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• W2093264695 abstract "It was previously shown that 1,N 6-ethenoadenine (εA) in DNA rearranges into a pyrimidine ring-opened derivative of 20-fold higher mutagenic potency in Escherichia coli (AB1157 lacΔU169) than the parental εA (Basu, A. K., Wood, M. L., Niedernhofer, L. J., Ramos, L. A., and Essigmann, J. M. (1993) Biochemistry 32, 12793–12801). We have found that at pH 7.0, the stability of the N-glycosidic bond in εdA is 20-fold lower than in dA. In alkaline conditions, but also at neutrality, εdA depurinates or converts into products: εdA → B → C → D. Compound B is a product of water molecule addition to the C(2)–N(3) bond, which is in equilibrium with a product of N(1)–C(2) bond rupture in εdA. Compound C is a deformylated derivative of ring-opened compound B, which further depurinates yielding compound D. Ethenoadenine degradation products are not recognized by human N-alkylpurine-DNA glycosylase, which repairs εA. Product B is excised from oligodeoxynucleotides byE. coli formamidopyrimidine-DNA glycosylase (Fpg) and endonuclease III (Nth). Repair by the Fpg protein is as efficient as that of 7,8-dihydro-8-oxoguanine when the excised base is paired with dT and dC but is less favorable when paired with dG and dA. Ethenoadenine rearrangement products are formed in oligodeoxynucleotides also at neutral pH at the rate of about 2–3% per week at 37 °C, and therefore they may contribute to εA mutations. It was previously shown that 1,N 6-ethenoadenine (εA) in DNA rearranges into a pyrimidine ring-opened derivative of 20-fold higher mutagenic potency in Escherichia coli (AB1157 lacΔU169) than the parental εA (Basu, A. K., Wood, M. L., Niedernhofer, L. J., Ramos, L. A., and Essigmann, J. M. (1993) Biochemistry 32, 12793–12801). We have found that at pH 7.0, the stability of the N-glycosidic bond in εdA is 20-fold lower than in dA. In alkaline conditions, but also at neutrality, εdA depurinates or converts into products: εdA → B → C → D. Compound B is a product of water molecule addition to the C(2)–N(3) bond, which is in equilibrium with a product of N(1)–C(2) bond rupture in εdA. Compound C is a deformylated derivative of ring-opened compound B, which further depurinates yielding compound D. Ethenoadenine degradation products are not recognized by human N-alkylpurine-DNA glycosylase, which repairs εA. Product B is excised from oligodeoxynucleotides byE. coli formamidopyrimidine-DNA glycosylase (Fpg) and endonuclease III (Nth). Repair by the Fpg protein is as efficient as that of 7,8-dihydro-8-oxoguanine when the excised base is paired with dT and dC but is less favorable when paired with dG and dA. Ethenoadenine rearrangement products are formed in oligodeoxynucleotides also at neutral pH at the rate of about 2–3% per week at 37 °C, and therefore they may contribute to εA mutations. 1,N 6-ethenoadenine 1,N 6-ethenodeoxyadenosine humanN-methylpurine-DNA glycosylase, truncated form 3,N 4-ethenocytosine 7,8-dihydro-8-oxoguanine high performance liquid chromatography 7MeG-2,6-diamino-5N-methyl-formamidopyrimidine E. coli formamidopyrimidine-DNA glycosylase E. coli endonuclease III polyacrylamide gel electrophoresis mass spectrometry 1,N 6-Ethenoadenine (εA)1 and other exocyclic DNA adducts such as 3,N 4-ethenocytosine (εC) or N 2,3-ethenoguanine (εG) are introduced to DNA by the human carcinogen vinyl chloride and related compounds (1Bartsch H. Barbin A. Marion M.-J. Nair J. Guichard Y. Drug Metab. Rev. 1994; 26: 349-371Crossref PubMed Scopus (151) Google Scholar). These DNA lesions are also formed during interaction with DNA of the peroxidation products of ω-6-polyunsaturated fatty acids (2Chung F.-L. Chen H.-J.C. Nath R.G. Carcinogenesis. 