From the Susan Lehman Cullman Laboratory for Cancer
Research, Department of Chemical Biology, College of Pharmacy, Rutgers,
State University of New Jersey, Piscataway, New Jersey 08854-8020 and the ¶ Laboratory of Bioorganic Chemistry, NIDDK, National
Institutes of Health, Bethesda, Maryland 20892-0820
Received for publication, November 13, 2002, and in revised form, February 12, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Site-specific mutagenicity of
trans-opened adducts at the exocyclic
N2-amino group of guanine by the
(+)-(7R,8S,9S,10R)- and
( Mutagenic effects of benzo[a]pyrene, a
carcinogenic environmental pollutant (1), are largely mediated through
stereoisomers of its reactive diol epoxide
(DE)1 metabolite,
7,8-dihydroxy-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene. Early studies (2) suggested that BPDE was responsible for the major DNA
adduct formed from benzo[a]pyrene. Two diastereomers, DE-1
and DE-2 (3) in which the benzylic 7-hydroxyl group and epoxide oxygen
are either cis or trans, respectively, are
metabolically possible. Each of these exists as a pair of enantiomers
(4). The predominantly formed
(+)-(7R,8S)-dihydroxy-(9S,10R)-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene ((+)-BPDE-2) (5) is the most mutagenic stereoisomer in cell cultures
(6) and the strongest carcinogen in mice (7, 8). Its enantiomer,
()-(7S,8R,9R,10S)-enantiomers of a benzo[a]pyrene 7,8-diol 9,10-epoxide (7-hydroxyl and
epoxide oxygen are trans, BPDE-2) has been determined in
Chinese hamster V79 cells and their repair-deficient counterpart, V-H1
cells. Four vectors containing single 10S-BPDE-dG or
10R-BPDE-dG adducts positioned at G0 or
G
1 in the analyzed 5'-ACTG0G
1GA sequence of the non-transcribed strand were separately transfected into
the cells. Mutations at each of the seven nucleotides were analyzed by
a novel primer extension assay using a mixture of one dNTP
complementary to the mutated nucleotide and three other ddNTPs and were
optimized to quantify levels of a mutation as low as 1%. Only G
T
mutations were detected at the adducted sites; the 10S
adduct derived from the highly carcinogenic (+)-diol epoxide was 40-50
and 75-140% more mutagenic than the 10R adduct in V79 and
V-H1 cells, respectively. Importantly, the 10S adducts, but
not the 10R adducts, induced separate non-targeted
mutations at sites 5' to the G
1 and G0
lesions (G0
T and C
T, respectively) in both cell
lines. Neither the T 5' to G0 nor sites 3' to the lesions
showed mutations. Non-targeted mutations may enhance overall
mutagenicity of the 10S-BPDE-dG lesion and contribute to
the much higher carcinogenicity and mutagenicity of (+)-BPDE-2 compared
with its (
)-enantiomer. Our study reports a definitive demonstration
of mutations distal to a site-specific polycyclic aromatic hydrocarbon adduct.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
)-7S,8R-dihydroxy-9R,10S-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene ((
)-BPDE-2), has little or no carcinogenic activity (6). Treatment of
cells with (+)-BPDE-2 or (
)-BPDE-2 causes DNA point mutations (mostly
G
T transversions) and DNA frameshifts (9-11) that may lead to
changes in expression or function of critical genes such as
proto-oncogenes and tumor suppressors and eventually to carcinogenesis. (+)-BPDE-2 is ~5-6 times more active in forming DNA adducts than (
)-BPDE-2 (12). Although the (
)-BPDE-2 enantiomer gives substantial amounts of dA adducts (12, 13), both enantiomers react largely with the
exocyclic N2-amino group of deoxyguanosine (dG)
residues by trans opening of the epoxide ring at C-10 to
produce 10S-BPDE-dG and 10R-BPDE-dG lesions (14)
as shown in Fig. 1. Both of these lesions
lie in the minor groove with the aromatic portion pointing toward the 5'- and 3'-ends of their respective adducted strands (15, 16).
View larger version (26K):
[in a new window]
Fig. 1.
Structures of the
trans-opened BPDE adducts at
N2 on the guanine base.
Mutagenicity of BPDE adducts depends upon efficiency of cellular repair
systems to remove the adducts from DNA (mainly nucleotide excision
repair (17, 18)) and fidelity of DNA polymerases replicating residual
adducted sites. Although the normal replicative DNA polymerases pol
and pol
are blocked by bulky adducts, members of newly discovered Y
superfamily of bypass DNA polymerases such as pol
, pol
, and
pol
(19) were found to replicate past sites of various DNA lesions,
albeit with low fidelity and low processivity. The frequency of
nucleotide misincorporation of pol
(20), pol
(21), and pol
(22) replicating an undamaged template in vitro is in the
range of 10
3-10
2. Assuming that the
enzymes replicate not only the site of a lesion but possibly also a
short stretch of DNA around the lesion with such a low fidelity
suggests that besides incorporation of mismatched nucleotides opposite
the adducted site, the bypass DNA polymerases could also introduce
secondary mutations in the region flanking the adducted site,
especially if its DNA structure is perturbed by the presence of the
adduct. Multiple mutations in a shuttle vector treated with (±)-BPDE,
believed to be generated by an error-prone polymerase, have been
observed (23) in random mutagenesis experiments.
In the present study, we have examined the mutagenicity of
10S-BPDE-dG and 10R-BPDE-dG lesions both at
adducted sites and at unmodified flanking sites. We constructed
double-stranded plasmid vectors bearing cDNA of the HPRT
gene of Chinese hamster V79 cells containing a single adduct positioned
at each of the two adjacent sites in the non-transcribed strand of the
gene and transfected them separately into V79 cells and also into their
nucleotide excision repair-deficient derivative, V-H1 cells (24). These cells are defective in the xeroderma pigmentosum complementation group
D/ERCC2 gene encoding for an
ATP-dependent DNA helicase (25), an essential subunit of
transcription and nucleotide excision repair complex TFIIH (26). They
are 9-fold more sensitive to cytotoxic effects of (+)-BPDE-2 than V79
cells and have ~50% lower capacity for removal of (+)-BPDE-2-induced
adducts from DNA compared with V79 cells (27). We qualitatively and
quantitatively evaluated mutagenic effects of both 10R and
10S adducts in a sequence of seven nucleotides by a novel
quantitative minisequencing method, and we compared their differences
with respect to 10R/10S stereochemistry, position
of the adducts at two adjacent sites, and also cellular DNA repair
status. The most striking observation was that, in addition to
mutations at the adduct sites, significant numbers of mutations were
induced 1 or 2 bases 5' to these sites by 10S but not
10R adducts.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Oligonucleotides--
All non-adducted oligonucleotides were
prepared at IDT (Coralville, IA). The 19-mer oligonucleotide and
primers for quantitative minisequencing (Fig.
