for short
simulations. Thus, some simulations may give geometries
that poorly represent the real ones. Perhaps less important
was the error of the solvation model (see for instance the
outlaying solvation energy of -CG*A- in Table 3), which
stems from having only three dissected base pairs for
solvation. However, attempting to include the sugar-
phosphate segments into the calculations would create
insurmountable methodological problems and reduce the
quality of the predictions. In view of all the approximations,
the complexity of the system, and relatively small enthalpy
differences, the observed nonideal correlation was not
surprising. Alternatively, the free energy difference could
be acquired by postprocessing the MD trajectories, using the
MM-PBSA method. Although this method is extensively
used in the literature, it is based on approximations and the
uncertainty of the results would be in the low kcal/mol range
The QM results described above indicate that both
electrostatics and solvation were important sequence effectors
as their sequence variation and magnitude were larger than
the variability of the dispersion energy contribution. Although
inclusion of these factors did not significantly improve the
correlation between the interaction energy (∆∆Etot(S-B))
and the S/B population ratios (Table 3), the enthalpy
differences calculated were similar to the values estimated
from the NMR and thermomelting measurements. Despite
the lack of quantitative correlation, we believe that our
present model, the QM description augmented by solvent
effects, is a good compromise and provides valuable qualita-
tive insights into the sequence effect.
The Dynamics of S/B Conformational Heterogeneity. The
dynamics of S/B heterogeneity were complicated by the
concomitant presence of the S/B equilibrium and the melting
process of the modified duplex. Nevertheless, valuable
kinetic and thermodynamic properties were obtained from
UV melting (van’t Hoff) and19F NMR (dynamic exchange)
experiments. Analysis of the van’t Hoff parameters indicated
that the thermal and thermodynamic stabilities of the FAF-
modified duplexes were dictated primarily by the polarity
and the numbers of hydrogen bonds associated with the
flanking base pairs. The small energy differences (∆G° <
0.5 kcal/mol) observed between the two conformers from
the dynamic19F NMR experiments suggest a possible facile
adduct conformation switch when in contact with repair
proteins or the active pocket of a polymerase (3, 8). Rapid
dynamics of S/B conformational heterogeneity were ob-
served. The exchange rate constants at the coalescent
temperature were found to be faster than average rates of
spontaneous base pair opening of the Watson-Crick base
pairs in B-DNA (69). The highly lipophilic carcinogenic
moiety in the S conformer was not involved with any
hydrogen bonds, but adopted a stable stacked conformation
within the helix.
The S/B exchange occurred in milliseconds and was
significant because it was in the physiological range of DNA
synthesis. Eckel and Krugh (18) have previously coined the
term “mutagenic switch” to describe S/B conformational
heterogeneity. We have shown recently the dynamic presence
of the S/B equilibrium at the replication fork, rationalizing
either error-free or error-prone replication (31). The results
also support a model in which the conformational hetero-
geneities at positions remote from the primer-terminus
influence polymerase function through a long-range DNA-
protein interaction (31). The S/B equilibrium situation is
reminiscent of the extensively studied base flipping by the
DNA repair enzyme uracil DNA glycosylase. An important
distinction is that the inner helix of the S/B model is always
occupied by either the hydrophobic carcinogen (i.e., S
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11273
conformer) or the modified dG (i.e., B conformer). Jiang et
al. (70) have studied the role of hydrogen bonding and steric
effects in uracil DNA glycosylase using difluorophenyl as a
model uracil. They concluded that the hydrophobic moiety
is stabilized within the helix despite a rapid exchange
between the stacked and unstacked states (kC> 1200 s-1),
which could also explain the S/B equilibrium in our
experiments. It has been shown that base-stacking interaction
dominates not only in the duplex stability but also contributes
into the dependence of the duplex stability on its sequence
(71). A recent theoretical study indicated that the fluorescence
quenching effect of 2-aminopurine is more pronounced when
it is stacked with a purine base than with a pyrimidine base,
indicating the importance of extended π-oribital conjugation
(72). The sequence dependent base stacking and unstacking
has been shown to be enthalpic in nature (71). Adduct
formation induced weakened base stacking, and the resulting
conformational flexibility could be an important determining
factor for many repair proteins, including the E. coli UvrABC
system (73, 74).
