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2-Amino-1-methyl-6-phenylimidazo-[4,5-b]pyridine (PhIP) is the most abundant heterocyclic amine mutagen formed during high temperature cooking of meats, accounting for ~70% mean dietary intake of heterocyclic amines in the United States (Keating and Bogan, 2004). PhIP has been identified in smoke condensates from the frying of meat (Thiebaud et al., 1995) and has been detected in airborne particles, diesel-exhaust particulates and incinceration ash from garbage-burning plants (Manabe et al., 1993). PhIP is found in cigarette smoke (Manabe et al., 1991) and is a urinary mutagen in smokers of black tobacco (Peluso et al., 1991). PhIP causes mammary, prostate, and colon tumors in the rat and is classified as a probable human carcinogen (Sinha et al., 2000; National Toxicology Program, 2005).
PhIP-induced DNA adduct formation and mutagenesis require metabolic activation, and one important pathway includes N-hydroxylation catalyzed by cytochrome P450s (Turesky 2004; 2007; Kim and Guengerich, 2005). Although many arylamine carcinogens undergo N-hydroxylation by cytochrome P4501A2, this isoform is primarily restricted to the liver and PhIP carcinogenesis is independent of this P450 in mice (Kimura et al., 2003). Arylamine carcinogens also are activated by cytochrome P4501A1 (CYP1A1) (Murray et al., 1993; Edwards et al., 1994; Hammons et al., 1997; Yamazaki et al., 2004; Kim and Guengerich, 2005; Turesky, 2007) and cDNA-expressed human CYP1A1 exhibited over 15-fold higher affinity for PhIP than human CYP1A2 (Crofts et al., 1998). Since PhIP induces tumors in extrahepatic organs and has relatively high affinity for CYP1A1, it is important to evaluate the role of CYP1A1 in PhIP-induced genotoxicity. N-hydroxy-PhIP can be further O-acetylated by N-acetyltransferase 2 (NAT2) to the acetoxy-derivatives that are highly unstable, leading to electrophilic intermediates that are mutagenic and form DNA adducts (Schut and Snyderwine, 1999).
PhIP-induced DNA adducts primarily form at the C8 position of deoxyguanosine (dG-C8) and can be removed by nucleotide excision repair (NER) (Schut and Snyderwine, 1999). Because DNA adduct levels are a function of environmental exposure and polymorphism in genes involved in carcinogen metabolism, DNA adducts are an informative biomarker for investigations of genetic variation in carcinogen metabolism.
Since heterocyclic amines require metabolic activation to exert their carcinogenic effects, genetic polymorphisms in the enzymes catalyzing the activation and/or detoxification pathways of carcinogen metabolism may account for differences in susceptibility to carcinogens between individuals (Turesky, 2004). Humans exhibit genetic polymorphism in NAT2 resulting in rapid and slow acetylator phenotypes. Relative to the NAT2*4 reference allele, NAT2*5B possesses three polymorphisms in the NAT2 open reading frame: T341C (I114T), C481T (synonymous) and A803G (K268R) and is associated with slow acetylator phenotype (Zang et al., 2004). Epidemiologic studies suggest a role for NAT2 genetic polymorphism in susceptibility to various cancers, but laboratory-based experiments are valuable for investigating metabolism of carcinogens that may contribute to these findings.
O-acetylation of N-hydroxy-PhIP has been shown to generate DNA adducts (Snyderwine et al., 2002) suggesting that PhIP- induced DNA damage and subsequent mutagenesis would be greater in rapid than slow NAT2 acetylators. On the other hand, several previous studies have shown that metabolic activation of N-hydroxy-PhIP did not require O-acetylation (Buonarati and Felton, 1990; Wild et al., 1995; Wu et al., 1997). Thus, we investigated the effects of human CYP1A1 and NAT2 genetic polymorphism on PhIP-induced DNA adducts and mutagenesis in nucleotide excision repair-deficient Chinese hamster ovary (CHO) cells transfected with human CYP1A1, human NAT2*4 (rapid acetylator allele) or human NAT2*5B (slow acetylator allele).
