Chem. Res. Toxicol. 2007, 20, 1075–1083
1075
Articles AlkB Influences the Chloroacetaldehyde-Induced Mutation Spectra and Toxicity in the pSP189 supF Shuttle Vector Min Young Kim,† Xinfeng Zhou,† James C. Delaney,† Koli Taghizadeh,‡ Peter C. Dedon,† John M. Essigmann,†,‡,§ and Gerald N. Wogan*,†,‡,§ Department of Biological Engineering, Center for EnVironmental Health Sciences, and Department of Chemistry, Massachusetts Institute of Technology, 77 Massachusetts AVenue, Cambridge, Massachusetts 02139 ReceiVed May 16, 2007
2-Chloroacetaldehyde (CAA), a metabolite of the carcinogen vinyl chloride, reacts with DNA to form cyclic etheno ()-lesions. AlkB, an iron-/R-ketoglutarate-dependent dioxygenase, repairs 1,N6-ethenodeoxyadenosine (A) and 3,N4-ethenodeoxycytidine (C) in site-specifically modified single-stranded viral genomes in ViVo and also protects the E. coli genome from the toxic effects of CAA. We examined the role of AlkB as a cellular defense against CAA by characterizing the frequencies, types, and distributions of mutations induced in the double-stranded supF gene of pSP189 damaged in Vitro and replicated in AlkB-proficient (AlkB+) and AlkB-deficient (AlkB-) E. coli. AlkB reduced mutagenic potency and increased the survival of CAA-damaged plasmids. Toxicity and mutagenesis data were benchmarked to levels of -adducts and DNA strand breaks measured by LC-MS/MS and a plasmid nicking assay. CAA treatment caused dose-dependent increases in A, C, and 1,N2-ethenodeoxyguanosine (1,N2-G) and small increases in strand breaks and abasic sites. Mutation frequency increased in plasmids replicated in both AlkB+ and AlkB- cells; however, at the maximum CAA dose, the mutation frequency was 5-fold lower in AlkB+ than in AlkB- cells, indicating that AlkB protected the genome from CAA lesions. Most induced mutations in AlkB- cells were G:C to A:T transitions, with lesser numbers of G:C to T:A transversions and A:T to G:C transitions. G:C to A:T and A:T to G:C transitions were lower in AlkB+ cells than in AlkB- cells. Mutational hotspots at G122, G123, and G160 were common to both cell types. Three additional hotspots were found in AlkB- cells (C133, T134, and G159), with a decrease in mutation frequency and change in mutational signature in AlkB+ cells. These results suggest that the AlkB protein contributes to the elimination of exocyclic DNA base adducts, suppressing the toxic and mutagenic consequences induced by this damage and contributing to genetic stability. Introduction Manmade chemicals that enter the cell as well as cellularderived chemicals that enter the cell may on their own or through bioactivation covalently bind to cellular macromolecules including DNA and RNA, thus altering replication, transcription, and translation. There are many types of DNA alkylation damage that can be studied, and this work focuses on the etheno ()DNA lesions, which contain a CdC bridge between the exocyclic and heterocyclic nitrogens of adenine, cytosine, or guanine. Manmade routes to -nucleobase lesions can arise from the process of making polyvinyl chloride (PVC). The main ingredient in making PVC is vinyl chloride (VC1), which is a human and rodent carcinogen (1) and has been linked to form angiosarcoma of the liver (2). VC can be epoxidized by mouse liver microsomes (3), and the cyto-chrome P450 monooxygenase CYP2E1 enzyme has been implicated in generating the highly reactive chloroethylene oxide (CEO) metabolite (4), which can * To whom correspondence should be addressed. Tel: 617-253-3188. Fax: 617-258-0499. E-mail:
[email protected]. † Department of Biological Engineering. ‡ Center for Environmental Health Sciences. § Chemistry Department. | Present address: Barclays Global Investors, 45 Freemont Street, San Francisco, CA 94105.
rearrange to form 2-chloroacetaldehyde (CAA) (5). Although CAA is often used as a vehicle for generating -lesions, CEO is an order of magnitude more efficient at generating such lesions (6, 7). CAA reacts with DNA bases in Vitro, resulting in the formation of exocyclic DNA adducts, including 1,N6-ethenodeoxyadenosine (A), 3,N4-ethenodeoxycytidine (C), 1,N2ethenodeoxyguanosine (1,N2-G), and N2,3-ethenodeoxyguanosine (N2,3-G) (8–11). In addition to exogenous sources of -adducts such as chemicals used in the plastics industry, the aforementioned lesions can arise from lipid peroxidation products generated under oxidative stress and chronic inflammation (12–17), which have been linked to a variety of human 1 Abbreviations: AlkB+, AlkB-proficient; AlkB-, AlkB-deficient; , etheno; VC, vinyl chloride; CEO, 2-chloroethylene oxide; CAA, 2-chloroacetaldehyde; A, 1,N6-ethenodeoxyadenosine; C, 3,N4-ethenodeoxycytidine; 1,N2-G, 1,N2-ethenodeoxyguanosine; N2,3-G, N2,3-ethenodeoxyguanosine; ANPG, alkyl-N-purine-DNA glycosylase; MAG, 3-methyladenine DNA glycosylase; AlkA, 3-methyl-adenine-DNA glycosylase II; MUG, mismatch-specific uracil DNA glycosylase; hTDG, mismatch-specific thymine-DNA glycosylase; cfu, colony forming units; TE, transformation efficiency; MF, mutation frequency; R-HOEG, R-hydroxyethanodeoxyguanosine (6-hydroxy-3,5,6,7-tetrahydro-9H-imidazo-[1,2-a]-purin-9-one); R-HOEC, R-hydroxyethanodeoxycytidine; R-HOEA, R-hydroxyethanodeoxyadenosine.
10.1021/tx700167v CCC: $37.00 2007 American Chemical Society Published on Web 07/20/2007
1076 Chem. Res. Toxicol., Vol. 20, No. 8, 2007
Kim et al.
Figure 1. Experimental design and structure of CAA-induced etheno adducts.
degenerative disorders, such as Wilson’s, Crohn’s, and Alzheimer’s diseases, primary hemochromatosis, ulcerative colitis, chronic pancreatitis, atherosclerosis, and colon cancer (18–25). The likelihood of these simple -adducts being genotoxic and/ or mutagenic has been established in Vitro through primer extension assays (26–33) and in ViVo by site-specific mutagenesis in cells (32, 34–41). Repair of these exocyclic lesions should eliminate the threat to genomic integrity, and indeed, two base excision repair pathways exist for -adducts in Escherichia coli (E. coli) and human cells using conserved DNA glycosylases. In one pathway, the E. coli 3-methyl-adenine-DNA glycosylase II (AlkA), mammalian alkyl-N-purine-DNA glycosylase (ANPG), or yeast 3-methyl-adenine DNA glycosylase (MAG) performs the excision (11, 39, 42–45), and in Vitro, AlkA has been shown to excise A (43) and N2,3-G (44). The other pathway utilizes the E. coli mismatch-specific uracil DNA glycosylase (MUG) or human mismatch-specific thymine-DNA glycosylase (hTDG) to excise the -adduct (46–48), and in Vitro, MUG has been shown to excise C (46) and 1,N2-G (47). Recently, AlkB has been shown to eliminate or reduce the genotoxicity and mutagenicity of A or C, respectively, in single-stranded viral genomes that have been passaged through E. coli (39). AlkB is an E. coli DNA (and RNA) repair enzyme that protects the cell from the toxic effects of several methylating agents (49–51). Mechanistically unlike the base excision repair enzymes, AlkB was originally found to remove the methyl group from DNA containing 1-methyladenine and 3-methylcytosine as formaldehyde (presumably through a hydroxylated intermediate), thus restoring the undamaged bases in DNA (52, 53). Soon after, AlkB was found to also repair 1-methylguanine (54, 55), 3-methylthymine (54–56), and 3-ethylcytosine (54). Although humans possess eight homologues of AlkB (57), only hABH2 and hABH3 have been shown to repair DNA that has been damaged by simple methylating agents (55, 58–60). Surprisingly, hABH3 has also been shown to repair RNA (55, 59–61) as well as A in Vitro (62).
