Investigation of the Adducts Formed by Reaction of Butenedioic Acids

1186

Chem. Res. Toxicol. 1997, 10, 1186-1191

Investigation of the Adducts Formed by Reaction of Butenedioic Acids with Adenosine Robert Franze´n,* Masatoshi Morita, Kiyoshi Tanabe, Hiroo Takagi, and Yasuyuki Shibata National Institute for Environmental Studies, 16-2 Onogawa, Tsukuba, Ibaraki 305, Japan Received March 5, 1997X

Several genotoxic butenedioic acids present in chlorine-disinfected drinking water were allowed to react with adenosine, guanosine, and cytidine in aqueous solution. HPLC analyses, with detection at 254 and 310 nm, showed that clearly detectable products were formed only in the reactions with adenosine. The major products from the reactions between either 2-chloro3-methyl-2-butenedioic acid (ox-MCF) or 2-chloro-3-(chloromethyl)-2-butenedioic acid (ox-CMCF) and adenosine were the same. This substance was isolated by C18 column chromatography and characterized by UV absorbance, 1H and 13C NMR spectroscopy, and mass spectrometry. It was identified as 3-(β-D-ribofuranosyl)-7-carboxy-7-formyl-8-[9′-(β-D-ribofuranosyl)-N6-adenosinyl]-1,N6-ethanoadenosine (cfA,A). The yields of cfA,A in reactions performed at pH 7.4 and 37 °C were 0.7% and 0.3% with ox-MCF and ox-CMCF, respectively.

Introduction Chlorine disinfection of drinking water is known to produce compounds which generate mutagenicity in the Ames assay (1-4). The presence of such compounds in chlorinated drinking water has raised concern over the possible health effects of long-term consumption of chlorine-disinfected drinking water. It is generally accepted that interaction of chemical carcinogens with DNA is of key importance in the events involved in the induction of malignant tumors. Alteration of the genetic material resulting from the covalent reaction of carcinogens with DNA changes the cell genome and in this way induces the malignant transformation of the cell (5). In work aimed at the identification of main mutagens in drinking water, the extremely potent direct-acting mutagen 3-chloro-4-(dichloromethyl)-5-hydroxy-2(5H)furanone (MX)1 was identified (6, 7). Recent studies have shown that, besides MX, several other structurally related compounds are present in chlorinated drinking water (8-10). Among these compounds are several oxidized forms of chlorinated hydroxyfuranones, i.e., butenedioic acids, namely, 2-chloro-3-(dichloromethyl)2-butenedioic acid (ox-MX), 2-chloro-3-(chloromethyl)-2butenedioic acid (ox-CMCF), 2-chloro-3-methyl-2-butenedioic acid (ox-MCF), and 2,3-dichloro-2-butenedioic acid (ox-MCA) (Chart 1). In recent work by Kronberg and Franze´n (9) all these butenedioic acids were found to be present in the water samples that were studied. The compounds were formed in reactions of chlorine with macromolecular organic constituents of water, i.e., lignin or humic substances. In a sample of chlorination-stage bleaching liquors (CBL) and in chlorinated humic water, the compound ox-MCA was clearly the most abundant diacid followed by ox-MCF. The sample of CBL was * Author for correspondence. E-mail: [email protected]. Phone: +358-9-70851. Fax: +358-9-70859556. X Abstract published in Advance ACS Abstracts, September 15, 1997. 1 Abbreviations: MX, 3-chloro-4-(dichloromethyl)-5-hydroxy-2(5H)furanone; ox-MX, 2-chloro-3-(dichloromethyl)-2-butenedioic acid; oxCMCF, 2-chloro-3-(chloromethyl)-2-butenedioic acid; ox-MCF, 2-chloro3-methyl-2-butenedioic acid; ox-MCA, 2,3-dichloro-2-butenedioic acid; cfA,A, 3-(β-D-ribofuranosyl)-7-carboxy-7-formyl-8-[9′-(β-D-ribofuranosyl)-N6-adenosinyl]-1,N6-ethanoadenosine; CBMA, carboxymalonaldehyde.

