Formation of a Fluorescent Adduct in the Reaction of 2

Dean J. Tuma, Mark L. Kearley, Geoffrey M. Thiele, Simon Worrall, Alvin Haver, Lynell W. Klassen, and Michael F. Sorrell. Chemical Research in Toxicol...
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Chem. Res. Toxicol. 1998, 11, 989-994

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Articles Formation of a Fluorescent Adduct in the Reaction of 2′-Deoxyadenosine with a Malonaldehyde-Acetaldehyde Condensation Product Frank Le Curieux, Donata Pluskota,† Tony Munter, Rainer Sjo¨holm, and Leif Kronberg* Department of Organic Chemistry, Åbo Akademi University, Biskopsgatan 8, FIN-20500 Turku/Åbo, Finland Received April 23, 1998

Malonaldehyde (malondialdehyde, MDA) was reacted with 2′-deoxyadenosine in buffered aqueous solution. HPLC analyses of the reaction mixtures showed that, besides the two previously characterized N6-propenal (M1dA) and N6-oxazocinyl (M3dA) adenine adducts, a third compound eluting at longer retention time was formed. The compound generated a strong peak in the chromatogram recorded by a fluorescence detector. The new compound was isolated by preparative C18 chromatography, and its structure was characterized by UV absorbance, fluorescence emission, 1H and 13C NMR spectroscopy, and mass spectrometry. The product was identified as 9-(2′-deoxyribosyl)-6-(3,5-diformyl-4-methyl-1,4-dihydro-1-pyridyl)purine (M2AA-dA). The yield of the product was 0.8% following 7 days of reaction at 37 °C and pH 4.6. Lower yields were obtained at higher pH conditions. By the addition of acetaldehyde, the yield increased about 10-fold at all studied pH conditions. The adduct was most likely formed by an initial condensation of two molecules of malonaldehyde with one molecule of acetaldehyde followed by reaction of the condensation product with the exocyclic amino group of 2′-deoxyadenosine. The identification of this adduct shows that acetaldehyde may react with DNA bases also through an initially formed malonaldehyde-acetaldehyde condensation product.

Introduction Malonaldehyde (malondialdehyde, MDA)1 is a byproduct of prostaglandin biosynthesis and the major carbonyl end product of lipid peroxidation (1, 2). Consequently, malonaldehyde is an ubiquitous natural product observed in substantial amounts in mammalian and human tissues. Malonaldehyde is mutagenic in bacteria and mammalian cells and carcinogenic in rats (3-6). At present, the possibility of malonaldehyde being an endogenous human carcinogen cannot be ruled out. Malonaldehyde is known to form adducts in reaction with deoxyguanosine, deoxyadenosine, and deoxycytidine (7-10). Recently, one of the deoxyguanosine adducts (M1dG) was detected in the DNA of human liver cells (11). Acetaldehyde is a compound formed endogenously through enzymatic oxidation of ethanol in the liver (12). * To whom correspondence should be addressed. Tel.: +358-2-21 54 183. Fax: +358-2-21 54 866. E-mail: leif.kronberg_abo.fi. † Present adress: Faculty of Chemistry, Adam Mickiewicz University, Grunwaldzka 6, 60-780 Poznan, Poland. 1 Abbreviations: MDA, propanedial, malonaldehyde, malondialdehyde; M1dA, 3-(2′-deoxyribo-N6-adenosinyl)propenal; M3dA, 9-(2′-deoxyribosyl)-6-[5*,7*-diformyl-(2*H)-3*,6*-dihydro-2*,6*-methano-1*,3*oxazocin-3*-yl]purine; M1dG, 3-(2′-deoxyribosyl)pyrimido[1,2-a]purine10(3H)-one; M2AA-dA, 9-(2′-deoxyribosyl)-6-(3,5-diformyl-4-methyl-1,4dihydro-1-pyridyl)purine; COLOC, CH shift correlation NMR spectroscopy via long-range coupling; PEG, poly(ethylene glycol).