The UV5-CHO cell line, a nucleotide excision repair-deficient derivative of the AA8 line, was obtained from the ATCC (Catalog number: CRL-1865). UV5-CHO is hypersensitive to bulky adduct mutagens and belongs to the excision repair cross complementation group 2. All cells were grown in alpha-modified minimal essential medium (Cambrex) without L-glutamine, ribosides, and deoxyribosides supplemented with 10% fetal bovine serum (Hyclone), 100 units/ml penicillin, 100 units/ml streptomycin (Cambrex), and 2 mM L-glutamine (Cambrex) at 37 °C in 5% CO2. Media were supplemented with appropriate selective agents to maintain stable transfectants. Construction of UV5/CHO cells expressing human CYP1A1 and NAT2*4 or NAT2*5B was recently reported and characterized (Bendaly et al., 2007). Briefly, the pFRT/lacZeo plasmid (Invitrogen) was transfected into nucleotide excision repair-deficient UV5 cell lines to generate a UV5 cell line containing a single integrated FRT site (UV5FRT). Purified human NADPH cytochrome P450 reductase (POR) and CYP1A1 polymerase chain reaction (PCR) products were digested and ligated into similarly treated pIRES vector and transformed into DH5α competent cells. The pIRES plasmid containing cDNAs of human CYP1A1 and POR was transfected into the newly established UV5FRT cell line. Those UV5 cells expressing a single FRT site, CYP1A1, and POR were expanded, and intact geneticin-resistant cells assayed for CYP1A1 activity by measuring 7-ethoxyresorufin O-deethylase activity as described previously (Metry et al., 2007). Cell lines with similar levels of 7-ethoxyresorufin O-deethylase activity were selected for additional transfection with either NAT2*4 or NAT2*5B.
The open reading frames of NAT2*4 and NAT2*5B were amplified by PCR, digested with NheI and XhoI (New England Biolabs), and inserted into the similarly prepared pcDNA5/FRT vector (Invitrogen). The pcDNA5/FRT plasmid containing human NAT2*4 or NAT2*5B was co-transfected with pOG44, a Flp recombinase expression plasmid, into UV5FRT cells. Integration of the pcDNA5/FRT construct into the FRT site was confirmed by PCR (Metry et al., 2007). The NAT2*4- and NAT2*5B-transfected cells were characterized for N-acetylation of sulfamethazine, a NAT2-selective substrate (Bendaly et al., 2007).
N-hydroxy-PhIP O-acetyltransferase assays were determined by high performance liquid chromatography (HPLC) as previously described (Metry et al., 2007). Briefly, reaction mixtures containing equal amounts of cell lysate protein, 1 mg/ml deoxyguanosine, 100 μM N-hydroxy-PhIP (Toronto Research Chemicals), and 1 mM acetyl-coenzyme A were incubated at 37°C for 10 min and stopped by the addition of water-saturated ethyl acetate. The reactions were centrifuged for 10 min and the organic phase was transferred, evaporated to dryness, and resuspended in 100 μl of 10% acetonitrile. HPLC separation was achieved using a gradient of 85:15 sodium perchlorate (pH 2.5)/acetonitrile to 0:100 sodium perchlorate (pH 2.5)/acetonitrile over 10 min. Baseline measurements using extracts of UV5 and UV5/CYP1A1 cells were subtracted from measurements in the NAT2*4- and NAT2*5B-transfected CHO cell lines.
Assays for cytotoxicity and mutagenesis were carried out as described previously (Metry et al., 2007). Briefly, cells were grown for 12 doublings, with selective agents in complete hypoxanthine-aminopterin-thymidine medium (30 μM hypoxanthine, 0.1 μM aminopterin, and 30 μM thymidine). Cells were plated at a density of 5 × 105 cells/T-25 flask and incubated for 24 h, after which media were changed and the cells were treated separately for 48 h with various concentrations of PhIP (Toronto Research Chemicals) or vehicle control (0.5% dimethyl sulfoxide). Survival was determined by colony-forming assay and expressed as percent of vehicle control. The remaining cells were replated and subcultured. After 7 days of growth, cultures were plated for cloning efficiency in complete media and for mutations in complete media containing 40 μM 6-thioguanine (Sigma). Dishes were seeded with 1 × 105 cells/100 mm dish (10 replicates) and incubated for 7 days; cloning efficiency dishes were seeded with 100 cells/well/six-well plate in triplicate and incubated for 6 days.