2) were gel-purified. Adducted
oligonucleotide 18-mers
(5'-AAACTG0G1GAAAGCCAAAT) containing a single
trans-opened BPDE adduct at one or the other of the numbered
dG residues were prepared by solid-phase synthesis using the mixed
10R/10S diastereomers of the appropriately
protected adducted phosphoramidite as described (28, 29), followed by
high pressure liquid chromatography separation of the resultant pair of
diastereomeric oligonucleotides. Synthesis and characterization of the
diastereomeric pair of oligonucleotides bearing trans-opened
BPDE adducts at G0 with 10R and 10S
configuration at the point of attachment of the base to the hydrocarbon
have been reported (28). A second R/S pair of 18-mers with
trans-opened BPDE adducts at G
1 was prepared
(3-µmol scale) by the same methodology and purified by high pressure
liquid chromatography as described for the adduct at G0
(28) (Hamilton PRP-1 column (7 × 305 mm), eluted with a gradient
from 0 to 35% solvent B in solvent A over 20 min, where solvent A is
0.1 M (NH4)2CO3, pH 7.5, and solvent B is a 1:1 mixture of solvent A with CH3CN
adjusted to the same pH). The late eluting oligonucleotide
(tR 19.0 min) was assigned as containing the
trans-opened 10S-BPDE dG adduct and the early
eluting oligonucleotide (tR 17.6 min) as containing
the 10R adduct, on the basis of the CD spectrum (14, 30) of
the known nucleoside adducts obtained upon enzymatic hydrolysis (12) of
the late eluting adduct. The CD spectra of the 18-mer oligonucleotides
themselves exhibited bands at 320-350 nm that were positive for the
early eluting (10R) isomer and weakly negative for the late
eluting (10S) isomer, consistent with previous observations
(31) of other oligonucleotides containing trans-opened BPDE
dG adducts of known absolute configuration as follows: 18-mer modified
with the 10S or 10R adduct at the position
G0, 5'-AAACTGGGAAAGCCAAAT; 18-mer
modified with the 10S or 10R adduct at the
position G
1, 5'-AAACTGGGAAAGCCAAAT; 19-mer,
5'-CATATTTGTGTCATTAGTG; 59-mer scaffold,
5'-GAATTCTCATCTTAGGCTTTGTATTTGGCTTTCCCAGTTTCAGTAATGACACAAATATG; primer
A, 5'-TGCGGGATCCCTCCTCACACCGCT; primer B, 5'-CTGCTTTCCCTGGTCAAGCGG; primer C, 5'-GAAATTAATACGACTCACTATAGGG; primer D,
5'-GCAGATTCAACTTGAATTCTCATC. Sequence of primers used for
mutation analysis by quantitative minisequencing is shown in Fig.
2.
|
Scaffold-directed Extension of the Adducted 18-Mers to the Adducted 37-Mers-- The adducted oligonucleotides and the non-adducted control were extended on their 5' termini with the 19-mer to the final 37-mers by scaffold-directed ligation. Adducted 18-mers or the corresponding non-adducted 18-mer (10 pmol), 5'-labeled with 32P, were incubated with the 19-mer and 5'-32P-labeled 59-mer scaffold in a molar ratio 1:2:1.5 at 65 °C for 5 min and cooled to room temperature over 10 min. Ligase reactions (30 µl) containing 66 mM Tris-HCl, 5 mM MgCl2, 1 mM dithioerythritol, 1 mM ATP, and 0.5 units of T4 DNA ligase were incubated at 25 °C for 2 h at pH 7.5 and terminated by adding 15 µl of stop solution containing 90% formamide, 10 mM NaOH, 20 mM EDTA, 0.05% bromphenol blue, and 0.05% xylene cyanol FF. Reaction products were separated on 10% PAGE and detected by autoradiography. The 37-mer bands were excised from the gel and briefly washed with water. The gel was repeatedly treated with water by three rounds of heating and cooling (65 °C for 5 min and 0 °C for 5 min) to elute the DNA, and the DNA was precipitated from the eluate with ethanol. The total yield of the ligation product was ~40% for the non-adducted oligonucleotide and ~25% for the adducted oligonucleotides.
Preparation of Plasmid Vectors Containing the Full-length Hamster HPRT cDNA-- The HPRT cDNA was prepared by RT-PCR from total mRNA of V79 cells using primer A and primer D as described before (32). Primer A and primer D create BamHI and EcoRI sites, respectively. Double-stranded pCR3 vector (5.1 kb, Invitrogen) containing the HPRT cDNA was prepared by TA cloning (Invitrogen) of the PCR product. The orientation of the HPRT cDNA insert was screened by HindIII digestion of the plasmid DNA. The construct with the antisense orientation of the insert with respect to the cytomegalovirus promoter (pCR3/HPRTantisense), used for preparation of adducted vectors, yields a 289-bp fragment, whereas the construct with the sense orientation (pCR3/HPRTsense) yields a 596-bp fragment. Sequence of the insert in pCR3/HPRTantisense was further verified by dideoxy sequencing using ThermoSequenase Cycle Sequencing Kit (U. S. Biochemical Corp.).
Preparation of Plasmid Vectors Containing Site-specific 10S- or
10R-BPDE Adducts in the HPRT cDNA--
Four plasmid vectors
adducted with BPDE and a non-adducted control were prepared by the
enzymatic extension of the BPDE-adducted and control 37-mer
oligonucleotides (see above for preparation) and annealed to the
single-stranded circular pCR3/HPRTantisense DNA
as shown in Fig. 3. The single-stranded
DNA was isolated from the supernatant of an Escherichia coli
culture transformed with pCR3/HPRTantisense
plasmid after co-infection with M13K07 helper phage (33).
