Sequence-specific detection of aristolochic acid–DNA adducts in the human p53 gene by terminal transferase-dependent PCR

Volker M. Arlt, Heinz H. Schmeiser2 and Gerd P. Pfeifer1

Division of Molecular Toxicology, German Cancer Research Center, Im Neuenheimer Feld 280, 69120 Heidelberg, Germany and
1 Department of Biology, Beckman Research Institute of the City of Hope, Duarte, CA 91010, USA


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The carcinogenic plant extract aristolochic acid (AA) is thought to be the major causative agent in the development of urothelial carcinomas found in patients with Chinese herb nephropathy (CHN). These carcinomas are associated with overexpression of p53, suggesting that the p53 gene is mutated in CHN-associated urothelial malignancy. To investigate the relation between AA–DNA adduct formation and possible p53 mutations, we mapped the distribution of DNA adducts formed by the two main components of AA, aristolochic acid I (AAI) and aristolochic acid II (AAII) at single nucleotide resolution in exons 5–8 of the human p53 gene in genomic DNA. To this end, an adduct-specific polymerase arrest assay combined with a terminal transferase-dependent PCR (TD-PCR) was used to amplify DNA fragments. AAI and AAII were reacted with human mammary carcinoma (MCF-7) DNA in vitro and the major DNA adducts formed were identified by the 32P-postlabeling method. These adducted DNAs were used as templates for TD-PCR. Sites at which DNA polymerase progress along the template was blocked were assumed to be at the nucleotide 3' to the adduct. Polymerase arrest spectra thus obtained showed a preference for reaction with purine bases in the human p53 gene for both activated compounds. For both AAs, adduct distribution was not random; the strongest signals were seen at codons 156, 158–159 and 166–167 for exon 5, at codons 196, 198–199, 202, 209, 214–215 and 220 for exon 6, at codons 234–235, 236–237 and 248–249 for exon 7 and at codons 283–284 and 290–291 for exon 8. Overall guanines at CpG sites in the p53 gene that correspond to mutational hotspots observed in many human cancers seem not to be preferential targets for AAI or II. We compared the AA–DNA binding spectrum in the p53 gene with the p53 mutational spectrum of urothelial carcinomas found in the human mutation database. No particular pattern of polymerase arrest was found that predicts AA-specific mutational hotspots in urothelial tumors of the current p53 database. Thus, AA is not a likely cause of non-CHN-related urothelial tumors.

Abbreviations: AA, aristolochic acid; AAI, aristolochic acid I (8-methoxy-6-nitrophenanthro[3,4-d]-1,3-dioxolo-5-carboxylic acid); AAII, aristolochic acid II (6-nitrophenanthro[3,4-d]-1,3-dioxolo-5-carboxylic acid); CHN, Chinese herb nephropathy; dA–AAI, 7-(deoxyadenosin-N6-yl)aristolactam I; dG–AAI, 7-(deoxyguanosin-N2-yl)aristolactam I; dA–AAII, 7-(deoxyadenosin-N6-yl)aristolactam II; dG–AAII, 7-(deoxyguanosin-N2-yl)aristolactam II; LM-PCR, ligation-mediated polymerase chain reaction; TD-PCR, terminal transferase-dependent polymerase chain reaction.


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The plant extract aristolochic acid (AA), a mixture consisting essentially of aristolochic acid I (AAI) and aristolochic acid II (AAII) (Figure 1Go), is a genotoxic mutagen (13) and a potent carcinogen in rodents (46) and man (7,8). These mutagenic and carcinogenic properties are believed to be based on the formation of DNA adducts by AA. After reductive metabolic activation, both AAs react with DNA preferentially at the exocyclic amino groups of adenine and guanine (9,10).



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Fig. 1. Structure of aristolochic acid I (R = OCH3) and aristolochic acid II (R = H).