Cells grown in 15- cm plates were treated separately with PhIP as described above for the cytotoxicity and mutagenesis assays. Cells were harvested after 48 h of treatment and DNA was extracted and quantified as previously described (Metry et al., 2007). dG-C8-PhIP and dG-C8-PhIP-D3 adduct standards (>95% purity) were obtained from Toronto Research Chemicals. One-tenth volumes each of proteinase K solution (20 mg/mL) and 10% SDS were added to the cell lysate, and the mixture was incubated at 37°C for 60 min. One volume of phenol, equilibrated with 10 mM Tris HCl (pH 8.0), was added to the mixture, which was then vortexed and centrifuged at 3,600 x g for 15 min. The aqueous layer was removed and added to 1 volume of phenol/chloroform/isoamyl alcohol (25:24:1) saturated with 10 mM Tris HCl (pH 8.0), which was vortexed and centrifuged. The aqueous layer was removed and added to 1 volume of cold (-20°C) isopropanol, and the mixture was vortexed and centrifuged. The DNA pellet was washed with 70% ethanol and redissolved in 5 mM Tris HCl (pH 7.4) containing 1 mM CaCl2, 1 mM ZnCl2, and 10 mM MgCl2. DNA was quantified by UV spectroscopy using A260 nm. DNA quality was monitored by UV spectroscopy using A260/280 nm and this ratio was consistently above 1.9. DNA samples (200 μg) added to 1 ng (3.3 adducts per 106 DNA bases) deuterated internal standard (dG-C8-PhIP-D3) were digested at 37°C with 10 units DNAse I (US Biological) for 1 h followed by 5 units micrococcal nuclease (Sigma), 5 units nuclease P1 (US Biological), 0.01 units spleen phosphodiesterase (Sigma), and 0.01 units snake venom phosphodiesterase (Sigma) for 6 h followed by 5 units alkaline phosphatase (Sigma) overnight. Two volumes of acetonitrile were added to the digest, which was then filtered and concentrated to 100 μl in a speed vacuum.
Samples were subjected to binary gradient HPLC and introduced into a Micromass Quattro LC triple quadrupole mass spectrometer using a custom-built nanospray as described previously (Metry et al., 2007; Neale et al., 2008). Samples were loaded onto a Inertsil C18 precolumn (5 mm × 300 μm i.d., 5 μm; LC Packings) using Perkin Elmer ABI 140D syringe pumps and a Hewlett Packard 1100 Series autosampler. Multiple reaction monitoring (dwell time, 0.5 s; span, 0.4 Da) was used to measure the [M+H]+ to [(M-116) + H]+ (loss of deoxyribose) mass transition. A Quattro LC micromass triple quadrupole mass spectrometer equipped with a nanoelectrospray ion source was used for PhIP-DNA adduct quantitation. Multiple reaction monitoring in the electrospray ionization-positive ion mode was carried out using argon as the collision gas. Capillary and cone voltages and collision energies were optimized for cleavage of the glycosidic bond. The dG-C8-PhIP adduct was monitored using the transition from m/z 490 to m/z 374 and the deuterated internal standard (dG-C8-PhIP-D3) was monitored using the transition from m/z 493 to m/z 377.
UvrABC endonuclease was cloned from UvrABca, UvrBBca, and UvrCTma, and those individual recombinant protein subunits were overexpressed in and purified from E.coli (Jiang et al., 2003; 2004). Plasmid pTHQB04 (5.45 kbp, which contains a 1.5 kbp fragment of the human β-globin gene) was propagated in XL1-Blue (Quan and States, 1996). Form I plasmid DNA was purified by CsCl-ethidium bromide sedimentation equilibrium ultracentrifugation. DNA concentrations were determined by measuring the A260. DNA quality was checked by agarose gel electrophoresis as well as measurement of the A260/A280 ratios (which were 1.8-2.0).