Nucleotide Excision Repair Versus S/B Conformational
Heterogeneity. The majority of NER studies to date have
assumed that a bulky lesion adopts a single conformation in
the DNA duplex (39, 40, 57-60). Clearly, this is not the
case for the AF adducts which equilibrated dynamically
between the S and B conformations. The sequence depen-
dence of NER has been documented in both bacterial and
mammalian systems (39, 40) and may account for the
presence of mutational “hot spots”, such as the E. coli NarI
exonuclease sequence (5′-CG1G2CG3C-3′) (11). Mekhovich
et al. (75) have shown that the rate of incision of both AF
and AAF adducts in the E. coli UvrABC system is
significantly faster when they are positioned in the NarI
sequence (5′-GGCG*CC-3′) than when located in a random
The key issue here is which conformer, S or B, is
recognized as a defect in the first step of E. coli NER. This
is an important question since S/B interconversion occurs
in physiologically relevant scales (see above). Our present
19F NMR/NER results provided structural/conformational
insights into this very intriguing matter. Control experiments
showed that the incision efficiencies of two different AF
adduct sequences were not affected by the presence of
fluorine, thus justifying the use of FAF as repair probes (45).
It has been shown previously that the exclusively stacked
AAF adduct is repaired more readily than the conforma-
tionally flexible AF adduct (39, 40). The S conformation,
which lacks Watson-Crick base pairs at the lesion site, is
also structurally similar to the base-displaced cis-benzo[a]-
pyrene-N2-dG (BPDE-dG) adducts. In contrast, the minor-
groove trans-BPDE adducts maintain Watson-Crick base
pairs at the lesion site (8, 9). The cis adducts have been
shown to be incised by UvrABC more efficiently than the
trans adducts (59). Thus, our NER results support growing
evidence that a base-displaced intercalated conformer is more
efficiently repaired than adducts with intact Watson-Crick
hydrogen bonds. However, it should be pointed out that
carcinogen stacking in certain intercalated S-like conforma-
tions is responsible for increased thermal and thermodynamic
stabilization of a duplex (9, 10), a known negative indicator
for NER (43). While the modified dG of the B conformer
maintains Watson-Crick bonds at the lesion site, the highly
hydrophobic carcinogen moiety in the major groove may
create a solvation environment favorable for repair. Zou et
al. (59) have shown that AF and AAF adducts in the -TG*T-
sequence context are incised more efficiently by UvrABC
than in the -CG*C- context. Similar results have been
obtained with the minor groove trans-BPDE adduct by
UvrABC proteins from both E. coli and the thermophilic
Bacillus caldotenax (76). The presence of the T:A flanking
base pairs adjacent to the modified dG* allowed for greater
local bend, flexibility, and conformational heterogeneity than
in the -CG*C- sequence. This is consistent with the lower
Tm and -∆G° values observed for the -TG*T- sequence.
Interestingly, we have shown in the present study that the
-CG*N- duplex series were more stable (greater Tm and
-∆G° values) than the -AG*N- counterpart, yet they were
more susceptible to repair (Figure 7). It appears that
important recognition factors of UvrABC NER are not
necessarily thermal and thermodynamic stabilities, but rather
the quality of Watson-Crick base pairs and conformational
flexibility at the lesion site. Taken together, these results
illustrated the complexity of the way multiconformeric
adducts interact with repair proteins, information that is
crucial to the first step in damage recognition in NER.
Several mechanisms have been proposed to explain how
the NER machinery recognizes lesions situated within an
expanse of intact genomic DNA (39-43). However, there
is no single one-fits-all mechanism. The concept of “thermal/
thermodynamic probing” has evolved in recent years to
involve more complex “bipartite” or “multipartite” mecha-
nisms (42, 43). The latter focuses on the quality of Watson-
Crick base pairs at or near the lesion site and the adduct
structures. Alternative possibilities include the indirect
readout mechanisms which emphasize the local conforma-
tional distortion of either the modified or the complementary
strand at the modification site (77, 78).
It has been shown that the DNA sequence itself has no
intrinsic or direct effect on the incision efficiency, as the
damage recognition factor UvrA2B also binds to undamaged
DNA in a sequence-independent manner (79). The DNA
sequence at a lesion may only affect the structure of the
lesion which is recognized and incised. Since all the
substrates examined in this study contained the same adduct
FAF, they should be recognized equally well by UvrB (80).