 
In 1991, a unique form of nephropathy was observed in Belgium, called Chinese herb nephropathy (CHN) (11). CHN, associated with the prolonged intake of Chinese herbs during a slimming regimen, has been related to the toxic effects of Aristolochia species, containing nephrotoxic AA (12,13). One prescribed Chinese herb, Stephania tetrandra, was accidentically replaced by Aristolochia fangchi (14), because both are used in Chinese folk medicine under the same name, Fangji. More than 100 CHN cases have been identified, half of whom need renal replacement therapy, mostly including renal transplantation. Since the first reports of these cases of renal failure in Belgium, similar clinical presentations have been described in Spain (15), Japan (16), France (17), the UK (18) and China (19). Recently, an increasing number of urothelial carcinomas has been reported in CHN patients, suggesting that AA also plays a role in the formation of these tumors (7,8). In a group of 39 CHN patients with end-stage renal disease, statistical analysis predicted that the cumulative dose of Chinese herbs, previously shown to contain AA (14), was associated with a significantly higher probability of developing urothelial carcinomas (8). Using the 32P-postlabeling method, specific AA–DNA adducts were detected in all urothelial tissues of CHN patients; this unambiguously showed that all CHN patients analysed so far had indeed ingested AA (8,20,21). However, no difference was found between the amounts of AA–DNA adducts in CHN patients with urothelial carcinomas and in tumor-free CHN patients (8). Thus, the role of AA–DNA adducts in CHN-related carcinogenesis and therefore AA-induced carcinogenesis in humans remains to be investigated.

Proto-oncogenes and tumor-suppressor genes are potentially critical targets for carcinogens (2224). The mutational specificity of a carcinogen often serves as indirect evidence for initiation of tumorigenesis due to interaction of the carcinogen with these specific DNA sequences (25,26). Likewise, a distinct molecular characteristic of AA-initiated carcinogenesis in rodents is the activation of the Ha-ras gene by a specific AT->TA transversion mutation at the first adenine in codon 61 (CAA) (27). To a certain degree, it is possible to associate these specific mutations with the DNA-binding specificity of the carcinogen in vitro. Recently we demonstrated by polymerase arrest assay that both adenines in codon 61 of the Ha-ras gene in a plasmid are AA–DNA binding sites (28), suggesting that the mutations observed in AA-treated rodents may originate from adduct formation in this codon. As the presumed guardian of the genome, the p53 gene is one of the most commonly mutated genes observed in human tumors and is mutated in >50% of all human cancers (29,30). In CHN patients, urothelial carcinomas as well as renal and ureteral atypia were associated with overexpression of p53, suggesting that the p53 gene is also mutated in CHN-associated urothelial malignancy (7). In many cancers the distribution of mutations along the p53 gene is tumor-specific and characterized by several mutational hotspots, which correspond to amino acids within the DNA-binding domain of p53 (29,30). Mapping the distribution of DNA adducts in the p53 gene formed by AA might therefore reveal which adducts are potentially mutagenic lesions and may be converted into mutations. Thus, comparison of the AA–DNA binding spectrum in the p53 gene with the not yet determined p53 mutational spectrum of tumors from CHN patients may help us to understand the molecular mechanism by which AA causes cancer in humans.

In the experiments reported here, we mapped the distribution of DNA adducts along exons 5–8 on the nontranscribed strand of the p53 gene in human DNA modified in vitro by AAI and AAII using an adduct-specific polymerase arrest assay combined with a terminal transferase-dependent polymerase chain reaction (TD-PCR) to amplify DNA fragments. We also investigated the relation between AA–DNA adduct formation in the p53 gene and p53 mutations found in urothelial carcinomas obtained from the human p53 mutation database.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
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 References
 
Chemicals and enzymes
Pure AAI and AAII as sodium salts were kindly provided by Madaus (Köln, Germany). All chemicals and enzymes used for 32P-postlabeling (31) and for TD- and ligation-mediated polymerase chain reaction (LM-PCR) (32) were as described previously.

Treatment of MCF-7 cells with AAI
The human mammary carcinoma cell line MCF-7 (original stock line was obtained from the tumor bank of the German Cancer Research Center) was grown in 75 cm2 cell culture flasks in a total volume of 15 ml of Dulbecco's modified Eagle's medium (DMEM; Biochrom KG, Berlin, Germany), high-glucose type (DMEM with 4.5 g D-glucose/l), supplemented with 10% fetal calf serum (Biochrom KG), 1 mM sodium pyruvate (Biochrom KG), 4 mM L-glutamine (Pan Biotech, Aidenbach, Germany), 100 U of penicillin/ml and 100 µg of streptomycin/ml (Biochrom KG) at 37°C, 7% CO2 and 90% saturated atmospheric humidity untill all reached ~70–80% confluence. For incubations with AAI, cells were seeded at a density of 2 x 106 cells per flask and were grown for 48 h. Then cells were exposed to 10, 100, 200, 400 and 1000 µM of AAI for 24 h at 37°C. Control cells were treated with solvent (water) only. DNA was isolated as described below.