1996; 17: 2105-2111Crossref PubMed Scopus (328) Google Scholar). Ethenoadenine has been found in the DNA of unexposed humans and rodents at highly variable levels, ranging from 0.043 to 31.2 εA molecules/108 of unmodified adenine residues (3Barbin A. Exocyclic DNA adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. International Agency for Research on Cancer Scientific Publication No. 150. IARC, Lyon, France1999: 303-313Google Scholar, 4Nair J. Exocyclic DNA Adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. IARC Scientific Publication No. 150. International Agency for Research on Cancer, Lyon, France1999: 55-61Google Scholar). Upon treatment of animals with vinyl chloride, the level of εA increased in the DNA of rat liver, lung, lymphocytes, and testis (in liver and lung severalfold) (3Barbin A. Exocyclic DNA adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. International Agency for Research on Cancer Scientific Publication No. 150. IARC, Lyon, France1999: 303-313Google Scholar). The level of ε-DNA adducts correlates with increased oxidative stress, such as observed during the accumulation of transient metal ions in Wilson disease, a human metal storage disease, and with increased content of polyunsaturated fatty acids in the diet (5Nair J. Sone H. Nagao M. Barbin A. Bartsch H. Cancer Res. 1996; 56: 1267-1271PubMed Google Scholar, 6Nair J. Carmichael P.L. Fernando R.C. Phillips D.H. Strain A.J. Bartsch H. Cancer Epidemiol. Biomarkers Prev. 1998; 7: 435-440PubMed Google Scholar, 7Bartsch H. Nair J. Owen R.W. Carcinogenesis. 1999; 20: 2209-2218Crossref PubMed Scopus (405) Google Scholar). Ethenoadenine is eliminated from DNA by N-methylpurine-DNA glycosylases. Eukaryotic glycosylases from yeast, rat, and human excise this lesion about 500-fold more efficiently than bacterial AlkA protein (8Saparbaev M. Kleibl K. Laval J. Nucleic Acids Res. 1995; 23: 3750-3755Crossref PubMed Scopus (214) Google Scholar). Molecular dosimetry experiments suggest, however, that it is a persistent lesion, because 2 weeks after exposure of rats to vinyl chloride, the εA level in liver DNA remains very similar to that obtained directly after treatment (9Swenberg J.A. Bogdanffy M.S. Ham A. Holt S. Kim A. Morinello E.J. Ranasinghe A. Scheller N. Upton P.B. Exocyclic DNA Adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. IARC Scientific Publication No. 150. International Agency for Research on Cancer, Lyon, France1999: 29-43Google Scholar). All known exocyclic DNA adducts are mutagenic. In bacteria, εA is recognized mostly as an unmodified adenine by DNA polymerases, infrequently giving rise to AT → TA substitutions (10Moriya M. Pandya F. Johnson F. Grollman A.P. Exocyclic DNA Adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. International Agency for Research on Cancer Scientific Publication No. 150. IARC, Lyon, France1999: 263-270Google Scholar). In mammalian cells, ε-DNA adducts are classified among lesions with the highest mutagenic potency. In site-directed mutagenesis either 70 (10) or 10% (11Levine R.L. Yang I.-Y. Hossain M. Pandya G.A. Grollman A.P. Moriya M. Cancer Res. 2000; 60: 4098-4104PubMed Google Scholar) of εA residues in DNA were replicated erroneously, giving rise mainly to AT → GC (10Moriya M. Pandya F. Johnson F. Grollman A.P. Exocyclic DNA Adducts in Mutagenesis and Carcinogenesis.in: Singer B. Bartsch H. International Agency for Research on Cancer Scientific Publication No. 150. IARC, Lyon, France1999: 263-270Google Scholar) but also to AT → TA (preferential mutation on the leading strand) and AT → CG substitutions (11Levine R.L. Yang I.-Y. Hossain M. Pandya G.A. Grollman A.P. Moriya M. Cancer Res. 2000; 60: 4098-4104PubMed Google Scholar). In the same studies only 0.3% of 7,8-dihydro-8-oxoguanine (8-oxoG) residues induced GC → TA transversions (11Levine R.L. Yang I.