Our repair results reinforced the importance of Watson-
Crick base pairing at the lesion site for the initial recognition
by the UvrA2B proteins. Our CD data indicated, however,
that neither the B nor the S conformer significantly altered
the global DNA conformation. So, what differentiates B and
S conformers in terms of NER recognition? Figure 8 shows
the surface models of B and S conformers and highlights
the polarity and shape differences in their major groove
area: i.e., the extrusion of the highly hydrophobic carcinogen
moiety in the B conformer Versus the adduct-displaced highly
polar dG and dC in the S conformer. It is plausible that this
subtle electrostatic difference may be responsible for the
observed conformation-specific repair. Direct identification
of the lesion structures at those sequence-dependent hot spots
in cells is almost impossible. However, our19F NMR/NER
results provided insights into the three-way relationship of
sequence, conformation specificity, and NER, a biological
consequence closely associated with mutagenesis (81).
11274 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.
S/B conformational heterogeneity may also play an
important role in modulating the mutational outcomes of the
AF adduct. Significant sequence effects were observed on
both frameshift and substitution mutation frequency of AAF,
AF, and PhIP adducts derived both in E. coli and mammalian
cells (44, 64). Shibutani et al. (66) have shown that the
mutation frequency and specificity of AF in COS-7 cells vary
significantly depending on the nature of the bases flanking
the lesion. For example, the base substitution mutation of
AF in the 5′-GG*C-3′ context was 50-fold higher than in
the 5′-TG*C-3′ context. The presence of a dG 5′ of the lesion
increases the overall mutational frequency dramatically.
These effects must originate from the sequence dependence
of S/B heterogeneity. A more complete set of19F NMR
results will be required to establish meaningful conforma-
Summary. In the present study we conducted a systematic
spectroscopic (UV, CD,
(VDW, QM) investigation in order to gain structural insight
into AF-induced S/B conformational heterogeneity. We found
that FAF-modified DNA duplexes equilibrated between S
and B conformations and interconverted slowly at room
temperature. The S/B population ratios were modulated by
the 3′-flanking sequence. Specifically, purine bases at the
3′-flanking site promoted the S conformation (G > A > C
> T). Van der Waals energy calculations indicated that the
stacking interaction between the carcinogen and neighboring
base pairs was a key to the observed sequence effect.
Quantum mechanics calculations provided a more complete
19F NMR) and computational
picture and stressed the importance of both electrostatic and
solvation forces. The19F NMR/NER results suggested that
the lesions with higher S/B ratio appeared to be more
susceptible to NER repair than most of the lesions with lower
S/B ratio (except CG*T). The results represented the first
quantitative structure-function investigation relating frac-
tions of S conformers with increased NER efficiencies and
provided valuable insights into how bulky DNA adducts are
accommodated by the NER protein in a conformationally
SUPPORTING INFORMATION AVAILABLE
Synthesis (Figure S1), HPLC purification (Figures S2, S3),
and ESI-TOF-MS characterization (Figures S4-S7) of FAF-
modified 12-mer oligodeoxynucleotides; imino proton
(Figure S8) and19F (Figure S9) NMR spectra of the -AG*N-
and CG*N- duplex series; simulated dynamic spectra (Figure
S10) of the -CG*N- duplexes; the atom type assignment of
FAF-dG (Figure S11); assumed enthalpy ordering diagram
(Figure S12); calculated and measured molecular ion mass
(Table S1);19F NMR H/D isotope effects (Table S2); van
der Waals parameters for B (Table S3) and S (Table S4)
conformers; force field parameters for dG-FAF (Table S5);
RESP charge results for B (Table S6) and S (Table S7)
conformers; QM calculations, DFT-D dimer interaction
energies (Table S8). This material is available free of charge
via the Internet at http://pubs.acs.org.
FIGURE 8: Surface model illustrations which highlight the polarity and shape difference in the major groove area of the B and S conformers;
i.e., the extrusion of the highly hydrophobic carcinogen moiety (red) in the B conformer Versus the adduct-displaced highly polar dG (cyan)
and dC (green) in the S conformer.
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11275
1. Hoeijmakers, J. H. J. (2001) Genome maintenance mechanisms
for preventing cancer, Nature 411, 366-374.
2. Luch, A. (2005) Nature and nature-lessons from chemical car-
cinogenesis, Nat. ReV. Cancer 5, 113-125.
3. Guengerich, F. P. (2006) Interactions of carcinogen-bound DNA
with individual DNA polymerases, Chem. ReV. 106, 420-452.
