Mutagenesis vol. 19 no. 2 pp. 91-97,
March 2004
© 2004 UK Environmental Mutagen Society/Oxford University Press
Comparison of mutagenic potentials and mutation spectra of benzene metabolites using supF shuttle vectors in human cells
1Department of Global Environmental Engineering, Graduate School of Engineering, Kyoto University, Sakyo-ku, Kyoto 606-8501, Japan 2Division of Radiobiology and Environmental Science, Research Institute for Advanced Science and Technology, Osaka Prefecture University, 1–2 Gakuen-cho, Sakai-city, Osaka 599-8570, Japan
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Abstract |
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Benzene is a human leukemogen and the metabolites are thought to be deeply involved in benzene leukemogenesis. In a previous study we reported the molecular analysis of p-benzoquinone (p-BQ) mutagenesis by using a supF shuttle vector plasmid and here we report the mutagenesis of the other metabolites, hydroquinone (HQ) and trans, trans-muconaldehyde (MUC). HQ is a precursor of p-BQ and MUC is produced by a ring-opening metabolic pathway. We found that the HQ redox cycle produced an oxidative lesion in plasmid DNA and significant differences among the mutagenic potentials of MUC, HQ and p-BQ. HQ has stronger mutagenicity than the others. It is about 20 and 600 times stronger than p-BQ and MUC, respectively. Furthermore, we found notable differences in each mutational feature. The MUC mutational type was characterized by a high frequency of tandem base substitutions that could be due to crosslinks produced by its aldehyde moieties, while HQ was characterized by frequent deletion. This HQ feature is the same as in vivo benezene mutagenesis of Big Blue mice reported by Provost et al. in 1996 and is also quite similar to a hydrogen peroxide mutational feature. Therefore, we presume that HQ and reactive oxygen species may play an important role in benzene carcinogenesis.
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Introduction |
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Large amounts of benzene have been used industrially for the synthesis of organic compounds and as an additive in automobile fuel. It is a ubiquitous environmental pollutant and classified as carcinogenic to humans (IARC 1977, 1987
) and experimental animals (National Toxicology Program, 1986).
Although benzene itself is not regarded as a mutagenic substance, some of its metabolites are genotoxic. Benzene is first metabolized to phenol in the liver, subsequently to hydroquinone (CAS no. 123-31-9) (HQ) in the bone marrow and then converted non-enzymatically to p-benzoquinone (CAS no. 106-51-4) (p-BQ) (Goldstein and Witz, 1992). These are considered to be ultimately toxic metabolites and generate reactive oxygen species such as O2– and H2O2 in redox cycling mediated by intracellular NADH and copper (Hiraku and Kawanishi, 1996). The alternative metabolic route of benzene is the ring-opening process. Trans,trans-muconaldehyde (CAS no. 18409-46-6) (MUC) is produced in this pathway. MUC is a potent bone marrow toxin in mice, since its administration results in a significant decrease in the red blood cell count, hematocrit and hemoglobin bone marrow cellularity (Witz et al., 1985). Additionally, it is weakly mutagenic to bacteria in the Ames assay in both the presence and absence of S9 mix (Glatt and Witz, 1990; Snyder et al., 1993), while manifesting remarkably in mammalian V79 cells (Glatt and Witz, 1990).
We previously examined p-BQ-induced mutations in mouse and human cells using supF shuttle vector plasmids (Nakayama et al., 1999, 2000) and found that p-BQ itself produces DNA adducts and induces base substitutions mostly at G:C base pairs.
In this study we have analyzed the mutations induced by the other metabolites, MUC and HQ, using shuttle vector plasmids pMY189 in human cells and compared their mutagenicity. We found significant differences among the mutagenic potentials and mutation spectra of MUC, HQ and p-BQ. These results will provide a better insight into understanding which metabolite is important in benzene leukemogenesis.
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Materials and methods |
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Chemicals and cells
MUC was kindly provided by Dr Witz (UMDNJ–Robert Wood Johnson Medical School, Rutgers University). HQ and other chemicals were obtained from Wako (Osaka, Japan). Human WI38-VA13 cells (Girardi et al., 1965
) were purchased from the American Type Culture Collection (Rockville, MD). The cells were cultured in Dulbecco’s modified minimum essential medium supplemented with fetal bovine serum (10%) in a humidified atmosphere of 5% CO2 at 37°C. The supF shuttle vector plasmids pMY189 (Matsuda et al., 1995) were used for the mutation analysis and the purified stock plasmid was prepared with a Qiagen plasmid purification kit (Qiagen, Hilden, Germany). Escherichia coli KS40/pKY241 (Akasaka et al., 1992) was used as a mutational indicator in this study.