-Y. Hossain M. Pandya G.A. Grollman A.P. Moriya M. Cancer Res. 2000; 60: 4098-4104PubMed Google Scholar). Treatment of mammalian cells with compounds inducing etheno-DNA adducts additionally triggers chromosomal aberrations and sister chromatid exchanges (1Bartsch H. Barbin A. Marion M.-J. Nair J. Guichard Y. Drug Metab. Rev. 1994; 26: 349-371Crossref PubMed Scopus (151) Google Scholar). Early investigations by Tsou et al. (12Tsou K.C. Yip K.F. Miller E.E. Lo K.W. Nucleic Acids Res. 1974; 1: 531-547Crossref PubMed Scopus (38) Google Scholar) and Basu et al. (13Basu A.K. Niedernhofer L.J. Essigmann J.M. Biochemistry. 1987; 26: 5626-5635Crossref PubMed Scopus (33) Google Scholar, 14Basu A.K. Wood M.L. Niedernhofer L.J. Ramos L.A. Essigmann J.M. Biochemistry. 1993; 32: 12793-12801Crossref PubMed Scopus (204) Google Scholar) have shown that in alkali, but also under physiological conditions, εA is rearranged into a pyrimidine ring-opened derivative, 4-amino-5-(imidazol-2-yl)imidazole, which has about 20-fold higher mutagenic potency in Escherichia coli(AB1157 lacΔU169) than the parental εA (14Basu A.K. Wood M.L. Niedernhofer L.J. Ramos L.A. Essigmann J.M. Biochemistry. 1993; 32: 12793-12801Crossref PubMed Scopus (204) Google Scholar). Because the secondary lesions arising from εA might contribute significantly to its mutagenesis, we undertook a detailed study of the chemical stability of εA in DNA, during which we found enzymes repairing a derivative formed during εA chemical rearrangement and identified excised lesion. 1,N 6-εdA was synthesized using the modified procedure of Barrio et al. (15Barrio J.R. Secrist III J.A. Leonard R.J. Biochem. Biophys. Res. Commun. 1972; 46: 597-604Crossref PubMed Scopus (296) Google Scholar) described for the synthesis of its ribo-cogener. The material obtained (needles, melting point, 163–164 °C) was 99% pure (HPLC). Its identity was confirmed by UV, NMR, and electrospray MS. Oligodeoxynucleotide (40-mer) containing a single εA at position 20 in the sequence 5′-d(GCT ACC TAC CTA GCG ACC TεAC GAC TGT CCC ACT GCT CGA A)-3′ was purchased from Eurogentec Herstal, (Herstal, Belgium). The purity and identity of this oligomer was verified by HPLC and mass spectrometry. The oligomer was digested enzymatically to deoxynucleosides, which were separated by HPLC (for the details see below and Fig. 5). The deoxynucleoside content calculated on the basis of peak areas was for dC, 16.1, dG, 7.5, dT, 8.0, dA, 7.5, and εdA, 0.9 residues/molecule, which is in good agreement with the previewed values of dC, 16, dG, 7, dT, 8, dA, 8, and εdA, 1 residue(s)/molecule. The identity of the εA-oligomer was confirmed by mass spectrometry using an electrospray Quadrupol-time of flight instrument (Q-TOF, Micromass). The measured molecular mass of εA-oligomer was 12,145.3 ± 0.4, which is consistent with the expected mass of 12,144.9. No significant traces of the compound, the molecular weight of which would correspond to the presence in the oligomer of compound B instead of εdA (M r 12,163), were detected (less than 2%). No peak corresponding to the presence of compound C in this oligomer (M r 12,134) was recorded. Both methods confirmed the identity and purity of εA-oligomer. Four complementary oligodeoxynucleotides containing T, C, G, or A opposite εA were either purchased from Eurogentec or synthesized according to standard procedures using a Beckman Oligo 1000m synthesizer (Oligonucleotide Synthesis Laboratory, Institute of Biochemistry and Biophysics, Polish Academy of Sciences). E. coli Fpg and Nth proteins were purified from overproducing strains (JM105 supE endA sbcB15hsdR4rpsL thiΔ(lac-proAB) carrying the pFPG230 plasmid and BH410 (as JM105 but fpg-1:Kn harboring the pNTH10) as previously described (16Boiteux S. O'Connor T.R. Laval J. EMBO J. 1987; 6: 3177-3183Crossref PubMed Scopus (250) Google Scholar, 17Dizdaroglu M. Laval J. Boiteux S. Biochemistry. 