Although previous studies have demonstrated that CAA treatment of plasmids increased the mutation frequency in the supF gene and produced predominantly G:C to A:T transitions and G:C to T:A transversions (63–65), the repair role of AlkB on CAA-induced mutations has not yet been defined. As outlined in Figure 1, here, we analyzed the formation of -adducts and other forms of DNA damage induced by CAA in the pSP189 shuttle vector carrying the supF gene. Their mutagenic properties were assessed in E. coli cells that were either AlkB deficient (AlkB-) (HK82) or AlkB proficient (AlkB+) (HK81). To our knowledge, direct comparisons of -adduct levels with their mutagenic properties and elucidation of the role of AlkBmediated repair in the supF shuttle vector system have not previously been reported.
Experimental Procedures Treatment of Plasmid with CAA. The pSP189 shuttle vector containing an 8-bp signature sequence was a gift from Dr. Michael M. Seidman (NIH, Bethesda, MD). As described previously (66), the pSP189 plasmid was amplified in E. coli AB2463 cells grown at 37 °C in LB media containing 50 µg/mL ampicillin (Sigma) for 12–14 h with shaking at 250 rpm, and plasmid DNA was isolated using a Maxi DNA isolation and purification kit (Qiagen). CAA was obtained as a 50% w/w aqueous solution from Aldrich Chemical Co. (Milwaukee, WI). On the basis of previous studies, 0, 0.15, 0.3, and 0.6 M CAA were used as test concentrations in this study (6365). Plasmid DNA (40 µg) was treated with each concentration of CAA in 0.3 M sodium acetate (pH 7) at 37 °C for 1 h in a total volume of 200 µL, followed by ethanol precipitation of the plasmid to remove unreacted CAA. LC-MS/MS Quantification of -Adducts. The -adducts were quantified by an LC-MS/MS method based on the work of a number of groups (6770), with modifications to accommodate the isolation and quantification of multiple -adducts in the same sample. Plasmid DNA (3 µg) was digested by incubation with 1 U of nuclease P1 for 3 h in 100 µL of sodium acetate buffer (30 mM, pH 5.6) containing zinc chloride (0.2 mM) and desferoxamine (0.2 mM).
AlkB Influences CAA-Induced Mutation and Toxicity Quantities (3.33 pmol) of isotopically labeled internal standards ([15N5]-1,N6-ethenodeoxyadenosine, A;[15N3]-3,N4-ethenodeoxycytosine, C; and[15N5]-1,N2-ethenodeoxyguanosine, 1,N2-G) were added to each DNA sample before the digestion. Lack of the N2,3G standard precluded its quantification. Following the addition of 100 µL of sodium acetate buffer (30 mM, pH 8.1), further hydrolysis and dephosphorylation was achieved with alkaline phosphatase (1 µL, 40 U, New England Biolabs) and phosphodiesterase I (1 µL, 1U, USB) by incubation at 37 °C for 3 h. The reaction mixture was passed over a Microcon YM-30 column, and the solvent was removed under vacuum. Samples were reconstituted in 35 µL of Milli-Q water and stored at –20 °C for subsequent HPLC prepurification. Adducts were prepurified using an Agilent 1100 HPLC system, equipped with a Varian C18 reversed-phase column (250 ∼ 4.6 mm). Following injection of reconstituted samples, the column was eluted at a flow rate of 0.5 mL/min with the following gradient of acetonitrile in 8 mM ammonium acetate buffer (pH 6.9): 0–20 min, 3–10%; 20–30 min, 10–20%; 30–40 min, 20–40%; 40–45 min, 40–100%. The unmodified and -deoxynucleoside adducts were well resolved, and the -adducts were isolated by collection of fractions bracketing their retention times: 1,N2-G, 25.6 min; C, 27.9 min; A, 28.8 min. All corresponding fractions were dried under vacuum for LC-MS/MS analysis. The dried fractions containing -adducts and their corresponding isotopomers were dissolved in 20 µL of water and analyzed by HPLC-ESI-MS/MS using an Agilent 1100 series HPLC system interfaced with an API-Sciex 3000 triple quadrupole mass spectrometer equipped with a TurboIonSpray source. Further chromatographic purification of -adducts was achieved by a 150 mm ∼ 1 mm C18 Agilent Eclipse XDB-C18 column, using acetonitrile (1% for C and A, 2% for 1,N2-G) in 0.1% acetic acid at a flow rate of 0.1 mL/min. Retention times were as follows: C, 10.7 min; A, 12.4 min; and 1,N2-G, 20.7 min. The mass spectrometer was operated in the positive ion mode, with all instrument parameters optimized for maximal sensitivity. Samples were analyzed in the multiple reaction-monitoring (MRM) modes, with the following ion pairs (MH|m+) used for -adducts and their isotopomeric standards: A, 276/160; [15N5]-A, 281/165; C, 252/136; [15N3]C, 255/139; 1,N2-G, 292/176;[15N5]-1,N2-G, 297/181. Quantification of Strand Breaks and Abasic Sites in Plasmid DNA by Nicking Assay. The quantity of strand breaks caused by CAA treatment was determined using a plasmid nicking assay (71). Samples of CAA-treated plasmid DNA were split into halves (500 ng DNA each), with one portion treated with 100 mM putrescine dihydrochloride (Sigma) for 0.5 h at 37 °C to convert all abasic sites to stand breaks (71); the other half was left untreated as a control for direct strand breaks. The plasmid topoisomers present in these DNA samples were resolved on 0.8% agarose gels in the presence of 0.2 µg/mL ethidium bromide at 2.5 V/cm for 2 h in Tris-borate-EDTA buffer (72). The gels were subjected to UV illumination (315 nm) and the quantity of DNA in each band determined by fluorescence imaging (Ultra-Lum, Claremont, CA). The quantity of strand breaks caused by CAA treatment was calculated from the net increase in the percentage of nicked (form II) plasmid (66). Determination of Transformation Efficiency in AlkB+ and AlkB- Cells. E. coli HK81 (AlkB+) and HK82 (AlkB-) cells were grown to log phase density (OD600 ≈ 0.5–0.6) and were washed twice with cold water to ensure the removal of all salts before being placed in a chilled 2 mm gap electroporation cuvette (VWR) with 10 ng of CAA-treated plasmid DNA and pulsed at 12.5 kV/cm, 50 µF, and 129 Ω using an electroporation system (Electro Cell Manipulator 600, BTX, USA). The transformation efficiency (TE) of plasmids into AlkB+ and AlkB- E. coli was determined by immediately adding SOC buffer after electroporation and allowing a 30 min room-temperature recovery time. The cells were then centrifuged at 3,-000g for 5 min and resuspended in PBS, pH 7.0, and plated onto 1.5% agar plates containing 50 µg/mL ampicillin. TE was defined as the number of colony forming units (cfu) produced by 1 µg of pSP189 DNA in a transformation reaction.