Double-stranded DNA was prepared as follows: a mixture (72 µl)
containing 0.5 pmol of single-stranded
pCR3/HPRTantisense, 0.5 pmol of 37-mer
oligonucleotide, 20 mM Tris-HCl, pH 7.4, 5 mM
MgCl2, and 50 mM NaCl was incubated at 95 °C
for 3 min and then at 65 °C for 3 min and cooled slowly (15 min) to
room temperature. After adding 1 mM DTT, 1 mM
ATP, 500 µM dNTPs, T4 gene 32 protein (5 µg), T4 DNA
polymerase (5 units), and T4 DNA ligase (2.5 units), the reaction (100 µl) was incubated at 25 °C for 2 h. Following extraction with
phenol/chloroform/isoamyl alcohol (25:24:1), DNA was precipitated with
2 volumes of cold ethanol after adding 0.3 M sodium
acetate, pH 5.2, and 2 µg/ml tRNA. After centrifugation, the DNA
pellet was dissolved in 50 mM Tris-HCl, pH 7.6, 5 mM MgCl2, 5 mM DTT, and 50 µg/ml
bovine serum albumin (100 µl) and digested with 20 units of
exonuclease III for 1 h at 37 °C to remove partially extended
reaction products. Extraction (phenol/chloroform/isoamyl alcohol) and
ethanol precipitation was repeated. The pellet was dissolved in 10 mM Tris-HCl, 150 mM NaCl, 10 mM
MgCl2, 1 mM DTT, 100 µg/ml bovine serum
albumin, pH 7.9 (25 µl). The DNA was cut with BamHI + EcoRI (5 units each) for 1 h at 37 °C. Reaction
products, including two BamHI-EcoRI fragments
(15- and 31-mer) and a 24-mer EcoRI-EcoRI
fragment, were separated on a 0.9% preparative agarose gel. Bands
corresponding to 713 bp (HPRT cDNA) and ~5 kb were eluted from the gel (QIAquick Gel Extraction kit, Qiagen, Valencia, CA)
and were re-ligated at 16 °C for 16 h in a 50-µl mixture
containing 66 mM Tris-HCl, 6.6 mM
MgCl2, 10 mM DTT, 66 µM ATP, and
1 unit of T4 DNA ligase. This step changes the orientation of the
insert from the antisense to sense and also separates residual
single-stranded DNA. The reaction was stopped by 500 mM
EDTA (1 µl), and products were applied on the Qiagen QIAquick column
(Nucleotide Removal kit, Qiagen). DNA was eluted in a sterile 10 mM Tris-HCl buffer, pH 8.5. The final concentration of
pCR3/HPRTantisense-BPDE and the non-adducted
control used for cell transfections was 20 ng/µl. All four plasmid
vectors were prepared at least three times.
|
Transfection of V79 and V-H1 Cells with Adducted Plasmid
Vectors--
The Chinese hamster V79 cells were obtained from the ATCC
and propagated in minimum Eagle's medium containing 10% dialyzed and
heat-inactivated fetal bovine serum in an incubator with controlled humidified atmosphere containing 5% CO2. Before
transfection, the cells were seeded at a density 2 × 105 per 60-mm dish. Following 24 h of incubation,
cells were transfected with 0.3 µg of one of the modified plasmids
(10S or 10R adduct at the G0 or
G1 position) or the control plasmid using Effectene
(Qiagen). The cells were incubated for 24 h, washed with
phosphate-buffered saline, and supplied with fresh medium. After
48 h following the transfection, the cells were harvested by
trypsinization and seeded in the same medium supplemented with 500 µg/ml G418 (Invitrogen) at a density 3 × 104 per
60-mm dish to select permanently transfected cells. Ten dishes were
prepared from each transfection. Cellular colonies developed after an
11-day incubation with G418 were harvested by trypsinization from each
dish and collected by centrifugation. To ensure analysis of at least
100 independent cellular clones from each plasmid transfection, 10 plates containing at least 60 cellular colonies in a random mixture
were harvested. Thus, each transfection experiment yielded 5 × 10 pooled samples, which were separately processed and analyzed.
The repair-efficient Chinese hamster V-H1 cells (kindly provided by Dr. M. Zdzienicka, Leiden University Medical Center, Leiden, The Netherlands) were grown and transfected in the same way with several changes. The cells were seeded at a density 2.6 × 105 per a 60-mm dish, transfected with 0.5 µg of the plasmid, and seeded after the transfection at a density 1 × 105 per 60-mm dish.
Preparation of DNA Samples for Mutation Analysis-- Chromosomal DNA from harvested cells was isolated using High Pure PCR Template Preparation kit (Roche Molecular Biochemicals). DNA fragment (337 bp) encompassing the examined region was prepared by PCR amplification using primer B corresponding to region 437-457 of the HPRT gene (exon 6) (32) and primer C corresponding to the T7 promoter region of the plasmid construct; the endogenous cellular HPRT gene is not amplified using this set of primers. The reaction was carried out in a 50-µl mixture containing 200 ng of DNA, 200 nM primers, 200 µM dNTP, and Taq DNA polymerase (0.025 unit/µl, Qiagen). Following 30 cycles (95 °C for 30 s, 55 °C for 10 s, and 72 °C for 10 s) with the initial 3 min at 95 °C and the final 7 min at 72 °C, the residual dNTPs were removed by a 30-min alkaline phosphatase treatment (1 unit per each 20 µl of the reaction). The product was purified using QIAquick PCR Purification kit (Qiagen). Concentration of the product was estimated based on absorbance at 260 nm, and the purity was checked on 1.5% agarose gel containing 0.5 µg/ml ethidium bromide. Standard 337-bp fragments used as positive and negative controls for quantitative minisequencing were amplified from plasmids pCR3/HPRTantisense containing all four nucleotides at each of the seven examined positions (Fig. 2). These plasmids were prepared by site-directed mutagenesis using a QuikChange Kit (Stratagene, La Jolla, CA).
Mutation Detection Assay Using Quantitative
Minisequencing--
The method is based on a single nucleotide primer
extension assay (34, 35) modified to quantify mutations in a known
sequence context at levels lower than 10%. A primer annealed to the
DNA template (337-bp fragments), one nucleotide before the analyzed site (Fig. 2), is extended with a mixture of one dNTP and three other
ddNTPs. When annealed to a fragment carrying a specific analyzed
nucleotide (mutation), the primer can be extended with the
complementary dNTP before being terminated by ddNTP incorporation at
the subsequent nucleotide, whereas elongation of the primer annealed to
fragments not containing the analyzed nucleotide is terminated
immediately with ddNTP. The level of mutation is determined by
calculating the ratio of differently extended primers following their
separation on denaturing polyacrylamide gel and analysis with
PhosphorImager (see Fig. 5). Primer extension was catalyzed by
ThermoSequenase DNA polymerase (0.4 units/µl, U. S. Biochemical Corp.) in a reaction mixture (10 µl) containing three ddNTPs (1 µM), one dNTP (10 µM) complementary to the
analyzed nucleotide, 26 mM Tris-HCl, pH 9.5, 6.5 mM MgCl2, 50 nM
32P-labeled primer, and 2.5 nM DNA template.
Twenty-five cycles (95 °C for 30 s, 50 °C for 10 s, and
72 °C for 10 s) with the initial 1 min at 95 °C were used.