Exogenous sources of acetaldehyde are, for example, tobacco smoke and automobile emissions (13, 14). Acetaldehyde has been shown to be mutagenic in several biological assays and is considered to be carcinogenic in experimental animals (15). In studies of reaction of acetaldehyde with nucleosides, it has been found that acetaldehyde forms unstable Schiff’s bases (16, 17) mainly with deoxyguanosine. The identity of these adducts has been elucidated following reduction of the imine group. In 1972, Reiss et al. (18) showed that some fluorescent compounds are formed both in the reaction of malonaldehyde with DNA and in the reaction of malonaldehyde with guanine and adenine bases. The formation of these fluorescent products (the structures of which were not determined) also correlated with the increase of UV absorption at 325 nm (18). Among the malonaldehyde adducts identified hitherto, only the M1dG may provide an explanation for the findings of Reiss et al. However, none of the identified adenine nucleoside adducts display fluorescence and thus cannot account for the fluorescence observed by Reiss in the reaction of malonaldehyde with adenine. Very recently, Tuma et al. (19) reported on the formation of highly fluorescent compounds in the liver of ethanol-fed rats. Later, Xu et al. (20) found that the

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990 Chem. Res. Toxicol., Vol. 11, No. 9, 1998

fluorescence properties originated mainly from a protein adduct that was formed from a condensation product of malonaldehyde and acetaldehyde. Previously, Go´mezSa´nchez et al. (21) reported that malonaldehyde prepared by hydrolysis of 1,1,3,3-tetramethoxypropane may be cleaved to acetaldehyde. Further, it was shown that malonaldehyde and acetaldehyde undergo condensation and, in the presence of a suitable amino group, form a similar N-substituted dihydropyridine derivative as the major fluorescent protein adduct of Xu et al. (20). In the current work, we show that the fluorescent dihydropyridine unit is obtained also when 2′-deoxyadenosine is reacted with malonaldehyde prepared from 1,1,3,3-tetraethoxypropane. In this reaction, the exocyclic amino group of the adenine unit acts as the appropriate amino group. Besides providing an explanation for the observation of Reiss et al. (18), our finding shows that in studies of the acetaldehyde in vivo reactions with DNA one has to take into account the formation of acetaldehyde and malonaldehyde condensation products. These products may form stable and possibly more harmful adducts than those formed by the direct reaction of acetaldehyde with DNA.

Materials and Methods Chemicals. Malonaldehyde was prepared by acid hydrolysis of 1,1,3,3-tetraethoxypropane, as described by Stone et al. (9). 2′-Deoxyadenosine and 1,1,3,3-tetraethoxypropane (97% purity) were purchased from Sigma Chemical Co. (St Louis, MO). Acetaldehyde (purity >99%) was obtained from J. T. Baker B. V. (Deventer, Holland). The water used was Aqua Sterilisata from Pharmacia AB (Stockholm, Sweden). Acetonitrile was gradient grade for chromatography from Merck (Darmstadt, Germany). 1,1,1-Trichloroethane was obtained from Fluka Chemie AG (Buchs, Switzerland). Chromatographic Methods. HPLC analyses were performed on a Kontron Instruments liquid chromatographic system consisting of a model 322 Pump, a 440 diode-array detector (UV), a JASCO FP-920 fluorescence detector, and a KromaSystem 2000 data handling program (Kontron Instruments S.P.A., Milan, Italy). The reaction mixtures were chromatographed on a 5-µm, 4- × 125-mm reversed-phase C18 analytical column (Spherisorb ODS2, Phase separation, U.K.). The column was eluted isocratically for 5 min with 0.01 M phosphate buffer, pH 7.1, and then with a gradient from 0% to 30% acetonitrile in 25 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, Bondesil, Analytichem International, Harbor City, CA). The products were further purified on a semipreparative 8-µm, 10- × 250-mm (Hyperprep ODS, Hypersil, Krotek, Tampere/Tammerfors, Finland) reversedphase C18 column. The column was coupled to a Shimadzu HPLC system, which consisted of two Shimadzu LC-9A pumps and a variable-wavelength Shimadzu SPD-6A UV spectrophotometric detector (Shimadzu Europe, Germany). Spectroscopic and Spectrometric Methods. The 1H and 13C NMR spectra were recorded at 30 °C on a JEOL JNM-A500 Fourier transform NMR spectrometer at 500 and 125 MHz, respectively (JEOL, Japan). The samples were dissolved in Me2SO-d6, and TMS was used as an internal standard. The 1H NMR signal assignments were based on chemical shifts and H-H and C-H correlation data. The determination of the shifts and the coupling constants of the multiplets of the proton signals in the deoxyribose unit were based on a first-order approach and are given with an accuracy of (0.3 Hz. Assignment of carbon signals was based on chemical shifts, C-H correlations, and carbon-proton couplings. The electrospray ionization mass spectra were recorded on a Fisons ZabSpec-oaTOF instrument (Manchester, U.K.). Ion-