Chem. Res. Toxicol., Vol. 20, No. 8, 2007 1077
Figure 2. Increase in etheno adduct levels (A), strand breaks (SB) and abasic sites (AP) (B) in pSP189 plasmid DNA treated with CAA. Data show mean ( SD of three independent experiments (symbols may occlude error bars).
Determination of Mutation Frequency in MBL50 Cells. For mutational analyses, cells were recovered after electroporation and grown in 10 mL of LB medium at 37 °C for 6 h with agitation. Plasmid DNA was harvested using Wizard Plus plasmid miniprep kits (Promega) and stored at 4 °C for re-transformation into MBL50 cells. MBL50 cells were transformed with 10 ng of the recovered DNA as described above. Aliquots (225 µL) of cell suspensions were plated onto medium A containing 50 µg/mL ampicillin, 20 µg/mL IPTG (Roche), and 10 µg/mL X-gal (Roche). The remaining portion of cell suspension was diluted appropriately and plated onto LB agar for the determination of the total number of transformants. This medium was supplemented with 2 g/L L-arabinose (Sigma) for the arabinose resistant selection method. All plates were incubated at 37 °C for 24–96 h. Mutation frequency (MF) was defined as the ratio of total mutants to total transformants. Background mutation frequencies were determined in plasmids exposed to 0.3 M sodium acetate, the solvent for CAA. Individual mutants were restreaked onto plates containing arabinose, ampicillin, IPTG, and X-gal for confirmatory identification and then isolated for sequencing. Analysis of Mutated supF Gene. Mutant plasmids (harvested from white and light blue colonies (73)) were isolated using a Wizard Plus plasmid miniprep kit. DNA sequencing was carried out by the Harvard University DNA Sequencing Facility (Cambridge, MA) using a 20-mer primer with the following sequence: 5′;-GGC-GAC-ACG-GAA-ATG-TTG-AA-3′; (IDT Coralville, IA). The signature sequences were identified using the Sequencer program (Gene Codes Corporation, version 4.1.4). Each sequence was unique, indicating that mutations arose from independent (nonsibling) events. Poisson distribution analysis was used to assess the randomness of the distribution of mutations, and hot spots were defined as described previously (66).
Results Quantification of E-Adducts and Strand Breaks in pSP189 Induced by CAA. Quantitative analysis revealed the dose-dependent formation of adducts in the order A > C >
1078 Chem. Res. Toxicol., Vol. 20, No. 8, 2007
Kim et al.
Figure 3. Transformation efficiency (A) and mutation frequency (B) induced by CAA in the supF gene of pSP189 replicated in E. coli AlkB+ (9) and AlkB- cells (|b7). Data represent mean ( SD for three experiments. Mutation frequencies of spontaneous mutation are subtracted from those induced by CAA (symbols may occlude error bars).
1,N2-G (Figure 2A), ranging from 0.5 to 2.5 per 100 nucleotides (approximately 50–250 per plasmid) at the 0.6 M CAA dosage. Other groups have found that high DNA adduct levels are needed to observe biological effects of CAA; therefore, the plasmid was treated with doses ranging from 0.15 to 0.6 M (63–65) in order to achieve this level of adduction. Although the lack of a standard precluded quantification of N2,3-G, the balance of our results agree with other studies showing DNA exposure to CAA forms A > C > N2,3-G > 1,N2-G (10, 11), which also holds when cells are treated with CAA (74). The formation of strand breaks and abasic sites (SB and AP sites) as measured by the plasmid nicking assay was small in our study; thus, the -load rather than SB and AP sites are responsible for the decrease in transformation efficiency seen in Figure 3A. The 0.6 M CAA treatment resulted in only 5 SB and AP sites per 106 nucleotides (Figure 2B), amounting to nicking of less than 3% of the parent supercoiled plasmid. This result is consistent with the known mechanism of CAA-induced DNA damage, which involves alkylation of DNA bases to form -adducts that are relatively stable toward depurination.
CAA-Induced Mutation Frequencies and Transformation Efficiencies. The MF of alkylated plasmid increased in both cell types (AlkB positive and negative cells) following treatment with CAA, compared to that of the control. The MF induced by the treatment of 0.6 M CAA was 7.5- and 40-fold higher than the spontaneous MF in AlkB+ (21.7 ∼ 10-5 vs 2.9 ∼ 10-5) and AlkB- cells (108 ∼ 10-5 vs 2.7 ∼ 10-5), respectively (Figures 3B and 4), corresponding to a 5-fold higher MF in AlkBcells. Consistent with these results, the relative TEs decreased in a dose-dependent manner following replication in both AlkB+ and AlkB- cells after treatment with CAA. At the 0.6 M CAA dosage (450 measured adducts per plasmid genome), TEs for supF vectors in AlkB+ cells (3.7 ∼ 107 cfu/ug) were also 5-fold higher than those in AlkB- cells (0.7 ∼ 107 cfu/ug), which was seen for all CAA concentrations (Figure 3A). Types of Mutations. We selected mutants induced by 0.6 M CAA in both cell types for the characterization of the mutation spectra. Spectra were first determined in 23 and 19 independent spontaneous mutants from untreated plasmids replicated in AlkB+ and AlkB- cells, respectively. The majority of spontaneous mutations observed were single base-pair substitutions (91% and 89%), and others included multiple sequence changes (9% and 11%) (Table 1 and Figure 4A). The most common single base-pair substitutions were G:C to T:A (33% and 35%) and A:T to T:A (29% and 29%) transversions (Table 2). Also found in both cell types were G:C to A:T transitions (19% and 18%) and G:C to C:G transversions (19% and 12%) (Table 2). We next analyzed 93 and 96 supF CAA-induced mutant plasmids replicated in AlkB+ and AlkB- cells, respectively (Figure 4B). Single base substitutions were found in 89% and 88% of the plasmids; 10% and 6% contained multiple sequence changes (Table 1, Figure 4C). The remainder (1% and 6%) contained deletions. In plasmids harboring single base-pair substitutions, the majority (78% and 84% in AlkB+ and AlkB- cells, respectively) were located at G:C base pairs (Table 2). In AlkB- cells, G:C to A:T transitions (56%) were the most frequent type of mutation, followed by G:C to T:A transversions (24%), A:T to G:C transitions (12%), and G:C to C:G transversions (4%) (Table 2). A comparison of AlkB- with AlkB+ cells shows a reduction of the G:C to A:T (56% vs 25%) and A:T to G:C (12% vs 5%) mutational signatures, and an increase in G:C to C:G (4% vs 23%), A:T to T:A (2% vs 15%), and G:C to T:A (24% vs 30%) mutational signatures (Table 2). Mutation Spectra. The spectra of mutations induced in the supF gene of plasmids replicated in AlkB- and AlkB+ cells following treatment with CAA are summarized in Figure 4B. The mutation spectra included six hotspots (G122, G123, C133, T134, G159, and G160) in AlkB- and five (G122, G123, A135, G141, and G160) in AlkB+ cells. Sites G122, G123, and G160 were common to both cell types.