The reaction was stopped with 1 volume of 95% formamide, 20 mM EDTA, 10 mM NaOH, 0.05% bromphenol blue,
and 0.05% xylene cyanol FF. Reaction products were separated on 15%
PAGE in 1× TBE buffer and visualized by autoradiography. To increase
the throughput of the assay, up to four sets of samples were
subsequently loaded into the same gel. For quantitative evaluation (Fig. 5), the signal from the gel was transferred onto a screen and
scanned by Cyclone PhosphorScreening System (Packard Instrument Co.),
and the amount of radioactivity associated with each spot was
quantified using software provided with the system.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Construction of Site- and Stereospecifically Adducted Plasmid
Vectors--
BPDE-adducted plasmid vectors were constructed by the
in vitro enzymatic extension of 37-mer oligonucleotides
after their annealing to a single-stranded plasmid template as shown in
Fig. 3 and described under "Materials and Methods." The plasmid
contains the HPRT gene inserted in the antisense orientation
toward the plasmid promoter. The 37-mers were prepared from 18-mer
oligonucleotides modified with 10S or 10R adducts
at the G0 or G1 sites (see "Materials and
Methods" and Fig. 4A). The
sequence of the 18-mers corresponds to region 629-646 of the
non-transcribed strand of the HPRT gene in Chinese hamster
cells. The use of 37-mers instead of 18-mers in the reaction
substantially increases the yield of the covalently closed circular
reaction products (data not shown). After the reaction, covalently
closed circular DNA was digested with restriction enzymes
EcoRI and BamHI and re-ligated yielding a vector
with the HPRT gene in the sense orientation and a single
BPDE adduct in the non-transcribed strand of the gene (Fig. 3). Purity
of the final products was checked by digestion with restriction enzymes
EcoRI and MslI and separation of the 32P-labeled restriction fragments on denaturing PAGE (Fig.
4B). The presence of BPDE adducts retards mobility of the
adducted fragments through the gel, which translates into the shift of the corresponding bands (Fig. 4B). Scanning of the gel using
the Cyclon system detected at least 99.3% of a band corresponding to
the adducted fragment in adducted samples. This indicates that the
adducts were very stable during the preparation of the plasmid vectors
and that the adducted vectors contain less than 0.7% of contaminants.
|
Transfection of Cells with Adducted Plasmid Vectors-- Four adducted plasmids and a non-adducted control were separately transfected into repair-proficient V79 cells and repair-deficient V-H1 cells. Each experiment was repeated at least twice with a different plasmid preparation. Doubling times of exponentially growing V79 and V-H1 cells were ~14 and ~18 h, respectively. After transfection, doubling times of V79 and V-H1 cells increased to ~19 and ~34 h, respectively. There was no difference in proliferation of the cells transfected with different plasmid constructs. For the selection with antibiotics, cells were seeded in a density ensuring approximately the same yield of cellular colonies per plate from both cell lines. Because the cells kept proliferating at a time between the transfection and seeding (48 h), and their proliferative rate was different, it is reasonable to expect that 5.8 of the V79 cellular colonies and 2.7 of the V-H1 cellular colonies on average originated from a single cell. Thus, samples harvested for mutation analysis contained a random mixture of cellular colonies (pools) that were not necessarily independent. A sufficient total number of colonies (~600) in random pools was harvested from each plasmid transfection to ensure that at least 100 of these were independent clones. Assuming that permanent transfection decreases the proliferative rate of the affected cells more than the non-affected cells, which are not selected, this number of analyzed independent colonies is the lowest possible estimate.
Quantitative Minisequencing and Calculation of the Mutation
Level at a Specific Site--
Chromosomal DNA from pooled cellular
samples was used for amplification of the 337-bp DNA template
encompassing a region originally containing the adducted nucleotide.
The amplified template is a mixture of fragments with a DNA sequence
reflecting the mutagenicity of the adduct at the region. Quantitative
minisequencing was used to assess both
the type and the level of mutations at the region as described under
"Materials and Methods" and Fig.
5.
|
|
To validate accuracy and sensitivity of the assay model, binary
mixtures of 337-bp fragments differing in a single nucleotide at each
site of the region (pseudo-pools) were analyzed. These fragments were
prepared by PCR amplification of 337-bp region from the HPRT
cDNA manipulated in plasmid vectors by site-directed mutagenesis.
The relative content of one fragment in the other was 50, 25, 12.5, 6.25, 3.13, 1.56, 0.78, 0.39, 0.19, and 0%. Composition of these
binary mixtures represented all potential single nucleotide mutations
in the analyzed region
(5'-A+3C+2T+1G0G1G
2A
3). The ratio between a level of the fragment detected by the assay (fmeasured) and a predicted level of the
fragment in the mixture (fpredicted) represents
a correction factor (k). The means ± S.D. of the
correction factor were calculated at each analyzed point from at least
three independent experiments and plotted. The relationship between
k and the level of the analyzed fragment was found linear between 0.78-50%. Fig. 8 is included as Supplemental Material and
shows the results from analyses of model binary fragment mixtures differing by a nucleotide in positions found mutated in the study (C+2, G0, and G
1). The average
k values (0.85-1.05) calculated for each mutation in the
region from model binary mixtures (the rest of the results not shown)
were used to re-calculate levels of mutations found in unknown samples.
The results demonstrate accuracy and linearity of the assay in tested
binary mixtures containing more than 1% of the analyzed fragment.
Accuracy of the assay decreases sharply when levels of the analyzed
fragment are lower than 1% and limits sensitivity of the assay to
0.3% (see Supplemental Material Fig. 8). Thus, mutagenic changes at levels higher than 0.3% can be qualitatively detected and at levels higher than 1% can be accurately quantified by the assay.
Mutagenicity of BPDE Adducts in Repair-proficient and
Repair-deficient Cells--
Mutagenicity of 10S-BPDE-dG
and 10R-BPDE-dG lesions in the sequence
5'-A+3C+2T+1G0G1G
2A
3 was examined at the adducted sites (G0 and
G
1) and also at their 5'- and 3'-flanking sites (from +3
to
3, Fig. 2 and also Fig. 6). G0 corresponds to the
hotspot (G-634) of the non-transcribed strand of the gene found in
random mutagenesis studies of (+)-BPDE-2 in V79 cells (10, 32).
In repair-proficient V79 cells, the guanine adduct derived from
(+)-BPDE-2, 10S-BPDE-dG, induced 2.5 ± 1.3 and
2.8 ± 1.1% of G T mutations of the adducted nucleotides at
the G0 and G
1 sites, respectively. The
guanine adduct derived from (
)-BPDE-2, 10R-BPDE-dG,
induced 1.8 ± 1.0 and 1.9 ± 0.7% of G
T mutations of
the adducted nucleotides at the G0 and G
1
sites, respectively (Fig. 7A).