DNA isolation
After incubation for 24 h, the medium was removed and cells were harvested by trypsinization with 2 ml of 0.025% trypsin and 0.01% EDTA (Biochrom KG) in PBS (PAA Laboratories, Linz, Austria). Four milliliters of medium supplemented with 10% fetal calf serum was added twice to stop trypsinization. Subsequently, centrifugation at 350 x g and two washing steps with phosphate-buffered saline (PBS) yielded a cell pellet, which was stored at –20°C until DNA isolation. To isolate the DNA from MCF-7 cells, the pellet was resuspended in 500 µl lysis buffer [10 mM Tris, 1 mM EDTA and 1% (w/v) sodium dodecyl sulfate (SDS); pH 8.0] and incubated overnight at 37°C with proteinase K (1 mg/ml) (Roche). The mixture was extracted once with 1 vol Tris-saturated phenol (Roti-phenol; Roth, Karlsruhe, Germany), 1 vol Tris-saturated phenol–chloroform–isoamyl alcohol (25:24:1 by volume) (Roti-phenol–chloroform; Roth) and 1 vol chloroform–isoamyl alcohol (24:1, v/v). The DNA was precipitated with 0.1 vol 5 M NaCl and 2 vol ethanol, washed with 70% ethanol, dried and dissolved in 250 µl TE buffer (10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0). Samples were treated with RNase by adding 5 µl RNase stock solution (RNase A 2 mg/ml + RNase T1 200 U/ml; Roche) for 1 h at 37°C. An equal volume (250 µl) of proteinase K buffer (20 mM EDTA, 40 mM Tris, pH 8.0) and 50 µg/ml proteinase K was added and incubation was continued for 1 h at 37°C. The mixture was extracted again as described above; precipitated DNA was resuspended in TE buffer (pH 8.0). DNA concentration in the solution was determined by A260nm and diluted to a final concentration of 0.5 µg/µl.

Modification of MCF-7 DNA by AAI and AAII in vitro
DNA isolated from MCF-7 cells (100 µg; final concentration 1.2 mM dNps) was treated with 0.24 mM AAI and AAII in 250 µl 50 mM potassium phosphate buffer, pH 5.8, in the presence of 5 mg zinc dust for activation. Control DNA was treated in the same way but without carcinogen. For enzymatic activation, AAI and AAII were added at a final concentration of 2 mM to a deaerated and argon-purged solution of 650 µg (final concentration 4 mM dNps) MCF-7 DNA containing 1 mM hypoxanthine (Sigma) and 1 U xanthine oxidase (Sigma) in 50 mM potassium phosphate buffer, pH 5.8 (total volume 1 ml). Solvent control DNA was treated in the same way but without carcinogen. The incubations were performed at 37°C for 3 h and DNA was isolated as reported previously (31). The integrity of the adducted genomic DNA was verified by alkaline agarose gel electrophoresis as described previously (33).

32P-postlabeling analysis
DNA adduct analysis of MCF-7 cells treated with AAI was performed by the nuclease P1-enrichment procedure of the 32P-postlabeling method as described previously (31). DNA adducts in MCF-7 DNA modified in vitro with AAI and AAII were determined by the standard 32P-postlabeling method as previously described (28).