S0893-228x(97)00036-2 CCC: $14.00

Chart 1

derived from softwood (pine) kraft pulp which had been prebleached with oxygen. Natural humic water, with a TOC content of 20 mg/L, was collected from Lake Savojarvi in Finland and was chlorinated at pH 7 at a Cl2/TOC weight ratio of 1.0. The concentration of oxMCA in the CBL sample was 1100 µg/L, and the other butenedioic acids were found in a concentration range of 1-130 µg/L. The predominant diacid in chlorinedisinfected drinking water (DW) was ox-MCF. This compound was found to represent a major chlorinated byproduct in DW and was detected at a concentration of 210 ng/L. All of these compounds, except ox-MX, are direct-acting mutagens, inducing about 0.1-2 revertant colonies/nmol in the Ames tester strain TA100. It is generally accepted that chemical reactions of genotoxic compounds with the DNA bases likely may cause gene mutations and contribute to cancer initiation (11). The characterization of reaction products of nucleosides and genotoxic agents provides information on the kinds of adducts that could be produced in DNA. Some data are available on the functional significance of the modification of various species of DNA by chemical carcinogens. However, it is not possible at this moment to understand completely if the modifications of DNA are involved in the mechanism of chemical carcinogenesis. In the current work, we have studied the reactions of several butenedioic acids with nucleosides. The nucleosides were used as chemically stable models (5, 11). We report on the isolation and structural elucidation of a previously unknown adduct, formed by reaction of either ox-MCF or ox-CMCF with adenosine (Chart 2). We also present a plausible mechanism for the formation of this modified nucleoside.

Materials and Methods Caution: All the hydroxyfuranones (MX, CMCF, MCF, and MCA) and the butenedioic acids (ox-CMCF, ox-MCF, and ox-

© 1997 American Chemical Society

Adducts via Butenedioic Acids + Adenosine Reaction Chart 2

MCA) have tested positive in the Ames mutagenicity assay with S. typhimurium (TA100) without metabolic activation. Therefore, caution should be exercised in the handling and disposal of the compounds. HPLC analyses were carried out using dual Shimadzu LC10AD pumps equipped with a variable wavelength Jasco UVIDEC-100V UV spectrophotometric detector. The reaction mixtures were chromatographed on a 5 µm, 4- × 125-mm reversed phase C18 analytical column (Spherisorb ODS2, GL Sciences, Japan). The column was eluted isocratically for 5 min with 5% acetonitrile in 0.01 M potassium dihydrogen phosphate (pH 4.6) and then with a gradient from 5% to 40% in 30 min at a flow rate of 1 mL/min. Preparative isolation of the products was performed by column chromatography on a 2.5- × 10-cm column of preparative C18 bonded silica grade (40 µm; GL Sciences, Japan). The 1H and 13C NMR spectra were recorded at 22 °C on a Jeol JNM-A500 Fourier transform NMR spectrometer at 500 and 125 MHz, respectively. The samples were dissolved in Me2SO-d6, and TMS was used as an internal standard. The determination of the shifts and the coupling constants of the multiplets of the proton signals in the ribose units was based on a first-order approach. Assignment of the carbon signals was based on chemical shifts and carbon-proton couplings. In the assignment of protons, proton-decoupled 13C NMR spectra were recorded. Assignments of the carbon signals were made using proton-coupled 13C NMR spectra and C-H shift correlation spectra. The 2-D NMR spectra were recorded using standard pulse sequences of the instrument. The assignments of the proton signals were supported by H-H shift correlation spectroscopy. Fast atom bombardment (FAB) mass spectra were recorded on a Jeol SX 102A mass spectrometer. Xenon was used as the ionization gas and glycerol as the matrix. The electrospray spectrometry system consisted of a JEOL SX 102A mass spectrometer fitted with an electrospray ion source for JEOL mass spectrometers. The sample introduction system consisted of a 250 µL syringe (2.3 mm i.d.), a digital infusion (syringe) pump (Harvard Apparatus, Inc.), and a fused silica capillary (150 µm i.d., 360 µm o.d. × 50 cm length; Polymicron Technologies, Inc.). The compounds (0.5 mg/mL) were introduced into the system dissolved in 10 mM ammonium hydrogen carbonate solution at pH 8.3. The UV spectrum of the isolated product was recorded with a Hitachi U-3200 spectrophotometer (Hitachi Corp., Japan). All the butenedioic acids were synthesized and purified according to the methods of Franze´n and Kronberg (9, 12). 3-(β-D-Ribofuranosyl)-7-carboxy-7-formyl-8-[9′-(β-D-ribofuranosyl)-N6-adenosinyl]-1,N6-ethanoadenosine (cfEA,A). ox-MCF (1 g, 6 mmol) and adenosine (400 mg, 2 mmol) were dissolved in 100 mL of 0.5 M aqueous phosphate buffer (KH2-