4. Lukin, M., and de Los Santos, C. (2006) NMR structures of
damaged DNA, Chem. ReV. 106, 607-686.
5. Pages, V., and Fuchs, R. P. P. (2002) How DNA lesions are turned
into mutations within cells?, Oncogene 21, 8957-8966.
6. Seo, K. Y., Jelinsky, S. A., and Loechler, E. L. (2000) Factors
that influence the mutagenic patterns of DNA adducts from
chemical carcinogens, Mutat. Res. 463, 215-246.
7. Seo, K. Y., Nagalingam, A., Tiffany, M., and Loechler, E. L.
(2005) Mutagenesis studies on four stereoisomeric N2-dG benzo-
[a]pyrene adducts in the identical 5′-CGC sequence used in NMR
studies: although adduct conformation differs, mutagenesis
outcome does not as GfT mutations dominate in each case,
Mutagenesis 20, 441-448.
8. Cho, B. P. (2004) Dynamic conformational heterogeneities of
carcinogen-DNA adducts and their mutagenic relevance, J.
EnViron. Sci. Health, Part C: EnViron. Carcinog. Ecotoxicol. ReV.
9. Geacintov, N. E., Cosman, M., Hingerty, B. E., Amin, S., Broyde,
S., and Patel, D. J. (1997) NMR solution structures of stereoiso-
metric covalent polycyclic aromatic carcinogen-DNA adduct:
principles, patterns, and diversity, Chem. Res. Toxicol. 10, 111-
10. Patel, D. J., Mao, B., Gu, Z., Hingerty, B. E., Gorin, A., Basu, A.
K., and Broyde, S. (1998) Nuclear magnetic resonance solution
structures of covalent aromatic amine-DNA adducts and their
mutagenic relevance, Chem. Res. Toxicol. 11, 391-407.
11. Hoffmann, G. R., and Fuchs, R. P. P. (1997) Mechanisms of
frameshift mutations: insight from aromatic amines, Chem. Res.
Toxicol. 10, 347-459.
12. Elmquist, C. E., Wang, F., Stover, J. S., Stone, M. P., and Rizzo,
C. J. (2007) Conformational differences of the C8-Deoxyguanosine
adduct of 2-amino-3-methylimidazo[4,5-f]quinoline (IQ) within
the NarI recognition sequence, Chem. Res. Toxicol. 445-454.
13. Heflich, R. H., and Neft, R. E. (1994) Genetic toxicity of
2-acetylaminofluorene, 2-aminofluorene and some of their me-
tabolites and model metabolites, Mutat. Res. 318, 73-114.
14. Beije, B., and Moller, L. (1988) 2-Nitrofluorene and related
compounds: prevalence and biological effects, Mutat. Res. 196,
15. Hsu, G. W., Kiefer, J. R., Burnouf, D., Becherel, O. J., Fuchs, R.
P., and Beese, L. S. (2004) Observing translesion synthesis of an
aromatic amine DNA adduct by a high-fidelity DNA polymerase,
J. Biol. Chem. 279, 50280-50285.
16. Dutta, S., Li, Y., Johnson, D., Dzantiev, J., Richardson, C. C.,
Romano, L. J., and Ellenberger, T. (2004) Crystal structures of
2-acetylaminofluorene and 2-aminofluorene in complex with T7
DNA polymerase reveal mechanisms of mutagenesis, Proc. Natl.
Acad. Sci. U.S.A. 101, 16186-16191.
17. Cho, B. P., Beland, F. A., and Marques, M. M. (1994) NMR
structural studies of a 15-mer duplex from a ras protooncogene
modified with the carcinogen 2-aminofluorene: conformational
heterogeneity, Biochemistry 33, 1373-1384.
18. Eckel, L. M., and Krugh, T. R. (1994) Structural characterization
of two interchangeable conformations of a 2-aminofluorene-
modified DNA oligomer by NMR and energy minimizations,
Biochemistry 33, 13611-13624.
19. Mao, B., Gu, Z., Hingerty, B. E., Broyde, S., and Patel, D. J. (1998)
Solution structure of the aminofluorene [AF]-intercalated con-
former of the syn [AF]-C8-dG adduct opposite dC in a DNA
duplex, Biochemistry 37, 81-94.