Mapping of AAI– and AAII–DNA adducts by TD-PCR
TD-PCR for exons 5–8 of the human p53 gene on the nontranscribed strand was performed essentially as described previously (32,34) with minor modifications. One microgram of modified and control DNA was used as template for TD-PCR. For the initial primer extension, primer 5-4 (5'-GGGCCAGACCTAAGAGCAATCAGT; Tm 59°C) was used for exon 5, primer 6-4 (5'-AGGCCACTGACAACCACC; Tm 60°C) for exon 6, primer 7–4 (5'-CAGGGGTCAGCGGCAAGCAGAG; Tm 65°C) for exon 7 and primer 8–4 (5'-CAAGGAAAGGTGATAAAAGTGAATCTGAG; Tm 58°C) for exon 8. The initial primer extension was performed using a 16:1 combination of Vent exo and Vent DNA polymerase (New England Biolabs) with a thermocycler protocol of 3 min at 95°C, 3 min at the primer annealing temperature, 5 min at 72°C, eight cycles of (1 min at 95°C, 3 min at primer annealing temperature and 2 min at 72°C), followed by 2 min at 95°C. After linear PCR amplification, DNA was precipitated in the presence of glycogen. Ribotailing and adapter ligation were performed as described (32). Adapter-ligated fragments were PCR-amplified after ethanol precipitation with Amplitaq Gold (Perkin Elmer) DNA polymerase according to the manufacturer's instructions. For the PCR amplification, primer 5-5 (5'-GGAATCAGAGGCCTGGGGACCCT; Tm 64°C) was used for exon 5, primer 6-5 (5'-ACTGACAACCACCCTTAACCCCTC; Tm 64°C) for exon 6, primer 7-5 (5'-AGAGGCTGGGGCACAGCAGGCC; Tm 67°C) for exon 7 and primer 8-5 (5'-AAAGTGAATCTGAGGCATAACTGCACCC; Tm 63°C) for exon 8. The amplified PCR products were separated by denaturing gel electrophoresis, electroblotted to a Genescreen Plus (New England Nuclear) nylon membrane and hybridized with 32P-labeled p53-specific probes as described (35). For preparation of the hybridization probes, primer 5-6 (5'-TGGGGACCCTGGGCAACCA; Tm 63°C) was used for exon 5, primer 6-6 (5'-TAACCCCTCCTCCCAGAGACCCC; Tm 66°C) for exon 6, primer 7-6 (5'-CACAGCAGGCCAGTGTGCAGGGT; Tm 65°C) for exon 7 and primer 8-6 (5'-CTTGGTCTCCTCCACCGCTTCTTG; Tm 62°C) for exon 8. All TD-PCR experiments were repeated at least once, giving very similar results. The nylon membranes were exposed to a PhosphorImager (Molecular Dynamics, Sunnyvale, CA).

Because the Maxam–Gilbert sequencing ladder created by TD-PCR is difficult to read, the sequencing was done by LM-PCR as described previously (32) with minor modifications. For the initial primer extension step, the same primers were used as for TD-PCR but with a different thermocycler protocol of 3 min at 95°C, 3 min at the primer annealing temperature and 5 min at 72°C. After DNA precipitation in the presence of glycogen, primer-extended fragments were ligated as described (32). PCR amplification and hybridization were done exactly as described for TD-PCR.

Stability of AAI– and AAII–DNA adducts during initial primer extension
To determine whether AAI– and AAII–DNA adducts were stable under the conditions used for TD-PCR, aliquots of MCF-7 DNA modified in vitro with AAI and AAII were analyzed after the initial primer extension by 32P-postlabeling. The conditions for the initial primer extension were as described above except that DNA polymerase and primer were omitted.


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 Materials and methods
 Results
 Discussion
 References
 
Adduct formation by AAI in MCF-7 cells
To mimic AA–DNA adduct formation observed in tissues of CHN patients, human MCF-7 cells were treated with various concentrations of AAI. Using the nuclease P1 enrichment procedure of the 32P-postlabeling method, specific AA–DNA adducts were detected. The adduct pattern was qualitatively similar to that found in CHN patients, consisting of two major spots (spots 1 and 2) and one minor one (spot 3) (8,20,21). As shown in Figure 2Go, the two major adducts were identified as reported previously (31) as 7-(deoxyadenosin-N6-yl)aristolactam I (spot 1) (dA-AAI) and 7-(deoxyguanosin-N2-yl)aristolactam I (spot 2) (dG-AAI) and the minor adduct as 7-(deoxyadenosin-N6-yl)aristolactam II (spot 3) (dA-AAII). Quantitative analysis obtained by 32P-postlabeling (31), shown in Table IGo, revealed a dose-dependent formation of AAI–DNA adducts. However, at the two highest doses of AAI (400 and 1000 µM), cell viability was too low to isolate enough DNA for analysis. Total modification levels ranged from 1 to 50 adducts in 107 nucleotides. In all CHN patients analysed so far, AA–DNA adduct levels ranged from ~0.01 to 1 adduct in 107 nucleotides (8,20,21). Since the lower detection limit of DNA adducts by TD-PCR is one adduct in 104 nucleotides (A.Riggs, personal communication) adduct levels generated in cell culture and in vivo were too low to determine AA–DNA binding sites.