Analysis of secondary mutations of nucleotides at sites flanking the
lesion on the 5'-end revealed that the 10S adduct, but not
the 10R adduct, induced C
T mutations (0.7 ± 0.4%) at the (+2) position when the G0 site was adducted,
and G
T mutation (1.3 ± 0.5%) at the G0 position
when the G
1 site was adducted. Interestingly, adducts at
the G0 or G
1 site did not induce mutations of
the T nucleotide located at the (+1) position. Also, no mutations were
found in the 3'-vicinity of the adducted sites (up to 3 nucleotides) or
further upstream in the 5'-direction (3 and 4 positions analyzed
overall from the adducted G0 and G
1 sites,
respectively).
|
The effect of repair deficiency in the V-H1 cells on the observed
frequency of mutations is relatively small (a factor of 2 or less) and
is only significant for the constructs containing 10S
adducts at G0 (Fig. 7B). For example, in the
VH-1 cells, the 10S adduct induced 4.6 ± 1.5 and
2.1 ± 0.6% of G T mutations of the adducted nucleotides at
G0 and G
1 sites, as compared with 2.5 and
2.8%, respectively, in the V79 cells (see above). The 10R
adduct induced 1.9 ± 0.8 and 1.2 ± 0.6% of G
T mutations at the adducted nucleotides when at the G0
and G
1 sites, respectively. Analysis of secondary
mutations of nucleotides at sites flanking the lesion on the 5'-end
showed the same pattern of mutations found in V79 cells; the
10S adduct, but not the 10R adduct, induced C
T mutations (1.7 ± 0.6%) at the (+2) position when the
G0 site was adducted and G
T mutations (1.5 ± 0.7%) at the G0 position when G
1 was the
adducted site. As with the V79 cells, no other mutation type or other
mutated nucleotide was found in the analyzed region. The formation of
secondary mutations depends on the presence of the 10S
adduct but probably not on the formation of primary mutations. If so,
tandem mutations (two mutations on the same analyzed DNA fragment)
would be detected. In the primer extension assay tandem T mutations
would give rise to multiple A incorporations in the primers used. No
such multiple extensions of the primer were observed. Although we
cannot exclude the possibility of formation of tandem mutations in
levels below the detection limit of the assay (0.3%), random formation
of secondary mutations independent of the primary mutations is a more
probable scenario. Large standard deviations of the means of the
presented data originate in large differences in the individual data
from each of the 10 samples containing pools of cellular colonies, not
from irreproducibility of the assay. Repeated analyses of the same
samples showed remarkable reproducibility (maximum scatter did not
exceed 10% of the calculated values, data not shown).
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
At the target sites (G0 and G1), the
10S adduct and the 10R adduct induced only G
T mutations. This mutation was found to be dominant in previous studies
examining mutagenicity of site-specific adducts both in prokaryotic
(31, 36, 37) and eukaryotic cells (37), although G
C and G
A
mutations have also been observed in some sequences. The sequence
context of our adducted G0 site is identical to the
~TGG~ sequence examined in simian kidney cells (37).
There was no significant difference between mutagenicity at the
adducted site for the 10S-BPDE-dG lesion relative to the
10R-BPDE-dG lesion, and a preponderance of G
T mutations
was observed, as in the present study. However, the level of mutation
induced by both adducts was substantially higher than in our study (13 versus 1.8-2.5%), and low levels (<1%) of G
C and G
A mutations were also detected. These differences may stem mainly
from differences in the experimental systems; the single-stranded
vector system used by Page et al. (31) and Moriya et
al. (37) eliminates the involvement of DNA repair, whereas the
double-stranded vector system used in our study is sensitive to DNA
repair (both in V79 and in V-H1 cells, which have ~50% remaining
capacity to remove adducts derived from the (+)-(7R,8S,9S,10R)-enantiomer
of BPDE from DNA compared with V79 cells (27)). Consequently, the
results of our study show substantially lower mutagenicity of the
examined adducts and reflect both DNA repair of the adducts and
fidelity of their bypass.
Results from DNA repair-proficient (V79) and DNA repair-deficient
(V-H1) cells show only minor differences. It is possible that the
defect of V-H1 cells in the xeroderma pigmentosum complementation group
D/ERCC2 gene (25) influences mainly transcriptional
coupled repair (18) and causes a decreased ability of the cells to
preferentially repair BPDE adducts from the transcribed strand of
active genes (27), whereas the efficiency of global genomic repair
(18), which is responsible for the repair of the non-transcribed strand (location of the adducts in this study), may be affected only marginally. Both lesions in both adducted G sites generated the same
kind of mutation (G T), and the 10S adduct induced
identical secondary mutations in these two cell lines. Quantitatively,
no significant difference in mutagenicity of the 10R adduct
was observed in corresponding sites in V79 and V-H1 cells. However,
mutagenicity of the 10S adduct was significantly higher at
the G0 site in the V-H1 cells than in the V79 cells
(p < 0.006), although similar at the G
1
site in both cell lines (p > 0.1). The data suggest that repair of the 10S adduct is more efficient than the
10R adduct in the non-transcribed strand but also that
repair efficiency of the 10S adduct may differ from site to
site. These results are in accordance with a study by Custer et
al. (29) who demonstrated a remarkable resistance of the
10R adduct to DNA repair in vitro compared with
10S adduct using a whole cell extract. However, in a
different sequence context, no difference in repair efficiency of these
two adducts in vitro was observed (38).
Mutations remote from a specifically modified base in DNA were first
described by Lambert et al. (39) for frameshifts induced by
an acetylaminofluorene adduct. The present study reports a definitive
demonstration of substitution mutations remote from the target site
induced by a single, site-specific polycyclic aromatic hydrocarbon DE
adduct. These non-targeted mutations were observed only when the adduct
has the 10S configuration and were found within two
nucleotides on the 5'-side of the adducted base, namely at
C2 when G0 was modified and at G0
when G1 was modified (for sequence see Fig. 2). Notably,
the absence of non-targeted mutations with 10R adducts at
either position provides strong internal evidence that these mutations
with the 10S adducts do not result from any artifact of the
oligonucleotide synthesis, since for each sequence the 10R
and 10S adducted oligonucleotides were prepared together
from the mixed 10R/10S diastereomers of the
phosphoramidite (see "Materials and Methods"), and thus the two
diastereomeric oligonucleotides underwent identical treatment prior to
final chromatographic purification. The non-targeted mutations at both
sites represented mutagenic changes of the original nucleotides to T,
suggesting their bypass with a mismatched A. Furthermore, the T that is
immediately 5' to the G0 lesion or separated by one base
from the G
1 lesion was not found to be mutated in the
presence of either lesion. An attractive explanation for these data is
that translesion synthesis results in A being inserted (either
correctly or incorrectly) as the preferential nucleotide at the
adducted site and at the two additional 5'-flanking nucleotides.