Discussion The mechanism of -adduct formation by CAA involves an attack of the exocyclic base nitrogen on the carbonyl (9, 7576).
Table 1. Types of Spontaneous and CAA-Induced Mutations in the supF Gene of pSP189 after Replication in AlkB+ and AlkB- E. coli number of mutations (% of total) AlkB mutation type single base pair substitutions deletions multiple sequence changes total
+
AlkB-
spontaneous 21 0 2 23
(91) (0) (9) (100)
CAA-induced 83 1 9 93
(89) (1) (10) (100)
spontaneous 17 0 2 19
(89) (0) (11) (100)
CAA-induced 84 6 6 96
(88) (6) (6) (100)
AlkB Influences CAA-Induced Mutation and Toxicity
Chem. Res. Toxicol., Vol. 20, No. 8, 2007 1079
Figure 4. Mutation spectra in spontaneous supF mutants from unexposed control pSP189 replicated in AlkB- or AlkB+ E. coli (A). Key: open square symbols (0) indicate one base pair deletion. Underlined (_) symbols indicate components of multiple sequence changes within one plasmid. Distribution of mutations induced by CAA in the supF gene of pSP189 replicated in AlkB- or AlkB+ E. coli (B). Key: 9, hotspot sites common to both the AlkB- and AlkB+ cells; O, unique. Deletions are denoted by open square symbols (0). Multiple sequence changes illustrated in C have been omitted and excluded from hotspot analysis. Distribution of multiple sequence changes in the mutated supF gene (C). The diagram shows the locations of base substitution mutations (unshaded columns), single base insertions (shaded columns), and single base-pair deletions (denoted by D) in the supF gene of pSP189 exposed to CAA after replication in AlkB- and AlkB+ E. coli cells.
Table 2. Types of Single Base Pair Substitutions in the supF Gene of Spontaneous and CAA-Induced pSP189 Mutants after Replication in AlkB+ or AlkB- E. coli (Excluding Multiple Sequence Changes) number of base substitution mutations(% of total) AlkB+ base substitution type transversions
transitions total
G:CtoT:A G:CtoC:G A:TtoT:A A:TtoC:G G:CtoA:T A:TtoG:C
AlkB-
spontaneous 7 4 6 0 4 0 21
(33) (19) (29) (0) (19) (0) (100)
CAA-induced 25 19 12 2 21 4 83
The endocyclic base nitrogen can attack the chloride, which would form the structures shown in Figure 5. Dehydration of these ethanohydrates (R-HOEA, R-hydroxyethanodeoxya-denosine; R-HO-
(30) (23) (15) (2) (25) (5) (100)
spontaneous 6 2 5 1 3 0 17
(35) (12) (29) (6) (18) (0) (100)
CAA-induced 20 3 2 2 47 10 84
(24) (4) (2) (2) (56) (12) (100)
EC, R-hydroxyethanodeoxycytidine; and R-HOEG, R-hydroxyethanodeoxyguanosine), which have half-lives of ∼1–48 h (8, 33), generates the respective -adducts (Figure 1).
1080 Chem. Res. Toxicol., Vol. 20, No. 8, 2007
Figure 5. Structures of ethanohydrates, which are intermediates in the CAA-induced formation of the etheno lesions shown in Figure 1.
The recent finding that AlkB can directly reverse A damage in DNA (39, 62) led us to speculate that other types of -lesions may also be repaired by AlkB. To investigate this possibility, a plasmid was treated with CAA, which is known to produce etheno lesions in model systems. Previous mass spectral evidence for the repair of A in DNA was consistent with a mechanism of epoxidation of the etheno CdC bond by the R-ketoglutarate non-heme dioxygenase AlkB enzyme, followed by epoxide hydrolysis and ultimate liberation of the dialhedyde, glyoxal (39). As described for A, this mechanism should hold similarly for the repair of C, 1,N2-G, and N2,3-G lesions, as long as the CdC bond can be situated near the high-valent Fe(4+)dO intermediate held in the active site of AlkB, whose crystal structure has recently been solved (77). The observed decrease in TE and increase in MF when CAAtreated plasmids were passaged through AlkB-deficient E. coli (Figure 3) suggested that AlkB repaired some of the CAAinduced lesions. Accordingly, the mutants were sequenced to gain insight as to lesions involved in the AlkB-sensitive biological activity of CAA. Figure 2A shows that at the 0.6 M concentration of CAA used for analysis, the amount of A formed was twice that of C, which, in turn, was twice that of 1,N2-G. Upon replication of the same duplex CAA-treated plasmid in ViVo, mutations occurred predominantly at guanines or cytosines in both AlkB- or AlkB+ cells (Figure 4B). The paucity of mutations at adenine sites, even in AlkB- cells, may seem at first glance to be surprising given the dominance of A in the adduct spectrum of plasmids entering cells. Our earlier work showed that A is very mutagenic in single-stranded DNA replicated E. coli cells lacking AlkB (39), but we have speculated that AlkB may be a backup to AlkA, which probably has the dominant role in repairing A in duplex DNA. AlkB is likely an important backup, however, in that its ability to repair a replication blocking and highly mutagenic lesion in singlestranded DNA presents an opportunity to repair promoter regions and replication forks, which have significant single-stranded character. Given that the globally modified plasmids used in the present study were duplex, it is likely that AlkA efficiently repaired the A lesions, allowing the mutations at G and C sites to dominate the mutational spectrum. Nevertheless, our data show that AlkB did indeed act on lesions in the CAA-treated plasmid because the mutational spectra in AlkB- and AlkB+ cells were different. The mutational spectra of CAA-treated plasmids replicated in AlkB- and AlkB+ cells (Figure 4B) were compared to our previous analysis of the mutagenesis of etheno adducts inserted one at a time into the genome of a bacteriophage and replicated in ViVo. This comparison was done in order to (a) nominate the specific lesions that are responsible for specific features of the mutational landscape of CAA and (b) propose how the action of AlkB on the global adduct population results in specific changes in the mutational spectrum. We note that the single variable in this experiment was the presence or absence of AlkB, and other repair enzymes for -lesions, such as AlkA and MUG,
Kim et al.