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Fig. 2. Autoradiographic profile of AA–DNA adducts obtained in MCF-7 cells modified with 200 µM AAI. The nuclease P1 enrichment procedure of the 32P-postlabeling was used. The origin, in the bottom left-hand corner, was cut off before exposure. Screen-enhanced autoradiography was at room temperature for 30 min. Chromatographic conditions: D1, 1 M sodium phosphate, pH 6.8; D3, 3.5 M lithium formiate, 8.5 M urea, pH 4.0; D4, 0.8 M LiCl, 0.5 M Tris–HCl, 8.5 M urea, pH 9.0; D5, 1.7 M NaH2PO4, pH 6.0. Spot 1, dA–AAI; spot 2, dG–AAI; and spot 3, dA–AAII.

 

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Table I. Quantitative analysis of AAI–DNA adducts formed in MCF-7 cells and in MCF-7 DNA modified by AAI and AAII in vitro
 
Adduct formation by AAI and AAII in MCF-7 DNA in vitro
In order to generate higher levels of adducts in the p53 gene, we modified isolated MCF-7 DNA in vitro with AAI and AAII using either chemical activation (by zinc) or enzymatic activation (by xanthine oxidase, a mammalian nitroreductase). Specific AAI– and AAII–DNA adducts were detected and identified by the standard procedure of the 32P-postlabeling method as reported previously (31) as dA-AAI (spot 1), dG-AAI (spot 2), dA-AAII (spot 3) and 7-(deoxyguanosin-N2-yl)aristolactam II (spot 4) (dG-AAII) (Figure 3Go). Besides the previously identified dA-AAII and dG-AAII adducts, another adduct (spot 5) was detected in the AAII-modified MCF-7 DNA, derived from reaction with cytosine (dC-AAII), confirming recent results (28). As shown in Table IGo, quantitative analysis revealed that chemical activation by zinc proved to be the most efficient activation system for both AAs in vitro. Total adduct level ranged from approximately seven to 55 adducts in 104 nucleotides for AAI and AAII by chemical activation and from one to six adducts in 104 nucleotides by enzymatic activation. Therefore these modified DNAs were used as templates for TD-PCR. In all in vitro incubations, adenine adduct formation was favoured for both AAs independent of the activation system used. Likewise, for both activation systems, DNA modification by AAII was much higher than that by AAI. The integrity of the modified genomic DNAs was verified by alkaline gel electrophoresis (data not shown). Modification of DNA by both AAs in the presence of zinc did not increase the number of strand breaks in genomic DNA whereas activation of both AAs by xanthine oxidase showed increased numbers of strand breaks and was therefore only qualitatively included in this study.



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Fig. 3. Autoradiographic profiles of AA–DNA adducts obtained in MCF-7 DNA modified with AAI (A and C) and AAII (B and D) by enzymatic activation with xanthine oxidase (A and B) and by chemical activation with zinc at pH 5.8 (C and D). The standard method of 32P-postlabeling was used. The origins, in the bottom left-hand corners, were cut off before exposure. Screen-enhanced autoradiography was at room temperature from 30 min to 3 h. Chromatography was performed as described in the legend to Figure 2Go. Spot 1, dA–AAI; spot 2, dG–AAI; spot 3, dA–AAII; spot 4, dG–AAII; and spot 5, dC–AAII.

 
Distribution of AAI– and AAII–DNA adducts in the human p53 gene
We mapped the distribution of DNA adducts formed by AAI and AAII in vitro along exons 5–8 of the p53 gene on the nontranscribed strand, which would be expected to be repaired relatively inefficiently according to the concept of transcription-coupled repair (36,37). To this end we used TD-PCR, a recently developed technique for detecting DNA adducts at single nucleotide resolution (34). The method is based on adduct-specific polymerase arrest of primer extension reactions, tailing of the resulting 3' ends with terminal transferase using rGTP, linker–adapter ligation and PCR to amplify DNA fragments. For some p53 exons we used primer sets different from those reported for LM-PCR (25,38,39). 32P-postlabeling analyses of AAI- and AAII-modified MCF-7 DNA performed before and after the initial primer extension step revealed no qualitative or quantitative changes in AAI– or AAII–DNA adduct profiles (data not shown). These 32P-postlabeling results indicate that AA–DNA adducts were stable during the initial primer extension reactions.