Chem. Res. Toxicol., Vol. 10, No. 10, 1997 1187 Table 1. 1H and 13C NMR Chemical Shifts (δ)a and H-H (JH,H) and C-H (1JC,H) Spin-Spin Coupling Constants (Hz) of Protons and Carbons in the Ribose Unit of cfEA,Ab proton

δ

multiplicity

JH,H

carbon

δ

multiplicity

H-1′ H-1′A H-2′ H-2′A H-3′ H-3′A H-4′ H-4′A H-5′ H-5′ H-5′A H-5′A OH-2′ OH-2′A OH-3′ OH-3′A OH-5′ OH-5′A

6.04 6.00 4.57 4.60 4.18 4.18 4.12 4.11 3.75 3.67 3.75 3.67 5.40 5.40 5.40 5.40 5.40 5.40

d d t t m m q q dd dd dd dd br br br br br br

5.2 5.2 5.2 5.2 na na 3.9 3.9 12.1, 4.1 12.1, 4.1 12.1, 4.1 12.1, 4.1

C-1′ C-1′A C-2′ C-2′A C-3′ C-3′A C-4′ C-4′A C-5′ C-5′A

87.9 87.8 74.2 73.9 70.2 70.3 85.6 85.8 61.3 61.4

d d d d d d d d t t

1J

C,H

165.5 165.5 148.9 146.9 149.0 149.0 147.9 146.9 141.4 140.7

a Relative to Me SO-d at δ ) 2.500 ppm. b The superscript A 2 6 designates the ribose unit in the adenosine moiety (see Chart 2).

Table 2. 1H NMR Chemical Shifts (δ)a in cfEA,A proton H-5 H-2 H-2A H-8A

δ

proton

δ

9.03 8.53 8.57 8.71

H-8 CHO COOH NH

7.29 9.32 nt obsb 8.77

a Relative to Me SO-d at δ ) 2.500 ppm. b nt obs ) not 2 6 observed.

PO4/Na2HPO4 in ratio adjusted to pH 6.0), and the resulting solution was stirred for 5 days at 50 °C. The product was isolated from the reaction mixture by column chromatography on a C18 preparative column. The column was washed with 100 mL of 0%, 5%, 10%, and 20% acetonitrile solutions in 0.05 M KH2PO4 (pH 4.6). Fractions of 20 mL were collected. The fractions containing the pure product (10% washes) were combined and concentrated to dryness. Most of the buffer was removed by redissolving the adduct in methanol and filtration. After evaporation of the solvent, the residue was dissolved in a small amount of water, and following a second purification on the preparative column and rotary evaporation to dryness, the residue was subjected to spectrometric studies. The yield of cfA,A was 5% based on the original amount of adenosine in the reaction mixture. The isolated compound (cfA,A) had the following spectral properties: UV spectrum (H2O) UVmax 325 nm ( 31 700 M-1 cm-1), UVmin 270 nm. The following ions were observed in the negative FAB mass spectra [m/z (relative abundance, formation)]: 379 (40, M-ribosyl-purinyl-H+), 363 (10, M-(8-adenosinyl)2H+), 289 (100, M-(8-adenosinyl)-COOH-CHO-H+). In the positive FAB mass spectra only protonated adenosine could be found as the most abundant fragment. In the negative ion electrospray mass spectra the following ions (m/z) were observed: 629 (0.1, M-H+), 379 (100, M-ribosyl-purinyl-H+), 289 (40, M-(8-adenosinyl)-COOH-CHO)-H+). The 1H and 13C NMR spectroscopic data are presented in Tables 1-3, and the results from a 1H-13C COSY experiment are outlined in Figure 4. Formation of cfEA,A in Reaction with ox-MCF or oxCMCF at pH 7.4 and 37 °C. ox-MCF (1 g, 6 mmol) or ox-CMCF (1 g, 5 mmol) and adenosine (400 mg, 2 mmol) were allowed to react in 50 mL of 0.5 M phosphate buffer at pH 7.4 and 37 °C. The formation of cfA,A was followed by HPLC analyses of aliquots of the reaction mixtures. At maximum yield the reactions were stopped and cfA,A was isolated and purified as described previously. The UV and 1H NMR spectra were recorded and compared with the data above. In every aspect they were identical. Small Scale Reactions of Butenedioic Acids with Adenosine, Guanosine, and Cytidine. The butenedioic acids