were operative. Hence, the signature changes observed were those purely created by the action of AlkB in ViVo on the CAAmodified genome substrate. As a technical point, the overall MFs need to be considered in the analysis of such data. The MF of CAA-treated plasmids in AlkB- cells was 5∼ that of AlkB+ cells, and this factor should be taken into account in assessing the effects of AlkB in Figure 4B; however, spontaneous mutations were negligible when comparing AlkB- and AlkB+ cells at the 0.6 M[CAA] used for analysis. In assessing possible lesions that may have caused the mutations, we utilized information from previous site-specific lesion-induced mutagenesis studies. Studies in AlkB- E. coli, performed on single-stranded DNA, showed that A resulted in 25% A to T, 5% A to G, and 5% A to C mutations, and C induced 50% C to A and 30% C to T mutations (39). In AlkB+ E. coli, N2,3-G induced 13% G to A transitions as the sole mutation (40) and 1,N2-G gave 2% G to A and 0.75% G to T mutations (in double-stranded DNA, using NER-, UV-irradiated cells) (32). Additionally, it is important to note that the local sequence context of a lesion, which is usually different in such studies from different groups, may vary the preference for certain base insertions as well as polymerase traversal, as has been found for A and N2,3-G (,79). Sequence context may also aid in preferential adduct formation or diminished adduct repair. In AlkB- cells, CAA-treated plasmids suffered an excess of G:C to A:T mutations, suggesting that an G or C adduct may be the premutagenic lesion (see, for example, positions 123, 159, and 160 in Figure 4B). The G to A transition is the mutational signature of N2,3-G. However, this lesion does not appear to have the proper geometry to be acted upon by AlkB, at least by its canonical mechanism; therefore, the reduction in mutation and change in mutational signature at these sites in AlkB+ cells may be due to a different lesion being repaired. A second attractive possibility to explain the source of the G:C: to A:T mutations is C, which induces C to T mutations and is repaired by AlkB. While it induces C to T mutations, it also induces C to A at about twice the C to T frequency. We did not observe an excess of C to A mutations, but it is possible that the relative amounts of the two mutations may be sequence context dependent. Hence, C, along with the G lesions, remain formal and not mutually exclusive candidates to explain the G:C to A:T mutations observed in the mutational spectrum. Although not measured analytically, a final possible causal lesion is the ethanocytidine hydrate. It is unknown if this precursor to C is repaired by AlkB. Another conspicuous feature of the mutational spectrum is the G:C to T:A at position 123 in AlkB+ cells. This mutation could be due to the 1,N2-G adduct, which may be a poor AlkB substrate in this sequence context or in general. The residual G:C to C:G signature at position 159 in AlkB+ cells could be due to the ethanoguanosine hydrate; its attempted replication by T7 DNA polymerase gives nearly equal incorporation of G and A (33). The residual mixture of G to T and G to C mutations at position 160 in AlkB+ cells could arise from a mixture of poorly repaired 1,N2-G and the ethanoguanosine hydrate. Sequence context, once again, may have modulated the relative amount of each lesion formed as well as lesion repair. Diminution of C to A (or G to T) mutations at position 133 in AlkB+ compared to those in AlkB– cells would be consistent with a mutational signal of C or 1,N2-G, which may be partially repaired by AlkB. Normalizing for the MF differences, position 122 had 15 (G to A) and 10 (G to T) mutations in AlkB- cells, which decreased in AlkB+ cells to 2 (G to A) and
AlkB Influences CAA-Induced Mutation and Toxicity
1 (G to T) but also to newly seen 2 (G to C) mutations. Because N2,3-G is known to produce exclusively G to A mutations, it is most likely not the lesion in question. Both C and 1,N2-G would give the mutational signature observed in AlkB- cells, and the drop in mutation load in AlkB+ cells would be suggestive of their repair by AlkB. The remaining G to C mutations, refractory to repair in AlkB+ cells, may be due to the ethanohydrate of guanine, which can give G to C, in addition to G to T mutations (Vide supra). Normalizing for MF differences, position 134 had 25 (T to C) mutations in AlkB- for every 2 (T to A) mutations in AlkB+ E. coli. Because one would expect reaction at adenine (T is not known to react with CAA), the preponderance of A to G mutations in AlkB- cells would come from an adenine-derived lesion that evaded repair by the known glycosylases but can somehow be repaired by AlkB. As indicated above, this is not a mutation of A, and there are other reasons to believe that A does not play a major role in the mutational spectra observed. However, the chemical precursor to A, the ethanohydrate of adenine, is a viable candidate. Ethano lesions are poorly repaired by base excision repair enzymes, compared to their unsaturated etheno counterparts (80, 81). We have recently shown that AlkB can metabolize ethanoadenine via hydroxylation off the N1 adenine position (82). A similar repair mechanism can be envisioned for the ethanohydrates shown in Figure 5, which would afford a mechanism of directly reversing the base damage in the form of the dialdehyde glyoxal, reminiscent of the repair of A by AlkB. Evidence from our current study may be interpreted within a broader perspective. Exocyclic -adducts are known to be produced in cellular DNA not only by reaction with metabolites of environmental/industrial chemicals such as vinly chloride but also from endogenous sources through interactions with DNA of lipid peroxidation-derived aldehydes and hydroxyal-kenals. Although present in normal human and rodent liver DNA, levels of exocyclic -adducts are significantly increased by lipid peroxidation and oxidative stress. Our findings provide supportive evidence regarding mechanisms through which exocyclic DNA adducts, together with other forms of oxidative DNA damage, may play a role in the well-documented association between chronic inflammation and multistage human carcinogenesis. Acknowledgment. We thank Laura J. Trudel for manuscript preparation. The LC-MS/MS analyses were performed in the MIT Center for Environmental Health Sciences Bioanalytical Facilities Core. This work was supported by National Cancer Institute Grants 5 P01 CA26731 and 5 RO1 CA80024, and NIEHS Center Grant ES02109.
References (1) Agency for Toxic Substances and Disease Registry (ATSDR). (1993) Toxicological Profile for Vinyl Chloride. U.S. Department of Health and Human Service, Public Health Services, Atlanta, GA. (2) Lelbach, W. K. (1996) A 25-year follow-up study of heavily exposed vinyl chloride workers in Germany. Am. J. Ind. Med. 29, 446–446. (3) Bartsch, H., Malaveille, C., Barbin, A., and Planche, G. (1979) Mutagenic and alkylating metabolites of halo-ethylenes, chlorobutadienes and dichlorobutenes produced by rodent or human liver tissues. Evidence for oxirane formation by P450-linked microsomal monooxygenases. Arch. Toxicol. 41, 249–249. (4) El Ghissassi, F., Barbin, A., and Bartsch, H. (1998) Metabolic activation of vinyl chloride by rat liver microsomes: low-dose kinetics and involvement of cytochrome P450 2E1. Biochem. Pharmacol. 55, 1445–1445. (5) Guengerich, F. P., Crawford, W. M., Jr., and Watanabe, P. G. (1979) Activation of vinyl chloride to covalently bound metabolites: roles of 2-chloroethylene oxide and 2-chloroacetaldehyde. Biochemistry 18, 5177–5177.