20. Mao, B., Gu, Z., Hingerty, B. E., Broyde, S., and Patel, D. J. (1998)
Solution structure of the aminofluorene [AF]-external conformer
of the anti [AF]-C8-dG adduct opposite dC in a DNA duplex,
Biochemistry 37, 95-106.
21. Mao, B., Gu, Z., Gorin, A., Hingerty, B. E., Broyde, S., and Patel,
D. J. (1997) Solution structure of the aminofluorene [AF] stacked
conformer of the syn [AF]-C8-dG adduct positioned at a DNA
template-primer junction, Biochemistry 36, 14491-14501.
22. Gorin, A., Gu, Z., Hingerty, B. E., Broyde, S., and Patel, D. J.
(1999) Solution structure of the aminofluorene [AF]-stacked
conformer of the syn [AF]-C8-dG adduct positioned opposite dC
at a template-primer junction, Biochemistry 38, 10855-10870.
23. Norman, D., Abuaf, P., Hingerty, B. E., Live, D., Grunberger,
D., Broyde, S., and Patel, D. J. (1989) NMR and computational
characterization of the N-(deoxyguanosin-8-yl)aminofluorene ad-
duct (AF)G opposite adenosine in DNA: (AF)G[syn]: A[anti]
pair formation and its pH dependence, Biochemistry 28, 7462-
24. Abuaf, P., Hingerty, B. E., Broyde, S., and Grunberger, D. (1995)
Solution conformation of the N-(deoxyguanosin-8-yl)aminofluo-
rene adduct opposite deoxyinosine and deoxyguanosine in DNA
by NMR and computational characterization, Chem. Res. Toxicol.
25. Mao, B., Cosman, M., Hingerty, B. E., Broyde, S., and Patel, D.
J. (1995) Solution conformation of [AF]dG opposite a -1 deletion
site in a DNA duplex: intercalation of the covalently attached
aminofluorene ring into the helix with base displacement of the
C8-modified syn guanine into the major groove, Biochemistry 34,
26. Mao, B., Hingerty, B. E., Broyde, S., and Patel, D. J. (1995)
Solution conformation of [AF]dG opposite a -2 deletion site in a
DNA duplex: intercalation of the covalently attached aminofluo-
rene ring into the helix with base displacement of the C8-modified
syn guanine and adjacent unpaired 3′-adenine into the major
groove, Biochemistry 34, 16641-16653.
27. Mao, B., Gorin, A., Gu, Z., Hingerty, B. E., Broyde, S., and Patel,
D. J. (1997) Solution structure of the aminofluorene [AF]-
intercalated conformer of the syn [AF]-C8-dG adduct opposite a
-2 deletion site in the NarI hotspot sequence context, Biochemistry
28. Zhou, L., Rajabzadeh, G., Traficante, D. D., and Cho, B. P. (1997)
Conformational heterogeneity of arylamine-modified DNA:
NMR evidence, J. Am. Chem. Soc. 119, 5384-5389.
29. Meneni, S. R., D’Mello, R., Norigian, G., Baker, G., Gao, L.,
Chiarelli, M. P., and Cho, B. P. (2006) Sequence effects of
aminofluorene-modified DNA duplexes: thermodynamic and
circular dichroism properties, Nucleic Acids Res. 34, 755-
30. Liang, F., Meneni, S., and Cho, B. P. (2006) Induced circular
dichroism characteristics as conformational probes for carcino-
genic aminofluorene-DNA adducts, Chem. Res. Toxicol. 19,
31. Meneni, S., Liang, F., and Cho, B. P. (2007) Examination of the
long-range effects of aminofluorene-induced conformational het-
erogeneity and its relevance to the mechanisms of translesion DNA
synthesis, J. Mol. Biol. 366, 1387-1400.
32. Prive, G. G., Yanagi, K., and Dickerson, R. E. (1991) Structure
of the B-DNA decamer C-C-A-A-C-G-T-T-G-G and comparison
with isomorphous decamers C-C-A-A-G-A-T-T-G-G and C-C-
A-G-G-C-C-T-G-G, J. Mol. Biol. 217, 177-199.
33. Sponer, J., Leszczynski, J., and Hobza, P. (1996) Nature of nucleic
acid-base stacking: nonempirical ab initio and empirical potential
characterization of 10 stacked base dimers. Comparison of stacked
and H-bonded base pairs, J. Phys. Chem. 100, 5590-5596.