In vitro studies with isolated bypass DNA polymerases (pol
,
,
, and
) on templates containing a single
10S-BPDE-dG or 10R-BPDE-dG lesion identified
pol
as involved in low fidelity insertions at these lesions in
vitro (40, 41). The enzyme incorporates mostly A and G opposite
the lesion site (40, 41) and thus is likely to be responsible at least
in part for mutations induced by BPDE-dG lesions in mammalian cells
(mostly G
T with minor G
C and G
A). Although the enzyme
extends the primer beyond the lesion site rather inefficiently, its
preference to extend mispaired primers containing a purine opposite the
adduct and its high misincorporation frequency on non-damaged templates (10
3-10
2) (41) are features that may
contribute to the formation of non-targeted mutations and would be
worthy of further exploration by studies in vitro and in
cells lacking functional pol
.
The 10S BPDE adducts at the G0 and
G1 sites induced identical non-targeted mutations in both
repair-proficient (V79) and repair-deficient (V-H1) cell lines, and the
frequency of the non-targeted mutations correlated with the frequency
of targeted mutations. The increased level of the targeted mutation at
the G0 site (G
T) in V-H1 cells was accompanied by a
similar increase in the level of the non-targeted mutations (C
T) in these cells compared with V79 cells, whereas no difference in the
frequency of non-targeted mutation (G
T at the G0 site)
between V-H1 and V79 cells was observed when the levels of the targeted
mutation (G
T at the G
1 site) were similar in both
cell lines. Assuming that the differences in the targeted mutation
frequency between these two cell lines reflect only the efficiency of
the 10S adduct removal from a particular site, it seems
likely that the adduct level determines the frequency of not only
targeted but also non-targeted mutations.
The present observation that mutations are induced 1 or more bases away
from the target site by a polyaromatic hydrocarbon DE lesion is
consistent with results of our recent study (42) of mutations in an
E. coli-M13 system induced by several cis-opened adducts derived from both BPDE-2 (the diastereomer used in the present
study, whose benzylic hydroxyl group and epoxide oxygen are
trans) and BPDE-1 (benzylic hydroxyl group and epoxide
oxygen cis). More limited data in E. coli by
Jelinsky et al. (36) using a single trans-opened
10S adduct of BPDE-2 had also led to the tentative
suggestion of non-targeted mutations at a base immediately adjacent to
this adduct. In our previous study (42) using the sequence
~G6C5G4G3G2G1G0~
with cis-opened adducts at G0,
non-targeted substitution mutations 4 and 6 bases remote from the
target site were described, but their significance was not fully
recognized. No significant non-targeted substitutions had been detected
in this DNA sequence containing trans-opened BPDE-1 or
BPDE-2 dG adducts in the same experimental system (31). The most
prevalent non-targeted mutations (frequency 2-4%) induced by the
cis-opened BPDE-1 and BPDE-2 dG adducts were T substitutions
at G6 and G4, and they occurred both in the
absence of mutations at the target site (G0) and in
combination with G0 T mutations at this site. These
non-targeted mutations were not observed with the control (non-adducted) sequence, and most significantly they were observed only
when the adducts at G0 had 10S but not when they
had 10R configuration, in analogy to our present
observations. Because no significant non-targeted mutations were
observed in a different (~G6C5G4T3T2C1G0~)
sequence containing BPDE-1 or -2 adducts at G0 (42), the
above mutations were most likely related to the run of 5 guanines in
the
~G6C5G4G3G2G1G0~
sequence. Despite the specific sequence effect, as well as the
differences in experimental systems, the similarity between these
results in E. coli and the present observations in mammalian
cells is intriguing and suggests that mutations remote from the lesion
site induced by polyaromatic hydrocarbon DE adducts may
constitute a not uncommon mechanism of polyaromatic hydrocarbon
mutagenesis whose significance has not been appreciated previously.
A limited number of site-specific mutagenesis studies with other types
of bulky DNA adducts has demonstrated induction of non-targeted
substitution mutations at non-adducted sites in the vicinity of the
lesion. They include studies of the N7-guanyl adduct of
aflatoxin B1 8,9-epoxide and the adduct's ring opened formamidopyrimidine form in SOS-induced E. coli (43, 44) as well as N-deoxyguanosin-8-yl)-2-acetylaminofluorene and
N(deoxyguanosin-8-yl)-2-aminofluorene in COS-7 cells
(45). Although it is difficult to make any generalization about the
mechanism of these non-targeted mutations by comparing the data of
these studies to those presented here (different experimental models,
structure of the adducts, and sequence context), it is clear at this
point that the formation of various non-targeted mutations is likely
related to the bulky character of the adducts and that the orientation
of the adducts may play a significant role. The hydrocarbon of the
10S BPDE-dG adduct orients toward the 5'-side of the
modified base in the minor groove of duplex DNA in the sequence
5'-CGC (15) as well as 5'-TGC (46) and could
thus cause structural perturbations 5' to the lesion in our study
(5'-TGG and 5'-GGG), resulting in the observed
non-targeted mutations 5' to the adduct. In contrast, the
10R BPDE-dG adduct, which orients in the opposite direction
toward the 3'-end of the modified strand in duplex DNA (16), was not
found to cause any mutations at non-target bases on either side of the
adduct. Interestingly, about 13% of the total mutations in SOS-induced
E. coli caused by a dG adduct of aflatoxin B1
8,9-epoxide (43), which intercalates into the helix on the 5'-side of
the modified G base (47, 48), were primarily C T mutations at the
dC immediately 5' to the lesion (G) in a 5'~CGA~
sequence. In contrast, an
N-(deoxyguanosin-8-yl)-2-acetylaminofluorene lesion,
whose hydrocarbon moiety displaces the modified G and intercalates in
its place opposite the complementary C (49), gave rise to base
misincorporation on the 3'-side of the lesion site (50).
The dependence of non-targeted mutations on adduct configuration is of
particular interest in light of the marked differences in mutagenicity
and carcinogenicity between (+)- and ()-BPDE-2 enantiomers. Higher
tumorigenic activity of (+)-BPDE-2 compared with (
)-BPDE-2 has been
demonstrated. (+)-BPDE-2 induced lung tumors and skin tumors when
injected intraperitoneally into newborn mice and applied topically to
the skin of adult mice, respectively; but (
)-BPDE had little or no
carcinogenic activity (7, 8). Although the present mutagenesis study
showed little difference in the mutagenicity of (+)- and (
)-BPDE-2 dG
adducts at specific sites, (+)-BPDE-2 was ~11 times more mutagenic
than (
)-BPDE-2 in the HPRT gene of V79 cells on a per dose
basis in a random mutagenesis study (11). Many factors can contribute
to these different results, including more efficient adduct formation
from (+)- relative to (
)-BPDE-2 (12), differences in relative
proportions of dG and dA adducts formed from the two enantiomers (12,
13), as well as sequence effects on adduct formation, repair (51), and
bypass (41). Furthermore, the higher mutagenicity of the (+)-BPDE-2
adducts observed in random mutagenesis studies may result in part from
their mutagenic effects on the whole region (i.e.
non-targeted mutations in the vicinity of the adduct) rather than only
at a particular adducted site. Because not all DNA mutations lead to
amino acid substitutions, and some substitutions may be silent in terms
of function, it is obvious that induction of a mutation at more than
one site by a single adduct increases the probability of an amino acid
change that could generate a protein with a compromised function.