Chem. Res. Toxicol., Vol. 20, No. 8, 2007 1081 (6) Guengerich, F. P. (1992) Roles of the vinyl chloride oxidation products 1-chlorooxirane and 2-chloroacetaldehyde in the in vitro formation of etheno adducts of nucleic acid bases. Chem. Res. Toxicol. 5, 2–2. (7) Guengerich, F. P., Persmark, M., and Humphreys, W. G. (1993) Formation of 1,-N2- and N2,3-ethenoguanine from 2-halooxiranes: isotopic labeling studies and isolation of a hemiaminal derivative of N2-(2-oxoethyl)-guanine. Chem. Res. Toxicol. 6, 635–635. (8) Kusmierek, J. T., and Singer, B. (1982) Chloroacetaldehyde-treated ribo- and deoxyribopolynucleotides. 1. Reaction products. Biochemistry 21, 5717–5717. (9) Leonard, N. J. (1984) Etheno-substituted nucleotides and coenzymes: fluorescence and biological activity. CRC Crit. ReV. Biochem. 15, 125– 125. (10) Kusmierek, J. T., and Singer, B. (1992) 1,N2-ethenodeoxyguanosine: properties and formation in chloroacetaldehyde-treated polynucleotides and DNA. Chem. Res. Toxicol. 5, 634–634. (11) Dosanjh, M. K., Chenna, A., Kim, E., Fraenkel-Conrat, H., Samson, L., and Singer, B. (1994) All four known cyclic adducts formed in DNA by the vinyl chloride metabolite chloroacetaldehyde are released by a human DNA glycosylase. Proc. Natl. Acad. Sci. U.S.A. 91, 1024– 1024. (12) El Ghissassi, F., Barbin, A., Nair, J., and Bartsch, H. (1995) Formation of 1,-N6-ethenoadenine and 3,-N4-ethenocytosine by lipid peroxidation products and nucleic acid bases. Chem. Res. Toxicol. 8, 278–278. (13) Chung, F. L., Chen, H. J., and Nath, R. G. (1996) Lipid peroxidation as a potential endogenous source for the formation of exocyclic DNA adducts. Carcinogenesis 17, 2105–2105. (14) Marnett, L. J. (2000) Oxyradicals and DNA damage. Carcinogenesis 21, 361–361. (15) Blair, I. A. (2001) Lipid hydroperoxide-mediated DNA damage. Exp. Gerontol. 36, 1473–1473. (16) Lee, S. H., Arora, J. A., Oe, T., and Blair, I. A. (2005) 4-Hydroperoxy2-nonenal-induced formation of 1,N2-etheno-2′;-deoxyguanosine adducts. Chem. Res. Toxicol. 18, 780–780. (17) Pang, B., Zhou, X., Yu, H., Dong, M., Taghizadeh, K., Wishnok, J. S., Tannenbaum, S. R., and Dedon, P. C. (2007) Lipid peroxidation dominates the chemistry of DNA adduct formation in a mouse model of inflammation. Carcinogenesis[Online early access], DOI: 10.1093/ carcin/bgm037. (18) Singer, B., and Bartsch, H. (1999) Exocyclic DNA Adducts in Mutagenesis and Carcinogenesis, Proceedings of the 2nd International Conference, Heidelberg, Germany, September, 1998, IARC Scientific Publications, Lyon, France, pp 1–1. (19) Nair, J., Carmichael, P. L., Fernando, R. C., Phillips, D. H., Strain, A. J., and Bartsch, H. (1998) Lipid peroxidation-induced etheno-DNA adducts in the liver of patients with the genetic metal storage disorders Wilson’s disease and primary hemochromatosis. Cancer Epidemiol., Biomarkers PreV. 7, 435–435. (20) Bartsch, H., and Nair, J. (2004) Oxidative stress and lipid peroxidationderived DNA-lesions in inflammation driven carcinogenesis. Cancer Detect. PreV. 28, 385–385. (21) Witztum, J. L., and Steinberg, D. (2001) The oxidative modification hypothesis of atherosclerosis: does it hold for humans. Trends CardioVasc. Med. 11, 93–93. (22) Arlt, S., Beisiegel, U., and Kontush, A. (2002) Lipid peroxidation in neurodegeneration: new insights into Alzheimer’s disease. Curr. Opin. Lipidol. 13, 289–289. (23) Owen, R. W. (2001) Biomarkers in colorectal cancer. IARC Sci. Publ. 154, 101–101. (24) Nair, J., De, F. S., Izzotti, A., and Bartsch, H. (2007) Lipid peroxidation-derived etheno-DNA adducts in human atherosclerotic lesions. Mutat. Res. DOI: 10.1016/j.mrfmmm.2007.02.013. (25) Nair, J., Gansauge, F., Beger, H., Dolara, P., Winde, G., and Bartsch, H. (2006) Increased etheno-DNA adducts in affected tissues of patients suffering from Crohn’s diseaseulcerative colitischronic pancreatitis. Antioxid. Redox Signaling 8, 1003–1003. (26) Hall, J. A., Saffhill, R., Green, T., and Hathway, D. E. (1981) The induction of errors during in vitro DNA synthesis following chloroacetaldehyde-treatment of poly-(dA-dT) and poly-(dC-dG) templates. Carcinogenesis 2, 141–141. (27) Barbin, A., Bartsch, H., Leconte, P., and Radman, M. (1981) Studies on the miscoding properties of 1,-N6-ethenoadenine and 3,-N4ethenocytosineDNA reaction products of vinyl chloride metabolitesduring in vitro DNA synthesis. Nucleic Acids Res. 9, 375–375. (28) Singer, B., and Spengler, S. J. (1986) Replication and transcription of polynucleotides containing ethenocytosineethenoadenine and their hydrated intermediates. IARC Sci. Publ. 70, 359–359. (29) Singer, B., Kusmierek, J. T., Folkman, W., Chavez, F., and Dosanjh, M. K. (1991) Evidence for the mutagenic potential of the vinyl chloride induced adductN2,3-etheno-deoxyguanosineusing a site-directed kinetic assay. Carcinogenesis 12, 745–745. (30) Simha, D., Palejwala, V. A., and Humayun, M. Z. (1991) Mechanisms of mutagenesis by exocyclic DNA adducts. Construction and in vitro