34. Reha, D., Kabelac, M., Ryjacek, F., Sponer, J., Sponer, J. E.,
Elstner, M., Suhai, S., and Hobza, P. (2002) Intercalators. 1. Nature
of stacking interactions between intercalators (ethidium, dauno-
mycin, ellipticine, and 4′,6-diaminide-2-phenylindole) and DNA
base pairs. Ab initio quantum chemical, density functional theory,
and empirical potential study, J. Am. Chem. Soc. 124, 3366-
35. Luo, R., Gilson, H. S. R., Potter, M. J., and Gilson, M. K. (2001)
The physical basis of nucleic acid base stacking in water, Biophys.
J. 80, 140-148.
36. Sponer, J., Jurecka, P., Marchan, I., Luque, F. J., Orozco, M., and
Hobza, P. (2006) Nature of Base Stacking: Reference Quantum-
Chemical Stacking Energies in Ten Unique B-DNA Base-Pair
Steps, Chem. Eur. J. 12, 2854-2865.
37. Spackova, N., Berger, I., Egli, M., and Sponer, J. (1998) Molecular
dynamics of hemiprotonated intercalated four-stranded i-DNA:
stable trajectories on a nanosecond scale, J. Am. Chem. Soc. 120,
38. McDowell, S. E., Spackova, N., Sponer, J., and Walter, N. G.
(2007) Molecular dynamics simulations of RNA: An in silico
single molecule approach, Biopolymers 85, 169-184.
39. Gillet, L. C. J., and Scharer, O. D. (2006) Molecular mechanisms
of mammalian global genome nucleotide excision repair, Chem.
ReV. 106, 253-276.
11276 Biochemistry, Vol. 46, No. 40, 2007
Meneni et al.
40. Truglio, J. J., Croteau, D. L., Van Houten, B., and Kisker, C.
(2006) Prokaryotic nucleotide excision repair: the UvrABC
system, Chem. ReV. 106, 233-252.
41. Sancar, A., and Reardon, J. T. (2004) Nucleotide excision repair
in E. coli and man, AdV. Protein Chem. 69, 43-71.
42. Dip, R., Camenisch, U., and Naegeli, H. (2004) Mechanisms of
DNA damage recognition and strand discrimination in human
nucleotide excision repair, DNA Repair (Amst) 2, 1409-23.
43. Geacintov, N. E., Broyde, S., Buterin, T., Naegeli, H., Wu, M.,
Yan, S., and Patel, D. J. (2002) Thermodynamic and structural
factors in the removal of bulky DNA adducts by the nucleotide
excision repair machinery, Biopolymers 65, 202-210.
44. Shibutani, S. (2004) Requirements for frame-shift deletion during
translesion synthesis, EnViron. Mutagen Res. 26, 135-141.
45. Meneni, S., Shell, S. M., Zou, Y., and Cho, B. P. (2007)
Conformation-specific recognition of carcinogen-DNA adduct in
E. coli. nucleotide excision repair, Chem. Res. Toxicol. 20, 6-
46. Friebolin, H. (1998) in Basic one- and two-dimensional NMR
spectroscopy, 3rd ed., pp 301-329, Wiley-VCH, New York.
47. Wang, J. M., Cieplak, P., and Kollman, P. A. (2000) How well
does a restrained electrostatic potential (RESP) model perform in
calculating conformational energies of organic and biological
molecules?, J. Comput. Chem. 21, 1049-1074.
48. Cheatham, T. E., III, Cieplak, P., and Kollman, P. A. (1999) A
modified version of the Cornell et al. force field with improved
sugar pucker phases and helical repeat, J. Biomol. Struct. Dyn.
49. Wang, J., Wolf, R. M., Caldwell, J. W., Kollman, P. A., and Case,
D. A. (2004) Development and testing of a general amber force
field, J. Comput. Chem. 25, 1157-1174.