Consequently, the mutagenic and tumorigenic potential of a BPDE adduct
leading to "multiposition" mutations would be significantly higher.
In summary, results of this study contradict the intuitive notion that
point mutations arise exclusively by erroneous replication of the
modified base and suggest that the higher mutagenic and carcinogenic
activity of (+)-BPDE-2 compared with (
)-BPDE-2 may partly stem from
the capability of its major dG adduct to induce DNA mutations at
multiple sites.
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The on-line version of this article (available at
http://www.jbc.org) contains Supplemental Fig. 8 documenting the
linearity and detection limit of the quantitative minisequencing method.
§ To whom correspondence should be addressed: Susan Lehman Cullman Laboratory for Cancer Research, Dept. of Chemical Biology, College of Pharmacy, Rutgers, State University of New Jersey, 164 Frelinghuysen Rd., Piscataway, NJ 08854-8020. Tel.: 732-445-3400 (ext. 238); Fax: 732-445-0687; E-mail: kramata@rci.rutgers.edu.
Present address: Dept. of Chemistry, City College of CUNY, New
York, NY 10031.
Published, JBC Papers in Press, February 20, 2003, DOI 10.1074/jbc.M211557200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: DE, diol epoxide; BPDE-1 and BPDE-2, the 7,8-diol 9,10-epoxide diastereomers in which the benzylic 7-hydroxyl group and epoxide oxygen are either cis or trans, respectively; 10S-BPDE-dG and 10R-BPDE-dG, trans-opened N2-dG adducts derived from the (+)-(7R,8S,9S,10R)- and (-)-(7S,8R,9R,10S)-enantiomers of BPDE-2, respectively; HPRT, hypoxanthine (guanine) phosphoribosyltransferase; DTT, dithiothreitol.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Dipple, A., Moschel, R., and Bigger, C. A. H. (1984) ACS Monogr. 2, 41-163 |
2. | Sims, P., Grover, P. L., Swaisland, A., Pal, K., and Hewer, A. (1974) Nature 252, 326-328[Medline] [Order article via Infotrieve] |
3. | Yagi, H., Hernandez, O., and Jerina, D. M. (1975) J. Am. Chem. Soc. 97, 6881-6883[Medline] [Order article via Infotrieve] |
4. | Yagi, H., Akagi, H., Thakker, D. R., Mah, H. D., Koreeda, M., and Jerina, D. M. (1977) J. Am. Chem. Soc. 99, 2358-2359[Medline] [Order article via Infotrieve] |
5. | Thakker, D. R., Yagi, H., Akagi, H., Koreeda, M., Lu, A. H., Levin, W., Wood, A. W., Conney, A. H., and Jerina, D. M. (1977) Chem. Biol. Interact. 16, 281-300[CrossRef][Medline] [Order article via Infotrieve] |
6. | Wood, A. W., Chang, R. L., Levin, W., Yagi, H., Thakker, D. R., Jerina, D. M., and Conney, A. H. (1977) Biochem. Biophys. Res. Commun. 77, 1389-1396[Medline] [Order article via Infotrieve] |
7. | Buening, M. K., Wislocki, P. G., Levin, W., Yagi, H., Thakker, D. R., Akagi, H., Koreeda, M., Jerina, D. M., and Conney, A. H. (1978) Proc. Natl. Acad. Sci. U. S. A. 75, 5358-5361[Abstract] |
8. | Slaga, T. J., Bracken, W. J., Gleason, G., Levin, W., Yagi, H., Jerina, D. M., and Conney, A. H. (1979) Cancer Res. 39, 67-71[Medline] [Order article via Infotrieve] |
9. | Chen, R. H., Maher, V. M., and McCormick, J. J. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 8680-8684[Abstract] |
10. | Wei, S. J., Chang, R. L., Bhachech, N., Cui, X. X., Merkler, K. A., Wong, C. Q., Hennig, E., Yagi, H., Jerina, D. M., and Conney, A. H. (1993) Cancer Res. 53, 3294-3301[Abstract] |
11. | Wei, S. J., Chang, R. L., Hennig, E., Cui, X. X., Merkler, K. A., Wong, C. Q., Yagi, H., Jerina, D. M., and Conney, A. H. (1994) Carcinogenesis 15, 1729-1735[Abstract] |
12. | Sayer, J. M., Chadha, A., Agarwal, S. K., Yeh, H. J. C., Yagi, H., and Jerina, D. M. (1991) J. Org. Chem. 56, 20-29 |
13. | Vepachedu, S. R., Ya, N., Yagi, H., Sayer, J. M., and Jerina, D. M. (2000) Chem. Res. Toxicol. 13, 883-890[CrossRef][Medline] [Order article via Infotrieve] |
14. | Cheng, S. C., Hilton, B. D., Roman, J. M., and Dipple, A. (1989) Chem. Res. Toxicol. 2, 334-340[Medline] [Order article via Infotrieve] |
15. | Cosman, M., de los Santos, C., Fiala, R., Hingerty, B. E., Singh, S. B., Ibanez, V., Margulis, L. A., Live, D., Geacintov, N. E., Broyde, S., and Patel, D. J. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1914-1918[Abstract] |
16. | de los Santos, C., Cosman, M., Hingerty, B. E., Ibanez, V., Margulis, L. A., Geacintov, N. E., Broyde, S., and Patel, D. J. (1992) Biochemistry 31, 5245-5252[Medline] [Order article via Infotrieve] |
17. |
Wood, R. D.
(1997)
J. Biol. Chem.
272,
23465-23468 |
18. |
de Laat, W. L.,
Jaspers, N. G.,
and Hoeijmakers, J. H.
(1999)
Genes Dev.
13,
768-785 |
19. | Ohmori, H., Friedberg, E. C., Fuchs, R. P., Goodman, M. F., Hanaoka, F., Hinkle, D., Kunkel, T. A., Lawrence, C. W., Livneh, Z., Nohmi, T., Prakash, L., Prakash, S., Todo, T., Walker, G. C., Wang, Z., and Woodgate, R. (2001) Mol. Cell 8, 7-8[Medline] [Order article via Infotrieve] |
20. |
Johnson, R. E.,
Washington, M. T.,
Prakash, S.,
and Prakash, L.