1082 Chem. Res. Toxicol., Vol. 20, No. 8, 2007
(31)
(32)
(33)
(34)
(35) (36)
(37)
(38)
(39)
(40)
(41) (42)
(43)
(44)
(45) (46)
(47)
(48)
template characteristics of an oligonucleotide bearing a single sitespecific ethenocytosine. Biochemistry 30, 8727–8727. Langouet, S., Muller, M., and Guengerich, F. P. (1997) Misincorporation of dNTPs opposite 1,-N2-ethenoguanine and 5,6,7,9-tetrahydro7-hydroxy-9-oxoimidazo-[1,2-a]-purine in oligonucleotides by Escherichia coli polymerases I exo- and II exo-T7 polymerase exo-human immunodeficiency virus-1 reverse transcriptaserat polymerase beta. Biochemistry 36, 6069–6069. Langouet, S., Mican, A. N., Muller, M., Fink, S. P., Marnett, L. J., Muhle, S. A., and Guengerich, F. P. (1998) Misincorporation of nucleotides opposite five-membered exocyclic ring guanine derivatives by Escherichiacoli polymerases in vitro and in vivo: 1,-N2-ethenoguanine5,6,7,9-tetrahydro-9-oxoimidazo-[1,2-a]-purine5,6,7,9-tetrahydro7-hydroxy-9-oxoimidazo-[1,2-a]-purine. Biochemistry 37, 5184–5184. Goodenough, A. K., Kozekov, I. D., Zang, H., Choi, J. Y., Guengerich, F. P., Harris, T. M., and Rizzo, C. J. (2005) Site specific synthesis and polymerase bypass of oligonucleotides containing a 6-hydroxy3,5,6,7-tetrahydro-9H-imidazo-[1,2-a]-purin-9-one basean intermediate in the formation of 1,-N2-etheno-2′;-deoxyguanosine. Chem. Res. Toxicol. 18, 1701–1701. Basu, A. K., Wood, M. L., Niedernhofer, L. J., Ramos, L. A., and Essigmann, J. M. (1993) Mutagenic and genotoxic effects of three vinyl chloride-induced DNA lesions: 1,N6-ethenoadenine 3,N4ethenocytosine4-amino-5-(imidazol-2-yl)-imidazole. Biochemistry 32, 12793–12793. Pandya, G. A., and Moriya, M. (1996) 1,N6-Ethenodeoxyadenosinea DNA adduct highly mutagenic in mammalian cells. Biochemistry 35, 11487–11487. Levine, R. L., Yang, I. Y., Hossain, M., Pandya, G. A., Grollman, A. P., and Moriya, M. (2000) Mutagenesis induced by a single 1,N6-ethenodeoxyadenosine adduct in human cells. Cancer Res. 60, 4098–4098. Moriya, M., Zhang, W., Johnson, F., and Grollman, A. P. (1994) Mutagenic potency of exocyclic DNA adducts: marked differences between Escherichia coli and simian kidney cells. Proc. Natl. Acad. Sci. U.S.A. 91, 11899–11899. Palejwala, V. A., Simha, D., and Humayun, M. Z. (1991) Mechanisms of mutagenesis by exocyclic DNA adducts. Transfection of M13 viral DNA bearing a site-specific adduct shows that ethenocytosine is a highly efficient RecA-independent mutagenic noninstructional lesion. Biochemistry 30, 8736–8736. Delaney, J. C., Smeester, L., Wong, C., Frick, L. E., Taghizadeh, K., Wishnok, J. S., Drennan, C. L., Samson, L. D., and Essigmann, J. M. (2005) AlkB reverses etheno DNA lesions caused by lipid oxidation in vitro and in vivo. Nat. Struct. Mol. Biol. 12, 855–855. Cheng, K. C., Preston, B. D., Cahill, D. S., Dosanjh, M. K., Singer, B., and Loeb, L. A. (1991) The vinyl chloride DNA derivative N2,3ethenoguanine produces GsA transitions in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 88, 9974–9974. Akasaka, S., and Guengerich, F. P. (1999) Mutagenicity of sitespecifically located 1,N2-ethenoguanine in Chinese hamster ovary cell chromosomal DNA. Chem. Res. Toxicol. 12, 501–501. Singer, B., Antoccia, A., Basu, A. K., Dosanjh, M. K., Fraenkel-Conrat, H., Gallagher, P. E., Kusmierek, J. T., Qiu, Z. H., and Rydberg, B. (1992) Both purified human 1,N6-ethenoadenine-binding protein and purified human 3-methyladenine-DNA glycosylase act on 1,-N6ethenoadenine and 3-methyladenine. Proc. Natl. Acad. Sci. U.S.A. 89, 9386–9386. Saparbaev, M., Kleibl, K., and Laval, J. (1995) Escherichia coli Saccharomyces cereVisiaerat and human 3-methyladenine DNA glycosylases repair 1,N6-ethenoadenine when present in DNA. Nucleic Acids Res. 23, 3750–3750. Matijasevic, Z., Sekiguchi, M., and Ludlum, D. B. (1992) Release of N2,3-ethenoguanine from chloroacetaldehyde-treated DNA by Escherichia coli 3-methyladenine DNA glycosylase II. Proc. Natl. Acad. Sci. U.S.A. 89, 9331–9331. Fortini, P., Parlanti, E., Sidorkina, O. M., Laval, J., and Dogliotti, E. (1999) The type of DNA glycosylase determines the base excision repair pathway in mammalian cells. J. Biol. Chem. 274, 15230–15230. Saparbaev, M., and Laval, J. (1998) 3,N4-ethenocytosinea highly mutagenic adductis a primary substrate for Escherichia coli doublestranded uracil-DNA glycosylase and human mismatch-specific thymine-DNA glycosylase. Proc. Natl. Acad. Sci. U.S.A. 95, 8508–8508. Saparbaev, M., Langouet, S., Privezentzev, C. V., Guengerich, F. P., Cai, H., Elder, R. H., and Laval, J. (2002) 1,N-(2)-Ethenoguaninea mutagenic DNA adductis a primary substrate of Escherichia coli mismatch-specific uracil-DNA glycosylase and human alkylpurineDNA-N-glycosylase. J. Biol. Chem. 277, 26987–26987. Jurado, J., Maciejewska, A., Krwawicz, J., Laval, J., and Saparbaev, M. K. (2004) Role of mismatch-specific uracil-DNA glycosylase in repair of 3,-N4-ethenocytosine in vivo. DNA Repair 3, 1579–1579.