Figure 4Go shows the profile of AA–DNA adducts produced after chemical activation within exons 5–8 of the p53 gene. The bands varied in intensity, indicating that some sites in the sequence were adducted more readily than others. As can be seen, arrest sites for AAI and AAII were qualitatively similar, but distinct differences between the two nitrophenanthrene carboxylic acids were also observed. DNA adduct profiles produced by AAI and AAII mediated by enzymatic activation showed a very similar pattern (data not shown), indicating that the number of DNA modifications is much higher than the number of DNA strand breaks induced during modification catalyzed by xanthine oxidase. The Maxam–Gilbert sequencing products were also loaded on the gel to determine the site in the p53 gene at which the DNA polymerase had arrested. The strongest signals were seen at codons 156, 158–159 and 166–167 for exon 5, at codons 196, 198–199, 202, 209, 214–215 and 220 for exon 6, at codons 234–235, 236–237 and 248–249 for exon 7 and at codons 283–284 and 290–291 for exon 8 when one corrects for the extra two or three nucleotides that are added as part of the TD-PCR procedure. Because the Maxam–Gilbert sequencing ladder is difficult to read by TD-PCR, the sequencing was done by conventional LM-PCR (32).



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Fig. 4. TD-PCR mapping of AAI– and AAII–DNA adducts in the untranscribed strand of exons 5–8 of the human p53 gene. MCF-7 DNA was modified in vitro with AAI and AAII by activation with zinc at pH 5.8. LM-PCR was done on chemically sequenced genomic DNA samples (first four lanes) to provide positions markers. AA–DNA adducts that block polymerase progression were detected by TD-PCR. TD-PCR-derived signals migrate two or three nucleotide positions more slowly than LM-PCR-derived sequencing bands. The positions of p53 codons are indicated on the right. Modified bases are marked by an asterisk after the base in the corresponding codon. C'G stands for methylated CpG sequences.

 
Assuming that DNA polymerase was blocked predominantly at the nucleotide 3' to the adduct, the arrest spectrum revealed that most of the arrests were associated with purine residues in the sequence. This preference for reaction with purine bases is consistent with the 32P-postlabeling results and confirms previous reports that purine adducts are the major reaction products of AAI and AAII with DNA (4,10,21,31,45). In the AAII-modified template, strong arrest bands corresponding to cytosine modifications were found, consistent with the 32P-postlabeling results and recent results obtained by 32P-postlabeling analysis on single-stranded homopolymers modified with AAII (28). Furthermore, arrest bands associated with cytosine residues have been detected in the Ha-ras gene in a plasmid using a polymerase arrest assay (28).

Comparison of AA–DNA binding in the p53 gene with the p53 mutational spectra of urothelial carcinomas
Figure 5Go shows the distribution of mutations along the p53 gene involved in urothelial carcinomas as reported in a p53 mutation database (40). In the current database, 61 mutations are recorded for urothelial carcinomas. The mutational spectrum is characterized by mutational hotspots in codons 175, 220, 248, 280 and 282. Three of these codons (175, 248 and 282) contain 5-methylcytosines within a CpG sequence (38) and represent mutational hotspots in many other tumors (29). Mutations at these codons may be formed by a methylation–deamination mechanism (23) but enhanced adduct formation at guanines in CpG sites by bulky carcinogens, such as benzo[a]pyrene (26, 41, 42), may also contribute to mutations at these mutational hotspots. Codon 220 is a mutational hotspot specific for urothelial carcinomas and does not occur as a hotspot in other cancers of the urinary tract (40). It is also the only mutational hotspot in urothelial carcinoma associated with adenine. The mutations are dominated by G->A (39%), followed by A->G (16%) and C->T (13%) transition mutations whereas A->T transversion mutations are rare (two out of 61(40)).



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Fig. 5. p53 mutational spectrum of urothelial carcinomas obtained from the human p53 mutation database (40).

 
As shown in Figure 4Go, no AA–DNA binding was observed at codon 175 in exon 5. At codon 220 in exon 6, moderate DNA binding was observed for AAI and a greater extent of binding for AAII. The major mutational hotspot for urothelial carcinomas at codon 248 in exon 7 was a strong AA–DNA binding site within a CpG site; binding of this site by AAII was also stronger. The other mutational hotspot within a CpG sequence at codon 282 in exon 8 was only a marginal AA–DNA binding site. Overall, guanines in CpG sequences seem not to be preferential targets for either AA. AA–DNA binding at codon 280 in exon 8 was also only marginal.