Le Curieux et al. ization was carried out using nitrogen as both nebulizing and bath gas. A potential of 8.0 kV was applied to the ESI needle. The temperature of the pepperpot counter electrode was 90 °C. The isolated compound was introduced by loop injection at a flow rate of 20 µL/min (H2O/CH3CN/acetic acid 80:20:1). PEG 200 was used as standard for the exact mass determination. The mass spectrometer was working at a resolution of 7000. The UV spectrum of the isolated compound was recorded with the diode-array detector as the peak eluted from the HPLC column. A Shimadzu UV-160 spectrophotometer (Shimadzu Europa, Germany) was used to determine the molar extinction coefficient (). The fluorescence spectrum was recorded with a Hitachi F-2000 fluorescence spectrophotometer (Hitachi Ltd., Tokyo, Japan). Preparation of 9-(2′-Deoxyribosyl)-6-(3,5-diformyl-4methyl-1,4-dihydro-1-pyridyl)purine (M2AA-dA). Malonaldehyde was prepared by the hydrolysis of 4 g of 1,1,3,3tetraethoxypropane (18 mmol) with 50 mL of 0.1 M HCl. Following the hydrolysis, the pH of the mixture was adjusted to 4.6 by the addition of 1 M NaOH. One gram of 2′deoxyadenosine (4 mmol) was dissolved in about 300 mL of a 0.5 M phosphate buffer solution (pH 4.6), and this solution was added to the 1,1,3,3-tetraethoxypropane hydrolysate. The reaction was performed at 37 °C, and the progress of the reaction was followed by HPLC analyses on the C18 analytical column. After 7 days, the reaction was stopped, and the reaction mixture was filtered and finally passed through the preparative C18 column. The column was eluted with 200 mL of water and then with 100 mL of 5%, 10%, 15%, 20%, 25%, 30%, and 35% acetonitrile solutions in water. Fractions of 30 mL were collected. The compound M2AA-dA eluted from the column with the 25 and 30% acetonitrile washes. The fractions containing the product were combined and concentrated by rotary evaporation to about 20 mL. Further purification of M2AA-dA was carried out by using the semipreparative HPLC column. The column was eluted with a gradient from 17% to 30% acetonitrile in 0.01 M phosphate buffer, pH 7.1, in 15 min at a flow rate of 4 mL/min. The fractions containing the pure compound were combined, concentrated to about 30 mL, and then desalted by use of the preparative C18 column. The desalted solution was rotary evaporated to dryness. The residue, a slightly yellow powder, was subjected to spectroscopic and spectrometric studies. The isolated amount of the compound was 4.0 mg. M2AA-dA had the following spectral characteristics: UV spectrum (HPLC eluent, 25% acetonitrile in 0.01 M phosphate buffer pH 7.1) UVmax 320 nm ( 46 500 M-1 cm-1), 224 nm, UVmin 272, 208 nm, shoulder between 348 and 378 nm; fluorescence spectrum (H2O) λex,max 319 nm, λem,max 444 nm. In the positive-ion electrospray mass spectrum, the following ions were observed (m/z (relative abundance, formation): 386 (100, MH+), 270 (54, MH+ - deoxyribosyl + H). High-resolution mass spectrometry gave the protonated molecular formula as C18H20N5O5 (MH+ 386.1460, calcd 386.1464). The 1H and 13C NMR spectroscopic data of M2AA-dA are presented in Table 1. Small-Scale Reactions of Malonaldehyde with 2′-Deoxyadenosine. Malonaldehyde hydrolyzed from 1,1,3,3-tetraethoxypropane (40 mg, 0.18 mmol) was reacted with 2′deoxyadenosine (10 mg, 0.04 mmol) in 3 mL of 0.5 M phosphate buffer solutions at pH 4.6, 5.5, 6.0, and 7.4. The reactions were performed at 37 °C. The progress of the reactions was followed daily by HPLC analyses of aliquots of the reaction mixtures using the C18 analytical column. Small-Scale Reactions of Malonaldehyde and Acetaldehyde with 2′-Deoxyadenosine. Malonaldehyde hydrolyzed from 1,1,3,3-tetraethoxypropane (40 mg, 0.18 mmol) and acetaldehyde (8 mg, 0.18 mmol) was reacted with 2′-deoxyadenosine (10 mg, 0.04 mmol) in 3 mL of 0.5 M phosphate buffer solutions at pH 4.6, 5.5, 6.0, and 7.4. Acetaldehyde was added to malonaldehyde (1,1,3,3-tetraethoxypropane hydrolysate), and