Kim et al. (49) Kataoka, H., Yamamoto, Y., and Sekiguchi, M. (1983) A new gene (alkB) of Escherichia coli that controls sensitivity to methyl methane sulfonate. J. Bacteriol. 153, 1301–1301. (50) Dinglay, S., Trewick, S. C., Lindahl, T., and Sedgwick, B. (2000) Defective processing of methylated single-stranded DNA by E. coli AlkB mutants. Genes DeV. 14, 2097–2097. (51) Falnes, P. O., and Rognes, T. (2003) DNA repair by bacterial AlkB proteins. Res. Microbiol. 154, 531–531. (52) Trewick, S. C., Henshaw, T. F., Hausinger, R. P., Lindahl, T., and Sedgwick, B. (2002) Oxidative demethylation by Escherichia coli AlkB directly reverts DNA base damage. Nature 419, 174–174. (53) Falnes, P. O., Johansen, R. F., and Seeberg, E. (2002) AlkB-mediated oxidative demethylation reverses DNA damage in Escherichia coli. Nature 419, 178–178. (54) Delaney, J. C., and Essigmann, J. M. (2004) Mutagenesisgenotoxicityrepair of 1-methyladenine3-alkylcytosines1-methylguanine3-methylthymine in alkB Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 101, 14051–14051. (55) Falnes, P. O. (2004) Repair of 3-methylthymine and 1-methylguanine lesions by bacterial and human AlkB proteins. Nucleic Acids Res. 32, 6260–6260. (56) Koivisto, P., Robins, P., Lindahl, T., and Sedgwick, B. (2004) Demethylation of 3-methylthymine in DNA by bacterial and human DNA dioxygenases. J. Biol. Chem. 279, 40470–40470. (57) Kurowski, M. A., Bhagwat, A. S., Papaj, G., and Bujnicki, J. M. (2003) Phylogenomic identification of five new human homologs of the DNA repair enzyme AlkB. BMC Genomics 4, 48. (58) Duncan, T., Trewick, S. C., Koivisto, P., Bates, P. A., Lindahl, T., and Sedgwick, B. (2002) Reversal of DNA alkylation damage by two human dioxygenases. Proc. Natl. Acad. Sci. U.S.A. 99, 16660–16660. (59) Aas, P. A., Otterlei, M., Falnes, P. O., Vagbo, C. B., Skorpen, F., Akbari, M., Sundheim, O., Bjoras, M., Slupphaug, G., Seeberg, E., and Krokan, H. E. (2003) Human and bacterial oxidative demethylases repair alkylation damage in both RNA and DNA. Nature 421, 859– 859. (60) Falnes, P. O., Bjoras, M., Aas, P. A., Sundheim, O., and Seeberg, E. (2004) Substrate specificities of bacterial and human AlkB proteins. Nucleic Acids Res. 32, 3456–3456. (61) Ougland, R., Zhang, C. M., Liiv, A., Johansen, R. F., Seeberg, E., Hou, Y. M., Remme, J., and Falnes, P. O. (2004) AlkB restores the biological function of mRNA and tRNA inactivated by chemical methylation. Mol. Cell 16, 107–107. (62) Mishina, Y., Yang, C. G., and He, C. (2005) Direct repair of the exocyclic DNA adduct 1,-N6-ethenoadenine by the DNA repair AlkB proteins. J. Am. Chem. Soc. 127, 14594–14594. (63) Matsuda, T., Yagi, T., Kawanishi, M., Matsui, S., and Takebe, H. (1995) Molecular analysis of mutations induced by 2-chloroacetaldehyde the ultimate carcinogenic form of vinyl chloridein human cells using shuttle vectors. Carcinogenesis 16, 2389–2389. (64) Chiang, S. Y., Swenberg, J. A., Weisman, W. H., and Skopek, T. R. (1997) Mutagenicity of vinyl chloride and its reactive metaboliteschloroethylene oxide and chloroacetaldehydein a metabolically competent human B-lymphoblastoid line. Carcinogenesis 18, 31–31. (65) Choi, J. H., and Pfeifer, G. P. (2004) DNA damage and mutations produced by chloroacetaldehyde in a CpG-methylated target gene. Mutat. Res. 568, 245–245. (66) Kim, M. Y., Dong, M., Dedon, P. C., and Wogan, G. N. (2005) Effects of peroxynitrite dose and dose rate on DNA damage and mutation in the supF shuttle vector. Chem. Res. Toxicol. 18, 76–76. (67) Doerge, D. R., Churchwell, M. I., Fang, J. L., and Beland, F. A. (2000) Quantification of etheno-DNA adducts using liquid chromatography on-line sample processingelectrospray tandem mass spectrometry. Chem. Res. Toxicol. 13, 1259–1259. (68) Chen, H. J., Lin, T. C., Hong, C. L., and Chiang, L. C. (2001) Analysis of 3,N-(4)-ethenocytosine in DNA and in human urine by isotope dilution gas chromatography/negative ion chemical ionization/mass spectrometry. Chem. Res. Toxicol. 14, 1612–1612. (69) Churchwell, M. I., Beland, F. A., and Doerge, D. R. (2002) Quantification of multiple DNA adducts formed through oxidative stress using liquid chromatography and electrospray tandem mass spectrometry. Chem. Res. Toxicol. 15, 1295–1295. (70) Ham, A. J., Engelward, B. P., Koc, H., Sangaiah, R., Meira, L. B., Samson, L. D., and Swenberg, J. A. (2004) New immunoaffinity-LCMS/MS methodology reveals that Aag null mice are deficient in their ability to clear 1,-N6-etheno-deoxyadenosine DNA lesions from lung and liver in vivo. DNA Repair 3, 257–257. (71) Dedon, P. C., Salzberg, A. A., and Xu, J. (1993) Exclusive production of bistranded DNA damage by calicheamicin. Biochemistry 32, 3617– 3617. (72) Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (1989) Current Protocols in Molecular Biology, John Wiley and Sons, New York.
AlkB Influences CAA-Induced Mutation and Toxicity (73) Juedes, M. J., and Wogan, G. N. (1996) Peroxynitrite-induced mutation spectra of pSP189 following replication in bacteria and in human cells. Mutat. Res. 349, 51–51. (74) Foiles, P. G., Miglietta, L. M., Nishikawa, A., Kusmierek, J. T., Singer, B., and Chung, F. L. (1993) Development of monoclonal antibodies specific for 1,-N2-ethenodeoxyguanosine and N2,3-ethenodeoxyguanosine and their use for quantitation of adducts in G12 cells exposed to chloroacetaldehyde. Carcinogenesis 14, 113–113. (75) Sattsangi, P. D., Leonard, N. J., and Frihart, C. R. (1977) 1,-N2ethenoguanine and N2,3-ethenoguanine. Synthesis and comparison of the electronic spectral properties of these linear and angular triheterocycles related to the Y bases. J. Org. Chem. 42, 3292–3292. (76) Guengerich, F. P., and Persmark, M. (1994) Mechanism of formation of ethenoguanine adducts from 2-haloacetaldehydes: 13C-labeling patterns with 2-bromoacetaldehyde. Chem. Res. Toxicol. 7, 205–205. (77) Yu, B., Edstrom, W. C., Benach, J., Hamuro, Y., Weber, P. C., Gibney, B. R., and Hunt, J. F. (2006) Crystal structures of catalytic complexes of the oxidative DNA/RNA repair enzyme AlkB. Nature 439, 879– 879. (78) Litinski, V., Chenna, A., Sagi, J., and Singer, B. (1997) Sequence context is an important determinant in the mutagenic potential of
Chem. Res. Toxicol., Vol. 20, No. 8, 2007 1083
(79)
(80)
(81)
(82)
1,N6-ethenodeoxyadenosine (epsilonA): formation of epsilonA basepairs and elongation in defined templates. Carcinogenesis 18, 1609– 1609. Mroczkowska, M. M., and Kusmierek, J. T. (1993) The effect of neighboring bases on miscoding properties of N2,3-ethenoguanine. Z. Naturforsch., C: Biosci. 48, 63–63. Guliaev, A. B., Singer, B., and Hang, B. (2004) Chloroethylnitrosoureaderived ethano cytosine and adenine adducts are substrates for Escherichia coli glycosylases excising analogous etheno adducts. DNA Repair 3, 1311–1311. Borys-Brzywczy, E., Arczewska, K. D., Saparbaev, M., Hardeland, U., Schar, P., and Kusmierek, J. T. (2005) Mismatch dependent uracil/ thymine-DNA glycosylases excise exocyclic hydroxyethano and hydroxypropano cytosine adducts. Acta Biochim. Pol. 52, 149–149. Frick, L. E., Delaney, J. C., Wong, C., Drennan, C. L., and Essigmann, J. M. (2007) Alleviation of 1,-N6-ethanoadenine genotoxicity by the Escherichia coli adaptive response protein AlkB. Proc. Natl. Acad. Sci. U.S.A. 104, 755–755.
TX700167V