50. Frisch, M. J., Trucks, G. W., Schlegel, H. B., Scuseria, G. E.,
Robb, M. A., Cheeseman, J. R., Montgomery, J. A., Jr., Vreven,
T., Kudin, K. N., Burant, J. C., Millam, J. M., Iyengar, S. S.,
Tomasi, J., Barone, V., Mennucci, B., Cossi, M., Scalmani, G.,
Rega, N., Petersson, G. A., Nakatsuji, H., Hada, M., Ehara, M.,
Toyota, K., Fukuda, R., Hasegawa, J., Ishida, M., Nakajima, T.,
Honda, Y., Kitao, O., Nakai, H., Klene, M., Li, X., Knox, J. E.,
Hratchian, H. P., Cross, J. B., Bakken, V., Adamo, C., Jaramillo,
J., Gomperts, R., Stratmann, R. E., Yazyev, O., Austin, A. J.,
Cammi, R., Pomelli, C., Ochterski, J. W., Ayala, P. Y., Morokuma,
K., Voth, G. A., Salvador, P., Dannenberg, J. J., Zakrzewski, V.
G., Dapprich, S., Daniels, A. D., Strain, M. C., Farkas, O., Malick,
D. K., Rabuck, A. D., Raghavachari, K., Foresman, J. B., Ortiz,
J. V., Cui, Q., Baboul, A. G., Clifford, S., Cioslowski, J., Stefanov,
B. B., Liu, G., Liashenko, A., Piskorz, P., Komaromi, I., Martin,
R. L., Fox, D. J., Keith, T., Al-Laham, M. A., Peng, C. Y.,
Nanayakkara, A., Challacombe, M., Gill, P. M. W., Johnson, B.,
Chen, W., Wong, M. W., Gonzalez, C., and Pople, J. A. (2004)
Gaussian 03, revision C.02, Gaussian, Inc., Wallingford CT.
51. Cornell, W. D., Cieplak, P., Bayly, C. I., Gould, I. R., Merz, K.
M., Ferguson, D. M., Spellmeyer, D. C., Fox, T., Caldwell, J.
W., and Kollman, P. A. (1995) A second generation force field
for the simulation of proteins, nucleic acids, and organic mol-
ecules, J. Am. Chem. Soc. 117, 5179-5197.
52. Simmerling, C., Elber, R., and Zhang, J. (1995) in Modeling of
Biomolecular Structure and Mechanisms (Pullman, A., Ed.) pp
241-265, Kluwer, The Netherlands.
53. Becke, A. D. (1993) Density-functional thermochemistry. III. The
role of exact exchange, J. Chem. Phys. 98, 5648-5652.
54. Jurec ˇka, P., C ˇerny ´, J., Hobza, P., and Salahub, D. R. (2007) Density
functional theory augmented with an empirical dispersion term.
Interaction energies and geometries of 80 noncovalent complexes
compared with ab initio quantum mechanics calculations, J.
Comput. Chem. 28, 555-569.
55. Tao, J., Perdew, J. P., Staroverov, V. N., and Scuseria, G. E. (2003)
Climbing the density functional ladder: nonempirical meta-
generalized gradient approximation designed for molecules and
solids, Phys. ReV. Lett. 91, 146401-146404.
56. Ahlrichs, R., Ba ¨r, M., Ha ¨ser, M., Horn, H., and Ko ¨lmel, C. (1989)
Electronic structure calculations on workstation computers: The
program system turbomole, Chem. Phys. Lett. 162, 165-169.
57. Zou, Y., Liu, T. M., Geacintov, N. E., and Van Houten, B. (1995)
Interaction of the UvrABC nuclease system with a DNA duplex
containing a single stereoisomer of dG-(+)- or dG-(-)-anti-BPDE,
Biochemistry 34, 13582-13593.
58. Yang, Z., Colis, L. C., Basu, A. K., and Zou, Y. (2005) Recognition
and incision of gamma-radiation-induced cross-linked guanine-
thymine tandem lesion G[8,5-Me]T by UvrABC nuclease, Chem.
Res. Toxicol. 18, 1339-1346.
59. Zou, Y., Shell, S. M., Utzat, C. D., Luo, C., Yang, Z., Geacintov,
N. E., and Basu, A. K. (2003) Effects of DNA adduct structure
and sequence context on strand opening of repair intermediates
and incision by UvrABC nuclease, Biochemistry 42, 12654-
60. Luo, C., Krishnasamy, R., Basu, A. K., and Zou, Y. (2000)
Recognition and incision of site-specifically modified C8 guanine
adducts formed by 2-aminofluorene, N-acetyl-2-aminofluorene and
1-nitropyrene by UvrABC nuclease, Nucleic Acids Res. 28, 3719-
61. SantaLucia, J., and Hicks, D. (2004) The thermodynamics of DNA
structural motifs, Annu. ReV. Biophys. Biomol. Struct. 33, 415-
62. Singer, B., and Hang, B. (2000) Nucleic acid sequence and
repair: role of adduct, neighbor bases and enzyme specificity,
Carcinogenesis 21, 1071-1078.