A Fluorescent Malonaldehyde-Acetaldehyde dA Adduct Table 1.

1H

and13C Chemical Shifts (δ)a and Spin-Spin Coupling Constants, JH,H and JC,H (Hz) of Protons and Carbons in M2AA-dA δ

proton

multiplicity

H-2/H-6

(2H)

9.06

s

CHO H-4 CH3

(2H) (1H) (3H)

9.59 3.70 1.08

s q d

H-2p

H-8p

(1H)

(1H)

8.78

8.89

s

s

H-1′ H-2′ H-2′′ H-3′ H-4′ H-5′ H-5′′

(1H) (1H) (1H) (1H) (1H) (1H) (1H)

6.51 2.75 2.40 4.46 3.92 3.64 3.56

t dt ddd dt dt dd dd

OH OH

(1H) (1H)

5.37 5.02

br br

a

Chem. Res. Toxicol., Vol. 11, No. 9, 1998 991

JH,H

carbon

δ

1,4-Dihydropyridine Unit C-2/C-6 141.3 C-3/C-5 125.9 CHO 190.7 6.7 C-4 22.7 6.7 CH3 21.1 Purine Unit C-2p C-4p C-5p C-6p C-8p 2′-Deoxyribosyl Unit 6.5 C-1′ 13.5, 6.5 C-2′ 13.5, 6.5, 4.0 6.5, 4.0 C-3′ 4.5, 4.0 C-4′ 11.5, 4.5 C-5′ 11.5, 4.5

151.2 152.5 121.6 146.3 143.7

multiplicity

1J C,H

dt dm ddd dm qd

183.6 176.1 135.5 128.8

d ddd dd dt dd

207.4

84.0 39.5

d t

171.2 133.4

70.3 88.0 61.3

d d t

148.4 146.9 141.5

217.0

>1J C,H

4.4 24.8 7.0, 2.0 6.1

11.4, 5.2, 2.6 11.9, 1.0 11.4, 2.6 4.4

Relative to TMS.

this mixture was then added to the solution of 2′-deoxyadenosine. The reactions were performed at 37 °C, and the progress of the reaction was followed daily by HPLC analyses of aliquots of the reaction mixtures. Determination of Product Yields. Quantitative 1H NMR analysis, using 1,1,1-trichloroethane as an internal standard, was performed on an aliquot of the pure M2AA-dA adduct. Then a HPLC standard solution was prepared by taking an exact volume of the NMR sample and diluting it with an appropriate volume of water. The quantitative determination of the amount of M2AA-dA in the reaction mixtures was made by comparing the chromatographic peak area (recorded at 320 nm) of the compound in the HPLC standard solution with the peak area of the compound in the reaction mixtures. The molar yield was calculated from the original amount of 2′-deoxyadenosine in the reaction mixtures.