1993; 32: 12105-12111Crossref PubMed Scopus (266) Google Scholar). HumanN-methylpurine-DNA glycosylase (ANPG-40) as well as bacterial strains overproducing Fpg and Nth glycosylases were a kind gift from Dr. Jacques Laval (Institut G. Roussy, Villejuif, France). T4 kinase was from TaKaRa, nuclease P1 from Amersham Pharmacia Biotech, snake venom phosphodiesterase from PL Biochemicals, and E. coli alkaline phosphatase from Sigma. HPLC was performed using a Waters dual pump system with a tunable UV/visible light absorbance detector managed by Millenium 2010SS (version 2.15) controller. All separations were performed on a Waters Nova-Pak® C18 reversed-phase column (60 Å, 4 μm, 4.6 × 250 mm). NMR spectra were measured on a UNITY 500plus (Varian) spectrometer equipped with a gradient generator unit Performa II, Ultrashims, and a high stability temperature unit using 5 mm 1H{13C/15N} pulsed field gradient triple probe. Electrospray ionization mass spectrometry of nucleosides was performed on a Mariner apparatus with time of flight detection. UV spectra were recorded on a Cary 3E spectrophotometer. Silica gel 60 F254 aluminum sheets (Merck, catalog no. 1.0554) were used for analytical purposes. For preparative purposes 20 × 20 cm plates were prepared using silica gel 60 PF254(Merck, catalog no. 1.07747). The following solvent systems were used: methanol/chloroform, 15:85 (I); isopropyl alcohol/25% aqueous NH3/water, 70:10:10 (II) and 70:5:5 (III). The solutions of εdA (1–2 mm) were incubated in pH 12 (0.02 nNaOH), pH 9.2 (0.1 mNa2B4O7), or pH 7.5 (0.1m phosphate buffer) at 37 °C or at room temperature (23 °C) for various periods (15 min to 30 days) and were analyzed by HPLC. The representative separations, retention times, and chromatographic conditions are given in Fig. 2. The neutral depurination of 2′-deoxyadenosine and εdA was studied in 0.1m phosphate buffer, pH 7.5, at 60 °C by HPLC. The HPLC analysis of depurination was performed similarly to the analysis of alkaline degradation of εdA with the exception that gradient was present for 30 min (the relevant retention times under these conditions (in minutes) were: dA, 11.5; A, 9.7; εdA, 13.0; εA, 11.9; and product B, 8.4). A reaction mixture contained ∼0.1 mmol of εdA in 1 ml of 0.05n NaOH. After 3–7 days at 37 °C, TLC in solvent II showed the presence of product B(R f = 0.52) and C(R f = 0.67), some nonreacted εdA (R f = 0.60), D(R f = 0.38), and other products. The separation of products was done using preparative silica gel plates run 2–4 times in the same direction in solvent III. The appropriate bands were eluted by methanol, and the purity of products was verified by HPLC. The final purification was done by preparative TLC in solvent I. The products obtained were more than 95% pure, and they were used for studies by UV, NMR, and MS. UV (λmax, nm): B, 259 (H20), 265 (pH 12), 267 (pH 1); C, 274 (H2O), 265 (pH 12), 282 (pH 1) (in conformity with spectra of ribo-cogener (18Yip K.F. Tsou K.C. Tetrahedron Lett. 1973; 33: 3087-3090Crossref Scopus (48) Google Scholar)); D, 247 (H2O), 247 (pH 12), 242 (pH 1). Electrospray-MS analytical data (m/z assignment and relative abundance in parentheses): εdA, 276.1 (MH+, 100), 160.1 (BH+, 35); B, 294.1 (MH+, 100), 178.1 (BH+, 35); C, 266.1 (MH+, 100), 150.1 (BH+, 15);D, 288.3 (not assigned, 100), 316.3 (not assigned, 40). NMR data are gathered in supplemental Tables 1S and 2S. Samples for the NMR measurements were prepared in D2O or Me2SO-d 6 at concentrations of ∼5 mm. Spectra were measured at 25 °C using proton 1D, TOCSY (19Aue W.P. Bartholdi E. Ernst R.R. J. Chem. Phys. 1976; 64: 2229-2246Crossref Scopus (3089) Google Scholar, 20Bax A. Freeman R. J. Magn. Reson. 