SupF shuttle vector mutation assay
The plasmids (20 µg/ml) were treated with MUC at final concentrations of 0–100 mM in Tris–HCl buffer (pH 8.0) for 1 h at 37°C. They were also treated with HQ at final concentrations of 0–50 mM in Tris–HCl buffer (pH 8.0) with or without NADH (250 µM) and Cu(II) (CuCl2, 20 µM) for 1 h at 37°C. After treatment with chemicals, the plasmids were precipitated with ethanol to remove unreacted excess chemicals and redissolved in 50 µl of Tris–HCl buffer (pH 8.0). The plasmids were subjected to 0.7% agarose gel electrophoresis to examine the DNA strand breaks caused by the chemicals. As a result of this electrophoresis, we determined the concentration of the chemicals used to treat the plasmids in the plasmid mutation assay.
After the purification of plasmids, transfection into human WI38-VA13 cells (3 x 107 cells) was conducted by electroporation. Then, after 3 days incubation, the plasmids were extracted with a QIAprep-spin Plasmid kit (Qiagen) (Nakayama et al., 2000) and introduced into the indicator E.coli KS40/pKY241 to select for the mutated supF gene.
The base sequences of the mutated supF gene were determined with a 373A automatic DNA sequencer (Applied Biosystems, Foster City, CA). Details of the experimental method were described previously (Nakayama et al., 2000).
Analysis of 8-OH-dG
Plasmid DNA treated with HQ/Cu(II)/NADH was digested with RNase to remove contaminated RNA and then hydrolyzed to nucleotides by nuclease P1 (4 U) (Yamasa, Japan) at 37°C for 1 h, followed by incubation with alkaline phosphatase (Type III; Sigma, St Louis, MO) at 37°C for 1 h.
The levels of 8-hydroxydeoxyguanosine (8-OH-dG) in the plasmids were measured with an HPLC-ECD system which consists of a liquid chromatograph LC10-AD equipped with a damper (Shimadzu, Japan) and EC detector LC-4C (Bioanalytical Systems, IN). A Cap Cell Pak C18 column (25 x 1 cm) (Shiseido, Japan) was used for HPLC. The eluent was 20 mM sodium acetate/10 mM citric acid buffer (pH 4.3) containing 10% methanol. The column temperature was 37°C and the flow rate was 1.0 ml/min. Determination of the nucleoside 8-OH-dG was performed with an EC detector (redox potential 0.6 V) and nucleoside deoxyguanosine (dG) was measured with a Waters M484 (Waters, MA) UV detector (wavelength 280 nm). Quantification of the nucleoside dG and 8-OH-dG was performed by comparison with corresponding standards purchased from Sigma.
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Results |
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DNA strand breaks by HQ
Figure 1 shows the electrophoresis pattern of the pMY189 plasmid treated with HQ. In the presence of Cu(II) and NADH, the amount of open circular plasmids (Form II) increased with HQ concentration and DNA fragmentation was observed with the 1 and 10 mM HQ treatments. On the other hand, no increase in open circular plasmids or DNA fragmentation was observed in the absence of Cu(II) and NADH. The electrophoresis pattern of MUC-treated pMY189 (maximum 100 mM) remained the same as control plasmids (data not shown).
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8-OH-dG in the plasmids
To determine whether HQ yields oxidative DNA lesions or not, we measured the 8-OH-dG level in plasmid DNA treated with 50 µM HQ in the presence of Cu(II) and NADH. Figure 2 shows the levels of 8-OH-dG per 105 dG of control and HQ-treated plasmids. The background level of 8-OH-dG was 0.98 per 105 dG and 50 µM HQ increased the amount of 8-OH-dG to 117 per 105 dG. A significant difference was seen compared with the control by t-test. This result suggests that HQ/Cu(II)/NADH generated reactive oxygen species and caused oxidative DNA damage.