Results and Discussion HPLC analyses of the small-scale reactions of malonaldehyde with 2′-deoxyadenosine revealed the formation of one product peak with longer retention time than 2′-deoxyadenosine and the previously characterized malonaldehyde adducts 3-(2′-deoxyribofuranosyl-N6-adenosinyl)propenal (M1dA) and the oxazocinyl adduct (M3dA) (9). The compound, marked M2AA-dA, eluted at 27 min from the analytical reversed-phase C18 column (Figure 1A). The fluorescence detector (λex 319 nm, λem 444 nm) gave a very intense signal for the compound (Figure 1B). The highest yield (0.8 mol %) of the compound was obtained in reactions carried out for 7 days at pH 4.6 and 37 °C. At higher pH conditions, the yields were markedly lower, and at pH 7.4 the yield was only about 0.03 mol % (Figure 2A). For the purpose of determining the structure of the compound, a large-scale reaction was performed at pH 4.6. After 7 days of reaction, the compound was isolated from the reaction mixture by preparative C18 column chromatography and finally purified by HPLC. On the basis of structural data from UV, fluorescence and NMR spectroscopy, and mass spectrometry, the structure of the adduct was assigned as 9-(2′-deoxyribosyl)-6-(3,5-

diformyl-4-methyl-1,4-dihydro-1-pyridyl)purine (M2AAdA in Scheme 1). The UV spectrum of M2AA-dA exhibited absorption maxima at 320 and 224 nm, absorption minima at 272 and 208 nm, and a weak shoulder between 348 and 378 nm (Figure 3A). Absorption maxima around 320 nm have been previously reported for adenine nucleosides where a propenal unit is attached to N6 (9, 22). The fluorescence spectrum of M2AA-dA showed an emission maximum at 444 nm when excited at 319 nm (Figure 3B). In the positive-ion electrospray mass spectrum of M2AA-dA, the protonated molecular ion peak was observed at m/z 386 and was the most abundant ion. The fragment recorded at m/z 270 (abundance 54%) corresponds to the cleavage of the ribosyl moiety from the protonated molecular ion (followed by the attachment of a proton to N-9). The 1H NMR spectrum of M2AA-dA displayed, besides the signals from the protons of the deoxyribose moiety, one-proton singlets at δ 8.89 and 8.78 ppm and twoproton singlets at δ 9.59 and 9.06 ppm (Table 1). Moreover, the spectrum displayed a three-proton doublet at δ 1.08 ppm and an one-proton quartet at δ ) 3.70 ppm. H-H connectivity was observed between the signal at δ 8.89 ppm and the signal of H-1′ of the deoxyribosyl unit and between the signals at δ 8.89 and 8.78 ppm. The signals were assigned to the purine protons H-8p and H-2p, respectively (Scheme 1). The two proton signal at δ 9.59 ppm was assigned to the two chemically equivalent formyl protons based on a one-bond H-C correlation with the carbon signal at δ 190.7 ppm. The signal of the formyl protons displayed an H-H long-range correlation with the two-proton signal at δ 9.06 ppm, which was assigned to the β-protons in two R,β-unsaturated aldehyde functionalities with the β-carbons bound to N6. Previously, we have observed similar chemical shifts for the formyl and β-protons in adenosines substituted with a propenal unit at N6 (22, 23). The lack of a signal from an N-bound proton supported the binding of two β-carbons to N6. On the

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Le Curieux et al.

Figure 2. Formation of M2AA-dA at 37 °C and various pH conditions in the reaction of 2′-deoxyadenosine with malonaldehyde (A) and with malonaldehyde and acetaldehyde (B).