1188 Chem. Res. Toxicol., Vol. 10, No. 10, 1997

Franze´ n et al.

Table 3. 13C NMR Chemical Shifts and One-Bond (1JC,H) and Long-Range (>1JC,H) C-H Spin-Spin Coupling Constants (Hz) of Carbons in the Base Units of cfEA,A C atoma

δ

multiplicity

1J C,H

C-5 C-3a C-9b C-9a C-2 C-8 C-7 CHO COOH C-2A C-4A C-5A C-6A C-8A

135.4 138.1 117.0 147.3 139.9 130.2 110.0 191.3 171.8 152.0 151.6 120.8 149.7 142.5

dm m d dd dd dm d dd s d br d br dd

192.5

214.2 192.4 176.9

>1J C,H

14.2 16.3, 2.1 3.8 22.1 7.2

204.9 12.3 218.3

3.7

Figure 2. UV spectra of cfA,A.

a The superscript A designates carbons in the adenine moiety; the other carbons belong to the ethenoadenine moiety.

Figure 1. C18 column HPLC separation of the reaction mixture of ox-MCF and adenosine held at 37 °C and pH 7.4 for 4 days. Abbreviations: A, adenosine; cfA,A, isolated reaction product presented in Chart 2. (0.01 mmol) were allowed to react separately with 0.005 mmol of adenosine, guanosine, and cytidine in 2 mL of 0.5 M phosphate buffer at pH 7.4 and 6.0. The formation of products was followed by HPLC analyses. Determination of Product Yields. The purity of cfA,A was determined by 1H NMR analyses. A standard solution was prepared for HPLC analysis by taking an exact amount of the compound and diluting it with an appropriate volume of water. The quantitative determination of the adduct in a reaction mixture was made by comparing the peak area of the adduct in the standard solution with the area of the adduct peak in the reaction mixture. The adduct (cfA,A) was quantified using UV detection at 325 nm. The molar yields of the adduct were calculated from the original amount of adenosine in the reaction mixture.

Results and Discussion The small scale aqueous reactions of the butenedioic acids with adenosine, guanosine, and cytidine were followed by HPLC analyses with UV detection at 254 and 310 nm. In the reactions of ox-MX, and ox-MCA with all of the nucleosides, no products could be observed. On the other hand, in reactions of both ox-MCF and oxCMCF with adenosine at pH 7.4 and 37 °C, a distinct product peak and several other unidentified peaks were observed at longer retention times than that of adenosine (Figure 1). The compound marked cfA,A was eluted at 12.2 min. In order to determine the structure of cfA,A, a large scale reaction was performed in water. The yield was

Figure 3. cA,A (13).