63. Hansen, P. E., Dettman, H. D., and Sykes, B. D. (1985) Solvent-
induced deuterium isotope effects on fluorine-19 chemical shifts
of some substituted fluorobenzenes. Formation of inclusion
complexes, J. Magn. Reson. 62, 487-496.
64. Broschard, T. H., Koffel-Schwartz, N., and Fuchs, R. P. P. (1999)
Sequence-dependent modulation of frameshift mutagenesis at
NarI-derived mutation hot spots, J. Mol. Biol. 288, 191-199.
65. Page, J. E., Zajc, B., Oh-hara, T., Lakshman, M. K., Sayer, J. M.,
Jerina, D. M., and Dipple, A. (1998) Sequence context profoundly
influences the mutagenic potency of trans-opened benzo[a]pyrene
7,8-diol 9,10-epoxide-purine nucleoside adducts in site-specific
mutation studies, Biochemistry 37, 9127-9137.
66. Shibutani, S., Suzuki, N., Tan, X., Johnson, F., and Grollman, A.
P. (2001) Influence of flanking sequence context on the mutage-
nicity of acetylaminofluorene-derived DNA adducts in mammalian
cells, Biochemistry 40, 3717-3722.
67. Zhuang, P., Kolbanovskiy, A., Amin, S., and Geacintov, N. E.
(2001) Base sequence dependence of in vitro translesional DNA
replication past a bulky lesion catalyzed by the exo- Klenow
fragment of Pol I, Biochemistry 40, 6660-6669.
68. Kolbanovskiy, A., Kuzmin, V., Shastry, A., Kolbanovskaya, M.,
Chen, D., Chang, M, Bolton, J. L., and Geacintov, N. E. (2005)
Base selectivity and effects of sequence and DNA secondary
structure on the formation of covalent adducts derived from the
equine estrogen metabolite 4-hydroxyequilenin, Chem. Res. Toxi-
col. 18, 1737-1747.
69. Krueger, A., Protozanova, E., and Frank-Kamenetskii, M. D.
(2006) Sequence-dependent base pair opening in DNA double
helix, Biophys. J. 90, 3091-3099.
70. Jiang, Y. L., McDowell, L. M., Poliks, B., Studelska, D. R., Cao,
C., Potter, G. S., Schaefer, J., Song, F., and Stivers, J. T. (2004)
Recognition of an unnatural difluorophenyl nucleotide by uracil
DNA glycosylase, Biochemistry 43, 15429 -15438.
71. Yakovchuk, P., Protozanova, E., and Frank-Kamenetskii, M. D.
(2006) Base-stacking and base-pairing contributions into thermal
stability of the DNA double helix, Nucleic Acids Res. 34, 564-
72. Hardman, S. J., and Thompson, K. C. (2006) Influence of base
stacking and hydrogen bonding on the fluorescence of 2-ami-
nopurine and pyrrolocytosine in nucleic acids, Biochemistry 45,
73. Yang, W. (2006) Poor base stacking at DNA lesions may initiate
recognition by many repair proteins, DNA Repair 5, 654-
74. Malta, E., Moolenaar, G. F., and Goosen, N. (2006) Base flipping
in nucleotide excision repair, J. Biol. Chem., 281, 2184-2194.
75. Mekhovich, O., Tang, M., and Romano, L. J. (1998) Rate of
incision of N-acetyl-2-aminofluorene and N-2-aminofluorene
adducts by UvrABC nuclease is adduct- and sequence-specific:
comparison of the rates of UvrABC nuclease incision and protein-
DNA complex formation, Biochemistry 37, 571-579.
76. Ruan, Q., Liu, T., Kolbanovskiy, A., Liu, Y., Ren, J., Skovraga,
M., Zou, Y., Lader, J., Malkani, B., Amin, S., Van Houten, B.,
and Geacintov, N. E. (2007) Sequence context- and temperature-
dependent nucleotide excision repair of a benzo[a]pyrene diol
epoxide-guanine DNA adduct catalyzed by thermophilic UvrABC
proteins, Biochemistry 46, 7006-7015.
Nucleotide Excision Repair
Biochemistry, Vol. 46, No. 40, 2007 11277