Scheme 1

Figure 1. C18 analytical column (column length 125 mm) HPLC chromatogram of the reaction mixture of malonaldehyde and 2′-deoxyadenosine held at 37 °C and pH 4.6 for 7 days: (A) chromatogram recorded by the UV diode-array detector and (B) by the fluorescence detector, λexc 319 nm. For analysis conditions, see Materials and Methods.

basis of the 1H NMR proton signal intensities, it was concluded that the two R,β-unsaturated aldehyde functionalities must be identical and be part of a symmetrical cyclic unit where N6 is one of the connective atoms. The lack of a proton at the R-carbon and the connectivity observed between the formyl protons and the quartet at δ 3.70 ppm suggested that the second bridge consisted of a methine group. The methine group was further bound to a methyl group as shown by the H-H connectivity with the three-proton signal at δ 1.08 ppm, which was split into a doublet. Moreover, H-H connectivities were observed between the methyl group and the formyl protons and the β-protons, respectively. The 13C NMR spectrum of M2AA-dA displayed, besides the signals from the purine and the deoxyribose moieties, carbon signals at δ 190.7, 141.3, 125.9, 22.7, and 21.1 ppm, respectively (Table 1). The signal at δ 190.7 was assigned to the formyl carbons (see above), and the signals at δ 141.3 and 125.9 ppm were assigned to C-2/ C-6 and C-3/C-5, respectively. The signal at δ 141.3 ppm showed C-H correlation with the proton signal assigned to the β-protons. The strong coupling (J ) 24.8 Hz) of the signal at δ 125.9 ppm indicated that the carbon was located at the R-position to the formyl group (24-26). In the long-range C-H correlation spectrum (COLOC),

connectivities were observed between the carbon signal at δ 125.9 ppm and the signal of the formyl protons (δ 9.59 ppm) as well as the signal of the methyl proton (δ 1.08 ppm), confirming the bridging character of the methine carbon atom. The signal of the methine carbon (C-4) was found at δ 22.7 ppm, and the signal at 21.1 ppm was assigned to the methyl carbon bound to C-4. The carbon signal at δ 146.3 ppm, assigned to C-6p, displayed a three-bond C-H triplet pattern (J ) 2.6 Hz), probably due to coupling to H-2/H-6 in the dihydropyridyl unit. The NMR, UV, and fluorescence spectroscopic and the mass spectrometric data were consistent with the structure of M2AA-dA presented in Scheme 1. 1H NMR data of various 3,5-diformyl-4-methyl-1,4dihydropyridine derivatives have been reported by Go´mez-Sa´nchez et al. (21) and Xu et al. (20). The chemical shifts reported in Table 1 are very close to the ones reported by these authors. The only significant difference

A Fluorescent Malonaldehyde-Acetaldehyde dA Adduct

Figure 3. (A) UV absorbance and (B) fluorescence emission spectra of M2AA-dA (λexc 319 nm). The UV spectrum was recorded with the diode-array detector as the compound eluted from the column.

is in the shift of the signal of the H-2/H-6 protons. These protons were observed at δ 9.06 ppm, whereas Go´mezSa´nchez et al. and Xu et al. reported chemical shifts of δ 6.6-7.0 ppm for the corresponding protons. The reason for the downfield shift in M2AA-dA may be explained by the anisotropy effect of the aromatic purinyl moiety. In the work of Go´mez-Sa´nchez et al. (21) and Xu et al. (20), the 1,4-dihydropyridines were substituted with saturated moieties. Go´mez-Sa´nchez et al. (21) reported on the formation of 2,4-dihydroxymethylene-3-methylglutaraldehyde (Scheme 1, MDA-AA) upon acid hydrolysis of 1,1,3,3tetramethoxypropane. They suggested that MDA-AA was formed by condensation of two molecules of malonaldehyde with one molecule of acetaldehyde that was generated through hydrolytic cleavage of malonaldehyde. The yield of MDA-AA could be drastically increased by reacting acetaldehyde with the sodium salt of malonaldehyde. Further, Go´mez-Sa´nchez et al. (21) showed that treatment of MDA-AA with an appropriate amine in water at room temperature yielded 3,5-diformyl-4-methyl-1,4-dihydropyridines. Nair et al. (27) reported on the formation of the corresponding N-substituted 1,4-dihydropyridines upon treatment of amino acids with malonaldehyde. The dihydropyridines were reported to be highly fluorescent, and the yields of the amino acid derivatives were found to increase by performing the reactions in the presence of acetaldehyde. In the current work, we show that a highly fluorescent N-substituted 1,4-dihydropyridine (M2AA-dA) is formed when 2′-deoxyadenosine is reacted with malonaldehyde derived from