1985; 65: 355-360Google Scholar, 21Braunschweiler L. Ernst R.R. J. Magn. Reson. 1983; 53: 521-528Crossref Scopus (3108) Google Scholar), ROESY (22Bothner-By A.A. Stephens R.L. Lee J.-M. Warren C.D. Jeanloz R.W. J. Am. Chem. Soc. 1984; 106: 811-813Crossref Scopus (1967) Google Scholar, 23Bax A. Davis D.G. J. Magn. Reson. 1985; 63: 207-213Google Scholar) and {1H/13C}gHSQC (23Bax A. Davis D.G. J. Magn. Reson. 1985; 63: 207-213Google Scholar, 24Kay L.E. Keifer P. Saarinen T. J. Am. Chem. Soc. 1992; 114: 10663-10665Crossref Scopus (2433) Google Scholar, 25Palmer III A.G. Cavanagh J. Wright P.E. Rance M. J. Magn. Reson. 1991; 93: 151-170Google Scholar, 26Kontaxis G. Stonehouse J. Laue E.D. Keeler J. J. Magn. Reson. Ser. A. 1994; 111: 70-76Crossref Scopus (94) Google Scholar) experiments under standard conditions and with standard parameters. All spectra were analyzed using VNMR 5.1A (Varian) software. Proton spectra in aqueous solution were calibrated against a water signal (27Gottlieb H.E. Kotlyar V. J. Org. Chem. 1997; 62: 7512-7515Crossref PubMed Scopus (2967) Google Scholar). In Me2SO-d 6 solution the residual solvent signal was used as a reference (28Hoffman R.E. Davies D.B. Magn. Reson. Chem. 1988; 26: 523-525Crossref Scopus (37) Google Scholar) in both proton and carbon dimensions in {1H/13C} correlation spectra. The kinetics of compounds B and C formation in oligodeoxynucleotides as well as the identification of substrates for repair glycosylases was performed by HPLC. εA-40-mer was incubated in 0.2 n NaOH for 1–4 h, neutralized by the addition of an equivalent amount of 1 n HCl and 1/10 volume of 1n Tris-HCl, pH 7.0, ethanol-precipitated, and washed. Then oligomers were digested enzymatically to nucleosides. The reaction mixture (50 μl) contained 1.5 nmol of oligomer, 1.5 units of nuclease P1, 0.075 units of snake venom phosphodiesterase, 0.3 units of E. coli alkaline phosphatase in 20 mm Tris-HCl, pH 8.5, and 10 mm MgCl2. After digestion (1 h, 37 °C), proteins were ethanol-precipitated, and the supernatant containing the nucleosides was evaporated to dryness, dissolved in water, and subjected to HPLC. The representative separations, retention times, and chromatographic conditions are given in Fig. 5. The 40-mer oligodeoxynucleotide containing a single εA or its rearrangement products obtained by oligomer incubation in 0.2 n NaOH for 1–16 h was radiolabeled with32P at the 5′-end and annealed to the complementary strand (double molar excess). The release of εA or its rearrangement products by glycosylases was assessed by measuring the cleavage of 40-mer at the site of lesion. The standard reaction mixture (20 μl) contained 5′ 32P-labeled duplex (1 pmol), 100 mm KCl, 1 mm EDTA, and 5 mmβ-mercaptoethanol in 70 mm Hepes-KOH, pH 7.6 for the Fpg protein or pH 7.8 for ANPG-40 or Nth glycosylases. The mixtures were incubated at 37 °C for a 10 min in the presence of excess of repair glycosylases (100–150 ng of each protein/sample) and then subjected to 20% PAGE in the presence of 7 m urea. To cleave the oligomer at the apurinic/apyrimidinic site remaining after excision of εA by ANPG-40 protein, the reaction mixture prior to PAGE was incubated in 0.2 n NaOH at 70 °C for 30 min. Gels were exposed to x-ray film, scanned in an LKB densitometer, and quantified using Microcal Origin. Kinetic constants were established in εA-oligomer incubated in 0.2 n NaOH for 4 h. The concentration range of the oligomer was 0.6–48 nm when the modified base was paired with dT, 4.8–65 nm when paired with dA, 0.3–75 nm paired with dC, and 2–160 nm paired with