N-hydroxy-PhIP-damaged DNA was prepared with 2 μg of pTHQB04 plasmid DNA, 2.5 mM acetyl coenzyme A and varying concentrations of N-hydroxy-PhIP. Water was substituted for acetyl coenzyme A in controls. The total reaction was incubated at 37°C for 5 min, and stopped with an equal volume of ethyl acetate. The aqueous layer was extracted from the ethyl acetate mixture 4 times. DNA was precipitated by adding 1/10 volume of 3 M sodium acetate (pH = 5.6), and 2.2 volume of cold ethanol. DNA concentrations were determined by measuring the A260. DNA quality was checked by agarose gel electrophoresis as well as measurement of the A260/A280 ratios (which were 1.8-2.0).
The plasmid relaxation assay was adapted from that described previously (Jiang et al., 2004). N-hydroxy-PhIP-treated plasmid (20 fmol, equivalent to 71 ng pTHQB04, 1 nM of final DNA substrate) was preincubated with UvrA and UvrB in 20 μl of UvrABC buffer (50 mM Tris-HCl, pH 7.5, 50 mM KCl, 10 mM MgCl2, 5 mM dithiothreitol, and 1 mM ATP) at 55°C for 30 min, UvrCTma was added and incubated at 55°C for additional 30 min. Reactions without UvrABC were performed as a no incision control. Reactions were terminated by adding 2 μl of stop buffer (1% SDS and 200 mM EDTA-Na2). Cleavage of damaged plasmid-DNA was monitored by following the conversion of Form I plasmid to Form II plasmid. A 20 ng sample of the UvrABCTma incised DNA was resolved by 1% agarose gel electrophoresis and visualized by staining with SYBR-Gold (Molecular Probes, Eugene Oregon, USA). Fluorescence of resolved bands was quantitated using fluorescence detection mode (Blue excitation, 537 nm) of a Molecular Dynamics Storm 860 Phosphorimager (Amersham Pharmacia Biotech, Piscataway, NJ).
Cell lysates from UV5 and each of the transfected CHO cell lines were tested for their capacity to activate N-hydroxy-PhIP to form dG-C8-PhIP adducts. N-hydroxy-PhIP O-acetyltransferase activity in CYP1A1/NAT2*4 -transfected cell line was 2.5-fold higher (P=0.0150) than in the CYP1A1/NAT2*5B-transfected cell line (Fig. 1). Low but detectable levels of N-hydroxy-PhIP activation were detected in the UV5 and the CYP1A1-transfected cell lines that were subtracted from the experimental measurements in the CYP1A1/NAT2-transfected cells.
The CYP1A1-, CYP1A1/NAT2*4- and CYP1A1/NAT2*5B-transfected CHO cell lines each showed concentration-dependent cytotoxicity (Fig. 2) and hypoxanthine phosphoribosyl transferase mutagenesis (Fig. 3) following PhIP treatment, whereas they were not observed in untransfected CHO cells.
The dG-C8-PhIP standards were characterized by HPLC-tandem mass spectrometry and used to verify the identity of DNA adducts formed in vitro. They were quantitated relative to dG-C8-PhIP-D3 internal standard. One principal PhIP adduct (dG-C8-PhIP), corresponding to m/z 490, was identified in CHO cells incubated with PhIP (Fig. 4). Following PhIP treatment, dG-C8-PhIP DNA adduct levels were dose dependent in cells transfected with CYP1A1 while the further addition of NAT2*4 or NAT2*5B did not have an appreciable effect (Fig 5). Adduct levels were undetectable in untransfected cells. The observation that cells transfected only with CYP1A1 were able to bioactivate PhIP raised the question of whether N-hydroxy-PhIP required further metabolism by phase II enzymes to generate the DNA reactive electrophile. In order to investigate whether N-hydroxy-PhIP could be activated by acetyl coenzyme A to form DNA adducts without catalysis by NAT2, we incubated N-hydroxy-PhIP and acetyl coenzyme A but without NAT2 in the presence of supercoiled plasmid DNA. We assayed for the formation of DNA adducts by UvrABC incision in a plasmid relaxation assay as outlined in Methods. UvrABC is a prokaryotic endonuclease that recognizes and excises bulky DNA adducts such as dG-C8-PhIP. DNA adduct formation, detected by UvrABC incision, was observed in the presence of acetyl coenzyme A despite the absence of catalysis by NAT2 (Fig 6).