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the hydrolysis of 1,1,3,3-tetraethoxypropane. Consistent with the work of Go´mez-Sa´nchez et al. (21) and Nair et al. (27), we observed that the yield of the product was significantly increased (about 10-fold) when the reaction was performed in the presence of acetaldehyde (Figure 2B). Acetaldehyde has been shown to be mutagenic in several biological systems and to induce carcinomas in experimental animals (15). In studies of the mechanisms behind the genotoxic effects of acetaldehyde, the compound has been reacted with nucleosides, and the products have been structurally characterized (16, 17). The outcome of these studies is that the acetaldehyde carbonyl group reacts mainly with exocyclic amino groups of the nucleosides forming Shiff’s bases (imines). Usually, the imines are unstable, and the hitherto identified adducts have been characterized after conversion of the imines to saturated amines. Recently, Tuma et al. (19) observed that in the presence of malonaldehyde, acetaldehyde formed stable bindings with proteins in a concentration-dependent manner. The bindings were found to be due mainly to the formation of a 3,5-diformyl-4methyl-1,4-dihydropyridine protein adduct (20). The same dihydropyridine protein adduct was reported to be formed also in the liver of ethanol-fed rats, and this finding was explained by the coexistence of malonaldehyde and acetaldehyde in the rat liver during ethanol metabolism. The current work shows that the MDA-AA conjugation product reacts with 2′-deoxyadenosine and forms a stable dihydropyridine adduct. The finding of the corresponding dihydropyridine protein adduct in the liver of ethanolfed rats indicates the presence of the MDA-AA conjugation product in experimental rats. Therefore, it seems possible that besides the formation of dihydropyridine protein adducts, dihydropyridine DNA adducts could also be formed as a consequence of ethanol metabolism or as a consequence of exogenous sources of acetaldehyde. This means that acetaldehyde interaction with DNA through conjugation with malonaldehyde may be an important mechanism for the expression of the genotoxicity of acetaldehyde. Acetaldehyde is known to cause DNA-protein and DNA-DNA intrastrand cross-linking, but the underlying chemical mechanism has not been elucidated (28). The initially formed conjugation product, MDA-AA, and the M2AA-dA adduct, having four and two reactive aldehyde groups, respectively, are good candidates for the explanation of the cross-linking capacity of acetaldehyde. Xu et al. pointed out that the fluorescent MDA-AAprotein adduct is not the only adduct formed when malonaldehyde and acetaldehyde coexist (20). This statement is supported by our findings. HPLC analysis of the reaction mixture containing malonaldehyde, acetaldehyde, and 2′-deoxyadenosine revealed the formation of fluorescent as well as nonfluorescent products of unknown structure. The M2AA-dA adduct and the hitherto unidentified adducts explain in part the observation made by Ohya (29) of the enhanced reactivity of malonaldehyde toward adenine nucleosides in the presence of acetaldehyde. In the work of Go´mez-Sa´nchez et al. (21), it was shown that conjugation of aliphatic aldehydes with malonaldehyde is a general reaction. This means that also other aldehydes present in physiological fluids may form reactive condensation products with malonaldehyde. Studies should be conducted in order to clarify whether such

994 Chem. Res. Toxicol., Vol. 11, No. 9, 1998

condensation products are involved in the cytotoxic and genotoxic effects generated by the aldehydes.

Conclusion The current work shows that acetaldehyde forms a condensation product with malonaldehyde and that the condensation product reacts with the exocyclic amino group of 2′-deoxyadenosine resulting in a strongly fluorescent 1,4-dihydropyridine adduct. The finding stresses the importance of the presence of reactive aldehyde condensation products in physiological fluids. The condensation products may form more stable and possibly more harmful adducts than those formed by a specific aldehyde.

Acknowledgment. We thank Mr. Markku Reunanen for the mass spectra. Financial support for the work was obtained from the European Commision (Contract No. ERBFMBICT 961394, F.L.C.), the Centre for International Movement in Helsinki, Finland (D.P.), and the Åbo Akademi University (T.M.).

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