dG. The amounts of pure Fpg protein used in the reaction were adjusted to obtain less than 50% utilization of substrate and equaled 0.4 ng when modified base was paired with dT and dG, 2 ng when paired with dA, and 0.2 ng with dC. In each experiment two control samples were set: negative without enzyme, to quantify nonspecific breakage of oligodeoxynucleotide; and positive with excess enzyme (150 ng of Fpg), to get 100% cleavage of oligomer. The reaction was performed precisely for 10 min and stopped by adding sequencing kit stop solution, and reaction products were separated by PAGE. Autoradiograms were scanned (LKB scanner), and the peaks on resulting plots, corresponding to cleavage product and nonreacted oligomer, were quantified using multi-peak Lorentzian fitting in Microcal Origin. In calculations the average substrate concentration (i.e. ([S]0 + [S] t )/2) and average velocity (i.e.([S]0 − [S] t )/t) were used (29Lee H.-J. Wilson I.B. Biochim. Biophys. Acta. 1971; 242: 519-522Crossref PubMed Scopus (173) Google Scholar).V max and K m values were calculated by two methods: a program using the Eisenthall-Cornish-Bowden nonparametric algorithm (30Kamiński Z.W. Domino E.F. Comput. Methods Programs Biomed. 1987; 24: 41-45Crossref PubMed Scopus (10) Google Scholar) and direct fitting of the hyperbolic Michaelis-Menten equation to the data points in Microcal Origin. Values obtained by both methods differed usually by no more than 3–6%. 1,N 6-Ethenodeoxyadenosine at pH 12 was degraded sequentially into three products: εdA →B → C → D (Fig.1), which could be separated by HPLC (Fig. 2, A and B) or by TLC. The first stage, εdA → B, was the fastest (t 12 at 23 °C was 2.5 h). The B→ C and C → D reactions at pH 12 were at least 10 times slower than the conversion εdA →B (Fig. 2). The rearrangement of εdA → B is strongly pH-dependent; a half-time of this reaction at 37 °C equals 1.5 h at pH 12, 7 days at pH 9.2, while at pH 7.5 about 1 year, as estimated on the basis of 30-day measurements (not shown). A half-time for B → C and C→ D reactions at pH 7.5, 37 °C is about 12 days, as judged by the HPLC analysis of the isolated compounds B andC. Thus, at physiological pH the first step of rearrangement in nucleosides occurs very slowly, and the subsequent steps are much faster. We were searching for other than high pH factors that could stimulate εdA rearrangements under physiological conditions. The degradation of εdA was tested in the presence of several amino acids (glycine,l-proline, l-lysine, l-serine, all at 80 mm, and 80% saturateddl-tyrosine), 50 mm glutathione (reduced form), and 80 mm mercaptoethanol, as well as 100 mmNaHS, NaN3, KF, and KI, all at 37 °C in 0.1m phosphate, pH 7.5. None of these compounds accelerated εdA → B conversion (not shown). HPLC analysis of neutral εdA and dA depurination at 60 °C shows that the initial rate of εA formation is 0.08%/h, whereas that of A is 0.004%/h. Depurination of εdA is concomitant with formation of productB with a rate of 0.2%/h (not shown); this shows that the glycosyl bond in εdA is 20-fold less stable than that in dA and that the pyrimidine ring opening in εdA is 2.5 times faster than the rupture of the glycosyl bond under neutral conditions. Composition of εA-oligomer incubated for 1–16 h in NaOH at 37 °C was analyzed by HPLC. Only εdA, productsB and C, could be identified, because compoundD was masked by components of the buffer used for enzymatic digestion of oligomer (Figs. 2 B and 5 D). To reach a rate of εdA → Bconversion similar to that in nucleoside, oligomer was treated with NaOH at a concentration 10-fold higher than that used for nucleoside. Similarly, the fluorescence loss, because of decomposition of εA (the only fluorescent component of the pathway) was