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The combination of adduct-specific endonucleases with LM-PCR has led to a sensitive method for detecting bulky DNA adducts in mammalian DNA at single-nucleotide resolution (43). The sequence position of adducts can be mapped whenever it is possible to convert the adduct into a DNA strand break with a ligatable 5'-phosphate group. Another technique for the sequence-specific detection of bulky DNA adducts in mammalian DNA is the recently developed TD-PCR (34). One major advantage of this technique is that no special enzymes are needed to convert DNA adducts into strand breaks. Instead, any lesion that blocks primer extension is detectable. This technique has been successfully applied for the sequence-specific detection of UV-induced damage (34), aflatoxin B1-induced adducts (44) and benzo[a]pyrene-induced adducts (V.Arlt and G.Pfeifer, unpublished results). There are advantages and disadvantages to each approach, although both methods give similar adduct distributions (34,44).

To mimic the AA–DNA adduct formation detected in tissues from CHN patients, we treated human MCF-7 cells with various concentrations of AAI. The observed AA–DNA adduct pattern was similar to those found in urothelial tissue of CHN patients (8,20,21) with an induced adduct level of one to 50 adducts in 107 nucleotides. The adduct level in DNA from CHN patients analyzed previously ranged from 0.01 to one adduct in 107 nucleotides (8,20,21). Since the lower limit of detection by TD-PCR is around one adduct in 104 nucleotides (A.Riggs, personal communication) adduct levels were too low to determine AA-binding sites in DNA isolated from cells or human tissue. Because reactive metabolites of AAI and AAII are not available, chemical activation by zinc and enzymatic activation by xanthine oxidase was used for both AAs to modify human (MCF-7) DNA in vitro to obtain adduct levels suitable for TD-PCR. Since DNA modification through AA catalyzed by xanthine oxidase led to strand breaks only, DNA modified by AA in the presence of zinc was a suitable template for TD-PCR. 32P-postlabeling analysis of the adducted DNA identified the four major purine adducts of AAI and AAII previously identified in rodents (45) and man (8,20,21).

Strong evidence that AA–DNA adducts are effective blocks for DNA polymerase immediately preceding AAI– and AAII–DNA adducts comes from several sources. Firstly, when one corrects for the extra two or three nucleotides that are added as part of the TD-PCR procedure, most of the arrests were associated with purine residues in the sequence. This preference for reaction with purine bases is consistent with the 32P-postlabeling results reported here and several in vitro and in vivo studies (8,9,31,45). Secondly, in primer extension studies with site-specifically mono-adducted oligonucleotides containing purine–AA adducts, DNA synthesis was blocked predominantly at the nucleotide 3' to the adduct by Sequenase DNA polymerase regardless of the type of AA adduct examined (46). Thirdly, arrest bands associated with cytosine residues in the AAII-modified DNA detected by TD-PCR were confirmed by the 32P-postlabeling results and by a recent report mapping AAII–DNA adducts in the Ha-ras gene using a plasmid polymerase arrest assay (28).

Thus, these results suggest that DNA polymerase is effectively arrested one base prior to most AA–DNA adducts and that the intensity of the arrest bands should, therefore, reflect the frequency of adduct formation at that sequence site. However, as discussed by Denissenko et al. (44), for TD-PCR it is generally difficult to extrapolate from the sites of polymerase arrest to the precise site of adduct formation because of one or two nucleotide variability in the termination of primer extension and/or addition of Gs by terminal transferase. The exact adduct assignment of neighbored purine bases is particularly difficult. Therefore the adduct distribution obtained by this technique cannot be considered as quantitative, but rather is semiquantitative. Nevertheless, the adduct is presumably near the site of polymerase arrest such that the latter can be probed to determine whether hotspots of mutations correlate with hotspots of adduct formation.