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Plasmid survival and mutation frequencies
Both MUC and HQ reduced the number of bacterial colonies whereas they increased the proportion of bacteria carrying the mutated supF gene in a dose-dependent manner (Figures 3 and 4). When the plasmids were treated with 100 mM MUC, the mutation frequency was 5.2 x 10–3, or 26-fold the spontaneous mutation frequency (2.0 x 10–4). On the other hand, 100 µM HQ in the presence of NADH and Cu(II), which is a 1000 times lower concentration than MUC, enhanced the mutation frequency from 2.0 x 10–4 to 4.5 x 10–3.
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Base sequence analysis
We determined the base sequences of the mutated supF gene retrieved from human WI38-VA13 cells and classified the identified mutations into seven categories according to the type of mutation: single, tandem (two adjacent) and multiple base substitutions, frameshift, deletion, insertion and others. Table I shows the types of mutations induced by MUC (100 mM) and HQ (100 µM), together with those of spontaneous mutations obtained in our previous study (Nakayama et al., 1999
). In the spontaneous and MUC-induced mutations, base substitutions were predominant (73.9 and 80.0%, respectively) and a distinctly higher frequency of tandem base substitutions (13.3%) was observed in the MUC-induced mutations. In the HQ-induced mutations, deletions were the predominant mutation (42.4%).
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The types of base substitutions are classified in Table II. Although the MUC-induced mutations mostly occurred at G:C sites (66.6%), the proportion of base substitutions at A:T sites was significantly higher (33.3%) than the proportion of spontaneous mutations (2.5%) (Fisher’s exact test, P < 0.01). In the HQ-induced base substitutions, 87.1% of the substitutions occurred at G:C sites, and the proportions of each substitution were very similar to those of each spontaneous substitution mutation (Table II).
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Mutation spectra
Figure 5 shows the distributions of the spontaneous, MUC- and HQ-induced base substitutions in supF. Most of the MUC-induced base substitutions were uniformly distributed in the gene, although many mutations were found around base position 160. In contrast, spontaneous and HQ-induced mutations were found at certain specific sites, i.e. positions 123 and 129.
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Deletions were frequently induced by HQ, and the deleted lengths are shown in Table III. Large deletions (>40 bp) were frequently found in HQ-induced mutations, as shown in Figure 6. We found that 9 of 32 large deletions (
30%) occurred between two short repeat sequences.
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Discussion |
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We have demonstrated that HQ with NADH and Cu(II) enhances the mutation frequency significantly more than MUC. Since MUC is also a relatively weak mutagen and a minor metabolite (Witz et al., 1990
), it may not contribute significantly to the initiation of benzene leukemogenesis. Previously we analyzed the mutagenesis of another benzene metabolite, p-BQ. It is also a weak mutagen compared with HQ. Figure 7 shows the dose–response curves for each metabolite and the mathematical expression of each regression line. The slope factors, representing mutagenic intensities, are 3.2 x 10–2 for HQ, 1.4 x 10–3 for p-BQ and 5.2 x 10–5 for MUC, respectively. From these values we could presume that HQ is about 20 and 600 times stronger than p-BQ and MUC, respectively, and that it is more important in benzene carcinogenesis.
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We also found that MUC and HQ have distinctive mutagenic features suggesting their involvements in benzene carcinogenicity. Among the MUC-induced mutations, the proportion of tandem base substitutions at GG sites is much higher (6 of 8 tandem base substitutions) compared with spontaneous (0 of 1 tandem base substitutions), p-BQ-induced (1 of 4 tandem base substitutions) or HQ-induced (no tandem base substitutions) mutations (Nakayama et al., 2000
). As aldehyde chemicals such as formaldehyde and acetaldehyde form crosslinks between two adjacent guanines (Matsuda et al., 1998), MUC may also readily bind to GG sites because of its two aldehyde moieties.
Large deletions are remarkable in the mutations induced by HQ in the presence of NADH and Cu(II). In other mutation studies using shuttle vector plasmids, base substitution mutations are predominantly induced by other chemical mutagens such as aldehydes (Matsuda et al., 1995, 1998; Kawanishi et al., 1998a,b), alkylating agents (Moriwaki et al., 1991; Wang et al., 1991; Murata Kamiya et al., 1997), aromatic hydrocarbons (Maher et al., 1989, 1990) and another benzene metabolite, p-BQ (Nakayama et al., 2000). However, hydrogen peroxide also induces many large deletions in shuttle vector plasmids (Moraes et al., 1990). The HQ- and H2O2-induced mutations resemble the types of base substitutions.