(2000)
J. Biol. Chem.
275,
7447-7450 |
21. |
Zhang, Y.,
Yuan, F.,
Xin, H.,
Wu, X.,
Rajpal, D. K.,
Yang, D.,
and Wang, Z.
(2000)
Nucleic Acids Res.
28,
4147-4156 |
22. |
Vaisman, A.,
Tissier, A.,
Frank, E. G.,
Goodman, M. F.,
and Woodgate, R.
(2001)
J. Biol. Chem.
276,
30615-30622 |
23. | Courtemanche, C., and Anderson, A. (1999) Mutat. Res. 430, 23-36[Medline] [Order article via Infotrieve] |
24. | Zdzienicka, M. Z., and Simons, J. W. (1987) Mutat. Res. 178, 235-244[Medline] [Order article via Infotrieve] |
25. | Kadkhodayan, S., Salazar, E. P., Ramsey, M. J., Takayama, K., Zdzienicka, M. Z., Tucker, J. D., and Weber, C. A. (1997) Mutat. Res. 385, 47-57[Medline] [Order article via Infotrieve] |
26. | Hoeijmakers, J. H., Egly, J. M., and Vermeulen, W. (1996) Curr. Opin. Genet. & Dev. 6, 26-33[Medline] [Order article via Infotrieve] |
27. |
Schiltz, M.,
Cui, X. X.,
Lu, Y. P.,
Yagi, H.,
Jerina, D. M.,
Zdzienicka, M. Z.,
Chang, R. L.,
Conney, A. H.,
and Wei, S. J.
(1999)
Carcinogenesis
20,
2279-2286 |
28. | Simhadri, S., Kramata, P., Zajc, B., Sayer, J., Jerina, D., Hinkle, D., and Wei, C. (2002) Mutat. Res. 508, 137-145[Medline] [Order article via Infotrieve] |
29. | Custer, L., Zajc, B., Sayer, J. M., Cullinane, C., Phillips, D. R., Cheh, A. M., Jerina, D. M., Bohr, V. A., and Mazur, S. J. (1999) Biochemistry 38, 569-581[CrossRef][Medline] [Order article via Infotrieve] |
30. | Moore, P. D., Koreeda, M., Wislocki, P. G., Levin, W., Conney, A. H., Yagi, H., and Jerina, D. M. (1977) in Drug Metabolism Concepts (Jerina, D. M., ed) , pp. 127-154, American Chemical Society, Washington, D. C.ACS Symposium Series no. 44 |
31. | Page, J. E., Zajc, B., Oh-hara, T., Lakshman, M. K., Sayer, J. M., Jerina, D. M., and Dipple, A. (1998) Biochemistry 37, 9127-9137[CrossRef][Medline] [Order article via Infotrieve] |
32. | Wei, S. J., Chang, R. L., Wong, C. Q., Bhachech, N., Cui, X. X., Hennig, E., Yagi, H., Sayer, J. M., Jerina, D. M., Preston, B. D., and Conney, A. H. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 11227-11230[Abstract] |
33. | Sambrook, J., and Russell, D. W. (2001) Molecular Cloning: A Laboratory Manual , 3rd Ed., Vol. I , pp. 3.42-3.52, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY |
34. | Syvanen, A. C. (1999) Hum. Mutat. 13, 1-10[CrossRef][Medline] [Order article via Infotrieve] |
35. |
Haff, L. A.,
and Smirnov, I. P.
(1997)
Genome Res.
7,
378-388 |
36. | Jelinsky, S. A., Liu, T., Geacintov, N. E., and Loechler, E. L. (1995) Biochemistry 34, 13545-13553[Medline] [Order article via Infotrieve] |
37. | Moriya, M., Spiegel, S., Fernandes, A., Amin, S., Liu, T., Geacintov, N., and Grollman, A. P. (1996) Biochemistry 35, 16646-16651[CrossRef][Medline] [Order article via Infotrieve] |
38. | Hess, M. T., Gunz, D., Luneva, N., Geacintov, N. E., and Naegeli, H. (1997) Mol. Cell. Biol. 17, 7069-7076[Abstract] |
39. | Lambert, I. B., Napolitano, R. L., and Fuchs, R. P. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1310-1314[Abstract] |
40. |
Rechkoblit, O.,
Zhang, Y.,
Guo, D.,
Wang, Z.,
Amin, S.,
Krzeminsky, J.,
Louneva, N.,
and Geacintov, N. E.
(2002)
J. Biol. Chem.
277,
30488-30494 |
41. |
Chiapperino, D.,
Kroth, H.,
Kramarczuk, I. H.,
Sayer, J. M.,
Masutani, C.,
Hanaoka, F.,
Jerina, D. M.,
and Cheh, A. M.
(2002)
J. Biol. Chem.
277,
11765-11771 |
42. | Pontén, I., Kroth, H., Sayer, J. M., Dipple, A., and Jerina, D. M. (2001) Chem. Res. Toxicol. 14, 720-726[Medline] [Order article via Infotrieve] |
43. |
Bailey, E. A.,
Iyer, R. S.,
Stone, M. P.,
Harris, T. M.,
and Essigmann, J. M.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
1535-1539 |
44. |
Smela, M. E.,
Hamm, M. L.,
Henderson, P. T.,
Harris, C. M.,
Harris, T. M.,
and Essigmann, J. M.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
6655-6660 |
45. | Shibutani, S., Suzuki, N., Tan, X., Johnson, F., and Grollman, A. P. (2001) Biochemistry 40, 3717-3722[CrossRef][Medline] [Order article via Infotrieve] |
46. | Fountain, M. A., and Krugh, T. R. (1995) Biochemistry 34, 3152-3161[Medline] [Order article via Infotrieve] |
47. | Gopalakrishnan, S., Harris, T. M., and Stone, M. P. (1990) Biochemistry 29, 10438-10448[Medline] [Order article via Infotrieve] |
48. | Giri, I., Jenkins, M. D., Schnetz-Boutaud, N. C., and Stone, M. P. (2002) Chem. Res. Toxicol. 15, 638-647[CrossRef][Medline] [Order article via Infotrieve] |
49. | O'Handley, S. F., Sanford, D. G., Xu, R., Lester, C. C., Hingerty, B. E., Broyde, S., and Krugh, T. R. (1993) Biochemistry 32, 2481-2497[Medline] [Order article via Infotrieve] |
50. | Suzuki, N., Ohashi, E., Hayashi, K., Ohmori, H., Grollman, A. P., and Shibutani, S. (2001) Biochemistry 40, 15176-15183[CrossRef][Medline] [Order article via Infotrieve] |
51. | Wei, D., Maher, V. M., and McCormick, J. J. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 2204-2208[Abstract] |