Amongst the adenines and guanines in exons 5–8 of the p53 gene, there was an uneven distribution of polymerase arrests, indicating that some purine sites in the sequence were more readily adducted than others and, therefore, that AA–DNA adduct formation is not random but sequence-specific. Recently we reported that 5'-nearest pyrimidine bases have a strong effect on AA–DNA binding in a plasmid, containing exon 2 of the Ha-ras gene (28) suggesting that the nearest-neighbor bases may affect the efficiency of AA–DNA adduct formation. It is clear, however, that such a general rule was not found for AA–DNA binding in the p53 gene. All of the 42 CpG dinucleotides in the p53 coding sequences are methylated, irrespective of the tissue (38). It has been shown by others (26,39,41,42) that adduct formation in the p53 gene by bulky carcinogens, such as benzo[a]pyrene and other polycyclic aromatic hydrocarbons, is enhanced by methylation at CpG sites, but we did not observe a preference for enhanced adduct formation at CpG sequences for AA. However, particularly strong AA–DNA binding was seen at the CpG site at codon 248, which is also associated with a mutational hotspot in many tumors (29).

In CHN patients, the major adenine adduct of AAI, dA–AAI, is detectable at relatively high levels even <=7 years after the patients stopped taking the herbal medicine (8). Since AT->TA mutations have been found in high frequency in activated Ha-ras oncogenes in tumors of rodents treated with AA (27) this adduct may also be the adduct primarily responsible for the carcinogenesis. This assumption is further supported by the apparently lifelong persistence of dA–AAI adducts in the target organ forestomach in rats, whereas dG–AAI adducts are removed continuously from the same DNA (47). Recently we demonstrated that both adenines in codon 61 of the Ha-ras gene are AA–DNA binding sites, suggesting that the mutations observed in tumors of AA-treated rodents may originate from adduct formation in this codon (28). Moreover, primer-extension studies with site-specifically adducted oligonucleotides containing purine–AA adducts suggested a higher mutagenic potential of adenine adducts than guanine adducts (46). To a certain degree, it is possible to relate the DNA-binding specificity of a carcinogen to specific mutations found in a target gene for tumor formation. For example, UV-induced damage in the p53 gene is correlated with p53 mutational hotspots in skin cancer (25) and selective adduct formation by benzo[a]pyrene and other polycyclic aromatic hydrocarbons at codons 157, 158, 248 and 273 of the p53 gene is correlated with p53 mutational hotspots in lung cancer associated with smoking (26,39,41). Since the p53 mutational spectrum of urothelial carcinomas from CHN patients has not yet been investigated, the AA–DNA binding spectrum in the p53 gene obtained in this study was compared with the p53 mutational spectrum of urothelial carcinomas found in the human mutation database (40). This mutational spectrum is characterized by mutational hotspots in codons 175, 220, 248, 280 and 282 (Figure 5Go). Strong AA–DNA binding was observed in codon 248, the major mutational hotspot of urothelial carcinomas. However, codon 248 is also a mutational hotspot in many other tumors (29). Codon 220 is a mutational hotspot specific for urothelial carcinomas and does not occur as a hotspot in other cancers of the urinary tract (40); it is also the only mutational hotspot in urothelial carcinomas associated with adenine. Polymerase arrest sites due to AA–DNA binding at this mutational hotspot were only moderate, but it is possible that mutations might be induced when DNA polymerase is not completely arrested by an adduct. Also, the mutation frequency depends not only on initial adduct formation, but also on the rate of repair and individual mutagenic activity of each adduct as well as on the sequence context (36,37,44). Certainly many intense arrest bands were present at other codons not found in the p53 mutational spectrum of urothelial carcinomas. Therefore, it is clear that no particular pattern of polymerase arrest predicts AA-specific mutational hotspots in urothelial tumors of the current p53 database. This might also be limited by the small number of mutations recorded for urothelial carcinomas in the database. Nevertheless, this may suggest that AA is not a likely cause of non-CHN related urothelial tumors. However, all CHN patients have been exposed to AA, so comparison of the AA–DNA binding spectrum in the p53 gene reported here with the p53 mutational spectrum of tumors from CHN patients, which is in progress, may help us to understand the molecular mechanism by which AA causes cancer in humans.


    Notes
 
2 To whom correspondence should be addressed Back


    Acknowledgments
 
We thank Dr Christian A.Bieler, Dr Leslie E.Smith, Young-Hyun You and Jung-Hoon Yoon for support and discussions. This work was supported by a stipend of the Boehringer Ingelheim Fonds to V.M.A.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received June 15, 2000; revised September 12, 2000; accepted September 12, 2000.