In addition, HQ- and H2O2-induced base substitutions bear a resemblance to spontaneous mutations, although in the spontaneous mutations deletions are not so remarkable. They mostly occurred at G:C base pairs and the proportions of G:C
T:A transversion were higher than other chemical agents as described above. This seems inconsistent with the report that mainly thymine damage was induced (Hiraku and Kawanishi, 1996), but it might be due to the ‘A rule’ (Sagher and Strauss, 1983). Endogenous reactive oxygen species are the major factors in spontaneous mutation. These facts imply that hydrogen peroxide is generated in the redox cycling, as reported by Hiraku and Kawanishi (1996), and is the direct causative agent of HQ mutagenesis.
Another feature also supports this idea. Most of the base substitutions, 53 of 62 HQ-induced base substitutions and 55 of 81 spontaneous base substitutions, were induced at cytosine or guanine residues at 5'-PyC-3' or 5'-GPu-3' sites, as shown in Table IV. This feature, i.e. that guanine in the 5'-GPu-3' sequence is a mutational hot-spot, is consistent with other reports showing that the oxidative damage 8-OH-dG is formed readily on the 5'-side of guanine (Hatahet et al., 1998; Watanabe et al., 2001a,b). In fact, HQ treatment yields oxidative damage to plasmid DNA. The levels of 8-OH-dG in the 50 µM HQ-treated plasmids increased to 117 per 105 dG, which is about 120 times higher than non-treated plasmids. In other words, 50 µM HQ formed 2.4 x 10–1 8-OH-dG per supF gene because the number of dG and dC in the supF gene is 208.
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8-OH-dG mainly causes G:C
T:A base substitution (Shibutani et al., 1991), so we could estimate the mutational contribution of 8-OH-dG as follows:
Contribution of 8-OH-dG (%) = [(NG:CT:A/supF)/ (N8-OH-dG/supF)] x 100
NG:CT:A/supF = mutation frequency x (no. of base substitutions/total mutations) x (no. of G:CT:A substitutions/total base substitutions) = 2.5 x 10–3 x 31/66 x 17/62 = 3.2 x 10–4
N8-OH-dG/supF = (8-OH-dG/dG) x (dG/supF) = 117 x 10–5 x 208 = 0.24
therefore, the contribution of 8-OH-dG (%) = (3.2 x 10–4)/0.24 x 100 = 0.13
This estimation is approximately consistent with reports that 8-OH-dG are turned to point mutations at a rate of 0.1–1% in E.coli (Wood et al., 1990; Cheng et al., 1992; Wagner et al., 1997). However, it is less than another report that estimates 2% in human NCI-H1299 lung cancer cells (Sunaga et al., 2001). The discrepancy might be due to less efficient DNA repair because NCI-H1299 lung cancer cells have a homozygous partial deletion of the p53 protein (Lai et al., 1998).
From this estimation the 8-OH-dG induced by HQ would not make a huge contribution to mutation, but we must examine another feature of HQ-induced mutations correlated with the reactive oxygen species hydrogen peroxide. HQ induced many deletions, the majority of which occurred between two short repeat sequences. Deletions at a short direct repeat sequence are characteristic of double-strand breaks in a human shuttle vector system (Dar and Jorgensen, 1995). Even if double-strand breaks are not formed directly by HQ, oxidized DNA lesions can be repaired by base excision repair and nucleotide excision repair which converts the damage to a double-strand break (Lindahl and Wood, 1999). Double-strand breaks can be repaired via homologous recombination, but this leads to chromosomal instability (Haber, 1999), which is closely related to leukemia.
Our data are from an in vitro study but reflect in vivo benzene mutagenesis. The peculiar feature of frequent deletion is also seen in benzene-exposed Big Blue mice (Provost et al., 1996). This could, perhaps, be the consequence of strand breaks induced by reactive oxygen species as described above. The redox cycling of HQ may play an important role in benzene leukemogenesis.
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Notes |
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3To whom correspondence should be addressed. Tel: +81 75 753 5156; Fax: +81 75 753 5066; Email: nakayama{at}risk.env.kyoto-u.ac.jp
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