CYP1A1 and NAT2 genotypes may increase risk of colorectal cancer from heterocyclic amines (Murtaugh et al., 2005). Recent studies have shown that lung PhIP-DNA adducts were significantly lower in lung tissue of CYP1A1 null mice but not cytochrome P4501A2 null mice (Ma et al., 2007). Our study tested the effect of human CYP1A1 in combination with rapid or slow acetylator NAT2 on mutagenesis and DNA adducts following PhIP exposure. Our study showed that PhIP induced cytotoxicity and mutagenesis in all CYP1A1-transfected cell lines but not in the untransfected UV5 cell line. Our plasmid relaxation studies suggest that enzymatic catalysis of N-hydroxy-PhIP is not required to generate a DNA reactive electrophile. We observed induction of UvrABC sensitive sites in plasmid DNA incubated with N-hydroxy-PhIP in the presence of AcCoA concentrations as low as 20 μM.
Heterocyclic amines can also be detoxified through glucuronidation and glutathione-S-transferase or via aromatic ring hydroxylation. Once the N-hydroxy-metabolite is formed, it can be reduced to the initial carcinogen, deactivated by glutathione-S-transferases or glucuronosyltransferases, or further activated by N-acetyltransferases and sulfotransferases. Although we observed differences in N-hydroxy-PhIP O-acetyltransferase catalytic activity between the CYP1A1/NAT2*4- and CYP1A1/NAT2*5B- transfected CHO cell lines, NAT2 did not affect PhIP-induced mutagenesis significantly, consistent with results previously reported in cytochrome P4501A2/NAT2-transfected CHO cell lines (Metry et al., 2007). Although the NAT2- effect was slight and not significant (p >0.05), N-hydroxy-PhIP-induced mutagenesis and DNA adduct levels were slightly higher in the CYP1A1/NAT2*4- than the CYP1A1/NAT2*5B-transfected CHO cell line. Previous studies have reported that PhIP DNA adduct levels in human breast tissue are higher in rapid than slow NAT2 acetylators (Zhu et al., 2003) and have suggested that rapid acetylator NAT2 phenotype increases colorectal (Lang et al., 1994; Chen et al., 1998; Le Marchand et al., 2001; Lilla et al., 2006; Ognjanovic et al., 2006; Cotterchio et al., 2008), breast (Deitz et al., 2000; Gallicchio et al., 2006), and lung (Chiou et al., 2005) cancer risk in individuals exposed to heterocyclic amine carcinogens. In contrast, other studies carried out with colorectal (Barrett et al., 2003; Barlak et al., 2006), breast (van der Hel et al., 2004; Ochs-Balcome et al., 2007; Mignone et al., 2009), and lung (Barlak et al., 2006) cancer do not support the association with rapid acetylator NAT2 phenotype. A much more robust effect of NAT2 phenotype has been noted for mutagenicity and DNA adduct formation from 2-amino-3,8-dimethylimidazo-[4,5-f]quinoxaline (MeIQx) (Bendaly et al., 2007). Thus, the effects of NAT2 phenotype in the gene-environmental studies may reflect heterocyclic amines such as MeIQx more than they do PhIP as noted for colorectal cancer (Ishibe et al., 2002). In summary, these results strongly support activation of PhIP by CYP1A1 with little effect of human NAT2 genetic polymorphism on mutagenesis and DNA damage.
Declaration of interest
This study was supported by United States Public Health Service grants R01-CA034627 and P30-ES014443.