observed at a 10-fold higher concentration of NaOH in polymer than in monomer (not shown). CompoundsB and C were already found in εA-oligomer not treated with NaOH (Fig. 2 D), although their amount was negligible and differed from batch to batch; usually they constituted 2–6% of the expected εdA amount (not shown). After 4 h of oligomer incubation in 0.2 n NaOH, εAB and C were found in comparable amounts (Fig.2 D), whereas a 4-h incubation of monomer in 0.02n NaOH resulted in the conversion of 80% of εdA intoB (Fig. 2 C). This suggests that in polymer the reaction is shifted toward the formation of compound Cunder conditions the same as those in the monomer rate of εA →B conversion. Proton chemical shifts of εdA compounds Band C were assigned using TOCSY and ROESY; the spectra are listed in supplemental Table 1S and carbon chemical shifts in supplemental Table 2S. The NMR signals in the spectra of εdA have been assigned following the assignment for the ribo-cogener (31Kronberg L. Sjöholm R. Karlsson S. Chem. Res. Toxicol. 1992; 5: 852-855Crossref PubMed Scopus (48) Google Scholar). The spectra of compoundB at room temperature contain two sets of signals (Fig.3) that are temperature-dependent, with the coalescence in Me2SO-d 6 at 80 °C. Each of the signal sets probably belongs to one of the isomers of compoundB (Fig. 1). We assigned the isomer B1 to be that in which the adenine ring has the hydroxyl group at position 2 and the hydrogen atom at position 3. The isomer B2 has the opened pyrimidine ring with the carbonyl group at position 2 and the proton at position 1. The ratio of B1 and B2 isomers in Me2SO d6 solution at 25 °C is 1:1, whereas in D2O the ratio is 13:87. The spectra of product C differ from those of compoundB in the values of chemical shifts of the H1′ and H2′ protons and lack the signal at ∼8.25 ppm corresponding to the proton H2. A comparison of the signals of all other non-exchangeable protons of the sugar moiety indicates that proton and carbon chemical shifts are almost identical for both the B and Cproducts as well as the parental compound εdA. This means that the sugar conformation remains very similar in all these compounds. The only differences observed are those indicating the shielding effects originated from the changes occurring within the base moiety. However, the NMR spectra of compound D are very different. The signals of the sugar moiety are absent, and only two very broad signals in the proton spectrum at ∼3.5 and 6.9 ppm were observed. We have not assigned these signals and do not propose any structure for the compound. The structures of the B and C compounds assigned by NMR were confirmed additionally by mass spectroscopy. The most abundant peaks in the B spectrum, the protonated deoxynucleoside (m/z = 294.1) and protonated base (m/z = 178.1), differ by 18 mass units from the corresponding peaks in the εdA spectrum (276.1 and 160.1, respectively). Then, the peaks in the C spectrum (266.1 and 150.1) differ by 28 units from the corresponding peaks in the B spectrum. These data are consistent with the following reaction scheme: εdA + H2O (18Yip K.F. Tsou K.C. Tetrahedron Lett. 1973; 33: 3087-3090Crossref Scopus (48) Google Scholar) → B − CO (28Hoffman R.E. Davies D.B. Magn. Reson. Chem. 1988; 26: 523-525Crossref Scopus (37) Google Scholar) → C. The presence in the mass spectrum ofD peaks of m/z 316.3 and 288.3, together with the NMR data showing lack of deoxyribose in this compound, would indicate that D is a dimeric form of base moiety of compoundC. The conclusion can be drawn that depurination is the final step of εdA rearrangements. Human glycosylase-ANPG-40 protein effectively excised εA from the εA:T pair in the 40-mer duplex. However, its capability to cleave