13C

NMR chemical shifts of compounds cfA,A and

shown to be better when ox-MCF interacted with adenosine than when the same reaction was performed with ox-CMCF. ox-CMCF is less stable than ox-MCF. Several degradation products from ox-CMCF are observed when the compound is dissolved in water. In addition an increase of the temperature from 37 to 50 °C results in a higher yield of cfA,A. The structure of the compound was assigned on the basis of data obtained by UV and NMR spectroscopy and mass spectrometry (Chart 2). The UV spectrum of the compound showed an intense absorption maximum at 325 nm (Figure 2). The spectrum was very similar to that obtained by Asplund et al. (13) of cA,A (Figure 3). This indicated that cfA,A was structurally related to cA,A. In the negative ion electrospray mass spectrum of the product, a weak molecular ion-H was observed at m/z 629 (0.1% of base peak). The fragment peak at m/z 379 was formed by cleavage of a ribosyl and a purinyl unit from M-H. At m/z 289 the fragment peak was due to cleavage of the 8-adenosinyl unit together with both COOH and CHO from M-H. Except for the deprotonated molecular ion, the same fragments could be observed in the negative FAB spectra. In the positive FAB mass spectrum only a peak representing protonated adenosine

Adducts via Butenedioic Acids + Adenosine Reaction

Chem. Res. Toxicol., Vol. 10, No. 10, 1997 1189

Figure 4. Partial 1H-13C chemical shift correlation spectrum of cfA,A. The sample was prepared by dissolving 13 mg of cfA,A in 1 mL of DMSO in a 10-mm NMR tube. The spectrum was obtained at 22 °C, and the 1H decoupler coil was used for observation. The partial 1H and 13C NMR spectra, with the proton and carbon assignments, are shown on the top and on the side of the 2-D plot. For abbreviations, see Chart 2.

could be observed where it was the most abundant fragment. No chlorine isotope patterns were observed in any of the mass spectra. Both the 1H and 13C NMR spectra of cfA,A displayed two sets of ribose signals (Table 1). This indicated that the compound contains two adenosine units and that the units were not identical in structure. Besides the ribose resonances, the 1H NMR spectrum of cfA,A displayed seven singlet signals (Table 2). The chemical shifts of the three signals at δ ) 8.71, 8.57, and 8.53 ppm were close to those of H-8A (δ ) 8.64 ppm), H-2A (δ ) 8.53 ppm), and H-2 (δ ) 8.80 ppm) of the previously identified “dimeric structure” of adenosine (cA,A) (13). The signal at δ ) 9.32 ppm was assigned to a formyl proton by the C-H correlation to the carbon resonance at δ ) 191.3 ppm and by the observation that this signal was a double of doublets in the coupled 13C NMR spectrum. A coupling (J ) 7.2 Hz) of the formyl carbon was observed to H-8. Thus it seemed likely that both a proton and an another adenosine unit were attached to C-8. Also the difference in the shift of the formyl carbon in cfA,A and in cA,A (δ ) 178.1 ppm) indicated that the structures were different. The proton at δ ) 7.29 ppm was assigned to H-8, and a signal assigned to an NH group was localized at δ ) 8.77 ppm. The shifts of both these protons were in accordance with previous data (14). The signal at δ ) 9.03 ppm was assigned to H-5 based on C-H correlation to the carbon resonance at δ ) 135.4 ppm. No signal

which could be assigned to the COOH proton was detected. This could be due to a fast exchange of protons assisted by the presence of some water in the solvent. The assignment of the carbon chemical shifts was based on comparison of carbon shifts of the known compound cA,A (Figure 3) on C-H couplings (Table 3), and on C-H correlations (Figure 4). In cA,A C-2A, C-4A, C-5A, C-6A, and C-8A were observed at δ ) 151.8, 151.6, 121.4, 149.7, and 142.0 ppm, respectively. For cfA,A only a downfield shift was observed in the corresponding carbons (δ ) 152.0, 151.6, 120.8, 149.7, 142.5 ppm). In the other adenosine unit, saturation of the C7-C8 double bond was reflected in the chemical shifts. In cA,A the carbon signals of C-2, C-5, C-3a, C-9a, and C-9b were observed at δ ) 142.1, 136.3, 141.9, 142.0, and 121.9 ppm. In the 13C NMR spectrum of cfA,A, signals which could be attributed to the above-mentioned carbons were observed at δ ) 139.9, 135.4, 138.1, 147.3, and 117.0 ppm, respectively. In the proton-coupled 13C NMR spectra (Table 3), the signal of C-2A displayed a one-bond C-H coupling and the signals of C-2, CHO, and C-8A were all observed as doublets of doublets (dd). In addition to the one-bond couplings, C-2 is coupled to H-5, the carbon of the aldehyde group is coupled to H-8, and C-8A is coupled to H-2A. From C-5 and C-8, long-range couplings could be observed in addition to the one-bond couplings. These could not be measured exactly and are reported as double multiplets. Differentiation of the signals was made on