εA-oligomer pretreated with NaOH decreased proportionally to the time of incubation in 0.2 n NaOH (Fig.4 A). The εA-oligomer incubated in NaOH was cleaved by E. coliformamidopyrimidine-DNA glycosylase (Fpg protein), leaving behind the β-δ-elimination product, and by endonuclease III (Nth protein), which worked by the β-elimination mechanism (Fig. 4 B). At the same enzyme concentration 40-mer was cleaved to a greater extent by the Fpg than the Nth protein. When, however, both enzymes were used, only the Fpg cleavage product was found (Fig. 4 B), which suggests that both enzymes recognized the same lesion, but/or Nth had a lower affinity to the lesion. Prolonged incubation of εA-oligomer in NaOH resulted also in a partial breakage of DNA at the εA site (Fig.4, A and B). This could be because of alkali-triggered hydrolysis of abasic sites formed after non-enzymatic depurination of either εA or D (Fig. 1) or to other, unknown mechanism. HPLC analysis shows that the εA-oligomer incubated for 4 h in 0.2 n NaOH contains εdA and both products of its conversion, compounds B and C (Fig.5 A). The amount of productB decreased substantially when oligonucleotide was digested with both the Fpg protein (Fig. 5 B) and the Nth glycosylase (Fig. 5 C), suggesting that both enzymes recognize and excise the modified base present in compound B. However, we were unable to discriminate between both isomers of compoundB. For the Fpg protein, the K m andk cat values for excision of the εA derivative when paired with dT and dC were very similar to the kinetic constants of known Fpg substrates 8-oxoG and Fapy-7MeG. Interestingly, theK m for excision of the εdA derivative paired with dA was an order of magnitude higher (∼60 nm) and when paired with dG two orders of magnitude higher than when it was paired with dT and dC (∼6 nm, TableI). For the Nth protein, theK m for excision of B from theB:T pair was 44 nm (k cat=13 min−1).Table IKinetic constants for the removal of modified bases from oligonucleotides by the Fpg proteinSubstrateK mk catk cat/K mnmmin−1ɛA derivative paired with:dA59.9 ± 20.50.49 ± 0.10.008dC6.2 ± 1.40.42 ± 0.020.068dG832 ± 3141-aEstimated values only.47.9 ± 13.11-aEstimated values only.0.058dT5.6 ± 2.10.46 ± 0.070.082Fapy-7MeG101-bValues are given according to Ref.37.0.501-bValues are given according to Ref.37.0.058-OxoG41-bValues are given according to Ref.37.0.431-bValues are given according to Ref.37.0.111341-cValues are given according to Ref.39.1.31-cValues are given according to Ref.39.0.01141-dValues are given according to Ref.42.0.131-dValues are given according to Ref.42.0.0092,2,4-Triaminooxazolone3161-cValues are given according to Ref.39.0.731-cValues are given according to Ref.39.0.002Apurinic/apyrimidinic41-bValues are given according to Ref.37.2.51-bValues are given according to Ref.37.0.61-a Estimated values only.1-b Values are given according to Ref.37Jurado J. Saparbaev M. Matray T.J. Greenberg M.M. Laval J. Biochemistry. 1998; 37: 7757-7763Crossref PubMed Scopus (63) Google Scholar.1-c Values are given according to Ref.39Duarte V. Gasparutto D. Jaquinod M. Cadet J. Nucleic Acids Res. 2000; 28: 1555-1563Crossref PubMed Scopus (102) Google Scholar.1-d Values are given according to Ref.42Tchou J. Bodepudi V. Shibutani S. Antoshechkin I. Miller J. Grollman A.P. Johnson F. J. Biol. Chem. 1994; 269: 15318-15324Abstract Full Text PDF PubMed Google Scholar. Open table in a new tab The εA-oligomer stored for several months in aqueous solutio" @default.
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• W2093264695 title "The Pyrimidine Ring-opened Derivative of 1,N 6-Ethenoadenine Is Excised from DNA by theEscherichia coli Fpg and Nth Proteins" @default.
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