1190 Chem. Res. Toxicol., Vol. 10, No. 10, 1997 Scheme 1

the basis of the observed C-H correlation. The protonated carbon assignments were established by a 2-D 1H13C heteronuclear correlated spectroscopy (COSY) experiment (Figure 4). This proton-carbon 2-D plot connects the proton signals to their associated carbons via one-bond scalar coupling. The assignments of the carbon resonances of the ribose units are based on C-H correlation spectroscopy and a comparison of the carbon shifts in cA,A. The NMR chemical shifts of cfA,A were further studied by NMR simulation2 of the structure. The simulation resulted in the correct assignment of the shifts in the proposed structure of cfA,A. The compounds ox-MCF and ox-CMCF react with adenosine and form an adduct similar to those isolated by Asplund et al. (13). A plausible mechanism of formation of cfA,A could involve a sequential attack of two molecules of adenosine on the butenedioic acids. In the initial step carboxymalonaldehyde (CBMA) is formed from the butenedioic acids (Scheme 1). This trans formation is similar to that proposed by Kronberg et al. (15) for mucochloric acid. All steps of this mechanism including proton transfer from methyl groups (16), formation of hydroxymethyl groups from chloromethyl groups (17), oxidation of a hydroxymethyl group to an aldehyde group (18), and replacement of chlorine β to the aldehyde group (19) were reinvestigated and confirmed as reported previously. Also a decarboxylation of a enolketo acid has been reported (15). In the next step (Scheme 2) the aldehyde of CBMA is attacked by the exocyclic amino group of two adenosine molecules and the N/N-acetal (A) is formed (13). The formation of cfA,A proceeds by Michael addition and rearrangement of A. Several alkylating agents have been shown to form cross-links between the base moieties in DNA. In many cases cytotoxicity is believed to correlate with crosslinking efficiency (20, 21). Malondialdehyde induces cross-links between complementary strands of DNA and produces stable adducts to guanosine, adenosine, and cytosine bases (22-25). Chloromalonaldehyde (a putative breakdown product of mucochloric acid) induces mutagenicity in the TA100 strain and produces a drastic increase of GC f AT transitions (26). Mutagenicity associated with acrolein and other R,β-unsaturated, bifunctional carbonyl compounds could be due either to 2 NMR simulation was performed using the gNMR simulation program (version 3.6), Cherwell Scientific Publishing Ltd., Oxford, U.K. (1995).

Franze´ n et al. Scheme 2

the formation of DNA cross-links or to exocyclic adducts of one or more of the nucleic acid bases (27). The current work shows that butenedioic acids, active in the Ames assay and present in drinking water, react with adenosine and form adducts. It is still unknown if butenedioic acids can produce “dimeric products” from other nucleosides also. This may only be revealed when the still unidentified minor peaks in our HPLC chromatograms of reaction mixtures are identified. In addition a further analysis with Ames tester strain TA104 will reveal whether the formation of adenosine adducts gives an explanation for the mutagenicity of chlorinated butenedioic acids. Also mutational spectra may be informative in regard to the types of mutations which might be induced by butenedioic acids in rodents and humans.

Acknowledgment. The authors are thankful to Dr. Tomoharu Sano and Mrs. Chieko Suzuki for conducting the NMR analyses. The experimental work was supported by the STA Fellowship program (to R.F.).

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